Enzyme Catalysis in Organic Synthesis: A Comprehensive Handbook 2nd

Enzyme Catalysis in Organic Synthesis

Edited by K. Drauz and H. Waldmann

Second Edition

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Applied Homogeneous Catalysis with Organometallic Compounds Second, Completely Revised and Enlarged Edition Three Volumes 2000, ISBN 3-527-30434-7

6. Cornils, W. A. Herrmann, R. Schlogl, C.-H. Wong (Eds.)

Catalysis from A-Z A Concise Encyclopedia 2000, I S B N 3-527-29855-X

I

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Fine Chemicals through Heterogeneous Catalysis 2001, I S B N 3-527-29951-3

Enzyme Catalysis in Organic Synthesis A Comprehensive Handbook

Edited by Karlheinz Drauz and Herbert Waldmann Second, Completely Revised and Enlarged Edition

@WILEY-VCH

Editors: Prot Dr. Karlheinz Drauz Degussa AG 1ZN Wolfgang, Bereich FC-TRM

Rodenbacher Chaussee 4 63457 Hanau-Wolfgang Germany

This book was carefully produced. Nevertheless editors, authors and publisher do not warrant the information contained therein to be free of errors. Readers are advised to keep in mind that statements, data, illustrations, procedural details or other items may inadvertently be inaccurate.

Prof. Dr. Herbert Waldmann

Max-Planck-Institut fiir Molekulare Physiologie Otto-Hahn-Strage 11 44227 Dortmund Germany Library of Congress Card No.: applied for British Library Cataloguingin-PublicationData A catalogue record for this book is available from

the British Librav. Die Deutsche Bibliothek - CIP CataloguinginPublication-Data A catalogue record for this publication is availa-

ble from Die Deutsche Bibliothek

0 Wiley-VCH Verlag GmbH, Weinheim 2002 All rights reserved (including those of translation into other languages). No part of this book may be reproduced in any form - by photoprinting. microfilm, of any other means - nor transmitted or translated into machine language without written permission from the publishers. In this publication, even without specific indication, use of registered names, trademarks, etc., and reference to patents or utility models does not imply that such names or any such information are exempt from the relevant protective laws and regulations and, therefore, free for general use, nor does mention of suppliers or of particular commercial products constitute endorsement or recommendation for use. Printed on acid-free paper. Printed in the Federal Republic of Germany Cover Gesine Schulte, Max-Planck-Institutfiir

Molekulare Physiologie, Dortmund Composition Typomedia, Ostfddern Printing Strauss Offsetdruck, Morlenbach Bookbinding Buchbinderei Schaumann GmbH,

Darm stadt ISBN

3-527-29949-1

I'

Foreword That biological systems are masterful chemists is a fact long appreciated by those who study how living things build complexity from simple compounds in the environment. Enzymes catalyze the interconversion of vast numbers of chemical species, providing materials and energy to fuel cell survival and growth. Enzymes build the intricate natural products, which, for their potential utility in treating disease, pose almost unlimited new challenges for ambitious synthetic chemists. But, unlike most industrial chemical processes, Nature's catalysts generate few waste products and effect their transformations under mild conditions-in water, at room temperature and atmospheric pressure. Biocatalysts are models of energyefficient, environmentally-consciouschemistry and will play a prominent role in the 21Stcentury's chemicals industry. The world of biocatalysishas undergone significant change in the eight years since the first edition of this handbook appeared. Most of the news is good, with enzymes showing up in many more organic syntheses and a number of important new industrial processes coming on line. Apart from continuing clever insights into how to integrate biocatalysis into synthetic chemistry, several forces are accelerating a move to biocatalytic processes. In the first place, the search for better, enantiomerically pure drugs has forced many chemists to turn to enzymes for assistance in their preparation. Ever increasing demands for environmentally acceptable processes push in the same direction. At the same time, rapidly-developing technologies for making better catalysts through genetic enginering and for discovering new catalysts are are offering new process opportunities which in the past were either not economical or not even conceivable. A plethora of new catalysts to choose from, as well as a high probability that a catalyst can be further improved during the process design and engineering phases, means that we can respond rapidly to new synthetic needs with biocatalybc solutions. The organization of these volumes into specific technologies and transformations provides a comprehensive coverage of practical biocatalysis that no other single source provides. The work of experts in each of the fields, the individual chapters review vast relevant literature and synthesize it in order to present key concepts and many illustrative examples. This coverage should give organic chemists immediate access to the wealth of experience that has accumulated in the biocatalysis world and allow them to identify the most promising ways to use biocatalysts in their own

VI

I

Foreword

syntheses. Biocatalysts should feature prominently in the repertoire of synthetic chemistry, and this handbook deserves a prominent place in the modern chemist’s library. Pasadena, January, 2002

Frances Arnold

Preface Nearly eight years have passed since we the First Edition of ,,Enzyme Catalysis in Organic Synthesis“ was issued but much of what we had written in its preface then still applies today. The application of biocatalysis in organic synthesis is a powerful technique. It has grown steadily and today this field is well-established in both academia and industry. With increasing application and acceptance the need for a comprehensive and up to date overview of the state of the art has grown. In addition numerous colleagues have approached us and asked for an update of “the Handbook”. In response to these demands and in recognition of the new and groundbreaking strides taken since the first half of the nineties the Second Edition which is now in the hand of the reader was prepared. In comparing it with the First edition one discovers that we have not changed the overall arrangement in the volumes. Therefore we continue to have a part that addresses general principles (Chapters 1-10) and another one which summarizes the application of enzymes in organic synthesis according to reaction type (Chapters 11-20). This arrangement was very well received by the readers before and we hope that it will be for the Second Edition as well. However, the entire text was streamlined and in many cases regrouped to ensure for a better presentation. Also a few chapters which in the long run turned out to be less relevant to organic synthesis were not included again. In contrast other aspects were now integrated and attention was given to techniques of enzyme evolution, bioinformatics and enzymatic reactions in low-water media, areas that have developed with great pace and that we believe to be of major importance in the time to come. We hope that the Second Edition of the “Handbook will be a plentiful source of information just as valuable as the First Edition was eight years ago. Dortmund and Hanau, February 2002

Karlheinz Drauz, Herbert Waldmann

I

Contents Foreword V Preface VII Volume I 1

Introduction 1 Maria- Regina Kulo

1.1 1.2 1.3 1.4 1.5 1.5.1 1.5.2 1.5.3 1.5.4 1 .G 1.7 1.8 1.8.1 1.8.2 1.8.3 1.8.4

Enzymes as Catalysts 1 Enzyme Structure and Function 4 Cofactors and Coenzymes 12 Enzyme Nomenclature 21 Enzyme Kinetics 23 Reaction Rate and Substrate Concentration 23 Inhibitors and Efl’ectors 26 Influence of pH and Buffers 27 Temperature 28 Organic Solvents as Reaction Media 31 Enzyme Handling: Quality Requirements 32 Biotransforrnation Using Whole Cells 33 General Aspects 33 Biotransformation with Growing Cells 36 Biotransformation with Resting Cells 37 Biotransforrnationswith Permeabilized or Dried Cells 37 Bibliography 38

2

Production and I d a t i o n of Enzymes 41 Yoshihiko Hirose

2.1 2.2 2.3 2.3.1 2.3.2 2.3.3

Introduction 41 Enzyme Suppliers for Biotransforrnation 44 Origins of Enzymes 45 Microbial Enzymes 45 Plant Enzymes 46 Animal enzymes 46

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2.4 2.4.1 2.4.2 2.4.3 2.5 2.5.1 2.5.2 2.5.3 2.6 2.7 2.7.1 2.7.1.1 2.7.1.2 2.7.1.3 2.7.1.4 2.7.1.5 2.7.1.6 2.7.1.7 2.7.2 2.7.2.1 2.7.2.2 2.7.2.3 2.7.2.4 2.7.3 2.7.4 2.7.5 2.7.5.1 2.7.5.2 2.8

Fermentation of Enzymes 46 Liquid Fermentation 46 Solid Fermentation 47 Extraction of Enzymes 47 Extraction of Enzymes 47 Microbial Enzymes 47 Plant Enzymes 48 Animal Enzymes 48 Concentration 48 Purification of Enzymes 49 Chromatography 49 Ion Exchange Chromatography (IEX) 49 Hydrophobic Interaction Chromatography (HIC) 54 Gel Filtration (GF) 56 Reversed Phase Chromatography 58 Hydrogen Bond Chromatography 59 Affinity Chromatography 59 Saltingout Chromatography 62 Precipitation 62 Precipitation by Salting out 62 Precipitation by Organic Solvents 63 Precipitation by Changing pH 63 Precipitation by Water-Soluble Polymer 63 Crystallization 64 Stabilization During Purification 64 Storage of Enzymes 64 Storage in Liquids 64 Storage in Solids 65 Commercial Biocatalysts 65 References 66

3

Rational Design of Functional Proteins Tadayuki lmanaka and Haruyuki Atomi

3.1 3.2 3.3 3.4 3.5 3.6 3.6.1

Protein Engineering 67 Gene Manipulation Techniques in Enzyme Modification 68 Protein Crystallization 70 Comparative Modeling of a Protein Structure 73 What is Needed to Take a Rational Approach? 75 Examples of Protein Engineering 76 Protein Engineering Studies: Providing a Rational Explanation for Enzyme Specificity 76 Enhancing the Thermostability of Proteases 78 Contribution of Ion Pairs to the Thermostability of Proteins from Hyperthermophiles 79

3.6.2 3.6.3

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Contents

3.6.4 3.6.5 3.6.6 3.6.7 3.6.8 3.6.9 3.6.10 3.7

4

4.1 4.2 4.2.1 4.2.2 4.3 4.3.1 4.3.1.1 4.3.1.2 4.3.1.3 4.3.2 4.3.2.1 4.3.2.2 4.3.2.3 4.4 4.4.1 4.4.2 4.4.2.1 4.4.2.2 4.4.2.3 4.4.2.4 4.4.3 4.5 4.5.1 4.5.2 4.5.3 4.5.3.1 4.5.3.2 4.6

Thermostability Engineering Based on the Consensus Concept 80 Changing the Optimal pH of an Enzyme 81 Changing the Cofactor Specificity of an Enzyme 82 Changing the Substrate Specificity of an Enzyme 84 Changing the Product Specificity of an Enzyme 85 Combining Site-directed Mutagenesis with Chemical Modification 86 Changing the Catalyhc Activity of a Protein 087 Conclusions 89 References 90 Enzyme Engineering by Directed Evolution 95

Oliver May, Christopher A. Voigt and Frances H.Arnold

Introduction 95 Evolution as an Optimizing Process 96 The Search Space of Chemical Solutions 97 The Directed Evolution Algorithm 98 Creating a Librarr of Diverse Solutions 99 Mutagenesis 99 Random Point Mutagenesis of Whole Genes 99 Focused Mutagenesis 104 Calculation of Mutagenesis Hot-Spots 105 Recombination 107 In Vitro Recombination 107 In vivo Recombination 110 Family Shuffling 111 Finding Improved Enzymes: Screening and Selection 112 You Get What You Screen For 113 Screening Strategies 113 Low-Throughput Screening 114 High-Throughpu t Screening 115 Choosing Low versus High Throughput 116 Analyzing the Mutant Fitness Distribution 117 Selection and Methods to Link Genotype with Phenotype 119 Applications of Directed Evolution 121 Improving Functional Enzyme Expression and Secretion 122 Engineering Enzymes for Non-natural Environments 127 Engineering Enzyme Specificity 129 Substrate Specificity 129 Enantioselectivity 131 Conclusions 132 References 133

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5

Enzyme Bioinformatics

139

Kay Hofmann

5.1 5.2 5.2.1 5.2.2 5.2.3 5.2.4 5.3 5.3.1 5.3.2 5.3.3 5.3.4 5.4 5.4.1 5.4.2 5.4.3 5.5 5.5.1 5.5.2 5.5.3 5.5.4 5.5.5 5.5.6 5.5.7 5.6 5.6.1 5.6.2 5.6.3 5.7 5.7.1 5.7.2 5.7.3 5.8

6

Introduction 139 Protein Comparison 140 Sequence Comparison versus Structure Comparison 140 Substitution Matrices in Sequence Comparisons 141 Profile Methods 142 Database Searches 144 Enzyme-specificConservation Patterns 145 General Conservation Patterns 145 Active Site Conservation Patterns 146 Metal Binding Conservation Patterns 146 Making Use of Conservation Patterns 148 Modular Enzymes 149 The Domain Concept in Structure and Sequence 149 A Classification of Modular Enzymes 150 Inhibitory Domains 151 Enzyme Databases and Other Information Sources 151 E. C. Nomenclature and ENZYME Database 152 BRENDA 152 KEGG and LIGAND database 153 UM-BBD 153 Structural Databases 153 Metalloprotein Databases 154 Databases for Selected Enzyme Classes 154 Protein Domain and Motif Databases 154 PROSITE 155 PFAM 156 Other Related Databases 156 Enzyme Genomics 156 Ortholog Search 157 Paralog Search 157 Non-homology Based methods 159 Outlook 159 References 161 Immobilization o f Enzymes 163 James Lalonde

6.1

6.2 6.2.1 6.2.2 6.2.2.1 6.2.3

Introduction 163 Methods of Immobilization 164 Non-Covalent Adsorption 165 Covalent Attachment 168 Carriers for Enzyme Immobilization 170 Entrapment and Encapsulation 171

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Contents xi’i

6.2.4 6.3 6.3.1 6.3.2 6.3.3 6.3.4 6.4 6.4.1 6.4.2 6.4.3

Cross-Linking 175 Properties of Iminobilized Biocatalysts 175 Mass Transfer Effects 176 Partition 176 Stability 177 Activity of Immobilized Enzymes 177 New Developments and Outlook 178 Cross-linked Enzyme Crystals (CLEC@) 179 Sol-Gel 181 Controlled Solubility “Smart Polymers” 181 References 182

7

Reaction Engineering for Enzyme-Catalyzed Biotransformations Manfed Biselli, Udo Kragl and Christian Wandrey

7.1 7.2 7.3 7.3.1 7.3.2 7.3.2.1 7.3.2.2

Introduction 185 Steps of Process Optimization 186 Investigation of the Reaction System 190 Properties of the Enzyme 190 Properties of the Reaction System 193 Thermodynamic Equilibrium of the Reaction 193 Complex Reaction Systems: The Existence of Parallel and Consecutive Reactions 195 Other Properties of the Reaction System 204 Application of Organic Solvents 204 Investigation of Enzyme Kinetics 208 Methods of Parameter Identification 209 The Kinetics of One-Enzyme Systems 210 THE Michaelis-Menten Kinetics 210 Competitive Inhibition 214 Non-Competitive Inhibition 215 Uncompetitive Inhibition 216 Reversibility of One-Substrate Reactions 217 Two-Substrate Re,ictions 218 Kinetics of Aminoacylase as Example of a Random Uni-Bi Mechanism 223 Kinetics of Multiple Enzyme Systems 230 Enzyme Reactors 232 Basic Reaction Engineering Aspects 232 Reactors for Soluble Enzymes 238 Reactor Optimization Exemplified by the Enzyme Membrane Reactor 241 Control of Conversion in a Continuously Operated EMR 249 Reactor Systems for Immobilized Enzymes 250 Reaction Techniques for Enzymes in Organic Solvent 251

7.3.2.3 7.3.2.4 7.4 7.4.1 7.4.2 7.4.2.1 7.4.2.2 7.4.2.3 7.4.2.4 7.4.2.5 7.4.2.6 7.4.2.7 7.4.3 7.5 7.5.1 7.5.2 7.5.2.1 7.5.2.2 7.5.3 7.5.4

185

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7.6

Conclusions and Outlook 253 References 254

8

Enzymic Conversions in Organic and Other Low-Water Media Peter Halling

8.1 8.2 8.2.1 8.2.2 8.2.3 8.2.4 8.2.5 8.2.6 8.3 8.3.1 8.3.2 8.3.3 8.4 8.5 8.6 8.6.1 8.6.2 8.6.3 8.6.4 8.6.5 8.6.6 8.7 8.7.1 8.7.2 8.7.3

Introduction 259 Enzyme Form 260 Lyophilized Powders 260 Immobilized Enzymes 261 Cross-Linked Crystals 261 Direct Precipitation in Organic Solvents 262 Additives in Catalyst Powders 262 Solubilized Enzymes 263 Residual Water Level 264 Fixing Initial Water Activity of Reaction Components 266 Control of Water Activity During Reaction 269 “Water Mimics” 273 Temperature 274 Substrate (Starting Material) Concentrations 274 Solvent Choice 276 Effects on Equilibrium Position 276 “Solvent Effects” that Really are Not 276 Solvent Polarity Trend and Recommended Choices 277 Solvent Parameters 279 Solvent Effects on Selectivity 280 No Solvent or Little Solvent Systems 280 Acid-Base Conditions 281 pHMemory 281 Processes Erasing pH Memory 282 Systems for Acid-Base Buffering 283 References 285

9

Enzymatic Kinetic Resolution 287 Jonathan M.J. Williams, RebeccaJ. Parker, and Claudia Neri

9.1 9.2 9.2.1 9.2.2 9.2.3 9.3 9.3.1 9.3.2 9.3.3 9.4

Introduction 287 Alcohols and their Derivatives 288 Cyanohydrins 289 Other Readily Racemized Substrates 290 Enzyme and Metal Combinations 293 Carboxylic Acids and their Derivatives 297 Readily Enolized Carboxylic Acid Derivatives 297 Amino-Esters and Related Compounds 301 Reactions of cyclic amino acid derivatives 302 Reduction of fi-Ketoesters 307

259

‘Ontents

9.5

Conclusion 309 References 310

10

Enzymes from Extreme Therrnophilic and HyperthermophilicArchaea and Bacteria 313 Costanzo Bertoldo and Carabed Antranikian

10.1 10.2 10.2.1 10.2.1.1 10.2.1.2 10.2.1.3 10.3 10.3.1 10.4 10.4.1 10.5 10.6 10.6.1 10.7

Introduction 31 3 Starch-ProcessingEnzymes 315 Thermoactive Annylolyhc Enzymes 316 Heat-StableAmylases and Glucoamylases 316 a-Glucosidases 317 Thermoactive Pullulanases and CGTases 317 Cellulose-Hydro1yzing Enzymes 0321 Thennostable Cellulases 321 Xylan-Degrading Enzymes 324 Thermostable Xylanases 324 Chitin Degradation 325 Proteolytic Enzyrnes 326 Stable Proteases 327 Intracellular Enzymes 329 References 331 Volume I1

11

Hydrolysis and Formation of C - 0 Bonds 335

11.1

Hydrolysis and Fmnation of Carboxylid Acid Esters 335 Hans-joachim Cars and Fritz Theil

11.1.1 11.1.1.1 11.1.1.2 11.1.1.3

11.2

Hydrolysis and Formation of Carboxylic Acid Esters 351 Hydrolysis of Carboxylic Acid Esters 351 Formation of Carboxylic Esters 472 Inter- and Intramolecular Alcoholysis 545 References 574 Hydrolysis of Epoxides 579

Kurt Faber and Romano V: A. Orru 11.2.1 Epoxide Hydrolases in Nature 581 11.2.1.1 Isolation and Characterization of Epoxide Hydrolases 582

11.2.1.2 11.2.1.3 11.2.2 11.2.2.1 11.2.2.2 11.2.2.3 11.2.3 11.2.3.1

Structure and Mechanism of Epoxide Hydrolases 584 Screening for Microbial Epoxide Hydrolases 587 Microbial Hydrolysis of Epoxides 588 Fungal Enzymes 588 Bacterial Enzymes 590 Yeast Enzymes 591 Substrate Specificity and Selectivity 592 Asymmetrization of meso-Epoxides 592

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11.2.3.2 11.2.3.3 11.2.4 11.2.5 11.2.6 11.3

Resolution of Racemic Epoxides 592 Deracemization Methods 596 Use of Non-Natural Nucleophiles 599 Applications to Asymmetric Synthesis GOO Summary and Outlook 604 References 605 Hydrolysis and Formation of Glycosidic Bonds

609

Chi-Huey Wong

1.3.1 11.3.2 11.3.2.1 11.3.2.2 11.3.2.3 11.3.2.4 11.3.3 11.3.4 11.3.4.1 11.3.4.2 11.3.4.3 11.3.5 11.3.5.1 11.3.5.2 11.3.6 11.3.6.1 11.3.6.2 11.3.7 11.4

Introduction 609 Glycosyltransferases of the Leloir Pathway 611 Synthesis of Sugar Nucleoside Phosphates 613 Substrate Specificity and Synthetic Applications of Glycosyltransferases 619 In Situ Cofactor Regeneration 626 Cloning and Expression of Glycosyltransferases 628 Non-Leloir Glycosyltransferases:Transfer of Glycosyl donors from Glycosyl Phosphates and Glycosides 631 Glycosidases 633 Equilibrium-controlledSynthesis 633 Kinetically Controlled Synthesis 634 Selectivity 634 Synthesis of N-glycosides 637 Nucleoside Phosphorylase 638 NAD Hydrolase 639 Biological Applications of Synthetic Glycoconjugates 639 Glycosidase and Glycosyl Transferase Inhibitors 639 Glycoprotein Remodeling 641 Future Opportunities 642 References 643 Natural Polysaccharide-degrading Enzymes 653 Constanzo Bertoldo and Carabed Antranikian

11.4.1

Introduction 653 Starch 6 5 3 11.4.2.1 Classification of Starch-degrading Enzymes 654 11.4.2.2 a-Amylase (1,4-a-~-Glucan,4-G1ucanhydrolase, E. C. 3.2.1.1) 655 11.4.2.3 P-Amylase (1,4-a-~-Glucan Maltohydrolase, E. C. 3.2.1.2) 656 11.4.2.4 Glucoamylases (1,4-a-~-glucan glucohydrolase,E. C. 3.2.1.3) 656 11.4.2.5 a-Glucosidase (a-6-GlucosideGlucohydrolase, E. C. 3.2.1.20) 657 11.4.2.6 Isoamylase (Glycogen 6-Glucanohydrolase,E. C. 3.2.1.68) 657 11.4.2.7 Pullulanase Type I (a-Dextrin 6-Glucanohydrolase,E. C. 3.2.1.41) 657 11.4.2.8 Pullulanase Type I1 or Amylopullulanase 658 11.4.2.9 Pullulan Hydrolases (Type I, Neopullulanase;Trpe 11, Isopullulanase, E. C. 3.2.1.57, Pullulan Hydrolase Type 111) 659 11.4.2.10 Cyclodextrin Glycolsyltransferase (1,4-a-~-Glucan 4-a-~-(1,4-a-~G1ucano)-Transferase,E. C. 2.4.1.19) 659 11.4.2

=Ontents

11.4.2.11 Biotechnological Applications of Starch-degrading Enzymes 659 11.4.3 Cellulose 661 11.4.3.1 Cellulose-degrading Enzyme Systems 663 11.4.3.2 Endoglucanase (I,4-P-~-Glucan-Glucanohydrolase, E.C. 3.2.1.4) 663 11.4.3.3 Cellobiohydrolase (1,4-P-~-Glucan Cellobiohydrolase, E. C. 3.2.1.91) 663 11.4.3.4 P-Glucosidase (P-D-GlucosideGlucohydrolase, E. C. 3.2.1.21) 664 11.4.3.5 Fungal and Bacterial Cellulases 664 11.4.3.6 Structure and Synergistic Effect of Cellulases 665 11.4.4 Xylan 667 11.4.4.1 The Xylanolytic Elnzyme System 668 11.4.4.2 Endoxylanase (1;l-P-D-Xylan Xylanohydrolase, E. C. 3.2.1.8) 670 11.4.4.3 P-Xylosidase (P-c-XylosideXylohydrolase, E. C. 3.2.1.37) 670 11.4.4.4 a-L-Arabinofuranosidase (E. C. 3.2.1.55) 671 11.4.4.5 a-Glucuronidase (E.C. 3.2.1.136) 671 11.4.4.6 Acetyl Xylan Esterase (E.C. 3.1.1.6) 672 11.4.4.7 Mechanism of Action of Endoxylanase 672 11.4.4.8 Biotechnological Applications of Xylanases 672 11.4.5 Pectin 673 11.4.5.1 Classification of Pectic Substances 675 11.4.5.2 Pectolpc Enzymes 675 11.4.5.3 Classification of Pectolytic Enzymes 676 11.4.5.4 Protopectinase 676 11.4.5.5 Pectin Methylesttxase 677 11.4.5.6 Pectin and Polygalacturonate Depolymerizing Enzymes 677 11.4.5.7 Pectin and Polygalacturonate Hydrolase 678 11.4.5.8 Pectin and Polygalacturonate Lyase 679 11.4.5.9 Biotechnological Applications of Pectolytic Enzymes 680 References 681 11.5 Addition of Water to C=C Bonds 686 Marcel Wubbolts

11.5.1 11.5.2 11.5.2.1 11.5.1.2 11.5.3 11.5.3.1 11.5.4 11.5.5 11.5.5.1 11.5.5.2 11.5.6 11.5.6.1 11.5.6.2 11.5.7

Addition of Water to Alkenoic Acids 686 Addition of Water to Alkene-Dioic Acids 687 L- and D-Malic A'cid 687 Substituted Malic Acids 688 Addition of Water to Alkene-TricarboxylicAcids 688 Citric Acid and Derivatives 688 Addition of Water to Alkynoic Acids 690 Addition of Water to Enols 690 Carbohydrates: Addition of Water to 2-Keto-3-Deoxysugars 690 Addition/Elimination of Water with Other Enols 691 Addition of Water to Unsaturated Fatty Acids 693 CoA and ACP Coupled Fatty Acid Hydratases 693 Hydratases Acting on Free Fatty Acids 695 Addition of Water to Steroids 695 References 690

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12

12.1 12.1.1 12.1.2 12.1.2.1 12.1.2.2 12.1.3 12.1.3.1 12.1.3.2 12.1.3.3 12.1.3.4 12.1.3.5 12.1.3.6 12.1.4 12.2

12.2.1 12.2.2 12.2.3 12.2.3.1 12.2.3.2 12.2.3.3 12.2.4 12.2.5

12.2.5.1

12.2.5.2 12.2.5.3 12.2.6 12.3 12.3.1 12.3.2 12.3.2.1 12.3.2.2 12.3.2.3 12.3.2.4 12.3.3

Hydrolysis and Formation of C-N Bonds 699

Hydrolysis of Nitriles 699 Birgit Schulze Introduction 699 Types of Nitrile Hydrolyzing Enzymes 700 Enzymatic Hydrolysis of Organic Nitriles 700 Enzymatic Hydrolysis of Cyanide 702 Examples of Enzymatic Nitrile Hydrolysis 703 Enantioselective Hydrolysis of Nitriles 703 Monohydrolysis of Dinitriles 705 Substrate and Product Inhibition of Nitrile Hydrolysis 708 Activation and Stabilization of Nitrile Hydratases 710 Nitrile Hydrolysis in Organic Solvents 710 Large Scale Production of Acrylamide 711 Availability and Industrial Future of Nitrile Hydrolyzing Biocatalysts 713 References 713 Formation and Hydrolysis of Amides 716 Birgit Schulze and Erik de Vroom Introduction 716 Enzymatic Formation of Amides 716 Enzymatic Enantioselective Hydrolysis of Amides 719 Hydrolysis of Carboxylic Amides 719 Hydrolysis of Amino Acid Amides 720 Hydrolysis of Cyclic Amides 727 Selective Cleavage of the C-Terminal Amide Bond 728 Amidase Catalyzed Hydrolytic and Synthetic Processes in the Production of Semi-syntheticAntibiotics 729 Enzymatic Production of 6-APA, 7-ADCA and 7-ACA Using Amidases: Hydrolytic Processes 730 A New Fermentation-based Biocatalytic Process for 7-ADCA 735 Enzymatic Formation of Semi-syntheticAntibiotics: Synthetic Processes 735 Conclusions and Future Prospects 737 References 738 Hydrolysis of N-Acylamino Acids 741 Andreas 5.Bommarius Introduction 741 Acylase I (N-AcylaminoAcid Amidohydrolase, E. C. 3.5.1.4.) 742 Genes, Sequences, Structures 743 Substrate Specificity 744 Stability of Acylases 746 Thermodynamics and Mechanism of the Acylase-catalyzed Reaction 748 Acylase I I (N-Acyl-L-Aspartate Amidohydrolase,Aspartoacylase, E.C. 3.5.1.15.) 749

Contents

12.3.4 12.3.5 12.3.6 12.3.7 12.4

Proline Acylase 1:N-Acyl-L-ProlineAmidohydrolase) 752 Dehydroamino k i d Acylases 753 D-Specific Aminoacylases 754 Acylase Process on a Large Scale 757 References 758 Hydrolysis and Formation of Hydantoins 761 Markus Pietzsch m d Christoph Syldatk

12.4.1 12.4.2 12.4.3 12.4.4 12.4.5 12.4.6 12.4.7 12.5

Classification and Natural Occurrence of Hydantoin Cleaving and Related Enzymes 761 D-Hydantoinases- Substrate Specificity and Properties 773 DN-Carbamoylases - Substrate Specificity and Properties 777 L-Hydantoinases- Substrate Specificity and Properties 784 L-N-Carbamoylases- Substrate Specificityand Properties 786 Hydantoin Raceinases 792 Conclusions 794 References 796 Hydrolysis and Formation of Peptides 800 Hans-Dieterjakulike

12.5.1 12.5.2 12.5.2.1 12.5.2.2 12.5.3 12.5.3.1 12.5.3.2 12.5.3.3 12.5.3.4 12.5.3.5 12.5.3.6 12.5.3.7 12.5.4 12.6

Introduction 800 Hydrolysis of Peptides 801 Peptide-CleavingEnzymes 801 Importance of Proteolysis 813 Formation of Peptides 818 Tools for Peptide Synthesis 818 Choice of the Ideal Enzyme 822 Principles of Enzymatic Synthesis 823 Manipulations to Suppress Competitive Reactions 831 Approaches to Irreversible Formation of Peptide Bond 840 Irreversible C-N Ligations by Mimicking Enzyme Specificity 842 Planning and Process Development of Enzymatic Peptide Synthesis 851 Conclusion and Outlook 858 References 859 Addition of Amines to C=C Bonds 866 Marcel Wubbolts

12.6.1 12.6.1.1 12.6.1.2 12.6.1.3 12.6.1.4 12.6.1.5 12.6.1.6 12.6.2 12.6.2.1

Addition of Ammonia to Produce Amino Acids 866 Aspartic Acid 866 Aspartic Acid Derivatives 868 Histidine Ammonia Lyase 869 Phenylalanine, Tyrosin and L-DOPA 870 Serine and Threonine Deaminases 871 Ornithine Cyclocleaminase 871 Ammonia Lyases that Act on Other Amines 871 Elimination of Ammonia from Ethanolamine 871 References 872

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12.7

Transaminations 873

J. David Rozzell and Andreas 5.Bommarius

tntroduction 873 Description of Transaminases 875 Homology and Evolutionary Subgroups of Aminotransferases 875 Mechanism of Transamination 875 Protein Engineering and Directed Evolution with Aminotransferases 876 Use of Aminotransferases in Biocatalytic Reactions 878 12.7.3 12.7.3.1 Synthesis of a-L-AminoAcids 878 12.7.3.2 Synthesis of Enantiomerically Pure Amines 880 12.7.3.3 Other Preparative Applications of Aminotransferases 881 Driving the Reaction to Completion 884 12.7.4 12.7.5 Production of L-Amino Acids Using Immobilized Transaminases 885 12.7.6 D-Amino Acid Transferases 889 Synthesis of Labeled Amino Acids 891 12.7.7 12.7.8 Availability of Enzyme 892 References 892

12.7.1 12.7.2 12.7.2.1 12.7.2.2 12.7.2.3

13

Formation and Cleavage of P - 0 Bonds 895 George M. Whitesides

Introduction 895 Enzymes Forming or Cleaving Phosphorous-OxygenBonds 896 Biological Phosphorylating Agents 899 Phosphorylation 901 Regeneration of Nucleoside Triphosphates 901 Regeneration of ATP from ADP and AMP 902 Regeneration of other Nucleoside Triphosphates 906 Applications 907 Phosphorylations with ATP as a Cofactor 907 P- 0 Bond Formation with other Nucleoside Triphosphates than ATP 909 13.2.2.3 Other Phosphorylating Agents 910 13.2.3 Tables Containing Typical Examples Ordered According to the Classes of Compounds 918 Cleavage of P - 0 bonds 918 13.3 13.3.1 Hydrolysis of Phosphate and Pyrophosphate Monoesters 919 13.3.2 Hydrolysis of S- and N-substituted Phosphate Monoester Analogs 920 13.3.3 Hydrolysis of Phosphate and Phosphonate Diesters 922 13.3.3.1 Nucleic Acids and their Analogs 922 13.3.3.2 Other Phosphate and Phosphonate Diesters 922 13.3.4 Other P - 0 Bond Cleavages 923 References 926

13.1 13.1.1 13.1.2 13.2 13.2.1 13.2.1.1 13.2.1.2 13.2.2 13.2.2.1 13.2.2.2

931

14

Formation o f C-C Bonds Chi-hey Wong

14.1 14.1.1 14.1.1.1 14.1.1.2

Aldol Reactions 931 DHAP-Utilizing Aldolases 931 Fructose 1,G-Diphosphate(FDP) Aldolase (E. C. 4.1.2.13) 931 Fuculose 1-Phosphate (Fuc 1-P) Aldolase (E.C. 4.1.2.17), Rhamnulose 1-Phosphate (Rh.a 1-P) Aldolase (E. C. 4.1.2.19) and Ragatose 1,6-Diphosphate (TDP) Aldolase 939 Synthesis of DihlydroxyacetonePhosphate (DHAP) 943 Pyruvate/Phosp:hoenolpyruvate-UtilizingAldolases 944 N-Acetylneurarninate (NeuAc)Aldolase (E.C. 4.1.3.3) and NeuAc Synthetase (E.C. 4.1.3.19) 944 3-Deoxy-~-mann.o-2-octu~osonate Aldolase (E. C. 4.1.2.23) and 3-Deoxy~-manno-2-octulosonate &Phosphate Synthetase (E. C. 4.1.2.16) 946 3-Deoxy-~-arabiii0-2-heptulosonic Acid 7-Phosphate (DAHP) Synthetase (E.C. 4.1.2.15) 947 2-Keto-4-hydroxyglutarate(KHG) Aldolase (E. C. 4.1.2.31) 948 2-Keto-3-deoxy-6-phosphogluconate (KDPG) Aldolase (E. C. 4.1.2.14) 949 2-Keto-3-deoxy-1~glucarate (KDG) Aldolase (E. C. 4.1.2.20) 950 2-Deoxyribose 5-phosphate Aldolase (DEW) (E. C. 4.1.2.4) 950 Ketol and Aldol Transfer Reactions 960 Transketolase (TK) (E. C. 2.2.1.1) 960 Transaldolase (TA) (E.C. 2.2.1.2) 962 Acyloin Condensation 962 C-C Bond Forming Reactions Involving AcetylCoA 963 Isoprenoid and Steroid Synthesis 965 p-Replacement of Chloroalanine 966 References 966 Enzymatic Synthesis of Cyanohydrins 974

14.1.1.3 14.1.2 14.1.2.1 14.1.2.2 14.1.2.3 14.1.2.4 14.1.2.5 14.1.2.6 14.1.3 14.2 14.2.1 14.2.2 14.3 14.4 14.5 14.6 14.7

Martin H. Fechtcr and Hefried Cringl

14.7.1 14.7.2 14.7.3 14.7.4 14.7.5 14.7.6 14.7.7 14.7.8 14.7.9

The Oxynitrilasm Commonly Used for Preparative Application 975 Oxynitrilase Caialysed Addition of HCN to Aldehydes 976 HNL-CatalyzedAddition of Hydrogen Cyanide to Ketones 978 Transhydrocyanation 978 Experimental Techniques for HNL-Catalysed Biotransformations 981 Resolution of Racemates 982 Follow-up Chemistry of Enantiopure Cyanohydrins 985 Safe Handling of Cyanides 985 Conclusions and Outlook 986 References 98.6

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Volume 111

15

Reduction Reactions 991

15.1

Reduction of Ketones

991

Kaoru Nakamura and Tomoko Matsuda

15.1.1 15.1.1.1 15.1.1.2 15.1.1.3

Introduction 991 Enzyme Classfication and Reaction Mechanism 991 Coenzyme Regeneration 992 Form ofthe Biocatalysts: Isolated Enzyme vs. Whole Cell 995 15.1.1.4 Origin of Enzymes 996 15.1.2 Stereochemical Control 997 15.1.2.1 Enantioselectivity of Reduction Reactions 997 15.1.2.2 Modification of the Substrate: Use of an “Enantiocontrolling” Group 998 15.1.2.3 Screening of Microorganisms 1000 15.1.2.4 Treatment of the Cell: Heat Treatment 1001 15.1.2.5 Treatment ofthe Cell: Aging 1001 15.1.2.6 Treatment of the Cell: High Pressure Homogenization 1002 15.1.2.7 Treatment of the Cell: Acetone Dehydration 1002 15.1.2.8 Cultivation Conditions of the Cell 1003 15.1.2.9 Modification of Reaction Conditions: Incorporation of an Inhibitor 1004 15.1.2.10 Modification of Reaction Conditions: Organic-Solvent 1005 15.1.2.11 Modification of Reaction Conditions: Use of a Supercritical Solvent 1006 15.1.2.12 Modification of Reaction Conditions: Cyclodextrin 1007 15.1.2.13 Modification of Reaction Conditions: Hydrophobic Polymer XAD 1007 15.1.2.14 Modification of Reaction Conditions: Reaction Temperature 1008 15.1.2.15 Modification of Reaction Conditions: Reaction Pressure 1009 15.1.3 Improvement of Dehydrogenases for use in Reduction Reactions by Genetic Methods 1010 15.1.3.1 Overexpression of the Alcohol Dehydrogenase 1010 15.1.3.2 Access to a Single Enzyme Within a Whole Cell: Use of Recombinant Cells 1011 15.1.3.3. Use ofa Cell Deficient in an Undesired Enzyme 1012 15.1.3.4 Point Mutation for the Improvement of Enantioselectivity 1012 15.1.3.5 Broadening the Substrate Specificity of Dehydrogenase by Mutations 1012 15.1.3.6 Production of an Activated Form of an Enzyme by Directed Evolution 1014 15.1.3.7 Change in the Coenzyme Specificity by Genetic Methods: NADP(H) Specific Formate 1014 15.1.3.8 Use of a Mutant Dehydrogenase for the Synthesis of 4-Amino-2-Hydroxy Acids 1014 15.1.3.9 Catalyhc Antibody 1015 15.1.4 Reduction Systems with Wide Substrate Specificity 1016

15.1.4.1 15.1.4.2 15.1.4.3 15.1.4.4 15.1.4.5 15.1.5 15.1.5.1 15.1.5.2 15.1.5.3 15.1.5.4 15.1.5.5 15.1.5.6 15.1.5.7 15.1.5.8 15.2 15.2.1 15.2.2 15.2.3 15.2.4 15.2.5 15.2.6 15.2.7 15.2.7.1 15.2.7.2 15.3 15.3.1 15.3.2 15.3.2.1 15.3.3 15.3.3.1 15.3.3.2 15.3.4 15.3.4.1 15.3.4.2 15.3.4.3 15.3.4.4 15.3.5 15.3.6 15.3.6.1 15.3.6.2

Bakers' Yeast 1016 Rodococcus Erythropolis 1016 Pseudomonas sp. !$trainPED and Lactobacillus Kefir 1017 Thermoanaerobium Brockii 1018 Geotrichurn Candidum 1019 Reduction of Various Ketones 1021 Reduction of Fluoroketones 1021 Reduction of Fluoroketones Containing Sulhr Functionalities 1024 Reduction of Chloroketones 1025 Reduction of Ketones Containing Nitrogen, Oxygen, Phosphorus and Sulfur 1028 Reduction of Diketones 1028 Reduction of Diary1 Ketones 1029 Diastereoslective Reductions (Dynamic Resolution) 1030 Chemo-enzymabc Synthesis of Bioaktive Compounds 1031 Reduction of Various Functionalities 1033 Reduction of Aldehydes 1033 Reduction of Peroxides to Alcohols 1034 Reduction of Sulfoxides to Sulfides 1034 Reduction of Azide and Nitro Compounds to Amines 1035 Reduction of Carbon-Carbon Double Bonds 1036 Transformation of a-Keto Acid to Amine 1037 Reduction of Carbon Dioxide 1038 Reduction of COz to Methanol 1038 Reductive fixation of COz 1039 References 1040 Reduction of C=lV bonds 1047 Andreas 5. Bomtmarius

Introduction 1047 Structural Features of Amino Acid Dehydrogenases (AADHs) 1049 Sequences and Structures 1050 Thermodynamics and Mechanism of Enzymatic Reductive Amination 1050 Thermodynamics 1050 Mechanism, Kinetics 1051 Individual Amino Acid Dehydrogenases 1052 Leucine Dehydrogenase (LeuDH, E. C. 1.4.1.9.) 1052 Alanine Dehydrogenase (AlaDH, E. C. 1.4.1.1.) 1053 Glutamate Dehydrogenase (GluDH, E. C. 1.4.1.2-4) 1054 Phenylalanine Dehydrogenase (PheDH, E.C. 1.4.1.20) 1054 Summary of Substrate Specificities 1056 Process Technology: Cofactor Regeneration and Enzyme Membrane Reactor (EMR) 1058 Regeneration of NAD(P)(H)Cofactors 1058 Summary of Processing to Amino Acids 1060 References 1001

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1065

16

Oxidation Reactions

16.1

Oxygenation of C-H and C=C Bonds 1065 Sabine Flitsch

16.1.1 16.1.2 16.1.2 16.1.4 16.1.4.1 16.1.4.1 16.1.4.3 16.1.4.4 16.1.5 16.1.5.1 16.1.5.2 16.1.5.3 16.1.5.4 16.1.6 16.1.7 16.1.7.1 16.1.7.2 16.2

Introduction 1065 Hydroxylating Enzymes 1066 Hydroxylating Enzymes 1068 Hydroxylation of Non-Activated Carbon Atoms 1069 Hydroxylation of Monoterpenes 1069 Hydroxylation of Monoterpenes 1075 Hydroxylation of Steroids 1078 Miscellaneous Compounds 1079 Epoxidation of Olefins 1084 Epoxidation of Straight-Chain Terminal Olefins 1084 Short-Chain Alkenes 1088 Terpenes 1090 Cyclic Sesquiterpenes 1096 Conclusions, Current and Future Trends 1097 Cis Hydroxylation of Aromatic Double Bonds 1099 Introduction 1099 Preparation of cis Dihydrodiols 1100 References 1103 Oxidation of Alcohols 1108 Andreas Schmid, Frank Hollmann, and Bruno Buhler

16.2.1 16.2.2 16.2.2.1 16.2.2.2 16.2.2.3 16.2.2.4 16.2.2.5 16.2.2.6

Introduction 1108 Dehydrogenases as Catalysts 1108 Regeneration of Oxidized Nicotinamide Coenzymes 1108 Dehydrogenases as Regeneration Enzymes 1109 Molecular Oxygen as Terminal Acceptor 1111 Electrochemical Regeneration 11 12 Photochemical Regeneration 1114 Oxidations Catalyzed by Alcohol Dehydrogenase from Horse Liver (HLADH) 1115 16.2.2.7 Alcohol Dehydrogenase from Yeast (YADH) 1120 16.2.2.8 Alcohol Dehydrogenase from Thernaoanaerobiurn brokii (TBADH) 1120 16.2.2.9 Glycerol Dehydrogenase (GDH, E. C. 1.1.1.6) 1122 16.2.2.10 Glycerol-3-phosphateDehydrogenase (GPDH, E. C. 1.1.1.8) 1124 16.2.2.11 Lactate Dehydrogenase (LDH, E.C. 1.1.1.27) 1125 16.2.2.12 Carbohydrate Dehydrogenases 1126 16.2.2.13Hydroxysteroid Dehydrogenases (HSDH) 1127 16.2.2.14 Other Dehydrogenases 1127 16.2.3 Oxidases as Catalysts 1129 16.2.3.1 General Remarks 1129 16.2.3.2 Methods to Diminish/Avoid H202 1129

Contents

16.2.3.3 16.2.3.4 16.2.3.5 16.2.3.6 16.2.3.7 16.2.3.8 16.2.3.9 16.2.4 16.2.4.1 16.2.4.2 16.2.4.3 16.2.4.4 16.2.5 16.2.5.1 16.2.5.2 16.2.5.3 16.2.6 16.2.6.1 16.2.6.2 16.2.6.3 16.2.6.4 16.2.6.5 16.2.7 16.2.7.1 16.2.7.2 16.3

Pyranose Oxidasa (P20, E.C. 1.1.3.10) 1132 Glycolate Oxidase (E.C. 1.1.3.15) 1135 Nucleoside Oxidase (E. C. 1.1.3.28) 1138 Glucose Oxidase (E. C. 1.1.3.4) 1138 Alcohol oxidase (E.C. 1.1.3.13) 1139 Galactose Oxidase (E. C. 1.1.3.9) 1141 Cholesterol Oxidase (ChOX, E. C. 1.1.3.6) 1142 Peroxidases as Catalysts 1142 Introduction 1142 Methods to Generate HzOz 1143 Chloroperoxidase (CPO, E. C. 1.11.1.10) 1145 Catalase (E.C. 1.11.1.6) 1145 Quinoprotein Dehydrogenases (QDH) 1146 General Remarks 1146 Methanol Dehydrogenase (E.C. 1.1.99.8) 1147 Glucose Dehydrogenase (E. C. 1.1.99.17) 1148 Whole-Cell Oxidations 1148 Stereoselective Oridation of (-)-Carve01 to (-)-Carvone 1148 Sugar Dehydrogenases Applied in Whole Cells 1149 Oxidation of Aromatic and Aliphatic Alcohols to Corresponding Aldehydes and Acids 1150 Enantiospecific Reactions 1154 Stereoinversions using Microbial Redox Reactions 1157 Miscellaneous I162 Biofuel Cells 1162 Biomimetic Analogs to Nicotinamide Co-nzymes 1163 References 1164 Oxidation of Phenols 1170 Andreas Schmid, Frank Hollmann, and Bruno Biihler

16.3.1 16.3.2 16.3.2.1 16.3.2.2 16.3.3 16.3.3.1 16.3.3.2 16.3.4 16.3.4.1 16.3.4.2 16.3.4.3 16.3.5 16.3.5.1 16.3.5.2 16.3.6 16.3.6.1

Introduction 1170 Oxidases 1170 Vanillyl oxidase (E. C. 1.1.3.38) 1170 Laccase (E.C. 1.10.3.2) 1174 Monooxygenases 1176 Tyrosinase (E.C. 1.10.3.1) 1176 2-Hydroxybiphen1yl-3-monooxygenase (HbpA, E. C . 1.14.13.44) 1179 Peroxidases 11115 Oxidative Coupling Reactions 1185 Hydroxylation of Phenols 1186 Nitration of Phenols 1187 Other Oxidoreductases 1188 4-Cresol-oxidorecluctase (PCMH, E. C. 1.17.99.1) 1188 4-Ethylphenol Oxidoreductas 1189 In vivo Oxidations 1190 Phenoloxidase of Mucuna pwriens 1190

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16.3.6.2 Monohydroxylationof (R)-2-PhenoxypropionicAcid and Similar Substrates 1191 16.3.6.3 Biotransformation of Eugenol to Vanillin 1191 References 1192 Oxidation of Aldehydes 1194 16.4 Andreas Schmid, Frank Hollmann, and Bruno 6uhler

16.4.1 16.4.2 16.4.3 16.4.4 16.4.4.1 16.4.4.2 16.4.5 16.4.5.1 16.4.6 16.5

Introduction 1194 Alcohol Dehydrogenases 1194 Aldehyde Dehydrogenases 1196 Monooxygenases 1198 Luciferase (E. C. 1.14.14.3) 1198 P 4 5 0 ~ ~ . 31199 Oxidases 1201 Xanthine Oxidase (E.C. 1.1.3.22) 1201 Oxidations with Intact Microbial Cells 1201 References 1201 Baeyer-Villiger Oxidations 1202 Sabine Flitsch and Gideon Grogan

16.5.1 16.5.1.1 16.5.1.2 16.5.1.3 16.5.1.4 16.5.2 16.5.2.1 16.5.2.2 16.5.2.3 16.5.3 16.5.4 16.5.5 16.6

Introduction 1202 Steroidal Substrates 1202 Aliphatic Substrates 1205 Alicyclic Substrates 1207 Polycyclic Molecules 1212 Baeyer-Villiger Monooxygenases 1213 Type 1 BVMOs 1214 Type 2 BVMOs 1216 Mechanism of the Enzymatic Baeyer-Villiger Reaction 1216 Synthetic Applications 1222 Models for the Action of Baeyer-Villiger Monooxygenases 1234 Conclusion and Outlook 1238 References 1241 Oxidation of Acids 1245 Andreas Schmid, Frank Hollmann, and Bruno Buhler

Introduction 1245 Pyruvate Oxidase (PYOx, E.C. 1.2.3.3) 1246 Formate Dehydrogenase (FDH, E. C. 1.2.1.2) 1247 Oxidations with Intact Microbial Cells 1247 Production of Benzaldehyde from Benzoyl Formate or Mandelic Acid 1247 16.6.4.2 Microbial Production of &,cis-Muconic Acid from Benzoic Acid 1248 16.6.4.3 Biotransformation of Substituted Benzoates to the Corresponding cis-Diols 1249 References 1249 16.6.1 16.6.2 16.6.3 16.6.4 16.6.4.1

Contents

16.7

Oxidation of C-N Bonds 1250 Andreas Schmid, thank Hollmann, and Bruno Buhler

16.7.1 16.7.2 16.7.2.1 16.7.2.2 16.7.3 16.7.3.1 16.7.3.2 16.7.4 16.8

Introduction L!50 Oxidations Catalyzed by Dehydrogenases 1251 L-Alanine Dehydrogenase (L-Ala-DH,E. C. 1.4.1.1) 1251 Nicotinic Acid Dchydrogenase (Hydroxylase)(E. C. 1.5.1.13) 1252 Oxidations Catalyzed by Oxidases 1254 Amino Acid Oxiclases 1254 Amine Oxidases 1256 Oxidations Catalyzed by Transaminases 1260 References 1261 Oxidation at Sulfur 1262 Karl-Heinz van Pke

16.8.1 Enzymes Oxidizing at Sulfur and their Sources 1262 16.8.2 Oxidation of Sulfides 1263 16.8.2.1 Oxidation of Sulfides by Monooxygenasesand by Whole Organsims 1263 16.8.2.2 Oxidation of Sulfides by Peroxidases and Haloperoxidases 1264 References 1266 16.9 Halogenation 1267 Karl-Heinz van Pke

16.9.1 16.9.1.1 16.9.1.2 16.9.2 16.9.2.1 16.9.2.2 16.9.3 16.9.3.1 16.9.3.2 16.9.4 16.9.4.1 16.9.5

17

17.1 17.2 17.2.1

Classification of lialogenating Enzymes and their Reaction Mechanisms 1;!67 Haloperoxidases and Perhydrolases 1267 FADH2-dependent Halogenases 1268 Sources and Production of Enzymes 1268 FADHz-dependentHalogenases 1268 Haloperoxidases and Perhydrolases 1269 Substrates for Hadogenating Enzymes and Reaction Products 1271 Halogenation of ,4romatic Compounds 1271 Halogenation of Aliphatic Compounds 1273 Regioselectivity and Stereospecificityof Enzymatic Halogenation Reactions 1275 FADH2-dependentHalogenases 1275 Comparison of Chemical with Enzymatic Halogenation 1277 References 1275 lsornerizations 1281 Nobuyoshi Esaki, 'Tatsuo Kurihara, and Kenji Soda

Introduction 12.81 Racemizations and Epimerizations 1282 Pyridoxal 5'-phosphate-dependentAmino Acid Racemases and Epimerases 1283 17.2.1.1 Alanine Racemase (E. C. 5.1.1.1) 1283

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17.2.1.2 Amino Acid Racemase with Low Substrate Specificity (E.C. 5.1.1.10) 1289 17.2.1.3 a-Amino-E-caprolactam Racemase 1292 17.2.2 Cofactor-independent Racemases and Epimerases Acting on Amino Acids 1293 17.2.2.1 Glutamate Racemase (E. C. 5.1.1.3) 1293 17.2.2.2 Aspartate Racemase (E. C. 5.1.1.13) 1297 17.2.2.3 Diaminopimelate Epimerase (E. C. 5.1.1.7) 1299 17.2.2.4 Proline Racemase (E. C. 5.1.1.4) 1301 17.2.3 Other Racemases and Epimerases Acting on Amino Acid Derivatives 1301 17.2.3.1 2-Amino-A2-thiazoline-4-carbo~ylateRacemase 1301 17.2.3.2 Hydantoin Racemase 1303 17.2.3.3 N-Acylamino Acid Racemase 1306 17.2.3.4 Isopenicillin N Epimerase 1308 17.2.4 Racemization and Epimerization at Hydroxyl Carbons 1310 17.2.4.1 Mandelate Racemase (E. C. 5.1.2.2) 1310 17.3 Isomerizations 1312 17.3.1 D-Xylose (Glucose) Isomerase (E.C. 5.3.1.5) 1313 17.3.1.1 Properties 1313 17.3.1.2 Reaction Mechanism 1314 17.3.1.3 Production of Fructose 1316 17.3.1.4 Production of Unusual Sugar Derivatives 1316 17.3.2 Phosphoglucose Isomerase (E. C. 5.3.1.9) 1318 17.3.3 Triosephosphate Isomerase (E. C. 5.3.1.1) 1320 17.3.4 L-Rhamnose Isomerase (E. C. 5.3.1.14) 1321 17.3.5 L-Fucose Isomerase (E. C. 5.3.1.3) 1323 17.3.G N-Acetyl-D-glucosamine 2-Epimerase 1324 17.3.7 Maleate cis-trans Isomerase (E. C. 5.2.1.1) 1324 17.3.8 Unsaturated Fatty Acid cis-trans Isomerase 1325 17.4 Conclusion 1326 References 1326 18

Introductionand Removal of Protecting Groups 1333 Dieter Kadereit, Reinhard Reents, Duraiswamy A. Feyaraj, and Herbert Waldmann

18.1 18.2 18.2.1 18.2.2 18.2.3 18.2.4 18.2.5 18.3

Introduction 1333 Protection of Amino Groups 1334 N-Terminal Protection of Peptides 1334 Enzyme-labile Urethane Protecting Groups 1338 Protection of the Side Chain Amino Group of Lysine 1341 Protection of Amino Groups in P-Lactam Chemistry 1341 Protection of Amino Groups of Nucleobases 1343 Protection of Thiol Groups 1343

Contents

18.3.1 18.4 18.4.1 18.4.2 18.5 18.5.1 18.5.2 18.5.3 18.5.4 18.5.5 18.5.6 18.5.7 18.5.8 18.6 18.6.1 18.6.2 18.7

19

19.1 19.2 19.3 19.3.1 19.3.1.1 19.3.1.2 19.3.1.3 19.3.2 19.3.2.1 19.3.2.2 19.3.2.3 19.3.2.4

Protection of the Side Chain Thiol Group of Cysteine 1343 Protection of Casboxy Groups 1344 C-Terminal Protection of Peptides 1344 Protection of the Side Chain Groups of Glutamic and Aspartic Acid 1352 Protection of Hydroxy Groups 1353 Protection of Monosaccharides 1354 Deprotection of Monosaccharides 1369 Di- and Oligosaccharides 1378 Nucleosides 1350 Further Aglycon Glycosides 1383 Polyhydroxylated Alkaloids 1386 Steroids 1388 Phenolic Hydroqq Groups 1390 Biocatalysis in Pclymer Supported Synthesis: Enzyme-labile Linker Groups 1392 Endo-linkers 13.93 Exo-linkers 1402 Outlook 1408 References 1400

Replacing Chemical Steps by Biotransformations: Industrial Application and Processes Using Biocatalysis Andreas Liese

1419

Introduction 1419 Types and Handling of Biocatalysts 1420 Examples 1421 Reduction Reactions Catalyzed by Oxidoreductases (E. C. 1) 1422 Ketone Reduction Using Whole Cells of Neurospora crassa (E.C. 1.1.1.1)~1422 Ketoester Reduction Using Cell Extract of Acinetobacter calcoaceticus (E.C. 1.1.1.1) 1423 Enantioselective Reduction with Whole Cells of Candida sorbophila (E.C. l.l.X.X) 1424 Oxidation Reactions Catalyzed by Oxidoreductases (E. C. 1) 1425 Alcohol Oxidation Using Whole Cells of Gluconobacter suboxydans (E.C. 1.1.99.21) 1425 Oxidative Deamin ation Catalyzed by Immobilized D-AminO Acid Oxidase from T'gonopsis variabilis (E. C. 1.4.3.3) 1426 Kinetic Resolution by Oxidation of Primary Alcohols Catalyzed by Whole Cells from Rhodococcus erythropolis (E. C. l.X.X.X) 1427 Hydroxylation of Nicotinic Acid (Niacin) Catalyzed by Whole Cells of Achrornobacter xylosoxidans (E. C. 1.5.1.13) 1428

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I

Contents

19.3.2.5 Reduction of Hydrogen Peroxide Concentration by Catalase (E.C. 1.11.1.6) 1428 19.3.3 Hydrolybc Cleavage and Formation of C-0 Bonds by Hydrolases (E.C. 3) 1430 19.3.3.1 Kinetic Resolution of Glycidic Acid Methyl Ester by Lipase from Serratia rnarcescens (E.C. 3.1.1.3) 1430 19.3.3.2 Kinetic Resolution of Diester by Protease Subtikin Carlsberg from Bacillus sp. (E. C. 3.4.21.62) 1431 19.3.3.3 Kinetic Resolution of Pantolactones and Derivatives thereof by a Lactonase from Fusariurn oxysporurn (E. C. 3.1.1.25) 1433 19.3.3.4 Hydrolysis of Starch to Glucose by Action of Two Wnzymes: a-Amylase (E. C. 3.2.1.1) and Amyloglucosidase (E. C. 3.2.1.3) 1433 19.3.4 Formation or Hydrolyhc Cleavage of C-N Bonds by Hydrolases (E.C. 3) 1435 19.3.4.1 Enantioselective Acylation of Racemic Amines Catalyzed by Lipase from Burkholderiaplantarii (E. C. 3.1.1.3) 1435 19.3.4.2 7-AminocephalosporanicAcid Formation by Amide Hydrolysis Catalyzed by Glutaryl Amidase (E. C. 3.1.1.41) 1436 19.3.4.3 Penicillin G Hydrolysis by Penicillin Amidase from Escherichia coli (E.C. 3.5.1.11) 1438 19.3.4.4 Kinetic Resolution of a-Amino Acid Amides Catalyzed by Aminopeptidase from Pseudornonasputida (E. C. 3.4.1.11) 1439 19.3.4.5 Production of L-Methionineby Kinetic Resolution with Aminoacylase of Aspergillus oryzae (E. C. 3.5.1.14) 1441 19.3.4.6 Production of D-p-Hydroxyphenyl Glycine by Dynamic Resolution with Hydantoinase from Bacillus brevis (E. C. 3.5.2.2) 1441 19.3.4.7 Dynamic Resolution of a-Amino-E-caprolactamby the Action of Lactamase (E.C. 3.5.2.11) and Racemase (E. C. 5.1.1.15) 1442 19.3.4.8 Synthesis of P-Lactam Antibiotics Catalyzed by Penicillin Acylase (E.C. 3.5.1.11) 1444 19.3.4.9 Synthesis of Azetidinone P-Lactam Derivatives Catalyzed by Penicillin Acylase (E.C. 3.5.1.11) 1444 19.3.4.10 Enantioselective Synthesis of an Aspartame Precursor with Thermolysin from Bacillus proteolicus (E. C. 3.4.24.27) 1446 19.3.4.11 Hydrolysis of Heterocyclic Nitrile by Nitrilase from Agrobacteriurn sp. (E.C. 3.5.5.1) 1447 19.3.5 Formation of C-0 Bonds by Lyases 1447 19.3.5.1 Synthesis of Carnitine Catalyzed by Carnitine Dehydratase in Whole Cells (E.C. 4.2.1.89) 1447 19.3.6 Formation of C-N Bonds by Lyases (E. C. 4) 1448 19.3.6.1 Synthesis of L-Dopa Catalyzed by Tyrosine Phenol Lyase from Enuinia herbicola (E. C. 4.1.99.2) 1448 19.3.6.2 Synthesis of 5-Cyano Valeramide by Nitrile Hydratase from Pseudornonas chlororaphis B23 (E. C. 4.2.1.84) 1449

Contents

19.3.6.3 Synthesis of the Commodity Chemical Acrylamide Catalyzed by Nitrile Hydratase from Rhodococcus rodochrous (E. C. 4.2.1.84) 1450 19.3.6.4 Synthesis of Nicotinamide Catalyzed by Nitrile Hydratase from Rhodococcus rodcchrous (E. C. 4.2.1.84) 1451 19.3.7 Epimerase 1452 19.3.7.1 Epimerization of Glucosamine Catalyzed by Epimerase from E. coli (E. C. 5.1.3.8) 1452 19.4 Some Misconceptions about Industrial Biotransformations 1453 19.5 Outlook 1454 References 1454 20

Tabular Survey ofCommercially Available Enzymes Peter Rasor

Index

1519

1461

P'

List of Contributors Carabed Antranikian

Manfred Biselli

Technische Universitat HarnburgHarburg Institut fur Biotechnologie KasernestraBe 12 21073 Hamburg Germany

Fachhochschule Aachen Abteilung Julich Labor fur Zellkulturtechnik Ginstenveg 1 52428 Julich Germany Andreas 5. Bommarius

Frances H. Arnold

California Institute of Technology Department of Chemical Engineering MC 210-41 Pasadena CA 91 125 USA

School of Chemical Engineering Georgia Institute of Technology 315 Ferst Drive Atlanta, GA 30332-0363 USA Bruno Buhler

Haruyuki Atomi

Department of Synthetic Chemistry and Biological Chemistry Graduate School of Engineering Kyoto University Yoshida, Sakyo-ku Kyoto, 606-8501 Japan

Institut f i r Biotechnologie ETH Honggerberg, HPT 8093 Zurich Switzerland Nobuyoshi Esaki

Institute for Chemical Research Kyoto University Uji

Kyoto-fu 611 Japan Constanzo Bertoldo

Technische Universitat HannburgHarburg Institut fur Biotechnologie KasernestraBe 12 21073 Hamburg Germany

Kurt Faber

Department of Chemistry Organic and Bioorganic Chemistry University of Graz Heinrichstrasse 28 8010 Graz Austria

XXXIV

I

List ofcontributors

Duraiswamy A. Feyaraj

Peter Halling

Max-Planck-Institutfur Molekulare Physiologie Abteilung Chemische Biologie Otto-Hahn-StraBe11 44227 Dortmund Germany

Department of Chemistry University of Strathclyde Glasgow G1 1XL United Kingdom

Martin H. Fechter

Institut f i r Organische Chemie Technische Universitat Graz Stremayrgasse 16 8010-Graz Austria Sabine Flitsch

Department of Chemistry The University of Edinburgh West Mains Road The King’s Building Edinburgh EH9 3J J United Kingdom Hans-Joachim Gais

Institut fur Organische Chemie RWTH Aachen Professor-Pirlet-StraBe1 52056 Aachen Germany Herfried Criengl

Institut fur Organische Chemie Technische Universitat Graz Stremayrgasse 16 8010-Graz Austria

Yoshihiko Hirose

Amano Enzyme Inc. Gifu R & D Center 4-179-35,Sue, Kakamighara Gifu 509-0108 Japan Kay Hofmann

Bioinformatics Group MEMOREC Stoffel GmbH Stockheimer Weg 1 50829 Koln Germany Frank Hollmann

Institut fur Biotechnologie ETH Honggerberg, HPT CH-8093 Zurich Switzerland Tadayuki lmanaka

Department of Synthetic Chemistry and Biological Chemistry Graduate School of Engineering Kyoto University Yoshida-Honmachi,Sakyo-ku Kyoto 606-8501 Japan Hans-Dieterjakubke

Cideon Crogan

Department of Chemistry The University of Edinburgh West Mains Road The King’s Building Edinburgh EH9 3JJ United Kingdom

Fakultat fur Biowissenschaften, Psychologie und Pharmazie Institut fur Biochemie Universitat Leipzig TalstraBe 3 04103 Leipzig Germany

List ofcontributors

Dieter Kadereit

Tomoko Matsuda

Johann-StrauB-StraBe18a 65779 Kelkheim Germany

Department of Materials Chemistry Faculty of Science and Technology Ryukoku University Otsu Shiga 520-2194 Japan

Udo Kragl

Universitat Rostock Fachbereich Chemie BuchbinderstraBe 9 18051 Rostock Germany

Oliver May

Degussa-Huh AG Rodenbacher Chaussee 4 63457 Hanau Germany

Maria-Regina Kula

Institut fur Enzymtechnologie Heinrich-Heine Universitai Dusseldorf im Forschungszentrum Jiilich Stetternicher Forst 52428 Jiilich Germany

Kaoru Nakamura

Tatsuo Kurihara

Claudia Neri

Institute for Chemical Research Kyoto University Uji Kyoto-Fu 611 lapan

Institute for Chemical Research Kyoto University Uji Kyoto 611-0011 Japan

School of Chemistry University of Bath Claverton Down Bath BA2 7AY United Kingdom Romano A. Orru

lamesJ. Lalonde

USA

Division of Chemistry Bio-organicChemistry Vrije University Amsterdam De Boelelaan 1083 1081 H V Amsterdam The Netherlands

Andreas Liese

RebekkaJ. Parker

Forschungszentrum Jiilich GmbH I BT Leo-Brandt-Strage D-52428 Jiilich Germany

School of Chemistry University of Bath Claverton Down Bath BA2 7AY United Kingdom

Altus Biologics Inc. 625 Putnam Avenue Cambridge MA 02139-4807

I-

xxxvt

I

List ofcontributors

Markus Pietzsch

Birgit Schulze

Institute for Bioprocess Engineering University of Stuttgart Department of Microbial Physiology Allmandring 31 70569 Stuttgart Germany

DSM Food Specialties Nutritional Ingredients P.O. Box 1 2600 MA Delft The Netherlands

Reinhard Reents

Max-Planck-Institutfur Molekulare Physiologie Abteilung Chemische Biologie Otto-Hahn-Strage 11 44227 Dortmund Germany

Christoph Syldatk

Institute for Bioprocess Engineering University of Stuttgart Department of Microbial Physiology Allmandring 31 70569 Stuttgart Germany

Peter Rasor

Fritz Theil

Industrial Biochemicals Business BB-PS, Roche Molecular Biochemicals Roche Diagnostic GmbH Nonnenwald 2 82372 Penzberg Germany

ASCA Angewandte Synthesechemie Adlershof GmbH Richard-Willstatter-Strage 12 12489 Berlin Germany

I. David Rozzell

Karl-Heinz van Pie

School of Chemical Engineering Georgia Institute of Technology 315 Ferst Drive Atlanta, GA 30332-0363 USA

Institut fur Biochemie Technische Universitat Dresden Mommsenstrage 13 01062 Dresden Germany

Kenji Soda

Christopher A. Voigt

Faculty of Engineering Kansai University Yamate-cho Suita Osaka-Fu 564 Japan

California Institute of Technology MC 210-41 Pasadena CA 91125 USA

Erik de Vroom Andreas Schmid

Institut f i r Biotechnologie ETH Honggerberg, HPT CH-8093 Zurich Switzerland

DSM Food Specialties Nutritional Ingredients P. 0. Box 1 2600 MA Delft The Netherlands

Herbert Waldmann

Jonathan M.J. Williams

Max-Planck-Institutfiir Mcllekulare Physiologie Abteilung Chemische Biologie Otto-Hahn-StraBe11 44227 Dortmund Germany

School of Chemistry University of Bath Claverton Down Bath BA2 7AY United Kingdom Chi-Huey Wong

Forschungszentrum Julich GmbH Institut fur Biotechnologie 52425 Julich Germany

Department of Chemistry The Scripps Research Institute 10550 Torrey Pines Road La Jolla, CA 92037 USA

George M. Whitesides

Marcel Wubbolts

Department of Chemistry Harvard University 12 Oxford Street Cambridge, MA 0213&2902 USA

Manager Research & Development DSM Biotech GmbH Karl-Heinz-Beckurts-Strage13 52428 Julich Germany

Christian Wandrey

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

1 Introduction Maria-Regina Kula

1.1 Enzymes as Catalysts

Enzymes are the catalysts evolved in nature to achieve the speed and coordination of a multitude of chemical reaction necessary to develop and maintain life. Chemical reactions are far too slow to be effective under the conditions prevalent in normal living systems - aqueous environments with neutral pH values and temperatures between 20 and 40 "C. Even catalysts developed in the chemical industry fall short; enzymes in comparison achieve up to lo7 - fold faster reaction rates. Mankind has utilized enzymes empirically since ancient times for the conservation or production of food, e. g. in cheese making or brewing. A historical background is given in Table 1-1.The catalytic properties of enzymes were recognized long before their chemical nature was known. We stil~use acceleration of reaction rate to search for unknown enzymes as well as to measure and quantify enzyme activity. As catalysts - true to the definition familiar in chemistry - enzymes alter the rate at which a thermodynamic equilibrium is reached, but do not change that equilibrium. This implies that enzymes work reversibly. The acceleration in reaction rate is achieved by lowering the activation energy of the overall process as shown schematically in Fig. 1-1. Enzymes bind their substrates by multiple non-covalent interactions on a specific surface. This way, a micro-heterogenization occurs and the local concentration of substrates is increased relative to the bulk solution. In addition, the chemical potential of specific groups may be drastically changed temporarily compared to aqueous solutions by the Iexclusion of water in the reactive site upon binding of substrate. Both aspects contribute to the observed phenomenon of high acceleration in reaction rate; some examples are presented in Table 1-2. Enzymes often bind the substrate in the transition state better than in the ground state, which lowers the activation energy. Since the pioneering work of Buchner (1897), it has been known that enzymes do not require the environmept of a living cell to be active. This opened the way to many applications in food technology, in the production of leather, textiles and paper, in

2

I

1 Introduction Table 1-1.

Brief history of enzymes and their applications.

-

BC

Chymosin from the stomach of young cattle, sheeps and goats was used for cheese production in many ancient cultures for approximately 7000 years.

1783

Hydrolysis of meat by gastric juice demonstrated.

Spallazani

1814

Starch degradation and sugar production by malted barley observed.

Kirchhoff

1833

The active principle of malt is called diastase and its application to industrial art described.

Payen and Persoz

1846

Invertase activity observed.

Dubonfout

1867

The term enzymes is coined to describe catalytic activity not bound to living cells (unorganised ferments). The name is extended later also to intracellular catalysts (organised ferments as defined by Pasteur).

Kiihne

1893

Definition of a catalyst including enzymes is given.

Ostwald

1894

Enzyme stereospecificity anticipated.

E. Fischer

1894

“Taka diastase” produced commercially with Aspergillus olyzae by surface culture

Takamine

1897

The conversion of glucose to ethanol demonstrated by a cell free extract from yeast.

Buchner

1906

Preparative separation of L-leucinefrom the racemate carried out by hydrolysis of the propyl ester with liver extracts.

Warburg

1908

Synthesis of optically active cyanohydrins described, using D-oxynirerilasefrom almonds as catalyst.

Rosenberg

1908

Application of pancratic enzymes in the leather industry for the bating of hides.

Rohm

1911-1913

Glucoside synthesis in the presence of high concentration of ethanol or acetone described.

Bourquelot, Bride1 and Verdon

1913- 1915 Application of pancreatic enzymes to clean laundry introduced, first commercial product sold to the public: Burnus.

Rohm

1916

Immobilization of invertase on charcoal demonstrated with retention of activity.

Nelson and Griffin

1926

Urease from Jack beans crystallized.

Sumner

1936

Enzymatic ester synthesis improved using pancreatic lipase in the presence of benzene.

S P

1953

The first primary sequence of a protein (Insulin) established, proving the chemical identity of proteins.

Sanger

1960

Cultivation of Bacillus lichenijomis in submerged culture started for protease production on a large scale.

NOVO

after 1980

Application of genetic engineering techniques to improve enzyme production and to alter enzyme properties by protein engineering and evolutionary design.

many

1.1 Enzymes as Catalysts 13

\

I

\

: C

[G

a a a

II

L

? 71

C

\ \ \\

-'i\ ,I

II

//

----

I

\

C

+

U

\\ \

ES

E+P

Course of r e a c t i o n Figure 1-1. Free-energy profile for the course o f an enzyme-catalyzed reaction involving the formation o f enzyme-substrate (ES) and enzyme-product (EP) complexes. The reaction pathway goes through the transition states TS,, TS,2, and TS,3 with standard free energy o f activation ACc. The rate limiting step would be the conversion o f ES into EP. The schematic profile for the uncatalyzed reaction is shown as the dashed line. The catalytic effect is due t o the lowering o f t h e standard free energy o f activation from A C u t o ACc and is not governed by the difference in free energy between S and P.

diagnostics and food analysis and, last but not least, in the production of chemicals by biotransformations. One or more of the following reasons could make enzymes the catalysts of choice: - a highly selective operation in complex mictures, - stereo- and regiospecificity of conversions, - absence of side reactions leading to simpler separation processes and higher

yields, or - savings in energy and waste treatment cost owing to mild reaction conditions.

Enzymes have limitations, SI; does any other highly specialized catalyst. Most notable is one consequence arising from the selectivity of enzymes with regard to the substrates bound and the type of reaction catalyzed. The price for such selectivity is that it may be difficult to satisfy the requirement for many special enzymes to cover the diversity of chemical rextions desired in organic chemistry. The enzyme needed in a specific case may not be readily available. However, there are new enzymes discovered all the time and an increasing number can be obtained commercially. Other limitations with regard to reaction conditions, pH and temperature tol-

4

I

1 Introduction

Relative rates of enzyme catalyzed and non-catalyzed reactions under conditions optimal for enzyme reaction.

Table 1-2.

Enzyme

Reaction

Ratioa

9

Triose phosphate Isomerase

@

Y

H

3x 108-109 %===

@

T

OH

O

H

0

105-1010

Urease

1014

0 H2NKNH2

C02+2NH3

Hexokinaseb

> 8 x 10" M

glucose + ATP

+

glucose G @ + ADP

Alcoholb

>2x1O1OM

dehydrogenase ethanol + NADa

-+

acetaldehyde + NADH + Ha

a ratio of enzyme catalyzed to non-catalyzed rate

b bimolecular reactions

@ = Phosphate

erated by enzymes are to some extent predictable by their chemical nature. In this Introduction, general aspects of enzyme structure, function and nomenclature will be discussed to guide the reader with little biochemical background into the field of enzyme application in organic chemistry.

1.2 Enzyme Structure and Function

All enzymes are proteins, with the exception of the recently discovered ribozymes. Ribozymes are special ribonucleic acids performing catalytic functions in the processing of RNA which will not be considered here. Proteins are polar macromolecules with molecular mass in the range 104-106. They are linear polymers, defined by the sequence of amino acids, which are linked by peptide bonds:

0 H2N

O0

The individual properties of a protein depend on the chemical nature of the side chains depicted as R in Scheme 1-1.In protein biosynthesis, 20 amino acids are

1.2 Enzyme Structure and Function 15

Table 1-3.

Amino acids for protein biosynthesis.

Name

Symbol

Glycine

GlY (G)

Alanine

Ala (A)

Valine

Val (V)

Leucine

Leu (L)

Isoleucine

Ile (I)

Serine

Ser (S)

Threonine

Thr (TI

Cysteine

CYS (C)

Methionine

Met (M)

Proline

Pro (P)

Phenylalanine

Phe (F)

Tyrosine

Tyr (Y)

Tiyptophane

Trp (W)

Asparagine

Asn (N)

Glutamine

Gln (Q)

Aspartate

ASP (D)

pK, of ionizing side chain"

Structure

/COOH I

9.1-9.5

9.7

HonCOOH 4.5

0

NH,

6

I

I Introduction Table 1-3.

(cont.).

Name

Symbol

Glutamate

Glu (E)

pK, of ionizing side chain"

Structure

4.6

"o&cooe NH2

Lysine

LYS

10.4

(K) H2N-cooH NH2

a-amino group

6.8-7.9

a-carboxylgroup

3.5-4.3

a The p& values depend on temperature, ionic strength and, especially on the microenvirronment of the

ionizable group

condensed according to information coded in the corresponding genes. The coded amino acids are summarized in Table 1-3. Some modified amino acids, for example 4-hydroxyproline, 5-hydroxylysine,y-carboxyglutamate,0-phosphorylated serine, or N-glycosylated asparagine are also found in proteins, usually in minor amounts, resulting from post-translational modifications. These modifications are important for the structure and function or the regulation of the activity of certain proteins. For 20 building blocks and a random sequence, the number of possible variations in the primary structure is 20"; for a protein of average size of 33 000 Da 6 300 amino acids, lo3" possibilities exist. The number is far beyond our perception, the known cosmic space is not large enough to accommodate a single copy of each variant. To generate a specific surface as part of the active center of an enzyme, the protein chain has to fold. From the observed length and angle of the C=O and C-N bonds in peptides it can be deduced that the peptide bond possesses partial double bond character, resulting in a planar arrangement as shown schematically in Fig. 1-2. Movement of the planes against each other occurs around the a-carbon, which serves as a joint (Fig. 1-3). Rotation around the C-C and C-N bond is restricted because of steric influences of the side chain R. By this feature of the peptide bond, two structural arrangements of a polypeptide become energetically favored: the a-helix and the ppleated sheet, which are further stabilized by hydrogen bonds between the peptide backbone (Fig. 1-4). Helices and pleated sheets are commonly found in proteins; these secondary structure elements are linked by p-turns or loops to build a domain or a subunit. This level of organization is called the tertiary structure of proteins, while the assembly of subunits into homo- or hetero-oligomers or multi-component systems is called quaternary structure. The hierarchy of structures is depicted in Fig. 1-5.

7.2 Enzyme Structure and Function 17

Figure 1-2. Special features ofthe peptide bond. Dimensions are given in A and represent average values from X-ray analyses. The peptide group has a rigid and planar structure.

The mechanism determining the folding of a given protein is presently the topic of intense research. For this discussion, it is sufficient to state that there exists a unique tertiary/quaternary structure for each native protein chain, determined as an energy minimum in aqueous solution. The information to reach this structure is thought to be encoded in the primary sequence in a way not yet understood completely. The

8

I

7 Introduction

7.2 Enzyme Structure and Function 19

Figure 1-5. Hierarchy o f protein structures. The three-dimensional structure of an enzyme is the result ofdifferent levels of folding and interactions of protein chains proceeding in ordered fashion from the primary structure after protein biosynthesis.

Primary structure: sequence of acid, e, g, - Gly - Glu - Ser - Lys - Phe secondary structure: a-helix

p-sheet

tertiary structure:

Single domain or multi domain folding

Quarternary structure: protein aggregate of like or unlike subunits

folded structure of a protein is stabilized by a network of non-covalent interactions, most notably hydrogen bonds, hydrophobic and van der Waals interactions, and ionic bonds. In the folding process, hydrophobic side chains of amino acids are

10

I preferentially oriented towards the interior of the molecule, thereby diminishing the I Introduction

surface area in contact with water and minimizing the free energy. Polar groups are preferentially oriented towards the surface interacting with water. In the compact inner core of a protein, water is virtually excluded or present as single HzO molecules (!) in defined places. The folding process generates a unique threedimensional surface of a protein defined in molecular dimensions by the specific side chains and the polypeptide backbone. Substrates and their transition states are also bound by multiple noncovalent interactions with such a surface. Since the strength of all these noncovalent bonds is strongly dependent on distance and angle of interaction, a highly selective binding may result. By a three-point attachment even discrimination between two enantiomers is possible. Steric constraints may also contribute to differentiation between similar structures during binding. The specific binding site of enzymes often is found in a cleft on an irregularly shaped surface. Substrate recognition is a dynamic process not only with regard to association and dissociation of the substrate; it may also involve movements of the polypeptide chain in response to the binding. An example of the latter is shown in Fig. 1-6. Carboxypeptidase A hydrolyzes proteins sequentially starting from the free carboxyl terminus. The enzyme preferentially cleaves hydrophobic amino acids. Already in 1967 the three-dimensional structure had been determined with high resolution by W. Lipscomb and his group. In the meantime much is known about the catalybc mechanism. The essential features are discussed here briefly to improve the understanding of how enzymes work. Two aspects of enzymatic catalysis can be illustrated by this example: 1. Substrate binding may be accompanied by changes in enzyme structure. 2. Substrate binding induces subtle but important shifts in electron distribution in the substrate, making it more susceptible to certain reactions (here hydrolysis). In Fig. 1-6,the tertiary structure of the free carboxypeptidaseA is presented as well as an enzyme-substrate complex with glycyl-tyrosine. Changes in the enzyme structure are most evident by looking at the position of tyrosine-248. The phenolic hydroxyl group of the side chain moves from a position near the surface of the enzyme 12 A toward the interior. A distance of 12 A represents about a quarter of the diameter of carboxypeptidase A. Tyrosine-248then covers the bound glycyltyrosine (Fig. 1-6 B) and the phenolic hydroxyl group is oriented toward the terminal carboxyl group of the substrate. The movement of the tyrosine-248 side chain is possible by rotation of the C - C bond at the j3 carbon. As a consequence of the rotation, the binding site of carboxypeptidase A is shielded from bulk water. Closer inspection of Fig. 1-6 A and B shows that the guanido group of arginine-145 as well as glutamate270 also move about 2 A upon substrate binding. Both residues are involved in the catalytic step. The second important point is the perturbation of the electron distribution in the substrate by the essential Zn” and specific side chains in the enzyme. During the binding process the substrate is oriented first by an electrostatic interaction of the carboxylate group with the positively charged arginine-145;in addition, tyrosine-248 forms a hydrogen bond with the amide nitrogen of the terminal peptide bond. The carbonyl group becomes coordinated to the Zn” displacing water as a ligand.

7.2 Enzyme Structure and Function Figure 1-6. The structure of carboxypeptidase A changes dynamically upon substrate binding. (A) Enzyme alone, (B) enzyme complex with glycyl-tyrosine. Tyrosine 248 moves 12 A after binding of substrate. Hydrolysis results as a concerted action o f ZnZ+, Glu, Tyr, and Arg side chains towards the carbonyl and nitrogen group i n the susceptible peptide bond

(C).

B

12

I

7 Introduction

The hydrophobic group in the substrate (tyrosine in the example illustrated here) is bound into an unpolar, large cleft by hydrophobic forces replacing at least 4 water molecules upon binding and inducing the movement of tyrosine-248 discussed above. The unpolar lining and the size of the cleft explains the preference of carboxypeptidase A for bulky, hydrophobic side chains of the terminal amino acid. The free amino group of glycyltyrosine is hydrogen bonded through a water molecule to glutamate-270. This bonding of glutamate is thought to slow down dramatically the hydrolysis rate of glycyltyrosine and related dipeptides (and make possible the X-ray analysis of the complex). Such a hydrogen bond is not found in productive enzyme-substrate complexes involving oligopeptides or proteins. The carboxylate group of glutamate-270 is thought to attack the carbon in the carbonyl bond of the substrate leading to a mixed anhydride. The carbonyl bond is already polarized by the Lewis acid Zn2+, the induction of the dipole is favored by the unpolar surrounding of the Zn2+ion and the tetrahedral intermediate is stabilized by the positive charge of nearby arginine-127. The hydrolysis of the peptide bond is completed by transfer of a proton form water to the nitrogen, releasing the Cterminal amino acid. Substrate binding in a defined manner is a prerequisite for enzyme catalysis. It exposes a chemical compound long enough to a unique chemical potential built into the system, which defines the type of reaction that will proceed, for example, hydrolysis, oxidation/reduction, or C - C bond formation. The mechanism most often is the same as that known from solution chemistry, for example, acid-base catalysis. The close proximity of reactants and the precise orientation, together with the effect of microheterogenization discussed above, lead to the outstanding performance of an enzyme as catalyst (examplesare given in Table 1-2). Often, transient covalent bonds are formed between substrate and enzyme or coenzyme (see below) during a catalyhc cycle. Serine, cysteine,histidine, lysine, aspartate or glutamate may donate en electron pair to a substrate, forming a covalent linkage as shown in Fig. 1-7 for the well-known charge-relay system in serine proteases. The highly reactive intermediates formed may be attacked by water or a second substrate to yield the characteristic products of the reaction.

1.3 Cofactors and Coenzymes

The chemical potential of side chains found in amino acids is limited; for example, there are no efficient electron acceptors. Therefore, enzyme catalysis incorporates if necessary additional chemical potential by specific metal ions, for example, Zn2+(see Fig. 1-6), Fe2+ Co2+, Cu2+ and others Examples are shown in Fig. 1-8 for the coordination of the transition metal ions in protein structures. Besides metal ions, cofactors or coenzymes serve to activate groups and participate in the catalytic process. A summary of cofactors and coenzymes is given in Table 1-4;the relation to vitamins is quite apparent. Chemical structures are presented in Table 1-5. Coenzymes and cofactors may act by nucleophilic or electrophilic attack on the sub-

7.3 Cofactors and Coenzymes 113

Tetrahedral transition state

Substrate Ser 195 -CH2-O

R'

-O\

O=C'

/

I

I

H.

-CH2-0

R-N--H

d

Acyl-enzyme intermediate

,R' C\ N-H / RH

-

0% C- R'

-CHz-O

I

R-N-H I

H

HC\IN-!" HC,lN +- i H N-C, H CH2

;-c,cI-lz

:

His 57

I

N-C, H

0I

o@-

Asp102 0."-

:

I

CH2

I

0I

O,C\ Acyl-enzyme intermediate

Tetrahedral transition state

0% C-R' -CH2-0 H , 0

-O\ -CH2-0

I

U ..

-HIS 5 / *

,.

-

I/

N-C, H

7H2 I

0I

,R' C\ 0-H H I

N-CH "C:'

/

Acid component of the substrate

N-CH

II lN-iH -

HC\\ + N-C, H

: - -u, /u C

I

CH2

HC:' N-C, H

:

I

CHz

I

nU

o'c\

I

Asp 102 O S c \ Figure 1-7. The catalytic triad in serine proteases. The reactive serine forms an acyl enzyme as a covalent intermediate during the proteolytic cleavage o f a peptide bond. During substrate binding a proton is transferred from serine 195 t o histidine 57, and the positive charge o f t h e iniidazole ring is stabilized by interaction with the carboxylate side chain o f aspartic acid 102. The numbering corresponds t o the structure o f chymotrypsin.

strate(s) to initiate a reaction. Cofactors are tightly (covalently)bound to the protein and may undergo cyclic reactions during the catalytic process but will return to the ground state at the end. Ifoxygen is the terminal electron acceptor in FAD, FMN or NAD(P)+ linked reactions,, these cofactors require a second reaction with the cosubstrate oxygen to regenerate the active form. In the older literature cofactors sometimes are called "prostetic groups". Coenzymes are bound in an association/ dissociation equilibrium to enzymes and have to be present in sufficient concentration to obtain maximal enzymatic activity. Some are regenerated in the catalytic cycle

14

I

7 Introduction H

H

C"rl 1

3

HisxJ H

@ 7

,

Typical co-ordination complexes o f transition metal ions in proteins. 1: M may be Fe", as i n rubredoxin, or Zn" as in aspartate transcarbamylase and alcohol dehydrogenase, 2: carboxypeptidase A, 3: carbonic anhydrase, 4: liver alcohol dehydrogenase, 5: azurin, 6: heme group, L is His and L'either H i s or Met i n cytochromes, 7: deoxy-heme group in hemoglobin and myoglobin, 8: oxyform o f 7, 9: superoxide dismutase. Figure 1-8.

R,6CooH 0

Leucine-Dehvdroaenase

-/-E.C. 1.4.1. /

Formate Dehydrogenase N A D H regeneration using formate dehydrogenase (FDH) in a coupled reaction with leucine dehydrogenase (Leu DH). Figure 1-9.

CO2

HCOONH4 ammonium formate

7.3 Cofactors and Coenzymes Table 1-4.

Cofactors and coerizymes.

Compound"

1:unction

Relation to vitamins

NAD+/NADH+ H'

redox reactions and hydrogen transfer

vitamin PP, (niacin)

NADP'/NADPH + H'

redox reactions and hydrogen transfer

vitamin PP, (niacin)

FAD

redox reactions and hydrogen transfer

vitamin Bz. (riboflavin)

FMN

redox reactions and hydrogen transfer

Haem

iiansfer of electrons

vitamin B2, (riboflavin) -

Coenzyme A

transfer of acyl groups

pantothenic acid

ATP

metabolic energy, phosphate-,pyrophosphatetransfer, adenylation

Pyridoxal phosphate

transamination, amino acid decarboxylation

vitamin Bg, (pyridoxine)

(PLP)

Thiamine pyrophosphate (TW Biotin

decarboxylation, transfer of Cz units

vitamin B1, (thiamine)

transfer of COZ

biotin

Tetrahydrofolicacid

transfer of C1 groups

S-Adenosylmethionine

methylation

folic acid -

Adenosyl-cobalamine

isomerisation (hydrogen-shift)

vitamin BIZ

Methyl-cobalamine

methylation

cyano-cobalamine

(SAM)

a The structure of the various compounds is summarized in Table 5.

while bound to the enzyme, for example, pyridoxal phosphate or thiamine pyrophosphate, so that catalytic amounts are sufficient to sustain the reaction. Others require one or more separate reactions with cosubstrates other than oxygen to regenerate the starting material. This holds true for example for NAD(P)', NAD(P)H, SAM, coenzyme A, ATP and other nucleotide triphosphates. In such instances, the coenzyme is consumed i n stoichiometric relation to product formation. This relation may render enzymatic synthesis quite expensive unless efficient coenzyme regeneration cycles can be devised. In situ regeneration processes have been successfully developed i n recent years, especially for the nicotinamide nucleotides. The stoichiometric relation with product formation is shifted from the expensive coenzyme to h e conversion of a cheap cosubstrate such as formate, as shown schematically i n Fig. 9. A detailed discussion of coenzyme regeneration is found i n another chaptei:

16

I

1 Introduction

Table 1-5.

Chemical structures of cofactors and coenzymes.

Nicotinamide nucleotides NAD' and NADP+and their reduced forms are involved in many dehydrogenase reactions within the cell. They are water-soluble, and are usually free to diffuse away from the enzyme, after conversion to the oxidized or reduced form to take part in another dehydrogenase reaction catalyzed by another enzyme.

R = H: N A D ~ R = P O ~ HNADP : @ = Phosphate (PO3H2-group)

Flavin nucleotides FAD is the coenzyme of a class dehydrogenases calledpauoproteins.The flavin moiety of the molecule is derived from riboflavin (vitamin Bz). Reduction of FAD involves the two unsubstituted N atoms of the isoalloxazine structure.

7.8-Dimethylisoalloxatine

'

CH, I HC-OH I

Hy-OH HC-OH ,I

5

OH I

CH2-0-P-OR It

0

Flavin adenine dinucleotide FAD

R = -@-cH

d OH OH

Flavin mononucleotide FMN R=H

1.3 Cofactors and Coenzymes 117

(cont.).

Table 1-5.

The electron transport chain Enzymes in the electron transport chain split hydrogens into H+ and ec. The electrons are then camed by enzymes called cytochrornes a, b, c, d. These enzymes are able to accept an alectron and then pass it on to another cytochrorne. The iron atom is bound, within the haern a, b, c, d group, to a porphyrin coenzyme identical with that found in haernoglobin, with the difference that in the cytochrornes the iron undergoes oxidation and reduction.

Haem b

I

82 Cytochrome c with haem c

I

Cytochrome b Fe'@

Cytochrome b Fe

'

@

1

18

I

1 Introduction Table 1-5.

(cont.).

CoenzymeA (CaA-SH) Coenzyme A is a complex molecule which contains a free sulfydryl (-SH) group. This group can react with a carboxyl group to form a thioester. In acetyl CoA, the thioester linkage can activate the methyl carbon as well as the acetyl group.

--Cysteamine P-Alanine Pantoic acid

w Pantothenic acid

R =H: CoA-SH R = Acetyl: Acetyl CoA

w OH

HO-P=O I

OH

Adenosine-3'-phoshate-5'-diphosphate

Adenine nucleotides ATP, ADP and AMP are coenzymes influencing the direction of flow in metabolic pathways. In addition ATP often functions as a donor of a phosphate to other molecules in reactions catalysed by kinases.

CH*-O-

@ - @ - @-OH

1.3 Cofactors and Coenzymes Table 1-5.

(cont.).

Pyridoxal phosphate Pyridoxal phosphate, a derivativ,: of vitamin B6, acts as coenzyme in transamination and decarboxylationreactions. In a transamination reaction the aldehyde group of pyridoxal phosphate first forms a Schiff base with the amino group of the amino acid, which is then converted to keto acid. Pyridoxal phosphate is thereby converted to pyridoxamine phosphate which transfers the amino group to another keto acid to form the corresponding amino acid.

H

o

CH2M2

e -@-OH

CH&-

OH

H3C

Pyridoxal phosphate

RYC*H

RKCmH 0

NHZ

Thiamine pyrophosphate All biochemical reactions with participation of thiamine start with C-C-bond cleavage of 2-0x0 carbonyl-compound and proceed with formation of an “activatedaldehyde”, TPD catalyzes decarboxylationof a-keto acid::, oxidative decarboxylationstogether with lipoic acid, and transketolase reactions.

Biotin Biotin containing enzymes catalyze CO2-transfer reactions: these are carboxylation,transcarbox~ lysine in an ylation and decarboxylations. The carboxy group of biotin is bound to an E - N Hof enzyme protein. 0

0

R = H : D(+)-Biotin R = COOH: N-1’-Carboxybiotin

20

I

7 Introduction Table 1-5.

(cont.).

Folate coenzymes The transfer of a Cl-group like methyl, methylene, formyl or formimino often involves folic acid in one of its substituted forms 2-Amino-4-hydroxy6-methylpteridine

p-Aminobenzoic acid

L-GIu

H

0

Tetrahydrofolic acid Compound Tetrahydrofolic acid

COOH

Cl-fragment

Structure H 4-10\

H N

-

N5-formyl-

formic acid

Po-methenyl-

formic acid

N5, N"-rnethenyl-

formic acid

N5-formimino-

formic acid

N5, N"-methylene-

formaldehyde

Ns-methyl-

methanol

S-Pdenosyl-1-methionine

YHz

S-Adenosyl-L-methionineas sulfonium compound could transfer its methyl group as CH3e to nucleophile centers of substrates in biochemical reactions.

7.4 Enzyme Nomenclature Table 1-5.

(cont.).

Cobalamine Adenosyl-cobalaminecatalyzes hydrogen shifts as a special isomerisation reaction. With exception of reduction of ribonudeotides the H-shift occurs intramolecularly. Methyl-cobalamine and tetrahydrofolic acid are the coenzymes in methylating homocysteine to methionine.

CH,-CH

I

I

y

2

NH

,CONHI

\

p

2

FH2 CONH2 : Adenosyl-cobalamine

R=CH,

: Methyl-cobalarnine

R=CN

: Cyano-cobalarnine (Vitamine

BI~)

1.4

Enzyme Nomenclature

The IUB has classified enzymes into 6 main classes according to the type of reaction catalyzed:

I

21

22

I

1 Introduction

1. Oxidoreductases

2.

3.

4.

5.

6.

These catalyze oxidation/reduction reactions, transferring hydrogen, oxygen, and/or electrons, between molecules. In this important class belong dehydrogenases (hydride transfer), oxidases (electron transfer to molecular oxygen), oxygenases (oxygen transfer from molecular oxygen), and peroxidases (electron transfer to peroxide) Transferases These catalyze the transfer of groups of atoms, e. g. amino-, acetyl-, phosphoryl-, glycosyl- etc. from a donor to a suitable acceptor. Reactions covered in class 1 , 3 , or 4 are excluded. Hydrolases These catalyze the hydrolytic cleavage of bonds. Many commercially important enzymes belong to this class, e. g. proteases, amylases, acylases, lipases, and esterases. Lyases These catalyze the non-hydrolytic cleavage of, for example, C - C, C - 0 or C - N bonds by elimination reactions leaving double bonds or, in reverse, adding groups to a double bond. Examples are fumarase, aspartase, decarboxylases, dehydratases, and aldolases; many lyases are important catalysts for organic synthesis. In older literature class 4 enzymes are often called synthases, e.g. tryptophan synthase. These should not be confused with synthetases, as class 6 enzymes are sometimes called. Isomerases These catalyze isomerization and transfer reaction within one molecule. The most prominent member of this group is D-xylose ketol-isomerase, commonly known as glucose isomerase. Ligases These catalyze the covalent joining of two molecules coupled with the hydrolysis of an energy rich bond in ATP or similar triphosphates. An example is y-Lglutamyl-L-cysteine:glycine ligase (ADP-forming),also found under the name glutathion synthetase. Ligases find limited applications only for synthetic purposes.

The main classes are further subdivided into subclasses and subgroups, as in part indicated above. A complete ordering system can be found in the publications from IUB. The systematic name of an enzyme is based on the equation of the chemical reaction taking place and the type of reaction, followed by the suffix-ase. By international agreement the catalytic reaction is expressed and identified by 4 groups of numbers according to the E. C. (enzyme classification)system introduced above. For example, an enzyme converting an alcohol to an aldehyde (or ketone) using NAD as coenzyme would be classified as oxidoreductase acting on CH - OH groups using NAD+ as acceptor alcohol NAD+-oxidoreductase

main class 1 sub class 1.1 sub group 1.1.1 E.C. 1.1.1.1

7.5 Enzyme Kinetics

The last number is the serial number of an enzyme identified by the first three entries. An alcohol could be converted to similar products also using oxygen as the electron acceptor by an oxidase. oxidoreductase acting on CH - OH groups using oxygen as acceptor alcohol: oxygen-oxidoreductase

main class 1 sub class 1.1. sub group 1.1.3 E. C. 1.1.3.13

For newly isolated enzymes the nomenclature committee of IUB assigns the correct E.C. number to avoid confusion. The last edition (1992) contains 3196 entries. This code system is used in the scientific literature, textbooks and catalogues to identify an enzyme on the basis of the chemical reaction it catalyzes. For a proper description the source has to be included. Besides the systematic name IUB also lists trivial names or recommended names, the two enzymes described above being better known as alcohol dehydrogenase or alcohol oxidase, respectively. The recommended name is shorter and preferred in discussion after the catalyst has been duly identified. It should be noted that the classification is not based on the enzyme source and in general not on a single substrate. The physical properties of the individual enzyme protein may vary, for example, pH optimum, K, values, stability, substrate range etc., but the systematic name and the number code are identical as long as the same type of reaction is catalyzed. Often it is worthwhile to test enzymes from different sources for the reaction of interest to find the optimal catalyst. Numerous successful applications of enzymes are described in the following chapters. Many more opportunities exist for innovative approaches in synthetic chemistry.

1.5

Enzyme Kinetics 1.5.1 Reaction Rate and Substrate Concentration

An enzymatic reaction rnay be described by the following steps: first, binding of enzyme E and substrate IS occurs; second, while bound to the enzyme the substrate will be converted to the product P;finally, the product is released from the enzyme and free enzyme becomcs available for the next cycle. In the simple case of a one substrate reaction this can be described by the following equations [ S l + [El

5 K,-'

[ES] =+[EP] 5

[El+ [PI

Michaelis and Menten derived a mathematical description for the reaction rate of an enzymatic process from this scheme

I

23

24

I

1 Introduction a

2.0 - v,,,

([El = 2.0 u n i t -

---------- -

VI c .-

C

3

K,,

Concentration of substrate IM1

-

b

0.8 C

0 .c

0

go6 L

I/

I

0.2

0 Figure 1-10. Reaction rate as a function o f substrate concentration: a) using two different enzyme concentrations in the assay, b) comparing low and high affinity substrates o f t h e same enzyme.

with the assumption that the binding of substrate and enzyme is reversible and fast compared to product release. Equation (1) represents a hyperbolic curve, relating reaction rate with substrate concentration as shown in Fig. 1-10. The hyperbola is described by two parameters: V,, and K,. K, the so-called Michaelis constant, is defined as the substrate concentration for which the observed reaction rate is half of V,, The K, value characterizes the affinity between substrate and enzyme and in a first approximation can be viewed as the dissociation constant of the enzyme-

1.5 Enzyme Kinetics

substrate complex ES. K, is independent of enzyme concentration and usually has is the maximal reaction rate possible if every values between and 3 0-2 M. V, enzyme molecule present is saturated with substrate and is a property of the particular enzyme. It may be related to the molecular mass of the enzyme and then is called turnover number, representing the number of substrate molecules converted per active site of an enzyme molecule per unit of time. The turnover number may and lo6 s-l; 103-4s-l is commonly found. have values between Another quantity used frequently for the characterization of an enzyme is the catalpc activity. The unit for the catalytic activity is the Katal (kat), as defined by the International Union of Eliochemistry (IUB), 1 kat corresponds to the amount of enzyme catalyzing the coriversion of one mole of substrate per second at 30 "C under specified conditions. In the biochemical literature, another quantity is often used, the international unit (IU);1 IU catalyzes the conversion of 1 pmole of substrate per minute under specified conditions. From the catalytic activity other values such as volumetric activity ]kat 1;'; IU ml-'1 or specific activity [kat kg-'; IU mg-'1 are derived. Catalpc activity can be determined unequivocally even in crude mixtures and if the molecular prop-rties of the enzyme are unknown. Therefore, enzymes are quantified measuring tht-ir catalytic activity and sold on the basis of activity. To ensure reproducible and meaningful results when measuring enzyme activity, the several points have to be taken into consideration. As shown in Fig. 1-10[Eq. (l)], reaction velocity depends on substrate concentration; for [S] 2 100 K, the reaction rate becomes zero order and so no longer depends on substrate concentration. In special cases, for example, lipases reacting at an interface, the reaction rate depends on the available interface rather than the concentration. Lipases are therefore preferentially analyzed in stable emulsions. The catalpc activity has to be determined at sufficiently high substrate concentration (>10 Km) to ensure pseudo-zero order rates. This may be difficult to achieve with substrates of low solubility. Furthermore, it is desirable to measure initial rates, when only a small amount of total substrate is converted; [S] remains essentially constant during the reaction time and [PI is small. In reactions involving more than one substrate all concentrations have to be considered. I f an unknown substrate or reaction is investigated two or more substrate levels should be employed. At low substrate concentration and high K, values the observed reaction rate may be small and not easily differentiated from background noise, while, at high substrate concentration, inhibition by surplus substrate (see below) may cause a substantial drop in the rate. The reaction rate is be5 t determined by analyzing product formation as a function of time by physical methods such as UV/VIS spectroscopy,optical rotation, potentiometry, etc. Alternatively, formation of a coproduct produced in stoichiometric relations may be followed, such as formation of NAD(P)H in dehydrogenase reactions, which is followed conveniently at 340 nm in a spectrophotometer. Product formation may be coupled to a second reaction using a surplus of an auxiliary enzyme producing an easily quantified signal, for example (NAD(P)+ or NAD(P)H+H+with a dehydrogenase. glutaminase

L-gluta:mine + H2O 4 r-glutamate + NH3

I

25

26

I

I lntroduction

L-glutamate + NAD' + H2O

.

glutamate-dehydrogenase

a-ketoglutarate + NADH + NH4+

A similar approach determines a quinone-imine dye formed by the reaction of HzOzcatalyzed by peroxidase. alcohol oxidddase

2 ethanol + 2 0 2 + _ _ _ j 2 acetaldehyde + 2 H202

2 Hz02 + 4-amino antipyrine + phenol

peroxidase

quinone-imine dye + 2 HzO

In such coupled systems, care must be taken in choosing reaction conditions, such that the enzyme of interest is catalyzing the rate-determining step. Special synthetic colorless substrates converted to colored products have been developed for hydrolases (esterases, phosphatases, glycosidases and proteases); 4-nitrophenol or 4-nitroanilide are used as the alcohol or amide component, which can be measured readily around 400-420 nm. phosphatase

4-nitrophenyl phosphate + HzO L4-nitrophenol + phosphate N-a-benzoylarginine-4'-nitroanilide') __+ N-a-benzoyl arginine + 4 nitroaniline protease

If direct physical measurements are not available or feasible, the enzymatic reaction can be stopped at predetermined times by rapid heating, acid treatment, or similar measures and the amount of product present at time t measured by available analytical techniques such as HPLC, GC, TLC (with or without prior derivatization). Controls are required to ensure that the conditions employed to stop the enzymatic reaction do not destroy the product and that the derivatization is complete. It may be more convenient to follow the decrease in substrate concentration over time as a measure of enzyme activity. This has the disadvantage that the difference of two large values is prone to error. If such an approach is adopted it has to be proven by independent experiments that the anticipated product is actually formed. 1.5.2 Inhibitors and Effectors

Chemical compounds negatively influencing the reaction rate of an enzymecatalyzed process are called inhibitors. Irreversible inhibitors might be reactive substrate analogs forming a covalent linkage to the enzyme after binding and in this way blocking the reactive site. Usually, such reactions are designed intentionally. Heavy metal ions present in trace amounts as contaminants in crude substrates may react with essential sulfhydryl groups and inactivate the enzyme. the situation is similar to the well known poisoning of a metal catalyst by sulfur compounds. Far more important for enzymatic processes are reversible inhibitors, forming specific enzyme inhibitor complexes and thereby influencing the reaction rate. It is important to note that substrates and especially products might inhibit an enzymatic

7.5 Enzyme Kinetics Figure 1-11. Reaction rate as a function o f substrate concentration illustrating allosteric regulation o f enzyme activity: a) rate in the presence o f an allosteric activator, b) rate in the absence o f effectors, c) rate in the presence of an inhibitory effector.

reaction as might substrate analogs. Inhibition by substrate and/or product(s) is important when considering how much of the activity added actually can be utilized at a given set of reaction conditions. Such reaction engineering aspects are treated in more detail in Chap. 4 of this book. Many enzymes may also be activated by inorganic ions such as Ca2+, K+, or C1- possibly raising V, by stabilizing certain protein conformations. If such an effect is noted, the activator should be added in saturating amounts. Special effects are observed in the kinetics of allosteric enzymes. A typical sigmoidal curve describing reaction rate as a function of substrate concentration is presented in Fig. 1-11. Binding of an effector to the regulatory center alters the reaction rate very efficiently and subtly and is often used in nature to divert the metabolic flow into different directions at branching points. Such a response is important for living systems, but rarely will be seen with enzymes employed in organic synthesis. The complex kinetics may be described by appropriate mathematical models, found in the specialized literature. 1.5.3

Influence of pH and Buffer!;

Enzymes contain many polar amino acids at the surface which may be protonated or unprotonated depending on the pH of the surrounding medium. Typikal pK, values are included in Table 1-3. Consequently, charges on the protein surface are altered will depend on pH. Fig. 1-12 illustrates that an optimum of the and K, and V,, reaction rate is observed ;is a function of pH. The optimal pH may vary slightly for different substrates, reflecting differential binding energies. The pH optima for the forward and reverse reaction of the same enzyme are not identical and may differ by 2-3 pH units. In the laboratory, the pH is usually set and maintained using buffers. Selection of buffer ions may influence the observed reaction rate as shown in Fig. 1-3. The reasons are not well understood and are thought to be related to the polarity of buffer molecules interacting with the protein, influencing simultaneously hydratization and solubility of substrates. On the preparative scale, pH is maintained better by a pH-stat arrangement, saving chemicals and separation cost. Also, on the

I

27

28

I

7 Introduction

Figure 1-12. Reaction rate as a function o f pH. Reductive amination of 2-keto caproate (O), 2-keto-isocaproate ( O ) ,2-keto-valerate (o), 2-keto-4-mercapto-butyrate (A),and 2-keto isovalerate g)by leucine dehydrogenase (Bacillus cereus) are shown as function of pH.

so0

.v 300 0

Ex 0,

N

;

200

100

Y

I

1

I

7

8

9

10

Y

71

PH preparative scale, high substrate levels are desired, impossible to buffer sufficiently in reactions involving release or consumption of protons. In switching from buffered to pH-stat operation one should be aware of changes in kinetics as discussed above. The pH of the solution is important not only for enzyme activity but also for enzyme stability. Unfortunately, the optimal pH values for enzyme activity and stability are not necessarily identical, as is well documented in the literature for the hydrolysis of penicillin G by penicillin acylase. In such cases, the method for controlling pH and mixing behavior of the reactor may become crucial. 1.5.4

Temperature

Another important factor for enzyme activity is temperature. In general, the reaction rate will increase with temperature (Fig. 1-14). From an Arrhenius type plot, the activation energy of the process may be calculated. With increasing temperature, however, the mobility of protein segments increases while the strength of hydro-

7.5 Enzyme Kinetics 129

-

5

6

7

1

8

I

9

10

11

PH Effesct o f p H on the activity ofsec-alcohol dehydrogenase (Candida boidinii) during the oxidation o f isopropanol in various buffers in 50 m M concentration: 0 sodium citrate, 0 potassium phosphate, Atriethanolamine/HCI, ATris/HCI, W glycine. Figure 1-13.

phobic interaction decreases. At first, this results in a decrease in catalytic activity, but, with further rise in temperature, in complete deactivation. Thermally induced denaturation of proteins often leads to aggregation which is not readily reversible. Denaturation may be expected in the temperature interval between 30 and 80 "C. The optimal temperature of operation has to be lowered if long reaction times or long service life of an enzyme are required. For enzyme assays, a defined (for example 30 "C) and constant temperature has to be maintained. Enzymes from extremely thermophilic microorganisms may be almost inactive at ambient temperatures and operate in the temperature interval between 80 and 120 "C.

30

I

1 introduction

1

.-2. 8 > ._ 5 m

1:

1

30

I

I

40

50

*

60

Temperature ("C)

/-

t 0 '

Y h c

,

20

0

60-

(u

; .-

a:

LO-

(u

20

2o

1 -

I

20

I

30

I

LC!

I

*

50

Temperature ("C) Figure 1-14. Temperature dependence o f the reaction rate A: L-2-hydroxysisocaproate dehydrogenase (L. confusus) 6: D-lactate dehydrogenase (L. confusus).

1.6 Organic Solvents as Reaction Media

1.6

Organic Solvents as Reaction Media

Enzymes as biocatalysts have been developed for aqueous reaction systems. Application of enzymes in the preijence of organic solvents is of interest to organic chemists because substrates may not be sufficientlysoluble in water, or the equilibrium of the desired reaction may be iinfavorable in aqueous solution. The following general approaches are used - to add increasing con'centrations of water miscible solvents to the reaction

system, - to work in two-phase systems composed ofwater and an immiscible solvent, - to work in nearly anhydrous organic solvents with minimal necessary amounts of

water. In the first two cases, the enzyme may be employed either in the soluble state or immobilized. In nearly anhydrous organic solvents the enzyme is present in the solid state only. The presence of organic solvents will influence activity as well as stability of enzymes. In recent years, work of various groups has shown that the majority of bulk water in a reaction system may be replaced by organic solvents. A certain low amount of resi dual water is needed for activity; 0.02 % may be sufficient. Organic solvents influence the dielectric properties of the reaction medium and to varying degree disrupt ordered water structures. This, in turn,will influence the non-covalent, weak forces responsible for the ordered structures of an enzyme. Protein structures may be stabilized by adsorption, crosslinking, or covalent binding to a hydrophilic surface. Immobilization may also help to avoid denaturation at the interface in two-phase systems. If an immobilized or solid enzyme preparation is used, it is important to provide sufficient surface area to catalyze the reaction. In nearly anhydrous systems, maintaining the pH in the optimal range is a problem. In such cases the enzyme has to be prepared (dried)under pH conditions providing the optimal activity. This way the dissociation of charged groups on the enzyme surface is fixed; there obviously exists a memory effect. The selection of a suitable solvent with regard to activity and stability may be guided by the log P concept, where Pis the partition coefficient of thr. solvent in an octanol/water biphasic system. Hydrophilic solvents with log P value:<< 2 often lead to enzyme deactivation if present in high concentrations; in contrast, apolar solvents with log P 2 4 are compatible with enzymes, leaving the essential layer of water molecules on the polar surface regions unperturbed. The results using solvents with intermediate values of log P (2-4) are unpredictable and depend very much on the individual case. Solvent selection and reaction conditions today are optimized empirically. Nevertheless, there are many examples which clearly demonstrate that enzymes can be employed successfully also in organic solvents.

I

31

32

I

1 Introduction

1.7

Enzyme Handling: Quality Requirements

An increasing number of enzymes are offered on the market by manufacturers and vendors. Enzymes are produced for different purposes and may differ widely in purity and price. In general, enzymes are sold on the basis of unit of activity. The catalogue or data sheet of the supplier should contain information or a reference on assay conditions and definition of the unit. Special requirements for storage (4 “C or -20 “C) are recorded on the label. Enzymes are shipped as dry powders, suspensions or (frozen) concentrated aqueous solutions. The sample may contain undeclared additives such as inert materials (for example, fillers or filter aids), salts or saccharides for stabilization, or precipitating agents such as ammonium sulfate or polyethyleneglycol. Crude preparations may also contain several enzymatic activities, for instance crude pig liver esterase is a mixture of esterases, lipases and other enzymes. The presence of other proteins may be inferred from the specific activity (U/mg protein) of the enzyme preparation, provided the catalytic activity of the pure protein of interest is known. All common protein assays are relative measurements only and therefore depend to some extent on the method and the protein used for calibration. If desired, compounds of low molecular weight can be removed from the enzyme by dialysis, ultrafiltration or gel filtration. For assay purposes, the necessary dilution of the catalyst may often be sufficient to avoid interferences. Wetting of dry proteins is not without problems, resolubilization of (freeze)dried material may take many hours and appropriate controls should be applied if the expected conversion or activity is found to be low. The requirements for purity of an enzyme are not very high from the point of view of application in the synthesis of fine chemicals. More importantly, enzyme quality can be accessed fairly accurately by establishing a mass balance. Only such activities responsible for undesired side reactions have to be removed. As an example, fumarase (adding water to fumaric acid) should be absent from an aspartase preparation (adding ammonia to fumaric acid), because the enzymatic reaction is performed in water. Aspartase, however, is “silent”in a fumarase preparation as long as ammonium ions are absent. Therefore, aspartase does not necessarily need to be removed from a catalyst employed for production L-malic acid. Very often one can exploit the fact that the reaction in synthetic applications is restricted to the available reactants. Such a situation is entirely different from analytical applications, where complex mixtures are introduced and the selectivity of the reaction depends solely on the catalyst. An enzyme should be purified only to the degree necessary for the application. Absence or low levels of proteases are also desired to protect the enzyme catalyst from degradation. Stability under operation conditions is usually more important for production than the initial cost of the enzyme. Stability will allow long reaction times at low enzyme levels or prolonged service life in continuous processes. Genetic engineering techniques can be employed for the production of large amounts if a certain enzyme is needed for application as a catalyst. In addition, specialized knowledge is available to bring the desired enzyme to an appropriate

1.8 Biotransformation Using Whole Cells

level of purity. Such develcipments will lower the cost for the catalyst considerably, which should be kept in mind when analyzing and evaluating process costs.

1.8 Biotransforrnation Using Whole Cells 1.8.1 General Aspects

Microbial cells can be employed also as biocatalysts to achieve a desired conversion instead of isolated, cell-free enzymes. In contrast to microbial cultivation exploiting the complex metabolism of cells to produce, for example, organic acids or antibiotica from sheap nutrients, biotransformation utilizes only one or a few enzymes to convert added educts to a desired product. Since many microbial enzymes will accept non-natural compounds biotransformation also gives access to many products not found in nature, while microbial cultivation yields only natural primary or secondary metabolites. The acetic acid generator developed in 1824 by C. Ham represents a suprisingly modem industrial biotransformation process oxidizing ethanol to acetic acid using strains of Acetc bacter immobilized on beech shavings in a current of air. About 100 years later the biotransformation process for the production of Rl-phenyl-l-hydroxypropane-2-one, the key chiral intermediate in the synthesis of ( l R , 2s) ephedrine was cclmmercialized using Saccharomyces cereuisiae to catalyze a stereoselective acyloin condensation. At the same time regiospecific oxidation of Dsorbitol to L-sorbose by Aretobacter suboxydans or related species was developed as a key step in vitamin C production. Biotransformations have since revolutionized the production and availability of steroid hormones and steroid related drugs including contraceptives, cortisone, prednisone etc. The modification of the steroid ring system as well as other cyclic structures by various microbial cultures is well documented in the literature. From this information, selection of potentially useful strains is possible for hydroxylation, oxidation, dehydration, reduction or dehydroxylation reactions. For a successful process development beyond a few grams and scale-up to an industrial ~~rocess extensive screening and biochemical engineering will be necessary demanding a close collaboration between biologists, chemists and engineers. There are far more rnicroorganisms available from culture collections than enzymes on the market. A list of important culture collections can be found in Table 1-6. Further information can be found on the internet:

http://wdcrn.nig.ac.jp/wfi:c/wfcc.html Fortunately, the majority of known microorganisms are classified as non-pathogenic and harmless to humans and the environment. These can be handled safely in the normal laboratory. Strains with desirable properties identified from the literature can often be ordered front catalogues of culture collections,just like chemicals from a supplier. There may be more than one strain of a given species listed in the catalogues as there are many “John Brown” found in a telephone directory. It is

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7 Introduction Table 1-6.

Important collections of microorganisms in the public domain.

American Type Culture Collection (ATCC), 12301 Parklawn Drive, Rockville, Maryland 20852, USA Centraalbureau voor Schimmelcultures (CBS), P. 0. Box 273, Oosterstraat 1, NL-3740 AG Baarn Japan Collection of Microorganisms (JCM)RIKEN, Wako, Saitama 351-01, Japan Institute of Applied Microbiology (LAM), University ofTokyo, Yavoi 1-1-1,Bunkyo-ku,Tokyo 113, Japan Culture Collection of the Institute for Fermentation (IFO), 17-85 Juso-Houmachi 2-chome, Yodogawa-ku,Osaka, Japan Agricultural Research Service Culture Collection (NRRL), Northern Regional Research Center, Agricultural Research Service, US Department of Agriculture, 1815 North University Street, Peoria, Illinois 61604, USA National Collection of Industrial and Marine Bacteria Ltd., 23 St Machar Drive, Aberdeen AB2 lRY, UK Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (DSM-Z), Mascheroder Weg lb, D-38124 Braunschweig

advisable to search for the strain identified by the unique catalogue number that was originally employed. If the particular strain is no longer available or cannot be identified from older literature on should screen for the desired activity in a number of strains from the same species to find a good substitute. Usually strains are mailed as lyophilized culture or as a freshly inoculated agar slant. The investigator has to master some basic skills to maintain and grow cells under aseptic conditions if he wants to utilize these biological resources. Detailed information can be found in text books of microbiology and laboratory manuals. For aerobic organisms (needing oxygen for growth) or microaerophilic organisms (tolerant against trace amounts of oxygen) a working knowledge of media preparation is required (useful compositions are found in the literature or in catalogues of culture collections) as well as sterilization techniques for media and equipment to remove or destroy all living cells by heat, microfiltration or chemical treatment before inoculation with a pure strain. Aseptic conditions have to be maintained when inoculating sterile media with pure strains and in sampling the culture later. This means that airborne particles containing contaminating microorganisms have to be excluded from the growing culture, which can be accomplished working in a laminar flow hood and using exclusively sterilized equipment in handling. The laminar flow hood provides a positive pressure of filtered air in the working area and was especially designed to allow easy handling under aseptic conditions. Aerobic microorganisms are grown in a capped Erlenmeyer flask with baffles on a rotary shaker at constant temperatures in the range between 25 and 40 "C. Microaerophilic cultures are purged with nitrogen and agitated slowly if at all. Growth is observed as turbidity; the uniformity of the culture is checked using a light microscope. Pure cultures can be used to prepare seed stocks taking aliquots at the mid or late

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1.8 Eiotransforrnation Using Whole Cells 35

logarithmic growth phase and freezing the suspension at -20°C or -80 “C if available. Glycerol (5-10%) or dimethylsulfoxide (2.5-5%) may be added as a protecting agent. Working cultures can be maintained on nutrient agar in a Petri dish or vial at 4 “C for some weeks. There are numerous other methods to preserve and maintain pure cultures. The culture collection or a microbiologist should be consulted in case of doubt. Anaerobic microorganisms cannot utilize oxygen as a terminal electron acceptor during growth and usually are very sensitive to oxygen in the environment. Instead they use a vast variety of organic and inorganic compounds as electron donors and acceptors in their energy metabolism. Strictly anaerobic suains are difficult to handle especially on a small scale without proper training and specialized equipment. Nevertheless anaerobic strains may contain interesting biocatalysts also for biotransformations, especially unusual redox enzymes. Some microorganisms produced as starter cultures in the food industry can be easily obtained without necessitating the ability to grow microbial cultures under defined conditions in the laboratory. The most notable example is “baker’s yeast”, Saccharomyces cerevisiae, which can be bought cheaply in large amounts from the regional supplier of bakeries or in smaller amounts from any local food store. Baker’s yeast is sold as pressed filter cake (usually containing a certain amount of starch granules), in the form of blocks or flakes or as a lyophilized preparation. The latter is rehydrated to a viable culture using water or buffer. Because of the ready ,availability and cheap price, baker’s yeast has been extensively used for the stereoselective reduction of carbonyl groups in numerous examples. Problems may arise from genetic and physiological differences in local strains of S. cerevisae beyond the control of the organic chemist. This leads to variation in the stereoselectivity and sometimes poor reproducibility of published results. The cells often contain more than one dehydrogenase/reductase accepting the substrate of interest and the result obtained is the sum of all parallel reactions. Besides, the biosynthesi:; of enzymes in general is strictly regulated in microorganisms. The activity oj-an enzyme in a given microbial cell may vary more than 1000-fold and depends 011 available nutrients and inducing agents, growth conditions, time of harvest, condition of storage etc. This biological variability (and the reasons behind them) should be kept in mind when planning and judging biotransformations. The relative merits of using whole cells versus cell-free enzymes as biocatalyst are summarized in Table 1-7. Obviously whole cell biotransformations are the method of choice if the enzyme involved is not stable enough in isolated form or is an integral part of the cell membrane and utilizes tk e electron transport chain in complex coenzyme regeneration schemes. Whole cell biotransformations are particularly important for hydroxylation and oxidation reactions involving mono- and dioxygenases or epoxidases. Depending on the type of the reaction investigated, whole cells are employed as growing culture, as viable resting cells or in a nonviable form simply as a “bag of enzymes”.

7 introduction Table 1-7.

Advantages and disadvantages using isolated enzymes or whole cell biocatalysts.

Isolated Enzymes

Advantages

Disadvantages

Catalyst concentration as free process variable

Limited stability

High catalyst concentration possible

Cofactor regeneration needed

No side reaction Simple product recovery No transport limitation

Multienzyme reactions possible Whole cell biocatelysts

Unlimited availability exploiting growth

Side reactions

Cofactor recycling by cellular machinery

Transport limitation

Multistep conversions possible

Complex product recovery In general low space time yield

1.8.2

Biotransformationwith Growing Cells

When growing cells are to be used for a biotransformation, e. g. of steroids, the cells are grown in a suitable growth medium, usually in batch culture. After an initial lag phase the cell will grow and multiply exponentially as observed by the turbidity of the culture. After one or more compounds become rate limiting for growth a period of linear growth may be observed before a stationary phase is reached, where viability of cells is maintained but growth stops. Eventually cells will die if energy sources are exhausted. Many h n g i but also bacteria produce spores in the stationary phase, which are able to withstand adverse conditions and secure the survival of the species in nature. The biosynthesis of a desired enzyme is often not parallel with growth: the specific activity will vary with the growth phase. Therefore timing of educt addition is crucial for product yield. The optimal conditions are usually determined experimentally. It may be advantageous to add small amounts of educt in the early logarithmic phase to help to induce the formation of the desired enzyme(s).Toxic substances are added best in the late logarithmic growth phase to minimize negative effects on growth and enzyme production. Repeated dosing helps to maintain low stationary levels of toxic or inhibitory educts but allow product accumulation to high levels provided the product is non-toxic. After batch or fed batch biotransformation processes, the biomass is separated and discarded after the conversion is completed or when side reactions become prominent. The product has to be isolated from whole broth or the spent medium depending on the yield. Fresh biomass has to be prepared from culture stocks for each successive experiment.

1.8 Biotransformation Using Whole Cells

1A.38

Biotransformationwith Resting Cells

Resting cells are non-growing viable cells retaining many enzyme activities of growing cells. Bakeri yeast, discussed above, essentially consists of resting cells of S. cerewisiae. In the laboratory, resting cells are obtained by growing the selected microbial culture under ,appropriate conditions until a high or maximal enzyme activity is reached in the cells. At this point in the growth cycle, cells are separated frorn the growth medium by centrifugation or filtration and washed with saline. Then cells are resuspended in the biotransformation buffer, and the conversion is followed by suitable analytical techniques. Cell concentration can be varied and a higher catalyst concentration applied in comparison with experimlmts using growing cells. The addition of small amounts of glucose or other energy sources helps to maintain the electrochemical potential across the cell membrane and the viability of resting cells. If the biotransformation step requires coenzyme regeneration, co-metabolites such as glucose or glycerol have to be added in sufficient amounts. Resting cells are convenient to use as one large cultivation yields a uniform biocatalyst for many parallel biotransformations. It may also be possible to store the cell cake at 4 "C for some days or weeks without detrimental losses in actikity. Otherwise resting cells may be conserved by lyophilizatiori using sucrose or trekialose as a protectant. The biotransformation buffer is less complex than spent medium, and thus product isolation is usually easier than from growing cells. Once the initial growth is completed the danger of infection is also less using resting cells because biotransformation buffers often lack essential nutrients for growth. This makes handling in the laboratory more convenient. 1.8.4

Biotransformations with Plermeabilized or Dried Cells

Microbial cells are separated from their surroundings by complex cell walls and one or more membranes. The cell wall provides the mechanical strength to withstand sudden changes is osmotic pressure while the membrane serves as an effective diffusion barrier. In addition, the membrane is important for the selective interaction of the cells with the environment and the maintenance of an electrochemical potential important for viability. Cell membranes may cause more or less severe mass transfer limitation:;, hindering educts from reaching biocatalysts inside the cells and the transport of products out of the cells. For lipophilic, uncharged and sufficiently soluble educi s, passive diffusion across the membrane@) may provide satisfymg reaction rates. Detergents or solvents can be used to enhance permeability and mass transport. This approach should be used with care in case membranebound enzymes or coenzyme regeneration via the electron transport chain are necessary for the particular biotransformation. Otherwise, the integrity of the membrane may be of no concern for simple reactions involving only intracellular enzymes. For example L- malic acid is produced from fumarate using Breuibacieriurn amrnoniagenes as a biocatalyst. The cells are permeabilized by treatment with bile to

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I improve transport of the charged reactants, and in addition a side reaction, the I introduction

conversion of fumarate to succinate by membrane-bound enzymes of the citric acid cycle, is abolished. Resting cells may be dried by treatment with a large excess of cold (-20 "C) acetone; the resulting "acetone powder" can be stored at -20 "C for many months, yielding a convenient enzyme source. It should be noted that acetone treatment removes lipids from the cell membranes, and therefore membrane-bound activities are irreversibly damaged. Disturbance of membrane function is the major factor in the toxicity of solvents towards microbial cells and needs to be considered in a case-by-caseevaluation.

Bibliography S. L. Neidleman (1990),The archeology of enzymology, in Biocatalysis (D. Abramovicz, van Nostrand, eds), pp. 1-23. H. Uhlig, Enzyme arbeitenfir uns, Technische Enzyme und ihre Anwendung, Carl Hanser Verlag, Miinchen, 1991. W. N. Lipscomb (1983), Structure and catalysis of enzymes, in: Ann. Rev. Biochem. 52, 17-34. D. W. Christianson, W. N. Lipscomb (1987), The complex between carboxypeptidase A and a possible transition state analogue, I. Am. Chem. Soc. 108,4998-5003. B. L. Vallee, A. Galdes (1984), The metallobiochemistry of zinc enzymes Adv. Enzymol. Rel. Areas Mol. Biol. 56, 284-430. W. N. Lipscomb (1971), Proc. Robert A. Welch Found. Con$ Chem. Res. 15, 141. G. E. Schulz, R. H. Schirmer: Principles of protein structure. Springer Verlag, New York, 1979. C. R. Cantor and P. R. Schimmel, Biophysical chemistry, part I: The conformation of biological macro-molecules. part 111: The behavior of biological macromolecules, W. H. Freeman and Co., New York, 1980. T. E. Creighton, Proteins, struaures and molecularproperties, W. H. Freeman and Co., New York, 1984. K. Buchholz und V. Kasche, Biokatalysatoren und Enzymtechnologie, VCH, Weinheim, 1997. D. Zaks, Enzymes in organic solvents, in: Biocatalystsfor Industry (J. Dordick, ed), Plenum Press, New York, 1991, pp. 161-180. Enzyme nomenclature, published for the I U B by Academic Press Orlando, 1992.

Enzyme chemistry, (C. J. Suckling, ed) Chapman and Hall, London, 1984. A.Schellenberger (ed),Enzymkatalyse, Springer Verlag Berlin, 1989. M. J. Page, 7'he chemistry ofenzymeaction, Elsevier, Amsterdam, 1984. M. J. Page and A. Williams, Enzyme mechanisms, Royal Society of Chemistry, London, 1987. H. Bisswanger, n e o r i e und Methoden der Enzymkinetik,Verlag Chemie, 3rd edn, 2000. W. W. Cleland, Enzyme kinetics as a toolfor determination ofenzymemechanism, in: Bernasconi (ed),Investigations of rates and mechanism of reactions, Wiley, 1986, pp. 792-821. A. Cornish-Bowden, Principles ofenzyme kinetics, Butterworth & Co., London, 1976. J.A. Robinson, J. Retey, Stereospecijkity i n organic chemistry and enzymology,VCH Verlag, Weinheim 1982. I . H. Segel, Enzyme kinetics, Behavior and analysis of rapid equilibrium and steady state enzyme systems, Wiley & Sons, New York. 1975. G. N. Wilkinson, Statistical estimations on enzyme kinetics, Biochem. J. 80, 1009-1012. A. Fersht, Enzyme structure and mechanism (2ndedn.), W. H. Freeman & Co., New York, 1985. B. E. Kirsop and A. Doyle (eds), Maintenance ofmicroorganisms and cultured cells, Academic press, London, 1991. J. C. Hunter-Cevera and A. Belt, Isolation of cultures, in: Manual of Industrial Microbiology and Biotechnology, (2ndedn.),A. L.

1.8 Biotransformation Using Whole Cells

Demain and J. E. Davies (eds.), ASM Press, Washington, DC 1999, pp. 3-;!0. R. L. Monoghan, M. M. Gagliardi and L. Streicher, Culture preservation and inoculum development, in: Manual ofhdustrial Microbiology and Biotechnology (2ndedn.), A. L. Demain and J. E. Davies (eds.) ASM Press, Washington, DC 1999, pp. 29-48.

M. D. Hilton, Small scale liquid fermentation, in: Manual oflndustrieal Microbiology and Biotechnology, (2ndedn.), A. L. Demain and J. E. Davies (eds.), ASM Press, Washington, DC 1999, pp. 49-GO. R. Leon, P. Fernandes, H. M. Pinheiro and J.M.S. Cabral, Whole-cell biocatalysis in organic media, EnzymeMicrob. Biotechnol. 23, 483-500.1998.

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Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

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2 Production and Isolation o f Enzymes Yoshihiko Hirose

2.1 Introduction

This chapter gives a brief review of the isolation and production of enzymes. More detailed information can be obtained from various published textbooks and review~['-~]. Most of the industrial enzymes used for chemical synthesis are supplied in a crude form with an active enzyme content of only a few percent. The other constituents are inorganic salts, polysaccharides and diatomaceous earth used as stabilizers and excipients. Purified enzymes for biotransformation are supplied by some manufacturers in a crystal or immobilized form. These enzymes, though expensive, are easy to apply for biotransformation in organic media. The use of more purified enzymes is increasing. Barriers to the production of industrial enzymes include economic factors, the availability of optimal enzymes and safety issues. Common fermentation and purification processes are described in Figs. 2-1 to 2-3. The process differs for extracellular and intracellular enzymes, liquid and solid culture, and enzyme application. The fermentation conditions are computer-controlled for optimization, e. g. of temperature, pH, agitation speed, aeration, demand oxygen etc. There are no internationally standard assay methods for industrial enzymes and the definition of enzyme activity unit is also different for each enzyme. The activity of industrial enzymes is shown by various methods depending on manufacturers. For instance, commercial lipase activities are measured by the hydrolysis of olive oil under the various conditions and these figures are not comparable with each other. When customers apply these biocatalysts for chemical synthesis in organic solvents, these figure are sometirnes reliable, and sometimes not. Users should not judge commercial enzymes based only on price and the activity shown in the table the manufacturer provides. Enzymes should be evaluated based on their practical performance under the conditions used. Most users of biotransformation are not experts in measuring enzyme activity, so the establishment of an assay method and practice are essential if one is to optimize the performance of enzymes. Several commercial enzymes are powders including diatomaceous earth or

Broth-out Filtration Concentration by evaporation or ultrafiltration Precise filtration Solvent precipitation Filtration or centrifugation

Dissolution Ion-exchangechromatography Salting-out Desalting Hydrophobic chromatography Gel filtration Desalting

V

Freeze-drying

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2.1 fntroduction 43

Preparation of fermentation medium ----- Starch, sugar, soybean powder, yeast extracts, minerals, inducer, etc. Dissolution Sterilization Inoculation

Fermentation in flask to in main tank

1

Centrifugation or Filtration

Disruption and extraction

I

Filtration Concentration by evaporation or ultrafiltration Further purification (Salting-out, chromatography, etc) Precise filtration

j.

Crystallization or freeze-drying

Product Figure 2-2.

Commori production process for special intracellular enzymes.

Sterilization Inoculation Extraction of enzyme Filtration Concentration by ultrafiltration Precipitation

44 2 Production and lsolation of Enzymes

I

dextrin. These enzymes should be used after immobilization on a suitable carrier. The activity of an immobilized enzyme usually is enhanced up to tenfold. Regulatory assessments for enzymes used in biotransformation are not clearly stipulated. At present, food assessments of microbial enzymes are provided by AMFEP, which has suggested microbial enzyme purity and immobilization as given below. Purity A chemical and microbial specification must be given. Based on FCC recommendations, AMFEP recommended the following. Arsenic Lead Heavy metals Mycotoxins Antibacterial activity Coliforms E. coli Salmonella Total viable count

3 PPm 10 PPm <40 ppm negative negative <30/g negative in 25 g negative in 25 g <5oooo/g

Immobilization The immobilization system should be described in detail. Tests to indicate the physicochemical stability of both the system and its carrier and enzyme are essential. These regulatory aspects would be acceptable for biocatalysts.

2.2

Enzyme Suppliers for Biotransformation

There are more than 400 companies dealing with enzymes all over the world and approximately 12 major producers with an increasingly distinct separation of product ranges. About GO companies produce substantial amounts of a small range and about 400 companies produce a very limited range of industrial enzymes. Japanese enzyme producers have a special range for industrial or in-house use and contribute to 12-15% of world production. There are 24 companies which supply special enzymes for biotransformation (Table 2-1).

2.3 Origins of Enzymes Table 2-1.

Main enzyme suppliers for biotransformation.

Company

Country

Altus Biologics Inc. Amano Enzyme Inc. Asahi Chemical Co. Biocatalysts Ltd. Biozyme Labs Ltd. Calbiochem Corp. Christian Hansen AS Diversa Fluka Chemicals Ltd. Genencor Int. Genzyme Ltd. DSM (Gist) Meito Sangyo Co. Merck KSA Nagase Biochemicals Novo Nordisk AS Oriental Yeast Co. Osaka Saiken KK Roche Diagnostics GmbH Rohm GmbH Shin Nihon Chem Co. Sigma Chemical Co. ThermoGen Toyobo Co.

USA Japan Japan UK UK USA. Denmark DK USA Germany, UK Finland, USA UK Holland Japan Germany Japan Denmark Japan Japan Germany Germany Japan USA USA Japan

2.3

Origins of Enzymes 2.3.1

Microbial Enzymes

More than 90% of enzymes are produced by fermentation by microorganisms, which are used to prepare industrial and special use enzymes. Prokaryotic cells and eukaryotic cells can be easily grown in culture, and the technology of scale-up is well established on an industrial scale. Various kinds of fungi, bacteria and yeast have been screened for the production of special enzymes. Extracellular enzymes, for instance hydrolytic enzymes, are secreted into liquid and solid culture and are relatively stable in cultivation media. Production by genetically modified organisms (GMO)is becoming popular in this field, and several kinds of GMO have been used to increase the productivity of biocatalysts. When one uses GMO enzymes for industrial use, one should know the origin of the microorganisms used and how the production method was changed. The new enzyme preparation is likely to have a different compositional spectrum of enzymes and side activities.The regulations for biocatalysts are not severe at present, but are likely to become more stringent.

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2 Production and /solation ofEnzyrnes

2.3.2 Plant Enzymes

Some proteases, such as papain, bromelain and ficin, lipoxygenases from soy bean and white germ, and peroxidase from horseradish, are typical plant enzymes. Plant proteases are extracted and partially purified to give a powder extract. Some are supplied as digestive enzymes or neutraceutical enzymes. These have the characteristics of an SH-enzyme (thiol protease) and work in the hydrolysis of racemic esters as a protease. Lipoxygenases are only available from soybean, but the activity is not high and the regiospecificity for unsaturated fatty acids is not severe. Lipoxygenases from other plants are relatively unstable and used in-house only. 2.3.3 Animal enzymes

Porcine liver esterase (PLE), porcine pancreas lipase (PPL) and arginase are well known as biocatalysts among industrial animal enzymes. PLE catalyzes very well the hydrolysis of certain kinds of prochiral diesters and is supplied as a suspension with ammonium sulfate or in liquid form. The substrate specificity of PLE is not wide, but this is a well-investigated enzyme. PPL is very cheap and a useful biocatalyst in industry. Commercial PPL is a mixture of many kinds of pancreas enzymes, and the name pancreatin is well known as a digestive enzyme. Pregastric esterase is applied for transesterification of triglycerides. Arginase from calf liver is used to produce Lornithine from the proteinogenic amino acid-arginene. The use of animal enzymes seems to be gradually decreasing because of disease and a variable supply. In the future, animal enzymes will no doubt be replaced by microbial enzymes of equivalent performance.

2.4 Fermentation of Enzymes 2.4.1 Liquid Fermentation

Liquid fermentation is useful for the production of enzymes as well as antibiotics. It is good for scale-up and reproduction. There are two types of enzyme produced: intracellular and extracellular enzymes. With advances in genetic engineering, Escherichia coli is now being used to produce enzymes. When E. coli is used, the enzyme is accumulated inside the cell. This method is very popular.

2.5 Extraction of Enzymes

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2.4.2 Solid Fermentation

In Japan, solid fermenta.tion is still used to produce many kinds of enzymes including lipases, proteases and acylases. Some glycotransferases are also produced by solid fermentation. In the production of proteases, solid fermentation is often used to increase the productivity in solids. On changing to liquid fermentation, the protease is not produced and its properties change. Solid fermentation is oldfashioned and difficult to scale up because of the expensive facilities needed. 2.4.3 Extraction of Enzymes

In order to improve the erttraction of enzymes, organic solvents and surfactants are sometimes used.

2.5

Extraction o f Enzymes 2.5.1 Microbial Enzymes

Extraction methods depend on the fermentation conditions and the microorganism, for example, liquid or solid culture, intracellular or extracellular enzyme, laboratory or production scale preparation, etc. Because enzymes are more soluble in buffer solution than water, enzymes in solid fermentation are extracted by stirring several times in a suitable buffer solution. The pH of the buffer solution is adjusted based on the stability and the PI of the enzyme. The solid medium is removed by filtration after extraction and the crude enzyme solution is concentrated at the next step. Liquid fermentation produces two types of product, intracellular and extracellular enzymes. In the case of extracellular enzymes, the crude enzyme solution is collected by filtration or centrifugation of the microorganism. In the case of intracellular enzymes, collection of the microorganism and extraction of enzymes after disruption is required. On a laboratory scale, ultrasound equipment or a French press is used for disruption of microorganisms. On a large or industrial scale, a mechanical grinding mill with glass beads (for instance, a Dynamill) is used. It can treat microorganisms suspended in buffer solution at a rate of 100 L/h. Another method is enzymatic disruption of microorganism with lysozyme or YL. This method is easy to apply on an industrial scale because it does not require special apparatus. The microorganism suspended in buffer is stirred for several hours in the presence of a suitable amount of lysozyme at room temperature. During the enzymatic treatment, freezing and thawing of the microorganism is effective for disruption. In this case, lysozyme should be added before freezing the micro-

48

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2 Production and Isolation offnzyrnes

organism. The combination of mechanical and enzymatic treatment is much more effective. After disruption of the cell wall, enzymes should be extracted with buffer solution. Sometimes enzymes are adsorbed by or adhered to the cell wall and must be extracted by adding a small amount of surfactant, such as Triton X-100. The cell wall is then filtered or centrifuged to obtain the crude enzyme solution. Recombinant heterogeneous enzyme produced by gene-modified organisms (GMO) such as E. coli are sometimes obtained as inclusion bodies which form insoluble aggregates and show less original activity. In order to regenerate the activity from the inclusion body, it is dissolved in the presence of denaturing agents, usually a highly concentrated guanidinium salt and urea and reducing agents, usually thiol compounds. Protein is allowed to refold into its original active conformation after removal of the agents. 2.5.2 Plant Enzymes

Some proteases, papain and bromelain, are derived from plants and are extracted from fruits. The fruits are ground by a grinder or cutter and the proteases are extracted by buffer solution. A diluted cooled buffer is more effective than water for extraction and the extracted solution including desired enzymes should be cooled during all treatments. The content varies depending on the time and place as well as the plant. 2.5.3 Animal Enzymes

Animal organs containing desired enzymes are stored frozen and then ground and crushed by a homogenizer. Enzyme stabilizers or protease inhibitors are sometimes added on homogenizing the organs, and a buffer is preferable for extraction. In order to improve the extraction, the residue is removed by filtration or by centrifugation to obtain crude extract. Animal organs contain various kinds of enzymes and a large volume of protein is extracted. Even relatively unstable enzymes retain their activity in crude extracts, but it is necessary to purify the enzymes step by step.

2.6 Concentration

After the extraction of extracellular enzymes of solid culture or the fermentation of extracellular enzymes of liquid culture, the fermentation medium including desired enzymes is centrifuged or filtered to remove the microorganism etc. The next step is concentration by evaporation under reduced pressure or ultrafiltration (UF) to reduce the volume of the enzyme solution. It is not easy to evaporate large amounts of water under reduced pressure. Evaporation should be carried out at <30 "C except

2.7 Purification of Enzymes 149

in the case of thermostable proteins, which can be evaporated at higher temperatures. The concentr,itions of the salt and other soluble materials of the concentrated solution are increased by evaporation. The most convenient and simple method for production is ultrafiltration. The method uses membrane tubes with pore sizes from about 6000 to 50000 A. Small molecules like salt ions as well as water pass through the pores of the membrane tube while large molecules like proteins remain inside the tube. The concentration of the buffer solution is the same before and after ultrafiltration. The leaking of desired proteins in permeates should be checked during the concentration stage. Regular maintenance is carried out by using a standard protein. The membrane tube is made of polyethylene, polypropylene etc., and the irreversible adsorption of desired proteins should be avoided. The materials for the membrane should be selected before use. The flow rate of ultrafiltration depends on the facility and the protein solution applied. The final protein concentration is up to about 100 g/L. Ultrafiltration is also used for desalting. Smaller membrane tubes are used for this purpose. Another concentration method is precipitation using organic solvents or salts as described in Sect. 2.7.2. Ethanol is especially useful for this purpose. Despite the volume of organic solvents, it is still frequently used as a first step in the purification.

2.7 Purification of Enzymes 2.7.1 Chromatography

Chromatography is the major purification method. The choice of technique is determined by the overall yield, efficiency, speed and convenience. To purify a desired enzyme in the broth, a combination of different types of chromatography is an effective approach. 2.7.1.1 Ion Exchange Chromatography (IEX)

Ion exchange chromatography (IEX) 171 is the most typical and frequently used method for separating enzymes. Some of the advantages of IEX are high resolution power, applicability, and ease of control and scale-up. There are two types of exchanger in IEX (Figs. 2-4 and 2-5). Positively charged exchangers have negatively charged counter-ions (anions) available for exchange and are called anion exchangers. Negatively charged exchangers have positively charged counter-ions (cations) and are called cation exchangers. The basic principle of separation in IEX is the reversible adsorption of charged protein molecules dissolved in a buffer solution by oppositely charged ion-exchanged groups on the inatrix (Fig. 2-6).

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2 Production and Isolation ofEnzyrnes

Anion exchanger with counter-ions Figure 2-4.

Cation exchanger with counter-ions

Two types o f ion-ex~hanger[~J.

c-

/

Figure 2-5. The principle of ion exchange equilibrium 141.

Proteins dissolved in buffer solution have different charges depending on the solution. When the pH of the buffer solution is below the isoelectric point (PI), the protein has a positive charge, and when the pH is above the PI, the protein has a negative charge (Figs. 2-7 and 2-8). An ion exchanger consists of a solid matrix covalently bound to a charged group. The matrix is made of an organic compound, synthetic resin or polysaccharide, such as Sepharose and Sephadex. A typical matrix is a round microbead. The characteristics of the matrix determine its chromatographic properties, for instance, eficiency, capacity, recovery, chemical stability, mechanical strength and flow proper-

2.7 Purification of Enzymes

0

nvo

v v

000 000 000 000 000

Substances to be separated Figure 2-6.

The principle of anion exchange chromatography[’].

ties. The properties of the matrix affect its behavior towards biological substances and the maintenance of hiological activity. The charged group determines the basic property of IEX, such as the type and the strength of the ion exchanger. The number of charged substituent groups per gram of dry ion exchanger or per mL of swollen gel affects its total ionic capacity. It can be measured by titration with a strong base or acid and is shown as pmol/mL gel. Typical functional groups are shown in Table 2-2 and are classified in two types. The functional groups of ani’mexchangers are substituted ammonium groups. Cation exchangers have sulfoql or carboxyl groups. Ion exchangers with sulfonic and quaternary amonium grcups are called “strong ion exchangers”.Those with carboxyl and diethylaminoethyl groups form “weak ion exchangers”. The terms strong and weak refer to the extent of the variation of ionization with pH and not the strength of binding. A strong exchanger is ionized over a broad pH range, a weak one over a narrower range.

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2 Production and Isolation of Enzymes

Ib ( K L J

tange of stability

Attached to anion exchangers

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\ 6

I PH

I

8

10

t

Figure 2-7.

+

I

The net charge o f protein as a function o f p H f71.

acid

PI

base

R anion exchange cation exchange NH2

Figure 2-8.

Relation between the charge of proteins and the p H 16]

Experimental Design The choice of matrix and functional group depends on the pH stability, molecular size and isoelectric points ( PI) of the protein, and on the requirements of the application. PI can be measured by electrophoresis or can be checked in the comprehensive lists of PI for proteins.

2.7 Purification of Enzymes

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53

Table2-2.

Functional groups used on ion exchangers and its structure.

Ion exchangers

Structure

Anion exchangers Dimethylaminoethyl (DE) Diethylaminoethyl (DEAE) Quaternary ammonium (QA) Quaternary ammonium (QAE) Quaternary ammonium (QMA)

-CHzCHzN(CH3)2 -CHzCHzN(CzHs)z -CHzN+(CH3)3 -CH2CH2NC(C2H5)2CH2CH(OH)CH, -N'(CH,)s

Cation exchangers Carboxymethyl (CM) Phosphate (P) Sulfonic ethyl (SE) Sulfonic propyl (SP)

The starting pH of buffer is chosen so that proteins to be bound to the exchanger are charged. So, the startjng pH is at least more than 1 unit above the PI for anion exchangers or at least less than 1 unit below the PI for cation exchangers to facilitate adequate binding. Proteiru begin to dissociate from ion exchangers at about 0.5 pH units from their PI at ionic strength 0.1 M. Most proteins have their PI within the acidic range, so they are usually negatively charged in neutral buffer solution and show the properties of as anion. Ion exchange separation is carried out using the following three procedures: column chromatography, a batch method and an expanded bed adsorption. Industrial- scale preparation is used.

Column Separation Ion exchangers are available for laboratory-scaleseparations, and factors such as cost and reproducibility etc. are not very important. For industrial separation, however, it is necessary to optimize the purification conditions. The DEAE exchanger is the most useful in terms of the PI and stability of most proteins. Choice of Exchanger Groujr and Buffer The choice of ion exchanger and buffer solution is limited by the stability of the proteins. Because most proteins have their PI in the acidic range, they have a slightly positive charge below the PI and can be easily absorbed on a cation exchanger (ex. Chl). On the other hand, they have a negative charge above the PI and an anion exchanger (ex. DEAE) is used. Choice ofpH and Ionic Strength The pH of the buffer depends on the PI of the proteins, and the ionic strength causes absorption on the ion exchanger and desorption from it. The required concentration of starting buffer depends on the nature of the buffering substance. It should be at

54

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2 Production and Isolation of Enzymes

least 10 mM. Suitable ion salts stabilize the proteins in solution and excess salts cause the denaturation and precipitation of the protein.

Batch Separation Batch separation is conducted in the same way as column development by a stepwise elution method. Batch separation is a better method for large sample volumes with low concentration protein. Large volumes take a long time. The initial conditions for batch separation are almost the same as for column chromatography, for instance, buffer pH and ionic strength etc. It is necessary that the conditions are rather strong so that the proteins completely bind to the adsorbent. In order to increase the recovery, the pH of the buffer is kept a couple of units away from the PI of the protein. Batch separation is simple and useful to concentrate a low protein solution, but the resolution is not high. Experimental Procedure Batch separation is a simple method whereby the protein solution is stirred together with the ion exchanger in an appropriate buffer for 1h. The slurry is collected by filtration and the ion exchanger is washed with fresh buffer solution. When no desired protein is observed in the filtrate, a couple of bed volumes of the elution buffer are added and stirred for 1 h to desorb the desired protein. The solution including the desired protein is collected by filtration. The change of pH or ionic strength of the elution is determined by gradient or stepwise chromatography. 2.7.1.2

Hydrophobic interaction Chromatography (HIC)

Hydrophobic interaction chromatographyf8Iis based on the hydrophobic properties of proteins and the hydrophobic ligands covalently bonded to the matrix. The chromatography has three forms: hydrophobic chromatography,where both absorption and desorption are based only on hydrophobic binding, hydrophobic-ionic chromatography,where absorption is based on hydrophobic binding and desorption on ionic exclusion by changing the pH of the buffer, and mixed function chromatography, based on hydrophobic, ionic and hydrogen binding. HIC and reversed phase chromatography (RPC)are very similar principles based on hydrophobic interaction. Adsorbents for RPC are much substituted with hydrophobic ligands, such as octadecyl,octyl or phenyl groups (Figs. 2-9 and 2-10).Protein binding to RPC adsorbents is usually very strong in spite of the concentration of salt, and some kinds of organic solvents, such as acetonitrile and is0 propyl alcohol, are used to desorb the protein. So, RPC is carried out for low molecular weight molecules such as peptides because most proteins are unstable in highly concentrated organic solvents. On the other hand, adsorbents for HIC are less substituted with similar groups, mainly butyl groups. Protein binding to HIC adsorbents requires a neutral salt like ammonium sulfate in the mobile phase, and the protein is desorbed by decreasing the concentration of salt. The slope of ionic strength in HIC is opposite to that of IEX.

2.7 Pur$cation of Enzymes

f-

OCH2CH(OH)CH20-R:

RC2H5-

Ethyl

C4H9-

Butyl

C6H 13-

Hexyl

C8H17-

Octyl

C1oH21-

Decyl

C6H5-

Phenyl

I

I

c4

c6

c8

Length of the n-alkyl chain Figure 2-10. The effect of alkyl chain length and degree of substitution on binding capacity in HIC['].

Figure 2-9.

Hydrophobic ligands attached to matrix.

The surface of a protein is relatively hydrophilic in the lower concentration buffer solution, but hydrophobic: interaction increases at high ionic strength (Fig. 2-11). It is estimated that 40-50% of the surface area of a protein is non-polar. The HIC parameters are type of ligand, degree of substitution, concentration of salt, and effect of temperature and pH. The immobilized ligands used are hydrocarbon groups like butyl and octyl groups and phenyl groups. The polarity of the ligand increases with alkyl chain length and its degree of substitution. The interaction of the phenyl group is not simple because of an aromatic effect as well as hydrophobicity. The most typical salt in HIC is ammonium sulfate. As the concentration of ammonium sulfate is increased, the amount of protein adsorbed on the ligand increases linearly up to the precipitation point. The effect of the ion used in HIC the precipitation of proteins is shown in Table 2-3. Sodium, potassium or ammonium sulfates have a relatively high saltingout effect and molar surface tension of water an increasing effect. Ammonium (1M)sulfate is a good starting point for experiments. If the protein does not retain the ligand, a more hydrophobic ligand should be selected. The recovery of protein in H I C should be S O % . When a small amount of miscible organic solvent is needed, the ligand should be changed to a less hydrophobic one. Hydrophobic interaction in HIC is diminished by increasing the pH and increased by decreasing the pH. The PI of protein is in the acidic range and the hydrophilicity increases in the basic range. Hydrophobic interaction strength changes strongly at a pH below 5 or above 8.5. Also, on increasing the temperature of HIC, hydrophobicity slightly increases. A small amount of miscible organic solvent affects the decrease in hydrophobicity of protein and facilitates elution in the buffer solution.

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2 Production and lsolation of Enzymes

The principle o f hydrophobic chromatography[']. Figure 2-11.

ow

P: S: L: H: W: S:

Polymer matrix Soluble molecule Ligand attached to polymer matrix Hydrophobic patch on surface of soluble molecule Water molecules in the bulk solution Salt (ammonium sulfate)

Table 2-3. The effect of some anions and cations in precipitating proteins141.

Protein

Antichaotropic effect * Chaotropic effect

Collagen Zelatin

SO4'- < CH3COO- < C1- < Br- < NOs- < CI04- < I- < SCNNH4' < Rb', K', Na', Cs' < Li' < MgZ' < Ca" Ba"

2.7.1.3

Gel Filtration (CF)

Gel filtration"] is a key method in the purification of enzymes as well as biological macromolecules. It is reliable and simple as a separation technique without adsorption and interaction on gel filtration media. In gel filtration, the principle of separation is very simple, and macromolecules in solution are separated based on differences in their size as they pass through a column (Fig. 2-12). Large molecules pass through the stationary phase first while smaller molecules move about the gel filtration medium slowly. Gel filtration is also called molecular-sieve chromatog-

2.7 Purifcation ofEnzyrnes

0

0 Figure 2-12.

.

.

Sample proteins of different molecular size Gel particles

The principle of gel filtration.

raphy. Molecules are eluted in the order large to small. Gel filtration is usually used at the final or latter stage for changing buffer and concentration. Gel filtration is carried out using a single buffer solution of appropriate pH and ionic strength. Some gel filtration media have a small number of ionic charged groups, such as carboxyl and sulfonic groups, which sometimes cause non-specific adsorption of basic proteins at low ionic strengths. In order to avoid the adsorption, gel filtration should be carried out at an ionic strength above 0.15 M. Non-ionic interactions between proteins and gel filtration media are negligible at buffer concentrations between 0.15 M and 1.5 M. An ionic strength below 0.15 M causes a slight retardation of basic proteins and exclusion of acidic proteins. The first gel filtration medium Sephadex, provided by Pharmacia, was a beadformed gel prepared by cross-linking dextran with epichlorohydrin. Gel filtration media with various particle size grades are now available and globular proteins can be separated between 700 and 4x10' A. The fractionation range of the medium determines the porosity of the gel and is measured using typical globular biological molecules or dextrans. The shape (its diameter and length) of biological molecules affects the theoretical separation. When the molecule is not globular but a linear string, the separation is quite different.

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2 Production and /solation of Enzymes

Choice ofColumn, Sample Volume and How Rate There are some factors affecting the choice of column equipment in order to obtain a good separation. The length of column is necessary more than 30 times the diameter of column, because the resolution increases at the square root of column length. That is why a longer column is used for gel filtration especially for an analytical fractionation. A bed length of more than 1m is not useful and effective for industrial separation. The dead volume at the inlet and outlet should be less than 0.1%. A sample volume of 0.5-5% of the bed volume is recommended for good resolution and depends on the gel filtration medium. The relationship between sample volume, medium and resolution is described; however, the actual sample volume should be determined by experiment. A smaller sample size is not good for resolution. Up to 30 % of the total bed volume can be applied for changing the buffer and salting out. An effective flow rate for resolution is the order of 5 mL/cm2h and up to about 25 mL/cm2h is allowed for industrial preparations. The length of the column and flow rate are basically in an inverse relation. 2.7.1.4

Reversed Phase Chromatography

Except for a few specific applications, reversed phase chromatography (RPC)[lo]is rarely used in biological purification. RPC is commonly used for the purification of organic compounds and low molecular weight peptides. The principle of RPC is similar to that of HIC. The ligand is stronger in RPC than in HIC, and includes octadecyl, octyl, butyl and phenyl groups (Fig. 2-13). The remaining silanols are quenched with trimethylsilyl groups. RPC medium consists of hydrophobic ligands chemically bonded to porous microbeads. The microbeads are made of silica gel or a synthetic organic polymer like polystyrene.

Figure 2-13.

Reacting a silanol with octadecyldimethylsilyl group.

2.7 Purijication of Enzymes

Table 2-4.

Solvents used in reversed phase chromatography.

Solvent

Acetonitrile Ethanol Methan o1 Propanol Isopropanol Water

Diielectric constant (20 "C)

Viscosity (cP at 20 "C)

38.8 24.3 3:i.G 20.1 18.3 80.4

0.36 1.20 0.60 2.26 2.30 1.00

bP ("C) 82 78 65 98 82 100

It is necessary to use water miscible organic solvents in order to elute proteins (Table 2-4). Most proteins are apt to be denatured in that case. RPC is useful for the purification of small samples or peptides and is usually carried out as high performance liquid chromatography (HPLC). 2.7.1.5 Hydrogen Bond Chromatography

There are three types of chemical interaction between the ligands and proteins: ionic, hydrophobic and hydrogen bond interaction. Hydrogen bond chromatography is not as popular as ionic and hydrophobic chromatography.Precipitation of proteins is sometimes observed in the presence of a water-soluble polysaccharide and polyethylene glycol. The complex with proteins easily forms via hydrogen bonding at high ionic strength. Ionic cellulose, such as DEAE-cellulose and CM-cellurose, as well as cellulose, is used as a matrix for this purpose. Hydrogen bond - ion chromatography is complicated because the ionic strength of the buffer solution used for each of the two methods is opposite to elute proteins. Protein is adsorbed on DEAD-cellulose in buffer solution with 3 M ammonium sulfate and is eluted by decreasing the concentration of ammonium sulfate or adding the releasing reagents, such as urea and sucrose etc. Sodium formate and sodium acetate are used instead of ammonium sulfate. On the other hand, small amounts of ethanol, glycerol and ethylene glycol are available for elution. 2.7.1.6 Affinity Chromatography

The principle of affinity cliromatography is shown in Fig. 2-14. Affinity chromatography["] is used for a biologically specific ligand bound to the matrix. The protein binds with ligand specifically in an active form and the rest passes through without adsorption upon washing with a buffer solution. The ligand should have specificityand reversibility for the protein and release it on affinity elution or change of ionic strength and pH. Interactions between proteins and ligands include ionic binding, hydrogen binding and hydrophobic binding. The factors necessary for iortic binding have been listed before. The effect of ionic

I

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2 Production and /solation ofEnzyrnes

A A

Adsorption

Desorption

___)

00

0 0

0 U0

A

0

Figure 2-14.

Affinity absorbent

Q

If GI

Q

Q

Protein to be separated

impure proteins

The principle of affinity chromatography.

strength on ionic binding is opposite to that on hydrophobic binding and the recovery of protein is sometimes not good. The use of a 1-3 M urea solution or 5-20% sucrose is a good idea in such a case. Affinity chromatography is carried out by the batch and column methods. The procedure involves (i)equilibration of the adsorbent, (ii) preparation of sample, (iii) application of the sample, (iv) washing away of unbound materials, (v) elution and (vi) regeneration of adsorbent. When the ligand has a simple specificity for protein, about 90% purified protein is obtained by one-step purification. So, affinity chromatography is a revolutionary

2.7 Purification of Enzymes

0

NH:

\ Figure 2-15.

Immobilized dye chrornatography[ll].

purification method. Adsorbents are relatively expensive and affinity chromatography is useful for small scale purification. There are several types of affinity chromatography.Two typical types, immobilized dye chromatography and metal chelate affinity chromatography, are described.

lmnzobilized Dye Chromatography Immobilized dye chromatography is the most useful affinity chromatography. Its ligands are synthetic polycyclic dyes. These structures are very similar to the cofactors NAD’ and NADP’ as a dinucleotide analog, which are apt to bind strongly with a protein like kinases, dehydrogenases, etc. (Fig. 2-15). Some of the proteins bind biospecifically with the dye because of its stmctual similarity to NAD and NADP. Some proteins like lipoproteins and albumin bind in a less specific manner by electrostatic and hydrophobic interactions with the aromatic anionic ligand. Bound proteins can be ‘eluted by affinity elution using low concentration free cofactors. On the other hand, non-specifically bound proteins need a much higher concentrate of cofactors or ionic strength. Metal Chelate Afinity Chromatography Metal chelate affinity chromatography is a kind of separation method which has, as a ligand, a metal ion. Some proteins and peptides are purified on the basis of affinity for metal ions immobilizl2d by chelation on the adsorbents. Histidine and cysteine form complexes with the chelated metals around neutral pH. Biological proteins include many histidines as well as recombinant proteins as polyhistidine fusions: for instance, His-tag protein:; have a specific metal chelate affinity. The adsorbent is prepared by coupling a metal chelate ligand with an iminodiacetic acid group, which forms a chelate with divalent metal ions such as Zn2+,Cuz+,Cd2+,Hg”, Co2+, Ni2+, Fez+,etc. Elution is carried out by reducing the pH and increasing the ionic strength of the buffer or by adding EDTA. to the buffer. The most typical method is to gradually add sodium chloride (0.5-1.0 M) or imidazole (0-0.0sM). This ligand is very expensive, so metal chelate affinity chromatography is used only for small scale purification.

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2 Production and lsolation of Enzymes

His-tag proteins produced by a recombinant are easily purified by metal chelating chromatography. His-tag proteins have about G histidines at the N- or C-terminal site and the His-tag easily forms a chelate with Ni2+,Zn2+and Cu2+Elution of His-tag proteins is carried out by increasing the concentration of imidazole in the buffer solution. 2.7.1.7 Salting-out Chromatography

Salting out is popular for the purification of proteins. Salting-outchromatography is a precise method based on the same principle; however, it is not popular. Both positive and negative salting-out chromatography are carried out. The former is a combination of molecular-sieving chromatography and salting out. Proteins in buffer solution are applied to a concentration gradient column of salts. With the latter method, the precipitation of proteins salted out in the presence of celite is filled in the column and proteins are eluted with a flowing buffer solution by decreasing the concentration of salt. 2.7.2 Precipitation

Among the methods of purifymg protein, precipitation is the most useful and typical for both small and large scale procedures. The precipitation methods are classified into 4 types, salting-out, organic solvent precipitation, pH changing precipitation and water-solubleprecipitation. The precipitation is usually carried out early and the total protein concentration should be >0.1mg/mL. 2.7.2.1 Precipitation by Salting out

The solubility of macromolecules like proteins in water generally increases in the presence of a suitable concentration of salt, so-called salting in. Furthermore, increasing the concentration of salt further leads to a decrease in the solubility of the proteins and their precipitation (salting out). Salting out much depends on the pH and temperature of the solution. Proteins show minimum solubility around their isoelectric point (PI) in water and a little lower solubility in buffer solution, in other words, in the presence of salt. Regarding temperature, the solubility of proteins generally decreases at higher temperature in buffer solution with higher ionic strength. Many salts are used for salting out, including ammonium sulfate, sodium sulfate, potassium phosphate, magnesium sulfate, sodium citrate, sodium chloride etc. The solubility of these salts is independent of temperature, and the salts do not affect the denaturation of the proteins. Ammonium sulfate is the most effective salt for salting out because of its high solubility at any temperature and low cost. Ammonium sulfate is also a useful stabilizer for proteins.

2.7 Purification of Enzymes

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63

2.7.2.2 Precipitation by Organic Solvents

Water miscible organic solvents, such as ethanol, isopropanol and acetone, reduce the solubility of proteins b'y decreasing the dielectric constant of aqueous solution and taking away water from around the proteins. Also, these organic solvents can remove the lipids bound to a protein. Precipitation by organic solvents is affected by temperature, ionic strength and the pH of the buffer solution. Common proteins are precipitated at about 40% in ethanol, but proteins with hydrophobic surface are soluble, like lipases, under the conditions. Alcohol concentrations of 80-90 % are necessary to precipitate lipases. The concentration of protein should be >lmg/mL and that of buffer solution <SO mM. The solution and organic solvent should be cooled and the mixture kept at below 0 "C during the addition of organic solvents. 2.7.2.3 Precipitation by Changing pH

There are two types of precipitation by pH change, isoelectricpoint (PI) precipitation and acidic precipitation. PI precipitation is suitable for a protein with very low solubility and is more effective in combination with salting-out and organic solvent precipitation. Anions bind with proteins more easily than cations, so the PI of proteins shifts a little to the acidic range. On the other hand, acidic precipitation is good when the protein is stable, but impure proteins are unstable in the acidic range. 2.7.2.4 Precipitation by Water-Sohble Polymer

Precipitation by water-soluble polymer is a simple method for the purification and crystallization of proteins. Many proteins are easily precipitated in the presence of water-soluble non-ionic polymers such as polyethylene glycols (PEG 2000, 4000, GOOO), methyl cellulose, pdyvinyl alcohol (PVA) and dextrans (DEX). These water-solublepolymers take away water from around proteins. The proteins bind with these polymers via hydrogen bonds, and then the complex precipitates as a solid or sometimes becomes a viscous liquid. Hydrogen bond chromatography is based on this principle. The complex contains .water-solublepolymers, which must be removed by ionic chromatography,salting out, ethanol precipitation, electrophoresis etc.

64 2 Production and Isolation of Enzymes

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2.7.3 Crystallization

Relatively purified proteins are easily crystallized at >1%, usually 5-lo%, of the protein concentration in buffer. So, crystallization is the final stage of purification, and useful for storage of proteins and X-ray crystal structure analysis. In protein chemistry, crystallization does not mean the protein is 100% pure even though it is in crystalline form. As described for salting out, a crystallized protein is in a solid state together with precipitation aids such as salts, organic solvents, water-soluble polymers etc. Freeze drying is one of the crystallization methods; however, denaturation, deactivation, or a slight change in the three-dimensional structure of a protein is sometimes observed. It is necessary to check the stability before freeze drying. 2.7.4 Stabilization During Purification

Care must be taken not to lose the activity during purification of the enzyme after fermentation. Enzymes are affective macromolecules influenced by changing the pH, temperature, the concentration of buffer and salts, metal ions, detergents, organic solvents and so on. In order to preserve their activity, enzymes should be kept under natural physiological conditions such as a low temperature of about 4 "C, natural pH for the enzyme, physiological buffer solution and concentration, etc. Some additives for enzyme stabilization are used during purification. Mercaptoethano1 and dithiothreitol work as antioxidants and EDTA works as a chelating agent to prevent inactivation by heavy metal ions and metalloproeases. Polysaccharides like dextrin, sugars, sugar-alcohols like sorbitol and mannitol, glycerol and ethylene glycol are sometimes used as stabilizers. Some peptides and amino acids are useful excipients for purification. Compounds with a similar structure to that of the substrate are generally effective as stabilizers and are used as fillers for storage. Degradation by proteases derived from the same microorganism or from contamination during purification must be avoided. Once a protease contaminates an enzyme solution, the desired enzyme is degraded during purification and might disappear. To prevent degradation by proteases, it is helpful to add protease inhibitors like PMSF (SH protease) and EDTA (metal protease). 2.7.5

Storage of Enzymes 2.7.5.1

Storage in Liquids

Common enzymes in liquid form should be stored below 4 "C in a refrigerator and kept with a stabilizer. Most enzymes keep their activity for several years under suitable conditions, especially thermostable enzymes.

2.8 Commercial Biocatalysts

Ammonium sulfate (2 h4) is a popular storage solution for commercial porcine liver esterase (PLE). Ammonium sulfate prevents microbial growth on the solution. Storage in 50% glycerol is also useful and this glycerol stock can be stored below 0 "C. 2.7.5.2

Storage in Solids

Solid forms for storage are preferred in commercial enzymes. Generally, an enzyme is much more stable in solid form than in liquid form even without a stabilizer. A solid form for storage is prepared by precipitation with organic solvents and freeze drying or spray drying depending on the purification stage. Precipitation using an organic solvent is convenient, but the purity is not so high. Freeze drying is very useful but expensive. Spray drying is preferable for commercial enzymes. Spray drying is commonly carried out at about 140-70 "C for which the enzyme needs moderate thermostability. Stabilizers are effective to avoid loss of activity and the typical stabilizers described above are used during precipitation and crystallization. Some enzymes in solid form are very stable and can be stored at room temperature for several years without loss of activity.

2.8

Commercial Biocatalysts

Among biocatalysts, hydrolases like lipases and proteases are the most popular. There are several types of biocatalysts in commercial products. Immobilized lipases and cross-linking enzymes are briefly described in this section. The most popular immobilization method is adsorption on a carrier such as diatomaceous earth or a synthetic polymer. The advantage of this method is that the original activity of the enzyme is maintained, but the disadvantage is that the enzyme cannot be used in an aqueous solution. Lipases immobilized 011 ceramics modified with a chemical silyl reagent adsorb strongly and can be used in aqueous solutions as well as organic solvents. The activity is sometimes ten times the original and the thermostability is increased. These products can be reused more than ten times depending on conditions. Cross-linked enzymes are commercial biocatalysts and can be reused in organic solvent and aqueous solution. They are purchased as crystals derived from a single cross-linked enzyme. Some screening kits an- provided for user convenience. Main suppliers are listed in Chap. 20.

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66 2 Production and Isolation of Enzymes

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References M. P. Deutscher (ed), Methods in Enzyrnology, vol. 182, Academic Press, San Diego, 1990. 2 W. B. Jakoby (ed), Methods in Enzymology, vol. 104, Academic Press, San Diego, 1984. 3 T. Godfrey (ed), Industrial Enzymology, Macmillan Press Ltd, London, 1996. 4 T. Horio (Ed.),Theory and Practice on Enzymes and Other Proteins, Nankodo, Tokyo, 1994. 5 K. Drauz (ed), Enzyme Catalysis in Organic Synthesis, VCH, Weinheim, 1995. 6 Amersham Pharmacia Biotech (ed), Purification for Proteins: Principles and Methods, APB, Uppsala, 1999. 1

Amersham Pharmacia Biotech (ed), Ion Exchange Chromatography: Principles and Methods, APB, Uppsala, 1999. 8 Amersham Pharmacia Biotech (ed), Hydrophobic Interaction Chromatography: Principles and Methods, APB, Uppsala, 1999. 9 Amersham Pharmacia Biotech (ed),Gel Filtration Chromatography: Principles and Methods, APB, Uppsala, 1998. 10 Amersham Pharmacia Biotech (ed),Reversed Phase Chromatography: Principles and Methods, APB, Uppsala, 1999. 11 Amersham Pharmacia Biotech (ed), Affinity Chromatography: Principles and Methods, APB, Uppsala, 1999. 7

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

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3 Rational Design of Fiunctional Proteins Tadayuki /manaka and Hatuyuki Atomi

3.1

Protein Engineering

One of the ultimate goals of protein engineeringrl] is the ability to design and synthesize a biocatalyst that meets the demand of any user. The enzyme would have to satisfy the desired activity, stability, specificity and so on in each individual case. Unfortunately at the present time, although we have the means to synthesize proteins of any desired primary structure, we are still a long way away from the de novo design and synthesis of enzymes. We will need to understand the structurefunction correlation of proteins, and the principles of protein folding and interaction much better. Even a protein of average size will contain thousands of atoms, and therefore the number of possible inter-atomic interactions will be in the millions, and the number of conformations accessible to a protein grows exponentially with chain length. Although ab initio structure prediction methods (predicting threedimensional protein structures from amino acid sequences alone) are steadily advancing, accurate predictions are still fairly limited to small proteins or structural domains. A completely contradictory approach that has been, and is still now a major method to obtain an ideal biocatalyst, is to simply find it. This strategy leaves most of the work to nature. Adaptation of a wide variety of organisms to diverse environments on our planet has led to a massive collection of enzymes from which we can select. As the number of crganisms identified keeps growing, so does the number of constituent enzymes. In particular, the recent studies on extremophiles (thermophiles, halophiles etc.), have significantly broadened the range of available biocatalystsL2].Hyperthermophiles, which grow at temperatures above 90 C, provide a complete set of thermostable proteins that are sufficient to maintain life at these temperatures L31. However,this approach does have its limitations. Enzyme activities towards substrates not found in nature, or properties that are not required for life such as protein stability against organic solvents, may be difficult to find. The two methods mentioned above are the extremes, and many methods that combine the two are now being developed, or have already been applied. One

67

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I popular method is the random mutagenesis of 3 Rational Design of Functional Proteins

a gene encoding a particular protein of interest, for example a lipase. A lipase that most satisfies the demands would be chosen rationally, to try to optimize the enzyme. Random mutagenesis, or recently, DNA shufflingF4], of the lipase gene produces a collection of proteins that resemble the original lipase from which we can select. If the structure of the lipase is available, it may be possible to define the region that should be subject to mutagenesis rationally, thereby raising the fraction of improved mutants. The structure may also allow us to pinpoint rationally one or more particular residues as targets for sitedirected mutagenesis. This is usually followed by biochemical and structural evaluation of the variant protein, comparison with the wild-type protein, interpretation of the data, and then the designing of a further modification. This will have to be repeated until the desired changes in a protein are obtained. When affinity to a particular molecule can be used as a means for selection, phage display['], cell surface engineering 16], catalytic antibodies "1, the two-hybrid system [I', Profusion technology['], and so on provide powerful tools for selection of a desired peptide from a de nouo synthesized library. The simplicity of selection, which in many cases is basically the binding of the peptide to a molecule immobilized on a matrix, allows multiple cycles of mutagenesis and selection in a relatively short time. Although this methodology had been considered an advantage enjoyed mainly by affinity screening, recent high-throughput technologies have enabled rapid analyses of tens of thousands of clones for various enzyme parameters such as stability and substrate specificity. Thus it is now possible to improve various parameters of biocatalysts by which is presented in Chap. 4. this methodology, called directed evolution In the present chapter, we will focus on the more rational approaches of enzyme engineering and design. Basic techniques for site-directed mutagenesis, protein crystallization,and comparative modeling will also be introduced. Some recent, key examples of rational protein engineering will be described in a somewhat detailed manner. There are also very informative reviews in the literature["? 15-171.

3.2

Gene Manipulation Techniques in Enzyme Modification

The repertoire of recombinant gene technology allows us to manipulate foreign or heterologous genes in a genetically well understood organism. There may be a few exceptions, but from a general viewpoint, Escherichia coli is the organism of choice for protein engineering. Thanks to the general availability of easy to use cloning kits tailor-made for mutagenesis, straightforward experiments can be carried out. Even if other organisms are superior for the production of a particular enzyme, mutagenesis procedures will be carried out in E. coli. A variety of multi-purpose plasmids for mutagenesis and expression are readily available from commercial sources. PCRbased methodologies now make it possible to incorporate, clone, isolate and confirm gene mutations within a couple of days. E. coli host cells have also been dramatically improved, and an abundant collection of strains are now commercially available for various needs. Some typical techniques will be described here.

3.2 Gene Manipulation Techniques in Enzyme Modification

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Many recombinant gene expression systems have been developed in the past years. Synthesis of the prot(einis controlled at the transcriptional level as in the wellknown lac and tac systems ["]. Chemicals such as isopropyl-0-D-thiogalactopyranoside (IPTG) in the lac and tac systems are used to initiate and induce high levels of transcription. Although expression of the genes is low when an inducer chemical is not present, this basal expression may inhibit experiments when the target proteins are lethal to the host cell. Some systems overcome this problem by controlling gene expression under the control of a T7 promoter["]. This promoter is specifically recognized by T7 polymerase, whose gene is introduced into the host cell. Expression of the T7 polymerase gene is regulated by an upstream lac promoter. When IPTG is not present, very little T7 polymerase is produced, consequently leading to minimal expression of the target gene. A lac operator can be inserted in between the T7 promoter and the target gene in order to achieve higher stringency. Furthermore, the gene encoding T7 lysozyme, a natural inhibitor of T7 polymerase, can also be introduced in the host cells to reduce target gene transcription under uninduced conditions further [l']. One of the major problems one might encounter when expressing foreign genes in E. coli is the formation of inclusion bodies when the proteins produced are hostlethal, or mis-folded. Thi:j will require the unfolding of the protein with various detergents or denaturants, followed by refolding experiments. Another problem often seen is the low levels of target gene expression in the host cells when these genes contain many codons that are not frequently used in E. coli. This is due to the depletion of rare tRNA :species in the host cells. There are now commercially available host cells transformed with extra copies of argU, ileY and leuW tRNA genes to allow high-level expression of genes with rare codons. Many other strategies, such as inactivatingthe Lon protease gene in the host cell[18],have been applied in order to maximize the production of diverse recombinant proteins that may be of interest. Site-directed mutagenesis methodology has also seen many advances in the recent years. Most strategies are described in detail in reference 18. In essence they all rely on synthetic oligonucleotides which contain the desired information for a modified protein sequence, be it replacement, insertion or deletion of amino acids. Classical cassette mutagenesis techniques are available, along with newly developed strategies utilizing PCR techniques. In cassette mutagenesie, synthetic complementary oligonucleotides including the modified sequence are hybridized to form a double-stranded DNA fragment. This fragment should span a region including two appropriate restriction enzyme sites on opposite sides of the mutation. It is then easily possible to exchange the native sequence with the mociified sequence after restriction enzyme digestion and ligation. When only a pariicular mutation is required, PCR-based methods should be less tedious and faster. However, when mutations span a relatively long region, or require completely different nucleotide sequences compared with the original gene, or when random sequences are to be introduced into a particular region, this classical method still has its advantages. To introduce mutations into genes via PCR four instead of the normal two primers are needed. The so-called outer primers bind at the beginning and the end of the

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target region, while the inner primers bind to the site where mutations are desired and consist of the modified base information. To obtain the complete sequence with the mutation, two rounds of PCR must be performed. The first reaction amplifies the DNA fragment from the beginning to the mutated region, the second one from the mutation to the end. Both PCR products carry the mutation either at the beginning or at the end of the DNA fragment. After purification of both PCR products the two amplificates are mixed, denatured and then the PCR will be started (without primers). To amplify the whole sequence with the mutation the outer primers are added and the PCR is started again. During the PCR both the mutated strand and the natural strand are amplified but after only a few cycles will there be much more DNA harboring the mutation. There are now also methods that can efficiently introduce mutations in a single PCR run (Fig. 3-1). The procedure utilizes a double-stranded DNA plasmid with the target gene isolated from a dam" E. coli strain. Two complementary oligonucleotides with the desired nucleotide substitutions are used for PCR along with the plasmid as a template. The product is a mutated plasmid with staggered nicks. This is treated with DpnI, an endonuclease specific for methylated and hemimethylated DNA. As dam+ strains methylate plasmid DNA, the parental strain harboring the original gene will be susceptible to DpnI treatment and digested, markedly enhancing the efficiency of mutant gene isolation. The interesting part begins when comparing the original enzyme with the mutant enzyme. Thus it is advisable to run the experiments in parallel. To interpret the results various options have to be considered. Either the enzyme activity is unchanged (a so-called silent mutation) or it is changed for the better or worse. The interpretation of these changes poses a serious problem. First it has to be asked whether the mutation has altered the overall enzyme topology or whether it influenced only the local geometry. Thus besides the usual kinetic analysis some structural determination is advisable. To date X-ray crystallography and NMR spectroscopy have given the most detailed picture, CD or IR spectroscopy are of less value.

3.3

Protein Crystallization

The three-dimensional structure of a protein is the most powerful basis from which a rational approach can be taken to modify a protein. When the structure of a highly homologous protein has been determined, one may attempt to obtain structural information by comparative modeling, or homology modeling. However, the reliability of a model is questionable when similarities of the compared proteins are not high, and we are almost helpless when a structurally novel protein is the one of interest. Although rapid progress is being made in the use of NMR spectroscopy,the orthodox methodology in elucidating a protein structure is still protein crystallization and X-ray diffraction. As detailed explanations of both methodologies appear in the literature[1'], we will just touch on some points concerning protein crystallization.

3.3 Protein Crystallization

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extension using a thermostable DNA polymerase

digestion of the parental, methylated DNA template with Dpn I

introduction into competent cells and sealing of the nicks

Figure 3-1. A rapid method to introduce mutations into a target gene. Thick lines represent the metiylated plasmid DNA harboring the wild-type target gene.

Although there exists a vast number of protein structures in the databases, there is as yet no rational procedure to crystallize a particular protein. The procedure is still mainly based on a trial arid error approach. The crystallization process itself is one of which the protein is slowly and orderly precipitated from a solution. As a general rule, the purity of the protein is the most important factor to be dealt with before attempting to crystallize a protein. If possible, care should be taken not only to remove contaminant proteins, but also to remove any structurally heterologous populations in the purified protein sample. This may be achieved by discarding tail

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fractions after chromatography. Ideally, all molecules of the protein should have identical surface properties, especially in terms of charge distribution, as this will influence the packing of the molecules in the crystal. SDS-PAGE (sodium dodecyl sulfate-polyacrylamideelectrophoresis), often used to display the homogeneity of a purified protein in biochemical studies, will not always provide sufficient information. Mass spectrometry can be recommended for a detailed examination of protein homogeneity. After the purity of a protein sample is confirmed, an appropriate solvent and precipitant must be chosen. Again, there are no rational means in deciding these components. Solvents are usually a water-buffer solution, and detergents or organic solvents may be added when necessary, such as in the cases of membrane proteins or lipases with a relatively hydrophobic surface. Typical precipitants are polyethylene glycol (PEG), ammonium sulfate, sodium or lithium chloride salts, or 2-methyl2,4-pentanediol (MPD).As multiple parameters must be considered, the search for an optimal crystallization condition may be complicated and tedious. Alternatively, convenient kits with an array of ready-to-usesolutions including various buffers and precipitants are commercially available. One may screen for an appropriate crystallization condition with these kits, and then optimize the conditions based on the results. Various techniques can be applied to crystallize proteins. Vapor diffusion using the hanging drop method is one of the popular ways to obtain crystals. In this method, a sample solution of 2-5 pL of protein solution is placed on a siliconized microscope cover glass. The same volume of precipitant solution is mixed, forming a small drop on the glass surface. The glass is then placed on a well, with the drop hanging down from the glass. Prior to this, 1 mL of the precipitant solution is poured into the bottom of the well, so that the surface does not make contact with the hanging drop. Vaseline or grease should be applied to the rims of the wells, so that an air-tight chamber is made when positioning the cover glass. In this example, the concentration of precipitant in the well is twice that of the drop. Equilibrium is reached by vapor diffusion, and the precipitant concentration in the hanging drop will gradually increase, possibly leading to crystallization. The sitting drop method can also be applied when there is a surface separated from the precipitant solution in the well. Drops are placed on the surface, and the chamber is sealed. Other methods include batch crystallization, liquid-liquid diffusion, and dialysis. Approximately 20 pg of protein are used in a single screen, therefore 50 to 100 tests will require roughly 1 to 2 mg of purified protein. In Fig. 3-2, some examples of protein crystal^[^^^^] are shown. Figure 3-2 A and B provide a good example of different crystallization conditions of a single protein, archaeal 06-methylguanine-DNA methyltransferase, leading to distinct forms of crystals [20, 21]. Crystals of archaeal . shown in Fig. 3-2 C and D, DNA polymerase[22,231 and archaeal R ~ b i s c o [251~ ~are respectively.

3.4 Comparative Modeling ofa Protein Structure

Figure 3-2. Crystals o f various proteins from the hyperthermophilic archaeon, Thermococcus kodakaraensis KODl. A, rod-like crystal of O6-methylguanine-DNA methyltransferase (MCMT); 6,plate-like crystal of MCMT; C, bar-like crystal o f DNA polymerase; D, cubic or hexagonal

crystals o f ribulose 1,s-bisphosphate carboxylase/oxygenase (Rubisco).

3.4 Comparative Modeling of ;a Protein Structure

Comparative modeling, or homology modeling, is the most powerful tool when a rational approach is taker1to engineer a protein with an unknown three-dimensional structure (the target protein). Through comparative modeling, the three-dimensional structure of the target protein can be predicted based on its alignment to one or more proteins of kncwn structure (the templates). The rapid accumulation of known protein structures and the advances in modeling software have significantly increased the accuracy of comparative modeling. It is now possible to model, with sufficient accuracy, significant parts of one third of all known protein sequences. Detailed and informative reviews[26-30] can be found in the literature, and we will only present a general overview in this chapter. In order to predict a structural model, similarity between the primary sequences of the target and template(!;)must be detectable. Furthermore, an accurate alignment of the two or more sequences must be calculated. If these requirements are met, one may proceed to model the target. The process of comparative modeling can be divided into four steps: (i) fold assignment and template selection, (ii) targettemplate alignment, (iii) model building, and (iv) model evaluation.

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3 Rational Design ofFunctional Proteins

Templates can be selected using the target sequence as a query for searching protein structure databases [e.g. Brookhaven Protein Data Bank (PDB): h q : / /www.rcsb.org/pdb/index.html;Structural Classification of Proteins (SCOP): scop.mrc-lmb.cam.ac.uk/scop/; DALI: www2.ebi.ac.uk/dali/; Class, Architecture, Topology and Homologous superfamily classification at CATH: www.biochem.uc1.ac.uk/bsm/cath/). Methods for protein comparison can be divided into three types. BLAST and FASTA represent the first type, where the target sequence is independently compared with each sequence in the databases, using pairwise sequence-sequence comparison. The second type is represented by PSI-BLAST, which expands the set of homologs of the target sequence. In a PSI-BLAST search, an initial set of homologs against the target sequence is collected, aligned with the target sequence, and a position-specific scoring matrix is constructed from the alignment. This matrix is then used to cany out another search for new homologs, and this is subsequently repeated until no new homologs are identified. It has been reported that PSI-BLAST identifies homologs of known structure for approximately twice as many sequences than a BLAST search. The third type of search is the 3D template matching method. The target sequence is threaded through a library of known three-dimensional protein folds, and a structure-dependent scoring function predicts the suitability between the protein and the fold. This method is useful when homologs of the target sequence cannot be found in terms of primary structure comparison. After a collection of candidate templates is obtained, one should take into account the relationship of each template to the target, the quality of the templates, and other factors (e.g. the presence of convenient protein-ligand structures) before choosing the template(s) to be used for alignment and modeling. Comparisons of the relationships between protein sequences can be determined by constructing a phylogenetic tree among the candidates [CLUSTALW at European Bioinformatics Institute (EBI): http://www.ebi.ac.uk/clustalw/ or DNA Data Bank of Japan (DDBJ): http://www.ddbj.nig.ac.jp/htmls/E-mail/clustalw-~.html]. The CLUSTAL programs can be further used for target-template sequence alignment. When protein sequences display over 40 % identity, the alignment is usually correct. When sequence identity is below 20%, multiple template structures should be used in order to identify specific regions or secondary structures that can be used as “guides” to construct an accurate alignment. Once a target-template sequence alignment has been constructed, a variety of methods and software is available for model building. Modeling methods are based on rigid-body assembly, by segment matching or coordinate reconstruction, or by satisfaction of spatial constraints. We will not introduce the details of each method, and readers should refer to the indicated literature. When used optimally, all three methods usually give similar results. Furthermore, the accuracy of the alignment used in modeling is crucial, as no current comparative modeling method can compensate for an incorrect alignment. On the other hand, the evaluation of a model is usually more reliable than the evaluation of an alignment. Therefore, when a choice among candidate alignments is difficult, one should generate models from each alignment, and choose the most promising one by evaluating the integrity of

3.5 What is Needed to Take a Rational Approach?

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the three-dimensional models. Information concerning modeling programs such as Insight I1 and QUANTA are available at http://www.accelrys.com, SYBYL/Base at www.tripos.com, and Internal Coordinate Mechanics (ICM) software at http:/ /www.molsoft.com. A detailed list of databases and software programs is shown in reference [261. Various programs are available to the public at the ExPASy Molecular Biology Server (http://wwvv..expasy.ch/). After generating a structure model, it is essential to evaluate its accuracy before one can use it for rational purposes. A wide variety of programs and servers exist that can be used to evaluate A model. The BIOTECH (biotech.embl-ebi.ac.uk:8400/), ERRAT and VERIFY3D (http://www.doe-mbi.ucla.edu/Services/), and PROVE (http://www.ucmb.ulb.ac.l)e/UCMB/PROVE/) are servers available to the public. Model evaluation programs check various features of the model, including bond length, bond angles, main-chain and side-chain torsion angles, peptide bond and side-chain ring planarities, chirality, and clashes between non-bonded pairs of atoms. After the evaluation step, the generated structure model is ready for use as a basis for site-directedmutagenesis. The information obtained from the biochemical evaluation of variant proteins will also contribute to improvements in the model for the rational design of muiants.

3.5 What is Needed to Take a I?ationalApproach?

It is quite obvious that the more information that is available on a particular protein, the easier it is to take a raiional approach to improving its performance. Biochemical analyses of a protein provides valuable information in terms of its activity, specificity, and stability under various conditions. Kinetic analysis of the enzyme reveals the kinetic mechanism of the reaction, in other words the order in which substrates enter and products are released from the enzyme. This gives us an idea as to what types of intermediates or complexes may be formed during the reaction. Cloning and sequencing of the genes provides the primary structure of the protein. The number of sequences available in public databases (e.g. GenBank/EMBL/DDBJ for genes, SwissProt and PIR for proteins) is enormous, and readily available (Entrez protein or nucleotide sequence search at NCBI; http://www.ncbi.nlm.nih.gov/entrez/query.fi:gi). Comparative analysis of these sequences, along with their biochemical properties, may in some cases provide enough information to modify a protein rationally through site-directed mutagenesis. Furthermore, there are now an ever increasing number of three-dimensional structures of enzymes in the databases, and these provide us wiih the precise architecture of various enzymes. Along with advances in protein crystallization methods and X-ray diffraction technology, rapid progress has also been made in solving protein structures by alternative tools, such as NMR spectroscopy. Using the comparative modeling mentioned above, it is also possible to predict the three-dimensional structure of a protein using the determined structure of a closely related protein. Modeling software can also calculate and predict in silico the local :structuralchanges of a protein after site-directed mutagene-

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sis, opening the way for rational protein design. In the following sections, we will introduce some successful examples of rational improvement or alteration of enzyme biocatalysts based on various degrees of available information.

3.6

Examples of Protein Engineering 3.6.1

Protein Engineering Studies: Providing a Rational Explanation for Enzyme Specificity

Tyrosyl-tRNA-synthetase from Bacillus stearothermophilus is an enzyme (MW = 2 x 47 500 Da) that as a dimer catalyzes the aminoacylation of tRNATrwith tyrosine in the following two steps.

E + Tyr + ATP = E.Tyr-AMP + PPi E.Tyr-AMP + tRNA = Tyr-tRNA + AMP This enzyme was one of the first and best case studies of protein engineering. Based on the X-ray structure, there are eleven hydrogen bond contacts between the protein and its reaction intermediate tyro~yl-adenylate[~'I. Of these, eight are hydrogen bonds involving amino acid side chains, the remaining three are with backbone C = 0 or N - H. A great deal of detailed information about the performance of these contacts has been obtained. Thus the amino acids Tyr 34, Cys 35, Gly 36, Asp 38, His 48, Thr 51, Tyr 169, Gly 192 and Gln 195 are the relevant hydrogen bond partners for the enzyme-bound intermediate (Fig. 3-3). The contributions that these individual hydrogen bonds can make have been (Table 3-1). Thus, deletion of a hydrogen bond donor or acceptor weakens the substrate-binding energy by 0.5-1.5 kcal mol-'. Deletion of the charged Asp 38 that binds to the substrate lowers the binding energy by -4 kcal mol-'. Based on Michaelis-Mentenkinetics, this amounts to the following barrier:

AG*T = RT(ks T/h) - RTln(kca,/KM) kB = Boltzmann constant h = Plancks constant and BAG for the mutant enzyme (mut) compared with the wild-type enzyme (wt): AA G*T = Rnn{ (kcat/ &)mut/ (kcat/ &I),} Hydrogen bonding energies in the range of 0.05-1.5 kcal mol-' give discrimination rates in kCa,/KM from 2-fold to 12-fold whilst 4 kcal mol-' amounts to 1000-fold discriminations. The latter point explains nicely the enormous specificity for tyrosine as compared with phenylalanine. Asp 176 in the binding pocket binds to the phenolic hydroxyl group of tyrosine (Fig. 3-3).Thus from the large amount of data presented we can learn about the binding energy of hydrogen bond donors and acceptors in proteins: deletion of a side chain between the enzyme and substrate to leave an unpaired, uncharged, hydrogen donor and acceptor weakens the binding energy by only 0.5-1.5 kcal mol-'. However, the presence of an unpaired and

3. G Examples of Protein Engineering

Figure 3-3. adenylate.

I

Hydrogen bonds between the tyrosyl-tRNA synthetase and tyrosyl

Table 3-1. Relative binding eiiergies of groups in tyrosyl-tRNA synthetase infered from comparison between mutant and wilde-type enzymes at 298 K. Compared residues and their numbering

Phe 34 Gly 35 Ala 51 Gly 48 Gly 48 Ser 35

Phe 169 Gly 195 Gly 35

Ala 51

Tyr 34 cys 35 cys 51 Asn 48 His 48" cys 35" Tyr 169" Gln 195" Ser 35 Thr 51"

Substrate

TYr ATP ATP ATP ATP ATP Tyr TYr ATP ATP

MGf (kcal mol-')

-

0.52 1.14

0.47 0.77 0.96 1.18

3.72 4.49 - 0.04 - 0.44

* Residues found in the wild-type protein

charged donor or acceptor weakens binding by a further 3 or more kcal mol-'. These values are much lower than the absolute strength of hydrogen bonds in vucuo and are the consequence of hydrogen bonding in aqueous solution being an exchange process.

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B

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1

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to

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20

Time (min)

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Figure 3-4. (A) Structure o f thermolysin from Bacillus thermoproteolyticus used t o determine candidate residues for site-directed mutagenesis o f t h e neutral protease from B. stearothermophilus. The structure was visualized with Ras-Mol software and slightly modified. Data were from M M D B ID: 3638, PDB ID: lLNF, submitted by Holland, and Matthews. The two domains are indicated with the bars on the left, and the Cly 144 residue is highlighted i n black. (B) Themostability o f t h e wild-type neutral protease (open circles), and the C144A variant (closed circles).

3.6.2

Enhancing the Themostability o f Proteases

An increase in thermostability of a neutral protease from Bacillus stearothemophilus (NprT) was achieved from a rational approach by comparing its sequence with the thermostable thermolysin from B. t h e m o p r o t e o l y t i ~ u s [ The ~ ~ ] . enzymes were 85 % identical, while the thermostability of NprT at 75 C was significantly lower than that

3.6 Examples of Protein Engineering

of thermolysin. Taking into account the statistical data of various amino acid substitutions that increase thermostability, and the three-dimensional structure of thermolysin, a single mutation G144A was chosen as a candidate to increase the thermostability of NprT. The glycine residue was supposed to be located in an a-helix that connected the N- and C-terminal domains of the enzyme (Fig. 3-4A). The mutation was expected to stabilize the a-helix, and increase internal hydrophobicity of the enzyme. Furthermore, the G144A mutation introduces only a small methyl group, minimizing any structural or functional interruption that may be caused by introduction of a new side chain. Indeed, this single mutation led to a significant increase in the thermostability of NprT (Fig. 3-4B). This is a good example of the fact that an increase in internal hydrophobicity of an enzyme and stabilization of a secondary structure a-helix leads to an increase in the thermostability of a protein. 3.6.3 Contribution of Ion Pairs to the Thermostability of Proteins from Hypertherrnophiles

Proteins found in hyperthermophiles display an astonishing resistance to thermal denaturation. Some are :stable for hours or even days at temperatures near to the boiling point. This has attracted much attention as these proteins are promising candidates themselves as stable biocatalysts, and also provide valuable hints to the understanding of the mechanisms of protein thermostability. The present authors have pursued attempts to elucidate the three-dimensional structures of various proteins from the hyperthermophilic archaeon Thennococcus kodakaraensis KOD1. These include DNA polymerase[22s231, homing endonuclease I1 [34], 0'-methylguanine-DNA methyltransferase (MGMT)[20s 211, aspartyl-tRNA synthetase[351, and ribulose 1,s-bisphosphate carboxylase/oxygenase (Rubisco) [24, 251. MGMT repairs alkylated DNA by suicidal alkyl transfer from guanine 0' to its own cysteine residue. We determined the three-dimensional structure of MGMT from T. kodakaraensis KODl (Tk-MGMT) at 1.8 A resolution[211.This structure was compared with its counterpart from Escherichia coli (AdaC, C-terminal fragment of Ada protein). It has been reported that helical conformation is stabilized by (i+ 4) or (i+ 3 ) glutamate-lysineintra-helixion-pairs in a short model peptide. We observed seven intra-helixion pairs in Tk-MGMT, while none were detected in AdaC. It is presumed that these intra-helix ion-pairs contribute to reinforcement of the stability of the a-helices. Furthermore, four extra inter-helix ion-pairs not found in AdaC were observed in the interior of Tk-MGMT, stabilizing the internal packing of the tertiary structure. The structure of Tk-MGMT strongly indicates that intra-helix and inter-helix ion-pairs provide a major contribution to the thermostability of the protein. As the importance of ion-pairs toward protein thermostability has been stressed in many cases, addition or removal of an ion-pair should have significant effects. A clear example is provided by mutagenesis studies of glutamate dehydrogenase from ?: kodakaraensis KODl (Tk-GDH)r3'1. The GDH from Pyrococcusfirriosus (P'GDH) and Tk-GDH are 83% identical in terms of primary structure. However, while ' P GDH displays a half-life of 12 h at 100 C, that of Tk-GDH is 4 h. The three-

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3 Rational Design ofFunctional Proteins

A

7

30 40 50 60 70 80 90 100 110

Temperature (“C)

Time (h)

Figure 3-5. Temperature profile (A) and thermostability ( 6 ) o f glutamate dehydrogenase from T. kodakaraensis KODl and its mutants. Data o f t h e wild-type enzyme (circles), the T138E

mutant protein (squares), and the El58Q mutant protein (triangles) are shown.

dimensional structure of P’GDH has been determined, and exists in a stable hexameric form. A structural model of Tk-GDH was constructed based on the structure of P’GDH. A difference was observed between the two structures at the monomer-monomer interface. In P’GDH, there is a large ion-pair network comprised of six residues, Arg 35, Asp 132, Glu 138, Arg 164, Arg 165, and Lys 166. Glu 138 is located at the center of the network, interacting with Arg 165 and Lys 166. In the case of 2%-GDH,Glu138 was replaced by a threonine residue. When a T138E mutation was introduced into Tk-GDH,an increase in both thermostability (2 to 3 h at lOOC) and optimal temperature (80 to 85C) was observed, confirming the importance of ion-pair networks (Fig. 3-5). At one of the two-fold axes of the proteins, Glu 158 is at the center of another ion-pair network, interacting with Arg 124 and Arg 128. An El58Q mutation would interrupt this network, and is presumed to destabilize the protein. As expected, the El58Q mutant protein of TkGDH displayed a lower optimal temperature for activity (80 to 60 C), and decreased thermostability (2 h to 50 min at 100 C, Fig. 3-5). 3.6.4

Thermostability Engineering Based on the Consensus Concept

The examples mentioned above have shown that in many cases, sequence comparisons between two homologous enzymes with different thermostabilities provide valuable clues as to the how to increase protein thermostability rationally. An interesting observation has recently been made that even a set of amino acid sequences of homologous, mesophilic enzymes provides sufficient information to

3.6 Examples of Protein Engineering

I

allow the rapid design of a thermostabilized variant of the family of enzymes[37]. Using myo-inositol hexakisphosphate phosphohydrolase (phytase) as the target enzyme, a sequence alignment of 13 homologous fungal phytases was used to calculate a consensus amino acid sequence. An amino acid that has already been proven to fit into the structure of at least one of the homologous enzymes used in the alignment is chosen as a (consensusresidue. A synthetic gene, corresponding to the consensus phytase sequence was expressed and the recombinant protein, consensus phytase-1, was characterized. Differential scanning calorimetry revealed that consensus phytase-1 displayed an unfolding temperature (T,) of 78.0 C, which was 15-22C higher than the T, values of all parent phytases used in its design. Furthermore, by including six more sequences in the alignment, a refined consensus sequence was calculated (consensus phytase-10). Consensus phytase-10 displayed even higher thermostability, with a T, value of 85.4 C. Further optimization through site-directed mutagenesis eventually led to consensus proteins with unfolding temperatures of up to 90.4 C. When the effects of individual substitutions were evaluated, all single mutations affected the thermostability by less than 3 C. This suggests that the increases in stability observed in the consensus phytases were due to the combination of multiple amino acid exchanges distributed over the entire sequence of the protein. Remarkably, in spite of the increase in thermostability, catalybc activity at 37 C was not compromised. Although further examination with other proteins will be necessary, the consensus concept may provide a powerful alternative as a means to enhancing the thermostability of proteins when the information available is limited. 3.6.5

Changing the Optimal pH o f an Enzyme

Various thermostable alcohol dehydrogenases have been studied for use in the industrial production of alcohol. Based on the three-dimensional structure of horse liver alcohol dehydrogenase and a multiple sequence alignment of alcohol dehydrogenases from variou:; sources, the optimal pH of a thermostable alcohol dehydrogenase (ADH-T) from Bacillus stearothermophilus NCA 1503 was rationally The amino acid residues responsible for the catalytic activity ofhorse liver ADH had been clarified on the basis of its three-dimensional structure. As the catalytic amino acid residues were fairly conserved in ADH-T and other ADHs, ADH-Twas presumed to harbor the same proton release system as horse liver ADH, and confirmed by site-directed mutagenesis. In ADH-T, catalysis was showri to be performed by a proton release system involving a zinc-bound water molecule, a hydroxyl group of Thr 40, and an imidazole ring of His 43 (Fig. 3-6)[391.Cys 38, which interacts with the zinc ion, along with Thr 40, and His 43 were the targets for site-directed mutagenesis, and C38S, T40A, T40S, and H43A mutants were produced. The C38S, T40A, and H43A mutations completely abolished the activity of ADH-T, while the T40S mutant displayed a slightly lower activity than the wild-type enzyme. As the pK, value of His 43 was presumed to play an important role in proton release, an H43R. mutation was incorporated in order to alter the optimal pH

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Active site zinc Water

\

i!! /

H -0

I

Figure 3-6. Mechanism for the proton release system of ADH-T. The

I

H-

-

I

H

H

1 R -0

NAD+

system is composed of a zinc-bound water molecule, and the side chains of residues Thr 40 and His 43. Proton release is induced by NAD' binding.

H

R

H

H I

H+

of the enzyme. As expected, the optimum pH of the mutant enzyme H43R was shifted from 7.8 (wild-type enzyme) to 9.0. Furthermore, at the optimum pH, the H43R enzyme exhibited a higher level of activity than the wild-type ADH-T. 3.6.6

Changing the Cofactor Specificity of an Enzyme

Nicotinamide adenine dinucleotide (NAD') and nicotinamide adenine dinucleotide phosphate (NADP') are ubiquitous redox cofactors involved in a huge variety of enzyme reactions. The two are similar in structure, with NADP' harboring a single additional phosphate group esterified to the 2'-hydroxyl group of the AMP moiety. However, enzymes found in nature usually display a clear preference for one of the two cofactors, providing an interesting example of molecular recognition by enzymes. Many studies have addressed this subject, and it is now possible to engineer and switch rationally the cofactor specificities of particular enzymes from NADP' to NAD' and vice versa. Pioneering work has been carried out with glutathione reductase, a member of the highly homologous flavoprotein disulfide oxidoreductase family[40].Most members of this enzyme family utilize NADP' as a cofactor with one exception, the NAD'-dependent dihydrolipoamide dehydrogenases. Using the three-dimensional structure of glutathione reductase from human erythrocytes (HGR), and sequence alignment of various enzymes in the flavoprotein disulfide oxidoreductase family (Fig. 3-7), the cofactor specificity of glutathione reductase from E. coli (E-GR,gor gene product) was switched from NADP' to NAD'. From the structure of H-GR, a Pap "fingerprint" motif was found in the NADP-binding domain ofthe enzyme. Two arginine residues in H-GR,Arg 218 and Arg 224, bind to the 2'-phosphate group of the NADP' molecule. These residues are conserved in virtually all NADP'-dependent enzymes in the flavoprotein disulfide oxidoreductase family, but not in the NAD'-dependent dihydrolipoamide dehydrogenases. Substitution of each corresponding arginine residue in E-GR (R198M, R204L) or both,

3.6 Examples ofprotein Engineering E-GR H-GR P-GR S-MR P-MR T-TR

114 194 172 251

E-DD

180 GGG ILGLEMGTVYHALG---SO I DWEMFDOVIPAAD 183 GGGYIGIELGTAYAhFG---TKVTILEGAGEILSGFE 209 GGGI IGLEMGSVYSRLG---SKVTVVEFQPQIGASMD 220 GAGv IGVELGSWQRIG---ADVTAVEFLGHVGGVG I

6-DD Y-DD H-DD

I

211 195

* * *

*

*

**

*

pj AD+

Sequence alignment of various enzymes in the flavoprotein disulfide oxidoreductase family. The sequences of the NADP’dependent enzymes are the glutathione reductase from E. coli (ECR), human (H-CR), Pseudomonas aeruginosa (P-GR), mercuric reductase from Staphylococcus atireus (S-MR), P. aeruginosa Tn 501 (PCR), and trypanothione reductase from Trypanosoma congolense (T-TR). The NAD+-dependent enzymes are dihydrolipoamide dehydrogenase from E. coli (E-DD), B. stearothermophilus (B-DD), yeast (Y-DD), and human (H-DD). Residue positions marked with an asterisk correspond to those that were targets o f site-directed mutagenesis in the text. Figure 3-7.

resulted in a modest fall in the kcat value of the NADP+-dependentactivity, but caused a large increase in KM toward NADP+ (-25-fold). Drastic effects were not observed for NAD+-dependentE-GR activity. Further mutations were introduced, focusing on the G-X-G-X-X-Gmotif, found in various NAD+-dependentdehydrogenases, including dihydrolipoamide dehydrogenase. In NADP+-dependent enzymes, including H-GR and E-GR, the third Gly residue is usually replaced by an Ala residue (Ala 179 in E-GR).Another Ala residue is also conserved four resi~duesfurther toward the C-terminus in NADP+-dependent enzymes (Ala 183 in E-GR), but substituted by a Gly residue in dihydrolipoamide dehydrogenases. The A179G mutation in E-GR led to a dramatic decrease in the KM toward NAD’ (-40-fold), with little change in the kcat value. The A183G mutation had little effect towards N AD+-dependentactivity. Another set of mutations were introduced centered on the Val 197 residue of EGR. In the NAD+-dependent dihydrolipoamide dehydrogenases, this residue is replaced by a Glu residue, whose negative charge interacts with the 2’-hydroxyl group of NAD’ via a hydrogen bond. In order to generate such interaction in E-GR, a V197E mutation, along with K199F and H200D mutations to remove residual positive charges that may interact with the 2’-phosphate group of NADP’, were introduced. The mutant protein with seven mutations, A179G/A183G/V197E/ R198M/K199F/H200D/R204P displayed a - 250-fold decrease in kcat/KM value for NADP+-dependentactivity, while that for NAD+-dependentactivity increased by a factor of - 70. The ratio of these two contrasting shifts is 18 000, indicating that the cofactor specificity of the enzyme was rationally switched. As all mutation sites ‘chosen in this study are limited to the pap “fingerprint” motif, the strategy appl led is applicable to other NAD+- and NADP+-dependent dehydrogenases. Indeed, a systematic replacement of amino acid residues in the pap “fingerprint” motif in tht? NAD’-dependent dihydrolipoamide dehydrogenase from E. coli converted its cofactor specificity from NAD+ to NADP+L4*]. A similar strategy

83

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l has been successfully applied

3 Rational Design of Functional Proteins

on inverting the cofactor specificity of NAD+-dependent malate dehydrogenase from Themusflavus, using the crystal structure of the NAD+-dependentporcine enzyme and alignment with the NADP+-dependentenzyme from chloroplasts[421. The engineered mutant protein displayed a 1000-fold improvement toward NADP' and a 600-folddecrease in efficiency with NAD'. Other key examples have been shown with decarboxylating dehydrogenases, isocitrate dehydrogenase (IDH)[43, 441 and isopropylmalate dehydrogenase (IMDH)[451. Although these enzymes do not bind the nucleotide cofactors in the pap binding motif mentioned above, conversion of an NADP'-dependent IDH into an NAD+-dependent enzyme (850-foldpreference) has been achieved[43].Engineering the secondary structure of NAD+-dependentIMDH from Themus themophilus led to a 1000-fold preference for NADP+L4']. 3.6.7 Changing the Substrate Specificity of an Enzyme

Recently, there have been an increasing number of reports where rational mutageneses of enzymes led to a dramatic change in their substrate specificity. One example is the study on cucumber linoleate 1 3-lipoxygenaseL4'1. Lipoxygenases constitute a family of non-heme, iron-containing dioxygenases catalyzing the regioand stereoselective dioxygenation of polyenoic fatty acids to form hydroperoxy derivatives. Enzymes from plants are classified into 9- and 13-lipoxygenasesaccording to their positional specificitytoward linoleic acid oxygenation. Multiple sequence alignments and structural modeling of enzyme-substrate interaction suggested that a single residue, His 608, played a key role in the regiospecificity of the 13-lipoxygenase. An H608V mutation was introduced, and resulted in an enzyme variant with specific 9-lipoxygenaseactivity. This was elegantly explained by the fact that an H608V mutation enables a positively charged guanidino group of Arg 758, masked by the bulky His 608 residue in the wild-type enzyme, to interact with the carboxyl group of the substrate linoleic acid. This interaction forces a reversal of the substrate in the active site. This explanation was strongly supported by the observations that an R758L/H608V double mutant protein exhibited a lower reaction rate and random positional specificity. Furthermore, the drastic alteration of positional specificity was not observed when substrates lacking a free carboxyl group were examined. Another example deals with the mammalian 3a-hydroxysteroiddehydrogenase14'1. Mammalian hydroxysteroid dehydrogenases convert potent steroid hormones into their cognate inactive metabolites and belong to the aldo-keto reductase superfamily. Although 3a- and 20a-hydroxysteroid dehydrogenases display 67 % amino acid sequence identity with one another, they differ in their regiospecificity and stereospecificity. 3a-Hydroxysteroid dehydrogenase converts 5-dihydrotestosteroneinto 3-androstanediol, while 20a-hydroxysteroid dehydrogenase converts progesterone into 20-hydroxyprogesterone, the two enzymes catalyzing the formation of secondary alcohols on opposite ends of steroid hormone substrates. The crystal structure of 3a-hydroxysteroid dehydrogenase complexed with testosterone indicated that 10 residues located on 5 loop structures were involved in the enzyme-substrate

3. G Examples of Protein Engineering

interaction. Multiple sequence alignment of various hydroxysteroid dehydrogeriases displayed that G of these 110 residues were substituted in the 20a-enzyme. Single and multiple replacements of the 3a-enzyme residues to the 20a-enzyme residues did not lead to an alteration in regiospecificity. However, when individual loops were exchanged, a drastic chan:gein regiospecificitywas observed. An exchange of loop A led to a protein variant with both 3a- and 17P-hydroxysteroiddehydrogenase activity. A double exchange of loops A and C resulted in 3a- and 20a-activity. Finally, a triple exchange of loops A, B and C completely converted the specificity of the enzyme into a stereospecific 20a-hydroxysteroid dehydrogenase with a resultant shift in kcat/ KM for the desired reaction of'2 x lo1'. 3.6.8

Changing the Product Specificity of an Enzyme

A rational approach can also be used to change the product specificityof an enzyme. Prenyl diphosphate synthases catalyze the condensations of isopentenyl diphosphate with allylic diphosphate to give linear hydrocarbons of various lengths and different stereochemistries. Heptaprenyl diphosphate synthase from B. stearothemophilus is a member of the medium-chain prenyl diphosphate synthases. The enzyme catalyzes the consecutive condensation of isopentenyl diphosphate with allylic diphosphate to produce (all-E)-C35 prenyl diphosphate as the ultimate product. The product specificity of short-chair1 prenyl diphosphate synthases has been shown to be regulated by a structure around the first aspartate-rich motif (FARM). Component 11' of heptaprenyl diphosphate synthase also harbors a FARM, suggesting that this structure in component 11' may also regulate elongation in this enzyme. Via sitedirected mutagenesis, a relatively bulky isoleucine residue eight positions before the FARM, was substituted by a small glycine residue (I7GG variant). As anticipated, the I7GG variant catalyzed condensations of isopentenyl diphosphate beyond the native chain length of C35. Furthermore, two small residues Ala79 and Ser80 were individually replaced with the bulky tyrosine and phenylalanine, respectively (A79Y and S80F variants). In contrast to the I7GG mutation, these variants mainly yielded a C20 product. The study demonstrates that in the wild-type enzyme, the elongation reaction is precisely blocked at the length of C35 by the bulky Ile 76 residue, and that the degree of elongation can be controlled by removal or introduction of a bulky residue in the enzyme (Fig. 3-8). A similar approach can be utilized with the geranylgeranyl diphosphate synthase from Sulfolobus acidocaidarius. The wild-type enzyme yields (all-E)-C20 prenyl diphosphate as a final product. The three-dimensional model of the enzyme suggested that the remotal of two bulky residues Phe 77 and His 114 would allow additional prenyl-chain elongation. F77G, F77G/H114A, F77G/H114G, H114A, and H114G variants gave C30, C(45), C50, C30 and C40 as the major maximum length products, respectively L4'1

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3 Rationo/ Design of Functional Proteins

Figure 3-8. Proposed mechanism of the chain-length determination of the wild-type and variant heptaprenyl diphosphate synthases based on the pocket mechanism. A, Wild-type enzyme; 6, 176C variant; C, A79Y variant; D, SSOF variant.

3.6.9

Combining Site-directed Mutagenesis with Chemical Modification

Combining site-directed mutagenesis strategies with chemical modification is a popular tool in both enzyme engineering and mechanistic studies. This has often been applied to the subtilisin from Bacillus lentus (SBL), or savinase. Subtilisins are

3. G Examples of Protein Engineering

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87

one of the most well-characterized and well-engineered proteins; mutational effects of more than half of the 275 amino acid residues have been reportedLs0I. A highresolution, three-dimensional structure of S BL is also available. Furthermore, wildtype SBL does not harbor any cysteine residues. Therefore, if a single cysteine residue were to be introduced by site-directedmutagenesis, treatment with methanethiosulfonate reagents would lead to specifically localized modification of the enzyme. This approach, the chemically modified mutant approach (CMM),has been utilized in altering the stability,specificity, kinetic properties, and pH profiles of SBL. The following example displays how the CMM approach can expand the specificityof the S 1 pocket of SBLL5l1. Wild-type SBL is known to prefer bulky, hydrophobic P1 residues in its S 1 pocket. The Phe P1 residue of the standard suc-AAPF-pNA (succinyl-alanyl-alanyl-prolylphenylalanyl-p-nitroanilide)substrate was shown to be preferred by a factor of 9500-fold over the small P1 residue of suc-AAPA-pNA, by a factor of 24-fold compared with the positively charged P1 residue of suc-AAPR-pNAand by a factor of 522-fold compared with the negatively charged P1 residue of suc-AAPE-pNAA.The Ser 166 residue, located at the bottom of the S 1 pocket and whose side chain points inward toward the pocket, was chosen for substitution by cysteine and subsequent chemical modification. In order to increase specificity toward small uncharged P1 residues such as Ala, bulky moieties, for example benzyl, decyl, cyclohexyl, and steroidyl groups, were incorporated at Sl66C so as to reduce the volume of the S 1 pocket and induce a betier fit for small P1 groups. Likewise, negatively charged groups such as an ethylsulfonatomoiety, a dicarboxylic aromatic group, and aliphatic mono-, di-, and tri-carboxyl groups were incorporated for higher specificity for positively charged P1 residues such as Arg. A positively charged ethylamino group was introduced to improve the acceptance of the negatively charged P1 residue Glu. In the case of a cyclohexyl group, the modified enzyme showed a 2-fold improvement in kcat/& with the suc-AAPA-pNA substrate and a 51-fold improvement in suc-AAPA-pNA/suc-AAPF-pNAselectivity relative to WT-SBL. The enzymes modified with mono-, di-, arid tricarboxyl groups displayed improved kcat/& values toward suc-AAPR-pNA. Furthermore, these values increased in parallel with the number of carboxyl groups introduced, and led to a 9-fold improvement in kc,,/ KM for the suc-AAPR-pNA substrate and a 61-foldimprovement in suc-AAPR-pNA/sucAAPF-pNA selectivity compared with the wild-type SBL. Conversely, the introduction of the positively charged ethylamino group led to a 19-foldimprovement in kcat/ KM for the suc-AAPE-pNA substrate and a 54-fold improvement in suc-AAPE-pNA/ suc-AAPF-pNA selectivity relative to the wild-type SBL. 3.6.10

Changing the Catalytic Adivity of a Protein

With the abundant number (> 16 000) of three-dimensional structures in the Brookhaven Protein Data Bank, a challenging but promising task in protein engineering is the synthesis of novel biocatalysts by assembling individual functional modules (substrat12 binding sites, catalytic centers etc.), or by introducing a

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designed functional environment into a known protein template structure. The following is an example of the latter strategy, taking advantage of the diverse functions of a protein superfamily sharing a common fold LS21. The 2-enoyl-CoA hydratase/isomerase enzyme superfamily is comprised of enzymes with various specificities and functions, including 4-chlorobenzoyl-CoA dehalogenase, 2-enoyl-CoA hydratase, carnitine racemase, dihydroxynaphthoate synthase, 2-ketocyclohexanecarboxyl-CoAhydrolase, A3,A2-enoyl-CoA isomerase, and even the proteolytic component of Clp protease. Structural comparison of these proteins indicated the possibility that a majority of the individual active sites were derivatives of a single active site structure. This environment provides a CoA binding site, an expandable acyl-binding pocket, an oxyanion hole for binding or polarizing the thioester carbonyl group, and numerous sites for strategic positioning of catalytic residues. In the study, the active site of one member of the 2-enoyl-CoA hydratase/ isomerase family, 4-chlorobenzoyl-CoAdehalogenase, was altered by site-directed mutagenesis to include the two glutamate residues functioning in acidlbase catalysis in a second family member, 2-enoyl-CoAhydratase. As a result, the syn hydration of 2-enoyl-CoA, absent in the wild-type 4-chlorobenzoyl-CoAdehalogenase, was observed in the engineered protein with kcat and KM values of 0.06 s-l and 50 M , respectively. Although the efficiency of the engineered protein is far from the native 2-enoyl-CoA hydratase, the study clearly demonstrates the possibility of exchanging catalytic functions of two enzymes within a structural enzyme family. It also sends an encouraging message that if an appropriate template is available, it is possible to obtain a desired enzyme activity by rationally designing a catalytic environment on the “template landscape”. Other studies have explored or resulted in even more drastic alterations in enzymatic characteristics. Tyrosine phenyl-lyase (TPL) and aspartate aminotransferase (AspAT) both belong to the a-family of vitamin B6-dependentenzymes. While TPL catalyzes the p-elimination reaction of L-tyrosine,AspAT catalyzes the reversible transfer of an amino group between dicarboxylic amino acids and their corresponding 2-0x0 acids. The double mutation R100T/V283R, leading to an AspAT-like sequence, was introduced into TPL. The protein obtained displayed a 104-fold increase in p-elimination activity towards dicarboxylic amino acids than the wild-type TPL. The activity towards L-aspartate was twice as high as that towards the native substrate L-tyrosine. The created enzyme can be considered a dicarboxylic amino acid p-lyase, an enzyme that is not found in nature[53].A further study attempted to design a protein with enzymatic activity, starting from a structurally homologous non-catalytic protein The nuclear transport factor 2 (NTF2) and scytalone dehydratase both share a common alp barrel structure. Four key catalytic residues, along with a C-terminal a-helix found in scytalone dehydratase, but not in NTF2, were introduced into the NTF2 protein. A mutant protein exhibited scytalone dehydratase activity with minimal kcat and KM values of 0125 min-’ and 800 p ~ , respectively. The study is one of the few examples of converting a non-catalytic protein scaffold into an enzyme.

I

3.7 Conclusions 89 3.7

Conclusions

The examples above represent some of the most successful studies in protein engineering. They show that it is possible to enhance protein thermostahility rationally, alter cofactor or substrate specificity, regiospecificity, and even change catalytic activity. Furthermore, the creation of enzymatic activity from a non-catalytic protein backbone, and tht creation of a biocatalyst with an unprecedented catalytic activity not found in nature, have also been achieved. However, the examples published in the literature are probably only a tiny fraction of the many studies that have been, or are still, in progress awaiting positive results. We are still at a premature stage in designating precise rules to engineer a variant protein with each and every desired property. It is still not easy to predict the outcome of even a single amino acid residue substitution. However in some cases, depending on the information available and the property desired, some basic guidelines are available. Whatever the position, the three-dimensional structure of the protein, or of a homologous protein is highly desired. Without any structural information, strategies will be limited, and the sense of rationality of the experiments will be low. When enhancement of protein (thermo)stabilityis desired, there are a number of strategies available, taking into account four major interactions within a protein; covalent bonds via disulfide bridges, ionic interactions, hydrogen bonds, and hydrophobic interaction (Fig. 3-9). Introducing a covalent disulfide bond in a region distant from the catalyticcenter of T4 lysozyme was reported to enhance dramatically the thermostability of the protein[55,561. With human lysozyme, introduction of Asp residues to generate a Ca2+ binding pocket rationally, and consequently ionic interactions, led to a calcium binding variant protein with an increase in thermoAlthough performed by a random approach, the effects of hydrogen bonds on protein thermostability has also been displayed with T4 lysozyme[581.A single T157I mutation, interrupting a hydrogen bond in the wild-typeenzyme, led to a temperature-sensitive mutant protein. The importance of hydrophobic interactions has been mentioned above. Addition of any of these four types of interactions may be considered in order to enhance the thermostability of a protein. Another alternative may be to introduce proline residues at P-turn structures (the proline rule). This has been clearly demonstrated with oligo-1,G-glucosidasesfrom various Bacillus species[58-6*1

When the aim is to aher the substrate or cofactor specificity of an enzyme, one should look for a homologous structure of an enzyme bound with the target molecule or a structurally similar compound (template structure). This will provide much more information than the structure of a homologous protein alone, even when the latter has been determined at a higher resolution. If the (modeled) structure of the target enzyme is also available, superimposing the structures of the two proteins will make the examination of the supposed interaction of the target enzyme and the binding molecule possible. Side chains that sterically or electrostatically interfere with binding may be identified, and subsequent mutations can be

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3 Rational Design of Functional Proteins Figure 3-9. Various interactions in a protein molecule. Increasing these interactions may enhance the thermostability of the protein, as described in some examples in the text.

dOOH

designed for their removal. On the other hand, residue substitutions that may possibly enhance affinity or increase specificity can also be designed. Even when a (modeled) structure of the target protein is not available, an accurate sequence alignment may also be sufficient, as long as the three-dimensional template structure is available. In some very well-studied cases, such as the pap binding motifs for NAD( I?) cofactor binding (mentioned above), primary sequence alignment may provide enough information to engineer the binding site. Recent studies, some mentioned here, convey new strategies and concepts for protein engineers. Combining rational design with directed evolution has also become a popular means of obtaining a protein with a desired function. The growing number of strategies will surely attract more scientists to become engaged in the field of protein engineering. This will hopefully accelerate the accumulation of information available to the engineer, ultimately enabling the de novo design of a biocatalyst.

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J. A. Bocanegra, N. S. Scrutton, R. N. Perham, Creation of an NADP-dependent pyruvate dehydrogenase multienzyme complex by protein engineering, Biochemistry 1993, 32(11), 2737-2740. 42 M. Nishiyama, J. J. Birktofi, T. Beppu, Alteration of coenzyme specificity of malate dehydrogenase from Thermuspavus by sitedirected mutagenesis, /. Biol. Chem. 1993, 268(7),4656-4660. 43 R. Chen, A. Greer, A. M. Dean, A highly active decarboxylating dehydrogenase with rationally inverted coenzyme specificity, Proc. Natl. Acad. Sci. USA 1995, 92(25), 11666-11670. 44 J. H. Hurley, R. Chen, A. M. Dean, Determinants of cofactor specificity in isocitrate dehydrogenase: structure of an engineered NAD' specificity-reversalmuNADP' tant, Biochemistry 1996, 35(18), 5670-5678. 45 R. Chen, A. Greer, A.M. Dean, Redesigning secondary structure to invert coenzyme specificity in isopropylmalate dehydrogenase, Proc. Natl. Acad. Sci. USA, 1996, 93(22),12 171-12176. 46 E. Hornung, M. Walther, H. Kiihn, I. Feussner, Conversion of cucumber linoleate 13-Iipoxygenase to a 9-lipoxygenatingspecies by site-directed mutagenesis, Proc. Natl. Acad. Sci. USA 1999, 96(7),4192-4197. 47 H. Ma, T. M. Penning, Conversion of mammalian 3a-hydroxysteroiddehydrogenase to 20a-hydroxysteroiddehydrogenase using loop chimeras: changing specificity from androgens to progestins, Proc. Natl. Acad. Sci. USA 1999, 96(20),11161-11166. 48 K. Hirooka, S. Ohnuma, A. Koike-Takeshita, T. Koyama, T. Nishino, Mechanism of product chain length determination for heptaprenyl diphosphate synthase from Bacillus stearothermophilus, Eur. /. Biochem. 2000, 267(14), 4520-4528. 49 K. Hirooka, T. Kato, J. Matsu-ura, H. Hemmi, T. Nishino, The role of histidine-114 of Sulfolobus acidocaldarius geranylgeranyl diphosphate synthase in chainlength determination, FEBS Lett. 2000, 481(1), 68-72. 50 P. N. Bryan, Protein engineering of subtilisin, Biochim. Biophys. Acta 2000, 1543(2), 203-222. 51 G. DeSantis, X. Shang, J . B. Jones, Toward tailoring the specificity of the S 1 pocket of subtilisin B. lentus: chemical modification 41

+

References 193 of mutant enzymes as a strategy for removing specificity limitations, Biociiemistry 1999, 38(40), 13391-13397. 52 H. Xiang, L. Luo, K. L. Taylor, I). DunawayMariano, Interchange of catalyic activity within the 2-enoyl-coenzymeA hydratasel isomerase superfamily based cn a common active site template, Biochemistry 1999, 38(24),7638-7652. 53 B. Mouratou, P. Kasper, H. Genring, P. Christen, Conversion of tyrosine phenollyase to dicarboxylic amino acid p-lyase,an enzyme not found in nature, 1.Biol. Chem. 1999.274(3), 1320-1325. 54 A. E. Nixon, S. M. Firestine, F. G. Salinas, S . J. Benkovic, Rational design of a scytalone dehydratase-like enzyme using a structurally homologous protein scaffold, Proc. Natl. Acad. Sci. USA 199!), 96(7), 3568-3571. 55 L. J. Perry, R. Wetzel, Disulfide bond engineered into T4 lysozyme: stabilization of the protein toward thermal inxtivation, Science 1984,226(4674), 555-557 56 R. Wetzel, L. J. Perry, W. A. Baase, W. J. Becktel, Disulfide bonds and thermal stability in T4 lysozyme, Proc. Natl. Acad. Sci. USA 1988,85(2),401-405. 57 R. Kuroki, Y. Taniyama, C. Seko, H. Nakamura, M. Kihchi, M. Ikehara, Design and

creation of a Ca” binding site in human lysozyme to enhance structural stability, Proc. Natl. Acad. Sci. USA 1989, 86(18), 6903-6907. 58 T. Alber, D. P. Sun, K. Wilson, J. A. Wozniak, S. P. Cook, B. W. Matthews, Contributions of hydrogen bonds of Thr 157 to the thermodynamic stability of phage T4 lysozyme, Nature 1987, 330(6143),41-46. 59 K. Watanabe, T. Masuda, H. Ohashi, H. Mihara, Y. Suzuki, Multiple proline substitutions cumulatively thermostabilize Bacillus cereus ATCC7064 oligo-l,6-glucosidase. Irre fragable proof supporting the proline rule, Eur. /. Biochem. 1994,226(2), 277-283. 60 K. Watanabe, K. Chishiro, K. Kitamura, Y. Suzuki, Proline residues responsible for thermostability occur with high frequency in the loop regions of an extremely thermostable oligo-l,6-glucosidase from Bacillus themoglucosidasius KP1006,/. Bid. Chem. 1991,266(36),24287-24294. 61 K. Watanabe, Y. Hata, H. Kizaki, Y Katsube, Y. Suzuki, The refined crystal structure of Bacillus cereus oligo-1,6-glucosidaseat 2.0 8, resolution: structural characterization of proline-substitution sites for protein thermostabilization, /. Mol. Bid. 1997, 269(1), 142- 153.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

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Enzyme Engineering by Directed Evolution Oliver May, Christopher A. Voigt and Frances H. Arnold

4.1

Introduction

Previous chapters have outlined the huge potential of enzymes as tools for organic synthesis. However, this potential is only slowly being realized in large-scale industrial applications. The main reason for this is that enzymes are often incompatible with the specific requirements of a synthesis, especially under economic constraints. Enzyme behaliors such as substrate or product inhibition, stability, and are all finely tuned by natural evolution to support catalytic efficiency (kc,&,) efficient reproduction of the organisms that make them. Product inhibition can be useful in a living cell, where it prevents the accumulation of undesired or even toxic products. But it is highly undesirable in a synthesis requiring high substrate concentrations and complete conversion into products. Similarly, an enzyme may naturally be highly substrate specific so as to prevent undesired side reactions with other chemically similar metabolites. But such an enzyme can only be used to synthesize a very limited range of products. Other properties that are highly desirable for chemical applications, such as long-term stability and activity in organic solvents, are simply not Iequired in nature and are therefore not found in natural enzymes. While it is possible to devise effective bioprocess engineering solutions to some of these problems, it will often be necessary or more effective to engineer the catalyst itself. The previous chapter reviews methods for structure-guided enzyme engineering. A prerequisite for this approach is knowledge of the enzyme structure and detailed insight into how this structure determines function. Then we must be able to predict how specific amino acid changes affect the desired properties. Despite rapid growth in the numbers of enzyme structures solved and the considerable progress made in cornputational methods, our understanding is still very limited and in most cases insufficient to obtain the desired features with an acceptable probability of success. The strategy nature uses to adapt organisms to new demands is evolution. According to Darwinian iheory, the fantastic diversity of life was created by random mutation and natural selection'']. The power and simplicity of the evolution

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algorithm has tempted scientists and engineers to try to implement this same approach for biomolecular design. In 1984,long after Eigen’s pioneering work on the theory of evolution [2231, Eigen and Gardiner suggested the following procedure that “should allow a new type ofevolutionary biornolecular engineering [41: 10 20 30 40 50 60 70

PRODUCE A MUTANT SPECTRUM OF SELF-REPRODUCINGTEMPLATES SEPARATE AND CLONE INDIVIDUAL MUTANTS AMPLIFY CLONES EXPRESS CLONES TEST FOR OPTIMAL PHENOTYPES IDENTIFY OPTIMAL GENOTYPES RETURN TO 10 WITH A SAMPLE OF OPTIMAL GENOTYPES

Scientists wishing to design useful proteins, peptides, or nucleic acids have picked up this evolutionary approach, which is now known as directed evolution, applied molecular evolution, in vitro evolution, or molecular breedingI5-l3I.Directed evolution combines a high probability of success (the possibility of obtaining an improved catalyst within months) with no requirement for detailed knowledge of structure, function or even mechanism. The basic evolutionary engineering approach outlined in Fig. 4-1has generated impressive results in a few short years, from enzymes that function in organic solvents[l4land at high temperature[151 to enzymes that are active towards non-natural substrates[16]or even carry out whole new reactions[17].It is now clear that directed evolution will drive biocatalysis into a growing number of commercial settings, including many synthetic applications. The aim of this chapter is to explain the concepts underlying directed evolution and to describe its application to engineering useful enzymes. In Sect. 4.2,we describe the principles of an evolutionary optimization algorithm. The tools and their implementation in different working strategies of directed evolution are then described in Sections 4.3and 4.4.The intention is to highlight the main practical and conceptual differences among the various approaches and to compare their strengths and limitations. Section 4.5 discusses specific examples of directed evolution, with a focus on enzymes and properties that are of interest in organic syntheses. Many other important and highly successful applications of directed evolution, such as the design of catalytic antibodies and nucleic acids (ribozymes)or peptides and proteins of pharmaceutical interest, are covered in recent reviews [13,

18-26]

4.2

Evolution as an Optimizing Process

Without an understanding of the theory of evolution, one may be tempted to consider in vitro evolution an irrational, trial-and-error approach to protein design. However, the beauty of the structural architectures and sophisticated functions that nature has created attest to the power of the evolutionary design strategy. Many theoretical studies of evolution explain this process based on physical principles. The

4.2 Evolution as an Optimizing Process

Choose parent gene@)

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t

-

P Create diversity by mutation / recombination

-7

/

Isolate gene(s)

z

\ a

\

1 , -

Couple genotype to phenotype e.g. by expression of the genes in recombinant cells

f

Screen or select for the property of interest

Figure 4-1. Evolutionary enzyme optimization. One or several parent genes are chosen and subjected to mutagenesis and/or recombination. The mutant gene library (genotype) is then expressed in vivo (or in vitro) where it is linked to the enzyme produced (phenotype). These enzymes are tested for the targeted property by screening or selection. DNA from the most fit clone(s) is isolated and subjected to a new cycle of mutagenesi:; and screening or selection. This procedure is repeated until the desired goal is reached or until no further improvements are observed.

principles that emerge are very different from those important in traditional “rational” design. Rather than trying to fully understand how mutations affect the structure and function of the enzyme (which is very difficult), the physics of evolution aims to understand the forces that make systems and problems evolvable. That is, what makes proteins so apt for evolution? Moreover, how can this be used to advantage in enzyme desi-gn? 4.2.1

The Search Space of Cherriical Solutions

To describe evolution as a search process, it is necessary to define the search space. It is convenient to define sequence space as the connected network of all possible amino acid combinations (for a fixed sequence length) [27].For a protein composed of A different amino acids and a sequence length of N residues, there are N sequences, connected by an N(A-1)-dimensionalnetwork. Each point in this vast space has an associated fitness, representing the combination of properties undergoing selection. Together, sequence space and a fitness description construct a fitness landscape on which an enzyme walks towards higher peaks under the influence of mutation and selection [28-301.

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Exhaustively searching all possible solutions is impossible as sequence space is extraordinarily large. The mass of all amino acid combinations for 285 residues (about possible sequences) would be lo3’’ times the mass of the universe, thus creating sequence spaces of a size greater than the power of our Nature could have explored only a small fraction of the sequence space of proteins during the age of earth. Nevertheless, excellent solutions to biological and environmental challenges have been found. Similarly, i n uitro evolution experiments have been successful in finding improved molecules, although a priori success probabilities may seem to be prohibitively small. There are many successful examples where only a very small fraction of the many possible sequences have to be explored to find mutants with improved properties. The key is to develop experimental algorithms that optimize exploration subject to practical limitations. 4.2.2

The Directed Evolution Algorithm

Spontaneous mutations, recombination and selection are the tools of evolutionary design. Plant and animal breeders who influence properties of the offspring by choosing parents with the desired traits have been successfully using these tools for millennia. Following their lead, molecular biologists first employed selection strategies that acted on spontaneously generated bacterial mutants with the goal of developing new metabolic pathways 132, 331. Solutions found by these approaches often resulted from complicated changes in regulatory genes, transport proteins, or the activation of silent genes. Whole pathways are targeted by the evolution experiment when such solutions are desired. However, if the target is a specific enzyme, it is not desirable to produce solutions that are not directly related to the enzyme. The milestone techniques of cloning and i n uitro recombination of genetic information[34]and other advances in molecular biology, such as the development of the polymerase chain reaction (PCR) 135], allow carefully controlled directed evolution experiments. Researchers are now able to specifically engineer the enzyme of interest and control the rate of mutagenesis and focus mutations towards specific regions within the gene. Furthermore, methods are now available to reconstruct “sexual” recombination i n vitr~[36-381 as well as in recombinant 401. In addition, using screening and in uitro selection methods, we can control the selection pressure independently of constraints in living cells, thus allowing acquisition of properties never required in nature. Technologies are available to create protein molecules and select them within a few hours or days[41]. libraries of up to All of these tools allow us to implement the evolutionary design algorithm i n uitro and accelerate it to create molecules with desired properties in a fraction of the timescale of natural evolution. The success of evolutionary protein design is highly dependent on the thoughtful combination of methods for creating diversity and searching the mutant population that is generated. Optimizing an evolutionary search has been well studied in the computer science genetic algorithm literature [42, 431. Some of these results can be applied to directed evolution, including

4.3 Creating a Library ofDiverse Solutions

determining appropriate mutation and recombination rates, optimal recombination parameters, and the appropriate screening effort

4.3

Creating a Library of Diverse Solutions

Given the high cost (both in terms of money and time) of analyzing a mutant library, the goal of the diversity-cn?atingstep is to produce mutant libraries that are rich in variants with improved properties. To achieve this, the few positive mutations that might occur on a gene cannot be diluted with many neutral or deleterious mutations. The level of mutant redundancy also affects the quality of the molecular diversity. Redundancy must be low because screening or selection efforts are wasted on testing identical mutants. In this section, we will first describe different approaches to creating mutant libraries, including mutation and recombination. 4.3.1 Mutagenesis

A commonly used strategy to create mutant libraries is to target the whole gene for random point mutagenesis. Nucleotide mutations are typically introduced by errorprone PCR, mutator strains, or by treatment of the isolated DNA with chemicals or UV light. The success of this approach depends critically on using an appropriate error rate. If the error rate is too low, inadequate diversity is created and screening is wasted on large numbers of redundant parent enzymes. On the other hand, if the error rate is too large, the haction of positive mutants also becomes very low and the search for improved mutants is wasted on screening inactive clones. A serious limitation of the random mutagenesis approach comes from the degeneracy of the genetic code and the biases of available methods, for example the preference for transitions over transversions. Together, these effects limit the amino acid substitutions that arc accessible by DNA point mutations. A combination of a stepwise random mutagenesis approach with methods of focused mutagenesis and recombination can circumvent some of these limitations. The different requirements, limitations, and advantages of the most commonly employed methods are summarized in Table 4-1.In practice, a good strategy is to use a combination of methods. 4.3.1.1 Random Point Mutagenesis of Whole Genes

Before the introduction ofthe polymerase chain reaction (PCR)[351, point mutations were usually produced by UV radiation, by chemical treatmentL4’1 or by using mutator strains that have an increased mutation rate compared to normal strains because of defects in their DNA-repair mechanisms [461. Chemical mutagenesis [471, mutator strains 491, and even spontaneous mutations coupled with selection in a [483

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4 Enzyme Engineering by Directed Evolution Table 4-1.

Comparison of methods for creating genetic diversity for directed evolution. Requirement

Advantage

Limitation

Random point mutagenesis

None

Exhaustive

No multiple simultaneous mutations; requires multiple rounds to accumulate beneficial mutations

Focused mutagenesis

Structural information or knowledge from previous generations

Reduced library size; multiple simultaneous mutations possible

Misses possible good sites

None

Recombine positive mutations; remove neutral and deleterious ones

No multiple simultaneous mutations: recreates large number of already known sequence

“Functional diversi-

Not exhaustive: limited to amino acid diversity in parental sequence space

Recombination - single gene

-

family shuming

Homologous genes

ty”: large jumps in sequence space

chemostat are still used for random point mutagenesis in directed evolution experiments. However, the dominant method is now error-prone PCR because the protocols are very straightforward, safe, fast and versatile, and allow simple adjustment of the error level. Point mutations are introduced by PCR because of erroneous incorporation of a nucleotide during the amplification of the target gene. Under normal reaction conditions, the error rate of the Tuq DNA polymerase is 521. Although this about 0.001 % to 0.02 % per nucleotide per replication error rate is sufficient to create libraries of large genes lS31, it is too low for efficient mutagenesis of small genes. However, the fidelity of the PCR can be reduced by changing the reaction conditions (e.g., increasing the concentration of MgC12, adding MnCl2 to the reaction mixture, increasing and unbalancing the concentrations of the four dNTPs, adding deoxyinosine triphosphate (dITP), increasing the concentration of Tuq polymerase, or increasing the extension time and cycle numbers [54-581. These methods can result in error rates as high as 2 % per nucleotide position. The possibility of using of low fidelity Tuq mutants has been described[59], but has yet to be explored experimentally. By changing some of the PCR conditions, the error rate can be adjusted according to the gene length to produce the desired average number of amino acid substitutions. A frequently used method to estimate the level of mutation (a quality check of the diversity) is to determine the fraction of inactive clones from small samplings of the generated mutant libraries L60, 61]. However, the statistical distribution of muta-

4.3 Creating a Library ofDiuerse Solutions

tions (which should be narrow) cannot be estimated by this method, and the relation between the fraction of inactive clones and average number of amino acid substitutions can differ for different enzymes. Therefore, the statistical distribution of mutations and the relationship between inactive clones and the number of mutations are determined by sequencing randomly picked mutants. Another consideration I S the distribution of mutation type. Typically, there is a strong bias for transitions (A-G or T+C) over transversions (C-G or G+C), which limits the accessible amino acid substitutions. There are protocols to reduce this bias, but they do not completely eliminate it[55, 62]. In addition, the structure of the genetic code limits the accessible amino acid substitutions. Depending on the specific codons, only 2 4 4 0 % of the possible amino acid changes are accessible by single base substitutions 631. Furthermore, the accessible substitutions are more likely to be conservative with similar physicochemical properties. For large genes and small error rates in whole-gene mutagenesis, it is very unlikely that two DNA mutations will occur in tkie same codon, dramatically reducing the possible amino acid substitutions. Although little is known of the cost of these constraints in directed (and natural) evolution, several studies have shown that the best mutations at specific sites that introduce required multiple substitutions in a single ~ o d o n [651.~ ~Methods , diversity at the codon level might therefore be preferable to methods that create point mutations at the nucleotide level [631. Methods available for codon-level mutagenesis of a few amino acid positims are unfortunately very cumbersome and expensive for mutagenizing whole genes, leaving room for future developments of improved mutagenesis methods. 3' '

4.3.1.1.1

Optimal Mutation Rates: Experimental

One rule-of-thumb has been to adjust the error rate according to the number of targeted amino acid residues and the size of the screen, such that a significant fraction of the total number of combinatorial possibilities can be sampled["]. For an enzyme of 300 amino acid residues, there are about six thousand possible single mutants, sixteen million double mutants, and thirty billion triple mutants. It is generally within our abi1ii.y to exhaustively screen a single-mutation library, whereas double-mutation libraries already exceed the throughput of most screening methods (typically < lo6). Standard selection methods allow a throughput of about lo8 mutants and can therefore sample most double mutants. In vitro selection methods might push this limit somewhat higher (10"). The assertion that it is desired to sample most of the conibinatorial possibilities is based on the assumption that beneficial mutations are rare and the probability is very small that multiple random amino acid substitutions are beneficial. An error rate resulting in single or double mutation libraries has been found to be a good compromise between creating adequate diversity while limiting the screening effort. In practice (e.g., Table 4-3), significant improvements in enzyme activity, stability, selectivity, folding, and expression have been achieved by the stepwise accumulation of single amino acid substitutions. Even large changes in substrate

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specificity[621 and inversion of enantioselectivity["I can be achieved by this conservative approach. The obvious disadvantage of this approach is that some properties will benefit from simultaneous mutation of multiple amino acids. These solutions will not be found. Arguments for low error rate mutagenesis were recently challenged by several researchers. Zaccolo and Gheradi produced p-lactamase libraries with error rates that they claim generated 5-16 amino acid changes per mutant enzyme per generation [67]. The libraries (- 104-105 clones) were selected for increased resistance to the antibiotic cefotaxime over three generations. However, the best mutant differed in only three effective amino acid substitutions from wild-type and is therefore much less mutated than would be expected from three generations of 25 amino acid changes per generation. This indicates that the applied average error rate was actually much lower, and a more suitable average mutation rate would have been three amino acid substitutions. Another recent study by Georgiou and coworkers indicates that a high mutation rate was appropriate in improving the affinity of a single-chain antibodyr6*].When screening for improved affinity, they found that the most improved mutants were observed in libraries created with moderate to high mutation rates (3.8-22.5 mutations/gene). In the next section, we discuss the circumstances under which a higher mutation rate is advantageous.

4.3.1.1.2

Optimal Mutation Rates: Theory

In nature, the spontaneous mutation rate is tightly controlled. During one replication round, about 3 ~ 1 0 -mutations ~ occur in the genome of human cells, which is 1 . viruses, such as HIV, have a much about the same level as in Escherichia c 0 l i [ ~ ~RNA higher mutation rate, typically on the order of one mutation per genome per replication Such a high mutation rate is crucial for the viruses to survive the attacks of the immune system. However, there is a maximum mutation rate, above which the requirement of inheritance for evolutionary optimization breaks down (referred to in quasi-species theory as the error At the mutation rate just before this threshold, the speed of evolution is highestL3. 721. Interestingly,it was demonstrated that the mutation rate of the replication machinery of fast For directed evolving RNA viruses is indeed close to this error evolution of enzymes, the optimal mutation rate maximizes the speed of the adaptive walk and is influenced by the number of mutants that can be screened and the structure of the fitness landscape. As discussed in the previous section, the general rule has been to use a mutation rate for which the permutations can be effectively sampled during screening. However, if the fitness landscape is amenable to a high mutation rate, fewer mutants must be screened in order to achieve the benefit of a higher mutation rate. For example, it would require a smaller library than all double mutant permutations to benefit from a mutation rate of two per sequence. In this section, we explore how the optimal mutation rate is influenced by the number of mutants that can be sampled, the fitness of the parents, and the ruggedness of the fitness landscape. The optimal mutation rate depends on fitness of the parental sequence. For a sub-

4.3 Creating a Library of Diverse Solutions

optimal sequence, a large mutation rate allows a greater sweep of sequence space. However, because the probability of finding improved mutations decreases as the fitness of the sequence increases, adaptation via a large mutagenesis rate is rapid at first, then slows. If the parent is highly optimized, the probability that a mutation is deleterious is higher. The accumulation of deleterious mutations is more rapid and these mutations quickly (erode the few positive mutations that occur. By using a Markov chain analysis to study genetic algorithms, Miihlenbein found that there should be approximately one amino acid substitution per sequence for highly optimized sequences [741. ]His analysis also suggested that that the optimal mutation rate should decrease as the fitness of the parent increases. In several independent studies, it was demonstrai ed that the solution of an evolutionary search is improved whm the mutation rate was decreased over time[75-781. A higher mutation rate dramatically increases the fraction of mutants in the library that contain stop codons, requiring a larger screening effort[7g].For instance, if the average number of DNA mutations per gene is five, over 20 % of the resulting library will contain stop codons. The quality of the mutant library can also be degraded by the accumulation of deleterious mutations, an effect that is exacerbated by the landscape rugged~iess[~~~]. For the mutation of a highly coupled residue to generate a fitness improvement, it is necessary to optimize all the other residues to which it is coupled. Ideally, the optimal mutation rate equals that of the maximum number of residues involved in a single coupled interaction, thus assuring that the sequence will not become trapped in a local optimum. However, the finite number of mutants that can be screened imposes an upper limit on the mutation rate. Therefore, the optimal mutation rate decreases as the landscape ruggedness increases. This observation is similar to the long-jump mutagenesis strategy suggested by KauffmanIs0I. By making moves that are larger than the correlation length (smoother landscapes have larger correlation landscapes), more space can be explored. Quasi-species theory also predicts that smoother landscapes have higher optimal mutation rates [811. Because real protein fitness landscapes are undoubtedly highly anisotropic, they contain many correlation lengths, and different regions of the sequence will have different optimal mutation rates 82]. A highly coupled region (such as the catalytic site) has a small correlatlon length; thus a smaller mutation rate is allowed with a limited mutant library. Based on some simplified simulations, it was found that the probability of picking a mutant that has a highly coupled mutation decreases significantly as the sequt-nce increases in fitness iU]. This effect intensifies as the number of interactions that are coupled to the mutated residue increases. From this observation, it follows that when the screening effort is limited, uncoupled regions of the protein should be targeted for mutation. More highly coupled residues require a larger rearrangement of amino acids than is likely given the limited mutation rate. Avoiding the regions of high coupling decreases the total number of residues undergoing mutagenesis. To utilize this observation, it is necessary to have experimental techniques to target specific positions as well as methods that can be used to predetermine the coupling of each residue. These goals are the subject of the following two sections.

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4.3.1.2

Focused Mutagenesis

Focused mutagenesis strategies are used with the intention of enriching a library for the targeted region can be desired mutants. To reduce screening efforts [83-851, reduced from 300 to only a few residues (Fig. 4-2). The library of quintuple mutants has a theoretical size of only -lo6 mutants, compared to 10“ if the entire gene is targeted. This reduced library can be searched exhaustively with currently available methods. Focused mutagenesis significantly reduces screening requirements for libraries of mutants with multiple amino acid substitutions and eliminates the codon bias of PCR. However, it imposes obvious limitations on the possible solutions and can fail to explore the most effective mutations. Targeting single amino acids (“saturation mutagenesis”) is straightforward because of available strategies that eliminate laborious subcloning steps. Several commercial kits are available, such as the TransformerTM(CLONTECH Laboratories, Palo Alto, CA, USA), Altered Site@ I1 (Promega, Madison, WI, USA), and QuickChangeTM(Stratagene, La Jolla,CA, USA) site-directed mutagenesis systems, which can produce targeted mutant libraries in one day. This approach has been used to target amino acid positions that random point mutagenesis identified as important for the targeted enzyme properties [64. 651. Variants with improved proper-

number of simultaneously changed amino acids Figure 4-2. Plot showing the number of possible protein variants (v) that can be created given the number of amino acid positions ( M ) changed simultaneously and the sequence length ( N ) (Y=lg’[N!/(N-M)!M!]). From a given standard screening throughput (
4.3 Creating a Library ofDiverse Solutions

ties were identified in saturation mutagenesis libraries that contained mutations not accessible by random point mutagenesis. Methods for randomization of specific regions of a gene typically employ randomized oligonucleotides or PCR mutagenesis of small stretches of the gene. The required diversity is introduced by randomized oligonucleotides that are produced by an automatic (programmable) DNA synthesizer. If only one amino acid position is targeted, the specific codon can be completely randomized (saturated) by adding an equal amount of all four bases (A, T, G ,C) during the oligonucleotidesynthesis at the first two positions of the triplet and with a mixture of G and C at the third position to exclude stop codons (referred to as NN(G,C)-mutagenesis). To target multiple amirto acid positions simultaneously within a small region of the gene, oligonucleotide-cassette mutagenesis is often used. This method was among the first applied for in vitro evolution of DNA sequences [“I and has also been used successfully for the evolution of various enzymesL5’, 87-901. When suitable restriction sites are not already present, they are introduced by site-directedmutagenesis adjacent to the targeted region. These restriction sites are then used to substitute the wild-type region with the synthetic randomized DNA duplex (cassette) [911. Depending on the number of targeted amino acid positions, randomization strategies are used to facilitate an exhaustive search of the library. Complete randomization of all targeted codons is preferred if few amino acid residues are targeted and can be done as described above for the randomization of single sites by using NN(C/G)during oligonucleotide synthesis. It was also reported that trinudeotide analogs such as 9-fluorenylmethoxycarbonyl (Fmoc) trinucleotide phosphoramidites can be used during DNA synthesis to achieve a codon-based mutagenesis 1921. If several distant positions are targeted simultaneously, cassette mutagenesis is technically cumbersome. Other methods allow the efficient assembly of several randomized oligonucleotides to whole genes. For example, recursive PCR1931,the ligase chain reaction[”, ‘)41, or in uitro assembly of whole genes[”] can be used to construct targeted mutant libraries. Partial randomization of many positions most frequently employs spiked oligonucleotides,which are produced by DNA synthesis using a mixture of the wild-type base and equimolar amounts of all four The oligonucleotides can also be produced by error-prone PCR. It is possible to calculate the appropriate compositions of the nucleotide mixture in order to encode whole sets of amino acid!; at certain 4.3.1.3 Calculation of Mutagenesis Hot-Spots

In focused mutagenesis experiments, the challenge is to identify the residues where mutagenesis is likely to be beneficial. Indeed, many successful directed evolution experiments show that mutations occur in regions that would be hard or impossible to predict (and difficult to explain that they do), even when a high-resolution structure and much information about the enzyme is available[14,9s1001 . 0ne possibility is to make use of knowledge gained from early rounds of random point

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mutagenesis that targeted the whole enzyme[", 101, lo21. The content of the mutant library can be improved by only mutating sites that do not severely disrupt stability. A structurally tolerant protein allows more mutations, and therefore more potentially beneficial ones, making it more likely that there is a connected path in sequence space of single mutations that leads to regions of higher fitness. By reducing the evolutionary search to regions of sequence space that retain structure, functional space can be explored more This concept can also be inverted: if the goal is to improve stability while retaining functionality, then eliminating the sequence space inconsistent with the function improves the search. Several groups have proposed targeting mutagenesis to residues where natural diversity is observed. Fersht and coworkers reengineered the tumor suppressor p53 by creating a small library of mutants where the hot-spots were determined from a sequence alignment of 23 homologous proteins [1031. The mutations were made in the wild-type sequence background, and several were found that improved stability. Using a similar methodology, Lehmann et al. constructed a thermostable phytase from the consensus sequence of 13 homologous The mutant phytase exhibited a 15-22 "C increase in melting temperature. Alanine scanning has been widely used to identify the residues which are contributing to various protein properties L105, lo6].Alanine substitutions are made at various positions and the perturbation in the property of interest is measured. This has several potential applications to directed evolution. For instance, it can be used to predetermine which positions are essential to the structure (or function) of the protein and therefore should be avoided. Conversely, positions that tolerate the alanine substitutions may be good candidates for saturation mutagenesis. Unfortunately, this procedure is tedious. To surmount this difficulty, Kollman and coworkers proposed a method to determine the effects of alanine substitutions c~mputationally['~~]. Kollman's method could be used to scan the protein structure for positions to mutagenize in directed evolution. The observation that some sequence positions are more tolerant to mutation initiated the application of information theory to studying the importance of these residues to structure and function["]. The sequence entropy can be calculated from the probability distribution of allowed amino acids substitutions at each residue [108-1101 . u sing simulations of evolution on fitness landscapes, Voigt et al. predicted that beneficial mutations are found by directed evolution at amino acids that are largely uncoupled to other sites (Figure 4.9) 1441. To test this prediction, they compared the calculated site entropies with mutations found from previous evolution experiments on subtilisin E and T4 lysozyme. The sequence space considered in amino acid combinations (274 the subtilisin E computation was enormous: residues). Seven out of the nine mutations that improved the thermostability of subtilisin E occur at positions computed to be highly tolerant. Mutations that improved activity in organic solvent similarly occurred at high-entropy positions. This calculation may be used to determine the positions where improvement will likely be found in an evolution experiment.

4.3 Creating a Librmy of Diverse Solutions

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4.3.2 Recombination

Another approach to creating genetic diversity is based on DNA recombination. Multiple positive variants are used to parent the next generation, which allows for recombination of beneficial mutations, elimination of deleterious mutations as well as creation of new diversity. Using recombination also has a practical advantage. More than one mutant may show improved fitness during a single screening effort. Allowing only the fittest mutant to continue to the next generation can be wasteful (Fig. 4-3). For highly non.additive (rugged) fitness landscapes, recombining positive mutations is less certain to be beneficial because combining individually good mutations can have deleterious effects. In this discussion, we focus on the experimental techniques for recombination and describe the theoretical basis for the optimal parameters.

,

4.3.2.1

In Vitro Recombination

In vitro recombination of’ DNA, often referred to as DNA shuffling, was introduced by Stemmer for evolutionary protein design[3G,371. The method is based on recursive As PCR, which allows for whole gene synthesis from several DNA outlined in Fig. 4-4, one or several parental genes are cut by enzymatic digestion using the endonuclease DNase I in the presence of Mg2+.This generates overlapping DNA fragments that are randomly distributed over the gene. The isolated DNA fragments are then reassembled in a PCR-like reaction with denaturation, annealing, and extension steps, during which recombination occurs through the reannealing of DNA fragments from different parents.

A

14-.--;A+B+c

B,

6-c+B+; f=:

-6 -C

--wt

1

2

---t

3

-wt

1

2

3

Figure4-3. Comparison o f the progress o f evolution for a random mutagenesis approach (A) where the best mutant is used as parent for the next cycle o f mutagenesis and screening and a DNA-recombination approach (B) where several improved mutants are used as parents for the next generations. N o t shown here is any additional screening cost associated with finding several improved variants in each generation.

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Diversity is created by combining parental mutations and random point mutations which are introduced at a rate of about 0.7 % because of the intrinsic error rate of Taq DNA polymerase [371. A high error rate, however, can mask the relationship between evolved phenotype and combined parental mutations. Using different DNA polymerases and substituting Mg2' with Mn2+during DNase I digestion reduces the mutation rate to as little as 0.05 % r61]. A high-fidelity protocol is also important if recombination is used to distinguish between functional and non-functional mutations or for structure-function studies of evolved sequences ['"I. In normal PCR, recombination can also occur at a low rate. This is caused by incomplete extension of primer during the extension cycle and annealing to a different template [112-1141. Increasing the recombination efficiency of incomplete primer extension motivated development of the staggered extension process (StEP)[1151. The basis of StEP is repeated switching of the template caused by fast annealing and extension cycles (Fig. 4-5). During each cycle, the growing oligonucleotide can randomly anneal to different templates and thereby combine information from different parents. A further method for in uitro DNA recombination is the random-priming method (RPR)f3'1, which involves production of gene fragments by annealing and extension of random-sequence primers. The fragments are reassembled as in the Stemmer procedure. StEP, RPR and Stemmer's method were compared based on their recombination efficiency of truncated GFP genes[''']. The Stemmer method using small fragments ( 4 0 0 bp) and StEP yielded the highest recombination efficiencies. However, the efficiencies may differ from gene to gene and are highly dependent on the experimental conditions chosen. Detailed protocols for RPR, StEP and the Stemmer method are given in [151.

parent genes

PCR-Iike

assembly

-2 - = =

.n-_y;

=

_ ._- _ .

*

'

~

.

+

-A'

Figure 4-4. In uitro recombination by DNA shuffling as described by Stemmer[37].Parent genes carrying mutations (indicated by X) are digested with DNase I and randomly reassembled in a cyclic PCR-like reaction t o yield a library of recombined genes. New point mutations are introduced (indicated by underlined X) during the reassembly reaction.

4.3 Creating a Library ofDiuerse Solutions PO9

In uitro recombination by the staggered extension process (StEP) I”’]. Only one primer and single strands from two parents are shown. Fast annealing and extension cycles during PCR of the parent genes cause template switching of the extending strand. After .full-sized recombined genes are synthesized, parent genes are removed by treatment with Dpnl. Figure 4-5.

F

parent genes

template switching during annealing and polyrnerizatior

Dpnl digestion of pareni

It is possible to recombine any number of parent genes with the available methods, which raises the question of what is the optimal number. Similar to determining the optimal mutation rate for random mutagenesis, the answer will depend on the number of screened mutants and the additivity of the combined mutations. It could be advantageous to screen all the permutations of mutations from the parents. Assuming independent and additive recombination, the probability pd that an offspring has d mutations is given by p d = - T! (T- d)!d![

M-1

T-d

h)*[7)

where M is the number of sequences and T is the total number of mutations[1’71. Unfortunately, only 25 74, of the sequences in the recombination library have novel combinations of mutations because there is a statistical disadvantage for the presence of new combinations of mutations. The probability of creating a mutant that contains all mutations is only (l/M)? If all mutations from four parents (each having two mutations) are recombined, the probability of creating an offspring gene with all the parental mutations is only 1.5~10-~. This probability decreases rapidly as the number of mutations and parent genes increase. The probability is further reduced if recombination is accompanied by a high error rate. It is clear that recombination of large pools of sequences quickly reaches the throughput limitation for available screening or selection methods if the mutant containing all mutations needs to be sampled. One strategy for reducing the screening requirements is to divide the recombination experiment into multiple generations L1l7]. The following example demonstrates the advantage of this procedure when the goal is to combine eight mutations onto a single gene from four dtmble-mutant parents. The experiment is divided such that

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4 Enzyme Engineering by Directed Evolution

two libraries are created using two parents each. In each library a mutant is created that contains all four mutations of its parents. The probability of creating this quadruple mutant is p = (1/2)4 = 1/16. All recombinant mutants at this step can be sampled by screening only 32 clones (some oversampling will be required). If the quadruple mutants from each library are then recombined to create a mutant with all eight mutations, the probability of creating it is (1/2)'. In sum, screening fewer than 300 clones (16 + 16 + 256) would be sufficient,whereas the simultaneous recombination of all four mutants to create the same mutant in one recombination experiment would require screening about 65 000 clones. The success of this procedure also relies on the additivity of the mutations. If some of the mutations are non-additive, then combining all mutations is not guaranteed to be optimal. In addition, not all combinations of mutations are screened. If a particular double or triple mutant is the fittest, it may not be found (see pooling strategies, Sect. 4.2.3). Note that if the mutations to be combined were discovered in previous generations, then it is likely that they are relatively additive (or uncoupled, see Mutagenesis Hot-Spots, Sect. 3.1.3).

All of the recombination methods described above require considerable sequence identity among the parents for crossovers to occur. For example, in the Stemmer method, the fragments require a minimum nucleotide sequence identity to reanneal and form an offspring gene. Non-homologous recombination methods seek to remove the sequence identity restriction from the recombination process. Ostermeier et al.["'* describe a method based on generation of N- or C-terminal fragment libraries of two genes by progressive truncation of the coding sequences with exonuclease 111 followed by ligation of the products to make a single-crossover hybrid library. An intrinsic problem of this approach is that random ligation of two DNA fragments results in two-thirds out-of-frame sequences, yielding mostly nonfunctional products. In addition, recombination of more than two parents is hard to realize. Another approach to create a mutant enzyme library is based on the permutation of modules or secondary structure units[120.l2'1. It is not yet known to what extent non-homologous recombination generates usefil genetic diversity, rich in improved functions. 4.3.2.2

In vivo Recombination

In vivo homologous DNA recombination mechanisms are known for various host cells such as E. c0li1~~1 or Saccharomyces c e r e v i ~ i a e [and ~ ~ ] have long been applied in protein engineering purposes, for example to shuffle mammalian P-450 substrate specificitiesf9'1. Because of its simplicity, it is an attractive tool for directed evolution of enzymes['22,1231 . The most common system is based on the ability of S. cerevisiae to rescue plasmids with a double-bond break by intermolecular homology-dependent re~ombination1'~~I. A plasmid is cut by restriction enzymes and transformed into yeast together with different genes or fragments thereof. Recombination at homologous positions on both sites of the gap and within homologous regions of the fragments yields a functional, self-replicatingcircular plasmid. The reconstitution of

4.3 Creating a Library ofDiverse Solutions

the functional plasmid allows for easy detection of recombination events based on the selection marker of the plasmid. Besides its simplicity, an important feature of in vivo recombination is that additional point mutations are extremely rare because of the high fidelity recombination mechanism in yeast. This is particularly advantageous when the aim is solely to recombine positive mutations or eliminate deletevivo methods are unlikely, however, to generate as many rious and neutral ones. IYL crossovers as i n uitro methods. Volkov et al. described a hybrid in uitro-in vivo recombination method involving formation of a heteroduplex between two homologous sequences in uitro and transformation into bacterial cells [1251. Mismatches present in the heteroduplex are randomly repaired by the host cell, creating recombinant sequences composed of the elements of each parent. This approach may be particularly useful in recombining large genes or entire operons. 4.3.2.3 Family Shuffling

Recombination of homo1ogous parent genes, referred to as “family shuffling” or “molecular breeding,” ini-roduces an additional dimension to creation of molecular While random point mutagenesis and local diversity for directed evolution.]‘’1L recombination explores only the sequence space covered by neighboring mutants, and focused mutagenesis covers only a small fraction of the whole sequence space of the protein, family shuffling explores more distant regions of the sequence space with a lower sampling density. Since the diversity is usually created by a combination of mutations from pareni s that were previously selected in nature to be functional, it is suggested that family shuffling provides functional diversity that could accelerate evolution [”‘I. Several recent studies ,attest to the power of this . Us ing a family of four genes from different species, a comparison to single-gene recornbination suggested an evolutionary advantage for family shuffling[”‘]. Recombination of four cephalosporinase genes (57%-82 % identity in amino acid sequence) and screening of 5x104 clones yielded $3 mutant with a 540-fold increased moxalactam resistance compared to wild-type. The mutant that was generated by family shuffling was an offspring of three out of the four parent sequences and contained 33 new amino acid substitutions. The large number of new point mutations raises the question of how so many could be tolerated. It is unclear whether these mutations contribute to the improved Ij-lactamasefunction. An important Characteristic of the family shuffling approach is that the diversity changes after each generation. During early generations, many different combinations of mutations are tested, and the best combination becomes fixed during subsequent generations After futation, new diversity is created only by additional random point mutations inherent to the recombination method. Thus, the most promising distant regions in sequence space are explored by recombination, followed by step-wise mutagenesis towards the fitness optimum. This hybrid methodology might take more than the maximum of four generations of recombina-

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4 Enzyme Engineering by Directed Evolution

tion that have been used so far in family shuffling Based on studies of genetic algorithms it was also proposed that the recombination process with additional mutations provides a powerful method for finding higher fitne~ses1~~~1. Family shuffling should gain increasing importance with the greater availability of homologous genes as a result of the rapid accumulation of new sequences in public databases. The restrictive licensing policies for the known recombination technologies might be limiting the commercial use of recombination methods at the moment. However, this situation and technical limitations of the existing methods are stimulating the development of new and improved recombination methods.

4.4

Finding Improved Enzymes: Screening and Selection

Given a thoughtful strategy to generate a mutant library, developing a method to detect positive mutants is probably the single most important step determining success (or failure) of directed evolution. Screening refers to a qualitative or quantitative assay of each single clone or few pooled clones of a mutant library, whereas selection refers to methods that enrich positives in a pool of all members of a mutant library or, even better, allow growth only of desired variants (extensively reviewed in[133-135]). In directed evolution, diversity is created on the genotype (DNA) level, whereas screening and selection acts on the phenotype (protein) level. A physical link between the genotype and phenotype is required because direct amino acid sequencing of positive mutants is not feasible and because DNA or RNA is required for replication of the desired mutants as well as for introduction of diversity in subsequent cycles of evolution. For the evolution of enzymes, the most simple, versatile and therefore most commonly used approach to couple genotype and phenotype employs recombinant cells. Genes are introduced (transformed) into cells and translated into proteins by the cell's natural transcription-translation machinery. For organisms such as E. coli, Bacillus subtilis, and S. cerevisiae, efficient transformation protocols are available that allow for the production of reasonably sized mutant libraries of 105-109different members. In addition, recombinant protein expression is well established for those organisms, making them the current preferred choices for directed evolution experiments. In this section, we focus on general principles for searching mutant libraries and discuss important characteristics of available systems such as throughput, error level and versatility. We also discuss theoretical approaches to determining the required screening effort, analyzing the immense amount of data that are generated during the screening step and the theoretical advantages of different screening strategies.

4.4 Finding lmproved Enzymes: Screening and Selection

4.4.1

You Get What You Screen For

The first rule of directed evolution is “you get what you screen for.” The importance of establishing appropriate screening or selection methods cannot be overestimated. Corlditions used for screening or selection should be as close as possible to the conditions where the enzyme is going to be applied. This includes the actual substrate, its concentration, pH, buffer, salt, temperature, co-solvents, and any other conditions that affect the enzyme behavior. For example, the screen can be established conveniently using an artificial substrate, the enzymatic products of which produce color or fluorescence. However, optimizing an enzyme on an artificial substrate bears the risk that gains in performance will disappear on the desired substrate. This is true for virtually all enzyme behaviors. If compromises in reaction conditions or substrates cannot be avoided, the risk of obtaining undesired solutions during several rounds of directed evolution can be reduced by using a secondary screen under more “real”reaction conditions or by testing the chosen mutant(s)with the actual substrate after each cycle of directed evolution[’’]. For in vivo selection, which uses the indirect measurement of cell growth as an indicator of enzyme performance, one has to he aware of the immense flexibility (and creativity)of living organisms to circumvenl selection pressures by inventing new solutions unrelated to the enzyme and its property we wish to optimize. Many such examples are well known from studies of metabolic acquisitions through laboratory selection, where strong selective pressure uncovered solutions to biochemical blocks [32*33, 1361. Directed evolution experiments using combinatorial mutant libraries have also found complementation that was caused by activating a novel gene locus rather than by the mutated enzyme[L37]. Starvation under selection conditions may induce lowfidelity polymerases and speed up the evolution of new s o l ~ t i o n s1[3’1~. ~ ~ ~ 4.4.2

Screening Strategies

Screening methods allow enzyme behavior to be measured independently of biological function. Novel enzyme properties can be explored, such as activity or stability in unnatural eiivironments (e.g., extreme pH, temperatures, or organic solvents) or activity on unnatural substrates. These properties are impossible to target using in vivo selection methods, because of the cell’s inability to survive under such biologically harsh conditions. The advantage of higher versatility and better control of the applied selection pressure comes at the cost of lower throughput (defined as the number of clones that can be tested in a given time period). Depending on the screening methods, libraries of about 104-10G clones can be screened within a few days, which is several orders of magnitude less than the lO’-lO” clones that can be tested with selection methods. High throughput is usually accompanied by a high error level in the measurement, which dictates the minimum change that can be detected. On one hand, the consequence of low throughput is missing r.ire beneficial mutants because they are not sampled. On the

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4 Enzyme Engineering by Directed Evolution

other hand, mutants will also be missed if the method cannot resolve the small fitness difference between a mutant and the parent (Fig. 4-6). Part of this difficulty stems from the fact that only a few mutations are made at each generation, and it is often only over multiple generations that large fitness improvements are observed. It is important to take both the throughput and the error level of the assay into account when setting up a screen. The intrinsic error level of a screen can be tested by screening a number of wild-type clones, which allows an estimate to be taken of the minimal change that can be resolved. Fig. 4-7 shows typical statistics of a screen using a 96-well microtiter plate pH-indicator assay, specifically applied for the evolution of a hydantoina~e[~~]. From the standard deviation (in this case, 5 %) and maximal deviation (20%), one can estimate that mutants differing in more than 50% activity can be detected with high confidence. Thus, this screening method is suitable for detecting small improvements in activity in a mutant library. 4.4.2.1

Low-Throughput Screening

Screening in 96- or 384-well plate formats allows precise fitness measurements. The accuracy of the detection system, kinetic assays (in contrast to end point assays),the possibility to normalize activity values (e.g., using measured cell concentrations), and better control over cell growth and protein expression contribute to the improved

A

>

clone #

6 Throughput (sampling size) versus error level of a screen. Low error level and small sampling size (A) might miss the best mutant because it is too rare to be sampled. Large sampiing size and high error level (B) might miss the mutant because it cannot be distinguished from wild-type background even though it was sampled Figure46

sampling of large library size <

*.

0

* .

......

missed mutant

.

.*. 0

clone #

P

.. ... .

______,

high

-

4.4 Finding Improved Enzymes: Screening and Selection

Statistics of a typical screen using a 96-well microtiter plate pH-indicator assay which was applied for the evolution of a hydantoin a ~ e l ~Total ~ ] . # clones = 192. The experimental data roughly follow a Gaussian distribution. Figure 4-7.

h

6o 50

0

40

2D

30

c

2

I

experimental data gaussian distribution

standard deviation: 5% maximal deviation: 20%

20

\c

10 0 0.8

0.9

1.0

1.1

1.2

1.3

activity

accuracy of microtiter plate-based screening systems compared to colony-based screens. Semi-quantitatiw visual screens are usually based on fluorescence[140k1421, color formation[16,1431, or formation of clear zones (halos)[144, 14’1 of colonies grown on agar plates (often filter membranes). The time-consuming process of gridding transformed colonies into microtiter pates is not required. Further, sample preparation steps are straightforward (and sometimes not necessary at all), which simplifies and accelerates the screening process. If coupled to digital imaging analysis, visual 14’1. However, the high throughput and screens can be used for simplicity of these methods are often balanced by a larger error. Standard analybc systems such as HPLC, GC or capillary electrophoresis allow very sensitive and precise measurement and are highly versatile. However, throughput is very limited to fewer than lo3 samples per week. This is too low for efficient screening of most mutant enzyme libraries. Further developments, such as integration of HPLC, GC, or capillary electrophoresis into automatic liquid handling systems, coupling of mass detection systems, and parallel separations will increase the throughput. 4.4.2.2

High-Throughput Screening

Complete automatic systems are now available that can screen up to a thousand 96-well plates per day (about lo6 samples per week). Systems have been developed to increase the density of the plates (number of samples per area) by reducing the required sample volume[’48,14’1. This reduces the cost of screening each mutant and increases the throughput of the assays. In screening enzyme libraries, the throughput seems to be more limited by the required step to transfer single clones from agar plates into the arrays of microtiter plates or silicon wafers rather than by the assay step. Robotic systems are sometimes used to speed up this transfer. An alternativeto this step might be a dilution that adjusts cell density to one cell within a certain liquid volume, which could be transferred into adequate plates (or onto silicon wafers) much faster. Although the theoretical throughput would be increased, the statistical problem arises. that a huge fraction of the transferred volume would be empty or contain more than a single cell. Other ultra-high-throughput systems that

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4 Enzyme Engineering by Directed Evolution

can directly analyze single cells or single proteins do not require a transfer step and thus have a potentially higher throughput. Confocal fluorescence coincidence analysis (CFCS) can analyze up to lo7 single molecules or cells per week[”0]. Although the reported sensitivity (
Choosing Low versus High Throughput

In this section, we will discuss the critical issues in deciding the balance between throughput and accuracy in the screen. A minimal screening requirement can be roughly estimated from the frequency of positive mutants found either in earlier rounds of directed evolution or from results reported in literature. The frequency of positive mutants for different enzymes and different properties varies, but is usually found in the range of about one out of lo2 to lo’ mutants (Table 4-2). However, it should be noted that the frequency of positive mutants will strongly depend on the fitness of the parent the property, and the strategy chosen to create the mutant library. A theoretical approach to determine the required number of screened mutants is based on the landscape paradigm. Following this paradigm, several studies have shown that, when the landscape is additive, the number of mutants that need to be screened in order to find fitness improvements increases linearly as the wild-type sequence increases in fitness 12’, 15*1. However, as the landscape ruggedness increases, the number of fitter neighbors decreases more rapidly as the sequence becomes 159, 160]. Thus, in order to discover improved mutants, the number of mutants screened has to increase more rapidly on rugged landscapes. This implies that a protein that is tolerant (corresponding to a smooth landscape) can undergo more rounds of mutation and improvement. There is a tradeoff between generating large libraries for a few generations and generating small libraries for many generations. In other words, if the total number of mutants that can be screened is fmed, what is the optimal number of generations? While the improvement in fitness increases with the size of the screening library, the benefit of accumulating stepwise positive mutations over multiple generations is compromised. Both experimental and theoretical studies have suggested that the best method may be short, adaptive walks utilizing small “‘1 . Hu simi ‘ and Aita further studied the effect of the screening cost on the optimal search strategy[78]. They found that screening multiple generations of small libraries has the advantage of evolving more rapidly; however, it has a greater potential of being

4.4 Finding Improved Enzymes: Screening and Selection

I

117

Frequencies of positive mutants found during directed evolution studies. Calculation is based on reported mutants and might not represent all positive mutants. Actual frequencies rnay be higher.

Table 4-2.

Frequency o f positive mutants

Evolved property / rnutagenesis procedure /test

Reference

(tested/positive) 3 ~ 1 0 -(1500/5) ~ 5 ~ 1 0 (2000/1) -~ 2xlO-' (1.5x106/34) 3x10-' (6.4x104/2) 3 ~ 1 0 (103/3) -~ 2xlO-' (2x106/35) 4 ~ 1 0 (2x105/7) -~ I x ~ O -(104/124) ~ 3 ~ 1 0 (300/1) -~ 2xlO-' (1.7x103/4) ~ X I O - ' (1.2x104/1) 1x 10-2 (10~112) 1x10-3 (75o/i) 4xlO-* (150/6)

Tiermostability / PCR / screen Tierrnostability / family shuffling / screen Therrnostability / chemical / selection Thermal and oxidative stability / PCR / screen Activity in organic solvent / PCR / screen Activity at elevated temperature / PCR / screen Activity / cassette rnutagenesis / selection Activity / family shuffling / screen Activity in organic solvent / PCR Activity / family shuffling Functional expression / PCR / screen Enantioselectivity/ PCR /screen Enantioselectivity/ rnutator strain / screen Substrate specificity / focused rnutagenesis / screen

-

trapped in a local optimum. If the cost of screening a mutant is high, then the walk should tend towards many generations of small libraries, whereas, if it is low, it should tend towards fewer generations of large libraries. A powerful strategy for balancing the limitations of throughput and sensitivity is tiered screening (Fig. 4-8). In this method, a series of screening or selection methods with decreasing throughput and increasing accuracy are combined. This strategy has been used successfully for the evolution of subtilisin towards a combination of and for the evolution of an esterase towards increased enantioselectivity[491. A computational method, such as the entropy calculation presented in Sect. 4.3.1.3,can be used to eliininate the portions of sequence space where improvement is unlikely (Figure 4.9)[82 . Pre-screening drastically reduces the number of mutants that have to be experimeritally screened. Another approach to increasing throughput is to use a pooling ~trategy["~-'~~I. This methodology is coiiceptually equivalent to the recombination strategy presented in Section 4.3.2, in which the recombination load is subdivided into multiple generations, thus reducing the required screening effort. Most screens, however, are not sufficiently sensitive to use a pooling strategy to find small improvements. 4.4.2.4 Analyzing the Mutant Fitn'ess Distribution

During the screening, a large amount of fitness data is generated, but only the fitness information of the improved mutants is used to continue to the next round of evolution. The large ensemble of less fit mutants provides a view of the local fitness landscape. By analyzing these data, certain statistical landscape parameters can be

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FACS, selection

Figure 4-8. Strategy o f tiered screening. Series of screening or selection methods with decreasing throughput and increasing accuracy are combined.

+

visual screening, microtiter plates

I

deduced, such as the fitness landscape ruggedness, which can then be used to guide in setting evolutionary parameters. In this analysis, sequencing is time-consuming and expensive, so a sequence cannot be assigned to each measured fitness. The lack of sequencing data means that only the probability distribution of mutant fitnesses can be analyzed. In this section, we discuss some methods that have been developed to extract useful information from the mutant fitness distribution. Several theoretical approaches based on the additivity of mutations have been proposed to analyze the screening data. Urabe and coworkers developed a model that captures additive and non-additive mutational effects in directed evolution and fit their theoretical model to the experimental fitness distribution of catalase I. By investigating the degree of non-additivity of specific properties, they tuned the parameters of the experiment to suit the fitness landscape['"]. In a similar approach, Aita and Husimi proposed that the additive model can be applied to give a rough estimate for the Hamming distance from the wild-type to the optimum, the fitness slope near the wild-type, and the height of the optimum['58].They calculated the expected fitness distribution and compared this to experimental data produced by the mutagenesis of a region of E. coli lac promoter. Based on a fit between the theoretical and experimental distributions, they estimated that the Hamming distance between the wild-type lac promoter and the optimum is 7-10 nucleotide substitutions and the activity could be improved 100-to 1000-fold. Mean-field theory can be used to predict the effects of mutation rate, landscape ruggedness, and parental fitness on the moments of the mutant fitness distribut i ~ n [ ~ 'In ] . this analysis, only the portion of the mutant distribution that is not dead (zero fitness) or parent (unmutated) is considered. The mutant effects are averaged over the transition probabilities. In order to obtain the fitness distribution, two sets of probabilities are required: (1) the probabilities Pi('i(a)that a particular amino acid identity a exists at a residue i, and (2) the transition probabilities that one amino acid

4.4 Finding lmproued Enzymes: Screening and Selection

will mutate into another, la+,. The probabilities PG(a)can be determined through a mean-field approach and the probabilities qa-b are calculated based on the genetic code["', 166168]. Using the mean-field solution, the change in the mutant fitness distribution is captured as the sequence ascends the fitness landscape. By increasing the coupling interactions between residues, the effect of the landscape ruggedness on the moments is calculated. As the fitness of the wild-type increases, the first and second moments increas'e (Fig. 4-9). In other words, as the sequence ascends the fitness landscape, the mutant distribution spreads out (difises) and becomes skewed towards less-fit rnutants (drifts). In addition, the dependence of the moments on mutation rate can be predicted. As the mutation rate increases, both the drift and the diffusion of mutants from the parent increases. Because rugged landscapes have less correlation between parent and offspring fitnesses, the driftdiffusion effect becomes exaggerated as the coupling between residues increases. Through this approach, ii may be possible to model the mutant fitness distribution to experimentally obtain :#tatisticalparameters that describe the fitness landscape. 4.4.3

Selection and Methods to ILink Genotype with Phenotype

The advantage of high throughput, together with minimal experimental and technical effort, makes selection an attractive tool for in vitro evolution. The most commonly used in vivo selection approach links a targeted enzyme property to cell growth through their contribution to resistance[36,37, 88, 126, 1691 or complementation of auxotrophy or genetic defects that block the metabolism of a host Most of these systems allow cells with improved enzymatic strain[", 89. 90. 1"'. properties to grow, while cells with wild-type properties do not. Such systems are especially powerful if the selection pressure can be adjusted as the evolution progresses, for example, by increasing antibiotic concentrations [36, 371, decreasing substrate concentrations or changing the enzyme expression level [ 6 2 ] . If the growth of cells with wild-type activity cannot be prevented and positive mutants contribute only to an increased growth rate, cells with improved variants are usually enriched by continuous culture techniques, dilution series, or detection of the size of tested colonies. A problem for selection arises if enzymes are secreted into the culture medium. This problem was avoided in one case by growing cells in hollow fibers that limit (cross-feedingbetween neighboring colonies [1701. Another limitation of in vivo selection methods is that selection conditions must be compatible with the requirements of the host organisms, which are often very different from conditions under which the enzyme is going to be applied. This is particularly true if the enzyme is to be use'd in an industrial reactor, where conditions often involve extreme temperatures and the substrates are suspended in organic solvents. It might be advantageous to use (other expression hosts that grow under different reaction conditions than the commonly used organisms (E. coli, S. cerevisiae or B. subtilis). Thermophilic hosts with reasonable transformation efficiencies, such as Bacillus steiarothemophilus or Tetrahymena themophilus, have been used for selection for Other in vivo increased thermostability by growth at elevated temperatures [171-1731.

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A

B

"*"I-

0

05

1.5

1

2

25

Si

Figure4-9. A histogram o f t h e residue entropies for the structure o f subtilisin EIg2]. The entropy is a measure o f t h e number o f amino acid mutations that can be made at a residue without disturbing the structure, as determined using a model o f stabilizing interactions (van der Waals, electrostatics, hydrogen-bonding, and the hydrophobic effect). When all amino acids are equally allowed at a residues, then s; = In 20 = 3.0. When a single amino acid is allowed

at a residue, then s; = 0.0 (marked by the arrow). The connected bar i n the center o f t h e graph marks the mean and standard deviation o f t h e histogram. The lines above this bar indicate where beneficial mutations that improved stability (top row) and activity in organic solvents (bottom row) occurred in directed evolution experiments. Most o f these beneficial mutations occurred at residues that are predicted t o have high entropies.

selection approaches are based on infectivity of phages['74, 17'1 rather than on cell growth. This method has been applied to select for proteins with improved Unfortunately, many of the thermo~tability['~~I and stability against proteolysis [l7'1. targeted enzyme properties and catalyzed reactions cannot be linked to cell growth or infectivity of phages. Even if selectable traits exist, it remains a tedious task to guarantee that the targeted enzyme property and not other factors such as substrate uptake or other metabolic steps is limiting for growth. Selections also face difficulties in controlling biological complexity. New in vitro selection methods might reduce some of the limitations imposed by biological complexity of living cells ['* 19* 241. In addition, much larger mutant libraries can be searched by in vitro selection methods if they are used together with in vitro genotype-phenotype coupling systems. However, such methods often select enzymes based on single turnover 17'1, binding of transition state analogs[15o,179,1 " ' or suicide inhibitors [lS1]and therefore do not necessarily reflect enzyme properties of highly active catalysts. Another in vivo approach to couple the genetic information with a screenable or selectable phenotype is phage display, which has been extensively reviewed elsewhere[lg, 21, 23, 182, 1831. Th e most commonly used approach is to fuse the mutage-

4.5 Applications ofDirected Evolution

nized target genes to a coat protein gene of filamentous bacteriophages. After transformation of bacteria with the recombinant DNA, bacteriophages are assembled that display the protein of interest on their surface fused to the coat protein and carry the genetic information for the displayed enzyme in the DNA. The typical diversity of a library produced by phage display is high: 107-1011 different sequences. However, it is difficult to screen for properties other than Furthermore, the folding of displayed proteins occurs in the periplasm of the bacteria, which is often problematic for the functional display of cytoplasmic enzymes. In addition, large and multimeric enzymes are difficult to display. However, a few examples exist where some of these limitations have been overcome, such as alkaline phosphatase [I', P-lactamase glutathione transferase [181] and penicillin G acylase, which is a 86 kDa heterodimeric enzyme[187].Several theoretical models have been proposed to capture the &pamics of phage All in vivo approaches require the transformation of the mutated genes into cells, which limits the library size that can be produced to a maximum of about 109-1010 members [lo2].In vitro approaches circumvent the transformation step by using cellfree transcription/translai ion systems that can produce the mutant proteins directly from the mutated The required link between the gene and the protein produced can be achieved with ribosome display['"] or mRNA-protein fusions [I9'* 1961. These elegant systems allow the generation of protein libraries that contain up to 1013different members, which is a 10 000-foldexpansion of accessible sequence space. Theoretically, such methods can cover all possible triple mutants of an enzyme with 300 amino acid residues if coupled to a suitable selection system. Another approach to coupling a phenotype with its genotype is based on In vitro transcription / translation reactions compartmentalized in water-in-oil emulsions [126, 1391. In principle, each aqueous compartment contains one single gene that is transcribed and translated into protein. In practice, however, many compartments will be empty or contain inore than one gene. All these approaches require suitable screening or selection technology to make use of the larger libraries.

4.5

Applications of Directed Evolution

With directed evolution we can engineer enzyme properties rapidly and with a high probability of success. Many enzymes that have been improved by directed evolution are listed in Tab. 4-3. This powerful biocatalyst engineering strategy creates new opportunities in organic synthesis: new and improved bioconversion processes can be developed and novel compounds that are otherwise inaccessible by classical chemistry can be synthesized. In addition, the molecules created by directed evcllution offer an excellent opportunity for improving our still poor understanding of sequence-structure-function relationships. The specific applications of directed evolution that are described below focus on properties that are important for efficient enzyme production as well as on those that are of special interest for applications in organic synthesis, including enzyme

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4 Enzyme Engineering by Directed Evolution

specificity, activity towards non-natural substrates, and function in non-natural environments. 4.5.1

Improving Functional Enzyme Expression and Secretion

Pharmaceutical or industrial applications of enzymes require their production at a large scale. This is usually done by overexpressing the enzyme in E. coli, Bacillus sp., yeasts or fungi. Many enzymes, however, fail to fold properly in heterologous hosts. This is a problem particularly for membrane-bound, highly glycosylated or disulfide bond-containing eukaryotic proteins. Eukaryotic enzymes such as glycosylases, peroxidases or cytochrome P-450s will require significant improvements in functional expression to make them available in large quantities and at low cost. Enzyme expression can be affected by mutations in the structural gene that may or may not change amino acid sequence or by mutations in regulatory elements that control expression. It was reported that horseradish peroxidase (HRP),a glycosylated heme enzyme, cannot be expressed in functional form in E. coli or yeast. We found, however, that we could obtain significant levels of HRP activity in the supernatant of yeast cultures by accumulating point mutations in the structural gene['56].Furthermore, mutants expressed at high levels in S. cereuisiae were also better expressed in Pichia pastoris. Although far from sufficient for commercial enzyme production, the activity level that was achieved enabled us to carry out further generations of directed evolution to tailor the enzyme for specific applications. Other reports show that intracellular expression of misfolded or unstable proteins can be dramatically improved by directed evolution. For example, directed evolution increased the expression of disulfide-containing antibody fragments in E. coli 50-fold,to reach a level of more than 0.5 g/L[2121. The expression of a wide spectrum amidase of B. stearothemophilus in E. coli was improved 23-fold by two mutations [210]. And, in uiuo fluorescence of the green fluorescent protein was improved 45-fold by increasing its solubility and native folding in E. c ~ l i [ ' ~ ~ I . Secretion into the culture medium is preferred for some industrial enzymes because it can facilitate purification. This is especially true if the protein reaches high concentrations. Schellenberger's group at Genencor devised a method to select for improved subtilisin-secreting mutants in Bacillus[170].Mutants secreting up to five times as much enzyme were found after one round of error-prone PCR and selection. Directed evolution can improve functional enzyme expression and even allow expression of enzymes that are otherwise difficult to produce in recombinant systems. Results obtained by directed evolution, however, will depend on the particular system, and high expression will undoubtedly be best achieved by a combined approach of molecular biology, fermentation optimization and directed evolution.

Activity towards new substrate

P-Lactarnase activity

Activity towards new substrate; substrate specificity

Protection against alkylating agents (gene therapy)

Arsenic resistance

Aminoacylation of a modified tRNA

P-Galactosidase

06-alkylguanine- DNA alkyltransferase

Arsenate detoxification pathway

Arninoacyl-tRNA synthetase Aspartate aminotransferase

:.----I-- :- ~ - 1 r i ; r . .

55-fold increase in activity

43-fold increase in sensitivity to gancyclovir in hamster cells 66-fold increased activity; 1000-foldincrease in substrate specificity 10-foldincreased protection against toxic methylating agent 12-foldincreased rate of arsenate reduction

60- to 150-foldincrease

I W W U - I W I U IIILICa.JC LII I I a u - I u c

* ~ A A r-12

rnethylformarnide 32 000-fold greater resistance to cefotaxime

- 170-foldincrease in 60% di-

>200-foldincrease in half-life at 60-65 "C

Changeeffected

Activity towards P-branched lo5 increase amino and 2-0x0 acids

Substrate specificity (gene therapy)

Thymidine kinase

-

Activity in organic solvents

Subtilisin E

p-Nitrobenzyl esterase

Thermostability

Sra'uiiiryin iiir dLxrlie of Ca2+ Activity towards pNB esters; activity in organic solvent

Target function

Kanarnycin nucleotidyltransferase

Examples of designed enzymes by directed evolution.

Target enzyme

Table 4-3.

DNA shuffling + selection

E. coli

E. coli

E. coli

DNA shuffling + selection DNA shuffling + selection

E. coli

E. coli

Cassette rnutagenesis + selection

DNA shuffling + screening

Cassette rnutagenesis + selection + screening

[621

1111

[ZOO1

~ 4 1

[161

~ 5 1

[99, 117, 1991

E. coli

Random rnutagenesis + DNA shuffling + screening

E. coli

[198!

E. s?!bti!k

DNA shuffling + selection Loci: remG7ed + cassette mutagenesis + screening

[14, 1971 [371

B. subtilis

[167]

Reference

E. coli

+

B. stearothennophilus

Mutator strain + selection Random mutagenesis screening

Organism

Approach

*

4A

a

w N

4

:

P

9

2

8'

9

a6, '

L,

Thermostability

Expression level (total activity of secreted enzyme) Activity at 10 "C

Thermostability

Subtilisin E

Subtilisin E

B. lentus subtilisin

3-Isopropylmalatede. hydrogenase Cephalosporinases

Activity towards moxalactam

Enantioselectivity of hydrolysis of a sterically hindered 3-hydroxy ester Thermostability

Esterase

Subtilisin BPN

Thermostability

pNB esterase

Lipases

Lipase

3.4-fold increase in activity at 70 "C 270- to 540-fold increased resistance

2-fold increase

17 "C increase in T,,, + increased activity at all temperatures 50 x increase in half-life at 65 "C 50 % increase

14 "C increase in T, + increased activity at all temperatures Increase in enantiomeric excess from 0% to 25 %

Improved performance in one-cyclewash Enantioselectivity in hydrol- Increase in enantiomeric exysis ofp-nitrophenyl2-me- cess from 2% to 81 % thyldecanoate Activity towards long-chain 3-fold increase p-nitrophenyl esters

Wash performance

Lipase

Change effected

Target function

(cont.)

Target enzyme

Table 4-3.

B. subtilis B. subtilis

DNA shuffling + screening Random mutagenesis + enrichment in hollow fibers Chemical mutagenesis + screening Spontaneous mutations + selection DNA "family" shuffling + selection

E. coli

Th. thermophilus

E. coli

B. subtilis

E. coli

E. coli

E. coli

Random mutagenesis + DNA shuffling + screening

Mutator strain + screening

In vivo recombination of homologous genes + screening Random mutagenesis + DNA shuffling + screening

S. cerevisiae

Mutagenesis + in vivo recombination + screening Random mutagenesis + screening

E. coli

Organism

Approach

Reference

b

-c 3

z

m

Conversion to monomeric enzyme (solubility)

In viwo recombination

E. coli

B. sub&

Random mutagenesis + DNA shuffling + screening Random mutagenesis + saturation mutagenesis + screening

More resistant to glutaraldehyde and formaldehyde 100-fold increase in half-life

Retention of function after glutaraldehyde cross-linking

Improved thermostability

B-Glucuronidase

Subtilisin S41

Yeast

E. coli

DNA “family”shuffling + screening 110 x greater thermal stability, Random mutagenesis + in viwo recombination + 2.8 x oxidative stability site-directed mutants + screening

Substrate specificity

Glutathione transferase Peroxidase

Stability to peroxide, thermostability

E. coli

DNA shuffling + selection

2.lx1OG-foldincrease in cat. efficiency towards valine

Substrate specificity

Aspartate aminotransferase Found range of specificities

E. coli

Random mutagenesis + step shuffling + screening

5- to 20-fold increase

Increased activity in peroxide shunt pathway, towards naphthalene

E. coli

Cytochrome P450cam

Targeted random mutagenesis + screening

E. coli

E. coli

E. coli

Becomes 10 bp cutter

Random mutagenesis + DNA shuffling + screening

Improved recombination efficiency in E. coli and mammalian cells

DNA “family” shuffling + screening

Oligonucleotide directed codon mutagenesis + selection

Extend recognition site

temperatures in E.coli and mammalian cells; in vitro thermostability

PfiriPnry a t elevated

Gained activity towards substrates poorly degraded by native enzymes; improved activity towards various substrates

Functional monomeric enzyme

EcoRV endonuclease

FLP recombinase

Biphenyl dioxygenases Degradation of polychlorinated biphenyls (PCBs)

Chorismate mutase

11541

P

L,

(cont.) Change effected

13-26 x more thermostable 70-fold activation without cofactor (fully active in the absence of cofactor) Production of tomlene in E.coli

Thermostability

Cofactor (fmctose-1,6-bisphosphate) requirement

Catechol2,3-dioxygenases Lactate dehydrogenase

Kanamycin nucleotidyl transferase B. stearothermophilus amidase Horseradish peroxidase Cytochrome P450 BM-3

Indole-3-glycerolphosphate synthase

Increase expression in E. Coli Increase activity/ expression in s. cereuisiae Substrate specificities

Hydroxylates indole

Increase total activity 40 x

Increase expression (23 x )

Increase 20 “C

7-44 fold improved specificity

Substrate specificity

Thymidine kinase

New carotenoid pathway (substrate and reaction specificity) Confer new catalytic activity (phosphoribosyl anthranilate isomerase) Thermostability

Two-fold improved E

Enantioselectivity

Esterase

Phytoene desaturase and lycopene cyclase

Improved activity stability

Various properties

Enantioselectivity + total ac- Inverted enantioselectivity, 5 x increase in total activity tivity

Target function

Subtilisins

Hydantoinase

Target enzyme

Table 4-3.

E. coli T thermophilus

Rational design + DNA shuffling + selection

DNA shuffling + screeninglselection Random mutagenesis + screening Random mutagenesis + screening Saturation mutagenesis + screening

E. coli

S . cereivisiae

E. coli

E. coli

E. coli

E. coli

E. coli

E. coli

B. subtilis

E. coli

Organism

DNA “family” shuffling + screening

Random mutagenesis + saturation mutagenesis + screening DNA “family” shuffling + screening Random mutagenesis + screening DNA “family” shuffling + screening DNA “family” shuffling + screening DNA shuffling + screening

Approach

Reference

5

?

m

A

-

4.5 Applications of Directed Evolution

4.5.2

Engineering Enzymes for Non-natural Environments

Bioconversion processes performed in organic solvents or at elevated temperatures impart such benefits as increased substrate solubility, decreased viscosity of the reaction medium, altered reaction selectivity and equilibria, higher rates, and reduced risk of microbial contamination. High thermostability also tends to translate to resistance to other denaturants and better long-term stability at lower temperatures. Most natural enzymes are poorly suited for function in organic solvents or at high temperatures, however, and their limited stability and activity in these environments can be a limiting factor for applications in organic synthesis. These properties are good targets for engineering by evolution. Directed evolution has generated a large number of thermostabilized enzymes (see Table 4-3 for example:<);there are too many reports for a comprehensive review here. We will discuss only the general picture that arises from those studies; the interested reader is referred to a recent review that deals with evolution of enzyme stability in greater detail[2131. The increase in thermostability imparted by single amino acid substitutions is usually small and is typically in the range of a 1-2 "C increase in melting temperature or optimal reaction temperature[152,17', 173, 214-21G]. Larger changes are possible, but rare. Significant changes in thermostability therefore require the accumulation of multiple substitutions, e. g., by sequential rounds of mutagenesis or recombination. This strategy has generated 20 "C and higher increases in thermo~tability['~~ "l]. The stabilization mechanisms are consistent with those found in naturally thermophilic ertzymes and include reduction of surface loop flexibility, new hydrogen bonds, altered core packing, helix stabilization and acquisition of surface salt bridges[217]. Although the mechanisms are familiar, most of the changes would have been difficult or impossible to predict. I n nature, thermophilic enzymes tend to be less active at low temperatures than their mesophilic counterparts, which in turn are less thermostable (Fig. 4-10). One popular explanation for this observation is that activity and stability make mutually exclusive demands on enzyme flexibility. Two properties coupled in this way cannot evolve independently. However, directed evolution experiments have shown that lS2,2021. A 17 "C increase in the these properties can evolve melting temperature of a mesophilic esterase was achieved at the same time as catalybc efficiency was increased several fold by random mutagenesis and screening over several generations I'S2, 2131. A similar approach taken with mesophilic subtilisin E generated a 17 "C increase in the temperature optimum and a >200-fold increase in half-life at 65 oC[202].The thermostable subtilisin was also more active than wild-typeover the whole temperature range. Most recently, directed evolution of a psychrophilic subtili~in1~~] led to a 500-fold increase in half-life at GO "C at no cost to its activity at low temperature. The evolved enzyme is more stable than homologous mesophilic subtilisins. The stabilized enzymes contained between 7 and 13 amino acid substitutions. In the studies described above, mutants were screened simultaneously for activity and thermostability, and mutations were accepted only when

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4 Enzyme Engineering by Directed Evolution

Figure 4-10. Enzymes isolated from organisms growing at different temperatures often exhibit a tradeoff between thermostability and catalytic activity measured at low temperature. Enzymes that are both highly thermostable and highly active at low temperatures are rare in nature but highly desired for various applications and can be obtained by directed evolution with relatively few

enhanced thermostability came at little or no cost to activity. If the selection pressure is not maintained, thermostability can easily be lost[218.2191. Creating enzymes that are both more thermostable and more active is particularly exciting for industrial applications. In addition, these studies nicely demonstrate that behaviors of natural enzymes may not necessarily be due to physical limitations intrinsic to proteins themselves, as is often assumed. Instead they reflect what is both relevant to the organism and accessible to natural evolution Directed evolution has also been very effective for increasing enzyme activity in organic solvents 991. For example, the serine protease subtilisin can catalyze specific peptide syntheses and transesterification reactions, but organic solvents are required to drive the reaction towards synthesis. Sequential rounds of error-prone PCR and visual screening yielded a subtilisin variant with twelve amino acid substitutions that was 471 times more active than wild-type in GO% dimethylformamide (DMF) [145, 2201; this enzyme is much more effective for peptide and polymer synthesis. The production of cephalosporin-derivedantibiotics requires a deprotection step usually catalyzed by zinc in organic solvents. Since this step produces large amounts of solvent- and zinc-containing waste material, scientists at Eli Lilly were interested in using an enzyme. Classic screening identified an esterase that catalyzed the desired reaction but performed poorly in the solvents required to solubilize the substrate. Directed evolution was therefore used to try to improve the performance of the enzyme for efficient hydrolysis of an antibiotic p-nitrobenzyl ester intermediate in aqueous-organic solvent mixtures [991. Four rounds of random mutagenesis by error-prone PCR and screening followed by one recombination step improved the esterase activity 50- to GO-fold in 25 % DMF and yielded mutants that performed

4.5 Applications of Directed Evolution

as well in 30% DMF as the wild-type enzyme in water. None of the six mutations found in the best mutant were in direct contact with the substrate and some were as far away as 20 A. Thus, focused mutagenesis in the substrate binding site may have overlooked important beneficial mutations. High product concentrations are important in organic synthesis but often detrimental to enzymes. Scientist at Celgene reduced product inhibition in transaminases1221] which are valuable for the production of chiral amines or amino acids. A single round of error-prone PCR and screening of 10000 clones revealed mutants with better product tolerance that translated to a four-fold increase in volumetric productivity for a substituted amphetamine. 4.5.3 Engineering Enzyme Specificity

Enzymes are particularly valuable for the production of enantiomerically pure compounds, as shown in examples throughout this book. However, the narrow range of substrates that some enzymes accept and the less than impressive enantioselectivities exhibited by others often frustrate attempts to develop new synthetic applications and to commercialize existing ones. Directed evolution can efficiently tune substrate specificity and catalytic efficiency towards non-natural substrates; it can also tailor enantioselectivity,as illustrated in the examples below. 4.5.3.1 Substrate Specificity

Zhang et al. evolved a fucosidase from a galactosidase[''I. Seven rounds of DNA shuffling and screening using a chromogenic fucose substrate yielded a mutant with GG-fold increase in fuco:;idase activity. Kinetic analysis of the purified enzyme revealed a 10-to 20-fold increase in kc,,&, for the fucose substrate and a SO-fold decrease for galactose ( a total of 1000-fold increased substrate specificity for fucose). Kumamaru et al. reco nbined two biphenyl dioxygenases (96% identical) and visually screened for mutants whose substrate range differed from the parents'. These mutants degraded various biphenyl compounds more efficiently and also exhibited oxygenation act]vity for single-ring aromatic compounds on which neither parent was active[13'1. Lanio et al. reported the tailoring of restriction endonucleases EcoRV specificit^[*^]. Focused combinatorial mutagenesis was used to make variants that cleave specific DNA sequences of eight or ten base pairs rather than the six recognized by the natural enzyme. Twenty-two amino acid residues were targeted by oligonucleotidr-directed mutagenesis within three different regions of the enzyme. Screening a total of only 500 colonies over three cycles of mutagenesis was sufficient to find several mutants with high activity and high specificity for AT- or GC-flanked GATATC cleavage sites. Aspartate aminotransferase catalyzes amino group transfer between acidic amino

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4 Enzyme Engineering by Directed Evolution

acids, aspartate and glutamate, and their corresponding 2-0x0 acids. The wild-type activity for P-branched amino acids is barely detectable, but was dramatically increased by directed evolution["! 1'" . The aspartate aminotransferase gene derived from E. coli was subjected to DNA shuming and introduced into an E. coli host lacking the branched-chain amino acid aminotransferase gene and therefore allowing selection by complementation with mutant aspartate aminotransferases. The stringency of the selection was increased during the progression of evolution by omitting the substrate (2-oxovaline)from the medium, shortening the incubation time and decreasing the expression level of the mutant enzymes by manipulating the construction of the plasmid. A mutant with 105-foldincreased catalytic efficiency (kcat&) for P-branched amino acids and 30-fold decrease for the natural substrate was created after five cycles of shuffling and This mutant was further improved to yield a mutant with a remarkable 2.1x10G-foldimproved catalyhc Analysis of the structure of the mutant enzyme efficiency compared to wild-type["l'. complexed with a valine analog provided detailed insight into how the mutations affected substrate binding and demonstrated the importance of cumulative effects of residues far from the active site. The P-450 monooxygenase from Pseudomonas putida was evolved for efficient utilization of hydrogen peroxide in lieu of 0 2 and NADH and for improved activity towards the non-natural substrate One round of error-prone PCR and screening of about 200 000 clones by high-throughput digital image analysis [1461 revealed several mutants with increased activity. Subsequent recombination of five improved mutants yielded several variants with about 20-fold improvements in naphthalene hydroxylation activity over wild-type using hydrogen peroxide as sole cofactor. Fructose 1,6 bisphosphate (FBP) is an allosteric activator of the thermostable L2-hydroxyacid dehydrogenase from B. stearothermophilus, which might be useful for the asymmetric synthesis of chiral compounds. Since FBP is quite expensive, Allen and Holbrook wished to create an FBP-independent Three rounds of shuffling and screening produced a mutant L-2-hydroxyacid dehydrogenase with three amino acid substitutions that is almost as active in the absence of FBP as the wild-type is in its presence. Recently, Schmidt-Dannert et al. reported the molecular breeding of carotenoid biosynthetic pathways in E. coli I2O91. Two different phytoene desaturases were shuffled and expressed in the context of a carotenoid biosynthetic pathway assembled from different bacterial species. Clones containing mutant desaturases were visually screened to identify new carotenoid products. One out of approximately 10 000 colonies turned pink and produced shuffled tetradehydrolycopene instead of lycopene. The new pathway was extended with a second library of shufled lycopene cyclases. Visual screening identified a cyclase with altered substrate specificity that produced the cyclic carotenoid torulene for the first time in E. coli. Complementary to the strategy of creating new polyketides by mixing and matching subunits in a multi-enzyme 2231, the combination of a rational pathway assembly and directed evolution is an exciting opportunity to create libraries of otherwise inaccessible biologically active compounds.

4.5 Applications of Directed Evolution

4.5.3.2 Enantioselectivity

Matcham and Bowen were among the first to apply an evolutionary approach to improve the enantioselectivity of an enzyme for use in chiral synthesis [2211. The wildtype enzyme (an s-selective transaminase) converts a particular p-tetralone to the corresponding amine at only 65 % ee. By screening a mutant library of 10000 variants in a microtiter plate-based assay, they identified 10 mutants that produced the (S)aminotetraline with 80-94 % ee. Sequencing the mutants revealed positions important for enantioselectivity and, interestingly,the existence of synergistic combinations of mutations. The lipase from Pseudoinonas aeruginosa (PAL) catalyzes the hydrolysis of 2-methyldecanoic acid p-nitroplienyl ester with only 2 % ee in favor of the (S)-acid.Keetz and Jaeger used four rounds of error-prone PCK and screening on enantiomerically pure R and S substrates to generate a more enantioselective variant that produced the desired (S)-acid with 81% Additional cycles of error-prone PCK in combination with saturation mutagenesis further improved the enantioselectivity of this enzyme, which hydra’lyzesthe 2-methyldecanoicacid p-nitrophenyl ester with 91 % ee (E= 25.8) in favor of the (S)-acid[121. Bornscheuer et al. improved the enantioselectivity of an esterase from Pseudomonasfluore~cens[~’,’071. In their first report, the enzyme was evolved for hydrolysis of a 3-hydroxy ester serving als a building block in epithilone ~ynthesis[~’1.Isolated plasmids obtained from a mutator strain were transferred into E. coli and plated onto two different kinds of agar plates. One plate contained a pH indicator which shows active clones by a color change. The other plate contained a minimal medium and a glycerol ester as the sole carbon source. Cleavage of the glycerol ester releases glycerol, which leads to growth of active cells. One clone that produced the desired enantiomer with 25 % ee was identified, compared to no enantioselectivity for wildtype. The screen allowed for detection of active clones, but is not sensitive to enantioselectivity; this mirght explain why further improvements in enantioselectivity were not reported. A subsequent report deaicribes the evolution of the same enzyme for the hydrolysis of 3-phenylbutyric acid re:sorufinester using both a mutator strain and error-prone PCK[2071.Mutants were sc:reened for improved enantioselectivity based on a microtiter plate assay using the optically pure R- or S-esters. Both mutagenesis methods generated first-generation mutants with higher enantioselectivity (E=6.6 and 5.8 compared to wild-type E=3.5). Recent results show that directed evolution can also invert enzyme enantioselectivity“”]. The hydantoinase derived from Arthrobacter sp. shows a substrate-dependent inversion of enantioselect-ivitywhich limits its use for the production of certain Lamino acids such as L-methionine (for applications of hydantoinases in organic syntheses see Chapter 12).By accumulation of mutations through sequential rounds of error-prone PCR and. saturation mutagenesis, the enantioselectivity of the hydantoinase was inverted from ee = 40% for the D-enantiomerto ee = 20% for the Lisomer at 30% conversion. Only one amino acid substitution was required for the

I

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4 Enzyme Engineering by Directed Evolution

inversion of enantioselectivity. Furthermore, mutant hydantoinases exhibiting high D-selectivity (ee = 90% at 30% conversion) were also found. The L-selective mutant, whose overall activity was improved 5-fold over wild-type, was co-expressed with a racemase and L-specific carbamoylase in E. coli. This yielded a recombinant wholecell catalyst with an improved hydantoin converting pathway. Application of this whole-cell catalyst for the production of L-methionine resulted in >S-foldimproved productivity for 290 % conversion of the racemic substrate into the optically pure product. The optimization of whole pathways by directed evolution and their introduction into recombinant whole-cell catalysts may offer the possibility of substituting complicated multi-step processes with straightforward single-pot processes. This, of course, is highly desired for industrial applications and a major advantage of biocatalysis over other competing technologies used in organic synthesis.

4.6

Conclusions

The power of directed evolution is now well documented. These methods are robust and are able to improve industrial enzymes in reasonably short times. The first laboratory-evolvedenzymes are now used commercially in laundry detergents [201]; other commercial applications are on the horizon. Directed evolution may well help move biocatalysis from an “enabling tool” to a “lowest cost approach. It also offers new opportunities to engineer multi-enzyme pathways and even whole microbes 224* 2251, which will lead to straightforward single-pot, multi-enzyme bioconversions and new fermentation processes based on “green”resources such as glucose or inexpensive waste materials. Sixteen years after Manfred Eigen and William Gardiner presented the basic algorithm for evolutionary molecular engineering; it is worth commenting on the conclusion of their paper: “... The clones have to be addressable; the analytical methods must combine parallel processing and automatic sampling with sensitivity and speed. With such elaboration and scale, experimental biology might well become ‘Big science’. ’ [751

Today’s tools of evolutionary engineering certainly fulfill these requirements, and directed evolution has in fact emerged as the method of choice for biocatalyst improvement. However, we are only beginning to explore the power of evolutionary design. The most obvious limitations of these methods are still related to the tools. Screening or selection methods require significant development time. This might be reduced by the development of versatile enzyme assays that can be adapted rapidly to specific conditions. The problem will also be reduced by integrating versatile standard analytic systems such as mass spectroscopy, HPLC or capillary electrophoresis into automatic high-throughput systems. The finite sampling capacity of most screening methods and the low versatility of

References I 1 3 3

selection methods will Frobably remain significant limitations. This makes it difficult, if not impossible, to generate surprising new functions that require multiple simultaneous amino acid substitutions. It is clear that more “rational” approaches, based on stmcturelsequence comparisons or computation, will be necessary to target key amino acid positions. Other limitations of directed evolution are inherent in the current mutagenesis and recombination methods, which strongly bias the combinatorial libraries. It is not yet clear how best to create molecular diversity for evolution. What is clear is that many of these questions and limitations can and will be addressed in the near future. The field of molecular evolution used to focus on the past and aimed to explain the existence of today’s fantastic array of biological molecules. Applied molecular evolution is changing this focus to the future, by creating molecules for a biotechnology industry of un limited opportunities.

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Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

I

5 Enzyme Bioinformatics Kay Hofmann

5.1 Introduction

Enzymes are a particular class of proteins which, through gradual developments, have been optimized extensively to catalyze a large variety of chemical reactions ‘1. The primary metabolism enzymes, whose role is the “housekeeping” catalysis of metabolically important reactions, are generally optimized for robustness and high turnover rate. By contrast, the products of secondary metabolism, which comprise e. g. colorants, odorants, hormones and toxins, frequently have complex chemical structures and require biosynthetic enzymes highly optimized for stereo- and regiospecificity. A number of enzymes, many of them from microbial sources, have already proved useful for ex vivo applications in synthetic chemistry. The main part of this book gives an extensive overview of biosynthetic applications of enzymes currently in use. The advent of genome sequencing, both of microbes and other organisms, has lead to a sharp increase in the information available on their enzyme repertoire and metabolic pathways. It is to be expected that these additional insights will soon find their way into biocatalytic applications, leading to a broadened base of synthetically useful enzymes. ( h e consequence of the increased amount of raw genomic data becoming available is the requirement for bioinformatical analysis in order to extract useful information. While there is an extensive literature on bioinformatics algorithms, on protein bioinformatics in general, and on the analysis of particular protein families, there are only a few publications dealing with enzyme-specific issues of bioinformatical analysis. This chapter tries to fill a gap by specifically addressing those aspects of protein sequence analysis that are important for identifying enzymes in genomic sequences, for understanding their mode of action, and for predicting some of their properties. ‘fie problem of understanding the mechanism of an enzyme, particularly when pertaining to the optimization of catalytic properties, is more suitably addressed by analysis of the enzyme’:; three-dimensional structure instead of its sequence 12, ’1. W.hile structural analysis is occasionallyconsidered a subtopic of bioinformatics, this

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chapter will focus exclusively on aspects of sequence analysis. No matter how fast the currently initiated projects on structural genomics proceed, the availability of a protein structure will always lag considerably behind the availability of its seq~ence[&’~. Thus, any piece of information that can be derived from the sequence alone will be useful in its own right. Moreover, many tasks that are commonly believed to require knowledge about an enzyme’s structure, can nowadays be performed by using the sequence alone, given that the appropriate tools are used and the analysis is done properly. Examples include the identification of active site residues or the establishment of extremely distant protein relationships with sequence similarity way below 20 % identical residues [*I. Since the major part of enzyme bioinformatics is based on the results of the comparison of evolutionarily related sequences, the following paragraphs will start (Sect. 5.2) with a brief survey of protein sequence comparison approaches. Comparison of multiple sequences belonging to a single family usually reveals a specific set of conserved residues. When dealing with enzymes, the nature and positioning of the resulting conservation patterns can be indicative of the enzymatic mechanism, of cofactors involved, or of other properties of that particular enzyme family. Thus, Section 5.3 will discuss the conclusions that can be drawn from this type of analysis. Section 5.4 elaborates on the “domain” concept, both in terms of structure and of sequence. Multi-domain organization of enzymes is frequently associated with multiple functionalities, which can occasionally be separated and used for overcoming undesirable regulation mechanisms or even for combinatorial biocatalysis. As in other areas of bioinformatics, specialized databases are of crucial importance for the field of enzyme bioinformatics. Section 5.5 provides an overview of publicly accessible databases that digest and store information on enzymes, pathways and metabolites and make it available for querying. Section 5.6, which also deals with databases, puts a focus on collections of pre-classified conservation patterns characteristic of protein families and domains, both of enzymes and non-catalytic proteins. These databases have become indispensable tools for the recognition and classification of novel enzymes, a task frequently encountered when dealing with genome sequences. Section 5.7 introduces and compares a number of strategies used to mine microbial and other genome sequences for enzymes. Finally, Sect. 5.8 attemps to give an outlook on possible future developments and on the impact ofbioinformatics on the identification and optimization of enzymes for biocatalytic applications.

5.2 Protein Comparison 5.2.1 Sequence Comparison uersus Structure Comparison

It is a widely held tenet that the three-dimensional structure of a protein family is better conserved than the sequence itself. In general, there is some truth to this assumption, although the methods of measuring structural or sequence similarity

5.2 Protein Comparison

are merely operational and the results are difficult do compare. Sequence similarity is fi-equentlyexpressed in terms of “% residue identity”, a measure that cannot be applied to structural comparisons. Conversely, structural similarity is usually expressed by the r.m.s. distance, i. e. the root of the mean square distance of atom pairs, which in turn cannot be applied to sequences. Nevertheless, there are a number of proteins with identical or related function, whose 3D-structures look similar to the skilled eye, while there is no apparent similarity in the amino acid sequence, at least no similarity that would exceed the ‘background noise’ expected from comparing two random sequences[” This apparent superiority of structural comparison has pervaded the recent literature and has fuelled the demand for large-scale projects in structural genomics. Whide such projects undoubtedly have their merits, it should not be neglected that several recently introduced or improved methods of sequence analysis come very close to the sensitivity of structural comparisons. Profile- or Hidden Markov Model-b,ised methods in particular can make use of the enzymespecific conservation patterns discussed in Sect. 5.3, and thus are very well suited to identifylng and classifying even the most distant evolutionary relationship between enzymes. A comprehensive treatment of issues in protein comparison would be beyond the scope of this chapter; the interested reader is referred to some recent ’I. reviews on this topic 5.2.2

Substitution Matrices in Sequence Comparisons

Most sequence comparison methods, including the modern profile techniques, are based on a “dynamicprograming” algorithm introduced by Smith and Waterman in 1981 [l4I. The method strives to find a mathematically optimal alignment from two given sequences. The scoring system used for comparing the alignment “quality”is a compromise between being a good model of biological reality and being computationally tractable. The airn is to maximize a composite score that is calculated from all positions in the alignment. The pairing of identical residues makes a positive contribution to the alignment score; the contribution of non-identical paired residues depends on their similarity as defined by a generally valid similarity table, the substitution matrix. Similar residues are associated with positive scores, while dissimilar residue pairings give a negative contribution to the alignment score. Insertions and deletions of residues in one sequence with respect to the other are allowed, but penalized. Given a proper choice of the substitution matrix and the deletion/insertion penalty, it can be assumed that the resulting mathematically optimal alignment will be close to an evolutionarily optimal one[”, ‘1 (see Fig. 5-1). While there are several possible ideas of what constitutes a “biologically correct” alignment [l’], the context of enzyme comparison would minimally require that corresponding active site residues of the two sequences are properly aligned. The concept of using a substitution matrix, i.e. a knowledge-derived table for judging amino acid sim larities, was introduced by Dayhoff et al. in 1978“’l. Most types of currently used substitution matrices are derived from the analysis of well established alignments, by counting which types of residues are frequently substi-

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I I I I I I A-EDF-ASKL

ADEFGAKL I I II AEDFASKL

Figure 5-1. Influence o f t h e scoring system o n the alignment appearance. The left half o f the figure shows an unreasonable alignment resulting from too low deletion/insertion penalties. The right half shows a better alignment, although the "% identity" score is worse.

tuted for particular amino acids [17-201. The resulting 20 x 20 table has high positive values for identical or highly similar residue pairs, since they can be easily exchanged by evolution without significantly altering a protein's structure or function. Dissimilar residue pairs, by contrast, have negative values, since they are rarely found in homologous positions of related proteins. It is interesting to note that not all identical residue pairings have the same positive value. For example, the Trp -Trp value is very high, while almost all combinations of Trp with non-Trp residues have negative scores. The most likely interpretation is that tryptophane tends to have a very specialized role in protein architecture, which can not really be fulfilled by any other amino acid. By contrast, the Ile Ile value is not nearly as high as the Trp selfscore and is only marginally higher than the Ile Leu score. The likely reason is that most functions of isoleucine can also be fulfilled by other residues such as leucine. All commonly used substitution matrices are derived from a large collection of protein alignments, containing both enzymes and non-enzymes. Thus, favorable residue groupings tend to reflect a structural compatibility rather than a functional equivalence. It would be expected that a substitution matrix derived from particular sets of enzymes would have quite different values for residues that are frequently found in active sites. Both the inequality of residue self-matching scores, and the above mentioned influence of the gap penalties on the alignment appearance show that the "% residues identity" value is not always a good measure for judging the similarity of two sequences. First of all, this value only makes sense when based on a biologically reasonable alignment. Moreover, identical alignment positions containing tryptophane or cysteine can be considered better indicators of sequence relatedness than conserved leucines or isoleucines.

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5.2.3 Profile Methods

The profile method, introduced in 1987 by Gribskov et al.12'] and improved more recently by various groups [22-251, can be considered a generalization of the Smith and Waterman method. The idea of this technique is to abandon the traditionally equal treatment of all positions in an alignment. When using profiles, it is possible to assign a specific weight, a specific substitution matrix and a specific set of gap penalties to each alignment position. The advantage of the additional degrees of freedom lies in the possibility to incorporate a priori knowledge into the calculation

5.2 Protein Comparison

of the alignment score. If, for example, a sequence region is known to be very important for a protein’s structure or function, it can be assigned a higher weight in the calculation of the overall alignment score. Similarly, if a position is known to be part of the active site, a particular substitution matrix could be used for that region. If the structure of a protein is known, sequence stretches corresponding to solventexposed regions could bc assigned “cheaper” gap penalties, since it is known that external loops in protein structures can accommodate deletions or insertions more easily than the structural core. The most typical field of profile application is the alignment of a single sequence to an already established protein family. Starting from a multiple alignment of the protein family, specialized profile construction programs (Table 5-1) look for regions with high conservation, implying a greater importance for the family’s structure or function, and assigns high weights to the preferred residues in these positions. Regions that already harbor gaps in the original family alignment are considered structurally variable and are assigned lower gap penalties. A mathematically very different approach, which is formally equivalent to the generalized profile method, uses so called Hidden Markov Models (HMMs).A more Table 5-1.

Unified resource locators (URLs) for online accessible information sources mentioned

in the text.

Section 2 Profile and HMM construction programs http://www.isrec.isb-sib.ch/ftp-server/pfiools pfiools HMMer http://hmmer.wustl.edu Setzion 5: Enzyme databases http://www.expasy.ch/enzyme ENZYME SWISS-PROT http://www.expasy.ch/sprot BRENDA http://www.brenda.uni-koeln.de KEGG http://www.genome.ad.jp/kegg http://www.genome.ad.jp/dbget/ligand.html LIGAND PDB http://www.pdb.org Enzymes Structures Databa5.e http://www.biochem.ucl.ac.uk/bsm/enzymes UM-BBD http://umbbd.ahc.umn.edu PROMISE http://bmbsgil l.leeds.ac.uk/promise MDB http://metallo.scripps.edu MEROPS http://merops.iapc.bbsrc.ac.uk ESTHER http://www.ensam.inra.fr/cholinesterase Section 6 Domain and motif databases PROSITE http://www.expasy.ch/prosite PFAM http://www.sanger.ac.uk/Pfam SMART http://smart.embl-heidelberg.de BL.OCKS http://www.blocks.fhcrc.org PRINTS http://bioinf.man.ac.uk/dbbrowser/PRINTS INTERPRO http://www.ebi.ac.uk/interpro PROCAT http://www.biochem.ucl.ac.uk/bsm/PROCAT/PROCAT.html - ~ _ _ Section 7: Genome resources http://wit.integratedgenomics.com/GOLD GOLD COG http://www.ncbi.nlm.nih.gov/COG STRING http://www.bork.embl-heidelberg.de/STRING

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extensive coverage of the construction of profiles and HMMs, as well as their application in sequence comparisons, is given elsewhere[10-131. 5.2.4

Database Searches

A frequently encountered task in sequence analysis is to screen a sequence database for relatives of a given protein. In general, all sequence comparison methods that assign alignment scores, including the Smith and Waterman method and the profile method, can be used to that end. A straightforward way is to compare the query sequence (or profile) to each single sequence in the database and sort the results by their respective alignment score. A major obstacle of this approach is the large amount of computation necessary for full dynamic programing algorithms, making these database searches very slow unless running on high-performance computers. Alternative database search methods, including the well known FASTA and BLAST programs[2G-28], are substantially faster through their use of heuristical approximations. While these methods cannot guarantee to find the optimal solution, the minor trade-offin sensitivity is more than compensated for by the immense gain in speed. The heuristical methods are nowadays routinely used for database searches, sometimes combined with a true Smith and Waterman post-processing step for the highest scoring matches. An additional problem that has to be faced when doing sequence database searches is the judgement of alignment score significance. Whenever comparing a query to every sequence in a database and sorting the results by score, it is inevitable that one database sequence will come out at the top of the list. However, this does not necessarily mean that the top-scoring sequence is a true relative of the query: it is quite possible that the database does not contain any relative at all. A number of strategies have been devised to address this question. A common basis is the statistical analysis of the score distribution that would be expected if the database contained only random sequences. For each score obtained in the actual sequence comparison an “expectation value” or “E-value” is then calculated, which corresponds to the probability that the given score is the result of a chance match alone. Low E-values are indicative of significant matches, a value of 0.01 would correspond to a 1 % chance of being a mere coincidence and thus a 99 % chance of being meaningful. Heuristical or exact database search methods, combined with a rigorous statistical analysis of the scores obtained, are very useful tools for identifylng relatives to given sequences, with the profile and HMM methods being the most sensitive ones. The two latter approaches have the additional advantage of allowing an “iterative refinement” process[l1]. In the first step, a profile or Hidden Markov Model is calculated from a starting family of proteins. In the second step, the resulting profile or HMM is used for a database search. Database sequences that give significant scores and are not members of the initial family can, after carrying out some additional tests, be considered new members of the family. The expanded family now contains more sequence diversity and thus offers more possibilities to discrim-

5.3 Enzyme-spec$c Conservation Patterns

inate the important regions from the less important ones. A new profile or HMM calculated from the expanded family can then be used for further rounds of database searches, until no new proteins with significant scores are found any more. This iterative process has becn very successful in detecting even extremely distant sequence relationships with residue identities €15 %.

5.3

Enzyme-specific Conservation Patterns

An iterative profile refinement search, as discussed in the previous section, frequently starts with a relatively small set of sequences with readily visible sequence homology. The multiple alignment will typically contain many conserved residues, some of them being conserved because of their crucial importance for the protein’s structure or function, and others being conserved just for the reason that the genes encoding the proteins did not have enough time to diverge. Subsequent runs of the refinement process will pick up more distantly related proteins, decreasing the number of “chance conservations”.When the iterative process has finished and the protein family under study contains a sufficient number of members and sequence diversity, most of the inkariant residues will have a particular reason for being that well conserved. ‘fie next paragraphs try to interpret frequently occurring classes of “conservation patterns’, meaning the set of totally invariant or at least highly conserved positions in a family alignment. The conservation patterns of enzyme families are different from those ofnon-catalytic proteins and can be used for enzyme identification and classification. 5.3.1 General Conservation Patterns

When analyzing a large number of solved three-dimensional protein structures, it becomes evident that the amino acids buried in the internal regions of the structure are mainly non-polar and form the so called “hydrophobic core”. By contrast, residues that are exposed on the surface and thus in contact with the solvent tend to be hydrciphilic. Data available on structural flexibility also indicate major differences between the rigid core region and the flexible surface. If the conservation patterns of typical protein families are analyzed, a further trend becomes visible. Residues contributing to the hydrophobic core tend to be much better conserved than residues exposed on the outside. As a consequence, highly conserved residues are mostly hydrophobic. There are two other classes of residues that are frequently found to be invariant: glycine and proline. The reason for this preference is again based on the structure. A number of secondary structure elements, such as p-turns, require a very small residue like glycine. Proline too is required for particular structural elements since it introduces some rigidity to the backbone. In general it can be stated that structural reasons contribute most to the

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determination of the average protein family’s conservation pattern. Hydrophobic residues tend to be highly conserved but not invariant, since they can frequently be replaced by related hydrophobic residues. Glycine, proline and, in the case of disulfide-bridges,also cysteine tend to be invariant or nearly invariant at structurally unique positions. The multiple alignment of a typical non-catalytic protein region is shown in Fig. 5-2A. 5.3.2 Active Site Conservation Patterns

The structure of enzymes is governed by the same principles as that of every other protein. However, in addition to the structurally important residue conservation, enzyme families also have the tendency to conserve their active site residues highly. As will be discussed below, most enzyme families have retained a common reaction mechanism and thus a common set of catalybcally important residues. As a consequence, active site positions are not only well conserved but mostly invariant. The set of residues found in catalytic centers of enzymes consist mainly of amino acids that can be protonated and/or deprotonated, or those able to form hydrogen bonds. The exact set of residues depends on the nature of the catalytic mechanism, but serine, cysteine, histidine and aspartate are particularly frequent. In addition, lysine, arginine, glutamate, threonine and tyrosine are occasionallyfound. In several enzyme classes, the high degree of conservation around the active center extends into a second layer, consisting of residues involved in orienting the catalytic side chains by forming a network of hydrogen bonds. As an example of a typical enzymatic conservation pattern, the multiple alignment of the duplicated but very compact catalytic region of phospholipase D type enzymes[2g]is shown in Fig. 5-2B. 5.3.3

Metal Binding Conservation Patterns

A number of proteins contain metal ions, which may serve either a structural or functional role, or even In some proteins, the metal is bound by a particular cofactor, such as haem; other enzymes use the side chains of amino acids for coordinating the metal ion. While bound metals are not restricted to enzymes, a substantial proportion of hydrolases contain Zn2+and other heavy metal ions, which typically contain one unoccupied coordination site that is used for binding and thus activating the substrate to be hydrolyzed. Similarly, a number of redox enzymes coordinate metal ions that are able to change their oxidation state, such as Fe2+/3+ or Cu+l2+.Prominent members among the proteins that bind metals for non-catalytic purposes are zinc-fingers, which frequently bind to DNA or to other proteins, and Ca2+-bindingEF-hand proteins, which serve mainly regulatory purposes. Not all amino acid side chains make good ligands for metal ions. Acidic residues such as aspartate and glutamate are frequently found to coordinate small metal ions like Ca” or Mg2+,while cysteine, histidine and aspartate are frequently involved in

5.3 Enzyme-specific conservation Patterns

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Typical conservation patterns o f three protein classes. Residues invariant or conserved in more than 80% o f t h e sequences are printed on a black or grey background, respectively. A Mainly nonpolar conservation in the UBA domain, a small protein domain that interacts preferentially with u b i q ~ i t i n [ ’ ~ IB:. invariant polar active site residues i n the phospholipase D family[29].C: Nearly invariant metal-binding residues i n the HtpXpteZ4 family o f Zn-containing metalloproteases. Figure 5-2.

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coordinating Zn2+or heavy metal ions. Just as with the amino acids participating in catalytic conversions, those coordinating metal ions fulfil a specialized role and tend to be invariant within protein families. If substitutions are observed, they normally stay within one class of coordinating residues, such as Cys-His or Cys-Asp. Since all side chain bound metal ions require multiple ligands, the corresponding protein families usually have a characteristic Conservation pattern consisting of several invariant positions of the mentioned residue classes. A typical example is shown in Fig. 5-2 C. 5.3.4

Making Use of Conservation Patterns

From what was said in the previous paragraphs, it appears that the specific conservation pattern of a protein family can be used to predict whether the proteins are enzymes, bind metal ions, or rather have a structural or regulatory role. If the proteins are known to be enzymes, the conservation pattern can be used to predict which residues are part of the active site, and possibly also which catalytic mechanism is being used. For example, it would be straightforward to submit a family of structurally uncharacterized proteases to that type of analysis in order to find out whether they are serine proteases, aspartate proteases, metalloproteases, or if they belong to a different class. Moreover, it is possible to compare the family’s conservation pattern with those of other, better characterized enzyme families; this approach will be discussed in more detail in Sect. 5.6. There are, however, a number of caveats that apply to the analysis of enzymespecific conservation patterns. As mentioned previously, the method can be expected to work only in those cases where the sequence family contains enough divergent sequences to discriminate between the important and non-important positions. The large amount of available sequence data from all phyla, in combination with sensitive comparison methods like the iterative profile technique, make it possible to meet this requirement quite frequently. In addition, the analysis is complicated by the presence of catalytically inactive members of enzyme families. There is a rapidly increasing number of reports on those “outsider” proteins, which in the course of evolution have acquired fundamentally different non-catalytic roles. Examples include the transferrin receptor, which is a metal-free and inactive member of an ancient metalloprotease and the neuroligins, which are inactive members of the choline esterase family[32].Those proteins have no selection pressure to preserve the non-functional active site residues and, as a consequence, they are typically replaced by various structurally compatible amino acids. The presence of inactive members in a family alignment means that one can no longer expect a total invariance of the active site residues. Since the non-catalytic proteins usually replace not only one active site residue but rather all of them, there is the chance to identify inactive members or even inactive subfamilies by the concerted loss of conservation in the presumed catalytic positions. Finally, there is a small number of cases, where members of an enzyme family have, in the course of evolution, assumed a different catalytic role, using a different

5.4 Modular Enzymes

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set of active site residues. An example of this situation is the enoyl-CoA isomerasel hydratase family (or crotonase family). The “inner family” comprises various enoylCoA hydratases, isomerases, epimerases and 4-chlorobenzoyl-CoA dehalogenases[331.While these reactions are catalytically distinct, they all share the feature of using CoA-activated substrates and all of them utilize the same set of residues for catalyzing the first common step of the reaction[34.351. However, sensitive sequence comparisons demonstrate a more distant but nevertheless highly significant relationship to the ClpP enzymes, a class of bacterial proteases. This latter family does not use C:oA activated substrates, catalyzes a totally different reaction and uses a distinct set of active site residues grafted onto a very similar structural core[3G* 371. In terms of conservation pattern analysis, this case can be treated similarly to the previous one, i.e. a coordinated loss or change in residue conservation has to be accounted for. It has to be said that a11 of the mentioned complications should be considered exceptions rather than the rule. Overall, an analysis of conservation patterns has been and will continue to be a valuable tool in the identification and classification of new enzyme families.

5.4 Modular Enzymes

A survey of known three-dimensional structure of proteins shows that a sizeable portion of them contain several apparently independent folding units, usually referred to as “domains”. 5.4.1 The Domain Concept in Structure and Sequence

A protein domain, in the structural sense, is a part of the whole protein that folds independently from the rest of the structure and has a hydrophobic core of its own. Residues lying within the domain are mainly in contact with other residues of the same dornain; there are only few interactions between residues within and outside of the domain. In evolutionary terms, genes encoding multi-domain proteins can be explained as fusion products of simpler genes. Nature’s main advantage of using a multi-domain organization of proteins is the possibility of having different functions assigned to different domains of a proteins, which can act more or less independently of leach other. Functional domains that have proven useful can then, by an evolutionary process involving exon shuffling or gene fission/fusion events, be reused in other proteins where they fulfil a similar function[38,391. Apparently, this modular approach to pmtein structure has been very successful: there are several functional domains that can, with only minor modifications, be found in more than 100 different proteins of one organism. While the original definition of a protein domain is based on the structure, it is also possible to detect “ re-usable modules” in protein sequences. Local regions of

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sequence similarity, which are typically found in several proteins per organism, are called “homology domains” and usually correspond roughly to structural domains [401. The self-sufficiency of protein domains makes it possible to insert them into almost any sequence context, thus giving rise to the sharp drop of sequence similarity at the domain boundaries. When comparing two sequences, the presence of a well-conserved homology domain, embedded into a totally unrelated context, makes it necessary to use “local” alignment methods as opposed to “global” ones. Local alignment algorithms do not require the total sequences to match with each other but rather score the best matching region within the sequences. All sequence comparison methods mentioned in Sect. 5.2 support a local alignment mode.

5.4.2

A Classification of Modular Enzymes

A modular architecture is no particular hallmark of enzymes, the highest degree of modularity is typically observed in structural proteins of the extracellular matrix and in proteins involved in intracellular signal transduction. Nevertheless, there are a number of modular enzymes including some which are of interest for biocatalytic applications. A recent review of modular mentions three different but not mutually exclusive types of modularity: i) the separation of substrate recognition and catalytic activity on different domains, ii) modularity in multi-substrate enzymes, which use different domains for binding the two or more substrates that are to react with each other, and iii) modular enzyme systems that catalyze several consecutive reactions of a metabolic pathway. When considering the biocatalytic usefulness of modular enzymes, the first type appears less relevant, since the typical substrate molecules are too small to allow a true spatial separation of recognition and reaction. In physiological situations, however, this type of modularity is highly important wherever the recognition and conversion of macromolecular substrates is concerned. The second type, i. e. modularity in multi-substrate enzymes, might offer some possibilities to change the components involved. If, in a family of transferases catalyzing different reactions, donor and acceptor moieties are recognized by different domains, it could be attempted to swap domains between family members and thus change the specificity of the enzyme, possibly even to one not observed in nature. Examples of this situation are the NDP-glycosyltransferases and the bacterial polyketide synthases. The third type of modularity, the multi-catalyticenzymes using substrate channelling, are of particular interest for synthetic applications. Prominent members are the fatty acid synthases, the polyketide synthases and the non-ribosomal peptide synthases 142-441. In these large proteins, a number of catalytic domains is combined with accessory domains and allows the catalysis of an entire pathway by a single polypeptide chain. Multi-catalyticenzymes frequently use a “swinging arm”, which is covalently attached to the intermediary product of one reaction step, and is subsequently able to present this molecule to the next catalytic domain for further

5.5 Enzyme Databases and Other Information Sources

processing 14’1. As an example, a typical non-ribosomal peptide synthase contains one or more domains forining the “swinging arm”, several domains for activating specific amino acids, several domains for catalyzing the transfer of the activated residues onto the growing chain, and one domain each for loading and unloading the swinging arm. Additional domains that catalyze further enzymatic steps such as redox reaciions or cyclizations are also found. These enzymes, as well as the bacterial polyketide synthases, are promising tools for the biosynthesis of antibiotics and other related natural products. Part of the promise stems from the specificity of the activation reaction and from the fact that the sequence of the reaction process is encoded by the domain arrangement. It has been shown that artificially swapped 471. domains can lead to active enzymes that now synthesize a different So far, not all domains occurring in those enzymes are fully understood by function, and not all attempted domain swappings have lead to viable enzymes. Nevertheless, multifunctional re-prograinable enzymes and other engineered hybrid-enzymeswill undoubtedly have an interesting future in biocatalyhc applications14’1. 5.4.3 Inhibitory Domains

Besides the three types of modularity mentioned, there is a fourth type that is very useful in a physiological setting but tends to be undesired ex uiuo. In a living cell, an uncontrolled enzymatic activity at the wrong place or the wrong time can be deleterious. To avoid is type of complication,many enzymes have acquired inhibitory domains, which are encoded by the same polypeptide as the enzyme itself. Whenever in the biological system the enzymatic activity is needed, the inhibitory region is cleaved off or is neutralized by other methods, e. g. by binding to an activator protein. Biocatalytic applications iypically require permanently active enzymes. Thus, it is desirable to recognize inhibitory domains and remove them before using the enzyme. ,4s mentioned above, bioinformatic methods such as sequence comparisons can help to find those domains and to determine, with some confidence, the likely domain boundaries.

5.5 Enzyme Databases and Other Information Sources

Now that the principles of enzymatic architecture and the corresponding analysis strategies have been highlighted and briefly discussed, an overview of the existing enzyme classes and their properties is needed. Given the more than 4000 different enzyme types, any attempt at only listing them would be far beyond the scope of this chapter. Fortunately, there are a number of specialized databases available, which aim to treat various aspects of enzyme structure and function comprehensively. All of these databases are accessible via the Internet, and a list of the relevant URL addresses,is given in Table 5-1.

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E.C. Nomenclatureand ENZYME Database

The widely accepted basis of all enzyme classifications are the recommendations of the Enzyme Committee (E.C.) of the International Union of Biochemistry and Molecular Biology (IUBMB) [491. Within this system, enzymatic activities are classified by a four-level hierarchy and each entry is described by a set of four numbers. The first number describes the top level and can be either “1”for oxidoreductases, “2” for transferases, “3” for hydrolases, “4” for lyases, “5” for isomerases or “6” for ligases. The meaning of the three lower hierarchy levels depends on the top level group. As an example, glycogen synthase is classified as 2.4.1.11; here, the “2” stands for transferases, the “4” for glycosyl-transferases,the “1” for hexosyl-transferasesand the “11”for the particular subfamily. The ENZYME database[50],maintained by the Swiss Institute for Bioinformatics (SIB), provides a comprehensive list of all IUBMB classifications, together with associated information such as systematic and alternative enzyme names, cofactor requirements, and pointers to the corresponding entry in the SWISS-PROTdatabase of protein sequences[51].In addition, there is a concise free-text description of the reaction catalyzed, together with a description of preferential substrates and products. Currently, the ENZYME database holds entries for approximately 3700 enzymes. 5.5.2

BRENDA

A much more ambitious database that builds on the IUBMB classification is BRENDA, maintained by the Institute of Biochemistry at the University of Cologne. In addition to the data provided by the ENZYME database, the BRENDA curators have extracted a large body of information from the enzyme literature and incorporated it into the database. The database format strives to be readable by both humans and machines. The categories of data stored in BRENDA comprise the EC-number, systematic and recommended names, synonyms, CAS-registry numbers, the reaction catalyzed, a list of known substrates and products, the natural substrates, specific activities, KM values, pH and temperature optima, cofactor and ion requirements, inhibitors, sources, localization, purification schemes, molecular weight, subunit structure, posttranslational modifications, enzyme stability, database links, and last but not least an extensive bibliography. Currently, BRENDA holds entries for approximately 3500 different enzymes. From the wealth of information presented, it is clear that BRENDA is a very important resource for enzymes in organic synthesis.

5.5 Enzyme Databases and Other Information Sources

5.5.3 KECC and LICAND database

The Kyoto Encyclopedia of Genes and Genomes (KEGG) is an effort to reconstruct biological pathways from the gene repertoire found in the genome sequencing The LIGAND database is an associated database of enzymes and their reactions, which is also hosted by the University of LIGAND consists of three different but interconnected segments. The COMPOUND section holds 5600 entries of various compound classes with relevance to enzymatic reactions (substrates, products, inhibitlxs etc.). The ENZYME section contains 3400 entries corresponding to the enzymes themselves. Finally, the REACTION section contains approximately 5200 reactions. In combination with the KEGG/PATHWAY database, the data stored in LIGAND are not only presented in static form but can also be used to calculate biological pathways between a given substrate and product. 5.5.4 UM-BBD

The University of Minnesota Biocatalysis/Biodegradation Database (UM-BBD) is a data repository providing curated information on microbial catabolic enzymes and their organization into metabolic At present, the UM-BBD stores information on approximately 100 pathways with 700 reactions, GOO compounds and 400 enzymes. The database does not try to cover every known enzyme but rather focuses 011 those used for the biodegradation of xenobiotics. UM-BBD is linked to the ENZYME, BRENDA and KEGG/LIGAND databases mentioned above. 5.5.5

Structural Databases

Although not being in the focus of this chapter, structural databases are a most useful resource for the slcientist interested in enzymes and reaction mechanisms. The Protein Data Bank (PDB) is the main repository for all three-dimensional structures of macromolecules including enzymes ['I. Nowadays, most journals accepting manuscripts that describe new structures require a simultaneous deposition of the structural coordinates with the PDB database. In addition to the structure of single protein molecules, the PDB also contains several entries of multi-plotein complexes, or proteins bound to small-moleculecompounds. Of the 14500 entries currently in PDB, there are roughly 7200 enzyme structures. The Enzymes Structures Database, maintained by University College, University of London, Focuses on this portion of PDB and offers links between the E.C. nomenclature of the IUBMB and the corresponding PDB entries.

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Metalloprotein Databases

As mentioned in Sect. 5.3.3, a number of enzymes contain metal ions that participate in the catalytic reaction. Two specialized databases store information on metal ions and other bioinorganic motifs in enzymes. PROMISE (prosthetic centers and metal ions in protein active sites) is maintained by the University of Leeds and focuses on six major groups of metal containing proteins: diiron-carboxylate proteins, haem proteins, iron-sulfur proteins, molybdopterin proteins, mononuclear iron proteins, and chlorophyll containing proteins ["I. The Metalloprotein Database and Browser (MDB) is maintained by the Scripps Research Institute and aims to collect quantitative information on all metal containing sites available from structures in the PDB database[57].The data stored comprises both structural and functional information on the metals bound and the ligands involved. The associated database server allows specific queries for particular site geometries and functions. 5.5.7 Databases for Selected Enzyme Classes

In addition to the above mentioned databases that try to cover the entire world of enzymes, there are a number of more topical databases focusing on particular enzyme families. The MEROPS database, maintained at the Babraham Institute in Cambridge, provides a catalog and a structure-based classification scheme for all proteolytic enzymes ["I. In addition to the classification,the database also provides a digest of published information on the peptidases as well as cladograms and multiple sequence alignments of the peptidase families. The ESTHER database, maintained at the INRA-ENSAM in Montpellier, follows a similar concept but focuses on the a/S fold family of esterases/lipases ["I.

5.6

Protein Domain and Motif Databases

As has been described in Sect. 5.3, the conservation patterns of enzymes are often indicative of the particular family they belong to and can be used for their classification. However, the iterative searches and multiple alignment methods used for their establishment require a certain bioinformatic infrastructure as well as some experience with these issues. If the goal of the analysis is not the detection of novel enzyme families, but rather the classification of a novel sequence into one of the already existing enzyme families, there are a number of protein domain and motif databases that will be useful in this respect[", "1. These databases do not store the sequences themselves but rather work with "descriptors" of protein families and protein domains. These descriptors can consist of the Profiles or Hidden Markov Models mentioned above, but other types are also being used. With a particular

5.6 Protein Domain and MotifDatabases

search engine, typically prsovided with the databases, it is possible to scan one or more unknown protein sequences against large libraries of pre-defined family or domain descriptors. These search engines are publicly accessible via the Internet; the relevant addresses are listed in Table 5-1. Currently, none of the available databases hlas a particular focus on enzymes. Nevertheless, a substantial proportion of the databases discussed below consist of enzyme families or of catalytic or regulatory enzyme domains. 5.6.1

PROSITE

The PROSITE database, maintained by the Swiss Institute of Bioinformatics (SIB), was the first database that tried to catalog functional motifs and domains of proteinslG21.Nowadays, PROSITE consists of two major parts storing different types of descriptors: the “pattern”library and the “profile” The pattern entries of the PROSITE database are based on a regular expression syntax, which emphasises only the most highly conserved residues in a protein family, corresponding approximately to what is termed a “conservation pattern” in Sect. 5.3. I n contrast to the other databases mentioned below, PROSITE patterns do not attempt to describe a complete domain or even protein, but rather try to identify the functionally most important residue combinations, which in enzymes typically correspond to the active site. As an example ofthe PROSITE syntax, “K-x(1,2)-[DE]” would mean a lysine residue, followed by one or two arbitrary residues, followed by a residue that is either aspartate or glutamate. When a sequence is compared with a library of such patterns, (my pattern is found to be either present or absent, no intermediate scores are assigned. Currently, the PROSITE pattern libraries contains approximately 1400 entries. A consequence of the riazid syntax of PROSITE patterns is the restriction that they work well only with those protein families that really contain invariant or at least highly conserved positions. When dealing with catalytic sites of enzymes, this requirement is usually met. However, a large number of protein families and domains are too divergent to be appropriately described in the framework of a regular erpression syntax. To circumvent that problem, the PROSITE curators introduced another secticln of the database using generalized profiles as descript o r ~ [ As ~ ~mentioned ]. above, profiles are based on preferences for particular amino acids rather than on strici requirements. Thus, profiles are suited better for highly divergent protein families and domains, but require a different search engine. An important factor contributing to the usefulness of PROSITE is the extensive documentation of the entries, discussing e.g. the active site residues or the phylogenetic scope of a motif, and also providing links to other databases and to the literature.

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PFAM

PFAM is a database of Hidden Markov Models of protein families and domains, maintained at the Sanger Centre in Cambridge[65].The concept of PFAM is comparable to that of the PROSITE profile section. Similar to the profiles, the HMMs in PFAM have been derived by the iterative refinement procedure mentioned in Sect. 5.2.4. Unlike the PROSITE profiles, which all have been created manually by the curators, the HMMs in PFAM are generated semi-automatically, which accounts for a slightly lower sensitivity. However, this lack is more than compensated for by the facilitated update procedure, allowing the database to grow much faster than PROSITE and to have a shorter generation cycle. Currently, PFAM holds 2727 entries. 5.6.3

Other Related Databases

A number of other protein motif databases should not be left unmentioned. The SMART database is conceptually very similar to PFAM, but the collection of Hidden Markov Models focuses on proteins involved in intracellular signal transduction (“1. The PRINTS and BLOCKS databases are similar to PROSITE and PFAM in that they do not have a thematic focus [67, “1. However, unlike the databases mentioned above, their motif descriptors recognize short non-gapped regions of the proteins. Several other protein motif- and domain-databases and their application in the classification of proteins have been reviewed recently’“, 61]. The INTERPRO consortium, consisting of the curators of various protein domain databases, is currently developing a non-redundant combination database, offering a common search interface [(jg]. A fundamentally different approach is used by PROCAT, which does not describe motifs in linear sequence but rather structural motifs, i. e. combinations of residues that occur in a similar position in the 3D-structure of protein family membersl70. 711

5.7

Enzyme Cenomics

The last few years have seen a rapid increase in the number of completely sequenced genomes; an even greater number of whole genome sequences is near completion. Currently,49 genome sequences have been published in the scientificliterature, and both their DNA sequence and the protein sequence of the predicted gene products are in publicly accessible databases. A taxonomic breakdown of the completed genome sequences shows that five of them belong to eukaryotes,nine to archaea and 35 to eubacteria. So far, the choice of the organism for the genome projects has been based mainly on the general scientific interest or on their biomedical importance. A number of organisms selected for their technological interest are being sequenced as

5.7 Enzyme Genomics

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well. However, the driving force behind those genome projects are mainly commercial entities, raising the question of when and to what extent the sequences will be made known to the public. Databases like GOLD provide information on both finished genomes projects and those What is the relevance of genome sequences to the search for biocatalytically applicable enzymes? At least two different avenues, in this context called “ortholog search and “paralog search have the potential to yield results that are immediately useful. 5.7.1

Ortholog Search

A number of enzymes from microbes or other organism are considered useful but

not totally satisfactory for synthetic applications. Frequently encountered problems include lack of stability,too low catalytic rate, too broad or too narrow specificity, and poor availability of the naiural or recombinant enzyme. In these cases, it might be favorable to replace the enzyme by an ortholog from another organism, i. e. by an enzyme that fulfils exactly the same role in another species and is related to the original enzyme in the same way as the corresponding organisms. One possible rationale for this approach is that not all orthologs in a family have exactly the same properties thus there is a certain likelihood of finding a “better”enzyme in another species by chance alone. A more targeted approach for finding “better”orthologs can also be envisaged if e. g. the goal is to increase the thermal stability of an enzyme, the orthologs from therrriophilic organisms are prime candidates for the desired improvement [731. In general, different life environments and slight differences in metabolic pathways give rise to certain variations in an enzyme’s properties, which can be exploited in the search for optimized enzymes. An obvious prerequisite for this type ofoptimization is being able to find orthologs to given enzymes. A second requirement is that the (sequence derived) ortholog pair has not evolved so far that they catalyze different reactions. When dealing with completely sequenced geiiomes, the search for othologs is frequently straightforward. A number of complications have been described [741, the most frequent being that the gene of interest has been duplicated in one of the lineages. In the cases of absent one-to-one ortholog relationships, it is more appropriate to speak of orthologous groups of genes rather than of ortholog genes. The COG database, maintained at the NCBI, has defined orthology clusters for the publicly available genome sequences and is updated whenever new genome sequences become available[75]. 5.72 Paralog Search

A more demanding problem in the bioinformatic mining of genome sequences is the search for truly novel enzymes. A possible starting point would be the knowledge

that a particular organism possesses an enzyme with the desired specificity, while the corresponding protein sequence is elusive. In order to address this type of

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I question, several approaches are conceivable.One of them is based on the analysis of conservation patterns and phylogenetic relationships in large, non-orthologous enzyme families, and will be discussed in the following paragraph. Other methods, which are not based on sequence homology at all, are highlighted in Sect. 5.7.3. When analyzing the sequence/function relationship in multiple enzyme families such as those collected in PROSITE and PFAM, a number of general rules emerge. It has been mentioned previously that the catalytic mechanism and the active site residues of an enzyme are better conserved than the overall sequences. In most cases, the same is true for the region ofthe substrate that is modified in the course of the reaction, in particular for the type of bond that is being broken or formed. There is a general trend that related enzymes catalyze identical or closely related reaction types but not necessarily with related substrates. While there are numerous examples of this trend, there are only a few counter examples. Almost all members of the u/p fold lipase family catalyze the hydrolysis of carboxyl-esters (or the reverse reaction), no matter whether the substrates are lipids or polar compounds. Similarly, there are several families of phosphoesterases, that act on substrates as diverse as phospholipids, phosphoproteins and nucleic acids, but invariantly cleave a phosphoester bond. Multiple families of acyltransferases exist, which have as a unifying criterion the nature of the acceptor atom (0,N, S) rather than a common recognition feature in the substrates. Among the very few counter examples are the enzymes of fatty acid B-oxidation. As mentioned in Sect. 5.3.4, the enoyl-CoA hydratase and isomerase catalyze different reactions but use a very similar substrate. This particular example can, however, be explained by a common activation step in both reactions. This knowledge can be exploited to search complete genome sequences for proteins that encode enzymes of a given specificity. In the first step, the enzymatic reaction under question has to be analyzed for the nature of the atoms involved and the bonds to be formed or broken. In the second step, the available knowledge base of enzymes and enzymatic reactions has to be screened for any relatives. Useful in this respect are the databases of Sect. 5.5, like ENZYME and BRENDA, which already have classified enzymatic reactions by the necessary criteria. In addition, the protein motif databases of Sect. 5.6 might already have assembled a family of enzymes that catalyze the desired reaction type. If, in this process, a known enzyme is found to catalyze a reaction with a similar mechanism to the desired one, this enzyme sequence can be used for a paralog search in the third step. The expression “paralog” applies to evolutionarily related proteins, either within one species or between species, that are not “orthologs”,i. e. that do not directly correspond to each other. Paralog pairs are expected to catalyze similar reactions instead of identical ones. Finally, in the fourth and last step, the found paralogs can be assumed to be candidates for the missing enzyme and their activity can be verified experimentally. Since paralogs are typically more distantly related than orthologs, their detection frequently requires sensitive protein comparison methods such as profiles or HMMs. Even the detection of orthologs can, under some circumstances, require sophisticated database searching methods, e. g. if the corresponding organisms belong to very distant phyla.

5.8 Outlook 1159

5.7.3 Non-homology Based methods

The methods described in the previous section are all based on homology, i.e. a recognizable sequence relationship caused by a common evolutionary descent. An additional approach to identify candidate genes for a given enzymatic function does not rely on homology, but rather on a peculiarity of bacterial genome organization. Bacteria tend to have proteins belonging to one metabolic pathway clustered in a contiguous stretch of the genome, all present in the same transcriptional orientation. The reason for this clustering is an economy of transcriptional regulation. In most cases, the components of a pathway have to be expressed in a coordinated fashion. This regulation is greatly facilitated by the “operon” arrangement, where multiple bacterial genes are under the control of a single promoter. Again, this knowledge can be exploited when searching for an unknown enzyme with a known involvement in a particular pathway. The first step is the identification of other proteins likely to work in the same pathway as the desired enzyme. In the second step, the genome cf the target organism is searched for genes encoding those upstream or downstream components. In the third step, other genes belonging to the same operon(s) are identified and treated as enzyme candidates unless a different function can be assigned to them. This “operon-approach”to enzyme identification is particularly useful in situations were the gene in question cannot be identified by sequence similarity, e. g. in cases of “non-orthologous gene displacement”. This expression d.escribes a phenomenon that is occasionally observed in bacterial genome compai isons [74s 761. Here, two organisms use similar pathways, where most but not all of the genes involved have a clear one-to-onerelationship. The remaining genes might catalyze exactly equivalent reactions, but are not related at all because the two organisms have recruited members coming from different protein families fix an identical task. Not all bacterial genes in general, and enzymes in particular, are organized in operons. A prerequisite ior the method described above is a reliable detection of operons and the participating genes. Again, evolutionary considerations can help: if related genes, preferably orthologs, occur in a conserved order in several bacterial genomes, this is a clear indication of an operon organization and thus most likely also of a functional coupling. Computer databases of genome organization, such as the STRING system maintained at the EMBL, are useful tools for detecting those relationships [771.

5.8 Outlook

In the recent years, at least two developments have made major contributions to the field of enzyme bioinforniatics. One of them, the advent of whole genome sequencing, is widely recognized for its impact on virtually every field of biochemistry and moleculair biology. By contrast, the development of sensitive sequence comparison

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methods has remained largely unnoticed, although it has made possible a new level of understanding genomic data. The most useful databases of protein families and domains, together with their associated search systems, would not have been possible without profile and Hidden Markov Model methods. These two achievements work synergistically.On one hand, the sequence comparison and classification approaches are required for an efficient functional assignment of genome sequences and also for inter-genome comparisons. On the other hand, the iterative refinement process relies on sequence diversity within protein families and can make use of the genomic data, even in its raw and functionally uncharacterized state. At present, only a fraction of known enzymatic domains and protein families is covered by databases such as PROSITE and PFAM. Within the next years, this fraction will increase, since more genome data will probably uncover a large amount of new enzymes, accompanied by only a minor increase in the number of truly new enzyme families. Eventually, we will see a nearly complete coverage of enzyme families, which will greatly facilitate the identification and classification of any new enzyme sequence that becomes available. A field that will most certainly gain influence in the next years is that of “structural genomics”. Several attempts have been initiated to elucidate the three-dimensional structure of an organisms entire protein complement, or at least a substantial fraction of it. While the results coming from these projects will open a straightforward path to fold recognition, the value for enzyme bioinformatics might not be as high as it might seem. The most useful structural information on enzymatic mechanisms comes from structures where the enzyme is analyzed while binding to a substrate analog or to an inhibitor. These studies, however, require a priori knowledge on the enzymatic properties and the nature of the substrate, which is not available in “blind”high-throughput studies. Another area of intensive research in the field of applied genomics is the gene expression analysis by DNA microarrays and similar methods. As of now, most applications of these techniques are either based on their scientific merits or on medial/pharmaceutical/toxicologicalapplications. It is probably only a matter of time until these methods find their way into research on biocatalysis. Possible applications include the analysis of coordinated regulation of enzymes not linked in operons, or the identification of new enzymes on the basis of their expression pattern. As in all other areas of bioinformatics, databases will play an increasingly important role in managing and integrating the data coming from various sources. A database system meant to be useful for the exploitation of enzymes for synthetic applications would have to encompass information on organisms, their genome sequences and their metabolic pathways, with a special emphasis on the enzymes involved, their reaction types and the nature of the substrates and products. Databases such as KEGG and others have already started to address these questions. However, none of the currently available genome- and pathway-databases are focused on biocatalysis, a fact that will certainly change within the next couple of years.

References I161

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Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

6 Immobilization of Enizymes James Lalonde and Alexey hlargolin

6.1 Introductio'n

Readers of this text are well aware of the promise of enzyme catalysis for the elegant synthesis of complex molecules. However, the practical application of enzymes as catalysts for organic synthesis is often limited by the inherent differences between the way that molecules are synthesized by biological systems and the way they are prepared on the laboratory bench. Nature has designed enzymes to catalyze reactions under physiological conditions, most often in aqueous media at ambient temperature and pressure and at neutral pH with dilute concentrations of reactants. Preparative chemical syntheses, in contrast, usually require high concentrations of reactants and the use of organic solvents to dissolve organic substrates and to shift reaction equilibria. Isolatilm of organic products from water can be complicated by the presence of an amphiphilic protein. While biological systems destroy and regenerate enzymes as they are needed, catalysts used in chemical manufacture must often be recovered and reused many times for economic viability. Immobilization of an enzyme is the most commonly used strategy to impart the desirable features of conventional heterogeneous catalysts onto biological catalysts. By definition, enzyme immobilization is the conversion of an enzyme to a form with artificially restricted mobility and retention of catalytic function [l]. This restricted mobility allows for containment and recovery of the enzyme and is often achieved by either conversion to an insoluble form (for example by linking to insoluble ]particles)or by containment within a semi-permeable barrier (for example entrapment within an ultrafiltration membrane). In the course of this immobilization, enzy:mes can acquire four advantageous properties: - Immobilized enzymes can be used repeatedly or continuously in a variety of -

reactors:. They can be easily separated from soluble reaction products and unreacted substrate, thus simplifjing work-up and preventing protein contamination of the final product.

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G lmrnobilization ofEnzyrnes - The catalytic properties, pH-activity profile and enzyme stability can be enhanced

in the immobilized form. - The control of microbial contamination in solid immobilized preparations is often

simpler than for soluble protein. There are, however, a number of practical limitations on the utility of immobilized enzymes. First, the yield of protein binding is rarely quantitative. Second, in many cases, the cost of the carrier can be quite significant and may even exceed the cost of the enzyme itself. Third, the activity of the resulting immobilized enzyme is usually reduced because of chemical modification of the protein, steric hindrance and mass transfer limitations. Finally, the proportion of active enzyme to the carrier material in immobilized enzyme preparations rarely exceeds 5-10 % w/w, and thus dramatically reduces catalytic activity per weight of solid. Despite the limitations, the great success of enzyme immobilization in diagnostics, pharmaceutical, food and chemical industries is undeniabler2>'1. The decision whether one should use a soluble enzyme preparation or an immobilized enzyme does not have a universal solution and can be decided only on a case by case basis. Ordinarily, if the cost of an enzyme represents a significant portion of the overall cost or if isolation of the final product is complicated by the presence of the soluble protein, the cost of immobilization can be offset by the gains in productivity and improved product quality. The intent of this section is to describe, in general terms with illustrative examples, the features and considerations of these broad classes of enzyme immobilization as they impact their application to biocatalysis. Detailed experimental protocols are available in the original literature and exemplary protocols for these methods are offered in many excellent reviews and texts 141.

6.2

Methods of Immobilization

Thousands of publications and patents detail the immobilization of specific enzymes using an impressive array of strategies. The majority of these immobilization techniques can be divided into four broadly defined groups: - non-covalent adsorption of an enzyme onto a solid support; - covalent attachment of an enzyme to a solid support; - entrapment of an enzyme in a polymeric gel, membrane or capsule; -

cross-linking of an enzyme with a polyfunctional agent.

The first three classes involve the use of a solid matrix to support or entrap the enzyme and to confer the desirable mechanical properties of the solid carrier (Fig. 6-1 A-D). The last method entails covalent linking of the enzyme to itself with no additional support (Fig. 6-1 E). Each ofthe covalent methods requires one or more covalent bonds between reactive groups on the enzyme surface with complementary groups on the carrier, either directly or through the action of a multivalent crosslinking reagent. Covalent attachment methods result in direct chemical modification

G.2 Methods oflmmobilization

Figure 6-1.

Classification o f Immobilization Methods.

of the protein molecule. Non-covalent methods are based on formation of an enz yme/c,arrier complex through simple physical confinement or by electrostatic attraction, hydrogen bonds, van der Waals interactions, and so-called hydrophobic interactions. Matrix entrapment (Fig. 6-1 C) and encapsulation (Fig. 6-1 D) are both considered to be methods of entrapment in this chapter. A summary of the advantages and disadvantages of each of these four classes of immobilization is given at the end of this section in Table 6-1. 6.2.1 Non-Covalent Adsorption

Adsorption of an enzyme to a solid carrier is characterized by the interaction of a protein with a solid surface through reversible, non-covalent binding. The interaction forces in adsorption processes range from relatively strong ionic and hydrogen bondiing to weaker van der Waals forces and “hydrophobic”interactions of the protein with the support. Electrostatic forces of ionic and hydrogen bonding are much stronger than purely hydrophobic ones, and so can afford a tightly bound protein, even in purely aqueous media. Immobilization by adsorption has the advantage of simplicity, is often inexpensive, and does not usually result in disruption of the catalytic protein structure. No chemical modification of the protein or

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I support occurs; however, binding to the carrier is reversible and leaching of the G Immobilization ofEnzymes

protein can be a problem. Moreover, in cases where the binding forces are weak, there is little stabilization of the enzyme tertiary structure relative to the solution form of the enzyme. Interaction of the reaction substrate or products with the support can cause desorption of the adsorbed enzyme. This reversibility of binding can in some situations be advantageous; if the protein catalyst has become inactivated from extended use, the resin can be regenerated by a change in pH or solvent to desorb the deactivated enzyme from the carrier and then fresh biocatalyst added under binding conditions. While more exotic or expensive proteins often warrant the use of covalent binding methods, adsorption is most often used in large scale industrial processes because of the low cost and simplicity. Electrostatic binding of enzymes to polyionic carriers is operationally simple, once appropriate binding conditions are identified. The carrier is first equilibrated using aqueous media at the appropriate pH, solvent composition and ionic strength. An aqueous solution of the enzyme is then treated with the adsorptive solid under conditions of protein concentration, pH, ionic strength, and temperature that have been determined experimentally to give efficient protein binding. If the binding protocol is relatively selective, the immobilization can also effect purification of the enzyme from cell debris and fermentation by-products.The immobilized biocatalyst can be recovered by filtration and then washed. Air drying or washing with a watermiscible organic solvent can be used to give a dried biocatalyst solid for use in nonaqueous media. A disadvantage of this method is that the support materials used in ionic adsorption are polyfunctional and charged and thus can dramatically change the microenvironment of the protein. Steric hindrance to diffusion of substrate and product can also be a problem, due to the short protein-support distance in tight ionic bonding. Ionic adsorption of proteins is one of the oldest methods of protein immobilization and has been used widely in industry. Chibata and co-workers developed one of the earliest industrial biocatalytic processes using an amino acylase adsorbed on diethylaminoethyl (DEAE) carbohydrate resin for the kinetic resolution of amino acids '1. Macroporous synthetic ion exchange resins, based on those originally developed for chromatography and water treatment, are among the most frequently used carrier materials. The protein is bound through association with the side chains of amino acids such as aspartate and glutamate (carboxylate) and lysine (ammonium) through oppositely charged groups on the carrier. The tightness of binding is dependent on the proximity and charge of the binding residues on the protein surface and the carrier. Protein binding can be quite tight if factors which affect ionization such as pH, counter-ion identity, hydrophobicity and ionic strength are optimal. A demonstration of the principles of ionic adsorption is found in the use of glucose isomerase bound to DEAE-cellulose[61 for the conversion of glucose syrup to high fructose corn syrup. Remarkably little enzyme desorption of glucose isomerase is observed over many months, despite the use of elevated temperatures and high flow rates through columns of the resin-bound enzyme. During the lifetime of the catalyst, 1 g of catalyst converts 15000 g (dry substance) of high fructose corn syrup.

G.2 Methods oflmmobilization

However, once inordinate activity has been lost, the protein can be easily removed by a simple shift in pH and then the resin regenerated in situ. Li.pases from Candida cintarctica, Humicola lanuginosa, and Mucor meihei, useful for enantioselective ester hydrolysis or tranesterification, have also been immobilized by ionic attachment to synthetic resins 1'1. For the interesterification of fats and oils, macroporous (>lo0A pore diameter) methacrylate resin cross-linked with divinylberizene gives virtually quantitative binding of the protein. The air-dried resin can be used to catalyze interesterification of oils in the absence of solvent[']. The preparatio'n of Candida anarctica B lipase has been widely used for the resolution of carboxylic acids and alcohols ['I. Ionic attachment to noriionic surfaces can be effected through the intermediacy of a polyvalent metal catiori[lO].Chelation of a transition metal by both the carrier surface and the enzyme results in binding to the surface. Inorganic oxides (such as silica) or polyhydroxylated biopolymers (such as polysaccharides) are used as solid supports in cornbination with polyvalent transition metals capable of binding multiple ligands such as Ti(V).This type of chelation binding is also used extensively in the isolation of genetically engineered proteins by incorporation of a polyhistidine tag sequence. The poly-His sequence chelates tightly to Cu(I1) or Ni(II), providing a selective means for selective recovery of the protein["]. Affinity binding is an important sub-group of ionic protein adsorption methods. Specific electrostatic and hydrophobic interactions between the target enzyme and an immobilized ligand allow for extremely tight, selective binding of the protein of interest. The ligand may be a small molecule or a large protein such as an antibody; however, the loading capacity tends to decrease with the effective molecular weight of the ligand. One of the most frequently used affinity binding systems is the cornbination of biotin with the protein avidin. Avidin is a tetrameric protein which binds four biotin ligands ! ~ specific y ion-pair interactions with a dissociation constant of .about lo-'' M. In a typical embodiment, biotin derivatives that are linked to a reactive fiinctional group are covalently attached to both the solid surface and to the protein. The biotinylated solid is first treated with avidin, and this is followed by treatment with the biotinylated enzyme [I2]. The expense and necessity for extensive ma.nipulakions make this method of affinity binding practical only for aqueous systems and those using very highly valued enzymes. Immobilized enzyme preparations that are bonded only through strictly noniordc, physical adsorption are rare; however, in non-aqueous systems physical adsorption can be a very effective approach. In purely hydrophobic binding, the protein molecule is not solvated by the bulk reaction solvent sufficiently to overcome the weak interaction forces with the solid surface, and so the protein does not desorb from the carrier. In many of these non-aqueous systems, there is thought to be activation of the protein !JY the support providing a more hydrophobic environment which facilitates wetting and interaction with the non-polar substrate and by distribution of the enzynie over a larger surface area. Alternatively, activation of the lipase enzyme by interaction of hydrophobic regions on the protein with the hydrophobic surfaces have been postulated. Dispersion of lipases over a highsurface hydrophobic polymeric carrier such as polypropylene or nylon has been

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shown to activate this enzyme in organic solvent media relative to particles of the untreated protein prepared by lyophilization. Hydrophobic binding of lipases is sufficiently strong to allow their use in purely aqueous media, presumably because of the affinity of this protein for waterloil interfaces. Pate1 has reported that a lipase immobilized on polypropylene could be reused for ten cycles without loss of activity in the kinetic resolution of a key intermediate for semi-syntheticT a ~ o l [ ~ ~ ] . Adsorption on polar, nonionic carriers represents the middle ground in noncovalent attachment, where a combination of hydrogen bonding and dipole interactions helps to bind the protein to the support. The immobilized lipase used in the upgrading of low value fats and oils by interesterification is a successful example of this mode of non-covalent adsorption. In one example, an aerosol of an aqueous solution of the lipase is sprayed onto finely divided silica and then the particles are agglomerated to give the particulate biocatalyst. The simplicity and effectiveness of this adsorption process afford a dried immobilized biocatalyst with sufficient productivity to be used on a manufacturing scale at relatively high temperatures [I4]. 6.2.2 Covalent Attachment

The immobilization of enzymes by covalent attachment to a solid carrier involves formation of a covalent bond between amino acid side chain residues of the protein with reactive groups on the support surface. Covalent attachment is often the method of choice where the protein value is high, minimal protein leaching from the support is required or rational control of the biocatalyst properties is desired. Because of the stronger carrier-protein linkage, the resulting heterogeneous biocatalyst can be much more stable than those prepared by adsorption or entrapment. The most common protein functional groups involved in covalent bonding are nucleophilic amino (lysine, hisitidine and arginine), thiol (cysteine) and hydroxyl groups (serine, threonine and tyrosine) and electrophiliccarboxylate groups (aspartic acid and glutamic acid). The reactivity of these functional groups can be modulated through chemical modification, but this can be detrimental to activity and the extra degree of complexity is not often warranted. Rational control of properties of the immobilized biocatalyst is possible with covalent binding; by choice of the reactive functional group on the support and control of its distribution, the practitioner can control the nature and degree of protein modification and the microenvironment of the immobilisate. Binding with minimal loss of catalytic activity is thought best to occur with residues on the surface of the protein and should involve groups that are remote from the active site of the enzyme to avoid deactivation. In the preponderance of cases, primary amino groups on the protein surface are coupled with electrophilicgroups on the support material. Surface-exposedlysine and arginine residues are allowed to react with electrophiles via alkylation, conjugate addition, imine formation or acylation. Alkylation and conjugate addition proceed with retention of the net protein charge. Less frequently, carboxylate residues on the enzyme are activated for reaction with nucleophilic functional groups on the carrier. Figure 6-2 depicts some of the more commonly

6.2 Methods oflmmobilization

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Figure 6-2.

Examples of Corrimon Carrier Activation Methods.

used combinations of reactive protein groups and activated supports['5]. The choice of reactive group is important; highly reactive groups may result in non-specificovermodification, and chemical functionalization that adds or removes charge from the protein can alter the actikity and stability. There are many examples of more limited and specific attachment methods using reagents selective for less common amino acids. No one support and linker is ideal and the large number of supports and binding method leads to an enormous number of possible permutations. The preparation of covalently bound immobilized enzymes involves treatment of a solution of the protein with the reactive support. Judicious choice of conditions including enzyme concentration, pH, and ionic strength can be used to increase the yield of bound activity. The loading capacity of the carrier can be estimated from the manufacturer's specifications or by titration with reagents specific for the reactive functional group. A competitive inhibitor or high concentration of substrate may be used during attachment to protect the active site and to maintain the active conformation of the enzyme. After incubation, the resin is washed repeatedly to remove unbound protein, and then the free reactive sites are quenched by treatment with an appropriate nucleophilic or electrophilic reagent (for example, glycine or acetic anhydride). Either the solid support or the enzyme may be activated, but to limit disruption of the enzyme tertiary structure the functional groups of the support material are most often actiwated. The activation may occur prior to the coupling reaction (pre-activated supports), or a bi-functional linking reagent may be used to form the bond between

G lmmobilization of Enzymes Table 6-1.

A Comparison of immobilization methods.

Immobilization Method

Advantages

Disadvantages

Adsorption

Simple No chemical modification of enzyme Reversible Often inexpensive

Weak binding, leaching of enzyme Little or no stabilization Non-specificbinding May limit mass transfer

Covalent

Tight binding Wide variety of supports and linkers available Rational control of enzyme loading, distribution and microenvironment

Chemical modification of enzyme Often expensive Activity diluted by carrier May limit mass transfer

Entrapment and Encapsulation

No chemical modification of enzyme Can be simple Efficient for whole cells

Little or no stabilization Environmental changes can disrupt network and cause leakage Often limits mass transfer

Cross-linking

High volumetric activity Compatible with elevated temperature and organic solvents No carrier required Tight binding Efficient for whole cells

Chemical modification of enzyme Little control of particle properties (especiallyfor precipitate and whole cell) Requires crystallization o f enzyme (for CLEC@) May limit mass-transfer

protein and support. A comparison of the various immobilization methods is given in Table 6-1. 6.2.2.1

Carriers for Enzyme Immobilization

The physical and chemical properties of protein molecules are often not compatible with the conditions used in most chemical syntheses, and so fixation to a solid carrier is one effective strategy to alter the properties of the biocatalyst. By binding of the protein to a proportionately large amount of a solid carrier, the bulk properties of the resultant solid biocatalyst are more derived from the carrier than from the protein. Enzymes are subject to denaturation conditions found in typical chemical processing such as high concentrations of organic reagents and high shear forces. Moreover, proteins are water soluble and amphipathic thus causing emulsions on extraction and being difficult to recover and reuse. The carrier-fixed biocatalyst is often more resistant to deactivation by organic reactants or shear and can be recovered by simple filtration. In many cases the recovered biocatalyst maintains catalytic function and may be reused many times. An enormous number of carriers are available for the immobilization of enzymes

6.2 Methods oflmmobilization Table 6-2.

Carrier types

Organic - synthetic polymer

Organic - biopolymer

Inorganic

Polv.amides Nylon Polyalkylene Polystyrene Polyacrylates Polyacrylamide Polyethylene Polypropylene Polyvinyl alcohol Polyvinyl.acetate Polyvinykhloride Polyethylene glycol Polyester Polycarbonate Pol)~~rethane Polysiloxane Phenol-fonnaldehyde

Polysaccharide

Minerals

Cellulose Starch Agarose Dextran Chitin Polyalginate Carrageenan

Sand Pumice Metal oxides Diatomaceous earth Clays

~~~~

~

Proteinaceous

Synthetic

Gelatin Collagen Silk Albumin Bone

Glass, controlled pore glass Zeolites Silica Sol-gel Alumina Metal Oxides Metals

and a wide range of methods have been used for fixation of protein to these carriers. For rapid preparation of laboratory samples, commercially available pre-activated macroporous resins are available. Considerations of the desired properties of the immobilized biocatalyst such as ease of use, mechanical strength, activity density, stability, intended application, cost, and availability help to determine which carriers and methods of attachment are appropriate. In most industrial applications, cost of the support and efficiency of immobilization are paramount, while in biomedical applications binding efficiency and ability to sterilize can be most important. Classification of materials used in solid carriers is given in Table 6-2. When the mass of carrier material is large relative to that of the enzyme, the physical and chemical properties of the carrier (Table 6-5) will, in large part, determine properties of the resultant immobilized enzyme. Often, the carrier will impart mechanical strength to the enzyme, allowing repetitive recovery by simple filtration of the solid particles and reuse of the enzyme. The degree of porosity and pore volume will determine the resistance to diffusion and molecular size selectivity of the biocatalyst. When used in non-aqueous media, dispersion of the enzyme over a large surface area can greatly increase its activity. Table 6-3 summarizes many of the key properties and considerations for enzyme carrier materials. 6.2.3

Entrapment and Encapsulation

Entrapment can be defhed as any system in which an enzyme or whole cell is physically restricted within a confined space or network. This class of immobilization is often extended to include systems where a combination of physical entrap-

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6 lrnrnobilization ofhzyrnes Table 6-3.

Summary of properties and considerations for enzyme carriers.

Property

Examples or typical range

Characteristics and considerations

Binding mode

Covalent, ionic or physical adsorption. Pre-activated or activated in situ.

Binding strength, enzyme stabilization, ease of use, protein charge

Shape

Bead, flat sheet or hollow fiber membrane, amorphous aggregate Crystal

Ease of filtration, Control of diffusion path length and flow properties Simple preparation

Surface area

>SO m2/g

Binding capacity,volumetric activity

Porosity, pore size

71 ml/g,

Resistance to diffusion, molecular weight sieving, flow properties, enzyme retention

Particle size and distribution

2-50 nm

1 pm to 1 mm

Ease of filtration, sedimentation velocity

Density

Resistance to diffusion, sedimenta tion velocity

Safety

Sterility, toxicity, regulatory approval for food and drug use, consumer and worker exposure

Mechanical strength

Resistance to shear, compression, tearing of membranes or particle fracture

Compressibility

Rigid particle to soft gel

Ease of filtration and handling, column packing Swelling, dissolution of carrier, enzyme desorption, controlled dissolution

Solvent compatibility

Hydrophobicity and charge

Hydrophobic to polyionic

Alteration of substrate selectivity, shift of pH/rate optimum, enzyme stabilization, binding force and capacity

Reactive site distribution

Evenly distributed or only on surface

Surface vs. bulk attachment, multipoint attachment, stabilization

Loading capacity

0.1% to 10% w/w

Enzyme/carrier ratio, volumetric activity

cost

“Free”to 1000s of USD/g

Economics, availability for scale-up, catalyst productivity and lifetime

and covalent binding is used. Entrapment immobilization includes enzymes contained within such diverse systems as polymeric matrices, hollow-fiber ultrafiltration membranes, liposomes, cross-linked arrays, or cross-linked whole cells. Depending on the density of the entrapment matrix, the environment of the protein can be similar to that of the protein in the bulk reaction media, and disruption of catalybc activity is relatively minor. The pore structure of the matrix used for merit

6.2 Methods oflmmobilization

entrapment is such that small molecules (substrates and products) are able to diffuse in and out of the matrix, while the macromolecular enzyme is maintained within the network. Mass transfer limitations are almost always an issue with entrapped enzymes and whole cells, since precise control of pore size is usually not possible. Often a certain fraction of the enzyme is able to diffuse from the network, and swelling of these molecular networks by a change in reaction conditions can accelerate this leakage of protein. Entrapment of enzymes,or whole cells in a cross-linkedpolymeric network can be achieved by a number of methods. The most common methods of gelation of a polymer or pre-polymer include: Cross-linking of a pre-formed polymer or formation of a polymer network in the presence of the biocatalyst; Solvent-,temperature-, 'or pH-induced precipitation; Addition of multivalent cations to a polyacid. Polyacrylate and po1yacry:lamide gels have been found to have favorable properties for the entrapment of enzymes and whole cell biocatalysts116]. These gels are sufficiently hydrophilic to provide an environment similar to that of the bulk aqu.eous :solution. Acryla mide or methacrylate monomers, for example, can be polymerized in the presence of enzymes and polyfunctional cross-linkers to form a gehentrapped biocatalyst preparation. The stiffness of the gel and pore size can be controlled by the amount and type of cross-linker used. Higher degrees of crosslinking and short spacer groups give a stiffer gel, while longer spacer groups give larger pores. The average molecular weight of the gel can be influenced by the amount of free radical initiator used, the reaction time and the temperature of polymerization. The particle size can be controlled by mechanically cutting the particles to the desired size or by performing the polymerization under emulsion polymerization conditions. Rhodoc,occus sp. microorganisms which express high levels of nitrile hydratase have been entrapped in polyacrylamide and polyacrylate resins for the conversion of acrylonitrile to acrylamide ["I. Limitations common in cell entrapment such as resin swelling, deactivation du:ring the entrapment, mass transfer limitations of substrate and product were addressed by the control of mixing rate during polymerization, gel density, particle size and resin hydrophobicity. Activation of carboxylate residues in the polymer matrix by conversion to the hydrazide improved retention of the enzyme, presumably through the covalent attachment of lysine side chains on the enzyme surface via amide linkages [*'I. Gelation of polyanionic or polycationic polymers by the addition of multi-valent counter-ions is a simple and common method of entrapment of enzymes and whole cells. In one common embodiment, whole cells or enzymes are entrapped by the drop-wise addition of an aqueous solution of sodium alginate and the biocatalyst to a concentrated solution of a Ca2+salt. The cation acts as a cross-linking agent towards the alginate biopolymer and the droplets precipitate as beads with the biocatalysts entrapped within the network. Although the beads are relatively soft and unstable, this method has been one of the preferred methods for entrapment of whole cells. A

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second commonly used example of this technique, gel formation using K-carrageenan in the presence of high concentrations of potassium salts, has been used for the immobilization of asparatase producing cells for the production of L-asparticacid ['I. Similarly,carrageenan entrapment of yeast cells has been used on an industrial scale for the production of malic acid by the action of fumarase on fumaric Leakage of enzyme is often a problem in these systems, especially on exposure to ion-complexing agents such as phosphate buffer. The mechanical properties and enzyme retention can be improved by treatment with glutaraldehyde or other covalent cross-linking reagents. The term encapsulation has been used to distinguish entrapment preparations in which the biocatalyst environment is comparable to that of the bulk phase and where there is no covalent attachment of the protein to the containment medium (Fig. 6-1 D)C2'1. Enzymes or whole cells may be encapsulated within the interior of a microscopic semi-permeable membranes (microencapsulation) or within the interior of macroscopic hollow-fiber membranes. Liposome encapsulation, a common microscopic encapsulation technique, involves the containment of an enzyme within the interior of a spherical surfactant bilayer, usually based on a phospholipid such as lecithin. The dimensions and shape of the liposome are variable and may consist of multiple amphiphile layers. Processes in which microscopic compartmentalization (cf. living cells) such as multienzyme systems, charge transfer systems, or processes that require a gradient in concentration have employed liposome encapsulation. This method of immobilization is also commonly used for the delivery of therapeutic proteins. Enzyme-membrane reactors represent an interesting subset of macroscopic enzyme entrapment [22]. A semi-permeable ultrafiltration membrane with a sufficiently low molecular weight cut-off restricts passage of the enzyme to the bulk substrate and product phase. In these reactors, the soluble enzyme can be used in a continuous fashion as the product isolation is isolated in a separate vessel from the enzyme. Progress in this application has been facilitated by the availability of solvent-resistant membranes with tighter pore size distributions. The membrane can be used to simply separate the enzyme and bulk substrate and product phases or to separate the aqueous enzyme phase from an organic phase containing substrate and product. The resolution of L-methionineby the enantioselectivehydrolysis of Nacetyl-L-methioninehas been performed on the scale of hundreds of tonnes/year in a continuous process using soluble amino acylase in a membrane reactor[231. An extension of this strategy to cofactor restriction was effected by coupling the cofactor nicotinamide adenine dinucleotide (NAD+) with polyethylene glycol to increase its molecular weight. Co-entrapment of the pegylated cofactor with the soluble enzymes leucine dehydrogenase and formate dehydrogenase in the asymmetric reductive amination of trimethylpyruvate to L-tert-leucine[241 allows thousands of turnovers of the expensive cofactor. In the synthesis of the key chiral intermediate for Diltiazem, a lipase entrapped an asymmetric hollow fiber membrane performs the kinetic destruction of the undesired enantiomer. The membrane serves to maintain an aqueous environment for the enzyme and an interface between the buffer phase and that of an organic phase which contains the substrate phenylglycidate ester [251.

G. 3 Properties oflmmobilized Biocatalysts

6.2.4 Cross-Linking

Imrnobilization by chemical cross-linking without the addition of an inert carrier or matrix can provide the means to stabilize and reuse a biocatalyst without dilution of volumetric activity. A major deficiency in all of the aforementioned immobilization methods is that a substantial amount of a catalytically inert carrier or matrix is used to bind or contain the biot atalyst. In many cases, the amount of carrier is two orders of magnihde higher than the protein catalyst. Unfortunately,direct cross-linking of the enzyme, followed by precipitation of an amorphous solid often results in low activity arid poor mechanically properties and so this method is not often used. Recently, however, cross-linked enzyme crystals have been reported to give many of the desirable properties of immobilized enzymes without the need for a support material (Sect. 6.4.1). Chemical cross-linking of an enzyme within its host cell is a simple and economical method to produce an entrapped or encapsulated biocatalyst, eliminating the need for isolation or purification of the enzyme. Whole cells may be lysed or left intact and then chemically cross-linked by the addition of polyfunctional reagents such as glutaraldehyde or toluene diisocyanate. The mechanical properties of such preparations are poor, but can be improved by the addition of support matrices such as gelatin or synthetic organic polymers (which, technically, are considered to be entrapment methods). Cross-linking of whole cells is an effective entrapment method for relatively stable enzymes that do not require additional stabilization of the support matrix. One of the largest industrial biocatalytic processes, that to produce high fructose corn syrup, can employ the biocatalyst as a crosslinked whole cell preparation. A patent assigned to Nov0[~'1 describes the immcibilization of glucose isomerase via entrapment of the lysed cells of the host organisml within a cross linked network of glutaraldehyde and, optionally, an alkyl diamine.

6.3

Properties of Immobilized Biocatalysts

As with most heterogeneous catalysts, it is often difficult to characterize immobilized enzymes at a molecular level. Most immobilized preparations are often complex mixtures with a distribution of chemically modified protein species. The gross caialytic properties observed are a composite of those of a range of differ-

entially modified individual proteins, often irregularly distributed within the sample. Mass transfer limitations and microenvironment effects further complicate characterization.

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6.3.1 Mass Transfer Effects

The catalytic behavior of enzymes in immobilized form may dramatically differ from that of soluble homogeneous enzymes. In particular, mass transport effects (the transport of a substrate to the catalyst and diffusion of reaction products away from the catalyst matrix) may result in the reduction of the overall activity. Mass transport effects are usually divided into two categories - external and internal. External effects stem from the fact that substrates must be transported from the bulk solution to the surface of an immobilized enzyme. Internal difhsional limitations occur when a substrate penetrates inside the immobilized enzyme particle, such as porous carriers, polymeric microspheres, membranes, etc. The classical treatment of mass transfer in heterogeneous catalysis has been successfully applied to immobilized enzymes [271. There are several simple experimental criteria or tests that allow one to determine whether a reaction is limited by external diffusion. For example, if a reaction is completely limited by external diffusion, the rate of the process should not depend on pH or enzyme concentration. At the same time the rate of reaction will depend on the stirring in the batch reactor or on the flow rate of a substrate in the column reactor. The extent of internal mass-transfer is usually expressed by the efficiency coefficient N, where V,, and Vsolare the rates of the reaction catalyzed by an immobilized and soluble enzyme, respectively. In order to find out whether a reaction is limited by diffusion one can calculate n as a function of the Thiel modulus (Fr)

Where R is the carrier radius, De%the effective diffusion coefficient of the substrate, E is the enzyme concentration in the carrier, and kcat and K, are the kinetic parameters of an enzyme. From a practical standpoint it is important to remember that there are no diffusional limitations as long as substrate concentration S exceeds K,. This condition normally exists at the beginning of many processes catalyzed by immobilized enzymes. At the end of the process, when a substrate is depleted and effective K, may increase because of the product inhibition, the whole reaction may be limited by diffusion. 6.3.2 Partition

The other important phenomenon that, in addition to the mass transfer, occurs when enzymes become heterogeneous catalysts, is the partitioning of substrates, products, inhibitors, metal and hydrogen ions between a bulk solution and a carrier. An elegant and simple theory describing the effect of microenvironment inside the particles of immobilized enzymes on their kinetics, has been developed by the group

G.3 Properties oflmmobilized Biocatalysfs

of K.atchalsky[28].In particular, the theory explains why one often observes shifts in pH profiles of activity with immobilized enzymes; it is due to the redistribution of hydrogen ions between a bulk solution and a carrier. As a practical consequence, one should use a negatively charged carrier if a shift of pH profile to a more alkaline pH is desired and a positively charged carrier if the opposite shift to an acidic pH region is necessary. However these electrostatic effects exist in solutions with low ionic strength and almost disappear when salt concentration increases. In general, the partitioning of substrates and products between a solution and a carrier may occur whenever the character ofa carrier (charge, hydrophobicity, etc.) differs from that of a bulk solution. As a result, the binding constants for substrates (K,) and for products (,KJ with immobilized enzymes may vary dramatically from those observed for free enzymes. 6.3.3

Stability

One of the chief benefits of enzyme immobilization is the ability to use them repeatedly in chemical reactors on a large scale. Usually this cannot be achieved without a significant increase of enzyme stability. It is clear that attachment of an enzyme to a solid surface greatly limits deactivation by intermolecular proteinprotein processes such a:: aggregation or proteolysis. In some cases, this is the only stabilization provided by immobilization. In other cases, immobilization leads to stabilization of the three-dimensional catalybc structure against intramolecular protein denaturation under conditions such as elevated temperature, extremes of pH, organic solvents and oxidants. Protein unfolding can be prevented by multipoint attachment of a protein to a support; however, it is not clear whether this increase in rigidity is generally beneficial to catalytic function. As an approximation, the optimal immobilization is given by the maximum functionalization which results in minimal activily loss. Over modification of the enzyme often results in loss of activity and stability. In some specific cases, covalent multipoint attachment of a prtotein to a solid carrier clearly enhances the resistance to chemical and thermal An increase in the number of both polar (electrostatic) and hydrophobic interactions among the protein molecules when a protein goes from a frce to immobilized environment may also significantly enhance stability of proteins against heat and other denaturants I3O1 by preventing unfolding, aggregation or dissociation of the proteins 13'1. Moreover, observed stabilization effects correlate w th the number of contacts involved[32].The intermolecular cross-linking of proteins by glutaraldehyde and other cross-linking may, in turn, lead to additional thermostabilization of proteins by preventing their unfolding. 6.3.4

Activity of Immobilized Enzymes

On the surface the activity assay of immobilized enzymes is quite simple and is not very dissimilar from measuring the activity of soluble enzymes. In both cases the

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G lrnrnobilization ofhzyrnes Table 6-4.

Parameters for characterization of immobilized biocatalysts.

General description Reactionscheme Enzyme and source Carrier type Method of immobilization

Preparation Method Detailed reaction conditions Dry weight yield Activity left in supernatant Enzyme leakage, conditions

Physical and chemical Kinetics characterization

*

Particle shape, diameter, swelling Compression in columns or abrasion in stirred vessels or sedimentation and abrasion in fluidized bed

Initial rate versus concentration for free and immobilized enzyme pH and buffer effects Diffusional limitations Effect of particle size and enzyme loading Degree of conversion versus residence time Storage stability Operational stability

activity is measured in pmol of substrate per minute per gram of a catalyst under defined conditions (concentrations, pH, temperature, etc.). Yet, the heterogeneous nature of immobilized enzymes poses additional challenges. First, special care should be taken in choosing a representative sample of an immobilized enzyme. Second, the activity of immobilized enzymes is much more sensitive to external parameters, such as stirring, and may be limited by diffusion (see above).Third, the determination of true catalytic parameters is more difficult, since the amount of the active enzyme attached to the carrier is not easy to measure. One has to realize that the k,,,, K, and Ki measured for immobilized enzymes often represent the effective parameters. This is further complicated by surface activation effects in lipases [341. The complexityof the physical and catalytic properties of immobilized biocatalysts and the difficulty in comparison of effectiveness based on literature descriptions has led to the publication of guidelines for the characterization of immobilized biocatalyst^^^^]. The authors suggest that description of parameters listed in Table 6-4 should be the minimum required for characterization of an immobilized preparation.

6.4

New Developments and Outlook

Opportunity for innovation and creativity still exists in the field of biocatalyst immobilization. Despite the tremendous volume of biocatalyst immobilization literature, there is no one technology that is universally applicable and no one technique that can be applied using a generic procedure. The limitations of individual immobilization techniques have been pointed out in each section. Operationally simple adsorption methods often are limited by the lack of stabilization and by protein leaching, especially under aqueous conditions. Restriction of diffusion can be severe for entrapped proteins and cells. Covalent methods often result in protein inactivation and a much higher carrier cost. The combined effects of

G.4 New Developments and Outlook

inefficiency in protein binding, carrier expense, protein inactivation on binding, restricted substrate diffusion, enzyme leaching, and enzyme denaturation during use can result in a tremendous overall activity loss and increase in cost when compared to the native biocatalyst. For example, with most current carrier-fixation technologies, Ra~or[~‘] estimates that a 10- to 25-fold overall increase in cost can be expected in converting an enzyme to its immobilized form. Recent work in the field of biocatalyst immobilization has focused on the development of more efficient systems that employ relatively inexpensive support materials (see for example [141), and in some cases, no support at all (Sect. 6.4.1). 6.4.1

Cross-linked Enzyme Crystals (CLEC@)

Early work in protein X-ray crystallographic structure determination demonstrated that protein crystals could be stabilized by cross-linking with glutaraldehyde13’1. More recently, cross-linked enzyme crystals (CLEC@)have been shown to be highly active and stable heterogeneous biocatalyst preparations 1381. In this method, a polyfunctional cross-linking agent is allowed to diffuse into a protein crystal such that the protein is cross-linked throughout the entire particle. In this case the enzyme is not linked to a carrier, but to adjacent enzyme molecules within the crystal. The protein itself is thus both catalyst and carrier. Electrostatic and hydrophobic contacts within the crystalline lattice, combined with added covalent crosslinkers, help maintain protein activity and stability in aqueous and organic media. It has been proposed that a higher degree of chemical functionalization is possible than with attachment to a two-dimensional surface because the added proteinprotein contacts within the crystal particle stabilize the tertiary structure. Iinmobilization by cross-linking of enzyme crystals appears to be a generic method; however, unique protocols must be developed for each individual protein. Preparation of a CLEC form of many types of proteins and classes of enzymes have been reported including hydrolases, oxidoreductases, carbon-carbon lyases and isomerases. Crystallization of the protein is a highly effective purification step, so

Figure 6-3. Graphic Comparison o f 6 A Zeolite B Channel (A) and 21 A Thermolysin Crystal Pore (B).

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G lmmobilization ofhzymes Figure 6-4. Cross-Linked Enzyme Crystals@ o f Thermolysin, Average Length 40 pm.

that undesired side activities can also be However, conversion of a soluble protein to a cross-linked enzyme crystal form requires development of procedures of protein crystallization that are specific for each protein, so that no generic protocol can be applied. The high degree of porosity (average of 50%) and large pore diameter (20-100 A) of most protein crystals allow relatively unrestricted diffusion of small organic molecules (<2000 Da) within the crystal. This unrestricted diffusion, combined with the absence of an inert carrier, results in maximal volumetric activity for CLEC immobilized enzymes. As with all immobilized biocatalyst particles, mass transfer limitations can become appreciable as the rate of catalysis increases, as molecular radius increases and when larger crystal particles are used. The rate of enzyme catalysis in organic media for hydrolases in CLEC form, long considered to be two or three orders of magnitude lower than catalysis in water, have been shown in some instances to be equal to or greater than that of the corresponding hydrolytic reaction f4O]. Cross-linked enzyme crystals of lipases r3’l, proteases and penicillin acylase [421 have been made on the multi-kilogram scale and used in industrial processes to produce tonnes of drug intermediate per Cross-linkedenzyme crystals, in contrast to untreated protein crystals or cross-linked amorphous precipitate, are mechanically tough. In high shear mixing or repetitive filtration cycles of the CLEC form of penicillin acylase, there was no observable particle attrition. Kasche and Tischer have compared the relative benefits of CLEC immobilization and traditional carrier fixation. The authors conclude that CLEC enzymes follow the same kinetic laws as traditionally immobilized enzymes and that the chief benefits are the lack of inert carrier, operational stability and high volumetric activity An alternative approach to CLEC technology, the cross-linking of protein in solution followed by precipitation or freeze drying has been used for some time to avoid the use of inert carrier (see Sect. 6.2.4). Recently it has been reported by Sheldon that a cross-linked enzyme aggregate (CLEA)formed by first precipitation of penicillin acylase using salt or alcohols followed by chemical cross-linking gives an insoluble enzyme preparation with excellent activity and stability in water and organic solvents l4’1.

6.4 New Developments and Outlook

6.4.21

Sol-Gel

Reetz [4G1 and others [471 have found that entrapment of lipases within a hydrophobic silica sol-gelcan result in ;i biocatalyst whose activity in organic media is enhanced in comparison to the corresponding lipase powder under the same conditions. A silica matrix is generated in the presence of an aqueous solution of the lipase by treating hydrophobic alkyl alkoxysilanes with a catalpc amount of sodium fluoride. The gel is allowed! to set, then dried and crushed to the desired average particle size. Optimization of lipase a’ctivity can be achieved through variation of the hydrophobicity of the gel by choice of the alkyl group of the silane monomer, the use of hydrophilk polymeric add-itives (polyvinyl alcohol or inert proteins), control of water activity and by the waterlsilane monomer ratio. Activation of up to two orders of magnitude in the rate of fatty acid esterification relative to the suspended protein powder h.ave been attributed to the combined effects of dispersion over a large surface area and the interaction of hydrophobic regions of the polysiloxane gel with hydrophobic domains of the lipase. The volumetric activity of these systems can be quite low: protein loadings of 0.1-1 % relative to the carrier are necessary for high activation factors. Moreover, the lipase is for the most part passively entrapped within the gel, so its use would be limited to non-aqueous systems. Organopolysiloxanes carrying pendant groups for covalent binding can be employed for immobilization catalysts intended foI aqueous systems [481. More recently, Ballesteros has extended the use of siloxane gel supports by developing support systems employing glyceryl poly(alkylsi1oxanes)L4’)] and poly(hydroxyniethylsiloxanes)for the gel entrapment of lipases. The authors point out that the higher water solubility of these polymers allow for better control of the protein-polymer ratio. Mechanical properties and protein retention of the preparation were improved by entrapping the poly(hydroxylmethylsi1oxane) lipase adsorllates within a solid cross-linked silicone rubber matrix. The versatile chemistry of silicone polymers allows the tailoring of the hydrophobicity and rigidity of the support matrix. Lipase loadings of 1-5% are described for the poly(hydroxymethyl)silox;ine/silicone polymer composites. Entrapment efficiencies are apparently sufficient to retard protein leaching; however, most aqueous reaction systems reported employed an organic co-solvent which limits enzyme dissolution. These lipase/polymer composites could be used for the kinetic resolution of rac:emic carboxylic acids and alcohols via ester hydrolysis or synthesis with negligible loss of activity over 10 reuse cyclles. 6.4.3

Controlled Solubility “Smart Polymers”

Enzymes are normally used as water-solublehomogeneous catalysts or immobilized onto water-insoluble solid supports. The many advantages of immobilized systems have been outlined in this chapter. Yet insoluble immobilized enzyme preparations can also have serious drawbacks. First, efficiency with macromolecular or insoluble

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lrnrnobihzation ofEnzyrnes

substrates such as proteins or polysaccharides is often limited by diffusion. Secondly, in many chemical processes a product of a reaction is less soluble in the reaction mixture than a substrate. In these cases, the precipitation of a product from the reaction mixture makes product isolation and reuse of immobilized enzyme difficult. Thus, it is highly desirable to create a catalyst which combines the advantages of both water-soluble (homogeneous) and water-insoluble (heterogeneous) catalysts. To combine the salient features of immobilized and soluble enzymes, immobilization can be performed on so-called “smart polymers”[5o]that undergo a reversible phase separation from water with small changes in the environmental parameters (pH, ionic strength or temperature). In the most effective systems, these phase transitions are fast, completely reversible and take place within a narrow range of environmental parameters. Several pH-controlled solubility systems have been developed. They include immobilization of enzymes on carboxymethylcellulose, synthetic polyelectrolytes or on polyelectrolyte complexesrS11. The drawbacks of these systems stem from the fact that precipitation normally occurs at low pH (5.5 or lower), which may lead to inactivation of many enzymes. In addition, the repeated use of acids and bases for pH adjustment leads to accumulation of salts and subsequent loss of enzyme activity or change of precipitation pattern. In this regard, temperature-sensitive polymers [521 may be advantageous if the precipitation can be achieved at temperatures near that of biological systems. One such system, based on oligomers of N-is0 propyl acrylamide [531, has been successfully used in the laboratory for repeated solution-precipitation cycling without significant loss of activity.

References

K. Buchholz, J.Klein, Methods Enzymol 1987, 135, 3-30. W. Hartmeier, Immobilisierte Biokatalysatoren, Springer Verlag, Berlin, 1986, 14-16. 2 A. Liese, K. Seelbach, C. Wandrey, Industrial Biotransformations, Wiley-VCH, Weinheim, 2000. 3 E. Katchalski-Katzir,Tibtech 1993, 11, 471-478. 4 Methods in Enzymology, Vol. 44,K. Mosbach, Editor, Academic Press, 1976. Immobilised Enzymes, 1. Chibata (ed),Wiley, New York, 1978. J. F. Kennedy and J. M. S. Cabral, Solid Phase Biochemistry, w. H. Scouten (ed),Wiley Interscience, New York, 1983. Methods in Enzymology, Vol. 135, 136 and 137, K. Mosbach (ed),Academic Press, 1987. Protein Immobilization, Fundamentals and Applications, R. F. Taylor (ed), Marcel Dekker, Inc., New York, 1991. Industrial Application of Immobilized Biocatalysts, A. Tanaka, T. Tosa, T. Kobayashi (eds),Marcel Dekker, New York, 1993. Methods in Bio1

technology, Vol. 1, Immobilization of Enzymes and Cells, G. F. Bickerstaff (ed), Humana Press, NJ, 1997. Progress in Biotechnology,Vol. 15, Stability and Stabilization of Biocatalysts,A. Ballesteros, F. J. Plou, J.L. Iborra, P. J. Halling (eds), Elsevier, New York, 1998. 5 T. Tosa, T. More, N. Fuse, I. Chibata, Enzymologia 1966, 31,225-238. 6 S. Pedersen, in: Industrial Applications of Immobilized Biocatalysts,A.Tanaka, T. Tosa, T. Kobayashi (eds), M. Dekker, New York, 1993. Antrim, R. L., Lloyd, N. E., Auterinen, A. L., Staerke 1989,41, 155. 7 P. Eigtved, EP 0 140,542 B2 1985. 8 P. Eigtved, US 5,156,963 1992. 9 E. M. Anderson, K. M. Larsson, 0. Kirk, Biocat. Biotrans. 1998, 16, 181-204. 10 J. F. Kennedy, J. M. S . Cabral, M. R. Kosseva, M. Paterson in: Immobilised Cells and Enzymes: A Practical Approach, j. Woodward (ed),IRL Press, Oxford, 1985, 345-359.

References I183 11 K. Ng, D. W. Pack, D. Y Sasaki F. H. Ar-

nold, Langmuir 1995, 11,4048-4055. 12 H. E. Swaisgood,X. L. Huang, M. K. Walsh in: Methods in Biotechnology,Vol. 1, Immobilization of Enzymes and ICells. G. F. Bickerstaff (ed),Humana Press, NJ, 1997, 13-20, 13 R. N. Patel, A. Banerjee, R. Y. KO, J. M. Howell, w. S. Li, F. T. Comezoglu, R. A. Partyka, F. T. Szarka, Biotechnol. Appl. Biochem. 1994,20,23-33. 14 M. W. Christensen, K. M. Larsson, H.-j. Deussen, 0. Kirk, Nachwachsende Rohst. 1998, 10,98-105. M.W. Christensen, 0. Kirk, C. Pedersen, WO 99/33964,1999. 15 For a more icomprehensive reliew, see J. M. S. Cabral, j. F. Kennedy in: R. F. Taylor, Protein Immobmilization: Fundamentals and Applicationij. Marcel Dekker, New York, 1991. 16 D. C. Cram R. j. Kazlauskas, B. L. Hirschbein, C.-H. Wong, 0. Abril, G M. Whitesides, Methods Enzymol.. 1987, 136, 263--280. G K. Skryabin, K. A. Koshcheenko, Methods Enzymol. 1987, 135, 198-216. 17 Y. A:jhina, EL. Suto, in: Industrial Application of Immobilized Biocatalysts. A. Tanaka, T. Tosa, T. K.obayashi (eds), Marcel Dekker, Inc. New Ycirk, 1993, 91-107. 18 M. I Yutaka, S. Isoji, S. Shuji, K. Yasuo, JP 58149680,1983. 19 T. Sato, S. Takamatsu, K. Yamamoto, I. Umemura, T. Tosa, I. Chibata, Enzyme Engineering 1982, 6, 271-272. 20 I. Takata, K Yamamoto, T. Tosa, I. Chibata, Eur, J . Appl. Microbiol. Biotechool. 1979, 161--172. 21 M. Kierstan, M. P. Coughlan, in: Protein Imnnobilizakion:Fundamentals and Applications. R. F. Taylor (ed), Marixl Dekker Inc., New York, 1991, 13-71. 22 L. Giomo, E. Drioli, Tibtech 2000, 18, 339--349. U. Kragl, in: Industrial Enzymology, 2nd edn. T. Godfrey, S. (ed),1996, 273--283. 23 A.S. Bomniarius, K. Drauz, IJ. Groeger, C. Waridrey, Membrane, in: Chiralty in Industry. .4. N. Collins, G. N. Sheldake, J. Crosby (eds), John Wiley & Sons, Chichester, U. K., 199.2. K. Drauz, Proceedings of'Chiral '93 USA, 1993. 24 A. S. Bomniarius, M. Schwarm, K. Drauz, J . Mol. Cat. B: Enzymatic 1998, .i, 1-11.

D. R. Dodds, j. L. Lopez, C. M. Zepp, S. Brandt WO 90/04643,1990. 26 S. Amotz, T. K. Nielsen, N. 0. Thiesen, US 3,980,521, 27 L. Goldstein, Methods Enzymol. 1976, 44, 397-443. 28 E. Katchalsky, I. Silman, R. Goldman, Adv. Enzymol.1971,34,445-536. 29 C. Mateo, 0.Abian, R. Fernandez-Lafuente, J. M. Guisan, Enzymol.Microb. Technol. 2000,27,509-515. 30 J , M. Guisan, Enzyme Microb. Technol., 1988, 10, 375-382. 31 A.M. Klibanov, Anal. Biochem., 1979,93, 1-25. 32 V. V. Mozhaev, Tibtech 1993, 1I , 88-95 and references cited therein. 33 S. S. Wong, Chemistry of Protein Conjugation and Cross-Linking, CRC Press, Boca Raton, 1993. 34 A. L. Pavia, V. M. Balcao, F. X. Malcata, Enzymol. Microb. Technol. 2000, 27, 187-204. 35 K. Buchholz, J. Klein, Methods Enzymol. 1987, 135,21-30. 36 P. Rasor, Immobilized Enzymes in Organic Synthesis, in: Chiral Catalyst Immobilization and Recycling. De Vos, Vankelecom, Jacobs (eds), Wiley-VCH,Weinheim, 2000, 97-122. 37 F. A. Quiocho, Methods Enzymol.197644, 546-558. 38 N. St. Clair, M. Navia,]. Am. Chem. Soc. 1992, 114,7314-7316. 39 J.J.Lalonde, C. Govardhan, N. Khalaf, A. M. Martinez, K. Visuri, A. M. Margolinj. Am. Chem. Soc. 1995, 117,6845-6852. J. J. Lalonde, A. L. Margolin, M. Navia, Methods Enzymol.1997,286,443-464. 40 N. K Khalaf, C. Govardhan, j. j. Lalonde, R. A. Persichetti, Y. F. Wang, A. L. Margolin, J. Am. Chem. SOC.1996,118, 5494-5495. 41 R. A. Persichetti, N . L. St. Clair, J. P. Griffith, M. A. Navia, A. L. Margolin,1.Am. Chem. SOC. 1995, 117, 2732-2737. 42 J. Lalonde, Cur. Opin. Drug Disc. DW. 1998, 1,272-277. 43 J. Lalonde, in: Enzyme Technologies 2000 Proceedings 5, IBC, Southborough, MA 2000 44 W. Tischer, V. Kasche, Tibtech 1999, 17, 326-335. 45 L. G o , F. van Rantwijk, R. A. Sheldon, Org. Lett. 2000, 2, 1361-1364. 46 M. T. Reetz, J. Simpelkamp, A. Zonta, Ger. 25

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Pat. Appl. DE-A 44,08152.9,1994. M. T. Reetz, A. Zonta, J. Simpelkamp, Angew.Chem. Int. Ed. Engl. 1995, 34, 301-303. 47 L. M. Ellerby, C. R. Nishida, F. Nishida, S. A. Yamanaka, B. Dunn, J. S. Valentine, J. I. Zink, Science, 1992, 255, 1113-1115. S. Braun, S. Shtelzer, S. Rappoport, D. Avnir, J. Non-Cryst. Solids 1992, 147,739-743. 48 F. Wedekind, A. Daser, W. Tischer, WO 9516773,1994. 49 I. Gill, A. Ballesteros,J . Am. Chem. Soc. 1998, 120,8587-8598.1. Gill, E. Pastor, A. Ballesteros,J. Am. Chem. SOC.1999, 121,

9487-9496. I. Gill, A. Ballesteros, Tibtech 2000, 18,282-296. 50 I. Y.Galaev, B. Mattiasson, Tibtech, 1999, 17, 335-340. 51 L. Cong, U. Dissing, R. Kaul, B. Mattiasson, J . Biotechnol 1995, 42, 75-84. K. Hoshino M. Taniguchi, H. Ueoka, M. Ohhwo, C. Chida, S. Morohashi, T. Sasakura, J . Ferment. Bioeng. 1996,82, 253-258. 52 J. I?. Chen, M. S. Hsu,J. Mol. Catal. B: Enzym. 1997,2,235-241. 53 G. Chen, A. S. Hoffman, Bioconjugate Chem. 1993,4,509-514.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

7 Reaction Engineeringfor Enzyme-CatalyzedBiotransformations Manfied Biselli, Udo Kragl and Christian Wandrey

7.1

Introduction

The application of isolated enzymes to preparative organic synthesis on an industrial scale is a matter of active research worldwide. Since the late sixties, immobilized enzymes have been used in amino acid production in continuous processes on a large scale['* 1' . In the late seventies, the use of soluble enzymes, especially in membrane reactors, broadened the scope of enzyme technology[3.41 and opened the way to simultaneous use of more than one enzyme for complex conversions especially coenzyme-dependentbiotransformations ['-I. In the early 1980sthe use of enzymes was extended further to involve organic solvents L8, 'I. Enzymes are used as catalysts for large scale bioconversions e.g. glucose isomerase in the high fructose corn syrup (HFCS) process[''], penicillinase in the synthesis of semisynthetic penicillins ["I, as well as aminoamidase[12,131 and aminoacylase in the production of L-amino acids. Additionally, a variety of processes for fine chemical synthesis has been developed, e. g. for amino acids, peptides and a broad spectrum of other optically active substances [15-231. Based on the classical methods of enzyme isolation and characterization and the screening for appropriate microorganisms, about 3200 different enzymes are known today and are listed with E. C. numbers[", 241. Modern methods of genetic engineering give access to sufficient quantities of enzymes by overexpression in microorganisms, thus reducing costs of enzymes [25-281. Enzyme reaction engineering allows further reduction of the amount of enzyme consumed per kilogram of product. Therefore, costs of enzymes do not necessarily dominate the overall cost of production, but they are often, still, a major factor. The productivity of enzyme reactors typically exceeds the level of 100 g product per litre of reactor volume per day (extreme values of 25 kgL-ld-l are reported)[*'] and consequently is no longer far below the productivity of classical chemical synthesis. The aim of this contribution is to illustrate some basic aspects of a strategy to optimize an enzymatic process, starting from considerations focusing on the enzymes used and the reactions involved. Additional investigation of the enzyme

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7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

Downstream

-

processing

Enzyme

El- Substrate

Microorganism

--

-

Chemical catalyst

Figure 7-1.

Process design.

kinetics yields a deeper insight into the process and is the basis of final optimization of process performance by reaction engineering methods. A detailed process optimization is of great significance especially for pilot and production scale.

7.2

Steps of Process Optimization

Considerations about process design precede any detailed process optimization. Process design includes the choice of an appropriate substrate, of a catalyst and of methods for downstream processing in order to gain the final product in a defined purity (Fig. 7-1). The situation may be exemplified by showing the different catalytic methods of asymmetric synthesis of L-phenylalanine, starting from six different substrates (Fig. 7-2). Additionally, fermentation processes, using glucose as the carbon and energy source, have been developed [30-321. Within the scope of this chapter, only enzyme-catalyzed biotransformations are considered, presuming a defined product (especiallyimportant in the context of this book is the demand for high optical purity of the product), - a defined enzyme (criteria for the usefulness of an enzyme are its activity, selectivity, stability, coenzyme dependency and its kinetic constants; process optimization studies have to cover these subjects and therefore may influence the choice of the enzyme, e. g. using an enzyme from a different source), - defined substrate(s) (the selection of a substrate including the method of enzymatic transformation is determined by availability and price of both enzyme and substrate (see Fig. 7-2). -

The aim of process optimization is to find process conditions defined by - high conversion of the substrate, -

high selectivity of the reaction, high optical purity of the product,

7.2 Steps ofprocess Optimization I187

COOH

0

Figure 7-2.

I

I

I

I

Asymmetric syntklesis o f L-phenylalanine.

- high space-time yield (productivity)of the process, -

low enzyme and coenzyme consumption per unit mass of product.

The overall process development basically consists of three steps (Fig. 7-3).

The jrsf step is the investigation of the reaction system. This can be further classified into two parts: First of all the enzyme properties have to be examined, considering the following aspects:

- reaction catalyzed, -

substrates, cosubstrate:;, coenzymes, effectors, inhibitors,

- dependency of enzyme activity on substrate concentration, temperature and pH

value,

- dependency of enzymestability on pH, temperature, oxidizing agents, - dependency of enzymeselectivity on reaction conditions, -

necessity of immobilization for enzyme stabilization,

- c:onsiderationson the use of organic solvent/cosolvents,

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7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations 1 Investigation of the reaction system

Enzyme properties *

activity

Parameters

*temperature

'

stability

*

selectivity

I I

Properties of reaction system

I

*

substrate product concentration

thermodynamic equilibrium

* complex reaction

system

*

other properties

" solvent

J 1 Selection of reaction conditions

2 Investigation of reaction kinetics

Measurement and mathematical modelling of enzyme kinetics

*

non-enzymic reaction kinetics

*

mass transfer phenomena

Selection of reactor

3 Investigation of reactor kinetics

Q@Ite@

_ Parameters _

*

conversion

*

selectivity

*

productivity

*

enzyme/ coenzyme consumption

*

of reactor performance

*

4)

Figure 7-3.

residence time concentration of

- substrate(s)

- enzyme(s) - coenzyme

Selection of reactor conditions

Steps of process optimization in enzyme reaction engineering

~

7.2 Steps of Process Optimization

Then the other properties of the reaction system have to be characterized. The following topics are of major concern: - Equilibrium constant of reversible reactions,

Structure of the reaction system. The term “structure” should include the sum of the chemical reactions occurring within the reaction system, e. g. parallel reactions, consecutive reactions (see Eq. (4)), coupled reactions such as in the case of non-catalyzed reactions occurring alongside coenzyme regeneration (seeEq. (49)), the enzymatic reaction. - Influence of the reaction conditions (pH, temperature, concentration of substrates, organic solvent/cosolvents)on the reaction system and on the equilibrium constant, - Other properties such as pH- and temperature effects of the reaction and solubility of substrates and products. -

There are reciprocal relationships between the parameters summarized above. On the one hand enzyme stability measurements strongly depend on the concentrations of substrates, coenzymes, buffers etc. in the assay. On the other hand the choice of an appropriate concentration ].eve1is a consequence of the enzyme kinetics investigated afterwards. A compromise has to be found between different optimization criteria e. g. a lower temperature leads to a reduced enzyme activity but results in a higher enzyme stability. In the example ofthe oxynitrilase reaction (Eq. (12)) a low pH value is a prerequisite for high enantiomeric purity of the product but lowers enzyme activity. As a consequence, only a rough optimization can be carried out at this level. The investigation of the reaction system ends with a (preliminary) selection of reaction conditions. This is a prerequisite for investigation of the reaction kinetics as it makes no sense to measure kinetics without knowing whether the kinetic assay conditions are favorable for the final process. The second step in process optimization is the investigation of the reaction kinetics. This step aims at a detailed analysis of the kinetics of the reaction system under the optimized reaction conditions chosen above. The following topics have to be covered: - Measurement and modeling of the kinetics of the enzyme-catalyzed reaction@), - Measurement and modeling of the kinetics of the non-enzyme-catalyzedreac-

tion(s), - Measurement and modeling of mass transfer phenomena. These will occur in

reaction systems which consist of two or more phases, such as in the case of immobilized enzymes or in the case of liquid/liquid two-phase systems. This aspect will not be covered and the reader is referred to the literatu~-e[~~-~’I. The second step results in ;I kinetic model for the whole reaction system and a choice of an appropriate reactor based on the reaction kinetics. Criteria for reactor choice will be disussed in Sect. 7.5.1. The Jinal step includes tlne reactor design and the prediction of reactor performance depending on operation conditions. The scope of points to consider is as follows:

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7 Reaction Engineeringfor Enzyme-Catalyzed Eiotransformations

- Formulation of the reactor kinetics by combining the kinetic model of enzymatic

and non-enzymatic reactions and mass balances of the reactor. Description of the reactor performance as a function of parameters such as substrate concentration, enzyme and coenzyme concentration and residence time. - Optimization of reactor performance based on the following criteria: substrate conversion, selectivity, space-time yield, enzyme and coenzyme consumption. -

The extent of optimization in the development of an enzymatic reaction may be described as follows: - Studies with one-enzyme systems on a laboratory scale with the aim of small scale

production or verification of the desired concept do not require a detailed kinetic analysis. The properties of the enzyme and of the reaction system should be investigated to choose convenient assay and reaction conditions (e.g. temperature, pH value, substrate and cosubstrate concentrations). Typically some essential enzyme data are provided by the enzyme supplier or can be taken from handbooks [411. - Also some basic kinetic data of the enzyme should be known, especially Michaelis and inhibition constants. For example, an enzyme suffering substrate inhibition, when used at a high substrate concentration, will exhibit only a little activity, whereas at low concentrations the reaction will run quite well. - Dealing with complex systems (two or more coupled enzymatic reactions or reactions with coenzyme regeneration) a complete kinetic investigation and computer simulation of the reaction system is very helpful to achieve the desired selectivity and yield of reaction (e.g. by choosing a sensible substrate and coenzyme concentration, enzyme ratio and reaction time). A case study is available [42, 431 exemplifylng the production of L-tert-leucine by reductive amination and simultaneous coenzyme regeneration. - To optimize a production process all topics listed above have to be treated in detail to achieve the optimum process performance.

7.3

Investigation of the Reaction System 7.3.1

Properties of the Enzyme

The purpose of studies on enzyme properties is to select favorable reaction conditions for the investigation of enzyme kinetics. The choice of assay conditions has to be performed very carefully, and it has to be proven that the assay conditions are as close as possible to the reactor conditions of the final process. This aspect cannot be stressed too much! The influence of all relevant reaction parameters on enzyme activity, selectivity and stability has to be considered. Parameters determining the enzyme properties are

7.3 Investigation ofthe Reaction System

I

191

listed below (the most important of them have been discussed already in Chap. 7.1): - Substrates, cosubstrates, coenzymes, effectors, inhibitors

pH value Temper(1ture Buffer Organic solvents/cosohents Ionic strength Viscosity of the medium Enzyme modification by - immobilization - covalent modification of the enzyme - Redox potential (oxygen sensitivity) - Heavy metal ions - Influences resulting from reactor conditions - shear stress - effects of the reactor material - surface effects (adsorption on reactor surfaces, liquid/liquid or gas/liquid interfaces).

-

-

Measurement and modeling of the influence of concentrations of substrates, cosubstrates, coenzymes and inhibitors on enzyme activity form the central subject of enzyme kinetics and are discussed in Sect. 7.4. A few of the aspects of practical importance are as follows: First of all the pH value and the temperature of the assay have to be chosen. The pH-optimum ofthe enzymc is determined by measuring the effect of pH on activity. It has to be recognized that the location of the pH-optimum depends on -

the substrate,

- the choice of buffer, - the reaction medium (organic cosolvents change dissociation equilibria at the

enzyme), the ionic strength, - chemical derivatizatiort or immobilization of the enzyme. -

In addition the temperature dependency of enzyme activity must be measured, also yielding an optimum curve. This “temperature optimum” depends on the assay corlditions,especially the incubation time, and is not, on its own, useM to identify a reasonable reaction temperature. Instead of this, the temperature stability of the enzymehas to be determined. To that end the enzyme is incubated with all relevant reaction components in test tubes, changing the temperature while keeping all other parameters constant. The assay conditions have to be as close as possible to the conditions relevant in the final process. In particular, stability measurements have to be performed in the presence of a relevant concentration of all substrates and coenzymes which have a stabilizing influence on the enzyme. Typically, the remaining activity is plotted versus incubation time and the deactivation rate can be taken from the slope of a half logarithmic plot (Fig. 7-4).

192

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Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

1

0.5

65°C

0.1

I

0 Figure 7-4.

5

10 Time (h)

15

20

Influence o f temperature on enzyme stability; example: a r n i n o a ~ y l a s e l ~ ~ 1

The activation energy of an enzymatic reaction (typicalvalues 20-GO kJ/mol) is far below the activation energy of thermal denaturation (values between 200 and 600 kJ/ mol) [451. Therefore, by lowering the temperature, the deactivation rate decreases more rapidly than enzyme activity and more product is obtained during the mean lifetime of the enzyme. In practice, the temperature is lowered until enzyme stability is acceptable or other denaturation effects become dominant. Lowering of reaction temperature is of course limited by the solubilities of the substrates and the freezing point of water. Similar measurements of enzyme activity and stability have to be performed, varying the other parameters mentioned above. The dependency ofenzymestability on p H and ionic strength are quite important. The same method may also be used for immobilized enzymes. In order to identify influences of the reactor operation, enzyme stability has to be Figure 7-5 demonstrates that the deactivameasured under the process tion rate may be greatly enhanced by the conditions of reactor operation and is influenced by the reactor material. The decrease in enzyme activity in a continuous reactor causes a decreasing conversion of the substrate. Only in the case of low conversion (compare to “initial reaction rate”, see Sect. 7.4.1),the decrease of enzyme activity is proportional to the decrease of conversion and may be calculated from these data. Alternatively, the remaining activity in the reactor may be measured by taking samples of the enzyme (especially when using membrane reactors) or by adding fresh enzyme, until the initial conversion is re-established. In stirred tank reactors, the product of enzyme activity and residence time determines the conversion (see Sect. 7.5.1). Therefore,

7.3 Investigation ofthe Reaction System

0

5

10

15

20

Time (day) Figure 7-5.

Deactivation o f Neu5Ac-Aldolase depending on the reactor material[47].

the flow rate can be lowered until the initial degree of conversion is reached again and the residual enzyme activity can be calculated from the flow rates. The question of enzymt: immobilization typically depends on the stability of the soluble en'zyme under process conditions. Not only activity and stability but also selectivity of reactions may become the most important criterion in the selection of the reaction conditions. The impacts of additional reactions and of selectivity problems will be discussed in the following section. 7.3.;! Properties of the Reaction System 7.3.2.1 Thermodynamic Equilibrium of the Reaction

In the case of a hypothetical reaction (Eq. (1)). kl

S-P k-I

Enzymes, as catalysts, accelerate the reaction rates (a kinetic factor). The forward reaction and the back reaction are accelerated to the same degree. The position of equilibrium (a thermodynamic measure), which is not influenced by the enzyme, yields information about the maximum conversion and therefore is of basic importance for process development. Two examples will serve to demonstrate this principle.

I

193

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I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

Example: Hydrolysis of acetyl-L-aminoacidsby aminoacylase Aminoacylase stereospecifically cleaves N-acetyl-L-aminoacidsto acetate and Laminoacid, a reaction well known for the production of optically pure L-aminoacids (Eel. (2)).

-

aminoacylase

N-acetyl-L-amino acid

L-amino acid + acetate

(2)

The equilibrium conversion for hydrolysis of the acetyl-L-aminoaciddepends on the initial oncentration. Only at zero concentration may one hundred percent conversion be attained (Fig. 7-6). 100

96 h

92 S

.-0

2

9

s S

88 84

80 0

100

200

300

400

500

[N-acetyl-L-methionine](mmol L-') Figure 7-6. Hydrolysis of N-acetyl-L-methionine by arninoacylase: equilibrium conversion as function of substrate concentration [441.

In the industrial acylase process, an increase of both substrate concentration and conversion is desired to reduce costs, but, as a thermodynamic principle, in cases of increasing mole number during the reaction, the equilibrium conversion decreases with rising substrate concentration.

Example: Synthesis of N-acetylneuraminic acid by aldolase N-Acetylneuraminic acid aldolase catalyzes the cleavage of N-acetylneuraminicacid (NeuSAc) to N-acetylmannosamine (ManNAc) and pyruvate (Pyr).The reverse reaction can be employed to synthesize N-acetylneuraminic acid, which plays an important physiological role as a terminal sugar residue of glycosylated proteins L4'] (Eel. (3)).

ManNAc+Pyr

-

NeuSAc-aldolase

-

NeuSAc

(3)

1.o

-,

,

-

-,

I

_ _. -. _ -.- -

I

’ ,

0.10 mol L-’ ManNAc

_.-.-.-

*.-,*

-.C.-. .C. .C.‘

0.01 mol L-’ ManNAc

. C .

0.4

-

-

O . O . C

0 ’

- 0 .

-

/

0.

0.2

”

25 “C pH 7.5

K = 29.7 L mol-’

0.0 .

I

Figure 7-7. Synthesis o f N-acetyl-neuraminic acid with NeuSAc-aldolase: equilibrium conversion as a function of the concentration o f both

To obtain a high conversion, high concentrations of both substrates have to be used since ManNAc is more expensive than pyruvate, the latter is used in excess (Fig. 77). Bloth examples demonstrate how the maximum conversion can be influenced by the choice of appropriate substrate concentrations (without changing the equilibriurn constant I&). Additionally, a disadvantageous equilibrium position can be overcome by -

continuously adding a mbstrate,

- continuously withdrawing a product, e. g. by crystallization, extraction, electrocomplexation (aspartame process [521), dialysis [49-511, - coupling to an irreversible reaction (see Eq. (49)), - choosing reaction conditions where kqis changed to a more favorable value: X,, depends on pH if acids or bases are involved in the reaction, X , depends on temperature,

k& may be influenced liy c o s ~ l v e n t s [ ~ ~ ~ .

7.3 2 . 2 Complex Reaction Systems: The Existence of Parallel and Consecutive Reactions

In this chapter, two examples demonstrate that, in addition to the desired enzymecatalyzed (conversionof the substrate S to the product PI, other reactions have to be considered, e. g. a parallel reaction of S to P2 or a consecutive reaction of P1 to P3.For a hypothetical reaction scheme, see (Eq. 4).

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7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

The following definitions are set up (all stoichiometric coefficients are set to one): Conversion x:

The conversion xs of the substrate S stands for the ratio of substrate consumed to the initial substrate: ([S]o-[S]= [PI]+[P,]+[P,]). Selectivity 0:

The selectivity opl,s is defined as the product P1 formed in relation to the substrate S consumed. Yield Y:

The yield YpI,s is defined as the ratio of the product PI formed to the initial substrate

so Sometimes, these definitions are mixed up in the literature. Eq. (8) describes the relationship between conversion, selectivity and yield: Yp,,, = X, .(Jq,s (yield = conversion. selectivity)

(8)

The values of yield and selectivity become equal at quantitative conversion of the substrate. The following diagram may elucidate the relation of these three parameters (Fig. 7-8). Additionally, the following measurements are defined: Space-time yield:

The space-time yield STYpl is the amount of product PI produced per reactor volume V and time with the dimension of kg L-ld-’. The space time yield is also referred to as “volumetricproductivity”. Enantiomeric excess:

eel,, =- iP,l-[P*l [PIl+[P21

7.3 Investigation ofthe Reaction System I197

l.O?

&= 1.0

1.04

1.c

1.(

TPlS=

YPlS=

0.8(

0.8:

X,= 0.6

c P l s = Ypls=

0.830.5

Figure 7-8.

Interrelation o f conversion, selectivity and yield in the case ofthe reaction scheme

Eq. (4) (PI being the main product; for definitions see Eqs. (5) to (7)).

The enantiomeric excess eepl of the enantiomer PI is defined as surplus of PI in relation to the sum of enantiomers. Kinetic parameters are defined in Sect. 7-4. Two examples will illustrate the implications of complex reactions in enzyme reaction engineering. Exumple: Enzyme-catalyzed peptide synthesis An instructive example ofthe occurrence of selectivity problems is available from the field of enzymatic peptide synthesis. For instance, the following complex reaction scheme shows where parallel reactions and consecutive reactions occur. It describes the synthesis of Tyr-Arg from the electrophile Tyr-OEt and the nucleophile Arg-NHz by simultaneous use of CarboxypeptidaseY (CPD-Y) and Peptide amidase (PA)from orange flavedo[54](Eq. (11.)).

?=: Tyr-OH

Tyt-OEI "Electrophile"

[Acetyl-enzyme]

PA

Tyr.Arg.NH,-

Arg-NH2

(2)

CPD-Y

Tyr-Arg

(3)

-

(11)

"Dipeptide"

Tyr-OH + Arg OH

(4)

"Nucleophile"

The protease CarboxypeptidaseY (CPD-Y)reacts with the tyrosine ester forming an acylenzyme activated complex, which is further converted in a competitive way. Aminolysis of the acylenzyme by Arg-NH2 (2) yields the dipeptide amide Tyr-ArgNH2. The competing hydrolysis (1)yields the non-activated tyrosine. The dipeptide amide Tyr-Arg-NHz is deamidated selectively by peptide amidase (PA)to the desired

198

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

100

80

> c ..-> 0 a -

60

.I-

a

cn

40

20 0

0

20

10

[Tyr-OEt] (mmol L-’)

30

40

40

30

20

2

Y

c

0

100

200

300

0

400

500

[Arg-NH2] (mmol L-’) Figure 7-9a, b. Kinetics o f t h e synthesis ofTyr-Arg-NHZ by CPD-Y: activity and selectivity as a function of the concentrations ofTyr-OEt (A) and Arg-NHz (B)154’.

dipeptide, which may be further hydrolyzed by CPD-Y to the amino acids Tyr and Arg (secondary hydrolysis). Secondary hydrolysis of Tyr-Arg-NHzto Tyr and Arg by CPD-Y can be neglected.

1 .o

--0.8

--

Yeld

[TyrOEt], = 40 mmol L-’

Selectivity

[ArgNH,], = 40 mmol L“

~-

h

v I

.-% .-

0.6

. I -

0 a,

Q)

v)

5 a, F

0.4-

0.2-

0.0---

c

c

c

c

e

c

c

c

c

c

c

c

c

c c

-

c

*

Figure 7-10. Synthesis ofTyr-Arg by CPD-Y and P A selectivity and yield as a function o f cotwersion for continuous operation in an enzyme membrane

The selectivity s of the formation of the dipeptide Tyr-Arg is reduced by the undesired parallel (1) and consecutive reaction (4). Measuring the kinetics of the aminolysi,~and the hydrolysis reactions gives an opportunity for process optimization (Fig. 7-9). As expected, increasing concentration of Tyr-0-Etinfluences the activity of CPD-Y bui not the selectivity of the competing reactions 1 and 2. On the other hand, increasing the concentraiion of Arg-NH2results in increased activity and selectivity favoring the aminolysis reaction compared to the competing hydrolysis reaction. The data in Fig. 7-9 were measured under initial rate conditions (see Sect. 7.4.1). Additionally, selectivity and yield were calculated as a function of conversion (Fig. 710). If CPD-Y and PA are used in one reactor the yield of dipeptide initially increases wiih conversion whereas the selectivity drops because of the consumption of the nucleophile (compare to Fig. 7-9B). At higher conversion, the yield reaches a maximum and selectivity drops steeply because of the subsequent hydrolysis reaction i(4)(Eq. (11)).At 100% conversion, yield equals selectivity, both being zero. For this reaction, a steady-state conversion of about 0.8 defines the best reactor performamce. The above complex reaction is an instructive example of the correlation of selectivity, conversion and yield. Example: Synthesis ofchiral cyanohydrins The “enantiomerically pure compound (EPC) synthesis” has become a major strategic concern in the synthesis of bioactive compounds especially for pharmaceutical and agrochemical use. EPC synthesis can be achieved very efficiently using

200

I

7 Reaction Engineeringfor Enzyme-CatalyzedBiotransformations

enzymes as chiral catalysts. Although enzymes mostly show a high degree of enantio- or diastereoselectivity,non-stereospecificside-reactionsmay occur lowering the optical purity of the product. In this case, the problem of the selectivity of the reactions has to be addressed. A suitable catalyst for the synthesis of (R)-cyanohydrins is the enzyme (R)oxynitrilase from bitter almonds. It catalyzes exclusively si-face addition of hydrogen cyanide to benzaldehyde or other aldehydes. A competing non-enzymatic parallel reaction lowers the enantiomeric excess of the product[55,561.

pD<; (R)-Oxynitrilase

(R)-Mandelonitrile

Re-face attack

(12) "O*,, No enzyme

*

O

c

'

\

NS

)CN H

+

\

Hydrogen cyanide

D

c

.

\

O H , H

/ Racernic product

Benzaldehyde

This example is very instructive in that it demonstrates methods to suppress a nondesired reaction. The investigation of the reaction system leads to the question of the influence of the reaction conditions on selectivity. Influence ofpH value upon selectivity The pH value has an influence not only on the activity of enzymes but also on chemical reactions. The chemical cyanohydrin reaction is base-catalysed, as, compared to HCN, the cyanide ion more easily attacks the carbonyl group. As a result, a distinct decrease of the reaction rate for the non-enzymatic synthesis of mandelonitrile occurs at lower pH values. Also, the enzyme activity decreases but not to the same extent; therefore, the enantioselective enzymatic reaction becomes dominant at lower pH (see Fig. 7-11). Influence of temperature upon selectivity The investigation of the temperature dependence of the competing parallel reactions gives information about their activation energy. According to the Arrhenius equation (Eel. (13112 k=k,,;e

_E.4_ R1

orIn(klk,,)=--

E.4

R. T

(13)

7.3 Investigation of the Reaction System

25000

20000

15000 10000 5000

30

20 10

0

3.0

3.5

4.0

4.5

5.0

PH Figure 7-11.

Initial reaction rate ofthe synthesis of rnandelonitrile as function of pH[".

561.

where

k k,, EA R

T

(s-l) (s-l) (J mol-l) (8#.314J K-lmol-') (K)

rate constant of the reaction frequency factor activation energy gas constant absolute temperature,

the activation energy EA can be determined from the following plot: In this case the non-enzymatic reaction, having an activation energy of 7 3 kJ mol-', dlecreases faster with temperature than the enzymatic reaction (activation energy 46 kJ mol-'). In general, at a lower temperature the reaction with the lower activation energy is favored. The enzymatic reaction therefore becomes more dominant at lower temperature.

Influence of substrate concentration With respect to benzaldehyde, (R)-oxynitrilaseexhibits saturation kinetics (Michaelis Menten kinetics, see Sect. 7.4.2.1) and a maximum reaction rate is reached above a concentration of about 5 mmol L-'. The chemical reaction presents a linear increase of the reaction rate with increasing benzaldehyde concentration, representing first order kinetics, when the concentration of HCN is kept constant (see Fig. 7-13). As a consequence the enzymatic reaction becomes more dominating at lower concentrations of the substrate benzaldehyde (for HCN as substrate the same kinetic behavior occurs, data not shown). Accordingly an enzyme reactor would be suitable that works under minimum average substrate concentrations. These requirements are satisfied by the continuous stirred tank reactor (CSTR). In Sect. 7.5.2.1 this aspect of enzyme reaction engineering wi I1 be discussed further.

I

201

202

I

7 Reaction Engineeringfor Enzyme-CatalyzedBiotransformations

25°C 100 80

\

60

\

40

10°C

I

20

0 Non-enzymatic reaction Enzymatic reaction 10 0.1 335

0.00340

0.00345

0.00350

0.00355

T-' (K-') Figure 7-12. Relative reaction rate o f t h e synthesis of mandelonitrile as a function o f t h e reciprocal temperaturelS71.

4000 2000

]/

f

1

0

I

Enzymatic reaction

0

[Benzaldehyde] (mmol L-') Figure 7-13. Initial reaction rate o f synthesis o f mandelonitrile as function o f benzaldehyde concentration '551.

7.3 Investigation of the Reaction System

Influence oiorganic solvents There are two arguments for the use of organic solvents in the synthesis of alphahydroxynitriles by oxynitrilase: - the best substrates of the enzyme are aromatic and straight chain aliphatic

-

aldehydes, both of which have a limited water solubility (e. g. benzaldehyde 80 mmol L-'). The solubility may be enhanced by organic solvents; the non-specific reaction is favored in water and may be suppressed by lowering the water activity of the medium.

Diflerent methods have been used, employing organic solvents in this reaction (reviewedin [58, 591): - the use of water-miscible organic cosolvents such as ethanol or methanol, - the application of aqueous two-phase systems, - iimmobilizationof the enzyme and the use of organic solvents (e.g. ethyl acetate or

others) which contain only traces of water to preserve the enzyme's activity, - the application of biphasic lyotropic liquid crystal systems.

In the oxynitrilase reaction, high optical purities can be reached by the use of organic solvents. Often a major drawback of these methods is a lowered enzyme activity, resulting in long reaction times and a reduced enzyme stability in continuous experiments.

Selection ofreaction conditions Summarizing the above effects, the high selectivity toward the enzyme-catalyzed reaction and suppression of the non-specificparallel reaction is achieved by -

lowpH, low tewcperature, low stationary substrate concentrations, additionally a high enzyme activity exclusively favors the enzyme-catalyzedreaction without influencing the non-enzymatic reaction.

Knowing that the enzyme stability is sufficiently high at pH 3.75 (50% deactivation after 150 h), water was chosen as the reaction medium and the reaction was peirformed at room temperature. This selection ofreaction conditions was followed by a detailed kinetic analysis of the system, the investigation of the reactor kinetics and the simulation of steady state conditions in continuous experiments [55. 'fie discussion about the choice of an appropriate reactor and optimized operation coiiditions will be continued in Sect. 7.5.2.1. Anticipating the results, the reaction can be performed effectively under the above reaction conditions in an enzyme membrane reactor yielding a product with an enantiomeric purity of higher than 99 %.

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7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

7.3.2.3 Other Properties of the Reaction System

Other properties that have to be considered in enzyme reaction engineering are: pH effects on the reaction A pH effect during the reaction occurs if proton shifts are involved, as is the case in most enzymatic reactions. As enzyme activity is strongly dependent on pH, a pH shift induced by the reaction has to be prevented by buffering or titrating the reaction medium. Problems arise especially in immobilized systems where pH gradients occur within the enzyme matrix if the buffering capacity of the medium is too low[61,621. In cofactor dependent reactions the influence of pH on the stability of NAD(P)+and NAD(P)Hhas to be c ~ n s i d e r e d [ ~ ~ l . The effect of temperature on the reaction The effects of the variation of temperature on enzymatic reactions usually do not require precautions, as in typical small scale reactors the capacity of heat exchangers is sufficient to maintain a constant temperature. Solubilities of substrates and products This information may be useful in cases where a product can be removed continuously from the reaction mixture by crystallization. 7.3.2.4 Application of Organic Solvents

The use of enzymes in non-aqueous solvents can be an advantageous alternative to reactions in water, especially for poorly water-solublesubstrates and products, e. g. in the synthesis of esters, lactones or selected peptides. However, the knowledge of how solvents influence enzyme activity and selectivity is still not profound. As it is impossible to cover all relevant aspects within this chapter, the reader is referred to some selected articles for further reading[*’*66751 . H ere some of the basics will be discussed briefly; in Sect. 7.5.4 reactor concepts for the use of enzymes in organic solvents will be presented. There are advantages in performing enzymatic transformations in non-aqueous media: - non-specific side-reactions such as the chemical addition of HCN to aldehydes

-

during the oxynitrilase reaction may be suppressed by performing the reaction in an organic solvent. This example has been presented in the previous section. the solubility of poorly water-soluble substrates or products may be increased in organic solvents or at least by addition of water-miscible solvents. If non-watermiscible solvents are used in combination with water, inhibitory effects such as substrate or product inhibition may be overcome if the organic phase contains most of the substrates and products and the enzyme remains in the water phase.

7.3 lnvestigation ofthe Reaction System

I

205

A very interesting alternative has been published detailing the use of cyclodextrins to enhance the solubility of 2-acetylnaphthalenein aqueous solution. The complex between the cyclodextrin derivative and the ketone is transformed fast enough to support high reaction rates in an enzymatic reduction.1'7r - equilibrium reactions may be shifted toward the desired product for one or both of the following reasons: (i) lowering of water activity by adding a water-miscible (ii)continuous extracorganic solvent (example: enzymatic peptide synthesis tion of the desired product into a non-water-miscibleorganic phase (compare to the multiple-compartment enzyme membrane reactor (see later, Fig. 7-

37,[77.781).

Both stabilizing and destabilizing effects of solvents on enzymes have been reported. A reasonably reliable measure of the compatibility of solvents with enzymes is the log P value, where P is defined as the distribution coefficient of a solvent between water and 1-octanolin a two-phase system[64*791. Solvents with a log P value above 4 are suitable (e.g. aromatics, aliphatics)whereas water-misciblesolvents with a log P value below 2 (short chain esters, DMF, short-chain alcohols) are not suitable for employment with biocatalysts. The latter solvents interfere with the water at the boundary of the protein itself and so disrupt the binding forces necessary to maintain an active form of the enzyme. Surprisingly, tert-butanol has a stabilizing effect on some oxidative enzymes , I ' ' [ despite its low log P value (0.35). Fig. 7-14 shows a classification for biotransformations in organic solvents into one-phase and two-phase systems. One-phase systems may consist mainly of water, water plus a water-misciblesolvent or a pure organic solvent. Most water-miscible solvents may be used in concentrations up to 20% before enzyme deactivation exceeds the benefits obtained by better solubility of substrates and/or increased selectivity. Using this approach, enzyme activity and enzyme stability have to be examined carefully to select appropriate reaction conditions. Using mainly water non-miscible solvents several approaches are possible. In most cases, the organic solvent has to be saturated with water in order not to remove the boundary water surrounding the enzyme, which otherwise results in deactivation. In such microaqueous systems the pH of this tiny amount of water should be carefully chosen for optimal enzyme activity. The control of water activity can be achieved by addition of salts or utilization of saturated salt solutions fS1, 82]. The simplest way of using an enzyme in organic solvents is to suspend the insoluble enzyme in the required solvent. This technique was first reported in 1900 [831 and has been extended over the last few years to encompass many systems (mainly proteases and lipases) L7', 84, 8'l. Organic solvents may be replaced by supercritical liquid carbon dioxide, which exhibits similar properties to hexane .'[ 14'1. To achieve true homogeneous catalysis, enzyme solubility may be increased by coupling polyethyleneglycol to its surface LS71. The coupling may alter the stability, activity and selectivity of the enzyme. By use of detergents and small amounts of water or buffer, reversed micelles can be formed containing the enzyme in the water phase while the organic solvent forms the bulk phase[", "1.

206

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

Figure 7-14.

Classification of enzyme catalysis in the presence of organic solvents.

Just like one-phase systems, two-phase systems may consist mainly of water or mainly of organic solvent. In an aqueous-based system an insoluble substrate is dispersed, using a non-miscible solvent (oil-in-wateremulsion). Hydrolysis reactions of poorly water soluble substrates (e.g. fat hydrolysis by esterases) may be performed in this way. If the aqueous phase is dispersed in a water-immiscibleorganic solvent a water-in-oil emulsion is obtained. This type of system may be advantageous if a condensation reaction has to be performed where the water content has to be kept low. A special form of a two-phase system involves lyotropicliquid crystals, which are obtained by mixing a detergent, water and an organic solvent["^ 911. Most of the water and the detergent form the liquid crystal wherein the enzyme is immobilized, whereas most of the organic solvent forms the second phase. Enzymes immobilized on an insoluble support also belong to two-phase systems. They have been mentioned earlier and are not discussed further at this point. Very recently ionic liquids emerged as a new class of solvents. They are salts with melting points below 100 "C and are non-volatile. Some typical structures are shown in Fig. 7-15. They possess good solvating properties for many substrates and catalysts[92294]. They can also be used to replace organic solvents in enzymatic For the lipase-catalyzed kinetic resolution of phenylethanol an reactions [95-981. improved enantioselectivity at higher temperatures was observed compared to the selectivity obtained when using methyl-tert-butylether(MTBE) as solvent [", 981. What has been neglected so far is a consideration of the location of the enzyme

7.3 lnvestigation ofthe Reaction System

I

207

Figure 7-15. Increased enantioselectivity foir lipases by reaction

h

N

0

G

.

20

pFg

\NQN,

40 60 80 Temperature ("C)

CFW;

100

conditions: 1 mol/L alcohol, 2.5 mol/L vinylacetate, water content 0.3% 12 g/L Chirazyme L6 (fseudomonas sp. lipase)

and hence the enzymatic reaction. Depending on the hydrophobic and hydrophilic properties of the enzyme it may be enriched in one of the phases (normally the aqueous one). For lipases, catalysis often occurs at interfaces. Therefore, the surface area per unit reactor volume available in such a system is an important quantity, not only for reaction performance but also for determination of kinetic data. As shown in Fig. 7-16, a larger surface area requires a higher enzyme concentration to be saturated. But a tenfold increase in the surface area - even if saturated with One reason for this enzyme - does not result in a tenfold increase of might be product inhibition. Wherleas kinetic studies are quite easy to perform in homogeneous solution the extent of the interface has to be taken into consideration for biphasic systems. The most reliable way of measuring the interfacial area is by use of Fraunhofer

208

I

7 Reaction Engineering for Enzyme-Catalyzed Biotransformations

0.0

0.4

0.8

1.2

1.6

2.0

2.4

2.8

3.2

Enzyme Concentration (mg Lipase/g Oil) Figure 7-16.

Saturation ofthe phase boundary surface with lipase.

diffraction[lo0I.All reaction velocities can be given based on the same surface area. A change of droplet size and surface area, which may occur with change of substrate or product concentration during the reaction, can be distinguished from true inhibition effects.

7.4

Investigation of Enzyme Kinetics

In this chapter, some principles of the kinetics of enzymatic reactions are discussed. A more detailed description of enzyme kinetics is covered in a number of textbooks

and articlesL45, 101-1041. F'irst of all, a few general definitions will be given. The reaction rate u of any chemical reaction (Eq. (14))

+

nAA n,B

+ n,C + n,D

is defined as follows (Eq. (15)): =nA

u

[A], [B] nA, nB

k

d[A1- function of k , [A1 [B], .. dt

(mmol L-' min-') (mmol L-') (-)

(mmol'-" L"'

min-')

reaction rate concentrations of A, B ... stoichiometric coefficients rate constant (nbeing the order of reaction)

7.4 Investigation ofEnzyrne Kinetics

Eq. (15) is the rate equation of the reaction (also called the "kinetic model'). The formulahon of such a differential equation for all reacting substances is the basic step in describing the kinetics of chemical/biochemical reactions. These rate equations include concentration values of the relevant reaction partners and kinetic parametcrs such as the rate constant k. An investigation of enzyme kinetics includes the measurement of reaction rates, the choosing of an appropriate kinetic model and the identification of the kinetic parameters. 7.4.1 Methods of Parameter identification

Kinetic rneasurements [loSl have to be carried out to examine the dependence of the reaction rate on the concentrations of all relevant components. As described in a previous chapter, for measuring enzyme kinetics initial reaction rates v, = j S ] are determined at optimal reaction conditions, which may be chosen according to the procedure outlined in Sect. 7.3. The initial reaction rates are measured by varying the concentration of only one component and keeping all other concentrations (e.g. of cosubstrates and inhibitors) at a constant level (for an example, see Figs. 7-19 and 7-20). The rate of conversion has to be smaller than 5-10%, essentially to keep all initial concentration values constant. The parameters of the kinetic model can be identified by fitting the kinetic data using methods of non-linear regression such as those described by Rosenbrock or Nelder Mead['OG 1'" (Fig. 7-17A). Methods of linear regression that are often used need a rearrangement of rate equations into a linear form (e.g. a double reciprocal plot according to Lineweaver-Burk ['08]). This gives different weight to the data points measurcd at different concentration levels 'lo]. For the correct calculation of the regression line this point must be considered, otherwise the Lineweaver-Burk double reciprocal plot is not acceptable["']. Initial rates are not significant in large-scale processes where high conversion of the substrate is desired. With rising conversion, the simultaneous effects of both substrate S and product P on the reaction rate have to be described. In the case of equilibrium reactions, the forward reaction and the back reaction have to be described by one rate equation: they can only be treated separately under initial rate conditions. The overall rate equation has to describe the reaction rate as aficnction of all relevant components at all relevant concentration levels. A correct fit of all initial reaction rate data gives no guarantee that the kinetic model will fit the overall reaction data! A proper fit of the time-courses of some batch reactor experiments at different starting concentrations represents an appropriate test of the rate equation. This implies that numerical integration of the rate equation (e.g. by the Runge Kutta yielding a simulated time-course, has to fit the data of the measured time-coiirse over the whole range of conversion (compare to Fig. 7-17 B). Examples of these methods will be given after the presentation of the basic kinetic models. A combination of the Runge Kutta method and methods of non-linear regression allows a parameter identification from the time-course data. This technique starts with a given set of parameters, performs the numeric integration of the rate equation [lo93

210

I

7 Reaction Engineeringfor Enzyme-CatalyzedBiotransformations

Figure 7-17. Methods of parameter identification: A by fitting initial rate kinetic data, and B by fitting the time-course of a reaction.

and compares the simulated with the measured time-course. Then the parameters are changed and the same steps are repeated until the simulation fits the measured data (Fig. 7-17 B). This method requires specially designed computer software,which is commercially available (ScientistTM,MathlabTM,MapleTM).The method is especially useful if components having an effect on the enzyme kinetics are not available or if they cannot be measured, but are in a defined stoichiometric relation to a measurable component. 7.4.2

The Kinetics of One-Enzyme Systems 7.4.2.1

Michaelis-Menten kinetics

In this section the basic kinetic model for enzyme-catalyzed bioconversions is presented. Understanding this model is the foundation for deriving more complex models. In their theory of enzyme catalysis, Michaelis and Menter~[''~] postulated the existence of an enzyme substrate complex (ES), which is built up in a reversible

7.4 investigation ofEnzyrne Kinetics

reaction of the substrate S and the enzyme E. The dissociation of this E S complex to E and P is assumed to be the rate-determining step (Eq. (16)). E+S

Rate constant kl (L mmol-’min-’) k.1 (min-’) kz (miin-’)

-

kl

k-1

-

ES

k2

E+P

Reaction Association of the ES complex Dissociation of the E S complex into E and S “Turnover number” (see below) (also refered as “kcat”)

If the rate constant k2 is much smaller than the rate constant k.1 of the enzyme, the substrate and the enzyme-substrate complex are in equilibrium, which is not disturbed by the decomposition of E S into E and P (“rapid equilibrium-assumption”). Based on this assumption, Michaelis and Menten derived the following rate equation (Eq. (17)). .[SI Ks + bl

- urnax

(nimol L-’min-’)

2)

urnax (nimol L-lmin-’) Ks

(mmol L-l)

(17)

Reaction rate Maximum reaction rate Dissociation constant of the E S complex

This is the classic form ofthe Michaelis-Mentenequation. Eq. (17)can be rearranged to

& KS

(min-’) represents the rate constant of the first order enzyme kinetics (seeFig.7-18)

bl

1+ KS

B1

__ - _[Ed Ks [El

(-)

denominator of the kinetic model :represents a dimensionless “adsorption term” (see below)

(.)

represents the dissociation of ES into E and S and may be simply derived from the dissociation equilibrium

The curve u = j S ] belonging to the Michaelis-Mentenfunction is shown in Fig. 718. As long as [ S ] is well below Ks‘ Michaelis-Menten kinetics transpose to (linear) first-order kinetics. In this case nearly all enzyme molecules are present in the free form and saturation of the enzyme with substrate is rate limiting. This degree of saturation is represented by the “adsorption tern”, which gives the sum of the values 1 + [ES]/[E] + [ESi]/[E] (if more substrates Si or inhibitors or products are involved). The adsovption term represents the dissociation equilibria of all relevant enzyme-substrate

I*”

212

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

Figure 7-18.

Michaelis-Menten kinetics: curve analysis

complexes, and reveals the “weight”of the equilibria involved. For example, for [S] = Ks the value of the adsorption term equals 2, so that 50% ofthe enzyme is in the ES form and the other 50% is in the free form. The adsorption term initially grows from a value of 1 (at [S] << Ks) slowly to become proportional to [S] and transforms the linear first-order rate law to the curve reflecting Michaelis-Menten kinetics. At concentration values above Ks it is not enzyme saturation but the catalytic turnover of the enzyme that becomes more and more limiting. Being proportional to the concentration of the ES complex, the reaction rate cannot rise above the u, value when all enzyme molecules are in the ES form. The enzyme kinetics turn to zero order. The Michaelis-Menten kinetics, represented by Eq. (18),may be extended to more complicated reactions by looking at the structure of the adsorption term. This procedure shown below is valid as long as the “rapid equilibrium” assumption is made. This is not valid in all cases. Briggs and Haldane[’I4]did not use the above assumption, but pointed out that within a very short time after starting the reaction, ES would build up to a nearly “steady state” level where the following assumption is valid: d[ES]/dt = 0. From this “steady state assumption” they derived the same rate equation:

K~

(mmol L-’)

Michaelis constant

7.4 Investigation ofhzyrne Kinetics

Here the KM value no longer is a dissociation constant but a kinetic constant which includes lil, Ll, k2 and for more complex reactions even more rate constants. This equation is the standard form of the Michaelis-Mentenequation. Although the derivation of reaction rates using the “steady state assumption” is more exact, often the rapid equilibrium assumption is used because it allows a simple derivation of the rate equation from the relevant enzyme-substrate complexes (see below) and allows fitting of the kinetic data. The following explanations are based on the rapid equilibrium assumption, and therefore allfollowing constants K are used as dissociation constants with the component dissociatingfr.om the enzyme as the subscript, e.g. KA, KB, and the component remaining at the enzyme as second subscript (e.g. KIs, .see below). Within Eq. (17)urnaxcan be expressed as shown in Eq. (20):

vmax (mmol L-’min-’) [Elc,’

(mmol L-’)

kz

(niin-’)

Maximum reaction rate Total enzyme conzentration in molar value: [El,’ = [El + [ES] Turnover number

The "turnover number‘‘ represents the moles of product formed per minute and per mole of active sites of enzyme. Usually the molar concentration of the active sites is not known, so the weight per volume of the enzyme preparation is used; Eq. (20) is changed to Eq. (21).

[El0 A,,,

Total enzyme concentration (in g L-‘) (mmol g-’ min-’) Maximum specific activity of the enzyme (g L-7

Using Eq. (21), Eq.(17)can be converted to Eq. (22).

and, with. Eq. (23),

A

(rnmol g-‘ min’)

Specific activity of the enzyme

to Eq. (24). ‘4-

ha.bl K , + [SI

Besides .;pecific activity, two other quantities are used for the characterization of enzymes: the International Unit [U](defined as the amount of enzyme which

I

213

214

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

produces 1 pmol of product per minute) and the katal (kat) (defined as the amount of enzyme which produces 1mol of product per second). The following correlations exist between the reaction rate u, the specific activity A and the definitions of unit U and katal (kat)(Eq.(25)): U 1-=1mL U l-=lmg

pmol mmol = 1(reaction rate = volumetric activity) mL.min L.min pmol mmol = 1-__ (specific activity) mg.min g.mm

1kat = 6.10’ U

(activity)

The use of values for the specific activity of an enzyme is only significant if the conditions of measurement are specified, especially temperature, substrate concentration and pH value. Additionally, enzyme activities are always measured at initial reaction rate conditions (see above). In the following sections the extension of Eq. (18) to more complex reaction schemes is described. Again the “rapid equilibrium assumption” is used to show how more complex rate equations are derived from simple Michaelis-Menten kinetics. Attention is focused on some typical rate equations that are useful to describe enzyme kinetics with respect to a desired process optimization. The whole complexity of enzyme kinetics is of importance for a basic understanding of the enzyme mechanism, but it is not necessary for the fitting of kinetic data and the calculation of reactor performance. 7.4.2.2

Competitive Inhibition

Substances that cannot be converted by the enzyme but are competing with the substrate for the active site of the enzyme are called “competitive inhibitors”. The following reaction scheme represents this situation: KS E+S=====

ES

‘ ’ w

E+P

t

I

llhl

El

I KI

(mmol L-’)

Inhibitor Dissociation constant of the EI-complex

The corresponding kinetic model again may be presented in two forms (Eqs. (27 and 28)). u

max

~,

u =+ 1+-+-

Ks

bl K,

7.4 fnvestigation ofEnzyrne Kinetics I215

Compared to Eq. (18)the rate equation (Eq. (27))for competitive inhibitors includes an additional term [I]/ K1 in the denominator representing the additional dissociation equilibrium of the EI complex ($ the above discussion about the “adsoqtion term”). Also, each alternative substrate S, ofa reaction would render such a term [Sn]/Ksnin the denominator. As a consequence of their affinity to the enzyme, alternative substrates and inhibitors block a part of the enzyme otherwise available for the reaction S P. Thus, the product P of enzymatic reactions is often a competitive inhibitor of the enzyme leading to “product inhibition” (compare Eqs. (27)and (37)).The influence of product in the case of reversible reactions will be discussed later. From Eq. (28) it can be concluded that the effect of a competitive inhibitor is to increase the apparent Ks value while the urnaxvalue is not affected. For example if [I] is chosen as KI, in the presence of the inhibitor it would take twice as much substrate S to reach umax/2as without inhibitor. -+

7.4.2.3

Non-Competitive Inhibition

“Non-competitiveinhibitors” bind at the enzyme E or at the ES complex and build up a ternary complex ESI, which cannot be converted to the product P (Eq. (29)): E+S=

KI KIS Ks I

(mmol L-’) (mmol L-’) (mmol L-’)

KS

ES

+

+

I

I

EI + S

ESI

A

E+P

Dissociation constant of the EI-complexwith release of I Dissociation constant of the ESI-complex with release of I Dissociation constant of the ESI-complexwith release of S

For ordinary non-competitive inhibition, the Kl and KIs values are identical and also the KS and Ks’ values. In this case rate equation (30) is valid:

216

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

u=

Compared to Eq. (27), the rate equation for non-competitive inhibitor includes another term for the equilibrium of decomposition of the ESI-complex into E, S and I. From Eq. (31) it can be concluded that a non-competitiveinhibitor has no effect on the KS value but lowers the vmaX value. This is because the inhibitor binds all enzyme species with the same affinity. 7.4.2.4

Uncompetitive Inhibition

Substances which only bind the ES complex and not the free enzyme E are “uncompetitive inhibitors”: t+s-

4

”*

ES

I

E+P

(32)

11; u=

mu

. [SI s.1

Ks+[S]+[llj

(34)

K,S

Compared to Eq. (30) the rate equation for uncompetitive inhibitors includes the same term for the equilibrium of decomposition of the ESI complex into E, S and I, but no term for an equilibrium of an EI complex.

7.4 Investigation of Enzyme Kinetics

I

217

One special form of uncompetitive inhibition is substrate inhibition. Here a second substrate molecule binds at the ES complex resulting in an inactive ESS complex. This form of inhibition is often found and will be discussed below (see acylase kinetics, Fig. 7-20A). 7.4.2.5

Reversibility of One-Substrate Reactions

Because of the “principle of microscopic reuersibility” each molecular process (in contrast to a macroscopic process) may occur in both forward and backward directions. As a consequence the end product P of an enzymatic conversion can act as a competitive inhibitor of the enzyme or, depending on the thermodynamic equilibrium, be transformed to the substrate S. If the interconversion of the ES to the EP complex is the rate-determining step the rapid equilibrium assumption is valid and the rate equation can be derived easily. KS

Ek-S-

ES

-

kF

-

KP

EP-

-

E+P

kB

’P

U

F ,

~

(mmol L-’ min-l) (mmol L-’ min-’) ~ (mmol L-’ min-’)

Reaction rate for the formation of the product P Maximum reaction rate of the forward reaction Maximum reaction rate of the backward reaction

Again the denominator represents the dissociation equilibria of the ES complex and the EP complexes. Both partial reactions, the forward and the backward reactions, are catalyzed simultaneously; both “substrates” S and P are competing for the same enzyme. If the value in Eq. (37) equals zero, the equation reflects competitive inhibition of the enzyme by the product P (see Eq.(27)). Eq. (371is a result of the rate equation of the forward reaction reduced by the rate equation of the backward reaction, both having the same denominator. The numerator represents the first-order kinetics of the forward and the backward reactions. If the equilibrium of the reaction is reached, the numerator becomes zero. From the equilibrium condition (Eq. (38)),

a correlation between the thermodynamic parameter “equilibrium constant” and the kinetic parameters can be derived (Eq. (39)):

218

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

This relation, representing the ratio of the rate constants of the first order kinetics is known as the “Haldane equation”. Haldane equations can be formulated for every kinetic model describing an equilibrium reaction, just by setting the numerator to zero. An example may illustrate this principle: Eq. (37) is the basic kinetic model for isomerizations and racemizations. For example, the kinetic constants of an alanine racemase of Bacillus stearothermophilus are found to be K D . A =~ ~3.42 mmol L-’ z ) D . A ~ ~ - L . A= ~ ~ 7.02 , ~ ~ U mL-’ = 11.82 U mL-’ K L . A = ~ ~5.76 mmol L-’ The corresponding rate constants of the first order kinetics are z ) D . A ~ ~ + L . A/ ~KD.A~= ~ , ~ ~ = 2.052 min-l z ) L . A ~ ~ - D . A /~ K ~ ,L~ . ~A= ~ ~2.052 rnin-l These rate constants are identical, resulting in Gq = 1, a predictable result of racemization of D- or L-alanine. This example shows that the rate constants of the first order kinetics determine the position of equilibrium of the reaction. In the case of reversible two-substrate reactions, the rate constants of the second order kinetics have to be used as shown below. 7.4.2.6 Two-Substrate Reactions

In most enzyme-catalyzed reactions two or more substrates are involved, such as in the enzyme-catalyzed aldol reaction, the cyanohydrin reaction or enzyme-catalyzed peptide synthesis (examples used before). For many reaction schemes kinetic models have been derived using the steady-state assumption. Some important reaction mechanisms and the corresponding rate equations are summarized in Table 7-1. An approach to the steady-state method and a detailed discussion of the resulting kinetic models is difficult and is not the aim of this chapter. For a practicable approach, the “rapid equilibrium” assumption is applied and the structure of kinetic models of two substrate reactions is demonstrated for the case of a “randombi-uni reaction”. The binding of two substrates A and B to an enzyme may occur in a compulsory order or in a random order. If one product (uni) is formed out of two substrates (bi), the corresponding mechanisms are the “ordered bi-uni“ mechanism and the “random bi-uni mechanism”,respectively (water is not regarded as a substrate). For a “random bi-uni” mechanism of an equilibrium reaction A + B +=P the following reaction scheme is valid (Eq. (40)): E+A + B

KA

-

EA

+ B

7.4 Investigation ofEnzyme Kinetics

If the interconversion of the central EAB complex to the EP complex is rate determining and all other reactions are in a “rapid equilibrium”,the following rate equation for the random bi-uni reaction may be derived according to the above method:

2),

___. 2, F,max [A].[B]-’”.””. [p] K, . K,” KP = A B P A.B I+-+-+-+-

K,

K,

K, KA.K;

Again the denominator represents all enzyme-substrate equilibria, e. g. Eq. (42):

The numerator represents the second-order kinetics of the forward reaction reduced by the firs-t-orderkinetics of the back reaction. A Haldane equation can be formulated by setting the numerator to zero, resulting in a relation for the equilibrium constant (Eq. (43)):

In the absence of the product P, Eq. (41)may be rearranged by multiplication of numerator and denominator by KAKBA (respectively K B K A ~ which , is identical to &KBA, compare to Eq. (40)). Then Eq. (44)results:

u p=

2, F.max

‘

[B1

K,.KBA+KBA.[A]+KAB.[B]+[A].[B]

Only if a further identity KA = K.A.~is assumed can Eq. (44) be rearranged to Eq. (45):

This rate equation is called “Michaelis-Mentendouble substrate kinetics”. It is a formal multiplication of two Michaelis-Menten models for both substrates A and B. This model can be used to describe rate kinetics of two substrate reactions in the absence of the product(s). The kinetic measurements have to be performed by varying the concentration of one substrate keeping the concentration of the second substrate at a constant value well above the KM value. The model cannot be used if back reactions occur and an equilibrium has to be described by an appropriate Haldane tquation.

I

219

220

I

7 Reaction Engineeringfor Enzyme-CatalyzedBiotransformations

U

G

a-

% II

7.4 investigation of Enzyme Kinetics

r

c.

+

w Q\

I**'

222

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations Footnote to Table 7-1. The above reaction schemes are described according to the notion of Cleland11161.The substrates binding to the enzyme are indicated as A and B according to the order of binding; the products are indicated as P and Q, respectively. The different mechanisms may be explained in short as follows (for a detailed discussion the reader is referred to textbooks, listed above). Random bi bi: Both substrates A and B may bind: also the products P and Q may dissociate from the enzyme in a random order. (Example: E. coli galactokinase) Random bi uni: Two substrates A and B and only one product P are involved (water is not considered). (Example: aminoacylase, see above) Ordered bi bi: The substrates and products bind to or dissociate from the enzyme in a specific order. (Example: in most dehydrogenase-catalyzedreactions the coenzyme has to bind first; hexokinase) Theorell-Chance: This special case of the ordered bi-bi mechanism occurs if the first product P dissociates from the enzyme very rapidly and an EAB-, EPQ-complex does not occur in a significant concentration. (Example: alcohol-dehydrogenase) Ordered bi uni: Two substrates and only one product are involved. (Example: NeuSAc-aldolase and other aldolases) Ping Pong bi bi: In this mechanism a product P is released between the addition of two substrates. This mechanism is often found in group transfer reactions. (Example: transaminases, yeast transaldolases) Above-mentionedand additional examples are given in [117,1181. mechanisms the following points can be identified:

Looking at the rate equations for the different

Corresponding to the above discussion about enzyme kinetics, the numerator is nearly identical for all different bi-bi-mechanisms (for bi-uni mechanisms, respectively),as the numerator characterizes the thermodynamic equilibrium of the reaction (which is independent of a kinetic mechanism). The denominator consists of terms characterizing all enzyme substrate equilibria. Depending on the mechanism every substrate A, B, P and Q requires one or two kinetic constants (designated as KTand KM)in order to describe its reciprocal action with the enzyme. According to the steady -state derivation of these rate equations, KI and KM are no longer simple dissociation constants (compare discussion about KS and KM). In some cases KI is identical to a dissociation constant as described before, but most often these steady state parameters are defined by three and more rate constants. A verbal distinction between an inhibition constant; a Michaelis constant and a dissociation constant does not have a corresponding mechanistic scenario in all cases. The random mechanism in both cases is the simplest mechanism requiring only 8 (random bi-bi) and 6 (random bi-uni) kinetic constants. This mechanism should be tried first to fit data of a two-substrate reaction. The denominator may be supplemented by additional terms e.g. for inhibitory substances, not being described by basic mechanism (compare to discussion of the acylase kinetics). A distinction between the different mechanisms is best done using initial rate kinetic measurements as described in detail in the literature. For reaction engineering purposes only a proper fit of reactor data is desired, using a minimum amount of kinetic parameters for statistical reasons.

If the concentrations of the products P and Q in the models summarized in Table

7-1 are set to zero, the resulting rate equations will be identical to Eq. (44) in all cases with the exception of the “ping-pong”mechanism. This means that the MichaelisMenten double substrate kinetics is valid under the above circumstances. An example of the use of this model will be shown for a two enzyme system (see Sect. 7.4.3).

7.4 fnvestigation ofEnzyme Kinetics 7.4.2.7

Kinetics of Aminoacylase as Example of a Random Uni-Bi Mechanism

Aminoacylase kinetics may be used as an example for demonstrating the measurement and modeling of enzyme kinetics. This reaction is of industrial importance in 12

I

I

I

10 - - _ _ _ ~

8

6

4

2 0

0

Measured Calculated I

0

50

100

150

200

Calculated 0

~

I

I

223

224

Ic

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

12

10

-

h

z

8

3 6 .2 > .c

2

4

2 0

I

I

I

0

200

400

I

600

I

800

1000

[D/L-Alanine] (mmol L-’) Kinetics of aminoacylase: effects on the activity of hydrolysis o f acetyl-~-alanine~”~] (initial rate measurements).

Figure 7-19.

the production of L-amino acids by hydrolysis of acetyl-L-arninoacids.Since the maximum conversion is limited by the equilibrium of the reaction (compare with Fig. 7 4 , kinetic measurements of both hydrolysis and synthesis have been performed and fitted by a kinetic model, using acetyl-L-alanine as an example. Additionally the influence of D-alaninewas measured. Fig. 7-19 shows influences of acetyl-L-alanine,acetate, D- and L-alanine on the hydrolysis activity of aminoacylase. With respect to acetyl-L-alanine, the substrate of the hydrolysis reaction, the enzyme exhibits Michaelis-Menten kinetics (Fig. 7-19A). The hydrolysis activity is inhibited by acetate (Fig. 7-19 B) and L-alanine (Fig.7-19C). These are the substrates of the synthesis reaction (see below) acting as inhibitory products. Figure 7-20 shows the influences of acetate, L-alanine, D-alanine and acetyl-Lalanine on the specific activity of arninoacylase synthesizing acetyl-L-alanine. With respect to acetate as the substrate (Fig. 7-20A)the enzyme exhibits substrate inhibition (compare Eq. (33). Concerning the second substrate, r-alanine, MichaelisMenten kinetics are apparent (Fig. 7-20B). D-Alanine slightly inhibits the enzyme (Fig. 7-2OC). On adding acetyl-L-alanine(as the product of the synthesis reaction) to a reaction mixture as specified in Fig. 7-20 D, the measured enzyme activity rapidly decreases, indicating product inhibition. Finally the measured activity reaches negative values. A negative activity here means a negative rate of acetyl-L-alaninesynthesis, that is, hydrolysis.

7.4 Investigation ofEnzyrne Kinetics

A 2.5

2.0 h 7

.5

E

2

$ 8 .-

.-

.o

I-,

0.5

0.0

B 3.0 2.5

2.0 r

b E

2 1.5 .-> .-> c)

ti 1.0

a

0.5 0.0

I

0

500

I

I

I

1000

1500

2000

[L-Alanine] (mmol L')

This cannot be described by product inhibition alone and means the hydrolysis reaction commences above a concentration of 16.5 mmol L-' acetyl-L-alanine.At this value the reaction is in its equilibrium position and the measurable enzyme activity

225

226

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

c

2.0

I

1.5

-

h

F

3

I

[L-Ala] = 500 mmol L-’ -

1.0

[Ac]

= 400 mmol L-’

0.5

Measured

0 0.0 0

400

200

600

800

1000

[D-Alanine] (mmol L-’) D 2.0 1.5

-i3-J

[L-Ala] = 500 mmol L-’ [Ac]

= 400 mmol L-’ I

h

E

1.0

\

3

Measured

0

v

0.0

-0.5

I

I

I

I

3

[Ac-L-Ala] (mmol L-’) Figure 7-20. Kinetics of aminoacylase: effects o n the activity of synthesis of acetyl-L-alanine (initial rate measurements) [1151.

7.4 Investigation of Enzyme Kinetics

equals zero. Of course, the enzyme is not inactivated but there is no macroscopic turnover of substances. Fig. 7-19 and Fig. 7-20 show a number of effects on the activity of aminoacylase, which cannot be described by a standard kinetic model alone. To derive a kinetic model for the aminoacylase and for describing all influences, a reaction scheme based on the rapid equilibrium random uni-bi model was used (compare with Eq. (40)). This model was supplemented, according to the above measured influences of substrate inhibition by acetate and inhibition by D-alanine. Consequently, the reaction scheme Eq. (40)had to be extended to yield equation (46) and the corresponding rate equation (47),which is modified by the additional equilibria: E( D-Ala)

L-Ala

L-Ala

+

k,

Ac

E( Ac)z(L- Ah)

The substrate inhibition of Fig. 7-20A is represented by the complexes E(Ac)2 and E ( A c ) ~ ( L - A . ~This ~ ) . means that binding of acetate to all enzyme species besides the E(Ac-L-Ala)complex and the E(D-Ala)-complexis assumed. The rate equation of the random bi-uni model (Eq. (41)) is enlarged by three terms in the denominator according to the additional enzyme substrate complexes E(D-Ala), E ( A C ) ~and E(Ac)$-Ala), whereas the numerator has the same structure as in Eq. (41). The rate equation corresponding to the above reaction scheme can be written as follows: Kinetic model ofaminoacylase d[Ac --L - Ala] --

(47)

dt

[ E ] , . L .[Ac-L-Aka]-[E],

-

KAF&Al.

[Ac] [D-Ala]

1+--+--+ KAC

~

KwAi,

“.““.[Ac].[L-Ala] KAc ‘ K : : A l ~

[L-Ala]+ [Ac-L-AlaIl [Ac].[L-Ala] +- [ A c ] ~ K , ~Aia KAc-[.-,a ’ KA, KLA-CAI~ K A ’ K:: ~ ’

+

[Acy .[L-Ala] K A ’ ~KL~CAL~ .K t :

I

227

228

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

(mmol L-'1

Maximum activity of the synthesis of Ac-L-Ala Maximum activity of the hydrolysis of Ac-L-Ala Dissociation constant of the EX complex with X = acetate X = L-alanine X = D-alanine X = acetyl-r-alanine dissociation constant of the E(Ac)z-complexwith release of acetate dissociation constant of the E(Ac)(l-A1a)complexwith release of L-alanine

This kinetic model contains 8 parameters: one parameter for each of the four components D,L-alanine, acetate and acetyl-L-alanine,one parameter ( KL.AlaAC) for the second substrate in the bimolecular synthesis reaction, one parameter describing the substrate inhibition and two parameters describing the hydrolysis and the synthesis rate. This is the minimum number of parameters necessary to describe the Table 7-1). above influences on enzyme activity (6 Eq. (47) was used for a simultaneous fit of all kinetic data measured under initial rate conditions (Figs. 7-19 and 7-20). Separate fitting of each curve gives a better coincidence in every single case, but the optimized kinetic parameter will vary from fit to fit. The optimized kinetic parameters are summarized in Table 7-2. As described above, the rate equation can be set to zero yielding the equilibrium condition ofthe reaction (Haldane equation). Using the kinetic data, k& is calculated to be 12.2 mol L-'.

As ultimate proof of the kinetic model, a fit of time-courses of batch reactor experiments was performed (Fig. 7-21). Initial concentrations of the components over a significant range were chosen to yield hydrolysis conditions (1)and synthesis conditions (2) respectively. Additionally, the equilibrium positions of corresponding experiments A, B and C were chosen to be identical. Figure7-21 shows a good correlation of calculated and measured data over the whole range of the conversion, for hydrolysis as well as for synthesis. Table 7-2.

Kinetic parameters o f aminoacylase.

Hydrolysis ofAc-L-Ala

AH.^^ KA~-L-AI=

11.2 Umg-' 8.9 mmol L-'

Synthesis ofAc-L-Ala

118.1 1070 26.9 K L - A I ~ ~ ~ 1070 KL-AI~ 177 KD-AI~ 270

&.ma

K A ~ ~ KkA'

U mg-' mmol L-' mmol L-' mmol L-' mmol L-' mmol L-'

7.4 Investigation o f h z y r n e Kinetics

I

229

6 50

0 Time (min)

230

I

7 Reaction Engineeringfor Enzyme-CatalyzedBiotransformations

0

50

100

150

200

250

300

350

Time (min) Figure 7-21. Time-course data o f hydrolysis and synthesis o f acetyl-L-alanineusing initial conditions as specified in Table 7-3[”’1.

Table 7-3.

Starting conditions for the experiments shown in Fig. 7-21.

Concentrations IACIO [D,L-A~~], [Ac-L-Ala], X [L-Ala], X [~-Ala],

(mmol L-’) (mmol L-’) (mmol L-’)

(mmol L-’) (mmol L-’)

4’)

A(2)

B(’)

175 450 25 200 250

200 500 0 200 250

150 1400 50 200 750

200 1500 0 200 750

C(’)

C(2)

300 1300 100 400 750

400 1500 0 400 750

7.4.3

Kinetics of Multiple Enzyme Systems

If two or more enzymes are involved in coupled reactions, the influence of all substances present in the reaction mixture on the activity of all enzymes has to be studied and considered in the kinetic model. Knowing the kinetic behavior of the single enzymes, coupling can be done by writing mass balances. As an example, the reduction of dihydroxyphenylpyruvic acid (DHPP) to dihydroxyphenyllacetic acid (DHPL), a precursor of rosmarinic acid, is presented in Eq. (49)‘lI9’.

7.4 Investigation of Enzyme Kinetics

I

231

H"E?OH Dihydroxyphenylpyruvic acid DHPP 0

Dihydroxyphenyllactic acid OHPL a-Hydroxyisocaproate

HO

PEG-NADH + H"'

PEG-NAD")

e-c

H

I

I

(49)

The reaction is catalyzed by D-hydroxyisocaproate dehydrogenase (D-HicDH).The essential cofactor PEG-NADH is regenerated from PEG-NAD' by a second enzyme, formate dehydrogenase (FDH).By coupling to water-solublepolyethyleneglycol with a molar inass of 20 000 g mol-', the cofactor can be retained, together with the enzymes, by an ultrafiltration membrane, and the whole process may be performed continuously in an enzyme membrane reactor. The kinetic models of D-HicDH and FDH are described as follows (Eqs. (50) and (51)): Kinetic model of D-HicDH

u - ~= [D - HicDH], A,-,,,,,,, I -

d[DHPL] dr

K,,

[DHPP] + [DHPP]

[PEG - NADH + H+]

.-

Kinetic model of FDH

u

d[€or] = -- = [FDH], . A,,,,, dt

.

.

[For1

\PEG - NAD' ,

-

,

K,, + [For1

1 _.

In this case Michaelis-Mentendouble substrate kinetics is chosen (compare to Eq. (45)). The system involving D-HicDH with DHPP and PEG-NADH as substrates exhibits Michaelis-Menten kinetics for both substrates and competitive product inhibition by PEiG-NAD'. FDH also shows Michaelis-Menten kinetics for both substrates formate and PEG-NAD' and competitive product inhibition by PEG-

232

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

NADH. Both inhibiting effects are taken into consideration by an additional term in the denominator. This formal procedure is valid as it allows a correct fit of all initial rate data (not shown). For a batch reaction (compare to Sect. 7.5) mass balances of all components of this system may be formulated as follows (Eqs. (52-54)): d[DHPL] - d[DHPP] =v, dt dt

d[For] -~ =v * dt

d[PEG-NADH+H+] - - d[PEG-NADt] =v, -v, dt dt

(52)

(53)

(54)

The two enzymatic reactions are coupled by PEG-NADH and PEG-NAD’. This stoichiometric coupling does not affect enzyme kinetics but has to be considered when writing the mass balances. The discussion of this system will be continued in Sect. 7.5 where some implications of coupled enzyme systems on reactor design are described.

7.5

Enzyme Reactors 7.5.1 Basic Reaction Engineering Aspects

First, some basic terms of chemical reaction engineering will be discussed before introducing the field of enzyme reactors. For further reading, textbooks are available [1*&1261 The mode of reactor operation can be classified as “batchwise”or “continuous”. Batch reactions are started by filling a reactor with the reaction mixture and stopped after reaching the desired conversion. A steady state is only reached at equilibrium conversion of the reaction. A typical batch reactor is represented by the stirred tank reactor. Continuous reactions are characterized by a continuous substrate feed and product output. A residence time of the reaction mixture within the total reactor volume V can be defined by Eq. (55):

V F

=5

V

F

(h) (L) (L h-’)

Residence time Total reactor volume Substrate feed rate

(55)

7.5 Enzyme Reacton

stirred tank reactor (STR)

plug flow reactor (PW

continuous stirred tank reactor (CSTR)

&I-dx

t

4

4

._ .-

L

”

m c

8

*

I .

*

* time

+

time

4

position

time

4

position

position

Figure 7-22. Comparison of stirred tank reactor, plug flow reactor and continuous stirred tank reactor (Reaction: S + P; asterisks indicate time or position of substrate entering the reactor).

After a certain time a steady state will be reached within the reactor, meaning that concentrations of substrates and products do not change. Typical continuous reactors are the plug flow reactor (PFR) and the continuous stirred tank reactor (CSTR).A comparison of these different reactor types is given in Fig. 7-22. The differences between the reactors may be described by showing concentration profiles of substrate and product as a function of reactor position and time respectively. - The stirred tank reactor (STR) in batch mode exhibits a decreasing substrate

concentration and increasing product concentration with time, independently of the position within the reactor (the reactor is “wellmixed”, meaning that there are no gradients within the reactor). - The plug flow reactor (PFR) presents the same concentration curve along the reactor length, which is shown for the tank reactor with reaction time. In the steady state, the concentrations of substrates and products at distinct positions of the reactor do not change with time. The reactor has plug flow characteristics,

I

233

234

I

7 Reaction Engineerhgfor Enzyme-Catalyzed Biotransformations

-

implying flow of reaction mixture through the reactor in the form of a plug without any axial mixing. The continuous stirred tank reactor (CSTR) in the steady state exhibits constant concentrations as a function of both time and position. The exit stream from this reactor has the same composition as the fluid within the reactor; the CSTR is also a well-mixed reactor.

A very common variation of the CSTR is a cascade of n CSTRs. With an increasing number of reaction vessels, the cascade approximates to the plug-flow reactor. The product concentration increases stepwise from vessel to vessel. For example, a twostage cascade can be used to overcome effects of product inhibition, e.g. in the synthesis of L-tert-leucine [421 or GDP-Man[’44s14’1. The basis for calculating reactor operation conditions is the formulation of mass balances for all reaction components for the distinct reactor type. The mass balances for the above reactors can be formulated as follows: Stirred tank reactor: as no fluid enters or leaves the reactor the mass balance of every compound is defined by the reaction rate only. To determine the time t necessary to reach the desired conversion x the reciprocal rate equation has to be integrated from zero to the desired conversion x: With the definition of conversion shown in Eq. (56),

the equation for the reaction time t can be derived from the reaction rate v (Eq. (57)): r-i

at

2)

L

2)

J”

x

r. .

.

(57)

PlugJow reactor: The change of reaction rate within a unit volume passing through the length of the reactor is equivalent to a change corresponding to the residence time z within the reactor. To determine the residence time z necessary to reach the desired conversion x, it has to be integrated again over the whole range of conversion. Eq. (57) can also be used for the PFR simply by replacing t by z (Eq. (58)). x

z =[S&.jllu-dx 0

Continuous stirred tank reactor: In the case of CSTR, the change of concentration of a substrate S within the reactor (“accumulation”)is brought about by two terms: - The “convection term”, which describes the change of concentration of S within

-

the reactor as an effect of the influx of the substrate into the reactor, reduced by the efflux out of the reactor. The “reaction term”,which describes the change of concentration of S as result of the reaction S P. +

7.5 Enzyme Reactorr

accumulation = convection + reaction

&I

~~

-

b3, - bl --u

(59)

z

dt

At steady state, the concentrations within the reactor will not change, meaning that the accumulation term equals zero. By using the definition of x (Eq. ( S ) ) , the residence time T necessary to reach the desired conversion x can be calculated (Eq. (60)):

T is given by a simple multiplication of x

by the reciprocal reaction rate at this steady state conversion x. The situation can be further illustrated by the following plot (Fig. 7-23): k

STR, PFR

X

1

CSTR

b

X

*

2 CSTRs in series

X

*

’

*

2

Figure 7-23. Reactor design from l / v = f(x) plots (shadowed area represents r[S]o’; asterisks represent reactor outlet conditions).

I

235

236

I

7 Reaction Engineeringfor Enzyme-CatalyzedBiotransformations

If the reciprocal reaction rate is plotted as a function of conversion the shadowed areas represent the residence time necessary to reach a given conversion x (Eqs. (61) and (62)). ~

T

[SL

= 11/11 .dx

2 ~

[SL

(for stirred tank and plug flow reactor)

(61)

(for continuous stirred tank reactor)

(62)

0

= I/II . dr

Obviously the residence time is higher in the case of the CSTR compared to the STR and the PFR if the reaction rate continuously decreases with conversion, this most often being the case in enzymatic reactions. Minimizing the necessary r value for reaching a defined conversion x (at constant [Sl0 value) can be done graphically using these l / v = f(x) plots (Fig. 7-23). For different reactors, the area may be determined according to the above method and the appropriate reactor is given by the minimal area. For example the r value of a CSTR can be minimized by using two CSTRs in series. The steady state reaction mixture of the first reactor (conversionxl) is fed into the second reactor, yielding an overall conversion x2. With the relation 11 = [El,

.A

(compare to Eq. 23)

Eq. (61) and Eq. (62) may be written as

[SI,

= 11/ A

[EL -T

.dx (for stirred tank and plug flow reactor)

___ = I / A . x

[SL

(64)

0

(for continuous stirred tank reactor)

This means that the conversion x of a reaction at a constant [S]o value depends on the extent of activity during the conversion on the one hand and on the product [El0 . T on the other. Further, this means that lowering the enzyme concentration by a factor y and raising the residence time by the same factor y gives the same conversion x. This is the so-called ‘‘[El0 . r-concept“, valid for all three reactors. Up to now general reaction engineering principles have been discussed. Now we turn to enzyme reactor design. The suitability of different reactors is demonstrated for two typical enzyme kinetic examples, involving substrate inhibition in one case and product inhibition in the other (Fig. 7-24). The kinetics within Fig. 7-24 do not represent initial rate kinetics, but the reaction rate during the conversion S -+ P plotted versus the remaining substrate concentration (initial concentration [Slo = 1 mmol L-l).

7.5 Enzyme Reactors

I

237

[Substrate] (mmol L-')

-0:o Figure 7-24.

0.2

0.6 [Substrate] (mmol L-') 0.4

0.8

1.o

Performance of a CSTR depending on enzyme kinetics: A substrate inhibition, and (bold points indicate the inlet and outlet concentrations o f a CSTR).

B product inhibition -

The ideal reactor to overcome substrate inhibition (Fig. 7-24A) is the continuous stirred tank reactor (possible in form of an Enzyme Membrane Reactor, see below). In spite of a high feed concentration of substrate a high reaction rate occurs, as the steady state substrate concentration within the reactor is low.

238

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7 Reaction Engineeringfor Enzyme-CatalyzedBiotransformations

- In the case of product inhibition (Fig. 7-24B),the continuous stirred tank reactor

is not beneficial, as a high reaction rate would occur at high substrate concentration and low conversion only. With increasing conversion the inhibiting effect of the product becomes dominating, yielding a reduced reaction rate. The CSTR as a whole operates at low steady state substrate concentration and therefore low reaction rate. In the case of product inhibition, the plug flow reactor is advantageous, as the concentration of the inhibiting product slowly increases along the reactor length, whereas in a CSTR the product concentration is at the maximum value in the whole reactor. Therefore, the average reaction rate is higher in the PFR. A plug flow reactor may be realized using immobilized enzymes within a column reactor or using soluble enzymes within a cascade of membrane reactors. A batch or a repetitive batch process with soluble enzymes (see below) has the same productivity as the plug flow reactor. The overall reaction rate not only determines the necessary reaction time to reach the desired conversion (see Fig. 7-23) but also the enzyme and cofactor consumption, both being strongly influenced by the reactor conditions. Enzyme and cofactor consumption can be defined as follows: - Enzyme consumption is defined as the number of units of enzyme consumed per unit weight of product ( U kg-’); - Cofactor consumption is specified with the “total turnover number”, defined as

mols cofactor consumed per mol of product formed. As in most cases reaction rate decreases with conversion, enzyme consumption increases to the same extent (an exceptional case is when strong substrate inhibition overcompensates product inhibition). Therefore enzyme consumption is minimal under initial reaction rate conditions (zero conversion) (Fig. 7-25).Approaching total or equilibrium conversion the reaction rate approaches zero and enzyme consumption increases rapidly. In contrast to enzyme consumption, substrate utilization, defined as kg substrate consumed per kg product produced, increases with increasing steady state conversion. Besides the classical engineering question of reactor choice, the most important point in enzyme reactor design is the aspect of enzyme reuse, either by immobilization or by separation from the product stream. Batch processes without enzyme reuse are only possible if the costs of the biocatalyst are negligible. Different reactor techniques addressing the aspect of enzyme reuse are discussed in the following sections. 7.5.2 Reactors for Soluble Enzymes

If soluble enzymes exhibit sufficient operational stability, their use is advantageous, as the effort of immobilization and resulting mass transfer limitations can be avoided. Different techniques have been developed to retain soluble enzymes.

7.5 Enzyme Reactors

I

239

-cn

h

n

-

a

h

Y

d c

w

Y

2 C

C

.-0

0 .c

e

4-

E 3

3 C

v)

C

v)

a, c

a,

8

0

2

5

0

c

N C

v)

3

w

cn

0.0

0.2

0.4

Conversion Figure 7-25.

-

0.6

0.8

1 .o

Substrate utilization and enzyme consumption as a function o f conversion.

For small-scale synthesis enclosure of enzymes in dialysis tubes has been described for several systems (membrane-enclosed enzyme catalysis or the MEEC technique In this case mass transport of the low-molecular-weightsubstrates and products across the membrane becomes rate limiting because mass transport only occurs by diffusion and not by convection as described below. For synthesis on a preparative scale, repetitive batch processing has proved to be an effective and easy-to-handle method[12*].The repeated use of the enzyme is possible after concentration of the solution by means of commercially available ultrafiltration equipment and adding fresh substrate solution. Some of the advantages given for the Enzyme Membrane Reactor (see below) are also valid for the repetitive batch technique. Compared to batch processes, continuous processes often show a higher spacetime yield. Reaction conditions may be kept within certain limits more easily. For easier scale-up of some enzyme-catalyzed reactions, the Enzyme Membrane Reactor (EMR) has been developed. The principle is shown in Fig. 7-26A. The difference in size between a biocatalyst and the reactants enables continuous homogeneous catalysis to be achieved while retaining the catalyst in the vessel. For this purpose, commercially available ultrafiltration membranes are used. When continuously operated, the EMR behaves as a continuous stirred tank reactor (CSTR) with complete backmixing. For large-scale membrane reactors, hollow-fiber membranes To prevent concentration polarization on or stacked flat membranes are the membrane, the reaction mixture is circulated along the membrane surface by a low-shear recirculation pump (Fig. 7-26 B).

240

I

7 Reaction Engineerhgfor Enzyme-Catalyzed Biotransformations

A

pressure

substrate o

+-J-

product B

-0-

0

ration module

Figure 7-26. A Principle o f t h e Enzyme Membrane Reactor. 6 Set-up for larger scale including pump for circulation and filtration module.

Advantages and disadvantages of the EMR are summarized as follows: Advantages Working under sterile conditions is easy to achieve; No loss of enzyme activity by immobilization; No mass transport limitation; High volumetric activity achievable; Use of multi-enzyme systems without mass transport limitation; Use of coenzymes without mass transport limitation; Simple addition of fresh enzyme to compensate for enzyme deactivation; Ultrafiltered (pyrogen-free)product solution. Disadvantages - Sometimes limited by enzyme stability in solution; - Back-mixingis not optimal for enzymes with product inhibition.

Figure 7-27 shows a flow chart of the experimental set-up of a two-stage membrane reactor cascade. A 10 mL version of the reactor for process development and small-

7.5 Enzyme Reactors

241

a 1 v m analyiics

1

7

@ pressure gauge @ thermometer

Figure 7-27.

I

4 sterile filter 5

ultrafiltration module

1 substrate reservoir

6 on-line analytics

2 pump (dosing, circulation) 3 bubble trap

7 fraction collector

(PH, uv, polarimeter u. a,)

Flow chart o f t h e experimental setup o f t h e EMR.

scale production has been commercialized[1301. Depending on the reactor and membrane material, sterilization by means of steam or chemicals (peraceticacid) is possible. 7.5.2.1

Reactor Optimization Exemplified by the Enzyme Membrane Reactor

The performance of the EMR may be calculated by means of the measured kinetics and the simultaneous calculation of mass balances of each reactant. The steady-state parameters of the reactor can be estimated by numerical integration of the differential mass-balance equations by means of the Runge-Kutta method. Simulation of the reactor performance is a useful tool to find suitable conditions for production. Therefore it is a method resulting in a saving of both time and costs by avoiding large numbers of different experiments. But it has to be proven that the model is able to describe the process within the range of interest. In the following discussion the method of reactor optimization will be demonstrated using two enzyme systems introduced earlier, namely the enzymatic synthesis of N-acetylneuraminic acid and the enzymatic synthesis of cyanohydrins using oxynitrilase. Exampb: Enzymatic synthesis of N-acetylneuraminicacid This example has been used earlier to discuss the influence of thermodynamics on the reaction conditions (Fig. 7-7). It was shown that a great excess of pyruvate is helpful in order to increase equilibrium conversion, but for practical reasons there is a limit, because

242

I

7 Reaction Engineering for Enzyme-Catalyzed Biotransformations

1

1.0 I 0.9 h

v

I

I

I

I 700 - 600

8 0.8

- 500

0.7

- 400

2 a 0.6-

- 300

x C

.-0

0.5-

1

0.3 0.4

3

3 - 200 o_

2

6

h

- 100 7

1

3

2

Ratio [Pyr],:[ManNAc],

-0

(-)

Figure 7-28. Steady-state concentration and conversion as functions o f initial substrate ratio in an EMR [471; conditions: [El0 = 4 g L-', t=3 h, [ManNAcIo = 300 rnmol L-'.

- to reach 91% conversion instead of 86% the amount of pyruvate has to be

increased by 50% (conditions specified in Fig. 7-28); - with increasing concentration of pyruvate, downstream processing is more

difficult and time consuming (Neu5Ac and pyruvic acid have similar pK, values (2.0 and 2.4), complicating separation by anion exchange chromatography).

To enable a quantitative description of the system, kinetic measurements of synthesis and cleavage of Neu5Ac were performed and expressed by a kinetic The overall reaction rate is given by Eq. (66).

Abbreviations: A=Pyr, B=ManNAc, P= NeuSAc, S=synthesis reaction, C=cleavage reaction. 2) &,ma &,ma

K M ~ K M ~

KM"

KIA

(kat L-') (kat g-') (kat g-') (mol L-'1 (mol L-'1 (mol L-') (mol L-')

Velocity of Neu5Ac synthesis Maximum specific activity of synthesis Maximum specific activity of cleavage Michaelis-Mentenconstant of Pyr Michaelis-Menten constant of ManNAc Michaelis-Menten constant of Neu5Ac Inhibition constant of Pyr

7.5 Enzyme Reactors

I

243

Table 7-4.

Kinetic parameters of Neu5Ac-aldolase.

Synthesis of NeuSAc AS,nlaX

KMA

KMB Kv

KIB [El0 Kv

Cleavage of Neu5Ac

230.5 13.8 0.13G 402.2

pkat g-’ U mg-’ mmol L-’ mmol L-’

Ac.rnax KMP

KIA KIB

141.8 pkat g-’ 8.51 Umg-’ 9.44 mmol L-’ 1.30 mmol L-’ 23.8 mmol L-’

mmol L-’

1556

(mol L-*) (g L-7 (L mol-’)

Inhibition constant of ManNAc Enzyme concentration (aldolase) Concentration-dependent inhibition

The first term in the denominator of Eq. (66) represents a non-competitive inhibition (compare to Eq. (31)) of the enzyme by the sum of concentrations of A, B and P. This non-specific inhibition could be correlated with an increased viscosity of the 1311. As the mutarotation of the reaction medium in the presence of A, B and carbohydrates is fast compared to the enzymatic reaction there was no need for a discrimination between the a,P-anomers or the open-chain form of the monosaccharides. A complete set of kinetic parameters was determined, summarized in Table 7-4. The mass balances for the substrates and products are given by Eqs. (67-69).

d[ManNAc] - [ManNac], - [ManNAc] -2) dt z d[NeuSAc] - [NeuSAc], - [NeuSAc] +2) dt 0 At steady state the mass balance of each component N may be rewritten as shown in Eq. (70). d“1 - 0 =+ -dt

2)

“I,

=-.x

z

meaning that the reaction term is equal to the convection term (compare to Eq. (59)). This identity can be used to determine the conversion of the substrate at a given residence time from a plot u=f(x) (Fig. 7-29). The point of intersection of the function u=f(x) and the straight line v = [N]@ . x (the “convection term”) determines the operating point of the reactor at given concentration “lo and residence time z. The operating point is characterized by substrate conversion and reaction rate under steady-state conditions. By changing the slope of the convection line, e. g. by change of the residence time, other operation

244

I

7 Reaction Engineering for Enzyme-Catalyzed Biotransformations

[ManNAc], = 300 mmol L-'

250

~

h

r

v)

j 200

7

-

E

'D

0

-2

150

v 7

Q)

2 100 S

.-0 a,

50

CT

0 0.0

0.2

0.4

0.6

0.8

1.o

Conversion xManNAc(-) Figure 7-29. Graphical method o f determination of steady-state operation conditions in an EM Rl4'1 (bold points indicate steady-state reactor conditions; asterisks indicate steady-state conversion).

points may be determined. Instead of increasing the residence time the enzyme concentration may be increased to the same extent, yielding the same result (compare to the [El . T concept). This method enables determination of operation conditions of an EMR from batch reactor data. For that purpose a differentiation of the function x(t) has to be performed and converted to the above curve v = f(x). Using the mass balances, optimum operating conditions for continuous production in the EMR were calculated. Concentrations of 300mmol L-' ManNAc and GOO mmol L-' pyruvate were found to be the most suitable to allow high conversion of ManNAc, high space-time yield, and easy product isolation. Figure 7-28 shows steady-state concentration and conversion as a function of substrate ratio in an EMR. With a ManNAc concentration of 300mmol L-' and an equimolar amount of pyruvate in the feed, G8% conversion is reached and the product solution contains 200 mmol L-' NeuSAc, 100 mmol L-' pyruvate and 100 mmol L-' ManNAc. At this low conversion, unreacted ManNAc should be recovered, requiring an additional purification step. Conversion increases with excess of pyruvate to almost 90 % at only twofold excess of pyruvate, but there is a larger amount of pyruvate remaining in the product solution which has to be separated. In Fig. 7-30, conversion and space-time yield for the continuous process in an EMR are depicted. Enzyme concentration and residence time were chosen to be within practical limits to ensure a reasonably high conversion, suitable for production.

7.5 Enzyme Reactors I

'

- 2500

Reactor - - p - - -

0 Figure 7-30.

500

1000 1500 2000 2500 [ManNAc], (mmol L')

3000

Conversion and space-time yield as function o f substrate ~oncentration1~~1.

The increase of-conversion with rising concentration results from an increase of equilibrium conversion as depicted in Fig. 7-7. The decrease of conversion at ManNAc concentrations >500 mmol L-' is due to the increasing effect of nonspecific inhibition (compare to Eq. (66)).The space-time yield decreases when the decrease of conversion exceeds the increase of ManNAc-concentration. According to these results a feed concentration of 300 mmol L-' ManNAc was chosen for NeuSAc production, allowing a high conversion (>85%) and a good space-time yield (650 g L-*d-'). The latter may be raised by increasing enzyme concentration and decreasing residence time ([El . T=constant), resulting in the same conversion within a shorter time. The model proved to be correct by a comparison of predicted and experimental conversions for several enzyme concentrations and residence times (Fig. 7-31). Biocatalyst consumption per unit weight of product was found to be about 6000 U/ kg at a conversion of 78%. For production purposes, enzyme membrane reactors with a working volume up to 500 mL were employed for the synthesis of approximately 2 kg of N-acetylneuraminic acid and other derivatives such as ketodesoxynonulosonic acid (KDN) 11*', 13*1. Downstream processing was achieved mainly by anion exchange chromatography on a 30 L column followed by reverse osmosis to concentrate solutions before lyophilization. Example: Enzymatic synthesis of benzaldehyde cyanohydrin In the case of the enzymatic synthesis of cyanohydrins, enantioselectivity is the most important criterion. Investigation of the kinetics of the whole system (enzymaticand non-enzymatic reaction) offers the possibility to optimize reaction conditions to

I

245

246

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

1.o

0.8

2 0.6 C

.-0

r

?

C

8

0.4 0.2

-Calculated 0

Measured I

0.0 0

20000

10000

[El Figure 7-31.

a s L-')

I

I

30000

40000

Comparison o f calculated and experimental c o n ~ e r s i o n ~ ~ ~ ] .

obtain products with high optical purity. The discussion within Sect. 7.3.2.2 is extended to a consideration of reactor design. From the kinetics of the enzymatic and the non-enzymatic reactions (Fig. 7-13) it is concluded that the side-reaction is suppressed very effectively by working with high enzyme concentrations and at a low benzaldehyde concentration. Benzaldehyde may react with amino functions of the enzyme to form Schiff bases resulting in deactivation of oxynitrilase, so low stationary benzaldehyde concentrations are also necessary with respect to enzyme stability. As the EMR behaves as a CSTR, conversion in the whole reactor becomes high and the stationary benzaldehyde concentration becomes low if the product [El . T is chosen to be sufficiently high. Enzyme concentration as well as residence time T have been varied and the resulting conversion and enantioselectivity calculated. Fig. 7-32 shows enantioselectivity as a function of conversion in continuous experiments. Different operating conditions were obtained by changing the enzyme concentration. The benzaldehyde concentration is determined by its maximum solubility. A fourfold excess of hydrogen cyanide is necessary to reach conversions greater than 90%. A larger excess will favor the non-enzymatic reaction yielding lower enantioselectivities. As already postulated from the initial rate measurements, enantioselectivity increases with increasing conversion. There is a good correlation between calculated and measured values, as well as for other substrate ratios not shown here. The reactor conditions and results are presented for one experimental run.

7.5 Enzyme Reactors

1.o

I

I

n

I

1

0 h

v I

..->

. I . I -

0

0 -

Q) v)

.-0

0.8

Results Benzaldehyde 0.048 mol/L Hydrogen cyanide 0.180 mollL Enzyme concentration 0.38 g/L

0.6 0.4

Residence time 10 min 90 Yo Max conversion Enantiomeric excess >99 Yo

0.2

Space-time yield

. I -

a C w C

I

Calculated

773 g/(L'd) I

0

Measured

1

I

1

In a production run,the space-time yield was enhanced to 2400 g L-'d-' (R)mandelonitrile (ee>99%) at a residence time of just 3.8 min using an enzyme concentration of 0.95 g L-'. A similar investigation for batch reactor systems has been published recently[133]. DSM Linz in Austria established a process for the synthesis of (S)-phenoxybenzalde1341. The reaction is performed hyde cyanohydrin using a cloned (S)-oxynitrila~e~~'~ in a two-phase system with the substrate dissolved in an organic solvent.

Example: Coupled enzyme systems In Sect. 7.4.3, the reduction of dihydroxyphenylpyruvic acid (DHPP) to dihydroxyphenyllactic acid (DHPL)was used as an example for discussion of the kinetics of multiple enzyme systems (Eq. (49)).The rate equations for the reduction reaction of DHPP to DHPL (u1) and the regeneration of PEG-NAD' to PEG-NADH (u2) have been introduced (Eqs. (50) and (51)). The mass balances of all components in the case of performing the reaction in a CSTR are formulated as follows (Eqs. (71)-(75)): d[DHPP] - [DHPP], - [DHPP] --'u dt z d[DHPL] [DHPL], - [DHPL] +-'u dt z d[For] - [For]" -[For] dt z

--

--2)

2

I

I

(72)

(73)

247

248

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

d[PEG - NADH+ H’] dt d[PEG - NAD’] dt

=I), -Ill

(74)

=I),

-v2

(75)

In the case of PEG-NAD’ and PEG-NADH,the convection term equals zero as both are completely retained within the reactor by the ultrafiltration membrane. The enzymatic reactions are coupled by PEG-NADH and PEG-NAD’. As stated previously, this stoichiometric coupling does not affect the enzyme kinetics, but has to be considered when writing the mass balances. Considerations about process optimization of coupled systems with coenzyme 13’1. One aspect may be illusregeneration are discussed in the literature[”, trated here - the question of enzyme ratio within the coupled enzyme system. Figure 7-33 represents the influence of the ratio of the two enzymes D-HicDH for the synthesis reaction and FDH for the cofactor regeneration. The enzyme ratio represents the ratio of the initial volumetric activities of the enzymes (dimensions U mL-l). For process conditions, enzyme activity under the actual steady-state reactor conditions is significant, and differs from initial rate conditions as determined by enzyme kinetics. Therefore, the optimum enzyme ratio, implying maximum conversion within minimum residence time, is not 0.5. As evident from Fig. 7-33, it does not make sense to add one enzyme, e.g. the cheaper one, in excess to the reaction mixture, as, a long way from the optimum of 42,433

‘I: = 60 min

0.0

0.2

0.6 Enzyme ratio (-) D-HiDH/(FDH+D-HicDH) 0.4

0.8

Figure 7-33. Dependence o f conversion on enzyme ratio a t different residence times[”91.

1.o

7.5 Enzyme Reactors

0

20

40

60

80

100

120

Time (h) Comparison of calculated and measured data for the enzymatic synthesis of dihydroxyphenyllactic acid in an EMR[”91. Figure 7-34.

enzyme ratio, only one enzyme determines the overall reaction rate. This situation might appear in coupled enzyme assays where care has to be taken that the enzyme of interest is rate determining. As proof of the kinetic model, fitting of initial rate data or time-course data of batch reactions have been introduced in Sect. 7.4. Additionally, a proper fit of continuous reactions in an enzyme membrane reactor (EMR) may serve as confirmation of the kinetic model. For this coupled enzyme system, calculated and measured conversions at different operating conditions (varying [El and T values, not further specified) are presented in Fig. 7-34. 7.5.2.2 Control of Conversion in a Continuously Operated EMR

As a consequence of enzyme deactivation, conversion may drop during the continuous operation of enzyme reactors. To maintain a constant degree of conversion, two methods can be employed according to the [El . T-concept (see above): -

addition of fresh enzyme,

- reduction of flow rate.

Both can be done very effectively by using methods of online analytics combined with an appropriate automatic controller. Useful methods for online analysis of enzymatic processes are -

polarimetry (useful for reactions where chiral reactants are involved)[1361,

- UV spectrometry,

I

249

250

I

7 Reaction Engineeringfor Enzyme-Catalyzed Biotransformations

- online-HPLC (may be used effectively for controlling complex reactions (e.g in

peptide or carbohydrate synthesis)) 7.5.3

Reactor Systems for Immobilized Enzymes

The choice of an appropriate reactor for applications of immobilized enzymes as well as for soluble enzymes depends on the kinetics of the reaction. Kinetics of immobilized enzymes are not only a function of enzyme activity but also of substrate transport to the enzyme, which is affected by the matrix used for immobilization. For a description of immobilized enzyme kinetics the reader is referred to the compre138-1401. Additionally, the use of immobilized hensive literature in this field enzymes is treated in Chap. 6 of this book. A brief overview of enzyme reactors used for application of immobilized biocatalysts in the laboratory and on the industrial scale is given in Fig. 7-35. Examples of industrial processes are given in [2] and [20]. To study the kinetics of immobilized enzymes a recirculation reactor may be used. This reactor allows one to perform kinetic measurements with defined external mass transfer effects, reached by establishing a high flow rate near the catalyst, minimizing mass transfer resistance. The reactor behaves as a differential “gradientless” reactor allowing initial-rate kinetic measurements to be made. The fixed bed reactor, behaving as a plug flow reactor, is most often used for immobilized enzyme reactions. Typically, the reactor is used with an upward direction of the flow to avoid compression of the bed and to release gas bubbles generated during the reaction. Reactor design may be done readily without knowing the detailed enzyme kinetics. Kinetic measurements are performed with a recirculation reactor and the data are plotted in the form l / u = f(x) (see above). From this plot, the residence time necessary to reach a desired conversion x can be calculated as described. The different enzyme concentrations in the recirculation reactor and in the plug flow reactor have to be considered. Fluidized bed reactors are advantageous if small particles which would give high flow resistance in a fixed bed reactor are used to minimize external mass transfer resistances. Often it is useful to install nets at different heights of the reactor to approach plug flow characteristics. If the immobilized enzyme particle size is so small that an effective retainment is not possible in fluidized bed reactors, a slurry reactor may be used. This reactor guarantees catalyst retainment using a filter or a microfiltration membrane. For a larger particle size, the use of a stirred tank reactor is not advantageous because the energy input necessary to give an optimal fluidization of the particles is much higher than in a fluidized-bedtype reactor. Besides the immobilization of enzymes on solid particles, enzymes may also be immobilized on the inner or outer surface of tubular supports such as on hollow fibers or flat membranes. Enclosure of enzymes by the use of an ultrafiltration or dialysis membrane is regarded as a form of immobilization. [35-40r

7.5 Enzyme Reactors

Fixed bed recirculation reactor

Fixed bed reactor

+ P

S

Slurry reactor

Fluidized bed reactor 71

S V 0

O

0

O

0

0

0

Figure 7-35.

0

0

sieve

Reactors for use of immobilized enzymes.

7.5.4

Reaction Techniques for Enzymes in Organic Solvents

The same reactors can be used for dealing with immobilized enzymes in organic solvents or with one-phase organic systems as for dealing with enzymes in aqueous solutions. For one-phase systems, the enzyme may be recovered from the solution by means of membrane filtration. Suspended enzyme particles may be retained in a slurry reactor (compare to Fig. 7-35) by microfiltration membranes or stainless steel sieves, whereas in other cases such as reverse micelles, ultrafiltration membranes have to be used[’’]. For some years ultrafiltration membranes have been available

I

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7 Reaction Engineeringfor Enzyme-CatalyzedBiotransformations

Figure 7-36.

Multi-compartment enzyme membrane reactor.

which are stable toward organic solvents (e.g. from polyaramide or cellulose). In these cases the enzyme membrane reactor, as described earlier for the pure aqueous system, may be used without modifications, if all materials (sealing rings, tubes) are stable toward the solvent used. For two-phase systems, special reactors may be used such as multi-compartment reactors as shown in Fig. 7-36[43, 14', 1421. In the latter case, the two phases are separated by a hydrophobic or hydrophilic membrane (solid supported interface). The enzyme is soluble in the aqueous phase and substrate is added up to its maximum solubility in the aqueous phase. Substrates and products are distributed according to their hydrophilic or hydrophobic properties. The membrane area has to be large enough to avoid mass transfer limitations. High membrane areas may be achieved by flat membrane stacks or by hollow fiber modules. If lipases are used they may be adsorbed at the interface on a hydrophobic membrane. Such a system has been developed by Sepracor, USA, for the enantioselective hydrolysis of racemic esters (Fig. 7-37)and is used by Tanabe Seiyaku Co. for the kinetic resolution of 3-(4-methoxyphenyl)glycidicacid methyl ester, which is an intermediate in the synthesis of diltiazem [20*77,1431. Here the racemate is circulated on one side of the membrane while the water necessary for hydrolysis is picked up from the other side. The resulting acid is extracted into the aqueous phase, where the pH is kept constant with NaOH. The enzyme is adsorbed in the pores of the membrane. For the pilot scale production the following data were obtained: reactor productivity 125 g d - k 2 , enzyme consumption 17 g per kg product, enantiomeric excess of product 99 %, product yield 43 %. Additional reactor systems, especially reactors for synthesis of oleochemicals are summarized in [99].

7.G Conclusions and Outlook

rac-Methyl-methoxyphenylglycidic acid methyl ester

(PS,BR)-Methyl-methoxyphenylglycidic acid

(2R, 3s) Ester

f

--&&

rac. Ester Figure 7-37.

(ZR,SS)-Ester

(2S, 3R) Acid

f

Q-J

M e m b r a n e reactor system for resolution o f a diltiazem intermediate.

7.6 Conclusions and Outlook

Some basic aspects of reaction engineering for enzyme-catalyzedbiotransformations have been presented within this chapter, using examples of processes investigated by the authors. A strategy was shown starting with considerations focusing on the appropriate enzymes, continuing with an investigation of enzyme kinetics, and developing a reactor model. Finally, with the aid of process optimization, the most favorable reactor conditions can be found in order to achieve a high space-time yield with high conversion, high selectivity and low enzyme consumption. Enzyme engineering has reached a status where many limitations have been overcome by interdisciplinary efforts. New enzymes are being sought and, indeed, found in natural environments which will perform biotransformations on unnatural substrates. Huge quantities of enzymes can be produced by genetic engineering methods, making availability and costs of enzymes no longer a barrier for commercial processes. Much progress has been made in the evolution of biocatalysts to improve their properties. But still the choice of solvent to perform the reaction and the configuration of the surrounding reactor are important issues to be considered as well, to obtain optimum results. Based on enzyme kinetics well established by biochemists, complex reactions can be modeled. By utilizing classical reaction engineering methods, a quantitative

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description of process performance can be achieved and an operational status of a reactor can be calculated, saving experimental effort. Of particular note is the development of continuous processes, which have the advantage of better controllable reaction conditions, allowing suppression of undesired side-reactions and yielding better selectivity. Space-timeyields of several kilograms of product per liter of reactor volume per day have been reached. The methods of enzyme reaction engineering have already shown their benefits in numerous industrial processes which are being established successfully. Hopefully it can be concluded from this article that process development for the application of enzymes in organic synthesis can be performed on a rational basis.

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Enzymol. 1976,44,746-759. 2 T. Sato, T. Tosa, in Industrial Application of Immobilized Biocatalysts. A. Tanaka, T. Tosa, T. Kobayashi (eds),Marcel Dekker, Inc. New York, 1993, 3-24. 3 C. Wandrey, E. Flaschel, Adv. Biochem. Eng. 1979, 12,147-218. 4 C. Wandrey, R. Wichmann, W. Leuchtenberger, M. R. Kula, Degussa AG/GBF, US Pat. 4 304 858,1981. 5 H. K. Chenault, E. S. Simon, G. M.Whitesides, Biotechnol. Genet. Eng. Rev. 1988, 6, 221-267. 6 M. R. Kula, C. Wandrey, Methods Enzymol. 1987, 136,9-21. 7 R.Wichmann, C. Wandrey, A. Biickmann, M. R. Kula, Biotechnol. Bioeng.1981,23, 2789-2802. 8 K. Martinek, A. N. Semenov, I. V. Berezin, Biochim. Biophys. A d a 1981, 658,7G-89. 9 A. Zaks, A.M. Klibanov, Science 1984,224, 1249-1251. 10 V. J . Jensen, S. Rugh, Methods Enzymol. 1987, 136, 365-70. 1 1 J.G. Shewale, H. Sivaraman, Process Biochem. 1989,24,146-154. 12 J. Kamphuis, W. H. J. Boesten, Q. B. Broxterman, H.F.M. Hermes, J.A.M. van Balken, E. M. Mejer, H. E. Schoemaker, Adv. Biochem. Eng. Biotechnol. 1990,42, 134ff. 13 DSM/Stamicarbon, US Pat., 4 172 846, 1979. 14 W. Leuchtenberger, M. Karrenbauer, U. Plocker, Ann. N . Y Acad. Sci. 1984,434, 78. 15 K. Faber, Biotrunsfomzations i n Organic Chemistry, Springer, Berlin, 2000.

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References I 2 5 5 G. H. Gil, S. R. Kim, J. C. Bae, J. H. Lee, Enzyme Microb. Technol. 1985,7, 370-372. 33 K. Buchholz, Adv. Biochem. Engin. 1982,24, 39-72. 34 E. Flaschel, In Biocatalytic Production of Amino Acids @ Derivatives. D. Rozzell, F. Wagner (eds), Hanser Publisher, Miinchen, 1992, Chapter 14. 35 U. Kragl, L. Greiner, C. Wandrey in Encyclopedia of Bioprocess Technology. M. C. Flickinger, S . W. Drew (eds), Wiley, New York, 1999, 1064. 36 U. Kragl in Basic Principles ofApplied Catalysis M. Baerns, in press. 37 W. Keim, B. DrieRen-Holscher in Handbook ofHeterogeneous Catalysis. G. E d , H. Knotzinger and J. Weinkamp (eds), WileyVCH, Weinheim 1997,231. 38 E. Katchalski-Katzir,I). M. Kraemer,J. Mol. Catal. B, 2000, 10, 157. 39 P. Rasor in Chiral Catalyst Immobilization and Recycling. D. E. DeVos, 1. F. J. Vankelecom, P. A. Jacobs (eds), Wiley-VCH,Weinheim, 2000,96. 40 W. Tischer, V. Kasche, Trends Biotechnol. 1999, 17, 326. 41 D. Schomburg, Enzyme Handbook, Springer Verlag, Berlin 1991; www.brenda.unikoeln.de. 42 U. Kragl, D. Vasic-Racki,C. Wandrey, Bioproc. Eng. 1996, 14,291-297 43 U. Kragl, W. Kruse, W. Hummel, C. Wandrey, Biotechnol. Bioeng. 1996,52, 309-319. 4-1 C. Wandrey, Reaktionstechnische Untersuchungen an Enzymkatalysatoren z u r Enhvicklung kontinirierlicher Verfahren, Habilitationsschrift, TU Hannover, 1977. 45 I. H. Segel, Enzyme kinetics, John Wiley & Sons, New York, 1975. 46 A. S. Bomniarius, K. H. Drauz, H. Klenk, C. Wandrey, Ann. N. !I Acad. Sci. 1992, 672, 126-136. 47 U. Kragl, Reaktionstechnik biokatalytischer Prozesse am Beispiel der kontinuierlichen enzymatischen Sythese uon N-Acetylneuraminsaure, Berichte der Kemforschungsanlage Jiilich Nr. 2583,1992. 48 R. Schauer, Trends Biochem. Sci. 1985, 10, 357-360. 49 C. Hsu, Europ. Pat. Appl. 86300161.6 1986. 50 M. Perry, Ger. Offenll. 29 07 450 1980. 51 H. S. Tichy, Zur Elektrodialyse als integriertes Trennvelfahren in der Enzymtechnologie, Dissertation TU Miinchen, 1988. 32

K. Oyama, in Chirality i n Industry. A. N. Collins, G. N. Sheldrake, J. Crosby (eds), John Wiley & Sons 1992, Chapter 11. 53 H. D. Jakubke, P. Kuhl, A. Konnecke, Angew. Chem. Int. Ed. Engl. 1985, 24,85-93. 54 C. Wandrey, A. Fischer, 8. Joksch, A. Schwarz, Ann. N. Y. Acad. Sci. 1992, 672, 528-539. 55 U. Kragl, U. Niedermeyer, M. R. Kula, C. Wandrey, Ann. N. Z Acad. Sci. 1990,613, 167-175. 56 U. Niedermeyer, M. R. Kula, Angew. Chem. 1990,102,424-425. 57 U. Niedermeyer, Untersuchungen z u r enzymatischen C-C Verkniipfung am Beispiel der chiralen Cyanhydrinbildung, Berichte des Forschungszentrums Jiilich Nr. 2343,1990. 58 C. G. Kruse, in Chirality in Industry. A. N. Collins, G. N. Sheldrake, J. Crosby (eds), John Wiley & Sons Chichester, 1992, Chapter 14. 59 H. Griengl, N. Klempier, P. Pochlauer, M. Schmid, N. Shi, A. Mackowa, Tetrahedron 1998,54,14477. 60 U. Kragl, Reaktionstechnische Untersuchungen z u r enzymkatalysierten Cyanhydrinsynthese, Diplomarbeit, Universitat Bonn, 1988. 61 E. Ruckenstein, P. Rajora, Biotechnol. Bioeng. 1985,27,807-817. 62 J. E. Bailey, M. T. C. Chow, Biotechnol. Bioeng. 1974,16,1345-1357. 63 M. R. Kula, U.Kragl in Stereoselective Biocatalysis. R. Pathel (ed), Marcel Dekker, 2000,839. 64 C. Laane, J. Tramper, M. D. Lilly, Biocatalysis i n Organic Media, Elsevier, Amsterdam, 1987. 65 P. Douzou, Biochemie 1971, 53, 1135-1145. 66 J. S. Dordick, Enzyme Microb. Technol. 1989, 11,194-211. 67 D. K. Eggers, H. W. Blanch, J. M.Prausnitz, Enzyme Microb. Technol. 1989, 11, 84-89. 68 E. Andersson, B. Hahn-Hagerdal, Enzyme Microb. Technol. 1990, 12, 242-254. 69 C. J. Sih, Q. M. Gu, X. Holdgrun, K. Hams, Chirality 1992, 4, 91. 70 H. W. Blanch, D. S. Clark, Applied Biocatalysis, Vol.1, Marcel Dekker, Inc. New York, 1991, Chapters 1, 2. 71 A. M. Klibanov, CHEMTECH 1986, 16,354. 72 A. Klibanov, Nature 2001,409, 241. 73 G . Carrea, S. Riva, Angew. Chem. Int. Ed. Engl. 2000, 112, 2312. 74 1. M. S. Cabral, M. R. Aires-Barros, H. Pin52

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heiro, D. M. F. Prazeres, J. Biotechnol. 1997, 59, 133. 75 U. T. Bornscheuer, R. J. Kazlauskas, Hydro-

lases in Organic Synthesis, Wiley-VCH,Weinheim, 1999. 76 T. Zelinski, M.-R. Kula, Biocatal. Biotrans. 77

78 79 80

1997,15,57-74. J. L. Lopez, S . A. Wald, S . L. Matson, J. A. Quinn, Ann. N. I:Acad. Sci. 1990, 613, 155-166. Sepracor, Inc., Marlborough, U. S. Patent, 1989, US89/04784. C. Laane, S. Boeren, K. Vos, C. Veeger, Biotechnol. Bioeng. 1987, 30, 81-87. M. P. J. van Deurzen, B. W. Groen, F. van Rantwijk, R. A. Sheldon, Biocatalysis 1994, 10,247-255.

R. H. Valivety, P. J. Halling, Biochim. Biophys. Acta 1992,1118,218. 82 P. Adlerkreutz, Biocatal. Biotrans. 1996, 14, 81

100

1992,659-666.

H. J.Fromm, Initial Rate Kinetics, Springer Verlag, Berlin, 1975. 102 A. Cornish-Bowden, Fundamentals o f E n zyme Kinetics, Butterworth, London, 1979, reprinted IRL Press, 1990. 103 H. Bisswanger, Theorie und Methoden der Enzymkinetik,Verlag Chemie, Weinheim, 101

1979.

J. Lasch, Enzymkinetik, Springer Verlag, Berlin, 1987. 105 R. Eisenthal, M. J. Danson, Enzyme Assays, A Practical Approach, IRL Press, Oxford, 104

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1-30.

83

7. H. Kastle, A. S . Loevenhart,J. Am. Chem. SOC.1900, 24,491.

84

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86 87 88

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4, 153.

B. Orlich, R. Schomacker, Biotechnol. Bioeng. 1999, 65, 357. 90 P. Miethe, R. Gruber, H. Voss, Biotechnol. Letters 1989, 1 1, 449-454. 91 P. Miete, I. Jansen, U. Niedermeyer, U. Kragl, R. Haftendorn, M.-R. Kula, C. Wandrey, K.-H. Mohr, H. W. Meyer, Biocatalysis 89

1992,7,61-73. 92 P.

Wasserscheid, W. Keim, Angew. Chem.

Int. Ed. Engl. 2000, 112, 3926. 93 K. R. Seddon, J. Chem. Tech. Biotechnol. 1997, 68, 351. 94 T. Welton, Chem. Rev. 1999, 99, 2071.

M. Erbeldinger, A. J. Mesiano, A. J. Russel, Biotechnol. Prog. 2000, 16, 1129. 96 R. Madeira Lau, F. van Rantwijk, K. R. Seddon, R. A. Sheldon, Org. Lett. 2000,2, 4189. 97 S. H. Schofer, N. Kaftzik, P. Wasserscheid, U. Kragl, Chem. Commun. 2001,425-426. 98 K.-W. Kim, B. Song, M.-Y. Choi, M.-J. Kim, Org. Lett. 2001, 3, 1507. 99 M. Biihler, C. Wandrey, Fat Sci. Technol. 95

1992,94,82-94.

H. Hofmann, U. Hofmann, EinjLhrung in die Optirnierung, Verlag Chemie, Weinheim, 1971.

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J. R.Rice, Numerical Methods, Software and Analysis, 3rd edn, McGraw-Hill, Singapore, 1987.

263,8017. 85

J. G. T. Kierkels, E. T. F. Gelade, L. F. W. Vleugels, D. P. Vermeulen, C. Wandrey, W. J. J. van den Tweel, J. Kamphuis, in Biocatalysis in Non-Conventional Media. J .Tramper (ed),Elsevier Science Publishers B. V.

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H. Lineweaver, D. Burk, ]. Am. Chem. Soc. 1934,56,658-666.

R. Eisenthal, A. Cornish-Bowden, Biochem. J . 1974, 139,715-20. 110 A. Cornish-Bowden, R. Eisenthal,. Biochem. J. 1974,139,721-30. 111 P. J. F. Henderson, in Enzyme Assays, A Practical Approach. R. Eisenthal, M. J. Danson (eds), IRL Press, Oxford,1992, Chapter 109

10.

R. Zurmiihl, Praktische Mathematikfirr Ingenieure und Physiker, Springer Verlag, Berlin, 1965. 113 L. Michaelis, M. L. Menten, Biochem. Zeit. 112

1913,49, 333-3G9. Briggs, J. B. S. Haldane, Biochem.]. 1925, 19, 338-339.

114 G. E.

M. Biselli, Enzymkatalysierte Racematspaltung von Aminosauren mit integrierter Riickfirhrung, Berichte der Kernforschungsanlage Jiilich Nr. 2581,1992. 116 W. W. Cleland, in The Enzymes, 3rd edn, Academic Press, 1970, Chapter 1. 117 1. H. Segel, Enzyme kinetics, John Wiley & Sons, New York, 1975, pp. 115

841-845. M. I. Page, Enzyme mechanisms, University Press, Belfast 1987. 119 K. Pabsch, M. Petersen, N. N. Rao, A. W. 118

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Alfermann, C. Wandrey, Red. Trav. Chim. Pays-Bas 1991, 110,199-205. 120 0. Levenspiel, Chemical Reaction Engineering, 3rd edn, John Wiley & Sons, New York, 1999. 121 J. E. Bailey, 1). F. Ollis, Biochemical Engineering Fundamentals, McGraw-Hill,New York, 1986. 122 K. Schiigerl, Bioreaktionstechnik, Verlag Sauerlander, Frankfurt 1985, Vol. I, 1991, Vol2. 123 B. Atkinson. Biochemical Reactors, Pion Ltd. London, 1974. 124 H. Chmiel, BioprozeJtechnik, Gustav Fischer Verlag, Stuttgart 1991, Vols. 1 , 2. 125 M. F. Chaplin, C. Bucke, Enzyme technology, Cambridge University Press, Cambridge, 1990. 126 S. Fukui, Journal ofBiotechnol. 1990, 14. 127 M. D. Bednarski, H. K. Chenault, E. S. Simon, G. M. Whitesides, J. Am. Chem. Soc. 1987,109,1283-85. 128 U. Kragl, A. Godde, C. Wandrey, W. Kinzy, J. J.Cappon, J. Lugtenburg, Tetrahedron: Asymmetry 1993,4,1193-1202. 129 C. Salagnad, A. Godde, B. Emst, U. Kragl, Biotechnol. Progr. 1997, 13, 810-813. 130 Bioengineering, Walcl, Switzerland and Jiilich Fine Chemicals, Julich, Germany. 131 B. Gavish, M. M. Werber, Biochemistry 1979, 18,1269-75. 132 U. Kragl, D. Gygax, 0. Ghisalba, C. Wandrey, Angew. Chem. Int. Ed. Engl. 1991, 30, 827-828. 133 W. F. Willeman, U. Hanefeld, A. J. J. Straathof, J. J . Heinen, Enzyme Microb. Tech. 2000, 27,413.

P. Pochlauer, Chemistry Today 1998, 16, 15. C. Wandrey, R. Wichmann, in Enzymesand Immobilized Cells in Biotechnology. Laskin (ed), Benjamin/Cummings Publ. Co. Inc., Menlo Park 1985, 177-208. 136 C. Wandrey, Chem.-hg.-Tech. 1976,48, 537. 137 S . E. H. Syed in, EnzymeAssays, A Practical Approach. R. Eisenthal, M. J.Danson (eds), IRL Press, Oxford, 1992, Chapter 4. 138 K. Buchholz, Characterization oflmmobilized Biocatalysts, DECHEMA Monographie 84, VCH Weinheim, 1979. 139 M. F. Chaplin, C. Bucke, Enzyme technology, Cambridge University Press, Cambridge, 1990,80-137. 140 L. B. Wingard, E. Katchalski-Katzir,L. Goldstein, Applied Biochemical Bioengineering, Academic Press, New York, 1976, Vol. 1. 141 W. Kruse, W. Hummel, U. Kragl, Recl. Trav. Chim. Pays-Bas 1996, 115,239-243. 142 A. Liese, M. Karutz, J. Kamphuis, C. Wandrey, U. Kragl, Biotechnol. Bioeng. 1996,51, 544-550. 143 J. W. Young, R. L. Bratzler, Proceedings ofthe Chiral90 Symposium, Manchester, U. K., 1990. 161 U. Kragl, A. Liese, in Encyclopedia ofBioprocess Technology. M. C. Flickinger, S. W. Drew (eds),Wiley, New York, 1999, 454-464. 145 S. Fey, L. Elling, U. Kragl, Carbohydr. Res. 1997,305,475-481. 146 T. Hartmann, E. Schwabe, T. Scheper in Stereoselective Biocatalysis. R. Pathel (ed), Marcel Dekker, 2000, 799. 134 135

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

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8 Enzymic Conversions in Organic and Other Low-Water Media Peter Halling

8.1 Introduction

It is now often preferred to carry out enzyme-catalyzed biotransformations in nonaqueous media. These are commonly based on organic solvents, but a variety of other low-water media can be useful. Popular reasons for using such systems are listed in Table 8-1. When a low-waterbiotransformation is planned, a variety of choices must be made about the precise reaction conditions to be used. As in all biocatalytic systems, the rate and yield obtained can be greatly affected by the choices made. Some of the factors that must be considered are the same as in conventional aqueous media: temperature, reactant concentrations, the form in which the enzyme is added. New factors must be taken into account such as solvent selection and the level of residual water in the system. Other factors become somewhat modified: acid-baseconditions remain important, but, usually, pH is no longer a useful parameter to characterize them. This chapter is written primarily for a reader who wishes to carry out a low-water biotransformation, and requires some general advice on the selection of reaction conditions. It will be a long time before our understanding of these systems is sufficient to predict confidently the optimal conditions for a novel reaction. Of course, we cannot do this for an aqueous biotransformation, or, for that matter, most non-enzymic chemical transformations. However, it is possible to give recommendaTable 8-1.

Common reasons for choosing low-water media for biotransformations.

Reactants don’t have to be (very)water soluble - use different solvent, or just undissolved solids Changes in solvation alter equilibria and kinetics - e. g. readily available hydrolytic enzymes catalyse synthetic reactions (including direct reversal of hydrolysis) Can tune specificityby changing medium etc. Enzymes can be more stable Suppression of unwanted processes in aqueous solution, e. g. microbial growth, side-reactions Better integration of biocatalytic with chemical steps in non-aqueous media

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tions that will often lead to a reasonable performance at first attempt, and will guide the design of experiments to approach closer to the optimum. In general I will not discuss in detail the evidence that lies behind the recommendations presented here. The reader who wishes to explore this further should These form a basis for an consult the following list of recent reviews of the individual assessment of the most useful choices. This chapter is more aimed at those who wish to rely on my judgments and is written in a rather brief and prescriptive style in many places, in order to minimize the length. Undoubtedly some of these recommendations will be found less than ideal in the light of further investigation. However, I am fairly confident that most of them will remain good choices.

8.2 Enzyme Form

In aqueous media, the most usual state of the enzyme molecules is dissolved in the reaction medium. In this case, the previous treatment of the enzyme has little effect on catalytic activity, provided irreversible inactivation is avoided. In contrast, in lowwater media, the enzyme molecules are usually present in solid particles. The way the solid biocatalyst is prepared will clearly affect the state of the enzyme molecules, and hence their catalytic properties. Furthermore, as hydration of the enzyme molecules is reduced, it is clear that conformational changes can become much slower. As a result the previous history of the enzyme has important effects, not just the final conditions. In other words, there may be pronounced hysteresis effects. Enzymes are normally isolated from their biological source as an aqueous solution. Hence preparation for transfer to a low-water medium requires removal of much of the water from the environment of the protein molecules. A variety of different drying methods can be considered. Although presented below are certain statements about the relative activity of these different forms, it must be acknowledged that there have been few direct comparisons of the same enzyme prepared in different ways but then used in the same medium. 8.2.1

Lyophilized Powders

Lyophilization (freeze-drying)is one of the most obvious ways of producing solid particles from an enzyme solution, and is probably the most used in preparing lowwater biocatalysts. However, I would generally not recommend it. It is of course a standard and usually reliable method of drying enzymes for storage. However, lyophilized enzyme powders used directly in low-water media typically show much lower specific activity (per enzyme molecule) than other preparations. At least two factors seem to contribute. Many of the enzyme molecules in lyophilized powders have changed conformations, probably due to the conditions in the concentrated unfrozen regions during freezing. This (partial) denaturation is reversed on return

8.2 Enzyme Form I261

to aqueous solution, but not in low-water media. Considerable attention has been given to methods that can increase the specific activity of lyophilized powders by orders of magnitude, such as drying in the presence of salts or lyoprotectants. But these large increases reflect primarily the very poor activity of the control lyophilized powders. 8.2.2 Immobilized Enzymes

Immobilization is an alternative route to solid particles containing the enzyme, typically by attachment to a pre-existing support. Immobilized forms of many enzymes are now commercially available. For use in low-water media, the attachment is usually done very simply by adsorption - the support particles are simply stirred for a time in the aqueous enzyme solution. The strength of the linkage to the surface is not really an issue, because enzyme desorption will not normally be possible in low-water media. Much of the water is removed by decanting the supernatant. The wet solid particles may then be dried further by evaporation in air. However, a better method seems to involve rinsing with a suitable solvent that is able to dissolve plenty of water (i.e. usually a water-miscible solvent). This exploits the hysteresis in enzyme behavior to give a higher specific activity catalyst, at least for initial rates and with some enzymes. Drying procedures are discussed further below in the context of water effects. More complicated immobilization methods have been described, including covalent attachment to a support, entrapment inside particles (such as silica made by a sol-gel process), and covalent incorporation into polymer particles (“biocatalytic plastics”). However, it is not clear that any of these methods is superior to simple adsorption, particularly for use in low-watermedia, where enzyme desorption is not an issue. A method first described around 1980 has remained popular - drying a slurry of support material (commonly celite) in an enzyme solution. The resulting powder necessarily contains both solid support and enzyme. However, the link between them is usually very weak, and the method is now usually described as co-drying or deposition rather than immobilization. At least some of the enzyme is probably present as large aggregates, not close to the support. My impression is that catalytic activity is usually poorer than with enzymes immobilized by methods more likely to produce an even protein layer. 8.2.3

Cross-Linked Crystals

One enzyme form that has received considerable attention is based on enzyme crystals. Production of protein microcrystals from aqueous solution is often quite easy, and is increasingly used as a step in the manufacture of enzymes on an industrial scale. (Many people have the impression that protein crystallization is very difficult, but this stems from the problems in growing large near-perfect crystals for

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8 Enzymic Conversions in Organic and Other Low-Water Media

Protein powder (e.g. lyophilized) Figure 8-1.

Cross-linked protein crystal

Support immobilized

Schematic illustration of how enzyme molecules are organized in different biocatalyst

forms.

X-ray diffraction.) The protein molecules in the microcrystals are then covalently cross-linked by treatment with an appropriate multi-functional reagent, usually glutaraldehyde. This renders the crystals insoluble on transfer to different aqueous media. The cross-linked crystals are effectively another form of immobilized enzyme, and can be dried for transfer to low-water media by the same methods (again see further details in the discussion of water effects below). Cross-linked crystals are available commercially for a number of enzymes. Figure 8-1shows a diagrammatic representation of the organization of the protein molecules in lyophilized powders, immobilized enzymes and cross-linked crystals. 8.2.4

Direct Precipitation in Organic Solvents

A good and even simpler method of preparation is available where reaction media are based on an organic solvent that has the ability to dissolve considerable amounts of water (or is water-miscible).An aqueous enzyme solution may be mixed directly with the organic liquid (dry or nearly so). Most of the water in the enzyme solution forms a molecular solution in the organic liquid. The enzyme and other solutes are precipitated, usually as fine particles. The catalpc activity obtained is usually quite good, certainly compared with lyophilized powders. In principle particles containing active enzyme prepared this way might be transferred to other solvents by centrifugation and re-suspension. 8.2.5

Additives in Catalyst Powders

A great deal of attention has been given to co-drying additives with enzymes for lowwater biocatalysis. Large improvements in rate as well as alterations in selectivity

8.2 Enzyme Form 1263

have been demonstrated. A variety of mechanisms have been suggested for these effects, including the following. - Denaturation during the drying process may be prevented (“lyoprotection”); - The additive may select particular conformational microstates that become fixed

on drying (“imprinting”); - The environment of the enzyme molecules may be favorably altered, for example

by being made highly polar (salt effects). There are undoubtedly some interesting effects here. However, in general I cannot recommend such approaches at present to those persons looking for practical options for biocatalysis. All the main studies of these effects have been made with lyophilized powders as the final catalyst. As noted above, these usually give a very poor specific activity compared with other forms of the same enzyme. This is probably why it is possible to demonstrate very large enhancements in rate - these are relative to a very low base for the control lyophilized powders. It would be very interesting to see whether these additive effects are found with other low-water catalyst forms. It may be that they have been looked for, not found, and hence the results are not reported. Because of the low base activity of lyophilized powders, it is not clear whether the “enhanced rates” resulting from these various additives are better than (or even as good as) those obtained with other catalyst forms (e.g. immobilized). It would be useful for some studies to make this comparison systematically, with the same enzyme and reaction. Until such a comparison has demonstrated superior performance for the lyophilized powder, I would recommend the use of other catalyst forms as a simpler and more certain route to good catalyhc activity. One exception would be where it is wished to exploit the alteration of specificity brought about by additives. 8.2.6 Solubilized Enzymes

All the enzyme forms discussed so far remain as visible suspended solids in organic reaction mixtures. However, it is possible to treat enzymes so that they become solubilized in organic media (or at least no longer form an obvious suspension). The known methods can be classified into three categories,although these are not clearly distinct at the boundaries: -

The enzyme molecule can be covalently modified with organic soluble groups;

- The enzyme can form a non-covalent complex with appropriate agents, usually

either surfactants or organic-solublepolymers; - The enzyme molecules can be contained in the water cores of hydrated reverse

micelles or microemulsion droplets. These systems have been extensively studied, particularly the last. They have considerable attractions for fundamental studies, because the more or less transparent and mobile systems permit spectroscopic studies. Thus catalytic activity may be easily followed by direct spectrophotometric measurement of substrate or product

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8 Enzymic Conversions in Organic and Other Low-Water Media

concentrations. Fluorimetry or spectrophotometry can be use to probe the protein molecules directly. In microemulsions and some of the other systems, it has been clearly shown that the enzyme is molecularly dispersed. In other cases, however, the individual units remain aggregates, often quite large, but dispersed in such a way that the suspension is more or less transparent. The specific activity of the enzymes is often good, comparable with the best alternative enzyme forms. The enzyme molecules should all be well accessible to the medium, and mass transfer limitations avoided. However, solubilized enzymes have not achieved widespread use by those mainly concerned with applications in synthesis. An extra step is required to separate the enzyme from the final reaction mixture containing the products. It may be even harder to separate solubilising additives, notably surfactants, from the products. Thus I in general would not recommend solubilized enzymes for synthetic applications. One exception is where it is wished to attack polymeric or solid-state substrates, where the enzyme molecules may need to be able to move to contact the substrate, rather than vice versa.

8.3

Residual Water Level

The level of water remaining in these systems usually has major effects on behavior. Some of these phenomena are due to mass action effects of water as a reactant. The equilibria of hydrolysis reactions become more favorable as water levels rise, and normally the rates increase as well. Low-water media are commonly selected in order to use hydrolytic enzymes to catalyze synthetic reactions. Hydrolysis will thus be an undesirable side reaction, or in many cases the direct reverse of what is wanted. There are also cases where hydrolysis reactions are wanted, and hence water mass action should be maximized (while keeping non-aqueous media for reasons of substrate solubility, for example). Water levels also have important general effects on enzyme behavior. If too little water is present, the catalytic activity of most enzymes falls dramatically. On the other hand, reduction in water levels often leads to an increase in enzyme stability. A decline in catalytic activity at high water levels is also commonly observed, with several possible explanations: The rate of the monitored reaction may fall as a result of competition from hydrolysis (as a side reaction or direct reversal); - Water promotes agglomeration of catalyst particles, leading to mass transfer limitation of rates; - Water may act as a competitive inhibitor for any of the substrates. -

It is often difficult to prepare systems of reproducible water content just by adding a known amount. Water from other sources may be significant as well as that added deliberately. Water may be introduced associated with reactants, solvents or the biocatalyst preparation. It can also enter from (or escape to) the environment. Furthermore, if the effects of other parameters are being studied, experiments at

8.3 Residual Water Level

-~ ~

~

Water dissolved in solvent m] Enzyme bound water

Non-polar solvent

Polar solvent (same water content)

Polar solvent (same water activity)

Figure 8-2. Quantities ofwater present in different phases when solvents are compared at equal total water content or equal water activity. The behavior o f t h e enzyme is likely t o depend only on the amount ofwater bound t o it.

fixed water content can lead to very misleading conclusions. The water present will end up distributed between several different phases in the reaction mixture. Some will be closely associated with the enzyme molecules, and it is this quantity that will mainly affect their behavior. But some water will be dissolved in the bulk phase of the reaction mixture (e.g. the organic liquid). More will be associated with other solid phases present, such as an immobilization support and some may be present as vapor in a gaseous headspace. Changing these other phases (e.g. changing the nature of the organic solvent or the immobilization support) will affect the amount of water they retain, and hence that available to the enzyme. So what may appear to be an effect of solvent etc. may in fact be an effect of water. Put another way, the optimal water content (on a mass or volume basis) will change when alterations are made in several other factors. This is illustrated in Fig. 8-2. It is increasingly accepted that a better parameter to characterize the water levels in these systems is its thermodynamic activity. This is defined as 1 in pure water, and will take on lower values in the various reaction media. Water will tend to transfer between the various phases present until they all reach equal water activity. Hence the whole reaction mixture will tend to come to a single equilibrium value of water activity. This will reflect the amount of water in each phase. In particular, a given water activity will tend to correspond to a particular quantity of water associated with the enzyme molecules. Hence their behavior will be most simply related to water

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8 Enzymic Conversions in Organic and Other Low-Water Media

activity of the system, and the relationship will often stay the same as other factors are changed (e.g. the solvent). Even in terms of water activity, hydration effects are not quite so simple however. As well as the current value, enzyme behavior depends on the history of hydration to which the catalyst has been exposed. In other words, there can be strong hysteresis effects. Nevertheless, water activity values are usually the best basis to define the previous history reproducibly. 8.3.1 Fixing Initial Water Activity of Reaction Components

The reaction mixture for biocatalysis will be prepared by combining several components. To ensure defined water conditions in the final mixture, all these components should preferably be brought to known water activity beforehand. (It may be safe to disregard this for a component that has a limited affinity for water and makes up only a small proportion of the final mixture.) Often the easiest way to set the initial water activity of components of the reaction mixture is by pre-equilibration with a saturated salt solution. The relative humidity or water activity is fixed above a saturated solution of a given salt at a known temperature. As water equilibrates in or out of the solution, solid salt will tend to dissolve or crystallize to maintain saturation and hence the futed water activity in the headspace. Any other material placed in contact with the headspace will eventually equilibrate to the same water activity. The reaction mixture component can simply be placed inside a closed vessel together with the salt solution, such that water can transfer between the two via the vapour phase. Wide-mouth screw cap jars are convenient, with salt solution over the base and an open vial containing the sample (Fig.8-3). The rate of equilibration depends on the surface areas exposed and the amount of water that must be transferred. Typically 1-2 days is sufficient for either solid biocatalyst preparations or liquid phases based on relatively non-polar organic solvents. The rate of equilibration may be checked by weighing or Karl Fischer analysis respectively. Table 8-2 shows the water activity values generated by a selection of salts we commonly use, taken from the best literature source. Most of these values are weakly temperature dependent. However, it is essential to use them at controlled tem-

Jar with reasonable seal

, Sample being equilibrated Saturated salt slush Figure 8-3.

Method of pre-equilibration of water activity of reaction mixture components.

8.3 Residual Water Level Table 8-2.

LiCl

KAC

MgCb K2CO3 Mg(N03)2 NaBr

Saturated salt water activities at 25 O C (frornI'*l).

0.113 0.225 0.328 0.432 0.529 0.576

KI NaCl KCI KN03 K2S04

0.689 0.753 0.843 0.936 0.973

perature, as fluctuations can cause the liquid phase to move away from saturation. The saturated solution is best prepared to have a lot of crystals surrounded only by thin layers of liquid, which will then re-equilibratewith the solid more quickly. Good sample purity is important when such large amounts of solid are used. For water activities below 0.05, drying agents are required rather than salt solutions. Agents suitable for exhaustive drying are described in many conventional reference sources. Molecular sieves are popular in applied biocatalysis, but I would note two cautions. Firstly, if they are reactivated by heating, about 350 "C is required to obtain maximal drying efficiency. Secondly, if placed in direct contact with a liquid phase, they can have significant acid-base effects. With water-miscible solvents, the organic phase can be prepared at the desired water activity more conveniently by simple addition of water to the dry solvent. The water concentration required will be significant, and the amount of water added will be much larger than unintentional exchanges with the environment or residual water levels in the dried solvent. The relationship between water concentration and activity will be more or less fked for a given solvent, and little affected by reasonably low concentrations of reactants. This will not be true for less polar solvents, where direct addition of water rarely gives reproducible hydration or water activity. Table 8-3 gives water contents of various solvents at different water activities. A decision must be made about the sequence and timing in which components are combined to make the final reaction mixture. The choices made can have large effects on the final hydration conditions and biocatalyst behavior. It is usually best initially to prepare as separate phases: (i) a non-aqueous solution or mixture of the reactants; and (ii) the solid biocatalyst preparation (lyophilizedpowder, immobilised enzyme, cross-linked crystal etc). The best treatment to apply then depends on the objective of the experiment. - If the aim is to make a fair comparison of the effect of other factors (e.g. different solvents),then it is desirable to produce reaction mixtures of defined water activity.

For this purpose, it is best if the two phases mentioned are separately preequilibrated to the target water activity before eventually combining them to start the reaction. In principle it is possible for the water activity to change somewhat from the pre-equilibrated value as components redistribute between the two phases. However, in practice such changes are small if the two phases noted are chosen. Another option is to pre-equilibrate the biocatalyst particles suspended in a non-aqueous fluid, and to add one final reactant at time zero. This reactant should be one added at fairly low concentration to prevent significant changes in water activity. These two options are illustrated in Fig. 8-4.

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8 Enzymic Conversions in Organic and Other Low-Water Media Table 8-3.

Water contents (v/v%) o f various solvents equilibrated at different water activities.

Water activity

0.05

0.1

0.2

0.4

0.6

0.8

1

Ethanol 2-Propanol tea-Butanol Dioxane Acetone Acetonitrile Tetrahydrofuran Ethyl acetate Methyl iso-Butyl ketone Methyl tert-Butyl ether iso-Propyl ether Toluene Hexane

0.449 0.280 0.166 0.172 0.188 0.154 0.089 0.066"

0.94 0.58 0.34 0.36 0.39 0.32 0.185 0.137"

2.06 1.26 0.75 0.82 0.86 0.72 0.402 0.298

5.2 3.06 1.86 2.25 2.18 1.86 0.98 0.73

11.0 6.0 3.7 5.8 4.56 4.19 1.94 1.44

28.5 12.2 8.2 27.3 11.7 15.3 4.02 2.99

M M M M M M M

0.0529"

0.1061"

0.214

0.43

0.66

0.89

1.12

0.0470* 0.0178" 0,0015" 0.0004"

0.0940" 0.0356" 0.0029" 0.0007"

0.189" 0.071" 0.0058" 0.0015*

0.38 0.143* 0.01174 0.0029"

0.58 0.216 0.0175" 0.0044"

0.78 0.290 0.0233" 0.0058k

0.98 0.364 0.0292" 0.0073"

-

Water contents are given as conventionally in terms of the volumes of pure liquids mixed to reach the required composition. With water-immiscible solvents, a water activity close to 1 is achieved in the mutually saturated system (as shown), but for miscible solvents (M),water activity 1means pure water! Water activity ofwatermiscible solvents are estimated using the correlations derived by Bell et al.1191.For water-immiscible solvents, and the approximation of constant activity coefficient they are based on water solubility measurements~20~2*l, up to saturation. * - It is not advised to try to obtain the water activities shown by adding these small amounts of water to dry water-immiscible solvents. The values are given purely for use in estimating water quantities present or required. Apart from usual errors, small water droplets can take a very long time to dissolve in such solvents. Attempting such a method with solvents like toluene or hexane is particularly disastrous.

i'[J --,']

10 mM Ac-Tyr-OEt

Toluene

1 M PrOH

Silica-enzyme

, ,,,/-,

Little effect Of pre-equilibrating

solid powder -_

--. -. --_

1

t d''

Mix and pre-equilibrate water activity

- _- _- _ - _- - _- - _

._- - _ . --_

------* +

Combine to start reaction

1

Pre-equilibrate water activity

Choices i n water activity pre-equilibration before a reaction i n organic medium. The solid and broken arrows indicate two alternative schemes that can be considered. Other conceivable schemes will probably lead t o a final reaction mixture ofwater activity very different from that used in pre-equilibration. Figure 8-4.

8.3 Residual Water Level -

On the other hand, if the aim is to achieve maximal catalytx activity in otherwise iixed conditions, it is often better to transfer the enzyme catalyst quickly and directly from an aqueous environment. Excess water can be removed if necessary by rinsing with a suitable polar solvent. It is best if the polar solvent contains enough water to bring it to the target water activity of the final reaction mixture. It is still best to ensure that the non-aqueous fluid phase starts with a defined hydration level, usually by pre-equilibration.

In either case, it is worth noting that achievement of a desired final water activity value is much easier in relatively polar solvents, where the substantial water concentration in the bulk phase will effectively buffer the whole system. Two factors should be mentioned that can lead to significant unintended (and rather irreproducible) changes in water activity. Firstly, exchange of water with the environment. When only very small quantities of water are present (e.g. in media based on non-polar solvents), significant changes are possible. To avoid them, reaction vessels should be carefully sealed, and it may even be necessary to sample through a septum or a similar membrane. Sealing sufficient to prevent noticeable losses of volatile organic solvent may still allow significant water exchanges,because of the much smaller total quantities involved. Secondly, water may distill to any cold surface in contact with the headspace above a reaction mixture. This can lead to significant removal of water from the liquid phase, especially if the temperature difference is large. For example, a surface at 20 "C will condense water away from a reaction mixture at GO "C until its water activity falls to 0.12 (the ratio of saturated vapor pressures). 1t is best to prevent this problem by eliminating all such cold spots. Reaction vessels may be surrounded by an air bath (taking account of explosion risk if flammable solvents escape), or completely immersed in a water bath. If individually jacketed vessels are used, unjacketed surfaces may be heated above the circulating water temperature. 8.3.2 Control of Water Activity During Reaction

Some methods offer the possibility of control of water activity during the reaction at the cost of greater complexity and/or the possibility of interference with the desired conversion. Most obviously a change in water activity may be due to water being produced or consumed in the reaction. However, it is also possible by water exchange with the environment (e.g. during sampling). With non-polar solvents water activity can also change because conversion of substrates to products will change the solvation of water. Hence water activity at a fixed concentration will be altered. One simple and convenient method is the addition directly to the reaction mixture of suitable pairs of solid salt hydrates. A given salt hydrate will give up its water at a characteristic water activity, transforming to a lower hydrate or an anhydrous form. If the pair are placed in a system of water activity below their characteristic transition value, the (higher) hydrate will tend to give up water to the rest of the system. Water release will continue until the whole system reaches the transition water activity (or

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8 Enzymic Conversionsin Organic and Other Low-Water Media

the higher hydrate is completely exhausted). If on the other hand the salt pair are added to a system of high water activity, the form with less (or no) water will tend to take up water, transforming back to the (higher) hydrate. Once again, this will continue until the system water activity has been reduced to the transition value, or the salt form with less water has been completely consumed. In principle these exchanges should be able to buffer the water activity of the systems at the transition value of the added salt pair, provided enough is added. Typically, the higher hydrate will give up water at the start, as the reaction mixture is prepared from dry ingredients. Later, the lower hydrate may take up water produced in a reverse hydrolysis reaction. An example system is represented by the equation: NazS04.10H20 (crystals)G+ Na2S04 (crystals)+ lOH2O

(1)

in which equilibrium with both solids present can only be reached at a single water activity value, 0.80 at 25 "C. Figure 8-5 illustrates the use of this hydrate pair in a synthetic reaction.

100

Salt hydrate pair:-

80

gives out water here

s

n

and takes it up here

\

/

c 60 .-0

W

2 Q) >

40

0

20

0

0

20

40

60

80

Reaction time (h) Figure 8-5. Control ofwater activity by adding salt hydrates to reaction mixture. Synthesis of butyl butanoate catalyzed by Candida rugosa lipase. Control reactions with catalyst relatively wet (0) or dry (0)initially. Reaction in the presence of Na2S04plus Na2S04.10H20 (0).Kvittingen et a1.[221.

8.3 Residual Water Level Table 8-4.

Selected salt pairs found useful for water activity control in biocatalysis.

Salt pair

Equilibrium water activity

Rate of water transfer

Maximum temperature (‘C)

NaI.2/0 Na2HP04.2/0 LizSO4.1/0 NaAc.310 NaBr.2/0 Na&0,.5/2 &Fe(CN)6.3/0 Na4P~07.10/0 CaHP04.2/0 Na~HP04.7/2 Na2HP04.1217 Na~S04.10/0

0.12 0.16 0.17 0.28 0.35 0.37 0.45 0.49 0.50 0.61 0.80 0.80

fast fast slow fast slow slow slow slow slow fast fast fast

68 95 233 c 58 50 48 87 c 80 > 100 48 35 32

The pairs used are identified by a shorthand notation: Na1.2/0 means a combination of NaI.2H20 and anhydrous Nal (i.e. OH20).Equilibrium water activity values are for 25 OC.“Fast”water transfer indicates equilibration in a few minutes, “slow”that several hours may be needed. There is only limited information on the behavior of hydrate pairs giving lower water activities, though some indication that they generally tend to equilibrate slowly. From Zacharis et al.‘”’.

All of this describes just the thermodynamically favored directions of water transfer, for ideal crystalline solids. Many salt hydrate pairs seem to behave approximately ideally. However, if water activity is to be controlled close to the transition value, the rates of water release and uptake must be sufficient. Different salt pairs have very different rates ofwater exchange. It is difficult to give quantitative values, because the rates will depend on the size and shape of the crystals in each of the salt hydrate forms. This will depend on how they have been crystallized and handled subsequently. For example, cycling between hydrate forms, with gain and loss of water, will usually lead to a reduction in crystal size, and hence more rapid water exchange in future cycles. The equilibrium water activity achieved depends on the choice of salt hydrate pair used and the temperature. In most cases the temperature dependence is higher than for saturated salt solutions. There is also a maximum temperature at which the higher hydrate will “melt” to give a liquid phase, so above this the biocatalyst will probably be seriously affected. Table 8-4gives water activity values for some pairs that can be recommended for biocatalysis, together with an indication of the rates of transfer, and the maximum temperature. A compilation from the gives information on temperature dependence, and notes some other hydrate pairs whose use has not been (fully)tested. Many chemists have adopted the direct addition of salt hydrates as a simple method of water activity control. However, it does require a little thought and care to make sure the desired water activity is really produced. In particular, it must be ensured that both solid salt forms really will be present at equilibrium. It is best to estimate a “water budget” for the system, to ensure that enough of the right salt forms are being added. Table 8-5 shows an example of this, for a system made up of

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8 Enzymic Conversions in Organic and Other Low-Water Media Table 8-5.

Example water budget for the use of salt hydrates.

Phase

10 mL toluene 10 mg immobilised enzyme (on silica) 100 mL gas headspace Water produced by esterification reaction from 10 mM substrates

Initial water content (pmol)

Estimated equilibrium Change (pmol) water content (pmol)

55 (0.01% w/v)

128

+73

500 26 (lab air, 20% RH)

430 104

-70 +78

-

+loo

Water is assumed to show ideal dilute behaviour in toluene up to the solubility limit (16 mM). Equilibrium water content of immobilised enzyme estimated from measured adsorption isotherm (mainly adsorbed by silica support).

the phases shown, to be controlled at water activity 0.80 at 25 “C using the pair Na2HP04.12/7. This example has been selected as one in which all four contributions are significant. More usually, one or two will dominate. In the example, the added salt hydrates will initially have to supply 73 + 78-70 = 81 pmol water to the reaction mixture. As the reaction proceeds, this will be all need to be taken up again, followed by another 19 pmol. Hence 81/5 = 16.2 pmol of Na2HP04.12H20should be able to supply the water required, transforming to the heptahydrate as it does so. To take up the last portion of the product water, 19/5 = 3.8 pmol of NazHP04.7Hz0 should also be added at the start. In practice, about 10 mg Na2HP04.12H20 and 2 mg Na2HP04.7H20might be sensible, to ensure excess. Even more might be wise if the reaction vessel is likely to be opened frequently, allowing loss of water to the surrounding air. In some cases, estimation of a water budget may indicate that buffering can be achieved by adding just one hydrate form, with the other formed in situ as water is given out or taken up. This is particularly attractive where one of the salt forms is not available. However, it is clearly wise to adopt this approach only when the direction of net water exchange with the reaction mixture has been very confidently determined. If not, the second hydrate form required can often be made fairly easily by hydration or drying of the one that is available. Some limitations of the addition of salt hydrates must be borne in mind. In some cases the added salts may have additional, undesirable effects. They may react chemically with compounds involved in the enzymic reaction. For example Na2CO3.1OH20 will neutralize carboxylic acids to their Na salts. More subtly, even quite weakly basic or acidic salts may exchange H+ with the enzyme molecules, affecting their behavior (see below). There have been some cases of confusion in the control of water activity between saturated salt solutions (see above) and salt hydrate pairs. These can both be useful methods, but the principles and recommended applications are quite different. Avoid phrases like “control of water activity using salts”,which do not make it clear which method is being used. Water activity can be controlled during the reaction via the vapor phase, as in pre-

8.3 Residual Water level

equilibration. Once again, saturated salt solutions are the best method of generating a vapor phase of controlled water activity. However, if the reaction produces or consumes water at significant rates, simple diffusion via the vapor phase will usually be too slow to maintain constant water activity. Forced circulation of the gas phase may give sufficient rates. For best water transfer, it can be bubbled through both the salt solution and the reaction mixture. There is an alternative method to achieve faster water exchange between a saturated salt solution and the reaction mixture. The two may be brought into contact across a membrane, so that only a very short diffusion path separates them (at the cost of a smaller diffusivity of water within the membrane). Microporous or ultrafiltration membranes may be best in principle, but for laboratory use one convenient solution is to use silicone This is resistant to most organic solvents, and offers reasonable water permeability. In some reactions the objective may be to remove water as vigorously as possible. This will lead to a low water activity, which would result in very poor catalytic activity of many enzymes. However, some enzymes are much more tolerant of low water activity. In this case, exhaustive dehydration may be the best policy, particularly to minimize hydrolysis reactions or maximize their reverse. In general, the methods adopted can be based on those used in conventional synthetic chemistry for handling water-sensitivematerials. However, many of the most powerful drying agents cannot be used when they might come into contact with the enzyme, because of catalyst inactivation. For direct addition to the reaction mixture, the usual choice is molecular sieve. Type 4A is most commonly used, and is effective because nearly all the solvents have sufficiently large molecules that they are completely excluded. One piece of practical information is not as widely known as it should be. If molecular sieves are to be reactivated after use, very severe treatments are necessary to restore their full water-adsorbingpower. If heating alone is used, a temperature of 350 "C is needed. It has recently become clear that molecular sieves can affect enzyme behaviour by acid-base effects as well as water removal. If the components of the reaction mixture are all relatively involatile (e.g. in solvent-freeesterification),water removal by evaporation can be another effective method. 8.3.3 "Water Mimics"

One approach to improving catalfic activity at low water activity should be mentioned. Small additions of certain very polar liquids have been reported to greatly enhance catalFc activity at low water activity. They are usually described as "water mimics". and seem able to replace at least some of the roles of water in facilitating enzyme activity. Most of them are strongly hydrogen-bonded associated solvents that show other behavior analogous to water, such as glycerol, glycols and formamide. However, strong effects have also been observed with methanol and dimethylsulfoxide, for example. Most of the studies with these additives have been made with lyophilized powders, and hence may in part reflect the low control activities of these preparations (see Sect. 8.2). However, some significant effects have been reported with other enzyme forms, so I would recommend that use of such water mimics be

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8 Enzymic Conversions in Organic and Other Low-Water Media

considered. They are clearly particularly attractive where very low water activity is desirable to prevent unwanted side reactions. Obviously the water mimic chosen must not promote analogous side reactions, such as with its hydroxyl groups.

8.4 Temperature

The pattern of temperature effects is the same as in aqueous media. The initial rate of reaction increases with temperature, in the usual Arrhenius fashion. However, the stability of the enzyme will decline with temperature, and at high enough values catalyhc activity will be lost rapidly before significant conversions are reached. Hence, for given conditions, there will be an optimum temperature to maximize product yield after a given time. This is rarely a real fixed optimum for the enzyme, and for example will usually become higher if the reaction time is reduced. Progressive enzyme inactivation will have less effect over a shorter reaction time. One important feature that can be exploited in low-water media is an increase in stability to temperature. Hence reactions may be carried out at temperatures higher than would be possible in aqueous media, often by many tens of degrees. It is fairly clear that the most important factor here is the amount of water in the molecular environment of the enzyme molecules, as determined primarily by the water activity of the system. The presence or nature of a solvent has little additional effect. Thus, beware of statements that “enzymes become more thermostable in organic solvents”. It is the reduction in hydration that increases stability. If anything, the presence of an organic solvent will be destabilizing (in a comparison at equal water activity). In an organic solvent at water activity close to 1 (i.e. water saturated), the stability will be no better than in water. If, however, water activity is reduced to substantially below 1, a very valuable increase in stability may be achieved. 8.5 Substrate (Starting Material) Concentrations

Substrate concentrations affect catalytic rates in the same general way as in aqueous solution. At low substrate concentrations the rate is roughly proportional to [S] (i.e. first order kinetics). At higher concentrations the enzyme becomes saturated with substrate and the activity approaches a maximum limiting value. The full dependence is often described quite well by the Michaelis-Mentenequation or its analogs for the more common two-substrate case (general two-substrate model, or the PingPong model). These equations include a K, parameter for each substrate, with units of concentration. When the actual substrate concentration is many times larger than its K, value, the enzyme will be saturated with that substrate. Further increase in its concentration will then have little effect on the rate of reaction. When the medium is changed, the K, values will change also. An important contribution to this change has nothing to do with the enzyme directly, but reflects

8.5 Substrate [Starting Material) Concentrations

a,

c

mX

0N

a,

10

1

1

0.1

L-

I

!&

a, t a,

3 -

:

100

10

u)

9

YE

0.1

b

0.01

10

100

s,

1

1000

(mM)

Figure 8-6. Kinetic parameters for subtilisin-catalyzed transesterification o f 2-AlaO N p in different solvents. Experimental Km (0)and Vm/Km (0) values are shown as a function ofsubstrate solubility. The filled symbols show the corresponding “corrected” values, after allowing for substrate solvation. The variation i n V,/K, is largely explained by solvation, while the “real” variation in K, is opposite t o the apparent trend. Reimann et

the changed solvation of the substrates in the different media. Often this effect accounts entirely for the observed change. A simple quantitative picture is based on the relationship of K, values to substrate solubility: the ratio of these will be approximately the same in each different medium. Figure 8-6 illustrates an example of this effect. Often experiments to screen different solvents will keep the same substrate concentration in each. Hence, if a solvent in which the substrate is more soluble is tested, the K, value will be increased, and the reaction rate may fall, as the enzyme is more limited by the availability of substrate. For preparative syntheses, good general advice is to use a saturated solution of the substrate(s)in any solvent tested. This will only be a poor choice in the relatively rare cases of substrate inhibition. It will certainly be a good policy to allow identification of any direct effects of the solvent. An obvious way to ensure that the medium is saturated with substrates is to include excess in the form of solid particles. This leads

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8 Enzymic Conversions in Organic and Other Low-Water Media

towards the mainly solid reaction mixtures mentioned elsewhere, and can be a good option in practice.

8.6 Solvent Choice

A large number of solvents might be chosen to form the basis of the low-water medium. The choice of solvent will usually have important effects on the rate and selectivity of the reaction, and on the stability of the biocatalyst. 8.6.1

Effects on Equilibrium Position

In many biocatalyzed reactions, the position of chemical equilibrium is important, because it will place a limit on the eventual yield. In such cases, the choice of solvent will usually have a significant effect on the equilibrium position. Because this simply reflects the differential solvation of reactants and products, these effects can be predicted fairly confidently, at least to a reasonable approximation[271. One of the equilibria most commonly of interest is esterification. It may be desired to hydrolyze an ester, or reverse this in condensation of an alcohol and acid. Alternatively the hydrolytic equilibrium may be an undesirable side-reaction during transesterification. In this case, at a given water activity, the equilibrium position is quite strongly solvent dependent. The fraction of ester will increase dramatically on going from a polar solvent to non-polar solvent (Fig. 8-7). Hence alkanes are preferred solvents for esterification, while acetonitrile, a ketone or tertiary alcohol would be best for ester hydrolysis. If the equilibrium constant is expressed in terms of concentrations (including that of water), it is relatively solvent independent. However, optimal enzyme behavior in the different solvents usually requires maintaining the same water activity. At futed water activity, the ratio of ester to acid and alcohol concentrations will be maximized in the least polar solvents. 8.6.2

“Solvent Effects” that Really are Not

Many apparent “solventeffects”reported in the literature are actually due to changes in the availability ofwater or substrate to the enzyme. It is commonly observed that activity appears to be highest in the least polar solvents. Sometimes the explanation will be added that these “have the least tendency to strip water from the enzyme”. This undoubtedly indicates a common mechanism, but in such cases the “solvent effect” will disappear completely if experiments are run at equal water activity, as recommended in the discussion above (Sect. 8.3.1 and Fig. 8-2). Many other observed “solventeffects” operate via changes in substrate solvation, as explained in Sect. 8.5. Hence, they are really effects of changing substrate availability when different solvents are compared with equal substrate concentrations.

8.6 Solvent Choice

P'

0

Y

CSI 0 2

0 -3

I

-2

I

-1

I

0

I

Figure 8-7. Correlation between equilibrium constant for esterification and solubility o f water in the solvent. Equilibrium constant was defined as [Ester]/([Alcohol].[Acid]), for reactions at fixed water activity (close t o 1). Solvents are: bb, butyl benzoate; be, bromoethane; bk, dibutyl ketone; bp, dibutyl phthalate; bz, benzene; ca, 1 ,1 ,1-trichloroethane; cf, chloroform; ct, carbon tetrachloride; cy, trichloroethylene; ee, ethyl ether; ek, diethyl ketone; ep, diethyl phthalate; hd, hexadecane; hx, hexane; mc, methylene chloride; mk, methyl iso-butyl ketone; nm, nitrornethane; oc, iso-octane; pe, ;so-propyl ether; tl, toluene. Valivety et al.[281.

8.6.3 Solvent Polarity Trend and Recommended Choices

A very commonly observed trend is that activity is highest in the least polar solvents. In many of these cases this is an effect ofwater or substrate availability, as just noted. Hexane is regularly identified as the best medium, because the low solubility of water and most substrates makes them most available to the enzyme, when comparisons are made at equal concentrations. Nevertheless, even when water and substrate availability have been allowed for, non-polar solvents seem to offer the highest activity. The probable explanation involves the tendency for solvent molecules to migrate from the bulk phase into the immediate environment of the enzyme. The picture is simplest when there is a discrete aqueous phase (albeit of very small volume) around the enzyme molecules. The more hydrophobic the bulk solvent, the lower will be the (saturating) concentration in the aqueous phase, which is what is experienced by the enzyme. Even in the absence of an identifiable aqueous

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8 Enzymic Conversions in Organic and Other Low-Water Media

phase, the immediate environment of the enzyme molecules will be more polar than the bulk. Hence, it is often best to select a non-polar or hydrophobic solvent, at least for initial trials. Some reasons why this might not be the best choice are: If the reaction wanted is a hydrolysis, the equilibrium will be less favourable than in a polar solvent (see above); - The reactants may be only poorly soluble; however, using a suspension of incompletely dissolved substrates may still be a good policy. Provided the rate of dissolution and concentration in solution are sufficient, a good reaction rate can still be achieved. -

The following list presents some choices for a more general solvent screening exercise: - An alkane: n-hexane is most commonly used, although on safety grounds

cyclohexane,heptane or isooctane would be preferred. - An aromatic hydrocarbon: toluene would usually be preferred over benzene on

safety grounds. An ether: diethyl ether is usually inconveniently volatile, and popular alternatives are di-iso-propylether or methyl tert-butylether. - A ketone: methyl iso-butyl ketone or acetone. Being miscible, the latter may not be suitable if a medium of high water activity is required - this will end up as a high water content mixture that may dissolve and denature the enzyme. - A tertiary alcohol: tert-pentanol or tert-butanol. These are useful because they do not react with most enzymes that accept alcohol substrates. - A water-miscible but aprotic solvent: one of tetrahydrofuran, dioxane or acetonitrile. - A small alcohol. Either ethanol or 2-propanol is probably best. These solvents must be avoided for many enzymes, as they will be reactive, for example as nucleophiles or reductants. Methanol as a pure solvent is often particularly inactivating. - Chlorinated solvents can have some distinctive properties but are usually avoided for two reasons. On safety and environmental grounds, they are increasingly disfavored for large scale applications. They also tend to be more inactivating to biocatalysts than other solvents of similar polarity. (In some cases this may in fact be due to the stabilizers added to most chlorinated solvents.) -

Supercritical fluids have advantages as reaction media for large scale applications, but the need for high pressure apparatus means they will not usually be favoured for laboratory syntheses. Volatile reactants can be supplied to a catalyst through a gas phase, and the higher temperature stability under low water conditions makes this applicable to more cases than might first be thought. However, the increased complexity of apparatus again makes this more likely to be favored only at an industrial scale.

8.6 Solvent Choice 279

I

8.6.4

Solvent Parameters

For preparative purposes, the idea of correlation with some qualitative idea of solvent polarity is often sufficient, as implied here. There are numerous parameters which can be used to quantify the difference between solvents, but they all show some correlation with each other. By almost any measure, we would obtain the order: hexane, toluene, methyl iso-butyl ketone, propanol. However, different parameters can give different rankings when more similar solvents are compared. For biocatalysis in non-aqueous media, there are few effects where the “correct” solvent scale can be confidently identified. However, it is useful to have an idea of two quite different classes of solvent scale. - Most of them describe features of how the bulk solvent behaves and is able to

interact with isolated solute molecules. These will be based on measurements on or in the solvent as a bulk medium. Different parameters measure different features of the interaction the solvent may have with solutes, e.g. dielectric, cohesiveness, acidity, basicity. When the behavior of the solvent as a bulk medium is being considered, it is appropriate to use scales from this group. - In contrast, some parameters are properties of individual solvent molecules. Examples are dipole moment and log P (the octanol-water partition coefficient). These parameters are appropriate where individual solvent molecules are engaged in interactions away from the bulk phase. Thus, log P is used sensibly to describe the tendency of solvents to interact with (and affect the functioning of) the enzyme molecules. However, these parameters are not good choices when bulk solvent behavior is important, such as its ability to solvate water or reactants (and hence affect their availability to the enzyme). Even when such mechanisms are important, it is quite common to see correlations presented against log P. However, any relationship probably reflects the correlation of log P with appropriate scales of bulk solvent behaviour. There is a tendency to use two different words that make a related distinction between different types of solvent parameter. The log P parameter can be called a measure of solvent “hydrophobicity”,which is an accurate description of what affects its value. This contrasts with other parameters such as dielectric,which measure the bulk “polarity”.One illustration of the difference is to consider homologous series of solvents. Adding extra methylene groups to an alcohol, for example, will cause a regular increase in hydrophobicity.The effect on polarity will be much less, however, as the hydroxyl groups can still be oriented to solvate a polar solute. Thus decanol is more hydrophobic (higher log P) than hexane, but will be more polar by almost any measure of bulk properties. One illustration of the difference between these two classes of measure comes in the treatment of mixed solvents. For parameters that relate to the ability of the solvent in bulk to interact with solutes, it is meaningful to define and measure a value for a mixture of solvents. Often this will be a simple function of the mole or volume fractions and the pure solvent values. However, for parameters that describe

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8 Enzymic Conversions in Organic and Other low-Water Media

the behavior of individual molecules of the solvent, a value for the mixture is meaningless. The two types of solvent molecule present will behave differently and essentially independently. 8.6.5 Solvent Effects on Selectivity

Solvent effects on enzyme selectivity or specificity are very important. One of the attractions of non-aqueous media is the ability to tune these key properties, and substantial effects can certainly be observed. Unfortunately, it is not yet possible to give confident predictions in most cases. Predictions can be offered for the effect on selectivity between two substrates. A major contribution here comes from differential solvation, and the selectivity at a fixed concentration ratio will depend on relative solubilities, as noted in Sect. 8.5. However, these effects are rarely of preparative relevance, as it is not common to use two competing substrates that differ greatly in solvation. Selectivity between enantiomers is often desired, but here solvation effects will not distinguish the two substrates (unless a chiral solvent is used). Changing the solvent can have important effects on the selectivity between enantiomers (up to 2 orders of magnitude, with inversion of stereopreference possible). The effects must by definition be based on differential solvent interaction with the two diastereoisomeric transition states. A model based on solvent interaction with exposed portions of the substrate moeities in these transition states can sometimes make correct predictions of the direction of the effect, although its generality needs more testing. 8.6.6 No Solvent or Little Solvent Systems

In many cases an attractive option is to use no “solvent”at all. In some cases at least one of the reactants will be liquid, so can be the basis of a fluid phase for transfer of reactants. If slightly raised temperatures are used, this condition will be met more often. (Remember that at reduced water activity, the enzyme will usually be stable to higher temperatures than in aqueous solution.) Another option is to abandon the usual idea that most or all of the starting materials should be dissolved in order to get effective reaction. Attractive reaction mixtures can be prepared containing mainly undissolved solid particles of substrates. The reaction actually takes place in a liquid phase containing the enzyme, but this can be totally hidden between the reactant particles. The system formed is illustrated in Fig 8-8. Usually the liquid phase will be generated by adding a small amount (e.g. 10% by weight) of a “solvent”.Often the best solvent is water itself, as it will usually give the highest catalytic activity. In these mainly solid systems, this may be combined with many of the advantages of non-aqueous media, notably the reversal of the equilibria of hydrolytic reactions.

8.7 Acid-Base Conditions

Liquid phase (e.g. aqueous)

Solid reactants and products Schematic illustration of mainly solid reaction system. Starting material crystals will progressively dissolve, while product crystals will grow, as the enzymic reaction happens in the liquid regions between them. Figure 8-8.

8.7 Acid-Base Conditions 8.7.1 pH Memory

It is well known that pH has a major influence on the behavior of enzymes in aqueous media. Most who use enzymes under low-waterconditions are aware of the phenomenon known as “pH memory”. The activity and other properties of the enzyme are affected by the pH of the last aqueous phase to which it was exposed before drying for use in low-water conditions. This phenomenon is usually attributed to the relative rigidity of enzyme molecules at low hydration, by analogy with the effects of co-drying with additives. (see Sect. 8.2.5) However, another picture may give a clearer view of pH memory and when it may prove insufficient to apply controlled acid-base conditions. In aqueous solution, pH influences enzymes by affecting the protonation state of acidic and basic groups in the molecule. At a given pH, the protein molecule will have a characteristic net charge. Electroneutrality requires that the surrounding solution contain an excess of oppositely charged counter-ions precisely to balance the protein charge. In aqueous solution these counter-ions are relatively far away, and their presence and identity has only limited effects on behavior. However, consider drying this portion of aqueous medium containing the enzyme. In general, the counter-ions present will remain, as only water is removed. So the net charge on the counter-ions, and hence the opposite net charge on the protein will be preserved. The requirement for electroneutrality means that the only possible changes in protonation state are internal H’ exchanges between groups in the protein. Each such exchange will create or destroy a pair of positive and negative groups, without altering the net charge. In summary, this picture shows that pH memory resides in the behavior of the counterions as much as the protein, and does not require any special rigidity of the latter. This is illustrated in Fig. 8-9. Also it should now be clear that pH memory is not a phenomenon unique to non-

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8 Enzymic Conversions in Organic and Other Low-Water Media

0

3

o\te r.,.

0 0

0

3

a

0

3 8

/

@

8

0

c

J

0

(net charge) 3

,

8

c

3 n

High dielectric

net charge

3

0

=+c

%%j/

\

'barge

0

'barge

Aqueous solution

(net charge]

Dried

3s = + c

Return to liquid medium

arge

Figures-9. Illustration ofthe relationship between protein net charge and that on the counter-ions, and how drying and re-suspension or dissolution cannot change it.

aqueous media. If a dried enzyme preparation is placed back into pure water, its behavior will be determined by the pH value before drying - the ionization state of the protein, and the counter-ions present will effectively buffer the water back to the original pH value. Of course, we would not normally think of doing an experiment like this in aqueous media. The pH value reached would be very weakly buffered, and might be greatly altered by traces of acid or base from impurities, reactants etc. In low-water media, it is more common that acid-base conditions will not be seriously affected by these unintended effects. Hence pH memory may be sufficient to control behavior. However, there are several common ways in which pH memory may fail, which at least should be carefully considered before deciding to rely on pH memory. In addition, there are now established some relatively simple methods to buffer acid-base conditions in some low-water media, making reliance on pH memory unnecessary. 8.7.2

Processes Erasing pH Memory

As the picture presented above suggests, the net charge on the protein may be affected by processes leading to preferential loss of counter-ions of one charge. This can happen if counter-ions undergo proton exchange reactions with the protein to produce a neutral species. The exchange may be driven to completion if the neutral species produced is then removed from the neighborhood of the protein. Such exchanges may be relatively easy if the counter-ions are derived from weak acids or bases. If the acid or base is then volatile, the counter-ions can be lost during drying under vacuum, with changes in protein net charge, as represented by reactions such as:

8.7 Acid-Base Conditions

Protein-NH3'.-OOCH --* Protein-NH2.HOOCH --* Protein-NH2 + HOOCH (gas)

(4

Protein-COO-.NH4' -+ Protein-COOH.NH3 Protein-COOH + NH3 (gas) (3) +

A similar process can occur if the acid or base can be extracted into the bulk phase of the reaction mixture (e.g. octanoic acid or triethylamine in an organic solvent). Other counter-ions may be exchanged with the bulk non-polar phase, provided something is able to solubilize them there. This will usually be in the form of an ionpair with a species better solvated by the medium. For example, an acid with a large hydrophobic group may form a Na' salt with sufficient solubility in the bulk medium. The protonated acid will carry H' to and from the enzyme in exchange, to maintain electroneutrality. A similar process with a hydrophobic amine, for example, can transfer H' and Cl-. Solubilization of the small ion may be aided by complexation, for example of Na' by a crown ether. The exchanges can be written as:

Protein-COO-.Na' + RCOOH (bulk phase) =+ Protein-COOH + RCOO-.Na' (bulk phase)

(4)

Protein-NH3+.C1-+ R3N (bulk phase) G= Protein-NH2 + R3NH'.CI- (bulk phase) (5) Acidic or basic species in the bulk phase may protonate or deprotonate the enzyme, becoming the necessary counter-ions in the process. So we might have equilibria such as: Protein-COOH + CH3NH2 (bulk phase) =+Protein-COO-.+NH3CH3

(6)

Protein-NH2 + CH3COOH (bulk phase) =+ Protein-NH3'. -0OCCH3

(7)

The protonation state of the enzyme may be affected by acidic or basic reactants (starting materials or products). These species could act as described by either of the two sets of equilibria just presented. Acidic or basic impurities in solvents could also be significant here. 8.7.3

Systems for Acid-Base Buffering

It should be clear that there are several possible mechanisms by which the protonation state of an enzyme may be altered in low-water media. It will often be desirable to try to maintain the optimal state by controlling acid-base conditions, rather than just relying on pH memory. This can be done by the addition to the reaction system of acid-base buffers, as in aqueous media. However, the details of these buffer systems and how they work is usually somewhat different. The equilibria represented by Eqs. (4) and (5) can be employed to set up buffering based on agents dissolved in the bulk non-aqueous phase. As the equilibria indicate, the state of ionizable groups in the enzyme will depend on the ratio of buffer forms added to the bulk phase: the acid and its ion-paired salt with Na' (or another cation); the base and its ion-paired hydrochloride salt (or similar). Also in analogy to aqueous buffers, a given pair will only be usable over a given range of acidity/basicity. The

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8 Enzymic Conversions in Organic and Other Low-Water Media

conditions where optimal buffering is found (analogous to aqueous pK) will depend on the solvent used. A number of such organic soluble buffer pairs have now been identified [29-311. Identification of buffers that can be dissolved in the bulk phase is restricted by solubility, usually of the ionized form. An alternative approach is to choose buffers expected to be almost completely insoluble in the reaction medium, which will remain as suspended crystals. Convenient choices are zwitterionic solids and their salts, which will give rise to equilibria as shown in Eqs. (8)and (9). Protein-COO-.Na' + TES'- (crystals, zwitterionic)=s Protein-COOH + TES-.Na' (crystals) Protein-NH3'.Cl- + Lys' (crystals, zwitterionic)+

(8)

Protein-NH2 + Lys'.Cl- (crystals) (9) Since the buffer compounds are now present as crystalline solids, the equilibrium position is independent of the quantity of each. A given pair sets a characteristic protonation state of the enzyme. This is analogous to the use of solid salt hydrate pairs to set a hydration state. Again, to cover the range of acid-base conditions that might be appropriate for different enzymic syntheses, a series of different buffer pairs is required. A number have been identified[32-34],but the known range probably needs extending. Of course, if such equilibria are to be established, a mechanism is required for the transfer of H' and counter-ions between the solid buffers and the enzyme molecules. Quite surprisingly, this usually does not seem to be a limitation. Only quite small quantities of ions must be exchanged, which will make equilibration easier. Probably traces of acids and bases soluble in the bulk phase can catalyze the transfers by equilibria such as Eqs. (4) and (5). If rates of equilibration are inadequate, deliberate addition of such transfer agents should help. Although the analogy to aqueous acid-base behavior is clear, there are important differences. In particular, the ionization of acidic and basic groups in the protein becomes to a considerable extent independent. Both are affected by the availability of counter-ions as well as of H+, as illustrated in equilibria like those shown in Eqs. (4) and (5). Hence in principle two different buffering systems should be used to fix the state of these two categories of protein groups. In a medium that is saturated with a simple salt (typically NaCl), these two different acid-base parameters become linked in a fixed relationship. In this case, the system reverts to having only a single acidbase variable, as in water. Only limited studies have been made so far of systems in which both classes of buffer are present, so it is not possible to say how often better performance can be obtained by optimizing both. Hence for the present, I would not advise those persons looking at practical syntheses to use more than one type of buffer.

References References

E. N. Vulfson, P. J. Halling and H. L. Holland, (eds.) Methods in Biotechnology: Enzymes in Nonaqueous Solvents. Humana Press, Totowa, NJ, USA, 2001. 2 A. M. Klibanov, Nature, 2001 409, 241-246. 3 G. Carrea, S. Riva, Angau. Chem. Int. Ed. Engl., 2000 39,2226-2254. Properties and synthetic applications of enzymes in organic solvents 4 P. J. Halling, C u r . Opin. Chem. Biol. 2000, 4, 74-80. 5 A. J. Mesiano, E. J. Beckman, A. J. Russell, Chem. Rev. 1999,99,623-633. 6 Y. L Khmelnitsky, J. 0. Rich, Cum Opin. Chem. Bid. 1999, 3,47-53. 7 M. Erbeldinger, X-W. Ni and P. J. Halling, Enzyme Microb. Technol. 1998, 23, 141-148. 8 R. D. Schmid, R. Verger, Angew. Chem. Int. Ed. Engl., 1998, 37, 1609-1633. 9 K. E. Jaeger, M. T. Reetz, Trends Biotechnol. 1998, 16,396403. 10 J. A. M. de Bont, Trends Biotechnol. 1998, 16, 493-499. 11 R. Leon, P. Femandes, H. M. Pinheiro, J. M. S. Cabral, Enzyme Microb. Technol. 1998,23,483-500. 12 M. T. De Gomez-Puyou, A. Gomez-Puyou, Cnt. Rev. Biochem. Mol. Bid. 1998, 33, 53-89. 13 Y. Okahata, T. Mori, Trends Biotechnol. 1997, 15, 50-54. 14 A M . Klibanov, Trends Biotechnol. 1997, 15, 97-101. 15 R. Lortie, Biotechnol. Adv. 1997, 15, 1-15. 16 J. Bosley, Biochem. Soc. Trans. 1997,25, 174- 178. 17 A. Koskinen, A. M. Klibanov (eds) Enzymatic reactions in organic media. Chapman 6 Hall, Andover, 1995. 18 L. Greenspan, J. Res. Nat. Bur. Standards 1977,81A, 89-96. 1

Bell, A. E. M.Janssen and P. J. Halling, Enzyme Microb. Technol. 1997,20,471-477. 20 J. A. Riddick, W. B. Bunger, T. K. Sakano, Organic solvents: physical properties and methods of purification, 4th ed, Wiley, New York, 1986. 21 R. M. Stephenson,J. Chem. Eng. Data 1992, 37, 80-95. 22 L. Kvittingen, B. Sjursnes, T. Anthonsen and P. J. Halling, Tetrahedron 1992.48, 2793-2802. 23 E. Zacharis, 1. C. Omar, J. Partridge, D. A. Robb and P. J. Halling, Biotechnol. Bioeng. 1997,55,367-374. 24 P. J. Halling, Biotechnol. Tech. 1992, 6, 271-276. 25 E. Wehtje, P. Adlercreutz in: E. N. Vulfson, P. J. Halling and H. L. Holland, Eds. (2001) “Methods in Biotechnology: Enzymes in Nonaqueous Solvents”, Humana Press, Totowa, NJ, USA. 2001, pp 127-134. 26 A. Reimann, D. A. Robb and P. J. Halling, Biotechnol. Bioeng. 1994,43, 1081-1086. 27 P. J. Halling, Enzyme Microb. Technol. 1994, 16, 178-206. 28 R. H. Valivety, G. A. Johnston, C. J. Suckling and P. J. Halling, Biotech. Bioeng. 1991, 38, 1137-1143. 29 K. Xu, A. M. Kliban0v.J. Am. Chem. Soc. 1996, 118,9815-9819. 30 M. Dolman, P. J. Halling and B. D. Moore, Biotechnol. Bioeng. 1997,55, 278-282. 31 N. Harper, B. D. Moore and P. J. Halling, Tetrahedron Lett. 2000,41, 4223-4227. 32 E. Zacharis, B. D. Moore and P. J. Halling, J. Am. Chem. SOC.1997,119,12396-12397. 33 N. Harper, M. Dolman, B. D. Moore and P. J. Halling, Chem. Eur. J. 2000, 6, 1923-1929. 34 J. Partridge, P. J. Halling and B. D. Moore ‘ J. Chem. Sac. Perkin I1 2000, 465-471. 19 G.

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Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

9 Enzymatic Kinetic Resolution Jonathan M. J. Williams, RebeccaJ. Parker, and Claudia Neri

9.1

Introduction

Conventional kinetic resolution procedures often provide an effective route for the preparation of enantiomerically enriched compounds. However, a resolution of two enantiomers will only provide a maximum of 50 % yield of the enantiomericallypure material. This limitation can be overcome in a number of ways, including inversion of the stereochemistry of the unwanted enantiomer, racemization and recycling of the unwanted enantiomer or dynamic kinetic resolution. A dynamic kinetic resolution reaction involves the interconversion of the enantiomers of a starting material under conditions where one enantiomer is converted selectively into product. This principle is shown in Fig. 9-1, where a conventional kinetic resolution reaction and a dynamic kinetic resolution reaction are compared. In both cases enantiomer A reacts to form product B more quickly than enantiomer A'. However, in the conventional kinetic resolution, enantiomer A' is simply left behind as unreacted starting material. In the dynamic kinetic resolution, Aand A' are in equilibrium, which allows for the possibility that all of the starting material will be converted into product B. The reaction conditions must be chosen that whilst the starting material enantiomers @/A')undergo rapid equilibration (racemization),the product B must be inert to racemization. Dynamic kinetic resolution reactions are not limited to enzyme-catalyzed processes, and there are reviews available that consider all aspects of such Conventional Kinetic Resolution

A

A' Figure 9-1.

+

-X-

Dynamic Kinetic Resolution

0

A

B'

A'

-

B

+

Comparison of conventional and dynamic resolution reactions.

B'

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9 Enzymatic Kinetic Resolution

In addition, reviews dealing with aspects of enzyme-catalyzed dynamic resolution and related processes such as stereoinversion and deracemisation have also been published [4-71. Details of the kinetic principles of dynamic kinetic resolution reactions have also been reported17-’]. Interestingly, a dynamic kinetic resolution reaction can provide a product with higher enantiomeric excess than the corresponding kinetic resolution. In a conventional kinetic resolution, the enantiomeric excess of the product often decreases as a function of conversion. This happens because as the reaction proceeds, the proportion of the preferred enantiomer of substrate decreases. Unless the enzyme is able to discriminate perfectly between the substrate enantiomers, it will catalyze the reaction of the less preferred enantiomer of substrate (the proportion of which grows as the reaction proceeds). However, in a dynamic kinetic resolution where the substrate enantiomers are interconverting rapidly, the ratio of substrate enantiomers will be constant at 1:l. Consequently, the enantiomeric excess of the product will not decrease as the reaction proceeds. The following sections consider dynamic resolution reactions of alcohols (and their derived esters), acids (and their derived esters) as well as dynamic resolution involving reaction catalyzed by dehydrogenase enzymes.

9.2 Alcohols and their Derivatives

In order to achieve a dynamic kinetic resolution of alcohols, procedures need to be found for the in situ racemization of these substrates. The racemization conditions need to be compatible with the enzyme-catalyzedstep, and the product must be inert to racemization. The general principles are identified in Fig. 9-2, where enzyme-catalyzedacylation selectively converts one of the equilibrating alcohols into the corresponding ester. Methods for racemization of the alcohol include substrates where the R or R’ group is a good leaving group, or where temporary dehydrogenation to the corresponding ketone can be achieved, as shown in Fig. 9-3.

H,

OH

,A R’

R

Acyl donor t

enzyme

HO H Figure 9-2.

R

alcohols.

Dynamic resolution i n the acylation of

9.2 Alcohols and their Derivatives

-HX

+HX

R

Figure 9-3.

R

8,

+ HX -HX

R

Racernization o f alcohols via carbonyl compounds.

9.2.1

Cyanohydrins

Cyanohydrins are readily racemized with base, and this has been exploited by Oda and co-workers in a dynamic kinetic resolution of these substrates [lo,'*I. In a typical procedure (Fig. 9 - 4 , the cyanohydrins were formed by transhydrocyanation with acetone cyanohydrin, catalyzed by the hydroxide form of an anion exchange resin (Amberlite IRA-904). The reversible nature of the cyanohydrin formation allows racemization to proceed during the course of the enzyme-catalyzed acetylation, and the choice of isopropenyl acetate as the acyl donor means that the only by-product is acetone. The immobilized lipase from Pseudomonas cepacia (Amano) afforded good enantioselectivities for the formation of a range of cyanohydrin acetates derived from aromatic aldehydes (Fig. 9-5). Polymer-supportedquinidine could also be employed

OH

w,

R

RACN

1

R

CN

(a) Acetone cyanohydrin, IRA-904 resin (HO-form) (b) Pseudomonas cepacia lipase, isopropenyl acetate 3A molecular sieves, i-Pr20, 40 "C,3-6 days Figure 94.

Racernization of cyanohydrins with in situ acylation.

I

289

290

I

9 Enzymatic Kinetic Resolution

OAc

W

C

OAc

N

\

/ O " C N CI

84% ee 96% yield

Figure 9-5.

<

r

C

N

\ 84% ee 83% yield

OAc

47% ee 57% yield

OAc

91% ee 81% yield

OAc

85% ee 88% yield

OAc

89% ee 80% yield

Examples o f cyanohydrin acetates formed by dynamic resolution.

as the base for racemization and formation of cyanhydrins, although the reactions were generally slower than with the Amberlite resin [I2]. 9.2.2 Other Readily Racemized Substrates

Kellogg, Feringa and co-workers have achieved successful dynamic kinetic resolution reactions using cyclic hemiacetals as substrates 1' 3, 141. The enzyme-catalyzed acetylation of 6-hydroxypyranone shown in Fig. 9-6 has been achieved with reasonable enantioselectivity with essentially complete conversion. The racemisation of the hemiacetal is presumed to proceed via reversible ring-opening of the pyran~ne['~I. The rate of reaction was found to greatly increase when the enzyme, lipase PS (Pseudomonas sp.) was immobilized on Hyflo Super Cell (HSC).

IIW.

Figure 9-6.

OH

Dynamic resolution o f cyclic hemiacetals.

9.2 Alcohols and their Derivatives

I

291

PS-HSC

0 O cyclohexanel butyl acetate (1:l) 18 h, r.t.

O

0Ac

100% conversion 89% ee

CAL (immobilised)

0

0a I

COMe

Figure 9-7.

0Ac ?J

cyclohexane 18 h, 69 "C

I

COMe 100% conversion >99% ee

Examples o f dynamic resolution o f furanone and pyrrolinones.

(a) HSR', SiOp (b) fseudornonas fluorescenslipase, t-BuOMe/vinyl acetate (3:1), 30 "C,4-11 days Figure 9-8.

Dynamic resolution o f hernithioacetals.

The related dynamic resolutions of the furanone and pyrrolinone substrates were achieved with higher selectivities (Fig. 9-7)[I4]. Again, these substrates underwent spontaneous racemization under the reaction conditions. Appropriate choice of enzyme afforded a good example of an essentially perfect dynamic kinetic resolution process in the case of the esterification of the hydroxypyrrolinone substrate. Rayner and co-workers have demonstrated that hemithioacetals can be racemized on exposure to sili~a['~1. In a typical experiment, an aldehyde and a thiol are combined to give a hemithioacetal. In the presence of silica, the enzyme-catalyzed acetylation proceeds under dynamic resolution conditions, as shown in Fig. 9-8.

292

I

9 Enzymatic Kinetic Resolution

OAc

Meo & ,

M e O TS-0SiEt3

SBu

0

0 63% yield >95% ee

83% yield 90% ee

OAc AcO

OAc

SCH(Me)2

A c O A -OSiEt3 S

65% yield >95% ee Figure 9-9.

Examples of acetylated hemithioacetals.

73% yield >SO% ee

0OH

+ H

A+-&H

75% yield 0OAc 97% ee

\ rneso intermediate

HO (a) Et3N, THF (b) lipase PS, vinyl acetate, 48 h, r.t. Figure 9-10.

Dynamic resolution involving a m e w intermediate

Representative products obtained using this procedure are given in Fig. 9-9. The acetylated products are inert to racemization under the reaction conditions. An interesting example of dynamic kinetic resolution of an alcohol has been reported by Taniguchi and Ogasawara~"].The a-hydroxy ketone in Fig. 9-10 under-

9.2 Alcohols and their Derivatives

goes racemization via a rneso enediol intermediate, and one of the enantiomers can be acetylated selectively with lipase PS (Pseudornonas sp. immobilized on Celite, Amano). The added triethylamine was required in order for the racemization to take place. Without triethylamine, the reaction proceeded under conventional kinetic resolution conditions. 9.2.3

Enzyme and Metal Combinations

Most substrates for enzyme-catalyzed kinetic resolution reactions do not undergo spontaneous racemization under conditions that are suitable for enzyme activity. One solution to this problem has been to design mild transition metal-catalyzed methods for in situ racemization[”]. In order to achieve this goal, the racemisation method must be able to function without an adverse effect on the enzyme. Additionally, the enzyme must not inhibit the racemization method. The first example of the use of enzyme and metal combinations to provide a dynamic resolution procedure was reported by Allen and Williams in 1996[”]. In this case, a palladium (11) catalyst was employed that was able to racemize the allylic acetate substrate, but did not erode the enantioselectivity of the product allylic alcohol (Fig. 9-11). For example, a cyclic acetate was shown to undergo a simple kinetic resolution, affording enantiomericallyenriched starting material and product at approximately 50 % conversion. However, performing the reaction in the presence of a palladium (11) catalyst facilitated a dynamic resolution by continuously racemizing the starting material as the reaction progressed. Similar methodology was applied to an acyclic allylic acetate by the group of Kim, who used Pd(0) catalysts[I9]. Acyclic allylic acetates are easier substrates for palladium-catalyzed racemization, and these workers were able to effect a dynamic resolution strategy within a more acceptable time scale (Fig. 9-11). The in situ racemization with palladium catalysts is limited in scope, since allylic acetates are required as substrates. In addition, not all allylic acetates are expected to undergo facile racemization .I‘’[ An alternative enzyme/transition metal combination employs transfer hydrogenation catalysts that are capable of racemizing secondary alcohols. The racemization procedure temporarily converts the alcohol into an achiral ketone, which is reduced back to the racemic alcohol. Coupling this racemization procedure to an enzyme-catalyzedacylation reaction affords a dynamic resolution process (Fig. 9-12). Several enzyme/transition metal combinations have been shown to be effective for these reactions, although ruthenium complexes 1-3appear to be especially effective for the in situ racemization of the alcohol. The product esters are not prone to racemization under the reaction conditions. Early results employing transfer hydrogenation catalysts to effect the racemization of alcohols required the use of added ketoner’l, 22]. However, it was subsequently shown that added ketone was not required when appropriate transition metal complexes were used as catalysts. Furthermore, the use of 4-chlorophenylacetate as the acyl donor afforded improved results.

293

I

294

I

9 Enzymatic Kinetic Resolution

enzyme

OAc

1

*

H20

OH

Pd(0) or Pd(1l)

,PR OAc

Ph

Pseudomonas fluorescens lipase 37-40 "C, pH 7.0, 19 days

I

Ph

w

U

0.1M phosphate buffer 5 mol% PdC12(MeCN)2 81% yield 96% ee

OAc

kPh

Candida antarctica lipase i-PrOHTTHF 16-18"C, 24 h

* then 5 mol% Pd(PPh& with 15 mol% dppf, 36 h

Ph 83% yield (HPLC) 98% ee

dppf = 1,l '-bis(dipheny1phosphino)ferrocene Figure 9-11.

Dynamic resolution using transition metal/enzyme combinations.

Backvall and co-workers have reported successful results for a wide range of substrates, some of which are identified in Table 9-1. The procedure works well for secondary alcohols containing aryl and alkyl groups [231, diols [241 and a-hydroxy esters 12'1. Although catalyst 1 requires no additional base, Kim, Park and co-workers used triethylamine to facilitate racemization using catalyst 2, Table 9-2 [ 2 6 ] . In their case, small quantities of oxygen were added to initiate the racemization procedure. In the case of allylic alcohols, careful choice of racemisation catalyst is required in order to minimize the amount of conversion of the substrate into saturated or

9.2 Alcohols and their Derivatives

vinyl acetatel CH2C12 (3:l)

* 1 equiv. PhCOMe 2 mol% Rh2(0Ac)4 6 mol% o-phenanthroline

PhM 'e

PhAMe

Pseudomonas fluorescens lipase 72 h, 20 "C

1

R

OAc

R'

OAc Ru catalyst 1 - 3 enzyme (see Figs 14 and 15)

Figure 9-12. Transition metal-catalyzed racemization of alcohols coupled with enantioselective enzyme-catalyzed acetylation.

unsaturated ketones. For allylic alcohols, Kim, Park and co-workers have used catalyst 3, which minimizes the formation of undesirable side products LZ71. In addition to the use of enzyme and transition metal combinations for the dynamic resolution of alcohols, there has been a brief report of the use of amines as substrates. In 1996, Reetz and Schimossek reported the combination of palladium on carbon with an immobilized lipase (from Candida antarctica) in the dynamic

I

295

9 Enzymatic Kinetic Resolution Tableg-1.

Examples of dynamic resolution of secondary alcohols with catalyst 1.

Substrate

Product

OAc

OH I

PhAMe

PhAMe

OH

m

OAc

\

OAc

PhOJ

Me

P

h

O

4

Me

C 6 b 3M e '

qH

OAc

OH

OH

PhAC02Me

OAc Ph n C 0 2 M e

Conditions

Yield ("h) ee ("A)

2 mol% Ru cat 1 Novozym 435 3 equiv pCIC6H40Ac toluene 70 "C, 46 h

80

> 99

2 mol% Ru cat 1 Novozym 435 3 equiv pCIC6H40Ac toluene 70 "C, 48 h

77

> 99

2 mol% Ru cat 1 Novozym 435 3 equiv pCIC6H40Ac toluene 70 "C, 46 h

88

> 99

2 mol% Ru cat 1 Novozym 435 3 equiv pCIC6H40Ac toluene 70 "C, 24 h

80

> 97

2 mol% Ru cat 1 Novozym 435 3 equiv pCIC6H40AC toluene 70 "C, 24 h

77

> 99

2 mol% Ru cat 1

(98:2 R,R/meso)

80

94

80

98

PS-C (type 11) 2 equiv pCIC6H40Ac cyclohexane 60 "C. 48 h 2 mol% Ru cat 1 PS-C (type 11) 2 equiv pCIC6H40Ac cyclohexane 60 "C, 48 h

Novozym 435 is Candida antarctua lipase B (Nova Nordisk A/S) PS-C (type 11) from Amano is Pseudornonas cepacia lipase

9.3 Carboxylic Acids and their Derivatives

I

297

Table 9-2.

Examples of dynamic resolution o f secondary alcohols with catalysts 2 and 3.

Substrate

Product

OAc PhAMe

PhM 'e

Yield

5 mol% Ru cat 2 5 mol% 0 2 PS-C (type 11) 3 equiv Et3N 3 equiv pCIC6H40Ac CHzClz, 60 "C, 43 h

85

96

5 mol% Ru cat 2 5 mol% 0 2 PS-C (type 11) 3 equiv EtjN 3 equiv pCIC6H40Ac CHzC12,60 "C, 4 3 h

98

99

84

> 99

4 mol% Ru cat 3

OAc

OH I

-

P

h

w Me

OAc

("A) ee (%)

Conditions

PS-C (type 11) 1 equiv Et3N 1.6 equiv pCIC6H40Ac CH2C12, r. t., 48 h

4 mol% Ru cat 3 90 PS-C (type 11) 1 equiv EtJN 1.6 equiv pCIC6H40Ac

95

CHzC12, r. t., 48 h

PS-C (type 11) from Amano is Pseudomonas cepacia lipase

resolution of phenethylamine[281.The N-acylated product was obtained with 99 % ee and with 75-77 % yield.

9.3 Carboxylic Acids and their Derivatives 9.3.1 Readily Enolized Carboxylic Acid Derivatives

Carboxylic acid derivatives that have a-substituents can exist as chiral compounds. The resolution of the enantiomers of such compounds is a useful process, leading to the preparation of a-amino acids, a-hydroxy acids and other a-substituted carboxylic acids and their derivatives in enantiomerically enriched form. In addition, the racemization of such compounds can be achieved by a deprotonation/reprotonation sequence, as shown in Fig. 9-13. The ease with which racemization of the carboxylic acid derivative occurs depends on the nature of the substrate. Carboxylic acids themselves are slow to racemize, since the carboxylic acid is initially deprotonated to form a carboxylate anion.

298

I

9 Enzymatic Kinetic Resolution

achiral enolate Figure 9-13.

Racemization o f a-substituted carboxylic acid derivatives by enolization.

S.griseus protease carbonate buffer (PH 9.7) 24 h, 22 "C Ketorolac 100% conversion 76% ee Figure 9-14.

Dynamic resolution i n the preparation of Ketorolac.

Subsequent deprotonation to afford the carboxylic acid enolate requires the formation of a doubly deprotonated species, which is disfavored relative to the formation of an ester enolate. In fact, activated esters such as phenyl estersI2'1 or thioesters[30]are especially prone to racemization, since enolization is easier than for simple esters. Fulling and Sih reported one of the earliest examples to exploit racemization of carboxylic acid derivatives in order to achieve a dynamic kinetic resolution[31].The anti-inflammatory drug Ketorolac was prepared by hydrolysis of the corresponding ester. Whilst most lipases afforded the undesired enantiomer preferentially, a protease from Streptornyces griseus afforded the required (S)-enantiomerof product with good selectivity. The substrate was particularly prone to racemization since the intermediate enolate is well stabilized by resonance effects, although a pH 9.7 buffer was required to achieve a useful dynamic resolution reaction. Thus the acid was formed with complete conversion and with 76 % enantiomeric excess. Drueckhammer and co-workers have published details of a successful strategy for dynamic resolution in the hydrolysis of suitable thioesters L30, 321. Trioctylamine was employed as the racemizing agent, which was effective for the racemization of a series of a-substituted thiopropionates. Specific examples include the hydrolysis of an ethylthioester using Pseudomonas cepacia lipase, the transesterification of an aaryloxy trifluoroethylthioester with butanol and PS-30, as well as hydrolysis of a trifluoroethylthioester using Subtilisin Carlsberg (Fig. 9-15). The ability to achieve dynamic kinetic resolution using thioester substrates has been recognized by other workers, and reports of dynamic resolution strategies

9.3 Carboxylic Acids and their Derivatives

PCL 0.5 equiv Oct3N

b

toluene, H20 65 h >99%conversion 96.3%ee

0

0

Subtilisin Carlsberg 0.5 equiv Oct3N *

97%conversion 83%ee

SCH2CF3

toluene, BuOH

Me Ar = 2,4-dichlorophenyl Figure 9-15.

lipase (PS-30) Et3N

OBu Me 98%conversion 75% ee

Dynamic resolution i n the hydrolysis/transesterification of thioesters.

leading to the anti-inflammatory drugs Napr~xen[~’] and S u p r ~ f e n l ~have ~ I been published. Trioctylamine is again used as the racemizing agent, as shown in Fig. 916. In addition to in situ racemization of a-substituted carboxylic acid derivatives by deprotonation/reprotonation, a procedure involving halide exchange has been developed[35,3G1. Whilst the a-halo esters undergo racemization at a reasonable rate, the corresponding carboxylates are almost inert to racemization under the reaction conditions. Using immobilized phosphonium halide and CLEC (cross-linked enzyme crystals),a dynamic resolution procedure has been developed for the hydrolysis of a-homo and a-chloro esters (Fig. 9-17).The enantiomeric excess in each case was similar to that achieved for simple kinetic resolution reactions using the same enzyme/substrate combinations. Nitriles can be hydrolyzed by various microorganisms, affording the corresponding carboxylic acids. A method has been reported for the hydrolysis of racemic mandelonitrile (PhCH(0H)CN) into (R)-mandelic acid using Alcaligenes faecalis

I

299

300

I

9 Enzymatic Kinetic Resolution

Candida rugosa lipase

Me

45 "C, 294h

OH Me Naproxen 70% conversion 92% ee

Candida rugosa lipase

*SCH2CFs

Oct3N isooctane Me

45 "C,20h

OH Me Suprofen 100% conversion 95% ee Figure 9-16.

Dynamic resolution in the synthesis o f Naproxen and Suprofen.

Br

$r

Polyrner-PPh3+ Brt

P h A 0 2 M e

CI

Polyrner-PPh3+ CI-

PhAC02Me Figure 9-17.

Candida rugosa lipase (CLEC)

H20, pH 7,4.5 h

Candida cylidfacea lipase (CLE;) H20, pH 7,24h

PhAC02H 80% conversion 79% ee

CI

phAco2~ 90% conversion 90% ee

Racemization of a-haloesters by halide exchange coupled with enzymatic hydrolysis.

9.3 Carboxylic Acids and their Derivatives Figure 9-18. Racemization o f a-aminoesters catalyzed by imine formation.

NH2

SLOW

NH2

7

RAC02Me

RAC02Me

R'

R' I

-

FAST-

RnC02Me

ATCC 8750. An isolated yield of 94% of the enantiomericallypure mandelic acid was obtained, indicating that a dynamic resolution process is occurring. 9.3.2 Amino-Esters and Related Compounds

Typical a-amino esters only undergo racemization slowly, but methods for accelerating this process have been devised[37.381. Temporary conversion of the amine to an imine lowers the pK, of the substrate, such that racemization becomes faster. A series of a-aminoesters has been hydrolyzed to a-amino acids using alcalase in the presence of pyridoxal S-pho~phate[~~]. During the course of these reactions, the amino acids precipitated from the reaction mixture, thereby protecting them from racemisation. The method was used to prepare enantiomerically enriched phenylalanine, leucine, tryptophan and norvaline with high selectivity (Fig. 9-19). A related ammonolysis of an amino ester has been reported using either pyridoxal or salicylaldehydeas the racemizing agent I4l1. The amino ester undergoes racemization more quickly than the amino amide, and an effective dynamic resolution could be achieved at -20 "C. Re-formed imino-esters have also been used as substrates for dynamic kinetic resolution reactions [421. The free amino acid precipitated from the reaction mixture as the reaction proceeded. a-Azido amides have been subjected to kinetic resolution reactions using whole cells of E. coli DHSa/pTrpLAP,affording hydrolysis to the corresponding acids[43].In the case of 2-azidophenylaceticacid amide, the substrate racemized in situ, and the acid product could be obtained with 98% ee at over 50% conversion.

I

301

302

1

9 Enzymatic Kinetic Resolution

Alcalase

y 2

20 mot% pyridoxal phosphate i-PrOHIH20(19:1), pH 8.5 t-BuOH/t-BuOMe (7:3) 40 "C, 3-4 h R

RnCO2yield

PhCH2 (CH3)2CHCH2CH2 (3-indolyl)CH2 CH3CH2CH2

NH

Ph

Novozym 435 (CAL-B) * pyridoxal NH3

t-BuOH/t-BuOMe (7:3) -20 "C, 66 h

N4 PhCH2A

Figure 9-19.

C02Et

chyrnotiypsin

*

H20/MeCN(1:19) 10 mol% DABCO added after 48h. Then 48 h at r.t.

92% 87% 95% 87%

ee 98% 93%

97%

91%

NH2

PhACONH2 85% conversion 88% ee

y42

PhCHzAC02H 87.5% yield

90% ee

Dynamic resolution of amino acids via imine formation.

9.3.3

Reactions of cyclic amino acid derivatives

There are several cyclic amino acids derivatives that are prone to racemization and have been used as substrates for dynamic kinetic resolution reactions. Oxazolinones were first used as substrates €or enzyme-catalyzed hydrolysis over 30years It was noted that spontaneous hydrolysis could be quite high, depending on the amino acid derivative being used and the pH of the reaction medium [451. Bevinakatti and co-workers demonstrated that oxazolinones could undergo racemization during a lipase-catalyzed enantioselective ring-opening with n - b u t a n ~ l [471.~ ~At, 100% conversion, they were able to obtain (S)-butylN-benzoylalaninate with 34% ee. This concept has been developed by the research groups of Sih and Turner. The oxazolinone derived from phenylalanine was subjected to lipase-catalyzed hydrolysis with ten lipases 14'1. Whilst several lipases gave good enantioselectivities, the lipase

9.3 Carboxylic Acids and their Derivatives

/

PL(Ferm1ipase) Phosphate buffer (PH 7.6)

I

303

phcH2H HN

OH

)C.

Ph

100% conversion 99% ee

r/c”

NQ

I

\

Ph

PhCH2

\

Lipase AP (from Aspergillus Niger) Phosphate buffer (pH 7.6), 17 h

HN

OH

Po \

Ph

100% conversion 99% ee Figure 9-20.

Dynamic resolution in the hydrolysis of oxazolinones.

phcH2H HN

OH

Ph>o

PhCH2Yfo P30

NYo Ph

PhCH2Ho

Lipase * 5 equiv. MeOH t-BuOMe 50°C,48h

OMe

HN

Prozyme 6

82% yield 295% ee

*

)=’

Ph

99% yield 65% ee

Po

Ph

15% yield

>95% ee Figure 9-21.

Two stage hydrolysis of oxazolinones.

from Aspergillus niger (AP) and porcine pancreatic lipase (PL Fermlipase) provided particularly good enantioselectivities,with an opposite sense of asymmetric induction from each other (Fig. 9-20). An additional strategy employed by Sih and co-workers involved sequential enzyme-catalyzedreactions. Pseudomonas lipases were found to tolerate a wide range of substrates although the enantioselectivity was generally only moderate. However, by first performing a methanolysis of the oxazolinone followed by a separate enzyme-catalyzed hydrolysis under kinetic resolution conditions, a highly enantiomerically enriched product could be obtained, as shown in Fig. 9-21.1’4

304

I

9 Enzymatic Kinetic Resolution

"w"

NYs Ph

Figure 9-22.

Prozyme 6

HN

phosphate buffer 10% MeCN pH 7.5 25 "C, 7-88 h

OH

Ph>s

R

yield

ee

Me

67%

90%(NoMeCN)

(CH&CHCH2 78% 94% CH3SCH2CH2 86% CH~CHZCH~CH~ H2NCOCH2 98%

97% 98% 99% 57% (No MeCN)

Dynamic resolution in the hydrolysis ofthiazolinones.

Lipozyme

NYo Ph

""w" NYo Ph

*

'""?--fO HN

OBu

25 mol% Et3N 2 equiv. BuOH toluene, 30 "C, 5 days ph>o 94% yield 99.5% ee

Novozyme (CAL-B) 25 mol% Et3N 2 equiv. BuOH toluene, 37 "C

HN

OBu

Po

Ph

79% yield 94% ee

i-prHo i-prxo Novozyme (CAL-B) c

2 equiv. MeOH MeCN, 37 "C

Ph Figure 9-23.

OMe

HN

Ph'

k o 83% yield 97% ee

Dynamic resolution i n the alcoholysis o f oxazolinones.

9.3 Carboxylic Acids and their Derivatives

I

305

Pseudomonas sp. AJ-11220 (whole cells) c

HN0 KNH

phosphate buffer (PH a), 30 h

H2N

OH

94% yield >99% ee

Agrobacterium radiobacter pH 8.4,40 "C, 48 h

79% yield

>92% ee

Agrobacterium radiobacter pH 8.4,40 "C, 48 h

71Yoyield >96% ee Figure 9-24.

Dynamic resolution in the hydrolysis of hydantoins.

Sih and co-workers also reported the dynamic resolution of a range of thiazolinones by enantioselective hydrolysis using proteases [491. In these cases, the product is the corresponding thioamide. Some of the higher enantiomeric excesses reported are identified in Fig. 9-22. Turner and co-workers identified conditions appropriate for the dynamic resolution of a 4-tert-butylsubstituted 0xazolinone[~~1. The ring-opened butyl ester could be obtained with high yield (94%) and enantiomeric excess (99.5%) using Lipozyme Mucor miehei and 0.25 equivalents of triethylarnine. Subsequent cleavage of the ester and amide groups afforded a route to enantiornerically pure (S)-tert-leucine.Whilst Lipozyme provided high selectivities for the sterically demanding tert-butyl group, Turner reported that Candida antarctica lipase B (Novozyme) was preferred for smaller groups, as shown in Fig. 9-23[511.

306

I

9 Enzymatic Kinetic Resolution

-

hydantoinase

""K""

buffer pH 8.5,50 "C

0

Figure 9-25.

HN

OH

0b N H 2

Dynamic resolution o f racemic hydantoins.

The other major class of cyclic amino acid derivative used in dynamic resolution reactions is the hydantoin group. Like oxazolinones, hydantoins readily undergo racemisation under mild conditions. Systems involving a two step procedure using D-hydantoinase and a carbamoylase were reported to provide a route to D-amino acids l S 2 . 531. Dynamic resolution of a p-hydroxyphenyl substituted hydantoin was reported in 1987[54].Using the intact cells of Pseudomonas sp. AJ-11220,the amino acid was prepared in over 90% yield, as shown in Fig. 9-24. This hydrolytic procedure leads directly to the amino acid, and the same enantiomer of product, the D-amino acid, was obtained independently of the stereochemistry of the substrate. A similar strategy has been used in the hydrolysis of hydantoins with pendant ureido groups, using the bacterial culture Agrobucterium r u d i o b ~ c t e r [ ~ ~ ] . D-Hydantoinaseshave also been isolated from thermophilic micro-organisms, and applied to the dynamic resolution of racemic hydantoins, where the isolated products are the N-carbamoyl D-amino acids (Fig. 9-25)[561. Subsequent transformation into D-amino acids could be achieved chemically or enzymatically. Representative examples using commercially available hydantoinases D-HYD-1 and D-HYD-2 are shown in Table 9-3. Variously ring-substituted D-phenylglycine derivatives have also been prepared by hydantoin hydrolysis using D-HYD-1and D-HYD-2,affording the amino acid with excellent levels of enantioselectivity and good yields LS71.

Table 9-3.

Dynamic resolution using hydantoinase enzymes.

R

Enzyme

Carbamoylateyield (%)

Me

D-HYD-1 D-HYD-2

71 73

94 34

PhCH2

D-HYD-1 D-HYD-2

67 12

> 99

i-Pr

D-HYD-1 D-HYD-2

66 71

> 99 > 99

(D-valine)

MeSchzCCHz

D-HYD-1 D-HYD-2

81

> 99 > 99

(D-methionine)

75

D-HYD 1 D-HYD-2

95 90

> 99

Ph

Amino acid

ee ("7)

(D-alanine) (D-phenylalanine)

> 99

96

(D-phenylglycine)

9.4 Reduction ofa-Ketoesten

Baker's yeast

OEt

spontaneous racemisation

Figure 9-26.

69% yield

Dynamic resolution in the reduction o f P-ketoesters.

9.4 Reduction of fi-Ketoesters

The reduction of ketones into alcohols can be achieved using biocatalytic methods. Amongst the most popular of the available methods is the use of Baker's yeast, BY (Saccharomyces cerevisiae). The use of P-ketoesters as substrates leads to the corresponding P-hydroxy esters, often with high enantioselectivity. In the particular case of a-substituted p-ketoesters, the substrates spontaneously racemize, and this provides the basis for many reports of dynamic resolution reactions, some of which are described in the following discussion. In 1976, Deol and co-workers showed that cycloalkyl fbketoesters could be reduced under dynamic resolution conditions (Fig. 9-26)1('. In fact, many microorganisms are able to achieve similar reductions on the same and related substrates. Azerad and co-workers have achieved higher selectivities using other microorganisms including Geotrichum candidum, Mucor racemosus, Kloeckera magna and Mucor circinelloides[5q-611.The opposite diastereomer of product (1S,2S instead of 1S,2R) was obtained using Penicillium chrysogenum and Colletotrichum gloeosporoides as the microorganism. A range of cyclic (3-hydroxyestershas been prepared, some ofwhich are identified in Fig. 9-27. The use of a P-ketothioester as a substrate has been reported to afford better stereoselectivity["I. Heterocyclic (3-ketoesters have also been used as substrates for reduction, where the products often have use in the synthesis of pharmaceutical agents or natural products. Representative examples of heterocyclic 0-hydroxyesters formed using Baker's yeast are given in Fig. 9-28[65-711. Acyclic (3-ketoestersare generally less predictable as substrates than their cyclic counterparts, with the selectivity depending on the nature of the groups attached to the dicarbonyl moiety (Fig. 9-29). Representative examples of acyclic p-hydroxyesters obtained by dynamic resolu-

I

307

308

I

9 Enzymatic Kinetic Resolution

OH

ii

Kloeckera magna [60]

Baker's yeast [62] X= OMe 93% cis 94% ee

X= SEt

OH

Mucor griseocyanus [60]

100% cis 99% ee

100% cis >96% ee

oo2 88% trans 95% ee

a OH

OH

OH

02Et

Beauveria bassiana [60]

Mucor griseocyanus [60]

100% cis 98% ee

?

?

U

Figure 9-27.

100% trans 88% ee

Baker's yeast [61] >98% cis 72% ee

Baker's yeast [63,64] 100% cis 98% ee Cyclic (3-hydroxyestersobtained by dynamic resolution.

tion are provided in Fig. 9-30, where Baker's yeast, as well as other microorganisms, have been employed in the reduction process [72-801. Improved stereocontrol has been obtained using recombinant E. coli strains expressing Gre3p or Gcylp (from Baker's yeast). Since fewer competing enzymes are present in the recombinant E. coli, the enantioselectivity and diastereoselectivity are found to be better than using Baker's yeast itself as shown in Fig. 9-31181].

I

9.5 Conclusion 309

OH C02Et

100% cis [65,66]

100% cis [651

85% ee

>85% ee

63% cis (671 100% ee

Boc

Boc

0 >99% cis [70] >93% ee

100% cis [68,69]

>90% ee Figure 9-28.

98% ee [71]

Heterocyclic 0-hydroxyesters obtained by dynamic resolution.

0

OH microorganism*

RV

O R'

Figure 9-29.

R

"

R

dOR''

E

R'

Dynamic resolution o f acyclic fl-ketoesters.

9.5 Conclusion

In summary, dynamic resolution strategies employing biocatalytic methods provide a useful synthetic route to a range of enantiomerically enriched building blocks. Over the last few years there has been a growing interest in finding new methods for the racemization of the starting material. The challenge is to discover racemization methods that are compatible with the biotransformation. Nevertheless, substrates that spontaneously racemize, such as (3-ketoesters,still provide the most practicable starting materials for biocatalytic dynamic resolution reactions.

310

I

9 Enzymatic Kinetic Resolution

OH

0

Me Baker'syeast [72] 73% de 97% ee

OH Me

Rhodotorula glutinis [73] 88% de 97% ee

Baker's yeast [74,75] 98% de 100% ee

S =

Me Baker's yeast [76]

Baker's yeast [77]

92% de >96% ee

79% ee

98% de 97% ee

96% de 91Yoee

Cl

Me Candida albicans [79]I

Geotrichurn candidurn [78]

Rhodotorula glutinis [80] 90% de 95% ee

Figure 9-30. Acyclic (3-hydroxyestersobtained

by dynamic resolution.

References 1

R. Noyori, M.Tokunaga, M. Kitamura, Bull.

Chem. Soc. Jpn., 1995,68, 36. 2 R. S. Ward, Tetrahedron: Asymmetry,1995, 6, 1475- 1490. 3 S. Caddick, K. Jenkins, Chem. Soc. Rev., 1996,447-456. 4 M.T. El Gihani, J. M. J. Williams, Cur. Opin. Chem. Biol., 1999, 3, 11-15. 5 R. J. Parker, J. M. J. Williams, Recent Res. Devel. Org. Bioorg. Chem., 1999, 3,47-64. 6 U. T. Strauss, U. Felfer, K. Faber, Tetrahedron: Asymmetry,1999, 10, 107-117. 7 H. Stecher, K. Faber, Synthesis,1997, 1-16. 8 M. Kitamura, M. Tokunaga, R. Noyori, Tetrahedron, 1993,49, 1853. 9 M. Kitamura, M Tokunaga, R. Noyori,J . Am. Chem. Soc., 1993, 115,144.

10 M.Inagaki, J. Hiratake, T. Nishioka, J. Oda, J . Am. Chem. Soc., 1991, 113,9360-9361. 1 1 M.Inagaki, J. Hiratake, T. Nishioka, J. Oda,

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12 M. Inagaki, A. Hatanaka, M. Mimura, J. Hiratake, Y. Nishioka, J. Oda, Bull. Chem. Soc. Jpn., 1992,65, 111-120. 13 M. van den Heuvel, A. D. Cuiper, H. van

der Deen, R. M. Kellogg, B. L. Feringa, Tetrahedron Lett., 1997, 38, 1655-1658. 14 H.van der Deen, A. D. Cuiper, R. P. Hof, A. van Oeveren, B. L. Feringa, R. M. Kellogg,J . Am. Chem. Soc., 1996, 118,3801-3803. 15 S. Brand, M.F. Jones,C. M. Rayner, Tetrahedron Lett., 1995, 36, 8493-8496. 16 T. Taniguchi, K. Ogasawara, Chem. Commun., 1997,1399-1400.

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References 311

E.coli

0

OH

recombinant *

Me Me

GCYlP Gre3p

MeY

O

E

t

>98% ee

>98% ee

>98% de

>98% de

recombinant E.coli *

Me GCY 1P Gre3p Figure 9-31.

>98% ee

>98% de

>98% de

Dynamic resolution using recombinant E. coli.

R. Stiirmer, A n g m . Chem. Int. Ed. Engf. 1997,36,1173-1174. 18 J. V. Allen, J. M. J. Williams, Tetrahedron Lett., 1996, 37, 1859-1862. 19 Y. K. Choi, J . H. Suh, D. Lee, I. T. Lim, J. Y. Jung, M-J. Kim, J. Org. Chem. 1999.64, 8423-8424. 20 C. G. Frost, J. Howarth, J. M. J. Williams, Tetrahedron: Asymmetry, 1992, 3, 1089-1122. 21 P. M. Dinh, J. A. Howarth, A. R. Hudnott, J. M. J. Williams, W. Hams, Tetrahedron Lett., 1996, 37,7623-7626. 22 A. L. E. Larsson, B. A. Persson, J-E Backvall, Angew. Chem. rnt. Ed. Engf. 1997,36, 1211-1212. 23 B. A. Persson, A. L. E. Larsson, M. L. Ray, JE. Backval1,J. Am. Chem. SOC.1999,121, 1645-1650. 24 B. A. Persson, F. F. Huerta, J-E. Backvall, J. Org. Chem., 1999,64,5327-5240. 25 F. F. Huerta, Y. R. S. Laxmi, J-E.Backvall, Org. Lett., 2000, 2, 1037-1040. 26 J. H. Koh, H. M. Jung, M-J. Kim, J. Park, Tetrahedron Lett., 1999, 40, 6281-6284. 27 D. Lee, E. A. Huh, M-J. Kim, H. M. Jung, J. H. Koh, J. Park, Org. Lett., 2000, 2, 2377-2379. 17

>98% ee

M. J. Reetz, K. Schimossek, Chimia, 1996, 50,668-669. 29 P. H. Dinh, J. M. J. Williams, W. Hams Tetrahedron Lett., 1999, 40, 749-752. 30 D. S. Tan, M. M. Gunter, D. G. Drueckhammer, J . Am.Chem. SOC.,1995, 117, 9093-9094. 31 G. Fulling, C. J. Sih,]. Am. SOC.1987, 109, 2845-2846. 32 P-J. Um, D. G. Drueckhammer,J. Am. Chem. SOC.,1998,120, 5605-5609. 33 C-S. Chang, S-W. Tsai, J. Kuo, Biotech. Bioeng., 1999,64, 120-126. 34 C-N. Lin, S-W. Tsai, Biotech. Bioeng., 2000, 69, 31-38. 35 M. M . Jones, J. M. J. Williams, Chem. Commun., 1998,2519-2520. 36 L. Haughton, J . M. J. Williams, Synthesis, 2001,943-946. 37 K.Yamamoto, K. Oishi, I. Fujimatsu, K-I. Komatsu, Appf. Environ. Microb., 1991, 3028-3032. 38 T. Fukumura, Agric. B i d . Chem., 1977, 41, 1327-1330. 39 M. Pugniere, A. Commeyras, A. Previero, Biotechnol Lett., 1983, 5, 447-452. 28

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40 S-T. Chen, W-H. Huang, K-T. Wang,]. Org.

Chem., 1994,59,7580-7581. 41 M. A. Wegman, M. A. P. J. Hacking, J. Rops, P. Pereira, F. van Rantwijk, R. A. Sheldon, Tetrahedron: Asymmetry,1999, 10, 1739-1750. 42 V. S. Parmar, A. Singh, K. S. Bisht, N. Kumar, Y.N. Belokon, K. A. Kochetkov, N. S. Ikonnikov, S. A. Orlova, V. I. Tararov, T. F. Saveleva,J . Org. Chem., 1996, 61, 1223-1227. 43 C. W. Tornee, T. Sonke, 1. Maes, H. E. Schoemaker, M. Medal, Tetrahedron: Asymmetry, 2000,11, 1239-1248. 44 J. de Jersey, B. Zerner, Biochem., 1969,8, 1967. 45 M. Pugniere, F. Kraicsovits, M-A. ColettiPreviero, A. Priviero, Biotech. Lett., 1985,7, 641. 46 H.S. Bevinakatti, R. V. Newadkar, A. A. Banerji,]. Chem. Soc. Chem. Commun., 1990,1091-1092. 47 H. S. Bevinakatti,A. A. Banerji, R. V. Newadkar, A. A. Mokashi, Tetrahedron: Asymmetry, 1992, 3, 1505. 48 R-L. Gu, I-S Lee, C. J. Sih, Tetrahedron Lett., 1992,33,1953-1956. 49 J.2. Crich, R. Brieva, P. Marquart, R-L. Gu, S. Flemming, C. J. Sih, j . Org. Chem., 1993, 58,3252-3258. 50 N. J.Turner, J. R. Winterman, R. McCague, J. S. Parratt, S. J. C. Taylor Tetrahedron Lett., 1995,36,1113-1116. 51 S.A. Brown, M-C. Parker, N. J. Turner, Tetrahedron:A.symmetry,2000, 11, 1687-1690. 52 A. Moller, C. Syldakt, M. S. Schulze, F. Wagner, Enzyme Microb. Technol., 1988, 10, 618. 53 S . Takahashi, T. Ohashi, Y. Kii, H. Kumagai, H. Yamada,]. Fement. Technol., 1979,57, 328. 54 K. Yokozeki, S. Nakamori, C. Eguchi, K. Yamada, K. Mitsugi, Agric. Biol. Chem., 1987,51,355-362. 55 K. Drauz, M. Kottenhahn, K. Makryaleas, H. Klenk, M. Bernd, Angew. Chem. Int. Ed. En&., 1991, 30, 712-714. 56 0.Kiel, M. P. Schneider, J. P. Rasor, Tetrahedron: Asymmetry,1995,6, 125771260, 57 M. J.Garcia, R. Azerad, Tetrahedron: Asymmetry, 1997,8,85-92. 58 B. S. Deol, D. D. Ridley, G. W. Simpson, Aust.]. Chem., 1976, 29, 2459.

59 D. Buisson ' R. Azerad, Tetrahedron Lett.,

1986,27,2631.

60 S. Danchet, C. Bigot, D. Buisson, R. Azerad,

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61 C. Abalain, D. Buisson, R. Azerad, Tetra-

hedron: Asymmetry,1996,7, 2983.

62 T. Sato, H. Maeno, T. Noro, T. Fujisawa,

Chem. Lett., 1988, 1739.

63 D. Seebach, S. Roggo, T. Maetzke, H.

Braunschweiger, J. Cercus, M. Krieger, Helv. Chim. Acta., 1987,70, 1605. 64 T. Kitahara, K. Mori, Tetrahedron Lett., 1985, 26,451. 65 R. W. Hoffmann, W. Helbig, W. Ladner; Tetrahedron Lett., 1982, 23, 3479. 66 K. Ghosh, W. J. Thompson, P. M. Munson, W. Liu, J. R. HuK Bioorg. Med. Chem. Lett., 1995,5,83. 67 K. Mori, M. Ikunaka, Tetrahedron, 1987, 43, 45. 68 J. Cooper, P. T. Gallagher, D. W. Knight, /. Chem. Soc. Perkin Trans.. 1, 1993, 1313. 69 M. P. Sibi, J. W. Christensen, Tetrahedron Lett., 1990, 31, 5689. 70 D. W. Knight, N. Lewis, A. C. Share, Tetrahedron: Asymmetry,1993,4,625. 71 N. Toyooka, Y. Yoshida, T. Momose, Tetrahedron Lett., 1995, 36, 3715. 72 R. W. Hoffmann, W. Ladner, K. Steinbach, W. Massa, R. Schimdt, G. Snatzke, Chem. Ber., 1981, 114,2786. 73 H. Akita, A. Furuichi, H. Koshiji, K. Horikoshi, T. Oishi, Chem. P h a m . Bull. 1983, 31, 4376. 74 G. Friter, U. Muller, W. Gunther, Tetrahedron, 1984,40, 1269. 75 K. Nakamura, T. Miyai, Y. Kawai, N. Nakajima, A. Ohno, Tetrahedron Lett., 1990, 31, 1159. 76 T. Itoh, Y. Yonekawa, T. Sato, T. Fujisawa, Tetrahedron Lett., 1986, 27, 5405. 77 K. Nakamura, T. Miyai, K. Ushio, S. Oka, A. Ohno, Bull. Chem. Soc. Jpn. 1988,61, 2089. 78 D. Buisson, C. Sanner, M. Larcheveque, R. Azerad, Tetrahedron Lett., 1987.28, 3939. 79 H. Akita, A. Furuichi, H. Koshiji, K. Horikoshi, T. Oishi, Chem. P h a m . Bull., 1984, 32, 1333. 80 0. Cabon, M. Larcheveque, D. Buisson, R. Azerad, Tetrahedron Lett., 1992, 33, 7337. 81 S. Rodriguez, K.T. Schroeder, M. M. Kayser, J. D. Stewart, ]. Org. Chem., 2000, 65, 2586-2587.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

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313

10 Enzymes from Extreme Thermophilic and Hyperthermophilic Archaea and Bacteria Costanzo Bertoldo and Garabed Antranikian

10.1 Introduction

Environments that are considered by man to be extreme, such as those affected by extremes of temperature, pH and salt content, are colonized by a diverse range of microorganisms. These include an interesting group which are adapted to growth at high temperatures[']. In the last two decades it has been possible to isolate microorganisms which can grow optimally even above 100 oC[2-51.The temperature range of growth can be used to define organisms as psychrophiles (-5 to 20 "C), mesophiles (20 to 45 "C), thermophiles (45 to 65 "C), extreme thermophiles (65 to 85 "C) and hyperthermophiles (85 to 110 "C). The majority of the last group, which thrive above the boiling temperature ofwater, belong to the Archaea. However, some of these microorganisms also belong to the bacterial kingdom. Based on comparisons of partial nucleic acid sequences derived from 16 S and 18 S rRNAs, the two primary kingdoms (prokaryotes and eukaryotes) are reclassified into three, namely Bacteria, Archaea and Eukarya[61.Archaea are a newly recognized group of organisms with a distinct evolutionary position and unique physiological,biochemical and genetic properties. The thermophilic representatives of the bacteria that optimally live above 65 "C comprise four genera, namely Themotoga, Themosipho, Fervidobacteriurn (Thermotogales order) and Aqu@feX (Aquificalesorder). The temperature optimum for growth of these microorganisms ranges between 65 and 90 "C. On the other hand the thermophilic representatives of the Archaea comprise more than 20 genera, which belong to the following orders: Sulfolobales, Pyrodictiales, Thermoproteales, Thermococcales, Archaeglobales, Thermoplasmales and the methanogens Methanobacteriales and Methanococcales. Table 10-1 describes some of the growth conditions and of the biochemical features of microorganisms capable of surviving at high temperatures [2-12]. The majority of the microorganisms described in Table 10-1 are heterotrophic and anaerobic; only a few are strict autotrophes. Organisms which belong to the Sulfolobales, Aquificales and Thermoplasmales can also live under aerobic conditions. None of these microorganisms, however, can grow optimally at 100 "C. An exception is the archaeon Pyrobaculurn aerophilum,

314

I

7 0 Enzymesfiom Extreme Thermophilic and Hyperthermophilic Archaea and Bacteria Table 10-1. Taxonomy and some biochemical features of bacteria and archaea growing at high

temperaturesa. Order

BACTERIA Themotogales

Genus

Optimal growth temperature ("C)

Heterotrophic (het) autotrophic (aut) facultative autotrophic (f)

Anaerobic (an) aerobic (ae)

Thermotoga Thermosipho Fervidobacterium

70-80 70-75 65-70

het het het

an an an

Aqu$ex

90

het

ae/an

Sulfolobus Metallosphaera Acidianus Desulfurolobus

65-80 75 88 80

f f aut het

ae/an ae ae/an ae/an

Pyrodictales

Pyrodictium Thermodiscus Hyperthemus

100-105 88 100

het, aut f het

an an an

Trtermoproteales

Thermoproteus Themotilum Desulfurococcus Staphylothemus Pyrobaculum

88 88 85 92 100

het, f, aut het

an an an an ae, an

Thermococcus Pyrococcus

70-87 100

Aqu$ecales ARCHAEA Sulfolobales

7'hemococcales

het het het, f het

het

an an

Archaeoglobales

Archaeoglobus

83

f

an

Themoplasrnales

Themtoplasma

GO

het

ae/an

a Methanogenic microorganisms (Methanobacteriales and Methanococcales) with thermophilic representa-

tives are not shown.

which also grows aerobically at 100 "C. Most of these exotic microorganisms have been isolated by Stetter, Zillig and co-workersfrom various geothermal habitats such as hot springs, sulfataric fields and deep-sea hydrothermal vents. Of great interest are the enzymes that are formed by extreme thermophilic and hyperthermophilic microorganisms. Some of the enzymes that have been recently studied are even active at 140 "C[l3].This short chapter will cover selected enzymes from extreme thermophilic and hyperthermophilic microorganisms that have been described recently. The enzymes from methanogens and thermophilic microorganisms that grow below 70 "C (such as Bacillus, Clostridium and Themus)will not be covered. For more detailed information of this rapidly developing field the reader should consult the following reviews] I7-lo, 12* 141.

70.2 Starch-Processing Enzymes

I

315

10.2 Starch-Processing Enzymes

Starch from cultivated plants represents an ubiquitous and easily accessible source of energy. In plant cells or seeds, starch is usually deposited in the form of large granules in the cytoplasm. Starch is composed exclusively of a-glucose units that are linked by a-1,4- or a-1,G-glycosidic bonds. The two high-molecular-weightcomponents of starch are amylose (15-25%), a linear polymer consisting of a-1,4-linked glucopyranose residues, and amylopectin (75-85 %), a branched polymer containing, in addition to a-1,4 glycosidic linkages, a-1,&linked branch points occurring every 17-26 glucose units. a-Amylose chains, which are not soluble in water but form hydrated micelles, are polydisperse, and their molecular weights vary from hundreds to thousands. The molecular weight of amylopectin may be as high as 100 million, and in solution such a polymer has colloidal or micellar forms. Because of the complex structure of starch, cells require an appropriate combination of hydrolyzing enzymes for its depolymerization to oligosaccharides and smaller sugars such as glucose and maltose. They can be simply classified into two groups: endo-acting enzymes or endo-hydrolases and exo-acting enzymes or exohydrolases. Endoacting enzymes, such as a-amylase (a-1,4-glucan-4-glucanohydrolase; E.C. 3.2.1.1), hydrolyze linkages in the interior of the starch polymer in a random fashion, which leads to the formation of linear and branched oligosaccharides. Exo-acting starch hydrolases include j3-amylase, glucoamalase, and a-glucosidase. These enzymes attack the substrate from the nonreducing end, producing small and well-defined oligosaccharides. P-Amylase (E. C. 3.2.1.2), also referred to as a-1,4-~glucan maltohydrolase or saccharogen amylase, hydrolyzes a-l,4glucosidic linkages to remove successive maltose units from the non-reducing ends of the starch chains, producing p-maltose by an inversion of the anomeric configuration of the maltose (Fig. 10-1). a-Glucosidase (E. C. 3.2.1.20), or a-D-glucoside glucohydrolase, attacks the a-1,4 linkages of oligosaccharides that are produced by the action of other amylolytic enzymes. Unlike glucoamylase, a-glucosidase liberates glucose with an a-anomeric configuration. Enzymes capable of hydrolyzing a-l,G glycosidic bonds in pullulan are defined as pullulanases. On the basis of substrate specificity and product formation, pullulanases have been classified into two groups: pullulanase type I and pullulanase type 11. Pullulanase type I (E. C. 3.2.1.41) specifically hydrolyzes the a-1,G-linkages in pullulan as well as in branched oligosaccharides (debranching enzyme), and its degradation products are maltotriose and linear oligosaccharides, respectively. Pullulanase type I is unable to attack a-l,4-linkagesin a-glucans. Pullulanase type 11, or amylopullulanase, attacks a-1,G-glycosidiclinkages in pullulan and a-l,4-linkages in branched and linear oligosaccharides, converting the latter to small sugars (Fig. 10-1B). In contrast to the previously described pullulanases, pullulan hydrolases types I and I1 are unable to hydrolyze a-1,G-glycosidic linkages in pullulan or in branched

316

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70 Enzymesfiom Extreme Thermophilic and Hyperthermophilic Archaea and Bacteria

substrates. They can attack a-1,4-glycosidic linkages in pullulan, leading to the formation of panose or isopanose. Pullulan hydrolase type I or neopullulanase (E. C. Pullulan hydrolase 3.2.1.135) hydrolyzes pullulan to panose (a-6-D-g~ucosylma~tose). type I1 or isopullulanase (E. C. 3.2.1.57) hydrolyzes pullulan to isopanose (a6-maltosylglucose).Recently, pullulan-hydrolase type I I1 was described, which attacks a-1,4-aswell as a-1,G-glycosidiclinkages in pullulan (Fig. 10-1). Cyclodextrin glycosyltransferase (CGTase, E. C. 2.4.1.19), or a-l,4-~-glucan a-4-~(a-l,4-~-glucano)-transferase, is an enzyme that is generally found in Bacteria and was recently discovered in Archaea. This enzyme produces a series of non-reducing cyclic dextrins from starch, amylose, and other polysaccharides. a-, p-, and ycyclodextrins are rings formed by 6, 7, and 8 glucose units that are linked by a1,4-bonds,respectively (Fig. 10-1). 10.2.1 Thermoactive Amylolytic Enzymes 10.2.1.1 Heat-Stable Amylases and Clucoarnylases.

Extremely thermostable a-amylases have been characterized from the hyperthermophilic Archaea Pyrococcusfuriosus, Pyrococcus woesei and Thennococcus profundus. The optimal temperatures for the activity of these enzymes are 100 "C, 90 "C and 80 "C, respectively. Thermoactive amylolybc enzymes have been also detected in hyperthermophilic Archaea of the genera Sulfolobus, Therrnophilum, Desulfirococcus, and Staphylothennus [15-191. Molecular cloning of the corresponding genes and their expression in heterologous hosts circumvent the problem of insufficient expression in the natural host. The gene encoding an extracellular a-amylase from P. firiosus has recently been cloned, and the recombinant enzyme has been expressed in B. subtilis and E. coli. This is the first report of the expression of an archaeal gene derived from an extremophile in a Bacillus strain. The high thermostability of the pyrococcal extracellular a-amylase (thermal activity even at 130 "C) in the absence of metal ions, together with its unique product pattern and substrate specificity, makes this enzyme an interesting candidate for industrial application. In addition, an intracellular a-amylase gene from P.furiosus has been cloned and sequenced. It was interesting to note that the four highly conserved regions usually identified in aamylases are not found in this enzyme. a-Amylases with lower thermostability and thermoactivity have been isolated from the Archaea Thennococcus profindus, Pyrococcus sp. KODl and the bacteria Themtotoga maritima and Dictioglomus themtophilum. The genes encoding these enzymes were successfully expressed in E. coli. Similar to the amylase from B. licheni$onnis, which is commonly used in liquefaction, the enzyme from 1 maritima requires Ca2+ for activity[2&26].Further investigations have shown that the extreme hyperthermophilic Archaeon Pyrodictium abyssi can grow on various polysaccharides and also secretes a heat-stable amylase (unpublished results). In contrast to a-amylase, the production of glucoamylase seems to be very rare in

70.2 Starch-Processing Enzymes

I

extremely thermophilic and hyperthermophilic Bacteria and Archaea. Among the thermophilic anaerobic Bacteria, glucoamylases have been purified and characterized from Clostridium themohydrosul&ricum 39E, Clostridium thermosaccharolyticumand Themoanaerobacterium themosaccharolyticurn DSM 571 [27-291. Recently, it has been shown that the thermoacidophilic Archaea Thermoplasma acidophilum, Picrophilus tomdus and Picrophilus oshimae produce heat- and acid-stable glucoamylases. The purified Archaeal glucoamylases are optimally active at pH 2 and 90 "C. Catalpc activity is still detectable at pH 0.5 and 100 "C. This represents the first report on the production of glucoamylases in thermophilic Archaea (unpublished results). 10.2.1.2 a-Clucosidases.

a-Glucosidasesare present in thermophilic Archaea and Bacteria. An intracellular aglucosidase has been purified from P.ficriosus. The enzyme exhibits optimal activity at pH 5.0 to 6.0over a temperature range of 105-115 "C; the half life at 98 "C is 48 h. An extracellular a-glucosidase from the thermophilic Archaeon Thermococcus strain AN1 was purified and its molecular characteristics determined13']. The monomeric enzyme (GO kDa) is optimally active at 98 "C. The purified enzyme has a half-life around 35 min, which is increased to around 215 min in the presence of 1% (w/v) dithiothreitol and 1% (w/v) BSA. The substrate preference of the enzyme is: paranitrophenyl-a -D-glucoside > nigerose > panose > palatinose > isomaltose > maltose and turanose. No activity was found with starch, pullulan, amylose, maltotriose, maltotetraose, isomaltotriose, cellobiose and P-gentiobiose. The enzyme is also active at 130 "C. The gene encoding a-glucosidase from Thermococcus hydrothermalis was cloned by complementation of a Saccharomyces cereuisiae maltase-deficient mutant The cDNA clone isolated encodes an open reading frame corresponding to a protein of 242 amino acids. The protein shows 42% identity to a Pyrococcus horikoshii, unknown ORF, but no similarities were obtained with other polysaccharidase sequences. 10.2.1.3 Thermoactive Pullulanases and CCTases.

Thermostable and thermoactive pullulanases from extremophilic microorganisms have been detected in Thermococcus celer; Desulficrococcus mucosus, Staphylothermus marinus and Thermococcus aggregans. Temperature optima between 90 "C and 105 "C, as well as remarkable thermostability even in the absence of substrate and calcium ions, have been observed. Most thermoactive pullulanases identified to date belong to the type I1 group, which attack a-1,4-and a-1,G-glycosidiclinkages. They have been purified from P.&riosus, T. litoralis, T. hydrothermalis and Pyrococcus strain ES4[32-371. Pullulanase type I1 from P.&riosus and P. woesei have been expressed in E. coli. The unfolding and refolding of the pullulanase from P. woesei has been investigated using guanidinium chloride as denaturant. The monomeric enzyme (90 kDa) was

317

318

I found to be very resistant to chemical denaturation and the transition midpoint for 70 Enzymesfrom Extreme Thermophilic and Hypertherrnophilic Archaea and Bacteria

guanidinium chloride-induced unfolding was determined to be 4.86 i 0.29 M for intrinsic fluorescence and 4.90 * 0.31 M for far-UV CD changes. The unfolding process was reversible. Reactivation of the completely denatured enzyme (in 7.8 M guanidinium chloride) was obtained upon removal of the denaturant by stepwise dilution; 100% reactivation was observed when refolding was carried out via a guanidinium chloride concentration of 4 M in the first dilution step. Particular attention has been paid to the role of Ca2+,which activates and stabilizes this archaeal pullulanase against thermal inactivation. The enzyme binds two Ca2+ions with a Kd of 0.080 * 0.010 mM and a Hill coefficient H of 1.00 i 0.10. This cation significantlyenhances the stability of the pullulanase against guanidinium chlorideinduced unfolding. The refolding of the pullulanase, on the other hand, was not affected by Ca" [381. Very recently, the genes encoding the pullulanases from T hydrothermalis, Desulfirococcus mucosus and 7: aggregans have been isolated and expressed in mesophilic hosts. Since the latter enzyme attacks a-1,4as well as a-1,G glycosidic linkages in pullulan, it has been classified as pullulan-hydrolase type 111. Pullulan is converted to maltotriose, maltose, panose and glucose [40-421. The aerobic thermophilic bacterium Thermus caldophilus GK-24 produces a thermostable pullulanase of type I when grown on starch. This enzyme debranches amylopectin by attacking specifically a-l,6-glycosidiclinkages. The pullulanase is optimally active at 75 "C and pH 5.5, is thermostable up to 90 "C, and does not require Ca" for either activity or stability. The first debranching enzyme (pullulanase type I) from an anaerobic thermophile was identified in the bacterium Fervidobacterium pennivorans Ven5, which was cloned and expressed in E. coli. In contrast to pullulanase type I1 from P.woesei (specific to both a-1,Gand a-1,4 glycosidic linkages) the enzyme from F. pennivorans Ven5 attacks exclusively the a-1,G-glycosidic linkages in polysaccharides. This thermostable debranching enzyme leads to the formation of long-chain linear polysaccharides from amylopectin IL4'1. Thermostable cyclodextrin glycosyltransferases (CGTases) are produced by Thermoanaerobacter species, Thermoanaerobacterium thermosulfirigenes and Anaerobranca g0ttschalkii1~"'~1. Recently, a CGTase, with optimal temperature at 100 "C, waspurified from a newly isolated Archaeon, Thermococcus sp. This is the first report of the presence of a thermostable CGTase in a hyperthermophilic Ar~haeon[~']. The enzyme from this strain has been cloned and sequenced. The gene of 2217 nucleotides encodes a protein with an MW of 83 kDa. The ability of extreme thermophiles and hyperthermophiles to produce heat-stable glycosyl hydrolases is summarised in Table 10-2. The finding of extremely thermophilic Bacteria and Archaea capable of producing novel thermostable starch-hydrolyzing enzymes is a valuable contribution to the starch-processing industry. By using robust starch-modifylngenzymes from thermophiles, innovative and environmentally friendly processes can be developed, aiming at the formation of products of high added value for the food industry. New and enhanced functionality can be obtained by changing the structural properties of starch. In order to prevent retrogradation, starch-modifyingenzymes can be used at higher temperatures. The use of the extremely thermostable amylolytic enzymes can

Pullulanase type 11

Pullulanase type I

100

StaphylothemtusmarinusIg01

100 100

Pyrodictium abyssi["]

85

pyrococcus woesei['OOl

~UCOSUS~~~]

75

Thermus caldophilus GK24[751

Desul&rococcus

90

nemotoga mantima MSB81"I

85-90

Themtotoga mantima MSB8["I 80

90

Therrnococcus aggregan~[~~l

Dyctyoglomusthemtophilum Rt46B.1[731

Fervidobacteriumpennavorans Ven517*1

6.5

95

Thermococcus projkndus[801

9.0

6.0

5.5

5.5

6.0

6

7.0

5.5

5.5 4.0-5.0

80 80

Thermococcus projkndus DT54321"I

5.5

90

5.5 -

5.0

5.5

6.5

Trtemococcus ceIeP51

Sulfolobussolfataricus1881

100

Pyrodictium abyssi["I

-

90

6.5-7.5 7.0

100 100 100

5.0

100

Pyrococcus woesi['OO1

Desulfurococcus m u c o s ~ s [ ~ ~ l

a-Amylase

Enzyme properties Optimal Optimal temperature pH

Pyrococcus sp. KODl

Organism'

Enzyme

-

90

65 74

190 (93) 93 (subunit)

61

75

-

42

42

240 -

-

-

68

49.5

129 68

-

Mw Wa)

Table 10-2. Starch hydrolyzing enzymes from extreme thermophilic and hyperthermophilicArchaea and Bacteria.

Crude extract

Purified/cloned/cell associated

Purified/cell associated Purified/doned

Purified/cloned Cloned/typs Ib

Purified/cloned/lipoprotein

Purified/cloned/cytoplasmic fraction

Cloned

Purified/"Amy L"

Purified/doned/"Amy S"

Crude extract

Extracellular

Crude extract

Purified/Extracellular Crude extractb

Purified/cloned/extracellular

Purified/cloned/intracellular Purified/cloned/extracellular

Purified/cloned

Remarks

3w

T

a 2. 2

: a

i? 3

h,

9

d

130 -

Thermococcus strain A N ~ I ~ ~ I Thermococcushydrothermalis["]

a Values in brackets give the optimal growth temperamre for each organism in 'C b Unpublished results; - not determined

a-Glucosidase

70

100

Anaerobranca g~ttaschallkii['~~

7.0

90

Picrophilus torridus["] Themtococcus SP.['~] 80

90

Thermoanaerobacterium thermosulfurigenes[601

90

Themtoplasma acidophilurn["l

Picrophilus oshimae["l

Glucoamylase

CGTase

2.0

100

Thermococcus aggregan~['~]

Pullulan-hydrolasetyp 111

63 -

Purified/extracell./glycoprotein Cloned

Purified

6G

-

Purified/cloned/crystallized

Purified Purified

Purified

Purified

Purified/cloned

Purified/extracell./glycoprotein

Purified/extracell./glycoprotein

Crude extract

Remarks

68

133 83

140

141

83

128

-

8.0

4.0-4.5

2.0

6.5

6.5

5.5

119

95

Therrnococcus hydrothermalis~80]

-

5.5

Mw (kW

5.5

90

Enzyme properties Optimal Optimal temperature pH

98

Organism"

Themococcus lit~ralis['~l

(cont.).

Thermococcus celerlss1

Enzyme

Table 10-2.

W

0 N

I

4

n

9

6. b 3 s

s-

3

z

$

n 3

4

3 3 9 a 2

a

3

-c

2 !

0

u

-

10.3 Cellulose-Hydrolyzing Enzymes

lead to valuable products, which include innovative starch-based materials with gelatin-like characteristics and defined linear dextrins that can be used as fat substitutes, texturizers, aroma stabilizers and prebiotics. CGTases are used for the production of cyclodextrins that can be used as a gelling, thickening or stabilizing agent in jelly desserts, dressing, confectionery, dairy and meat products. Because of the ability of cyclodextrins to form inclusion complexes with a variety of organic molecules, they improve the solubility of hydrophobic compounds in aqueous solution. This is of interest for the pharmaceutical and cosmetic industries['', 'l1. Cyclodextrin production is a multistage process in which starch is first liquefied by a heat-stable amylase, and in the second step a less-thermostable CGTase from Bacillus sp. is used. The application of heat-stable CGTase in jet cooking, where temperatures up to 105 "C are achieved, will allow liquefaction and cyclization to take place in one step.

10.3

Cellulose-Hydrolyzing Enzymes

Cellulose commonly accounts for up to 40% of the plant biomass. It consists of glucose units linked by P-1,Cglycosidicbonds with a polymerization grade of up to 15000 glucose units in a linear mode. Although cellulose has a high affinity to water, it is completely insoluble. Natural cellulose compounds are structurally heterogeneous and have both amorphous and highly ordered crystalline regions. The degree of crystallinity depends on the source of the cellulose, and the more highly crystalline regions are more resistant to enzymatic hydrolysis. Cellulose can be hydrolyzed into glucose by the synergistic action of at least three different enzymes: endoglucanase, exoglucanase (cellobiohydrolase)and P-glucosidase. Synonyms for cellulases (E. C. 3.2.1.4) are P-l,4-~-glucanglucano-hydrolases, endo-f3-1,4-glucanasesor carboxymethyl cellulases. This enzyme is an endoglucanase which hydrolyzes cellulose in a random manner as endo-hydrolase producing various oligosaccharides, cellobiose linkages in and glucose. The enzyme catalyzes the hydrolysis of P-1,4-~-glycosidic cellulose but can also hydrolyze 1,Clinkages in P-D-glucans containing 1,3-linkages. Exoglucanases, P-1,4-cellobiosidases, exocellobiohydrolases or P-1,kellobiohylinkages in cellulose and cellotedrolases (E. C. 3.2.1.91) hydrolyze P-l,4-~-glycosidic traose, releasing cellobiose from the non-reducing end of the chain. P-Glucosidases (E. C. 3.2.1.21), gentobiases, cellobiases or amygdalases catalyze the hydrolysis of terminal, non-reducing P-D-glucoseresidues releasing 0-D-glucose. These enzymes have a wide specificity for P-D-glucosides. They are able to hydrolyze P-D-galactosides,P-L-arabinosides,P-D-xylosides, and P-D-fucosides. 10.3.1

Thermostable Cellulases

Thermostable cellulases active towards crystalline cellulose are of great biotechnological interest. Several cellulose-degrading enzymes from various thermophilic

I

321

322

I

10 Enzymesfrom Extreme Thermophilic and Hyperthermophilic Archaea and Bacteria

organisms have been cloned, purified, and characterized. A thermostable cellulase from Thermotoga maritima MSB8 has been ~haracterized1~~1. The enzyme is rather small, with a molecular weight (MW)of 27 kDa, and is optimally active at 95 "C and between pH 6.0 and 7.0. Two thermostable endocellulases, CelA and CelB, were purified from Thermotoga neapolitana. CelA (MW of 29 kDa) is optimally active at pH 6 at 95 "C, while CelB (MW of 30 kDa) has a broader optimal pH range (pH 6 to 6.6) at 106 "C. The genes encoding these two endocellulases have been Cellulase and hemicellulase genes have been found clustered together on the genome of the thennophilic anaerobic bacterium Caldocellum saccharolyticum, which grows on cellulose and hemicellulose as sole carbon sources. The gene for one of the cellulases (celA)was isolated and was found to consist of 1751 amino acids. This is the largest cellulase gene described to A large cellulolytic enzyme (CelA) with the ability to hydrolyze microcrystalline cellulose was isolated from the extremely thermophilic bacterium Anaerocellum t h e r m o p h i l ~ m [The ~ ~ ]enzyme . has an apparent molecular weight of 230 kDa, exhibits significant activity towards Avicel and is most active towards soluble substrates such as CM-cellulose (CMC) and P-glucan. Maximal activity was observed at pH 5-6 and 85-95 "C. The thennostable exoacting cellobiohydrolase from Thermotoga maritima MSB8 has an MW of 29 kDa and is optimally active at 95 "C at pH 6.0-7.5 with a halflife of 2 h at 95 "C. The enzyme hydrolyzes Avicel, CM-Cellulose and b-glucan forming cellobiose and cellotriose. A thermostable cellobiase is produced by Thermotoga sp. FjSS3-Bl The enzyme is highly thermostable and shows maximal activity at 115 "C at pH 6.8-7.8. The thermostability of this enzyme is salt dependent. This cellobiase is active against amorphous cellulose and CM-cellulose. Recently, a thermostable endoglucanase, which is capable of degrading P-1,4 bonds of p-glucans and cellulose, has been identified in the Archaeon Pyrococcus firiosus. The gene encoding this enzyme has been cloned and sequenced in E. coli and has significant amino acid sequence similarities with endoglucanases from glucosyl hydrolases family 12. The purified recombinant endoglucanase hydrolyzes P-1,4- but not P-1,3-glycosidiclinkages and has the highest specific activity with cellopentaose and cellohexaose as substrates 15'1. In contrast to this, several 0glucosidases have been detected in Archaea. In fact, archaeal P-glucosidases have been found in Sulfolobus solfataricus MT4, S. acidocaldarius, S. shibatae and P. JiLriosus[s8-"1. The enzyme from the latter microorganism (homotetramer, 56 kDa/ subunit) is very stable and shows optimal activity at 102 "C to 105 "C with a half-life of 3.5 days at 100 "C and 13 h at 110 oC[60].The P-glucosidase from S. solfataricus The enzyme is a homotetramer (56 MT4 has been purified and kDa/subunit) and very resistant to various denaturants with activity up to 85 "C["l. The gene for this P-glucosidase has been cloned and overexpressed in E. c01i[63-6s1 (Table 10-3). Cellulose-hydrolyzing enzymes are widespread in Fungi and Bacteria. Less thermoactive cellulases have already found various biotechnological applications. The most effective enzyme of commercial interest is the cellulase produced by Trichoderma sp. [66]. Cellulolytic enzymes can be used in alcohol production to improve juice yields and effective color extraction of juices. The presence of cellulases in

Pyrococcus kodakaraensi~['~]

Thermotoga thermar~m['~I

Thermotoga neapolitana[801

Thermotoga sp. strain FjSS3-B.1[80,85]

Thermotoga mantima MSB8 Thermotoga sp. strain FjSS3-B.1(801 Anaerocellum thermophilum Rt46B. lllool Pyrococcus&riosus['Oo1 Pyrococcus&riosusl'ool SulfolobussoEfataricus[881 Thermotoga mantima MSB8 Thermotoga sp. strain FjSS3-B.1[801 Pyrodictium a b y s ~ i [ ~ ~ l Dyctyoglomus thermophilum Rt46B.1[73] Thermotoga mantima MSB8[801

Thermotoga mantima MSB8[801 Thermotoga neapolitana[801

Organism"

95 95 106 95 115 85 100 102-105 105 75 80 110 85 92 105 105 85 85 95 80 90-100 85

- not determined

5.3 6.2 7.0 5.5 6.5 6.2 5.4 5.3 6.3 5.5 5.5-6.0 6.0 7.0 5.0

6.0-7.5 6.0 6.0-6.5 6.0-7.5 6.S7.8 6.5 6.0 -

pH

Enzyme properties Optimal

temperature

Optimal

a Values in the brackets give the optimal growth temperature for each organism in "C

Chitinase

Endoxylanases

P-Glycosidase

Exoglucanase

Endoglucanase

Enzyme

31 120 40 31 40 37 119 105/150 35 135

27 29 30 29 36 31 35.9 230/58 240/56 95(47) lOO(75) -

Mw (kW

Purified/cloned/cellulase I Purified/cloned/Cell A Purified/cloned/Cell B Purified/cellulase I1 Purified/cell-associated Cloned Cloned Purified/cloned Purified/cloned Purified/cloned Purified/ toga-associated Crude extract Purified/cloned Pur./toga associated/XynA Pur./toga-associated/XynB Pur./cloned/ toga-associated Pur./cloned Purified Purified/cloned Pur./toga-associated/Endoxylanase1 Pur./Endoxylanase 2 Purified/cloned

Remarks

Production o f thermoactive cellulases (exoglucanase,B-glycosidase),xylanases (endoxylanase)and chitinase by some representatives of extreme thermophilic and hvpertherrnoDhilicArchaea and Bacteria.

Table 10-3.

W W N

2 -

3

-z

9

4

r' 5.

a

SL

:

2 -=

E

2-

LJ

P d

324

I

70 Enzymesfrom Extreme Jhermophilic and Hyperthermophilic Archaea and Bacteria

detergents causes color brightening, softening and improvement of particulate soil removal. Cellulase (Denimax@Novo Nordisk) is also used for the “biostoning” of jeans instead of using stones. Other applications of cellulases include the pretreatment of cellulosic biomass and forage crops to improve nutritional quality and digestibility, enzymatic saccharification of agricultural and industrial wastes and production of fine chemicals.

10.4 Xylan-Degrading Enzymes

Xylan is a heterogeneous molecule that constitutes the main polymeric compound of hemicellulose, a fraction of the plant cell wall which is a major reservoir of fked carbon in nature. The main chain of the heteropolymer is composed of xylose residues linked by P-1,4-glycosidicbonds. Approximately half of the xylose residues have substitution at 0 - 2 or 0 - 3 positions with acetyl, arabinosyl and glucuronosyl groups. The complete degradation of xylan requires the action of several enzymes (for a detailed description see reviews[67] The endo-P-1,4-xylanase (E. C. 3.2.1.8), or P-1,4-xylanxylanohydrolase,hydrolyzes P-1,4-xylosydiclinkages in xylans, while P-1,4-xylosidase,P-xylosidase, P-1,4-xylan xylohydrolase, xylobiase or exo-p1,4-xylosidase(E. C. 3.2.1.37) hydrolyzes P-1,4-xylansand xylobiose by removing the successive xylose residues from the non-reducing termini. a-Arabinofuranosidase or arabinosidase (E. C. 3.2.1.55) hydrolyzes the terminal non-reducing a-r-arabinofuranoside residues in a-L-arabinosides.The enzyme also acts on a-L-arabinofuranosides [a-L-arabinanscontaining either (1,3) or (1,5)-linkages].Glucuronoarabinoxylan endo-P-1,4-xylanase,feraxan endoxylanase or glucuronoarabinoxylanP-1,4-xylanohydrolase (E. C. 3.2.1.136) attacks ~-1,4-xylosyllinkages in some glucuronoarabinoxylans. This enzyme also shows high activity toward femloylated arabinoxylans from cereal plant cell walls. Acetyl xylan esterase (E. C. 3.1.1.6) removes acetyl groups from xylan. 10.4.1 Thermostable Xylanases

So far, only a few extreme thermophilic microorganisms are able to grow on xylan and secrete thermoactive xylanolybc enzymes (Table 10-3). Members of the order Thermotogales and Dictyoglomus themophilum Rt46B. 1 have been described to produce xylanases that are active and stable at high temperatures. The most thermostable endoxylanases that have been described so far are those derived from Themtotoga sp. strain FjSS3-B.l, Themtotoga maritima, T. neapolitana and T. thermarum. These enzymes, which are active between 80 and 105 “C, are mainly cellassociated and most probably localized within the toga. Several genes encoding xylanases have already been cloned and sequenced. The gene from T. maritima, encoding a thermostable xylanase, has been cloned and expressed in E. coli. Comparison between the T. maritima recombinant xylanase and the commercially

70.5 Chitin Degradation

available enzyme, PulpenzymeTMindicates that the thermostable xylanase could be of interest for application in the pulp and paper industry. A xylanase has been found in the Archaeon Thermococcus zilligii strain AN1, which grows optimally at 75 "C. The enzyme has a molecular weight of 95 kDa and a unique N-terminal seq~ence["~'1. The pH optimum for activity is 6.0, and the half-life at 100 "C is 8 min. Another archaeal xylanase with a temperature optimum of 110 "C was found in the hyperthermophilic Archaeon Pyrodictiurn abyssi. Xylanases from Bacteria have a wide range of potential biotechnological applications. They are already produced on an industrial scale and are used as food additives in poultry, for increasing feed efficiency diets [7G. 771 and in wheat flour for improving dough handling and the quality of baked In recent years, the major interest in thermostable xylanases is found in enzyme-aided bleaching of paper [791. More than 2 million tons of chlorine and chlorine derivatives are used annually in the United States for pulp bleaching. The chlorinated lignin derivatives generated by this process constitute a major environmental problem caused by the pulp and paper industry[79].Recent investigations have demonstrated the feasibility of enzymatic treatment as an alternative to chlorine bleaching for the removal of residual lignin from pulp[8o].Treatment of craft pulp with xylanase leads to a release of xylan and residual lignin without undue loss of other pulp components. Xylanase treatment at elevated temperatures opens up the cell wall structure, thereby facilitating lignin removal in subsequent bleaching stages. Xylanases from moderate thermophilic microorganisms are rapidly denatured at temperatures above 70 "C. Several of the non-chlorine bleaching stages used in commercial operations are performed well above this temperature; consequently,the pulp must be cooled before treatment with the available enzymes and reheated for subsequent processing steps c7'1.

10.5

Chitin Degradation

Chitin is a linear p-1,4homopolymer of N-acetyl-glucosamineresidues and is one of the most abundant natural biopolymers on earth. Particularly in the marine environment, chitin is produced in enormous amounts, and its turnover is due to the action of chitinolytic enzymes. Chitin is the major structural component of most fungi and invertebratesLs1. 8 2 ] , while for soil or marine Bacteria chitin serves as a nutrient. Chitin degradation is known to proceed with the endo-acting chitin hydrolase (chitinase A: E. C. 3.2.1.14) and the chitin oligomer-degrading exo-acting hydrolases (chitinase B) and N-acetyl-D-glycosaminidase (trivial name: chitobiase; E.C. 3.2.1.52). Chitobiase degrades only small N-acetyl-D-glucosamineoligomers (up to pentamers), and the released N-acetyl-D-glucosaminemonomers retain their C1 anomeric configuration. Chitin and its derivatives exhibit interesting properties that make them a valuable raw material for several applications[83-871.It has been estimated that the annual world-wide formation rate and steady state amount of chitin is in the order of lo1' to

I

325

326

I

10 Enzymesfram Extreme Thermophilic and Hyperthermophilic Archaea and Bacteria

lo1' tons per year. Therefore, application of thermostable chitin-hydrolyzing enzymes (chitinases) is expected for effective utilization of this abundant biomass. Although a large number of chitin-hydrolyzingenzymes have been isolated and their corresponding genes have been cloned and characterized, only few thermostable chitin-hydrolyzingenzymes are known. These enzymes have been isolated from the thermophilic microorganisms Bacillus lichenformis X - ~ U ,Bacillus sp. BG-11 and Streptomyces thennoviolaceus OPC-520[", 891. The extreme thermophilic anaerobic Archeon Thermococcus chitonophagus has been reported to hydrolyze chitin I.'[ This is the first extremophilic Archaeon which produces chitinase(s) and N-acetylglucosaminidase(s);however, sequence and structural information for archaeal chitinases have not yet been reported. Very recently, the gene encoding a chitinase from a hyperthermophilic archaeon Pyrococcus kodakaraensis KODl was cloned, sequenced and expressed in E. coli. The purified recombinant protein is optimally active at 85 "C and pH 5.0. The enzyme produces chitobiose as the major end product (Table 10-3).

10.6 Proteolytic Enzymes

Proteins are the most abundant organic molecules in living cells and constitute more than 50% of their dry weight. The molecular weight of proteins that are made up of one or more polypeptide chains can vary from a few thousands to more than one million daltons. All proteins are constructed from a basic set of 20 amino acids that are covalently linked by peptide bonds. The three-dimensional conformation of proteins may vary. Globular proteins (spherical or globular) are soluble and usually have dynamic function. Fibrous proteins on the other hand occur as sheets or rods, are insoluble and serve as structural elements. The enzymes which hydrolyze the peptide bonds in proteins are defined as proteases. They are also called endopeptidases because they hydrolyze peptide bonds inside the polypeptide chain. Exopeptidases (either carboxypeptidases or aminopeptidases) on the other hand can split off the terminal residues of the polypeptide chain. Proteases (endopeptidases) play an important role in the utilization of proteins by various microbes. They are classified into four groups depending on the nature of their active center. Serine proteases have a serine residue in their active center and are inhibited by DFP (diisopropylphosphofluoride)and PM SF (phenylmethylsulfonylfluoride). 11. Cysteine proteases have a SH groups in their active center and are inhibited by thiol reagents, heavy metal ions, alkylating agents and oxidizing agents. 111. The activity of metal proteases depends on tightly bound divalent cations. They are inactivated by chelating agents. IV. Aspartic proteases (acid proteases) are rare in Bacteria and contain one or more aspartic acid residues in their active center. Inactivation of the enzyme can be achieved by alkylation of the aspartic acid residues with DAN (diazoacetyl-DLnorleucine methyl ester) [911.

I.

10.6 froteolytic Enzymes

10.6.1 Stable proteases

A variety of heat-stable proteases have been identified in hyperthermophilic Archaea belonging to the genera Desulfirrococcus, Sulfolobus, Staphylothemus, Themococcus, Pyrobaculurn and Pyrococcus. It has been found that most proteases from extremophiles belong to the serine type and are stable at high temperatures even in the presence of high concentrations of detergents and denaturing agents (Table 10-4).A heat-stable serine protease was isolated from cell-free supernatants of the hyperA cell-associated serine thermophilic Archaeon Desulfirrococcus strain Tok12S1 protease was characterized from Desulfirococcus strain SY that showed a half-life of 4.3 h at 95 oC[93].A globular serine protease from Staphylothemus marinus was found to be extremely thermostable. This enzyme, which is bound to the stalk of filiform glycoprotein complex, named tetrabrachion, has a residual activity even at The properties of extracellular serine proteases 135 "C after 10 min of incubati~n['~I. from a number of Themococcus species have been analyzed["]. The extracellular enzyme from T. stetteri has a molecular weight of 68 kDa and is highly stable and resistant to chemical denaturation, as illustrated by a half-life of 2.5 h at 100 "C and retention of 70 % of its activity in the presence of 1% SDS (961. A novel intracellular serine protease (perinilase) from the aerobic hyperthermophilic Archaeon Aeropyrurn pernix K 1 was purified and characterized. At 90 "C, the pernilase has a broad pH profile and an optimum at pH 9.0 for peptide hydrolysis. Several proteases from hyperthermophiles have been cloned and sequenced, but in general their expression in a mesophilic host is difficult. A gene encoding a subtilisin-like serine protease, named aereolysin, has been cloned from Pyrobaculurn aerophilurn, and the protein was modeled based on structures of subtilisin-typeproteases [971. Multiple proteolytic activities have been observed in P. firriosus. The cell-envelope associated serine protease of P.firriosus, called pyrolysin, was found to be highly stable, with a half-life of 20 min at 105 oC[981.The pyrolysin gene was cloned and sequenced, and it was shown that this enzyme is a subtilisin-likeserine protease["]. A serine protease from Aqu@x pyrophilus was cloned and weakly expressed in E. coli. The activity of the enzyme was highest at 85 "C and pH 9. The half-life of the protein (G h at 105 "C) makes it one of the most heat-stable proteases known to date. Proteases have also been characterized from the thermoacidophilic Archaea Sulfolobus solfataricus and S. acidocaldarius. In addition to the serine proteases, other types of enzymes have been identified in extremophiles: a thiol protease from Pyrococcus sp. KOD1, a propylpeptidase (PEPase)and a new type of protease from P. firiosus. An extracellular protease, which is designated aeropyrolysin, was purified from Aeropyrurn pernix K 1 (JCM 9820). The enzyme activity is completely inhibited by EDTA and EGTA, indicating that it is a metalloprotease. The enzyme is highly resistant to denaturing reagents and highly thermostable, showing a half-life of 2.5 h at 120 "C and 1.2 h at 125 "C in the presence of 1mM CaC12. These results indicate that this enzyme is one of the most thermostable extracellular metallo-proteases reported to date. Thermostable serine proteases were also detected in a number of extreme thermophilic Bacteria belonging to the genera Themotoga and Fervido-

I

327

6.3

-

85

Pyrococcusfuriosus[loo]

95 85

Themtococcus celerI8'1

Themtococcuslitoralis[901

Themtococcus stetteri [751

Solfolobus solfataricus[881

Aminopeptidase I Aminopeptidase I1 Endopeptidase I, 11,111 Carboxypeptidase

- not determined

a Values in the brackets give the optimal growth temperature for each organism in "C

Sulfolobus acidocaldariu~[~~]

9.0-9.5

85

Thermobacteroidesproteolyticus I1'

Acidic protease

10

80

Fervidobacterium pennavorans I7O1

-

-

-

-

-

2.0 -

90

7

7.0-9.0

85

110

9.0

90

Aquij%xpyrophilus19001

170 115,32,27 160

> 450

44 -

130 -

50 43

118(52)

Crude extract Crude extract Crude extract Crude extract

Cloned

Crude extract Purified

Purified/keratin hydrolysis

Purified Purified

Purified

Stable up to 135 "C

6.5-8

Pur./doned 68

140

Aeropyrurn Pernix K1 I9O1

Pyrococcus sp. KODl [')'I

Crude extract Crude extract

-

-

Crude extract

-

-

Protease I/purified Pyrolysin/pur./cloned Cloned

Purified

9.0

-

Thiol protease

52 124(29) 105/80

Remarks

8.5

Sulfolobus solfataricuds8]

Staphylothermus marinus [901

7.5

95

Themtococcus aggregans L7'1 9.5

7.0

90

Pyrobaculum aerophilum 19'1

7.5

95

Desulfurococcus mucosus

Serine protease

Enzyme properties Optimal Mw pH (kW

Organism"

Optimal temperature

Properties of thermoactive proteolytic enzymes from extreme thermophilic and hyperthermophilicArchaea and Bacteria.

Enzyme

Table 10-4.

-6'

2.

H

0

a m

Q

$f! s

6' b

4z a-2

a z

Q 3

'"z -_

2

B $m

a

3

-c

9

u

0

W 00 l4

10.7 lntracellular Enzymes

I

329

bacterium (unpublished results). The enzyme system from Fervidobacteriurn pennivorans is able to hydrolyze feather keratin forming amino acids and peptides. The enzyme is optimally active at 80°C and pH 10.0['09].The amount of proteolytic enzymes produced worldwide on a commercial scale exceeds that of the other biotechnological enzymes used. Heat-stable proteases have great potential for various applications including the textile and pharmaceutical industries. Serine alkaline proteases are currently used as additives to household detergents for laundering, where they have to resist denaturation by detergents and alkaline conditions. Proteases showing high keratinolytic and elastolyhc activities are used for soaking in the leather industry. Proteases are also used as catalysts for peptide synthesis using their reverse reaction [100-1091.

10.7 lntracellular Enzymes

A number of intracellular enzymes from extreme thermophilic and hyperthermophilic microorganisms have been investigated. The majority of the intracellular enzymes known to date show slightly less thermostability than the extracellular enzymes. Some of these enzymes, which have been characterized from Archaea belonging to the order Sulfolobalses and Themococcales, include alcohol dehydrogenase, glucose dehydrogenase, glyceraldehyde-3-phospho-dehydrogenase, NADH dehydrogenase, j3-galactosidase, citrate synthase, malic enzyme, fumarase, sadenosylmethionine synthetase, ATPase, ATP sulfurylase, aspartate aminotransferase, DNA polymerase, RNA polymerase, topoisomerase and polyphosphate kinase (for review Other reports are available on extremely thermoactive intracellular enzymes that are even active above 100 "C. Glyceraldehyde-3-phosphate dehydrogenase from the Archaeon P. woesei was characterized and the gene was cloned in E. coli. This enzyme is strictly phosphate dependent and utilizes either NAD+ or NADP+; the half-life ofthe enzyme at 100 "C is 4 4 min['lO].The amino acid composition of glyceraldehyde-3-phosphate dehydrogenase from P. woesei was determined and compared with mesophilic and thermophilic Archaea. The primary structure of this enzyme exhibits a high proportion of aromatic amino acid residues and a low proportion of sulfur-containing residues. The glutamate dehydrogenase (GDH)from P. woesei and P.firriosus was purified and characterized. This enzyme is probably involved in the first step of nitrogen metabolism. GDH from P. woesei was purified in a single-affinity chromatography step['"]. It utilizes both NAD' and NADP+ as cofactors with a preference for the phosphorylated form. The purified enzyme from both strains is a hexamer with identical subunits of 45 kDa each['1*, "31 .Twenty-four N-terminal residues of GDH were determined and used to The construct gene-specific DNA probes via the polymerase chain reaction ["'I. GDH gene was cloned in E. coli. Its nucleotide sequence and amino acid composition were determined. A highly thermoactive glucose isomerase with maximal enzymatic activity at 105 "C was purified from T. maritima and ~haracterized["~]. This enzyme could play an important role in the industrial bioconversion of glucose

330

I

70 Enzymesfrom Extreme Thermophilic and Hyperthermophilic Archaea and BaGteria

to fructose. Other remarkable thermoactive enzymes such as hydrogena~e["~], aldehyde ferredoxin oxidoreductase[1'6],and acetyl-Co A synthetase (ADP forming) were detected in P . ~ r i o s u ~ ~ ' ~ ~ - ' ~ ~ ] . DNA polymerases (E. C. 2.7.7.7.) are other important intracellular enzymes that play a key role in the replication of cellular information present in all life forms. They catalyze, in the presence of Mg2+ ions, the addition of a deoxyribonucleoside 5'-triphosphate onto the growing 3'-OH end of a primer strand, forming complementary base pairs to a second strand. Thermostable DNA polymerases play a major role in a variety of molecular biological applications, e. g. DNA amplification, sequencing or labeling. More than 100 DNA polymerase genes have been cloned and sequenced from various organisms, including thermophilic Bacteria and Archaea. One of the most important advances in molecular biology during the last ten years is . The PCR procedure the development of a polymerase chain reaction (PCR)['20-1221 first described utilized the Klenow fragment of E. coli DNA polymerase I, which was heat labile and had to be added during each cycle following the denaturation and primary hybridization steps. Introduction of thermostable DNA polymerases in PCR facilitated the automation of the thermal cycling part of the procedure. The DNA polymerase I from the bacterium Themus aquaticus, called Taq polymerase, was the first thermostable DNA polymerase characterized[123, 1241 and applied in PCR. A thermostable DNA polymerase from Themotoga maritimal'251was reported to have a 3'-5'-exonudease Archaeal proofreading polymerases such as Pwo pol [1271 from Pyrococcus woesei[128],Pfi pol [1291fromPyrococcus f i r i o s ~ s ~ 'Deep ~~~, 1341 from T h e m o VentTM from Pyrococcus strain GB-D[132] or Vent: coccus litorali~['~~] have an error rate that is up to ten times lower than that of Taq polymerase. The 9"N-7 DNA polymerase from Themococcus sp. strain 9"N-7 has a fivefold higher 3'-5'-exonuclease activity than T. litoralis DNA polymerase [1361. However, Taq polymerase was not replaced by these DNA polymerases because of their low extension rates, among other factors. DNA polymerases with higher fidelity are not necessarily suitable for amplification of long DNA fragments because of their potentially strong exonuclease activity [1371. The recombinant KODl DNA polymerase from Pyrococcus sp. strain KODl has been reported to show low error rates high processivity (persistence of sequential nucleo(similar values to those of P'), tide polymerization) and high extension rates, resulting in an accurate amplification of target DNA sequences up to 6 kb[1381.In order to optimize the delicate competition of polymerase and exonuclease activity, the exo-motif 1 of the 9"N-7 DNA polymerase was mutated in an attempt to reduce the level of exonuclease activity without totally eliminating it [13'. 139. l4Ol. Similarly, the PCR performance was optimized by site-directed mutagenesis of the DNA binding motif of the DNA polymerase from Themococcus aggregans and Sulfolobus solfataricus['41].

References I331

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70 Enzyrnesfrorn Extreme Therrnophilic and Hypertherrnophilic Archaea and Bacteria

A. S. Kaledin, A. G. Sliusarenko, S. Goroetskii, Biokhimiya 1980,45, 644-651. 125 R. Huber, T. A. Langworthy, H. Konig, M. Thomm, C. R. Woese, U. B. Sleytr, K. 0. Stetter, Arch. Microbiol. 1986, 144, 324-333. 126 D. A. Bost, S. Stoffel, P. Landre, F. C. Lawyer, J. Akers, R. D. Abramson, D. H. Gelfand, FASEB J 8,1994, A1395. 127 B. Frey, B. Suppmann, Biochemica, 1995, 2, 34-35. 128 W. Zillig, 1. Holz, H. P. Klenk, J. Trent, S. Wunderl, D. Janekowic, E. Imsel, B. Haas, SystemAppl. Microbiol, 1987, 9, 62-70. 129 K. S. Lundberg, D. D. Shoemaker, M. W. W. Adams, J. M. Short, J. A. Sorge, E. J. Marthur, Gene, 1991, 108,l-6. 130 G. Fiala, K. 0. Stetter, Arch. Microbiol. 1986, 145,56-61. 131 F. B. Perler, S. Kumar, H. Kong, Adv. Protein. Chem. 1996,48,377-435 132 H. W. Jannasch, C. 0. Wirsen, S. J. Moly124

neaux, T. A. Langworthy,Appl. Environ. Microbiol. 1992, 58, 3472-3481. 133 N. F. Cariello, J. A. Swenberg, T. R. Skopek, Nucleic Acids Res., 1991, 19, 4193-4198. 134 P. Mattila, J. Korpela, T. Tenkanen, K. Pitkanen, Nucleic Acids Res., 1991, 19, 4967-497 3. 135 A. Neuner, H. W. Jannasch, S. Belkin, K.O. Stetter, Arch. Microbiol. 1990, 153, 205-207. 136 M. W. Southworth, H. Kong, R. B. Kucera, J. Ware, H. W. Jannasch, F. B. Perler, Proc. Natl. Acad. Sci. U S A1996,93, 5281-5285. 137 W. M. Barnes, Gene 1992, 112,29-35. 138 M. Takagi, M. Nishioka, H. Kakihara, M. Kitabayashi, H. Inoue, B. Kawakami, M. Oka, T. Imanaka, Appl. Environ. Microbiol. 1997, 63,4504-4510. 139 L. Blanco, A. Bernad, M. A. Blasco, M. Salas, Gene 1991, 100, 27-38. 140 A. Morrison, J. B. Bell, T. A. Kunkel, A. Sugino, Proc. Natl. Acad. Sci. U S A1991, 88, 9473.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

I335

11 Hydrolysis and Formation of C - 0 Bonds 11.1

Hydrolysis and Formation of Carboxylid Acid Esters HansJoachirn Gais and Fritz Theil

Catalysis of the hydrolysis and formation of the C - 0 bond of an ester, lactone or carbonate by hydrolases are amongst the most useful enzyme-catalyzed reactions in organic synthesis ( i n vitro) Today hydrolases are established tools for organic synthesis on a laboratory scale as well as on a industrial scale L3’1, The reason for this lies in the nature of hydrolases. Hydrolases are chiral catalysts, which are easy to function without a coenzyme, are commercially available in quite a number and and frequently feature low substrate specificity, high enantiotopic selectivity, and enantiomer selectivity. In addition, directed evolution, chemical modification and most importantly site-directedmutagenesis allow for the attainment of enzymes with improved activity, selectivity and stability, in particular toward organic solvents [35f]. This adds considerably to the versatility of hydrolases. The most important application of hydrolases lies in the field of asymmetric synthesis, which is therefore solely dealed with in this chapter. For the application of hydrolases in chemo- and regioselective transformations and in particular in protecting group chemistry, see Chapter 18. Among the hydrolases, the most widely used are, in first place, the lipases (E. C. 3.1.1.3),as for example pig pancreas lipase, Pseudornonas sp. lipases and Candida antarctica lipase (see Sect. 11.1.1.1.5, Tables 11.1-10to 11.1-25), in second place the carboxylesterhydrolases (E. C. 3.1.1.1), as for example pig liver esterase (see Sect. 11.1.1.1.1., Tables 11.1-1 to 11.1-6 and Sect. 11.1.1.2.3, Table 11.1-27),and in third place the proteases (E.C. 3.4.m.n), as for example subtilisin (see Sect. 11.1.1.1.4, Table 11.1-8 and Sect. 11.1.1.2.2, Table 11.1-26)and a-chymotrypsin (see Sect. 11.1.1.1.2, Table 11.1-7).Today, lipases are the most versatile hydrolases, primarily because of their ability to be highly active not only in water but also in water in the presence of an organic cosolvent and, most importantly, even in organic solvents of low water content. Another reason for the versatility of lipases is their accessibility in quite large numbers (see Sect. C). Some confusion has developed in the literature concerning the origin and names of some

336

I

1 1 Hydrolysis and Formation ofC-0 Bonds

CO,H

C0,Me

Scheme 11.1-1. Enantiotopos-differentiating hydrolysis of dicarboxylic diesters.

pig liver esterase HzO, PH 7

C0,Me

C0,Me

[37-42, 271

298% ee, 98% yield

Ph

1,

Ph C0,Me

I..'ACO,H

a-chymotrypsin

c

"AC0,Me Me

HZO, PH 7 [431

Me

C0,Me

298% ee, 95% yield

microbial lipases; this topic is dealt with in Sect. 11.1.1.1.5. Appropriate substrates for hydrolases are principally those compounds which bear enantiotopic ester groups with the prochirality contained either in the dicarboxylic acid (Schemes 11.1-1and 11.1-8)or in the diol part (Schemes 11.1-2 and 11.1-12) ofthe molecule, or those which carry enantiotopic hydroxyl groups (Schemes 11.1-3 and 11.1-12). A second and no less important class of substrates is the racemates, as for example esters of racemic carboxylic acids (Scheme 11.1-4) or esters of racemic alcohols (Schemes 11.1-5 and 11.1-7),racemic alcohols (Scheme 11.1-6),and racemic hydroxy carboxylic acid esters (Scheme 11.1-8). The hydrolase-catalyzedreactions utilized most for the selective transformation of such substrates are hydrolysis (Schemes 11.1-1,11.1-2, 11.1-4, 11.1-5 and 11.1-11), acylation (transesterification) (Schemes 11.1-3, 11.1-6 and 11.1-11)and alcoholysis (transesterification)(Schemes 11.1-7, 11.1-8 and 11.1-15). Hydrolase-catalyzedesterification of an alcohol with a carboxylic acid, although highly useful in some cases ["], has been utilized to a lesser extent. Catalysis of formation and cleavage of the C - 0 bond of an ester or lactone by pig liver esterase, most lipases, a-chymotrypsin and subtilisin, which are all serine hydrolases, involves the following steps (Scheme 11.1-9). Formation of an enzyme-substrate complex, attack of the hydroxyl group of

=%:

N3,,"

pig pancreas lipase H,O,pH7 1441

OAc

Pseudomonas cepacia lipase

OAc

HzO,pH7 145, 461

Scheme 11.1-2.

-

N3. I I

,coA OH

91% ee. 85% yield

OAc 96% ee, 85% yield

Enantiotopos-differentiating hydrolysis of diol diacetates.

7 7 . 7 Hydrolysis and Formation of Carboxylid Acid Esters

I

337

-

Pseudomonas cepacia lipase

/7/0yoH

vinyl acetate

uoLOAc

[471

298% ee, 78% yield

Pseudomonas cepacia lipase

*

vinyl acetate 97% ee, 77% yield

*P OAc

HO

1

Pseudomonas cepacia lipase

OCH,Ph

OCH,Ph vinyl acetate

/

"W

HO

HO 197% ee, 89% yield

Enantiotopos-differentiating transesteritication of diols.

Scheme 11.1-3.

/J3°f02Me

CI

~

\

\o

~

c

o

CI

87% ee, 49% yield

+

pig pancreas lipase

+

b

HzO, PH 7

[50-531

CI

299% ee, 49% yield Scheme 11.1-4.

Enantiotopos-differentiatinghydrolysis o f carboxylic acid esters.

the serine residue in the active site of the enzyme on the carbonyl group of the substrate or reagent with formation of a covalent acyl-enzyme-productcomplex and its transformation to the free acyl-enzyme and H-X-R2 [Eq. (l)]["3];reaction of the acyl-enzymewith a nucleophile, as for example water or an alcohol with formation of an acyl-enzyme substrate complex, which reacts with deacylation and formation of another enzyme product complex, which finally gives the free enzyme and the product [Eq. (2)]. The overall equilibrium, the attainment of which is catalyzed by the

z

H

338

I

1 1 Hydrolysis and Formation of C - 0 Bonds

299% ee, 43% yield

Pseudomonas sp. lipase

+

+

*

HZO, PH 7

WI SiMe, /

298% ee, 46% yield

OAc

OH

-

299% ee, 45% yield

Pseudomonas fluorescens lipase

+

H,O. PH 7

+

[551

OAc Scheme 11.1-5.

OAc 100% ee, 45% yield Enantiotopos-differentiating hydrolysis of acetates.

enzyme, is depicted in Eq. (3). All steps are in principle reversible. Formation of the acyl-enzyme and its reaction with a nucleophile involves the enzyme-bound tetrahedral intermediates A and B. In these processes a triad of three amino acids of the active site of the enzyme, Ser, His and Asp(Glu),which are specifically orientated in a three-dimensional way, together with other amino acids is involved. Crucial to the catalytic function of the enzyme are, besides the interplay of the residues of these amino acids, the stabilization of the oxy anion intermediates A and B and the corresponding transition states through hydrogen bonds provided by amide bonds or other amino acid residues of the active site. Hydrolysis of C - 0 bonds of esters and lactones [Eqs. (1)to (3), X = 0 and YR3 = OH] is usually carried out at room temperature in aqueous solution or in mixtures of water and either a water-miscible or water-immiscible solvent. Because of the large excess of water, equilibrium usually is mainly if not completely on the side of the

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters

Ph

I

339

82% ee, 46% yield

Pseudomonas sp. lipase

+

+

vinyl acetate tetrahydrofuran 1561

Ph

297% ee, 39% yield

OH

OH

YPh

Ph

* +

295% ee, 43% yield Pseudomonas sp. lipase

+

-

vinyl acetate [571

<

Ph

Ph

295% ee, 42% yield

OH

Pseudomonas cepacia lipase

+

OH 297% ee, 44% yield

+

c

vinyl acetate 1581

1 OH Scheme 11.1-6. of alcohols.

Me

phy$

Me'

Me

OAc

Enantiotopos-differentiating transesterification

297% ee, 26% yield

340

I

7 1 Hydrolysis and Formation of C-0 Bonds

(11

(11

OAc

I

I

Ph

OH

Ph

95% ee, 47% yield Pseudomonas cepacia lipase

+ OAc

+

nPrOH fed-pentyl alcohol [591

OAc

fl

(‘1

(11

OAc

OAc

I

I

Ph

Ph

Scheme 11.1-7. of acetates.

Enantiotopos-differentiatingalcoholysis

95% ee, 50% yield

-

Pseudomonas cepacia lipase Me 0

nBuOH diisopropyl ether [GO1

93% ee, 90% yield

74% ee

+

pig pancreas lipase z

+

diethyl ether [Ell OH

Scheme 11.1-8. Enantiotopos-differentiating lactonization of hydroxy esters.

298% ee

11.1 Hydrolysis and Formation ofcarbowylid Acid Esters

Rl-CO-X-R2

+ enzyme

R1-CO-enzyme + H-Y-R3

11

11 Rl-CO-X-R2

R1-CO-enzyme

enzyme

11

11 R1-CO-enzyme

H-Y-R3

Rl-CO-Y-R3

H-X-R2

enzyme

ci

11

enzyme + R1-CO-Y-R3

R1-CO-enzyme + H-X-R2

(Eq. 2)

(Eq.1) R1-CO-X-R2 + H-Y-R3

Rl-CO-Y-R3

-&-Asp-&

+ H-X-R2

(Eq.3)

-&Asp-&

f

f I

k

H-Nk

1

k

H-Nk

A

-R1C02H

6 Scheme 11.1-9.

Mechanism of hydrolase-catalyzed reaction.

carboxylic acid and the alcohol [Eq. (3), X, Y = 0, R3 = H] and the reaction is practically irreversible. Low solubility of liquid substrates normally presents no problem. In fact lipases are designed by nature to work at the liquid-liquid interface. In the case of crystalline and also liquid substrates of low solubility,solubility may be

I

341

342

I

7 7 Hydrolysis and Formation ofC-0 Bonds

enhanced by addition of an organic solvent. Most commonly used cosolvents are alcohols, as for example methanol and tert-butanol, and acetonitrile, acetone, tetrahydrofuran, ether, dimethylformamide and dimethyl sulfoxide[9, **, 30, 34, 361. The choice of the cosolvent, miscible or immiscible with water, depends on the enzyme. Lipases frequently show higher activity and selectivity in an emulsion of water and tert-butylmethyl ether or diethyl ether than in water. At higher concentrations of the organic solvent, the stability and the activity of the enzyme may be very low and the presence of the cosolvent can also alter the enantioselectivity of the enzyme in both directions. Many hydrolases, except lipases, show an interfacial deactivation. In aqueous solution, the parameters which influence the rate of hydrolysis most are the pH value and the temperature. In most cases, hydrolysis is run at room temperature at pH 7.0. However, at lower temperatures the selectivity of hydrolysis may be higher. For an easy recovery of the enzyme from the aqueous solution and for other purposes, it can be immobilized by various techniq u e ~ [ ~34, ' * 36. 64a1. One of the most popular for laboratory scale synthesis is the covalent immobilization of the enzyme on Eupergit C, a polymer with reactive epoxide groups. However, the nature of the polymer and the method of fmation may greatly influence the stability and activity of the immobilized enzyme. Another very useful technique, especially for synthesis on a larger scale, is the use of a continuous flow membrane reactor [64b]. Recovery of the enzyme from aqueous solution in smalland large-scale batch synthesis can be done by ultrafiltration [64c1. Finally, crosslinked enzyme crystals (CLECs), which are crystals of a pure enzyme cross-linked with glutaraldehyde, can be used for this and other purposes because of their insolubility in water and organic solvents L6&]. Formation of C-0 bonds of esters and lactones is accomplished by exploiting the transferase activity of hydrolases in acylation and alcoholysis (transesterification) reactions in organic media of low water content[30,34, 361. Especially lipases show a high activity in organic solvents, as for example ethers, hydrocarbons, chlorinated hydrocarbons, vinyl acetate, isopropenyl acetate and ethyl acetate. Lipases are thus by far the most useful hydrolases for asymmetric synthesis in organic media (Sect. 11.1.1.2.1). Subtilisin (Sect. 11.1.1.2.2.), pig liver esterase (Sect. 11.1.1.2.3) and ctchymotrypsin (Sect. 11.1.1.1.2), which also show activity in organic media of low water content, have been applied only to a lesser extent in asymmetric synthesis through C - 0 bond formation. Lipases, like other hydrolases, are generally not soluble in organic solvents. They rather form suspensions of protein aggregates. Enyzmes are only active if a certain amount of water, which is adsorbed or bound by the enzyme or dissolved in the organic solvent, is present. Organic solvents with a high solubility for water are generally not well suited for a hydrolase-catalyzed transesterification, perhaps because of a dehydration of the enzyme. It is frequently observed that the enantioselectivity of the acylation of an alcohol, catalyzed by a lipase, in an organic solvent is higher than the hydrolysis of the corresponding ester catalyzed by the same lipase in water. To provide for extreme equilibrium positions in acylation reactions of alcohols catalyzed by lipases in organic media, fatty acid trifluoro- or trichloroethyl esters, vinyl esters, oxime esters or carboxylic acid anhydrides have been used with much success. They provide for extreme equilib-

1 7 . 1 Hydrolysis and Formation ofcarboxylid Acid Esten

rium positions and practically irreversible reactions. Among these acylation reagents, vinyl esters are the most useful. However, the formation of acetaldehyde in transesterification with vinyl esters may cause a deactivation of the enzyme, most probably because of a reaction with its lysine amino groups[65,66]. Generally, the lipases used for the formation of C - 0 bonds under these conditions are crude preparations which may contain only a few percent of the actual lipase. These crude lipases usually also contain other proteins (which may be even other enzymes), additives such as carbohydrates, and salts, which stabilize the enzymes. Usually fair amounts of these crude materials are used as suspension in organic solvents together with the acyl transfer reagent. The water content of the system is in most cases only ill defined, and it can decrease through a competing hydrolase-catalyzed hydrolysis of the acylation reagent[65,“I. Because of the insolubility of hydrolases in most organic solvents, catalysis is carried out under heterogeneous conditions, which restrict not only the mobility of the enzyme but also of the substrate and products, and can thus cause mass transfer limitations. Immobilization on solid support, addition of hydrated salts, covalent attachment of methoxpoly(ethy1ene glycol) (MPEG) residues, lyophilization with organic polymers, cross-linking of enzyme crystals, sol-gel entrapment, coating with amphiphilic molecules and entrapment in reverse micelles are the most important techniques that have been shown to improve the performance of the enzyme[36].Some of these measurements serve to increase the surface area of the enzyme and thus to enhance its activity. Among the various types of hydrolase-catalyzed reactions, those involving the differentiation of enantiotopic ester groups (-COzR), acyloxy groups (-OCOR) and hydroxyl groups or the differentiation of the enantiomers of esters of racemic carboxylic acids and racemic alcohols are by far the most important ones. Under non-equilibrium conditions (e.g. hydrolysis in the presence of a large excess of water, alcoholysis with a large excess of an alcohol in organic solvents of low water content, or acylation with a vinyl ester in organic solvents of low water content), the decisive steps of differentiation are as follows. In the case of differentiation of enantiotopic ester groups or enantiomeric esters starting with the enzyme it may be the formation or cleavage of the acyl-enzyme; in the case of enantiotopic acyloxy groups or enantiomeric esters of racemic alcohols starting with the enzyme it may be the formation or break-down of intermediate A with generation of the acyl-enzyme; and in the case of enantiotopic hydroxyl groups or the enantiomers of racemic alcohols starting with the acyl enzyme it may be the formation or break-down of intermediate Awith formation of the enzyme. Numerous rneso-configured or otherwise prochiral substrates, preferentially containing enantiotopic methoxycarbonyl groups, have been converted by a pig liver esterase- or lipase-catalyzedenantioselectivehydrolysis in water to chiral monoesters (see Sect. 11.1.1.1.1.,Tables 11.1-1to 11.1-4 and Sect. 11.1.1.1.5, Tables 11.1-10 to 11.1-12). In nearly all cases investigated thus far the pig liver esterase-catalyzed hydrolysis of the substrate diester S terminates at the stage of the enantiomeric monoesters P and ent-P. In this case, where the products P and ent-P are not transformed further, the irreversible enantiotopos-differentiationmay be described by the process depicted in Scheme 11.1-10[G7“’].

I

343

344

I

I 1 Hydrolysis and formation ofC-0 Bonds

P

Hydrolase catalyzed enantiotoposdifferentiating irreversible S: substrate; P, ent-P: enantiorneric products; k,, kz: apparent first-order rate constants o f t h e irreversible process; E: selectivity factor; ee: enantiomeric excess. Scheme 11.1-10.

enf-P

kl

E=-=

k2

[PI

___

[ent-P]

E-1 ee(P)= __ E+l

Thus, in this case the ee value of the monoester P (or ent-P) is determined by the selectivity of enantiotopos-differentiation. It is not depended on the extent of the hydrolysis but only on the ratio of the two apparent first-order rate constants kl and k2 if one assumes an irreversible reaction and the absence of product inhibition, the former assumption being reasonable because of the large excess of water. In case of an insufficient degree of differentiation, selectivity can be raised only by a suitable temporary or permanent modification of the structure of the substrate S, by choice of another hydrolase, by addition of an organic solvent or by variation of the temperature or pH value, but not by stopping the reaction at various degrees of conversion. In practice the situation is somewhat different in the case of mesoconfigured or otherwise prochiral diacylated diols having enantiotopic acyloxy groups. For these diesters, hydrolysis by pig liver esterase and lipases in water usually does not stop completely at the stage of the enantiomeric monoester P and ent-P, but proceeds further - although at a significantly lower rate - to the achiral diol Q (Scheme 11.1-11)[67-G91. Thus enantiotopos-differentiation expressed through the apparent first-order rate constants kl and k2 is accompanied by an enantiomerdifferentiation as expressed by the apparent first-order rate constants k3 and k4. In this case the ee value of the monoester P (or eat-P) depends on the extent of the conversion of the diester S to the monoesters P and ent-P and of the conversion of the latter to the achiral diol Q, and thus on all four rate constants. From the fact that a hydrolase usually retains the ( R ) - or (S)-group preference of the enantiotopos differentiation in the enantiomer-differentiating hydrolysis, i. e. the hydrolysis of the faster formed monoester P to the diol Q is slower than the hydrolysis of the slower formed monoester ent-P to the diol Q ( k l > k2 and k4 > k j or vice versa), it follows that the ee value of the monoester P (or ent-P) can be raised upon carrying the hydrolysis further to the diol Q, at the expense of the yield. This can be advantageously used to raise the ee value of the monoester to the point where it can be isolated enantiomerically pure (for practical purposes). The diol can in most cases be converted to the diester. A mathematical model for the prediction of the ee value of the monoester and the quantity of the individual products in such a combined enantiotopos- and enantiomer-differentiating hydrolysis, which allows one to find the optimum in regard to the ee value and the yield, has been developed on the basis of an irreversible reaction and the absence of product inhibition (Scheme 11.1-11),[' 67-691 . Required are the kinetic constants a, E l and E2, which can be derived from a determination of

7 1 . 7 Hydrolysis and Formation ofcarbowylid Acid Esters

I

345

Scheme 11.1-11. Hydrolase catalyzed enantiotopos- and

enantiomer-differentiating irreversible transformationsL67-691.

[Q] = [So] - [S]- [PI - tent-PI [PI -[ent-PI ee (P) =

[PI + [ent-PI

100 90 80

4-

ent-P

[%I

60

50 40

30 20 10 0

ee[%I Figure 11.1-1. Dependence of ee value of monoester (P) on yield o f monoester in combined enantiotopos- and enantiomer-differentiation with different sets o f kinetic parameters.

346

I

7 7 Hydrolysis and Formation of C-0 Bonds

the amounts of S , P and ent-P as well as the ee values at various stages of the hydrolysisI7O1. The ee value of the monoester is a function of the conversion, which is generally expressed in curves as schematic depicted in Figure 11.1-1 for two sets of different kinetic constants a, E l and E2[67-691. The validity of this has been verified several times[']. A quite similar situation is encountered in the reverse hydrolysis, i. e. the hydrolase-catalyzedacylation of a prochiral diol with, for example, vinyl acetate in an organic solvent of low water content, conditions which render the reaction irreversible,with formation of a chiral monoester. Here the ee value of the monoester can also be raised at the expense of the yield through further acylation of the monoester with formation of the achiral diacylated diol. Normally and not surprisingly the hydrolase exhibits in the hydrolysis of the prochiral diacetate and in the acylation of the corresponding prochiral diol the same enantiotopic group recognition despite the fact that chemically different species are involved. This leads to the synthetically favorable situation that generally, through acylation of a prochiral diol in an organic solvent and hydrolysis of the corresponding diacetate in water, both enantiomers of the corresponding monoacetate are accessible with one enzyme (Scheme li.l-12)11-361.The validity of this approach has been demonstrated in numerous cases. Chiral monoesters, obtained either from a prochiral diol or diester, may be converted by a suitable series of chemoselective transformation to either enantiomer 401. of a given target compound (enantiodivergentsynthesis) (Scheme 11.1-13)[10~ Because of the results with numerous prochiral diesters and diols, which have been subjected successfully to hydrolase-catalyzed enantioselective hydrolysis and acylation, respectively, and because of the desire to predict the sense of the asymmetric induction in the conversion of a new substrate, active-site or substrate models have been developed for the hydrolases pig liver esterase [71-731,pig pancreas

pig pancreas lipase celite vinyl acetate

pig pancreas lipase H,O/ether pH 7.0

1 9"

Aco-LfoH

"U 298% ee, 89% yield

1141

93% ee, 86% yield (78% ee, without ether)

Scheme 11.1-12. Synthesis o f both

enantiomers of a monoacetate through transesterification and hydrolysis with a hydrolase.

-

7 7 . 7 Hydrolysis and Formation ojcarboxylid Acid Esters

I

347

1. CIC0,Et

acozH C0,Me

Scheme 11.1-13.

2. NaN,

NHCO,CHzPh

3. CH ,, A 4. PhCH,OH

C0,Me

I Kbutene

Synthesis o f both eantiomers from a given starting material (enantiodivergent synthesis).

C0,tBu

1

C0,Me NaOH

C0,tBu

1. CIC0,Et 2. NaN,

3. xylene, A

D

4. MeOH [39a-c]

aCOzH

-NHCO,Me

1. Na, EtOH, NH,

2. H+

I(yCOztBu

/

/

doo 298% ee, 82% yield

C0,Me 298% ee

3. H+ 1401

U

298% ee, 92% yield

lipase [741, Pseudomonas cepacia lipase r7', 761, Candida rugosa lipase [761, Candida antarctica lipase [77], Pseudomonas fluorescens lipase rS7, 78], Pseudomonas aeruginosa lipase [791, cholesterol esterase [761, subtilisin I,'[ and a-chymotrypsinL1, 'l]. The development of such models is greatly aided by X-ray crystal structure analyses of subtilisin a-chymotrypsin[831, Candida rugosa lipase, pig pancreas lipase, ["I Candida antartica lipase, '(lb] Pseudomonas cepacia lipase [871, and cholesterol esterase[86c].To a certain extent these models allow for a rationalization of the enantiotopic group and enantiomer preferences observed with the various substrates

348

I

11 Hydrolysis and Formation of C - 0 Bonds

and for a prediction in the case of new substrates. Interestingly, X-ray structure analyses show the active site of some lipases in the crystal to be blocked by a helical segment, called a lid or flap. In complexes of those lipases with transition state analogs the lid is opened, permitting access to the active site. Lipases in water usually show a lower activity toward water-soluble substrates than toward water-insoluble, liquid substrates. Thus, interfacial activation of lipases may be caused by a opening of the lid upon contact with a hydrophobic phase["]. One of the most valuable and much exploited features of hydrolases is their ability not only to differentiate between enantiotopic groups but also to differentiate between enantiomers [1-361. When, for example, a racemic alcohol or ester is subjected to a hydrolase-catalyzed acylation, alcoholysis or hydrolysis, respectively, a kinetic racemate separation (resolution)can take place, leading, if the process would be completely selective, at the point of 50 % conversion, to a mixture of the ester and the corresponding alcohol or acid of opposite configuration. In such a case both the unreacted enantiomer (substrate, S) and the newly formed ester, alcohol or acid (product, P) are enantiomerically pure, and their theoretical yield is 50% based on the racemic substrate. Hydrolase-catalyzedhydrolysis in water, acylation with vinyl acetate in an organic solvent of low water content and alcoholysis (provided that a large excess of alcohol is used) are, all three, practically irreversible, and the efficiency of the racemate separation only depends on the differentiation by the enzyme. When the selectivity of the enantiomer-differentiatinghydrolase-catalyzed transformation is insufficient, the enantiomeric purities of the product and of the unchanged substrate can be raised to a certain degree by changing the extent of conversion. Here too a mathematical model for the prediction of the ee value of the product and the unreacted substrate as function of the degree of conversion and the yield based on the simple classical homocompetitive model, assuming irreversibility and the absence of product inhibition, has been developed (Scheme 11.1-14) [S, 67-70]

By determining the E value from pairs of experimentally determined c and ee(P) values or c- and ee(ent-P) values, the ee values for the product and the substrate, depending on the degree of conversion, can be calculated and thus the optimum in terms of ee value and yield be f o ~ n d [ ~ ~ I . T hequations ree for E, called the enantiomeric ratio, allow one to calculate the inherent enantioselectivity of an enzyme, i. e. its ability to differentiate between enantiomers. Thus, E values can advantageously be used to compare the inherent enantioselectivitiesof different enzymes. E values are calculated by one of the equations of Scheme 11.1-14 on the basis of the determination of the conversion c and the enantiomeric excess ee of the remaining substrate or of the product. Alternatively, E values may be calculated on the basis of the ee values of both the remaining substrate and the product. Since ee values are often more accurately measured than conversion, the third equation is preferred[34]. It should be noted, however, that high E values (> 100)are less accurately determined than moderate E values, because of the enantiomeric ratio being a logarithmic function of the enantiomeric excess. Small changes in the measured enantiomeric purities gives large changes in the E values[34, Figure 11.1-2 indicates how to proceed practically in cases where the enzyme used exhibits only moderate selectiv-

7 1 . 1 Hydrolysis and Formation ofcarboxylid Acid Esters

I

349

k,

s

Scheme 11.1-14.

- P

+

k2

ent-S

E=

irreversible

Hydrolase-catalyzed enantiomer-differentiating c: conversion.

ent-P

[SI In [Sol [ent-S] In ___ [ent-So]

1 - ee (P)

E=

(1 + ee (P)] ; E= In[l - c (1 - ee (P)]

ln[l

-c

(for c <50%) c=l-

[S] + [ent-S] [So]+ [ent-So]

--

In[(l - c)(l - ee (S)]

1 + ee ( S )

In[(l - c)(l + ee (S)]

(for c >50%)

1 + (ee(S)/ee(P)

ee 6) ee(S)+ ee(P)

[PI -[ent-PI ee ( P ) =

[ent-S] -IS]

[PI + [ent-PI

ee (S) =

[ent-S]+ [S]

100

90 80 70 60

30 20

10

0 0

10

20

30

40

50

60

70

80

90

100

conversion [t]

Dependence of ee value of substrate (S) and product (P) on conversion in kinetic resolution with different E values. Figure 11.1-2.

1 1

1 + (ee(S)/ee(P) ; E=

350

I

I I Hydrolysis and Formation ofC-0 Bonds

R

O

L

C

I

+

R

88% ee, 43% yield or 295% ee

OH O A C

Scheme 11.1-15. Enatiomer preference in hydrolysis and transesterification by a hydrolase.

I

295% ee, 41% yield or 295% ee

Pseudomonas cepacia lipase H,O, pH 7.0 or nBuOH, diisopropyl ether

I

I

R= R

O

L

C

I

OH

+

R

O

A

C

I

Pseudomonas cepacia lipase vinyl acetate

or AqO, diisopropyl ether 1

OAc

R

O

L

C

I

295% ee, 47% yield [561

+

R

O

A

C

I

295% ee, 45% yield

ity. The product is isolated at the point of < 40% conversion and the substrate is isolated at > GO% conversion. In the case of not very high E values, the substrate but not the product can be obtained enantiomerically pure following this procedure. If the ee values of the thus obtained remaining substrate and the product are still not sufficiently high, both compounds may be isolated and subjected to another cycle of hydrolase-catalyzedtransformations, either hydrolysis or transesterification, guided by the above principles. Attempts have been made to carry out the second cycle of resolution without the isolation of the product and substrate obtained in the first cycle[”]. In the case of an insufficient selectivity of the resolution of Cz-symmetric substrates, the same approach as for substrates having enantiotopic groups can be applied (see above). For example, the Pseudomonas cepacia lipase-catalyzedhydrolysis of racemic trans-l,2-diacetoxycyclohexanein a three-phase system composed of

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters

I

351

water, n-hexane and sodium chloride gave the corresponding (R,R)-diolwith 2 99% ee in 42% yield and the (S,S)-diacetatewith 2 99% ee in 38% yield. Critical to the success of this resolution were the following three factors [921. The enzyme showed a preference for the acetoxy group attached to the (R)-centerin both the diacetate and in the monoacetate, the monoacetate was completely hydrolyzed to the diol, and an equalization of the rates of the hydrolysis of the diacetate and the monoacetate was be achieved through a favorable partitioning of both between water and n-hexane. A synthetically highly interesting method of converting a racemate completely to one enantiomer is dynamic kinetic resolution, a topic which is dealt with in Sect. 11.1.1.1.5 (Sect. 11.1.2.1.2,Table 11.1-24)on lipases. Kinetic resolution of alcohols and esters with hydrolases has opened up a new dimension for the synthesis of enantiomericallypure alcohols, esters and carboxylic acids, and in consequence the importance of resolution as a method for the attainment of enantiomerically pure compounds has been increased considerably. Hydrolase-catalyzed resolution is amenable to large-scale production [33-351, as was impressively demonstrated much earlier by the acylase-catalyzed racemate separation of N-acyl amino acids (not discussed in this chapter)[641. In lipase-catalyzed racemate separation of alcohols the same enantiomer preference is usually observed in acylation and hydrolysis (Scheme 11.1-15)[561. 11.1.1

Hydrolysis and Formation of Carboxylic Acid Esters 11.1.1.1

Hydrolysis of Carboxylic Acid Esters 11.1.1.1.1

Pig Liver Esterase

Pig liver esterase (PLE, E.C. 3.1.1.1) is one of the most useful hydrolases for the enantiotopos-differentiating hydrolysis of dicarboxylic diesters and diacetates of diols as exemplified by the hydrolysis of dimethyl cis-cyclohex-4-ene-l,2-dicarboxylate [37-421, which yields the corresponding cyclohexenoid monoester in nearly quantitative yield with an ee value of 2 98 % even on a 100 mol scale (Scheme 11.1-1 and Table 11.1-1)c4l1. Monoesters of the above type, which are in principle accessible by enzymatic hydrolysis on a large scale, are very useful chiral starting materials for 13, 271 the synthesis of biologically active natural and non-natural compounds 19, including p-amino acids [39a-dl. Up to now approximately 400 substrates for pig liver esterase have been described in the literature. Pig liver esterase, like other hydrolases, does not require a coenzyme, is commercially available, and often combines a low substrate specificity with high enantioselectivity. Pig liver esterase, which is a serine esterase, is isolated as a mixture of isoenzymes composed of the three subunits a (58.2 kDa), p (59.7kDa) and y (61.4 kDa), which behave more or less differently in regard to substrate specificity, pH dependence, inhibition or activation ~~~~~~~. by organic solvents or other compounds, and e n a n t i o s e l e c t i ~ i t yCommercially available are the natural isoenzyme mixture and several isoenzyme mixtures, enriched in one isoenzyme. These enzyme preparations may contain other proteins 9 '

352

I

7 7 Hydrolysis and Formation of C - 0 Bonds

and perhaps even other hydrolases. However, despite this variability in composition, the pig liver isoenzyme mixture has been applied with high success in almost all the cases reported. Even a rather crude acetone extract of pig liver, called pig liver acetone powder (PLAP), was successfully applied to enantioselectivehydrolysis. The cloning, functional expression and characterization of recombinant pig liver esterase has been de~cribed[’~”]. The recombinant pig liver esterase prepared by this method seems to be a single isoenzyme. It was reported that recombinant pig liver esterase, in the kinetic resolution of (l-phenyl-2-butyl)-acetate,shows a much higher selectivity than the isoenzyme mixture[’7b].Pig liver esterase frequently exhibits a reversal in enantiotopic selectivity such as changing from a (R)-centerto a (S)-centerester group preference of hydrolysis within a series of structurally closely related diester substrates. An active-site model for predicting the sense of the enantiotoposdifferentiation, which accounts for this reversal, has been p r o p ~ s e d [ ~ Usually ~-~~l. the best results are achieved with the dimethyl esters of dicarboxylic acids and with the diacetates of diols. Frequently the enantioselectivity of the hydrolysis and the yield of the monoester may be raised by changing the achiral alcohol component of the ester from methanol to ethanol or isopropanol. In the case of dicarboxylic diesters as substrates, pig liver esterase-catalyzedhydrolysis usually stops completely at the stage of the monoester formed. Further hydrolysis of the monoester to the dicarboxylic acid is in almost all cases thus far investigated extremely slow. This has been attributed to the presence of a charged group (carboxylate)in the molecule. In the case of the diacetates of diols, the rate difference is usually not so great. This, however, in the case of a moderate selectivity of the enantiotopos-differentiating hydrolysis, can be used to enhance the ee value of the monoacetate via a subsequent enantiomer-differentiating hydrolysis if the same stereochemical preference is maintained for the enantiomeric monoacetates; i. e., the faster formed enantiomeric monoacetate is more slowly hydrolyzed to the achiral diol, which is usually observed (Scheme 11.1-11).A mathematical model for the prediction of the ee value of the monoacetate in such a combined enantiotopos- and enantiomer-differentiating hydrolyses, which allows one to find the optimum in regard to the ee value and yield, has been developed for pig liver esterase-catalyzed hydrolyses [67-691. Pig liver esterase-catalyzed hydrolyses are generally carried out in aqueous phosphate buffer solution at pH 6-8 at room temperature. Equilibrium is under such conditions well on the side of the hydrolysis products because of the large excess of water. Normally, in the case of liquid substrates, low water solubility represents no problem. In the case of crystalline and liquid substrates of low solubility in water, up to 20% of organic cosolvents such as acetone, methanol, tert-butanol or dimethyl sulfoxide may It should be noted, however, that the enantioselectivity and the rate be added[9*8-’001. of the hydrolysis as well as the yield of the product may be influenced in either direction by organic cosolvents. Since pig liver esterase is a mixture of isoenzymes, it has been speculated that in the presence of organic cosolvents one or more isoenzymes might be deactivated. Organic cosolvents can in some cases be advantageously used to enhance the ee value of the chiral monoester or monoacetate. Pig liver esterase can be recovered from the aqueous solution by ultrafiltration[411 or by immobilizing the enzyme covalently on oxirane-activatedacrylic beads (Eupergit

7 7 . 7 Hydrolysis and Formation ofCarboxylid Acid Esten

Pig liver esterase-catalyzedenantiotopos-differentiating hydrolysis of prochiral cyclic dicarboxylic acid diesters in aqueous solution.

Table 11.1-1.

C0,Me

1 [1-4, 51

C0,Me Ph,ot,J

C ' 02H

C ' 02H

100% ee, 99 % yield

88 % ee, 99 % yield

a""."

C0,Me Me

U

2 [6, 71

~

Me .,,, Me

H

C ' 02H

91 % ee, 90% yield

a""."

Me ..,, Me

C0,Et

4 [I,3, 61

74 % ee, 95 % yield

5 [GI

45 % ee, 80 % yield

4

C0,Me

4

C0,Me

Ph,,,, Ph

[61

C0,Me

no hydrolysis

C0,nPr

Me .,,, Me

C0,nPr

7 [61

no hydrolysis

o
8 [4,8]

31 % ee, 65 % yield

H I N

occo2H 9 [91

C0,Me

10 [lo]

H02CP\C02Me

99 % ee, 90 % yield

92% ee

FO,CH,Ph N

C0,Me

11 [lo]

12 [l-4, 71

H02CP\C02Me

C '0 H .

38% ee

94 % ee, 98 % yield

13 [ll] IdCozMe -CO,H

doZH

86 % ee, 96 % yield

9 %, 80 % yield

C0,Me

I

353

354

I

II

Hydrolysis and Formation o f C - 0 Bonds

Table 11.1-1.

(cont.).

oaco C0,Me

15 [12, 71

C0,Me

16 [12, 131

88 % ee, 85 % yield

72 % ee, 83 % yield 82 % ee, 85 % yield, 10% MeOH 60% ee, 78% yield, 10% acetone

C0,Me 18 [13]

17 [12, 14)

CO,H

90% ee, 99% yield

22 % ee, 63% yield

c:aCozMe caCozH 20 1121

19 [12]

CO,H

C0,Me

48 % ee, 87 % yield (isolated as ketone)

0 % ee, 92 % yield (isolated as ketone)

Hoaco2H 22 [12]

21 [12]

C0,Me

22 % ee, 80 % yield

80 % ee, 76 % yield

23 (121

AcO

C0,Me

wCO,H 28% ee, 91 % yield

52% ee, 76% yield

26 (121

25 (121

58 % ee, 88 % yield

24 [12]

84% ee, 76% yield

7 1. 1 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.1-1.

(cont.).

73% ee

M

e

64% ee

T

xCO,H

29[14] 30 (151

0

CO,H C0,Me

CH,Ph

6% ee

38 % ee, 71 % yield

31 [15]

32 [15]

no hydrolysis

C0,Me

d

33 [15]

I

CO,H

C0,Me

34% ee, 82 % yield

qH

34 1161

42 % ee, 98 % yield

35 (161

36 117, 18, 191

C0,Me

46 % ee, 83 % yield

C0,Me

17%ee,85%yield 100% ee, 39% yield, 25% DMSO 100% ee, 10% MeOH 61 % ee, 10% DMSO 39% ee, 10% MeCN

I

355

356

I

I I Hydrolysis and Formation ofC-0 Bonds Table 11.1-1.

(cont.).

‘~‘‘,Cl ,+. CO,Me

Ho2C

N I CH,Ph

Ho2c””’,o \$..

37 [20]

Y

S0,Tol

27% ee. 96% conversion

39 [20]

Me%Yo2H

40 1211

Me

COPh

C0,Et 41 [22]

92;

42 [22]

HN >C02H

20 % ee, 83 % yield

6 % ee, 83 % yield

PhCH,N

CO,CH,CF,

295% ee, 62% yield

55 % ee, 16 % conversion 34% ee, 33% conversion

9 : ;

38 [20]

14% ee, 31 % conversion 11% ee, 88% conversion

44% ee, 28% conversion

S

CO,Me

44 11, 2, 7, 23,

43 [22]

241

10% ee, 63% yield

(Icoz 46 11, 23, 24,26,

45 [25]

27, 28, 291

C0,Et

C0,Me

298 % ee, 99 % yield

0% ee

47 [30]

aCozH 48 [30]

C0,nPr

25 % ee, 68 % yield

27 % ee, 67 % yield

49 [30]

a C O C0,iPr z H 2 % ee, 5 % yield

aoZH 50 [30]

C0,nBu

13% ee, 18 % yield

7 7 . 7 Hydrolysis and Formalion ofcarboxylid Acid Esters

I

357

Table 11.1-1.

(cont.).

C0,Me C0,Me

51 [12, 311

Mexo""~oH Me

CO,H

cC02H

295 % ee, 42 % yield

68 % ee, 84 % yield

C0,Me

:x;::+

: x;: : q

C0,Me

53 [34] C0,Me

Mexo,.,,(yH

54 [35]

CO,H

R = tBu no hydrolysis R = H 95 % ee

60% ee

55 [36]

0""

Me

52[32, 331

o,.-s

C0,Me 72 % ee, 86 % yield

96 % ee, 72 % yield (isolated as derivative)

C0,Me 57 191 O

a

O

z

no hydrolysis

98% ee, 82% yield

64% ee, 87% yield

C0,Me

58 [38, 391

75 % ee, 86 % yield

59 [38]

&02HC0,Me

&CO,H

H

4

C0,Me C0,Me

GO [38]

no hydrolysis

61 [38]

62 [40] Me 0 0% ee, 10% yield

358

I

11 Hydrolysis and Formation of C - 0 Bonds

(cont.).

Table 11.1-1.

no hydrolysis

36% ee, 77 % yield

Me*'OZH 0

65[43,44]

Me Hz'&' 0

66 [40,45]

CO,R

C0,Me 77 % ee, 96 % yield

R = Me

80% ee, 10% yield 100% ee, 37% yield R = nPr 45% ee, 15% yield R = iPr 39% ee, 22% yield R = nBu 73 % ee, 4 % yield R = Et

o&02H

67 [40, 45, 461

68 [47]

C0,Me

C0,Me 295 % ee, 99 % yleld

77% ee, 100%yield

C0,Et

GI) (481

O &

C0,Et

C0,Me 88%ee

65 % ee, 95 % yield

I

CO,R

71 [SO, 511

CiCO), R = Me 94% ee R=Et 99%ee 1 P. Mohr, N.Waespe-Sarcevic, C . Tamm, K. Gawronska, J. K. Gawronski, Helv. Chim. Acta 1983,66,2501. 2 G. Sabbioni, M. L. Shea, f. B. Jones,J . Chem. SOL., Chem. Commun.1984,236. 3 M. Schneider, N. Engel, P. Honicke, G. Heinemann, H. Gorich, Angew. Chem.1984,96, 55; Angew. Chem., lnt. Ed. Eng. 1984,23,67.

4 G.Sabbioni, I. B. Jones,J. Org. Chem.1987,52, 4565. 5 H.Ito, N . Imai, S. Tanikawa, S. Kobayashi, Tetrahedron Lett. 1996,37, 1795. 6 P. Walser, P. Renold, V.N'Goka, F. Hosseinzadeh, C.Tamm., Helv. Chim.Acta 1991,74,1941. 7 P. Renold, C.Tamm, Biocatal. Biotransform. 1995, 12,37.

7 7 . 1 Hydrolysis and Formation ofcarboxylid Acid Esters I359 30 K. Adachi, S. Kobayashi, M. Ohno, Chimia 1986, 40,311. 31 Y. Nagao, M. Kume, R. C. Wakabayashi, T. Nakamura, M. Ochiai, Chem. Lett. 1989, 239. 32 j. Zemlicka, L. E. Craine, M. J. Heeg, J. P. Oliver, J. 0%.Chem. 1988,53,937. 33 E. J, Hutchinson, S . M. Roberts, A. I. Thorpe, j . Chem. Soc., Perkin Trans. 1 1992, 2245. 34 I. C. Cotterill, P. B. Cox, A. F. Drake, D. M. Le Grand, E. J. Hutchinson, R. Latouche, R. B. Pettman, R. J. Pryce, S . M. Roberts, M. Stanley,j . Chem. SOC.,Perkin Trans. 1 1991, 3071. 35 M. Arita, K. Adachi, Y. Ito, H. Sawai, M. OhnoJ. Am. Chem. SOL 1983, 105,4049. 36 P. G. Hultin, F. J. Muesler, J. B. Jones,]. Org. Chem. 1991,56,5375. 37 T. Kuhn, C. Tamm, Tetrahedron Lett. 1989, 30, 693. 38 R. Bloch, E. Guibe-]ample, C. Girard, Tetrahedron Lett. 1985,26,4087. 39 G. Guanti, L. Banfi, E. Narisano, R. Riva, S. Thea, Tetrahedron Lett. 1986, 27,4639. 40 Y Ito, T. Shibata, M. Arita, H. Sawai, M. Ohno, J . Am. Chem. Soc. 1981,103,6739. 41 M. Ohno, Nucleosides and Nucleotides, 1985,4, 21. T. Norin, P. Szmulik, K. Hnlt, Bioorg. Med. Chem. 42 P. Metz, Tetrahedron 1989,45,7311. 1994, 2, 501. 43 H.-J. Gais, T. Lied, Angew. Chem. 1984, 96,495: 20 B. Danieli, G. Lesma, D. Passarella, A. Silvani, Angav. Chem., In#. Ed. Engl. 1984, 23, 511. Tetrahedron: Asymmetry 1996, 7, 345. 44 K. Adachi, S . Kobayashi, M. Ohno, Chimia 1986, 21 A. Salgado,T. Huybrechts, A. Eeckhaut, J. Van der 40,311. Eycken, 2. Szakonyi, F. Fulop, A. Tkachev, N. De 45 M. Ohno, Y. Ito, M. Arita, T. Shibata, K. Adachi, Kimpe, Tetrahedron 2001, 57, 2781. H. Sawai, Tetrahedron 1984,40, 145. 22 Y. Morimoto, K. Achiwa, Chem. Pharm. Bull. 1987, 46 S. Niwayama, S. Kobayashi, M. Ohno,J. Am. 35, 3845. Chem. SOC.1994,116,3290. 23 H:J. Gais, K. L. Lukas, Angau. Chem. 1984, 96, 47 S. Kobayashi, M. Sato, Y. Eguchi, M. Ohno, 140; Angau. Chem., Int. Ed. Engl. 1984,23, 142. Tetrahedron Lett. 1992, 33, 1081. 24 H:J. Gais, K. L. Lukas, W. A. Ball, S. Braun, H. J. 48 S. Niwayama, S. Kobayashi, M. Ohno, Tetrahedron Lindner, Liebigs Ann. Chem. 1986,687. Lett. 1988,29,6313. 25 F. Bjorkling, J. Boutelje, S . Gatenbeck, K. Hult, 49 I. C. Cotteril, S. M. Roberts, S. J. 0.William, T. Norin, Appl. Microbiol. Biotechnol. 1985,21, 16. J. Chem. Soc., Chem. Commun. 1988,1628. 26 S. Kobayashi, K. Kamiyama,T. limori, M. Ohno, 50 B.Malezieux, G. Jaouen, J. Salatin, j. A. S. Howell, Tetrahedron Lett. 1984.25, 2557. M. G . Palin, P. McArdle, M. OGara, D. 27 H. A. Sousa, J. G. Crespo, C. A. M. Afonso, Cunningham, Tetrahedron: Asymmetry 1992, 3, Tetrahedron:Asymmetry 2000, 11, 929. 375. 28 H. A. Sousa, C. A. M. Afonso, J. G . Crespo, 51 J. A. S. Howell, M. G. Palin, G. Jaouen, j . Chem. Technol. Biotechnol. 2000, 75,707. B. Malezieux, S . Top, J. M. Cense, J. Salauen, 29 H. A. Sousa, ). P. S . G. Crespo, Prog. Biotechnol. P. McArdle, D. Cummingham, M. O'Gara, 1998, 15,673. Tetrahedron: Asymmetry 1996,7, 95. 8 D. Habich, W. Hartwig, Tetrahedron Lett. 1987,28, 781. 9 P. Mohr, L. Rosslein, C. Tamm, Helu. Chim. Acta 1987.70, 142. 10 P. Renold, C. Tamm, Tetrahedron: Asymmetry 1993, 4, 2295. 11 I. Harvey, D. H. G. Crout, Tetrahedron: Asymmetry 1993.4, 807. 12 H.-J. Gais, B. Biilow, A. Zatorski, M. Jentsch, P. Maidonis, H. Hemmerle, J. Org. Chem. 1989, 54, 5115. 13 Y.Nagao, M. Kume, R. C. Wakabayashi, T. Nakamura, M. Ochiai, Chem. Lett. 1989, 239. 14 P. Renold, C. Tamm, Tetrahedron:Asymmetry 1993, 4, 1047. 15 S. Iriuchijima, K. Hasegawa, B. Tsuchihashi, Agric. Biol. Chem. 1982,46, 1907. 16 J. B. Jones, R. S . Hinks, P. G. Hultin, Can. I. Chem. 1985,63,452. 17 M. Kurihara, S. Kamiyama, S. Kobayashi, M. Ohno, Tetrahedron Lett. 1985, 26, 5831. 18 F. Bjorkling, J. Boutelje, H. Hjalmarsson, K. Huh, T. Norin, J. Chem. SOC.,Chem. Commun. 1987, 1041. 19 A. Mattson, J. Boutelje, I. Osoeregh, M. Hjalmarsson, U. Jacobsson, M. Lindbaeck,

C) [lol], BrCN-Sepharose['"], silica or in a hollow fiber ultrafiltration membrane[42].The Eupergit C immobilized pig liver esterase retains 68% of the specific activity of the soluble enzyme. It is easily removed by filtration from the reaction mixture and can be reused several times when stored at 7 "C. In large-scale experiments with pig liver esterase in aqueous solution the enzyme can be stabilized, if necessary, by the addition of inexpensive bovine serum For a determination of the ee value of the monoester, different methods can be used

360

I

7 7 Hydrolysis and Formation ofC-0 Bonds

addition of an enantiomerically pure chiral amine as for example ephedrine r401 or aphenylethylamine and 'H NMR spectroscopy of the diastereomeric salts formed thereby, conversion of the dicarboxylic acid monoester to the a-phenylethylamide and analysis by 'H NMR spectroscopy or HPLC1'041,or conversion to the tert-butyl ester and 'H NMR spectroscopy in the presence of a chiral shift Determination of the ee value of monoacetates has been carried out for example through conversion to the Mosher-esterand analysis by 'H NMR spectroscopy in the presence or absence of a shift reagent or by HPLC["'], or more directly either by 'H NMR spectroscopy in the presence of a chiral shift reagent or by HPLC and GC on chiral columns [Io7]. Cyclic dicarboxylic acid diesters, which bear enantiotopic ester groups, are substrates par excellence for a pig liver esterase-catalyzedhydrolysis under formation of the corresponding monoesters (1-71)(Table 11.1-1).The examples listed in Table 11.1-1are a good demonstration of the scope and limitation of pig liver esterasecatalyzed hydrolysis and illustrate general trends. The enantioselectivity and the yield can be influenced to a certain extent by the structure of the alcohol moiety of the diester as exemplified by the heterocyclic monomethyl and monopropyl esters 30 and 31.The corresponding isopropyl ester 32 is not a substrate for pig liver esterase. Extreme examples for the influence of the alcohol moiety are the cyclohexanoid methyl ester 44,which is formed with an ee value of 80% and the ethyl ester 45, which is produced as racemate. Usually the dimethyl esters are the best substrates. This trend, however, is not general. In the series of the cyclohexenoid monoesters 4650,the methyl ester 46 is the one formed with the highest ee value, and, in the series of the bicyclic monoesters 65 having a norbornene skeleton, it is the ethyl ester which has the highest ee value. However, the methyl ester 65 (R = Me) is formed much faster and is obtained in higher yield. Addition of an organic cosolvent such as dimethyl sulfoxide or methanol can lead to the monoester with a higher ee value. "his has been impressively demonstrated in the case of the benzyl-protected heterocyclic monoesters 36.A generalization of the effects of organic solvents upon the enantioselectivity of the PLE-catalyzed hydrolysis of diesters is difficult. As seemingly exemplified by the series of cyclopentanoid monoesters 14,15,17,19-29, small structural changes may invert the enantiotopic recognition through the enzyme. This can be advantageously used in certain cases to raise the ee value of a given target monoester by a suitable substrate modification. Branching in the aposition of the ester group is no prerequisite for high enantioselectivity. The monoesters lG and 18,the ester groups of which are separated by a methylene group from the ring, are obtained with comparable enantioselectivities. Interestingly, enantiotopic recognition is reversed in the series of diesters corresponding to the monoesters 15 and 16 but not with the diesters corresponding to the monoester 8,9, 17 and 18.This seemingly unpredictable behavior of pig liver esterase may at a first glance detract from its use in asymmetric synthesis. However, successful attempts have been made to rationalize this observation as well as the sense of asymmetric induction observed with the various substrates within an active-site model of the enzyme. Furthermore, one should bear in mind the experimental simplicity of a pig liver esterase-catalyzed hydrolysis and the synthetic advantages gained if the diester

I

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters 361 Table 11.1-2. Pig liver esterase-catalyzed enantiotopos-differentiating hydrolysis o f prochiral acyclic dicarboxylic acid diesters in aqueous solution.

A

R' CO,R~ R1

R'

R'

R2

1

2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 25 26 26 27 27 28 28 29 30 31 32 33 34 35 36 37 38 39 40 41

cop4 R4

ee (%)

yield VO) Ref.

73" 52" 19b 58" 46= 87= 88" 16" 16b

90-98 90-98 87 90-98 90-98 90-98 90-98 90-98 61 85-100 85-100

82'

93' 96

6 21 67 2

96 95 46b 84b 87b 96b 4Gb 84b no hydrolysis 88b 98b,' 15= 20

-

37 86 90 90 49

76 81 95 46 -

86 95 90-98

-

loa

90-98

8 25" 38 10" 5"

-

81

86b 92 82 78 97 97

-b

1Ob 80b

69

90-98 -

90-98

90-98 90 -

100 92 100 96 83 81 92 70 66

362

I

7 7 Hydrolysis and Formation of C-0 Bonds Table 11.1-2.

(cont.).

R'

cop3

i(CO,R~ - R'

R2

R3

42 43 44 45 46 47 48 49 50 51 52 53

ee (%)

yield (%)

80 78 69 79 CH3 CzHs C2HS C2HS H H H H

78 66 84 77 89 94 96 97 19 74 88 92

87 70 45 98 99 95 97 99

a In the presence of 25 % dimethylsulfoxide

b Absolute configuration not determined c In the presence of 50% dimethylsulfoxide

TR4

Rj

54 54 54 55 56 57 58 59 60 61 61

61 62 63 64 65 66 67 68 69 70 71 72

cop3

R'

R2

R'

R4

ee (%)

yield (?A)

Ref.

CH3 CH3 CH3 C2Hs n-C3H7 X3H7 G-CGHII C6H5 H C6HsCH2 C6HsCH2

H H H H H H H H p-FC6H4 H H

H H H H H CH3 H CH3 H CH3 CH3

CH3 CH3 CH3 CH3 CH3 H CH3 H CH, H H

90 79 92d 50 25

86 94

[20,211 [221 1221

C6HsCH2 C6H5(CHz)z CsHs(CHz)3 (E)-C~HSCH=CH-CH~ CGH~CH~O(CH~)Z (E)-HOCHz-CH=CH-CH2 (E)-THPOCHz-CH=CH-CHz (E)-CsHs-CH=CH NH2 CH3CONH C2HsCONH n-C3H7CONH

H H H H H H H H

CH3 CH3 CH3 CH3 CH3 CH3 CH3 CH3 H H H H

H H H H H H H H CH3 CH, CH3 CH3

H H H H

38 17 42

-

67 78 98 95 98 86 95

95 54 81d. e

-

73 44 88 88 54 18 74 93 41 93 6 15

98 98 97 95 100 100 95 100 94 81 50 52

1221 P I 1221 1221

P I 1231

1221 P I P I ~ 4 1 ~ 4 1 ~ 4 1 1241 ~ 4 1 1241 ~ 4 1 1251 ~ 5 1 1251 [251

1 1 . 1 Hydrolysis and Formation ofCarboxytid Acid Esters Table 11.1-2.

F+z:;:

(cont.).

R’ R’

R2

R3

R4

ee (%I

yield (“9) Ref.

73 74 75 76 77 78 79 80 81 82 83 84 85

~L-C~H~CONH i-C3H7CCONH (CHs)jCCONH c-C~HIICONH CH3CH=CH-CONH CbHsCONH CHz=CHCONH CbHsCONH CzHsCONH (CH,)3COCONH CbHsCHzOCONH C~HSCH~NH HO

H H H H H H H H H H H H H

CHj CH, CHs CH, CH3 CH, H CH, CH3 CH, CH, CH3 CH,

H H H H H H CH3 H H H H H H

2 93 79 100 72 8 20 40 53 93 33 15

48 55 50 52 60 59 50 60 70 93 93 58 100

86 87 88 89 90 91 92

HO COzC2H5 CsHsCHz0 CHsOCHzO CcjHsC00 CHjCOO CH30(CHz)zOCHzO H

CH3 OH H H H H H SCH2-p-CH30CrjH4

CH3 C2Hs CH, CH3 CH3 H H H

H H H H H CH3 CH3 CzHS

99 90 40 14 12 90 39 71

54

62 -

87 100 78 38 92 81

HoXcozH MeXcozMe 94 [20]

Me

CO,H

95 I201

HO

18% ee, 94 % yield

C0,Me

48% ee, 92 % yield

C0,Me 96 [20, 35, 361

97 [20]

CO,H Me

64% ee, 85 % yield d In the presence of 20% methanol; e At -10 “C

Me

98 % ee, 95 % yield

PI ~ 5 1 (251

PI (251 ~ 5 1 ~ 5 1 [251 1251 ~ 5 1 ~ 5 1 ~ 5 1 [18,2631, 321 1211 [401 [27, 321 [28] I281 [311 [27, 321

P I

I

363

364

I

11 Hydrolysis and Formation of C-0 Bonds Table 11.1-2.

(cont.).

99 1371

10% ee, 96% yield

‘C0,Me

79% ee, 70% yield

100 1381

82% ee

MeoXCo2Me

Me0

CO,H

101 1391

90 % ee, 90 % yield (20% MeOH)

102 1401

90-92% ee, 80-85 % yield 1 F. Bjorkling, J. Boutelje, S. Gatenbeck, K. Hult, T. Norin, P. Szmulik, Tetrahedron 1985, 41, 1347. 2 M. Ohno, M. Otsuka, Org. Reac. 1989, 37, 1. 3 M. Lyten, S. Muller, B. Herzog, R. Keese, Helv. Chim. Acta 1987,70,1250. 4 T. Kitazume, T. Sato, T. Kobayashi, J. T. Lin,J. Org. Chem. 1986,51,1003. 5 F. Bjorkling, J. Boutelje, S. Gatenbeck, K. Hult, T. Norin, Tetrahedron 1985, 26, 4957. G A. Fadel, J, L. Canet, J . Salaun, Tetrahedron Lett. 1989,30,6687. 7 M. Lyten, S. Muller, B. Herzog, R. Keese, Helv. Chim. Acta 1987,70, 1250. 8 H. Heidel, G. Hutmer, R. Vogel, G . Helmchen, Chem. Ber. 1994, 127, 271. 9 S. Mueller, A. Wolleb, L. Walder, R. Keese, Helv. Chim. Acta 1990,73,1659. 10 K. Osakada, M. Obana, T. Ikariya, M. Saburi, S . Yoshikawa, Tetrahedron Lett. 1987, 22,4297. 11 B De Jeso, N. Belair, H. Deluze, M. C. Rascle, B. Maillard, Tetrahedron Lett. 1990, 31, 653. 12 M. Schneider, N. Engel, H. Boensmann, Angew Chem. 1984,96, 54; Angew. Chem., In!. Ed. Engl. 1984,23, 66. 13 E. J. Toone, J. B. Jones, Tetrahedron: Asymmetry 1991,2, 1041. 14 K. Imchijima, K. Hasegawa, G. Tsuchihashi, Agnc. Biol. Chem. 1982,46, 1907.

15 M. Breznik, A. Mrcina, D. Kikelj, Tetrahedron: Asymmetry 1998,9,1115. 16 M. Breznik, D. Kikelj, Tetrahedron: Asymmetry 1997.8.425. 17 B. Klotz-Berendes,W. Kleemiss, U. Jegelka, H. J. Schaefer, S. Kotila, Tetrahedron: Asymmetry 1997.8, 1821. 18 R. N. Patel, A. Banejee, L. Chu, D. Brozozowski, V. Nanduri, L. J. Szarka, /. Am. Oil Chem. SOC. 1998,75,1473. 19 S. Kobayashi, M. Nakada, M. Ohno, Indian /. Chem., Sect. B 1993,328,159. 20 P. Mohr, N. Waespe-Sarcevic,C. Tamm, K. Gawronska, J. K. Gawronski, Helv. Chim. Acta 1983,66,2501. 21 P. Herold, P. Mohr, C. Tamm, Helv. Chim. Ada 1983,66,744. 22 L. K. P. Lam, R. A. H. F. Hui, J. B. Jones,I. Org. Chem. 1986,51,2047. 23 M. S. Yu, 1. Lantos, Z:Q. Peng, J. Yu, T. Cacchio, Tetrahedron Lett. 2000,41, 5647. 24 M. Nakada, S. Kobayashi, M. Ohno, Tetrahedron Lett. 1988,29, 3951. 25 K. Adachi, S. Kobayashi, M. Ohno, Chimia 1986, 40, 311. 26 P. Mohr, L. Rosslein, C. Tamm, Helv. Chim. Acta 1987,70, 142. 27 L. K. P. Lam, J. B. Jones, Can./. Chem. 1988,66, 1422.

11.1 Hydrolysis and Formation of Carbowylid Acid Esters 28 R. Roy, A. W. Rey, Tetrahedron Lett. 1987,28, 4935. 29 E. Baader, W. Bartrnann, G. Beck, A. Bergmann, H. Fehlhaber, H. Jendralla,K. Kessler, R. Saric, H. Schussler, W. Teetz, M. Weber, G. Wess, Tetrahedron Lett. 1988,29, 2563. 30 F. C. Huang, L. F. H. Lee, R. S. D. Mittal, P. R. Ravihmar, J. A. Chan, C. J. Sih, E. Capsi, C. R. Eck,]. Am. Chem. SOC.1975,97,4144. 31 E. Santaniello, M. Chiari, P. Ferraboschi, S. Trave, I . Org. Chem. 1988,53, 1567. 32 L. K. P. Lam, J. B. Jones,Can.]. Chem. 1988,66, 1422. 33 S . Kobayashi, K. Kobayashi, K. Hirai, Synlett 1999, 909.

34 S. Johnson, S. R. Kesten, L. D. Wise,]. Org. Chem. 1992,57,4746. 35 C. S. Chen, Y. Fujirnoto, C. J. Sih,]. Am. Chem. Soc. 1981, 103, 3580. 36 G. Guanti, L. Banfi, E. Narisano, R. Riva, S . Thea. Tetrahedron Lett. 1986,27,4639. 37 M. Mikolajczyk, P. Kielbasinski, R. Zurawinski, M. W. Wieczorek, J. Blaszczyk, Synlett 1994,127. 38 D. J. Horgan, J. K. Stoops, E. C. Webb, B. Zemer, Biochemistry 1969, 8, 2000. 39 H. I. Bestmann, U. Ch. Philipp, Angnu. Chem. 1991, 103,78 Angnu. Chem., Int. Ed. End. 1991, 30, 86. 40 R. Chenevert, B. Ngatcha, R. Tchedarn, S. Yannick, D. Goupil, Tetrahedron: Asymmetry 1998, 9,4325.

in question is a substrate. Cyclic diesters, the ester groups of which are not in the 1,2-position,seem to be less appropriate substrates for pig liver esterase (33-39 and 41-43). Finally, it seems noteworthy that transition metal complexes containing enantiotopic esters groups are also amenable to a highly selective pig liver esterasecatalyzed hydrolysis (71). Cyclic monoesters of Table 11.1-1,which can be obtained with other hydrolases as such or of opposite configuration, are contained in Tables 11.1-7 and 11.1-12. For synthetic and mechanistic reasons, a large number and variety of prochiral malonates have been subjected to pig liver esterase-catalyzed hydrolysis with formation of chiral malonates (1-53) (Table 11.1-2). Hydrolysis of dimethyl or diethyl malonates bearing a methyl group and another small alkyl or functionalized alkyl group leads preferentially to the monoester with the (S)-configuration. Upon an increase in the size of the second group, enantiotopic recognition is inverted and the monoester with the (R)-configurationis formed. Hydrolysis of dimethyl hydroxymethyl methyl malonate provides an excellent example of the strategy to enhance the enantioselectivity of pig liver esterase by the introduction of a protecting group on the substrate. While the parent compound itself yields the corresponding (S)configured monoester 13 with an ee value of only 6 %, the introduction of a tert-butyl or tert-butyldimethylsilylprotecting group allows the isolation of the corresponding (R)-configuredmonoesters (16 and 17) with ee values of 96% and 95%, respectively. An equally large number and variety of prochiral glutarates have been subjected to pig liver esterase-catalyzedhydrolysis with formation chiral glutarates (54-93) (Table 11.1-2). Among the synthetically most useful glutarates are the 3-amino-glutarates. The parent compound methyl amino glutarate 69 itself is obtained only with an ee value of 41%. The introduction of an amino protecting group improves the enantioselectivity of the hydrolysis of the corresponding diester dramatically. Pig liver esterase-catalyzedhydrolysis delivers methyl N-acetylaminoglutarate 70 of ( S ) configuration with an ee value of 93 % and methyl N-crotonylamino glutarate 77 of the opposite (R)-configurationwith an ee value of 100%.Thus, both enantiomers of methyl amino glutarate are accessible with one enzyme by a synthetically simple substrate modification. To a limited extent and with only moderate success, rneso-configured glutarates and succinates have been subjected to pig liver esterase-catalyzed hydrolysis with

I

365

366

I

I 7 Hydrolysis and Formation of C-0 Bonds Pig liver esterase-catalyzed enantiotopos-differentiating hydrolysis of prochiral cyclic diol diacetates in aqueous solution.

Table 11.1-3.

44% ee, 54% yield

37 % ee, 70 % yield

4 [l, 3,4-61

3 [l,21

OH 86% ee, 83 % yield

4 % ee, 44 % yield

5

PI

0+YH OAc

NI

6 [71

CH2Ph

90 % ee, 70 % yield

8 % ee, 40 % yield

0

7 (81 OH 13% ee, 75% yield

77% ee

CCC

-OH

10 [1,9]

55 % ee, 60 % yield 96 % ee, 78 % yield, tBuOH

4% ee, 31% yield

12 [lo]

11 (10, 111

OAc

OAc

13% ee, 75% yield

295 % ee, 80 % yield

1 1 . 1 Hydrolysis and Formation ofcarboxylid Acid Esten

Table 11.1-3.

(cont.).

,

M e a o !,.,NO H

13 [lo]

OAc 295% ee, 70% yield

me^

'"'NO,

vjn.NO

15 [lo] Me

16 [lo]

OAc

[ale - 1.3",60% yield

18 [lo]

O$oHN02

OAc

OAc

[a]o+ 14.7", 20% yield

295 % ee, 60 % yield

maoAc NO,

19 [lo]

@OAC

OAc

NO*

20 [lo]

OAc

no hydrolysis

no hydrolysis

FMe N3

14 [lo]

,

f ( eM

17 [lo]

SgoHNO,

NO,

OAc

[alD+ 9.8", 68 % yield

OH

86 % ee, 88 % yield

22 (121

OH

R=Me 99 % ee, 89 % yield 99% ee, 100% yield R = CHzPh R = CHzOMe 99% ee, 86% yield

OAc 23 (121

OH

99% ee, 89 % yield

367

EtoaoH 295 % ee, 60 % yield

OAc

I

24 [21

@OMe OH

99 % ee, 100% yield

368

e M&% )

I

I I Hydrolysis and Formation o f C - 0 Bonds Table 11.1-3.

(cont.).

"

25 [12]

HO

MxI~i~e

Me

26 [12]

OH

OAc

no hydrolysis

89% ee, 73 % yield OAc

OH

OH

OAc

Q

92 % ee, 100% yield

99 % ee, 70 % yield

R = nPr, 295 % ee, 83 % yield R = Me, slow hydrolysis, racem at e

87 % ee, 62 % yield

Me Me

v ..

32 [20]

31 [17, 18, 191 AcO

+OH Me Me

OCOR

298 % ee, 92 % yield

R = Me 96% ee, 86% yield R = Et 42% ee, 54% yield R = iPr 94% ee, 62% yield R = tBu 55% ee, 72% yield

P-4

0

-

34 [22]

33 [91 OAc

57% ee 68% ee 1 K. Laumen, M. Schneider, Tetrahedron Lett. 1985, 26, 2073.

2 J. B. Jones, C. J. Francis, Can. J . Chem. 1984, 62, 2578. 3 Y. F. Wang, C. S. Chen, G. Girdaukas, C. J. Sih, j . Am. Chem. Sac. 1984, 106,3695.

33 % ee, 52 % yield 47% diacetate 4 K. Laumen, M. Schneider, Tetrahedron Lett. 1984, 25, 5875. 5 Y. F. Wang, C. S. Chen, G. Girdaukas, C. J. Sih, F. S. Ciba, j . Am. Chem. SOL 1985, 11 I, 128. G K. Laumen, E. H. Reimerdes, M. Schneider, Tetrahedron Lett. 1985, 26, 407.

11.1 Hydrolysis and Formation ofcarboxylid Acid Esters 7 Y. F. Wang, C. J. Sih, Tetrahedron Lett. 1984, 25, 4999. 8 H. Suemune, T. Harabe, 2. F. Xie, K. Sakai, Chem. Pharm. Bull. 1988, 36,4337. 9 G. Guanti, L. Banfi, E. Narisano, R. Riva, S . Thea, Tetrahedron Lett. 1986, 27,4639. 10 M. Eberle, M. Egli, D. Seebach, Helu. Chim. Acta 1988, 71, 1. 11 D. Seeach, M. Eberle, Chimia 1986, 40,315. 12 M. Falk-Heppner,M. Keller, H. Prinzbach, Angew. Chem. 1989,101,1281:Angew. Chem., In!. Ed. Engl. 1989,28, 1253. 13 G. Baudin, B. J. Glanzer, K. S . Zwaminathan, H. Vasella, Heb. Chim. Acta 1988,71, 1367. 14 M. Carda, J. Van der Eycken, M. Vandewalle, Tetrahedron: Asymmetry 1990, I , 17.

15 H. Suemune, M. Takahashi, S. Maeda, Z. F. Xie, K. Sakai, Tetrahedron: Asymmetry 1990, 1, 425. 16 L. Dumortier, M. Carda, J. Van der Eycken, G. Snatzke, M. Vandewalle, Tetrahedron: Asymmetry 1991,2,789. 17 K. Naemura, N. Takahashi, H. Chikamatsu, Chem. Lett. 1988, 1717. 18 H. Estermann, K. Prasad, M. J. Shapiro, 0. Repic, M. J. Hardtmann, J. J. Bolsterli, M. D. Walkinshaw, Tetrahedron Lett. 1990, 31,445. 19 K. Naemura, R. Fukada, N. Takahashi, M. Konishi, Y. Hirose, Y. Tobe, Tetrahedron: Asymmetry 1993,4, 911. 20 A. Krief, D. Surleaux, N. Ropsan, Tetrahedron: Asymmetry 1993,4,289. 22 D. B. Berkowitz, J:H. Maeng, Tetrahedron: Asymmetry 19967,1577.

formation of the corresponding chiral succinates (94 and 95) and glutarates (96) (Table 11.1-2). Exceptions are methyl 3-hydroxy-2,4-dimethylglutarate (97) and methyl 1,2-dimethoxysuccinate (101),which can be obtained with ee values of 98% and of 90 %, respectively. Most interesting examples are the citric acid derivatives 87 and 102.While the pig liver esterase-catalyzedhydrolysis of triethyl citrate proceeded with high enantioselectivity but low regioselectivity, that of the diester derivative of 102 occurred with both high regio- and enantioselectivity. Acyclic monoesters ofTable 11.1-2, which can be obtained with other hydrolases as such or of opposite configuration, are contained in Tables 11.1-7 and 11.1-12. meso-Configured mono- and bicyclic diacetates, bearing primary or secondary acetoxy groups, are frequently hydrolyzed by pig liver esterase under standard Table 11.1-4. Pig liver esterase-catalyzed enantiotopos-differentiatinghydrolysis o f prochiral acyclic diol diacetates in aqueous solution.

Me

OAc

80 % ee, 36 % yield 95 % ee, 15% yield

29 % ee, 43 % yield 39 % ee, 5 4 % yield

Me

Me” 290% ee, 50 % yield 1 Y. F. Wang, C. S. Chen, G . Girdaukas, C. J. Sih, J. Am. Chem. SOC.1984,106, 3695.

2 H. Suemune, Y. Mizuhara, H. Akita, K. Sakai, Chem. Pharm. Bull. 1986,34, 3440. 3 D. Seebach, M. Eberle, Chimia 1986,40,315.

I

369

370

I

11 Hydrolysis and Formation of C-0 Bonds

conditions at pH 7.0 to give the corresponding monoacetates (1-34) in high yields and with high enantioselectivities(Table 11.1-3).The examples listed in Table 11.1-3 demonstrate once again the low substrate specificityof pig liver esterase. At the same time the series of nitro substituted cyclohexanoid mono and diacetates 11-20 reveals that seemingly small changes in the structure of the substrate can suppress hydrolysis. Strategies to improve the enantioselectivity of pig liver esterase-catalyzed hydrolysis of a dialkyl dicarboxylate are as follows: a synthetically tolerable and meaningful substrate modification in the dicarboxylic acid part, a modification of the alcohol part of the substrate or the addition of an organic cosolvent. These strategies can also be applied in the case of acylated prochiral diols. While the polycyclic monoester 29, which bears a butyryl group, is obtained with an ee value of 2 95 %, the derivative 29, which cames an acetyl group, is formed as racemate. Enantioselectivity in the pig liver esterase-catalyzedformation of the cyclohexenoid monoacetate 10 can be dramatically improved if the hydrolysis of the corresponding diacetate is carried out in the presence of tert-butanol.The heterocyclicmonoesters 31 demonstrate how, in the case of an diacylated diol, the achiral carboxylic acid part of the substrate influences the enantioselectivity of the hydrolysis. A series of highly functionalized cycloheptane derivatives (21-26) have been obtained through a pig liver esterasecatalyzed hydrolysis of the corresponding diacetates, with the same enantiotopic group recognition in all cases. Cyclic monoacetates of Table 11.1-3,which can be obtained with other hydrolases as such or of opposite configuration, are contained in Tables 11.1-9, 11.1-11 and 11.1-18.

Only a very few acyclic prochiral acylated diols have been subjected with moderate success to pig liver esterase-catalyzed hydrolysis with formation of the corresponding chiral monoacetates (1-3) (Table 11.1-4).For this kind of compounds, lipases are the hydrolases of choice. Acyclic monoacetates of Table 11.1-4,which can be obtained with other hydrolases as such or of opposite configuration, are contained in Tables 11.1-10and 11.1-17. Enantiomer-differentiating hydrolysis with pig liver esterase has, as with other hydrolases, become an important method for resolution (Table 11.1-5). Kinetic resolution of oxirane mono- and dicarboxylic acid esters with pig liver esterase proceeds effeciently with good selectivities, as demonstrated in the cases 14 and 15. Resolution is of course not restricted to enantiomers with central chirality.Axial and planar chiral racemic ester have been resolved with moderate to good results with pig liver esterase (33-36). Resolution of esters, the ester group of which is attached to a carbon atom bearing three other substituents, even when contained in a bi- or tricyclic ring system (67), represents no problem (Table 11.1-5).It seems interesting to note that these esters, which might be otherwise difficult to hydrolyze because of steric hindrance, are hydrolyzed readily via enzyme catalysis. The cyclopentanoid esters 65 and 66 nicely illustrate how a seemingly remote functional group can significantly influence the enantioselectivity.Pig liver esterases allow for the kinetic resolution of a-hydroxy acids (3-7) and a-amino acids (8-13) which have a quaternary Ca-atom.

11. 1 Hydrolysis and Formation ofCarboxylid Acid Esters Table 11.1-5. Pig liver esterase-catalyzed enantiomer-differentiating hydrolysis o f racemic carboxylic acid esters and lactones i n aqueous solution (HLE horse liver esterase).

Ho Me Me&CO,H

Me OH

e M,O C, , ) ,e M

low ee 88 % conversion

298 % ee, 12% yield

low ee 67 % conversion

94 % ee, 26 % yield

HO

1b 111

Me

& C H /O ,

low ee

94% ee, 11% yield

HO C O H

Et0,C

%

OH

&

Ph

Ph

75% ee 50 % conversion

64% ee 50% conversion

“0ywh PhL

Ph

38% ee

OH P h

40% ee

Ph “ O 80% ee

m

Ph

86% ee

HDXCo,H

7 b I21

Me

Ph

51% ee

40% ee

Ph J.

Me

Me HN ,

4b [2, 3,4]

CO,H

31%ee,6%yield

8a 151

5% ee, 67 % yield

I

371

I I Hydrolysis and Formation ofC-0 Bonds Table 11.1-5.

(cont.).

Et

9b 151 25 % ee, 28 % yield

10a [5]

-

Ph

10b [S]

H,NXCO,Et

95 % ee, 41 % yield

72% ee, 57% yield

I l b (51

l l a (51 20 % ee, 50 % yield nBu

nBu

Ph

97%ee, 31%yield

93 % ee, 41 % yield

ph-Ah HN ,

CO,H

Ph-

13a [5]

14a [GI

&CO,H

15a [7] Me0,C 97 % ee, 40 % yield 50 % conversion

-C 9. O,C,H,

141, [6]

9.

MeO,C-

16a [8]

Me

Me

E = 17

17a [9] z

H

L

Me C

Me 0 2

OH

64% ee

50% conversion

15b[7]

16b (81

Me&CO,H

E=17

COHO

C0,Me

: ',

295 % ee, 40 % yield

+co2Me

G

13b [5]

H,NXCO,Et

82 % ee, 40% yield

74% ee, 30% yield 50% conversion

o.,,

Ph

61 % ee, 10% yield

54% ee, 39% yield

Me

121, [5]

12a (51

HN , /C ' O,H

63 % ee

H

171, [9]

7 1 . I Hydrolysis and Formation ofCar6oxylid Acid Esters Table 11.1-5.

(cont.).

18a [lo]

O_o~CozH

18b [lo]

\

7 % ee

10% ee

19a [lo]

19b [lo]

&oxc"2H \

5% ee

5% ee

5% ee

G%ee

26% ee

22% ee

22a [lo]

[lo]

boxco2 22bH

CI

CI

11%ee

10% ee

24 [ll] Et

( M ~ o c H , c H , ) , N ~Me0 M~ \e

/5

299 % ee, 25-30 % yield

nPr

299 % ee, 25-30 % yield

M e O + W I

\ /

(MeOCH,CH,),N~" Me0 299% ee, 25-30% yield

Me0

I

373

374

I

7 I Hydrolysis and Formation ofC-0 Bonds Table 11.1-5.

(cont.).

0 II

tBu

/s-CO,Me

26a [12]

::

.,,S-CO,Me

271, [12]

27a [12] 34 % ee, 40 % yield

21% ee, 52% yield 0 II

28a [12]

C O ,M $ 3 ,e To1 '

C M O ,eS ,, Tot

R

28b [12]

46% ee, 58% yield

80 % ee, 32 % yield

0 Ph .,,I1

26b [12]

38% ee, 38% yield

48 % ee, 53 % yield

Ph

0 I1 tB"'S-CO'H

29b [13]

29a [131

Me/P-CozMe 82 % ee, 42 % yield

73% ee, 50% yield

30b [13]

30a [ 131 81 % ee, 41 % yield

96% ee, 45 % yield

31a [13]

4

HC O ,, ,P; ' Ph

R

31b [13]

18 % yield

299 % ee, 40 % yield

32b (131

32a [I31 79% ee, 43%yield

80 % ee, 46 % yield

Ph.,;/CO,Me 33a [14]

f7

Me

Ph

Et

L?

r7C0,Me Et

96 % ee, 82 % yield

Me

61 % ee, 50% yield

90% ee, 33 % yield Ph

33b [14]

34a [15]

C0,Me

L:.

Etf E't 83 % ee, 80 % yield

34b [15]

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.1-5.

(cont.).

Fe

70% ee

75% ee C0,Me

//c”

36a [14]

@

36b [14]

Fe(CO),

85% ee 40 % conversion

85% ee GO % conversion

n 37 [18]

86% ee 25 % conversion

38a [19] M Me e$

38b [19]

CO,H

C0,Me Me

36% ee

HO,C

40% ee

ye

39a 1191

C0,Me

Me0,C’

60% ee

x

C0,Me

50% ee, 50% yield

A C H N . , , , ~.CO,H

-

40a [20, 21, 221

AcHN--sr C0,Me

97 % ee, 47 % yield

87 % ee, 43 % yield

a:

a

CO,H

Me

297 % ee

39b [19]

40b [20,21, 221

C0,Me

41a [23]

Me

297 % ee

41b (231

I

375

376

I

I 1 Hydrolysis and Formation ofC-0 Bonds Table 11.1-5.

(cont.).

50 % ee, 50 % yield

50% ee, 47 % yield

OTMe

44a [25]

441, [25]

0 o,COzEt

72% ee, 53% yield

96 % ee, 42 % yield

&OZEt''-.R 45 46 47 48 49

R=Me R=Et R = n-CsH1, R = (CH&CH=CH* R = ~C9H19

0% ee 93 % ee, 6G % yield 97% ee, 50% yield 99 % ee, 60 % yield 81 % ee, 62 % yield

51 [27]

LJ 299 % ee, 88 % yield

70 % ee, 10% yield

53 [27]

299 % ee, 40% yield

299 % ee, 58 % yield

299% ee, 60% yield

7 7. I Hydrolysis and Formation ofCarboxylid Acid Esters Table 11.1-5.

(cont.).

56a [28]

56b [28]

&Me OMe 93 % ee, 36% veld

32 % 0

5% [28]

57a [28]

-0Me 96 % ee, 42 % yield, HLE

34%, HLE

0 \LOH

58b [28]

58a [28]

0*OMe

0

30 %

94% ee, 36% yield

OMe

0*OMe

92% ee, 38% yield, HLE

a"""

34% yield, E = 65, HLE

60a [23]

Me

a::

0:: C0,Me

Me

17% ee

22% ee

61b [23]

61a [23]

o.,, 297 % ee

297 % ee

....C0,Me

62a 1231

Me

297 % ee

601,[23]

297 % ee

62b 1231

I

377

I I Hydrolysis and Formation of C - 0 Bonds Table 11.1-5.

(cont.).

O"" +..C0,Me

63b (231

CH,Br

297 % ee

297 % ee

O,-.,Br ....CO,Me

e

0

o

64a [23]

64b [23]

297 % ee

297 % ee

C0,Me 65a [29]

C0,Me 49% ee, 54% yield

59% ee, 43 % yield

<.

65b [29]

C0,Me

CO,H

661, [30]

GGa 1301

C0,Me

O = C l C0,Me

95 % ee, 45 % yield

95 % ee, 34% yield 50 % conversion

671,[31] 0 299 % ee, 40 % yield

73 % ee, 45 % yield

68a [32, 33, 341

C0,Me

C0,Me

82 % ee, 45 % yield

68b [32, 33, 341

C0,Me 70% ee, 45 % yield 32 % ee, 95 % yield

73 % ee, 45 % yield 64 % conversion 53 % ee, 55 % yleld

WH

C0,Me

69a [32, 331

C0,Me C0,Me 90% ee, 40% yield

69b [32, 33)

11.1 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.1-5.

(cont.).

WozH

70a [32, 331

C0,Me C0,Me

C0,Me

82 % ee, 45 % yield

70b [32,33]

95 % ee, 40 % yield

71a [35]

A

71b [35b]

Et02C&o

72b [36]

C0,Me

CO,H 36 % ee, 40 % yield

57 % ee, 48 % yield

72a [3G]

nPen 47 % ee, 35 % yield

96 % ee, 35 % yield

73 [37]

98% e, 41 % yield, HLE

(yJ: j,

80 % ee, 42 % yield, HLE

75 [37]

HLE

H

GO% ee, HLE

76 [37] HLE

47 % ee, 40% yield, HLE

&

74 [37]

95% ee, 34% yield, HLE

77 [37]

I

379

380

I

1 I Hydrolysis and Formation ofC-0 Bonds Table 11.1-5.

(cont.).

R

78 79 80 81

82 83 84

R=Et R=Pr R=nBu R=nPen R=nHex R=nHep R=nOct

PLE 98% ee 62% ee 88% ee 77% ee 33% ee 60% ee 65% ee

R R R R R R R

HLE 22 4 38 53 90 60 63

R

R S S S S S

[38] [38] [38] [38, 391 [38] 1381 [38]

85b [40]

85a [40]

46 % ee, 79 % yield, PLE 64% ee, 83% yield, HLE

70% ee, 97 % yield, PLE 95 % ee, 90% yield, HLE

A

OH

86a [40, 38)

HO,C(CH,),

44% ee, 78% yield, PLE 47 % ee, 84 % yield, HLE

83 % ee, 86 % yield, PLE 76% ee, 80% yield, HLE

87a (401

6

M

86b 140, 38)

Me

HO,C(CH,),

e

pr ”

8% [40]

Me

84% ee, 65% yield, PLE 42% ee, 71% yield, HLE

295% ee, 70% yield, PLE 295% ee, 74% yield, HLE

?H 88b [40]

88a [40]

G

M

e 295 % ee, 80% yield, PLE 295% ee, 84% yield, HLE

295% ee, 78% yield, PLE 295% ee, 94% yield, HLE

Go Me

299 % ee, 88 % yield, HLE

89a [40]

HO,C(CH,),

JH

89b (401 Me

299 % ee, 86 % yield, HLE

11.1 Hydrolysis and Formation ofCarboxylid Acid Esters

(cont.).

Table 11.1-5.

91 [37]

90 [37]

uC

7 H15

92% ee, HLE

78% ee, HLE 18% ee, PLE

M 0 e& Me

C0,Me

92a [41]

82 % ee, 32 % yield

54% ee, 25% yield

FO,CH,Ph

H

93a [42] Me0,C

5

Me0,C

C0,Me

C0,Me

28% ee

27% ee

94a [43]

k

94b [43]

Ph

Ph

>94% ee, 90% yield

mo

93b [42]

after lactonization 94% ee, 80 % yield

0

95 [43]

[a];' + 31.9 290 % ee

\y/

0

NMe

/ '-. R CH,CO,Me

96a [44]

0 NMe \\ I/ .-.S \ d CH,CO,H

96b [44]

R = Ph, 10% ee, 45 % yield R=Tol, 13%ee, 70%yield

R = Ph, 12% ee, 25% yield R = Tol, 290 % ee, 12% yield

NH \\ 4 .S .\ R' Me 0

/ Me

5

R

R = Ph, 14% ee, 50% yield

97a [44]

R = Ph, 41 % ee, 12% yield

97b [44]

I

381

382

I

I I Hydrolysis and Formation ofC-0 Bonds Table 11.1-5.

(cont.).

98a [45]

98b [45]

95% yield [aIDzs-15.4(1.02,MeOH)

99b [46]

295% ee 1 W. K. Wilson, S. B. Baca, Y. J. Barber, T. J. Scallen, C. J. Morrow,]. Org. Chem. 1983,48,3960. 2 H. Moorlag, R. M. Kellogg, Tetrahedron: Asymmetry 1991, 2, 705. 3 H. Moorlag, R. M. Kellogg, M. Kloosterman, B. Kaptein, J. Kamphuis, H. E. Schoemaker, J . 0%.Chem. 1990,55,5878. 4 H. E. Schoemaker, W. H. J. Boesten, B. Kaptein, E. C. Roos, Q. B. Broxterman, W. J. J. van den Tweel, J. Kamphuis, Ada, Chem. Scand. 199650, 225. 5 B. Kaptein, W. H. J. Boesten, Q. B. Broxterman, P. J. H. Peters, H. E. Schoemaker, J. Kamphuis, Tetrahedron: Asymmetry 1993,4, 1113. 6 D. Bianchi, W. Cabri, P. Cesti, F. Francacalanci, M. Ricci,]. Org. Chem. 1988, 53, 104. 7 P. Mohr, L. Rasslein, C. Tamm, Helv. Chim. Acta 1989, 30, 2513. 8 P. Mohr, L. Roesslein, C. Tamm, Tetrahedron Lett. 1989,30, 2513. 9 C. S. Chen, Y. Fujimoto, G. Girdaukas, C. J. Sih, 1. Am. Chem. SOC.1982,104,7294. 10 R. Chenevert, 1.DAstous, Can.]. Chem. 1988,GG, 1219. 11 D. J. Bennett, K. 1. Buchanan, A. Cooke, 0. Epemolu, N. M Hamilton, E. J. Hutchinson, a. Mitchell,]. Chem. Soc., Perkin Trans. I 2001,4, 362. 12 P. Kielbasinski, Tetrahedron: Asymmetry 2000, 11, 911. 13 P. Kielbasinski, R. Zurawinski, K. M. Pietrusiewicz, M. Zablocka, M. Mikolajczyk, Tetrahedron Len. 1994, 35,7081. 14 M. Pietzsch, 0. Vielhauer, D. Pamperin, B. Ohse, H. Hopf,]. Mol. Catal. B: Enzym. 1999,G, 51.

15 S. Ramaswamy, R. A. F. Hui, J. B. Jones,]. Chem. Soc., Chem. Commun. 1986,1545. 16 T. Izumi, T. Hino, A. Ishihara,]. Chem. Technol. Biotechnol. 1993, SG, 45. 17 N. W. Alcock, D. H. G. Crout. C. M. Henderson, S. E. Thomas, ]. Chem. Soc., Chem. Commun. 1988, 746. 18 C. Ariente-Fliche, J. Braun, F. Le Goffic, Synth. Commun. 1992,22,1149. 19 M. Schneider, N. Engel, H. Boensmann, Angew. Chem. 1984,96,52;Angew. Chem., In!. Ed. Engl. 1984, 23,64. 20 C. Sicsic, M. Igbal, F. Le Goffic, Tetrahedron Lett. 1987,28,1887. 21 R. Csuk, P. Doerr, Tetrahedron: Asymmetry 1994.5, 269. 22 R. Dernoncour, R. Azerad, Tetrahedron Lett. 1987, 28, 4661. 23 E. J. Toone, J. B. Jones, Tetrahedron: Asymmetry 1991, 2, 207. 24 A. Basak, T. Mahato, G. Bhattacharya, B. Mukherjee, Tetrahedron Lett. 1997, 38,643. 25 M. Tarkov, M. Bolli, B. Schweizer, C. Leumann. Helu. Chim. Acta 1993,76,481. 26 B. Westermann, 1. Kortmann, Biocatalysis 1994, 10, 289. 27 B. Westermann, H. Scharmann, 1. Kortmann, Tetrahedron: Asymmetry 1993,4, 2119. 28 C. Tanyeli, B. Sezen, A. S. Demir, R. B. Alves, A. Arseniyadis, Tetrahedron:Asymmetry 1999, 10, 1129. 29 Y. Morimoto, Y.Terao, K. Achiwa, Chem. Pharm. Bull. 1987, 35, 2266. 30 H. Suemune, M. Tanaka, H. Obaishi, K. Sakai, Chem. Pharm. Bull. 1988,3G, 15.

11. I Hydrolysis and Formation ofcarboylid Acid Esters I383 31 A. J. H. Klunder, W. B. Huizinga, A. J. M. Hulschoff, B. Zwanenburg, Tetrahedron Lett. 1986, 27, 2543. 32 A. J. H. Klunder, F. J. L. Van Gastel, B. Zwanenburg, Tetrahedron Lett. 1988,29, 2697. 33 F. J. C. Van Gastel, A. J. H. Klunder, B. Zwanenburg, Red. Trav. Chim. Pays-Bas 1991 110, 175. 34 J . Van der Eycken, M. Vandewalle, G . Heinemann, K. Laumen, M. P. Schneider, J. Kredel, I. Sauer, ]. Chem. SOC.,Chem. Commun. 1989,306. 35 C. H. Tran, D. H. g. Crout, J. Chem. SOC.,Perkin Trans. 11998, 1065. 36 S. Drioli; F. Felluga; C. Forzato; P. Nitti; G . Pitacco; E. Valentin]. Org. Chem. 1998,63, 2385. 37 C. Guibe-Jampel,G . Rousseau, L. Blanco, Tetrahedron Lett. 1989, 30,67.

38 R. Fellous, L. Lizzani-Cuvelier,M. A. Loiseau, E. Sassy, Tetrahedron: Asymmetry 1994, 5, 343. 39 L. Blanci, E. Guibe-Jampel, G . Rousseau, Tetrahedron Lett. 1988,29, 1915. 40 E. Fouque, G. Rousseau, Synthesis 1989,661. 41 D. Moelm, N. Risch, Liebigs Ann. 1995,1901. 42 P. Renold, C. Tarnm, Tetrahedron: Aqmmety 1993, 4, 2295. 43 P. Barton, M. I. Page,]. Chem. SOC., Perkin Trans. 2 1993, 2317. 44 P. Kielbasinski, Polish]. Chem. 1999,73,735. 45 G . H. Hakimelahi, N.-W. Mei, A. A. Moosavi-Movahedi,H. Davari, S . Hakimelahi, K.-Y. King, J. R. Hwu, Y.3.Wen, J . Med. Chem. 2001,44,1749. 46 T. G . Back, K. Nakajirna,I. Org. Chem. 1998,63, 6566.

Phosphorus- and to a lesser extent sulfur-containing racemic esters could be resolved with pig liver esterase (29-32 and 26-28) as well. Particularly interesting examples are the P-keto esters (45-54), which were frequently obtained with high enantioselectivities.The enantiomer, which was hydrolyzed preferentially, suffered a decarboxylation with formation of the corresponding ketone. Interesting examples, showing the strategy in the application of hydrolases in kinetic resolution, are the cyclohexadiene-carboxylates5 6 5 9 . The use of both pig liver esterase and horse liver esterase (HLE) allows for the attainment of both enantiomers of both the acid and the ester. In the resolution of the racemic esters 60b-G2b,pig liver esterase shows in regard to the configuration of the C-atom to which the ester group is bound the same preference as in the case of the corresponding rneso-diesters (Table 11.1-1).This also holds true for the resolution of the racemates of esters 63b and 64b, which carry a bromomethyl group instead of the methyl group. Here, the enantiomer, which carries the carboxy group, suffers lactonization with formation of lactones 63a and 64a. A most remarkable example of a pig liver esterase-catalyzed reaction is the apparent enantioselective hydration of alkene 98a with formation of the hydroxy lactone 98b. The mechanism of this reaction is not known. It seems interesting to note in this context that reaction of 98a with aqueous ammonia affords racemic 98b. Finally, the pig liver esterase-catalyzedhydrolysis of the racemic diester derivativesof 99a and 99b shows the remarkable feature that the two enantiomers are hydrolyzed with different regioselectivities. A large number of mono- and bicyclic lactones (73-91)have been obtained by using pig liver esterase in combination with horse liver esterase for the enantiomerdifferentiating hydrolysis of the corresponding racemic lactones. Interestingly, in the series of methyl-substituted lactones (85-89), both enzymes show toward the sevenmembered lactone (86) the opposite enantiomer selectivity as compared to the other lactones. Acids or esters of Table 11.1-5, which can be obtained with other hydrolases as such or ofopposite configuration, are contained in Table 11.1-13. Kinetic resolution by pig liver esterase is not restricted to mono- and dicarboxylic acid ester derivatives. Acetates of racemic secondary alcohols are also excellent

384

I

I 1 Hydrolysis and Formation ofC-0 Bonds Table 11.1-6. Pig liver esterase-catalyzed enantiomer-differentiating hydrolysis of esters of racemic alcohols i n aqueous solution.

Ph

l b [l, 21

l a (1,21

( Y O A c

98 % ee, 53 % yield

98 % ee, 40% yield

0

CMe,Ph

CMe,Ph

<$..OH

G O A C 95% ee, 44% yield

67 % ee, 36% yield

tBu 3a [31

UH 299 % ee after crystallization

299 % ee after crystallization

OCOnPr

72 % ee, 99 % yield

93 % ee, 96% yield

5a [41

nPrCOO

A/

94% ee, 77% yield

86 % ee, 79 % yield

76% ee

62% ee

3b [31

I

J 7. J Hydrolysis and Formation ofcarboxylid Acid Esters 385

Table 11.1-6.

(cont.).

4 \q \q o\ 6 fiMe Q"' Me OH

98% ee

Me OH

ACO Me

98% ee

OH

Me

Me

295 % ee, 12% yield

295 % ee, 28 % yield

OAc

10a [7]

10b [7]

Me Me

M& h e

41 % ee

100% ee

Ilb (81

l l a [8]

MeJ o H f Me 295 % ee, 43 % yield 43% conversion

0U

M

288% ee, 40% yield 43% conversion

e

12a [9]

12b [9]

CH,

27% ee

CH,

94% ee, 32 % yield

386

I

7 7 Hydrolysis and Formation ofC-0 Bonds

Table 11.1-6.

(cont.).

13a [lo]

8

13b [lo]

91% ee, E = 14

'"'Ph

14a [ l l , 121

14b [ l l ,121

A c~' 85% ee

84% ee

15a [ll, 121

151, [ l l ,121

'"'OH dlAC

OH

82% ee

78% ee

OCOPh 16b 1131

16a [13] I

I R

R 100% ee, 69% yield 95 % ee, 40% yield

99 % ee, 55 % yield 40% ee, 38% yield

OCOPh 17a [13] I Me

Me 97 % ee, 45 % yield

99% ee, 71 % yield

18b (14,151

18a [14, 151

Me*Me

'

I

OH OAc 96 % ee, 43 % yield 50 % conversion

OAc

OAc

86 % ee, 46 % yield

I

11.1 Hydrolysis and Formation ofCarboxylid Acid Esters 387

Table 11.1-6. (cont.).

+

19a [14, 151

$7

AcO

OAc

19b [14, 151

OAc

31 % ee, 43 % yield

31 % ee, 43 % yield 50% conversion

20b [16]

20a [16] AcO

Acokb-OA~

10% ee

13% ee

21a [16]

21b [16]

‘

I

OAC OH

85% ee

87% ee

4

21c [16]

OAc OAc

8% ee

AAC

22a [16]

OAc

4

221, [16]

OAcOH

19% ee

15% ee

23b [16] AcO &OH

21 % ee

23a [16]

Aco&OAc

26% ee

24b [16] - .- L - - J

ba,

OH

AH

36% ee

84% ee

388

I

II

Hydrolysis and Formation ofC-0 Bonds

Table 11.1-6. (cont.).

24c [16] Ac OAc

66% ee

&OAC

25a [16]

OH

48% ee

&

26a [16]

Ac

4"

Aco4 26b [16]

OAc

17% ee

17% ee

27a 1161

I

27b [16] OAc

OAc

82% ee

85% ee

A A

28a [16]

28b [lG]

OAc OAc

OH

81 % ee

OH

2% 1161

I OAc

87% ee

HO

A 0 Ac

98% ee

28c [16, 171

OAc

73% ee

A0

OH

kOH

43% ee

83 % ee

Ac

29a [l6, 171

29b [16, 171

OAc

I

71. I Hydrolysis and Formation OfCarboxylid Acid Esters 389 Table 11.1-6.

(cont.).

OCOnPr

30a [18]

97 % ee, 36 % yield 35 % conversion

AcOCH,

30b [18]

99 % ee, 40% yield 56% conversion

CH,OAc

31a [19]

58 % ee, 48 % yield

58 % ee, 47 % yield

PhTPh Phy-Ph 32a 1201

32b [20]

OH

OAc OAc

84% ee, 43 % yield 5 % monoacetate

OH

92 % ee, 40 % yield

OAc

33a [20] OAc

295 % ee, 41 % yield 30 % conversion

d O A C OH

33b [20]

295 % ee, 49% yield

OH

33c 1201 OH

295 % ee, 10% yield

cri""' OAc

34a [20]

54% ee, 54% yield 25 % conversion

47 % ee, 53 % yield 74% conversion

OH

50 % ee, 43 % yield 25 % conversion

34c 1201 ( L O HOH

aoAC

341, [20]

1 1 Hydrolysis and Formation ofC-0 Bonds

(cont.).

Table 11.1-6.

35b [20,21]

35a [20, 211 0 % ee, 26 % yield

295 % ee, 33 % yield 25 % conversion

0::

35c [20, 211

295 % ee, 41 % yield \,..OAC

[22,23]

122,231 Q

3Ga 37a 38a 39a 40a 41a

R=Ph R = p-MeCsH4 R = p-tBuCsH4 R = p-PhCsH4 R = o-MeOCJI4 R = 2,4-Me2C~&

98% ee, 32% yield 299% ee, 30% peld 299 % ee, 33% yield 299 % ee, 34 % yield 92 % ee, 47 % yield 90 % ee yield

O

R

3Gb 37b 38b 39b 40b 41b

85% ee, 42% yield 70% ee, 41 %yield 90% ee, 36% yield 88 % ee, 37 % yield 77 % ee, 45 % yield GO%ee

\/.ON02

42a [24]

421,[24]

QOAc

71 % ee, 52% yield, 3d, P M P 299 % ee, 36% yield, 6d, P U P

299% ee, 35% yield, 3d, PLAP 66% ee, 55% yield, 6d, PLAP

,... OAc 43a [25]

( Y O H '"'NHPh

O

" "

44a [25]

299 % ee, 38 % yield

441,[25]

80% ee, 51% yield, E >477

ooH NH-m-BrC,H,

o\

%...OAc N(Me)Ph

N(Me)Ph

299% ee, 48% yield

"*,,

Q N H F 91 % ee, SO%, E 2637

299 % ee, 48 %

D

43b [25]

45a [25]

0, (...OAc

H-rn-BrC, H,

57% ee, 61 % yield, E >425

45b [25]

J1.1 Hydrolysis and Formation ofCar60xylid Acid Esters Table 11.1-6.

(cont.).

mO"

0, *... OAc

4Ga [25]

4Gb [25]

NH-pCIC6H4

96 % ee, 49 % yield, E >989

47b [25]

47a 1251 NH-pN02C6H4

57 % ee, 63 % yield, E >57

94 % ee, 32 % yield

D

O "'.NH-pMeOC, H H,

"',NH-pMeOC, H,

97% ee, 46% yield, E >39

79% ee, 52% yield

(yoH

48b 125)

4% [25]

0, ....OAc

49a [25]

,,NH-eMeC€, H,

49b [25]

NH-eMeOC,H,

42 % ee, 56 % yield

73%ee, 37%yield

50a 1251

( Y O'''.H NH-P-naphthyl

50b [25] NH-P-naphthyl

52% ee, 62% yield, E >335

299 % ee, 34%yield

0

\...OAc

""

N(Et)Ph

51a [25]

5 1 % ee, 51 % yield, E = 65

95 % ee, 32 % yield

52b [25]

52a 1251

( Y O "NHPh

87% ee, 40% yield, E = 13

65% ee, 60% yield

53a [25]

52% ee, 50% yield

51b [25]

N(Et)Ph

53b [25] 68% ee, 58% yield, E = 6

I

391

7 I Hydrolysis and Formation ofC-0 Bonds

(cont.).

Table 11.1-6.

aoH NHPh

54a [25]

N(Me)Ph

541,[25]

55b [25]

55a [25]

u N ( M e ) P h 68% ee, 64% yield, E = 67

94% ee, 33% yield

cyoH

0 N H P h

99% ee, 52% yield, E = 126

92 % ee, 45 % yield

0""

\,..OAc

0

*...OAc

5Ga 1251

N(Et)Ph

561, [25]

N(Et)Ph

70% ee, 55% yield, E >419

299 % ee, 38 % yield

\...OAc G

O

NHPh H

571, [25]

57a (251 0 N H P h

89% ee, 34% yield

aoH

57 % ee, 52 % yield, E = 22

58b (251

58a [25]

N(Me)Ph

39% ee, 50% yleld, E >292

299% ee, 25% yield

(-YOH

N(Et)Ph

59b [25]

59a [25]

37% ee, 73 % yield, E >295

299 % ee, 25 % yield

OH

OCOEt

TMe

Gob [26]

GOa [2G]

N /

98% ee, 43 % yield

88% ee, 44% yield

Glb (271

Gla [27]

21% ee, 60% yield

32% ee, 36% yield, E = 1,7

11.1 Hydrolysis and Formation ofCarboxylid Acid Esters

Table 11.1-6.

d

(cont.).

O ,,..H H

62a [27]

Me &TOnBu

621, [27]

I

86 % ee, 48 % yield

83 % ee, 49 % yield, E = 40

nBu ,s OAC

63a [27]

&

63b [27]

30% ee, 54% yield, E = 7,G

@j:H

nBu O B un& ;

64a [27]

40 % ee, 40 % yield

H$)Q

30% ee, 44% yield, E

65a [27]

39 % ee, 49 % yield

ll*.

sp

=

3,s

nBu ,...OAc

,s..H

H$Q

64b (271

65b [27]

33 % ee, 49% yield, E = 3,8

nBu e f C O n B u

H GGa [27]

GGb (271

I GO % ee, 45 % yield

AcO

50% ee, 48% yield, E = 8,s

OAc

67 [28]

56% ee

68 I281

64% ee

69a [28] OH

24% ee

69b (281

OAc 3 1 % ~

I

393

394

I

I I Hydrolysis and Formation ofC-0 Bonds

FkH

Table 11.1-6.

(cont.).

OAc

70a [29c]

70b [29c]

100% ee, 24 % yield

100% ee, 24 % yield

71a [30]

4

71b [30]

OAc

HO 299 % ee after hydrolysis, acylation and hydrolysis -5 to -10 "C, MeOH

299 %

Me-N

Me-N

twofold hydrolysis

72a (31, 321

&02Me

72b [31, 321 OH

OCOPh

71 % ee, 35 % yield

82% ee, 45 % yield

C0,Me OCOPh

73b [31, 321

73a [31, 321 H O Me0,C . &

95% ee, 85% yield

299% ee, 91% yield

Me\ C0,Me

C0,Me

74a [31, 321

74b [31,32]

MeO& Meo*OH

OCOPh

95 % ee, 3 % yield

82% ee, 35% yield

Me \ N M & e.

N

75b [31, 321

75a [31, 321 C0,Me OCOPh

99 % ee, 60 % yield

Meo%OH

C0,Me

99 % ee, 30 % yield

11.1 Hydrolysis and Formation ofCar6oxylid Acid Esten Table 11.1-6.

(cont.).

Me

Me

\

N

\

7Ga [31, 321 C0,Me

76b [31, 32)

h 0 C O P h

O H .$ Me0

97 % ee, 55 % yield

95 % ee, 26 % yield

1 po

0

77a [33]

C02Me

P

HO

OH

Ph

Ph

78% ee

97 % ee

78a [34]

77b [33]

78b [34]

R = H 299% ee R = F 93 % ee 50%conversion OH PhnP(O)(OMe),

79a [35]

7 % ee, 32 % yield 46 % conversion

0"

R"

Ph

OAc I C0,Et

80a [3G]

90% ee, 81 % yield

8Ob [36]

54% ee

OH

OH

81 [37]

93% ee 47 % conversion

79b [35]

37 % ee, 3 % yield 46 % conversion

OH

C0,Et

P(O)(OMe),

(398% ee 45 % conversion

82 [37]

I

395

396

I

7 I Hydrolysis and Formation ofC-0 Bonds

Table 11.1-6.

(cont.).

96% ee 45 % conversion

22% ee 42% conversion

G

P

85 [37]

h

22% ee 48 % conversion

moH

wocon

R = Me 36% ee, 55% yield R = nPr 1%ee, 33% yield

78% ee, 26% yield, E = 5 2 % ee, 56 % yield, E = 1

8Ga (381

w

8Gb [38]

w

87b [38]

87a [38]

%OHFe

94% ee, 43 % yield, E = 75

R = Me 91% ee, 43% yield R = nPr

88a [39]

Ph

75% ee, 50% yield, P U P 1 J. K. Whitesell, H. H. Chen, R. M. Lawrence, j . Org. Chem. 1985,50,4663. 2 J. K. Whitesell, R. M. Lawrence, Chimia 1986,40, 318. 3 P. Esser, H. Buschmann, M. Meyer-Storck, H.-D. Scharf, Angew.Chem.1992,104,1254; Angew. Chem., In#. Ed. Eng. 1992.31, 1190. 4 H:J. Gais, C. Griebel, H. Buschmann, Tetrahedron: Asymmetry2000, 1 1 . 917.

~I~""oA 88b [39]

Ph

87% ee, SO% yield, P U P 5 H.Hirohara, S. Mitsuda, E. Ando, R. Komaki, In Biocatalysts in Organic Synthesis, J. Tramper, H. C. van der Plaas, P. Linko, Eds., Elsevier, Amsterdam, 1985,p. 119. G K. Mori, J. 1 . J. Ogoche, Liebigs Ann. Chem.1988, 903. 7 K. Mori, P. Puapoomchareon. Liebigs Ann. Chem. 1991,1053. 8 C. Tanyeli, A. S. Demir, E. Dikici, Tetrahedron: Asymmetry1996,8,2399.

1 7 . 7 Hydrolysis and Formation ofCarboxylid Acid Esters 9 j. Y. Lallemand, M. Ledaire, R. Levet, G . Aranda, Tetrahedron: Asymmetry 1993,4,1775. 10 ]:P, Bamier; L. Blanco: G. Rousseau: E. Guib&)ampel/. Org. Chem. 1993, 58, 1570. 11 K. Naemura, H. Miyabe, Y. Shingai,/. Chem. Soc., Perkin Trans. I 1991,957. 12 K. Naemura, H. Miyabe, Y. Shingai, Y. Tobe, /. Chem. Soc., Perkin Trans. I 1993, 1073. 13 M. Roberti, R. Rondanin, R. Ferroni, B. Baruchello, F. P. Invidiata, V. Andrisano, C. Bertucci, V. Bertolasi, S. Grimaudo, M. Tolomeo, D. Simoni, Tetrahedron: Asymmetry 2000, 11,4397. 14 K. Naemura, T. Matsumura, M. Komatsu, Y. Hirose, H. Chikamatsu, Bull. Chem. Soc./pn. 1989,62,3523. 15 K. Naemura, T. Matsumura, M. Komatsu, Y. Hirose, H. Chikamatsu, /. Chem. SOC.,Chem. Commun.1988,239. 16 K. Naemura, N. Takahashi, S. Tanaka, H. Ida, /. Chem. Soc., Perkin Trans. I 1992, 2337. 17 K. Naemura, N. Takahashi, S. Tanaka, M. Ueno, H. Chikamatsu, Bull. Chem. Soc./pn. 1990,63, 1010. 18 J. F. Coope, B. G. Main, Tetrahedron: Asymmetry 1995,6, 1393. 19 K. Naemura, R. Fukuda, N. Takahashi, M. Konishi, Y. Hirose, Y. Tobe, Tetrahedron: Asymmetry 1993,4, 911. 20 K. Yamamoto, H. Ando, H. Chikamatsu, /. Chem. Soc., Chem. Commun. 1987,334. 22 D. Basavaiah, P. R. Krishna, T. K. Bharathi, Tetrahedron: Asymmetry 1995,2,439. 21 G . Caron, R. J. Kazlauskas, /. Org. Chem. 1991,56, 7251.

23 D. Basavaiah, P. R. Krishna, T. K. Bharathi, Tetrahedron Lett. 1990, 31,4347. 24 D. Basavaiah, S. Pandiaraja, K. Muthukumaran, Tetrahedron: Asymmetry 1996, 1, 13. 25 G . Sekar, R. M. Kamble, V. K. Singh, Tetrahedron: Asymmetry 1999,10,3663. 26 P. Rasor, C. Ruchardt, Chem. Ber. 1989,122,1375. 27 T. Izumi, 0. Itou, K. Kodera, /. Chem. Techno. Biotechnol. 1996, G7,89. 28 K. Naemura, A. Furutani, J . Chem. Res., Synop. 1992,174. 29 Y. K. Rao, C. K. Chen, J. Fried,j . Org. Chem. 1993, 58, 1882. 30 Y. Yokoyama, H. Takikawa, K. Mori, Bioorg. Med. Chem. 1996,4,409. 31 A. P. Kozikowski, D. Simoni, P. G. Baraldi, I. Lampronti, S. Manfredini, Bioorg. Med. Chem. Lett. 1996, G, 441. 32 D. Simoni, M. Roberti, V. Andrisano, M. Manferdini, R. Rondanin, F. P. Invidiata, Farmaco 1999, 54, 275. 33 P. Barton, M. I. Page, Tetrahedron 1992,48, 7731. 34 L. K. Hoong, L. E. Strange, D. C. Liotta, G. W. Koszalka, C. L. Burns, R. F. Schinazi, I. Org. Chem. 1992,57,55(13. 35 Y. F. Li, F. Hammerschmidt, Tetrahedron: Asymmetry 1993,4,109. 36 P. S. Vankar, I. Bhattacharya,Y. D. Vankar, Tetrahedron: Asymmetry 1996, 6, 1683. 37 C. Tanyeli, A. S. Demir, A. H. Arkin, 1. M. Akhmedov, Enantiomer 1997, 2,433. 38 T. Izumi, S. Aratani,/. Chem. Technol. Biotechnol. 1994,59,403. 39 T. Ganesh, S. K. Kamalesh, K. G. L. David, Indian /. Chem., Sect. B: Org. Chem. Incl. Med. Chem. 1999,388,397,

substrates for pig liver esterase in terms of an efficient resolution. In particular, pig liver esterase is, as well as lipases, the hydrolase of choice for the kinetic resolution of secondary cyclic alcohols and in particular of cyclohexane derivatives. This is impressively demonstrated by the many examples contained in Table 11.1-6. Quite a number of enantioenriched p-amino alcohols (43-59)having different ring size have been obtained in this manner. Pig liver esterase seems to be especially well suited for the kinetic resolution of cyclic 1,2-diols (14,15, 33-42). In the case of 1,2-diols, having Cz-symmetry, sequential kinetic resolution can be applied for ee enhancement. Resolutions of the racemates of 421,and 88b show the successful use of a crude extract of pig liver (PUP).Further interesting examples are the resolution of cocaine derivatives (72-76) and of amino alcohols (4-6). In the case of the resolution of the racemate of Ga, the remote butyroxy group attached to the aromatic ring is hydrolyzed. Within the series of racemic acetates which have been subjected to liver esterase-catalyzed hydrolysis (Table 11.1-6),the cyclohexanoid compounds 1-3 are particularly interesting since they are valuable chiral auxiliaries. Somewhat puzzling results were recorded in the case of cyclic 1,z-diacetateswith homotopic acetoxy groups. Selectivity is lowest in the case of the five-membered compound, and, not

I

397

398

I

I 1 Hydrolysis and Formation of C-0 Bonds

surprisingly, diol formation is in all cases significant. As observed in the case of cyclic rneso-dicarboxylic acid diesters (Table 11.1-1), there is a reversal in the sense of asymmetric induction on going from the four-membered to the six-membered diacetates. Especially noteworthy is the observation that racemic tertiary acetates are also amenable to kinetic resolution with pig liver esterase (7, 8, 13, and 30). Kinetic resolution of esters of acyclic alcohols with pig liver esterase has been studied only to a minor extent. However, some of the examples described proceeded highly selectively (60, 62, 81, 82, and 84). Acylated alcohols and alcohols of Table 11.1-6, which can be obtained with other hydrolases as such or of opposite configuration, are contained in Tables 11.1-15, 11.1-16, 11.1-20and 11.1-21.

11.1.1.1.2

a-Chymotrypsin

a-Chymotrypsin (CHT, E.C. 3.4.21.1) is one of the most thoroughly studied hydrolases['. 4. 9* 12, 21* 23. 26s 28. 33, 341, and its crystal structure has been It is a serine protease with a pH optimum of 7.8, which acts in vivo as an endopeptidase and catalyzes with great specificity the hydrolysis of non-terminal amide bonds adjacent to phenylalanine, tyrosine or tryptophan. The enzyme has been widely used for the kinetic resolution of racemic amino acid esters. From the results of these studies and based on the crystal structure of the enzyme a useful active site model for a-chymotrypsin has been developed['*81]. Hydrolyses catalyzed by a-chymotrypsin are usually carried out in aqueous buffer solution in a pH range of 7-8. In the case of a low solubility of the substrate in water cosolvents such as methanol, ethanol, dimethylformamide or dimethylsulfoxide, up to 20 % may be used. However, it should be noted that primary alcohols used as cosolvents may react with the acyl-enzyme intermediate with formation of the corresponding ester (Scheme 11.1-9).Diethyl ether, even in low concentrations, is a strong inhibitor of the enzyme. Immobilization of a-chymotrypsin by different methods has been described and the immobilized enzyme is commercially available. a-Chymotrypsin has been found to be active in organic solvents of low water content also['08]. A limited number of prochiral malonates and glutarates are hydrolyzed by achymotrypsin to the corresponding monoesters with synthetically useful enantioselectivities(1-9) (Table 11.1-7). Examples of enantioselective hydrolysis of cyclic diesters by a-chymotrypsin are comparatively rare (10-14) (Table 11.1-7).Interestingly, the cyclopentanoid and the cyclohexenoid monoesters 11 and 12 have the opposite absolute configuration to those obtained by the pig liver esterase-catalyzed hydrolysis of the corresponding diesters (Table 11.1-1).The keto ester 14, which is a valuable building block for the synthesis of prostacyclin analogs, has been obtained from the corresponding a,a'keto diester via a-chymotrypsin-catalyzed hydrolysis followed by a decarboxylationof the keto acid. Monoesters and monoacetates of Table 11.1-7,which can be obtained with other hydrolases as such or of opposite configuration, are contained in Tables 11.1-1, 11.1-2, 11.1-3,11.1-9, 11.1-11and 11.1-18.

11.1 Hydrolysis and Formation of Carboylid Acid Esters Table 11.1-7. a-Chymostrypsin-catalyzed enantiotopos-differentiating hydrolysis of prochiral cyclic dicarboxylic acid esters, acyclic dicarboxylic acid esters and cyclic diol diacetates and enantiomer-differentiating hydrolysis of racemic carboxylic acid esters in aqueous solution.

PhCH,;.

x

Me 1 [I]R = CH3 2 [l]R = C2H5

3 4

5 G

7 3

6

8 9

CO,H CO,R

298% ee, 90-98% yield, DMSO 298% ee, 90-98% yield, DMSO

R’

R2

ee (%)

yield (“h)

Ref.

HO C6HsCOO CcjHsCH2 CHsOCHz CH3COO HO CHsCOO CHjCONH

CH3 CH3 CH3 CH3 CH3 CH3 CzHs CzHs

64 84 92 93 90 55 95 295

100 68

[2-51

86

[41 [41

100 38 78 84 79

[51 [51 151 [GI

(41

OH

42 % ee, 73 % yield

83 % ee, 87 % yield

JfJ

acoZH HO

nPrOCb]

C0,Me

OH

86 % ee, 90 % yield

40%

14 [lo]

OH 295 % ee, GO % yield

ee, -

13 [91

I

399

400

I

I 1 Hydrolysis and Formation of C - 0 Bonds Table 11.1-7.

(cont.).

R2-CH-C02R3 I

NHR’

(4 R’

R2

R3

15a, b COCH,

16a, b CHO

e

c

L-acid D-ester Ref. op (“A),yield (“A) op (“A), yield (“A)

CH3 295,88

295.68

CH3 ~ 295,92 ~

-

-

17a.b COCH3

HoeCH CH3

-

295, ti9

-

295,50

OH

295.78

19a, b H

HO

20a, b H

295,78

21a, b H

>95,40

22a,b H

295, ti0

23a. b H

295,60

Hd

11. I Hydrolysis and Formation ofcarboxylid Acid Esters

Table 11.1-7.

(cont.).

R2-CH-C0,R3 I

NHR’

(4 R’

R2

R’

L-acid

D-ester

op VO), yield VO)op (%), yield (“7)

24a, b H

a

CH3 295,87

>95,70

CH3 97,GO

-

CH3 295,40

295, 50

CH3 high, 42

high, 45

CH3 295,53

295, GO

C2Hs 84,28

high, 93

CH3 >98,61

86,58

CH3 290,GS

295,86

CH3 295,GS

295,81

H

25a.b COCH3

100

H

28a, b COCH,

29a, b COCH3

30a, b COCH3

0 0 D ( C H . , *

Ref.

I

*l

402

I

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1.7.

(cont.).

R2-CH-C02R3 I

NHR’ R’

(4 R’

R2

L-acid

D-ester

Ref.

op (“A), yield VO)op (“A), yield (“A) ~

~~

33a, b COCH3

CH3

CzH5 >95,91

295,92

P I

34a, b COCH3

C2H50COCH2

C2Hs 295,97

295,83

P I

35a, b COCH3

C2HsOCO(CH2)2

CzH5 >95,74

295,68

~ 3 1

36a, b COCH3

NO2

CH3

295,-

~ 4 1

wester

Ref.

R3 R2+CO,R~

R’ (*) R’

R2

R3

0%

L-acid op YO), yield

YO)op (%), yield (“A)

H

low, 100

low, 40

(251

38a, b CH3

H

295,533

>95,88

P61

39a, b OH

CH3 high, 100

75,91

~ 7 1

40a, b CH3

NH2 >95,72

>95,88

[281

41a, b CH3

NH2 295,66

295,78

[281

42a, b CH3

NH2

37a, b OCOCH,

I H

-11.2, 78 (1.0, MeOH)

11.1 Hydrolysis and Formation ofCarboxylid Acid Esters

Table 11.1-7.

(cont.).

CH,-CH-C02Rz

I

OCOR'

(4

D-acid (R2 = H) op (%), yield (77)

L-ester (R2 = C2H5) op (%), yield (%)

43a, b C6Hs

high, 28

82,88

44a, b CH3

73,85

73,85

R'

Ref.

(4 R'

D-acid (R2 = H) op (%), yield (%)

L-ester (R2 = C2H5)

45a, b NHCOCH, 84,28

high, 93

4Ga, b OH

high, 84

high, 100

47 [30]

42 % ee,

Ref.

op (%), yield (%)

48 [30]

50% ee C02Me 49a [30]

5" 44% ee

49b [30]

47% ee

50 [30]

270% ee 50% conversion

KC' 0

51b [31]

C02Me

86% ee, 38%yield

82 % ee, 35 % yield

I

403

404

I

7 7 Hydrolysis and Formation of C-0Bonds Table 11.1-7.

(cont.).

52a [32] I

OMe

OMe

295% ee, DMF

295 % ee, 71 % yield, DMF no hydrolysis of the 2-isomer

53 [33]

54 [33]

Me high op, 38% yield

high op, 36% yield

55a [34]

!’ JH

PhAS

55b [34]

Ph’ 91 % ee, tBuOMe 58% conversion absolute configuration unknown

65% ee, tBuOMe 58 % conversion absolute configuration unknown

R=

R=Ph

56 [35]

57 (351

I

H 295 % ee

295 % ee, 27% yield

P C H ,

XCH2

295 % ee

85% ee

58 [35]

GO (351

59 [35]

7 1.1 Hydrolysis and Formation ofcarboxylid Acid Esters 1435 1 F. Bjorkling, J. Boutelje, S . Gatenbeck, K. Hult, T. Norin, P. Szmulik, Tetrahedron 1985,41, 1347. 2 S. J. Cohen, E. Khedouri,]. Am. Chem. SOL.1961, 83,4228. 3 P. Mohr, L. Rosslein, Ch. Tamm, Helv. Chim. Ada 1987, 70, 142. 4 R. Roy, W. Rey, Tetrahedron Lett. 1987.28.4935. 5 E. Santaniello, M. Chiceri, P. Ferraboschi, S . Trave,]. Org. Chem. 1988, 53, 1567. G S. G. Cohen, E. Khedouri,]. Am. Chem. SOL.1961, 83, 1093. 7 K. Laumen, M. Schneider, Tetrahedron Lett. 1984, 25, 5875. 8 H.-J.Gais, G . Bulow, A. Zatorski, M. Jentsch, P. Maidonis, H. Hemmerle,]. Org. Chem. 1989, 54, 5112. 9 G. Baudin, B. I. Glanzer, K. S. Swaminathan, A. Vasella, Helv. Chim. Ada 1988, 71, 1367. 10 K. Petzold, H. Dahl, W. Skuballa, M. Gottwald, Liebigs Ann. Chem. 1990, 1087. 11 H. T. Huang, C. Niemann,]. Am. Chem. SOC.1951, 73, 3228. 12 G. E. Hein, C. Niemann, ]. Am. Chem. SOC.1962, 84,4487. 13 R. L. Bixler, C. Niemann,]. Am. Chem. SOL.1958, 80, 2716. 14 G. E. Clement, R. Potter,]. Chem. Ed. 1971,48, 695. 15 J. H. Tong, C. Petitclerc, A. D’lorio, N. L. Benoiton, Can.]. Biochem. 1971,49,877. 16 H. T. Huang, C. Niemann, J. Am. Chem. SOC.1951, 73, 1541. 17 D. W. Thomas, R. V. Mac Allister, C. Niemann, I . Am. Chem. SOC.1951,73,1548. 18 1. B. Jones, T. Kunitake, C. Niemann, G. E. Hein, I. Am. Chem. SOC.1965,87,1777.

19 Y. Hayashi, W. B. Lawson,]. Biol. Chem. 1969,244, 4158. 20 T. N. Pattabiraman, W. B. Lawson, Biochem. J. 1972, 126, 659. 21 S. G. Cohen, Y. Sprinzak, E. Khedouri, 1.Am. Chem. SOC.1961,83,4225. S. G. Cohen, S. Y.Weinstein, ibid 1964,86, 725. 22 S. G. Cohen, 1. Crossley,]. Am. Chem. SOC.1964, 86, 4999. 23 S.G. Cohen, J. Crossely, E. Khedouri, Biochemistry 1963, 2, 820. 24 R. Chenevert, R. Letumeau, Chem. Lett. 1986, 1151. 25 S. G. Cohen, S . Y. Weinstein, J. Am. Chem. SOL. 1964,86,5326. 26 S. G. Cohen, A. Milovanovic,]. Am. Chem. Soc. 1968,90,3495. 27 S. G. Cohen, L. W. Lo,]. Biol. Chem. 1970,245, 5718. 28 G. M. Anantharamaiah, R. W. Roberts, Tetrahedron Lett. 1982, 23, 3325. 29 S. G. Cohen, J. Crossley, E. Khedouri, R. Zand, L. Klee,]. Am. Chem. SOC.19G3,85,1685. 30 J. B. Jones, P. W. Man, Tetrahedron Lett., 1973,34, 3165. 31 J. H. Udding, J. Fraanje, K. Gaubitz, H. Hiemstra, W. N. Speckamp, K. Kaptein, H. E. Schoemaker, J. Kamphuis, Tetrahedron: Asymmetry 1993, 4, 425. 32 G. Gcan, N. Satyamurthy, J. R. Barrio, Tetrahedron: Asymmetry 1995, 6, 525. 33 N. L. Dirlam, B. S. Moore, F. J. Urban,]. Org. Chem. 1987,52, 3587. 34 C. Cardellicchio, F. Naso, A. Seilimati, Tetrahedron Lett. 1994, 35,4635. 35 J. J. Calonde, D. E. Bergbreiter, D.-H. Wong, /. Org. Chem. 1988,53, 2323.

a-Chymotrypsin has been used most frequently and with much success for the enantiomer-differentiating hydrolysis of a wide range of natural and non-natural amino acid ester derivatives (Table 11.1-7), which usually leads in both cases to a mixture of the L-amino acid derivative and the D-amino acid ester derivative (15-36, 52,and 53). Excellent substrates for a-chymotrypsin are aromatic amino acid ester derivatives, but those amino acid esters which carry aliphatic or functionalized aliphatic chains of a certain length are excellent substrates also. Even a methyl or a nitro group as substituent is tolerated by the enzyme. Upon placement of a methylene group between the ester group and the stereogenic center, enantiomer differentiation is reverted and the D-acid derivative and the L-acid ester derivative are formed (14 vs 45). Interestingly, a-chymotrypsin also exhibits high enantiomer selectivity toward aromatic amino acids which bear a methyl group at the Ca-atom (40-42). Nitro analogs of methyl-substituted amino acids were also found to be suitable substrates for an a-chymotrypsin-catalyzed resolution (5660).Enantiomerdifferentiating hydrolysis of a-hydroxy acid ester derivatives is also feasible (43and 44).As organic cosolvents, dimethyl sulfoxide, dimethylformamide and tert-butanol have been used without significant deactivation of the enzyme. The enantiomer

1 I Hydrolysis and Formation ofC-0 Bonds

6

Table 11.1-8. Acetylcholine esterase-catalyzed enantiotopos-differentiating hydrolysis of prochiral cyclic diol diacetates and of racemic monoacetates in aqueous solution.

.-OAc

OH

2 [2, 31

OAc

96 % ee, 93 % yield

62 % ee, 62 % yield

OH 1 3 141

1

OAc

6

OAc

98% ee, 795 yield

295 % ee, 95 % yield

O -)(H

4 [51

Me .ex::;;:$'

5 161

OH

OH

100% ee, 39% yield

295 % ee, 79 % yield

8 [81

7 [71 OAc

92 % ee, 77 % yield

n o hydrolysis

&

bS*..OH 71% ee 50% conversion

82% ee 50% conversion

10 [lo]

I

NHC0,Me 72% ee,

9b 191

9a [91

11. I Hydrolysis and Formation ofCar6oxylid Acid Esten Table 11.1-8.

(cont.).

g

OAc

11b [lo]

I

NHAc

NHAc 92 % ee,

92 % ee,

after a second hydrolysis 1 H. Suemune, T. Harabe, Z. F. Xie, K. Sakai, Chem. 6 C. R. Johnson, C. H. Senanayke,]. Org. Chem. Pharm. Bull. 1988, 36, 437. 1989,54, 735. 7 A. J. Pearson, H. S . Bansal, Y.S. h i , ]. Chem. Soc., 2 D. R. Deardoff, A. J . Matthews, D. S . Mc Heekin, Chem. Commun. 1987,519. C. L. Carney, Tetrahedron Lett. 1986, 27, 1255. 8 H. E. Schink, J. E. Baeckvall,]. Org. Chem. 1992, 3 D. R. Deardofi, C. Q.Windham, C. L. Craney, Org. Synth. 1996, 73. 57, 1588. 4 D. M. Legrand, S. M. Roberts,]. Chem. SOC.,Perkin 9 J. V. Allen, J. M. J. Williams, Tetrahedron Lett. 1996, Trans. 11992, 1751. 37, 1859. 5 C. R. Johnson, T. D. Penning,]. Am. Chin. SOC. 10 M. J. Mulvihill, J. L. Cage, M. J. Miller,]. Org. 1988, 110,4726. Chem. 1998,63,3357.

differentiation by a-chymotrypsin can rather accurately be explained by the active site model for the enzyme.

11.1.1.1.3

Acetylcholine Acetylhydrolases

Acetylcholine acetylhydrolase (E. C. 3.1.1.7) or acetylcholine esterase is a well characterized hydrolase [lOgl which is commercially available. Acetylcholine esterase-catalyzed hydrolyses have been reported only for a small number of prochiral diacetates (Table 11.1-8).However, several of secondary monoacetates, which are valuable synthetic building blocks, have been obtained with high enantioselectivity (2-6 and 11)by using this enzyme. Acetylcholine esterase should be considered for the hydrolysis of diacetates which are not substrates for lipases and pig liver esterase. Monoacetates of Table 11.1-8,which can be obtained with other hydrolases as such or of opposite configuration, are contained in Tables 11.1-3, 11.1-7 and 11.1-18.

11.1.1.1.4

Subtilisin

Subtilisins are a family of serine proteases, the most important members of which are subtilisin Carlsberg (from Bacillus lichenijomis) and subtilisin BPN’ (from Bacillus amyloliquefaciens)[llol. Both enzymes are alkaline proteases with a pH optimum of 6-9. Because of their industrial importance, both subtilisin Carlsberg and subtilisin BPP’ have been studied intensively and are produced on a large scale. The crystal structures of both subtilisins have been determined[82].Directed evolution and site-directed mutagenesis and chemical modification of subtilisin were carried out in order to influence the stability, activity and enantioselectivity of the enzyme, in particular in organic solvents[111].As in the case of other enzymes,

I

407

408

I 7 1 Hydrolysis and Formation ofC-0 Bonds I Table 11.1-9.

Subtilisin-catalyzedhydrolysis of racernic and prochiral esters.

100% ee, 85% yield

90% ee, 96 % yield

2lJ[1] \

C0,Me

100% ee, 86 % yield

91 % ee, 95% yield

'UH, MeACO,H 86 % ee, 98 % yield

4a [31 N~ /

CO,H 98 % ee, 50 % yield

93% ee, 50% yield

%,

5a [31

\

GO,H 98 % ee, 50 % yield

93% ee, 50% yield

"2

~a [41

PhnC0,H 97% ee, 93% yield

91 % ee, 100% yield

CO2Et

Me0 90 % ee, 89 % yield

OMe

95 % ee, 101% yield

71, [4]

7 1. I Hydrolysis and Formation ofCar6oxylid Acid Esters

I

409

Table 11.1-9.

(cont.).

8 2 % ee, 100% yield

93% ee, 95% yield NHBoc

NHBoc

CO,H

CO,Me(Et)

9a [5, 61 R = SO,Na, 10a [5, 61 R = P(O)(OEt)z l l a [5,6] R = COZCHJ’h, 12a [5, 61 R = COzMe, 13a [S, 6]R = CONHCHzPh,

295% ee, 47% yield 295% ee, 40% yield 295% ee, 37% yield 295% ee, 49% yield 295% ee, 43% yield

9b (5, 61 295% ee, 48% yield 101, [5, 61 295% ee, 46% yield 11b [5, 61 295% ee, 40% yield 12b [S, 61 295% ee, 47% yield 13b [S, 61 295% ee, 46% yield

OH

297 % ee, 96 % yield

M

e

o

\2

Meozs

297 % ee, 99 % yield NHAc

s

L

15b [8]

CO,H

~

15a (81

C0,Et

OH

OH

298 % ee 47 % conversion

84% ee 47 % conversion

NHAc

NHAc

1Ga [8]

1Gb [8]

CO,H

C0,Me

96% ee 51 % conversion CLEC-Subtilisin

299 % ee 51 % conversion

NHBoc 17a [8]

17b [S]

CO,H

C0,Me

299 % ee 46 % conversion

85% ee 46 % conversion 18a (91

Br

97% ee,

CO,H

18b [9] Br

96% ee

410

I

I I Hydrolysis and Formation of C - 0 Bonds Table 11.1-9.

(cont.).

Me

MevMe BOC-HN&EyCO2H

0

19a [lo]

BOC-HN&iyC02Me

0

OCH,Ph

19111101

OCH,Ph

299 % de, 50 % yield starting from a 1:I mixture

299 % ee, 50 % yield starting from a 1:1 mixture

20b[11] CI

C0,Et

45 % yield

9G % ee, 45 % yield Me NHAc

21 [I21 Et0,C X C 0 2 H

PhK

C

81 % ee, 290% yield

vs/vK = 6.8, dioxane

$

I

23b [14]

...

”‘ / OAc

OH

OAc

86 % ee, 25 % yield

80 % ee, 29 % yield

0

OAc

74% ee, 11% yield

55% ee, 25% yield acetone

0

C02Me 24 [15]

CO,H

88 % ee, 85 % yield 90% ee, 95% yield, CLEC-subtilisin

22 [13]

k:

Table 11.1-9.

11.1 Hydrolysis and Formation ofCarboxylid Acid Esters I411

(cont.).

25 [16]

&k

CN

C0,Et

93% ee guanidinium chloride

99 % ee, 47 % yield

Me

26 [16]

Me

27 [17]R = CHzCeCH, 74% ee, HzO, MeCN 43 % conversion

80% ee, MeCN, H20, trioctyl amine 95 % conversion (withdynamic kinetic resolution)

R o z c D c o z H

29 [18]

28 [17] R = CHzCF3,73% ee, H20, MeCN

35 % conversion

Me

I H

83% ee, H20, MeCN, triodyl amine 97 % conversion (with dynamic kinetic resolution)

Me

R = (CH,),NH-CO

299% ee, 41 % yield CO H CO,H

30 [19]

Me

Ph&SO,tB” 298% ee,

A

Ph&SO,CON~O

31 (19)

Me

298 % ee,

7 R. Chevenert, S.Thiboutot, Synthesis 1989,444. Chem. Comrnun. 1986,1514. 8 Y.-F. Wang, K. Yakovlevsky, B. Zhang, A. L. Margolin, J. 0%.Chem. 1997,62,3488. 2 E. E. Ricks, M. C. Estrada-Valchs, T. L. McLean, G. A. Jacobucci, Biotechnol. Prog. 1992,8,197. 9 M. R. Leanna, H. E. Morton, Tetrahedron Lett. 1993,34,4485. 3 B. Imperiali, T.J. Prius, S. L. Fischer, S. L. Fister, J. Org. Chem. 1993.58,1613. 10 M. Borgenstatter, W. Steglich, Tetrahedron 1997, 4 J. Morgan, J. T. Pinhey, C. H. Sherry,J. Chem. Soc., 53,7267. 11 A. SalladiXavallo, J. Schwarz, V. Burger, Perkin Trans. 1997,613. 5 C. Garbay-Janreguibeny, I. MeCort-Tranclepain, Tetrahedron: Asymmetry 1994,5, 1621. B. Barbe, D. Ficheux, B. P. Roques. Tetrahedron: 12 S. Iriuchijima, K. Hasegawa, G. Tsuchihashi. Agnc. Biol. Chem. 1982,46,1907. Asymmetry, 1992,3,637. G K. Baczko, W.-Q. Liu, B. P. Roques, C. 13 G. Ottilina, R. Bovora, S. Riva, G. Carrea, Garbay-lanreguibeny, Tetrahedron 19%,52,2021. Biotechnol. Lett. 1994,16,923-928. 1 S.-T. Chen, K.-T. Wang, C.-H. Wong, J. Chem. Soc.,

412

I

11 Hydrolysis and Formation ofC-0 Bonds 14 F. Theil, H. Schick, P. Nedkov, M. Bohme, B. Hafner, S . Schwarz,]. prakt. Chemie 1988, 330, 893. 15 R. ChPnevert, R. Martin, Tetrahedron: Asymmetry 1992, 3, 199. 16 B. Wirtz, M. Soukup, Tetrahedron:Asymmetry 1997, 8, 187.

17 P. I. Urn, P. G . Drueckhammer,]. Am. Chem. SOC. 1998, 120, 5605. 18 T.Adachi, M. Ishii, Y. Ohta, T.Ota, T. Ogawa, K. Hanada, Tetruhedron:Asymmetry 1993,4, 2061. 19 S. Doswald, H. Estermann, E. Kupfer, H. Stadler, W. Walther, F. Weisbrod, B. Wirz, W. Wostl, Bioorg. Med. Chem. 1994,2,403.

CLECs of subtilisin have been prepared r1l2]. Autohydrolysis of subtilisin-CLECs seems to suppressed as compared to subtilisin. Subtilisin has been widely used for the kinetic resolution of amino acid esters. Hydrolyses catalyzed by subtilisin are usually carried out in aqueous buffer solution in a pH range of 7-8. In the case of a low solubility of the substrate in water, cosolvents such as methanol, ethanol, dimethylformamide or dimethyl sulfoxide may be used. subtilisin is the hydrolase of choice for the racemate separation of natural and non-natural amino acid esters (1-18 and 20) (Table 11.1-9). Generally, the L-amino acid ester is preferentially hydrolyzed. Free amino acid esters as well as N-protected amino acid esters are substrates for subtilisin. The utility of subtilisin for the synthesis of enantioenriched amino acids is impressively demonstrated by the highly selective resolution of amino acid esters, the side chains of which contain functional groups (9-15).Frequently, subtilisin is preferred in the large scale resolution of amino acid esters rather than other hydrolases because of its lower price. Not only racemic amino acid esters but also other racemic carboxylic acid esters have been resolved with subtilisin. Impressive examples in terms of selectivity and efficiency are the hydrolyses yielding the functionalized esters 25, 26,and 29-31, which were prepared on a large scale. Activity and selectivity of the enzyme in the kinetic resolution of the racemic esters of 30 and 31 could be improved by addition of dimethyl sulfoxide or guanidinium chloride, which was used at a concentration of 10 mM. A particularly interesting example of a kinetic resolution with subtilisin is that of the racemic thioesters derivatives of 27 and 28.This was carried out in the presence of a base to a ensure a dynamic kinetic resolution (see Sect. 11.1.1.2.1.2) through base-catalyzed racemization of the non-hydrolyzed enantiomeric thioester.

11.1.1.1.5

Lipases

Lipases (triacyl glycerol acyl hydrolases, E.C. 3.1.1.3) are a unique class of hydrofor asymmetric synthesis based on prochiral or racemic substrates. The lases [113-1151 application of lipases as biocatalysts has been reviewed emphasizing different aspects in a number of books[', 21, 22, 2 6 28, 30. 322 34, 351 and jourrials[". 14. 18, 19, 24, 25, 27, 29, 31. '171. Lipases are catalytically active in water, in mixtures of water and water-immiscible or miscible organic solvents, in almost anhydrous organic solvents, and in supercritical fluids [34, 361 and ionic liqUidS["8, 119l . They are available from plants, mammals, and microorganisms in considerable numbers, which explains in part their versatility for asymmetric synthesis. Lipases are typical induced-fit enzymes, accepting non-natural substrates of enormous structural diversity. There is some confusion in the literature regarding the origin and the name of

1 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters

I

413

some microbial lipases. The lipase from Pseudomonas cepacia from Amano was formerly called lipase from Pseudomonas Jluorescens, and was most recently reidentified as Burkholderia cepacia lipase. Candida cylindracea lipase was re-identified as C. rugosa lipase and Mucor miehei lipase was re-identified as Rhizomucor miehei lipa~eI~~1. Candida antarctica produces the two lipases A and B that are/were available either as a mixture or in both individual forms. In order to avoid any further confusion in this text, by and large the names from the original papers have been used, but the special supplier names have been translated into names referring to the biological origin so far as unambiguously possible. The lipases most used until now are the commercially supplied pig pancreas lipase (PPL), Pseudomonas cepacia lipase (PCL) or P. juorescens lipase (PFL), Candida cylindracea (CCL) or C. rugosa lipase (CRL),Pseudornonas sp. lipase (PSL), increasingly Candida antarctica B lipase (CAL-B) and to a lesser extent further lipases mentioned in Tables 11.1-10 to 11.1-25, and cholesterol esterase (CE). CAL-B is a recombinant protein produced in Aspergillus oryzae accepting a broad range of substrates and conditions. A special group of hydrolases, which are considered as lipases, are the cholesterol esterases (CE), found in mammals and microorganisms (ll31. About 70 different lipases are commercially available. Most of these are presumably serine hydrolases containing a serine residue in their active site and featuring presumably the triad Ser ... His .... Asp. The crystal structures of the 13 different lipases have been determined[84-871. The molecular weight of the known lipases in their active, native form ranges from 30 to 65 kDa. Lipases are generally soluble in water and insoluble in organic solvents, and may be strongly adsorbed at the air/water interface. Lipases are available and applied as lyophilized powders, in covalently and noncovalently immobilized form on inorganic or organic carriers, in sol-gel material['20plZ1l and as CLECs["', lZ2].Most mammalian lipases exhibit pH optima ranging from 8 to 9 and most microbial lipases from 5.6 to 8.5. The temperature range for optimal activity is between 30 and 50 "C. In the case of labile substrates or insufficient enantiomer selectivity, hydrolysis may be carried out in water-saturated water-immiscibleorganic solvent such as diisopropyl ether, hexane or cyclohexane. Most lipases are applied as crude materials consisting of a mixture of proteins that may even contain other hydrolases together with stabilizing solid supports. Pig pancreas lipase is a glycoprotein which exists as a mixture of isoenzymes differing in the glycan moiety of the enzyme. Crude pancreas lipase contains presumably another carboxyl esterase that may be responsible at least in part for the high enantioselectivity frequently observed with this enzyme in hydrolysis and esterification[44, 123-1253. Therefore, an isolated lipase of the same origin may have different activities and selectivities depending on the isolation and purification procedures of the individual suppliers. Some of these problems can be overcome, however, by the application of purified lipases, which are also commercially available. Lipases exhibit high catalytic activity in water and an even higher activity in two-phase systems composed of water and a water-immiscible organic solvent or water and a liquid substrate. In two-phase systems like water and tert-butyl methyl ether or water and

414

I

11 Hydrolysis and Formation of C - 0 Bonds

Table 11.1-10. Lipase-catalyzed enantiotopos-differentiating hydrolysis of prochiral acyclic diol diacetates in aqueous solution (MJL Mucorjauanicus lipase, PFL Pseudomonasfluorescens lipase, PPL ig pancreas lipase, PCL Pseudomonas cepacia lipase).

1

1

1 2

3 4 5 5 6 7 8 8 9 9 9 10 10 10 11 11 12 13 14

15

yield (99)

R'

R2

R3

Lipase

ee [%)

(CH3)zCH (CH3)zCH (CH3)2CH CHzPh CHFCH-CH~ CHz=CH-(CH2)2 Ph Ph C-CBHII (E)-n-Pent-CH=CH (E)-n-Pent-CH=CH (E)-n-Pent-CH=CH (2)-n-Pent-CH=CH (2)-n-Pent-CH=CH (2)-n-Pent-CH=CH (E)-i-Pr-CH=CH (E)-i-Pr-CH=CH (E)-i-Pr-CH=CH (2)-i-Pr-CH=CH (2)-i-Pr-CH=CH n-Hep i-Pent n-Pent-C=C

H H H H H H H H H H H H Ac Ac Ac H H H Ac Ac H H H

Ac Ac Ac Ac Ac Ac Ac Ac Ac Ac Ac Ac H H H Ac Ac Ac H

PPL,crude PPL" PPL, pure PPLa PPL PPLa PPL" PFL PPLa PPL PPLb PPLc PPL PPL' PPLb PPL PPLc PPLb PPL PPLc

-

Ac Ac Ac

PPL'

PPLc PPL PPLC PPLb PPL PPL' PPLb

37 75 very slow 61 95 295 295 94 60 84 95 93 50 53 55 90 97 88 21 15 70 72 78 80 82 82 85 88

i-Pr-CIC

H

H

Ac

91 hydrolysis 65 34 80 91 41 96 49 63 59 43 31 44 70 75 71 25 20 56 47 57 50 61 67 65 71

Ref.

111 111 (11 11, 21

[41 121

PI [31 121 [5,61 [5,61 [5,61 161 [61

PI [GI [61 [GI 161 [61 [61 [61

FI I61 [61

PI [61 (61

16

H

Ac

PPL

67

29

161

17

H

Ac

PPL

2

45

[GI

Hc,,

Me0,C

OCOR

18

PhCH,O

;CoAc OH

l9

11.1 Hydrolysis and Formation ofcarboxylid Acid Esters

I

415

Table 11.1-10.

(cont.).

R

Ref.

nPr, 65 % ee, 29 % yield, PPLd nBu, 68 % ee, 36 %yield, PPLd nC5HI1, 65 % ee, 29 %yield, PPLd nC6H13,70 % ee, <17% yield, PPLd nC7H15, 84 % ee, 48 %yield, PPLd.'

[7] [7] [7] [7]

Ref.

60 % ee, 40 % yield, PPL 80 % ee, 40 %yield, PPLd 88 % ee, 45 %yield, PPLc 91 % ee, 75 %yield, PFL

[81 [81

PI PI 21 [lo]

20 [lo] CbzHN

297 % ee,55 %yield, PPL

73 % ee, 46 %yield, PDL 91 % ee, 75 %yield, PPLg R = ME-(CH*)ls 57 % ee, 66 %yield, PPL 87 % ee, 49 % yield, PPLg

22 [lZ]

\o
23 (131

OH

89 % ee, 82 % yield, PPL

x0

0

95 % ee, -, MJL

24 [14, 151

298 % ee, 98 % yield, PFL

97 % ee, 83 % yield, PFL

96 % ee, 85 % yield, PFL

299 % ee, 33 % yield, PFL

~ ' " ' . "[211 " ~ ~ OH 2g

28 [20]

&OH \OAc

78 % ee, 80 % yield, PPLh

OAc

98 % ee, yield 63 %, PCL'

FI

95 % ee, yield 65 %, PCL'

416

I

I 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-10.

(cont.).

a Protein fraction from chromatography of

crude PPL b 10 % tee-BuOH c Diisopropyl ether d Absolute configuration unknown 1 G. M. Ramos Tombo, H:P. Schar, X. Fernandez I Busquets, 0. Ghisalba, in: C. Laane, J. Tramper, M. D. Lilly (Eds.), Biocatalysis in Organic Media, p. 43, Elsevier, Amsterdam, 1987. 2 G. M. Ramos Tombo, H:P. Schar, X. Femandez 1 Busquets, 0. Ghisalba, Tetrahedron Lett. 1986,27, 5707. 3 S. Atsumi, M. Nakano, Y. Koike, S. Tanaka, M. Ohkubo, T. Yonezawa, H. Funabashi, J. Hashimoto, H . Morishima, Tetrahedron Lett. 1990, 31, 1601. 4 Y.-F. Wang, C . J. Sih, Tetrahedron Lett. 1984, 25, 4999. 5 G. Guanti, L. Banfi, E. Narisano, Tetrahedron Lett. 1989,30,2697. 6 G. Guanti, L. Banfi, E. Narisano, Tetrahedron: Asymmetry 1990, I, 721. 7 J. Ehrler, D. Seebach, Liebigs Ann. Chem. 1990, 379. 8 D. Breitgoff, K. Laumen, M. P. Schneider, J. Chem. Soc., Chem. Commun. 1986,1523. 9 V. Kerscher, W. Kreiser, Tetrahedron Lett. 1987,28, 531. 10 Y.-F. Wang, J. J. Lalonde, M. Momongan, D. E. Bergbreiter, C.-H. Wong, J. Am. Chem. SOC. 1988, 110,7200.

e 30 % MeOH f 15 % Tetrahydrofuran g Hexane h 15 % t-BuOH i 30 % Diisopropyl ether 11 K. Prasad, H . Estermann, C:P. Chen, 0. Repic, G. E. Hardtmann, Tetrahedron: Asymmetry 1990, 1, 421. 12 Y.-F. Wang, C . 4 . Chen, G. Girdaukas, C. J . Sih, J . Am. Chem. SOC.1984, 106, 3695. 13 B. Wirz, R. Schmid, J. Foricher, Tetrahedron: Asymmetry 1992,3,137. 14 C. Bonini, R. Racioppi, L. Viggiani, G. Righi, L. Rossi, Tetrahedron: Asymmetry 1993, 4, 793. 15 C. Bonini, R. Racioppi, G. Righi, L. Viggiani, J . Org. Chem. 1993, 58, 802. 16 N. Adje, 0. Breuilles, D. Uguen, Tetrahedron Lett. 1993, 34,4631. 17 T. Itoh, H. Ohara, Y. Takagi, N. Kanda, K, Uneyama, Tetrahedron Lett. 1993, 34, 4215. 18 Y-B. Seu, Y:H. Kho, Tetrahedron Lett. 1992, 33, 7015. 19 2:F. Xie, H. Suemune, K. Sakai, Tetrahedron: Asymmetry 1993,4,973. 20 G. Guanti, E. Narisano, R. Riva, Tetrahedron: Asymmetry 1997,8,2175. 21 T. Yokomatsu, T. Minowa, T. Murano, S . Shibuya, Tetrahedron 1998, 54, 9341.

hexane, much higher reaction rates and enantioselectivities are most frequently observed. A synthetically useful alternative to the lipase-catalyzed hydrolysis of an ester in aqueous solution for the attainment of a chiral carboxylic acid or alcohol is the lipase-catalyzed alcoholysis of an ester described in Sect. 11.1.1.3. Generally, through (a) hydrolysis or alcoholysis of a prochiral dialkanoate or racemic alkanoate by water in homogeneous and heterogeneous aqueous/organic mixtures or by an alcohol in an organic solvent and (b) acylation of the corresponding prochiral diol or racemic alcohol in an organic solvent, access to both enantiomers of the corresponding monoacetate and alcohol, respectively, is provided with one enzyme [(a)Tables 11.1-10to 11.1-12, 11.1-14 to 11.1-16,and 11.1-22; (b) Tables 11.1-17 to 11.1-211(Scheme 11.1-12).Hydrolysis and ester formation are enantiocomplementary, which means that one and the same lipase for a given substrate reacts preferentially with the same enantiotopic group or the same enantiomer irrespective ofwhether it is an ester or an alcohol. Enantioselectivities and yields may differ in hydrolysis and alcoholysis, respectively, and in acylation. A combination of these routes may be advantageous in a given case for the attainment of both enantiomers in high enantiomeric purity (Schemes 11.1-11 and 11.1-14). One cannot expect, however, that a lipase will transform a non-natural substrate in an optimal manner. Therefore, different, more or less empirical, strategies have been

7 1.1 Hydrolysis and Formation ofCarboxylid Acid Esters

I

417

developed to improve reactivity and/or selectivity. The substrate structure and the origin of the lipase mainly determine reactivity and selectivity in a lipase-catalyzed reaction. Hence, the first step of an envisaged lipase-catalyzed kinetic resolution or desymmetrization of a given substrate comprises rapid screening of available lipases [35e1. There is no simple relationship, however, between structural parameters of a given substrate, the origin of the lipase and the outcome of the reaction. On the other hand, models have been developed in order to understand and predict the behavior of certain lipases toward structural properties of substrates with regard to reactivity and selectivity[57,74-79, 126-1321 that help to optimize the reaction by modifying the substrate structure. As mentioned before, lipase activity and selectivity are strongly influenced by the medium used for the desired reaction. Having identified a suitable lipase, variation of the medium - solvent engineering - might be the next step in order to optimize the outcome of the reaction. An additional fine tuning of the reaction conditions is achievable by using certain additives, which may have a beneficial influence on the microenvironment of the lipases and hence on their selectivity and/or reactivity['33].Finally, an increase of the selectivity of the lipase may be gained by lowering the temperature of the reaction mixture. Primary acyclic diacetates are substrates par excellence for lipases (1-24,2630) (Table 11.1-10).Prochiral 1,3-propane diol derivatives have been most thoroughly studied in terms of the influence of the substrate structure, the composition of the reaction medium and the origin and purity of the lipase on the enantioselectivity.A comparison of the hydrolysis of 2-alkyl substituted 1,%propane diol diacetates with crude pig pancreas lipase, highly purified commercial pig pancreas lipase and a carboxyl esterase fraction isolated from crude pig pancreas lipase showed that the latter gave the monoacetate 1 with higher enantioselectivity and reaction rate than the first two (Table 11.1-10).Imitating the in vivo conditions for the action of lipase on triglycerides by addition of diisopropyl ether to the aqueous solution and carrying out the hydrolysis in the two-phase system leads in some but not all cases to a higher enantioselectivity and reaction rate, as demonstrated for the monoacetates 7 and 8. Unsaturation in the alkyl chain frequently leads to the monoacetate of a higher ee value as exemplified with 16 and 17. Comparison of the enantioselectivitiesof the hydrolysis of diacetates to the corresponding monoacetates is often complicated by the lack of information on the amount of diol formed. The later arises from the hydrolysis of the monoacetate that may proceed under enantiomer differentiation, and thus the ee value of the monoacetate will be a composite of two enantioselective processes. Interestingly, upon changing the configuration of the double bond of the substituent R from (q to (Z) the enantiotopic group recognition by pig pancreas lipase inverts, as demonstrated by the monoacetates 8 and 9 as well as 11 and 12 (Table 11.1-10). Group-selective and enantioselective hydrolyses as well as the influence of the alkyl group of the acyl function and the cosolvent upon the enantioselectivity are demonstrated by the pig pancreas lipase-catalyzedhydrolysis of the methoxycarbonyl substituted diacylated propanediol corresponding to the monoesters 18. Protected glycerol monoacetate 19 can be prepared in acceptable enantiomeric purity and yield by crude pig pancreas lipase or Pseudomoms Juorescens lipase-

418

I

1 1 Hydrolysis and Formation of C - 0 Bonds Table 11.1-11. Lipase-catalyzed enantiotopos-differentiating hydrolysis of prochiral cyclic diol dialkanoates in aqueous solution (CCL Candida Gylindracea lipase, PFL PseudomonasPuorescens lipase, M M L Mucor miehei lipase, CVL Chromobacterium uiscosum lipase, PPL pig pancreas lipase, M J L Mucorjauanicus lipase, RSL Rhizopus sp. lipase, PCL Pseudomonas cepacia lipase, CCL, Ceotricum candidum lipase, ANL Aspergillus niger lipase, FSPC Fusarium solani pisi cutinase, CRL Candida rugosa lipase, CAL-B Candida antarctica B lipase, LIP Pseudomonas sp. lipase-Toyobo, RDL Rhizopus delemar lipase, MSL Mucor sp. lipase, CAL Candida antarctica lipase, not specified).

1

R

72 % ee, 94 % yield, PPL 93 % ee, 74 %yield, PPL 90 % ee, 83 %yield, PFL n-Pr 94 % ee, 84 %yield, PPL 299 % ee, 100 %yield, PPL

Me

[l]

4-11 40 % ee, 75 %yield, PPL

[2] [3] [2] [4]

4

3 OH R

88 % ee, 94 % yield, PPL 96 % ee, 78 %yield, PPL 295 % ee, 87 % yield, PFL n-Pr 96 % ee, 80 % yield, PPL Me

[l] [2] [3]

R

n-Pr

88 % ee, 94 % yield, DPL 89 % ee, 90 % yield, PPL 295 % ee, 86 %yield, PFL 96 % ee, 80 %yield, PPL

C K : 60 % ee, 75 %yield, PPL

[6] [7]

O

6

[l]

5

Me

86 % ee, 57 %yield, PPL 297 % ee, 75 %yield, PFL

[l]

[2] 131 [l]

C

o

A OHc

50 % ee, 70 %yield, PPL 77 % ee, 78 %yield, PPL (purified PPL)

7 151 50 % ee, 71 %yield, PPL

[5] [6]

11.1 Hydrolysis and Formation of Carborylid Acid Esters

I

419

Table 11.1-11.

(cont.).

30 % ee, 75 % yield, PPL

78 % ee, 77 % yield, PPL 93 % ee, 86 %yield, PPL (purified PPL)

R’

R’

HO 0-t-Bu OMe OEt H H H H H H

H H H H OAc

C1 C1 SPh SOzPh N3

68 % ee, 80 yield, PPL 26 % ee, 80 yield, PPL 52 % ee, 65 yield, PPL 66 % ee, 78 yield, PPL 90 % ee, 60 yield, PPL 88 % ee, 81 yield, PPL 95 % ee, 88 yield, PPL 96 % ee, 65 yield, PPL 68 % ee, 72 yield, PPL 91 % ee, 85 yield, PPL

41 % ee, 73 % yield, PPL 81 % ee, 85 % yield, PPL 98 % ee, 91 %yield, PFL

(purified PPL)

14

13 [7-91

13 % ee, 70 %yield, PPL

R

0 % ee, -, PFL 97% ee, 88 yield, PFL

Me

n-Pr

78 % ee, 81 yield, PPL 87 % ee, 67 yield, PPL 50 % ee, 54 yield, PPL 84 % ee, 24 yield, PPL

coAc OH

298 % ee, 96 %yield, PPL 92 % ee, 90 %yield, PFL

15

[l,101

[31 R

Me Et i-Pr

tBu

41 % ee, 35 % yield, PPL 33 % ee, 57 %yield, PPL 32 % ee, 68 %yield, PPL 16 % ee, 71 %yield, PPL

[l]

[2] [3] (21

420

I

7 7 Hydrolysis and Formation ofC-0 Bonds

Table 11.1-11.

(cont.).

R

80 % ee, 20 %yield, CCL 26 % ee, 69 % yield, CCL 28 % ee, 75 % yield, CCL 4 % ee, 71 %yield, CCL n-Bu 299 % ee, 75 %yield, MJL 68 % ee, 85 %yield, CCL

Me Et i-Pr t-Bu

99 % ee, 84 %yield, PFL 99 % ee, 66 % yield, MJL 85 % ee, 50 %yield, PPL 75 % ee, 37 %yield, CCL

-CoconPI

20 [S, 141

>< :OAC

OH

LOH

55 % ee, 77 %yield, PPL

96 % ee, 79 % yield, PFL

(IoA 22 [16-181

OH

83 % ee, 26 % yield, CCL

299% ee, 39% yield, PFL

0

BnOQoH

PH

OAc

50 % ee, 82 % yield, CCL 66 % ee, 83 % yield, RSL 92 % ee, 87 %yield, PPL 92 % ee, 84 %yield, PFL 95 % ee, 85 % yield, MML 91 % ee, 76 %yield, CVL >99 % ee, 85 %yield, MSL 99 % ee, 90 % yield, CAL-B

24 123)

OAc

36 % ee, 21 %yield, ANL 18 % ee, 1 3 % yield, PFL 84 % ee, 16 %yield, PFL

7 1 . I Hydrolysis and Formation ofCarboxylid Acid Esters

I

421

Table 11.1-11.

(cont.).

q ‘b0n .

R R

25 [24]

G

O

B

n

26 [25]

OAc N3

OAc

H 44 % ee, 40 %yield, CCL Me 100 % ee, 61 %yield, CCL

297 % ee, -, GCL

NM le 27 [25]

AcO

6

28 [25]

OAc

297 % ee, GO % yield, PCL

r O H

6

298 % ee, 100 % yeld, ANL

29 [26, 271

30 [28]

OAc

95 % ee, 90 %yield, PPL

79 % ee, 64 %yield, PCL

0

r O A C

ent-30 [29]

OH 297 % ee, -, CRL

31 [30, 311

O ‘H 55 % ee, 98 %yield, PPL 30 % ee. 45 % yield, CCL

32 [32]

33 [32]

6 OCOPh ; O ”’x0

295 % ee, 94 % yield, FSPC

34 [33]

HO

35 % ee, -, PCL

o::b

295 % ee, 87 %yield, FSPC

AcO 298 % ee, 89 %yield, PCL

35 [33]

422

I

1 1 Hydrolysis and Formation of C-0 Bonds Table 11.1-11.

(cont.).

emoH / 3

0

OAc 36[34]

37 [35]

0

95 % ee, 66 % yield, PPL

70 % ee, 80 % yield, PCL

38 [35]

>98 % ee, 88% yield, PCL

39 [36]

96 % ee, 72 % yield, PFL OTBDMS

I

40 [371

41 [38]

LOA~

>95 % ee, 89 % yield, PSL

HO

A 0

>95 % ee, 70 % yield, CAL

42 [38]

AcO &

L

Ac

O

H

43 [39]

I

Cbz

86 % ee, 100 %yield,CAL

>98 % ee, 82 %yield,ANL

OMOM

I

C0,Me

44 [391

45 [40]

AcO&OH

Cbz

HO ""

>98 % ee, 76% yield, ANL

O,,

77 % ee, 39 % yield, PFL

C0,Me

HO""

"OAc

C0,Me

46 1401

47 [40]

"OAc

Me 88 % ee, 49 %yield, PFL

Me

>99 % ee, 89 %yield, DSL

7 7 . 7 Hydrolysis and Formation of Carboxylid Acid Esten

I

423

Table 11.1-11.

(cont.).

C0,Me

C0,Me

48 [40]

48 [40] Et

Et

87 % ee, 49 % yield, PPL

>99 % ee, 88 % yield, PFL

50 [41]

49 [41] AcO "*

>99 % ee, 51 % yield, LIP

>99 % ee, 53 %yield, LIP

"::0$

51 [42]

co(:;

52 [42]

OH 95 % ee, 86 %yield, RDL 33 % ee, 73 %yield, PFL

OH >99 % ee, 95 % yield, RDL >99 % ee, 61 %yield, PFL 94 % ee, 60 % yield, PPL

rOAc

rOAC

54 [42]

53 [42]

95 % ee, 95 % yield, RDL 16 % ee, 60 %yield, PPL

95 % ee, 64 % yield, RDL 87 % ee, 39 %yield, PPL

55 [43]

I

56 [43]

Cbz

Bn

>98 % ee, 77 % yield, PFL

>98 % ee, 73 %yield, PFL

57 [441

>99 % ee, 70 %yield, CAL-B

Ac""''Q

58 [45]

O -w R: CHz-CH=CHz, CHI-CH=CHCH~(E), CH2-CH=C(Cl)CH,(E),CH2-CrC-CH3, CHI-Ph >98 % ee, 62-80 % yield, CCL

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters 40 Y. Zhao, Y Wu, P. De Clercq, M. Vandewalle, P. Maillos, J.-C. Pascal, Tetrahedron: Asymmetry 2000,l I , 3887-3900. 41 H. Konno, K. Ogasawara, Synthesis 1999, 1135. 42 M. Tanaka, Y. Norimine, T. Fujita, H. Suemune, K, Sakai,]. Org. Chem. 1996,61,6952. 43 B. Danieli, G. Lesma, D. Passarella, A. Silvani, /. Org. Chem. 1998.63, 3492.

44 F. Theil, S . Ballschuh, M. von Janta-Lipinski, R. A. Johnson,I. Chem. Soc., Perkin Trans. 1 1996, 255. 45 P. Renouf, J:M. Poiner, P. Duhamel, ]. Chem. SOC., Perkin Trans. 1 1997, 1739. 46 T. Taniguchi, R. M. Kanada, K. Ogasawara, Tetrahedron: Asymmetry 1997,8, 2773. 47 K. Toyama, S. Iguchi, T. Oishi, M. Hirama, Synlett, 1995,1243.

catalyzed hydrolysis of the corresponding diacetate. Here, too, enantioselectivity of the hydrolysis by crude pig pancreas lipase is considerably improved if the reaction is run in the two-phase system composed of water and diisopropyl ether. Glycerol diacetate derivatives with chain type substituent in the 2-position are hydrolyzed with crude pig pancreas lipase in a two-phase system composed of water and hexane to the monoacetates 21 with good enantioselectivity. Hydrolysis in aqueous solution alone is much less selective. The 2-benzyloxycarbonylaminosubstituted propanediol monoacetate 20 is also accessible with a high ee value by pig pancreas lipase-catalyzed hydrolysis. Monoacetates 23-27 can serve as a good illustration of the scope of lipases because of the number of different species available. The monoacetate 24 is a notable example since it documents the surprising ability of Pseudomonas fluorewens lipase to differentiate between enantiotopic groups located relatively far from the stereogenic ring atoms. Monoacetates 27 and 29 are of opposite configuration compared to those obtained from the same achiral diacetates via pig liver esterase-catalyzedhydrolysis (Table 11.1-3). Monoacetates of Table 11.1-10 which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-4 and 11.1-17. One of the most successful applications of lipases lies in the hydrolysis of cyclic meso-configured dialkanoates, mainly diacetates, to the corresponding chiral monoalkanoates (1-61) (Table 11.1-11). However, the attainment of high enantioselectivity is not restricted to primary dialkanoates. Cyclic secondary dialkanoates are good substrates too (Table 11.1-11).There seems to be no restriction in regard to the ring size. Heterocyclic systems are tolerated by the various lipases. Reversal of enantiotopic group recognition in a series of structurally closely related substrates as frequently observed in the case of pig liver esterase-catalyzed hydrolysis is usually not observed with a lipase. This is illustrated by the series of cyclopentanoid dimethanol diacetates 7-12. Enantioselectivitycan be enhanced in many cases with a given lipase by either resorting to hydrolysis in a two-phase system, addition of a cosolvent, or changing the nature of the acyl group (17). If these measures fail, resorting to another lipase may lead to success. This is exemplified by the cyclopentenoid monoacetate 23, which is obtained by Candida cylindracea lipase with an ee value of 50 %, by Pseudomonasfluorescens lipase with an ee value of 92 % and by Mucor sp. lipase or by Candida antarctica B lipase with ee values of 2 99%. A frequently encountered synthetically very attractive situation is illustrated by the synthesis of the enantiomeric monoacetates 30 and ent-30. The two enantiomers are accessible with two different lipases. Tetrahydropyran derivatives 37, 38, 41 and 42 as well as the piperidine derivatives 44, 45, 55 and 56 can be prepared with high enantiomeric purity. Bi- and tricyclic

I

425

426

I

11 Hydrolysis and Formation ofC-0 Bonds Lipase-catalyzed enantiotopos-differentiating hydrolysis of prochiral acyclic and cyclic dicarboxylic acid diesters in aqueous solution (CCL Candida cylindracea lipase, PPL pig pancreas lipase, PSL Pseudomonas sp. lipase, CVL Chromobacterium viscosum lipase, CE cholesterol esterase).

Table 11.1-12.

R?. CO,H

R'

1

2 3 4

5 5 6 7

8

("5)

yield (%)

R'

R2

R3

Lipase

ee

CF, F

H

Me Me Me Et Et Et Et Et Et

CCl CCL cc1 CCL

n o hydrolysis 99" 95 91 61 62a 93" 33" lla

F F F F F F F

Et Me Me H H Et n-Pr

n-Bu

PPL CCL CCL CCL CCL

Ref.

PI PI PI PI PI PI PI PI PI

-

87 74 87 23 70 87 30

78

a Absolute configuration n o t determined

S/\/CO,H R~*sc/\ozR~ /

R'

CI

R2

Me CH2CONEt2

298 % ee, 90 %yield, PSL, CVL 298 % ee, 90 %yield, CCL

10 151 97 % ee, 97 %yield, PPL

[31 [3,41

11 [51 6 % ee, 48 %yield, PPL

MeoxcozH 13 161

Me0

PPL, n o hydrolysis

C0,Me

92 % ee, 90 % yield, CCL

11.7 Hydrolysis and Formation ofcarboxylid Acid Esters

I

427

Table 11.1-12.

(cont.).

aNo2 14 [7-91

H O z C ~ ~ ~ H z O C O t B u Me

I

CH,OMe

ens14 [9]

HO,C

Me

I

Me

CH,OMe

299 % ee, 80 % yield, lipase B,

89 % ee, 87 % yield, PSL,

diisopropyl ether/H20 299 % ee, 80 % yield, PSL, diisopropyl ether/HzO

diisopropyl ether/H20

90 % ee, 95 % yield, CE 1 T. Kitazume, T.Sato, N. Ishikawa, Chem. Lett.

G H. J. Bestmann, U. C. Philipp, Angew. Chem. 1991, 103,78;Angew. Chem. Int., Ed. Engl. 1991,30,86. 1984,1811. 7 H. Ebiike, Y.Terao, K. Achiwa, Tetrahedron Lett. 2 T. Kitazume, T.Sato, T. Kobayashi, J. Tain Lin, j . Org. Chem. 1986,51,1003. 1991,32,5805. 3 D.L. Hughes, J. J. Bergan, J. S. Amato, P. J. Reider, 8 H. Ebiike, K. Maruyama, K. Achiwa, Tetrahedron: E. J. J. Grabowski, j . Org. Chem. 1989,54,1787. Asymmetry 1992,3,1153. 9 Y. Hirose, K. Kariya, J. Sasaki, Y. Kurono, 4 D. L. Hughes, 2. Song, G . B. Smith, I. J. Bergan, G. C. Dezeny, E. J . J . Grabowski, P. J. Reider, H. Ebiike, K. Achiwa, Tetrahedron Lett. 1992,33, Tetrahedron: Asymmetry 1993,4,865. 7157. 5 Y. Nagao, M. Kume, R. C. Wakabayashi, 10 R.Chenevert, R. Martin, Tetrahedron: Asymmetry 1992,3,199. T. Nakamura, M. Ochiai, Chem. Lett. 1989,239.

derivatives such as 36, 40, 49, and 50 are obtained from the corresponding mesodiacetates. The monoacetates 58, 59 and GO (Table 11.1-11)are products of the hydrolysis of prochiral enol diacetates. Monoalkanoates of Table 11.1-11which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-3, 11.1-7, 11.1-9 and 11.1-18. A limited number of acyclic and cyclic prochiral dicarboxylic acid diesters were found to be good substrates for hydrolysis catalyzed by lipases (Table 11.1-12). Notable examples which give a good illustration of the potential of hydrolases as well as of the trial and error approach one relies on to a certain extent are the dithio acetal derivative 9 and the fluoro alkyl malonates 1-8. The dithio monoester 9 is obtained with different lipases with high enantioselectivities and yields despite its remote chiral center. Candida cylindracea lipase is the enzyme of choice for the synthesis of fluoro alkyl malonates with small alkyl groups. An astonishing observation was

428

I

I 1 Hydrolysis and Formation of C - 0 Bonds Table 11.1-13. Lipase-catalyzed enantiomer-differentiating hydrolysis of racemic carboxylic acid esters and lactones in aqueous solution (PPL pig pancreas lipase, PSL Pseudomonas sp lipase, PFL PseudomonasPuorescens lipase, CCL Candida cylindracea lipase, ANL Aspergillus niger lipase, PCL Pseudomonas cepacia lipase, CAL-A Candida antarctica A lipase, CRL Candida rugosa lipase, CAL Candida antarctica lipase, not specified).

0 I)\/CO,H

l a PI

0:&CO,nC,H,

1b 111

295 % ee, -

-, - PPL, 60 % conversion

0 -_-H C O ,S ,,

R

R = Ph, p-NOzCsH4, p-MeOC&h, PhCH2, c - C ~ H I I 80-100 % ee, 20-30 %yield, PSL 50 % conversion

Me ,,,, CO,Me 3a [3,4]

Mey""."e C0,Me

(C0,H

296 % ee, -

95 % ee, 47 % v e l d , PPL 50 % conversion

\CO H 98 % ee, 45 %yield, PPL 50 % conversion

...C0,Me Me0

__

' C O Me

5a 151

Me0

CO,H

(15 % ee, 55 % yield

95 % ee, 40 % yield, PSL 42 % conversion

OH

rcoz 6a 161

/

92 % ee, 33 %yield, PFL 35 % conversion

99 % ee, 43 % yield 55 % conversion

7 7 . 7 Hydrolysis and Formation of Carboxylid Acid Esters

I

429

Table 11.1-13.

(cont.).

R MedC02H

R = OH:79 % ee, 48%yield, PFL 50 % conversion R = F: 69% ee, 53 %yield, PFL GO % conversion R = Br: 69% ee, 48% yield, PFL

7a

95% ee, 43 % yield

7b [61

8a

299 % ee, 37 % yield

8b [61

9a

73 % ee, 40YOyield

9b161

10a [7]

CF3

O2N3COzH

02NdC02Bn

298 % ee, -, PFL, 35 % conversion

98% ee, -

ye

10b [7]

llb [S]

lla [8]

DCozH

Me0

/

/

Me0

98% ee, -, CCL, 39 % conversion

63 % ee, -

96 % ee, 43 % yield, CCL 89 % ee, 49 % yield, PPL 93 % ee, 31 % yield, CRL

[I61

94% ee, 48 % yield 299 % ee, 49 % yield 94% ee, 46 % yield

14a [17]

)~).~*‘co2nBu “‘Me

93 % ee, 35 % yield, CCL 50% conversion

94% ee, 47% yield

[161

14b [17]

430

I

7 I Hydrolysis and Formation of C-0 Bonds Table 11.1-13.

(cont.).

xTC

C0,nBu 15a [171

15b [17]

*‘

OZH

71 % ee, 53 %yield, PPL 57 % conversion

295 % ee, 40 % yield

42 % ee, 38 % yield, CCL 69 % conversion

95 % ee, 19 % yield

ypF ”Me

77 % ee, 35 %yield

95 % ee, 41 %yield, CCL 55 % conversion

18a [6]

HOm

C

O

,

18b [6]

HO

H

299 % ee, 45 % yield

75 % ee, 50 %yield, PFL 55 % conversion

19a (181

-, -, PCL 55 % conversion

~

o

~

c o Me NO2 90 76 ee, 52 % yield

z

M19b [18] e

Me

R7?CozCH2CN 0 Et 80 % ee, 49 %yield, PPL 20a CsHll 85 % ee, 49 %yield, PPL 21a PhCH2 85 % ee, 49 %yield, PPL 22a

293 % ee, 44 % yield 298 % ee, 49 %yield 295 % ee, 49 % yield

23b [20]

23a [20]

299 % ee, -, PPL

20b [19] 21b [19] 22b [19]

2e99 % ee, -

7 7 . 7 Hydrolysis and Formation of Carboxylid Acid Esten

I

431

Table 11.1-13.

(cont.).

PhL C 0 , H

24a [2l]

93 % ee, -, PSL

OH /\/CO,Et Ph

24b [21]

98 % ee, OAc

25a [21]

94 % ee, -, PSL

96 % ee, -

BnO Me

Me OBn

x

x

R CO,H R Et n-Pr ally1 n-CsHt3 n-CgH19

R CO,H

81 % ee, 41 % yield, CCL 26a 95 % ee, 40 %yield, CCL 27a 299 % ee, 46 % yield, CCL 28a 299 % ee, 38 % yield, CCL 29a 30a 94 % ee, 46 % yield, CCL

HO,C-(CH,),

R Me Me

25b [21]

L C O , E t Ph

GO % ee, 54 % yield 70 % ee, 18 % yield 82 % ee, 52 % yield 67 % ee, 35 % yield 67 % ee, 8 % yield

L,,

OAc

RO,C-(CH,),

n 4 28%ee,-,CCL 8 68%ee,-,CCL n-Bu 8 299%ee,-,CCL

31a 32a 33a

26b [22] 27b [22] 28b [22] 29b [22] 30b [22]

-,-,-,-

‘61

31b [23] 32b [23] 33b [23]

OH

R

R LCOzMe

CO,H

R

n-Pr n-Bu n-C5H1l n-C6H13 n-C7Hls n-CsH17

84 % ee, 39 %yield, PPL 82 % ee, 40 % yield, PPL 74 % ee, 50 %yield, PPL 82 % ee, 48 % yield, PPL 85 % ee, 43 %yield, PPL 83 % ee, 48 %yield, PPL

34a 35a 36a 37a 38a 39a

R

R=Et R = n-C7H15

75 % ee, PPL 76 % ee, PPL

40 [25] 41 [25]

75 % ee, 47 % yield 77 % ee, 46 % yield 95 % ee, 40 %yield 93 % ee, 43 % yield 85 % ee, 45 % yield 88 % ee, 5 % yield

34b [24] 35b [24] 36b [24] 37b [24] 38b [24] 39b [24]

432

I

7 7 Hydrolysis and Formation of C-0 Bonds Table 11.1-13.

(cont.).

Me0,C

A

0

>99 % ee, 30 % yield, CAL-A >85 % ee, 54 % yield, CAL-A

42a [26] 42a [27]

GO % ee, 56 %yield >99 % ee, 41 %yield

43a [28]

P

Me0,C

CO,H

42a [26] 42a [27]

43b [28]

92 % ee, -

-, 60 % conversion, CRL

44a [29]

R o ~ C ~ N H A c 441,[29] R n-Bu >99 % ee, 42 % yield n-Hex >99 % ee, 44 % yield

>99 % ee, -, CCL

45b [30]

45a [30] Ph

PhkCoZMe

89 % ee, -, PCL

>98 % ee, 46 % yield, PCL

L C O , M e

46b [31]

0

84 % ee, 80 % yield, CCL

-, 97 % yield CO,H

C0,Et

47a [32]

ACN

p-Me-Ph s''s'/hN

p-Me-Ph

99 % ee, 41 % yield, CRL

98 % ee, 46 % yield

F

47b [32]

F

C0,nOctyl

48a [33]

73 % ee. -

93 % ee, 19 %yield, ANL (purified)

HN ,

.nQ CO,H

96 % ee, 40 % yield, lipase L

OH

48b [33]

49a [34] H,N

l *

C02Et

>99 % ee, 50 % yield

49b [34]

1 1 . I Hydrolysis and Formation ofcarboxylid Acid Esters

I

433

d., do

Table 11.1-13.

fc0nt.l.

-

5Oa [35]

"COOH

96 % ee, 42 % yield, CCL

A~....x-coo,, 5Ob [35]

91 % ee, 42 % yield

51 1361

HO,C

51b [36]

35 % ee, 22 % ee, -

89 % ee, -, PPL, 28 % conversion 82 % ee. -; CRL, 21 % conversion

P 0

II

52 1371

82 % ee, CAL, 47 % conversion

53 [37]

72 % ee, CAL, 59 % conversion

54 [37]

c;-

55 [37]

92 % ee, CAL, 50 % conversion

94 % ee, CAL, 50 % conversion

R O L C 0 2 H

OH RO&CO,Me

R

4-Me0-C6H4 2-AIlyl-C~H4 2-Naphthyl

99 % ee, -, PCL 98 % ee, -, PCL 99 % ee, -, PCL

56a 58a

97 % ee, -, PCL 98 % ee, -, PCL 98 % ee, -, PCL

59a [39]

H, C 5 y C 0 , E t

57a

,

56b [38] 57b [38] 58b 1381

59b [39]

0 98 % ee, 13 %yield, PPL 93 % ee, 13 %yield, PCL

-

1 D. Bianchi, W. Cabri, P. Cesti, F. Francalanci,

M. R i d , / . Org. Chem. 1988,53,104. 2 K. Burgess, I. Henderson, Tetrahedron Lett. 1989, 30,3633. 3 E. Guibe-lamuel, G. Rousseau, 1. Salaiin, 1. Chem. .~ Soc., Chem. Commun. 1987,1080.

26 % ee, 67 % yield 10 % ee, 80 % yield 4 J. Salaiin, B. Karkour, Tetrahedron Lett. 1987,28,

4669. 5 J.-P. Barnier, L. Blanco, E. GuiK-jampel, G. Rousseau, Tetrahedron 1989,45,5051. G P. Kalaritis, R. W. Regenye, J. J. Partridge, D. L. Coffen, j . Org. Chem. 1990,55,812.

434

I

1 1 Hydrolysis and Formation ofC-0 Bonds 7 T. Yamazaki, T. Ohnogi, T: Kitazume, Tetrahedron: Asymmetry1990, 1, 215. 8 Q:M. Gu, C:S. Chen, C. J. Sih, Tetrahedron Lett. 1986,27,1763. 9 Q:M. Gu, D. R. Reddy, C. J. Sih, Tetrahedron Lett. 1986,27,5203. 10 R. Demoncour, R. Azerad, Tetrahedron Lett. 1987, 28,4661. 11 B. Cambou, A. M. Klibanov, Biotechnol. Bioeng. 1984,2G, 1449. 12 R. Chhevert, L. D'Astous, Can.]. Chem. 1988,66, 1219. 13 2:W. Guo, C. J. Sih,]. Am. Chem. SOC.1989, 111, 6836. 14 S.-H. Wu, Z.-W. Guo, C. J. Sih,]. Am. Chem. SOC. 1990, 112,1990. 15 B. Loubinoux, C. Viriot-Chauveau. J. L. Sinnes, Tetrahedron Lett. 1992, 33, 2145. 16 1. J. Colton, S . N. Ahmed, R. 1. Kazlauskas, J. Org. Chem. 1995, GO, 212. 17 M. Pottie, J. Van der Eycken, M. Vandewalle,J. M. Dewanckele, H. Roper, Tetrahedron Lett. 1989, 30, 5319. 18 S . KnesoviS, V. Sunjit, A. Lhai, Tetrahedron: Asymmetry1993,4,313. 19 L. Blanco, G. Rousseau, J:P. Bamier, E. Guibe-Jampel,Tetrahedron: Asymmetry1993,4, 783. 20 R.-L. Gu, C. J. Sih, Tetrahedron Lett. 1990, 31, 3283. 21 N. W. Boaz,]. Org. Chem. 1992,57,4289. 22 T. Sugai, H. Kakeya, H. Ohta,]. Org. Chem. 1990, 55,4643. 23 U. T. Bhalerao, L. Dasaradhi, P. Neelakantan, N. W. Fadnavis,J . Chem. SOC., Chem. Commun. 1991.1197.

24 P. Allevi, M. Anastasia, P. Ciuffreda, A. M. Sanvito, Tetrahedron: Asymmetry1993,4, 1397. 25 L. Blanco, E. GuibC-Jampel,G. Rousseau, Tetrahedron Lett. 1988, 29, 1915. 26 J. Kingery-Wood, 1. S . Johnson, Tetrahedron Lett. 1996,37,3975. 27 E. W. Holla, H:P. Rebenstock, B. Napierski, G. Beck, Synthesis1996, 823. 28 C. M. Schueller, D. D. Manning, L. L. Kiessling, Tetrahedron Lett., 1996, 37, 8853. 29 R. Csuk,.'l Don; Tetrahedron: Asymmetry1994,5, 269. 30 G. Varadharaj, K. Hazell, C. D. Reeve, Tetrahedron: Asymmetry1998,9,1191. 31 Y. R. Santosh h i , D. S. Iyengar, Synthesis,1996, 594. 32 Y. Takeuchi, M. Konishi, H. Hori, T. Takahashi, T. Kometani, K. L. Kirk, Chem. Commun.1998, 365. 33 M. C. Ng-Youn-Chen,A. N. Serreqi, Q. Huang, R. J. Kazlauskas,]. Org. Chem. 1994,59, 2075. 34 D. M. Spero, S. R. Kapadia,]. Org. Chem. 1996, 61, 7398. 35 A. Bhaskar Rao, H. Rehman, B. Krishnakumari, J. S . Yadav, Tetrahedron Lett. 1994, 35,2611. 36 G. Pitacco, A. Sessanta o Santi, E. Valentin, Tetrahedron: Asymmetry2000, 11, 3263. 37 K. Shioji, A. Matsuo, K. Okuma, K. Nakamura, A. Ohno, Tetrahedron Lett. 2000, 41,8799. 38 K. Wiinsche, U. Schwaneberg, U. T. Bornscheuer, H . H . Meyer, Tetrahedron: Asymmetry1996,7, 2017. 39 F. Benedetti, C. Forzato, P. Nitti, G. Pitacco, E. Valentin, M. Vicario, Tetrahedron: Asymmetry 2001, 12, 505.

made in the case of the dihydropyridine ester 14 and ent-14. Both enantiomers are obtained with high ee values and in high yields by Pseudomonas sp. lipase-catalyzed hydrolysis merely upon changing the reaction medium from diisopropyl ether to cyclohexane, both saturated with water. The limitations of the lipase-catalyzed hydrolysis of carboxylic acid esters are evident too. Whereas the cyclohexenoid diester 10 is obtained through pig pancreas lipase-catalyzed hydrolysis with high enantioselectivity,the cyclopentanoid monoester 11 is formed only with low selectivity and the cyclopentanoid diester 12 is not a substrate for pig pancreas lipase. An interesting example for the use of a cholesterol esterase is the cyclopentanoid monoester 15. Monoesters of Table 11.1-12 which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-1,11.1-2 and 11.1-7. The usefulness of lipases for the enantiomer-differentiating hydrolysis of carboxylic acid esters and lactones is impressively demonstrated by examples 1-59 of Table 11.1-13. This broad substrate spectrum is covered mainly by lipases from Candida cylindracea (rugosa), pig pancreas and several Pseudomonas sp. lipases. Carboxylic acid esters having the alkoxycarbonyl group attached to a secondary, tertiary or even quaternary carbon atom are substrates. Thus, in contrast to

I

7 7 . 7 Hydrolysis and Formation ofcarboylid Acid Esters 435 Table 11.1-14. Lipase-catalyzed enantiomer-differentiating hydrolysis of esters of racemic primary alcohols in aqueous solution (PPL pig pancreas lipase, PCL Pseudomonas cepacia lipase, PCL-A Pseudomonas cepacia lipase, Sumitomo, PSL Pseudomonas sp. lipase, PAL Pseudomonas aeruginosa lipase, HLL Humicola lanuginosa lipase, CAL-B Candida antarctica B lipase, CRL Candida rugosa lipase).

C I T O A c

CI 90 % ee, -, pancreatin 60 % conversion" NHC0,Et Me

90 % ee, 30 % yield, pancreatin

Me&OAc 295 % ee. -

4 [31

O P o H ClLlHZ, 295 % ee, 32 % yield, PPL 60 % conversion",

295 % ee, 20 % yield, PPL 20 % conversiona, OCOnPr

~[4]

MeToconPr 295 % ee, -, PPL 60 % conversiona

295 % ee, -, PPL 58 % conversiona

Me

8 [41

, + o c o n P r

nPrToConPr Me

56 % ee, -, PPL 60 % conversion"

T O C o n P r

77 % ee, -, PPL 60 % conversion"

73 % ee, -, PPL 60 % conversion"

E

t

F

n

P

82 % ee, -, PPL 60 % conversiona

GOAC 295 % ee, 30 %yield", PPL

r

436

I

1 1 Hydrolysis and Formation of C - 0 Bonds Table 11.1-14.

(cont.).

-

12a [6]

1Zb [GI

+Ac

\OAc

295 % ee, 47 % yield

91 % ee, 45 %yield, PSL

.-“OH

n ,-,KO

0

0

R =H R= i-Pr R = t-Bu R = Ph

97 % ee, -, PAL 297 % ee, -, PAL 297 % ee, -, PAL 96 % ee, -, PAL

13a 14a 15a 1Ga

13b [7] 14b [8,91 15b [8, 91 1Gb [7]

99 % ee, 297 % ee, 297 % ee, 100 % ee, -

nProco”‘,*~Q

17b [lo]

17a [lo] I Boc

I Boc

91 % ee, 42 % conversion

94 % ee, -, PCL 53 % conversion

18a [Ill

99 % ee, 37 %yield

61 % ee, 54 % yield, CAL-B

b

19a [12]

TOnPr

N\

ND:::::;)

87 % ee, -, PCL, 46 % conversion

>98 % ee, -, PCL, 61 % conversion

20a [12]

87 % ee, -, PCL, 47 % conversion


>98 % ee, -, PCL, 60% conversion

19b [12]

20b [12]

7 7.7 Hydrolysis and Formation of Carboxylid Acid Esters Table 11.1-14. (cont.).

OCOnPr

OH 21a [12]

90 % ee, -, PCL, SO % conversion

21b [12]

90 % ee, -, PCL, 50 % conversion

/OConPr

22a [12]

N oJ ):! 93 % ee, -, PCL, 46 % conversion

22b [12]

"0

>98 % ee, -, PCL, 55 % conversion

bOH 23a [13]

>99 % ee, 11 %yield, PCL

95 K % ee, O41 %yield, H PCL

91 % ee, 35 % yield, PCL 86 % ee, 29 % yield, PCL 88%ee, 31 % yield, PCL

R

,OAc

p H

. .

23b [13]

16 % ee, 85 % yield

24a [14]

broAc 24b [14]

...".,.,

96 % ee, 41 % yield, PCL

Me

25a [15]

n-Pr

2Ga [15]

n-Bu

27a [15]

>98 % ee, 27 % yield 92 % ee, 34 % yield 9G % ee, 33 % yield

R Me

25b [lS]

n-Pr

2Gb [15]

n-Bu

27b [lS]

I

437

438

I

I 7 Hydrolysis and Formation of C - 0 Bonds (cont.).

Table 11.1-14.

H O Y S R

28a [16]

A c O T S R

OAc

28b [16]

OAc

47-939 % ee, 3 4 5 7 % yield, PCL

48->96 % ee, 34-50 % yield, PCL

Q

\CO,Me

-CO,Me

\CO,nBu

NHCbz +CO,Me

OEt

>99 % ee, 50 %yield, PPL

>99 % ee, 43 % yield, PPL

30b [18]

30a [18]

iPr0,C

H

~ ; ; A C H

98 % ee, 45 % yield

96 % ee, 50 % yield, HLL

31a [19]

31b [19]

..H ,O ,,

AcO

91 % ee, 26 % yield, PPL

37 % ee, 72 %yield, PPL

7 1 . 7 Hydrolysis and Formation of Carboylid Acid Esters Table 11.1-14.

(cont.).

96 % ee, 23 % yield >98 % ee, 21 %yield >98 % ee, 34 % yield >98 % ee, 25 % yield 95 % ee, 28 % yield 97 % ee, 27 % yield 92 % ee, 12 % yield 92 % ee, 28 % yield

R'

R2

H

COzMe

H

H

OMe H R'

=

lipase

32a[20] PCL PSL 33a 1201 PCL PSL 343 [20] PCL PSL 35a [20] PCL PSL

33%ee, 72%yield 44 % ee, 72 %yield 51 % ee, 57 % yield 34 % ee, 73 % yield 49 % ee, 65 %yield 40 % ee, 67 % yield 18 % ee, 72 %yield 52 % ee, 61 %yield

32b[20] 33b [20] 34b [20] 35b [20]

kMe

?U?

R ~ = H

OAc

OH

/

( 36a [211

,,,OMe

>98 % ee, 37 % yield, PPL, 38 % conversion -,-, PPL, 70 % conversion 79 % ee, -, PSL, 36 % conversion -,-, PSL, 69 % conversion

198 % ee, 29 % yield __

>98 % ee

37a [22]

NHCOC,,H,,

H 0 A / C 1 3 H 2 7

OAc

49 % ee, 54 % yield, PCL-A 96 % ee, 41 % yield, PCL-A (immobilized)

3Gb [2l]

NHCOCF,

NHCOCF,

NHCOCl,H,5

(--J

OH 98 % ee, 7 % yield

3% [22]

I

439

1 I Hydrolysis and Formation ofC-0 Bonds Table 11.1-14.

[cont.).

NHCOC,&,

37c [22] Ac0-?13H27

OAc

87 % ee, 38 %yield, PCL-A G9 % ee, 58 %yield, PCL-A (immobilized) 38a 1231

HOr Q - 0

38b [23]

/'+,'

AcO

79 % ee, -, PCL, 50 % conversion

84 % ee, -

39a [23] HO

391, [23] AcO 94 % ee, -

89 % ee, -, PCL, 53 % conversion

40a [24]

/"""

40b [23]

AcO 95 % ee, -

HO 96 % ee, -, PCL, 50 % conversion

.(3;.

41b [24]

41a [25] PhAOAc 53 % ee, 78 % ee, -

Ph-OH 78 % ee,-, PCL 82 % ee, -, CAL-B

"17

R2

O K0N d o H

75 % ee, 42 % yield 62 % ee, 50 % yield 89 % ee, 51 % yield

R

R'

RZ

Et

Ph

H

42a [25]

PCL

lipase

70%ee, 46%yield,

42b [25]

t-Bu

Ph

H

42a [25]

PCL

67 % ee, 47 %yield

431, [25]

Et

H

Ph 44a [25]

PCL

93 %ee, 42 %yield

44b[25]

7 7 . 7 Hydrolysis and Formation ofCarboxylid Acid Oten

I

441

Table 11.1-14.

87 %ee, 4 2 % yield 97%ee,44% yield 94%ee,43% yield 72%ee,46% yield 69%ee, 52% yield 97%ee, 50% yield, 0 "C

1

(cont.).

H

Ph

44a

PSL

90 % ee, 46 %yield

44b [25]

n-Pent H

Ph

44a

PCL 92 % ee, 50 %yield

45b [25]

Et

H

Bn

46a

PCL 91 % ee, 46 %yield

461,[25]

Et

H

Bn

46a

PSL

76 % ee, 44 %yield

461,[25]

Et

H

Et

47a

PCL 98 % ee, 40 %yield

47b [25]

Et

H

Et

47a

PCL 87 % ee, 47 % yield

47b [25]

Et

F 89 % ee, -, CRL 90 % ee, -, lipase MY 96 % ee, 54 % yield, lipase OF-360 91 % ee, -, CCL

48a [26]

1

F

481,[2G]

87 % ee, 77 % ee, 84 % ee, 59 % yield 80 % ee, -

a Theotherproduct (alcoholor ester) wasnot isolated. b Acetate was hydrolyzed.

1 S. Iruchijima, A. Keiyu, N. Kojima, Agric. Bid. Chem. 1982,46,1593. 2 F. Francalanci, P. Cesti, W. Cabri, D. Bianchi, T. Martinengo, M. Foi,J. Org. Chem. 1987,52,5079. 3 D. Bianchi, W. Cabri, P. Cesti, F. Francalanci, F. Rama, Tetrahedron Lett. 1988, 29, 2455. 4 W. E. Ladner, G. M. Whitesides, J. Am. Chem. SOC. 1984,106,7250. 5 F. Van Middlesworth, D. V. Patel, J. Donaubauer, P. Gannett, C. J. Sih, J. Am. Chem. Soc. 1985, 107, 2996. 6 J. Van der Eycken, M. Vandewalle, G. Heinemann, K. Laumen, M. P. Schneider, J. Kredel, J. Sauer, J. Chem. Soc., Chem. Commun.1989, 306. 7 S. Hamaguchi, H. Yamamura, J. Hasegawa, K. Watanabe, Agric. Biol. Chem. 1985,49, 1509. 8 S. Hamaguchi, M. Asada, J. Hasegawa, K. Watanabe, Agric. Biol. Chem. 1985,49, 1661. 9 S. Hamaguchi, M. Asada, J. Hasegawa, K. Watanabe, Agric. Biol. Chem. 1984, 48, 2331. 10 B. Wirz, W. Walter, Tetrahedron:Asymmetry1992, 3, 1049. 11 H:J. Gais, 1. von der Weiden, Tetrahedron: Asymmetry1996,7, 1253. 12 M. De Amici, C. De Micheli, G. Carrea, S. Riva, Tetrahedron: Asymmetry1996, 7, 787. 13 H. Tanimoto, T. Oritani, Tetrahedron: Asymmetry 1996, 7, 1695.

14 H. Nakano, K. Iwasa, Y. Okuyama, H. Hongo, Tetrahedron: Asymmetry1996,7, 2381. 15 0. Goj, A. Burchardt, G. Haufe, Tetrahedron: Asymmetry1997,8,399. 16 S. Brand, M. F. Jones, C. M. Raper, Tetrahedron Lett. 1997, 38, 3595. 17 G. D. Gamalevich, B. N. Morozov, A. L. Vlasyuk, E. P. Serebryakov, Tetrahedron 1999,55, 3665. 18 B. Schnell, U. T. Strauss, P. Verdino, K. Faber, 0.C. Kappe, Tetrahedron:Asymmetry2000, 11, 1449. 19 S. V. Ley, S. Mio, B. Meseguer, Synlett 1996,787. 20 H. Hongo, K. Iwasa, C. Kabuto, H. Matsuzaki, H. Nakano,j. Chem. Soc., Perkin Trans. 1, 1997, 1747. 21 S. Erbeck, X. Liang, R. Krieger, H. Prinzbach, Eur. J. Org. Chem. 1998, 481. 22 M. Bakke, M. Takizawa, T. Sugai, H. Ohta, J. Org. Chem. 1998,63,6929. 23 H.-J. Ha, K:N. Yoon, S:Y. Lee, Y.3. Park, M.-S, Lim, Y.G. Yim,J. Org. Chem. 1998,153,8062. 24 J. Pietruszka, T. Wilhelm, A. Witt, Synlett, 1999, 1981. 25 H. Wakamatsu, Y. Terao, Chem. Pharm. Bull. 1996, 44,261. 26 V. Khlebnikov, K. Mori, K. Terashima, Y. Tanaka, M. Sato, Chem. Pharm. Bull. 1995,43, 1659.

442

I

1 I Hydrolysis and Formation of C-0 Bonds

uncatalyzed ester hydrolysis, steric hindrance, at least for the known examples 14-17 and 26-30 in the enzyme-catalyzed hydrolysis, poses no problem. In substrates containing two alkoxycarbonyl groups, one attached to a secondary carbon and the other one to a tertiary carbon, the former is hydrolyzed more readily, as shown for 3-5. Esters with the alkoxycarbonyl group attached to quaternary carbon are readily hydrolyzed, as demonstrated for 17,26-30,47,49,51 (Table 11.1-13). Group selectivity is also observed between an ester group and a thioester group or an ester and a lactone moiety, as exemplified by 12 and 51, respectively. Acyclic as well as cyclic carboxylic acid esters are substrates for enantiomer-selectivehydrolysis catalyzed by lipases. High enantioselectivitiesare observed not only for those esters having a chiral center in a-position but also for those having the chiral center in pposition. A spectacular example in this regard is the acetoxy-substituted carboxylic acid 33,where the chiral center is separated by eight methylene groups from the carboxyl group. This acid is obtained by a Candida cylindracea lipase-catalyzed hydrolysis of the corresponding racemic butyl ester with very high enantioselectivity. Surprisingly, the hydrolysis of the corresponding methyl ester proceeds with a much lower enantioselectivity. Lipase-catalyzed enantiomer-differentiating hydrolysis has been utilized with much success for the synthesis of a-hydroxy and a-acetoxy carboxylic acids (6,7, 24 and 25).A series of vinylogous a-hydroxy carboxylic acids 34-39 is also accessible. The two a-amino acids 48 and 49 with unprotected amino groups are hydrolyzed with high enantioselectivity. The series of methyl-substituted seven-membered lactones 52-55 (Table 11.1-13) are converted in the presence of Candida antarctica lipase yielding the slow-reactinglactones with ee values between 72 and 94%. Acids, monoesters and lactones ofTable 11.1-13which can be obtained with other hydrolases as such or of opposite configuration are contained in Table 11.1-5. Lipase-catalyzed enantiomer-differentiating hydrolysis of acylated racemic primary alcohols covers a broad range of substrates (1-48) summarized in Table 11.1-14,including epoxy alcohols (3-lo),amino alcohols (2,17,36,37)and acylated y-hydroxymethyly-lactones (38-40).By means of incorporating the amino and the secondary hydroxyl group into a heterocyclic ring system, selectively protected amino diols are accessible by Pseudomonas aeruginosa lipase-catalyzed hydrolysis (13-1G).3-Hydroxymethyl-D2-isoxazoline butyrates 19-22 (Table 11.1-14) have been resolved with high selectivity in the presence of Pseudomonas cepacia lipase. Monoacetates and alcohols of Table 11.1-14 which can be obtained with other hydrolases as such or of opposite configuration are contained in Table 11.1-19. Given the experimental simplicity and the potential scale of reaction, lipasecatalyzed enantiomer-differentiating hydrolysis of racemic acylated secondary alcohols is today one of the best methods for the synthesis of optically active secondary alcohols. From the list of the tabulated examples 1-170 (Table 11.1-15)one gets the impression that there is almost no restriction in regard to the substrate structure. Because of the number of lipases available either as isolated enzymes or contained in the various organisms, it seems possible to find the right lipase for almost every substrate. Highly enantiomer-selectivehydrolysis and alcoholysis of esters of a wide structural range of secondary alcohols by the different lipases are possible. Not only

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.1-15. Lipase-catalyzedenantiomer-differentiating hydrolysis of esters of racemic acyclic secondary alcohols in aqueous solution (CCL Candida cylindracea lipase, PSL Pseudomonas sp. lipase, PFL Pseudomonasfluorescens lipase, PAL Pseudomonas aeruginosa lipase, ASL Alcaligenes sp. lipase, ANL Aspergillus niger lipase, PCL Pseudomonas cepacia lipase, ROL Rhizopus oryzae lipase, M M L Mucormiehei lipase, CAL-B Candida antarctica B lipase, LIP Pseudomonas sp. lipase Toyobo, HSL Hurnicola sp. lipase).

OH

R'

RIAMe

la 2a 3a 4a 5a Ga 7a 8a

R' Et

90 % ee, 39 % yield 299 % ee, 48 % yield 97 % ee, 47 % yield 80 % ee, 46 % yield 95 % ee, 47 % yield 95 % ee, 46 % yield 299 % ee, 43 % yield 95 % ee, 50 %yield

m = 0, n = 1 m = 0, n = 2

m = 1,n = 1 m = 1, n = 2

99 % ee, 37 %yield, PFL 46 % conversion 99 % ee, 35 %yield, PFL 46 % conversion 95 % ee, 36 %yield, PFL 48 % conversion 99 % ee, 35 %yield, PFL 48 % conversion

n=5

n = 10

7 ,

93 % ee, -, PSL

RZ

lipase

n-Pr Me Me Me Me Me Me CHzCl

CCI PSL PSL PSL PSL PSL PSL PSL

9a

88 % ee, 40 % yield 299 % ee, 48 % yield 99 % ee, 45 %yield 80 % ee, 47 %yield 97 % ee, 48 %yield 89 % ee, 47 %yield 299 % ee, 46 %yield 9G % ee, 44 %yield

95 % ee, 40 % yield 95 % ee, 40 % yield

9b I31 lob [3]

95 % ee, 40 % yield

l l b [3]

95 % ee, 40 % yield

12b [3]

96 % ee, -

13b [4]

10a lla 12a

13a [4a] 14 [4] 15 14) 16 [4]

299 % ee, -, PSL 299 % ee, -, PSL 299 % ee, -, PSL 299 % ee, -, PSL

n=O n=l

Me0,C

Ph 4-Me-C& 4-MeO-CsH4 PhCH2 4-Pyridyl 2-Naphthyl Ph

x,"'

-

-

OCOCH,CI

OH

Me

17a [5]

MeozC&Me OW0

299 % ee, -

1% [51

444

I

1 1 Hydrolysis and Formation ofC-0 Bonds Table 11.1-15.

R

(cont.).

jl"

OAc

CF,

R

Ph CHzPh (CH2)zPh 2-styryl CHzCO2Et CHZCOZHex

R

c1 Br

R-CF,

57 % ee, -, CCL 94% ee, -, CCL 98 % ee, -, CCL 93 % ee, -, CCL 96 % ee, -, CCL 90% ee, -, CCL 28-50 % conversion

18a [6] 19a [6] 20a[6] 21a [6] 22a[6] 23a [6]

-,-

100 % ee, 24 % yield, PSL 94% ee, 24 %yield, PSL 50 % conversion 92 % ee, -, PSL 98% ee, -, PSL 97% ee, -, PSL 42-53 % conversion

24a 25a

100 % ee, 29 %yield 100 % ee, 11 %yield

24b [7] 25b [7]

26a 27a 28a

299 % ee, 299 % ee, 98% ee, -

26b [8] 27b [8] 28b [8]

98% ee, -

19b [6]

-, -,-,-, -

- -

OCOR'

R ' A O T s R'

Me Et CHzCl

299 % ee, 40 % yield, PAL 299 % ee, 46 % yield, PAL 299 % ee, 46 % yield, PAL

OH

32a [ll]

RZ

Me, Pr299 % ee, 35 % yield 29b [9] Me, Pr299 % ee, 44% yield 30b [9] Me, Pr299 % ee, 45 %yield 31b [lo]

29a 30a 31a

OAc

Ph*CN

PhACN

- -

298% ee, 42 %yield, PSL

oH

33a [ll,121

~o'o""N /

98% ee, -

/

98 % ee, 40 %yield, ASL (PH 4-5) 87 % ee, -, PSL

32b [Ill

33b [ll,121

11.1 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.1-15.

(cont.).

34a [13]

-

34b (131

87 % ee, 39 %yield, PSL

&

OCH,SMe

nCe.H, 7

R L C N

C0,U

CI

R

Me Ph

298 % ee, -, PFL 298 % ee, -, PFL 60-64 % conversion Ph(CH2)z 298 % ee, -, PFL 2-styryl 298 % ee, -, PFL

35 [14] 36 (141

94 % ee, 31 %yield, PFL

39 [15]

37 [14] 38 [14]

OAc R LC02Me

R -CO,Me

R = Me 295 % ee, 37 %yield, PFL 40a [16] R = Et 295 % ee, 44 %yield, PFL 41a [ l G ]

91 % ee, 39 %yield 295 % ee, 45 %yield

OH ThexylMe,SiO-

.

’

C0,Me

72 % ee, 57 % yield, PFL

97 % ee, 41 %yield, PFL

ThexylMe,SiO &CO,Me

42a (161 295 % ee, 35 % yield

(yJ

C0,Me 43a [17]

40b [16] 41b [16]

421, [lG]

C0,Me

96 % ee, 26 %yield

43b [17]

COnPr

OH R&CO,Me

R

C0,Me

R

Et 74 % ee, -, CCL ClIH23 84 % ee, -, CCL (CH2)4CH(CdHs)z 92 % ee, -, CCL 40 % conversion

44a 45a 46a

42 % ee, 75 % ee, 50 % ee, GO % conversion

44b I181 451, [18] 46b [l8]

I

446

I

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-15.

(cont.).

OH

OAc

C0,Me

.I&,,

47a 48a 49a 50a 51a 52a

R’ W O z M e R R2

66 % ee, 56 % yield, ANL 67 % ee, 64 %yield, ANL 64 % ee, 51 % yield, ANL 75 % ee, 51 % yield, ANL 85 % ee, 53 %yield, ANL 79 % ee, 51 % yield, ANL

-

R’

R2

R’

2-fury1 2-thiophenyl 2-(2-butenyl) 2-fury1 2-thiophenyl 2-(2-butenyl)

H H H Me Me Me

Me Me Me H H H

R’

R2

R3

lipase

t-Bu

H H Et H H

H H H Et n-Pr

CCL PFL PFL PFL PFL

Ph Et 5Ga [21] 298 % ee, - Et 57a[21] 298%ee,- n-Pr 30-40 % conversion

299%ee, 33%yield 91 % ee, 35 %yield 299 % ee, 38 % yield 98%ee, 32%yield 299 % ee, 47 %yield 299 % ee, 44 % yield

47b(19] 48b [19] 49b [19] 5Ob[19] 51b [19] 52b [19]

298 % ee, -, ANL 298 % ee, -, ANL 298 % ee, -, ANL

53b (201 54b [20] 55b (201

52-60 % conversion

other product not isolated

R = Et R = n-Pr R = (CHZ)zCO*Et

100 % ee, 50 % yield, PPL 60 % ee, 50 %yield, PPL 56 % ee, 22 %yield, PPL

h L O N E t ,

Gla [22]

58a (211 59a (211 GOa (211

WF CONEt,

61b [22]

OAc

OH

96 % ee, -

58 % ee, -, CCL

GZa (231

Cbz

>97 % ee, 58 %yield, CCL

G2b [23]

Cbz

297 % ee, 42 % yield

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esten Table 11.1-15.

O

(cont.).

OH

h

63a [24]

C

q ’’ O

I

OH C, 3H27AMe

C13H27

52 % ee, 40 % yield, PCL 95 % ee, 23 % yield, PCL

4 7

63b [24]

OAc

L

C

l

88 % ee, 43 % yield

297 % ee, 41 % yield, PCL

x:,“ 54 % ee, 41 % yield 35 % ee, 42 %yield

R = Ac R = OCHzCCl3

64a 65a

64b 1251 65b 1251

-, 48 % yield

295 % ee, 27 % yield, PCL

(hydrolysiswith PFL gives the (S)alcohol with 295 % ee) 674271

, b OAc

\ G O C 6 H 4 0 M e

298 % ee, -, PCL 298 % ee, -, PCL 88 % ee, -, PCL 298 % ee, -, PCL 296 % ee, -, PCL 296 % ee, -, PCL 296 % ee, -, PCL 296 % ee, -, PCL 296 % ee, -, PCL

299 % ee, 42 % yield

R’

R2

R’

H Me H Me H Me n-Pi n-Pent n-Non

0-t-Bu 0-t-Bu SPh SPh

CHzCl CH2Cl Me Me CHzCl CHzCl CHzCl CHzCl CHzCl

SPh S-t-Bu S-t-Bu

S-t-Bu S-t-Bu

OH

68a 69a 70a 71a 72a 73a 74a 75a 76a

298 % ee, 298 % ee, 90 % ee, -, 91 % ee, 296 % ee, 296 % ee, 296 % ee, 296 % ee, 296 % ee, -

68b [28,29] 69b [28,29] 70b [28, 291 71b [28, 291 72b [28,29] 73b [28,29] 741, [28,29] 75b [28,29] 76b [28,29]

OCOC15H3, R o,,!,T~,o

T s O h O R

99 % ee, 44 %yield, PCL 99 % ee, 47 %yield, PCL 99 % ee, 45 % yield, PCL

67b [27]

OC,H,OMe

98 % ee, 53 %yield, PCL (buffer : acetone = 9 : 1)

R

C16H31 CloHzl C4H9

77a 78a 79a

I

295 % ee, 46 %yield 295 % ee, 42 % yield 295 % ee, 43 %yield

771, [30] 78b [30] 79b [30]

448

I

7 7 Hydrolysis and Formation ofC-0 Bonds

Table 11.1-15.

(cont.)

OAc

80 [31]

96 % ee, 27 % yield, PPL

other product not isolated

boxcl 81b (321

w (R = H, Me, OMe, NO*, ally], 0-allyl, c-C5HI1 39-99 % ee, 31-54 %yield, PSL

67-99 % ee, 31-52 %yield

OH

82a [33]

82b [33]

Ph-CI

Ph

97 % ee, 50 % yield, PSL

299 % ee, 50 % yield

83a [34]

"

&Me

(2-4) 295 % ee, 37-55 %yield, PSL

295 % ee, 37-45 % yield

84a 1351

OCOCH,CI

84b [35]

100 % ee, 48 % yield

89 % ee, 53 %yield, PSL

OAc

299 % ee, 31 %yield, ROL 299 % ee, 37 % yield, ROL 50 % ee, 21 %yield, ROL

R'

R2

Ph Ph Me

Me i-Pr Me

85a 86a 87a

90 % ee, 35 %yield 86 % ee, 46 % yield 9 % ee, 29 %yield

85b [36] 86b [36] 87b (361

299 % ee, 43 %yield 299 % ee, 43 % yield

88b (371 89b (371

nPrOCO

&o

R R

299 % ee, -, CCL 299 % ee, -, CCL

i-Pr Ph

88a [37] 89a[37]

7 1.1 Hydrolysis and Formation of Carboxylid Acid Esten Table 11.1-15.

tBu

(cont.).

OH &CI

9Oa [38) 298 % ee, 42 % yield

91 % ee, 38 %yield, PSL

P

OH O A

h

90b [38]

tBu

P

R

h

O

z

R

R

92 % ee, 31 %yield, PSL 86 % ee, 46 %yield, PSL 91 % ee, 48 %yield, PSL

C1 Br N3

97 % ee, 41 % yield 97 % ee, 41 % yield 84 % ee, 48 % yleld

91a 92a 93a

91b [39] 92b [39] 93b [39]

OCOEt

0

94a [40]

OBn 32 % ee, 66 %yield, CCL (isolated as alcohol obtained with NaBH4)

94b [40]

OBn 299 % ee, 22 % yield

OCOR2 R

295 % ee, -, PSL 295 % ee, -, PSL 295 % ee, -, PSL 295 % ee, -, PSL 295 % ee, -, PSL

R'

R2

Me Et CH2CI CH=CH2 CH20CH=CH2

n-Pr n-Pr n-Pr ClCH2 ClCHz

lOOa [41] Me 295 % ee, -, PSL

295 % ee, 295 % ee, 295 % ee, 91 % ee, 51 % ee, -

95a 9Ga 97a 98a 99a

OCH,CI

95b [41] 9Gb [41] 97b [41] 98b [41] 99b [41]

lOOb [41]

Me-OTBDMS 295 % ee, -

lOla [42] P h C 0 2 q g"'OH -,-,MML

,&OTBDMS

/',n..,(yOCOEt PhCO, S 76 % ee, 14 % yield

l0lb[421

I

449

450

I

7 7 Hydrolysis and Formation of C - 0 Bonds

Table 11.1-15.

(cont.).

0

0 Me\o+s\R

could not be isolated

be\O%\R]

OAc R

-CH2CH(OMe)2 -CHzCH(OEt)z -CH2CH(OBn)2 -(CH2)2CH(OMe)2 n-Bu -(CH2)20SiEt3 -(CH2)30SiEt3

>95 % ee, 44 % yield, PCL

>95 % ee, 42 %yield, PCL

R

Ph Ph Bn Bn -(CH2)2Ph

102 [43] 103 1431 104 [43] 105 [43] 106 [43] 107 [43] 108 1431

>95 % ee, 45 %yield, PFL >95 % ee, 49 %yield, PFL 65 % ee, 48 %yield, PFL >95 % ee, 45 %yield, PFL >95 % ee, 47 %yield, PFL 81 % ee, 47 %yield, PFL 85 % ee, 48 %yield, PFL

llOa

98 % ee, -, PCL 99 % ee, -, CAL-B 95 % ee, -, PCL 97 % ee, -, CAL-B 98 % ee, -, CAL-B

99 % ee, -, PCL 99 % ee, -, CAL-B 97 % ee, -, PCL 97 % ee, -, CAL-B 97 % ee, -, CAL-B

llla 112a

1lOb [45]

lllb (451 112b [45]

OR NPht ; H I OHH NPht

113a [4G] Et

R = CO(CH,),Me

>99 % ee, 49 % yield, ASL

R

O

OH A C

I

R

2-Naphthyl 95 % ee, -, CAL-B 4-AcNH-GH4 95 % ee, -, CAL-B 4-[MeO-(CHZ)2]-C6H495 % ee, -, CAL-B C6H4

>99 % ee, 50 % yield

R

114a 115a 116a

O

L

C

I

95 % ee, 47 %yield 79 % ee, 40 %yield 70 % ee, 37 %yield

114b 1471 115b [47] llGb (471

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters

I

451

Table 11.1-15.

F

(cont.).

/~

C

N 117a [481

‘WcN F

117b [48]

F

F 93 % ee, 40 %yield, PFL >98 % ee, 41 % yield, LIP

82 % ee, 37 %yield, PFL >99 % ee, 521 % yield, LIP

OH RL

R

R

89 % ee, -, PCL 93 % ee, -, PCL >95 % ee, -, PCL

Ph 4-F-C& 4-t-BU-C&

C

l

80 % ee, -, 78 % ee, -, >95 % ee, -,

118a 119a 120a

MeoL 121a [50]

\

.

C02Me

118b [49] 119b [49] 120b [49]

121b [SO]

C0,Me

OH

OAc

94 % ee, 51 %yield, ANL

>99 % ee, 48 % yield

F*N

F

R

/

F

F

94 % ee, 39 %yield, LIP 97 % ee, 38 % yield, LIP 89 % ee, 32 % yield, LIP

Me Et Ph

OAc

122a 122a 122a

R‘

not separated

R’

122b [51] 123b [51] 124b [51]

Aco*J5

OH

“R2O r ’

Me Et i-Pr Me

85 % ee, 39 %yield 92 % ee, 46 %yield 96 % ee, 43 %yield

R2

H H H Me

96%ee,CAL-B >98%ee,CAL-B >98%ee,CAL-B >98%ee,CAL-B

97 % ee 93% ee >98 % ee >96 % ee

125a,b 12Ga,b 127a,b 128a,b

>98 % ee, 42 % yield >98 % ee, 37 %yield 63 % ee, 39 %yield 96 % ee, 38 % yield

125c [52] 12Gc [52] 127c 1521 128c [52]

7 7 Hydrolysis and Formation of C-0 Bonds

(cont.).

Table 11.1-15.

OH 129a [531

o
129b [53]

\

O r M e

93 % ee, -

85 % ee, -, CAL-B 52 % conversion

OAc

Me

Me 96 % ee, -, CAL-B 49 % conversion

92 % ee, -

O -R

-OR OCOCH,CI

OH R SiMezThex

97 % ee, -, PCL 94 % ee, -, CAL-B 94 % ee, -, HSL

Bz

W

O

130b [53]

O r M e

130a [53]

? ‘Me

T

s

133a 1541

99 % ee, 95 % ee, 97 % ee, -

131a 132a

T

O

T

S

131b [54] 1321, [54]

133b [54]

OAc 99 % ee, -, PCL

OH 99 % ee, -, PCL

OAc

““T“

134a (551

134a [55]

/

H2N

HA

CI 96 % ee, 42 % yield

LI

99 % ee, 42 % yield, CAL-B

4

C0,Et

OAc 135a [56]

46% ee, 33 %yield

>99 % ee, 19 % yield, PSL

13Gb [57]

136a (571 J O : T C0,Et B D M S OH >99 % ee, 43 % yield, PCL

135b [56]

OCOCH,CI 90 % ee, 46 % yield

11.1 Hydrolysis and Formation ofcarboxylid Acid Esters

I

453

Table 11.1-15.

(cont.).

80 % ee, 45 % yield

87 % ee, 39 %yield, PPL

138a[58] Ph >96 % ee, 40 %yield, PPL

1- N a p h - O Y O M e

wph I

:

138b [S8]

70 % ee, 48 % yield

13% [58]

1-Naph-O/\/\OMe

OH

13% [59l

OAc

82 % ee, 47 % yield, CAL-B

90 % ee, 38 % yield

0 Me\o+o.R OAc abs. config. unknown R -(CHz)7CH3 -CHZCH=CHz -(CHz)*CH=CH2 -CHZCH=CHPh -CHZPh -(CHZ)C02Et -CHzCH(OEt)z

OH

racemic

>95 % ee, 43 % yield, PFL >95 % ee, 32 %yield, PFL 90 % ee, 37 % yield, PFL >95 % ee, 39 %yield, PFL >95 % ee, 35 %yield, PFL >95 % ee, 27 %yield, PFL >95 % ee, 37 %yield, PFL

140 [GO] 141 [60] 142 [GO] 143 (601 144 [GO] 145 [60] 146 (601

* d R* R2 R'

>99 % ee, 38 %yield, PCL >99 % ee, 37 % yield, CAL >99 % ee, 22 % yield, CRL >99 % ee, 36 %yield, ASL Me Me >99 % ee, 36 % yield, PCL >99 % ee, 23 % yield, CAL Et

R'

R2

Et

146a 147a 148a 149a 150a 151a

94 % ee, 42 % yield 59 % ee, 57 %yield 39 % ee, 48 % yield 81 % ee, 40 % yeld 91 % ee, 54 % yield 73 % ee. 21 %yield

14Gb [61] 147b [61] 148b [61] 149b 1611 150b [61] 151b [61]

454

I

1 1 Hydrolysis and Formation of C - 0 Bonds Table 11.1-15.

(cont.).

R’ = aromatic, heteroaromatic, R2 = Me, Et, LPr, R’ =Me, CH2CI 14-95 % ee, ANL or ROL 9-70 % ee

OH

Rl

Ap/(0R’),

153a[63]

II 0 in some cases R R’ = alkyl, R2 = Me, Et, LPr, R’ 7.5-98 % ee, ANL or ROL

OCOR~

= Me, CHlCl

3-80 % ee

OAc OEt

154a [64] a

O

E

153b (631

=

RlAp/(OR’)2 I1 0 in some cases S

t

1541, [64]

OEt

>98 % ee, 41 %yield, PSL

OAc &CO,Et

I

I

U

95 % ee, 38 %yield, PCL

156a[66] 80 % ee, 44 % yield, PSL

1561, [66]

9AC

87 % ee, 44 % yield OAc

R’+cop2 Cl

R’

R2

Me Et Et n-CsHI7 n-C8H17 n-CgH17

Et Me Et Et Me Et

>99 % ee, 29 %yield, PCL 96 % ee, 29 %yield, PCL 86 % ee, 29 %yield, PCL 98 % ee, 29 % yield, PCL 87 % ee, 35 % yield, PCL 94 % ee, 31 %yield, PCL

157a 158a 159a lGOa 161a 162a

-, 30 %yield

-, 43 %yield -, 22 %yield -, 18 %yield -, 31 %yield -, 22 % yield

15% 1671 158b 1671 159b [67] lGOb [67] 161b 1671 162b [67]

1 7 . 7 Hydrolysis and Formation ofCarboxylid Acid Esters

I

455

Table 11.1-15.

(cont.).

&

4-MeO-C6H,

OH

C0,Et

97 % ee, 42 % yield, PCL

OH P

h

164b [69]

M

C

l

95 % ee, 42 %yield, CAL-B

165a[70]

165b [70]

OH

OCOPh

91 % ee, -, CRL 48 % conversion

97 % ee, -

166a[70]

CL ,

N

OH

166b [70]

OCOPh

83 % ee, -, CRL 49 % conversion

80 % ee, -

167a[70]

167b [70]

OH

OCOPh

84 % ee, -, CRL 43 % conversion

65 % ee, -

168a [71]

168b [71]

Me0

Me0

>99 % ee, 29 %yield, PCL/Celite 98 % ee, 44 % yield, PCL/ ENTP-4000 (prepolymer)

48 % ee, 67 % yield 81 % ee, 52 % yield

456

I

1 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-15.

(cont.).

168a [71]

1681, [71]

OCOCH,CI

Me0

Me0

95 % ee, 49 %yield, PCL/ ENTP-4000 (prepolymer)

94 % ee, 48 % yield

R = H, 2-Me, 3-Me, 4-Me, 3-F, 4-F, 4-CN 80-97 % ee, 30-50 %yield, PSL

80-99 % ee, 30-50 % yield

OCOR

4;

CF,

CN

95 % ee, 23 %yield, CRL 99 % ee, 37 % yield, CRL 97 % ee, 36 % yield, CRL 99 % ee, 30 % yield, CRL 99 % ee, 40 % yield, CRL

R

R’

Me

Me

170a [73]

n-Bu

Me

170b [73]

n-Pent

Me

170c [73]

n-Hept

Me

170d [73]

Me

Et

170e [73]

1 B. Cambou, A. M. Klibanov, Biotechnol. Bioeng. 1984,26,1449. 2 K. Laumen, M. P. Schneider, J. Chem. Soc., Chem. Commun. 1988, 598. 3 D. Bianchi, P. Cesti, P. Golini, Tetrahedron 1989, 45, 869. 4 A. Scilimati, T. K. Ngooi, C. J. Sih, Tetrahedron Lett. 1988,29,4927. 5 T. K. Ngooi, A. Scilimati, Z.-W. Guo, C. J, Sih, /. Org. Chem. 1989, 54, 911. 6 J . T. Lin, T. Yamazaki, T. Kitazumi, /. Org. Chem. 1987,S2, 3211. 7 A. Kutsuki, I. Sawa, J. Hasegawa, K. Watanabe, Agric. Biol. Chem. 1986, SO, 2369. 8 K. Mori, R. Bernotas, Tetrahedron: Asymmetry 1990, I , 87. 9 S. Hamaguchi, T. Ohashi, K. Watanabe, Agric. B i d . Chem. 1986,50,1629.

10 S . Hamaguchi, T. Ohashi, K. Watanabe, Agnc. B i d . Chem. 1986,50,375. 11 A. van Almsieck, J. Buddrus, P. Honicke-Schmidt, K. Laumen, M. P. Schneider, /. Chern. Soc., Chem. Commun. 1989,1391. 12 H. Hirohara, S . Mitsuda, E. Ando, R. Komaki, in: J. Tramper, H. C. van der Plaas, P. Linko (Eds.), Biocatalysts i n Organic Synthesis: Amsterdam, Elsevier, 1985, p 119. 13 N. Matsou, N. Ohno, Tetrahedron Lett. 1985, 26, 5533. 14 T. Itoh, Y. Takagi, S. NishiyamaJ. Org. Chem. 1991,56, 1521. 15 T. Itoh, Y Tagaki, Chem. Lett. 1989, 1505. 16 K. Burgess, I. Henderson, Tetrahedron: Asymmetry 1990, I , 57. 17 H. Suemune, Y. Mizuhara, H. Akita, K. Oishi, Chem. Pharm. Bull. 1987,35,3112.

1 7 . 1 Hydrolysis and formation ofCarboxylid Acid Esters

I

457

18 C. Feichter, K. Faber, H. Griengl, Tetrahedron Lett. 1989, 30, 551. 19 H. Akita, H. Matsukara, T. Oishi, Tetrahedron Lett. 1986,27,5241. 20 E. Foelsche, A. Hickel, H. Honig, P. Seufer-Wasserthal.J. Org. Chem. 1990, 55, 1749. 21 B. A. Marples, M. Rogers-Evans, Tetrahedron Lett. 1989, 30, 261. 22 T. Tsukamoto, T. Yoshiyama, T. Kitazume, Tetrahedron: Asymmetry 1991.2, 759. 23 T. R. Nieduzak, A. L. Margolin, Tetrahedron: Asymmetry 1991,2,113. 24 H. S. Bevinakatti, A. A. Baneji, ]. Org. Chem. 1991,56,5372. 25 Y. Naoshima, Y. Munakata, S. Yoshida, A. Funai, J. Chem. Soc., Perkin Trans. 11991, 549. 26 T. Yamazaki, N. Okamura, T. Kitazume, Tetrahedron: Asymmetry 1990, I , 521. 27 S. Takano, M. Setoh, K. Ogasawara, Tetrahedron: Asymmetry 1993,4,157. 28 U. Goergens, M. P. Schneider,]. Chem. Soc., Chem. Commun. 1991,1064. 29 U. Goergens, M. P. Schneider, Tetrahedron: Asymmetry 1992,3,1149. 30 R. ChCnevert, R. Gagnon,J. Org. Chem. 1993.58, 1054. 31 M. Treilhou, A. Fauve, J.-R. Pougny, 1.-C. Prom&, H. Veschambre,]. Org. Chem. 1992,57, 3203. 32 U. Ader, M. P. Schneider, Tetrahedron: Asymmetry 1992, 3, 521. 33 M. P. Schneider, U. Goergens, Tetrahedron: Asymmetry 1992,3,525. 34 R. Seemayer, M. P. Schneider, Tetrahedron: Asymmetry 1992,3,827. 35 S. Liang, L. A. Paquette, Tetrahedron: Asymmetry 1990, I, 445. 36 Y:F. Li, F. Hammerschmidt, Tetrahedron: Asymmetry 1993,4,109. 37 M. Banziger, J. F. Mc Gamty, Th. Meul, J. Org. Chem. 1993,58,4010. 38 A. Chadha, U. Goergens, M. P. Schneider, Tetrahedron: Asymmetry 1993,4, 1449. 39 U. Ader, M. P. Schneider, Tetrahedron: Asymmetry 1992, 3, 205. 40 K. Matsumoto, N. Suzuki, H. Ohta, Tetrahedron Lett. 1990, 31,7163. 41 U.Goergens, M. P. Schneider,]. Chem. Soc., Chem. Commun. 1991,1066. 42 R. P. C. Cousins, M. Mahmoudian, P. M. Youds, Tetrahedron: Asymmetry 1995, 6, 393. 43 J. Milton, S. Brand, M. F. Jones, C. M. Rayner, Tetrahedron: Asymmetry 1995, 6, 1903. 44 B. Charpentier, 1.-M. Bemardon, P. Diaz, M. Vion, C. Millois, B. Bernard, B. Shroot, Bioorg. Med. Chem. Lett. 1995,5,2801. 45 B. H. Hoff, V. Waagen, T. Anthonsen, Tetrahedron: Asymmetry 1996,7,3181. 46 M. Seki, T. Furutani, T. Miyake, T. Yamanaka, H. Ohmizu, Tetrahedron: Asymmetry 1996, 7 , 1241.

47

I. L. Bermudez, C. del Campo, L. Salazar, E. F.

Llama, J. V. Sinisterra, Tetrahedron: Asymmetry, 1996, 7, 2485. 48 T. Sakai, T. Takayama, T. Ohkawa, 0. Yoshio, T. Ema, M. Utaka, Tetrahedron Lett. 1997, 38, 1987. 49 D. Bianchi, P. Moraschini, A. Bosetti, P. Cesti, Tetrahedron Lett. 1994,5,1917. 50 H. Akita, C. Y. Chen, S. Nagumo, Tetrahedron: Asymmetry 1994,51207. 51 T. Sakai, Y. Miki, M. Nakatani, T. Ema, K. Uneyama, M. Utaka, Tetrahedron Lett. 1998,39, 5233. 52 W. Adam, M. T. Diaz, C. R. Saha-Moller, Tetrahedron: Asymmetry 1998, 9, 589. 53 W. Adam, M. T. Diaz, C. R. Saha-Moller, Tetrahedron: Asymmetry 1998, 9, 791. 54 T. Ziegler, F. Bien, C. jurisch, Tetrahedron: Asymmetry 1998,9,765. 55 S. Conde, M. Fierros, M. I. Rodriguez-Franco,C. Puig, Tetrahedron: Asymmetry 1998,9, 2229. 56 N. Hayashi, K. Yanagihara, S, Tsuboi, Tetrahedron: Asymmetry 1998,9,3825. 57 T.Akeboshi, Y. Ohtsuka, T. Sugai, H. Ohta, Tetrahedron 1998,54, 7387. : . Yu, 0. Me&-Cohn, Tetrahedron Lett. 1999,40, 58 CY 6665. 59 L. Salazar, J. L. Bermudez, C. Ramirez, E. F. Llama, J. V. Sinisterra, Tetrahedron: Asymmetry, 1999, 10,3507. GO S . j. Fletcher, C. M. Rayner, Tetrahedron Lett. 1999, 40,7139. 61 T. Itoh, K. Kudo, N. Tanaka, K. Sakabe, Y. Takagi, H. Kihara, Tetrahedron Lett. 2000,41,4591. 62 G.Eidenhammer, F. Hammerschmidt, Synthesis, 1996,748. 63 M. Drescher, F. Hammerschmidt, H. Kahlig, Synthesis, 1995, 1267. 64 M:J. Kim, I.T. Lim, Synlett, 1996, 138. 65 M. Kamezawa, M. Kitamura, H. Nagaoka, H. Tachibana, T. Ohtani, Y.Naoshima, Liebigs. Ann. Chem. 1996,167. 66 K. Mori, H. Ogita, Liebigs Ann. Chem. 1994, 1065. 67 S. Tsuboi, J. Sakamoto, H. Yamashita, T. Sakai, M. Utaka,]. Org. Chem. 1998,63,1102. 68 S. B. Desai, N. P. Argade, K. N. Ganesh,]. Org. Chem. 1996,61,6730. 69 H:L- Liu, B. H. Hoff, T. Anthonsen,J. Chem. Soc., Perkin Trans. 1 2000, 1767. 70 F. Bellezza, A. Cipiciani, G.Cruciani, F. Fringuelli,J. Chem. Soc., Perkin Trans. 1 2000, 4439. 71 H. Akita, I. Umezawa, H. Matsukura, Chem. Pharm. Bull. 1997,45, 272. 72 C. Waldinger, M. Schneider, M. Botta, F. Corelli, V. Summa, Tetrahedron: Asymmetry 1996, 7, 1485. 73 K. Konigsberger, K. Prasad, 0. Repic, Tetrahedron: Asymmetry 1999, 10,679.

458

I

11 Hydrolysis and Formation of C-0 Bonds

secondary alcohols of the aryl alkyl or dialkyl type are accessible but also those containing all kinds of functional groups in the various positions. An inspection of Tables 11.1-15 and 11.1-13 reveals that in cases where an alkoxycarbonyl group is present as well as the secondary hydroxyl group, two possibilities for enantiomerdifferentiation may exist, hydrolysis of the acylated alcohol or hydrolysis of the carboxylic acid ester. Changing the acyl group from acetate to butyrate, chloroacetate, ethylthioacetate or hexadecanoate may have a beneficial effect on the enantioselectivity of the hydrolysis. The use of chloroacetates in many cases facilitates the separation of the ester and the alcohol formed. A series of cyanohydrin acetates have been prepared. Isolation of the cyanohydrin itself is usually not possible because of the alkaline pH. With Alcaligenes sp. lipase, which has its pH optimum between 4 and 5, isolation of the cyanohydrin acetate 331, as well as the cyanohydrin 33a becomes possible. Enantiomer separation of a-benzyloxy ketones can be accomplished via lipasecatalyzed enantiomer-differentiating hydrolysis of the corresponding enol esters with formation of a mixture of the resulting ketone and the unchanged enol ester (94a,b). a-Acetoxysulfides (102-108), a-acetoxyethers (140-146) and a-acetoxyphosphonates (152-153) (Table 11.1-15)are useful substrates for lipases too. Acylated alcohols and alcohols of Table 11.1-15 which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-20 and 11.1-22. A broad structural range of racemic secondary mono-, bi- and tricyclic acylated alcohols are substrates in lipase-catalyzed enantiomer-differentiating hydrolysis as the examples 1-90 of Table 11.1-16 reveal. A large number of cis- and transcycloalkanols bearing a functional group in 2-position (1-20, 25, 26, 58-62) is thereby available in enantiomericallypure form. Enantiomer selectivity in the case of cyclic allylic alcohols where the double bond bears no other substituent in the aposition is frequently low. Through a temporary substrate modification such as mono- or dibromination, enantiomerically pure cyclic allylic alcohols may also be obtained in these cases (51, 52). Prochiral diketones or racemic ketones, like enol esters, are also amenable to a hydrolase-catalyzed asymmetric transformation. The enol acetates and ketones 63 and 64, respectively, may be obtained by Pseudomonas cepacia lipase-catalyzed and Candida qlindracea lipase-catalyzed hydrolysis of the corresponding racemic enol esters or prochiral bis enol ester, respectively, with high enantioselectivity and yield. A variety of allylic monocyclic alcohols (50, 54, 56, 57, 68-70,77-79 and 81) (Table 11.1-16)have been obtained mainly by Pseudomonas cepacia lipase-catalyzedhydrolysis. The planar chiral [2,2]paracyclophane87 was readily resolved by two different lipases, yielding both enantiomers in almost enantiomerically pure form. The Candida cylindracea lipase-catalyzed, Candida rugosa lipase-catalyzed and cholesterol esterase-catalyzed hydrolyses of acetates 88b-1021, are examples of the utilization of a remote phenolic ester group as the site of enzymatic attack. For such cases, cholesterol esterase seems to be particularly well suited. Acylated alcohols and alcohols of Table 11.1-16which can be obtained with other

7 1 . I Hydrolysis and Formation ofCarboxylid Acid Esters

Table 11.1-16. Lipase-catalyzed enantiomer-differentiatinghydrolysis of esters of racemic cyclic secondary and tertiary alcohols in aqueous solution (PFL PseudomonasPuorescenslipase, PSL Pseudomonas sp. lipase, CCL Candida cylindracea lipase, ABL Arthrobacter sp. lipase, PCL Pseudomonas cepacia lipase, CRL Candida rugosa lipase, CE cholesterol esterase).

e;

,,st

R' OAc

COzEt N3

OCOR2 R' R2

Me Me n-Pr

lipase

299 % ee, 30 %yield 299 % ee, 42 % yield 92%ee,44%yield

PFL PFL PFL

la

2a 3a

0OH

30 % ee, 65 %yield 90 % ee, 50 % yield 298%ee,-

l b [l]

2b [l]

3b PI

OAc

0

1

C0,Et

>99 % ee, 43 %yield, PFL

95 % ee, 42 % yield

OCOR2

5a Ga

7a 8a 9a 10a lla 1h 13a 14a 15a

299 % ee, 33 % yield 84 % ee, 48 % yield 299 % ee, 41 % yield 96 % ee, 40 % yield 298 % ee, 40 % yield 93 % ee, 40 % yield 298 % ee, 38 % yield 95 % ee, 44 % yield 295 % ee, 47 % yield 98 % ee, 45 % yield 299 % ee, 42 % yield

R' OAc OAc

C02Et N3

NO2 CN CN Ph PhCH2 OMe OPh

R2

lipase

Me Me Me n-Pr n-Pr n-Pr n-Pr CHzCl Me Me Me

PFL PSL PFL CCL CCL CCL PSL PSL PSL PSL PSL

5c02Et 0

48% ee, 51 %yield 94 % ee, 41 %yield 55 % ee, 59 %yield 298%ee,85%ee,-

93%ee,95%ee,97 % ee, 43 %yield 295 % ee, 45 %yield 96 % ee, 49 % yield 96 % ee, 45 % yield

51, [31

~b [41 7b [31 8b PI 9b 121 10b [2] l l b [2] 12b 141

13b [4] 14b [4] 15b [4]

OAc

1Ga [3]

** *

>99 % ee, 32 %yield, PFL

co*Et

70 % ee, 63 % yield

16b [3]

I

459

460

I

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-16.

(cont.).

W R'

17a 18a 19a

299 % ee, 45 % yield 299 % ee, 38 % yield 89 % ee, 40 %yield

OAc COzEt N,

R2

W lipase

Me Me n-Pr

PFL 55% ee, 55 %yield PSL 68 % ee, 58 %yield CCL 91 %ee,-

17b [3] 18b [3] 19b [3]

OAC

45 % ee, 64 % yield

>99 % ee, 36 % yield, PFL

bR'

acetate was not isolated

R'

HO

99 % ee, -, ABL 98 % ee, -, ABL 79 % ee, -, ABL 30 % ee, -, ABL

R' R'

= CH2C=CH,

R2 = Me

= CH2CH=CH2'R2 = Me

R' = CH~CGCH,R2 = H R' = R2 = H 20-50 % conversion

6

/N3

21 [5-71 22 [5-71 23 [S-71 24 [S-71

OCOnPr

25a [2]

sYN3

25b 121

298 % ee, -

88 % ee, 40 % yield, CCL

OCOnPr

2Ga [2] 298 % ee, 40 % yield, CCL

(YN3

94 % ee, -

261, [2]

7 1.1 Hydrolysis and Formation ofcarboxylid Acid Esten I461 Table 11.1-16.

(cont.).

6iMe2

OAc

27a [8]

95 % ee, 27 % yield, CCL

OiMe2

2%

PI

57 % ee, 50 % yield

HO

AcO

28b [9, 101

28a [9, 101 298 % ee, 46 % yield, PFL

298 % ee, (further hydrolysis of the acetate)

29a [lo]

29b [lo]

81 % ee, -, PFL

H

30 [31

OH

299 % ee, 13 %yield, PFL

03

acetate was not isolated

OH

/

31a [ll]

299 % ee, 46 %yield, PSL

299 % ee, 47 % yield

OH

03 /

299 % ee, 47 % yield, PSL

31b [ll]

03 OAc 1

32a [ll]

299 % ee, 47 % yield

32b [ll]

462

I

11 Hydrolysis and Formation o f C - 0 Bonds Table 11.1-16.

(cont.).

OCOnPr

1

R = H 33a [12] 295 % ee, 40 %yield R = Me 34a [12] 44 % ee, 69 %yield

295 % ee, 43 %yield, CCL 295 % ee, 40 %yield, CCL

33b [I21 34b [12]

Y

OH

ConPr

R R = H 35a [12] 36 % ee, 46 %yield R = Me 3Ga [12] 77 % ee, 50 %yield

31 % ee, 48 %yield, CCL 84 % ee, 37 %yield, CCL

>:5npr

OH

x.3

35b (121 3Gb [12]

37a [12]

371, [12]

295 % ee, 50 % yield

295 % ee, 35 %yield, CCL

OCOnPr

1

38b 1121

295 % ee, 48 % yield

295 % ee, 35 %yield, CCL

OCOR

39a 40a 40a 41a 42a

x-Y CH=CH CH=CH CH=CH CH=CH CH2-CH2

2

R

90%ee,88%ee,97%ee,93%ee,75%ee,-

CH2 CH2 CH2 0 CHI

Me n-Pr n-Pr n-Pr Me

296%ee,89%ee,-, PSL

43a

94%ee,-

CH-CH

CH2

n-Pr

t97%ee,-

297 % ee, 52%ee,-

39b [13] 40b[14] 40b 1141 41b I151 42b [ll,141 43b [ll,141

J 1.1 Hydrolysis and Formation of Carboxylid Acid Esters

Table 11.1-16.

(cont.).

44a

CH2

n-Pr

83%ee,-

44b (171

4Sa

0

n-Pr

85 %ee,-

45b [15]

CH2

n-Pr

14%ee,-

46b [14]

4Ga

22 % ee, all CCL

x

0

0

~~

CH-CH

MeomNMe OH

47a 1161

4% [16]

Me0

Me0 Meo&NMe

93 % ee, 44 % yield, CCL

94 % ee, 40 % yield

48a [17]

Ac2L

48b [17]

96 % ee, 95 % ee, (further hydrolysis of reacylated alcohol), CCL (further hydrolysis of acetate)

49a [18] AcO

49b [18] ACO

81 % ee, 36 %yield, CCL

95 % ee, 46 % yield

299 % ee, 46 % yield, PSL

299 % ee, 43 % yield OCOnPr

Sla [20]

298 % ee, 46 %yield, PCL

oBr

298 % ee, 46 % yield

51b [20]

464

I

I 1 Hydrolysis and Formation ofC-0 Bonds Table 11.1-16.

(cont.). OCOnPr

52a [20]

ogr

298 % ee, 41 %yield, PFL

298 % ee, 45 % yield

295 % ee, 43 %yield, PFL

99 % ee, 49 % yield

52b [20]

54b [22]

AcO 295 % ee, 72 % yield

HO 295 % ee, 34 % yield, PSL

>'r)OB"

55a 1231

55b 1231

BnO 299 % ee, 50 % yield

299 % ee, 48 % yield, PCL

C0,Et

C0,Et

btoH hoAc 99 % ee, 44 % yield, PCL 100 % ee, 43 %yield, PCL

n=1 n=2

56a [24] 57a [24]

HobR 93 % ee, 45 %yield, PSL 90 % ee, 43 % yield, PSL

R Ph

58a [25] PhCHz 59a [25]

100 % ee, 45 %yield 91 % ee, 52 %yield

56b [24] 57b [24]

Aco'GR 298 % ee, 42 %yield 93 % ee, 45 %yield

58b [25] 59b [25]

11.1 Hydrolysis and formation ofCarboxylid Acid Esters Table 11.1-16.

(cont.).

84 % ee, 45 % yield, PSL 80 % ee, 43 %yield, PSL 51 % ee, 38 %yield, PSL

PhO 6Oa [25] PhCH20 61a [25] OAc 62a [25]

298 % ee, 45 %yield 298 % ee, 40 %yield 46 % ee, 45 %yield

0 II

Gob [25] Glb [25] 62b [25]

OAc I

63b [26]

24 % ee, 71 % yield, PCL

Aco&c

299 % ee, 20 % yield

64 [27]

298 % ee, 80 % yield CCL

0gHoH

H ~ 0s m

65a [28]

90 % ee, 34 % yield, PCL

-

O

65b [28]

-

d

94 % ee, 40 % yield

G6a 1291

>99 % ee, 37 % yield, CAL-B (in the presence of PdClZ(MeCN)zand air)

hc

661,[29]

>99 % ee, 42 % yield

.OAc 67a [30]

99 % ee, 35 %yield, PFL

67b [30]

99 % ee, 31 %yield

I

465

466

I

I 7 Hydrolysis and Formation ofC-0 Bonds Table 11.1-16.

(cont.).

AcO 68a [31]

68b [31] NO2

42 % ee, 34 %yield, PCL

88 % ee, 45 % yield

OAc

G9a [31]

G9b 1311 NO2

>99 % ee, 32 %yield, PCL

>99 % ee, 43 % yield

70a (321 TBDMSO

TBDMSO""

92 % ee, 50 % yield, PSL 87 % ee, 40 % yield, PCL

97 % ee, 44 % yield 99 % ee, 39 % yield

Me,N.,?H

70b 1321

?H nPrOCO

71a [33] 37 % ee, -

, %O ee, -, q CRL, o 28 % M conversion e 89

72a [33] +HO M

71b [33]

nPrOCO*OMe

72bI331

Me

>98 % ee, 40 % yield

>98 % ee, 40 % yield, CRL

73a [33]

90 % ee, 45 % yield, CRL

73b[33]

58 % ee, 46 % yield 33 % conversion >99 % ee, 35 % yield 60 % conversion

11.1 Hydrolysis and Formation of Carboxylid Acid Esters

I

467

Table 11.1-16.

(cont.).

ROA

A

c

R

2-Me-naphthyl >99 % ee, 46 % yield, 74a [34] >99 % ee, 46 % yield PCL >99 % ee, 48 % yield, 75a [34] >99 % ee, 49 %yield CHZPh PCL >99 % ee, 48 %yield, 76a [34[ >99 % ee, 48 % yield TBDMS PCL

74b [34] 75b [34) 76b [34]

OAc &SPh

77a [35]

100 % ee, 48 % yield, PCL

WPh

77b [35]

100 % ee, 46 % yield

eph

78a [35]

100 % ee, 45 %yield, PCL

OAc

us'" fj

78b [35]

100 % ee, 48 % yield

Bra..,,

Br

OAc

90 % ee, 47 % yield, PCL >99 % ee, 38 % yield, PPL, after recrystallization

79a [36a] 79a [36b]

80a [37]

97 % ee, 51 %yield, PSL

Q /

OH 94 % ee, 46 %yield, PCL

>98 % ee, 26 % yield >99 % ee, 38 % yield, after recrystallization

0h

79b [36a] 79b [3Gb]

80b [37] A

c

98 % ee, 48 % yield

81a [38]

rn

OAc

>99 % ee, 45 % yield

81b [38]

468

I

7 7 Hydrolysis and Formation of C-0 Bonds Table 11.1-16.

(cont.).

.,,\OH

0

n

AcO

1 >99%ee, 42 %yield, PCL 2 >99% ee, 44 % yield, PCL

82a [39] 83a [39]

>99% ee, 40 % yield >99% ee, 48 % yield

H

82b [39] 83b [39]

H

_ -

>99 % ee, 30 %yield, PFL

85b [41]

>99 % ee, 51 % yield

>99 % ee, 49 % yield, PCL

AcOo-&R EtO

8Ga [42]

R = n-Bu, n-C5HI1,n-C6H13, Ph 95-98 % ee, 17-35 % yield, PCL

AcO

A. . .,,

R

ent-8Ga [42]

E td

HOA.,.,,,,R Etd

unstable

HO',/&R

EtO

R = ~ - B u ,n-CsH11, n-C6H13, Ph >99 % ee, 32-49 % yield, CAL-B

unstable

>98 % ee, 46 % yield, CCL 90 % ee, 51 %yield, CRL

>98 % ee, 43 % yield >99 % ee, 44 % yield

87a [43] 87a [43]

8Gb [42]

ent-8Gb [42]

87b [43] 87b [44]

1 1 . 1 Hydrolysis and Formation ofcarboxylid Acid Esters

I

469

Table 11.1-16.

"

O

(cont.).

D

Aco

&

n

""a.

1 13 % ee, 55 %yield, CCL 2 20 % ee, 65 % yield, CCL

88b [45] 89b [45]

95 % ee, 13 %yield 98 % ee, 20 % yield

88a [45] 89a [45]

90a [4G]

90b [4G]

\

\

Me

O

55% ee, 50% yield, CE

R

76% ee, 42 % yield, CE

Me

R=

OH R @

\ Me ""Ph

91a [47]

90 % ee, 48 % yield, CRL 99 % ee, 42 % conversion CE (porcine pancreas), sodium taurocholate

53 % ee, 52% conversion CE (porcine pancreas), sodium

taurocholate

H?

49 % ee, 40 % conversion CE (porcine pancreas), sodium taurocholate

91b [47]

88 % ee, 45 % yield

G1% ee, 42% conversion

CE (porcine pancreas), sodium taurocholate

49 % ee, 52 % conversion CE (porcine pancreas), sodium taurocholate

AcO

33 % ee, 40 % conversion CE (porcine pancreas), sodium taurocholate

470

I

I 1 Hydrolysis and Formation ofC-0 Bonds Table 11.1-16.

(cont.).

Ac?

Act?

94b [47]

44% ee, 51 % conversion

43 % ee, 51 % conversion CE (porcine pancreas), sodium taurocholate

OH

CE (porcine pancreas), sodium taurocholate

R

0 95a [48]

E = 10-15, CE (porcine pancreas), sodium taurocholate

95b [48]

E = 10-15, CE (porcine pancreas), sodium taurocholate

ba,..CHCI

9Gb [48]

/

E = 4.6, CE (porcine pancreas), sodium taurocholate

E = 4.6, CE (porcine pancreas), sodium taurocholate

AcO

97b [48] nBu

E = 14, CE (porcine pancreas), sodium taurocholate

E = 14, CE (porcine pancreas), sodium taurocholate

98b [48]

E = 10, CE (porcine pancreas), sodium taurocholate

E = 10, CE (porcine pancreas), sodium taurocholate

g:

Table 11.1-16.

7 1 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters

I

471

(cont.).

s's*,,ph

/

AcO +jPh,,:

99b [48]

99a [48]

\

\

E = 19, CE (porcine pancreas), sodium taurocholate

E = 19, CE (porcine pancreas), sodium taurocholate

moH moAC lOOa [48]

OAc

1OOb [48]

OAc

E > 400, CE (porcine pancreas), sodium taurocholate

E > 400, CE (porcine pancreas), sodium taurocholate

101b [48]

lOla [48]

OAc

OAc

c=b"

&OH

E > 10, CE (porcine pancreas), sodium taurocholate

E > 10, CE (porcine pancreas), sodium taurocholate

0 I

N3

.3

0-

I

102b [49]

102a 1491

....Ph HO

AcO

83 % ee, 51 % conversion CE (Pseudomonaspurorescens) 51 % ee, 35% conversion CE (porcine pancreas)

80% ee, 51 % conversion

1 Z.-F. Xie, H. Suemune, K. Sakai,]. Chem. Soc., Chem. Commun. 1987,838. 2 H. Honig, P. Seufer-Wasserthal, F. Fiilop,]. Chem. Soc., Perkin Trans. 1 1989,2341.

CE (Pseudomonaspurorescens) 96% ee, 35 % conversion CE (porcine pancreas) 3 Z.-F. Xie, 1. Nakamura, H. Suemune, K. Sakai, J. Chem. SOC.,Chem. Commun. 1988,9GG. 4 K. Laumen, D. Breitgoff, R. Seemeyer, M. P. Schneider,]. Chem. Soc., Chem. Commun. 1989, 148.

472

I

7 7 Hydrolysis and Formation of C-0 Bonds 5 S. Mitsuda, S. Nabeshima, H. Hirohar, Appl.

Microb. Biotechnol. 1989, 31, 334. 6 H. Danda, A. Maehara, T. Umemura, Tetrahedron Lett. 1991, 32, 5119. 7 H. Danda, T. Nagatomi, A. Maehara, T. Umemura, Tetrahedron 1991, 47, 8701. 8 K. Fritsche, C. Syldatk, F. Wagner, H. Hengelsberg, R. Tacke, Appl. Microbiol. Biotechnol. 1989, 31, 109. 9 N. Klempier, K. Faber, H. Griengl, Biotechnol. Lett. 1989,685. 10 N. Klempier, P. Geymayer, P. Stadler, K. Faber, H. Griengl, Tetrahedron: Asymmetry 1990, I, 111. 11 K. Laumen, M. P. Schneider,]. Chem. Soc., Chem. Commun. 1988,598. 12 L. Dumortier, J. Van Eycken, M. Vandewalle, Tetrahedron Lett. 1989, 30, 3201. 13 G. Eichberger, G. Penn, K. Faber, H. Griengl, Tetrahedron Lett. 198G, 27, 2843. 14 T. Oberhauser, M. Bodenteich, K. Faber, G. Penn, H. Griengl, Tetrahedron 1987, 43, 3931. 15 R. Saf, K. Faber, G. Penn, H. Griengl, Tetrahedron 1988,44,389. 16 0. Hoshino, K. Itho, B. Umezawa, H. Akita, T. Oishi, Tetrahedron Lett. 1988, 29, 567. 17 Y Hirose, M. Anzai, M. Saitoh, K. Naemura, H. Chikamatsu, Chem. Lett. 1989, 1939. 18 K. Naemura, T. Matsumara, M. Komatsu, Y. Hirose, H. Chikamatsu, Bull. Chem. Soc.]pn. 1989,62, 3523. 19 S. Takano, M. Suzuki, K. Ogasawara, Tetrahedron: Asymmetry 1993,4,1043. 20 A. K. Gupta, R. J. Kazlauskas, Tetrahedron: Asymmetry 1993,4,879. 21 Z:F. Xie, H. Suemune, K. Sakai, Tetrahedron: Asymmetry 1990, I , 395. 22 P. Washausen, H. Grebe, K. Kieslich, E. Winterfeldt, Tetrahedron Lett. 1989, 30, 3777. 23 X. Chen, S. M. Siddiqi, S. W. Schneller, Tetrahedron Lett. 1992, 33, 2249. 24 S. Takano, T. Yamane, M. Takahashi, K. Ogasawara, Tetrahedron: Asymmetry 1992, 3, 837. 25 R. Seemayer, M. P. Schneider, J. Chem. Soc., Chem. Commun. 1990, 2359. 26 T. Izumi, F. Taura, K. Sasaki, Bull. Chem. Soc. Jpn. 1992,65,2784.

27 P. Duhamel, P. Renauf, D. Cahard, A. Yebga, 1. M. Poirier, Tetrahedron:Asymmetry 1993,4, 2447. 28 A. K. Gosh, Y Chen, Tetrahedron Lett. 1995, 36, 505. 29 H. Nagata, K. Ogasawara, Tetrahedron Lett. 1999, 40, 6617. 30 L. Aribi-Zouioueche, J.-C. Fiaud, Tetrahedron Lett. 2000,41,4085. 31 J. Doussot, A. Guy, R. Garreau, A. Falguisres, C. Ferroud, Tetrahedron: Asymmetry 2000, 11, 2259. 32 K. Sugawara, Y. Imanishi, T. Hashiyama, Tetrahedron: Asymmetry 2000, 1I , 4529. 33 H:J. Gais, C. Griebel, H. Buschmann, Tetrahedron: Asymmetry 2000, 11, 917. 34 T. Taniguchi, M. Takeuchi, K. Kodata, A. S. EIAzab, K. Ogasawara, Synthesis 1999, 1325. 35 S. Takano, 0. Yamada, H. lida, K. Ogasawara, Synthesis 1994, 592. 3G a) C. R. Johnson, M. W. Miller,]. Org. Chenr. 1995, 60,6674 b) 0. Block, G. Klein, H.-J. Altenbach, D. J. Brauer, ]. Org. Chem. 2000, 65, 716. 37 K. Kadota, A. S. ElAzab, T. Taniguchi, K. Ogasawara, Synthesis 2000, 1372. 38 M. Takahashi, R. Koike, K. Ogasawara, Chem. Pharm. Bull. 1995,43, 1585. 39 7. Taniguchi, R. M. Kanada, K. Ogasawara, Tetrahedron: Asymmetry 1997. 8. 2773. 40 R. A. MacKeith, R. McCague, H. F. Olivo, S. M. Roberts, S. J. C. Taylor, H. Xiong, Bioorg. Med. Chem. 1994,2, 387. 41 H. Nagata, N. Miyazawa, K. Ogasawara, Synthesis 2000, 2013. 42 B. Westermann, B. Krebs, Org. Lett. 2001, 3, 189. 43 A. Cipiciani, F. Fringulli, V. Mancini, 0. Piermatti, A.M. Scappini, Tetrahedron 1997,53,11853. 44 D. Pamperin, C. Schulz, H. Hopf, C. Syldatk, M. Pietzsch, Eur.]. Org.Chem.,1998, 1441. 45 J. Y. Goujon, F. Zammattio, B. Kirschleger, Tetrahedron: Asymmetry 2000, I I , 2409. 46 E. Mizuguchi, M. Takemoto, Tetrahedron: Asymmetry 1993,4,1961. 47 A. N. Serreqi, R. J . Kazlauskas, ]. Org. Chem. 1994, 59, 7609. 48 A. N. Serreqi, R. J. Kazlauskas, Can. J. Chem. 1995, 73, 1357. 49 X. Yang, A. R. Reinhold, R. L. Rasati, K. K. C. Liu, Org. Lett. 2000, 2, 4025.

hydrolases as such or of opposite configuration are contained in Tables 11.14 and 11.1-16.

11.1.1.2 Formation o f Carboxylic Esters 11.1.1 2.1

Lipases

Of the many hydrolases known, only the lipases, subtilisin and to some extent a-

chymotrypsin, pig liver esterase, and thermolysin [G4aJ show a sufficiently high

1 1 . 7 Hydrolysis and Formation ofCarboxylid Acid Esters

I

473

catalytic activity in organic solvents of low water content to be of practical value for asymmetric synthesis through acylation of prochiral or racemic alcohols, alcoholysis of prochiral or racemic acylated alcohols and prochiral anhydrides, and cyclization of racemic hydroxy carboxylic acids. Lipases, as stated previously, are unique for organic synthesis, since they exhibit not only a high catalytic activity in water or in two-phase systems composed of water and a water-immiscibleorganic solvent or the liquid substrate, but most importantly also in water-miscibleor immiscible organic solvents of low water content. This allows for the attainment of favorable equilibria not only in asymmetric hydrolysis but also in esterification reactions. In the formation of carboxylic esters in an anhydrous organic solvent, its hydrophobicity and the water activity have a major influence on the reacti0n[~'9 36* 1341. Hence, the organic solvent used can significantly influence the selectivity of a lipase-catalyzedenantiotopos- or enantiomer-differentiating reaction. Furthermore, the acyl donor may influence reactivity and selectivity. Lipases are most advantageously used for the acylation of prochiral diols or racemic alcohols and for the alcoholysis of racemic acylated alcohols. Generally, through acylation of a prochiral diol or racemic alcohol in an organic solvent such as diethyl ether, diisopropyl ether, tert-butyl methyl ether, tetrahydrofuran, dichloromethane, pentane, hexane, toluene or tert-pentyl alcohol with acylating reagents such as vinyl acetate, vinyl butyrate, vinyl propionate, vinyl laurate, vinyl palmitate, vinyl chloroacetate,isopropenyl acetate, oxime esters, ethyl acetate, ethyl propionate, trifluoroethyl butyrate, trichloroethyl butyrate, trifluoroethyl acetate, ethyl octanoate, ethyl methoxy acetate, ethyl thiooctanoate, acetic anhydride, succinic anhydride or 2-phenyloxazolin-5-one and hydrolysis of the corresponding prochiral diacetate (dipropionate,dichloroacetate) or racemic acetate (chloroacetate)in water or in water and a water-immiscible organic solvent, access to both enantiomers of the corresponding monoacetate and alcohol, respectively, is provided with one enzyme (Tables 11.1-10 to 11.1-12 and 11.1-18).This is because of the same enantiotopic group and enantiomer recognition shown in general by the enzyme in both reactions (Scheme 11.1-12), and favorable opposite equilibria. In many cases vinyl acetate, isopropenyl acetate, ethyl acetate and propionyl acetate not only serve as acylating reagents but also as solvents. For the acylation of prochiral diols, ee values of monoacetates (about 90%) can be raised considerably in most cases by a higher degree of conversion at the expense of a lower chemical yield to the point where an enantiomer-differentiating formation of the diacetate can take place (Scheme 11.1-11,Figure 11.1-l),because in most cases the enzyme preferentially catalyzes the acylation of the minor enantiomer. The enantioselectivity and thermostability of lipases is frequently enhanced in organic solvents of low water content. A minimum amount of water is required for the catalyhc activity of the lipase. In most cases lipase preparations with a residual water content of approximately 1 % in anhydrous organic solvents are employed. Frequently in organic solvents of low water content the thermostability of lipases is much higher than that in aqueous solution [361. The use of lipase in other forms than lyophilized powders, as for example on different kinds of solid supports, entrapped in sol-gelmaterials or as CLECs, has the

474

I

I I Hydrolysis and Formation ofC-0 Bonds Lipase-catalyzedenantiotopos-differentiating acylation of prochiral acyclic diols in organic solvents (CCL Candida cylindracea lipase, PFL PseudomonasPuorescens lipase, PPL pig pancreas lipase, CVL Chromobacterium viscosum lipase, PSL Pseudomonas sp. lipase, RJL Rhizomucorjavanicus lipase, A N L Aspergillus niger lipase, CAL Candida antarctica lipase, not specified, PCL Pseudomonos cepacia lipase, CRL Candida rugosa lipase).

Table 11.1-17.

R' 1 2 3 4 5

5

R2

R3

Me H CHFCH-CH~ AC CHz=CH-(CH2)2AC Ph Ac CHzPh Ac CHzPh Ac

Ac H H H H H

CH,

G

I

7 8 9

10 11

11 11 12

Ac

H

L

i-Pr C-CGHLI c-CaHiICH2 Cbz OCH2Ph

Ac Ac AC Ac

H

H H H H Ac

OCH2Ph OCH2Ph OEt

H H H

Ac Ac Ac

Lipase

Acyldonor

PFL PFL PPLa PPL" PPLa PFL PFL PFL PFL

vinyl acetate vinyl acetate ethyl acetate ethyl acetate ethyl acetate vinyl acetate vinyl acetate vinyl acetate vinyl acetate

PFL PPLa PPL" PPL PFL PFL

vinyl acetate ethyl acetate ethyl acetate vinyl acetate isopropenyl acetate vinyl acetate phenyl acetate phenyl acetate

PFL PFL

ee ("A) 60 81 90 92 13 294 97 86 90

yield

Ref.

70 89 70 98 90 100 96 93 95

[l]

61

58 10 97 96

85 90 90 77 53

[I] [2] [2] [4] [4]

92 90 90

92 88 90

[5] [5] [5, 61

("A)

[I] [2] [2] [2] [3] 111 [3] [l]

0011 0 B n AcO-OH

X

13 [71

OBn

O

14 PI

HOAOAc

298 % ee, 51 %yield, PFL vinyl acetate

295 % ee, 70 % yield, CCL vinyl acetate

R. r O H

Si

M~/L OCOiPr R = Ph

70 % ee, 80 %yield, CCL methyl isobutyrate R = n-octyl 75 % ee, 63 %yield, CCL methyl isobutyrate

15 [9] 16 [9]

70 % ee, 50 % yield, CVL methyl isobutyrate 76 % ee, 70 %yield, CVL methyl isobutyrate

ent-15 [9] ent-16 [9]

1 1. I Hydrolysis and Formation ofcarboxylid Acid Esters

I

475

Table 11.1-17.

(cont.).

MeO-0

CZ;OCQH,,

17 [lo]

18 [ll]

MeO-0

OTBDMS

95 % ee, 90 % yield, PPL C~HI~COCH~CCI~

97 % ee, 94 % yield, CRL, vinyl acetate

89 % ee, 80 % yield, PPL, vinyl acetate

>99 % ee, 76 % yield, PPL, vinyl acetate 21 (131

HO*OAC

AcO

0 98 % ee, 86 %yield, PFL, vinyl acetate

98 % ee, 75 % yield, PCL, vinyl acetate

22 [14]

92 % ee, 55 %yield, PPL, methyl acetate

94 % ee, 86 %yield, PPL, vinyl acetate

25 [16]

99 % ee, 86 %yield, PSL, vinyl acetate

t B u 0 / R \ c O AOH C

R = H , 91 % ee, 97 % yield, PFL, vinyl acetate R = Me, 88 % ee, 80 % yield, PFL, vinyl acetate

OH 28 [18] Me,PhSi

>99 % ee, 98 % yield, PFL, vinyl acetate 95 % ee, 98 % yield, PCL, vinyl acetate

26 [17] 27 [17]

29 [19] BZO

96 % ee, 63 % yield, PPL, vinyl acetate

11 Hydrolysis and Formation ofC-0 Bonds Table 11.1-17.

;

(cont.).

+ N

30 [20]

9

0Ac

31 [20]

OH

96 % ee, 84 % yield, PPL, vinyl acetate

&O

98 % ee, 81 % yield, PPL, vinyl acetate

q

Ac

\

0Ac OH

OH

84 % ee, 21 % yield, ANL, vinyl acetate 98 % ee, 42 % yield, CAL, vinyl acetate

33 (211

32 1211

97 % ee, 46 %yield, PPL, vinyl acetate

ent-32 [211

WoAc OH

2

CH2 99 % ee, 97 %yield, PCL, vinyl acetate CF2 99 % ee, 82 % yield, PCL, vinyl acetate

34 [22]

&b /

38 [23]

61 % ee, 84 % yield, CRL,

85-92 % ee, 35-78 % yield, CRL, 1-ethoxyvinyl2-hroate abs. config. not determined

1-ethoxyvinyl2-furoate abs. config. not determined

Ph

Ph

HOAc

36 [22]

35 [22]

37 [23]

, .Ph

87 % ee, 95 %yield, PCL, vinyl acetate

39 [24]

Ph

nOH

HO

AcO

90 % ee, 20 %yield, RJL, vinyl acetate

90 % ee, 28 % yield, CCL, vinyl acetate

ent-39 [24]

7 7 . I Hydrolysis and Formation ofCarboxylid Acid Esters

I

477

Table 11.1-17.

(cont.).

R

Me Et

>99 % ee, 26 % yield, PCL phenyl acetate 94 % ee, 43 % yield, PCL phenyl acetate

40 [25] 40 [25]

3

AcO

HO

R’

N’ I

R’ R‘

R2

Me

Me

Me

Me

Et

Et

Et

Et

97 % ee, 42 % yield, PCL vinyl acetate 96 % ee, 77 % yield, PCL phenyl acetate 96 % ee, 52 %yield, PCL vinyl acetate 84 % ee, 43 %yield, PCL phenyl acetate

41 [25] 41 [25] 42 [25] 42 [25]

a purified PPL

1 K. Tsuji, Y. Terao, K. Achiwa, Tetrahedron Lett. 1989,30,6189. 2 G.M. Ramos Tombo, H:P. Schar, X. Femandez I Busquets, 0. Ghisalba, Tetrahedron Lett. 1986, 27, 5707. 3 S. Atsumi, M. Nakano, Y. Koike, S. Tanaka, M. Ohkubo, T. Yonezawa, H. Funabashi, J. Hashimoto, H. Morishima, Tetrahedron Lett. 1990,31,1601. 4 Y. F. Wang, J. J. Ialonde, M. Momongan, D. E. Bergbreiter,C.-H. Wong. ]. Am. Chem. Soc. 1988, 110,7200. 5 Y.Terao, M. Murata, K. Achiwa, T Nishio, M. Akamtsu, M. Kamimura, Tetrahedron Lett. 1988,29,5173. 6 M. Murata, Y.Terao, K. Achiwa, T. Nishio, K. Seto, Chem. Phann. Bull. 1989,37,2670. 7 K. Burgess, I. Henderson, Tetrahedron Lett. 1991, 32, 5701. 8 C. Bonini, R. Racioppi, L. Viggiani, G. Righi, L. Rossi, Tetrahedron: Asymmetry1993,4, 793. 9 A:H. Djerourou, L. Blanco, Tetrahedron Lett. 1991, 32,6325. 10 H. J. Bestmann, U. C. Philipp, Angau. Chem. 1991, 1 0 3 , 7 8 Angew. Chem., Int. Ed. Engl. 1991,30,86.

11 R. ChCnevert, G.Courchesne, Tetrahedron: Asymmetry1995,6,2093. 12 T. Bando, Y. Namba, K. Shishido, Tetrahedron: Asymmetry1997,8,2159. 13 C. Bonini, R. Racioppi, L. Viggiani, Tetrahedron: Asymmetry1997,8,353. 14 J. C. Anderson, S. V. Ley, S. P. Marsden, Tetrahedron Lett. 1994, 35, 2087. 15 C. J. Bamett, T. M. Wilson, S. R. Wendel, M. J. Winningham, J. B. Deeter,]. Org. Chem. 1994,59, 7038. 16 A. Avdagit, M. Gelo-PujiC, V. Sunjit, Synthesis 1995,1427. 17 F:R. Alexandre, F. Huet, Tetrahedron :Asymmetry 1998, 9, 2301. 18 B. Danieli, G. Lesma, S. Macecchini, D. Passarella, A. Silvani, Tetrahedron: Asymmetry1999, 10, 4057. 19 V. B6dai. L. Novik, L. Poppe, Synlett 1999, 759. 20 G. Guanti, E. Narisano, R. Riva, Tetrahedron: Asymmetry1997,8,2175. 21 L. Banfi, G . Guanti, A. Mugnoli, R. Riva, Tetrahedron: Asymmetry1998, 9, 2481. 22 T. Yokomatsu, T. Minowa, T. Murano, S. Shibuya, Tetrahedron 1998, 54,9341.

478

I

I 7 Hydrolysis and formation ofC-0 Bonds 23 S . Akai, T. Naka, T. Fujita, Y. Takebe, Y. Kita,Chem. 25 K. Takabe, Y. Iida, H. Hiyoshi, M. Ono, Y. Hirose, Commun. 2000,1461. Y. Fukui, H. Yoda, N. Mase, Tetrahedron: 24 G. Nicolosi, A.Patti, M. Piatelli, C. Sanfilippo, Asymmetry 2000,II,4825. Tetrahedron: Asymmetry 1994,5, 283.

advantage of easy recovery by filtration and reuse. Furthermore, these lipases have higher stability, and, most importantly, their activity and selectivity are often much higher than with the lyophilized powders. One should bear in mind, however, that in nearly all cases lipase preparations are used, which contain, as well as a large amount of mostly unspecified material such as proteins, carbohydrates and solid support materials, only a minor amount of the lipase and in several case even additional mostly unidentified hydrolases. The solid material contained in the crude lipase preparation may have an important stabilizing function in organic solvents, in which the lipase preparation is insoluble. Crude lipase preparations supplied commercially contain up to 7 % of water. Drying the solid material in vacuum may reduce the water content. Acylating reagents such as vinyl acetate and isopropenyl acetate are very useful since they allow for an extreme equilibrium position in acylation because of the tautomerization of the vinyl and isopropenyl alcohol formed to acetaldehyde and acetone, respectively. The possible harmful effect of acetaldehyde on the enzyme with the crude lipase preparation used poses practically no problem in most cases because the low price of the enzyme enables relatively large amounts of it to be used. Synthetically, lipase-catalyzed acylations are convenient to carry out and, in contrast to the corresponding hydrolyses, catalysts are easy to recover and can be reused. A series of alkyl, alkoxy or acylamino 1,3-proanediol derivatives substituted in 2-position have been subjected to lipase-catalyzed acylation, and the monoacetates (1-12,19, 20, 23-38, 40-42) were obtained with moderate to high enantiomeric excess (Table 11.1-17).For the monoacetates 1-12,reactions with and in ethyl acetate are usually slower than those with and in vinyl acetate. As in the hydrolysis of the corresponding diacetates, much higher selectivities were recorded with the yet unidentified carboxyl esterase from crude pig pancreas lipase. An excellent lipase for the enantioselective acylation of 3-benzyloxy-l,3-propanediol is Pseudomonas Juorescens lipase, which gives high selectivity with vinyl acetate, isopropenyl acetate and ethyl acetate. By carrying the acylation further, to a certain extent to the diacetate, the enantiomerically pure monoacetate should be obtainable. Sterically demanding substituents in 2-position such as in 20, 25,26,28,29 and 34-36 guarantee high enantioselectivity and yield for the monoacetates. Sila propanediol derivatives (15,16),and butanediol, pentanediol, hexanediol and heptanediol derivatives (17,18, 21,22)(Table 11.1-17)have also been prepared. Monoacetates of Table 11.1-17which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-4and 11.1-10. Cyclic dimethanol derivatives have been extensively studied not only in lipasecatalyzed hydrolysis (Table 11.1-11) but also in lipase-catalyzed enantioselective acylation for synthetic and mechanistic reasons (1-16,20, 30, 32, 33, 37, 40, 45, Generally, enantioselectivities in acylation of 47-53, 57-62, 66,72) (Table 11.1-18). the diol and hydrolysis of the corresponding diacetate yielding enantiomeric com-

11.1 Hydrolysis and Formation ofCarboxylid Acid Esters

I

479

Lipase-catalyzedenantiotopos-differentiatingacylation of prochiral cyclic diols in organic solvents (PPL pig pancreas lipase, PFL Pseudomonasfluorescens lipase, PCL Pseudomonas cepacia lipase, CCL Candida cylindracea lipase, MSL Mucor sp. lipase, CVL Chrornobacteriurn viscosum lipase, CCL Ceotrichum candidum lipase, CRL Candida rugosa lipase, M M L Mucormiehei lipase, CAL-B Candida antarctica B lipase, LIP Pseudomonas sp. lipase-Toyobo). Table 11.1-18.

1 111 295 % ee, 82 % yield, PFL, vinyl acetate

2 111 88 % ee, 87 % yield, PFL,vinyl acetate 295 % ee, 82 % yield, PFL,ethyl acetate

3 I11

295 % ee, 85 % yield, PFL vinyl acetate

94 % ee, 64 % yield, PPL,vinyl acetate

25 % ee, 52 % yield, PPL,vinyl acetate

298 % ee, 87 %yield, PPL,vinyl acetate

a::c

R?,, R' R'

295 % ee, 87 %yield, PPL vinyl acetate

OH Ph3CO H H H H

8 [2, 31

R2

H

H

c1 SPh SO2Ph N3

no?""

PPL,vinyl acetate 98 % ee, 92 %yield 45 % ee, 72 % yield 296 % ee, 84 % yield 296 % ee, 85 % yield 68 % ee, 69 % yield 295 % ee, 86 % yield

10 PI

9 [31

W'o&OAc

77 % ee, 91 % yield, PPL,vinyl acetate 298 % ee, 78 % yield, PFL,vinyl acetate 9 % ee, 71 % yield, PFL,acetic anhydride

7 % ee, 44 % yield, PFL, vinyl acetate

f 80 % ee, 60 % yield, PFL,vinyl acetate

OAc

100 % ee, 32 % yield, CCL, vinyl acetate

480

I

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-18.

(cont.)

0

80 % ee, 80 % yield, GCL, vinyl acetate, CHzClz 95 % ee, 72 % yield, GCL, vinyl acetate, EtzO

99 % ee, 68 %yield, PPL, ethyl acetate 99 % ee, 92 %yield, PPL, vinyl acetate 0

0

6

96 % ee, 71 % yield, CCL, isopropenyl acetate 76 % ee, 70 %yield, CCL, vinyl acetate

17[7-111

OH

8 % ee, 38 % yield, PPL, vinyl acetate 87 % ee, 72 % yield, CCL, vinyl acetate

I

18 [lla]

r - i OH rofl

R = n-Pr, n-C7H15’ CHlCl 98 % ee, -, PFL, vinyl acetate >99-80 % ee, 58-39 % yield, pancreatin, 98 % ee, 52 % yleld, PPL, vinyl acetate trichloroethyl alkanoate 295 % ee, 48 % yield, pancreatin, trichloroethyl acetate >99 % ee, 50 % yield, PPL, trichloroethyl acetate >99 % ee, 65 % yield, pancreatin, vinyl acetate 94 % ee, 85 %yield, MSL, vinyl acetate

84 % ee, -, PPL, vinyl acetate

95 % ee, 80 %yield, PFL, vinyl acetate AcO

1

59 % ee, 60 %yield, PSL, vinyl acetate

295 % ee, 95 %yield, PCL, isopropenyl acetate

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters

I

481

Table 11.1-18.

(cont.).

AcO

1 23 [16]

@OMe(OBn)

A

24 [16]

"0H

HO

N3

297 % ee, 89 % yield, PCL, vinyl acetate

fj

no acylation with vinyl acetate and several 1ipases

25 1171

26 1181

/

OH

AcO

95 % ee, 51 % yield, PCL, vinyl acetate

"?:I,

.::0

299 % ee, 38 %yield, PPL 94 % ee, 75 % yield, PCL, vinyl acetate

27 [18]

AcO

AcO

2 % ee, 60 %yield, MSL, vinyl acetate

299 % ee, 94 % yield, PPL 299 % ee, 87 % yield, DCL, vinyl acetate

AcO \

29 [18]

HO

/-OH

30 1191

O ' AC

86 % ee, 75 % yield, PPL, vinyl acetate

31 [20]

OAc

97 % ee, 68 % yield, CCL 18 % ee, 57 %yield, PPL, vinyl acetate

o\;b

295 % ee, 90 % yield, PCL, isopropenyl acetate 100 % ee, 80 % yield, PCL, vinyl acetate

Gc;c;; a Fe

33 [21]

100 % ee, 80 % yield, CVL, vinyl acetate

34 [22]

HO

298 % ee, 46 % yield, PCL, isopropenyl acetate

482

I

11 Hydrolysis and Formation ofC-0 Bonds Table 11.1-18.

(cont.).

35 [22]

+o

b

36 [22]

AcO

HO

84 % ee, 92 %yield, PCL, vinyl acetate

17 % ee, -, PCL, isopropenyl acetate

38 [24]

%OH HO "*'

299 % ee, 81 % yield, PFL, vinyl acetate

299 % ee, 87 % yield, PCL, vinyl acetate

C0,Et

OAc

299 % ee, 96 % yield, PSL, vinyl acetate

95 % ee, 98 % yield, CCL, vinyl acetate

41 [27]

299 % ee, 99 %yield, PSL, vinyl acetate

98 % ee, 70 % yield, CRL, vinyl acetate 91 % ee, 64 %yield, PCL, vinyl acetate

43 [29] AcO

>98 % ee, 90 % yield, PCL, isopropenyl acetate

bOH 44 [30]

R = Cbz: >98 % ee, 91 % yield, PCL, isopropenyl acetate R = Boc: >98 % ee, 92 % yield, PCL, isopropenyl acetate

7 7.7 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.1-18.

(cant.).

46 [32]

45 [31]

U

H

298 % ee, 89 %yield, PCL, vinyl acetate 95 % ee, 93 %yield, MML, vinyl acetate 70 % ee, 89 % yield, CAL-B, vinyl acetate

>99 % ee, 88 % yield, PPL, ethyl acetate

RO

O

OR

R

TBDMS >99 % ee, 58 %yield, PCL, vinyl acetate >99 % ee, 65 % yield, PSL, vinyl acetate TIPS 299 % ee, 77 %yield, PSL, vinyl acetate CH2Ph 299 % ee, 81 %yield, PSL, vinyl acetate

47 [33] 48 [33] 49 [33]

51 [33]

50 [33]

HO

91 % ee, 82 %yield, PSL, vinyl acetate

92 % ee, 92 %yield, PSL, vinyl acetate

Ro$2: 52 [33]

AcO

O OAc H

88 % ee, 73 %yield, PSL, vinyl acetate

Ac

53 [34]

Bn

>99 % ee, 65 % yield, PPL, vinyl acetate

R = 4-MeOCbH4CHz 94 % ee, 76 %yield, PSL, vinyl acetate

a

HOO +

54 [35]

aoAc OH

n = 2: >98 % ee, 94 %yield, MML, vinyl acetate n = 3 : 80 % ee, 85 %yield, PSL, vinyl acetate

55 [35] 56 [35]

I

483

7 7 Hydrolysis and Formation of C-0 Bonds

(cont.).

Table 11.1-18.

OR

1

R = Boc: >98 % ee, 74 % yield, PFL, vinyl acetate R = Cbz: >98 % ee, 78 % yield, PFL, vinyl acetate

57 [36] 58 [36]

R = TBDMS: >98 % ee, 70 %yield, 59 [37] CAL-B, isopropenyl acetate R = MOM: >95 % ee, 68 %yield, GO [37] CAL-B, isopropenyl acetate

C0,Me

0.

AcO'"'

Cbz 61 [38] 62 [38]

C0,Me

65 [39]

"'OAc

'I.'

"OH

R

R = H: 95 % ee, 80 %yield, CAL-B,vinyl acetate R = OMOM: 96 % ee, 83 % yield, CAL-B, vinyl acetate

HO

I

>99 % ee, 97 %yield, PPL, vinyl acetate

R = Me: >99 % ee, 42 %yield, PSL, 63 [39] vinyl acetate R = Me: 93 % ee, 50 %yield, PSL, vinyl acetate 64 [39] R = Et: >99 % ee, 53 %yield, PSL, vinyl acetate

&F

66 [40]

>95 % ee, 89 %yield, PSL, vinyl acetate

68 [41]

w

HO"'

>99 % ee, 88 % yield, LIP, vinyl acetate

>99 % ee, 82 % yield, LIP, vinyl acetate

R

Bn 4-MeOCsH4CHz 2-NaphthylCHz

93 % ee, 93 % yield, LIP, vinyl acetate 84 % ee, 85 % yield, LIP, vinyl acetate 93 % ee, 93 %yield, LIP, vinyl acetate

69 [42]

70 [42] 71 [42]

I I, I Hydrolysis and Formation of CarboxylidAcid Esters Table 11.1-18.

(cont.).

72 [43]

92 % ee, 47 % yield, PPL, vinyl acetate >98% ee, 75 %yield, PCL 90 % ee, 73 %yield, PSL

1 U.Ader, D. Breitgoff, P.Klein, K. E. Laumen, M. P. Schneider, Tetrahedron Lett. 1989,30,1793. 2 H. Hemmerle, H.-J. Gais, Tetrahedron Lett. 1987, 28,3471. 3 H. Hemmerle, Ph. D. 7'hesis,Universitat Freiburg 1990. 4 M. Ihara, M. Suzuki, K. Fukumoto, C. Kabuto, 1.Am. Chem. SOC.1990,112,1164. 5 M.Murata, S. Ikoma, K. Achiwa, Chem. Pharm. Bull. 1990,38,2329. 6 C. Andreu, J. A. Marco, G . Asensio, /. Chem. Soc., Perkin Trans. 1 1990,3209. 7 S.-H. Hsu, S.4. Wu, Y.-F. Wang, C.-H. Wong, Tetrahedron Lett. 1990,31,6403. 8 K. A. Babiak, J. S . Ng, J. H. Dygos, C. L. Weyker, Y.-F. Wang, C.-H. Wong,]. 0%.Chem. 1990,55, 3377. 9 F. Theil, S . Ballschuh, H. Schick, M. Haupt, B. Hafner, S. Schwarz, Synthesis1988,540. 10 G. Jommi, F. Orsini, M. Sisti, L. Verotta, Gazz. Chim. Ifal. 1988,118,863. 11 a) F. Theil, H. Schick, M. A. Lapitskaya, K. K. Pivnitsky, LiebigsAnn. Chem. 1991,195;b) F. Theil, H. Schick, D. Weichert, K. Tannenberger, G. Klappach,]. Prakt. Chem. 1991,333,497. 12 H. Pottie, J . Van der Eycken, M. Vandewalle, H. Roper, Tetrahedron: Asymmetry1991,2, 329. 13 K. Naemura, A. Furutani,]. Chem. SOC.,Perkin Trans. 11991,2891. 14 C. R. Johnson, A. Golebiowski,T. K. McGill, D. H. Steensma, Tetrahedron Lett. 1991,32,2597. 15 C. R. Johnson, A. Golebiowski, D. H. Steensma, 1. Am. Chem. SOC.1992,114,9414. 16 C.Hoenke, P. Kliiwer, U. Hugger, R. Krieger, H. Prinzbach, Tetrahedron Lett. 1993,34,4761. 17 K. J. Hams, Q.-M. Gu, Y.-E. Shih, G. Girdaukas, C. J. Sih, Tetrahedron Lett. 1991,32,3941. 18 F. Theil, H. Schi&, G. Winter, G. Re&, Tetrahedron 1991,47,7569. 19 M. Mekrami, S. Sicsic, Tetrahedron: Asymmetry 1992,3,431. 20 C. R. Johnson, P. A. PI&,J. P. Adams, /. Chem. Soc., Chem. Commun.1991,1006. 21 G. Nicolosi, R. Monone, A. Patti, M. Piatelli, Tetrahedron: Asymmetry1992,3,753. 22 S.I. Bis, T. Whitaker, C. R. Johnson, Tetrahedron: Asymmetry1993,4,875. ~~

23 M. Tanaka, M. Yoshioka, K. Sakai, Tetrahedron: Asymmetry1993,4,981. 24 S. Takano, M. Moriya, Y. Higashi, K. Ogasawara, 1.Chem. Soc., Chem. Commun. 1993,177. 25 N. Toyooka, A. Nishino, T. Momose, Tetrahedron Lett. 1993,34,4539. 26 M. Sato, H. Ohuchi, Y. Abe, C. Kaneko, Tetrahedron: Asymmetry1992,3,313. 27 M. Sato, T.Hirokawa, H. Hattori, A. Toyota, C. Kaneko, Tetrahedron: Asymmetry1994,5,975. 28 K. Toyama, S. Iguchi, T. Oishi, M. Hirama, Synlett 1995,1243. 29 C. R. Johnson, L. S . Harikrishnan, A. Golebiowski, Tetrahedron Lett. 1994,35,7735. 30 C. R. Johnson, S . J. Bis,/. Org. Chem. 1995,60, 615. 31 B. Danieli, G.Lesma, M. Mauro, G. Palmisano, D. Passarella,]. Org. Chem. 1995.60,2506. 32 A. Patti, C.Sanfilippo, M. Piatelli, G. Nicolosi, /. Org. Chem. 1996,61,6458. 33 T. Oishi, M. Maruyama, M. Shoji, K. Maeda, N. Kumahara, S . Tanaka, M. Harima, Tetrahedron 1999,557471. 34 a) G. Guanti, R. Riva, Tetrahedron: Asymmetry 1995,G,2921;G. Guanti, R. Riva, Tetrahedron: Asymmetry2001,12,605. 35 G. Nicolosi, A. Patti, M. Piatelli, C. Sanfilippo, Tetrahedron: Asymmetry1995,6,519. 36 B. Danieli, G. Lesma, D. Passarella, A. Silvani, /. Org. Chem. 1998,63,3492. 37 R. ChCnevert, D. Goupil, Y. S . Rose, E. Bedard, Tetrahedron: Asymmetry1998,9,4285. 38 R. ChCnevert, G. M. Ziarini, M. P. Morin, M. Dasser, Tetrahedron: Asymmetry1999,10, 3117. 39 Y. Zhao, Y.Wu, P.De Clerq, M. Vandewalle, P. Maillos, 1.C.Pascal, Tetrahedron: Asymmetry

2000,11,3887. 40 M. Ranchoux, J.-M. Brunel, G. Iacazio, G. Buono, Tetrahedron: Asymmetry1998,9,581. 41 H. Konno, K. Ogasawara, Synthesis1999,1135. 42 T.Taniguchi, K. Ogasawara, Tetrahedron Lett. 1999, 40,4383. 43 C.Cinquin, 1. Schaper, G. Mandville, R. Bloch, Synlett, 1995,339.

486

I

1 1 Hydrolysis and Formation ofC-0 Bonds

pounds differ but not to a large extent (Tables 11.1-11and 11.1-18).In many cases the enantioselectivity of acylation is higher than that of the hydrolysis. Acylation of the three-, four- and five-membereddimethanol derivatives proceeds uniformly with the same enantiotopic group recognition to the monoacetates 1-8 with good to high enantioselectivity and yield. Acylation of the cyclohexanoid dimethanol system is erratic, giving 10 with low enantioselectivityand low yield. The cyclohexenoid system 11 however is obtained with the same lipase with good enantioselectivity.Acylation of cyclopentanoid dimethanol derivativeswith a functional group in %positionby pig pancreas lipase has been intensively investigated (4-8).The enantioselectivitycan be influenced (5 and 6) by the choice of the appropriate protecting group. The heterocyclic dimethanol monoacetate 9, which is a derivative of the parent compound meso-butane tetrol, is obtained with high enantioselectivity by Pseudomonas Jluorescens lipase instead of pig pancreas lipase. Acylation of meso-exo-oxa-norbornane dimethanol with pig pancreas lipase and with Candida cylindracea lipase provides access to both enantiomeric monoacetates 14 and 15.A further example of the attainment of both enantiomers by changing the lipase is provided by the acylation of 1,2-bis(hydroxymethyl)ferrocene with vinyl acetate catalyzed either by Pseudomonas cepacia lipase which gives the (S)-enantiomer 32 or by Chromobacterium viscosurn lipase which gives the (R)-enantiomer 33. Acylation catalyzed by lipases is, as in the case of the hydrolysis of the corresponding acetates, not restricted to substrates containing primary hydroxyl groups, as demonstrated by the successful synthesis of the monoacetates 17-19, 22-29, 31,34-36, 38,39, 41-44, 46,63-65. These examples give a good illustration of the scope of lipases as catalysts. Comparison of the bicyclo[3.l.0]cyclohexane derivatives 28 and 29 shows that changing the configuration of the cyclopropane ring is accompanied by a switch of enantiotopos-selectivity under identical reaction conditions. Lipases are the hydrolases of choice for the kinetic enantiomer separation of racemic primary, secondary and tertiary alcohols through acylation. Acylation of the racemic alcohols is complementary to the hydrolysis or alcoholysis of the corresponding esters. Monoacetates of Table 11.1-18 which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-3, 11.1-7, 11.1-9 and 11.1-11.

A large number of enantiomerically pure primary alcohols carrying additional nitrogen, oxygen and sulfur functionalities can be prepared by lipase-catalyzed enantiomer-differentiatingacylation with the usual acylating reagents (1-130)(Table 11.1-19).Most remarkably, a series of primary alcohols whose chiral center bears only alkyl or alkenyl groups (23-30)has been obtained with high enantioselectivity through Pseudomonas Jluorescens lipase-catalyzed acylation with vinyl acetate in dichloromethane. For the attainment of chiral primary alcohols, lipase-catalyzed acylation seems to be more efficient in terms of selectivity and yield than lipasecatalyzed hydrolysis of the corresponding esters. A comparison ofTables 11.1-19and 11.1-14 shows that enantiomer-differentiating hydrolysis of acetates and enantiomer-differentiating acylation of the corresponding alcohols catalyzed by one and the same lipase are complementary. Enantiomer-differentiatingacylation with succinic

7 1. I Hydrolysis and Formation of Carboxylid Acid Esters

I

487

Table 11.1-19. Lipase-catalyzed enantiomer-differentiating acylation of racemic acyclic primary alcohols in organic solvents (PPL pig pancreas lipase, PFL PseudomonasPuorescens lipase, PCL Pseudomonas cepacia lipase, CCL Candida cylindracea lipase, M M L Mucor miehei lipase, PSL Pseudomona sp. lipase, CAL-B Candida antarctica B lipase, CAL Candida antarctica lipase, not specified, CLL Candida lipolytica lipase, SM L Serratia marcensens lipase, HLL Humicola lanuginosa lipase).

NHCbz

NHCbz

RL O A c R Me Et

73 % ee, 78 % ee, 99%ee,95%ee,all PPL, ethyl acetate

n-Pr n-Bu

R&OH

la 2a 3a 4a

85 % ee, 83 % ee, 99 % ee, 95 % ee, -

Me

R

RA

-0Ac

R = PhS 98 % ee, R = PhS0298 % ee, -

O

H

5a Ga

98 % ee, 98 % ee, 60 % conversion

26 % ee, -, PCL, vinyl

7a

299 % ee, 37 % yield

acetate R = OMe 81 % ee, 43 %yield, PCL,acetic anhydride

7b I31

8a

83 % ee, 44 % yield

8b [41

9a [51

&OH

9b 151

all PCL,vinyl acetate

40 % conversion

u

5a PI Ga 121

Fi

R=H

OH -0COnPr

90 % ee, 40 % yield, CCL tributyrin

89 % ee, 36 % yield

NHC0,Et R&OH R = Me R = Et

90 % ee, 31 %yield 295 % ee, 31 %yield

all PPL,ethyl acetate

1Oa lla

295 % ee, 30 % yield 92 % ee, 32 % yield

lob [6] I l b [6]

488

I

I 7 Hydrolysis and Formation ofC-0 Bonds Table 11.1-19.

(cont.). ~~~~

R

0 3

.’,,,,, OCO(CH,),CO,H

, r O

60 % ee, 40 % yield 92 % ee, 41 % yield

R = Me R = Ph

~

R& fOH

12a 13a

61 % ee, 40 % yield 70 % ee, 32 % yield

12b [7] 13b [7]

all PFL, succinic anhydride

..**-OCO(CH,),CO,H

n’

,

14a [7]

I

14b [7]

npr”yo

npr/Nyo

0 75 % ee, 46 %yield, PFL

98 % ee, 38 % yield

0

succinic anhydride

n

15a [8] 0

0 R = t-Bu, i-Pr 295 % ee, 40-43 %yield, PFL acetic, propionic or butyric anhydride

295 % ee, 42-45 % yield

eoH

(\/OAc

R

15b [8]

R 4 f 0

R/NKO

R

‘0

R = CIO& R = (CH&CHMez

PPL ethyl acetate

1Ga 17a

Lon,

295 % ee, 31 % yield 295 % ee, 36 % yield

161, [9, 101 17b (9, 101

0

R

R = PhCH2 R = C9H19 R = Vinyl(CH2)3

298 % ee, 32 % yield 96 % ee,38 % yield 99 % ee, 38 % yield

all PFL, vinyl acetate 40 % conversion

RE 18a 19a 2Oa

O

H

298 % ee, 34 % yield 96 % ee, 36 % yield 98 % ee, 38 % yield 60 % conversion

18b [lla]

19b [lla] 20b (llb]

11.1 Hydrolysis and Formation of Carboxylid Acid Esters

I

489

Table 11.1-19.

kont.).

M

o

e

~

o

A

c

Meo OH

OH

21a [12]

94 % ee, 23 %yield, PSL isopropenyl acetate

22a [ 131

XZ:OCH,NHCOPh

97 % ee, 27 %yield

NHC0,Et A

O

H

21b [12]

22b 1131

63 % ee, -,

93 % ee, -, MML

Ph

2 R'

R2

n-Pr n-Bu n-Bu n-Hex (CHs)2CHCH2 n-Oct CH3CH=CHCH2 Ally1

Me Me Et Me Me Me Me Me

98 % ee, 17 % yield 99 % ee, 22 % yield 97 % ee, 23 % yield 96 % ee, 20 % yield 98 % ee, 26 % yield 98 % ee, 26 % yield 96 % ee, 33 % yield 97 % ee, 25 % yield

all PCL vinyl acetate. The corresponding acetates were of low ee.

23 [14] 24 [14] 25 [14] 26 [14] 27 [14] 28 [14] 29 [14] 30 [14]

OAc

boLoAc

.O\/\/OAc

R

H 2-Me 3-OMe 4-OMe 4-C1

4-t-Bu

79 % ee, 46 % yield 80 % ee, 48 % yleld 95 % ee, 47 % yield 94 % ee, 52 %yield 92 % ee, 49 % yield 93 % ee, 50 % yield all PCL, vinyl acetate

31a 32a 33a 34a 35a 36a

85 % ee, 48 % yield 93 % ee, 45 % yield 91 % ee, 49 % yield 96 % ee, 48 % yield 94 % ee, 48 % yield 99 % ee, 50 % yield

31b [15] 32b [16] 33b [15] 34b [15] 35b [15] 36b [15]

490

I

J J Hydrolysis and Formation ofC-0 Bonds Table 11.1-19.

(cont.).

B

9

R

82 % ee, 48 % yield 90 % ee, 46 % yield 84 % ee, 47 % yield 79 % ee, 40 % yield 90 % ee, 48 % yield all PSL, vinyl acetate

H Me i-Pr F OMe

04 .

.

O

A

297 % ee, 39 % yield 90 % ee, 48 % yield 294 % ee, 38 % yield 90 % ee, 33 % yield 298 % ee, 42 % yield

42a [18]

A,..' X

37a 38a 39a 40a 41a

37b [17] 38b [17] 39b [17] 40b [17] 41b [17]

421, [ 181

c

HO 86 % ee, 52 %yield, PCL vinyl acetate

99 % ee, 44 % yield

OAc R

A

O

A

c

R

Z

O

H

R

>98 % ee, 37 % yield >98 % ee, 39 % yield >98 % ee, 34 % yield all PFL, vinyl acetate

i-Pr t-Bu Ph

43a 44a 45a

57 % ee, 53 %yield 81 % ee, 52 %yield 67 % ee, 57 % yield

43b [19] 44b [19] 45b [I91

4Ga [20]

PhL

46b [20]

OAc Ph

&OAc

96 % ee, 48% yield, PCL, vinyl acetate

87 ee, 52% yield

47a [21]

F R = 4-OMeC6H4CH2 >99 % ee, 33 % yield, lipase OF (Meito Sangyo), vinyl acetate

O A c

47b [21]

I

11.1 Hydrolysis and Formation ofCarboxylid Acid Esten 491 Table 11.1-19.

(cont.).

,OH

OAc

48b [22]

OH 97 % ee, 20 % yield, PCL vinyl acetate

27 % ee, 76 %yield

97 % ee, 35 %yield, PCL vinyl acetate 93 % ee, 35 %yield, PSL vinyl acetate

62 % ee, 60 % yield >99 % ee, 41 %yield

OH 50a[24]

"

f

"OTBDMS

90 % ee, 51 %yield, CAL-B, isopropenyl acetate

PhE 1 0

A

0

2

Sob [24]

OTBDMS

-, 49 % yield

c

R

R = CH2Ph: >97 % ee, -, PSL vinyl acetate, 50 % conversion R = 4-MeCsH4CHz:94 % ee, -, PSL, vinyl acetate, 49 % conversion R = 4-MeC6H4CHz: 73 % ee, -, PSL, vinyl acetate, 58 % conversion R = CH2I: 94 % ee, -, PSL vinyl acetate, 43 % conversion

Ph 94 % ee, -, PSL, vinyl acetate 51 % conversion

51a

>97 % ee, -

51b 1251

52a

90 % ee, -

52b [25b]

53a

>99 % ee, -

53b [25b]

54a

70 % ee, -

54b [26]

55a [26]

+OH Ph >97 % ee, -

55b [26]

492

I

I 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-19.

(cont.).

'it

R2 R' = C1, R2= Ph: 95 % ee, -, PPL, vinyl acetate, 41 % conversion R' = I, R2= Ph: 96 % ee, -, PPL, vinyl acetate, 51 % conversion R' = I, R2= CH2-CH2Ph 55 % ee, -, PPL, vinyl acetate, 64 % conversion R' = I, R2= %Me3:94 % ee, -, PPL, vinyl acetate, 51 % conversion R' = I, R2= n-Bu: 89 % ee, -, PPL, vinyl acetate, 49 % conversion R' = I, R2= n-Hex: 51 % ee, -, PPL, vinyl acetate, 64 % conversion R' = I, R2= t-Bu: >97 % ee, -, PPL, vinyl acetate, 49 % conversion

56a

67 % ee, -

56b (261

57a

>97 % ee, -

57b [26]

58a

>97 % ee, -

58b [2G]

59a

>97 % ee, -

59b [26]

GOa

88 % ee, -

Gob [26]

Gla

93 % ee, -

Glb [26]

62a

94 % ee, -

621, [26]

R = 4-MePh: 72 % ee, 55 %yield, CRL, vinyl acetate R = Me: 62 % ee, -, PSL, vinyl acetate, 62 % conversion

63a

>99 % ee, 42 % yield

631, [27]

64a

96%ee,-

64b [28]

OH

OAc

O+i

OR R = Me: 90 % ee, 36 %yield, PCL, vinyl acetate R = Me: 90 % ee, 38 % yield, PSL, vinyl acetate R = MOM: 94 % ee, 40 % yield, PCL, vinyl acetate R = MOM: 97 % ee, 46 %yield, PSL, vinyl acetate

G5a

OR 69 % ee, 54 %yield

65b [29]

96 % ee, 45 % yield 661, [29]

66a

75 % ee, 51 %yield

99 % ee, 48 % yield

1 1 . 1 Hydrolysis and Formation ofCarboxylid Acid Esters Table 11.1-19.

\

(cont.).

...\'\ OAc

X = C1>95 % ee. 43 % yield, PCL, vinyl acetate X = F: >95 % ee, 44% yield, PCL, vinyl acetate X = H: >95 % ee, 39% yield, PCL, vinyl acetate

OAc

67a

>95 % ee, 37 % yield

6% (301

68a

295 % ee, 33 % yield

68b [30]

69a

>95 % ee, 33% yield

69b [30]

70a 1301 "-OAc -, 50% yield, PCL, vinyl acetate

0"" "

70b [30]

>95 % ee, 43% yield

71a [31]

N HBOC

71b [31] NHBoc 925 % ee, 38% yield

91 % ee, 45% yield, PCL, vinyl butyrate

72b [32]

72a [32] ( T O"OAc A c

>99 % ee, 48 % yield

90 % ee, 51 %yield, PCL, vinyl acetate

R k

R2

OKNdoAc 0 R'

R2

Ph

H

H

H

H

73 % ee, 47 % yield, PCL, vinyl propionate Ph 92 % ee, 43 %yield, PCL, vinyl propionate Ph 99 % ee, 44 % yield, PCL, vinyl propionate CHzPh 71 % ee, 52 %yield, PCL, vinyl propionate

73a

78 % ee, 42 % yield

73b [33]

74a

81 % ee, 48 % yield

7413 [33]

75a

74 % ee, 51 % yield

75b [33]

76a

87 % ee, 43 %yield

76b [33]

I

493

494

I

1 1 Hydrolysis and Formation o f C - 0 Bonds Table 11.1-19.

(cont.).

IFoAc

94 % ee, 38 %yield, PCL, vinyl acetate 89 % ee, 31 %yield, PSL, vinyl acetate

89 % ee, 40 % yield 91 % ee, 40 % yield

0

78a [35] mlOAc 99 % ee, 40 % yield, PCL, vinyl butyrate

COAc A3(4): 99 % ee, 34 % yield, PCL,vinyl butyrate A4(5):99 % ee, 34 %yield, PCL,vinyl butyrate

96 % ee, 47 % yield, PCL, succinic anhydride

51 % ee, 61 %yield, PSL, vinyl acetate

78b [35] C O H 97 % ee, 36 % yield

79a

94 % ee, 32 % yield

79b [35]

80a

97 % ee, 25 % yield

80b [35]

81a [36] -, 42 %yield

>99 % ee, 36 % yield

81b [36]

1 1 . 1 Hydrolysis and Formation of Carboxylid Acid Esters

I

495

Table 11.1-19.

(cont.).

I

I

R Me CH2Ph

92 % ee, -, CRL, vinyl acetate, 59 % conversion 96 % ee, -, CRL, vinyl 50 % conversion

83a

-, -

84a

-. -

>97 % ee, 52 % yield, CLL, Ac2O 88 % ee, 49 % yield, CLL, Ac~O >95 % ee, 46 %yield, olipase 4SD (Amano),AczO 73 % ee, 50 % yield, CLL, AczO

83b [38] 84b [38]

85a

-, 22 %yield

85b [39]

8Ga

-, 25 %yield

8Gb [39]

87a

-, 34 % yield

8% [39]

88a

-, 38% yield

88b [39]

LOAC 75 % ee, 55 %yield, PFL, vinyl acetate 98 % ee, -, PFL, vinyl acetate, 39 % conversion

R

A

O

A

89a

96 % ee, 42 % yield

89b [40]

89a

>99 % ee, -, 57 % conversion

89b [41]

c

R:

a

b

R = a: 97 % ee, -, PCL, vinyl acetate, 39 % conversion R = b: 95 % ee, -, PCL, vinyl acetate, 38 % conversion R = c: 97 % ee, -, PCL, vinyl acetate, 31 % conversion R = d 94 % ee, -, PCL, vinyl acetate, 41 % conversion

d

C

90a

_-

90b [42]

91a

-_

91b [42]

92a

__

92b [42]

93a

_ -

93b (421

496

I

I 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-19.

(cont.).

Boc

Boc I

I

W

O

A

C

-, -, PCL, vinyl acetate 58 % conversion

OH

O , Ac

YBn OAc

95a

88 % ee, 50 % yield, PCL, vinyl acetate 80 % ee, 21 %yield, PPL, vinyl acetate

R

\

&N--Bn \

95c

OH

78 % ee, 21 %yield

>99 % ee, 29 % yield

[&a]

83 % ee, 26 %yield, for ent95b

>99 % ee, 53 %yield

[&b, c]

24 % ee, 64 % yield

OAc R

0

>99 % ee, -, CAL, vinyl acetate, 32 % conversion 4-Br-Ph 98 % ee, -, CAL, vinyl acetate, 38 % conversion Me 91 % ee, -, PPL, vinyl acetate, 29 % conversion Ph 82 % ee, -, CAL, vinyl acetate, 54 % conversion

Ph

NMe

95b

OAc

X

0

OH

d - B n

91 % ee, 18 %yield, PCL, vinyl butyrate

0

96 % ee, -

97a

46 % ee, -

9% [46]

98a

59 % ee, -

981,1461

99a

38 % ee, -

99b [46]

lOOa

>98 % ee, -

1OOb [4G]

N lOla [47] Ph&OAc 96 % ee, 38 % yield, PCL, vinyl acetate, -50 "C

"US 0

'..

',-OAc 62 % ee, 55 %yield, PFL, vinyl acetate

102a[48]

+ O ,.,H Ph 62 % ee, 60 % yield

101b [47]

102b [48]

0 OH

94 % ee, 40 % yield

7 7.7 Hydrolysis and Formation of Carboylid Acid Esters Table 11.1-19.

(cont.).

R

Me CHzPh

73 % ee, -, CAL-B, vinyl acetate, 57 % conversion 81 % ee, -, CAL-B, vinyl acetate, 57 % conversion

Me PhA O A c 95 % ee, 26 %yield, PCL, vinyl acetate

103a

92 % ee, -

103b [49]

104a

99 % ee, -

104b [49]

105a [SO] Me
105b [SO]

Ho-)\,,

10Gb 1511

1OGa [511

HZ!3C,, COzMe 54 % ee, 52 %

83 % ee, 25 %yield, PFL, vinyl acetate OAc R O A O A c

R O L O A c

R C16H33

107a

94 % ee, -

10% [52]

108a

89 % ee, -

108b 1521

109a

93 % ee, -

109b [52]

80 % ee, -, PSL, vinyl acetate, 55 % conversion, 4 "C 30 % ee, -, PSL, vinyl acetate, 51 % conversion, 4 "C 24 % ee, -, PSL, vinyl acetate, 61 % conversion, 22 "C

C18H37 (9Z)-C&35

A c O & ~ ~ ~

11Oa [531

OH A C O + ~ ~ ~ N3

N3

91 % ee, -, PCL, vinyl acetate, 50 % conversion

1lOb [53]

>99 % ee, -

111b [54]

89 % ee, 39 % yield, PFL, vinyl acetate 86% ee, 44 % yield, PCL, vinyl acetate

80 % ee, 44 % yield 92 % ee, 42 % yield

I

497

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-19.

(cont.).

-OH

a 112 1551

O

H

&OH \

ent-112 [55] \

acylated enantiomers not shown >99 % ee, 45 % yield, HLL, vinyl hexanoate

acylated enantiomers not shown >99 % ee, 29 %yield, SML, vinyl hexanoate

(CH,),-OAc

-h P

’ /

HO-,(H,C) absolute configuration unknown n = 2: 113a 90 % ee, 41 % yield, CAL-B, vinyl acetate n = 3: 114a >99 % ee, 9 % yield, PSL, vinyl acetate

-h P

I

(CH,),-OAc

Ph*

AcO-,(H,C)

HO-,(H,C) absolute configuration unknown 113b 35 % ee, 27 %yield

absolute configuration unknown 113c [SG] 99 % ee, 32 % yield

114b >98 % ee, 18 % yield

114c (561 30 % ee, 73 % yield

I--\

HOH,C

CH,OH

(P)-115a [57] 98 % ee, 45 % yield, PCL, vinyl acetate (M)-115a [57] 92 % ee, 44 % yield, CAL-B, vinyl acetate

(M)-115b (571 80 % ee, 38 % yield

(M)-115~[57] 95 % ee, 13 % yield

1 1.1 Hydrolysis and Formation of Carboxylid Acid Esters

I

499

Table 11.1-19.

(cont.)

~ C H , O A C

& R

Me

Ph

t-Bu

81 % ee, -, MML, vinyl acetate, 46 % conversion 84 % ee, 47, CAL-B, vinyl acetate, 46 % conversion 88 % ee, -, CAL-B, vinyl propionate, 45 % conversion 90 % ee, -, MML, vinyl acetate, 35 % conversion

llGa

81 % ee, -

l l 6 b [58]

83 % ee, 47 % yield

117a

72 % ee, -

11% [58b]

118a

48 % ee, -

118b [58b]

119a[591

Fe

CH,OH

&

& '

89 % ee, -, CAL-B, vinyl acetate, 52 % conversion

96 % ee, -

92 % ee, 49 % yield, PCL, vinyl acetate

90 % ee, 51 %yield

121a [61] >97 % ee, 22 % yield, PCL, vinyl acetate

121b [61] 10 % ee, 55 %yield

121c [GI] >97 % ee, 18 % yield

122b [61]

122a [61] *OH

(oz> OAc >95 % ee, 42 %yield, PCL, vinyl acetate

119b (591

58 % ee, 59 % yield

500

I

1 7 Hydrolysis and Formation of C - 0 Bonds

Table 11.1-19. (cont.).

80 % ee, 30 % yield

75 % ee, 35 %yield, PCL, isopropenyl acetate The two further regioisomers show lower selectivity.

,OH /OAC

OAc

56 % ee, -

94 % ee, -, PCL, vinyl acetate, 38 % conversion 97 % ee, -, PFL, vinyl acetate, 38 % conversion

60 % ee, -

125a[64]

OH

125b [64]

LOA~

-0Ac

91 % ee, 44 %yield, CAL-B, vinyl acetate

91 % ee, 44 %yield

4

126b [65]

nPrOCO BocHN

99 % ee, 36 % yield

89 % ee, 51 %yield, CAL-B, vinyl butyrate

A

4

127b [65]

127a [65]

nPrOCO BOCHN'

&OH

95 % ee, 18 %yield, PPL, vinyl butyrate. 37 % conversion

96 % ee, 43 % yield, 5 3 % conversion

NHBoc

128b [65]

128a [65] 95 % ee, 40 %yield, PPL, vinyl butyrate, 44 % conversion

99 % ee, 36 % yield, 56 % conversion

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters

Table 11.1-19.

(cont.).

92 % ee, 22 % yield, PPL, vinyl butyrate, 30 % conversion

90 % ee, 32 % yield, 53 % conversion

83 % ee, 44 % yield, PCL, vinyl acetate

95 % ee, 34 % yield

1 S. Fembndez, R. Brieva, F. Rebolledo, V. Gotor, /. Chem. Soc., Perkin Trans. 1 1992, 2885. 2 P. Ferraboschi, P. Grisenti, A. Manzocchi, E. Santaniello,]. Org. Chem. 1990,55, 6214. 3 S. Antus, A. Gottsegen, J. Kajtir, T. Kovbcs,T. S. Toth, H. Wagner, Tetrahedron: Asymmetry1993,4, 339. 4 M. D. Ennis, D. W. Old, Tetrahedron Lett. 1992, 33, 6283. 5 B. Cambou, A. M. Klibanov,/. Am. Chem. Soc. 1984,106,2687. 6 F. Francalanci, P. Cesti, W. Cabri, D. Bianchi, T. Martinengo, M. Foi,]. Org. Chem. 1987,52,5079. 7 Y Terao, K. Tsuji, M. Murata, K. Achiwa, T. Nishio, N. Watanabe, K. Seto, Chem. Pharm. Bull. 1989, 37, 1653. 8 D. Bianchi, P. Cesti, E. Battistel,]. Org. Chem. 1988,53,5531. 9 D. Bianchi, W. Cabri, P. Cesti, F. Francalanci, F. Rarna, Tetrahedron Lett. 1988,29, 2455. 10 L. T. Kanerva, E. Vanttinen, Tetrahedron: Asymmetry1993,4,85. 11 a) P. Ferraboschi, D. Brernbilla, P. Grisenti, E. Santaniello;]. Org. Chem. 1991, 56, 5478 b) P. Ferraboschi, S. Casati, P. Grisenti, E. Santaniello, Tetrahedron: Asymmetry1993,4, 9. 12 H. Akita, 1. Umezawa, M. Nozawa, S. Nagumo, Tetrahedron: Asymmetry1993,4, 757. 13 H. S. Bevinakatti, R. V. Newadkar, Tetrahedron: Asymmetry1993,4,773. 14 S. Barth, F. Effenberger, Tetrahedron: Asymmetry 1993,4, 823. 15 a) F. Theil, J. Weidner, S. Ballschuh, A. Kunath, H. Schick,/. Org. Chem. 1994.59, 388; b) F. Theil, J. Weidner, S. Ballschuh, A. Kunath, H. Schick, Tetrahedron Lett. 1993, 34, 305. 16 F. Theil, S. Ballschuh, A. Kunath, H. Schick, Tetrahedron: Asymmetry1991,2, 1031. 17 B. Herradbn, S. Cueto, A. Morcuende, S. Valverde, Tetrahedron: Asymmetry1993,4, 845.

18 J. Weidner, F. Theil, A. Kunath, H. Schick, Liebigs Ann. Chem. 1991,1301. 19 G . Egri, E. Baitz-Gasc,L. Poppe, Tetrahedron: Asymmetry1996,7,1437. 20 K. Lemke, F. Theil, A. Kunath, H. Schick, Tetrahedron: Asymmetry1996,7, 971. 21 V. Khlebnikov, K. Mori, K. Terashirna, Y. Tanaka, M. Sato, Chem. Pharm. BULL. 1995,43,1659. 22 H. Tanimoto, T. Oritani, Tetrahedron: Asymmetry 1996,7, 1695. 23 T. Yokomatsu, N. Nakabayashi, K. Matsurnoto, S. Shibuya, Tetrahedron: Asymmetry1995,6, 3055. 24 C. R. Johnson, Y. Xu, K. C. Nicolaou, 2. Yang, R. K. Guy, J. G . Dong, N. Berova, Tetrahedron Lett. 1995,36,3291. 25 a) R. P. Hof, R. M. Kellogg, Tetrahedron: Asymmetry 1994,5565: b) R. P. Hof, R. M. Kellogg,]. Chem. Soc., Perkin Trans. 1, 1996, 2051. 26 S:T. Chen, J:M. Fang,J. Org. Chem. 1997, 62, 4349. 27 G . Laval, G. Audran, S. Sanchez, H. Monti, Tetrahedron: Asymmetry1999, 10, 1927. 28 G. Laval, G . Cadillio, H. Monti, A. Tolomelli, G . Audran, J:M. Galano, Tetrahedron: &mmetry 2000,II. 1289. 29 H. Miyaoka, Y. Kajiwara, M. Hara, A. Suma, Y.Yamada, Tetrahedron: Asymmetry1999, 10, 3189. 30 K. Miyazawa, D. S . Yufit, j. A. K. Howard, A. de Meijere, Eur. ]. Org. Chem. 2000,4109. 31 M. Peter, J. Van der Eycken, G . Bernith, F. Fiilop, Tetrahedron: Asymmetry1998,9,2339. 32 J. Weidner, F. Theil, H. Schick, Tetrahedron: Asymmetry1994,5,751. 33 H. Wakarnatsu, Y. Terao, Chem. Pharm. Bull. 1996,44,261. 34 H. Nakano, Y. Okuyama, K. Iwasa, H. Hongo, Tetrahedron: Asymmetry1994, 5, 1155. 35 J. K h a n , E. Forr6, F. Fiilop, Tetrahedron: Asymmetry2000,I1,1593.

I

501

502

I

1 7 Hydrolysis and Formation ofC-0 Bonds 36 J. A. Hyatt, C. Skelton, Tetrahedron: Asymmetry 1997,8, 523. 37 E. Mizuguchi, T. Suzuki, K. Achiwa, Synlett 1994, 929. 38 T. A. Ayers, R. A. Schnettler, G. Marciniak, K.T. Stewart, R. K. Mishra, D. J. Krysan, B. R. Bernas, P. Bhardwaj, T. L. Fevig, Tetrahedron: Asymmetry 1997, 8, 45. 39 K. Mizuno, S. Sakuda, T. Nihira, Y. Yarnada, Tetrahedron 1994,50,10849. 40 F. Bracher, T. Papke, Tetrahedron: Asymmetry 1994, 5, 1653. 41 0. Nordin, E. Hedenstrorn, H.-E. Hogberg, Tetrahedron:Asymmetry 1994, 5, 785. 42 0. Nordin, B.-V. Nguyen, C. Vorde, E. Hedenstrorn, H:E. Hogberg, ]. Chem. Soc., Perkin Trans. 1, 2000, 367. 43 F. Schieweck, H:J. Altenbach, Tetrahedron: Asymmetry 1998,9,403. 44 a) M. P. Sibi, J. L. Lu, Tetrahedron Lett. 1994, 35, 4915; b) G . Guanti, R. Riva, Tetrahedron: Asymmetry 1995, 6, 2921; c) G. Guanti, R. Riva, Tetrahedron: Asymmetry 2001, 12,605. The given absolute configurations in b and c are contradictory. 45 M. Lemaire, J. Bolte, Tetrahedron: Asymmetry 1999, 10,4755. 46 R. P. Hof, R. M. Kellogg,]. Org. Chem. 1996, 61, 3423. 47 T. Sakai, I. Kawabata, T. Kishimoto, T. Ema, M. Utaka, j . Org. Chem. 1997,62,4906. 48 M. Pallavicini, E. Valoti, L. Villa, 0. Piccolo, J. Org. Chem. 1994,59,1751. 49 H.-J. Gais, I. von der Weiden, Tetrahedron: Asymmetry 1996,7,1253. 50 0. Goj, A. Burchardt, G. Haufe, Tetrahedron: Asymmetry 1997,8,399. 51 0. J i m hez , M. P. Bosch, A. Guerrero,]. Org. Chem. 1997,62,3496.

52 G. G. Haraldsson, P. Thordarson, A. Halldorsson, B. Kristinsson, Tetrahedron: Asymmetry 1999, 10, 3671. 53 G . Iacazio, D. Martini, S. Sanchez, B. Faure, Tetrahedron: Asymmetry 2000,II, 1313. 54 P. Kielbsinski, J. Ornelanczuk, M. Mikolajczyk, Tetrahedron: Asymmetry 1998, 9, 3283. 55 T. Furutani, M. Hatsuda, R. Imashiro, M. Seki, Tetrahedron:Asymmetry 1999, 10,4763. 56 F. Theil, H. Sonnenschein, T. Kreher, Tetrahedron: Asymmetry 1996,7,3365. 57 a) K. Tanaka, Y. Shogase, H. Osuga, H. Suzuki, K. Nakamura, Tetrahedron Lett. 1995, 36, 1675; b) K. Tanaka, H. Osuga, H. Suzuki, Y. Shogase, Y. Kitahara,j . Chem. Soc., Perkin Trans. 1 1998, 935. 58 a) D. Lambusta, G. Nicolosi, A. Patti, M. Piattelli, Tetrahedron Lett. 1996, 37, 127; b) A. Patti, D. Lambusta, M. PiatteUi, G. Nicolosi, P. McArdle, D. Cunningham, M. Welsh, Tetrahedron 1997,53, 1361. 59 A. Patti, D. Larnbusta, M. Piattelli, G. Nicolosi, Tetrahedron:Asymmetry 1998,9, 3073. GO A. Patti, G. Nicolosi, Tetrahedron:Asymmetry 1999, 10, 2651. 61 G. Nicolosi, A. Patti, M. Piatelli, ].Org. Chem. 1994,59, 251. 62 J. H. Rigby, P. Sugathapala, Tetrahedron Lett. 1996, 37, 5293. 63 Y. Kawanarni, N. lizuna, Ke. Maekawa, Ky. Maekawa, N. Takahashi, T Kawada, Tetrahedron 2001,57, 349. 64 1. Izquierdo, M. T. Plaza, M. Rodriguz, J. A. Tarnayo, A. Martos, Tetrahedron: Asymmetry 2001, 12, 293. 65 J. KAmAn, J. Van der Eycken, A. Peter, F. Fiilop, Tetrahedron:Asymmetry 2001, 12,625. 66 E. Forr6. J. Arva, F. Fiilop, Tetrahedron: Asymmetry 2001, 12, 643.

acid anhydride (12-14) instead of the usually employed vinyl acetate may facilitate the separation of the remaining substrate and the ester formed because of the carboxyl group in the latter. Mixed primary secondary diols are very often separated into their enantiomers in a sequential two-step acylation wherein the first step - acylation of the primary hydroxyl group - shows high regio- but very poor enantioselectivity. The useful enantiomer-differentiating step is realized in the second step by acylation of the already monoacylated diol (31-36, 46, 72, 107-110, 120,125).On the other hand, as expected, mixed primary tertiary diols are not acylated at the tertiary hydroxyl group (47, 48, 51-54, 5 6 6 2 , 106). Regarding the structural diversity of the primary alcohols, which have been successfully resolved, there seems to be almost no limitation, as demonstrated by the axial-chiral diols 112-114, the ferrocene alcohols 116122 and the chiral chromium carbonyl complex 123, and most remarkably the helicenediolll5 (Table 11.1-19).

11.1 Hydrolysis and Formation ofCar6oxylid Acid Esters Table 11.120. Lipase-catalyzed enantiomer-differentiating acylation o f racemic acyclic secondary alcohols in organic solvents (PSL Pseudomonas sp. lipase, PPL pig pancreas lipase, PFL PseudomonasPuorescens lipase, PCL Pseudomonas cepacia lipase, CCL Candida cylindracea lipase, CAL-B Candida antarctica B lipase, CAL-A Candida antarctica A lipase, LIP Pseudomonas sp. lipase-Toyobo, BSL Burkholderia sp. lipase, GLL goat liver lipase, RML Rhizomucor sp. lipase, CRL Candida rugosa lipase).

1 [ l , 21

OAc R*CN R

ee (“A)

Yield (“A)

89

80

96

84

84

83

91

64

91

81

85

88

70

70

Amberlite IRA-904, aldehyde, acetone cyanohydrin, all PSL, vinyl acetate (dynamic kinetic resolution)

I

503

504

I

I I Hydrolysis and Formation of C - 0 Bonds

4

(cont.).

Table 11.120.

2a [31

Me

EtCO;""'

2

2b [31

Me

295% ee, -, PPL, EtCOzMe

35 % ee, -

3b [31

3a [31 OH

OCOEt

298% ee, -, PPL, EtCOzMe

18% ee, -

OCOR2

R

'

~

M

R'Me '

~

R'

R2

Et

n-Pr

4a

89 % ee, 35 % yield

4b [4, 51

5a

90 % ee, 30 %yield

5b [6]

5a

95 % ee, 38 %yield

5b [7]

Ga

95 % ee, 44 % yield

Gb [6]

7a

299 % ee, 43 %yield

7b [6]

8a

298%ee,-

81, [71

80 % ee, 38 % yield, PPL, F~CCHZOCOCIIHZ~

9a

297 % ee, 43 %yield

9b [8]

n-PR

87 % ee, 31 %yield, PPL, C13CCHzOCOnPr

10a

92 % ee, 26 %yield

l o b [6]

Ph

Me

lla

93 % ee, 41 % yield

l l b [9]

Ph

Me

lla

295 % ee, 43 %yield

11b [lo]

PhCHz

Me

12a

66 % ee, 43 %yield

12b [9]

PhCH2

Et

13a

92 % ee, 43 % yield

13b [lo]

1-Naphthyl

n-Pr

14a

295 % ee, 46 %yield

14b [ll]

2-Naphthyl

Me

299 % ee, 45 %yield, PSL, vinyl acetate 295 % ee, 39 %yield, PCL, acetic anhydride 299 % ee, 30 % yield, PSL, vinyl acetate 295 % ee, 39 % yield, PCL, propionic anhydride 95 % ee, 47 %yield, PPL, C13CCHZOCOnPr 299 % ee, 41 %yield, PSL, vinyl acetate

15a

95 % ee, 48 %yield

15b [9]

93% ee, 38% yield, CCL, tributyrin 95 % ee, 35 % yield, PPL, C13CCHzOCOnPr 92 % ee, 41 % yield, CCL, tributyrin 299 % ee, 44 % yield, PPL, C13CCHzOCOPr 98 % ee, 42 %yield, PPL, Cl3CCHZ0COPr -, -, PPL, vinyl acetate

YCH* YCH* Me

C11H23

Me

I 1.1 Hydrolysis and Formation of Carboxylid Acid Esten

I

505

Table 11.1-20.

(cont.).

OAc 1Ga [9, 101

&Me Ph

292 % ee, 43 % yield

295 % ee, 39 % yield, PCL, acetic anhydride

R' n-BuO n-BuO n-Bu PhCH2CH2

1Gb [9, 101 Ph

RZ Me Pr Me Me

295 % ee, 41 % yield 295 % ee, 23 %yield 295 % ee, 23 %yield 295 % ee, 41 %yield all PCL, vinyl acetate

17 [12] 18 [12] 19 [12] 20 [12]

NTBr 21 [13]

B; 97 % ee, -, PSL, C F ~ C H ~ O C O C I I H Z ~

R R n-C6Hls Ph c-C6HI1

297 % ee, 97 % ee, 95 % ee, all CAL-B, C7H1sCOSEt

22a 23a 24a

1

Me

298%ee,98%ee,298%ee,-

22b [14] 23b [14] 24b [14]

OH R LC02Me R Me Et n-Pr

R-C02Me

91 % ee, 39 %yield 295 % ee, 45 % yield

74 % ee, 54 % yield all PSL, isopropenyl acetate

25a 26a 27a

295 % ee, 38 % yield 295 % ee, 44 % yield 295 % ee, 42 % yield

251, [15] 26b [15] 27b [15]

OAc R -CO,Me

RLCO,Me

R

i-Pr 19 % ee, 43 % yield Me2ThexSi- 295 % ee, 35 %yield O(CH2)z all PSL, isopropenyl acetate

28a 29a

295 % ee, 38 % yield 72 % ee, 57 % yield

28b [16] 29b [16]

506

I

7 J Hydrolysis and Formation ofC-0 Bonds Table 11.1-20.

(cont.).

OCOCI, PhO-NHPh

OH

30a [17]

68 % ee, 26 % yield, PPL, (cc13c0)20

PhO

NHPh

301, [17]

96 % ee, 28 % yield

MeR -

OH

OAc R

80 % ee, 34 % yield 295 % ee, 25 % yield 295 % ee, 30 % yield all PSL,vinyl acetate

SiMeJ SiMeztBu SiMezPh

34a [19]

OAc O

A

C

295 % ee, 38 % yield 295 % ee, 44 % yield 295 % ee, 42 % yield

31a 3h 33a

34b [19]

I

295 % ee, 47 % yield

295 % ee, 45 %yield, PCL, vinyl acetate or acetic anhydride OAc

OH

35a [20] C,H,~/YO,H 77 % ee, 55 %yield, PCL, vinyl acetate

ACO,H CzzH45 299 % ee, 45 % yield

OCOnBu

R' n S n ( R z ) ) ,

R'

RZ

Me Et n-Pr Me Et

Me Me Me Et Et

97 % ee, 31 %yield 99 % ee, 36 % yield 97 % ee, 7 % yield 99 % ee, 35 % yield 97 % ee, 14 %yield all PPL, n-BuCOzCH2CF3

4

36a 37a 38a 39a 40a

71 % ee, 42 %yield 56 % ee, 36 % peld 68 % ee, 7 % yield 51 % ee, 47 %yield 57 % ee, 14 %yield

36b [21] 371, [21] 38b [21] 39b [21] 40b [21]

OH

41a [22]

Ph 295 % ee, 47 %yield, PSL, vinyl acetate

42a 1221 p

35b [20]

OH

R'ASn(R2),

MeL

31b [18] 32b [18] 33b [18]

h

295 % ee, 50 %yield, PSL, vinyl acetate

v

41b [22]

Ph 295 % ee, 32 % yield OH

Me-Ph

295 % ee, 47 % yield

42b [22]

1 1.1 Hydrolysis and Formation of Carboylid Acid Esters Table 11.1-20.

(cont.). ~

UHi7

43b [22]

43a [22]

Me

Me

31 % ee, 46 %yield, PSL, vinyl acetate

56 % ee, 35 % yield

OH

44b [22]

44a [22]

Me=C8H,,

Me

33 % ee, 63 %yield, PSL, vinyl acetate

CP,,

295 % ee, 22 % yield

OH

d

p

45a [22]

h

46 % ee, 32 %yield, PSL, vinyl acetate

&

Ph 295 % ee, 49 %yield, PSL, vinyl acetate

45b [22] w

p

h

295 % ee, 33 % yield

OH

4Ga [22]

P-h 295 % ee, 41 % yield

46b [22]

4% [22]

47a [22] Ph 68 % ee, 44 %yield, PSL, vinyl acetate

q

295 % ee, 28 % yield

48a [22]

Ph 295 % ee, 42 %yield, PSL, vinyl acetate

48b [22]

Ph 295 % ee, 43 % yield

49a [22] 81 % ee, 47 %yield, PSL, vinyl acetate

Me&.)

\

-

*.

OH

'

Ph

295 % ee, 38 % yield

50b [22]

50a [22] SiMe,

295 % ee, 38 %yield, PSL, vinyl acetate

49b [22]

SiMe, 81 % ee, 31 %yield

I

507

508

I

7 7 Hydrolysis and Formation of C - 0 Bonds

Table 11.1-20.

(cont.).

OH

R

‘

R’

q R2 R2

Me Me Me Me Me

Ph n-Bu n-CSH17 SiMej CH20CH2CGH40Me Et n-Bu C(Me)=CH2 C(Me)=CH2

295 % ee, 48 % yield 87 % ee, 41 % yield 295 % ee, 50 %yield 295 % ee, 45 % yield 78 % ee, 48 %yield

51a 52a 53a 54a

82 % ee, 36 % yield 295 % ee, 47 %yield all PSL, vinyl acetate

55a

295 % ee, 47 %yield 295 % ee, 31 %yield 295 % ee, 30 %yield 295 % ee, 27 % yield 295 % ee, 46 %yield

51b [22] 52b [22] 53b [22] 54b [22] 55b [22]

5Ga 57a

295 % ee, 46 % yield 295 % ee, 36 % yield

5Gb 1221 5% [22]

OH

R

CHZPh n-CsHlt

R’

77 % ee, 52 % yield -, 49 %yield all PSL, vinyl acetate

58a 59a

58b [22] 59b [22]

95 % ee, 31 % yield 23 % ee, 24 % yield

PR2

R’

R2

Me Et Me Et

SiMe3 Ph Et Et

295 % ee, 20 % yield 50 % ee, 63 % yield 295 % ee, 44 % yield 38 % ee, 33 % yield all PSL, vinyl acetate

\dPh

GOa Gla

295 % ee, 26 % yield 295 % ee, 33 %yield 54 % ee, 21 %yield 61 % ee, 27 %yield

62a G3a

OH

G4b [22]

G4a [22]

Ph 295 % ee, 34 % yield

72 % ee, 54 % yield, PSL, vinyl acetate

< \OCH ,O , Me

[231

299 % ee, 47 % yield, PSL, vinyl acetate OCOnPr Ph02S\j\Me

65 % ee, 20 % yield, PPL, CC13CH20COnPr

OH \ L O C 6 H 4 0 M e

G5b [23]

97 % ee, 49 % yield

OH

GGa [24]

P h 0 2 S dMe

95 % ee, 25 % yield

GGb [24]

Gob (221 61b [22] G2b [22] G3b [22]

11.1 Hydrolysis and Formation ofcarboxylid Acid Esters

I

509

Table 11.120.

(cont.).

OAc

Ph0,S

PhOS , -R

A \ I R

R

Me 295 % ee, 47 % yield Et 295 % ee, 49 % yield i-Pr 295 % ee, 48 % yield n-C6H13 295 % ee, 47 % yield PhCHZO(CH2)Z295 % ee, 47 %yield all PPL, vinyl acetate

298 % ee, 46 % yield 298 % ee, 49 % yield 94 % ee, 46 % yield 298 % ee, 48 % yield 88 % ee, 48 % yield

67a 68a 69a 70a 71a

6% [25, 261 68b 125, 261 69b [25, 261 70b (25, 261 71b [25, 261

0C0nC1,H3j

Tso L

T s O A O R

O

R

R

n-Cl6H33 n-CloH21 n-Bu

295 % ee, 45 % yield 295 % ee, 43 % yield 295 % ee, 43 % yield all PSL, (C1~H31C0)20

72a 73a 74a

295 % ee, 43 % yield 294 % ee, 42 % yield 296 % ee, 45 % yield

72b [27] 73b [27] 74b [27]

OH I

R = H, F, C1, Br, OMe 299 % ee, all PSL, vinyl acetate

RE O C P h ,

R

OH &OCPh,

R

ClCHz Me Et n-Pr

298 % ee, 43 % yield 295 % ee, 37 % yield 298 % ee, 43 % yield 70 % ee, 44 % yield all PCL, vinyl acetate

76a 77a 78a 79a

72 % ee, 54 % yield 78 % ee, 40 % yield 298 % ee, 48 % yield 56 % ee, 52 % yield

76b [29] 77b [29] 78b [29] 79b [29]

OAc

DoAcl 80a (301

R R R = H, Me, OMe, Allyl, c-CsHI1, CHZCN,NOz, 0-Ally1 83-87 % ee, 36-53 % yield 86-96 % ee, 38-48 % yield PSL, vinyl acetate OH

81 [31] PhACO,nBu

299 % ee, 43 %yield, PCL, vinyl acetate

80b [30]

510

I

I I Hydrolysis and Formation ofC-0 Bonds Table 11.1-20.

(cont.).

OAc 82a [32]

82b [32]

QMe 6-41 295 % ee, 41-50 %yield, PSL, vinyl acetate

295 % ee, 33-38 % yield

297 % ee, 23 % yield 297 % ee, 40 % yield 297 % ee, 41 % yield 92 % ee, 15 % yield 297 % ee, 32 % yield 97 % ee, 47 % yield all PSL, vinyl acetate

,

83a 84a 85a 8Ga 87a

88a

&

OAc i

89a

B~~NH88 % ee, 49 % yield, PCL, vinyl acetate

R

n-C7H15 n-ClsH31 i-Pr

Ph

83b [33] 84b [33] 85b (331 86b [33] 87b [33] 88b [33]

89b [34]

BocNH 94 % ee, 43 % yield

OH

OAc PhO&OCOR

Me

297 % ee, 38 %yield 74 % ee, 33 %yield 297 % ee, 34 % yield 18 % ee, 56 % yield 297 % ee, 30 %yield 85 % ee, 267 %yield

PhO&OCOR

98 % ee, 49 % yield, PCL, vinyl acetate 92 % ee, 45 %yield, PCL, vinyl acetate 98 % ee, 48 %yield, PCL, vinyl acetate 96 % ee, 42 % yield, PCL, vinyl acetate 95 % ee, 17 %yield, PCL, vinyl acetate

90a

93 % ee, 51 % yield

90b [35]

91a

82 % ee, 49 % yield

91b [35]

92a

89 % ee, 52 % yield

92b 1351

93a

52 % ee, 58 % yield

93b [35]

94a

19 % ee, 78 %yield

941, [35]

7 7 . 7 Hydrolysis and Formation ofCar6oxylid Acid Esters

(cont.).

Table 11.1-20.

OAc

R " O A C I

RnO&CI R2 = Naphthyl

R3 =

Ac

/

R4 =

+(CH,),OMe

R5= O N H n P r

>98 % ee, 45 %yield, PSL, vinyl acetate 95 % ee, -, CAL-B, vinyl acetate 95 % ee, -, CAL-B, vinyl acetate 95 % ee, -, CAL-B, vinyl acetate 95 % ee, -, CAL-B, vinyl acetate

R'

RZ R' R4 R5

cy

95a

91 % ee, 50 % yield

95b [36]

96a

27 % ee, -

961, [37]

97a

18 % ee, -

9% (371

98a

20% ee, -

98b [37]

99a

28 % ee, -

99b [37]

OAc

R( y 0 L C O * M e

O&C02Me

R

4-OMe

99 ee, -, PCL, vinyl acetate, 49 % conversion 99 ee, -, PCL, vinyl acetate, 49 % conversion 99 ee, -, PCL, vinyl acetate, 33 % conversion

2-Ally1 2,3-C&4

R

99ee,99 ee, -

lOOa

1OOb (381 101b [38]

lOla

49 ee, 102b [38]

102a

R y o T s

F O T s OAC

OH

R

CH=CH2

96 % ee, -, PCL, vinyl acetate 95 % ee, 48 % yield, PCL, vinyl acetate 93 % ee, -, PCL, vinyl acetate 92 % ee, -, PCL, vinyl acetate 80 % ee, -, PCL, vinyl acetate

Me CHzCl Et

T

O

B

107a

99%ee,>99 % ee, 98%ee,-

104a 105a lO6a

m

n

OAc 79 % ee.45 %yield, PCL, vinyl acetate

103a 98%ee,enf-103a 84 % ee, 49 % yield

O

OH

B

103b [39] enf-103b [40] 104b [39] 105b [39] lO6b [39]

n

70 % ee, 55 % yield

10% [40]

I

511

512

I

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.120.

(cont.).

P

y

h

W

qR

Ph

OH

OAc R

98 % ee, 43 %yield, PCL, vinyl acetate 98 % ee, 47 % yield, PCL, vinyl acetate

CH20Bz

COzMe

108a

88 % ee, 49 % yield

108b [41]

109a

98 % ee, 47 %yield

10% [41]

QMe OH

OMe OH

I

I

Me0

Me0

.eCN FwcN 1101, [42] 72 % ee, 58 % yield

llOa [42] >99 % ee, 42 % yield, PFL,

vinyl acetate

l l l a [43]

111b [43]

/

F

F

F

F

>99 % ee, 42 % yield

>99 % ee, 45 % yield, LIP, vinyl acetate OAc

NcTr l l 2 a [44]

112b [44]

/

HZN

CI

H2N

CI 97 % ee, 44 % yield

90 % ee, 46 % yield, PCL, vinyl

acetate OAc

113a [45]

O2N)+Br

113b [45]

BnO 96 % ee, 46 % yield

BnO 86 % ee, 48 % yield, PCL, vinyl acetate

r OH

114a [46] F 93 % ee, -, PCL, isopropenyl acetate, 46 % conversion

F

/

90 % ee, -

"'

114b 1461

11.1 Hydrolysis and Formation of Carboxylid Acid Esters

I

513

Table 11.120.

(cont.).

d l

115b [47]

R l i2

R

R' = Ph, 1-naphthyl, 2-naphthyl, benzyl, n-hexyl, R2= Me; R', R2= 50-99 % ee, 39-48 %yield, PCL. PSL, CAL-B,diketene 95-99 % ee, 30-43 % yield

I

OAc RI .&

R+cI R

n

Ph

2

Ph

2

4-t-Bu-C~H4

3

4-t-BU-c~H4

3

1-o-Naphthyl

1

Ph

3

4-F-CsH4

3

4-F-CsH4

3

92 % ee, 49 % yield, PCL, isopropenyl acetate 97 % ee, 31 % yield, CAL-B, vinyl butanoate 99 % ee, 47 % yield, PCL, isopropenyl acetate >95 % ee, -, PCL, vinyl acetate, 50 % conversion 89 % ee, 41 %yield, PCL, isopropenyl acetate 79 % ee. -, PCL, vinyl acetate, 55 % conversion 295 % ee, -, PCL, vinyl acetate, 50 % conversion 97 % ee, 44 % yield, PCL, vinyl acetate

OAc

A CX, R

R

R

X

2-Naphthyl

F

2-Naphthyl

H

1-Naphthyl

H

OCOCH(CH,)=CH,

124a [53]

99 % ee, 44 % yield

llGb [48]

llGa

96 % ee, 33 % yield

llGb [49]

117a

99 % ee, 47 % yield

117b [48]

117a

>95 % ee, -

117b [SO]

118a

99 % ee, 44 % yield

118b [48]

119a

>95 % ee, -

1191, [SO]

120a

>95 % ee, -

120b [SO]

120a

85 % ee, 48 % yield

1201,[51]

121a

>99 % ee, 51 % yield

121b [52]

122a

99 % ee, 43 % yield

122b [S2]

123a

69 % ee, 40 % yield

123b [52]

1

85 % ee, 37 %yield, PCL, vinyl acetate 97 % ee, 37 %yield, PCL, vinyl acetate >99 % ee, 32 %yield, PCL, vinyl acetate

A+ 85-97 % ee, -, PCL, 2,3butanedione monooxime methacrylate, 43-47 % conversion

llGa

CX,

Ar 87-95 % ee, -

124b [53]

514

I

11 Hydrolysis and Formation of C - 0 Bonds

Table 11.1-20.

(cont.).

OAc

Ph,

X

i\

CO,H

X (CH2)2 (E)-CH=CH

84 % ee, 35 % yield, PCL, vinyl acetate 94 % ee, 34 % yield, PCL, vinyl acetate

2

Ph

125a

>99 % ee, 45 %yield

125b [54]

12Ga

>99 % ee, 42 % yield

12Gb [54]

OH 127a [55]

98 % ee, 48 % yield, PCL, vinyl acetate >99 % ee, 49 % yield, PCL, vinyl acetate, 1,4,8,11-

-

Ph-

/

1271, [55]

92 % ee, 48 % yield 98 % ee, 51 % yield

tetrathiacyclo-tetradecane

as additive

R i q R ‘

R’

:

OAc R’

R2

Ph

Me

Me

Ph

OH 95 % ee, -, PSL, isopropenyl acetate, 48 % conversion 99 % ee, -, PCL, isopropenyl acetate, 19 % conversion 98 % ee, -, PSL, isopropenyl acetate, 48 % conversion 98 % ee, -, PCL, isopropenyl acetate, 46 % conversion

89%ee,-

128a

l28b [56]

89 % ee, 129a

92%ee,-

129b [56]

83 % ee, -

OAc 0 130a [57]

130b [57]

Ar >96 % ee. 26-44 % yield, CCL, vinyl acetate

Ar 33-70 % ee, 55-73 %yield

R ’ V O M e

R’

3

OCOR‘ R’

RZ

Ph

Me

Ph

Ph

Alkyl

Me

OMe

OH 93 % ee, 45 %yield, PCL, vinyl acetate >99 % ee, 12 %yield, lipase SL (Meito),vinyl benzoate 30-98 % ee, 42-74 %yield, PCL, vinyl acetate

131a

95 % ee, 50 % yield

131b [58]

132a

61 % ee, 41 %yield

131b [58]

133a

53-99 % ee, 17-43 % 133b [SO] yield

11.1 Hydrolysis and Formation ofcarboxylid Acid Esters

I

515

Table 11.120.

(cont.).

phvoMe

134b [58]

Ph=OMe.

134a [58]

OH >99 % ee, 46 %yield

OAc 93 % ee, 45 % yield, PCL, vinyl acetate 97 % ee. 31 %yield, lipase SL (Meito),vinyl acetate

2

42 % ee, 46 % yield 0

135a [58]

P h vOAc O M e

Ph

OAc

OH

RnACO,Et

/

/

Me R'

tBu

R'

R2 R3

R4 Rs

135b [58]

OH 80 % ee, 44 % yield

76 % ee, 38 %yield, PCL, vinyl acetate

$6

OMe

RnACO,Et

& @

tBu

R2

97 % ee, 46 %yield, PCL (PS-C), vinyl acetate, 48 % conversion 97 % ee, 41 %yield, CRL, vinyl acetate, 45 % conversion 95 % ee, 45 %yield, PCL (PS-C), vinyl acetate, 47 % conversion 97 % ee, 48 % yield, CRL, vinyl acetate, 51 % conversion 97 % ee, 40 %yield, PCL (PS-C), vinyl acetate, 46 % conversion

/

/

OH R4

R3

0

MeM %

R5

13Ga

-, -

13Gb [GO]

137a

-,-

13% [GO]

138a

-, -

138b 1601

139a

-,-

139b 1601

140a

-, -

140b [60]

141b [Gl] >98 % ee, 42 % yield, lipase TL, vinyl acetate

>94 % ee, 40 % yield

516

I

1 1 Hydrolysis and Formation of C - 0 Bonds Table 11.120.

(cont.)

OAc

142a [62] /

Nc*F

F F W N142b [62] F 96 % ee, 46 % yield

F 98 % ee, 50 %yield, LIP, vinyl acetate

143b 163)

143a [63] OH

65 % ee, 49 % yield

95 % ee, 45 %yield, CAL-B, vinyl acetate

‘GH13

a4-

[63]

6Ac 94 % ee, 42 %yield, CAL-B, vinyl acetate

i

144b[63]

OH

81 % ee, 51 %yield

R

Me Et n-Pr n-Bu CHzOPh

>99 % ee, -, CAL-B, vinyl acetate, 25 % conversion 97 % ee, -, CAL-B,vinyl acetate, 35 % conversion >99 % ee, -, CAL-B, vinyl acetate, 25 % conversion >99 % ee, -, CAL-B, vinyl acetate, 32 % conversion >99 % ee, -, CAL-B, vinyl acetate, 14 % conversion

145a

33%ee,-

14% [64]

146a

52%ee,-

146b [64]

147a

33%ee.-

147b [64]

148a

12%ee,-

148b [64]

149a

25%ee,-

14% [64]

94 % ee, 48 % yield, PCL, acetic anhydride 90 % ee, 38 % yield, PCL, acetic anhydride

150a

66 % ee, 35 %yield

150b [65]

151a

54 % ec, 44 %yield

151b [ G S ]

n‘&C02Me 6 7

7 1. I Hydrolysis and Formation ofcarboxylid Acid Esters

I

517

Table 11.120.

(cont.).

OCOnC,H,,

OH

R

R"R2

R l i2

R'

R2

n-C6H13

97 % ee, -, CAL-B, n-C7H1=,COSEt, 50 % conversion 95 % ee, -, CAL-B, C=CH n-C7HIsCOSEt,50 % conversion CH=CH2 96 % ee, -, CAL-B, n-C7HI5COSEt,49 % conversion 96 % ee, -, CAL-B, ET n-C7HIsCOSEt,51 % conversion

Me

n-CsHl;r n-C6H1, ~-C~HI,

152a

98 % ee, -

152b [66]

153a

96 % ee, -

153b [66]

154a

93 % ee, -

154b 1661

155a

>99 % ee, -

155b [66]

AcO-

156a [67]

j--k

SiMe, -, 39 % yield, PCL,vinyl acetate

156b [67] SiMe, 295 % ee, 43 % yield

SiMe,

SiMe,

157a (681

HO

157b (681

8,.

93 % ee, -,

98 % ee, -, BSL, vinyl acetate, 49 % conversion

AcoynprH o d n P r Me

Br

158a [69]

Br

Me

84 % ee, 46 % yield

94 % ee, 30 % yield, PSL, vinyl acetate

HO, d

AcO

i

e

159b [69]

159a (691 Me Br 95 % ee, 32 %yield, PSL, vinyl acetate

158b [G9]

Me Br >98 % ee, 21 % yield

OCOnC,H,, X

d

R

X

R

c1

Me

Br

Me

Br

Et

97 % ee, -, CAL-B, n-C7HISCOSEt,42 % conversion 98 % ee, -, CAL-B, n-C7HIsCOSEt,47 % conversion 96 % ee, -, CAL-B, n-C7H15COSEt,30 % conversion

l6Oa

71 % ee, -

lGOb[70]

l6la

88 % ee, -

1611, (701

lG2a

41 % ee, -

1621,1701

518

I

7 7 Hydrolysis and Formation of C-0 Bonds Table 11.1-20.

(cont.). OCOnC,H,,

Me$

Me$ p J R

/R

R

Me Et n-CI3H2,

99 % ee, -, CAL-B, n-C7Hi&OSEt 81-96 % ee, 24-49 % yield, PCL, vinyl acetate

163a

95%ee,-

163b [70]

164a

36->98 % ee, 41-71 %yield

164b [71]

165b [72]

98 % ee, -

71 % ee, -, PSL, succinic anhydride, 58 % conversion

yo?

l66b[73]

0

OH

>99 % ee, 45 % yield, PCL, vinyl acetate

167a [74] >98 % ee, -, CAL-B, isopropenyl acetate, 49 % conversion

167b [74]

OH

94 % ee, -

0

foEt

OAc

99 % ee, 35 % yield, BSL, vinyl acetate

Me Et

Ph 99 % ee, 36 % yield

q

QOEt R/ R

168b [75]

168a [75]

O

E

t

R*OH

'"OAc

87 % ee, 39 %yield, BSL, vinyl acetate 98 % ee, 38 %yield, BSL, vinyl acetate

169a

99 % ee, 35 %yield

169b [76]

170a

98 % ee, 42 % yield

170b [76]

7 7 . 7 Hydrolysis and Formation of Carboxylid Acid Esters

I

519

Table 11.1-20.

(cont.). RvCO2H

"Y OAc Co2H

OH

R

98 % ee, -, BSL, vinyl acetate, 48 % conversion 98 % ee, -, BSL, vinyl acetate, 48 % conversion

(CH2)13Me CHzPh

171a

91 % ee, -

171b (771

172a

89 % ee, -

172b [77]

173b [78]

173a [78] -NO,

&NO2 91 % ee, 31 %yield, GLL, vinyl acetate

1

__

i

C02Me

C0,Me

nPrOCO

174a [79]

C0,Me

174b [79]

HO "" C0,Me 96 % ee, 35 % yield

82 % ee, 49 % yield, CAL-A, vinyl butanoate

175a [80]

175b [80] 73-99 % ee, 47-56 % yield

>99 % ee, 35-44 %yield, PCL, CAL-B, PFL, vinyl acetate OCOEt CI,&C02R R

96 % ee, 29 % yield, RML, vinyl propionate 77 % ee, 43 %yield, RML, vinyl propionate 89 % ee, 45 %yield, RML, vinyl propionate >97 % ee, 48 % yield, RML, vinyl propionate

Et CHzPh c-C6H11 t-Bu

-,

177a

96 % ee, 29 %yield

177b [81]

178a

96 % ee, 44 %yield

178b [Sl]

179a

99 % ee, 42 % yield

179b [Sl]

-

180a [82]

o

f

y

o

180b [82]

0 OH

OAc

95 % ee, -, CAL-A, vinyl acetate, 25 % conversion, 55 "C 83 % ee, -, CAL-A, vinyl acetate, 56 % conversion, 22 "C

17Gb [81]

17Ga

32 % ee, -

>99 % ee, -

520

I

7 7 Hydrolysis and Formation of C-0 Bonds Table 11.1-20.

(cont.).

OH S0,Ph

181a [83] >99 % ee, 49 % yield, PCL, vinyl acetate

.

BocNH-

/

S0,Ph

1811, [83] >99 % ee, 46 % yield

Rq cop

R

97 % ee, 43 %yield, PCL, isopropenyl acetate 299 % ee, 37 % yield, PCL, vinyl acetate

Me Et

182a

>97 % ee, 42 %yield

1821, [84]

183a

70 % ee, 50 %yield

183b [84]

184a

41 % ee, 69 % yield

1841, [85]

185a

44 % ee, 61 %yield

18% [85]

Jg ph

0

'R

R H

>97 % ee, 24 % yield, PCL, vinyl acetate >97 % ee, 29 % yield, PCL, vinyl acetate

PMP

yPhgPh : H

0

0

R

HI

R

R

H TBDMS

>97 % ee, 49 % yield, PCL, vinyl acetate >97 % ee, 42 %yield, PCL, vinyl acetate

186a

>97 % ee, 47 % yield

186b [85]

187a

>97 % ee, 49 % yield

1871, [85]

R&R HO

EtOCO R'

R=

CF3

TMS

CHFz C2F5

>99 % ee, -, CAL-B,vinyl propionate, 42 % conversion TBDMS >99 % ee, -, CAL-B, vinyl propionate, 36 % conversion TMS 98 % ee, -, LIP,

188a

72 % ee, -

188b [86]

189a

53 % ee, -

189b [86]

190a

22%ee,-

190b (861

1 1.1 Hydrolysis and Formation of Carboxylid Acid Esters

(cont.).

Table 11.1-20.

191a [87]

~ 0

O :

=

H,R2 = N 0 2 , X = C1

95 % ee, 41 % yield, PCL, vinyl acetate R'= H, R2 = NO2, X = F >98%ee, 41 % yield, PCL, vinyl acetate R' = H, R2 = NO2, X = Br 94% ee, 21 %yield, PCL, vinyl acetate R'= NO2, R 2 = H, X = C1 92% ee, 36 % yield, PCL, vinyl acetate R'= NO2, R 2 = H, X = F 96% ee, 34 %yield, PCL, vinyl acetate

192a

89% ee, 43 %yield

192b I881

193a

95 % ee, 43 % yield

193b [88]

194a

39% ee, 45 % yield

194b [88]

195a

95 % ee, 43 % yield

195b [88]

196a

48 % ee, 45 % yield

196b [88]

OAc s

)-N

v

C 191b [87]

OH

OAc

R'

A

OH >99 % ee, 47 % yield

OAc 82 % ee, 52 % yield, PCL, vinyl acetate

OH

1

R Me

Mi

Me

CH2CH=CH2 -, -, PSL, vinyl acetate CHzCsCH -, -, PSL, vinyl acetate CH=CH2 -, -, PSL, vinyl acetate

197a 198a 199a

R

88 % ee, 46 %yield 94 % ee, 40 % yield 90 % ee, 48 % yield

19% 1891 198b (891 199b [89]

OAc

S&-J

200a [90]

93 % ee, -, PFL, vinyl acetate, 31 % conversion

42 % ee, -

200b [90]

I

521

522

I

1 1 Hydrolysis and Formation of C - 0 Bonds Table 11.120.

a

(cont.).

201a [91] 89 % ee, 36 %yield, CAL-B, vinyl acetate

Me

/

62 % ee, 23 % yield

202a [92] R'

R'

202b [92] R'

OAc = H,

OH

Br, Me, TBDMS-OCHz, TrOCH2, Ph, 2-Py

R2 = Me, Et, Vinyl 92-99 % ee, 31-49 % yield, CAL-B, vinyl acetate

78-99 % ee, 43-58 %yield

R = Me, n-Pr, n-C5HII, n-C7H15 98->99 % ee, 22-48 %yield, 203a [93] PCL or CAL-B, vinyl alkanoate

35-96 % ee, 25-45 %yield

M

201b[91]

203b [93]

Hz

>98 % ee, 44-46 % yield,

204a

89->98 % ee, 36-47 % yield

204b [94]

Zn

PCL, CAL-B or LIP, vinyl acetate >98 % ee, 12-24 %yield, PCL, CAL-B or LIP, vinyl acetate

205a

16->95 % ee, 72-82 % yield

205b [94]

1 7 . 1 Hydrolysis and Formation ofcarboylid Acid Esters

I

523

Table 11.1-20.

(cont.).

n

H

c

5

1

1

~20Ga R[95]

n

c

5

H

1

1

~ R 20Gb [95]

OH

OAc n = 1-3, TBDMS, TBDPS 94-95 % ee, 35-42 % ee, CRL, vinyl acetate

58-77 % ee, 33-46 % ee

AA 0

0

0

1

R' +i2

R'

Ph Ph(CH& PhCH=CH 2-Naphthyl

R2

Me Vinyl Me Me

96 % ee, 41 % yield 92 % ee, 42 %yield 97 % ee, 41 % yield 93 % ee, 40 % yield all CAL-B, methyl acetoacetate

207a 208a 209a 210a

98 % ee, 45 % yield 96 % ee, 44 % yield 90 % ee, 38 % yield 96 % ee, 46 %yield

20% [96] 208b [9G] 209b[9G] 210b [96]

OAc

211b [97]

R = H. 2-Me, 4-Me, 4-C1,4-Br, 2,3-C4&, 3,4-C4H4 89- >99 % ee, 34-55 % yield, PCL, vinyl acetate

35-94 % ee, 41-62 %yield

Meog ?Me

\

?Me

212a [98]

212b [98]

Meo$ 92 % ee, 30 %yield, PCL, vinyl acetate

47 % ee, 68 % yield

213a [98]

92 % ee, 38 % yield, PCL, vinyl acetate

213b [98]

76 % ee, 51 % yield

524

I

1 7 Hydrolysis and Formation of C-0 Bonds 1 M. Inagaki, I. Hiratake, T. Nishioka, J. Oda,]. Am. Chem. SOC.1991, 113,9360. 2 M. Inagaki, J. Hiratake, T. Nishioka, 7. Oda,]. Org. Chem. 1992,57,5643. 3 A. 1. M. Janssen, A. J. H. Klunder, B. Zwanenhurg, Tetrahedron 1991,47, 7645. 4 B. Cambou, A. M. Klibanov,]. Am. Chem. SOC. 1984,106,2687.

5 D. A. Abramowicz, C. R. Keese, Biotechnol. Bioeng. 1989, 33, 149. 6 G. Kirchner, M. P. Scollar, A. M. Klibanov,]. Am. Chem. SOC.1985, 107, 7072. 7 YF :. Wang, J. J. blonde, M. Momongan, D. E. Berghreiter, C.-H. Wong,]. Am. Chem. SOC.1988, 110,7200. 8 T. M. Stokes, A. C. Oehlschlager, Tetrahedron Lett. 1987,28, 2091. 9 K. Laumen, D. Breitgoff, M. P. Schneider,]. Chem. Soc., Chem. Commun. 1988, 1459. 10 D. Bianchi, P. Cesti, E. Battistel,]. Org. Chem. 1988,53,5531. 11 P. D. Theisen, C. H. Heathcock,]. Org. Chem. 1988,53,2374. 12 K. Burgess, L. D. jennings,]. Org. Chem. 1990, 55, 1138. 13 M. De Amici, C. De Micheli, G. Carrea, S. Spezia, 1. Org. Chem. 1989,54, 2446. 14 H. Frykman, N. Ohmer, T. Norin, K. Hult, Tetrahedron Lett. 1993, 34, 1367. 15 K. Burgess, J. Cassidy, I. Henderson,]. Org. Chem. 1991,56,2050. 16 K. Burgess, I. Henderson, Tetrahedron: Asymmetry 1990, 1, 57. 17 A. Kamal, M. V. Rao, Tetrahedron: Asymmetry1991, 2, 751. 18 M. A. Sparks, 1. S. Panek, Tetrahedron Lett. 1991, 33,4085. 19 H. S. Bevinakatti, A. A. Baneji,]. Org. Chem. 1991,56,5372. 20 T. Sugai, H. Ohta, Tetrahedron Lett. 1991, 32, 7063. 21 1. M. Chong, E. K. Mar, Tetrahedron Lett. 1991, 32, 5683. 22 K. Burgess, L. D. Iennings, J. Am. Chem. SOC. 1991, 113,6129. 23 S. Takano, M. Setoh, K. Ogasawara, Tetrahedron: wmmetry 1993,4,157. 24 R. Chinchilla, C. Nijera, J. Pardo, M. Yus, Tetrahedron: Asymmetry1990, I, 575. 25 E. Dominguez, J. C. Carretero, A. Femandez-Mayoralas, S. Conde, Tetrahedron Lett. 1991, 32, 5159. 26 1. C. Carretero, E. Dominguez,]. Org. Chem. 1992, 57, 3867. 27 R. ChPnevert, R. Gagnon,]. Org. Chem. 1993,58. 1054. 28 T. Miyazawa, S. Kurita, S. Ueji, T. Yamada, S. Kuwata,]. Chem. SOC.Perkin Trans. 1 1992, 2253. 29 M..]. Kim, Y. K. Choi,]. Org. Chem. 1992, 57, 1605. 30 U. Ader, M. P. Schneider, Tetrahedron: Asymmetry 1992, 3, 521.

31 C. Ebert, G. Ferluga, L. Gardossi, T. Gianferrara, P. Linda, Tetrahedron: Asymmetry1992, 3, 903. 32 R. Seemayer, M. P. Schneider, Tetrahedron: Asymmetry 1992, 3,827. 33 V. Fiandanese, 0. Hassan, F. Naso, A. Scilimati, Synlett 1993, 491. 34 H. Takahata, Y. Uchida, T. Momose, Tetrahedron Lett. 1992, 33, 3331. 35 F. Theil, K. Lemke, S. Ballschuh, A. Kunath, H. Schick, Tetrahedron: Asymmetry1995, 6 , 1323. 36 G. Di Bono, A. Scilimati, Synthesis1995, 699. 37 1. L. Bermudez, C. del Campo, L. Salazar, E. F. Llama, J. V. Sinisterra, Tetrahedron: Asymmetry1996, 7, 2485. 38 K. Wunsche, U. Schwaneherg, U. T. Bomscheuer, H. H. Meyer, Tetrahedron: Asymmetry1996,7, 2017. 39 N. W. Boaz, R. L. Zimmerman, Tetrahedron: Asymmetry 1994, 5, 153. 40 F. Marguet, J:F. Cavalier, R. Verger, G. Buono, Eur.]. Org. Chem. 1999,1671. 41 T. Ziegler, F. Bien, C. lurisch, Tetrahedron: Asymmetry 1998, 9, 765. 42 F. M. Hauser, D. Sengupta, S. A. Corlett,]. Org. Chem. 1994,59,1967. 43 T. Sakai, T. Takayarna,T. Ohkawa, 0. Yoshio, T. Ema, M. Utaka, Tetrahedron Lett. 1997, 38, 1987. 44 S. Conde, M. Fierros, M. I. Rodriguez-Franco, C. Puig, Tetrahedron: Asymmetry1998, 9, 2229. 45 F. Campos, M. P. Bosch, A. Guerrero, Tetrahedron: Asymmetry2000,l I, 2705. 46 R. L. Hanson, A. Banejee, F. T. Comezoglu, K. D. Mirfakhrae, R. N. Patel, L. 1. Szarka, Tetrahedron: Asymmetry1994,5,1925. 47 K. Suginaka, Y. Hayashi, Y. Yamamoto, Tetrahedron: Asymmetry1996, 7, 1153. 48 S. B. Raju, T.-W. Chiou, D.-F. Tai, Tetrahedron: Asymmetry1995,6, 1519. 49 H.-L. Liu, B. H. Hoff, T. Anthonsen, /. Chem. Soc., Perkin Trans. 1 2000, 1767. 50 D. Bianchi, P. Moraschini, A. Bosetti, P. Cesti, Tetrahedron: Asymmetry1994,5, 1917. 51 N. Gil, P. Bosch, A. Guerrero, Tetrahedron 1997,53 15115. 52 1. Gaspar, A. Guerrero, Tetrahedron: Asymmetry 1995, 6, 231. 53 V. Athawale, N. Manjrekar, Synlett 2000, 225. 54 A. Chadha, M. Manohar, Tetrahedron: Asymmetry 1995, 6,651. 55 Y. Takagi, 1. Teramoto, H. Kihara, T. Itoh, H. Tsukube, Tetrahedron Lett. 1996, 37, 4991. 56 W. Adam, M. T. Diaz, R. T. Fell, C. R. Saha-Moller, Tetrahedron: Asymmetry1996,7, 2207. 57 M. S. Nair, S. Ioly, Tetrahedron: Asymmetry2000, 11, 2049. 58 H. Hamamoto, V. A. Mamedov, M. Kitamoto, N. Hayashi, S. Tsuboi, Tetrahedron: Asymmetry 2000, 11,4485. 59 S. Tsuboi, N. Yamafuji, M. Utaka, Tetrahedron: Asymmetry1997,8,375.

7 1 . 1 Hydrolysis and Formation ofcarboxylid Acid Esters GO W. Zhang, P. G. Wang, J. 0%.Chem. 2000,65,

4732. 61 Y. Aoyagi, N. Agata, N. Shibata, M. Horiguchi, R. M. Williams, Tetrahedron Lett. 2000,41, 10159. 62 T. Sakai, Y. Miki, M. Tsuboi, H. Takeuchi, T. Ema, K. Uneyama, M. Utaka,J. 0%.Chem.2000,65, 2740. 63 K. Morishita, M. Kamezawa, T. Ohtani, H. Tachibana, M. Kawase, M. Kishimoto, Y.Naoshima, J. Chem. SOL, Perkin Trans. 11999,513. 64 H. Hamada, M. Shiromoto, M. Funahashi, T. Itoh, K. Nakamura,J. 0%.Chem. 1996,61,2332. 65 A. Sattler, G. Haufe, Tetrahedron: Asymmetry 1995, 6,2841. 66 C. Orrenius, N. Ohrner, D. Rotticci, A. Mattson, K. Huh, T. Norin, Tetrahedron: Asymmetry 1995,6, 1217. 67 C. Meert, J. Wang, P. J. De Clerq, Tetrahedron Lett. 1997,38,2179. 68 W. Adam, C. Mock-Knoblauch,C. R. Saha-Moller, Tetrahedron: Asymmetry1997,8,1441. 69 W. Adam, L. Blancafort, C. R. Saha-Moller,Tetrahedron: Asymmetry1997,8,3189. 70 D. Rotticci, C. Orrenius, K. Hult, T. Norin, Tetrahedron: Asymmetry1997,8,359. 71 P. Allevi, P.Ciuffreda, M. Anastasia, Tetrahedron Asymmetry1997,8,93. 72 K. Nakamura, K. Tikenaka, A. Ohno, Tetrahedron: Asymmetry1998,9,4429. 73 S . Vrielynk, M. Vandewalle,A. M. Garcia, J. L. Mascarefias, A. Mourifio, Tetrahedron Lett. 1995,36,9023. 74 W. Adam, C. R. Saha-Moller,K. S. Schmid, Tetrahedron: Asymmetry1999,10,315. 75 W. Adam, P. Groer, C. R. Saha-Moller,Tetrahedron: Asymmetry2000,11,2239. 76 W. Adam, P. Groer, H.-U. Humpf, C. R. Saha-Moller,J. Ox . Chem. 2000,65,4919. 77 W. Adam, M. Lazarus, A. Schmerder, H.-U. Humpf, C. R. Saha-Moller,P. Schreier, Eur.1. Org. Chem. 1998,2013.

78 D. Kalita, A. T. Khan, N. C. Barua, G. Bez, Tetrahedron 1999,55,5177. 79 A. Liljeblad, L. T. Kanerva, Tetrahedron: Asymmetry 1999,10,4405. 80 I. Petschen, M. P. Bosch, A. Guerrero, Tetrahedron: Asymmetry2000,11,1691. 81 B. H. Hoff, T. Anthonsen, Tetrahedron: Asymmetry 1999,10, 1401. 82 B. Henkel, A. Kunath, H. Schick, Liebigs Ann. Chem. 1995,921. 83 J.de Vicente, R. G. Arrayas, J. C. Carreteo, Synlett 2000,53. 84 N. Hayashi, K. Yanagihara, S . Tsuboi, Tetrahedron: Asymmetry 1998,9,3825. 85 M. Bucciarelli, P. Davoli, A. Forni, I. Moretti, F. Prati,]. Chem. Soc., Perkin Trans. 1 1999,2489. 86 M. Takeda, T. Ishizuka, K. Itoh, T. Kitazume, /. fluorine Chem. 1995,75,111. 87 J.E. Kaminska, K. Smigielski, D. tobodzidska, J. G6ra, Tetrahedron:Asymmetry2000,1 1 , 1211. 88 R. Skupin, T. G. Cooper, R. Frohlisch, J. Prigge, G. Haufe, Tetrahedron: Asymmetry 1997,8,2453. 89 B. Zhu, J.S . Panek, Tetrahedron Lett. 2000,41, 1863. 90 L. Di Nunno, C. Franchini, A. Scilimati, M. S . Sinicropi, P. Tortorella, Tetrahedron: Asymmetry 2000,11, 1571. 91 J. A. Fuentes, A. Maestro, A. Testera, J. M. Bbiiez, Tetrahedron: Asymmetry 2000, 11, 2565. 92 J. Uenishi, T.Hiraoka, S . Hata, K. Nishiwaki, 0.Yonemitsu, J. Org. Chem. 1998,63,2481. 93 T.Kijima, T. Moriya, E. Kondoh, T. Izumi. Tetrahedron Lett. 2000,41,2125. 94 T.Ema, M.Jittani, T. Sakai, M. Utaka, Tetrahedron Lett. 1998,39,6311. 95 J. S. Yadav, S . Nanda, A. Bhaskar Rao, Tetrahedron: Asymmetry2001,12,53. 96 A. Chdova, K. D. Janda,J. Org. Chem. 2001,66, 1906. 97 A. Kamal, G.B. R. Khanna, Tetrahedron: Asymmetry 2001,12,405. 98 E. Brenna, C. Fuganti. P. Grasselli, S . Serra, Eur. /. Org. Chem. 2001,1349.

Monoacetates and alcohols of Table 11.1-19 which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-6 and 11.1-14.

For a wide structural range of racemic secondary alcohols, lipase-catalyzed enantiomer-differentiatingacylation has been reported (1-213) (Table 11.1-20).The results show that this is a general method for the attainment of enantiomerically pure secondary alcohols that is complementary to the lipase-catalyzedhydrolysis of the corresponding acylated alcohols (Table 11.1-15). It is especially worth mentioning that secondary alcohols of the alkyl-alkyl, alkyl-aryl or alkyl-heteroaryltype, but also those bearing the various firnctional groups including stannylated derivatives, are accessible too. Acylation has been utilized in depth for the synthesis of allylic, homoallylic, propargylic and homopropargylic and allenylic alcohols (17-20,25-27,

I

525

526

I

1 1 Hydrolysis and Formation of C - 0 Bonds Table 11.1-21. Lipase-catalyzed enantiorner-differentiating acylation of racernic cyclic secondary alcohols in organic solvents (CCL Candido cylindracea lipase, PSL Pseudomonos sp. lipase, CAL-B Candida antarctica B lipase, PPL p i g pancreas lipase, PCL Pseudomonos cepacia lipase, LIP Pseudomonas sp. lipase-Toyobo, ASL Alcaligenes sp. lipase, PFL PseudomonasPuorescens lipase, BSL Burkholderia sp. lipase, CRL Candida rugosa lipase, MML Mucor miehei lipase).

U 96 % ee, 48 % yield

95 % ee, 48 % yield, CCL, triacetin

OAc

n 1 2

299 % ee, 46 % yield 97 % ee, 49 % yield, all PSL, vinyl acetate

2a 3a

95 % ee, 48 % yield 299 % ee, 44 % yield

95 % ee, 52 %yield, PSL, vinyl acetate

89 % ee, 48 % yield

299 % ee, 48 %yield, PSL,

299 % ee, 47 % yield

2b I21 3b 121

vinyl acetate

6 6

OH

OCOnC,H,, ..+Me

97 % ee, -, CAL-B, n-C,H&OSEt

OCOnPr

87 % ee, 95 % ee, -, (triple resolution) PSL,vinyl acetate

Ga[51

uMe 97 % ee, -

OH

98 % ee, -

P P j % 8P ‘aJ % L67 HO [6 ‘81 9f I

JlelaJe 1Xup ‘13d ‘PIJj % LE ‘aJ % L67

16 ’81 921

PPP % 8P ’Ja % L67 HO

0

Yd

PIJd % LE ‘JJ % LG7

avo [6 ‘81 901

b

Y

d

P I J j % LP ‘JJ % L67 HO

avo

PlJlX % GE ‘JJ % 8667 Ho<

HO

[LI 98 $%03v

7 I Hydrolysis and Formation ofC-0 Bonds Table 11.1-21.

(cont.).

Me phQ

Me

14a [S, 91

OAc

OH

70 % ee, 59 % yield, PCL, vinyl acetate

Me phQ

14b [8,91

Me

297 % ee, 41 % yield

15a [8, 91

OAc

15b [8, 91 OH

296 % ee, 26 % yield, PCL, vinyl acetate

297 % ee, 44 % yield

0sitBuPh,

n~*~'.OSitBuPh,

16a [lo] "OAc

OH

98 % ee, 50 % yield

90 % ee, 50 %yield, PCL, vinyl acetate

H

O

16b [lo]

Q

A

0

100 % ee, 48 % yield, CCL, acetic anhydride

17a [ll]

17b [ll]

98 % ee, 51 % yield

Q

P

OH

96 % ee, 50 % yield, CCL, acetic anhydride

100 % ee, 48 % yield

11.1 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.121.

(cont.).

FO,Et

C0,Et 19a [12, 131 &H

298 % ee, 48 % yield, PCL, vinyl acetate C0,Et

19b [12, 131

298 % ee, 48 %yield

C0,Et

bss.oActYH 20a [13]

99 % ee, 40 %yield, PCL, vinyl acetate

A

95 % ee, 53 %yield

21a [14]

OCOnPr

4

21b [14]

OH 87 % ee, -

87 % ee, -, PPL, n-PrC02CH2CC13

HO22a 1151

"

20b [13]

O

.*-R

0%

22b [15]

K R 0 0 R = Ph, 3-MeoC~H4,3,4-(MeO)&H3, 3,4-(methylenedioxy)CGH3 299 % ee, -, PCL, 299 % ee, vinyl acetate OCOR

R Me 99 % ee, 38 % yield n-Pr 99 % ee, 47 %yield PCL, vinyl acetate or butyric anhydride

R

OH

23a 24a

71 % ee, 51 %yield 94 % ee, 48 % yield

23b [16] 24b [16]

OH

25a (171

.,.'"\

cTo~..Ms

OTBDMS

94 % ee, 38 % yield, PCL, vinyl acetate

51 % ee, 57 %yield

25b [17]

I

529

530

I

7 7 Hydrolysis and Formation of C-0 Bonds

(cont.).

Table 11.1-21.

SiMe,

:T5 1

93 % ee, -, PCL, vinyl acetate, 50 conversion 97 % ee, -, BSL, vinyl acetate, 50 conversion 99 % ee, -, BSL, vinyl acetate, 50 conversion

2

SiMe,

2Ga

Ho,8*b

95 % ee, -

2Gb [18]

>99 % ee, 27a

98 % ee, -

2% [18]

28b [19]

R = Me, Et, n-Pr, n-CsH11, n-C7H15, n-CgH19, ClCH2 70-97 % ee, 41-51 %yield, PCL, vinyl alkanoate

69-95 5 ee, 40-51 %yield

OCOR

OH

1

29b [19]

R = Me, n-Pr, n-CgH19, ClCH2, Ph 98- >99 % ee, 40-46 % yield, PCL, vinyl alkanoate

67- >99 % ee, 50-58 % yield

Rb

AcO R

Ph PMP t-Bu

CMezPh TBDMS Bn THP

>95 % ee, 45 %yield, PCL, isopropenyl acetate 87-90 % ee, 40 %yield, PCL, isopropenyl acetate 97 % ee, 48 % yield, PCL, vinyl acetate 76 % ee, 51 %yield, pancreatin, vinyl acetate >99 % ee. 43 % yield, PCL, vinyl acetate 98 % ee, 48 % yield, pancreatin, vinyl acetate 30 % ee, 74 % yield, pancreatin, vinyl acetate 91 % ee, 45 %yield, pancreatin, vinyl acetate

30a

>95 % ee, 49 % yield

30a [20]

31a

-, 41 %yield

31b [20]

32a

91 % ee, 53 % yield

32b 1211

32a

98 % ee, 40 % yield

32b [23b]

33a

>99 % ee, 50 % yield

33b [22]

34a

98 % ee, 47 % yield

341,1231

35a

98 % ee, 20 % yield

35b (23bj

36a

94 % ee, 50 % yield

3Gb [23b]

1 1.1 Hydrolysis and Formation ofCarboxylid Acid Esters

Table 11.1-21.

I

531

(cont.). HO

R

91 % ee, 30 %yield, PSL, vinyl acetate 96 % ee, 20 % yield, PSL, vinyl acetate 91 % ee, 21 %yield, PSL, vinyl acetate

Me n-CsH19 CHzPh

37a

63 % ee, 59 % yield

3% [24]

38a

40 % ee, 75 % yield

38b [24]

39a

43 % ee, 63 % yield

39b [24]

ACooNHCbz 40a [25]

C

o

o

FN N e

40b [25]

__

92 % ee, 40 % yield, PSL, vinyl acetate

A

HO,,..

’

41a [26]

H O , , . . O& ,“.I,‘

FN

41b [26]

-

“Y “H,

“Y” NH,

>90 % ee, 35 %yield, PCL, vinyl acetate

>90 % ee, 33 % yield

OnP(0)(OEt),

OnP(0)(OEt),

42a [271

AcoQ OH 72 % ee, 42 % yield, PCL, vinyl acetate

HO,,,,b

42b [27]

OAc 95 % ee, 40 % yield

OAc

1

n

1

2

93 % ee, 50 % yield, PCL, vinyl acetate 98 % ee, 48 % yield, PCL, vinyl acetate

43a

100 % ee, 47 % yield

43b [28]

44a

100 % ee, 47 % yield

44b [28]

11 Hydrolysis and Formation of C - 0 Bonds

Table 11.121.

(cont.). NHCbz

NHCbz

OH

"OAc n

98 % ee, 47 % yield, PCL, vinyl acetate >99 % ee, 31 % yield, PCL, vinyl acetate

1

2

45a

>99 % ee, 57 % yield

45b [29]

46a

49 % ee, 63 % yield

46b[29]

NHCbz Q N H C"OAc b2

OH

n

1

99 % ee, 50 % yleld, PCL, vinyl acetate 99 % ee, 47 % yield, CAL-B, isopropenyl acetate

2

47a

99 % ee, 50 % yield

4% [30]

48a

70 % ee, 47 % yield

48b [30]

( W N R 2 "OAc

OH

n R

93-98 % ee, -, CAL-B or PCL, vinyl acetate 2 Me 98 % ee, 42 % yield, CAL-B,vinyl acetate 3 Me 95 % ee, -, CAL-B or PCL, vinyl acetate 1 -(CH45- 97-99 % ee, -, CAL-B or PCL, vinyl acetate 2 -(CH2)5- 99 % ee, 46 % yield, CAL-B,vinyl acetate 2 CH2Ph >99 % ee, 40 %yield, PCL, vinyl acetate 1 Me

49a

97-98 % ee, -

49b [31]

50a

96 % ee, 34 %yield

50b [32]

51a

92-95 % ee, -

51b [31]

52a

99 % ee, -

52b [31]

53a

97 % ee, 49 %yield

53b [32]

54a

99 % ee, 50 % yield

54b [32]

(oc"". OH n

R

1 Me

2 Me 2 Me 2 -(CH2)5-

95-99 % ee, -, PCL or CAL-B, vinyl acetate 99 % ee, 38 % yield, PCL, vinyl acetate 94-96 % ee, -, PCL or CAL-B, vinyl acetate 98 % ee, 44 % yield, PCL, vinyl acetate

55a

94-99 % ee, -

55b [31]

56a

96 % ee, 49 % yield

56b 1321

57a

37-53 % ee, -

57b [31]

58a

97 % ee, 46 %yield

58b 1321

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esten

I

533

Table 11.1-21.

(cont.).

Me I

(aN"HBoc "OAc

OH

n

91 % ee, 45 % yield, CAL-B,vinyl acetate 99 % ee, 20 % yield, CAL-B, vinyl acetate

1

2

59a

99 % ee, 31 %yield

59b [33]

60a

99 % ee, 22 % yield

Gob [33]

NHBoc OH

'"OAc

n 1 2

99 % ee, 46 % yield, PCL, vinyl butyrate >99 % ee, 43 % yield, PCL, vinyl acetate

c

6la

99 % ee, 27 % peld

6 l b [33]

G2a

98 % ee, 45 % yield

G2b [33]

Ho

3

AcO

C0,Et

SPh

63a 1341

G3b [34] Ho

>99 % ee, 42 % yield, PSL, vinyl acetate

SPh

299 % ee, 48 % yield

(yPh

G4b [35]

" OAc

>99 % ee, 50 % yield, PCL, vinyl acetate

>99 % ee, 49 % yield

()'TLoph

65b 1361

>99 % ee, 44 % yield, PCL, vinyl acetate

aO" >99 % ee, 48 %yield

G6a [37]

G6b [37]

C0,Et

>99 %, 45 %yield, PFL, vinyl acetate, 45 % conversion

>99 % ee, -

534

I

1 1 Hydrolysis and Formation of C- 0 Bonds Table 11.1-21.

(cont.).

,-G.l "

R2

R'

Me t-Bu c-C6HI1 Ph -(CH2)3-(CH2)4-(CHz)s-

H

H H H

91 % ee, -, CAL-B, vinyl acetate 93 % ee, -, CAL-B, vinyl acetate 85 % ee, -, CAL-B,vinyl acetate 89 % ee, -, CAL-B, vinyl acetate 95 % ee, -, CAL-B, vinyl acetate 96 % ee, -, CAL-B,vinyl acetate 98 % ee, -, CAL-B, vinyl acetate

OAc

__

67a 68a 69a 70a 71a 72a 73a

- -

__

_-

__ __ __

67b [38] 68b [38] 69b [38] 70b 1381 71b [38] 72b [38] 73b [38]

OH

74a [39]

oBr

95 % ee, -, PPL, vinyl acetate, 44 % conversion

75 % ee, -

99 % ee, -, CRL, isopropenyl acetate, 34 % conversion

60 % ee, -

74b [39]

OH

hCozEt 7Ga [41]

&

bBrpBr OH

OH

77a[42]

Br

OAc >98 % ee, 10 % yield, MML, vinyl acetate

7Gb [41]

95 % ee, 53 %yield

99 % ee, 40 % yield, PCL, vinyl acetate

OAc

~ c o ~ E t

77b[42]

.

Br

OAc 298 % ee, 37 % yield

77c[42]

"

Br

OH >98 % ee, 48 % yield

11.1 Hydrolysis and Formation ofCarboxylid Acid Esters

I

535

Table 11.1-21.

(cont.).

0

H?

AcO

@

OAc 96 % ee, 38 %yield, PCL, vinyl acetate >99 % ee, 32 % yield, pancreatin, AcOCH2CCls

78a [43] 78a (431

H

OH 98 % ee, 44 %yield

78b [43]

55 % ee, 57 % yield

78b [44]

6

R'OCO

R3

R'

R2

Me

H

Me

H

Me

Me

n-C5HI1 H

R3

84 % ee, -, CAL-B, 54 % conversion Me SG%ee,-,CAL-B, 53 % conversion H 90 % ee, -, CAL-B, 52 % conversion H, alkyl, 87-- >99 % ee, 43-51 % CH#h yield, CAL-B, isopropenyl hexanoate

H

79a

98 % ee, -

79b [45]

80a

99%ee,-

80b[45]

81a

97 % ee, -

81b [45]

83a [47]

95 % ee, 27 % yield, PSL, vinyl butanoate, double resolution

(f%

83b (471

83 % ee, -

94 % ee, -, CRL, vinyl acetate

nBuOCO

82b [4G]

OH

OAc

&o

75- >99 % ee, 41-53 % yield

82a

84a[48]

&o HO

91 % ee, 29 % yield

84b [48]

536

I

1 I Hydrolysis and Formation of C-0 Bonds Table 11.121.

(cont.).

AcO &co2Me

c;

H

>97 % ee, 42 % yield

>97 % ee, 43 % yield, PCL, vinyl acetate

&

HO

AcO

861,[SO]

86a [SO] H

H

98 % ee, 38 % yield

99 % ee, 44 % yield, PSL, vinyl acetate 99 % ee, 41 % yield, PCL, vinyl acetate

92 % ee, 32 % yield

n

86 % ee, -, PCL, vinyl acetate, 52 % conversion 79 % ee, -, PSL, vinyl acetate, 57 % conversion

1

2

87a [51]

95 % ee, -

87b[S1]

88a [Sl]

99 % ee, -

88b [Sl]

05/

98 % ee, 48 %yield, PCL, vinyl acetate >96 % ee, 44% yield, PCL, isopropenyl acetate

m.,,

89a [52] 89a [53]

89b [52]

>96 % ee, 46 % yield

89b [53]

NHCbz

NHCbz I

>99 % ee, 48 %yield

901, [ 541

,OAc

/

>99 % ee, 43 % yield, PSL, vinyl acetate

g0a'541

78 &OH % ee, 28 % yield

7 7 . 7 Hydrolysis and Formation ofcarboxylid Acid Esters

(cont.). NHCbz

NHCbz

~

-

~

~

~

O

A

c >99 % ee, 37 % yield

98 % ee, 41 %yield, PSL, vinyl acetate

OH d

N

H

C

b

z

>99 % ee, 41 %yield, PSL, vinyl acetate

96 % ee, 38 % yield

OH NHCbz

d I i

95 % ee, 38 %yield, PSL, isopropenyl acetate

20 % ee, 40 % yield

QAc D /

B

OH

94b [55]

r

100 % ee, 31 %yield

93 % ee, 35 % yield, CAL-B, vinyl acetate

OMe I 95a [56]

95b [56] HO

88 % ee, 48 %yield, CAL-B, vinyl acetate

98 % ee, 40 % yield

9Gb [57] HO"' 92 % ee, 41 % yield, LIP,

vinyl acetate

I

537

Table 11.1-21.

>99 % ee, 46 % yield

538

I

1 1 Hydrolysis and Formation of C - 0 Bonds

Table 11.1-21.

(cont.).

97a [SS]

>99 % ee, 44 %yield, CAL-B, vinyl acetate

97b I581

98 % ee, 47 % yield

n

99 % ee, 42 %yield, LIP, vinyl acetate 94 % ee, 36 %yield, LIP, vinyl acetate

1 2

h

98a [59]

95 % ee, 43 %yield

98b I591

99a [59]

55 % ee, 43 %yield

99b [59]

1OOb 1601

lOOa [60]

pOAc

\.

HO >99 % ee, 50 % yield

299 % ee, 49 % yield, PCL, vinyl acetate

HO n

1 2

299 % ee, 50 % yield, PCL, vinyl acetate >99 % ee, 46 % yield, PCL, vinyl acetate

s

CI CI

1Ola [61]

>99 % ee, 49 %yield

1Olb [61]

102a [61]

>99 % ee, 49 %yield

102b [61]

103a [62]

103b [62]

CI

>95 % ee, -, CRL, vinyl acetate, 44 % conversion

77 % ee, -

11.1 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.121.

(cont.).

@O.TO

104a (631

104b [63]

OAc

OH

87 % ee, 52 %yield, PSL, vinyl acetate

>98 % ee, 48 % yield

OAc

lO5a [64] 94 % ee, 41 % yield, PFL, isopropenyl acetate

105b [64]

97 % ee, 34 % yield

lOGa [65] I Ph

Ph

88 % ee, 46 % yield, PCL, vinyl acetate

LNFrnoc

1OGb [65] 99 % ee, 46 % yield

107a [66]

96 % ee, 46 % yield, PCL, vinyl acetate

107b [66] 99 % ee, 43 % yield

uoAc Y

108a [67]

Cbz

>99 % ee, 47 % yield, PCL, vinyl acetate

1081,[67] I Cbz

>99 % ee, 48 % yield

YJNH OAc

50 % ee, 53 %yield, lipase PL, vinyl acetate

109b [68] OH

98 % ee, 35 % yield

I

539

540

I

1 1 Hydrolysis and Formation ofC-0 Bonds

(cont.).

Table 11.1-21.

R2 " g N - B n OAc R'

R2

H

H

Me

H

OH

>99 % ee, 50 %yield, PCL 110a or CAL-B, vinyl acetate >99 % ee, 49 %yield, CAL-llla B, vinyl acetate

&..

>99 % ee, 48 % yield

llOb [69]

>99 % ee, 48 % yield

l l l b [69]

0

112a [69]

OAc

1121, [G9]

&-Bn OH

88 % ee, 49 % yield, PCL, vinyl acetate

99 % ee, 37 % yield

113a [70]

o'soH 113b [70]

o

TBDMS

TBDMS-'."'

78 % ee, 52 % yield, PCL, vinyl acetate 78 % ee, 54 %yield, PSL, vinyl acetate

97 % ee, 40 % yield 99 % ee, 42 % yield

phsL-cx 114a [71]

AcO

>99 % ee, 49 %yield, PCL, vinyl acetate

114b [71] HO ""

>99 % ee, 49 % yield

115b [72]

93 % ee, 96 % yield

62 % ee, 52 %yield, DCL, vinyl acetate

llGa [73]

I

1

OAc R' = H, Me, R2 = H, OMe, R' H, Me 71-100 % ee, -, CCL, vinyl acetate, 19-57 % conversion R2

"'yyJR3

llGb [73]

I

R2

i

OH

=

22-100 % ee, -

11.1 Hydrolysis and Formation of Carboxylid Acid Esters Table 11.1-21.

(cont.).

AcO

117a(74] I Ts

-r\ I

11%[74]

I

Ts 32 % ee, 75 %yield

96 % ee, 25 %yield, ASL, vinyl acetate

&o

118a [75]

H 92 % ee, 50 %yield, PFL, vinyl acetate

118b [75] H

100 % ee, 47 % yield

119a [76] RO R = 2-Naphthylmethyl,CHZPh, TBDMS >99 % ee, 47-50 % yield, PCL, vinyl acetate

120a[77]

119b [76] RO >99 % ee, 47-49 % yield

0

12Ob [77]

a Ack

B H >99 % ee. 47 % yield

>99 % ee, 48 % yield, PSL, vinyl acetate

H?

H

121a[78]

l 2 l b [78]

0 ; o

H

H 87 % ee, 45 % yield, PCL, acetic anhydride

dQ

95 % ee, 42 % yield

0

OAc

>99 % ee, -, PCL, vinyl acetate, 45 % conversion

122a[791

d,D

l22b [79]

_-

OH

I

541

542

I

1 7 Hydrolysis and Formation o f C - 0 Bonds

(cont.).

Table 11.121.

~~

do.

OH (OAc) 6 /0

/ &cOAc

123a [86] 100 % ee, 38 % yield PCL, vinyl acetate

A

c

123b [86] 100 % ee, 36 %yield

(OH)

123c [80] 100 % ee, 10 %yield

Brv OAc

Br"'"

R

dAc

OAc

R

H Br Me

>98 % ee, 50 % yield 94 % ee, 49 % yield 87 % ee, 50 % yield all PCL, vinyl acetate

124a 125a 12Ga

92 % ee, 50 % yield 87 % ee, 50 % yield 85 % ee, 50 % yield

~ A c

"'-4

124b [81] 125b [81] 126b [81]

OH

127a [81]

Br

OAC OTBDMS >98 % ee, 47 % yield, CAL-B vinyl acetate

AAC

ATBDMS

92 % ee, 50 % yield

128b [82]

128a [82] HO 95 % ee, 49 % yield

96 % ee, 49 % yiel, PCL, vinyl acetate

cdj

129a [83]

129b [83]

OH

OAc

>99 % ee, 45 %yield, CAL-B, vinyl acetate, 50 "C 1 K. Fritschke, C . Syldatk, C. Wagner,

>99 % ee, 46 % yield -

H. Hengelsberg, R. Tacke, Appl. Microbial Biotechnol. 1989,31, 107. 2 K. Laumen, D. Breitgoff, M. P. Schneider, j.Chem. Soc.. Chem. Commun. 1988,1459.

3 M. Inagaki, J. Hiratake, T. Nishioka, I. Oda, Agric. Bid. Chem. 1989,53,1879. 4 S. Takano, M. Suzuki, K. Ogasawara, Tetrahedron: Asymmetry1993,4,1043.

11. I Hydrolysis and Formation ofcarboxylid Acid Esters

I

543

5 H. F r y h a n , N. Ohmer, T. Norin, K. Hult, Tetrahedron Lett. 1993,34, 1367. 6 B. D. Johnson, B. Morgan, A. C. Oehlschlager, S . Ramaswamy, Tetrahedron:Asymmetry 1991, 2, 377. 7 R. Bovara, G. Canea, L. Ferrara, S . Riva, Tetrahedron:Asymmetry 1991,2,931. 8 D. B. Berkowitz, S . J. Danishefsky, Tetrahedron Lett. 1991, 32, 5497. 9 D. B. Berkowitz. S . J. Danishefsky, G. K. Schulte,]. Am. Chem. Soc. 1992,114,4518. 10 M. Sato, H. Ohuchi, Y. Abe, C. Kaneko, Tetrahedron:Asymmetry 1992, 3, 313. 11 L. Ling, Y. Watanabe, T. Akiyama, S . Ozaki, Tetrahedron Lett. 1992, 33, 1911. 12 S. Takano. T. Yamane, M. Takahashi, K. Ogasawara, Synlett, 1992,410. 13 S. Takano, T. Yamane, M. Takahashi, K. Ogasawara, Tetrahedron:Asymmetry 1992, 3, 837. 14 M. Meltz, N. S. Saccomano, Tetrahedron Lett. 1992, 33, 1201. 15 J. A. Gaboury, M. P. Sibi,]. Org. Chem. 1993,58, 2173. 16 T. Isumi, F. Tamura, Bull. Chem. Soc. Jpn. 1992, 65, 2784. 17 F. Theil, S. Ballschuh, Tetrahedron:Asymmetry 1996,7, 3565. 18 W. Adam, C. Mock-Knoblauch, C. R. Saha-Moller, Tetrahedron:Asymmetry 1997,8, 1441. 19 T Ema, S . Maeno, Y. Takaya, T. Sakai, M. Utaka, J. 0%.Chem. 1996,61,8610. 20 a) C. R. Johnson, B. M. Nerurkar, A. Golebiowski, H. Sundram, J. L. Esker, J. Chem. Soc., Chem. Commun. 1995,1139 b) T. Biadatti, J. L. Esker, C. R. Johnson, Tetrahedron:Asymmetry 19967, 2313. 21 T Sugahara, Y. Kuroyangi, K. Ogasawara, Synthesis 1996,1101. 22 H. Nagashima, M. Sato, T. Taniguchi, K. Ogasawara, Splett 1999,1754. 23 a) T. T. Curran, D. A. Hay, Tetrahedron:Asymmetry 1996,7,2791;b) T. T. Curran, D. A. Hay, C. P. Koegel, Tetrahedron1997,53,1983. 24 K. Kato, H. Suzuki, H. Tanaka, T. Miyasaka, M. Baba, K. Yamaguchi, H. Akita, Chem. Pharm. Bull. 1999,47,1256. 25 M. J. Mulvihill, J. L. Gage, M. J. Miller, ]. Org. Chem. 1998,63, 3357. 26 V. Merlo, S . M. Roberts, R. Storer, R. C . Bethell,J. Chem. Soc., Perkin Trans. 11994, 1477. 27 D. M. Coe, A. Garofalo, S . M. Roberts, R. Storer, A. J. Thorpe, ]. Chem. Soc., Perkin Trans. 1 1994, 3061. 28 S. Takano, 0. Yamada, H. Iida, K. Ogasawara, Synthesis 1994, 592. 29 A . Maestro, C. Astorga, V. Gotor, Tetrahedron: Asymmetry 1997,8,3153. 30 A. Luna, C. Astorga, F. Fiilop, V. Gotor, Tetrahedron:Asymmetry 1998, 9,4483. 31 E. Fon6, F. Fiilop, Tetrahedron:Asymmetry 1999, 10, 1985.

32 E. Forro, L. T. Kanerva, F. Fiilop, Tetrahedron: Asymmetry 1998,9,513. 33 E. Forro, 2. Szakonyi, F. Fiilop, Tetrahedron: Asymmetry 1999, 10,4619. 34 D. F. Taber, K. Kanai, Tetrahedron1998,54, 11767. 35 B. E. Carpenter, I. R. Hunt, B. A. Keavy, Tetrahedron:Asymmetry 1996,7, 3107. 36 P. Crotti, V. Di Bussolo, L. Favera, F. Minutolo, M. Pineschi, Tetrahedron:Asymmetry 1996, 7, 1347. 37 M. Panunzio, R. Camerini, A. Mazzoni, D. Donati, C. Marchioro, R. Pachera, Tetrahedron:Asymmetry 1997, 8, 15. 38 M. Barz, H. Glas, W. R. Thiel, Synthesis 1998, 1269. 39 P. Noheda, G. Garcia, M. C. Pozuelo, B. Henadon, Tetrahedron:Asymmetry 1996,7, 2801. 40 H:J. Gais, C. Griebel, H. Buschmann, Tetrahedron:Asymmetry 2000, 11, 917. 41 T. Yamane, K. Ogasawara, Synlett 1996,925. 42 C. Sanfilippo,A. Patti, G. Nicolosi, Tetrahedron: Asymmetry 2000, 11,1043. 43 K. Lemke, S. Ballschuh, A. Kunath, F. Theil, Tetrahedron:Asymmetry 1997,8, 2051. 44 M. A. Djadchenko, K. K. Pivnitsky, F. Theil, H. Schick,J. Chem. Soc., Perkin Trans. 1 1989, 2001. 45 J . P. Bamier, V. Rayssac, V. Morisson, L. Blanco, TetrahedronLett. 1997, 38, 8503. 46 J. P. Bamier, V. Morisson, 1. Volle, L. Blanco, Tetrahedron:Asymmetry 1999, 10, 1107. 47 M. C. R. Franssen, H. Jongejan, H. Kooijman, A. L. Spek, R. P. L. Bell, I. B. P. A. Wijnberg, A. de Groot, Tetrahedron:Asymmetry 1999, 10, 2729. 48 K. Mori, A. Horinaka, M. Kido, Liebigs Ann. Chem. 1994,817. 49 T Yoshimitsu, Y. Oshiba, K. Ogasawara, Synthesis 1994,1029. 50 T. Imori, 1. Azumaya, Y. Hayashi, S . Ikegami, Chem. Pharm. Bull. 1997,45, 207. 51 W.Adam, M. T. Diaz, R. T. Fell, C. R. Saha-Moller, TetrahedronLett. 1996,7, 2207. 52 M. Takahashi, K. Ogasawara, Synthesis 1996,954. 53 A. K. Gosh, J. F. Kincaid, M. G. Haske, Synthesis 1997,541. 54 A. Luna, A. Maestro, C. Astorga, V. Gotor, Tetrahedron:Asymmetry 1999, 10, 1969. 55 Y. Igarashi, S . Otsutomo, M. Harada, S . Nakano, S. Watanabe, Synthesis 1997, 549. 56 Y. Fujiwara, T. Yamato, T. Bando, K. Shishido, Tetrahedron:Asymmetry 1997, 8, 2793. 57 N. Yoshida, T. Kamikubo, K. Ogasawara, Tetrahedron Lett. 1998, 39, 4677. 58 N. Yoshida, H. Konno, T. Kamikubo, M. Takahashi, K. Ogasawara, Tetrahedron:Asymmetry 1999, 10, 3849. 59 K. Hiroya, H. Zhang, K. Ogasawara, Synlett 1999, 529. GO K. Tanaka, K. Ogasawara, Synthesis 1995,1237. 61 T. Taniguchi, R. M. Kanada, K. Ogasawara, Tetrahedron:Asymmetry 1997, 8, 2773.

544

I

1 I Hydrolysis and Formation of C - 0 Bonds 62 V. E. U.Costa, 1. Alifantes, 1. E. D. Martins, 74 J. Matsubara, K. Kitano, K. Otsubo, Y. Kawano, T. Tetrahedron: Asymmetry1998, '9 2579. Ohtani, M. Bando, M. Kido, M. Uchida, F. Tabusa, 63 K. Hirayama, K. Mori, Eur. J. Org. Chem. 1999, Tetrahedron 2000, 5G,4667. 2211. 75 R. A. MacKeith, R. McCague, H. F. Olivo, S . M. 64 L. Aribi-Zouioueche, J:C. Fiaud, Tetrahedron Lett. Roberts, S. 1. C. Taylor, H. Xiong, Bioorg. Med. 2000,41,4085. Chem. 1994,2, 387. 65 P. Camps, S. Gimenez, M. Font-Bardia, X. Solans, 76 T. Taniguchi, M. Takeuchi, K. Kadota, A. S . Tetrahedron: Asymmetry1995, 6,985. ElAzab, K. Ogasawara, Synthesis 1999,1325. 66 1. D. Scott, R. M. Williams, Tetrahedron Lett. 2000, 77 K. Kadota, A. S . Ehzab, T. Taniguchi, K. 41, 8413. Ogasawara, Synthesis 2000, 1372. 67 a) H. Sakagami, T. Kamikubo, K. Ogasawara, 78 A. K. Ghosh, Y. Chen, Tetrahedron Lett. 1995, 36, Synlett 1997,221; b) H. Sakagami, K. Ogasawara, 505. Synthesis 2000, 521 79 A. Kamal, K. V. Ramana, M. V. Rao, /. 0%.Chem. 68 N. Mase, T. Nishi, Y. Takamori, H. Yoda, K. 2001, G6, 997. Takabe, Tetrahedron: Asymmetry1999, 10,4469. 80 S . Nakano, Y. Igarashi, H. Nohira, Tetrahedron: 69 K. Takabe, M. Suzuki, T. Nishi, M. Hiyoshi, Y. Asymmetry2001,12,59. Takamori, H. Yoda, N. Mase, Tetrahedron Lett. 81 J. Gu, M. I. Heeg, C. R. Johnson, Tetrahedron Lett. 2000,41,9859. 2001,42,1213. 70 K. Sugawara, Y. Imanishi, T. Hashiyama, 82 H. Nagata, N . Miyazawa, K. Ogasawara, Syrtthesis Tetrahedron: Asymmetry2000, 11,4529. 2000,2013. 71 0. Yamada, K. Ogasawara, Synthesis 1995, 1291. 83 F. Fernindez, X. Garcia-Mera, 1. E. 72 T. Sugai, H. Ikeda, H. Ohta, Tetrahedron 1996, 52, Rodriguez-Borges, Tetrahedron: Asymmetry2001, 8123. 12. 365. 73 M. Majerit, M. Gelo-Pujii-,V. Sunjit, A. Levai, ' P. Sebok, T. Timar, Tetrahedron: Asymmetry1995, 6, 937.

28, 29, 31-33, 41-65, 67-71, 83-89, 107-109, 127, 143, 144, 153, 154, 156, 157, 163-170,181-183,201) (Table 11.1-20).A very good illustration for the potential of enantiomer-differentiating acylation catalyzed by lipases is provided by the highyield synthesis of a series of aromatic cyanohydrin acetates (la-g) from aldehydes, acetone cyanohydrin and vinyl acetate in the presence of Pseudomonas cepacia lipase and a basic anion-exchangeresin in diisopropyl ether which proceeds under kinetic resolution coupled with in situ formation and racemization of the cyanohydrin representing a dynamic kinetic resolution. For further examples see Table 11.1-24. To the secondary aliphatic alcohols, which have been resolved into their enantiomers, belong a variety of hydroxy carboxylic esters and acids (35,100-102,125,126, 131-140, 150,151, 166,168-172, 174,182,183),some hydroxy ketones (128-130, 141) and a crown ether derivative (203)(Table 11.1-20). Even the tetraphenylporphyrin derivatives 204 and 205 were substrates for different lipases. Diketene is useful acyl donor also, yielding acetoacetates with very high enantiomeric excess (115,207-210). Monoacetates and alcohols of Table 11.1-20 which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-6 and 11.1-15. Table 11.1-21 lists cyclic secondary alcohols that have been synthesized by lipasecatalyzed enantiomer-differentiating acylation (1-1 29).The compounds that have been obtained by the alternative route of hydrolysis are listed in Table 11.1-16. The complementary nature ofthe two routes is obvious. For the series ofthe glycals 9-15, Pseudomonas cepacia lipase-catalyzedacylation works with good to high enantiomer selectivity and yield. myo-Inositol derivatives 17 and 18 may be prepared enantiomer-

7 7 . 7 Hydrolysis and formation ofCarboxylid Acid Esters

I

ically pure by Candida cylindracea lipase-catalyzedacylation with acetic anhydride in diethyl ether not only with high enantiomer but also with high group selectivity. Axial-chiral enantiomerically highly enriched binaphthols 4, which are highly useful chiral auxiliaries, are accessible either through acylation of the racemic diol with vinyl acetate or deacylation of the racemic diacetate with butanol (Table 11.1-22), both catalyzed by Pseudomonas cepacia lipase. Among the many other cyclic secondary alcohols that have been obtained by lipase-catalyzed enantiomer-selective acylation with high enantiomeric excess are aminohnctionalized cycloalkanols (40, 45-62, 75), bicyclo[3.3.0]octanoIs (78, 84-86), different types of tri- and tetracyclic alcohols (96104),substituted indanols (87-94,123),hydroxy lactams (106,109-112) and brominated cyclohexenol derivatives (74,77,124-127) (Table 11.1-21). Monoacetates and alcohols of Table 11.1-21 which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-6 and 11.1-16. 11.1.1.3 Inter- and Intramolecular Alcoholysis

Hydrolase-catalyzed enantiomer-differentiating alcoholysis of esters of racemic alcohols with achiral alcohols in organic solvents of low water content is a valuable alternative to hydrolysis (Table 11.1-22). Lipase-catalyzed enantiomer-differentiatinginter- and intramolecular alcoholysis of acylated alcohols and lactones in organic solvents may most advantageously be used instead of hydrolysis in aqueous solution in those cases where insufficient stability, high solubility or low functional group selectivity is observed or may be anticipated in the latter case (1-16)(Table 11.1-22). For lipase-catalyzed intermolecular alcoholysis as alcohols, by and large more lipophilic ones such as n-propanol, n-butanol, n-hexanol, n-octanol, cyclohexanol or benzylalcohol are used whereas methanol (44)or ethanol (52,53,61) are used rarely. Typical solvents are n-hexane, diisopropyl ether, tert-pentyl alcohol, toluene, tetrahydrofuran or acetonitrile. In many cases the enantioselectivity and yield are higher for the alcoholysis than for the hydrolysis catalyzed by one and the same lipase, provided that a large excess of the alcohol is used. Enantiomer-differentiating alcoholysis of an acylated thiol (10)has also been described. Alcoholysis of y-and p-lactones gives access to enantiomerically pure y-hydroxyesters and y-lactones (17-24)and P-hydroxyesters and P-lactones (55-63),respectively. Enantiomerically pure enol acetates 67a and the y-acetoxybutenolide 51/ent-51 have been obtained by hydrolysis of the corresponding racemic substrates. As well as the above-described intermolecular alcoholysis of esters, the intramolecular version has been successfully utilized for the synthesis of lactones from racemic hydroxy carboxylic acid esters (25-41,64-66) (Table 11.1-22).High selectivity in the pig pancreas lipase-catalyzedenantiomer-differentiating lactonization of yhydroxy carboxylic acid esters with formation of butyrolactones substituted in

545

546

I

I I Hydrolysis and Formation of C - 0 Bonds

Lipase-catalyzedenantiorner- and enantiotopos-differentiatinginter- and intramolecular alcoholysis o f esters and lactones in organic solvents (PCL Pseudomonas cepacia lipase, PSL Pseudomonas sp. lipase, PPL pig pancreas lipase, M M L Mucor miehei lipase, HLL, Humicola lanuginosa lipase, PFL PseudomonasPuorescens lipase, CCL Candida cylindracea lipase, CAL-B Candida antarctica B lipase, CRL Candida rugosa lipase, PRL Penicillium roqueforti lipase, CAL-A+BCandida antarctica A+B lipase).

Table 11.1-22.

295 % ee, 45 % yield 295 % ee, 45 % yield 295 % ee, 45 % yield all PCL

BuOH BuOH/i-Pr20 H2O

90 % ee, 45 % yield 295 % ee, 45 % yield 88 % ee, 45 % yield

2b [*I

OH &NHCOnPr Ph

Phr r H C O n P r

295 % ee, 34 % yield 295 % ee, 39 % yield 295 % ee, 42 % yield 295 % ee, 43 % yield 295 % ee, 45 % yield 295 % ee, 36 % yield 295 % ee, 44 % yield 295 % ee, 43 % yield all PCL

BuOH, t-pentanol BuO H , toluene HexOH, toluene OctOH, toluene BuOH, n-Bu2O HexOH, n-Bu2O BuOH, THF BuOH, MeCN

295 % ee, 33 %yield 295 % ee, 40 % yield 295 % ee, 46 % yield 295 % ee, 41 % yield 295 % ee, 39 % yield 295 % ee, 41 % yield 79 % ee, 45 % yield 85 % ee, 46 % yield

3a [31

PhN?

95 % ee, 47 %yield, PCL n-PrOH CCL, t-pentanol, i-PrzO

OAc

3b [31

95 % ee, 50 %yield

OCOEt

R

L

O

T

R A O T s

S

R

ClCH2 n-Bu n-CloH21 Ph PhOCH2

96 % ee,-, PFL, BuOH, n-hexane 90 % ee,-, PFL, BuOH, n-hexane 298 % ee,-, PFL, BuOH, n-hexane 95 % ee,-, CCL, BuOH, n-hexane 84 % ee,-, HLL, BuOH, n-hexane

4a 5a Ga

7a 8a

96 % ee,-, 31 % ee,43 % ee,70 % ee,42 % ee,-

4b [41 5b [41 61, [41 7b [41 8b 141

1 1 . 1 Hydrolysis and Formation ofCarboxylid Acid Esters Table 11.1-22.

(cont.).

R = H, F, C1, Br, OMe 97-99 % ee, CCL, BuOH, i-Pr2O (fromthe butyrate) lob [7]

10a [7]

Me

Me

95 % ee, 42 % yield

88 % ee, 39 %yield, n-PrOH, PPL,

hexane

Oh

CICH2C00.,, " O S R

v

R

H Me Et n-Bu n-CbH13

79 % ee. 44 %yield 72%ee,80 % ee,86 % ee,66 % ee,all MML, PrOH, i-PrzO

lla

12a 13a 14a 15a

84 % ee, 45 % yield 95 % ee,91 % ee,90 % ee,86 % ee,-

Ilb [S] 12b [8] 13b [8] 141,[8] 15b [8]

16b [9]

98 % ee, 47 %yield, PSL, BuOH

96 % ee, 50 % yield

R

RQO

R

n-C5HII 70 % ee, n-C6H13 90 % ee, n-C7H15 93 % ee, n-CsH17 298 % ee, n-C9Hl9 298 % ee, n-CloHzl 298 % ee, n-C11H23 298 % ee, n-ClzHzs 298 % ee, all PPL,n-PrOH

17a 18a 19a 20a 21a 22a 23a 24a

55 % ee, 78 % ee, 80 % ee, 77 % ee, 71 % ee, 74 % ee, 76 % ee, 77 % ee, -

1%

[lo]

18b [lo] 19b [lo] 20b [lo] 21b [lo] 22b [lo] 23b [lo] 24b [lo]

I

547

548

I

7 7 Hydrolysis and Formation of C-0 Bonds Table 11.1-22.

(cont.).

298 % ee, 88 % ee, 82 % ee, 91 % ee, 92 % ee, 94 % ee, 88 % ee, 94 % ee, all PPL, Et20 or hexane or THF (intramolecular lactonization of the methyl ester)

78 % ee, -, PPL, Et2O (from methyl ester)

R

n-C5HI1 n-C15H,,

R

Ph c-C6HI3

25 [11] 26 [ l l ] 27 [ l l ] 28 [ll] 29 [11] 30 [ l l ] 31 [ll] 32 [11]

R = Me, Et, PhCHz 295 % ee, -, PPL, Et2O

86 %ee, 25 %yield 298 % ee, 15 %yield, PPL, Et20 (from the pentyl ester)

35 [13] 36 [13]

298 % ee, 35 % yield 298 % ee, 35 % yield all PPL, Et2O

37 [14] 38 [14]

1 1 . 1 Hydrolysis and Formation ofcarboxylid Acid Esters

(cont.)

Table 11.1-22.

U x-Y

CHzCHz 299 % ee, 14 %yield (E)-CH=CH 299 % ee, 18 %yield (2)-CH=CH 98 % ee, 20 %yield PSL,isooctane, molecular sieves

39 (151 40 [15] 41 I151

GBr

421, [lG]

/

94 % ee, 47 %yield, CAL-B, cyclohexanol

03

92 % ee, 49 % yield

NHAc

43a (171

,mttOH

/

>99 % ee, 43 %yield, CAL-B, n-BuOH

43b [17] OAc

>99 % ee, 48 % yield

44b [18]

>99 % ee, 44 % yield, PCL, MeOH

80 % ee, 55 % yield

/

R

>99 % ee, -, PCL, n-BuOH, 45a 49 % conversion 3-OPh 98 % ee, -, CAL-B, n45a PrOH, 45 % conversion R = H, 3-Me, 4-Me, 3-OMe, 4-OMe, 3-C1,4-C1 82-98 % ee, -, CAL-B, n46a PrOH, 45-53 % conversion 3-OPh

97 % ee, -

45b [19]

79 % ee, -

45b [20]

79- >9 % ee, -

46b [20]

I

549

550

I

1 1 Hydrolysis and Formation ofC-0 Bonds Table 11.1-22.

(cont.).

4% [21]

Ph F

82 % ee, -, CAL-B, n-BuOH, 34 % conversion

0

A

48b [22]

48a [22]

? H

AcO

o

13 % ee, 37 % yield

96 % ee, 60 %yield, PFL, n-PrOH

NcwB ~ A c

NcTBr 49b [23]

49a [23]

/

H2N

H*N

CI

CI

99 % ee, 42 % yield

86 % ee, 50 % yield, CAL-B, n-BuOH

50 [24]

>95 % ee, 93 %yield, PSL, n-BuOH

AcO ' " ' G O

51a [25] racemate

"$N-Bn

51b [25]

"R2g N - B n

:

OAc

OH

R'

'R

Me

H

H

H

ent-5lb [25] 70-98 % ee, -, CCL, lipase PSL, PRL, CRL, PCL, nBuOH, 49-61 % conversion

>99 % ee, 45 %yield, PCL, EtOH >99 % ee, 26 % yield, PCL, EtOH

521

53a

90 % ee, 49 %yield 43 % ee, 62 % yield

52b (261

53b [26]

6 6

Table 11.1-22.

*\\OAc

*“oAc 54b [27] 6

54a [27]

:

OAc

O

A

OAc

c

5 k [27]

“‘OAc

OH

>98 % ee, 24 % yield, CCL, n-BuOH

OAc

68 % ee, 58 % yield

-, 18 %yield

R’T C 0 2 R 2

6H R’

R2

Me

CH2Ph

n-Pr CH2Ph

i-Pr

CH2Ph

96 % ee, 36 % yield 75 % ee, 42 % yield 95 % ee, 41 % yield

85 % ee, 51 %yield, PPL, 55a PhCH20H 69 % ee, 45 %yield, PCL, 56a PhCHzOH 90 % ee, 43 %yield, PCL, 56a PhCHzOH

OH 72 % ee, 24 % yield, PCL, PhCHzOH

55b [28] 56b [28] 56b [28]

5% [28b]

57a [28b]

(-‘CO,Bn

70 % ee, 38 % yield

R’ R2Y C O 2 B n OH R’

R2

n-Pr 87 % ee, 34 % yield, PCL, PhCHzOH n-Pr Me 98 % ee, 24 % yield, PCL, PhCH2OH Me

58a 59a

92 % ee, 50 % yield 79 5% ee, 16 %

yield

58b [28b] 59b [28b]

6Ob [28b]

84 % ee, 39 % yield, PCL, PhCHzOH

I

551

(cont.).

OAc

I

1 1 . 7 Hydrolysis and Formation ofcarborylid Acid Esters

85 % ee, 13 %yield

552

I

I 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-22.

(cont.).

81 % ee, 40 % yield, PCL, EtOH 73 % ee, 36 %yield,

Et

n-C6Hls

Gla

97 % ee, 43 %yield

Gla

>99.9 % ee, 39 %yield Glb [29]

Glb [29]

PCL, n-CGH13OH

9,

G2b [30]

C0,Bn

OH

95 % ee, -, CAL-B, PhCHzOH, 51 % conversion

131

G3b [30]

CO,Bn

:

OH 99 % ee, -, CAL-B, PhCH20H, 50 % conversion

"

O

99 % ee. -

U

>99 % ee, -

0 R

R R

E-CH=CH-Ph

C=CH-Ph

80 % ee, 33 %yield, G4a pancreatin, >99 % ee, double resolution 61 % ee, -, CAL-A+B, 98 % 65a ee, by crystallization from the reaction mixture

-, 47 % yield

G4b [31]

70 % ee, 30 %

6% [32]

yield

GGb [33] I Ph 96 % ee, 28 % yield,

CAL-A+B

I Ph - -

7 7 . 1 Hydrolysis and Formation of Carboxylid Acid Esters

I

553

Table 11.1-22.

(cont.).

6N

A

67a [34]

67b [34]

Ar%N

100 % ee, 38 % yield (Ar =

Ph), 30 %yield (Ar = 3,4C12Ph),PFL, n-BuOH

HOA

O

A

c

R

Ts Cbz

95 % ee, 76 %yield, PCL, n-BuOH 72 % ee, 56 % yield, CCL, n-BuOH 98 % ee, 68 % yield, PCL, n-BuOH 49 % ee, 30 %yield, PPL, n-BuOH

68 [35] 69 [35]

70a [36] O

G

O

70b [36] '-8OAc

H

racemic

>99 % ee, 65 %conversion, PCL, n-BuOH, n-hexane

O

O

O

A

C

enb70b 1361

>99 % ee, 57 % conversion, CCL, n-BuOH, n-hexane 1 H. S. Bevinakatti, A. A. BanejiJ. Org. Chem. 1991,565372. 2 L. T. Kanerva, K. Rahiala, E. Vanttinen, J . Chem. SOC.,Perkin Trans. 11992, 1759. 3 D. Bianchi, A. Bosetti, P. Cesti, P. Golini, Tetrahedron Lett. 1992, 33, 3231. 4 C:S. Chen, YC : . Liu, Tetrahedron Lett. 1989, 30, 7165. 5 7. Miyazawa, S . Kurita, S. Ueji, T. Yamada, S. Kuwata,]. Chem. Soc., Perkin Trans. 11992, 2253. G H. S. Bevinakatti, A. A. Baneji, R. V. Newadkar, j . Org. Chem. 1989, 54,2453. 7 D. Bianchi, P. Cesti,]. Org. Chem. 1990,55, 5657. 8 J:B. Barnier, L. Blanco, G. Rousseau, E. Guibe-Jampel,I. Fresse,j. Org. Chem. 1993,58, 1570. 9 a) Y Tamai, T. Nakano, S. Koike. K. Kawahara, S. Miyano, Chem. Lett. 1989, 1135; b) M. Inagaki,

10 11 12 13 14 15 1G 17

J. Hiratake, T. Nishioka. J. Oda, Agnc. Bid. Chem. 1989,53,1879. M. Huffer, P. Schreier, Tetrahedron: Asymmetry 1991, 2, 1157. A. L. Gutman, K. Zuobi, T. Bravdo, j . Org. Chem. 1990,55,3546. T. Sugai, S. Ohsawa, H. Yamada, H. Ohta, Synthesis 1990, 1112. C. Bonini, P. Pucci, R. Racioppi, L. Viggiani, Tetrahedron: Asymmetry 1992, 3, 29. C. Bonini, P. Pucci, L. Viggiani,J. Org. Chem. 1991,56, 4050. H. Yamada, S. Ohsawa, T. Sugai, H. Ohta, S . Yoshikawa, Chem. Lett. 1989, 1775. Y. Igarashi, S. Otsutomu, M. Harada, S. Nakano, Tetrahedron: Asymmetry 1997, 8, 2833. A. T. Anilkumar, K. Goto, T. Takahashi, K. Ishizaki, H. Kaga, Tetrahedron: Asymmetry 1999, 10, 2501.

554

I

1 1 Hydrolysis and Formation ofC-0 Bonds 18 K. Tanaka, K. Ogasawara, Synthesis 1995,1237. S. Ageishi, H. Isobe, Y. Hayashi, Y. Yamamoto, 19 J. Roos, U. Stelzer, F. Effenberger, Tetrahedron: /. Chem. Soc., Perkin Trans. 12000, 71. Asymmetry 1998,9,1043. 29 T. Ito, M. Shimizu, T. Fujisawa, Tetrahedron 1998. 20 U. Hanefeld, Y. Li, R. A. Sheldon, T. Maschmeyer, 54, 5523. Synktt 2000, 1775. 30 W. Adam, P. Groer, C. R. Saha-Moller,Tetrahedron: 21 R. Morrone, G. Nicolosi, A. Patti, M. Piatelli, Asymmetry 1997,8,833. Tetrahedron:Asymmetry 1995, 6. 1773. 31 B. Henkel, A. Kunath, H. Schick, Liebigs Ann. 22 M. Pallavicini, E. Valoti, L. Villa, 0. Piccolo, /. Org. Chem. 1992,809. Chem. 1994,59,1751. 32 B. Henkel, A. Kunath, H. Schick, Tetrahedron: 23 S. Conde, M. Fierros, M. I. Rodriguez-Franco, Asymmetry 1994,5,17. C. Puig, Tetrahedron: Asymmetry 1998, 9, 2229. 33 B. Henkel, A. Kunath, H. Schick, Tetrahedron: 24 M. Ranchoux, J.-M. Brunel, G. lacazio, G. Buono, Asymmetry 1993,4,153. Tetrahedron: Asymmetry 1998, 9, 581. 34 a) A. J. Carnell, M. L. Escudero Hernandez, 25 H. van der Deen, R. Hof, A. van Oeveren, A. Pettman, J. F. Bickley, Tetrahedron Lett. 2000, B. L. Feringa, R. M. Kellog, Tetrahedron Lett: 1994, 41, 6929: b) G. Allan, A. J. Carnell, M. L. Escudero 35, 8441. Hernandez, A. Pettman, j . Chem. Soc., Perkin 26 K. Takabe, M. Suzuki, T. Nishi, M. Hiyoshi, Trans. 12000, 3382. Y. Takamori, H. Yoda, N. Mase, Tetrahedron Lett. 35 K. Fuji, T. Kawabata, Y. Kiryu, Y. Sugiura, 2000,41,9859. Tetrahedron Lett. 1990, 31, 6663. 27 C. Sanfilippo, A. Patti, G. Nicolosi, Tetrahedron: 36 M. van den Heuvel, A. D. Cuiper, H. van der Asymmetry 1999,10,3273. Deen, R. M. Kellogg, B. L. Feringa, Tetrahedron 28 a) Y. Koichi, K. Suginaka, Y. Yamamoto,j . Chem. Lett. 1997, 38, 1655. Soc., Perkin Trans. 1 1995, 1645; b) N. Sakai,

4-position and the unchanged y-hydroxy carboxylic acid esters of opposite configuration were observed (25-34). Pig pancreas lipase in diethyl ether is the combination of choice. Formation of the corresponding monosubstituted y-valerolactones was unselective. y-Valerolactoneswith a hydroxyl group in 4-position however could be obtained with high selectivity from the corresponding dihydroxy carboxylic acid pentyl or methyl ester (35-38). In order to suppress the competition between the methanol formed during lactonization and the intramolecular hydroxyl group, reactions were run in the presence of molecular sieve. Otherwise, conversion and ee value of the lactone were poor because of the reversibility of the reaction. Interestingly, macrocyclic lactones may be prepared by this method too. Treatment of racemic ricinoleic acid methyl ester, its racemic trans-isomer and the saturated racemic derivative with Pseudomonas sp. lipase in isooctane in the presence of molecular sieve gave the corresponding (R)-configurated 13-membered lactones 39-41 (Table 11.1-22) in fair yields with high ee values. Acylated alcohols, alcohols and lactones of Table 11.1-22 which can be obtained with other hydrolases as such or of opposite configuration are contained in Tables 11.1-14to 11.1-16and Tables 11.1-19 to 11.1-21. Lipase-catalyzed enantiomer- and enantiotopos-differentiating alcoholysis may also be extended to carboxylic acid esters, anhydrides and oxazolin-2-ones (1-22) (Table 11.1-23).Alcoholysis of methoxy malonic acid dimethyl ester with benzyl alcohol catalyzed by Candida cylindracea lipase gave, at 50 % conversion, the mixed diester 2 with high enantioselectivity. At higher conversion the ee values are lower because of the reversibility of alcoholysis.The enantiomeric mixed diester ent-2 may be obtained by methanolysis of the corresponding dibenzyl ester. Through catalytic hydrogenolysis the monobenzyl ester can be converted into the corresponding acid. It remains to be shown if this is an alternative to the pig liver esterase or lipasecatalyzed hydrolysis of the corresponding prochiral diester (Table 11.1-2).

7 1.1 Hydrolysis and Formation ofcarboxylid Acid Esters

I

555

Lipase-catalyzed enantiomer- and enantiotopos-differentiating alcoholysis of carboxylic acid esters and anhydrides, alcoholysis or hydrolysis of oxazolin-2-ones, and esterification of carboxylic acids (PPL p i g pancreas lipase, PCL Pseudomonas cepacia lipase, ANL Aspergillus niger lipase, CSL Candida sp. lipase, Candida cylindracea lipase, CAL-B Candida antarctica B lipase, CRL Candida rugosa lipase). Table 11.1-23.

CI,CH,OOC

Po

,

PEG-O,C’

Me0,C

/J

289 % ee, 46 % yield

296 % ee, 43 % yield, PPL PEG, i-PrzO

HxCozMe 2 121 C0,Bn

Me0

ent-2 [2] HxCozBn C0,Me

Me0

absolute configuration

absolute configuration

unknown

unknown 90 % ee, -, CCL equlibrium (70 %)

296 % ee, -, CCL PhCHZOH, hexane ( f r o m the dimethyl ester)

MeOH, hexane, 50 % conversion (from the dibenzyl ester)

8 PI

R’ HO,C A C O , n B u

R’

R2

H

Me Et n-Pr i-Pr

H H H

H

C1

1b PI

CO,H C0,iBu

93%ee 87%ee 60%ee 76%ee 62%ee 65-95 % y i e l d PCL, BuOH, i-Pr20 V

3 [3, 41 4 [3,4] 5 [3,4] 6 [3,4] 7 [3,41

90 % ee, 72 % y i e l d CSL, i-BuOH, C-C~HI~

,PI h

f Ph

9 [GI

ent-9 [6]

99 % ee

556

I

7 7 Hydrolysis and Formation ofC-0 Bonds Table 11.1-23.

(cont.).

11

Ph

Ph

N H

CO,Me(H)

R

MezCH MezCHCHz MeS(CH2)2 2-Naphthylmethyl 4-MeCsH4CHz Ph PhCHz Ph(CHz)z Ph(CHz)3

+f

iBu0,C

19b [lo]

19a [lo] CO,H

HO,C

"'YY

C0,iBu

92 % ee, 29 % yield

74 % ee, 30 % yield, CAL-B, 2-methylpropanol

HO,C

10 [7-91 11 [7-91 12 [7-91 13 [7-91 14 [7-91 15 [7-91 1 G 17-91 17 [7-91 18 [7-91

77 % ee, 47 % yield (H2O) 78 % ee, 82 % yield 82 % ee, 31 %yield (H2O) 75 % ee, 90 % yield 66 % ee, 86 % yield 75 % ee, 46 %yield (H20) 69 % ee, 93 % yield 93 % ee, 61 %yield (H20) 84 % ee, 91 % yield all PCL, MeOH ( or HzO), t-BuOMe

20b [11]

20a [11] iBu0,C

C0,iBu

C0,iBu

90 % ee, 48 % yield

90 % ee, 40 % yield, CAL-B, 2-methylpropanol

21b [12]

21a 1121 iBu0,C

HO,C

CO,H

C0,iBu

99 % ee, 29 % yield

88 % ee, 28 %yield, CAL-B, 2-methylpropanol 22a [13]

/\/CO,CH=CH,

/yC02nC6H13 Ph Ph 74 % ee, 45 % yield

99 % ee, 38 %yield, CAL-B, n-hexanol

1

23a [14] R

82-90 % ee, -, CRL, hexadecan-1-01,25-39 % conversion. esterification

221, [ 131

&CO,H

30-50 % ee, -

23b [14]

17.1 Hydrolysis and Formation ofCarboxylid Acid Esters

I

557

Table 11.1-23.

(cont.).

24b [14]

RnCO,nC,,H,, 85-93 % ee, -, CRL, hexadecan-1-01, 15-33 % conversion. esterification

64 % ee, 58 %yield, CAL-B,

>98 % ee, 39 % yield

n-PrOH, HC(OnPr),

2Gb [15]

-CO,nBu 53 % ee, 65 %yield, CRL, n-BuOH, H C (0nBu)S 1 J. S. Wallace, K. B. Reda, M. E. Williams, C. J. Morrow,]. Org. Chem. 1990,55,3544. 2 A. L. Gutman, M. Shapira, A. Boltanski,]. Org. Chem. 1992,57,1063. 3 Y. Yamamoto, K. Yamamoto, T. Nishioka, 1. Oda, Agric. Bid. Chem. 1988,52,3087. 4 Y. Yamamoto, T.Nishioka, J. Oda, Tetrahedron Lett. 1988,29,1717. 5 R. Ozegowski, A. Kunath, H,Schick, Tetrahedron: Asymmetry 1993,4,695. 6 R.-L. Gu,1 . 4 . Lee, C. J. Sih, Tetrahedron Lett. 1992, 33, 1953. 7 J.2. Crich, R. Brieva, P. Marquart, R:L. Gu, S. Flemming, C. 1. Sih, j . Org. Chem. 1993,58,3252. 8 H. S. Bevinakatti,A. A. Baneji, R. V. Newadkar, A. A. Mokashi, Tetrahedron: Asymmetry1992,3, 1505.

>97 % ee, 35 % yield 9 H. S. Bevinakatti, R. V. Newadkar, A. A. Baneji, j . Chem. Soc., Chem. Commun. 1990,1091. 10 R. Ozegowski, A. Kunath, H. Schik, Liebigs Ann. Chem. 1996,1443, 11 R. Ozegowski, A. Kunath, H. Schick, Liebigs Ann. Chem. 1994,215. 12 R. Ozegowski, A. Kunath, H. Schick, Liebigs Ann. Chem. 1994,1019. 13 H. Yang, E. Henke, U. T. Bornscheuer, j . Org. Chem. 1999,64,1709. 14 B.-V. Nguyen, E. Hedenstrom, Tetrahedron: Asymmetry1999,10,1821. 15 R. Morrone, M. Piatelli, G. Nicolosi, Eur.]. Org. Chem. 2001,1441.

Alcoholysis of prochiral glutaric anhydrides under the usual conditions gives, with moderate selectivities, the monoesters 3-8. Lipase-catalyzed enantiomer-differentiating hydrolysis of racemic phenyl benzyl oxazolin-2-one in aqueous solution in combination with an uncatalyzed in situ racemization of the unchanged enantiomer of the heterocyclic system, with two different lipases, gives access to D- and L-N-benzoyl-phenylalanine9 and ent-9, respectively. Enantiomer-differentiating alcoholysis and in situ racemization in organic solvents in the presence or absence of added water under the catalysis of lipase can in some cases furnish amino acid derivatives (10-18)with good selectivities and yields.

558

I

11 Hydrolysis and Formation of C - 0 Bonds

During alcoholysisof racemic substituted glutaric acid anhydride one is faced with regio- and enantioselectivity.These two processes may not cooperate in a matching sense. Despite this fact, the monoalkyl glutarates 19-21 have been obtained with moderate to good enantiomeric excess by lipase-catalyzed alcoholysis of the corresponding anhydrides in the presence of Candida antarctica B lipase with 2-methylpropanol. Alcoholysis of alkyl carboxylates is due to the competition of the two alcohols characterized by reversibility and associated with low conversion and poor enantioselectivity. The alcoholysis of vinyl carboxylates in the presence of Candida antarctica B lipase with n-hexanol as demonstrated for 22 can be regarded as an alternative in order to overcome these difficulties. Esterification of carboxylic acids (25, 26) (Table 11.1-23) in the presence of an orthoester as water-trapping agent may have advantages.

11.1.2.1.2

Dynamic Kinetic Resolution

The success of an enzyme-catalyzed kinetic resolution is limited by the maximum chemical yield of 50% for each enantiomer. However, this drawback can be overcome by a process called dynamic kinetic resolution. The key idea of this principle is to racemize the slow reacting enantiomer continuously reproducing the faster one. In an ideal case at the end of the conversion one enantiomer is formed in 100% yield with 100% of enantiomeric excess [135-1371. The kinetic requirements for a dynamic kinetic resolution are shown in Scheme ll.l-1G[8b]. The in situ racemization can be achieved by different means either spontaneously or catalytically. Due to their chemical properties certain substrates may racemize spontaneously under the reaction conditions. Useful catalysts could be ordinary chemicals such as bases, transition metal complexes and in theory another type of biocatalyst. Having identified a suitable enzyme promoting the enantiomer-differentiating process by hydrolysis or alcoholysis of a carboxylic ester or by acylation of an alcohol one has to find the appropriate racemizing catalyst. Lipase and catalyst must tolerate each other; they must work under identical conditions. The product must be chemically and configurationally stable in the presence of the catalyst. Table 11.1-24 lists lipase-catalyzeddynamic kinetic resolutions by different means. 4-Substituted oxazolin-5-ones racemize spontaneously by hydrolysis or alcoholysis caused by enolization to yield amino acid derivatives as outlined in the transformations (I), (2) and (3). Triethylamine may promote this type of transformations as

(S)-substrate kR

’ks

kS

kr,,

”k R

(S)-product krac

”ks

Scheme 11.1-16.

Dynamic kinetic resolution

11.1 Hydrolysis and Formation ofCar6oxylid Acid Esters

I

559

Lipase-catalyzeddynamic kinetic resolution (PCL Pseudomonas cepacia lipase, PPL pig pancreatic lipase, A N L Aspetgillus niger lipase, MML Mucor miehei lipase, CAL-B Candida antarctica 8 lipase, PSL Pseudornonas sp. lipase, PFL Pseudomoasfluorescens lipase, CAL Candida antarctica lipase, not specified).

Table 11.1-24.

Reaction

Type

Racemization

Ph

R = i-Pr, Me2CHCH2,MeSCHzCH2,

2-Naphthyl-CH2,4-MePhCH2, Ph, PhCH2, Ph(CH2)2,Ph(CHz)3 R = Me2CHCH2:78 % ee, 82 % yield, PCL, MeOH; 90 % ee, 85 % yield, MeOH+H20 R = Ph(CHz),: 84 % ee, 91 %yield, PCL, MeOH 95 % ee, 76 % yield, MeOH+H20

hydrolysis

spontaneous

hydrolysis

spontaneous

alcoholysis

NEt3

alcoholysis

NEt3

0 Ph

99 % ee, -, PPL Bn

-

p

h

*

I

i

q

H

0

Ph

99 % ee, -, ANL

-

4

H

,Yo

Ph P A 0 CoznBu >99 % ee, 67 % yield, MML, n-BuOH, toluene

Bnwo NYo

B")iH HN

Ph Ph AO R = Me: 94 % ee, 79 % yield R = Et: 97 % ee, 82 % yield R = n-Pr: 97 % ee, 83 % yield

CO,R

( 5 ) [41

560

I

I J Hydrolysis and Formation ofC-0 Bonds Table 11.1-24.

(cont.). ~

Reaction

Type

~~

Racemization

R = n-Bu: 95 % ee, 81 %yield all CAL-B, toluene R = Me: 97 % ee, 71 %yield, CAL-B, THF R = Me: 98 % ee, 88 % yield, CAL-B, MeCN

Hop AcO ...,

acylation

spontaneous

(6)(51

R2

R' R2 R' R' = R2 = H; R' = R2= Me; R' = Me, R2 = H; R'= H, R2 = Me 78-86 % ee, 100 % yield, PCL, vinyl acetate

A$:o&++: OH

OH

OAc

a: acylation

OH

a: acetate 290 % ee, 45 %yield, alcohol >90 % ee. 39 % yield, PCL, vinyl acetate b: 98 % ee, 100 %yield, PCL, n-BuOH, CHzCl2

spontaneous

(8) [71

spontaneous at >40 "C

( 9 ) (81

76 % ee, >99 % conversion, PCL, vinyl acetate 79 % ee, 93 conversion, PSL, vinyl acetate

H

I

o

-

e

R

'

C

R' = Me, Et, R2 = COMe, COEt >99 % ee, 100 %yield, CAL-B, n-hexane/CHzClz

0

I

,

e

acylation

7 7. 7 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.1-24.

(cont.).

Reaction

OH RACN

---

TYPe

Racernization

acylation

basic ion-exchange resin (10)[9]

acylation

silica gel

hydrolysis

N(n-C8H17)3

OAc R

~

C

N

R = 3-PhOCsH4, Ph, 4-Cl-CsH4, ~,~-OCHZO-CG 2-naphthyl, H~, 1-naphthyl 70-96 % ee, 6 6 8 8 % yield, PSL, vinyl acetate

OH

-

R’vS,~~ OAc

R’ = COzMe, BnOCH2, AcOCHr R2 = n-Bu, Et3SiO(CH2)2,i-Pr, n-Octyl 87->95 % ee, 63-87 % yield, PFL, vinyl acetate, t-BuOMe

,,+SEt

phs+oH

96 % ee, >99 % conversion, PCL, HzO/toluene

n = 1: 97 % ee, 75 %yield, PCL, vinyl acetate n = 2: 99 % ee, 67 % yield, PCL, vinyl acetate

OCOR

Mitsunobu inversion R2/iR1

R = n-Pr, R’ = Me, R2 = n-C6H1:+ 94 % ee, 100 % yield, PPL, vinyl propionate R = Me, R’ = Me, R2 = Ph 97 % ee, 97 % yield, PCL, vinyl acetate R = n-Pr, R’ = Ph, R2 = CH2NCOnPr, 97 % ee, 100 %yield, PCL, n-PrOH R = Me, R’ = Aryl, R2 = CN 61-97 % ee, 68-92 %yield, PCL, n-PrOH

acylation acylation alcoholysis of the (S)-acetate alcoholysis of the (S)-acetate

(14) [13]

1

561

562

I

I I Hydrolysis and Formation of C - 0 Bonds Table 11.1-24.

(cont.).

Reaction

Ph

Type

Racernization

hydrolysis

PdClz(MeCN)z

Ph (15) ~ 4 1

96 % ee, 81 %yield, PFL

OAc

(16)[151

acylation Ph

Ph\i

80 % ee, 76 % conversion, PFL, vinyl acetate, ortho-phenanthroline, PhCOMe, KOH 98 % ee, GO % conversion, PFL, vinyl acetate, ortho-phenanthroline, PhCOMe

[Rh(cod)C1]2 &(OAc)4

OAc Ph

acylation

Ph\i

A

A

>99 % ee, 92 % yield, CAL-B, 4-C1-C6H40Ac, PhCOMe, t-BuOH

Rlx",,-

OAc ,lA

2

R

= 4-Br-C6H4, R2 = Me; R' = 1-Naphthyl, acylation R2 = Me; R' = 2-naphthy1, RZ = Me; R' = PhOCH2, R2 = Me; R' = c-C6H11,R2 = Me; R' = n-C&13, RZ = Me; R' = Ph, R2 = E t >98 % ee, 65-80 %yield, CAL-B, 4-CI-CsH40Ac, PhCOMe, t-BuOH R' = 4-OMe-C&4, R2 = Me 91 % ee, GO % yield, CAL-B, 4-Cl-CsH4 acetate, PhCOMe, t-BuOH R' = PhOCH2, R2 = CHiCl 79 % ee, 68 % yield, CAL-B, 4-C1-C6H40Ac, PhCOMe, t-BuOH

R'

A cf(17)

1 1 . 7 Hydrolysis and Formation of Corboxylid Acid Oters

I

563

Table 11.1-24.

(cont.).

Type

Racemization

n = 1, 2: >99 % ee, 65 and 77 %veld, CAL-B, 4-C1-C6H40Ac,PhCOMe, t-BuOH

OH

rac/meso-mixhlre (-5O:SO)

(R,R/meso) (7G26100:O) (3852 for X = CH2) X = CH2, (CH2)2,(CH2)3,(E)-CH=CH,1,3C6H4, 1,4-C6H4,2,6-pyridylene, CH2N(Bn)CH2 >96->99 % ee, 47-90 % yield, CAL-B, 4-Cl-C6H40AC,tOlUene

OH L C O , E t R

-

OAc

i\/CO,Et

(17)

acylation

A, cf

alcoholysis

Pd(PPh3)4dppf

R

R = Ph: 95 % ee, 73 %yield R = 4-OMe-C&: 99 % ee, 69 %yield R = PhCH2: 96 % ee, 75 % yield R = C-C~HII: 70 % ee, 71 %yield all PCL, ~ - C ~ - C ~ H ~ cyclohexane OAC,

R R = Ph, 4-cl-C~H4,4-Me-c~H4,2-Fu~1, 1-Naphthyl 97->99 % ee, 70-87 % ee, CAL or PCL, GPrOH, THF

(23)[211

564

I

I 1 Hydrolysis and Formation of C - 0 Bonds Table 11.1-24.

(cont.)

Reaction

Type

Racemization

acylation

racemase

B R = Ph, 4-Cl-C6H4,4-Me-C&4, 4-OMe-CsH4, t-Bu, 2-Fury1, 1-naphthyl, C-C~HII, CHz-CHMe2,i-Pr, n-Pr 95->99 % ee, 8 4 9 1 %yield, PCL, 4-Cl-CsH40Ac,CHzClz OH PhAC02H

-

OAc Ph*CO,H

(25)~ 3 1

>98 % ee, 80 %yield, 1. PSL, vinyl acetate, 2. mandelate racemase 1 J. 2. Crich, R. Brieva, P. Marquart, R.-L- Gu, 11 D. S . Tan, M. M. Giinter, D. G. Drueckhammer, J. Am. Chem. SOC. 1995, 117,9093. S. Flemming, C. J. Sih, J. Org. Chem. 1993, 58, 12 T. Taniguchi, R. M. Kanada, K. Ogasawara, 3252. Tetrahedron: Asymmetry 1997, 8, 2773. 2 R.-L. Gu, 1:s. Lee, C. J. Sih, Tetrahedron Lett. 1992, 13 E. Vanttinen, L. T. Kanerva, Tetrahedron: 33, 1953. 3 N. J. Turner, J. R. Winterman, R. McCague, Asymmetry 19956,1779. J. S. Parratt, S. J. C. Taylor, Tetrahedron Lett. 1995, 14 J. V. Allen, J. M. J. Williams, Tetrahedron Lett. 1996, 37, 1859. 36, 1113. 15 P. M. Dinh, J . A . Howarth, A. R. Hudnott, 4 S. A. Brown, M.-C. Parker, N. A. Turner, Tetrahedron: Asymmetry 2000, 11,1687. J. M. J. Williams, W. Harris, Tetrahedron Lett. 1996, 37, 7623. 5 J. W. J, F. Thuring, A. J. H. Klunder, G. H. L. 16 A. L. E. Larsson, B. A. Persson, J:E. Backvall, Nefkens. M. A. Wegman, B. Zwanenburg, Angew. Chem. 1997, 109,1256; Angew. Chem. lnt. Tetrahedron Lett. 1996, 37,4759. Ed. Engl. 1997, 36, 1211. 6 J. W. J. F. Thuring, G. H. L. Nefkens, M. A. Wegman, A. J. H. Klunder, B. Zwanenburg, 17 B. A. Persson, A. L. E. Larsson, M. Le Ray, J.-E. J. Org. Chem. 1996, 61, 6931. Backval1.J. Am. Chem. SOC.1999, 121, 1654. 7 M. van den Heuvel, A. D. Cuiper, H. van der 18 B. A. Persson, F. F. Huerta, J:E. Backval1.J. Org. Chem. 1999,64,5237. Deen, R. M. Kellog, B. L. Feringa, Tetrahedron Lett. 1997, 38. 1655. 19 F. F. Huerta, Y R. S. Laxmi, J.-E. Backvall, Org. Letters 2000, 2, 1037. 8 A. D. Cuiper, M. L. C. E. Kouwijzer, P. D. J . Grootenhuis, R. M. Kellog, B. L. Feringa, J. Org. 20 F. F. Huerta, J:E. Backvall, Org. Letters 2001, 3, 1209. Chem. 1999,64,9529. 21 Y. K. Choi, 1. H. Suh, D. Lee, I.T. Lim, 1.Y. lung, 9 a) M. Inagaki, J. Hiratake, J , Oda,J. Am. Chem. M:J. Kim,J. Org. Chem. 1999, 64, 8423. SOC. 1991, 113,9360 b) M. Inagaki, J. Hiratake, 22 D. Lee, E.A. Huh, M.-J. Kim, H. M. Jung, J. Oda. j.Org. Chem. 1992, 57, 5643. J. H. Koh, J. Park, Org. Letters 2000, 2, 2377. 10 S. Brand, M. F. Jones, C. M. Raper, Tetrahedron Lett. 1995, 36,8493. 23 U. T. Strauss, K. Faber Tetrahedron:Asymmetry 1999, 10,4079.

7 1.1 Hydrolysis and Formation of Carboxylid Acid Esters

shown for (4) and (5). The hemiacetal and hemiaminal derivatives formed by the reactions (6)-(9) have been obtained by lipase-catalyzed acylation of the configurationally unstable hemiacetals or-aminal, respectively, or by alcoholysis of an acylated hemiacetal as shown for (7).The latter case is not a dynamic kinetic resolution but a normal kinetic resolution (acylation) followed by a spontaneous epimerization by lipase-catalyzed alcoholysis. Remarkably, the non-acylated hemiaminals are configurationally stable at temperatures below 40 "C and dynamic kinetic resolution proceeds according to reaction (9) at higher temperatures. Cyanohydrins are unstable under basic conditions regenerating the starting materials aldehyde and hydrogen cyanide. This property was used to prepare enantiomerically enriched cyanohydrin acetates according to transformation (10) by reaction of the corresponding aldehydes with acetone cyanohydrin in the presence of vinyl acetate, Pseudomonas sp. lipase and a strong basic ion-exchange resin as racemizing catalysts. The latter transformation demonstrates that lipase-catalyzed-acylationof one of the cyanohydrin enantiomers is faster than conversion of the cyanohydrin into aldehyde and hydrogen cyanide. The 2-acetoxysulfides are obtained according reaction (11)by in situ formation of the configurationally unstable hemithioacetals and subsequent lipase-catalyzed acylation. Racemization of the hemithioacetals was achieved by silica gel. The 2-phenylthiocarboxylicacid was formed by hydrolysis of the corresponding thioester in the presence of trioctylamine [reaction (141. The formation of the 2-acetoxy ketone in the reaction (13)was achieved by shifting an enediol-hydroxyketoneequilibrium with triethylamine. The formation of the enantiomericallyenriched or pure esters by reaction (14)was not a result of a dynamic process but a one-pot two-step procedure consisting of lipase-catalyzedresolution and a subsequent inversion of the slow reacting enantiomeric alcohol by Mitsunobu reaction. Enantiomerically pure allylic alcohols were prepared by a dynamic kinetic resolution in the transformations (15) and (23)by lipase-catalyzedhydrolysis or alcoholysis of the corresponding acetates in the presence of palladium complexes racemizing the slow reacting enantiomeric acetates. 1-Phenylethanol was converted in an acylation process into the corresponding acetate in reaction (16) by two different types of rhodium catalysts. On the other hand, 1-phenylethanoland a variety of the further secondary alcohols were obtained with high enantiomeric excess under in situ racemization with the ruthenium catalystA with very high efficiency [reactions (17)-(19)].Moreover, this catalyst was used also for the racemization/epimerization procedure (20) converting an approximately 1:1 mixture of racernic/rneso-diols into the corresponding enantiomeric diacetates under consumption of the meso-diol as well as for the dynamic kinetic resolution of 2- and 3-hydroxy carboxylic esters as shown in the reactions (21) and (22), respectively, under acylation conditions. The ruthenium catalyst was not compatible with vinyl acetate and therefore, 4-chlorophenyl acetate was found to be the acylating agent of choice. A redox process can explain the racemization of the slow reacting enantiomeric alcohol. Another ruthenium catalyst was used for the dynamic kinetic resolution of allylic alcohols [reaction (24)]by acylation yielding allylic acetates. Again a redox process should be responsible for the racemization.

I

565

566

I

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.1-25. The beneficial influence of additives on lipase-catalyzed enantiorner- and enantiotopos-differentiatingreactions (PCL Pseudomonas cepacia lipase, CCL Candida cylindracea lipase, CAL Candida antarctica lipase, not specified, LIP Pseudomonos sp. lipase-Toyobo,PFL Pseudomoasjuorescens lipase, CAL-B Candida antarctica B lipase, M M L Mucor rniehei lipase). Product(s)

Additive

Influence ofthe additive

NEt3

reaction rate

NEt3

selectivity

NEt3

selectivity

2 [21

2,6-lutidine or KHC03

selectivity

3 [31

NEt3

selectivity

4 [41

NEt,

reaction rate

5 [51

HO I

AcO pancreatin, AcOCH2CC13, THF no reaction without NEts pancreatin, vinyl acetate, no further solvent from 72 to >99 % ee

RY O T s

' Y 0 T . s OH

OAC

PCL, vinyl acetate, t-BuOMe R = Vinyl from E = 50 to >200 R = CH2Cl:from E = 39 to > 200 R = Et: from E = 7 to 60

Me0

OMe

Me0

OMe

OH

AcO

CCL, Ac20, toluene from E = 19 to 180with 2,6-lutidine from E = 19 to 240 with KHCO3

CAL, vinyl acetate, cyclohexane

OH

LIP, vinyl acetate, THF from 10 d to 3 h

1 1 . 7 Hydrolysis and Formation OfCarboXylid Acid Esters Table 11.1-25.

(cont.).

Produdlsl

Additive

Influence ofthe additive

NEt3

selectivity

161

NEb

selectivity

7 [71

NEt,

selectivity and reaction rate

8 [81

NEt3

selectivity and reaction rate

9 191

Dextromethorphan (DM) or its enantiomer Levomethorphan (LM)

selectivity and reaction rate

10 [lo]

AcO

PFL on sawdust, vinyl acetate, THF

Me0 OH

OAc

PCL on Hyflo Super Cell@,vinyl acetate, t-BuOMe, from 85 to 93 % ee (CH,),-OH

(CH,),-OAc

HO-,(H,C)

AcO-,(H,C)

absolute configuration unknown CAL-B,vinyl acetate, THF RxH

Ph1

0

MML or CAL-B, R'OH, n-hexane or toluene R = t-Bu, R' = n-Bu: from 80 to >99 % ee OAr

OAr

AC02H

ACO,Me

Me

I

567

7 7 Hydrolysis and Formation of C - 0 Bonds

(cont.).

Table 11.1-25. Product(s)

Additive

Influence ofthe additive

Dextromethorphan (cf 10) 01 (2S)-2-amino4-methylthio1-butanol

selectivity

11 [ll]

Cf3

selectivity

12 [12]

cr3

selectivity and reaction rate

13 [13]

Triton X-100

selectivity

14 1141

CaClz

selectivity

15 [ l S ]

NaCl

selectivity

16 [16]

CCL, phosphate buffer pH 7 Ar: 2-Me-4-Cl-C~Hj: from E = 1 to 37 with DM and to E = 81 with LM

OH EtL

OAc

C

N

Et/\/CN

PCL, aqueous buffer, pH 7.2

OH

Ph

Ph-

:

/

U

PCL, vinyl alkanoates, n-hexane or i-PrzO

OAc

RE

C

N

R&CN

U

and further crown ethers PCL, acetone/water

CCL, aqueous buffer pH 7.2

nC8H,,AC0,H

nC8H,,n ,\lC8H7 ,

CCL, aqueous buffer pH 8

PCL, HzO

J1.l Hydrolysis and Formation ofcarboxylid Acid Esterz Table 11.1-25.

(cont.).

Product(s)

Additive

Influence ofthe additive

aqueous LiCl

selectivity a n d reaction rate

H

"'A C0,nBu

0

I

X

17 [17]

I

X

CCL, n-BuOH, i-PrzO X = Et: from E = 3.8 to 201 X = CF,: from E = 1.3 to 56 1 a) F. Theil, S. Ballschuh, H. Schick, M. Haupt, B. Hafner, S. Schwarz, Synthesis1988, 540.; b) F. Theil, H. Schick, M. A. Lapitskaya, K. K. Pivnitsky, LiebigsAnn. Chem. 1991, 195. 2 N. W. Boaz, R. L. Zimmerman, Tetrahedron: Asymmetry1994,s. 153. 3 B. Berger, C. G. Rabiller, K. Konigsberger, K. Faber, H. Giengl, Tetrahedron: Asymmetry1990, I , 541. 4 P. Stead, H. Marley, M. Mahmoudian, G. Webb, D. Noble, Y. T. Ip, E. Piga, T. Rossi, S. M. Roberts, M. J. Dawson, Tetrahedron: Asymmetry1996,7, 2247. 5 a) T. Sugahara, K. Ogasawara, Synlett 1996, 319; b) T. Sugahara, Y.Kuroyanagi, K. Ogasawara, Synthesis1996, 1101. 6 W. Kreiser, A. Wiggemann, A. Krief, D. Swinnen, Ztrahedron Lett. 1996, 37, 7119. 7 A. Fadel, P. Arzel, Tetrahedron: Asymmetry1997, 8, 283. 8 F. Theil, H. Sonnenschein, T. Kreher, Tetrahedron: Asymmetry1996,7,3365. 9 a) N. J. Turner, J. R. Winterman, R. McCague, 1. S. Parratt, S. 1. Taylor, Tetrahedron Lett. 1995, 36, 1113; b) M.-C. Parker, S . A. Brown, L. Robertson, N. J. Turner, Chem. Commun.1998, 2247; c) S. A.

Brown, M.-C. Parker, N. A. Turner, Tetrahedron: Asymmetry2000,II,1687. 10 2.-W. Guo, C. J. Sih,/. Am. Chem. Soc. 1989,111, 6836. 11 T. Itoh, E. Ohira, Y. Takaki, S. Nishiyama, K. Nakamura, Bull. Chem. Soc./pn. 1991, 64,624. 12 a ) Y Takaki, 1. Teramoto, H. Kihara, T. Itoh, H. Tsukube, Tetrahedron Lett. 1996, 37,4991; b) Y. Takaki, R. Ino, H. Kihara, T. Itoh, H. Tsukube, Chem. Lett. 1997,1247. 13 a) T. Itoh, Y.Takaki, T. Murakami, Y. Hiyama, H. Tsukube,/. Org. Chem. 1996,61,2158; b) T. Itoh, K. Mitsukara, W. Kanphai, Y. Takaki, J. Teramoto, H. Kihara, H. Tsukube,/. Org. Chem. 1997,62,9165. 14 A. Bashkar Rao, H. Rehman, B. Krishnakumari, J. S. Yadav, Tetrahedron Lett. 1994, 35, 2611. 15 E. Holmberg, M. Holmquist, E. Hedenstrom, P. Berglund, T. Norin, H.-E. Hogberg, K. Hult, Appl. Microbiol. Biotechnol. 1991, 35, 572. 16 a) H. Tsukube, A. Betchaku, Y. Hiyama, T. Itoh, /. Chem. Soc., Chem. Commun. 1992,1751; b) H. Tsukube, A. Betchaku, Y. Hiyama, T. Itoh. 1.Org. Chem. 1994, 59, 7014. 17 T. Okamoto, S. Ueji, Chem. Commun. 1999, 939.

In reaction (25) racemization was realized by madelate racemase. However, this transformation is still a process carried out in two batches and therefore, not a dynamic kinetic resolution but certainly the starting point for further investigations by combining a lipase- and a second enzyme-catalyzed reaction in order to perform real dynamic kinetic resolution.

Enhancement of Selectivity and Reactivity of Lipases by Additives It has been shown that additives have a great potential for fine-tuning the reaction conditions for lipase-catalyzed reactions (1331. Certain additives such as tertiary amines, thiacrown ethers or inorganic salts may increase the selectivity and/or reaction rate. The reason for these effects are little understood and only a few systematic investigationshave been undertaken. From a synthetic chemist's point of 11.1 2 . 1 . 3

I

569

570

I

1 1 Hydrolysis and Formation of C - 0 Bonds

view treatment of the reaction mixture with an additive is a convenient way to improve the outcome of the reaction. Table 11.1-25 list examples in which certain additives have an unambiguous beneficial influence on selectivity and/or reaction rate. The enantiomerically enriched or pure compounds 1-9 have been prepared under the influence of mainly triethylamine or other bases. In case of 1 for the acetylation with 2,2,2-trichloroethyl acetate there was no reaction without triethylamine. For the formation of 5 the reaction time was shortened dramatically from ten days to three hours for 100% of conversion. In most cases there is no rationale for the effects of bases except the formation of ion-pairs between the added bases and traces of acids present in the reaction mixture. Only for the synthesis of 9 a systematic investigations demonstrates that triethylamine besides its racemizing properties (cf. Table 11.1-24)has a significant influence on the water activity of the reacting mixture['38].In other cases, triethylamine has been used as an additive without comparing its influence with the results in its Table 11.1-26.

R

Subtilisin-catalyzedacylation of racernic alcohols in organic solvents.

,r

Me

R = Et, VS/VR = 3.9, dioxane R = nBu, E = 28 R = (CH2)zCH=CMe2,E = 11 R = nHex, VS/VR = 100, dioxane R = nDec, E = 100 R = Ph, VS/VR = 50, dioxane R = 2-naphthyl, VS/VR = 58, dioxane vinylbutyrate, vinylacetate

PI 1

2

3 4

5

6

7

8a [2]

u L e

Me=Me

Me

298% ee 64% conversion

54% ee 64 % conversion

8b [2]

vinylacetate

OH

WMe

9b PI

9a [21

\

40% ee 30% conversion vinylacetate

1 P. A. Fitzpatrick, A. M. Klibanov,J. Am. Chem. SOC. 1991,131,3166.

92% ee 30% conversion 2 Y.-F. Wang, K. Yakovlevsky, B. Zhang, A. L. Margolin,/. Org. Chem. 1997, 62, 3488.

7 1.1 Hydrolysis and Formation of Carboxylid Acid Esters

I

571

The enantiomerically pure tertiary bases dextro- and levomethorphan were used for the preparation of 2-aryloxy propionic acid derivatives 10 by increasing the selectivity dramatically based on the enantioselectiveinhibition of the slow reacting enantiomer. The kinetic resolutions yielding the enantiomers 12 and 13 were conducted in the presence of thiacrown and some further crown ethers. Inorganic salts as shown for 15-17 were suitable modulators of selectivity and/or reaction rate. Particularly, in case of 17 a strong increase of the enantiomer-selectivitywas found by the addition of a defined amount of aqueous lithium chloride solution to the reaction mixture. The E value was increased by factors between ten to fifty depending on the substrate structure.

11.1.1.2.2

Subtilisin

A beneficial feature of subtilisin, and in particular subtilisin-CLECs, is their high catalytic activity in polar and non-polar organic solvents, allowing for transesterifications of alcohols in the presence of small amounts of water. Transesterifications catalyzed by subtilisin were mostly done with vinyl acetate. Apparently, the acetaldehyde formed during transesterification is not harmful to the enzyme as it is in the case of some lipases and pig liver esterase. Although resolution of such alcohols either through hydrolysis of the corresponding esters or transesterification is the domain of lipase, in some cases useful selectivities were achieved with subtilisin (1-9) (Table 11.1-26).

11.1.1 2 . 3

Pig Liver Esterase

Pig liver esterase-catalyzedenantioselectiveacylation of prochiral or racemic alcohols in organic solvents has not nearly gained the importance of the lipase-catalyzed acylation method. This is due to the fact that pig liver esterase shows only very low activity in organic solvents. The esterase differs considerably in this respect from lipases and subtilisin, which are both highly active in organic media. Attempts to confer activity to pig liver esterase in organic solvents by entrapment in water-filled porous supports 11391, covalent attachment of MPEG residues [140, 14'] , immobilization on and E ~ p e r g i t [ ' ~'@I,~ , or entrapment in polymers were met with various degrees of success. It was found, however, that colyophylizationof pig liver esterase with MPEG significantly enhances the activity and stability of the enzyme in organic solvents of low to medium polarity and low water content[", 14', 147j. The colyophilizate of pig liver esterase and MPEG was successfilly applied to the kinetic resolution of racemic glycerol derivatives through acylation with vinyl and isopropenyl esters in toluene containing less than 1% water (1, 3,and 5) (Table 11.1-27).Medium selectivities were recorded for alcohols having a primary hydroxyl group, and a high selectivity was found in the case of the glycerol derivative with a secondary hydroxyl group. Pig liver esterase-catalyzed hydrolysis of the corresponding racemic esters in water occurred with lower selectivities (2, 4 and 6 ) . A number of functionalized secondary alcohols have been resolved with high '

572

I

7 7 Hydrolysis and Formation of C-0 Bonds Table 11.1-27.

Pig liver esterase-catalyzed acylation of racemic alcohols in organic solvents

E = 24 vinyl propionate, toluene

E = 3-5 hydrolysis in water

E = 30 vinyl propionate, toluene

E=Z hydrolysis in water

E >lo0 vinyl propionate, toluene

E=l hydrolysis in water

11.7 Hydrolysis and Formation ofcarboxylid Acid Esters Table 11.1-27.

(cont.).

OCOEt M e O A P h

y 7a [ l , 21

M e O w P h

7b [1,2]

E = 100 vinyl propionate, toluene

E >lo0 vinyl propionate, toluene

E = 50 vinyl propionate, octane OCOEt

10b [ l , 21

E t S A P h

E >lo0 vinyl propionate, toluene OCOEt F

A

P

h

l l a [l,21

F

A

P

h

F

L

P

h

l l b [ l ,21

E >lo0 vinyl propionate, toluene OCOEt F

A

P

h

12a [ l , 21

12b 11.21

E = 50 vinyl propionate, toluene

y

OCOEt Me

&OPh

13a [ l , 21

E-100 vinyl propionate, toluene

Me

&OPh

13b [ l , 21

I

573

574

I

1 7 Hydrolysis and Formation of C-0Bonds Table 11.1-27.

(cont.).

OCOEt M e O h O P h

14a [ l , 21

OH MeO-OPh

14b [ l , 21

E >lo0 vinyl propionate, toluene 1 H:J. Gais, M. lungen, V. ladhav,J. Org. Chem. 2001,66,3384.

2 M. Jungen, H:].

1999,10,3747.

Gais, Tetrahedron: Asyrnrnstry

selectivity by pig liver esterase-catalyzed acylation with vinyl propionate in toluene. As in the case of lipases, a competing pig liver esterase-catalyzedhydrolysis of vinyl propionate and a partial deactivation of the enzyme by the acetaldehyde formed in transesterification had been observed. Critical to the activity and selectivity of pig liver esterase in the presence of MPEG in organic solvents is the water content of the system, which should be lower than 1 %. In general, the activity of pig liver esterase in the presence of MPEG in organic solvents is lower than that of lipases and subtilisin under comparable conditions. The colyophilizate of pig liver esterase and MPEG can be recovered from organic media with a minor loss of activity through a spontaneous immobilization on an ultrafiltration membrane placed in the reaction mixture r6'1.

Acknowledgement

The authors thank Carsten Griebel and Gabriele Bertrand for their help in the preparation of the manuscript.

References J. B. Jones, C. J. Sih, D. Perlman (eds), Applications of Biochemical Systems in Organic Chemistry, Wiley, New York, 1976, parts I and 11. 2 R. Porter, S. Clark (eds), Enzymes in Organic Synthesis, CIBA Foundation Symposium 111, Pitman, London, 1985. 3 J. Tramper, H. C. van der Plas, P. Limko (eds), Biocatalysts in Organic Synthesis, Elsevier, Amsterdam, 1985. 4 G. M. Whitesides, C. H. Wong, Angew. Chem. 1985,97,617; Angew. Chem., Int. Ed. Engl. 1985, 24, 617. 5 J. B. Jones, Tetrahedron 1986, 42, 3351. 6 M. P. Schneider (ed), Enzymes as Catalysts in Organic Synthesis, NATO AS1 Series C, 1

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References I 5 7 5 11 C. S. Chen, C. J. Sih, Angew. Chem. 1989,

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Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

1 1.2 Hydrolysis offpoxides

11.2

Hydrolysis of Epoxides Kurt Faber and Romano V A. Orru

Chiral epoxides and 1,2-diols,which are central building blocks for the asymmetric synthesis of bioactive compounds, can be obtained via the asymmetric hydrolysis of epoxides using enzymes - i.e. epoxide hydrolases (EHs) [EC 3.3.2.Xl. Enzymes from mammalian sources - such as rat liver tissue - have been investigated in great detail for several decades during detoxification studies L1]; however, their application for biotransformations on a preparative scale was hampered because of the limited supply of these enzymes, and, as a consequence, the examples reported rarely surpass the millimolar range[*, 1’. During the past few years, highly selective epoxide hydrolases were identified from a wide range of microbial sources, which allows for an (almost) unlimited supply of these enzymes for preparative-scale applications. These valuable biocatalysts have recently gained considerable attention, and their scope and limitations have been reviewed[”*]. Microbial epoxide hydrolases were found to be more abundant than previously expected, and numerous sources, predominantly among bacteria, fungi and (red) yeasts are known to date. The mechanism of enzymatic hydrolysis of epoxides can be compared to that of base-catalysis, i.e. it resembles an SNZ-type opening of the epoxide by the nucleophile (i.e. water), which leads to the formation of the corresponding trans-configurated 1,2-diol. Any chiral center present in the substrate oxirane can be “recognized”, thus effecting kinetic resolution or asymmetrization of racemic or meso-epoxides, respectively. The data available to date indicate that the enantioselectivities of enzymes from certain microbial sources can be correlated to the substitutional pattern of various types of substrates: red yeasts (Rhodotorula or Rhodosporidiurn sp.) give best enantioselectivities with monosubstituted oxiranes; hngal cells (e.g. from Aspergillus and Beauveria sp.) are best suited for styrene oxidetype substrates, whereas bacterial enzymes (in particular from Actinornycetessuch as Rhodococcus and Nocardia sp.) are the biocatalysts of choice for more highly substituted 2,2- and 2,3-substituted epoxides. In order to overcome the disadvantage of the classic kinetic resolution pattern, i.e. the formation of two enantiomers in each 50 % yield, various deracemization methods based on chemo-enzymatic or purely enzymatic protocols have been developed. The latter led to the highly desirable formation of a single stereoisomer of the diol in 100% theoretical yield. The synthetic potential of epoxide hydrolases for asymmetric synthesis has been proven by the preparation of a number of bioactive compounds. Chiral epoxides and vicinal diols (employedas their corresponding cyclic sulfate or sulfite esters as reactive intermediates) are extensively employed high-value intermediates in the synthesis of chiral compounds because of their ability to react with a broad variety of nucleophiles (Figs. 11.2-1 and 11.2-2). In recent years, extensive efforts have been devoted to the development of chemo-catalytic methods for their production['^ “1. Thus, the Sharpless methods allowing for the asymmetric epoxida-

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I 1 Hydrolysis and Formation ofC-0 Bonds Figure 11.2-1. SR'

R'

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RA

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nucleophiles.

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11

OR'

tion of allylic alcohols [''I and the asymmetric dihydroxylationof alkenes [lo]are now widely applied reliable procedures. In addition, asymmetric catalysts for the epoxidation of non-functionalized olefins [l2-l4]have been developed more recently. Although high stereoselectivityhas been achieved for the epoxidation of cis-alkenes,the results obtained with trans- and terminal olefins were less satisfactory using the latter method. More recently, two highly selective methods for the opening of terminal mono- and 2,2-disubstitutedepoxides have been published. These methods are both based on a kinetic resolution using cobalt-salen complexes and water[15]or chromium-salen complexes and azide respectively and have great potential in asymmetric synthesis [171. On the other hand, a number of biocatalytic methods have been reported to provide a useful arsenal of methods as valuable alternatives to the above-mentioned techniques [18-231. Prochiral or racemic synthetic precursors of epoxides, such as halohydrins, can be asymmetrized or resolved using hydrolytic enzymes [24, 251. In particular, esterases and lipases have been used for such a enantioselective ester hydrolysis or esterification. This methodology is well developed, and high selectivities have been achieved in particular for esters of secondary alcohols, but it is impeded by the requirement of regioisomerically pure halohydrins. Furthermore, it is known that a-haloacid dehalogenases catalyze the SN2-displacementof a halogen atom at the a-position of carboxylic acids with a hydroxy function. This process leads to the formation of the corresponding a-hydroxy acid with inversion of configuration[26].However, a-haloacid dehalogenation incurs two drawbacks: (i)the instability of the substrates, particularly the a-bromoacids, in aqueous systems, and (ii) the limited substrate tolerance, as only short-chain haloacids are accepted [271. Asymmetric biocatalyhc reduction of a-keto-acids12'] using D- or L-lactate dehydrogenase or a-keto-alcohols[291 by glycerol dehydrogenase provides access to chiral a-hydroxyacids or 1,2-diols,which can be converted into the corresponding epoxides using conventional chemical methodology. Although excellent selectivities are generally

11.2 Hydrolysis of Epoxides

r OH

I

R u 0 4 cat

L

TosCVPy

I

581

\

NalO

0

U,O

,x5 Figure 11.2-2.

Syntheses from chiral 1,2-diols.

achieved, the need for the recycling of redox-cofactors such as NAD(P)H has restricted the number of applications. Likewise, biocatalytic asymmetric epoxidation of alkenes catalyzed by mono-oxygenasescannot be performed on a preparative scale with isolated enzymes, because of their complex nature and their dependence on a redox cofactor such as NAD(P)H. Thus, whole microbial cells have to be used instead. This method is not trivial and requires high bioengineering skills [301. On the other hand, haloperoxidases are independent of nicotinamide-cofactors, as they produce hypohalous acid from H202 and halide, which in turn yields a halohydrin from an alkene. These enzymes are rare in Nature and exhibit usually low selectivities due to the fact that the formation of halohydrins can take place not only in the active site of the enzyme but also without enzyme Similar low selectivities have been observed with halohydrin epoxidases, which act like a "biogenic chiral base" by converting a halohydrin into the corresponding e p o ~ i d e [ ~ ~ ] . On the other hand, peroxidases, such as chloroperoxidase (CPO), are cofactorindependent and can be used in isolated form for the enzymatic epoxidation of alkenes [33-351. An attractive alternative to the methods mentioned above is the use of cofactorindependent epoxide hydrolases, which are readily available from microbial sources in sufficient quantities. 11.2.1 Epoxide Hydrolases in Nature

In eukaryotes,microsomal and cytosolic epoxide hydrolases mainly play a key role in the detoxification of mutagenic, poisonous and carcinogenic epoxidesf3', 371, which are formed by the action of P4so-dependentmonooxygenases[381. In addition, they are involved in the biosynthesis of hormones (e.g. leukotrienes). In plants, epoxide hydrolases are responsible for the generation of chiral aroma compounds and, in the biosynthesis of cutin, a wax-type polyester, which protects plants against microbial attack[3']. Insect epoxide hydrolases degrade juvenile hormones and pheromones bearing an oxirane moiety L31. On the other hand, in microorganisms these enzymes

582

I

11 Hydrolysis and Formation of C - 0 Bonds

are multi-functional: (i) they can function as detoxifylng agents, (ii) they can play a role in biosynthetic routes of complex (secondary)metabolites, or (iii) they may be crucial for the degradation of epoxides during the metabolism of alkenes and aromatics I4OI. The degradation of aromatics in eukaryotes occurs via two different pathways (Fig. 11.2-3):(i) dioxygenase-catalyzedcycloaddition of molecular oxygen to the C=C bond yields a (putative) dioxetane species, which is then detoxified via reductive cleavage of the 0-0 bond yielding a physiologically more innocuous cis-1,2-diol; (ii) The formation of a highly reactive arene oxide via the introduction of a single 0 atom (from molecular oxygen) into the aromatic system is catalyzed by a mono-oxygenase. The latter epoxy species is further metabolized via hydrolysis catalyzed by an epoxide hydrolase to yield a transb1,2-diol. In lower organisms, alkenes can be metabolized in an analogous fashion, i. e. via an epoxide intermediate. In an analogous fashion, this intermediate is hydrolyzed to the corresponding 1,2-diolby an epoxide hydrolase. The latter product is degraded either by oxidation or by elimination of water under catalysis of a diol dehydratase, yielding an aldehyde [411. Alternatively, such aldehydes are obtained via direct rearrangement of the epoxide catalyzed by an epoxide i~omerase[~']. For a long time it was generally assumed that epoxide hydrolases are predominantly found in mammals[" 1', although epoxide hydrolase activities had been detected in bacteria[43.441 or fungi i4', 461 quite some years ago. This early view was certainly too simplistic, and enzymes of this type have now been detected in many bacteria [47-491, fungi and red yeasts ['*I. Moreover, epoxide hydrolase activity has been demonstrated in plants ['*Iand insects FS31. 11.2.1.1

Isolation and Characterization o f Epoxide Hydrolases

Several membrane-bound and soluble epoxide hydrolases from mammalian origin have been purified and (at least partially) sequenced. Some of them have also been cloned and overexpressed, which is the case for the soluble EH from rat liver which "I. This enzyme (as well as its has been overexpressed in Escherichia microsomal analog) was shown to share an amino acid sequence similarity to a region around the active center of a bacterial haloalkane dehalogena~e['~]], an enzyme with known three-dimensional structure that belongs to the a/ P-hydrolase f0ld-family['~1.Rat soluble EH forms a dimer from two complete structural monomeric units, both possessing a distinct active site. The EH activity is known to be located close to the C-terminal unit, while the function of the N-terminal unit remains u n k n o ~ n [ ~ ~ l . To date, several epoxide hydrolases from microbial sources have been purified. For instance, from Bacillus megaterium La], Corynebacterium sp. 471 and Pseudomonas sp. [48, 6ol, but also from dematiaceous fungi such as Ulocladium atrum and Zopfcella karachiensis["I. However, some of these were only partially purified, or their enantioselectivitieswere low or not investigated. In contrast, highly enantioselective epoxide hydrolases from Rhodococcus sp. NCIMB lK!lC~[~'land Nocardia sp. EH1 ["I c0li['~2

11.2 Hydrolysis ofEpoxides

Rkx:

bk Oxygenase

-

cis -diol

dioxetane

R

R

Epoxide Hydrolase

Oxygenase

f *

-

H20

arene-oxide

trans -diol

1

OH Epoxide Hydrolase

/=

R

-

I

further metabolism

u: +

RLoH

DiolH20 4 D e h y d r a t a s e

H20

MonoOxygenase

~

PI

p Epoxide Isomerase

further metabolism

lsponf.

R

T

o

H Figure 11.2-3. involvement of epoxide hydrolases in the biodegradation of aromatics and alkenes.

were purified to homogeneity. Both (monomeric) proteins exhibit several common features: they are of similar size (= 34 kDa) and do not possess any metal ion or any UV-absorbing prosthetic group. The catalyhc power of both enzymes was found to be in the same range (=ZOO0pmol mg-' h-'), as was the optimum temperature (33-37 "C) and pH (7.5-9.0). The only notable difference between the two enzymes is the high instability of the Nocardia epoxide hydrolase (which completely loses its activity within hours, even when stored at -18 "C), whereas the Rhodococcus enzyme was shown to be relatively stable. It should be noted that the former enzyme could be stabilized by immobilization through ionic binding onto DEAE-cellulose. This resulted in a doubling of the activity (compared to the native enzyme) albeit at a

I

583

584

7 7 Hydrolysis and Formation of C-0 Bonds

I slight reduction in enantioselectivity

["I. The poor stability of the Nocardia enzyme and the fact that the N-terminus of the Rhodococcus epoxide hydrolase was unspecifically blocked precluded their N-terminal sequencing. In contrast, an epichlorohydrin-degrading epoxide hydrolase from Agrobacterium radiobacter AD1 could be isolated, characterized and sequenced after cloning and overexpression in E. coli BL21 (DE3)["]. This enzyme showed an amino acid sequence similarity to eukaryotic epoxide hydrolases, haloalkane dehalogenase and bromoperoxidase, which indicated that it belongs to the a/P-hydrolase fold family. Most epoxide hydrolases from pro- or eukaryotic sources seem to belong to this group of enzymes. Another bacterial epoxide hydrolase from this family has been isolated from Corynebacterium sp. C12 when grown on cyclohexene oxide["]. The purification to homogeneity was achieved in two steps. The enzyme is (partly) membrane bound and multimeric (probably tetrameric) which is in contrast to the enzymes described above. The subunit-size is ca. 32 kDa and amino acid sequence comparison showed that it is related to mammalian and plant (soluble) EH. Furthermore, it showed striking similarilties with the Agrobacterium enzyme, particularly around the catalytic site. Epoxide hydrolases from fungal sources were purified recently: an epoxide hydrolase from Aspergillus niger was purified to homogeneity["] and appears to be a tetramer composed of four identical subunits of molecular mass 45 kDa. The Nterminus was blocked, the pH optimum lies at 7.0 and the temperature optimum at 40 "C. From red yeasts (Rhodotorula g l ~ t i n i s [ and ~ ~ ] Rhodosporidium toruloides CBS0349 [68]), two epoxide hydrolases have been purified. Both membrane-bound enzymes are medium-sized (45 and 54 kDa, respectively)and are structurally related to other microsomal epoxide hydrolases. They probably belong to the a/ P-hydrolase fold family as well. An epoxide hydrolase with an unprecedented low molecular mass (only 17 kDa) was isolated from Rhodococcus erythropolis DCL14[69].The cofactor-independent enzyme is efficiently induced when the microorganism was grown on monoterpenes, such as limonene, reflecting its special role in the limonene degradation pathway. The low molecular mass, the unusually broad pHoptimum (6.0-11.0) and the elevated optimum temperature (50 "C), together with the fact that the N-terminal amino acid sequence revealed no homology with any other protein, led to the conclusion that this protein does not belong to the alphydrolase fold family. 11.2.1.2

Structure and Mechanism of Epoxide Hydrolases

The first X-ray structure of an epoxide hydrolase (from Agrobacterium radiobacter AD1) has been reported recently (Fig. 11.2-4)f7O1. The nearly globular protein consists of a core-domain with typical features of a/P-hydrolase fold enzymes and a so-called "cap-domain",which is located on top of the core domain. All epoxide hydrolases known to date require neither any prosthetic group nor a metal ion, and the mechanism by which these enzymes operate was long debated. Formerly, it was assumed that a direct nucleophilic opening of the oxirane ring by a histidine-activatedwater molecule would be the key step C7l1. However, convincing

17.2 Hydrolysis ofEpoxides

il Figure 11.2-4. X-Ray structure o f Agrobacterium radiobacter epoxide hydrolase (PDB-1EHY). The catalytic residues (Asp107 and His275) are located on top o f t h e core-domain: at some distance Asp246 is shown, which is presumably involved in proton transfer. The a-helices at top left constitute the “cap-domain”, which is covering the active site.

evidence was later provided which showed that the reaction occurs via a covalent glycol-monoester-enzymeintermediate[72,731 (Fig. 11.2-5). For the Agrobacterium enzyme, the proposed active-site residues (Asp107 and His275) are located in the predominantly hydrophobic internal cavity between the core- and cap-domains[64,701. Furthermore, a tunnel filled with water molecules has been located, which leads to the back of the active site cavity. It is perfectly suited to deliver the catalytic water molecule within hydrogen-bonding distance to His275. Since the water is positioned at the back, the epoxide probably enters the active site from the front. In addition, Asp246 has been proposed as the third member of the catalytic triad (not shown in Fig. 11.2-S), since replacement of AsplO7, His275 and Asp246 resulted in a dramatic loss of activity[“].

I

585

586

I

1 1 Hydrolysis and Formation ofC-0 Bonds

cy HO

H

H

OH

’glycol-monoester intermediate’

T

T

A0-

o/

H

..

..

H

H

‘alkyl-enzyme intermediate’ Figure 11.2-5. Schematic representation of the mechanism of epoxide hydrolase and of haloalkane dehalogenase.

Structure and mechanism show striking similarities to that of haloalkane dehalogenase from Xanthobacter autotrophicus (whose structure and mechanism has been substantiated by X-ray crystallography)[74, 781. Both enzymes have an Asp-His-Asp catalytic triad which superimpose very well, their side chains point in a similar relative direction and they form analogous hydrogen bonds. In haloalkane dehalogenase, it has been shown that a halide is displaced from the substrate by an aspartate residue via a nucleophilic attack, thus leading to an “alkyl-enzyme intermediate” which is further hydrolyzed in a second step[75,761. Similar mechanisms have been proposed for other epoxide hydrolases belonging to the a/P-hydrolase fold family[% 72, 771 A consequence of the above-mentioned mechanism is a trans-specific opening of the epoxide with one oxygen from water being incorporated into the product diol[60]. For instance, (+)-trans-epoxysuccinatewas converted into meso- (not D/L-) tartrate by an epoxide hydrolase isolated from Pseudomonas putida[601. In a complementary fashion, cis-meso-epoxysuccinategave D- and L-tartrate (with a Rhodococcus sp.) albeit in low optical purity[’’]. The fact that only one 0-atom originates from water was proven by 180-labelling experiments using bacterial, fungal i80] and mammalian epoxide hydrolases I8l]. Although two cases for reactions proceeding via a formal cis-

7 7.2 Hydrolysis of Epoxides

4"

S

k rac

L Inversion

"3,

+

R

(S) -Enantiomer reacts faster

R

R

R!

Figure 11.2-6. Microbial hydrolysis o f epoxides proceeding with retention or inversion of configuration.

hydration process have been reported in the mid-1970s[**.831, they seem to be rare exceptions and - given the present knowledge of the enzyme mechanism - attempts to explain this phenomenon remain rather speculative[83].It is interesting to note that several P-glycosidases act via formation of a covalent glycosyl-enzyme intermediate by retaining the configuration at the anomeric centre[84].This suggests that these enzymes may also be mechanisticallyrelated to epoxide hydrolases. The above-mentioned facts have important consequences for the stereochemical outcome of the kinetic resolution of asymmetrically substituted epoxides. In the majority of enzymatic transformations following a kinetic resolution pattern (e.g. by ester hydrolysis and synthesis using lipases, esterases and proteases) the absolute configuration at the stereogenic centre(s) always remains the same throughout the reaction, since it is not directly involved in the reaction. In contrast, the enzymatic hydrolysis of epoxides may take place via attack on either carbon of the oxirane ring (Fig. 11.2-6) and it is the structure of the substrate and of the enzyme which 8s-891 . A s a consequence, the determine the regioselectivity of the process absolute configuration of both the product and the substrate from a kinetic resolution of a racemic epoxide has to be determined in order to elucidate the stereochemical pathway. To facilitate the determination of this regioselectivity, a mathematical approach has been suggested, which only necessitates the study of the biohydrolysis of the racemic mixture ["I. 11.2.1.3

Screening for Microbial Epoxide Hydrolases

In spite of the considerablevalue of epoxide hydrolases for fine chemical synthesis, it was only recently that a detailed search for epoxide hydrolases from microbial sources was undertaken by the groups of F ~ r s t o s s [ ~and ~ ~ Faber[23s79, 911, bearing in mind that the use of microbial enzymes allows an (almost) unlimited supply of biocatalyst. The screening was based along the following considerations: on the one hand, the catabolism of alkenes often implies the hydrolysis of an epoxide inter-

I

587

588

I

7 7 Hydrolysis and Formation of C - 0 Bonds

mediate and, on the other hand, detoxification of the highly reactive epoxyintermediates is achieved via hydrolysis. As a consequence, it was anticipated that bacteria and fungi which were known to be able to epoxidize alkenes in an efficient manner should also possess a matching epoxide hydrolase activity. This proved to be true for the fungi Aspergillus niger and Beauueria bassiana, which were able to achieve the enantioselective hydrolysis of different types of epoxides derived from geraniol, limonene [)'I or substituted styrene derivatives 931. In an extensive follow-up study, seven additional fungal strains (from a total of 42) were selected for exhibiting promising epoxide hydrolase activity[94].The guidelines mentioned above were also successfully applied to the screening of bacteria, and strains were selected after a careful literature search based on the capabilityfor alkene-epoxidation.Following the work described above, the occurrence of epoxide hydrolases in yeasts has been in~estigatedl~'. "1. From a screening of 187 different yeast strains belonging to 25 different genera, 8 strains (Trichosporon, Rhodotorula and Rhodosporidium sp.) were identified by using 1,2-epoxyoctane as substrate [971. The membrane-associated epoxide hydrolases from these yeasts show good enantioselectivitiesand high initial rates, especially for monosubstituted aliphatic epoxides L9'1. It is noteworthy that, in contrast to mammalian systems, the majority of bacterial and fungal strains exhibited sufficient activity even when the cells were grown on a non-optimized standard medium. Since enzyme induction is still a largely empirical task, cells are usually grown on standard media in the absence of inducers. Furthermore, all attempts to induce epoxide hydrolase activity in Pseudomonas aeruginosa NCIMB 9571 and Pseudomonas oleovorans ATCC 29 347 by growing the cells on an alkane (decane) or alkene (decene) as the sole carbon source failedL4]. Epoxide hydrolases from Corynebacterium["] and Rhodococcus DCL4['"] seem to be exceptional with respect to their inducibility. 11.2.2 Microbial Hydrolysis o f Epoxides 1 1 2.2.1 Fungal Enzymes

One of the first observations on microbial epoxide hydrolysis on a preparative scale was reported from the terpene field: thus, racemic geraniol N-phenylcarbamate was efficiently hydrolyzed by the fungus Aspergillus niger, yielding 42 % of the remaining ((5s)-epoxidein 94% ee. Interestingly, from the preparative point of view, this could easily be conducted on 5 g of substrate using a 7 L fermentor r91'. Similar results were obtained with styrene oxide, which was again very efficiently hydrolyzed by A. niger, thus affording the (S)-epoxide in 99% ee within a few hours[85].In contrast, the fungus Beauveria bassiana (formerly B. sulfirescens) showed opposite enantioselectivity,leading to the (R)-epoxidein 99% ee (Fig. 11.2-7). In addition, interesting information concerning the mechanism implied in these transformations [*I' and the scope of the substrates admitted could be established. Thus, it was shown that cyclic styrene analogs like para-substituted styrene oxide

17.2 Hydrolysis ofEpoxides

I

589

e.e. >99%

e.e. 62%

OH

d

n

.

A. niger +

B.b.

&G

B. bassiana

89% e.e., 92% yield

rac Beauveria bassiana (inversion) e.e. >99% Figure 11.2-7.

e.e. 84%

Resolution and deracernization o f styrene oxide by fungal cells.

derivatives[991 or P-substituted analogs [921 were accepted by one - or both - of these fungi. During a subsequent study, seven additional fungi were tested on more than ten styrene oxide derivatives bearing various substituents[lOO]. It was shown that an increase of the size of the substituent resulted in a selectivity-enhancement,e. g., from E = 3 for styrene oxide to E = 39 for para-nitrostyrene oxide. However, a methyl substituent at Ca did not improve the enantioselectivity of the reaction. Racemic epoxyindene was rapidly hydrolyzed when submitted to a culture of B. sulfirescens, leading to a 20% yield of recovered enantiomerically pure (ee >98%) (lR,ZS)-epoxide,and to a 48% yield of the corresponding (lR,ZR)-trans-diolshowing G9 % ee [loll. The latter product is of considerable importance for the synthesis of the HIV protease inhibitor indinavir. This prompted Merck Co. to perform a more extensive study of this biotransformation [Io2. lo3],during which 80 fungal strains were evaluated for their ability to enantioselectivelyhydrolyze racemic epoxyindene. In a similar fashion, epoxydihydronaphthalenewas successfully hydrolyzed to the corresponding (1R,2R)-diol in excellent enantiomeric purity [lo']. Many of these fungal epoxide hydrolases were found to be soluble enzymes, which could be obtained as crude cell-free extracts and which could be stored at +4 "C without significant loss of activity. In this way, easy-to-use water-soluble catalysts were developed,which circumvented the problems often encountered when working with whole-cellmycelia ['04, 1"'.

590

I

7 7 Hydrolysis and Formation ofC-0 Bonds

0-

Lyophilized Bacterial Cells

large

c)

buffer pH 7-8

large

+v small

small

large

rac

0 small

OH

Figure 11.2-8.

Resolution o f 2,2-disubstituted oxiranes by bacterial cells.

11.2.2.2

Bacterial Enzymes

The use of bacterial cells for preparative biotransformations is particularly attractive for the following reasons: (i) they do not tend to form dense mycelia, which may impede agitation of large-scale reactions when whole-cell (fungal) systems are employed, and (ii) cloning of bacterial enzymes is generally less problematic. However, disappointingly low selectivities were observed with monosubstituted or benzyl glycidyl ether ( E <2) 831. aliphatic epoxides such as 1-epoxyoctane( E 4) On the other hand, the sterically more demanding 2,2-disubstitutedoxiranes turned out to be much better substrates (Fig. 11.2-8,Table 11.2-1). Especially the substrates bearing a straight alkyl chain were transformed with virtually absolute selectivity, and functional groups such as a C=C-double or a terminal bromogroup [lo7]were well tolerated. As a consequence of the exquisite enantioselectivity, the reactions ceased and did not proceed beyond a conversion of 50%. Interestingly, the enantiopreference was found to depend on the substrate structure, but not on the strain When the epoxide bears a synthetically useful phenyl moiety (mimicking a masked carboxyl function) at the o-position of the alkyl chain, the selectivity was slightly reduced but still in a useful range (Table 11.2-1, last entry, E = 123)['07,lo9]. Unexpectedly, when the carbon chain was extended by an Table 11.2-1. Selectivities in the resolution of 2,2-disubstituted oxiranes by bacterial cells (see Fig. 11.2-8).

Small substituent

Large substituent

CH3 CH3 CH3 CH3 CH3 CH3 CH3

n-CSH11 n-C4Hs (CHz),CH n-C7H15 n-CgH19 (C&)4-Br CHzPh

Biocatalyst

Selectivity (E)

Nocardia sp. EH1 > 200 Nocardia sp. TB1 > 200 = CH2 Nocardia sp. EH1 > 200 Rhodococcus equi I F 0 3730 > 200 Mycobactenurn paraflnicum NCIMB 10420 > 200 Nocardia sp. H8 > 200 Nocardia sp. EH1 123

1 1.2 Hydrolysis of Epoxides Table 11.2-2. Selectivities i n the resolution of 2,2-disubstituted oxiranes by whole cells of Rhodococcussp. NClMB11216 (seeFig.11.2-8). Small substituent

Large substituent

Selectivity (E)

2.Sa

105 7 125 > 200 111 9.5 a With opposite absolute configuration.

additional CHz-unitthe selectivity declined (Table 11.2-2, last entry, E = 9.5). From an extensive study on Rhodococcus sp. NCIMB 11216 it was concluded that the enantioselectivity largely depends on the relative difference in size of the two alkyl substituent groups (Table 11.2-2). Increasing this size difference resulted in enhanced selectivities. The fact that the substrates bearing a phenyl group behave differently might be attributed to electronic effects, such as n-n-stacking.All of these biohydrolyses can be performed on a multigram-scale. In contrast to the rather flexible bacterial epoxide hydrolases mentioned above, limonene-1,2-oxide hydrolase from Rhodococcus erythropolis DCL14, an enzyme involved in the limonene degradation pathway, has a rather narrow substrate specificity. Of the compounds tested, only the natural substrate limonene-1,2-oxide and several highly substituted (alicyclic) epoxides were substrates for the enzyme. The enantioselectivities were usually low, except for the natural substrate [l1O1. Styrene oxide, various derivatives thereof and phenyl glycidyl ether were obtained in high ee and reasonable yield using a recombinant epoxide hydrolase from Agrobacterium radiobacter ADl["']. Interestingly, the Tyrl52Phe and Tyr215Phe mutants showed a considerableincrease in stereoselectivity.For example, the Evalue for parachlorostyrene oxide increased fourfold from E = 32 for the wild-type enzyme to E >130 for the Tyr215Phe mutant enzyme[78]. 11.2.2.3

Yeast Enzymes

Yeasts are generally very sturdy microorganisms which are easy to cultivate on a large scale. These features make them interesting for preparative use r9'1. Enantioselective epoxide hydrolysis by (red) yeasts has been studied only recently; the first report demonstrated the epoxide hydrolase activity of Rhodotorula gl~tinis['~] on several aryl, alicyclic and aliphatic epoxides. In follow-up studies, additional yeast strains exhibiting good activities and sufficient enantioselectivitieswere found, and the application of these biocatalysts has great potential[97.981. Given the data available to date, it seems to be a general phenomenon that epoxide hydrolases from fungi and from bacteria generally possess an opposite enantiopreference. Whereas epoxide hydrolases from fungi of matching opposite enantio-

I

591

592

I

7 7 Hydrolysis and Formation of C-0 Bonds

preference are not known, an extensive screening showed that bacteria seem to be more flexible in this respect[’’*]. This allows one to control the stereochemical outcome by a simple choice of the appropriate microorganism. 112 . 3 Substrate Specificity and Selectivity

1 1 2.3.1 Asymmetrization o f meso-Epoxides

The asymmetrization of a meso-epoxide via regioselective attack at one of the (enantiomeric) stereocenters of the oxirane would be an elegant application of epoxide hydrolases since it leads to a single trans-diolin 100% theoretical yield. Such asymmetrization reactions have been demonstrated with epoxide hydrolases from mammalian origin, which afforded the enantiomerically enriched corresponding dio1[113.1141. u nfortunately, few such reactions have been reported with microbial enzymes. For instance, cyclohexene oxide was hydrolyzed using Corynesporium albeit with disappointingly low ee cassiicola cells yielding trans-cyclohexane-l,2-diol, (27%) It was only after further metabolism involving an oxidation-reduction sequence by dehydrogenases present in the cells that the reaction product was In a related experiment, transformed into optically pure (S,S)-cyclohexane-1,2-diol. asymmetric hydrolysis of cis-epoxysuccinate using a crude enzyme preparation derived from Rhodococcus sp. led to D- and L-tartaric acid in almost racemic form[79]. Similar discouraging results were obtained using baker’s yeast [IiG]. Recently, encouraging progress was made in the hydrolysis of cyclopentene oxide and cyclohexene oxide using the yeast Rhodotorula glutinis[”]. The corresponding (R,R)-trans-diolswere obtained in over 90% optical and chemical yields. However, asymmetric hydrolysis of meso-epoxides by bacterial and fungal epoxide hydrolases is still impeded by insufficient selectivities. 11L3.2 Resolution of Racemic Epoxides

Monosubstituted Epoxides. Monosubstituted oxiranes [Fig. 11.2-9(a), Table 11.2-31 represent highly flexible and rather “slim”molecules, which make chiral recognition a difficult task. As a consequence, the majority of attempts using epoxide hydrolases from bacterial and fungal origin to achieve highly selective transformations failed[i1s]with one exception [li7].Most interestingly, the only selective enzymes were found among red yeasts, such as Rhodotorula araucarae CBS 6031[”], Rhodo-

7 7.2 Hydrolysis of Epoxides Table 11.2-3. R

Enzymatic hydrolysis of monosubstituted epoxides, see Fig. 9(a). Selectivitya Enantiopreference Enzyme Sourceb Reference f

R

BEH

117

-

n. d.

BEH

8

-

R R R

BEH FEH YEH YEH YEH YEH

23 50 51 51 97 96

+ -

to f

++ ++

S

R R

a Selectivity denoted as (-) =low (E < 4), (3=moderate (E (E > SO).

b BEH = bacterial epoxide hydrolase; FEH n. d. = not determined.

= fungal

= 4-12),

(+) = good (E = 13-50), (++) =excellent

epoxide hydrolase; YEH =yeast epoxide hydrolase.

sporidiurn toluloides CBS 0349197],Trichosporon sp. UOFS Y-01118[971 and Rhodotorula glutinis CIMW147 ["I. These yeasts' epoxide hydrolase seem to have a preference for monosubstituted oxiranes with a chain length of approximately six carbon atoms or more (E up to 200). Furthermore, olefinic side chains are sometimes hydrolyzed selectively (E up to 100) as well[98].Based on the rules of the kinetic resolution of a racemate, diols of high ee could only be obtained at low conversions[95].With few exceptions, the enantiopreference for the (R)-configuratedoxirane was predominant regardless of the enzyme source['18]. Styrene Oxide-Type Epoxides. Styrene-oxide-type oxiranes [Fig. 11.2-9(b), Table 11.2-41have to be regarded as a special group of substrates, as they possess a benzylic carbon atom. This facilitates the formation of a carbocation which is stabilizedby the adjacent aromatic moiety. As a consequence, nucleophilic attack at the benzylic position is electronically favored. On the other hand, the benzylic position is sterically more demanding, which favors the non-benzylic position. As a consequence, either oxirane carbon atom is easily attacked in this class of substrates and mixed regiochemical pathways are common. Since this results in reactions occurring with inversion and retention of configuration, E-values reported on these type of oxiranes have to be regarded with great caution whenever the regioselectivityhas not clearly been elucidated. In order to achieve optimal enantioselectivities, the biocatalysts of choice for styrene oxide-type oxiranes are derived from red yeasts such as Rhodutorula glutinis CIMW 147 951 and particularly from fungal epoxide hydrolases, e.g. Aspergillus niger LCP 521 and Beauveria bassiana ATCC 7159['". '"]. The first entry in Table 11.2-4is given solely for reason of comparison, since mammalian hepatic epoxide hydrolase was used. This enzyme source is not applicable to preparative-scale reactions. Interestingly, the bacterial epoxide hydrolase from Agrobacteriurn radiobacter AD 1 seems to hydrolyze para-substituted styrene oxides with opposite enantiopreference when compared to EHs from fungi or yeast["8]. Although initial selectivities were 1"s

I

593

594

I

I I Hydrolysis and Formation ofC-0 Bonds Table 11.2-4.

Enzymatic hydrolysis of styrene oxide-type epoxides, see Fig. 9(b).

Ri

RP

R3

X

Selectivitf EnantioEnzyme Reference preference sourceb

CH3, CzHs, n-C3H7, n-C4H9, n-C6HI3 H

H

H

H

++

H H

1 S‘

mEHd

127

R

BEH

111

R

BEH YEH

111 93

YEH YEH YEH YEH FEH FEH FEH

93 93 51

+

n. d. R 2s S n. d. 2s

92 92 92

++

2R

FEH

92

p-CH3, o-C~, * p-c1 H CH3 H * H H p-F,p-Cl, + p-Br, p-CH3 H H p-CH3 + H H o-CH3, o-Hal H H H f CH3 H H ++ H H H ++ H CH3 H H

H

H H H H H H indene oxide, dihydronaphthalene oxide H CH3 H

H

S

S

51

a Selectivity denoted as (-) =low ( E < 4). (i)= moderate ( E = 4-12), (+) =good ( E = 13-50), (++) = excellent ( E >50).

b BEH =bacterial epoxide hydrolase; FEH = fungal epoxide hydrolase; mEH = microsomal epoxide hydrolase from liver tissue; YEH =yeast epoxide hydrolase. n.d. = not determined. c Enantioconvergent process (i.e. a single stereoisomeric diol was formed as the sole product). d Performed on a microgram-scale only.

rather low[”1], the E-values could be significantly increased by using specific mutants of the Agrobacterium enzyme[”]. Disubstituted Epoxides. Among the sterically more demanding substrates, 2,2-disubstituted epoxides were hydrolyzed with virtually absolutly enantioselectivities (E >200) using enzymes from bacterial sources [Fig. 11.2-9(c),Table 11.2-51.In particular, Actinomycetes such as Rhodococcus and (closely related) Nocardia sp. are the biocatalysts of choice for this class of oxiranes [”*I. Epoxide hydrolases from Chryseomonas l u t e ~ l a [ ’and ~ ~ ]several 941 were less useful. Also for yeasts a 2-alkyl substituent resulted in a dramatic decrease in enantioselectivity[”].In several cases, the regioselectivityof the reaction has been determined to be absolute. Attack occurs exclusively at the less hindered unsubstituted oxirane C-atom with complete retention at the stereogenic center. Most bacterial epoxide hydrolases showed a preference for the (S)-enantiomer.Only recently, it was shown that several methylotrophic bacterial strains exist, which show an opposite preference (i.e. for the ( R ) epoxy enantiomer), albeit in moderate selectivitiesI l l 2 ] . On the contrary, mixed regioselectivities were common when 2,3-disubstituted oxiranes were hydrolyzed and ring opening ocurred at both positions of the oxirane ring at various ratios (Table 11.2-6)[lls]. This is understandable, bearing in mind that the steric requirements are similar at both positions. As a consequence, E-values are not applicable to the description of stereoselectivities. Again, Actinomycetes were found to be the catalysts of choice for this group of Most remarkably, in selected studies, it was proven that the hydrolysis proceeded in an enantio-

7 1.2 Hydrolysis ofEpoxides

I

595

Table 11.2-5.

Ri

Enzymatic hydrolysis o f 2,2-disubstituted epoxides, see Fig. 9(c) Selectivitya

Rz

f f

f

+ ++

++ ++

Enantiopreference

Enzyme Sourceb

Reference

n. d. Ror Sc S S S S S S

BEH FEH BEH BEH BEH BEH BEH BEH

117 50 8 8 8 8 8 8

a Selectivity denoted as (-) =low (E < 4). (3=moderate (E = 4-12), (+) = good (E = 13-50), (++) =excellent IE > 50). I

,

b BEH = bacterial epoxide hydrolase; FEH = fungal epoxide hydrolase; YEH =yeast epoxide hydrolase. c Depending on the strain, the enantiopreference varied. n. d. = not determined. Table 11.2-6.

Enzymatic hydrolysis of 2,3-disubstituted epoxides, see Fig. 9(d).

Ri

Rz

R3

R4

H H H H H H H CH3 H H H C2HS H CH3

n-C4H9, n-CSH17 n-C4H9 n-CsH17 CH3,C2H5 (CHz)20H (CH2)zOCH3 CH3 H CH3 CH3 C2HS H CH3 H

H H H H H H H H CH3 H H H H H

n-C&17, n-CloH21 ++ ++ (CH2)ioOH (CH2)7C02H ++ n-C4H9, n-CsH11 ++ n-CsHl1 ++ n-C5H11 + n-CsH11 f n-CsH11 f H ++ CH3 ++ n-C4Hr, f L3H7 f n-C4H9 ++ n-C4Hg, n-CsHiI, ++ n-C~Hi3

Selectiviw EnantioEnzyme Reference preference Sourceb

2s 2s 2s 2s

2s 2s 2s 2R/2Sd

S

S 2s

1s 2s 1s

mEH mEH mEH mEH‘ mEHC mEH‘ FEH FEH YEH YEH BEH BEH BEHC BEH

137 137 137 138 138 138 50 50

51 51 8 8 89 8

a Selectivity denoted as (-) =low (E c 4), (3=moderate (E = 4-12), (+) = good (E = 13-50), (++) =excellent (E > 50). b BEH =bacterial epoxide hydrolase; FEH =fungal epoxide hydrolase; mEH = microsomal epoxide hydrolase

from liver tissue; YEH =yeast epoxide hydrolase. c Enantioconvergent process (i.e. a single stereoisomeric diol was formed as the sole product).

d Depending on the strain, the enantiopreference varied.

convergent fashion, and only one stereoisomeric diol was formed as the sole product. In contrast, fungi seem less appropriate for the hydrolysis of 2,3-disubstituted oxiranes Lg4], whereas Rhodotorula glutinis was more effective on the cisconfigurated analogs of this substrate class[’’]. Interestingly, in contrast to the bacteria, this yeast seems to operate via a classic kinetic resolution rather than an enantioconvergent pathway. In this way, the simple choice of the appropriate microorganism gives access to either the optically pure epoxide (yeast) or the optically pure diol (bacteria).

596

I

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.2-7.

Enzymatic hydrolysis of trisubstituted epoxides, see Fig. 9(d).

Ri

R2

H

(CHZ)~C(OAC)(CH~)C CH3 CH3 H=CH2 Ph CH3 CH3 1,2-limonene oxide 1-methylcyclohexeneoxide

H

R3

Rq

Selectivitya EnantioEnzyme preferenceb Source'

Reference

+

1s

BEH

119

-

-

++ ++

S

FEH YEH BEH

92 51 121

2s

Selectivity denoted as (-) =low (E < 4), (+) =moderate (E = 4-12), (+) =good (E = 13-50), (++) = excellent (E > 50). b Configuration of preferentially attacked oxirane carbon atom. c BEH =bacterial epoxide hydrolase; FEH = fungal epoxide hydrolase; YEH =yeast epoxide hydrolase. a

Trisubstituted Epoxides. To date, only a limited set of data are available on the enzymatic hydrolysis of trisubstituted epoxides (Table 11.2-7).Regardless of their steric bulkiness, however, they seem to be accepted by epoxide hydrolases from bacterial['". '*'I, fun gal["^ 941 and yeast["] sources, as long as the access to one side of the substrate is not too severely restricted (e.g. a 2,2-dimethyl-3-alkyloxirane). Further data are required to depict a general selectivity pattern within this group of substrates. 11.2.3.3 Deracemization Methods

In contrast to the asymmetrization of meso-epoxides, which would lead to the highly desirable formation of a single stereoisomeric vicinal diol in 100% theoretical yield, the kinetic resolution of racemic epoxides by fungal and bacterial cells has proven to be highly selective (see above). However, this latter technique forms both the unreacted epoxide and the corresponding vicinal diol in equal amounts. This socalled classic kinetic resolution pattern of the biohydrolysis is often regarded as a major drawback, since the theoretical chemical yield can never exceed 50% based on the racemic starting material. As a consequence, methods that offer a solution to this intrinsic problem are highly advantageous[120]. Several procedures which overcome this drawback have been reported in the last few years. For instance, based on the finding that styrene oxide could be resolved by whole cells of Aspergillus niger and Beauveria bassiana via two different pathways showing matching enantio- and regioselectivities a deracemization was developed thus, combination of both biocatalysts in a single reactor led to (R)-phenylethane-l,2-diol as the sole product in 98% ee and 8 5 % isolated yieldLssl (Fig. 11.2-7). Another strategy for the achievement of an enantioconvergent process was set up using the combination of bio- and chemo-catalysis[107r*09, 12', 1221 . For instance, 2,2-disubstituted epoxides were selectively resolved by lyophilized whole cells of Nocardia sp. The biohydrolysis proceeds via attack at the less substituted C-atom with excellent regioselectivity thus leading to retention of configuration at the stereogenic center. On the other hand, acid-catalyzed hydrolysis of such epoxides usually proceeds at the more substituted oxirane carbon with inversion. Careful

11.2 Hydrolysis ofEpowides

Lyophilized cells of Nocardia sp. EH1 or H8 rac

*

E >lo0

R O

A

+

:A R'

(S) -diol e.e. >95%, yield >90%

(R)-epoxide

4

I HpSO

cat. I dioxane I trace H 2 0

Figure 11.2-10. Resolution-inversion sequence f o r t h e deracemization of 2,2-disubstituted oxiranes involving t h e remaining epoxide.

combination of both catalytic steps (Fig. 11.2-10) in a resolution-inversion sequence yields the corresponding (S)-1,2-diolsin virtually enantiopure form and in high yields (> 90%) [Io7. In a similar fashion, racemic para-nitrostyrene oxide was deracemized using a crude enzymatic extract from Aspergillus niger (Fig. 11.2-11). In this case a 4:l water-DMSO solvent mixture was used, showing that this enzyme is operative in the presence of water miscible organic solvents. The resolution step was followed by the careful addition of acid, leading to (R)-para-nitrostyrenediol in good yield (94%) and ee (80%).Because of the reduced enantioselectivity and the fact that racemization occurred to a certain extent during the acidic hydrolysis, it was necessary to tune both catalytic steps very carefully. A mathematical method was therefore developed that made it possible to select the optimum conversion at which the acid hydrolysis step should be initiated[122.1231. Careful mechanistic analysis of the acidic hydrolysis reaction, using different solvents and mineral acids, made it possible to select general conditions for the resolution-inversionprocedure['071.As a consequence, large scale deracemization became feasible The compatibility of microbial epoxide hydrolases with organic solvents deserves a special comment. It has been reported that in the majority of cases, the addition of water-miscible or -immiscible organic (co)solvents has negative effects on the activity. This is particularly true for bacterial enzymes, which showed total deactivat i ~ n [ ' ~On ~ ]the . other hand, several epoxide hydrolases from yeasts and fungi seem

-- fl- fl enzymatic hydrolysis

recryst.

chemical hydrolysis

O2N

( R)-nifenalol overall yield 58%, e.e. 99%

yield 78%

e.e. 80% yield 94%

O2N

e.e. 99%

cyclisation yield 89%

e.e. 99%

Figure 11.2-11. Deracemization ofpara-nitrostyrene oxide by a chemoenzymatic process. Application t o t h e synthesis o f (R)-Nifknalol@.

I

597

598

I

1 1 Hydrolysis and Formation of C - 0 Bonds

-

p 4

Lyophilized Bacterial Cells

R1

HO

OH

Figure 11.2-12. Resolution and deracernization of 2,3-disubstituted oxiranes by bacterial cells.

R' (R,S)- diol

rac-trans

Lyophilized Bacterial Cells

R1

R' (R,R)-diol

rac-cis

to be less sensitive and are able to tolerate water-misciblecosolvents, such as DMSO at a low Cases for a non-classic deracemization of racemic epoxides using one single biocatalyst impose high requirements on matching regio- and enantioselectivities, and are therefore rare. For instance, the enantioconvergent hydrolysis of (*)-3,4-epoxytetrahydropyran I'[ and several cis-p-alkyl substituted styrene oxide~[''~] by hepatic microsomal epoxide hydrolase has been reported on an analytical scale. Similarly, soybean epoxide hydrolase converted (*)-cis-9,1O-epoxy-l2(Z)-octadecenoic and (~)-cis-12,13-epoxy-9(Z)-octadecenoic acid into the corresponding (R,R)dihydroxy acids as the sole products [lzsl. However, enantioconvergent hydrolysis on a synthetically useful scale was only reported recently. Thus, the fungus Beauveria bassiana transformed (*)-cis-P-methylstyrene oxide in an enantioconvergent manner to afford (lR,2R)-l-phenylpropane-1,2-diol in 85% yield and 98% eeI9'1. In a related fashion, 2,3-disubstituted epoxides were hydrolyzed by using the Nocardia EH 1 (Table 11.2-8)[89s12'1 . Thus, the biohydrolysis of cis-2,3-epoxyheptanefurnished (R,R)-threo-2,3-heptane diol in 79% isolated yield and 91 % ee on a gram scale. In the latter study, the four stereochemical pathways and the enzyme mechanism were elucidated by "0Hz-labeling experiments. The hydrolysis was shown to proceed by attack of a (formal) hydroxyl ion at the (S)-configuratedoxirane carbon atom with concomitant inversion of configuration at both enantiomers with opposite regioselectivity. In addition, a mathematical model for the kinetics which allows the optimization of such enantioconvergent processes in preparative applications was developed. Table 11.2-8. Selectivities in the deracemization of 2,3-disubstituted oxiranes by bacterial cells, see Fig. 9(d).

Ri

Rz

R3

Rq

Biocatalyst

Configuration ofdiol

ee [%]

n-C4H9 n-C3H7 n-CSHi1 n-GjH13 n-C4H9 n-C,H7

H H H H H H

H H H H CH3 C2H5

CHI C2H5 CH3 CH3 H H

NocardiaEHl Arthrobacter sp. DSM 312 Rhodococcus SP. NCIMB 11 216 Rhodococcus SP. NCIMB 11 216 Nocardia EH1 NocardiaTBl

2R, 3 s 2R, 3s 2R, 3s 2R, 3 s 2R. 3R 2R, 3R

90 63 77 78 97 77

17.2 Hydrolysis ofEpoxides

11.2.4 Use of Non-Natural Nucleophiles

In reactions catalyzed by hydrolytic enzymes of the serine-hydrolase type, which form covalent acyl-enzyme intermediates during the course of the reaction, it has been shown that the “natural” nucleophile (water) can be replaced with “foreign” nucleophiles [1301 such as an alcohol, amine, hydroxylamine, hydrazine and even hydrogen peroxide. As a consequence, a wealth of synthetically useful reactions, which are usually performed in organic solvents at low water content, can be performed in a stereoselective manner. Although one requirement is fulfilled by epoxide hydrolases - i. e. a covalent enzyme-substrate intermediate is formed - the sensitivity of epoxide hydrolases to most of the water-miscible or -immiscible organic solvents [49, 1241 poses a general problem in the use of non-natural nucleophiles in enzymatic epoxide hydrolysis. However, two types of transformations, i.e. the aminolysis and azidolysis of an epoxide have been reported for selected cases (Fig. 11.2-13). When racemic aryl glycidyl ethers were subjected to aminolysis in aqueous buffer catalyzed by hepatic microsomal epoxide hydrolase from rat, the corresponding (S)configurated amino-alcohols were obtained in 5 1 4 8 % On the other hand, when azide was employed as nucleophile for the asymmetric opening of 2-methyl1,2-epoxyheptane in the presence of an immobilized crude enzyme preparation derived from Rhodococcus sp., which contains an epoxide hydrolase activity, the reaction revealed a complex The (S)-epoxide from the racemate was hydrolyzed (as in the absence of azide), and the less readily accepted (R)-enantiomer was transformed into the corresponding azido-alcohol (ee >GO %). Although at present only speculations can be made about the actual mechanism of both the aminolysis and azidolysis reaction, in both cases it was proven that the reaction was catalyzed by a protein and that no reaction was observed in the absence of biocatalyst

“ 7 Ph

R =ti, CI

rac

Figure 11.2-13.

e.e. 51-88%

e.e. >60%

Enzyme-catalyzed arninolysis and azidolysis of epoxides.

Ph

I

599

600

I

11 Hydrolysis and Formation ofC-0 Bonds

or by using a heat-denatured preparation. However, a recent related report on the aminolysis of epoxides employing crude porcine pancreatic l i p a ~ e lmay ~~~ likewise ] be explained by catalysis of a chiral protein surface rather than true lipase catalysis, since the latter enzyme - being a serine hydrolase - is irreversibly deactivated by epoxides. In view of these facts, it remains questionable whether the use of nonnatural nucleophiles will be of general applicability with epoxide hydrolases. 11.2.5

Applications to Asymmetric Synthesis

Although the use of an epoxide hydrolase for the asymmetric hydrolysis was reported for industrial synthesis of L- and meso-tartaric acid as early as 1969 r60], it was only recently that applications to asymmetric synthesis appeared in the literature. This fact can be attributed to the limited availability of these biocatalysts from sources such as mammals or plants. Since the production of large amounts of crude enzyme is now feasible, preparative-scaleapplications are getting within reach of the synthetic chemist. For instance, fermentation of Nocardia EH1 on a 70-L scale afforded >700 g of lyophilized cells ['*I. One of the first applications of the microbial hydrolysis of epoxides for the synthesis of a bioactive compound is based on the the resolution of a 2,3-disubstituted epoxy-fatty acid having a cis configuration (Fig. 11.2-14).Thus, by using an enzyme preparation from Pseudomonas sp., the (9R,lOS)-enantiomerwas hydrolyzed in a trans-specificfashion (i.e. via inversion of configuration at C-10) yielding the (9R,lOR)-threo-diol.The remaining (9S,lOR)-epoxidewas converted into (+)-disparlure, the sex pheromone of the gypsy moth in >95% Another illustration of the use of such a biocatalytic approach was the synthesis of either enantiomer of a-bisabolol. One of the stereoisomers is of industrial value for the cosmetic industry. This approach was based on the diastereoselective hydrolysis of a mixture of oxirane-diastereoisomers obtained from ( R ) - or (S)-limonene (Fig. 11.2-15) r9O]. Thus, starting from (S)-limonene,the biohydrolysis of the mixture of (4S,8RS)-epoxidesled to unreacted (4S,8S)-epoxideand (4S,8R)-diol.The former

Pseudomonas sp.

buffer

-CO*H rac-cis

Ho&co2H HO 9R

+

.

(+)-Disparlure Figure 11.2-14.

Resolution of a cis-2,3-disubstitutedepoxide and synthesis of disparlure

I

71.2 Hydrolysis ofEpoxides

601

(p yield 63%

(-)-(4S) -1irnonene e.e. 99%

(-)-(4 S,8S)-a-bisabolol e.e. 99%, d.e. 94%

(4S, 8 R)

d.e. 94%

I

steps

I Aspergillus

OH

(4s,8S)

(4 S, 8 R)

Figure 11.2-15.

(-)-(4 S ,8R)-epi -a-bisabolol e.e. 99%, d.e. >95%

CulnHF

d.e. 98%

d.e. 94%

Chemoenzymatic synthesis ofa-bisabolol using fungal epoxide hydrolase.

29%

72%

OH

1) Nocardia EH1 2) H2SO,, dioxane

1 ) Pd

+2fcat./CuCIq/DME

'1,

L

o,

2) HCI, r.t. 89%

99% e.e.

(S)-(-)-frontalin 99% e.e

Figure 11.2-16.

Chemoenzymatic synthesis of (S)-(-)-frontalin using bacterial epoxide hydrolase.

showed a high diastereomeric purity (de >95%) and was chemically transformed into (4S,SS)-a-bisabolol. The formed diol (de >94%) could be cyclized back to the corresponding (4S,BR)-epoxide,thus affording access to another stereoisomer of abisabolol. In addition, the two remaining stereoisomers of bisabolol could be prepared in a similar manner starting from (R)-l'imonene. (Fig. 11.2-16) Based on the deracemization of (*)-3-methyl-2-(4-pentenyl)-oxirane using Nocardia EH1 and sulfuric acid in dioxane containing a trace amount ofwater (see above), (S)-2-methyl-hept-6-ene-1,2-diol was obtained in 97% yield and 99% ee["']. This intermediate was successfully applied in a short synthesis of (S)-

602

I

7 1 Hydrolysis and Formation of C-0Bonds Rhodococcus OAc

NCIMB 11216 epoxide hydrolase

OAc OH

9

I

MsCI, Et3N, 0 OC

OH

trans- linalool oxide d.e. 94%

Figure 11.2-1 7. Synthesis of cis- and

d.e. 98%

trans-linalool oxide. +

I

(R R) d.e. 98% MeOH, K2C03 reflux

I OH

cis- linalool oxide d.e. 98%

(-) -frontalin, a central aggregation pheromone of pine beetles of the Dendroctonus family [loGI. Enantiopure cis- and trans-linalooloxides are found in several plants and fruits and constitute the main aroma components of oolong and black tea. These compounds were prepared from 2,3-epoxylinalyl acetate (Fig. 11.2-17)[1191. The key step consists of a separation of the diastereomeric mixture of the starting epoxide by employing an epoxide hydrolase preparation derived from Rhodococcus sp. NCIMB 11216, which furnished the product diol and remaining epoxide in excellent diastereomeric excess (de >98%). Further follow-up chemistry gave both linalool oxide isomers on a preparative scale in excellent diastereomeric and enantiomeric purities. Both enantiomers of the biologically active Bower's compound, a potent analog of insect juvenile hormone[135],were prepared using Aspergillus sp. cells in 96% ee (Fig. 11.2-18).Subsequent biological tests showed that the (6R)-antipodewas about ten times more active than the (GS)-counterpartagainst the yellow meal worm Tenebrio molitor. Aspergillus niger was the biocatalyst of choice for the biohydrolysis of paranitrostyrene oxide (see above).A selective kinetic resolution using a crude enzyme extract of this biocatalyst, followed by careful acidification of the cooled crude reaction mixture, afforded the corresponding (R)-diolin high chemical yield (94%) and good ee (80%).This key intermediate could then be transformed via a four-step sequence into enantiopure (R)-nifenalol,a molecule with p-blocker activity, which was obtained in 58% overall yield (Fig. 11.2-11) r8', 1221. The natural (R)-(-)-isomer of mevalonolactone, a key intermediate in a broad spectrum of cellular biological processes and their regulation, was synthesized via

7 7.2 Hydrolysis ofEpoxides 1a3 &OR

Aspergi//us niger

k

e.e. 96% yield 36%

rac

e.e. 70% yield 48%

e.e. 96% Bower‘s compound

Synthesis of Bower’s compound.

Figure 11.2-18.

1) Nocardia EHl TRIS-buffer pH 7.5 _____)

P

H

2) H2S04 cat.

rac

v

C

H20/dioxane

o

2

H

1) LiAIH4TTHF

TsCW,

OH e.e. 94%

,

2) AC 20lDMAP

ho

HO 1) Na1O4/RuCl3 cat./ MeCN/CCI4/H 20 _____)

H O p C W o A c ,.’ OAc

2) HCI aq.

Figure 11.2-19.

1) K2C03/MeOH ____)

2) HCI aq.

(-)-( R)-rnevalonolactone e.e. >99%

Synthesis of (R)-(-)-mevalonolactone.

eight steps in 55% overall yield and >99% ee (Fig. 11.2-19). In the key step, the aforementioned enantioconvergent chemoenzymatic deracemization route was applied. Thus, 2-methyl-2-benzyl-oxiranewas deracemized on a 10 g scale using lyophilized cells of Nocardia EH1 and sulfuric acid. The product (S)-diolwas isolated in 94% chemical yield and 94% optical During the scale-up of this

604

I

I 7 Hydrolysis and Formation ofC-0 Bonds external recycling:

$- ,& J.

1) HBr/AcOH 2) KOH/MeOH

+ ~

Aspergillus niger

0

0

cell-free extract

i-Bu

,

OH OH

i-Bu

.

0

i-Bu

KMn04/H SO,

i-Bu

'

Figure 11.2-20.

(S)-ibuprofen Chemoenzymatic synthesis o f (S)-ibuprofen

biotransformation it was observed that the increase of the substrate concentration led to a fourfold enhancement of the enantioselectivity as compared to analytical scale test reactions Finally, a chemoenzymatic enantioconvergent procedure led to (S)-ibuprofen in four steps and 47 % overall yield (Fig. 11.2-20). The latter compound is a widely used antiinflammatory drug and pain remedy and is one of the top ten drugs sold worldwide[loo]. In the key step, the conditions for the enantioconvergent hydrolysis of para-iso-butyl-a-methylstyreneoxide was optimized (elevated substrate concentration at +4 "C) to afford the non-reacted epoxide in 295 % ee[l3']. After separation from the epoxide, the formed diol (70% ee) was recycled via a two-step sequence via the corresponding bromohydrin, which was cyclized back to give (+)-epoxide.The latter material was subjected to repeated biocatalyhc resolution in order to improve the economy of the process. 11.2.6

Summary and Outlook

Over the past few years, an impressive array of epoxide hydrolases has been identified from microbial sources. Due to the fact that they can be easily employed as whole-cellpreparations or crude cell-free extracts in sufficient amounts by fermentation, they are just being recognized as highly versatile biocatalysts for the preparation of enantiopure epoxides and vicinal diols. The future will certainly bring an increasing number of useful applications of these systems to the asymmetric synthesis of chiral bioactive compounds. As for all enzymes, the enantioselectivity of

References I605

microbial epoxide hydrolysis depends on the substrate structure and the type of enzyme involved. The data available to date indicate that the enantioselectivites of enzymes from certain microbial sources can be roughly correlated to the substitutional pattern of various types of substrates: red yeast give best selectivities with monosubstituted oxiranes, fungal cells are suited for styrene oxide-type substrates and bacterial enzymes are the catalysts of choice for more highly substituted 2,2- and 2,3-disubstituted epoxides. Since the first three-dimensional X-ray structure of an epoxide hydrolase has recently been solved, more will follow, which will improve the predictability of stereoselectivities. Given the data presented above, possible industrial applications of microbial epoxide hydrolases can be anticipated in the near future.

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7 7 Hydrolysis and Formation of C-0 Bonds

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Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

7 7.3 Hydrolysis and Formation ofclycosidic Bonds

I

11.3

Hydrolysis and Formation of Clycosidic Bonds

Chi-Huey Wong 1.3.1

Introduction

Carbohydrates are found in nature as components of a broad range of molecular structures [l-sO1. Attached to cell surface glycoproteins and glycolipids,they play vital roles in cellular communication processes ['-231, function as points of attachment for proteins such as antibodies, and serve as receptor sites for bacteria and viral particles [12, 13, 521. For example, the sialyl-Lewis X tetrasaccharide mediates the adhesion of neutrophils to the endothelial layer, an initial event in the inflammatory responsel". 45. 5 3 - 5 6 ] . ~1ycoprotein glycans can modulate protein folding and are involved in the sorting and trafficking of proteins to appropriate cellular sites[l. 21, 40, 41, 57, 581 Nature employs two groups of enzymes in the biosynthesis of oligosaccharides, namely those of the L e l ~ i r [ ~ ' -and ~ ~ ]non-Leloir pathways. Leloir enzymes are responsible for the synthesis of most glycoproteins and other glycoconjugates in mammalian systems. Glycoprotein glycans are typically classified as either N-linked or 0-linked, based on the linkage between the carbohydrate and the protein. Nlinked glycans are characterized by a j3-glycosidic linkage between GlcNAc and an asparagine 8-amide nitrogen. The majority of 0-linked glycans contain an aglycosidic linkage between GalNAc and a serine or threonine hydroxyl group. The addition of oligosaccharide chains to glycoproteins occurs post- or co-translationally in the endoplasmic reticulum and the Golgi apparatus r60]. N-linked oligosaccharides all contain the same basic core structure composed of GlcNAc and mannose residues. N-linked glycan biosynthesis involves the initial construction of a dolichyl pyrophosphoryl oligosaccharide intermediate in the endoplasmic reticulum catalyzed by GlcNAc-transferases and mannosyltransferases. This structure is then glucosylated, presumably as a signal for transfer of the oligosaccharide to the polypeptide. The entire oligosaccharide moiety is then transferred en bloc to an Asn residue of the growing peptide chain catalyzed by oligosaccharyltransferase[60. 6 3 , 641. ~h e Asn is typically part of the amino acid sequence Asn-X-Ser(Thr), where X#Pro or Asp[3o,60. 65-671 . B efore transport into the Golgi apparatus, trimming of the glycan by glucosidases I and I1 and a mannosidase reveals a core pentasaccharide (peptide-A~n-(GlcNAc)2-(Man)~). This structure is further processed by mannosidases and glycosyltransferases in the Golgi apparatus, resulting in either a high-mannose, complex, or hybrid type oligosaccharide. Sequential addition of monosaccharides then provides the fully-elaborated oligosaccharide chain. In contrast, the more structurally diverse 0-linked glycans are assembled within the Golgi apparatus by glycosyltransferases[*lS60]. In the most common route, GalNAc is initially appended to serine or threonine catalyzed by a UDP-Ga1NAc:polypeptide GalNAc-transferase. Monosaccharides are then added individually to the growing oligosaccharide chain by glycosyltransferases.

609

610

I

7 7 Hydrolysis and Formation ofC-0 Bonds

R HOO

&AcHN o G Ho

AcHNo-b(-O-)(-O 0 0

I 0-

I 0-

Dolichylpyrophosphate-linked oligosaccharide R = oligosaccharide, n = 9-15 R20 HO

HO OH

OH OH

Ganglioside GMI: R' = Gal 1,SGalNAc -, R2 = H Ganglioside GM2: R' = GalNAc -, R2 = H Ganglioside GM3: R' = H, R2 = H Ganglioside GD3: R' = H, R2 = NeuAc -

H ow& ; o& OH AcHN

OR

HO OH

+OH

Sialyl Lewis X antigen

NHAc

OH HO

0

Gal 1,2Gal 1,6Gal 1 Gal 1 f

Glycosyl phosphatidylinositol

2

j-;o&)Ho-..-&& NHAc

COZH

-03SHN0

OH

Heparin pentasaccharide

NeuAc 2,3Gal 1.4GlcNAc 1,PMan 1

Fuc

.N 6 \6 GlcNAc 1 - - - - - + 4Man 1,4GlcNAc 1.4GlcNAc 1N A s n NeuAc 2,3Gal 1,4GlcNAc 1,PMan 1 y 3

Figure 11.3-1.

Typical structure of N-linked complex glycan

oso3

7 1.3 Hydrolysis and Formation of Glycosidic Bonds

All mammalian cells, with the exception of erythrocytes, contain the necessary elements for glycosylation. In certain secretory cells, however, the preponderance of glycosyltransferases is greater fG81. The structures of some typical naturally-occurring glycoproteins, glycolipids,and oligosaccharides are illustrated in Fig. 11.3-1. The major classes of cell-surfaceglycolipids include the glycosphingolipids (GSLs) and glycoglycerolipids. Gangliosides IG91, or sialic acid-containingglycosphingolipids, are especially abundant on neural cell surfaces I7O]. These compounds play a role in the differentiation of cell types and in the regulation of cell growth. Additionally, sphingosine, the lipid component of GSLs, has been suggested to function as an intracellular second messenger [711. The mammalian glycosyltransferases of the Leloir pathway utilize monosaccharides activated as glycosyl esters of nucleoside mono- or diphosphates as glycosyl donor substrates Primarily eight nucleotide sugars serve as glycosyl donors for the synthesis of most oligosaccharides: UDP-Glc, UDP-GlcNAc, UDP-Gal, UDPGalNAc, GDP-Man, GDP-Fuc, UDP-GlcUA, and CMP-NeuAc (Fig. 11.3-1). Many other monosaccharides, such as the anionic or sulfated sugars of heparin and chondroitin sulfate, are also found in mammalian systems, but usually are the result of post-glycosyl transfer modifications 272 72a,b1. Non-Leloir glycosyltransferases typically employ glycosyl phosphates as activated donors. A diverse array of monosaccharides (e.g. xylose, arabinose, KDO) and oligosaccharides is also present in microorganisms, plants, and invertebrates[33* G2, 73-7G1 .The enzymes responsible for their biosynthesis, however, have not been extensively exploited for synthesis, though the same principles as in mammalian systems apply. Some sugar nucleotides used by enzymes of other pathways are also shown in Fig. 11.3-2. Chemists have employed glycosyltransferases from the Leloir and non-Leloir pathways for the synthesis of oligosaccharides and glycoconjugates[77-831. Glycosidases have also been exploited for synthesis [77-841. The function of glycosidases in vivo is to cleave glycosidic bonds; however, under appropriate conditions, they can be useful synthetic catalysts. Each group of enzymes has certain advantages and disadvantages for synthesis. Glycosyltransferasesare highly specific in the formation of glycosides, but the availability of many of the necessary enzymes is limited. Glycosidases have the advantage of wider availability and lower cost, but they are not as regio-specificor high-yieldingin synthetic reactions. Therefore the chemist must choose the enzyme which is best suited for the application at hand. Other enzymatic methods used to synthesize glycoconjugateswill also be discussed. 11.3.2 Clycosyltransferases o f the Leloir Pathway

Glycosyltransferases are highly regiospecific and stereospecific with respect to the formation of new glycosidic linkages. Although also usually substrate-specific, minor chemical modifications are tolerated on both the donor and acceptor components. The preparative use of glycosyltransferaseshas been somewhat limited in the past because of a lack of enzyme availability. Additionally, because glycosyltransferases are membrane-bound enzymes, they are relatively unstable and can be

I

612

I

7 1 Hydrolysis and Formation ofC-0 Bonds 0

n

NHz

HO

OH

HO

OH

a-GDP-Mannose (GDP-Man)

a-UDP-Glucose (UDP-Glc)

VD

HO& HO

HO

AcHN

HobUDP

a-UDP-Galactose (UDP-Gal)

HO OH OUDP

P-GDP-L-Fucose (GDP-FUC)

a-UDP-KAcetylglucosarnine (UDP-GIcNAc)

$ +

0

HO HO

HoOUDP

AcHN HO OH

a-UDP-KAcetylgalactosarnine (UDP-GalNAc)

a-UDP-Glucuronic acid (UDP-GlcUA)

P-CMP-N-Acetylneurarninic acid (CMP-NeuAc) 0

HO

HO

P-TDP-L-Rharnnose

OCMP

OH

b-CMP-KDO

a-UDP-N-Acetylrnuramicacid

Figure 11.3-2.

difficult to handle in solution. However, the recent isolation of many of these enzymes, as well as advances in genetic engineering and recombinant techniques, are rapidly alleviating these drawbacks. Glycosyltransferases utilize nucleotide sugars as activated glycosyl donors [Go]. Most of these sugar nucleoside phosphates are biosynthesized in uiuo from the corresponding monosaccharides (Fig. 11.3-3). The initial step is kinase-mediated phosphorylation to produce a glycosyl phosphate. This glycosyl phosphate then reacts with a nucleoside triphosphate (NTP), catalyzed by a nucleoside diphosphosugar pyrophosphorylase, to afford an activated nucleoside diphosphosugar [Eq. (l)]. Some sugar nucleoside phosphates, such as GDP-Fuc and UDP-GlcUA, are biosynthesized by further enzymatic modification of existing key sugar nucleotides.

Gal

Glc

-

11.3 Hydrolysis and Formation of Clycosidic Bonds

-

Gal-I-P

Glc-6-P

Glc-1-P

11 1

-

Fru-6-P

Man-6-P -Man-I-P

GkN-6-P

Man

1

GlcNAc-6--

I--

I

613

-4DP-Xyl

GIcNAc

ManNAc-6-P-N&AC-~-P

Figure 11.3-3.

In contrast, CMP-NeuAc is formed by the condensation of NeuAc with CTP [Eq. (2)]. Some of the enzymes involved in the biosynthesis of sugar nucleotides also accept unnatural sugars as substrates. In general, however, the rates are quite slow, thus limiting the usefulness of this approach. Sugar-1-P+ NTP NDP-Sugar + PPi (1) NeuAc + CTP CMPNeuAc + PPi (4 -+

+

11.3.2.1

Synthesis of Sugar Nucleoside Phosphates

Chemical syntheses of some sugar nucleoside phosphates have been reported[85]. Most of these methods involve the reaction of an activated NMP[8G”]with a glycosyl phosphate to produce a sugar nucleoside diphosphate (Fig. 11.3-4). Of the commonly used activated NMP derivatives, phosphoramidates such as phosphorimidazolidates [92-941 and phosphoromorpholidates [86-911 are considered the most effective. A recent improvement in coupling methodology employing 1 H-tetrazole as catalyst has been reported[”]. These activated NMPs may also be used to prepare

614

I

11 Hydrolysis and Formation of C-0 Bonds .OH

0 I1

-0-P

0

0 I1

I1

-0 - P -0 -P - 0

0I

*-

0. I

0. I

d

Figure 11.3-4.

XMP

XDP * *

XTP

OP

XTP

0

Jco, 0

A 0 ;

OP

A o 2

1. Adenylate kinase (EC 2.7.4.3, X = A,C,U) Guanylate kinase (EC 2.7.4.8, X = G) Nucleoside monophosphate kinase (EC 2.7.4.4, X = U) 2. Pyruvate kinase (EC 2.7.1.40)

Figure 11.3-5.

NTPs by reaction with pyrophosphate (Fig. 11.3-5)[88].A number of chemical methods are available for the synthesis of glycosyl phosphates. Reactions of phosphates with activated glycosyl donors [", 961 or chemical phosphorylation of anomeric hydroxyl groups [89-92, 971 have proven to be convenient. Additionally, routes via glycosyl phosphites are useful["]. Enzymatic procedures include employing glycogen phosph~rylase['~] and sucrose phosphorylase["'] for the production of aglucose-1-phosphate. Phosphoglucomutase can also be used to prepare glucose1-phosphate from glucose-G-phosphate[lO1], the latter generated from glucose by hexokinase catalysis. Preparative-scale synthesis of nucleoside triphosphates. Nucleoside triphosphates are utilized as substrates for the biosynthesis of sugar nucleoside phosphates. Practicalscale biosynthesis-based enzymatic preparation of NTPs for use in glycosylations is therefore required. Most preparative-scale enzymatic syntheses of NTPs use commercially available NMPs as starting materials. Alternatively, NMPs can be obtained from yeast RNA digests at low cost["*], or can be easily prepared chemi~ally~''~]. In general, enzymatic methods involve the sequential use of two kinases to transform NMPs to NTPs, via the corresponding NDPs. Several kinases have been utilized to synthesize NTPs from the corresponding NDPs, each employing a different phosphoryl donor: pyruvate kinase (E. C. 2.7.1.40) uses phosphoenolpyruvate "1' as a phosphoryl donor, acetate kinase (E. C. 2.7.2.1) uses acetyl phosphate, and nucleoside

11.3 Hydrolysis and Formation ofClycosidic Bonds

1

XTP phosphoglyceromutase

chemical synthesis

diphosphate kinase (E. C. 2.7.4.6) uses ATP. Pyruvate kinase has generally been the enzyme of choice because it is less expensive than nucleoside diphosphate ki94, lo7], and because PEP is more stable and provides a more thermodynamically favorable driving force for phosphorylation than does acetyl phosphate (Fig. 11.3-5). A recently described polyphosphate kinase uses polyphosphate as donor, providing a potentially cheaper kinase route [lo6]. The preparation of NDPs from NMPs is more complicated, and requires different enzymes for each NMP. Adenylate kinase (E. C. 2.7.4.3) phosphorylates AMP[88]and CMP[1081,and also slowly phosphorylates UMP. Guanylate kinase (E. C. 2.7.4.8) catalyzes the phosphorylation of GMP. Nucleoside monophosphate kinase (E. C. 2.7.4.4) uses ATP to phosphorylate AMP, CMP, GMP, and UMP; however, the enzyme is relatively expensive and Both CMP and UMP kinases exist but are not commercially available. For those kinases requiring ATP as a phosphorylating agent, ATP is usually used in a catalFc amount and recycled from ADP using pyruvate kinase/PEP or acetate kinaselacetylphosphate[77, lo9].Phosphoenolpyruvate may be prepared chemically from p y r u ~ a t e [or ~ ~generated ~] enzymatically from D-3-phosphoglycericacid['''] (Fig. 11.34). When chemical and enzymatic methods for NTP synthesis are enzymatic techniques provide the most convenient route to CTP and GTP, whereas chemical deamination of CTP is the best method for preparing UTP[941.ATP is relatively inexpensive from commercial sources, although it has been synthesized enzymatically from AMP on 50 mmol scale. Mixtures of NTPs can be prepared from RNA by sequential nuclease PI, polynucleotide phosphorylase, and pyruvate kinasecatalyzed reactions [llO]. This mixture can be selectively converted to a sugar nucleotide using a particular sugar nucleoside diphosphate pyrophosphorylase['lo]. UDP-glucose ( UDP-Glc) and UDP-galactose ( UDP-Gal). UDP-glucose has been prepared from UTP and glucose-1-phosphate under catalysis by UDP-glucose '13, '141. UDP-Ga1 can be synthesized in an pyrophosphorylase (Fig. 11.3-7)[94, analogous fashion using UDP-Gal pyrophosphorylasellO1],or from UDP-Glc by epimerization of C-4 with UDP-glucose epimerase['O'l (Fig. 11.3-7). Though the epimerase equilibrium favors UDP-Glc, the reaction can be coupled to an in situ glycosylation with galactosyltransferase to shift toward UDP-Gal production. The latter process has been applied to large-scale synthesis of N-acetyllactosamine (L~CNAC)['~']. UDP-Gal has been prepared from UMP and Gal using dried cells of 2' ' '

I

615

616

I

7 7 Hydrolysis and Formation of C - 0 Bonds

/&

HO HO

UTP, UDP-Glc DvroDhosDhorvlase

HO ~

Ho 0PO32.

K = 0.17

HO

ROH, GalT

7

UDP-Glc 4-epimerase

OR

HO OH

Figure 11.3-7.

E2: pyruvate kinase

Pyruvate

PEP

E3: phosphoglucomutase Ed: UDP-Glc pyrophosphorylase

E5 inorganic pyrophosphatase

Figure 11.3-8.

Towlopsis candida["'l. In this system, Gal-1-phosphate and UTP were generated in situ as substrates for UDP-Gal pyrophosphorylase. Gram quantities of UDP-Gal, as well as the 2-fluoro-UDP-Gal derivative have been synthesized by an enzymatic method employing Gal-1-P uridyltransferase UDP-N-acetylglucosamine ( UDP-GlcNAc). UDP-GlcNAc has been synthesized by reaction between GlcNAc-1-phosphate and UTP, catalyzed by UDP-GlcNAc pyrophosphorylase [lll].Although this enzyme is currently not commercially available, a whole-cell process using baker's yeast can be employed ["'I. Another procedure exploits UDP-Glc pyrophosphorylase to catalyze a condensation between UTP and glucosamine-1-phosphate (GlcN-I-P) to afford UDP-glucosamineI'121 (Fig. 11.3-8). The product UDP-GlcN can then be selectively N-acetylated to provide UDP-GlcNAc. GlcN-1-Phas been synthesized from GlcN by phosphorylation of the 6-position with hexokinase to give GlcN-G-P, followed by a phosphoglucomutase-catalyzed isomerization to provide GlcN-1-P. UDP-GlcNAc also serves as an acceptor for p1,4GalT to provide UDP-L~CNAC[~"]. UDP-N-acetylgalactosumi~e ( UDP-GalNAc). The biosynthetic enzymes UDP-GalNAc pyrophosphorylase and UDP-GlcNAc 4-epimerase are not readily available for facile synthesis of UDP-GalNAc.An alternative synthetic procedure based on UMP exchange between UDP-Glc and GalN-1-P, catalyzed by commercially available UDP-Glc: galactosylphosphate uridyltransferase (E. C. 2.7.7.12) has been reported (Fig. 11.3-9)[62. 'I3]. Galactose-1-P is the natural substrate for the enzyme, but 2-deoxygalactose-l-P, 2-deoxyglucose-l-P, and galactosamine-1-P are also tolerated.

HHO O

G

+

11.3 Hydrolysis and Formation ofclycosidic Bonds I617

Hr$

HoOUDP

H2N

El

H HO O

G

+

Hr$

lACZO lE2 H2NOUDP

H00P032-

0PO32.

E, = UDP-G1c:galactosylphosphate

HO

uridyltransferase

E2 = phophoglucomutase

H HOO

G

o

OH

Ho%

H

HO

AcHN

OUDP

Figure 11.3-9.

As the equilibrium constant for the exchange reaction is close to unity, phosphoglucomutase was required to relieve product inhibition and shift the equilibrium. The UDP-GalN thus produced was then chemically acetylated to give UDP-GalNAc. A modification of the latter procedure has been adapted to large-scalesynthesis of UDP-GalNAc["8]. In this procedure, UDP-Glc was regenerated in situ from UTP and the product Glc-1-P under catalysis by UDP-Glc pyrophosphorylase. This also shifts the equilibrium toward the formation of UDP-GalN. Alternatively, UDP-GalNAc can be prepared from UMP and sucrose employing sucrose ~ y n t h a s e [ ~Large-scale ~~]. production of UDP-GalNAc in yeast has also been accomplished[120]. GDP-mannose (GDP-Man).GDP-mannose has been prepared from Glc and GMP using dried baker's yeast cells ['"I. The procedure involves the biocatalytic conversion of glucose to Man-1-P and subsequently to GDP-Man using GDP-Man pyrophosphorylase. A cell-free extract from baker's yeast has also been used to synthesize GDP-Man from mannose"'']. A direct synthesis from chemicallyprepared Man-1-P and GTP, catalyzed by GDP-Man pyrophosphorylase (E. C. 2.7.7.13) is useful for large scale producation (Fig. 11.3-10)Lg4]. Alternative strategies for continuous GDP-Man production["'], the synthesis of GDP-Man directly from and other routes have also been pursued[124]. mannose GDP-ficose (GDP-Fuc). GDP-fucose is biosynthesized in vivo from GDP-Man by an NADPH-dependentoxidoreductase enzyme system. Such systems have also been utilized for in vitro syntheses of GDP-Fuc. For example, the synthesis of GDP-Fuc was accomplished using a crude enzyme preparation from Agrobacterium radioba~ter[l'~]. NADPH was regenerated in situ from NADP using glucose-6-phosphate dehydrogenase and Glc-6-P[12G]. Employing a similar procedure, GDP-Fuc has been Baker's Glc + GMP

yeast

HFq

[o*EGDP]

NADPH

HO

OGDP GTP, GDP-Man Man-1-P

t

DvroDhosoholvlase

Fucose

fucokinase

- A GDP-Fuc pyrophosho lase

p-tucose-,-p

GTP

Figure 11.3-10.

VDP pp,

618

I

1 1 Hydrolysis and Formation of C - 0 Bonds Figure 11.3-11. HO

2 NADH

L-LDH

0

OH

H HO O

h

o H NHAc

-HraoH

El: NeuAc aldolase

EZ CMP-NeuAc synthetase

HO-

Ho

~

A El0 2 - *

H o AcHN

a

C

O

z

H

HO OH

XCDP f

CTP

E4 E3 Adenylate Pyruvate kinase kinase

Pyruvate

PEP

H z@ H AcHN o: HO OH

Figure 11.3-12.

generated in situ for use in a glycosylation reaction with a-1,3-f~cosyltransferase~'~~]. Enzymes from a minor biosynthetic pathway which synthesize GDP-Fuc from Lfucose have also been exploited for Fucose was phosphorylated by fucokinase (E. C. 2.7.1.52)to produce Fuc-1-P, which subsequently underwent a GDP-fucose pyrophosphorylase-catalyzed reaction with GTP to provide GDP-Fuc. Several practical chemical syntheses of GDP-Fuc have also been reported[95,12']. UDP-glucuronic acid ( UDP-GkUA).UDP-Glucuronic acid is biosynthesized by C6 oxidation of UDP-Glc with UDP-Glc dehydrogenase, an NAD-dependent enzyme. Enzyme preparations from bovine liver have been employed for gram-scale syntheses of UDP-GlcUA (Fig. 11.3-11)[944. 1291. The NAD cofactor was regenerated with lactate dehydrogenase in the presence of pyruvate. Additionally, extracts from guinea pig liver have been used to generate UDP-GlcUA in situ for use in enzymatic glycosylations with glucuronyltransferases [1301. CMP-N-acetylneuraminic acid (CMP-NeuAc). CM P-N-acetylneuraminic acid has been prepared enzymatically on small scales (> 0.5 mmol) from CTP and NeuAc, under catalysis by CMP-NeuAc synthetase (EC 2.7.7.43)[l3l].An improvement in this procedure, involving in situ production of CTP from CMP under adenylate kinase and pyruvate kinase catalysis, is suitable for multigram-scale synthesis Adenylate kinase catalyzes the equilibration of CTP and CMP to CDP, which is subsequently phosphorylated by pyruvate kinase to provide CTP. A one-pot synthesis of CMP-NeuAcbased on this procedure involves the in situ synthesis of NeuAc from Nacetylmannosamine and pyruvate, catalyzed by sialic acid aldolase (Fig. 11.3-12)[1081. Chemical syntheses of CMP-NeuAc have also been

17.3 Hydrolysis and Formation of Glycosidic Bonds

The gene encoding E. coli CMP-NeuAc ~ y n t h e t a s e “ 1341 ~ ~ ,has been cloned into the Lambda ZAP vector and overexpressed in E. coli at a level 1000 times that of the wild type[307,3081. The enzyme from calf brain has also been cloned and overexpressed. CMP-NeuAc synthetase was shown to accept several NeuAc derivatives as substrates. For example, 9-deoxy-,7,9-dideoxy-,and 4,7,9-trideoxy-NeuAcare all converted to the corresponding CMP-NeuAc derivatives[137]. On the other hand, the 4-ox0,~-0xo, and 8-0x0 NeuAc derivatives are not substrates for CMP-NeuAc syntheta~e[~’~I. However, the enzyme accepts a variety of modifications at the 9-position, and the hydroxyl group can be replaced with several different groups with little effect on the KM value[139-141].CMP-NeuAc can also be obtained on the large scale by fermentation [1431 or by coupling of metabolically engineered bacterial cells I1&]. 11.3.2.2

Substrate Specificity and Synthetic Applications of Clycosyltransferases

For each sugar nucleotide glycosyl donor, many glycosyltransferases of varying substrate specificities exist. These enzymes are generally considered to be specific for a given glycosyl donor and acceptor, as well as for the stereochemistry and the linkage position of the newly formed glycoside bond. This specificity has led to the . I n other words, the specificity of the “one enzyme-one linkage” concept[28,142, lG1] glycosyltransferasesensures fidelity in oligosaccharide sequences in vivo without the use of a template scheme. Though systematic investigations of the in vitro substrate specificity of most glycosyltransferases have not been carried out, some deviations from this picture of absolute specificity have been observed in the tolerated modifications of both glycosyl donors and acceptors. Additionally, studies toward the design of inhibitors of glycoprotein biosynthesis[2051have also shown that the specificities of glycosyltransferases are not absolute. Galactosyltransferase (GalT). Because of its availability, P1,4-GalT (E. C. 2.4.1.22)[lG21471 is one of the most extensively studied mammalian glycosyltransferases with regard to synthesis and substrate specificity. The X-ray crystal structure of the bovine enzyme has recently been reported[148].P1,4-GalT catalyzes the transfer of galactose from UDP-Gal to the 4-position of p-linked GlcNAc residues to produce the Galpl,4GlcNAc (LacNAc) structure. In the presence of lactalbumin, however, both a- and P-linked substrates are allowed, and glucose is the preferred acceptor. P1,4-GalT has been employed in the in vitro syntheses of LacNAc and ~ ~ ~ ]11.3-1). glycosides thereof, as well as other g a l a c t o s i d e ~ [(Table P1,4-GalT also tolerates 2-deoxyglucose,D-xylose, 5-thioglucose,N-acetylmuramic acid, and myoinositol as acceptor substrates [147]. Modifications at the 3- or 6-position of GlcNAc are also accepted. For example, Fuca1,GGlcNAc and NeuAca2,bGlcNAc are substrates Acceptor substrates which are derivatized at the 3-position include 3-0-methyl-GlcNAc 3-deoxy-GlcNAc,3-O-allyl-GlcNAc~OBu, and 3-0x0G ~ C N A C D-Mannose, [~~~]. D-allose, D-galactose, D-ribose, and D-xylose do not serve as substrates. Monosaccharides which have a negative charge, such as glucuronic acid and a-glucose-1-phosphate, are also not accepted. Fig. 11.3-13 illustrates several disaccharideswhich can be synthesized with p1,4-GalT[’47]. A particulary interesting

620

I

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.3-1.

Products of galactosyltransferase reactions.

UDP-Gal (or analogs) + CalT

GalPl,4Glc Galpl,4GlcNAc Galp1,4GlcNAc-Agarose Gal~l,4GlcNAc-hexylarnine Galp1,4GlcNAcpl,4Gal Gal~l,4GkNAcpl,6Gal Galpl,4GlcNAcp1,3Gal Gal~1,4Glc~OCH2C~H~(NO~)-CONH-Polymer Gal~1,4Glc~l,4Glc~OCH~C~H~(NO~)-CONH-Polymer Gal~l,4Glc~1,4Glc~OCH~NH-~-Phe-CONH-Polymer Gal~1,4GlcNAc~l,3(Gal~l,4GlcNAc~l,G)Gal~l,4Glc~OMe Gal~1,4GlcNAc~1,6(GlcNAc~l,3)Gal~l,4Glc~OMe Galp1,4(Fucal,6)GlcNAc~O(CH~)~CO~Me Ga~~l,4(NeuAc(OMe)a2,G)GlcNAc~O(CHz)sCOZMe Galpl,4GlcNAcpR;R = N-Ac-Asn(0Me) Gal~1,4GlcNAcpl,4GlcNAc Gal!.31,4GlcNAc~1,4GlcNAc~R; R = N-Ac-Asn(0Me) G~~~~,~G~CNAC~O(CH~)~CO~M~

Scale"

Ref.

C A

[147] [142,1451 [147] 11471 (1471 [150] (1501 11511 [151] [151] [152] [152] [150] [lSO] [153] [153] I1531 [154] [154] [154] [154]

C

C

C

C C D D D C

C

D D C

C

C

D D GalNAc~l,4GlcNAc~1,2Man~O(CH~)~CO~Me D Ga~NAc~l,4GlcNAc~l,2Manal,6(GalNAc~1,4GlcNAc~1,2Manal,3) D GalNAcpl,4GlcNAc~O(CH~)&O2Me

Manp0(CH~)sCOzMe GlcNAc~1,4GlcNAc~O(CH~)~CO~Me Galpl,4GlcNAcpR R = GlyGlyAsnGlyGly or N-Alloc-PheAsnSerThrlle Galpl,3Galp1,4Glc Galal,3Galp1,4GlcNAc

a A, > 1

D C D D

g; B, 0.1-1 g; C, 10-100 mg; D, < 10 mg

+

HO

HO

NHAcOR

p1,4-GalT,Mn"

*

[154] [155] [156] 11561

H:go& HO

Ho

HoOUDP

OR

NHAc

H

Y

G HO ~

Ho & H

":Go--&

OH

HO

OH

Ho

NHAcOR

.:go&

R' = H, OH, OCH3,OCH&H=CH,

HO

OH

HO

R'

NHAcOR

Figure 11.3-13.

example is the P,P-l,l-linked disaccharide, in which the anomeric hydroxyl of 3-acetamido-3-deoxyglucose serves as the acceptor m ~ i e t y ~ 'The ~ ~ ]acetamido . function apparently controls the position of glycosylation.

17.3 Hydrolysis and Formation ofClycosidic Bonds

I

621

I

Fmoc-Thr(a-0GlcNAc)-COOH 1

* * SPPS, 2. deprotection

Ac-Lys-Pro-Pro-Asn-Thr-Thr-Ser-Ala 0

G O Y'NHAC

H(

'COOH

1. pl,4-GalT,

2. a2,3-SiaT, 3. a1, ~ - FucT 4. Pd(PPh3)4

Figure 11.3-14.

AcO

1. SPPS (RINK Amide Resin) HOHO &o:h 2 TFNH20

NHAc

AcHN

* Fmoc-HN

OH

0

3. NaOH, or

I

SOs-pyr, then NaOH

0

OR

1. p1,4-GalT, 2. a2,3-SiaT, 3. al,S-FucT

HA OH

OH GIu-NH,

Ac-Tyr-Asp-Phe-Leu-Pro-Glu-HN I

OR

0

Figure 11.3-15.

NHAc

Figure 11.3-16.

GIu-NH,

Ac-Tyr-Asp-Phe-Leu-Pro-Glu-HN

Protein

7 7 Hydrolysis and Formation of C-0 Bonds Table 11.3-2.

Relative rates of bl.4-CalT catalyzed transfer of donor substrates.

Donor substrate

Relative Rate

UDP-Gal

100

UDP-Glc

0.3

1165, 1541

5.5

~

UDP-4-deoxy-Glc

no

Ref.

5

1

UDP-Ara HoOUDP

UDP-GalNAc

UDP-GlcNAc

HO

UDP-GlcN

HO% HO

0

0.09 HzNOUDP

UDP-5-thio-Gal

H

O

q

5

HoOUDP

pl,4-GalT has also been employed in solid-phase oligosaccharide synthesis, and has been used to galactosylate gluco and cellobio subunits of polymer-supported oligosaccharides and polysaccharides[160]. The resulting oligosaccharides can then be removed from the support by either a photochemical cleavage or a chymotrypsinmediated hydrolysis. GlcNAc-amino acids and peptides have also been used as substrates for p1,4-GalT to afford galactosylated glycopeptides (Fig. 11.31 6 ) [ 1 5 3 , 155, 160. 1611 ~h . e carbohydrate chain can then be further extended with other “‘1, as was shown in the glycosyltransferases, such as SiaT and F u c T ~ ’ ~ ~ , enzymatic solid-phase synthesis of glycopeptides from MAdCAM-1 (Fig. 11.314)[“lb]. Furthermore, solid- and solution-phase techniques can be employed “‘9

11.3 Hydrolysis and Formation ofClycosidic Bonds

I

623

together for the synthesis of complex sulfated glycopeptides such as those from PSGL-1 (Fig. 11.3-15)['"d]. In terms of glycolipids, P1,4-GalT was utilized in the preparation of a ceramide-linked LacNAc glycoside that was further enzymatically sialylated to provide a GM3 analog['55r162]. With regard to the donor substrate, P1,4-GalT also transfers glucose, 4-deoxygalactose, arabinose, glucosamine, galactosamine, GalNAc, and 2-deoxyglucose from their respective UDP-derivatives.This flexibility provides an enzymatic route to oligosaccharides which terminate in Pl,4-linked residues other than galactose[326], such as 5-thiogalactose[1641 (Table 11.3-2).Although the rate of enzyme-catalyzed transfer for many of these unnatural donor substrates is quite slow, this method is useful for milligram-scale synthesis. Besides P1,4-GalT, other GalTs are also of interest synthetically.Recently, al,3-GalT has received a heightened focus because of its role in xenotransplantation studies. Several studies of substrate specificity and synthetic potential have also been carried Sialyltransferase (SiaT). a2,G- and a2,3-~ialyltransferasehave been used for oligosaccharide synthesis [166168] . Sialyltransferases generally transfer N-acetylneuramink acid (NeuAc)to either the 3- or 6-position of terminal Gal or GalNAc residues Table 11.3-3.

Products of sialytransferase reactions.

CMP-NeuAc + a2,6-SiaT

Scale"

Ref

D D C NeuAca2,6Galp1,4GlcNAc~OMe NeuAca2,6Gal~1,4GlcNAc~l,3GaI~l,4Glc NeuAca2,6Gal~l,4GlcNAc~1-N-Asn NeuAca2,6Gal~l,4GlcNAc~l,2ManaOMe NeuAca2,6Ga~~1,4GlcNAc~1,3(Ga~~1,4GlcNAc~1,6)Cal~l,4Glc~OMe NeuAc(9-O-Ac)a2,6Gal~1,4GlcNAc NeuAca2,6Galpl,4GlcNAc~R R = OH, NHs,GlyGlyAsnGlyGly or N-Alloc-PheAsnSerThrIle NeuAca2,6Ga~~l,4G1cNAc~l,4(NeuAca2,6Gal~l,4GlcNAc~l,2/3)Gal~O D (CH2)5C02Me

CMP-NeuAc + a2,S-SiaT

NeuAca2,3Galpl,4GlcpOMe NeuAca2,3Gal~l,4GlcNAc~OMe NeuAca2,3Galpl,3GlcNAc~OR; R = Me,Ph,(CHz)sC02Me NeuAca2,3Gal~1,3GlcNAcpl,3Gal~l,4Glc NeuAca2,3Gal~l,3GlcNAc~l,3Gal~O(CH2)8C02Me

NeuAca2,3Gal~l,3GlcNAc~1,6Gal~O(CH2)8C02Me NeuAca2,3Galpl,3GlcNAcpOR (R= Et) (R= H,(CH2)5COzMe) NeuAca2,3Galp1,3(NeuAca2,6)GalNAc~OPh

3-O-Me-Gal~l,4Glc~l,6(NeuAca2,3Gal~l,4)GlcNAc~l,3Gal~l ,4Glcp1,6(NeuAca2,3Gal~1,4)GlcNAc~OMe a A, > 1 g; B, 0.1-1 g; C, l(L-100mg;

D,< 10 mg

D D D D D

D

C D D D

[166] [166] [166] [166] [254] [254] [I771 11671 [178] [179]

7 7 Hydrolysis and Formation of C - 0 Bonds

(Table 11.3-3)[1691. Some SiaTs accept CMP-NeuAc analogs which are derivatized at the 9-position with amino, fluoro, azido, acetamido, or benzamido groups[139, 140. 142c,d, 168-1701 A . zido-, phthalimido-, carbamate, and pivaloyl analogs of LacNAc and Galpl,3GalNAc are also substrates for the enzymes[172].Sialyltransferases have been used to append NeuAc to galactose on the terminus of glycopeptides [‘“I, glycolipids and glycoproteins [‘“I. Fucosyltransferase (FucT). Fucosyltransferases are involved in the biosynthesis of many blood-group substances and tumor-associated antigens. al,3-FucT L-fucosylates the GlcNAc 3-position of LacNAc and sialyla2,3LacNAc to provide the Lewis X and sialyl Lewis X structures, respectively Several other acceptor substrates with modifications in the GlcNAc residue [lactose, Galpl,4Glucal, Galp1,4(5-thioGlc)]can also be fucosylated by various FucT isozymes (Table 11.3-4)[182]. a-1,3/4-FucT fucosylates either the GlcNAc 3-position of Galpl,4GlcNAc or the GlcNAc 4-position of Galpl,3GlcNAc (as well as the sialylated versions) to afford (sialy1)Lewis X or . Furthermore, a1,3/4-FucT will transfer a (sia1yl)Lewis A, respectively[174,17’, fucose residue which is substituted on C-6 by a very sterically demanding structure. Notably, a synthetic blood group antigen can be attached, and the resulting “oligosaccharide” can be transferred to an acceptor from its GDP derivative[’86].This approach has been employed to alter the antigenic properties of cell-surface glycoproteins. The Lewis A al,4-FucT has been used to transfer unnatural fucose derivativesfrom their GDP esters. 3-Deoxyfucose and L-arabinose are transferred to LacNAcpO(CH2)8C02CH3 at a rate of 2.3% and 5.9%, respectively, relative to ~ - f u c o s e [ ’ ~ ~ ] . Moreover, al,3-FucTs have been extensively employed as the final step in an enzymatic cascade for the synthesis of complex oligosaccharides [lS71, glycopeptides and glycoproteins [1811 in which the sLeX structure is formed. N-Acetylglucosaminyltransrase (GlcNAcT).In viuo, the N-acetylglucosaminyl transferases control the branching pattern of N-linked glycans[188, . Each of these enzymes transfers a p-GlcNAc residue from UDP-GlcNAc to a high mannose-based acceptor. The GlcNAc transferases I-VI, which catalyze the additon of the GlcNAc residues to

Table 11.3-4. CDP-Fuc

Products of fucosyltransferase reactions

+ a1,2 or a1,3/4

FucT

Fucal,2GalpOR;R = CH2CH3,(CH2)6NH2

Scale”

C C Fucal,3(NeuAca2,3Gal~l,4)GlcNAc~O(CH2)~C02Me C,D C Fuca1,3(Galp1,4)-5-thio-Glc Fucal,4(Gal~l,3)GlcNAc C Fuca1,3(NeuAca2,3Gal~l,4)Glucal D Fucal,4(NeuAca2,3Ga~~l,3)GlcNAc~l,6Gal~O(CH~)~CO~Me D Fucal,4(NeuAca2,3Gal~l,3)GlcNAc~l,3Gal~O(CH2)~CO~Me D Fucal,4(Gal~l,3)GlcNAc~O(CH~)~CO~Me D 3-deoxy-Fucal,4(Gal~1,3)GlcNAc~O(CH~)~CO~Me D S-desmethyl-Fucal,4(Gal~1,3)GlcNAc~O(CH~)~CO~Me D Fucal,2Gal~l,4GlcNAc~OR; R = H,(CH2)6NH2

a A, > 1 g; B, 0.1-1 g; C, 10-100 mg; D,< 10 mg

Ref.

[184] [184] 1175,1831 [183] [183] [163] [lSS] [185] [185] [184] [lSS]

11.3 Hydrolysis and Formation ofClycosidic Bonds

I

625

Table 11.3-5.

Products of ClcNAc-transferase reactions. ~

~~

UDP-ClcNAc (or analogs) + ClcNAcTase ~

~~

Scale”

Ref.

C D

[59, 1921 [190]

~

UDP-GIcNAc + GlcNAcT I

G~cNAc~1,2Manal,3(Manal,G)Man~O(CH~)~CO~Me 3-deoxy, 4-deoxy, or G-deoxy-GlcNAc~l,2Manal,3(Manal,G)Man~O (CH2)8C02Me

UDP-CICNAC+ CICNACTII

GlcNAc~l,2Manal,G(GlcNAc~l,2Manal,3)Man~O(CH,~)~CO~MeD

[192]

UDP-CICNAC+ CICNACT

G~cNAc~l,G(Gal~l,3)GlcNAc

D

[193]

C,D

[194]

UDP-CIC+ CICT

GlcPOR; R = C H ~ C H J(CH&NH2 , a A, > 1 g;

B,0.1-1 g: C, 10-100 mg; D,< 10 mg

the core Asn-linked pentasaccharide of glycoproteins (Fig. 11.3-16), have been identified and characterized[Iss, lS9l. GlcNAc transferases have been utilized for the synthesis of natural and nonnatural oligosaccharides (Table 11.3-5).In addition to transferring GlcNAc, GlcNAcT I from human milk catalyzes the transfer of 3-, 4-, or 6-deoxy-GlcNAc from its 19‘1. respective UDP derivative to Man al,3(Manal,6)Man~O(CH2)sC02CH3[”o~ The 4- and 6-deoxy-GlcNAcanalogs can also be transferred by GlcNAcT 11, although UDP-3-deoxy-GlcNAcis not a substrate for this The core 2 GlcNAcT can employ UDP-trifluoro-GlcNAcas a s~bstrate1”~I. GlcNAcT has also been used to attach the terminal GlcNAc of GlcNAcpl,4GlcNAca dolichyl pyrophosphate, a substrate of oligosaccharyltransferase[’gll.Mannosyltran~ruse(ManT). Various mannosyltransferases have been shown to transfer mannose and 4-deoxymannose from their respective GDP adducts to acceptors [1971. al,2-ManT transfers mannose to various derivatized a-mannosides and a-mannosyl peptides to produce the Manal,2Man structural This method has also been extended to whole cells as a source of al,2-ManTL”’]. Mannosyltransferases from pig liver accept GlcNAcpl,4GlcNAc phytanyl pyrophosphate, an analog of the natural substrate in which the phytanyl moiety replaces dolichol [200]. Overexpression of P1,CManT has also been instrumental in the synthesis of an N-glycan core structure L2011 as well as p1,CManT is especially valuable synthetically, as pthe bacterial 0 mannosyl glycosides are exceedingly difficult to form chemically. Sucrose synthetase. The fructose derivatives 1-azido-1-deoxy-,1-fluoro-1-deoxy-, 6-deoxy-,6-fluoro-6-deoxy-,and 4-fluoro-4-deoxy-fructosehave been used as glycosyl acceptors in the sucrose synthetase-catalyzed synthesis of sucrose analogs [*031. 6-Deoxy- and 6-fluoro-6-deoxy-fructose were generated in situ from the corresponding glucose derivativesunder catalysis by glucose i s o m e r a ~ e [ ~Sucrose ~ ~ I . synthetase has also been extensively employed in the synthesis of nucleotide sugars l2O4l.

626

I

7 1 Hydrolysis and Formation ofC-0 Bonds

HO

H O W OOH * OHo H OH

NHAc

f E j : pl,4-GalT E2: pyruvate kinase E3: UDP-Glc pyrophosphotylase E4: UDP-Glc 4-epimerase E5: pyrophosphotylase

E6:phosphoglucomutase

Figure 11.3-17.

11.3.2.3 In Situ Cofactor Regeneration

Though analytical and small-scale synthesis using glycosyltransferases is extremely powerful, the high cost of sugar nucleotides and the product inhibition caused by the released NMP or NDP present major obstacles to large-scale synthesis. A simple solution to both of these problems is to regenerate the sugar nucleotide in situ from the released NDP[2051. The first example of this strategy was the pl,4-GalT-catalyzed synthesis of LacNAc[2011(Fig. 11.3-17). Only a catalyhc amount of UDP-Gal is initially used to glycosylate GlcNAc. However, UDP-Gal is regenerated from the product UDP and galactose using an enzyme-catalyzed reaction sequence which requires stoichiometric amounts of a phosphorylating agent. Several oligosaccharides have been prepared using routes based on this concept[150]. Another regeneration system for UDP-Gal, which is based on the use of galactose-1-phosphate A third, which employs sucrose uridyltransferase, has also been synthetase for recycling of UDP-Glc/UDP-Gal from sucrose and UMP has recently been A very recent approach to recycling systems employs coupling metabolically engineered bacterial cells for large scale sugar nucleotide production, to date including UDP-Gal (Fig. 11.3-18)L2O9I and CMP-NeuAcI'441. In situ cofactor regeneration offers several advantages. First, a catalytic amount of NDP and a stoichiometric amount of monosaccharide are used as starting materials rather than a stoichiometric quantity of sugar nucleotide, thus tremendously reducing costs. Second, product inhibition by the released NDP is minimized

I

7 7.3 Hydrolysis and Formation ofClycosidic Bonds 627

UTP-

UDP C- UMP

Globotriooe

&

-0 HO

OH

OH

Figure 11.3-18.

because of its low concentration in solution. And third, isolation of the product is greatly facilitated. A multi-enzyme regeneration system for CMP-NeuAc is illustrated in Fig. 11.319[135,2081, n'IS system follows the same basic principles as the UDP-Gal recycling system. A CMP-NeuAc synthetase/a2,3-SiaTfusion enzyme with increased stability has also been applied to this The development of these regeneration systems, as well as those for G D P - M ~ ~ I [ ~GDP-FucL'~~], '~], and UDP-G~CUAL~~'] should facilitate the widespread use of glycosyltransferases for oligosaccharide synthesis. Notably, when UDP-GlcUA and UDP-GlcNAc recycling systems are New combined with hyaluronic acid synthase, HA polymers can be

/

CTP

"n

c

NHAc

HO OH

Aco; E, : a2,3-sialyltransferase; E,: nucleoside monophosphate kinase; E,: pyruvate kinase; E,: CMP-NeuAc synthetase; E5: pyrophosphatase Figure 11.3-19.

628

I

1 1 Hydrolysis and Formation of C-0 Bonds

HO AcNH

0

0

It

II

o=s-0-PI I 0-

0-

PAPS

0

O H \

/

o=po

/ Sulfotransferase IV \

Figure 11.3-20.

systems for the recycling of PAPS for the synthesis of complex sulfated carbohydrates have also recently been developed (Fig. 11.3-20)f2l2I. 11.3.2.4 Cloning and Expression o f Clycosyltransferases

While many glycosyltransferases catalyze similar reactions and use the same donor substrate, there appears to be little sequence homology among the different enzymes of this class (i.e GalT vs SiaT, etc.). There is, however, a significant cross species homology for the same glycosyltransferase. For instance, one finds 86% identity when comparing the P1.4-GalT protein sequence from humans to that from rat. The different glycosyltransferases do exhibit some similarity in that their cDNA sequences encode regions consistent with a short N-terminal tail, a hydrophobic transmembrane sequence, a short stem sequence, and a large C-terminal catalytic domain [2131. In addition to the membrane-bound form of the glycosyltransferases, soluble enzymatic forms have also been identified in body fluids such as blood, milk, and colostrum. Indeed, these fluids have been sources for the purification of specific glycosyltransferasesL2lk2l7].A comparison of the cDNA sequences of these soluble enzymes with full-length glycosyltransferasegenes suggests that the stem region has been cleaved to release the large catalytic domain from the membrane. Presumably, this theme of signal sequence cleavage is consistent for all the glycosyltransferases (Fig. 11.3-21)[2191. The amount of a glycosyltransferase that can be isolated from natural sources is often limited by the low concentrations of these enzymes present in most tissues and

7 1.3 Hydrolysis and Formation ofclycosidic Bonds Figure 11.3-21.

body fluids. The purification of glycosyltransferases is further complicated by their relative instability["]. For this reason, a great deal of interest has been directed toward the cloning of glycosyltransferase genes into convenient expression sys11.3-6).The general strategy involved is outlined in Fig. 11.3-22. t e m ~ [2191~ ~(Table , The glycosyltransferase gene must first be identified and isolated from the mRNA pool via the cloning of the cDNA to make a cDNA library. This library is then screened to identify the glycosyltransferase gene of interest among - lo6 different sequences present. Once identified, the gene is sequenced and a more complete cloning strategy is developed in order to incorporate the gene into an expression vector. This laborious path has been successfully employed by several groups, many ofwhom are referenced in Table 11.36 The nuances to the general cloning scheme used by these groups are discussed below. Among the organs that have been used for the isolation of glycosyltransferase mRNA are the liver [220, 2211, placenta [222], mammary gland[223],testis [2241, and Table 11.3-6.

Cloned glycosyltransferaseso f the glycoprotein and glycolipid pathways.

Enzyme

Source

Ref.

UDP-Glucuronosyltransferase

murine liver rat liver yeast rat liver porcine submaxillary gland bovine placenta bovine mammary gland murine mammary gland bovine liver murine F9 cells bovine kidney epithelial cells murine testes human placenta calf thymus murine F9 cells human A431 cells human A431 cells

POI PI1 ~321 ~291 12331

Mannosyltransferase a2,G-Sialyltransferase a2,3-Sialyltransferase ~1,4-Galactosyltransferase

~1,3-Galactosyltransferase

a1,2-Fucosyltransferase al,3/4-Fucosyltransferase

P I ~231

WI PSI

12361

WI ~241 ~381 12251

WI ~ 7 1 12391

I

629

630

I

1 7 Hydrolysis and Formation ofC-0 Bonds

(liver, testis, placenta, mammary gland, thymus, with DNA probes, antigenic response,

cDNA insert

or lectin screenings) A

Extraction of mRNA

-

I

I

/ Synthesis of cDNA

Figure 11.3-22.

Subclone into

/

‘ I

u

expression system L

Restriction map and redesign of expression system

I

1

I

In addition, tissue cultures have been used in place of the organ[22G, 2271. From these sources, cDNA is synthesized, and the double stranded cDNA is ligated in h phage via a convenient linker and packed into bacteriophages. The bacteriophages are then plated onto a lawn of E. coli, and screened for the desired gene or gene product. Identification of the glycosyltransferase gene has most frequently been achieved by the hybridization of the gene to specific radiolabeled DNA probes [220-2231. Screening in this manner obviously requires a previous knowledge of the gene sequence - information that in some cases may be obtained by extrapolation from a partial protein sequence or from the DNA sequence of the glycosyltransferase from a related source. Two other approaches have been used to screen glycosyltransferase cDNA libraries, both requiring successful transcription and translation of the gene product. In the cloning of the aZ,(i-SiaT from rat liver, Weinstein et al. used polyclonal antibodies raised to the purified enzyme to screen The approach used by Larsen et al. alleviated the need for a the X previous knowledge of the sequence[226]. This method made use of the specificity of a lectin that recognizes the surface-expressed glycoconjugate product of al,3-GalT. The transfected cells were then panned in dishes coated with the lectin. The adherent cells were isolated and re-panned for further purification. Each of these techniques makes use of libraries in which there are very few copies of the desired gene. A greater chance of success may be possible if the number of copies of the genes could be amplified. The introduction, in 1985, of an in vitro amplification method based on the polymerase chain reaction (PCR)fulfilled this need[229,2301. Of course PCR (and ECPCR)[2301, like the hybridization screening, requires a specific knowledge of the sequence. Once identified, the genes are sequenced using standard procedures. Recloning of the gene into an expression vector is then used to develop an expression system. This recloning has been performed on only a few of the glycosyltransferases.Toghrol et al. have inserted the mouse liver GlcUAT gene into the yeast vector pEVPll and expressed the enzyme in Saccharomyces cerevisiae[2201.The rat liver GlcUAT, on the

7 7.3 Hydrolysis and Formation of Clycosidic Bonds

other hand, has been expressed in COS cells using the SV40 Expression in COS cells using SV40 was also applied to the cloning of bovine j31,4-GalT[222]. A noteworthy approach toward the expression of glycosyltransferases in E. coli has been developed by Aoki et al. to obtain human j31,4-GalT[231].A unique RsrII restriction site in the P1,4-GalT gene allowed the dissection of the sequence at the location of signal peptidase cleavage. The cohesive terminus was digested with Klenow fragment, and the blunt end ligated to pINIII-ompA2[232] at a Klenow fragment treated EcoRI site. This generated the code for a soluble fusion protein of j31,4-GalT with the ompA signal sequence. Transcription and translation of this sequence in E. coli produced an active enzyme that was released into the periplasmic space. Purification and N-terminal sequencing of the enzyme verified the expression of the soluble form of j31,4-GalT with an additional tripeptide N-terminal tail. The kinetic parameters of this enzyme appear to be identical to the isolated native enzyme. To date, over a hundred glycosyltransferaseshave been clonedLZ1l.Expression and production in quantities sufficient for enzymatic synthesis is, however, another matter. Only a handful of glycosyltransferases are currently commercially available. Given the advantages of enzymatic synthesis of oligosaccharides over traditional schemes, research into the overexpression of glycosyltransferases will undoubtedly continue to be developed. 11.3.3

Non-Leloir Clycosyltransferases: Transfer o f Clycosyl donors from Clycosyl Phosphates and Glycosides

Oligosaccharides can also be prepared using non-Leloir glycosyltransferases. Phosphorolysis is reversibly catalyzed by glucan phosphorylases for the synthesis of polysaccharides. For example, and t r e h a l ~ s e [ ~have ~ ~ ”been ] synthesized by the corresponding p h o s p h ~ r y l a s e [ ~Sucrose ~ ~ ” ] . phosphorylase has also been used in the recycling of UDP-Ga1[241].Other enzymes of this class are involved in the synthesis of dextrans and levans [2421. Modified polysaccharides may provide materials with more desirable physical and biological properties than their natural counterparts. Approaches to controlling glycopolymer characteristics have included the control of genes encoding the enzymes responsible for their production, regulation of the activity of these enzymes, or the influence of their in vitro synthesis [2431. Potato phosphorylase has been used in vitro to prepare maltose oligomers,[”] as well as a family of linear, star, and comb-shaped polymers [2441. This enzyme will synthesize polysaccharides in the presence of primers A coupled potato phophorylase/sucrose phosphorylase system, where glucose1-phosphate is generated in situ from sucrose and inorganic phosphate, has been employed for polysaccharide synthesis [loo].The inorganic phosphate liberated by potato phosphorylase is used by sucrose phosphorylase to drive the formation of polymer, thereby increasing the yield. Regulation of the molecular weight of the polysaccharide product can be controlled by the concentration of the primer.

632

I

4

7 1 Hydrolysis and Formation of C-0 Bonds

HO HO

Sucrose phosphorylase

+ HOOP032HO OH

OH

Potato phosphorylase Primer --Pnrner

HoYxon

OH

n

OH

Sucrose, P,, sucrose

ln>61 Figure 11.323.

Unnatural primers bearing functional groups can also be used to prepare tailormade polysaccharides for further manipulation, e. g. attachment to protein or other compounds (Fig. 11.3-23). Cyclodextrin al,4-glucosyltransferase (CD a1,4-GlcT, E. C. 2.4.1.19) from Bacillus macerans catalyzes the cyclization of oligomaltose to form a-, 0- and &cyclodextrin, and the transfer of sugars from cyclodextrin to an acceptor to form oligosaccharides[245. 2461. is enzyme can transform a-glucosylfluoride into a mixture of a- and

n'

P-cyclodextrinsand malto-oligomers[2471. When immobilized on a silica gel support, CD a1,CGlcT was very stable, with no loss of activity observed after 4 weeks when stored at 4 "C. This type of enzymatic catalysis may provide a new route to unnatural cyclodextrin analogs and novel oligosaccharides, as glucose analogs are also substrates. For example, oligoglucosyl deoxynojirimycin and N-substituted derivatives were produced under CD al,4-GlcTcatalysis. Subsequent hydrolysis by glucoamylase gave glycosylazasugars like 4-O-a-~-glucopyranosyl deoxynojirimycin in - GO % yield (Fig. 11.3-24)[2481 The N-methyl derivative was reported to be a potent inhibitor of glucosidase. In spite of the progress that has been made, several difficulties limit the use of cellfree enzymes for the synthesis of polysaccharides. The major problem is the complexity of many polysaccharide-synthesizing systems. Isolation, purification, and stabilization of the required enzymes is often difficult, as many enzymes lose activity when they are no longer membrane-associated. Enzyme isolation from eukaryotic sources is tedious, because of low cellular enzyme concentration. It is unlikely that cell-free enzymatic synthesis will provide better routes to most natural polysaccharides than do fermentation and isolation. The use of genetic engineering, OH

+ HHO o

e

(glucose),

1. Cyclodextrin glucosyltransferase

*

Ho HO %&R

R OH

2. Glucoamylase

HO

OH

Figure 11.3-24.

7 7.3 Hydrolysis and Formation ofClycosidic Bonds

&

HO HO

ROH, glycosidase

X

OH

*& HO

HO

OH

OR

Equilibrium conditions: X = OH Kinetic conditions: X = F, o or p N 0 2 , OR’

Figure 11.3-25.

both using classical genetics and recombinant DNA technology, is an approach being used to prepare modified carbohydrate polymers [2501. 11.3.4 Clycosidases

Glycosidases catalyze the hydrolysis of glycosidic linkages, typically with retention of configuration at the anomeric carbon (P-galactosidaseand lysozyme),but sometimes with inversion (trehalase and (3-amylase)[2511. Enzymatic hydrolysis is thought to be mechanistically similar to acid-catalyzedhydrolysis of glycosides. Both proceed via A an oxonium ion intermediate or a transition state having oxonium ~haracter[~”]. proximal carboxylate in the enzyme active site appears as a common structural motif among glycosidases, and presumably acts to stabilize this intermediate or transition state. Whether the oxocarbenium ion exists as a stabilized ion pair or is trapped by the carboxylate to form a glycosyl ester has been the subject of debate. However, an aglycosyl enzyme intermediate has been observed by ”F NMR in the P-glucosidasecatalyzed hydrolysis of 2-deoxy-2-fluoro-~-glucosyl fluoride, and was shown to be a catalytically competent species[2521. Glycosidase-catalyzedglycoside synthesis is quite analogous to protease-catalyzed peptide synthesis. As with proteases, glycosidases may be used under either equilibrium or kinetically controlled conditions for synthetic purposes (Fig. 11.325) [84.

633

I

2531

11.3.4.1 Equilibrium-controlledSynthesis

The obvious approach to glycosidase catalyzed synthesis of glycosidic linkages involves reversing the catabolic role of the enzyme. Indeed, examples of equilibriumcontrolled synthesis were reported by Bourquelot at the early part of this century[253]. Synthesis by this approach involves an endergonic process with the free energy change under ambient aqueous conditions favoring bond cleavage by approximately 4 kcal/mol. Reaction conditions must therefore be manipulated in order to drive the reaction to produce glycoside. In efforts to shift the equilibrium toward product, the addition of water-miscible organic cosolvents was investigated, but this usually results in enzyme inactivation and a decrease in K, for the glycoside acceptor[254].The use of high substrate concentrations and elevated reaction temperatures have also been explored. Johansson et al. [2551 reported the synthesis of mannose disaccharideswith jack bean amannosidase, while Ajisaka et al. [2561 utilized almond P-glucosidase with glucose

634

I

11 Hydrolysis and Formation of C-0 Bonds

concentrations as high as 90 % w/v to obtain glucose disaccharides. Carbon-celite[2573 and active carbon columns have been developed as molecular traps which selectively absorb the product as the reaction mixture is circulated through the column. Yields, however, are still only about 15%. Though quite simple in theory, the equilibrium approach generally provides poor yields and the formation of side products, which make purification difficult. 11.3.4.2

Kinetically Controlled Synthesis

Kinetically controlled synthesis relies on the trapping of a reactive intermediate generated from an activated glycosyl donor to form a new glycosidic 2541. The trapping agent is generally an exogenous nucleophile. Suitable glycosyl donors for this transglycosylation reaction include di- or oligosaccharides, aryl glycosides, and glycosyl fluorides (Table 11.3-7).The reactive intermediate must be trapped by the glycosyl acceptor more rapidly than by water[258].Under the proper conditions, glycoside formation may be favored kinetically, but hydrolysis is always favored thermodynamically. The reaction must therefore be carefully monitored, and arrested when the glycosyl donor is consumed in order to minimize subsequent glycoside hydrolysis. Recently, mutant glycosidases have been engineered to avoid competing product hydrolysis. Because these enzymes lack a catalytic nucleophile in the active site, they can synthesize but not hydrolyze glycosides [293g*h1. In a comparative study of kinetically vs thermodynamicallycontrolled synthesis of Galpl,GGalNAc, the kinetic approach afforded of 10-fold increase in product yield (20% vs 2 %) [2591. Yields in kinetically controlled synthesis generally range from 20 to 40%. Although addition of organic solvent has not generally been observed to increase product yields, increase of acceptor or donor concentration seems to be quite effective. As an exception, though, polyethylene glycol-modified P-galactosidase is soluble in organic solvents and seems to be suitable for transglycosylation [2601. The kinetically controlled approach has primarily been applied to the retaining glycosidases. However, using glycosyl fluorides as glycosyl donors, an inverting glycosidase has been used to afford products having the configuration at the anomeric position which is opposite to that of the For example, the a,alinkage of a-D-glucopyranosyl-a-D-xylopyranoside has been prepared utilizing 0glucosylfluoride and a-trehalase 11.3.4.3 Selectivity

The primary goal of enzymatic glycoside formation or oligosaccharide synthesis is to achieve selectivity which is difficult to achieve by chemical methods. Glycosidasecatalyzed chemoselective reaction of one hydroxyl group of an unprotected sugar with the glycosyl donor has been observed, although the selectivity is not necessarily absolute or predictable. Kinetically controlled synthesis has been more successful

11.3 Hydrolysis and Formation ofclycosidic Bonds

I

635

Table 11.3-7.

Synthesis o f oligosaccharides and other glycosides using glycosidases.

~

Produd

Scalea Ref.

Raffinose + CHI = CHCHzOH

GalaOCH2CH = CHZ

A

[263]

Gala0-p-PhNO2+ GalaOCHZCH = CHz

Galal,3GalaOCHzCH = CHZ B

[263]

Galal,3/6GalaOMe

B

[263]

Galal,2/3GalaO-p-PhNOz

C

[263]

GlcPOPh + ROH (R = alkyl)

GlcPOR

C

[267]

GalP1,4Glc + GlcNAc-R (R = O H or SEt)

Galpl,3GlcNAcR

B

[265]

GalP1,4Glc + GlcNAc

GalPl,4GlcNAc

A

[269]

+ GlcNAc

Galp1,3-GlcNAc

B

[259,268]

+ GalNAc + ROH (R = allyl, benzyl,

GaQ31,G-GalNAc

B

[259, 2681

GalPOR

A, B

[263]

GalP1,3/6GalPOR

B

(2631

B

[270]

Substrate aCaladosidase [252, 4621

+ Gal(a or p)OMe Gala0-o-PhNO2+ Gal(a or P)O-p-PhNOz

fi-Caladosidase[252, 4631

TM S (CH2 ) 2 ) GalPOPh +

b

R'

1. R', 2. R', 3. R', 4. R',

R4 = H , R2 = OH, R3 = CHJ (Gitoxigenin) RZ, R4 = H, R' = CH3 (Digitoxigenin) RZ= 0, R' = CH', R4 = H (16P, 17P-epoxy-17a-digitoxigenin) R2 = H , R' = CHO, R4 = OH (Strophanthidin)

GalPO-o-PhNOZ + GalaOMe

GalP1,bGalaOMe

C

[263]

Galpl,6/3GalPOMe

C

[263]

GalbOPh + ROH (R = alkyl)

GalPOR

B

[271]

GlcPOPh + BnOH

GlcPOBn

B

[271]

GalP1,4Glc + sucrose

Galbl,6al,2Fru

E

[272]

E

[273]

E

[273]

E

[273]

+ GalPOMe

GalP1,4Glc or GalPODh +

HO

n=lor2

GalPO

(89-90% de)

GalPO

(75 % de)

Ho

R = (CH& or CH=CH

"O

(50% de)

636

I

1 1 Hydrolysis and Formation of C - 0 Bonds

Table 11.3-7.

(cont.).

Product

Scale” Ref.

Galpl,4Glc or GalPOPh + ROH

GalPOR

A, B

[274, 2751

GlcPOPh + ROH

GlcPOR

B

[275, 153, 1541

Galactal + ROH

2-deoxy-GalPOR

E

Galactal + Galactal

2-deoxy-Galpl,3/6Galactal+2-deoxy- C Gal~l,3-2-deoxy-Gal~l,6Galactal

Substrate fi-Calactosidase [252,463]

[159] (1601

GalpO-o-PhNO2 + 2-Ser-OR

GalPO-2-Ser-OR

C

[lSS]

GalPO-o-PhNO2 + Ser

GalpOSer

E

[279]

GalP1,4Glc + 2-Ser-OMe

GalpO-2-Ser-OMe

B

12801

Manal,2/6ManaOR (R = Me or p-PhNO2)

B

[263]

Glca1,lFru

D

12631

C

[154, 2671

a-Mannosidase[252,464] ManaO-p-PhNO2 + ManaOR a-Glucosidase [252,465] Glc + Fru Glcp1,4Glc + HO

x)

HO

GlcaO

fi-Glucosidase[252,4651 Glc

Glcpl,4/6Glc

C

[284]

Glcpl,4Glc

Glc~l,4Glc~l,4Glc

C

(2841

Gal(Glc)NAcpO-p-PhNOz+ Glc (NAc)POMe

Gal(Glc)NAc~l,3/4Glc(NAc)POMe C

[285]

Gal(Glc)NAcpO-p-PhN02+ Glc (NAc)aOMe

Gal(Glc)NAc~l,4/6Glc(NAc)aOMe C

[285]

Fucal,3Gal(a or p)OMe

E

~ 4 1

NeuAca-p-PhN02 + Gal(a or p)OMe NeuAca2,3/6Gal(a or p)OMe

D

[289]

NeuAca-p-PhN02 + Galpl,4GlcNAc NeuAca2,3/6GalPl,4GlcNAc

D

[289]

8-N-Acetylhexosaminidase [252, 46151

a-Fucosidase [252,467] FucaO-p-PhNOz + Gal(a or p)OMe Neuraminidase[252,468]

a A, > 1 g; B, 0.1-1

g; C, 10-100 mg; D, < 10 mg; E, not reported

7 7.3 Hydrolysis and Formation ofClycosidic Bonds

HO

I

637

chitinase polymerization

L Figure 11.3-26.

than thermodynamicallycontrolled synthesis in achieving selectivity. In general, the primary hydroxyl group of the acceptor reacts preferentially over secondary hydroxyl groups, resulting in a 1,G-glycosidic linkage. Some control of selectivity has been demonstrated by the selection of an appropriate donorlacceptor combination. For example, the a-galactosidase-catalyzed reactions of a-Gal-OPh-p-NO2with a-GalOMe and P-Gal-OMeform predominantly a-1,3 and a-l,G linkages, re~pectivelyl~"]. The substituent at the anomeric center of the acceptor controls the position of glycosylation to some extent. aGal-OPh-p-NO2'acting both as donor and acceptor, forms preferentially the a-1,3 linkage, whereas the ortho-nitrophenylglycoside reacts in a similar fashion to form predominantly the a-1,2 linkage [2631. With P-galactosidase, P-1,3-linkeddisaccharides were formed preferentially when benzyl or ally1 pgalactosidewas used as acceptor[84.2631. The use of glycals as acceptors has also been employed as a means of controlling ~electivityI~~~1. One can also use glycosidasesfrom different species to control the regioselectivity. For example, the 0-galactosidase from testes catalyzes the formation of Galpl,3GlcNAc[2651 from lactose and GlcNAc. The minor products produced in this preparation were then hydrolyzed by the E. coli p-galactosidase, which preferentially hydrolyzes P-1,G-linked galactosyl residues. The overall yield of the P-1,3-linked disaccharides was around 10-20 %. Synthesis of polysaccharides based on kinetically controlled glycosidase reactions have been accomplished, as exemplified by the cellulase-catalyzed reaction of pcellobiosyl fluoride to form cellulose, with degree of polymerization c 22L266]. In another strategy, employing a chitin hydrolysis transition state analog, chitinase catalyzed polymerization was accomplishedwithout competing hydrolysis (Fig. 11.326) [2491. Glycosyl transfer to non-sugar acceptor has also been demonstrated. These reactions are especially interesting with chiral, racemic, or meso alcoholic acceptors, as one might expect some degree of diastereoselectivity due to the asymmetric microenvironment of an enzyme active site. Such selectivity has indeed been observed, with diastereoselectivities ranging from moderate to exceptional, as illustrated in Table 11.3-7. 11.3.5

Synthesis of N-glycosides

Nucleosides and their derivatives are ubiquitous in nature, and are involved in a myriad of biochemical phenomena, most notably the storage and transfer of genetic information. Interest in this class of compounds has been stimulated by the efficacy

NHAc

1"

638

I

7 7 Hydrolysis and Formation of C - 0 Bonds

of certain nucleosides as antiparasitic [2971 and antiviral agents [298, 2991. Nucleosides have traditionally been prepared by chemical methods (3001 requiring multiple protecting group manipulations and glycosyl activation procedures. Problems encountered include control of anomeric configuration and regiospecific C-N glycoside formation when there are several possible nucleophilic groups in the purine or pyrimidine base. 11.3.5.1

Nucleoside Phosphotylase

Enzymatic preparations of both natural and unnatural nucleosides have been reported using nucleoside phosphorylases as catalysts[3011. These enzymes catalyze the reversible (but highly favorable) formation of a purine or pyrimidine nucleoside and inorganic phosphate from ribose-1-phosphate (R-1-P) and a purine or pyrimidine base. Nucleoside synthesis has relied on the transfer of the ribose moiety of a readily available nucleoside to a different purine or pyrimidine base or analogs through the intermediacy of R-1-P. This work has been done primarily with isolated but whole cells have also been employed in a few cases[3o3].The deleterious hydrolases present in whole cells could be largely neutralized by conducting the reactions at 60 "C, a temperature at which the nudeoside phosphorylases maintain < 70 % of their activity for 3-5 days [3031. The first synthetic strategy toward nucleosides employed involves isolation of R1-P,which can be prepared in good yield from a nucleoside in the presence of a high concentration of phosphate[3o4]. The isolated R-1-P is then used as the glycosyl donor in an enzymatic coupling reaction with added purine or pyrimidine bases or analogs. By this method, generally any heterocycle which is a substrate for a nucleoside phosphorylase can be glycosylated. The second strategy involves a one-pot exchange of one base for another in the presence of a catalytic amount of inorganic phosphate without isolation of R-1-P. At best, this procedure results in an equilibrium mixture of substrate and product nucleosides, from which the product must be isolated. In less favorable cases, the natural purine or pyrimidine base released from the glycosyl donor may be a potent competitive inhibitor versus the purine or pyrimidine analog. For example, competitive inhibition by hypoxanthine ( K , = 5.6 mM) was the cause (TCA, the aglycon for the lack of glycosylation of 1,2,4-triazole-3-carboxamide component of virazole, K, = 167 mM) when inosine was used as the ribosyl donor and purine nucleoside phosphorylase (PNPase) as the catalyst [25G1. It was, however, possible to synthesize virazole by isolating R-1-P and subsequently using it as the ribosyl donor l3O5]. An alternative way to circumvent the inhibition problem is to employ a pyrimidine nucleoside as the glycosyl donor and a purine (or purine analog) as the acceptor, since the released pyrimidine base does not inhibit the purine nucleoside phosphorylase[306]. By this method, both pyrimidine nucleoside phosphorylase and purine nucleoside phosphorylase are required. Direct purine-topurine exchange reactions have been conducted without isolation of R-I-P using activated purine derivatives as the ribosyl donors r3071. The nucleoside phosphorylases accept a wide range of nucleoside analogs as

7 7.3 Hydrolysis and Formation ofClycosidic Bonds

I

639

substrates, with modifications in both the base and glycosyl components. The use of unnatural bases has met with success using both natural and unnatural glycosyl donors. However, a few limitations have been observed, such as loss of appropriate regio-specificitywith unnatural bases I3OG]. The synthesis of sugar-modified nucleosides has made use of glycosyl donors which are prepared by chemical modification of readily available nucleosides, such as uridine and cytidine. Good yields of L3091 have also been obtained enzyarabino r3O3] and 2'-amino-2'-deoxyribonucleosides matically, although the enzymatic synthesis of 3'-amino-2',3'-dideoxyribonucleosides has given only low yields L3lo, 3111. The synthesis of ribosides of unnatural purine and pyrimidine bases and the synthesis of nucleosides containing modified glycosyl moieties are summarized in Table 11.3-8.Most of these reactions have been carried out in one step without isolation of the intermediate sugar phosphate, although involvement of the sugar phosphate intermediate has been demonstrated. In summary, the nucleoside phosphorylases provide a regio- and stereo-specific route for nucleoside synthesis which is applicable to nucleoside analogs which are modified in either the base or the sugar moiety. These processes provide good yields of products in most cases without the extensive protection and deprotection steps involved in traditional chemical synthesis of nucleosides. Application of this strategy to the synthesis of 2'-deoxy-and 2',3'-dideoxynucleosides was reported with the use of N-deoxyribosyltransferase from Lactobacillus species1301, 3121. 11.3.5.2

NAD Hydrolase

The enzyme NAD glycohydrolase has been used in exchange reactions for the preparation of NAD analogs (3181. The enzyme accepts nicotinamide analogs with modification at the amide functionality as substrates. Depending on the structure of the nicotinamide analogs used, the reaction may be either reversible or irreversible. NADH and its 6-hydroxyl derivative are not substrates for the enzyme. When 4-amino, 4-methylamino, or 4-dimethylamino nicotinamide or nicotinate was used as substrate, the product NAD analog existed as a 1,4-dihydro-typet a ~ t o m e r [ ~ ~ ' ] . 11.3.6 Biological Applications of Synthetic Clycoconjugates 11.3.6.1 Clycosidase and Clycosyl Transferase Inhibitors

Carbohydrate analogs and derivatives are valuable in studying the biosynthesis and modification of oligosaccharides: deoxynojirimycin, swainsonine, and castanospermine inhibit trimming of the N-linked oligosaccharides of glycoproteins[3201; tunicamycin and streptovirudin inhibit protein glycosylation in the Leloir pathway[32']; acarbose inhibits amylase [3221. These inhibitors provide a way of exploring cellsurface oligosaccharide chemistry, a topic of central interest in differentiation, development, and disease. Most are relatively easily understood as transition state

640

I

11 Hydrolysis and Formation ofC-0 Bonds Table 11.3-8.

Nucleoside phosphorylase-catalyzed synthesis with various heterocycles as acceptors or sugar-modified nucleosides as donors. Donor

Acceptor

Method" Yield

Ref.

(%)b ~~

Uridine

X = MeS, Y = H X = NHz, Y = C1 X = Me2N, Y = H

Thymidine 7-N-MethylGuanosine Inosine Uridine Thymidine 7-N-Methyl Guanosine

X = C ~ H I I SY, = H X

X = NH2, BnNH

Uridine

Inosine 7-N-Methyl Guanosine

k

X = OH, PhCONH

.GN N."

N

~~

B

59-76

[313, 3141

B B A

81 100 59

[313, 314) [307] 13131

B B B

18-79 [313, 3061 18-71 [313,306] 53 [307]

B

23-63

[313, 3151

A

47 44

[305] [307]

B

X = NHz, Y = H, NH2, C H j X = OH, Y = C1, H , NH2, CH, B

3 6 9 2 [316]

HO

,!+X = S H , Y = N H z

1-(P-o-arabinosyl)uracil

HO

X = NH2, Y = H X=OH,Y=H,NHz,CI

NH2

B

20-50

[308, 309, 3171

B

7-29

[310]

B

12-17

[306]

2'-amino-2'-deoxyuridine 0

deoxythymidine H

G HO

e

" R

R = H (5'-deoxyuridine)or R = OH (5'-deoxythymidine)

?.

a Method A: a-Glycosyl-1-phosphate generated and isolated prior to addition of acceptor heterocycle Method B: In situ generation of a-glycosyl-1-phosphate. b Yields are based on the initial amount of heterocycle acceptor.

analogs, and the design of new sugar analogs to inhibit other glycosidases and glycosyltransferases [Is2. 3231 can be accomplished. The syntheses of these types of structures are not straightforward using classical synthetic methods. Enzymatic methods have already been proven to be very useful in

7 1.3 Hydrolysis and Formation ofClycosidic Bonds

syntheses of deoxynojirimycinand related materials [3241, and are widely applicable to other similar structures. 11.3.6.2

Clycoprotein Remodeling

A number of the proteins employed as human pharmaceuticals (tissue plasminogen activator, juvenile human growth hormone, CD4, EPO) are glycoproteins. There is substantial interest in developing methods that will permit modification of oligosaccharide structures on these glycoproteins by removing and adding sugar units (“remodeling”) and in making new types of protein-oligosaccharide conjugates [325. 32G1. Modification of the sugar components of naturally occurring or unnatural glycoproteins might increase serum lifetime, increase solubility,decrease antigenicity, and promote uptake by target cells and tissues. Enzymes are plausible catalysts for manipulating the oligosaccharide content and

Fmoc-AA-0

11

0

I!

Fmoc-SPPS

t

Frnoc

--f+

Sugar

H TFA cleavage

Frnoc+T+o

PG removal

TFA,

PEPTIDE

1

0

Pd(O), nucieophile

Sugar

Engineered Subtilisin. Fmoc removal (morpholineIDMF)

PS = polystyrene PG = acid-sensitive protectin

Figure 11.3-27.

I

641

642

I

I 1 Hydrolysis and Formation of C - 0 Bonds

"."-:'

COOH

Expression as a C-terminal inteinfusioytein

HS H.N-

0

L

N

Protein

Protein of interest

k

y

k corn

Intein-mediated thioester formation

t

lntein

H.N-E;;~COOH

structure of glycoproteins. The delicacy and polyfunctional character of proteins and the requirement for high selectivity in their modification indicate that classical synthetic methods will be of limited use. Major problems in enzymatic glycoprotein remodeling and generation are the unavailability of many of the glycosyltransferases and the uncertainty in glycosyltransferase specificity on the surface of novel proteins. Recent advances in this area have provided several new methods for the synthesis of homogeneous glycoproteins. Proteases have been utilized for glycopeptide bond ligation (Fig. 11.3-27)[3271, specifically in the generation of a homogeneous RNase glycoform. Endo-glycosidases are capable of transforming heterogeneous glycans to homogeneous species in a single trans-glycosylation reaction 13281. Furthermore, intein-mediated splicing reactions allow modification of a protein Cterminus with carbohydrates or other molecular probes (Fig. 11.3-28) [3291. 11.3.7

Future Opportunities

In general, the development of carbohydrate-derived pharmaceutical agents has occurred at a slower pace than that of other biomolecules, undoubtedly because of difficulties in their synthesis and analysis. However, distinct areas of biology and medicinal chemistry have directed attention at carbohydrates. Interfering with the assembly of bacterial cell walls L7, 3301 remains one of the most successful strategies for the development of antimicrobials. As bacterial resistance to antibiotics of last resort (i. e. vancomycin) becomes more widespread, interest in developing new antipathogenic agents is increasing. Those based on carbohydrate components of the cell wall, such as KDO, heptulose, and Lipid A, represent novel targets. Cell-wall constituents are also relevant to vaccines and as leads toward non-protein immunomodulating compounds. Furthermore, cell-surface carbohydrates are central to differentiation and development, and may be relevant to abnormal states such as those characterizing some malignancies L71. The broad interest in diagnostics has begun to generate interest in carbohydrates as markers of human health. In addition, there are a number of other possible applications of carbohydrates, for example as dietary constituents, in antivirals, or as components of liposomes for

drug delivery. Enzymatic methods of synthesis, by rendering carbohydrates more accessible, will contribute to further research in all of these areas.

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Parks, Jr. Proc. Fed. Am. Soc. Exp. Bid. 1986, 45, 2773. 312 (a) J. Holguin, R. Cardinaud, Eur. J. Biochem. 1975,54, 505. (b) J. Holguin, R. Cardinaud, Eur. J. Biochem. 1975,54, 575. (c) D. A. Carson, D. B. Wasson, E. Beutler, Proc. Natl. Acad. Sci. U S A1984, 81, 2232. (d) D. A. Carson, D. B. Wasson, Biochem. Biophys. Res. Commun. 1988, 155, 829. (e) D. Betbeder, D. W. Hutchinson, A. 0. L. Richards, Nucleic Acids Res. 1989, 17, 4217. 313 T. A. Krenitsky, J. L. Rideout, G. W. Koszalka, R. B. Inmon, E. Y. Chao, G. B. Elion, J. Med. Chem. 1982,25, 32. 314 T. A. Krenitsky, G. W. Koszalka, J. B. Tuttle, Biochemistry 1981, 20, 3615. 315 J. L. Rideout, T. A. Krenitsky, G. W. Koszalka, N. K. Cohn, E. Y. Chao, G. B. Elion, V. S. Latter, R. B. Williams,/. Med. Chem. 1982,25,1040. 316 (a) T. Utagawa, H. Morisawa, F. Yoshinaga, A. Yamazaki, K. Mitsugi, Y. Hirose, Agnc. Bid. Chem. 1985,49, 1053. (b) H. Morisawa, T. Utagawa, T. Miyoshi, F. Yoshinaga, A. Yamazaki, K. Mitsugi, Tetrahedron Lett. 1980, 21, 479. (c) T. Utagawa, H. Morisawa, T. Miyoshi, F. Yoshinaga, A. Yamazaki, K. Mitsugi, FEBS Lett. 1980, 109, 261. 317 T. Utagawa, H. Morisawa, A. Nakamatsu, Agric. Biol. Chem. 1980, 119, 101. 318 (a) F. Schuber, Bioorg. Chem. 1979,8,83. (b) T. Imai,]. Biochem. 1995, 118, 196. 319 F. Tono-oka, Bull. Chem. Soc. j p n . 1982, 55, 1531. 320 B. Winchester, G. W. J. Fleet, Glycobiology 1992, 2, 199. 321 (a) A. D. Elbein, Annu. Rev. Biochem. 1987, 56,497. (b) R. T. Schwarz, R. Datema, Trends Biotechnol. 1984, 932. 322 L. Muller, in: Biotechnology, H.-J. Rehm, G . Reed (eds),VCH Verlagsgesellschaft,Weinheim, Vol. 4, Chapter 18, 1985. 323 (a) Y.-F. Wang, D. P. Dumas, C.-H. Wong, Tetrahedron Lett. 1993, 34,403. (b) P. Sears, C.-H. Wong, Angav. Chem. Int. Ed. Engl. 1999, 38, 2300. (c) G. S. Jacob, CUT. Bid. 1995, 5, 605. 324 (a) T. Ziegler, A. Straub, F. Effenberger, Angew. Chem. Int. Ed. Engl. 1988, 29, 716. (b) R. L. Pederson, M. J. Kim, C.-H. Wong, Tetrahedron Lett. 1988, 29, 4645. (c) C. H. von der Osten, A. J. Sinskey, C. F. Barbas, R. L. Pederson, Y.-F. Wang, C.-H. Wong, /. Am. Chem. Soc. 1989, 1 1 1, 3924. (d) T. Kaji-

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

I 1.4 Natural Polysaccharide-degrading Enzymes

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rnoto, K. K.C. Liu, R. L. Pederson, Z. Zhong, Y. Ichikawa, J. A. Porco, Jr., C.-H. Wong,]. Am. Chem. SOC.1992, 113,6187. (e) G. C. Look, C. H. Fotsch, C. H. Wong, Acc. Chem. Res. 1993,26,182.(f) H. J. M. Gijsen, L. Qiao, W. Fitz, C.-H. Wong, Chem. Rev. 1996,96,443. 325 (a) H. S. Conradt, H. Egge, J. PeterKatalinic, W. Reiser, T. Siklosi, K. Schaper, J. Biol. Chem. 1987, 262, 14600. (b) S. P. Little, N. U. Bang, C. S. Harms, C. A. Marks, L. E. Mattler, Biochemistry 1984,23,6191. 326 B. D. Livingston, E. M. D. Robertis, J. C. Paulson, Glycobiology 1990, 1 , 39. 327 K. Witte, P. Sears, R. Martin, C.-H. Wong, /. Am. Chem. SOC.1997, 119, 2114.

328 (a) L.-X. Wang, M. Tang, T. Suzuki, K. Kita-

jirna, Y. Inoue, S . Inoue, J.-Q. Fan, Y. C. Lee,

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11.4 Natural Polysaccharide-degrading Enzymes

Constanzo Bertoldo and Garabed Antranikian 11.4.1 Introduction

Polymeric substrates such as starch, cellulose, hemicellulose and pectin are abundant in nature and provide a valuable and renewable source of carbon and energy. A diverse range of fungi, yeast, bacteria and archaea are capable of attacking such complex polymeric substrates by producing extracellular enzymes with a wide range of specificity. In this chapter we summarize the current state of knowledge on polysaccharide-degradingenzymes, and attempts are made to show their biotechnological significance. 11.4.2 Starch

Starch is the most economically important reserve polysaccharide in the plant kingdom and is in addition the major source of carbohydrates in human nutrition. In contrast to non-starch reserve polysaccharides, which are outside the cell and the plasmalemma, starch is located in the so-called plastids or in vacuoles within the plant cells"]. In seeds, the highest starch content can be found in the endosperm, whereas its content in the embryo and the pericarp is very low. In general, the starch content of seeds or fruits varies with the degree of maturation[']. Starch occurs in semicrystalline form in granules. The size and the shape of the granules is dependent on the plant species and may reach about 175 mm.

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I I Hydrolysis and Formation of C - 0 Bonds Structure ofthe branching point in arnylopectin. Figure 11.4-1.

linkage

a-1,4linkage Y 0;

... Starch is composed of amylose (15-25%) and amylopectin (75-85%). Amylose is a linear macromolecule consisting of 1,4-linked a-D-glucopyranose residues. The chain length varies from several hundred to GOO0 residuescl].The direction of the chain is characterized by the reducing and the non-reducing end. The reducing end is formed by a free C-1 hydroxyl group. Like amylose, amylopectin is composed of a1,Clinked glucose molecules, but in addition branching points with a-1,G linkages occur. The branching points occur at every 17-26 glucose molecules, so that the content of a-l,Glinkages in amylopectin is about 5%131.With its molecular mass of 10' to lo', amylopectin is one of the largest biological molecules (Fig. 11.4-1). The iodine-binding capacity of starch is dependent on the degree of polymerization (DP). Amylose forms with iodine a helical inclusion complex with an intense blue colour, which possesses an absorption maximum at wavelengths between 620 and 680 nm. Amylopectin has much less iodine-binding capacity because of its branched character, leading to a red-violet colour with absorption maximum of 540 nm i31. 11.4.2.1

Classification of Starch-degrading Enzymes

Starch-degradingenzymes can be divided into two classes according to their reaction mechanism: the glucosidases and the glycosyl-transferases. The first class, the glucosidases, are classified as hydrolases, which catalyze an irreversible hydrolytic cleavage of the glycosidic bond. The group of glucosidases is further subdivided, according their point of attack, into endoacting and exoacting enzymes. Endoacting enzymes hydrolyze linkages in a random manner in the inner part of the starch molecule, releasing linear and branched oligosaccharides with various chain length. a-Amylaseis classified as an endoacting enzyme. In contrast to endoacting enzymes,

11.4 Natural Polysaccharide-degrading Enzymes

the exoacting enzymes hydrolyze linkages from the non-reducing end of the polysaccharide chain. This group includes P-amylase, glucoamylase and a-glucosidase. Isoamylase, pullulanases type I and pullulanase type I1 are classified as debranching enzymes. Glycosyltransferases transfer glycosyl groups from a starch chain to an acceptor. The acceptor may be another starch molecule, phosphoric acid or nucleotides. Most enzymes in this class catalyze reversible reactions; some enzymes are involved in the starch biosynthesis. The only glycosyltransferaseresponsible for starch degradation is the cyclodextrin glycosyltransferase. 11.4.2.2

aAmylase (l14-a-~-CIucan14-CIucanhydro~asel E.C. 3.2.1 .l)

Amylases are widely distributed in plants, mammalian tissues and microorganisms. The endoacting enzymes produce oligosaccharides and glucose as end products by hydrolyzing the a-1,4-glycosidiclinkages in a random manner. The enzyme catalyzes multichain attack as well as multiple attack on the same chaini41. Amylose is hydrolyzed to maltose and glucose. The anomeric carbon in all products formed has ~ a-Amylase is not able to attack a-1,6 linkages in amylopectin the a - configuration. and glycogen. The a-1,4 linkages in the vicinity of branching points are also not attacked by this enzyme[']. In spite of this, the enzyme is capable of bypassing the branching points. Therefore, the action of a-amylase on branched substrates results in the formation of a-limit dextrins. The structure of the a-limit dextrin is dependent on the source of a-amylase. a-Amylases are also described as liquefying and saccharifylng enzymes. The saccharifying a-amylases reduce the viscosity less than liquefymg enzymes and attack the substrate repetitively[']. Most enzymes have an absolute requirement for calcium ions, and the temperature optima as well as the temperature stability of a-amylases are significantly enhanced in the presence of calcum ions and substrate. a-Amylases are widely distributed among microorganisms, including aerobic and anaerobic bacteria and archaea as well as actinomyces, fungi and yeasts. a-Amylases are produced by a variety of Bacillus species, like B. ' ~ ] . Bacillus enzymes are amyloliquefaciens, B. cereus, B. circulans, or B. s ~ b t i l i s [ ~ - The characterized by a wide range of temperature and pH optima [l].The a-amylase from B. acidocaldarius shows optimal activity at pH 3.5 and 75 "C; the enzyme from Bacillus sp NRRL B2881 prefers alkaline conditions (pH optimum at pH 9.2) and 50 "C [11, 12]. Anaerobic microorganisms belonging to the genera Clostridium, Thermoanaerobacter, Themoanaerobiurn and Themobacteroides have also been reported to synthesize extracellular, amylolytic enzymes [131. Also the a-amylases from archaea have been characterized (e.g.: Pyrococcusfiriosus, Themococcusprofindus). Some of these enzymes are optimally active above 100 0C[14,"1.

I

655

656

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I 1 Hydrolysis and Formation of C - 0 Bonds 11.4.2.3

fi-Amylase (1,4-a-~-GlucanMaltohydrolase, E. C. 3.2.1.2)

P-Amylases occur in most higher plants and a number of microorganisms, and are absent in mammalian tissues. The exoacting P-amylaseshydrolyze a-1,4linkages by the stepwise removal of maltosyl residues from the non-reducing end of polysaccharides [16].During the hydrolysis an inversion of the anomeric configuration occurs leading to P-maltose as end product. Unlike a-amylase, P-amylasecannot bypass the a-l,Glinkages in branched substrates and stops two or three glucose units before the branching point. In amylopectin or glycogen, hydrolysis occurs only in the outer chains, and therefore maltose and a-limit dextrins of high molecular weight are the endproducts. p-Amylase is produced by few Bacillus species. The pH optima determined for the B. megateriurn and the B. polymyxa enzymes are in the neutral and slightly alkaline region and the enzymes are unstable above GO "C. The 0amylase from Clostridium themosulfirogenes ATCC 33 743 was characterized as a thermoactive enzyme with a temperature optimum of 75 "C So far, this is the only P-amylaseproduced by an anaerobic microorganism. 11.4.2.4

Glucoarnylases (1,4-a-o-glucan glucohydrolase, E. C. 3.2.1.3)

Glucoamylases, also termed amyloglucosidase or y-amylase, are produced predominantly by fungi, especially by species of Aspergillus, Rhizopus and Endomyces. They are rare in procaryots and absent in plants or in mammalian tissues. The enzyme acts similarly to p-amylase, but attacks a-1,4as well as a-l,Glinkages from the non-reducing end. 0-D-glucose is released as an end product. Glucoamylases are not specific for a-1,4 and a-l,Glinkages; hydrolysis of a-1,3linkages has also been reported["]. The enzyme prefers polysaccharides for rapid hydrolysis and has lower affinity to oligosaccharides or maltose. Because of the lower affinity to a-1,G linkages, the rate of starch hydrolysis decreases subsequentially. In practice, a complete degradation of amylopectin or branched substrates could not be observed["]. Pullulan hydrolysis from the non-reducing end by glucoamylases to glucose was also reported[21,22]. (For a description of pullulan see 1.2.1.5.). The glucoamylases produced by Aspergillus species and yeast are active in the acidic range (pH 4-5 for the fungi and pH 2.5-5.5 for the yeast).The enzymes are unstable above GO "C. The presence of glucoamylase in the thermophilic anaerobic bacterium Clostridium thermosaccharolyticum was reported by Specka et al. (1992) and very recently in the thermoacidophilic archaea Picrophilus oshima, Picrophilus torridus and Themoplasma acidophil~m['~]. These enzymes are optimally active at 90 "C and pH 2.0.

I 7.4 Natural Polysaccharjde-degrading Enzymes

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11.4.2.5

a-Clucosidase (a-Wlucoside Clucohydrolase, E. C. 3.2.1.20)

a-Glucosidase catalyzes the hydrolysis of terminal a-1,4 linkages from the nonreducing end in different substrates. The product released is a-D-glucose. The enzyme prefers short-chain oligosaccharides as substrates and has very low affinity to polysaccharides. In addition, many a-glucosidases show activity towards maltose, acylglucoside, alkylglucoside and isomaltose [241. aGlucosidases are produced by many Bacillus and Aspergillus strains and may be present in several industrial enzyme preparations as side activities. Intracellular a-glucosidases are produced by many microorganisms and are also widely distributed among animals and plants. a-Glucosidases formed by various Bacillus species (B. subtilis, B. amyloliquefaciens, B. cereus), Pseudomonas (P. amyloderamosa, P. Jluorescens W) and lactic acid bacteria ( Lactobacillus acidophilus, Streptococcus pyogenes) are active at slight acidic pH at temperatures up to 75 "C. (for review a-Glucosidases produced from thermophilic Clostridia and archaea are extremely thermostable and thermoactive. The highest activity for clostridial and archaeal enzymes is determined from 65 to 9O"C, and from 105 to 115"C, respectively. a-Glucosidases vary in their substrate specificity. In addition to a1,4-hydrolyzingactivity, some enzymes show low a-1,G-hydrolyzing activity and are capable of hydrolyzing isomaltose. Interestingly, the a-glucosidase from B. t h e m o glucosidasius KP lOOG is unable to hydrolyze maltose but attacks isomaltose with high affinityr2']. The enzymes from B. cereus and from P. amyloderamosa are reported to hydrolyze besides a-1,4and a-l,Glinkages also a-1,2 and a-1,3glucosidic bonds[28]. 11.4.2.6

Isoamylase (Glycogen 6-Clucanohydrolase, E.C. 3.2.1.68)

Isoamylase hydrolyzes with high specificity the a-l,Glinkages in branched substrates such as amylopectin or glycogen. The enzyme cannot catalyze a complete degradation of a- or P-limit dextrins, although the smallest subtrate is 26-a-D-mah0 triosylmaltose r21. Branched substrates are completely debranched, but isoamylase is unable to attack pullulan. Pullulan is an a-D-glucan synthesized by the yeast Aureobasidium pullulans and consists of about 480 maltotriose units linked by a-1,6-D bonds (Fig. 11.4-2). Pullulan is used as a model substrate for starch debranching enzymes, because the a-l,G linkages seem to imitate to a certain degree the a1,&branching points in substrates like amylopectin. Isoamylase has higher affinity to large branched polysaccharides. The enzyme is very rare among microorganisms and has been detected in Pseudomonas amyloderamosa and Cytophaga sp. [29, 301. 11.4.2.7

Pullulanase Type I (a-Dextrin 6-Clucanohydrolase, E.C. 3.2.1.41)

Pullulanase type I hydrolyzes a-l,G linkages in amylopectin, pullulan or limit dextrins with high specificity. Pullulan is completely degraded in a random fashion

*

658

I

11 Hydrolysis and Formation of C - 0 Bonds

OH

f-

a-1,6 linkage

*

+-.-

Figure 11.4-2.

a-1,6 linkage

Structure o f pullulan.

to maltotriose, whereas native glycogen is not attacked by the enzyme. Substrates with short branches such as P-limit dextrin are hydrolyzed at a higher rate than amyl~pectin[~~l. Pullulanase I requires at least two a-1,4linked glucose units in the The smallest substrate for pullulanse I was reported vicinity of the a-1,6 by Marshall to be 26-a-~-maltosylmaltose[211. Pullulanase I catalyzes a condensation reaction in the presence of maltotriose or maltose at high enzyme concentration; the Pullulanases type I are predocondensation products contain a-1,6 minantly formed by mesophilic microorganisms such as Klebsiella pneumoniae, Bacillus acidopullulyticus, B. cereus var mycoides, B. macerans ' B. polymyxa and Streptomyces mitisi2].Fervidobacterium pennivorans is one of the few anaerobic bacteria which produce heat-stable pullulanase type I [14]. 11.4.2.8

Pullulanase Type II or Amylopullulanase

Unlike pullulanase I, pullulanase I1 hydrolyzes a-1,6linkages in pullulan and in , enzymes with addition is capable to cleave a-1,4linkages in a m y l ~ s e [351.~ ~These dual specificity belong to a new class of pullulanases, termed pullulanase type I1 or amylopullulanase~'3~ 36, 371. Pullulanase I and 11 are absolutely unable to hydrolyze substrates like dextran or isomaltotriose, which contain exclusively a-1,6linkages. They possess an absolute requirement for a-1,4linkages in the vicinity of the a-1,6 linkages [13, 26]. Pullulanase type I1 is widely distributed in anaerobic microorganisms including species of the genera Clostridium, Dictyoglomus, Thermoanaerobacter, Thermoanaerobium, Thermobacteroides and Pyrococcus. These enzymes are extremely

7 1.4 Natural Polysaccharide-degrading Enzymes

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659

thermostable and are optimally active in the temperature range between 75 and 105 0 ~ 1 1 4 .381. 11.4.2.9

Pullulan Hydrolases (Type I, Neopullulanase; Type II, Isopullulanase, E.C. 3.2.1.57, Pullulan Hydrolase Type 111)

Pullulan hydrolase type I (neopullulanase) and pullulan hydrolase type I1 (isopullulanase) hydrolyze the a-l,4 linkages in pullulan, liberating panose and isopanose, respectively. Both enzymes are unable to hydrolyze a-1,6 glycosidic bonds in branched substrates or pullulan. Due to this fact, the classification of pullulan hydrolases into the group of debranching enzymes is misleading. Pullulan hydrolases type I have been described from B. stearothemophilus and B. stearothemophilus KP 1064 and pullulan hydrolase type I1 from Arthrobacter globij-omis T6[3g,581. Recently, pullulan-hydrolasetype I11 from T.aggregans has been detected cloned and expressed in mesophilic hosts. This enzyme attacks a-1,4 as well as a-1,6 glycosidic linkages in pullulan, producing maltotriose, maltose, panose and glucose f5'1. 11.4.2.1 0 Cyclodextrin Clycokyltransferase (1,4-a-~-Clucan4-a-~-(1,4-a-~-C~ucano)-Transferase, E.C. 2.4.1.19)

Cyclodextrin glycosyltransferases are produced predominantly by Bacillus species (B.circulans, B. stearothemophilus, B. macerans, B. megaterium), Klebsiella pneumoniae and Micrococcus sp. (for review see[40]).The extracellular enzymes produced by Bacillus macerans and B. megaterium catalyze the transformation of linear chains of starch into cyclic oligosaccharides, the Schardinger cyclodextrins. The glucose residues in cyclodextrins are linked by a-1,4-glycosidicbonds and, because of the ring structure, reducing ends are absent. a-,p- and y-cyclodextrinsconsist of 6,7 and 8 glucose units, respectively. The specificity, the source and the type of the enzyme are responsible for the ratio of different cyclodextrins formed. In principle, all cyclodextrin glycosyltransferases produce a-, p- and y-cyclodextrins simultaneo u ~ l y571.[ ~Thermostable ~~ cyclodextrin glycosyltransferases (CGTases) are produced by Themoanaerobacter species, Thermoanaerobacterium thermosul&rigenes and Anaerobranca gottschalkii. Recently, a CGTase, with optimal temperature at 100 "C, was purified from a newly isolated Archaeon, Thermococcus sp. This is the first report of the presence of a thermostable CGTase in a hyperthermophilic A r ~ h a e o n [ ~ ~ ] . The occurrence of different starch-degradingenzymes in microbes is summarized in Table 11.4-1. 11.4.2.1 1

Biotechnological Applications of Starch-degrading Enzymes

Starch-degrading enzymes are applied in the starch bioprocessing, sugar, alcohol and brewing industries. The commercially most important application of starch-

660

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1 7 Hydrolysis and Formation of C - 0 Bonds Table 11.4-1.

Occurrence of different starch hydrolyzing enzymes in microorganisms.

Enzymes

Substrate (enzyme action)

Products

Organisms

a-Amylase

starch (endoacting a-1,4)

a-limit dextrins branched oiligosaccharides, glucose maltose, linear oligomers

Bacillus amyloliquefaciens B. cereus Tnermotoga maritima Pyrococcusfuriosus

P-Amylase

starch (exoactinga-1,4)

p-maltose limit dextrins

Bacillus megaterium B. polymyqa Clostridium thermosulfurogenes

Glucoamylase polysaccharides [exoactinga-l,4,(a-l,6)]

0-D-glucose

Aspergillus niger, A. oryzae Rhizopus nodosus Clostridium acetobutylicum Picrophilus torridus

a-Glucosidase oligosaccharides [exoactinga-l,4,(a-l,6)]

a-D-glucose

Bacillus subtilis B. cereus Streptococcus pyogenes Thermococcus strain AN1 'I: hydrothermalis

Isoamylase

branched polysaccharides linear polysaccharides Pseudomonas amyloderamosa Flavobacterium odorratum [endoacting a-1,6]

Pullulanase Type I

pullulan, branched polysaccharides [endoacting a-1,6]

maltotriose, linear oligosaccharides

Klebsiella pneumoniae Bacillus acydopullulyticus Fervidobacterium pennivorans Thermotoga maritima

Pullulanase Type I1

pullulan, branched polysaccharides [endoacting; a-1,6 in pullulan; a-1,6 + a-1,4 in branched poly- and oligosaccharide]

maltotriose, linear oligosaccharides

B. subtilis C. thermohydrosulfuricum Pyrococcus woesei Desulfurococcus mucosus

Pullulanhydrolase Type I

pullulan [a-1,4]

panose

B. stearothermophilus

Pullulanhydrolase Type I1

pullulan [a-l,4]

isopanose

Atthrobacter globi&ormis

Pullulanhydrolase Type I11

pullulan [a-1,4]

glucose, maltose maltotriose

Thermococcus aggregans

a-P-y-cyclodextrin

B. circulans B. macerans Anaerobranca gottschalkii Thermococcus sp.

Cyclodextrin- branched glycosylpolysaccharides transferase [endoacting a-1,4]

*Values in brackets () indicate low enzyme affinity

17.4 Natural Polysaccharide-degrading Enzymes

degrading enzymes is the production of syrups and sweeteners. The conversion of corn starch to fructose begins with a liquefaction step carried out with a-amylase from B. lichenifomis at 105-115 "C and 90-95 "C at pH 6 . Amylose and amylopectin are hydrolyzed to dextrins and some oligosaccharides. The saccharification follows the liquefaction in the presence of glucoamylase from Aspergillus niger and debranching enzymes. The process conditions in the saccharification step have to be changed since the enzymes are optimally active at pH 4-4.5 and 55-65 "C. A dextrose solution of 95% results from this step. The dextrose solution can be crystallized or subsequently further isomerized. The isomerization from glucose to fructose again requires variation of the process conditions (55-GO "C, pH 7-8) r4lS421. The finding of different amylolytic enzymes that are active under the same conditions will certainly improve the starch bioconversion process. Recently, it was found that hyperthermophilic microorganisms are a good source of such enzymes. a-Amylase, pullulanase and a-glucosidasefrom Pyrococcus sp. are optimally active at pH 4-5 and 100-110 "C'2GI. In the baking industry a-amylase from fungi is used in order to release dextrin and fermentable sugars for yeast metabolism. Exhaustive dextrin formation, however, will lead to undesirable properties like loaf stickness and dark color. In the process of fuel alcohol production different grains and tubers serve as raw material in the fermentation process. The liquefaction and saccharification steps are carried out in the presence of a-amylase and glucoamylase, respectively. This saccharified feedstock forms the substrate for the ethanol fermentation with yeastsL4'].Pullulanases are also used in the production of "light beer", which has low carbohydrate content. During the fermentation process the pullulanase is added together with fungal a-amylase or glucoamylase to the The production of branched and more water-soluble cyclodextrins can also be carried out with pullulanase. The pullulanase catalyzes the transfer reaction of malto-oligosaccharidesto cyclodextrins. Branched cyclodextrins are more water-soluble than linear cyclodexCGTases are used for the production of cyclodextrins that can be used as a gelling, thickening or stabilizing agent in jelly desserts, dressing, confectionery, dairy and meat products. Because of the ability of cyclodextrins to form inclusion complexes with a variety of organic molecules, cyclodextrins improve the solubility of hydrophobic compounds in aqueous solution. This is of interest for the pharmaceutical and cosmetic industries. Cyclodextrin production is a multistage process in which starch is first liquefied by a heat-stable amylase, and in the second step a less thermostable CGTase from Bacillus sp. is used. The application of heat-stable CGTase in jet cooking, where temperatures up to 105 "C are achieved, will allow liquefaction and cyclization to take place in one step. 11.4.3

Cellulose

Cellulose is the principal component of plant cell walls, and thus represents the worlds most abundant organic polymer, with an annual production of 4 x 10" tonnes per year. Cellulose is found in nature as an unbranched insoluble polymer

I

662

I

1 7 Hydrolysis and Formation of C-0 Bonds

[Cellulose]

Endoglucanase

n

OH

OH

OH

Cellobiohydrolase

n

n

B-Glucosidase

Figure 11.4-3.

Action of cellulolytic enzymes on cellulose.

(Fig. 11.4-3) containing up to 14000 glucose units linked together by p-1,4-~glycosidic bonds 15’), ‘1. The hydrogen capacity between individual chains in cellulose is quite high, with each residue contributing up to three OH groups. The individual chains of cellulose tend to form microfibrils as a result of inter- and intramolecular hydrogen bonding. The microfibrils associate in a similar way to form fibers [l,“. “1. Cellulose contains both crystalline and amorphous regions. The term “crystalline” refers to those regions in which a high degree of order is found within and between the fibrils. In the amorphous regions, however, a lesser degree of order is predominant [62, 631. Crystalline regions are more resistant to degradation than amorphous regions [64, 651. Thus, the fraction of crystalline regions found in cellulose is an important factor affecting the rate and enzymatic hydrolysis of cellulose 16’, “1. Cellulose is found in nature as a principal structural element in cell walls of higher plants, in association with hemicellulose, lignin and other polysaccharidesiG7]. Cellulose is also found in some seaweeds and can be synthesized by some bacteria[“]. Cellulose occurs in an almost pure form (98%)in cotton fibers, while in flax (SO%), jute (GO-70%), wood (40-SO%), and forages ( 2 4 3 6 % ) a less pure form of cellulose is found[’, 6 2 , 691.

1 1.4 Natural Polysaccharjde-degrading Enzymes

11.4.3.1 Cellulose-degrading Enzyme Systems

Cellulases are a group of enzymes capable of hydrolyzing insoluble cellulose to its monomer glucose L7O]. Because of the crystalline and insoluble nature of cellulose, its degradation is very slow. Cellulose degradation requires a multienzyme complex involving at least three major enzymes, namely 1,4-P-~-glucan glucanohydrolase cellobiohydrolase (exoglucanase, E. C. (endoglucanase, E. C. 3.2.1.4), l,CP-~-glucan 3.2.1.91), and P-D-glucoside glucohydrolase (P-glucosidase, E. C. 3.2.1.21) [711 (Fig. 11.4-3). Mainly two types of enzyme systems have been recognized to be involved in cellulose hydrolysis. The first is the non-aggregating system, in which the three main cellulolyhc enzymes are produced and mainly found to be secreted into the growth medium as separate entities. In this system, endoglucanase, cellobiohydrolase and P-glucosidase act in synergy[129* 721. The second enzyme system is referred to as the aggregating system. This system is found mainly in anaerobic bacteria, where cellulases are secreted as a high molecular weight multienzyme complex. This complex is generally found on the cell surface, where it mediates the attachment between the cells and the substrate[129]. The most studied system is that of Clostridiurn thermocellum [731; the complex is named “cellulosome”. 11.4.3.2 Endoglucanase (1,4-fJ-~-Clucan-Clucanohydrolase, E. C. 3.2.1.4)

Endoglucanase hydrolyzes cellulose randomly, producing oligosaccharides, cellobiose and glucose as end products (Fig. 11.4-3). Endoglucanase attacks mainly the amorphous regions in cellulose and soluble derivatives of cellulose [741. The action of endoglucanase results in a decrease in the chain length of carboxymethylcellulose (CMC),acid-swollen cellulose and soluble barley glucan, producing mainly glucose, cellobiose, cellotriose and other oligosaccharideslG4,75, 7G]. Substrates like p-nitrophenyl-P-D-cellobiosideand methylumbelliferyl-P-o-cellobiosideare hardly attacked by endoglucanase. Low activity is also observed with microcrystalline cellulose[77].In contrast to this, the endoglucanase of Trichoderma viride shows high activity towards crystalline cellulose but only weak activity towards CMC I8O]. 11.4.3.3 Cellobiohydrolase (1 ,Cfi-~-ClucanCellobiohydrolase, E. C. 3.2.1.91)

Cellobiohydrolases are exoglucanases that attack the non-reducing end of the cellulose polymer chain to produce cellobiose (Fig. 11.4-3). Recent reports have indicated that the attack on cellulose is not restricted to the end of the chain. Thus, the cellobiohydrolase I from Trichoderma reesei is capable of degrading the p-glucan from barley in a manner typical of an endoglucanase[81].Cellobiohydrolases comprise the major part of fungal cellulase systems that are capable of degrading crystalline cellulo~e[~~1. Up to 80% of microcrystalline cellulose can be degraded by this enzyme[”]. Earlier studies have indicated the absence of cellobiohydrolases in

I

663

664

I

7 7 Hydrolysis and Formation of C - 0 Bonds

bacterial cellulase systems [751. However, Langsford et al. [821 reported the presence of this enzyme in Cellulomonasjmi. This enzyme has also been found in Ruminococcus albus and R. Jlavefaciens[841. Bacterial cellobiohydrolases are capable of hydrolyzand methylumbelliferyling model substrates such as p-nitrophenyl-P-D-cellobioside P-D-cellobioside. They release cellobiose from microcrystalhe cellulose and show low activity towards CMC [771. 11.4.3.4

fi-Clucosidase (b-d-ClucosideClucohydrolase, E. C. 3.2.1.21)

Cellobiase or P-glucosidase acts mainly on cellobiose and cellodextrins (up to DP of 6 ) to produce P-glucose (Fig. 11.4-3); cellulose and higher cellodextrins are not

hydrolyzed by this enzyme[“]. P-Glucosidase acts also on sophorose and cellobiose to produce monosaccharides. In addition, model substrates such as p-nitrophenyl-PD-glucosides or methyhmbelliferyl-~-D-glucoside are attacked. Because of the action of P-glucosidase,the inhibitory effect of cellobiose on cellobiohydrolaseand endoglucanase can be removed. As shown for the P-glucosidase from Penicillium finiculosum, this enzyme also acts synergistically with endoglucanases and cellobiohydrolases [”I. 11.4.3.5

Fungal and Bacterial Cellulases

Most of the studies on cellulases have been conducted using fungal cellulolytic systems. Relatively few cell-free cellulases have been reported to degrade crystalline cellulose. Such fungal systems contain extracellular endoglucanase and cellobiohydrolase activities that convert crystalline cellulose to cellobiose[861. The conversion of cellobiose to P-glucose is catalyzed by 0-glucosidase,which has been found in the cultures of T’choderma T. reesei[881,7: k~ningii[~’] and Talaromyces emersonnii[’OI (Table 11.4-2).Compared to fungal systems, cell-free supernatants from cultures of cellulolytic bacteria seems to lack activity against crystalline cellulose f8‘I. Several cellulolyhc bacteria have been isolated, but their cellulases have not been fully characterized[”, 921. The system from Cellulornonas sp. is one of the most studied cellulolytic systems in bacteria[93,941. Many species which belong to the genera Bacillus, Pseudomonas, Streptomyces, Thermoactinomyces and Thermomonospora are capable of producing cellulolytic enzymes [911. Several endoglucanases were detected in the culture fluid of many of these microorganisms. However, cellobiohydrolase has not been detectedrS6l.The hydrolysis of cellulose by bacteria involves the action of cellulolytic enzyme complexes consisting of different multicomponents. These complexes are associated with the cell wall of the bacterium and are often tightly bound to the cellulosic substrate I7Ol. They are released into the culture fluid only after extensive hydrolysis of cellulose. The most thoroughly studied cellulolytic enzyme complex, referred to as “cellulosome”, is that of Clostridium thermo~ e l l u m 1951. ~~-

7 7.4 Natural Polysaccharide-degrading Enzymes

I

665

Table 11.4-2.

Microbial cellulolytic enzymes

Organism

Endoglucanase

Fungi Aspergillus niger Humicola insolens Tricoderma koningii 1: reesei 1: viride

Cellobiohydrolase

fi-Clucosidase

-

+ +

+

Bacteria: Cellulornonas frmi Clostridiurn thermocellum C. stercorarium Cytophaga sp. Fibrobacter succinogenes Ruminococcus albus Trtermotoga maritima ‘I?termotoga neapolitana Archaea Pyroccusfuriosus Suljblobus solfataricus

+

-

+ +

11.4.3.6

Structure and Synergistic Effect of Cellulases 11.4.3.6.1

The “Cellulosome” Concept

The cellulolytic enzyme sytem of bacteria forms aggregates, which are associated with the cell wall forming catalytically active “protuberances”. Electron microscopy studies revealed that these “protuberances” are found on the surface of all cellulolytic bacteria studied, whereas they are absent on the surfaces of non-cellulolytic bacteria[73,96, 971. In addition, they are not present during growth in the absence of ~ e l l u l o s e [ The ~ ~ ~ best . characterized aggregation system is the cellulosome of Clostridium thennocellum~”~9 5 , 731. The cellulosome binds to the substrate and is active towards crystalline cellulose. A 200 kDa polypeptide seems to be responsible for the substrate binding. In the early stages of growth the cellulosomes of C. therrraocellum form polycellulosomes, which appears as protuberances on the cell surface. In the late growth phase the cellulosomes are released into the culture Cellulases present in the culture fluid seem to represent only Other cellulolytic bacteria fragments of cellulosomes and polycellulosomes.I‘’[ which express cellulosome-like structures are Ruminococcus albus, R. flavefaciens, Fibrobacter succinogenes [96, 981, Acetivibrio cellulolyticus, Bacteroides cellulosolvens, Clostridium cellobioparum, C. cellulovorans and Cellulomonas sp. [96, 991. Cellulose-degrading enzymes from various thermophilic organisms (Thennotoga maritima, Thennotoga neapolitana, Caldocellum saccharolyticum and Anaerocellum thermophilum) have been cloned, purified, and characterized. Recently, a thermostable archaeal endoglu-

666

I

7 7 Hydrolysis and Formation ofC-0 Bonds

canase which is capable of degrading p-1,4 bonds of P-glucans and cellulose has been characterized from Pyrococcus&riosus [l4I.

11.4.3.6.2

Multiple Forms o f Cellulases

The cellulolytic enzymes from bacteria and fungi (endoglucanase,cellobiohydrolase and 0-glucosidase)exist in multiple forms. Multiple forms of these enzymes seem to arise through post-translationalmodification by physiologically regulated processing activity or through post-secretional modification by proteolytic digestion["]. Diversity of endoglucanases and cellobiohydrolases have been reported by several investigators[G4, 'O0, loll. However, Penicillium notatum and Stereum sanguindentum produce a single cellulase and are still able to degrade cellulose['02.'031. Wilson[104] isolated five different endoglucanases from a protease negative mutant of Thermomonospora fisca; cellobiohydrolase activity, however, was not detected. Similarly, Shoemaker and B r ~ w n [ ' ~identified ~l four endoglucanases from Trichoderma uiride. Further studies with T. viride proved the presence of six endoglucanases (Endo I, 11, 111, IV, V and VI), three cellobiohydrolases (Ex0 I, 11 and 111) and one P-glucosidase L8O1.

11.4.3.6.3

Synergism

It has been recognized that the rate of hydrolysis of crystalline cellulose by the combination of endoglucanase and cellobiohydrolaseis much faster than the sum of the individual actions of the components[70].The rationale for the synergy of cellulase has been postulated as follows. The attack is initiated by a randomly-acting endoglucanase in the amorphous areas of the cellulose creating numerous new nonreducing ends that are attacked by cellobiohydrolase, resulting in the release of cellobiose. The P-glucosidase is needed for the removal of cellobiose, a strong inhibitor of both endoglucanase and cellobiohydrolase[72, loGI. Studies with the cellulases from T. reesei and bacteria showed that cellobiohydrolase 11 was only able to attack one end of the microcrystalline cellulose. However, in the presence of their endoglucanase, several sites of cellobiohydrolase attack at the amorphous region were ob~ervedI'~~1. Another observation showing the synergism between endoglucanase and cellobiohydrolase has been reported with the fungus Neocallimastix jontalis. Heat inactivation of the cellulases of this fungus resulted in loss of its ability to degrade crystalline cellulose. Interestingly, the endoglucanase activity was still measurable. The additon of cellobiohydrolasefrom Trichodema koningii restablished the ability ofthe system to degrade cotton Synergism between P-glucosidase and cellobiohydrolase or endoglucanase has also been observed. P-Glucosidase produced by T. koningii has shown synergism with cellobiohydrolase but not with endoglucanase['08].Synergism between a-glucosidase and cellobiohydrolase can be explained by the ability of P-glucosidaseto hydrolyze cellobiose, a strong inhibitor of cellobiohydrolase[log].

7 7.4 Natural Polysaccharide-degrading Enzymes

11.4.3.6.4

Biotechnological Applications o f Cellulases

Cellulase preparations have found different biotechnological applications in several industrial processes. The most effective commercial cellulase is the one produced by Trichodema species. Other cellulases of commercial interest are obtained from strains of Aspergillus, Penicillium and Basidomycetes. Fungal cellulases have been recommended for use in alcohol production. The alcohol yield from cassava is significantly increased if cellulases from Trichodema sp. are added [‘lo]. Cellulolytic enzymes can also be used to improve juice yields and effective color extractions of juices. Cellulolytic enzymes also improve the silage-making process [1301. The cellulase from Trichodema reesei has been reported to accelerate the rate of ensilage processing when treating grass, lucerne and red clover[’”]. The presence of cellulases in detergents causes colour brightening, softening and improved particulate soil removal^"*]. A novel application of cellulases in textil industry is the use of Denimax (Novo Nordisk) for the “biostoning”of jeans instead of the classical stones in stone-washed jeans [1131. Another application of cellulases includes the pretreatment of cellulosic biomass and forage crops to improve nutritional quality and digestibility, enzymatic saccharification of agricultural and industrial wastes and production of fine chemicals [130]. 11.4.4

Xylan

Hemicelluloses are non-cellulosic low molecular weight polysaccharides that are found together with cellulose in plant In the cell walls of land plants, xylan is the most common hemicellulosic polysaccharide, representing more than 30% of the dry weight[132].Most xylans are heteropolysaccharides which are composed of 1,Clinked 0-D-xylopyranosylresidues [133, 134, 13’1 .Th’is backbone chain is substituted with acetyl, arabinosyl, and glucuronosyl residues Homoxylans, on the other hand, consist of xylosyl residues exclusively and have been isolated from esparto grass tobacco stalks [I3’], and guar seed husk[138]. The xylan of hardwoods (0-acetyl-4-0-methylglucuronoxylan) consists of at least 70 P-xylopyranose residues (average degree of polymerization between 150 and 200) linked by P-1,4-glycosidicbonds (Fig. 11.4-4) Every tenth xylose residue carries a 4-0-methylglucuronic acid attached to the C-2 of xylose [1311. In addition, hardwood xylans are highly acetylated;e. g. birchwood xylan contains more than 1mol of acetic acid per 2 mols of xylose [1401. Acetylation occurs usually at the C-3 rather than the C-2 position of xylose. Acetylation at both positions has also been 14’1 . Th e presence of these acetyl groups is responsible for the partial solubility of xylan in water”33].The alkali extraction of xylan leads to the deacetylation of this substrate [I4’]. Softwood xylans (arabino-4-0-methyl-glucuronoxylans) are composed of shorter chains with a degree of polymarization between 70 and 130 (Fig. 11.4-5). Unlike hardwood xylan, the softwood xylan has a higher content of 4-0-methyl-glucuronic acid. The acetyl groups are replaced by a-L-arabinofuranoseunits, which are linked by a-1,3-glycosidicbonds to the C-3 position of xyl0se1~~~1.

I

667

668

I

I 1 Hydrolysis and Formation of C-0 Bonds

Ac

Ac

4

4

a-4-0-Me-GlcA

Ac

r' 2

1

2 3 3 4-D-Xylp1~4-B-Xylp-1~4-B-Xyl~14B-Xyl~l~4-~-Xyl~l~4-B-Xyl~l~4-R-Xyl~l~4-RXyl~l 2 3 2

t

t

t

Ac

Ac Figure 11.4-4.

Ac

0-Acetyl-4-0-methyl-glucuronoxylan from hardwood.

cli'

0

HOH,C

HOH,C

OH

a-4-0-Me-GlcA

OH

a-4-0-Me-Glc A

1

1

1

a-Araf Figure 11.4-5.

1 a-Araf

Arabino-4-0-methyl-glucuronoxylanfrom softwood.

11.4.4.1

The Xylanolytic Enzyme System

Because of the heterogeneity of xylan, its hydrolysis requires the action of a xylanolytic enzyme system which is composed of P-1,4-endoxylanase (E. C. 3.2.1.8), P-xylosidase (E. C. 3.2.1.37), a-L-arabinofuranosidase (E. C. 3.2.1.55), a-glucuronidase (E.C. 3.2.1.-) and acetylxylan esterase (E. C. 3.1.1.6) activities (Table 11.4-3).The concerted action of these enzymes converts xylan to its constituent sugars (Fig. 11.46). Xylan-degrading enzymes have been reported to be present in marine and

7 1.4 Natural Polysaccharide-degradingEnzymes

I

669

Table 11.4-3.

Microbial xylanolytic enzymes.

Organism

Endoxylanase B-Xylosidase a-L-Arabino furanosidase

Fungi Aspergillus awamori Furiarum oxysporum Tricoderma reesei

+

a-Glucuro- Acetyl xylan nidase esterase

+

i

+

+ +

+

+ +

+ +

-

N.D.

+

N.D.

+

+

+

N.D.

+

+

+

N.D. N.D. N.D.

+

+

N.D. N.D.

+

N.D.

N.D.

N.D.

N.D.

-

+ +

Bacteria:

Bacillus subtilis Streptomyces olivochromogenes Thermoactinomyces vulgaris Thermoanaerobacter saccharolyticum Thermonosporafusca Thermotoga maritima Thermotoga neapolitana Archaea

Thewnococcuszilligii

+

+

N.D. N.D.

+

N.D. N.D.

N. D.: not determined

A a-Araf 1

Ac

1

iQ

3

3

2

ArabinofuranosidaseCZ)i

a-Araf

a-Me-GlcA a-Glucuronidase

i

3

-+4~-Xylp-14-B-Xyip-14-~-xylp-l4B-Xylp-l-.4-B-Xylp-l4B-Xytp-l~B-Xylp-l 2

t

a

Acetyl Xylan Esterase

Ac

0

Endoxylanase

--f

2

t Ac

I'r

Endoxylanase

B BXylosidase

Figure 11.4-6. (A) Action ofxylanolytic enzymes on an hypothetical xylan structure. (B) Action o f 0-xylosidase on xylobiose. Ac, acetyl residue; a-Araf, a-L-arabinofuranose; a-Me-ClcA, 4-O-rnethylo-glucuronic acid; fl-Xylp, 0-D-xylopyranose.

670

I

1 1 Hydrolysis and Formation o f C - 0 Bonds

terrestrial bacteria, rumen and ruminant bacteria, fungi, marine algae, protozoa, snails, crustaceans, insects and seeds of terrestrial plants [la]. Among the different functions of xylanases is the utilization of xylan as a carbon and energy source, degradation of cell wall components and degradation of xylans during germination of 11.4.4.2

Endoxylanase (1,4+~-Xylan Xylanohydrolase, E. C. 3.2.1.8)

P-1,4-Endoxylanase cleaves the internal glycosidic linkages of the heteroxylan backbone, resulting in a decreased DP (degree of polymerization) of the substrate (Fig. 11.4-GA). The attack of the substrate is not random, and the bonds to be hydrolyzed depend on the nature of the substrate, e.g. length, presence of substituents and degree of branching[145].During the early course of hydrolysis of xylan the main products formed are xylooligosaccharides.As hydrolysis proceeds, these oligosaccharides are hydrolyzed to xylotriose, xylobiose and x y l ~ s e [ ~ ~Diffe~~~']. rentiation of endo-acting xylanases has been made according to the end products formed i. e. xylose, xylobiose and xylotriose, and/or arabinose. Thus, xylanases may be classified as non-debranching (arabinose non-liberating) or debranching (arabinose-liberating) enzymes [1452 1461 . M any organisms are able to produce both debranching and non-debranching xylanases, resulting in a maximum efficiency of xylan hydrolysis "'I. The production of multiple forms of xylanases has been reported for many organisms such as Aspergillus niger and Fibrobacter succinogenes[150*l5l]. The endoxylanase I from F. succinogenes possesses debranching activity and liberates arabinose from xylan. This is followed by the action of endoxylanase 11, which converts unbranched xylans to xylooligosaccharides['"I. This may indicate that the removal of the arabinose substituents, which act as a hindrance, is a requirement to permit the access of endoxylanase to the xylan backbone. This also demostrates the synergistic relation between debranching and non-debranching xylanases. Arabinose-cleaving endoxylanases have been purified from Streptornyces roseiscleroticus [1521 and T'chodema k~ningii["~I. 11.4.4.3 fi-Xylosidase (fi-o-Xyloside Xylohydrolase, E. C. 3.2.1.37)

P-D-Xylosidases are exo-glycosidases that hydrolyze short xylooligosaccharidesfrom the non-reducing end forming xylose as end (Fig. 11.4-GB). 0-Xylosidases appear to be mainly cell-associated (found in the cytosol) in bacteria and yeast [1341. However, extracellular P-xylosidases have also been reported [1541561. In the yeast Cryptococcus albidus, xylooligomers (xylobiose and xylotriose)enter the cells through a P-xylosidepermease transport system and are converted by 0-xylosidase to x y l o ~ e [ ' ~P-Xylosidases ~]. are in most cases unable to hydrolyze xylan. However, there are some reports of xylosidases that are capable of attacking xylan and producing xylose Such exo-xylanases would have a limited hydrolysis activity towards heteroxylans, as their action would end at the branch points [I4']. 0-Xylosidase activity

7 1.4 Natural Polysaccharide-degrading Enzymes

I

671

may play a role in relieving the end product inhibition of endoxylanase. This has been reported for the enzyme system of Themtornonospora&sca [ls71. Transferase activity is a typical feature of most P-xylosidases, resulting in products of higher molecular weight than the Transfer reaction may result in the formation of both p-1,3 and p-1,4 bond^['^^-'^^]. 11.4.4.4

a-L-Arabinofuranosidase(E. C. 3.2.1.55)

a-L-Arabinofuranosidasesare active against branched arabinoxylans, arabinans, arabinose-substituted xylooligosaccharides and p-nitrophenyl-a-L-arabinofuranoside. Their action on arabinoxylan results in the release of arabinose residues (Fig. 11.4-6A).The production of a-L-arabinofuranosidasein several actinomycetes seems to be induced among others by xylan, arabinan, and wheat bran['", 161].a - ~ Arabinofuranosidases from A. niger and S. purpurascens are also capable of hydrolyzing both 1,3-and 1,s-a-L-arabinofuranosyl linkages in arabinanr'", 163].TheAspergillus niger enzyme attacks first the a-~-1,3-linkedarabinofuranosyl residues to the extent of 30% and then proceeds with a slow attack of the a-L-l,S-arabinan['"]. Synergism between a-L-arabinofuranosidaseand endoxylanase has been reported. A significant increase in xylose, xylobiose and arabinose production was observed when both enzymes are used simultaneously['"]. 11.4.4.5

a-Glucuronidase (E.C. 3.2.1.136)

a-D-Glucuronidasesare required to hydrolyze the a-1,2linkages between glucuronic acid and xylose residues in glucuronoxylan (Fig. 11.4-GA). Because of the lack of aglucuronidase activity in many fungal hemicellulase preparations [13'1, this enzyme was not described until 1986[165]. Only a few a-glucuronidases have been purified so far; these include the enzymes from Trichoderrna reesei, Tnemoascus aurantiacus and Agaricus bisporus[166].Thus, most of the studies on a-glucuronidases have been performed using partially purified enzymes. These enzymes release 4-0-methylglucuronic acid from 4-0-methyl-glucuronicacid-substituted xylooligomers,but not from the polymer [13'1. Simultaneous hydrolysis of acetyl-4-0-methyl-glucuronoxylan with the endoxylanase from A. oryzae and the acetyl xylan esterase from T. longibrachiaturn resulted in the production of non-substituted xylan fragments as well as substituted xylooligomers. These products were further treated with a Pxylosidase from T. reesei and an a-glucuronidase from A. bisporus. The a-glucuronidase was not active against these oligomers, indicating that the acetyl groups next to the glucuronosyl substituent may hinder the action of the a-glucuronidase [1431.

672

I

11 Hydrolysis and Formation of C - 0 Bonds

11.4.4.6 Acetyl Xylan Esterase (E.C. 3.1.1.6)

Acetyl xylan esterase removes the 0-acetyl substituents at the C-2 and C-3 positions of xylose residues in acetylxylan (Fig. 11.4-GA). The importance of acetyl xylan esterase in the hydrolysis of xylan was demonstrated recently[l6'I. It is mainly due to the fact that most of the xylan preparations used to study xylanolyhc enzymes systems are alkali extracted xylans. Under these conditions mainly deacetylated xylans are obtained['68].Nowadays, acetyl xylan esterase activity has been recognized as a part of the xylanolytic enzyme system of many organisms such as T reesei, T. uiride, A. niger, Schizophilum commune['69]and Streptomyces sp. [161]. The importance of this enzyme in the hydrolysis of xylan has been clearly demonstrated. Incubation of endoxylanases with acetylated glucuronoxylan resulted in the production of small amounts of xylose, xylobiose, xylotriose and large amounts of substituted oligomers. The addition of acetyl xylan esterase to the hydrolyzed mixture significantly increases the production of xylotriose and xylotetr~se['~~]. Similarly, an enzyme mixture of endoxylanase and p-xylosidase results in a limited hydrolysis of acetylated xylooligoThus, mers. The addition of acetyl xylan esterase enhanced xylose complete hydrolysis of acetylated xylans by xylanases will require the deacetylation of the substrate by acetyl xylan esterases 11.4.4.7

Mechanism o f Action of Endoxylanase

Most of the studies on the mechanism of action of endoxylanase arise from the work of Biely et al. [172,1731 using the yeast Cryptococcus albidus. The reaction of the enzyme with 5 mM [ U-14C] xylotriose resulted in a constant product ratio of xylobiose to xylose throughout the reaction. However, when the concentration of [ U-I4C] xylotriose was increased, the major product formed was xylobiose. Xylotetrose is cleaved at the middle glycosidic bond to form xylobiose. Xylopentose when present in low concentrations is converted to xylobiose and xylotriose in a ratio of 2:1. However, at higher concentrations xylotetrose is also produced. The action of endoxylanase on xylotriose, xylotetrose and xylopentose is usually accompanied by the formation of xylooligosaccharides larger than the original substrates. These studies also revealed that xylose and xylobiose can act as acceptors for the transferase reaction of xylanase. Although the acidic endoxylanase produced by Aspergillus niger differs from that of C. albidus, the mechanism of action of the enzyme is similar to the yeast enzyme. The mechanism of action of endoxylanase appears to be analogous to that reported for lysozyme and a-amylase 11.4.4.8 Biotechnological Applications o f Xylanases

Plant polysaccharides are a major source of renewable substrates for the chemical, pharmaceutical and feed industries [12'1. Xylan-degrading enzymes have considera-

7 1.4 Natural Polysaccharjde-degrading Enzymes

ble potential in several biotechnological applications. Two main areas for the application of xylanolytic enzymes have been discussed by Biely['341.The first is the use of xylanolytic enzymes in the presence of cellulolytic enzymes for the effective conversion of paper pulp and agricultural wastes into xylose, for the clarification of juices and must, and for the pre-treatment of cellulosic biomass to improve digestibility of ruminant feeds or to facilitate c o m p ~ s t i n g [ ~The ~ ~ second ]. area of application involves the use of xylanolytic enzymes in the absence of cellulases Most attention has been paid to the incorporation of xylanases as pre-bleaching agents for kraft pulps. Here the use of xylanases will help in reducing the kappa numbers (measure of residual lignin) of the pulp, thus reducing the requirement for chlorine during pulp b l e a ~ h i n g [ ' ~Most ~ l . of the studies on the effect of xylanases in the pre-bleaching of pulp have been conducted with enzyme preparations from Trichodema sp. The reduction of chlorine required during chlorination of pulp has been reported to be 35-41 % for hardwoods and 10-26 % for softwoods [17'3. Additional applications of xylanases are as flour improvers for bakery products, in the extraction of coffee, plant oils and starch[177],for the saccharification of biomass, and in the production of fuel and chemical feedstocks[173,17', '*'I. 11.4.5 Pectin

Pectic substances are widespread in the plant kingdom. The dry substance of primary cell walls of plants consists of up to 90% polysaccharides and their derivatives. These polysaccharides are composed of approximately equal parts of cellulose, hemicellulose and pectic substances. The exact proportion depends on the kind of plant (plant species) and the plant texture['*']. In fruits and vegetables, pectic substances are often found between the cells in intercellular regions. To the large, heterogenous group of pectic substances belong rhamnogalacturonans, galacturonans, arabinans, galactans and arabinogalactans [182].Pectins are designated as rhamnogalacturonane with the structure shown in Fig. 11.4-7:molecules of galacturonic acid are linked by a-1,4glycosidic linkages forming a helically wound chain. This chain is interrupted by rhamnose molecules which are bound by u-1,2 glycosidic linkages to the galacturonic acid [lS3,lS41. The number of galacturonic acid molecules varies according to the origin of the pectin. For instance, between two rhamnose molecules in citrus pectin there are 25 galacturonic acid molecules, whereas in tomato pectin there are 16 galacturonic acid Pectic substances have no definite molecular weight. The molecular weight may range from 23 000 for citrus pectin to 360 000 for apple or lemon pectin['85. 18'1 . Th e break of the galacturonic acid chain by rhamnose leads to a break in the regular helical structure. In these regions, molecules are substituted to a high degree. The C-2 or C-3 atoms of the galacturonic acid and the C-4 atom of the rhamnose molecules are preferentially substituted. The substituents are acetate, Larabinose, L-rhamnose, L-fucose, D-galactose, D-xylose or D-glucose. These substituents give to the pectin a complex and branched configuration['87, 188].Furthermore, the main galacturonic acid chain is substituted with polymers of L-

I

673

674

I

7 7 Hydrolysis and Formation of C - 0 Bonds

I

0

R

O

ORq

a(l,:o+

0

G OR

R': H (= Polygalacturonic acid), CH3 (=Pectin) R : H, Acetate, L-Arabinose, L-Fucose, D-Galactose, D-Glucose, D-Xylose Araban. Galactan Rh: Rhamnose G : D-Galacturonic acid Figure 11.4-7.

0 I

COOR'

Roq

Structure o f pectin.

arabinose (1,s-linked arabinan) and D-galactose (p-1,4- or p-1,3-linked galactans). Also, arabinogalactan I, which contains p-1,4galactan, has been reported to form side chains[']. These various side chains account for the complexity of pectic substances. The degree of substitution and the kind of substituents is dependent on the source of the pectin. In addition to the modifications on the C-2/C-3 of galacturonic acid and the C-4 of rhamnose, a large number of carboxyl groups of the galacturonic acids are esterified with methanol [1891. The degree of esterification varies with the source of the pectin. Apple pectin is esterified to the extent 80-90 % and citrus pectin to 45-60 %[l9O].

-

7 7.4 Natural Polysaccharide-degrading Enzymes

1 Cellusose fibers

I 1

Xyloglucan sugar side chains of pectin

Rhamnogalacturonan (pectin)

Figure 11.4-8.

Structure of protopectin

11.4.5.1 Classificationof Pectic Substances

Protopectin is composed of water-insolublepectic substances, which are fixed to the middle lamella and primary cell walls of plant cells. The neutral sugar side chain of the pectin is attached to the xyloglucan residues, which are bound to the cellulose Protopectin fibers. The protopectin includes polyvalent such as calcium (Fig. 11.4-8). is present in unripe fruits. During the maturation process of fruits or after harvesting, the protopectin is converted to soluble pectin [1851. The insolubility of protopectin may be due to the polymerization of the molecule and to the crosslinking with divalent cations [186]. Pectin (pectinate) consists of rhamnogalacturonan molecules that are modified with neutral sugar side chains. The carboxyl groups of the galacturonic acid molecules are partially esterified with methanol. The concentration of pectin in fruits varies with the degree of ripeness and the storage conditions. The average pectin concentration in fruits (not citrus fruits) varies between 0.5 and 1%[186]. The completely demethoxylated pectin is designated as polygalacturonic acid (polygalacturonate) or pectate. 1 1.4.5.2 Pectolytic Enzymes

Pectolytic enzymes are widespread in nature, as they have been found in plants, fungi, insects, nematodes, protozoa and bacteria. During fruit development, ripening and leaf abscission, pectin-degrading enzymes play an important role [192-1951. Furthermore, plant pectinases are important in the defensive mechanisms preventing attack of the plant by pathogenic microorganisms. Microorganisms, especially plant pathogenic microorganisms, produce a wider spectrum of pectolybc enzymes than plants themselves. Many of these extracellular enzymes occur in multiple forms, which enhance the adaptation of the plant pathogens to different hosts[196,19’1 . The most important enzyme in the plant pathogenesis process is the endo-polygalacturonase(for review see [1981). Pectinases synthesized by microorganisms also take part in symbiotic processes and in the

I

675

676

I

7 7 Hydrolysis and Formation ofC-0 Bonds

Protopectin

methylesterase Pectin

Polygalacturonic acid (PGA)

hydrolase

lyase

Methyloligogalacturonates Figure 11.49.

unsaturated methyloligogalacturonates

oligo- and monogalacturonates

unsaturated oligoand digalacturonates

Action of pectolytic enzymes.

rotting of plant material. Therefore, pectolytic enzymes are widespread in pathogenic, symbiotic microoganisms, saprophFc soil bacteria and rumen bacteria. To this group belong members of the genus Envinia, Pseudomonas, Xanthomonas, Agrobacteriurn, Corynebacterium, Lactobacillus, Arthrobacter, Bacillus, Flavobacterium, Azospirillum, Actinomyces, Yersinia, Klebsiella, Clostridium, Cytophaga, Bacteroides and Lachnospira [199-205, 231-2341 11.4.5.3 Classification o f Pectolytic Enzymes

One can distinguish between three different types of enzymes acting on pectic substances (Fig. 11.4-9):protopectinases, which degrade protopectin, pectin methylesterases, which release methanol from the galacturonic acid, and depolymerizing enzymes. The group of depolymerizing enzymes is further divided into four subgroups according to the reaction mechanisms (hydrolases and lyases) and the substrates being used (pectin and polygalacturonic acid). 11.4.5.4 Protopectinase

Protopectinasesare enzymes acting on the water-insolubleprotopectin. By the action of protopectinases the protopectin is solubilized, and water-solublehighly polymerized pectin is released. These enzymes were first described by Sakai and Okushima in 1978[206]. Further investigations of protopectinases have been r e p ~ r t e d [ ~ ~ ~ - ~ ~ ’ I . Protopectinases (or pectin-liberating enzymes) have two points of attack in the protopectin (Fig. 11.4-8):the polygalacturonic regions of the protopectin (A-type of protopectinases) and the sugar side chains, which connect the protopectin to the xyloglucans and to the cellulose fibers of the cell walls (B-type of protopectinases) [2101. A-type protopectinases are produced by yeast, Kluyveromycesfiagilis, Galactomyces reesei I F 0 0288 and Trichosporon penicillatum SNO 3. Some of these extracellular enzymes have been purified from the concentrated culture broth[211,212]. Basedon

1 1.4 Natural Polysacchan'de-degrading Enzymes

its ability to hydrolyze the polygalacturonic acid backbone, protopectinase A is classified in the group of endo-polygalacturonases(E. C . 3.2.1.15, see also 4.2.5.1.). The protopectinase A hydrolyzes the glycosidic linkages in polygalacturonic acid if at least three unmethoxylated galacturonic acid molecules are present at a short distance. According to this, the molecular mass of pectic products increases with the increasing degree of esterification of the glucoronic acid residue l2l0]. B-type protopectinases, on the other hand, are unable to degrade the polygalacturonic acid chain. These enzymes were first detected in the culture filtrate of B. subtilis I F 0 12113 by Sakai and Ozaki in 1988[2131. Many strains of Bacillus species, including B. amyloliquefaciens, B. cereus, B. circulans, B. coagulans, B. firmus, B. lichen$ormis, B. macerans and B. pumilus, have been found to be good sources of Btype protopectinases[210].The production of B-type enzymes is repressed in the presence of glucose and enhanced in the presence of starch and soybean flour extract containing arabinogalactan. 11.4.5.5 Pectin Methylesterase

Pectin methylesterases (E. C. 3.1.1.11) deesterify the galacturonic acid methylester in pectins liberating pectic acid and methanol (Fig. 11.4-10a).The hydrolysis is characterized by high specificity and a high yield (98%)L2l4]. The deesterification proceeds from the reducing end of the pectin molecule in a linear mode along the chain"]. Pectin methylesterases are produced by molds, yeasts and bacteria [1851. In general, pectin methylesterases are active in the pH range 5.0-8.0. In contrast to fungal enzymes, which are active at low pH, the bacterial esterases prefer alkaline conditions. In fruits and vegetables, especially in citrus fmits and tomatoes, high pectin methylesterase activities have also been found. 11.4.5.6 Pectin and Polygaladuronate Depolyrnerizing Enzymes

The activity of pectin-depolymerizing hydrolases, especially endoacting enzymes, can be followed by a rapid decrease in the viscosity of the pectin solutions. By the cleavage of only 2-3 % of the glycosidic bonds the viscosity diminishes to about 50%. In addition, the increasing amount of reducing ends can be determined. The last stage of pectin depolymerization on an industrial scale is proved by the alcohol test. The depolymerizing reaction is complete when the addition of 50 % alcohol to the reaction mixture does not lead to flocculationL2l4]. The activity of trans-eliminases (lyases) can be followed photometrically by measuring the UV adsorption of 4,5-dehydrogalacturonicacid at 232 nm[2151.

I

677

678

I

1 1 Hydrolysis and Formation of C - 0 Bonds

+ n20

r

a. Pectin methylesterase

-

n

0

H O Q n

-%

+ H,O

+

HO =OH

OH

%

HO

HO

HO

?

?

q0

%

b. Pectin and polygalacturonicacid hydrolase 0

HO

HO O

W

O

+

H

OH

H

O

4

HO

?

0

c. Pectin and polygalacturonic acid lyase (R'= H: polygalacturonicacid; R'= CH3: pectin) Figure 11.4-10.

Reaction mechanisms of pectolytic enzymes.

11.4.5.7 Pectin and Polygalacturonate Hydrolase

Pectin hydrolase and polygalacturonate hydrolase (polymethylgalacturonase, polygalacturonase) catalyze the cleavage of the polysaccharide backbone of pectin and polygalacturonate. Pectin hydrolases prefer pectin, and polygalacturonases prefer polygalacturonic acid as substrates (Fig. 11.4-lob). According to the mode of action, these enzymes can be defined as endo- or exoenzymes. Exoenzyrnes are able to split mono-, di- or trimers from the reducing end of the polysaccharide chain (pectin or polygalacturonic acid). Endoacting enzymes, on the other hand, attack the complex polysaccharide in the inner part of the chain backbone, resulting in a rapid decrease of viscosity of pectin- or polygalacturonate solutions. Endoacting enzymes prefer long polysaccharide chains of pectin or polygalacturonic acid. The activity decreases with decreasing chain length. Endopolygalacturonatehydrolases (E. C. 3.2.1.15) are widespread in fungi, in most plant pathogens, in some bacteria, in plant organs and in the digestive tracts of some insects [*lG]. The enzyme catalyzes the random hydrolytic cleavage of a-1,4linkages of

7 7.4 Natural Polysaccharide-degrading Enzymes

galacturonan and requires free carboxyl groups for their catalytic activity. The activity therefore decreases with increasing degree of esterification of the polygalacturonic acid substrate [2171. Endopolygalacturonases have been purified from several plant and microbial sources and are optimally active under acidic conditions (PH 2.5-6.5). Most exopolygalacturonases release D-monogalacturonic acid from the non-reducing end of the chain (E. C. 3.2.1.67). The enzymes produced by Erwinia aroideae and Pseudomonas sp. (E. C. 3.2.1.82) are able to release digalacturonic 218]. Exopolygalacturonases from fungi exhibit optimal activity between pH 4.0 and pH 6.0, whereas the enzymes from Clostridium multijirmentans show highest activity at pH 7.2. In addition to exo- and endopolygalacturonases a number of microorganisms produce oligogalacturonases which hydrolyze oligogalacturonate chains forming short oligomers and galacturonate. The oligogalacturonases have higher affinity to low molecular weight oligogalacturonates than to polygalacturonates. The activity decreases with increase of the chain length of the substrate. The oligogalacturonases from Bacillus species and A. niger attack the substrate from the non-reducing end, whereas the enzymes produced by Erwinia carotovora and E. aroideae hydrolyze the substrate from the reducing end['8G]. 11.4.5.8

Pectin and Polygalacturonate Lyase

The reaction mechanism of lyases is characterized by a trans-elimination reaction resulting in 6 4,5-unsaturated galacturonic acid molecules. The lyases are calcium dependent and attack either pectin (pectin lyases) or pectic acid (polygalacturonate lyases) from the non-reducing end (Fig. 11.4-1Oc). Endopolygalacturonate lyase (E. C. 4.2.2.2) has been detected in many bacteria and some pathogenic fungi. These enzymes show highest activity under alkaline conditions in the pH range 8-10. The enzyme activity depends exclusively on the presence of calcium ions and decreases with decreasing chain length of the polygalacturonic acid. Exopolygalacturonatelyases (E. C. 4.2.2.9) have been detected in only a few bacteria which belong to the genera of Clostridium, Erwinia, Streptomyces and Fusarium['861. The majority of these enzymes are active under alkaline conditions (pH 8-9.5) and require calcium ions for activity. Erwinia carotovorans and E. aroideae have been found to synthesize oligogalacturonate lyase (E. C. 4.2.2.6) [219, 220]. The enzyme releases unsaturated monomers from the reducing end of the oligogalacturonate substrates. Endopectin lyases (E.C. 4.2.2.10) are widespread in fungi and prefer long polymethylgalacturonate chains (pectin) as substrates, resulting in decreasing activity with decreasing chain The distribution of different pectolytic enzymes in microorganisms is shown in Table 11.4-4. For review see [lSG.23G1.

I

679

680

I

7 7 Hydrolysis and Formation of C - 0 Bonds Table 11.4-4.

Occurence of different pectolytic enzymes in microbes.

Organism

Fungi: Aspergillus niger Aspergillus alliaceus A.flavus A. fumigatus Botrytis cinerea Fusarium tricinctum

PME

Pectin Pectin hydrolases lyases

+

-

+

+

-

+

i

-

PCA lyases

Ref.

+

-

+ +

+

-

-

i

-

-

+

+

-

-

+

-

-

+ +

+

[664] [665] [666] [666] [667, 6681 [669]

+ +

-

[670] [671]

-

[672] 16731 [674] [675] [676] [677] [678] [679] 16801

Yeasts:

Candida pseudotropicalis Saccharomyces vini

PCA hydrolases

-

-

-

+

-

-

Bacteria:

Clostridium pectinofmentans C. thermosulfurogenes 4B C. thermosaccharolyticum Bacillus stearothemophilus Corynebacterium michiganenese Enuinia chrysanthemi Pseudomonas marginalis Streptomyceskadiae Xanthomonas campestris

+

-

+ +

-

-

+ -

+ + +

-

-

-

-

-

-

-

+

+

-

+

+

-

-

-

-

+ + -

-

+

-

+

-

+

+

-

+

-

+

PME: Pectin methylesterase; PGA: Polygalacturonicacid.

11.4.5.9

Biotechnological Applications of Pectolytic Enzymes

Enzymes with pectolytic activity have been used since 1930 in the clarification of fruit juices. In freshly pressed apple juice, pectin acts as a stabilizing colloid for the insoluble cell debris. After hydrolysis of the pectin, the insoluble particles floc out. Also, in white wine production, a clarification process for the removal of insoluble particles suspended in the grape must is The commercial enzyme preparations for industrial application may contain, as well as pectolytic enzymes, cellulases, hemicellulases, xylanases and proteases. All these enzymes solubilize the cell wall constituents to form soluble products such as galactose, mannose, rhamnose, arabinose, galacturonic acid and methanol [222* 2231. Similar processes are in use for the maceration of vegetables and the extraction of olive oil. The preincubation of sugar beet with pectolytic enzymes (1-2 h at 54 "C or 6-8 h at 18 "C) before the pressing procedure improves the yield significantly[224]. Pectin methylesterases are also used in the production of apple cider. After demethoxylation of pectin, the product formed (polygalacturonic acid) can be easily removed from the fermenting apple juice by precipitation with calcium ions['"]. Pectolytic enzymes are also involved in natural fermentation processes. The coffee seeds (coffeebeans) are directly surrounded by the so-called seed coat or silver skin, followed by the endocarp (hull), the mesocarp (mucilage layer) and the exocarp (skin). One of these envelopes, the mesocarp, consists of 30% pectic substances.

References I681

Table 11.4-5. Organism

A. niger

Bacillus sp. Penicillium sp. Rhizopus sp.

Microorganisms used for the commercial production of pectolytic enzymes. Pectin methylesterase

Pectin hydrolase

Pectin lyase

PCA hydrolase

PCA lyase

Oligogalacturonase

-

-

+

-

-

+ -

+ -

PGA: Polygalacturonic acid.

This polysaccharide is degraded by the pectolytic enzymes that are produced by the epiphyhc microbial flora of the coffee fruits, i.e. Envinia and Enterobacter spec i e ~ [ ~ After ~ ' ] . 1-4 days the digestion is complete and a mechanical depulping step of the coffee fruits can take place. Also, by the cocoa fermentation during the first 1-2 days, pectolytic enzymes from yeasts aid in the maceration of the cocoa pulp and the draining of the fluid. The fermentation of cocoa and coffee fruits can be enhanced by the addition of commercial enzyme preparations containing pectindepolymerizing enzymes [2351. Protopectinases are also used in the production of pectin from mandarin orange peel. Pectin can be used as an additive in the food and cosmetic industries[22G]. In all applications described above involving conventional pectolFc enzymes, the rhamnogalacturonan backbone of pectic substances is not degraded ~ o m p l e t e l y [ ~ ~ ~ l . It has been reported that Aspergillus aculeatus produces an enzyme complex consisting of 10 to 15 different enzymes. This enzyme complex has the potential for the complete hydrolysis of complex polysaccharides and may support liquefaction processes with plant material, fruits or vegetables[2271. For the commercial production of pectolytic enzymes, Aspergillus niger or related species are mainly used. In these fermentations, low value agricultural products containing pectin are used as Table 11.4-5shows some of the microorganisms that are used for substrates [228-2301. the industrial production of pectolytic enzymes.

References I 0. P. Ward, M. Moo-Young, CRC Crit. Rev.

Biotechnol. 1989, 8, 237-274. z A. Guilbot, C. Mercier i n 7'he Polysaccharides,Vol. 3, G. 0. Aspinall (ed),Academic Press, Inc., Orlando USA, 1985,210282. 3 J. F. Kennedy, J. M. S . Cabral, S. A. Correia, C. A. White in Starch: Properties and Potential, T. Galliard (ed), John Wiley & Sons., Chichester, England, 1987, 115-148. 4 D. French, in Trends i n the Biology ofFernentationfor Fuels and Chemicals. A. Hollaender (ed), Plenum Press, New York, USA, 1981, 151.

J. J. Marshall, Adv. Carbohydr. Chem Biochem. 1974, 30,257. 6 J. D. Allen, J. A. Thoma,. Carbohydr. Res. 1978,Gl. 377-385. 7 P. E. Granum,]. Food. Biochem. 1979,3, 1-12. 8 N. Yoshigi, T. Chikano, M. Kamimura, Agric. Biol. Chem. 1985,47,2193-2199. 9 G. Takasaki, Agnc. Biol. Chem. 1983,47, 2193-2199. 10 J. Robyt, D. French, Arch Biochem Biophys. 1979,100,451-467. 11 V. Buonocore, C. Caporale, M. de Rose, A. Gambacorta,J. Bacteriol. 1976, 128,515. 5

682

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1 1 Hydrolysis and Formation of C-0 Bonds

E. W. Boyer, M. B. Ingle,]. Bacteriol. 1972, 110,992. 13 G. Antranikian, FEMS Microbiol. Rev. 1990, 75,201-218. 14 F. Niehaus, C. Bertoldo, M. Kahler, G. Antranikian. Appl. Microbiol. Biotechnol. 1999, 51,711-729. 15 S. H. Brown, H. R. Costantino, R. M. Kelly, Appl. Environ. Microbiol. 1990, 56, 1985-1991. 16 N. K. Matheson, B. V. McCleary in The Polysaccharides, Vol. 3, G. 0. Aspinall (ed),Academic Press, New York, USA, 1985. 17 H. H. Hyun, J. G. Zeihs, Appl. Environ. Microbiol. 1985, 49, 1162-1170. 18 G. J. Shen, B. C. Saha, Y. E. Lee, L. Ghatnagar, J . G. Zeel, Biochem.J. 1988, 254, 835-840. 19 J . H. Pazur, K. Kleppe,J . Biol. Chem. 1962, 237,1002. 20 J. J. Marshall, W. J. Whelan, FEBS Lett. 1970,9,85-88. 21 J. J. Marshall, Starch/Stdrke 1975, 27, 377-383. 22 B. C. Saha, T. Mitsue, S. Ueda, StarchlStarke 1979,231,307-314. 23 U. Specka, F. Mayer, G. Antranikian, Appl. Environ. Microbiol. 1991, 57, 2317-2323. 24 W. M. Fogarty, in Microbial Enzymes and Biotechnology. W. M. Fogarty (ed), Applied Science Publishers, London, England, 1983, pp. 71-132. 25 C. T.Kelly, W. M. Fogarty, Process. Biochemistry 1983, 18, 6. 26 G. Antranikian, in Microbial Degradation of Starch. G. Winkelmann (ed), VCH, Weinheim, Germany, 1992,27-56. 27 Y. Suzuki, T. Yuki, T. Kishigami, S. Abe, Biochim. Biophys. Acta 1976,445, 386-397. 28 A. Amemura, T. Sugimoto, T. Harada, J . Ferment. Technol. 1974, 52, 778-780. 29 A. Amemura, Y. Konishi, T. Harada, Biochim. Biophys. Acta 1980,611, 390-393. 30 R. M. Evans, D. J. Manners, J. R. Stark, Carbohydr. Res. 1979,76,203-213. 31 K. R.Adams, F. G. Priest, FEMS Microbiol. Lett. 1977, I , 269-273. 32 K. Kainuma, S. Kobayashi, T. Harada, Carbohydr. Res. 1978, 61, 345. 33 B. E. Norman, Starch/Starke 1982, 34, 340-346. 34 A. R. Plant, R. M. Clemens, R. M. Daniel, H. W. Morgan, Appl. Microbiol. Biotechnol. 1987,26,427-433. 12

35 A. R. Plant, R. M. Clemens, H. W. Morgan,

R. M. Daniel, Biochem. J.

36 H. Melasniemi, Biochem. J. 1988,250,

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120T.K. Ng, J. G. Zeikus, Biochem.]. 1981, 199,

341-350.

121 N. Creuzet, C. Frixon, Biochemie 1983,65,

149-156.

122 N. Creuzet, J.-F. Berenger, C. Frixon, FEMS

Microbiol. Lett. 1983,20, 347-350.

123 W. T. H. Chang, D. W. Thayer, Can. J. Micro-

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124 M. McGavin, C. W. Forsberg,]. Bacteriol.

1988,170,2914-2922.

125 L. Huang, C. W. Forsberg. D. Y. Thomas,

J. Bacteriol. 1988,170,2923-2932.

126 K. Ohmiya, K. Maeda, S . Shimizu, Carbo-

hydr. Res. 1987,166,145-155.

127 K. Ohmiya, M. Shirai, Y. Kurachi,

S. Shimizu,]. Bacteriol. 1985, 161, 432434. 128 C. E. Ware, A.M. Lachke, K. Greg, Biochem. Biophys. Res. Comm. 1990, 171, 777-786. 129 H.J. Gilbert, G. P. Hazlewood,]. Gen. Microbiol. 1993, 139, 187-194. 130 H.J. Gilbert, G. P. Hazlewood, Proc. Nutr. SOC.1991,50, 173-186. 131 J. Woodward in Topics in Enzyme and Fermentation Biotechnology A. Wiseman (ed), Ellis Horwood, Chichester, England, 1984, pp. 9-27. 132 J. P.Joseleau, J. Comtat, K. Rue1 in Xylans and Xylanases J. Visser, G. Beldman, M. A. Kusters-van Someren, A. G. J. Voragen (eds),Elsevier, Amsterdam, The Netherlands, 1992, pp. 1-15. 133 R. L. Whistler, E. L. Richards in The Carbohydrates- Chemistry and Biochemistry,Vol. 2A, 2nd edn., W. Pigman, D. Horton (eds), Academic Press Inc., New York, USA, 1970, pp. 447-469. 134 A. Sunna, G. Antranikian, Crit. Rev. Biotechnol. 1997, 17, 39-67. 135 J. Puls, K. Poutanen in Enzyme Systemsfor Lignocellulose Degradation M. P. Coughlan (ed),Elsevier Applied Science, London, England, 1989, pp. 151-165. 136 S. K. Chandra, E. L. Hirst, J. K. N. Jones, E. G. V. Percival,J. Chem. Soc. 1950, 1289-1297. 137 S. Eda, A. Ohnishi, K. Kato, Agric. Bid. Chem. 1976,40,359-364. 138 R. Montgomery, F. Smith, H. C. Srivastava, ].Am. Chem. Soc. 1956,78,2837. 139 T.E. Timell, Adv. Carb. Chem. 1964, 19, 247-302. 140 R.F. H. Dekker, A C S Symp. Ser. 1989, 399, 616-629.

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H. 0. Bouveng, Acta Chem. Scand. 1961, 15, 96-100. 142 B. Lindberg, K:G. Rossell, S. Svensson, Svensk. Papperstidn. 1973, 76, 30-32. 143 J. Puls, J. Schuseil in Hemicelluloses and Hemicellulases M. P. Coughlan, G. P. Hazlewood (eds),Portland Press, London, England, 1993, pp. 1-97. 144 M. Zinbo, T. E. Timell, Svensk. Papperstid. 1967,70,597-606. 145 P. J. Reilly in Trends in the Biology ofFernentations A. Hollaender (ed), Plenum Press, New York, USA, 1981, pp. 111-129. 146 R. F. H. Dekker, G. N. Richards, Adv. Carbohydr. Chem. Biochem. 1976,32, 277-352. 147 K. K. Y. Wong, L. U. L. Tan, J. N. Saddler, Microbiol. Rev. 1988, 52, 305-317. 148 M. Debeire-Gosselin, M. Loonis, E. Samain, P. Debeire in Xylans and Xylanases J. Visser, G. Beldman, M. A. Kusters-van Someren, A. G. J. Voragen (eds), Elsevier, Amsterdam, The Netherlands, 1992, pp. 463-466. 149 M. Debeire-Gosselin, J. P. Touzel, P. Debeire in Xylans and Xylanases J. Visser, G. Beldman, M. A. Kusters-van Someren, A. G. J. Voragen (eds),Elsevier, Amsterdam, The Netherlands, 1992, pp. 471-474. 150 S. Takenishi, T. Tsujisaka, Agnc. Bid. Chem. 1975,39,2315-2323. 151 A. Matte, C. W. Forsberg, Appl. Environ. Microbiol. 1992, 58, 157-168. 152 A. C. Grabski, T. W. Jeffries, Appl. Environ. Microbiol. 1991, 57, 987-992. 153 T. M. Wood, S. 1. McCrae, Carbohydr. Res. 1986, 148,321-330. 154 F. Deleyn, M. Claeyssens, J. Van Beeumen, C. K. De Bruyne, Can. /. Biochem. 1978,56, 43-50. 155 W. Zimmermann, B. Winter, P. Broda, F E M S Microbiol. Lett. 1988, 55, 181-186. 156 J. Dobberstein, C. C. Emeis, Appl. Microbiol. Biotechnol. 1991, 35, 10-215. 157 S. L. Bachmann, A. J. McCarthy, Appl. Environ. Microbiol. 1991, 57, 2121-2130. 158 D. Conrad, W. Noethen in Proc. 3rd Eur. Congr. Biotechnol. 1984, 2, pp. 169-177. 159 M. Claeyssens, E. Van Leemputten, F. G. Loontiens, C. K. De Bruyne, Carbohydr. Res. 1966, 3, 32. 160 M. Claeyssens, F. G. Lonntiens, H. Kersters-Hilderson, C. K. De Bruyne, Enzymologia, 1971,40,177-198. 141

C. R. MacKenzie, D. Bilous, H. Schneider, K. G. Johnson, Appl. Environ. Microbiol. 1987,53,2835-2839. 162 A. Kaji, K. Tagawa, Biochem. Biophys. Acta 1970,207,456-464. 163 K. Komae, A. Kaji, M Sato, Agnc. Bid. Chem. 1982,46,1899-1905. 164 K. Poutanen,/. Biotechnol. 1988,7, 271-292. 165 J. Puls, K. Poutanen, 0. Schmidt, M. Linko in Proc. 3rd Int. Con5 Biotechnol. Pulp Paper Industry K. E. Eriksson, P. Ander (eds),STFI, Stockholm, Sweden, 1986, pp. 93-95. 166 J. Puls in Xylans and Xylanases J. Visser, G. Beldman, M. A. Kusters-van Someren, A. G. J. Voragen (eds), Elsevier, Amsterdam, The Netherlands, 1992, pp. 213-224. 167 P. Biely, C. R. MacKenzie, J. Puls, H. Schneider, Bio/TechnoL 1986,4, 731-733. 168 M. Tenkanen, K. Poutanen in Xylans and Xylanases J. Visser, G. Beldman, M. A. Kusters-van Someren, A. G. J. Voragen (eds), Elsevier, Amsterdam, The Netherlands, 1992, pp. 203-212. 169 P. Biely, J. Puls, H. Schneider, FEBS Lett. 1985, 186,80-84. 170 J. Puls, M. Tenkanen, H. E. Korte, K. Poutanen, Enzyme Microb. Technol. 1991, 13, 483-486. 171 K. Poutanen, J. Puls, A C S Symp. Ser. 1989, 388,630-639. 172 P. Biely, 2. Kratky, M. Vrsanska, Eur. /. Biochem 1981, 119,559-564. 173 P. Biely, M. Vrsanska, 2. Kratky, Eur. /. Biochem. 1981, 119, 565-571. 174 M. Vrsanska, 1. V. Gorbacheva, 2. Kratky, P. Biely, Biochem. Biophys. Acta 1982,704, 116122. 175 L. Viikari, J. Sundquist, J. Kettunen, Paperi j a Puu 1991,73,384-388. 176 Cultor, Albazyme@,Pulp and Paper Enzymes (product data sheet, Cultor, UK Ltd.), 1991. 177 P. Biely, A C S Symp. Ser. 1991, 460,408-416. 178 M. Linko, K. Poutanen, L. Viikari in Enzyme Systemsfor Lignocellulose Degradation M. P. Coughlan (ed), Elsevier Applied Science, London, England, 1989, pp. 331-346. 179 K. Poutanen, M. Ratto, J. Puls, L. Viikari,/. Biotechnol. 1987, 6, 49-60. 180 Y.-E. Lee, E. E. Lowe, J. G. Zeikus, Appl. Environ. Microbiol. 1993, 59, 763-771. 161

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

686

I

1 1 Hydrolysis and Formation of C-0 Bonds

11.5 Addition of Water to C=C Bonds

Marcel Wubbolts

The addition of water to carbon-carbondouble bonds is a reaction that is catalyzed by lyases belonging to the subclass of the hydro-lyases (E. C. 4.2.1.-),which have been grouped under the carbon-oxygen lyases. Not all members of this subgroup are capable of water addition to carbon-carbon double bonds. Nitrile hydratase (E. C. 4.2.1.84, discussed in Section 12.1) for instance, is categorized in this subclass and catalyzes the addition of water to nitriles. The nomenclature of the hydro-lyases subgroup, which contains hydratases and dehydratases, does not preclude any direction of the reaction, but rather reflects the context in which the enzyme was originally discovered. The addition of water to carbon-carbon double bonds is very common to biology, and a large variety of enzymes from different sources representing almost a hundred different hydro-lyase types have been characterized biochemically. Hydro-lyases are for instance involved in the metabolism of a variety of carbohydrates and play a prominent role in fatty acid synthesis and degradation as well. Despite the abundant presence of hydro-lyases in nature, however, applications of these enzymes in organic chemical synthesis are not as widespread. This is mainly due to the limited availability of these enzymes and the fact that many of the enzymes cannot easily be stably maintained during catalysis. 11.5.1 Addition of Water to Alkenoic Acids

The catabolic enzyme 2-oxopent-4-enoatehydratase (E. C. 4.2.1.80) is involved in Lphenylalanine metabolism and in the degradation of a number of aromatic hydrocarbons as well[']. It catalyzes the selective addition ofwater to a terminal C-C double bond of cis-2-hydroxypent-2,4-dienoic acid and forms 4-hydroxy-2-oxopentanoic acid. The enzyme also accepts cis-2-hydroxyhex-2,4-dienoic acid as a substrate, but is not active on the trans-isomer121. D-Tartaric acid dehydratase (E. C. 4.2.1.81) and the stereochemical counterpart Ltartaric acid dehydratase (E.C. 4.2.1.32) are able to catalyze the conversion of oxaloacetic acid to D- and L-tartaric acid respectively. The actual addition of water to the C-C double bond is most likely to occur at the enol tautomer, and the resulting tartaric acid has the 2S,3S (D-stereo isomer made by E.C. 4.2.1.81) or 2R,3R (Ltartaric acid dehydratase)configuration. Despite the stereochemistry of the reactions catalyzed, the lack of available enzyme and the instability of the enzymes in presence of oxygen[3]have hampered their application in organic synthesis thus far.

11.5 Addition of Water to C=C Bonds

I

687

Furnarate hydratase

-

1

HOOC-COOH

H

O

3

-

COOH

COOH

-

-

YCOOH

2

COOH

HOOC

-

-

OH

Citraconate hydratase

5

HOOC/\\/

-

C

OH

Maleate hydratase

H O O C T

O

H O O C T C o o H

0

OH

Mesaconate hvdratase

3-isopropylmalate dehydratase

COOH HOOC

10

HOOC

COOH

I

dH Dirnethylrnaleate hydratase

Scheme 11.5-1.

11.5.2 Addition of Water to Alkene-Dioic Acids 11.5.2.1 L-

and D-Malic Acid

The production of L-malic acid (2)from fumaric acid (1)is carried out by the enzyme fumarate hydratase (E.C. 4.2.1.2). which is part of the tricarboxylic acid cycle and ubiquitous in nature (Scheme 11.5-1). The product is used in food, pharmaceutical and cosmetic industries and is produced at a multi-ten-tonne scale. Although the enzyme can be applied in isolated form, as performed by Tanabe, the use of whole cells of Corynebacteriurn glutarnicurn has been reported by Amino GmbH as well[4].

688

I

11 Hydrolysis and Formation of C-0 Bonds

The conversion of fumaric to L-malic acid is brought to completion by forcing the product to precipitate as calcium salt [41. The synthesis of D-Malic acid (4) from maleic acid (3) by maleate hydratase (E. C. 4.2.1.31) has been described as early as 1969, using an enzyme from rabbitI5]. Maleate hydratase from various other, more accessible sources such as Pseudomohave been used for the same purpose. The combined use of calcium-counter ions and maleate hydratase (E. C. 4.2.1.31) from Pseudomonas pseudoalcaligenes has been an elegant method to produce on multi-kilogram scale D-malate that in complex with calcium precipitated out from solution, thereby eliminating the reverse reaction[6]. He et al. used a similar enzyme from Arthrobacter pascens DMDCl2, which is called (R)-2-methylmalatedehydratase, citraconate hydratase or citraconase (E.C. 4.2.1.35), to produce D-malate as well as D-citramalate or ( R ) 2-methylmalic acid (6) from 2-methylmaleate ( S ) , using an enzyme membrane reactor The demand for D-malate is limited: it merely serves as a general synthetic building block for chiral 1' and as a resolving agent. 11.5.1.2 Substituted Malic Acids

The enzyme of opposite selectivity relative to citraconase (E. C. 4.2.1.35) mentioned above, is (S)-2-methylmalatedehydratase or mesaconate hydratase (E. C. 4.2.1.34), which has, among others, been found in Clostridium t e t a n o m ~ ~ h u m [Pseudomo~]], nasl"], Citrobacter and Morganella["], and is of use to convert 2-methylfumarate (7) to the (S)-isomerof citramalate (8). Interestingly, both citraconase and mesaconate hydratase have a broader substrate range and are also able to produce the respective stereo-isomers of malic acid and 2-ethylmalic acid [', 91. The 3-iso-propylmalate dehydratase (E. C. 4.2.1.33) from Neurospora crassa and numerous other prokaryotic strains are involved in synthesis of L-valine, L-isoleucine and L-leucine. The enzyme accepts the iso-propyl group as a substituent during the reaction and converts 2-isopropylmaleate (9) to 3-iso-propylmalate(10)[l21. Dimethylmaleate hydratase (E. C. 4.2.1.85) has been described as the enzyme that catalyzes the addition of water to dimethylmaleate (11) to yield a molecule with two chiral centers, (2R,3S)-2,3-dimethylmalate (12)[131. 11.5.3 Addition o f Water to Alkene-Tricarboxylic Acids 11.5.3.1 Citric Acid and Derivatives

Other C-0 lyase enzymes include aconitate hydratase or aconitase (E.C. 4.2.1.3), an enzyme that catalyzes two tricarboxylic acid cycle steps from isocitric acid to citrate (14)[141 or vice versa, via the intermediate cis-aconitate(13). Citrate dehydratase (E. C. 4.2.1.4) is only capable of converting citrate to cis-aconitate and does not act on isocitrate (15)[151.

11.5 Addition of Water to C=C Bonds

COOH

I

689

Citrate dehydratase or Aconitase

.

VooH 13

H O O C T c o o H

COOH

\COOH

Aconitase

13

COOH

l""OO"

16

OH

I

HoocY 15

.

Homo-aconitate hydratase

14

H O O C ~ c o o H

\7

17

COOH

COOH COOH

2-Methylcitrate dehydratase

COOH

18

19

\COOH

\COOH

2-Methylisocitrate de hydratase

COOH

18

20

\COOH

4-Oxarnesaconate hydratase

Hooc-= Hooc 22

21

COOH

COOH

Scheme 11.5-2.

A similar reaction is catalyzed by homoaconitate hydratase (E. C. 4.2.1.36),which is an enzyme from the L-lysine synthesis that forms homocitric acid (2-hydroxybutane-1,2,4-tricarboxylic acid, 17) from homo-cis-aconitate (16)[16]. The enzyme 2-methylcitratedehydratase (E. C. 4.2.1.79) catalyzes the addition of water to (Z)-but2-ene-1,2,3-tricarboxylic acid (18) to yield 2-methylcitric acid (2-hydroxybutane-

690

7 7 Hydrolysis and Formation of C-0 Bonds

I 1,2,3-tricarboxylicacid,

19) [l']. 2-Methylisocitrate dehydratase (E. C. 4.2.1.99) from Yarrowina lipolytica does not accept isocitrate (15) as substrate, but rather acts on (2)but-2-ene-1,2,3-tricarboxylic acid (18) to produce 2-methylisocitrate(20)["I. Lastly, 4-carboxy-2-oxohexenedioatehydratase (4-oxamesaconate hydratase, E. C. 4.2.1.83) adds water to (E)-4-oxobut-l-ene-l,2,4-tricarboxylic acid (21) and results in the formation of 2-hydroxy-4-oxobutane-1,2,4-tricarboxylic acid (22)[191 (Scheme 11.5-2). 115.4 Addition o f Water to Alkynoic Acids

Interestingly,two enzymes have been described that catalyze the addition of water to alkynes, resulting in the formation of alkenols: acetylene carboxylate hydratase from Pseudomonas (E. C. 4.2.1.71), which converts propynoic acid to 3-hydroxypropenoate f20]. The latter tautomerizes to malonic semialdehyde. Acetylene dicarboxylate hydratase (E. C. 4.2.1.72) converts acetylene dicarboxylic acid to 2-hydroxyethylenedicarboxylic acid, which spontaneously decarboxylatesto pyruvate 121]. 11S.5 Addition o f Water to Enols 11.5.5.1 Carbohydrates: Addition o f Water to 2-Keto-3-Deoxysugars

Hydro-lyasesplay a prominent role in the metabolism of sugars and of sugar-derived carboxylic acids in particular. The eliminationladdition ofwater to sugar carboxylates proceeds via an enol intermediate[22],as depicted in Scheme 11.5-3.The elimination or addition of the water molecule is highly specific, and a large variety of hydrolyases have been characterized examples include Pseudomonas saccharophila D-arabinoate dehydratase (E. C. 4.2.1.5) [231, Pseudomonas sp. and E. coli galactonate dehydratase (E. C. 4.2.1.G)[241,E. coli altronate dehydratase (E. C. 4.2.1.7)[251,E. coli mannonate dehydratase (E. C. 4.2.1.8) L2' ] , L-arabinoate dehydratase (E. C. 4.2.1.25) from Rhizobium[2G1, phosphogluconate dehydratase (E. C. 4.2.1.12) from various organisms [271, gluconate dehydratase (E. C. 4.2.1.39) from various organisms 12'], D-fuconate hydratase (E. C. 4.2.1.67) from Pseudornonas sp. [291, Mammalian L-fuconatehydratase (E. C. 4.2.1.68) i3O1, D-xylonate dehydratase (E. C. 4.2.1.82)I3l ], and fungal L-rhamnonate dehydratase (E. C. 4.2.1.90). The elimination of water from glucarate, a 1,G-dicarboxylic hexose, by glucarate dehydratase (E. C. 4.2.1.40) results in the formation of 5-dehydro-4-deoxy-~-glucarate[32].The reaction is however identical to that of the other dehydratases and the seemingly different specificity is only due to IUPAC rules (Scheme 11.5-3).The enzyme belongs to the enolase superfamily, and the structure of the enzyme from Pseudomonas putida has been resolved [331. Similarly, galactarate dehydratase from E. coli (E. C. 4.2.1.42) produces 5-dehydro-4-deoxy-~-galactarate [321.

7 7.5 Addition of Water to C=C Bonds

I

691

-

Dehydratase HokCOOH

~

HO O F C O O H

4 -

R

/

t R C O O H

-

-

Galactonate Dehydratase

HO

f:-

HO

CHpOH

CHpOH

-

D-Galaconate

'enol'

F

R

CHzOH I-OH

2-dehydro-3-deoxyD-Galactonate

-

COOH

Glucarate Dehydratase

Hoji

=

COOH

D-Glucarate

COOH -

'enol'

COOH 5-dehydro-4-deoxyD-Glucarate

Scheme 11.5-3.

11.5.5.2 Addition/Elirnination of Water with Other Enok

Dihydroxyacid dehydratase (E.C. 4.2.1.9) is a ubiquitous enzyme that is involved in the biosynthesis of the branched-chain amino acids (Ile, Leu and Val) and of pantothenic acid and coenzyme A. The enzyme catalyzes the elimination of water from 2,3-dihydroxyalkanoic acids (23) to 2-hydroxy-2-alkenoic acids (24), which tautomerize to 2-ketoalkanoic acids (25).The enzyme from spinach has the highest activity towards 2,3-dihydroxy-3-methylbutanoic acid (Val precursor, Scheme 11.5-4) but also accepts other substrates [341. Thus, 2,3-dihydroxybutanoic acid, 3-cyclopropylacid are 2,3-dihydroxybutanoic acid as well as 2,3-dihydroxy-3-methylpentanoic substrates. With the latter substrate a slight preference for (2R,3S)-2,3-dihydroxyy-

692

I

7 7 Hydrolysis and Formation of C - 0 Bonds

JCOOH

Dihydroxyacid dehydratase [
~

~

HO

23

24

25

Enolase 26

p l o ~ C O O H

27

Scheme 11.5-4.

3-methylpentanoate over the (2R,3R)-2,3-dihydroxy-3-methylpentanoate was observed [341. The glycolytic enzyme phosphoenolpyruvate (PEP) hydratase (enolase, E. C. 4.2.1.11) catalyzes the addition of water to 2-phospho-~-glycerate(26). The enzyme also accepts 3-phospho-~-erythronate(28) and thereby forms phosfrom E. c0li1~~1 phoenol-4-deoxy-3-tetrulosonate (29 in Scheme 11.5-4).Both PEP and phosphoenol4-deoxy-3-tetrulosonaterepresent “fixed enolates that can be isolated. The enzymes 1,2-propanedioldehydratase (E. C. 4.2.1.28) and glycerol dehydratase (E. C. 4.2.1.30) from the facultative anaerobic microorganism Klebsiella pneumoniae [36, 371 and other sources have recently gained interest, since these enzymes can be of use for the synthesis of 1,3-propanediol (PDO) starting from glycerol. PDO is of use for the synthesis of polyesters, and Dupont is currently developing a biological production method based on fermentation. Glycerol dehydratase (E. C. 4.2.1.30) catalyzes the elimination of water from a number of polyols: ethylene glycol to acetaldehyde, glycerol to 3-hydroxypropanal and 1,2-propanediol to propionaldehyde, all of which reactions proceed via an enol intermediate.

7 7.5 Addition of Water to C=C Bonds

11.5.6

Addition ofwater to Unsaturated Fatty Acids 11.5.6.1

CoA and ACP Coupled Fatty Acid Hydratases

Hydratases that add water to unsaturated fatty acids coupled to coenzyme A (CoA)or acyl carrier protein (ACP) cannot be used in vitro, and consequently have to be applied in whole-cellbiotransformations. Prohibitive as this may seem to production on a commercial scale, Kanegafuchi has developed a process, making use of whole cells of Candida rugosa, to produce (R)-2-hydroxybutanoicacid (31) from butanoic acid (30) (Scheme 11.5-5).The series of reactions catalyzed by these cells include coupling of butanoic acid to CoA, desaturation of butyryl-CoA to 2-butenyl-CoA and water addition catalyzed by enolyl-CoA hydratase (enoylase, unsaturated enoyl-

Y

COOH

30

32

Candida rugosa fatty acid metabolism

ll

Enoyl-CoA hydratase

Thioesterase

1 ' L C O O H

Scheme 11.5-5.

33

693

I

694

I

1 1 Hydrolysis and Formation ofC-0 Bonds

Agrobacterium fatty acid metabolism

35

tl

I

Carnitine de hydratase Carnitine dehydrogenase

36

SCoA

37

T hioesterase

38

Scheme 11.5-6.

coenzyme A hydratase, E.C. 4.2.1.17). Removal of the CoA group liberates the phydroxy acid, which is of use for the synthesis of carbapenems[381.Similarly, the Candida rugosa system has been used by Kanegafuchi to produce (R)-2-hydroxyisobutyric acid (33),an intermediate for the synthesis of the ACE inhibitor Captopril from the starting compound isobutync acid (32)r4, 38, 391. The production of L-carnitine by Lonza is also camed out by whole cells that make use of CoA-coupled fatty acid degradation or the p-oxidation pathway. L-Carnitine (38),or (R)-3-hydroxy-4-trimethylaminobutyric acid, serves as a fatty acid carrier and plays an important role in the metabolism of fats. In addition to clinical applications, such as for the treatment of disorders in fat metabolism, it is also a popular over-thecounter product in fitness and anti-obesity formulations. Lonza carries out the production of L-carnitineon a multi-tonne scale, in a whole-cellprocess. The wholecell process utilizes intact Agrobacteriurn cells that are fed with glucose and 4-butyrobetaine (34)as a precursor. The key enzyme that catalyzes the addition of water to crotonobetaine is L-carnitine dehydratase (crotonobetainyl-CoAhydratase, E. C. 4.2.1.89),which adds a water molecule to the fermentation product, crotonobetainyl-CoA (35,see Scheme 11.56).The resulting product, L-carnityl-CoA (36)is not oxidized to the corresponding B-keto acid 37, since the cells lack the enzyme

7 1.5 Addition of Water to C=C Bonds

I

695

carnitine dehydrogenase. Instead, the CoA coenzyme is hydrolyzed off by a thioesterase resulting in the release of L-carnitine (38) in the medium[40]. Despite the fact that numerous enzymes have been characterized that catalyze the addition of water to unsaturated fatty acids that are coupled to CoA or ACP, such as methylglucatonyl-CoA hydratase (E. C. 4.2.1.18), lactoyl-CoA dehydratase (E. C. 4.2.1.54), 3-hydroxybutyryl-CoA dehydratase (E. C. 4.2.1.55), itaconyl-CoA dehydratase (E. C. 4.2.1.56),isohexenylglutaconyl-CoA hydratase (E. C. 4.2.1.57),farnesylCoA dehydratase (E. C. 4.2.1.57), long-chain enoyl-CoA hydratase (E. C. 4.2.1.74), 3-hydroxydecanoyl-ACP dehydratase (E. C. 4.2.1.60) and 3-hydroxypalmitoyl-ACP dehydratase (E. C. 4.2.1.61), these enzymes are seldomly applied in organic synthesis. An unusual coenzyme A-coupled hydratase reaction occurs during the anaerobic degradation of benzoic acid by Thauera aromatica, where cyclohexa-1,5,-diene1-carboxylate CoA hydratase (E.C. 4.2.1.100) adds a water molecule to the cyclohexadiene functionality, resulting in the formation of 6-hydroxycyclohex-1-enecarbonyl COAI4l1. 11.5.6.2

Hydratases Acting on Free Fatty Acids

The enzyme oleate hydratase (E. C. 4.2.1.53) from Pseudomonas catalyzes the elimination ofwater from (R)-10-hydroxystearateor the addition ofwater to a number of free unsaturated fatty acids, yielding (R)-10-hydroxyfatty acids. Substrates that have been identified include linoleic acid, oleic acid and palmitoleic acid, which are converted to the corresponding 10-hydroxy-fattyacids [421. 11.5.7

Addition o f Water to Steroids

Hydratation of unsaturated carbon-carbon bonds in the steroid nucleus has been ascribed to hydrolyases such as 5-a-hydroxysteroiddehydratase (E. C. 4.2.1.62) from Saccharomyces cerevisiae, which catalyzes the interconversion of 5-a-ergostaSimilarly, the formation of 16-a-hydro7,22-diene-3+-5-diol and progesterone from 16-dehydroprogesteroneis catalyzed by 16-dehydroprogesterone dehydratase. The same enzyme catalyzes the addition of water to 16-a-hydroxyprogesterone or 16-a-hydroxy-pregnenolone, yielding the corresponding 16,17-didehydroprogesterone and 16,17-didehydropregnenolone (E. C. 4.2.1.86 or E. C. 4.2.1.98 La]).

696

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1 1 Hydrolysis and Formation of C-0 Bonds

References

E. J. Hughes, R. C. Bayly, R. A. Skurray, J. Batted. 1984, 158, 79-83. 2 R. Burlingame, P. J. Chapman,J. Bacteriol. 1983,155,426426. 3 S. K. Reaney, C. Begg, S. J. Bungard, J. R. Guest, J . Gen. Microbiol. 1993, 139, 1523-1530. 4 A. Liese, K. Seelbach, C. Wandrey, Industrial Biotransfonnations, Wiley-VCH Verlag GmbH, Weinheim 2000. 5 J. S. Britten, H. Morell, J. V. Taggart, Biochim. Biophys. Acta 1969, 185, 220-227. 6 M. J. van der Werf, W. J. van den Tweel, S. Hartmans, Eur. J. Biochem. 1993,217, 1011-1017. 7 B. He, T. Nakajima-Kambe,T. Ozawa, T. Nakahara, Process Biochem. 2000,36, 47-414. 8 M. J. van der Werf, W. J. van den Tweel, S. Hartmans, Appl. Environ. Microbiol. 1992, 58,2854-2860. 9 C. C. Wang, H. A. Barker, J. Biol. Chem. 1969,244,2527-2538. 10 Y. Oda, S.Suzuki, H. Katsuki, Biochem. Int. 1987, 14,871-878. 11 Y. Kato, Y. Asano, Arch. Microbiol. 1997, 168, 457-463. 12 S. R. Gross, H. E. Umbarger, Biochemistry 1963,2,1046-1052. 13 G. Hartrampf, W. Buckel, Eur. J . Biochem. 1986, 156,301-304. 14 P.K. Agrawal, G. K. Garg, K. G. Gollakota, Biochem. Biophys. Res. Commun. 1976,70, 979-986. 15 N. E. Neilson, Biochim. Biophys. Acta 1955, 17, 140. 16 G. Weidner, B. Steffan, A. A. Brakhage, Mol. Gen. Genet. 1997,255, 237-247. 17 T. Tabuchi, H. Aoki, H. Uchiyama, T. Nakahara, Agric. Biol. Chem. 1981,45, 2823-2929. 18 J. V. Schloss, M. H. Emptage, W. W. Cleland, Biochemistry 1984,23,4572-4580. 19 K. Maruyama, Biochem. Biophys. Res. Commun., 1985, 128,271-277. 20 E. W. Yamada, W. B. Jakoby,/. Biol. Chem. 1959,234,941-945. 21 E. W. Yamada, W. B. jakoby,J. Biol. Chem. 1958,233,706-711. 22 H. P. Meloche, W. A. Wood, J . Biol. Chem. 1964,239,3505-3510. 1

23 R. Weinberg, M. Doudoroff,/. Biol. Chem.

1955,217,607-624.

24 D. R. J. Palmer, S. J. Wieczorek, B. K. Hub-

bard, G. T. Mrachko, J. A. Gerlt, J. Am. Chem. Soc. 1997,119,9850-9581. 25 J. Robert-Baudouy,J. Jimeno-Abendano, F. Stoeber, Methods Enzymol. 1982,90 288-294. 26 M. J. Dilworth, R. Arwas, J. A. McKay, S. Saroso, A. R. Glenn, J. Gen. Microbiol. 1986,132,2733-2742, 27 E. L. 0.O’Connell, H. P. Meloche, Methods Enzymol.1982,89,98-101. 28 R. Bender, G. Gottschalk, Anal. Biochem. 1974,61,275-279. 29 A. S . Dahms, R. L. Anderson,]. Biol. Chem. 1972,247,2233-2237. 30 R. Yuen, H. Schachter, Can.J. Biochem. 1972,50, 798-806. 31 A. S . D. A. Dahms, Methods Enzymol.1982, 90 302-305. 32 B. K. Hubbard, M. Koch, D. R. J. Palmer, P. C. Babbitt, J. A. Gerlt, Biochemistry 1998, 37,14369-14375. 33 A.M. Gulick, D.R. j. Palmer, P. C. Babbitt, J.A. Gerlt, I. Rayment, Biochemistry 1998, 37,14358-14368. 34 M. C.Pinung, C. P. Holmes, D. M. Horowitz, D. S. Nunn,]. Am. Chem. Soc. 1991, 113,1020-1025. 35 H. K. Dannelly, H. C. Reeves, Cur. Microbiol. 1988, 17, 265-268. 36 K. W. Moore, J. H. Richards, Biochem. Biophys. Res. Commun. 1979,87, 10521057. 37 R. Daniel, T. A. Bobik, G. Gottschalk, FEMS Microbiol. Rev. 1999,22, 553-566. 38 R. A. Sheldon, Chirotechnology, Marcel Dekker Inc., New York 1993. 39 J. Hasegawa, M.Ogura, H. Kanema, N. Noda, H. Kawaharada, K. Watanabe, /. Ferment. Echnol. 1982,60, 501-508. 40 Th. P. Zimmermann, K. T. Robins, J. Werlen, F. H. Hoeks, in: Chirality i n Industry. A. N. Collins, Sheldrake G. N., J. Crosby (eds), John Wiley and Sons, New York, 1997, pp. 287-305. 41 Hanvood CS, Gibson J,J. Bacteriol. 1997, 179,301-309. 42 A. Kisic, Y. Miura, G. J. Schroepfer, Lipids 1971, 6, 545.

References I697 43

R.W. Topham, J. L. Gaylor, Biochem. Biophys. Res. Commun.1972,47,180-1%.

44 T.L. B. C. 2. Glass, J. Steroid Biochem. 1984, 21,65-72.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

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12 Hydrolysis and Formation of C-N Bonds

12.1

Hydrolysis of Nitriles

Birgit Schulze 12.1.1

Introduction

Organic nitriles are used extensively industrially as precursors for the production of a wide variety of amides and acids by chemical synthesis. In recent years, considerable attention has been paid to enzymatic hydrolysis of nitriles as an alternative route to the chemical synthesis of amides and carboxylic acids. Conventional chemical conversion of nitriles suffers from several disadvantages,including the requirement for highly acidic or basic reaction conditions, high energy consumption, formation of undesirable by-products, low yields and environmental problems due to the generation of waste salts. Biocatalysis, on the other hand, may be performed under mild conditions (low temperatures, neutral pH) thus affording high conversion yields and selective hydrolysis of the -CN functionality of compounds containing acid or base labile groups. Furthermore, enzyme hydrolysis is, in some instances, enantio- and/or regioselective. This chapter does not aim to give a complete treatise on the extensive literature on nitrile bioconversions but rather aims at presenting a brief overview of enzymatic nitrile hydrolysis with a selection of examples. Several reviews on the bioconversion of organic nitriles and its potential technological application have been published [1-71.

700

I

72 Hydrolysis and Formation ofC-N Bonds

12.1.2 Types of Nitrile Hydrolyzing Enzymes 12.1.2.1

Enzymatic Hydrolysis of Organic Nitriles

A variety of organic nitriles, such as cyanoglycosides and cyanolipids, occur naturally in biological materialrl, 1' . It has been shown that nitrile-hydrolyzing activity is widespread in bacteria and fungi but has recently also been identified in insects and humans 1('. In microorganisms the hydrolysis of organo-nitriles (hereafter denoted as nitriles) is effected by two distinct enzymes, nitrilases or nitrile hydratases. Nitrilase (E. C. 3.5.5.1.) catalyzes the hydrolysis of nitriles to the corresponding acids and ammonia in a one step reaction without the formation of a free amide intermediate:

Scheme 12.1-1.

whereas nitrile hydratase (E. C. 4.2.1.84) catalyzes the hydrolysis of nitriles to the corresponding amides:

0

Scheme 12.12.

Microorganisms which produce a nitrile hydratase also seem to synthesize one or more amidase enzymes [linear amide hydrolase (E. C. 3.5.1)] thus enabling them to hydrolyze nitriles to the corresponding acids in a two-step reaction: 0 R-cN

+

H20

nitrile hydratase)

0

R -,K,, lN am?

+

NH3

Hi0

Scheme 12.1-3.

Nitrilases and nitrile hydratases are distinct enzymes, apparently differing both with respect to prosthetic groups and reaction mechanisms.

12.1.2.1.1 Nitrile Hydratases

In recent years nitrile hydratases have been studied intensively. Enzymes from a wide range of microorganisms have been isolated and characterized and the corresponding genes have been cloned and overexpressed. Nitrile hydratases can be subdivided into two classes: enzymes requiring Fe3' (for example: from Rhodococcus sp. 19-121, Pseudomonas chlororaphis B23 [13] and Cornamonas testosteroni NII [I4]) and

12.1 Hydrolysis of Nitriles

OH@

Figure 12.1-1. Two o f the mechanisms discussed for nitrile hydratase c a t a l y s i ~ [ ~ ~ - ~ ~ ] . (I) Direct coordination o f t h e substrate t o the metal ion in the active site. (11) Attack o f

a water molecule activated by a metal-bound hydroxide ion.

enzymes requiring Co3+(for example: Rhodococcus rhodocrous J 1[15,1G1 and Pseudomonasputida NRRL 18GG8[171)for catalysis. The metal clusters of the former group have been studied intensely by EPR, ENDOR, EXAFS and other spectrophotometric techniques, revealing a unique coordination of a non-heme iron cluster [18-241. Nitrile hydratases show a high degree of homology. They consist of from two up to twenty subunits L2] containing one metal ion per a,p-dimer with the only exception described to date being the nitrile hydratase from P.chlororaphis which appears to be a hom~tetramer[~~l. The first X-ray structure of a nitrile hydratase was determined for the enzyme from Rhodococcus sp. R312[25].In this study an a&-tetramer conformation has been found in the native enzyme. Characterization of the highly related Rhodococcus sp. N771 nitrile hydratase, however, revealed a dimeric species in solution for this enzyme[l21. Based on the X-ray structure mechanisms have been proposed which are depicted in Fig. 12.1-1 [25-271. Earlier studies suggested the involvement of the cofactor pyrroloquinoline quinone (PQQ) X-ray studies and spectral characterization [24, 251 have, however, recently discarded this.

12.1.2.1.2

Nitrilases

Nitrilases have been studied less than the nitrile hydratases. The enzymes appear as homomultimers, exhibiting a wide range of molecular masses. The reaction mechanism depicted in Fig. 12.1-2 has been proposed recently by Kobayashi et al.[281. Several nitrilases have been found to be inhibited by reagents which bind to thiol groups, indicating that sulfhydryl groups are essential for the catalytic activity of

I

701

702

I

72 Hydrolysis and Formation of C-N Bonds

R-CGN Enz-SH

0

R-C\

4

OH Figure 12.1-2.

Reaction mechanism proposed for nitrilase catalysisIz8I.

these enzymes. This has been confirmed by the work of Kobayashi et al. who cloned the nitrilase gene from Rhodococcus rhodocrous J 1 and have proved that Cys 165 is crucial for nitrilase activity. The group also found that the nitrilase does catalyze the formation and hydrolysis of amides, although with lower activity[28]. All the nitrile hydrolyzing enzymes described so far are intracellular and differ considerably with respect to substrate specificity, stereoselectivity,molecular mass and substrate and product inhibition characteristics. 12.1.2.2 Enzymatic Hydrolysis of Cyanide

Microorganisms appear to have evolved separate metabolic pathways for the hydrolysis of inorganic cyanide. Thus most nitrilases and nitrile hydratases investigated so far do not display activity towards cyanide. However recently, as a first exception to this, a nitrilase has been reported that hydrolyses potassium cyanide as well as organic nitriles 12’* 301. Enzymes catalyzing the hydrolysis of cyanide had already been identified earlier. Here too, two different types can be distinguished. The enzyme cyanide hydratase [formamide hydrolase (E. C. 4.2.1.66)] can be found in various fungi[31].It catalyzes the hydration of cyanide to formamide: 0

Scheme 12.1-4.

A direct hydrolytic activity on cyanide, yielding formate and ammonia without forming formamide as an intermediate, has been identified in different bactefia[32,331.

-,AoH 0

HCN

+

2H20

Scheme 12.1-5.

cyanidase

+

NH3

72.1 Hydrolysis ofNitriles

This new type of enzyme, tentatively named ‘cyanidase’,has also recently been cloned, overexpressed and characterizedP4*351. Cyanidase tolerates high cyanide concentrations and is able to deplete cyanide in aqueous solutions to less than 0.01 PPm. Cyanide hydrolyzing enzymes (i.e. reaction schemes 12.1-4and 12.1-5) have, with the exception mentioned above, not been reported to hydrolyze organic nitriles and thus appear to be highly specific for inorganic cyanide. Therefore, they are mainly of interest in waste water treatment as a biological alternative to conventional chemical detoxification of cyanide by alkaline chlorination and will not be treated further. Comprehensive reviews on biological cyanide transformations may be found in the literature [36, 371. Reviews on microbial cyanide metabolism have been published by Knowles [38, 391 and Knowles and Bunch[40]. 12.1.3

Examples of Enzymatic Nitrile Hydrolysis 12.1.3.1

Enantioselective Hydrolysis of Nitriles

Several groups of investigators have reported on the production of optically active acids from racemic mixtures of nitriles. A comprehensive overview has recently been presented I4l]. Fukuda and coworkers have described one of the first applications of the enantioselective hydrolysis of nitriles (Scheme 12.14). Using a whole cell biocatalyst optically pure a-hydroxy acids (1-a-hydroxyisovalericacid and L-a-hydroxyisocaproic acid) have been prepared from the racemates of the corresponding a-hydroxynitriles [421.

Scheme 12.1-6.

In recent years, the enantioselectivehydrolysis of nitriles has been studied in more detail. Whereas in the past only whole cell catalysts had been investigated, it is now possible to assign the activities to specific enzymes occurring in the cell. These enzymes are nitrilases, nitrile hydratases and/or amidases.

12.1.3.1.1

Enantioselective Nitrilases

First indications of stereoselective nitrilases have been given by Macadam and Knowles describing the production of L-alanine from racemic a-aminopropionitrile by a stereoselective nitrilase produced by an Acinetobacter sp. [43j. Bhalla et al. [441 reported the stereoselective hydrolysis of a-aminonitriles by a

I

703

704

I

12 Hydrolysis and Formation ofC-N Bonds

nitrilase from Rhodococcus rhodochrous PA-34. The nitrilase exhibited relaxed substrate specificity hydrolyzing both mono- and dinitriles and its stereospecificity differs from that of the Acinetobacter sp. since it produced D-alanine from D,L-c~aminopropionitrile. Enzymatic production of (S)-(+)-ibuprofen,an anti-inflammatory drug, from racemic 2-(4'-isobutylpheny1)propionitrileusing various bacterial strains has been published by Yamamoto et al. (Scheme 12.1-7)[45. 461. One of the strains, Acinetobacter sp. AK226 produced (S)-(+)-ibuprofenby means of a nitrilase in whole cell experiments. High optical purity of the product (about 95 % ee) was obtained at a low percentage of hydrolysis but the optical purity of the product decreased at the later stages of the reaction. This was in accordance with other results which showed that (R)-(-)-Ibu-CNwas indeed also hydrolyzed by the nitrilase, although at a 108-fold lower rate than the (S)-nitrile. Also Rhodococcus butanica ATCC 21 197 has been shown to produce a nitrilase suitable for the enantioselective hydrolysis of racemic 2-aryl-propionitriles;however, selectivitieswere rather low [471.

Scheme 12.1-7.

An enantioselectivenitrilase has also been shown to be applicable in the dynamic kinetic resolution of mandelonitrile. Using the nitrilase produced by Alcaligenes faecalis ATCC 8750 Yamamoto et al. showed that they could derive (R)-(-)-mandelic acid from mandelonitrile in 91 % yield with an ee of 100%. Under the reaction conditions used non-reacting (S)-mandelonitrile undergoes spontaneous racemization leading to the high yield (see Scheme 12.1-8)L4'I. Currently (R)-mandelic acid and (R)-chloromandelicacid are produced using nitrilases on an industrial scale by the Mitsubishi Rayon Corp.

Scheme 12.1-8.

12.1.3.1.2 Nitrile Hydratases

Enantioselective hydrolysis of nitriles by the nitrile hydratase/amidase system has often been attributed to the combination of a non-selective nitrile hydratase and a selective amidase. However, more recently several enantioselectivenitrile hydratases have also been identified and studied in detail. For example, a nitrile hydratase from

12.7 Hydrolysis ofNitriles

I

705

Pseudomonas putida NRRL-18668 has been found to hydrate 2-(4-chlorophenyl)3-methylbutyronitrilewith high enantioselectivity (Scheme 12.1-9).The whole cell preparation was found to transform the (S)-enantiomer preferentially until approximately 90 % has been consumed. Successively the (R)-enantiomer also reacted but at a 6-fold lower ratel4’]. Using a purified enzyme preparation on the enantiomerically pure substrates the reaction rate towards the (S)-enantiomer has been determined to be more than 50 times higher than for the (R)-enantiomer[”l. ~ l Pichia Recently the enzyme has also been cloned and overexpessed in E . ~ o l i [ ~and pa~toris[~~]. The whole cells from Pseudomonas putida NRRL-18668 have also been investigated in the hydrolysis of a-substituted arylpropionitiriles. A stereoselective hydrolysis was found. However, in these cases the enantioselectivity must be attributed to the combined reactions of the hydratase and the amida~e[~’]]. This is also the case for several other whole cell catalysts containing the hydrataselamidase system. Hence the enantioselectivehydrolysis of a-substituted phenylacetonitrilesby Rhodococcus sp. AJ 270[52]and by Rhodococcus sp. (SP361)[531 is based on the highly (S)-selective amidases. However, evidence for enantioselective hydratases has also been provided by studying the transformations using a whole cell catalyst in the presence of amidase inhibitors. Thus, the hydratase of Rhodococcus equi exhibits a preference for (S)-nitriles whilst the hydratase from Agrobacterium tumefaciens is selective towards the (R)-nitriles[54, 551.

Scheme 12.1-9.

12.1.3.2

Monohydrolysis o f Dinitriles

Several investigators using enzymes of different microbial origin have reported on the monohydrolysis of dinitriles to the corresponding cyano-carboxylic acids in high yields. Selective hydrolysis of one cyano group of a dinitrile is very difficult to carry out by chemical hydrolysis and is, therefore, an intriguing aspect of nitrile bioconversion. Using whole cells of Rhodococcus rodochrous NCIB 11216 Bergis-Garber and Gutman demonstrated complete monohydrolysis of fumaronitrile and almost complete monohydrolysis of succinonitrile, 1,3-dicyanobenzeneand 1,4-dicyanobenzene (Scheme 12.1-10,12.1-11) rS6, ’1. Analysis of the reaction products showed that during conversion of succinonitrile into 3-cyanopropionic acid, succinamic acid (H~NOC-CH~CHZ-COOH) was detected as a free intermediate in the reaction mixture, suggesting that enzymatic activities other than nitrilases (i.e. nitrile hydratase) were present in the cell

706

I

-

72 Hydrolysis and Formation ofC-N Bonds

Nc*CN

0

Scheme 12.1-10.

CN

CN

Scheme 12.1-11.

preparation. In parallel experiments with the same bacterial strain glutaro-, adipoand pimelodinitriles were hydrolyzed further to glutaric, adipic and pimelic acid, respectively rS71. Kobayashi et al. ['*I on the other hand obtained complete conversion of glutarodinitrile into cyanobutyric acid without any formation of glutaric acid using a nitrilase from another strain of Rhodococcus (Rodococcus rodochrous K22). Turner et al. [591 have studied the hydrolysis of aromatic dinitriles using an immobilized preparation of Rhodococcus sp. (Novo SP 361). Only after esterification of the carboxylic acid formed was the second nitrile group hydrolyzed by repetitive use of the catalyst (Scheme 12.1-12).

g g 0

CN sp361_

OH

CN

CN

OH

0

Scheme 12.1-12.

The chemoselecivity of nitrile hydrolyzing enzymes has also been used by Tani and coworkers to examine the formation of trans-4-cyanocyclohexane-1-carboxylic acid (t-MCC) from trans-1,4-dicyanocyclohexane (t-DCC) using a resting cell system of Corynebacteriurn sp. C5 (Scheme 12.1-13)[601.The reaction was shown to be

12.7 Hydrolysis ofNitriles

I

707

catalyzed by the sequential action of a nitrile hydratase and an amidase, both of which were purified and characterizedIG1]. The nitrile hydratase exclusively hydrolyzed truns-l,4-dicyanocyclohexane (t-DCC) into trans-4-cyanocyclohexane-1-carboxyamide (t-MCMA), which could be detected in the reaction mixture[G1].The nitrile hydratase did not attack the nitrile groups in t-MCC and t-MCMA. This conversion has also been described using the nitrile hydratase from Rhodococcus rhodochrous AJ270 giving a 99% yield[62s63]. nitrile

H&rz

amidase

H

a

H

hydratase)

CN

CN

CN

Scheme 12.1-13.

In addition to the outstanding chemoselectivity,high regioselectivity can also be found in the hydrolysis of dinitriles and has been used for example in the chemoenzymatic production of lactams from aliphatic a,w-dinitriles. Using a nitrilase from Acidovoraxfacilis or a nitrile hydrataselamidase system from Comamonus testosteroni high yields of the lactams have been achieved (see Scheme 12.1-14)cG4].

Scheme 12.1-14.

The chemo- and enantioselectivity of nitrile-hydrolyzing enzymes gives rise to products of high optical purity with a theoretical yield of 100%, when starting from prochiral substrates. Thus several whole cell catalysts have been studied in the conversion of 3-substituted glutaronitriles. The enantioselectivity was found to be highly dependent on the substituent at the 3-position (Scheme 12.1-15, Scheme 12.1-16) 651. Using this technology (R)4-hydroxy-S-cyanopentene,a precursor for the protected lactone moiety of the 6-lactone pharmacophore of the mevinic acids, becomes available in high yields [GG]. Recently, an industrial application of the chemoselective hydration of a dinitrile, adiponitrile, has been introduced. A Pseudomonas chlororuphis B23 nitrile hydratase N

C

A

C

N

SP361

9 NC-COOH R= OBn OBz OH OMEM' OAc

e.e. = 83% e.e. = 84% e.e. = 22% e.e. = 61% e.e. = 0

73% yield 25% yield 52% yield 19% yield 45%yield

*MEM = methoxyethoxymethyl

Scheme 12.1-15.

708

I

72 Hydrolysis and formation ofC-N Bonds

R= Bn OBn OBz

e.e. = 29% e.e. = 90% e.e. = > 99%

59% yield 68% yield 71% yield

Scheme 12.1-16.

immobilized in alginate beads is used for the production of 5-cyanovaleramide.The biocatalyst is extremely stable and has been used in almost GO consecutive batches producing more than 13 metric tons in the production of the precursor of a new herbicide 12.1.3.3

Substrate and Product inhibition of Nitrile Hydrolysis

Substrate and/or product inhibition may seriously reduce the productivity of nitrilehydrolyzing enzymes. Already nitrile concentrations higher than 200-500 mM have been reported to be inhibitory, often causing rapid and irreversibleinactivation of the biocatalyst[Gg-741. Substrate inhibition may be overcome by running the enzymatic reaction constantly at a low substrate concentration using periodic or continuous feeding of the substrate. Product inhibition/inactivation, on the other hand, is considerably more difficult to tackle in a large scale industrial process and may prevent implementation of enzymatic hydrolysis for a particular reaction. Thus it appears that the success of the commercial acrylamide process of the Mitsubishi Rayon Corp. (the former Nitto Corp.) is the result of extensive and elegant efforts within the areas of process optimization and the development of improved biocatalysts which are less susceptible to product inhibition. Currently the acrylamide production is run optimally using a highly efficient nitrile hydratase catalyst at low temperature (5-10 "C) thereby avoiding substrate inhibition which occurs at higher 751. For details see Sect. 12.1.3.5. The same whole cell catalyst can be used in the hydration of 3-cyanopyridine to nicotinamide (Scheme 12.1-17).This vitamin, broadly applied in animal feeding, is currently produced biocatalybcally on an industrial scale (> 3000 t/a) by the Lonza AG. For this substrate Yamada and Kobayashi showed that the whole cell catalyst of Rhodococcus rhodocrous J 1, containing a nitrile hydratase induced with crotonamide, can even tolerate substrate concentrations up to 12 M [ ~(see ] Fig. 12.1-3). Mauger et al. also succeeded in achieving high final product concentrations of various amides when using the Rhodococcus rhodochrous J1catalyst (see Table 12.1-1).

Scheme 12.1-17.

12.1 Hydrolysis ofNitriles

I g -

90

.O

80

C

(? 0

I

after 5(*), 9 (B) and 22 h (A)of incubation at various substrate concentrations.

70 1 60 7

9

11

13

15

substrate concentration [MI Table 12.1-1.

Nitriles hydrolyzed by Rhodococcus rhodocrous J l 1’. 761.

Substrate

Amide

3-Cyanopyridine 4-Cyanopyridine 2,G-Difluorobenzonitrile 2-Cyanopyrazine 2-Cyanopyridine 2-Cyanothiophen 3-Indolylacetonitrile Benzonitrile 2-Cyanofuran

nicotinamide isonicotinamde 2,G-difluorobezmide pyrazinamide picolinamide 2-thiophencarboxamide indole-3-acetamide benzamide furanecarboxamide

I

709

Figure 12.1-3. Conversion o f 3-cyanopyridine

Product concentration

(g L-7

1465 1099 30G 985 977 210 1045 489 522

The hydrations were carried out either at low substrate concentrations with slow feeding of the substrate (for example: benzonitrile, 2,G-difluorobenzonitrile and 3-indoleacetonitrile)or, in the case of less toxic substrates, by direct incubation at high substrate concentrations (for example: 3-indolylacetonitrile and 2-cyanopyrazine[2. 7611. In addition, high substrate levels have been used in the industrial production of 5-cyanovaleramide(see Sect. 12.1.3.2)using the nitrile hydratase from Pseudomonas chlororaphis B23. Starting at a substrate concentration of 1.5 M, high above the solubility level (0.45 M). The hydration was carried out in a two phase system. The nitrile hydratase showed outstanding stability at these high substrate concentrations. Sequential addition of the substrate, instead of starting at a high concentration, only slightly improved the stability. Increased stability could be achieved by the addition of butyric acid to the medium. However, the higher stability has to be traded off with a lower activity caused by the inhibition of the nitrile hydratase by butyrate (see Sect. 12.1.3.4).

710

I

72 Hydrolysis and Formation ofC-N Bonds

12.1.3.4 Activation and Stabilization of Nitrile Hydratases

Iron-dependent nitrile hydratases, for example from Rhodococcus R3 12 or Pseudomonus chlororuphis, exhibit a remarkable dependency on light. The enzymes, after being inactivated by aerobic incubation in the dark, regain their activity when exposed to light irradiation[12,771. Using different spectrophotometric techniques (ENDOR, EXAFS, FTIR, UV-VIS and X-ray) this phenomenon has been studied extensively in recent years. It has now been confirmed that the deactivation is caused by the reversible binding of nitric oxide to the non-heme iron center in a 1 :1 stoichiometric complex. Upon irradiation the complex is destroyed and the activity of the nitrile hydratase is restored[‘4, 27, 78-811. Another interesting characteristic of the iron-dependent nitrile hydratases is their stabilization during purification and storage by alcanoic acids such as butyric acid, hexanoic acid and valeric acid. The effect has already been described by Nagasawa et al. in 1987[13].However, only in recent years has the role of the acids been clarified by spectroscopic studies. Studying the EPR signals of the nitrile hydratase from Brevibucterium R312, Kopf et al. showed that butyric acid interacts with the iron in the active site of the nitrile hydratase, stabilizing the enzyme but, at the same time acting as a competitive inhibitor[82]. 12.1.3.5 Nitrile Hydrolysis in Organic Solvents

Most nitrile bioconversions published have been conducted in aqueous media and consequently few data are available on the effect of solvents on enzymatic nitrile hydrolysis. Such studies seem highly justified in order to investigate the effects of different solvents or co-solvents on substrate specificity, conversion rate, stereoselectivity, and catalyst half-life.

‘w

Rhodococcussp. NCIMB 12218

.

Scheme 12.1-18.

De Raadt et al. reported on the inhibition of nitrile hydrolysis by various solvents [831. However, production of 2,G-difluorobenzamide(Scheme 12.1-18)was effected in 99.5 % n-heptane using the nitrile hydratase from Rhodococcus sp. NCIMB 12 218[84]. The enzymatic reaction was found to be activated by light (see 12.1.3.4). More recently, Layh and Willetts have studied nitrile transformations in various organic solvents and biphasic mixtures using a nitrilase from Pseudowonus sp. DSM 11 387 and a nitrile hydratase from Rhodococcus sp. DSM 11 397[”]. The enzymes exhibited good stabilities in biphasic mixtures with hydrophobic solvents when dispersed in

12.1 Hydrolysis ofhlitriles

the buffer-saturated higher alcohols 1-hexanol,1-heptanol, 1-octanoland 1-decanol, respectively. The nitrilase still retained 58 %, 49 %, 44% and 47 % activity, while the nitrile hydratase only showed low activities (2-5 %). 12.1.3.6

Large Scale Production o f Acrylamide

Acrylamide monomer is an important chemical commodity produced on a multihundred thousand ton scale for the production of polymers and copolymers. The preferred manufacturing process is by the catalytic hydration of acrylonitrile at 70-120 "C using reduced Raney copper as the catalyst; the initial concentration of acrylonitrile being around 4 M. There are several shortcomings to this process, among which are the high level of acrylic acid formed and byproduct formationI2. '1. An enzymatic acrylonitrile hydration was first patented in 1981["I. Many nitrile hydratases of different origin have been shown to be able to convert acrylonitrile into acrylamide. However, a major problem associated with biocatalysis for production of acrylonitrile is the short half-life of the enzyme due to substrate and product inhibition. Acrylonitrile is a strong alkylating agent which reacts by Michael addition with the sulfhydryl groups of proteins[", 691. The Mitsubishi Rayon Corp. (the former Nitto Chemical Industry Co.) established the industrial production of acrylamide in 1985 using immobilized cells of Rhodococcus sp. N-774L3.". 741. In 1988 a hyperproducing mutant strain of Pseudornonas chlororaphis B23 was chosen for production. As in Rhodococcus sp. N-774, the active Table 12.1-2.

Operating conditions for acrylamide production.

Reaction conditions

Productivity

pH 7.5-8.5 Temperature 0-5 "C Acrylonitrile concentration in the reactor 1.5-2.0%

Conversion acrylonitrile Yield of acrylamide Acrylamide concentration from the reactor

Table 12.1-3.

> 99.9% > 99.9% 27-30%

Comparison of enzyme data ofthree types of nitrile hydratases.

Parameter

Tolerance to acrylamide (%) Acrylic acid formation Cultivation time (h) Activity of culture broth (units mL-I) Specific activity (units per mg cells) Cell yield (g L-') Acrylamide productivity (g per g cells) Total amount of production (t per year) Final concentration of acrylamide ("h) First year of production scale

Rhodococcus sp.

N-774

Pseudomonas chlororaphis 823

Rhodoccus rhodochrous J l

27 very little 48 900 60 15 500 4000 20 1985

40 barely detected 45 1400 85 17 850 6000 27 1988

50 barely detected 72 2100 76 28 > 7000 30.000 40 1991

I

711

712

I

12 Hydrolysis and Formation of C-N Bonds Table 12.1-4.

Comaprison of enzyme data ofthree types of nitrile hydratases. Rhodoccus sp.

Parameter

N-774

Molecular mass Subunit molecular mass

70.000

Pseudomonas chlororaphis 823

Rhodoccus rhodochrous I1 505.000

100.000

a 27.000

a 25.000

Fe"'

Fe"'

co

p 27.500

a 26.000

p 25.000

p 25.000

Metal Optimum temperature ("C) Heat stability ("C) Optimum pH pH stability Substrate specificity

35 30 7.7 7.0-8.5

20 20 7.5 6.0-7.5

35-40 50 6.5 6.0-8.5

aliphatic nitriles

aliphatic nitriles

Activation by light irradiation Formation type

+

aliphatic and aromatic nitriles

-

-

constitutive

inducible (methacrylamide)

inducible (urea)

Biocatalyticalprocess immobilization of microorganism

1 Acrylonitrile

Water

iLI Spent catalyst

Cu-catalyticprocess

Acrylonitrile

Water

Figure 12.1-4. Comparison of the biocatalytic and the conventional chemical process for acrylamide production.

biocatalyst in Pseudomonas chlororaphis B23 is also a nitrile hydratase containing ferric ion as the cofactor[2s741. Current acrylamide production at Mitsubishi using bioconversion is around 40 000 tonnes per year. Using a highly improved cobalt-containingnitrile hydratase from Rhodococcus rhodochrous 71, final product concentrations of around 700 g L-'

References

can be obtained[87].The reaction is performed at 5-10 “C in order to reduce cell degradation and enzyme inhibition. No data have been published on the half-life of the Rhodococcus rhodochrous J1 nitrile hydratase under production conditions. A good summary of the biocatalyhc production of acrylamide has been given by Yamada and c o - ~ o r k e r s [871~ ~ , Tables 12.1-2 to 12.1-4). (see In Fig. 12.1-4 a comparison is presented of the enzymatic and the conventional chemical processes for acrylamide production. 12.1.4

Availability and Industrial Future o f Nitrile Hydrolyzing Biocatalysts

Although nitrile-hydrolyzingenzymes have attracted considerable interest as promising “green catalysts”,none of these enzymes are presently available as commercial products. Thus studies on nitrile biotransformations have been conducted with a variety of enzyme preparations ranging from resting cells, immobilized whole cells, cell-free extracts, immobilized enzymes and pure soluble enzymes. However, nowadays several nitrile hydratases and nitrilases have been cloned and overexpressed, giving rise to highly efficient and well defined catalysts 71. This not only provides commercial access to even more interesting catalysts, but also opens the way for the application of modern molecular biological methods for further optimization. Within the recent years several industrial processes based on nitrile hydrolyzing enzymes have been introduced, as has been discussed above. This number is now expected to increase rapidly, due to the better availability of these biocatalysts. 12 ‘

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12 Hydrolysis and Formation ofC-N Bonds

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Meth-Cohn, J. Colby, Tetrahedron: Asymmetry2000, 11, 1123-1135. 53 T. Beard, M. A. Cohen, J. S. Parratt, N. J. Turner, Tetrahedron: Asymmetry1993,4, 1085-1104. 54 L. Martinkova, A. Stolz, H.-J. Knackmuss, Biotechnol. Lett. 1996, 18, 1073-1076. 55 R. Bauer, H.-J. Knackmuss, A. Stolz, Appl. Microbiol. Biotechnol. 1998,49, 89-95. 56 C. Bengis-Garber,A. L. Gutman, Tetrahedron Lett. 1988, 29, 2589-2590. 57 C. Bengis-Garber,A. L. Gutman, Appl. Microbiol. Biotechnol. 1989, 32, 11-16. 58 M. Kobayashi, N. Yanake, T. Nagasawa, H. Yarnada, Tetrahedron Lett. 1990,4G, 5587-5590. 59 J. Crosby, J. Moilliet, J. S. Parratt, N. J. Turner, J . Chem. Soc. Perkin Trans. I 1994, 1679-1687. 60 Y. Tani, M, Kurihara, H. Nishise, K. Yamamoto, Agnc. Bid. Chem. 1989,53, 3 143-3 149. 61 Y. Tani, M, Kurihara, H. Nishise, K. Yarnarnoto, Agnc. Biol Chem. 1989,53, 3151-3158. 62 0. Meth-Cohn, M-X.Wang,J. Chem. Soc., Chem. Commun. 1997,1041-1042. 63 0. Meth-Cohn, M.-X. Wang,J. Chem. Soc., Perkin Trans. 1 1997,21, 3197-3204. 64 J. E. Gavagan, S. K. Gager, R. D. Fallon, P. W. Folsorn, F. E. Herkes, A. Eisenberg, E. C. Hann, R. DiCosimo,J. Org. Chem. 1998,63,4792-4801. 65 H. Kakeya, N. Sakai, A. Sano, M. Yokoyarna, T. Sugai, H. Ohta, Chem. Lett. 1991, 1823-1824. 66 S. J. Maddrell, N. J. Turner, A. Kerridge, A. J. Willetts, J. Crosby Tetrahedron Lett. 1996,37,6001-6004. 67 E. C. Hann, A. Eisenberg, S. K. Fager, N. E. Perkins, F. G. Gallagher, S. M. Cooper, J. E. Gavagan, B. Stieglitz, S. M. Hennessey, R. DiCosimo, Bioorg. Med. Chem. 1999,7, 2239-2245. 68 K. Ingvorsen, B. Yde, S. E. Godtfredsen, R. T. Tsuchiya, Microbial Hydrolysis of Organic Nitriles and Amides, in: Cyanide Compounds in Biology. Ciba Foundation Symposium, (Eds.: D. Evered, S. Harnett) 1988, I40,16-31.

I. Watanabe, Y. Satoh, K. Enomoto, S. Seki, K. Sakashita, Agric. Biol. Chem. 1987, 51, 3201-3206 70 C. Y. Lee, Y. B. Hwang, H. N. Chang, Enzyme Microb. Technol. 1991, 13,53-58 71 J. Mauger, T. Nagasawa, H. Yarnada, Tetrahedron 1990,45, 1347-1354. 72 K. Bui, M. Maestracci, A. Thiery, A. Araud, P. Galzy, J. Appl. Bacterial. 1984, 57, 183-190. 73 T. Nagasawa, T. Nakamura, Y. Yamada, Appl. Microbiol. Biotechnol. 1990, 34, 322-324. 74 M. Kobayashi, T. Nagasawa, H. Yamada, TIBTECH 1992,10,402-408. 75 C. Y. Lee, S. K. Choi, H. N. Chang, Enzyme Microb. Technol. 1993, 15, 979-984. 76 J. Mauger, T. Nagasawa, H. Yarnada, J . Biotechnol. 1988,8,87-96. 77 T. Nagamune, H. Kurata, M. Hirata, J. Honda, A. Hirata, I. Endo, Photochem. Photobiol. 1990, 51, 87-90. 78 T. Noguchi, J. Honda, T. Nagarnune, H. Sasabe, Y. Inoue, I. Endo, FEBS Lett. 1995, 358,9-12. 79 T. Noguchi, M. Hoshino, M. Tsujirnura, M. Odaka, Y. Inoue, I. Endo, Biochemistry 1996,35,16777-16781. 80 R. C. Scarrow, B. S. Strickler,J. J. Ellison, S. C. Shoner, J. A. Kovacs, J. G. Cummings, M. J. Nelson,J. Am. Chem. Soc. 1998, 120, 9237-9245. 81 S. Nagashima, M. Nakasako, N. Dohmae, M. Tsujimura, K. Takio, M. Odaka, M. Yohda, N. Kamiya, I. Endo, Nature Struct. Bid. 1998, 5, 347-351. 82 M. A. Kopf, D. Bonnet, I. Artoud, D. Petre, D. Mansuy, Eur. J . Biochem. 1996,240, 239-244. 83 A. de Raadt, N. Klempier, K. Faber, H. Griengl, J . Chem. Soc., Perkin Trans. 1 1992, I , 137-140. 84 Shell and Gist Brocades, European Patent Application 1988,252 564. 85 N. Layh, A. Willetts Biotechnol. Lett. 1998, 20, 329-331. 86 I. Watanabe, Y. Soto, T. Takano, US Patent (1981) 4248968. 87 H. Yamada, Chimia 1993,47, 5-10. 69

I

715

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002 716

I

72 Hydrolysis and Formation ofC-N Bonds

12.2

Formation and Hydrolysis of Arnides Birgit Schulze and Erik de Vroom 12.2.1

Introduction

Organic amides and acids are versatile precursors to the production of various commercial products; nowadays these compounds are mainly produced chemically. However, owing to environmental considerations and the increasing demand for chiral amides and acids there is a strong tendency to explore and develop biocatalytic production processes. This section gives a global overview of the potential of microorganisms and enzymes to catalyze the regio- and enantioselective hydrolysis and formation of amides. The biocatalytic hydrolysis of amides and also the enzyme catalyzed formation of amides and the synthesis of (semisynthetic)antibiotics is included in this section. 12.2.2

Enzymatic Formation of Arnides

Currently two biocatalytic methods are known for the production of amides: The hydrolysis of nitriles using nitrile hydratases (for example: acrylamide production). The enzymatic ammoniolysis of carboxylic esters and amidation of carboxylic acids. In the previous Section 12.1 the formation of amides from the corresponding nitriles is well addressed. The latter method, the enzyme catalyzed reaction of carboxylic esters or acids with ammonia or amines yielding amides, has only recently been studied in depth. Encouraging results have been described, especially in the field of amidation of esters, a technology now being used by BASF to produce optically pure amines (vide infa). Lipases and esterases comprise a versatile group of enzymes that catalyze the hydrolysis of esters, esterifications and transesterifications via an acyl enzyme intermediate (Chapter 11). Various other nucleophiles can attack this acyl enzyme complex in addition to water. In recent years several authors have also shown that NH3 and amines can act as nucleophiles leading to the formation of amides 1' . Initially De Zoete et al. showed that fatty acid esters could be converted into the corresponding amides by bubbling gaseous NH3 through the reaction mixture containing lipase B from Candida antarctica (SP 435 from Novo-Nordisk) in tee-butyl alcohol. As shown in Scheme 12.2-1 very good yields can be achieved. This enzyme has also been used in related studies by other authors. A conclusive summary can be found in L31.

72.2 Formation and Hydrolysis of Amides

Conversion 100% 95% Octanarnide 5% Octanoicacid

0 L

O

E

t

SP435

Conversion 100% 95% Hexanarnide 5% Hexanoic acid

Scheme 12.2-1.

Further to this kinetic approach, the thermodynamic ammoniolysis has also been studied. Here the amide is formed directly by the reaction of a carboxylic acid with ammonia. Because these reactions are governed by the equilibrium concentrations of substrates and products, the solubility of the reactants is of major importance. The optimal conditions for an efficient ammoniolysis of butyric acid and oleic acid have now been reported. Oleamide, which has been reported to have a pharmacological application as a sleep inducing has been derived from the acid by direct ammoniolysis in an efficient manner with good yields (Scheme 12.2-2) 1’1.

94% yield

Scheme 12.2-2.

An interesting effect was found whilst studying the enantioselectivity of these reactions. Although several esters (ethyl-2-chloropropionate, ethyl lactate, ethyl2-hydroxy hexanoate and ethyl-2-methylbutyrate) were converted into the amides with only low to moderate ee values, the ammoniolysis of ibuprofen (2’-chloroethy1)ester was highly enantioselective.At 56% conversion the ee of the remaining (S)-esterwas 96% (Scheme 12.2-3),corresponding with an Evalue ofthe ammoniolysis of 28. In comparison, the ester hydrolysis of ibuprofen (2’-chloroethy1)ester catalyzed by the same enzyme proceeded with an E value of only 3.5 [*I. The same phenomenon has been observed in the ammoniolysis of 4-methyloctanoic acid. Here an E value of 76 was determined for ammoniolysis, whereas in the transesterification reaction an E-value of only 23 was found[’, 1‘.

I

717

718

I

72 Hydrorysis and Formation ofC-N Bonds

(S)(+)-ester

(R) (-)-amide

56% conversion

e.e. = 96%

Scheme 12.2-3.

It has now been reported that the amidation of esters will be used by BASF in the industrial production of amines. A broad range of amines become available in their optically pure form by kinetic resolution using a lipase from Pseudomonas sp. DSM 8246 in the amidation of methoxyacetic acid ethyl ester (see Scheme 12.2-4) 171.

- -hydrolysidracemisation - - - - I

-

I 0

I _ _ _ _ _ _ I racernisation

Scheme 12.24.

A comparitive study of a variety of lipases and lipase preparations in such alkoxycarbonylation reactions of amines has been presented by Sinisterra and coworkers [I'.

72.2 Formation and Hydrolysis of Amides I719

12.2.3

Enzymatic EnantioselectiveHydrolysis of Amides 12.2.3.1

Hydrolysis o f Carboxylic Amides

Although amidase activities have been known for quite some time, it is only in recent years that the increasing demand for chiral drugs and herbicides has triggered their exploitation as biocatalysts to a great extent. Many, but not all, amidases have been identified in microorganisms also exhibiting nitrile hydratase activity. In some cases where the enantioselectivetransformation of the nitrile to the acid can be observed, the selectivity is based on the high selectivity of the amidases rather than on the discrimination by the nitrile hydratase. Thus, using these enzyme combinations, both the (R)-amides and the (S)-acids can be obtained by such a double enantiomeric selection. The first catalytic step is carried out by a nitrile hydratase with a slight preference for the (R)-enantiomer. At high conversions this will lead to a mixture of (S)-nitrile and (R)- and (S)-amide, the latter being subsequently hydrolyzed with high selectivity by an (S)-selectiveamidase to yield the ( S ) - a ~ i d [ ~ ~ , ~ ~ l Scheme 12.2-5).

(S)-selective amidase

Scheme 12.2-5.

Other examples of (S)-selectiveamidases are described for the production of (S)2-(4'-chlorophenyl)-3-methyl butyric acid 191, (S)-ibuprofen[lo],(S)-naproxen and L-carnitine[I2'131. In addition to the more common (S)-amidase activities, (R)-specificenzymes have also been identified. Thus (S)-ketoprofenamidehas been derived from the racemic mixture using a biocatalyst from Cornamonas acidovorans KPO 2771-4 to hydrolyze the (R)-enantiomer with high selectivity['4](see Scheme 12.2-6). Lonza AG has reported on the use of enantioselectiveamidases for the resolution of piperazine-2-carboxamide and piperidine-2-carboxamide using whole cell biocatalysts from Klebsiella terrigena, PseudornonasJluorescence and Burkholderia sp., the last containing an (R)-selective amidase (Scheme 12.2-7)[''I. Furthermore, several amidases exhibiting high selectivities [either (S)- or (R)-] towards 2-arylpropiona-

720

I

12 Hydrolysis and Formation ofC-N Bonds

C. KPO-2771-4 acidovorans

&c02H

49% yield

\

e.e. = 99%

Scheme 12.2-6.

H

Q\ H

H

CoNH2

(SJ-spec. amidase

4

t

(R)-spec. amidase

c X H C O O H

99.4% e.e. 41% yield

0 Klebsieiia

99.0% e.e. 20% yield

@Burkho\deria sp. DSM 9925

97.3% e.e. 20% yield

Bpseudornonas DSM 9924

terrigena DSM 9174

H

(;I... H

"'"COOH

(A

COOH

Scheme 12.2-7.

mides have been identified. A good overview has been given by Wieser and Nagasawa [791. In recent years the availability of several amidases has been improved by cloning and overexpression[16191 resulting in biocatalysts of high activities which can readily be used for industrial purposes. Furthermore, homology studies have been carried out to identify the common features of this class of enzymes[20]. 12.2.3.2

Hydrolysis o f Amino Acid Amides

The conversion of amino acid amides into chiral amino acids has been the subject of a large number of monographs and reviews [21-29]. In this section information will be given on amidases and aminopeptidases that have been reported for the stereoselective hydrolysis of amino acid amides.

12.2 Formation and Hydrolysis ofAmides

12.2.3.2.1

Production of Chiral a-H-a-Amino Acids

At DSM a very efficient and universally applicable industrial process for the production of both optically pure L- and D-amino acids has been commerciali ~ e d [ ~271. ’ , Pivotal in this process is the enantioselective hydrolysis of D,L-aminoacid amides. The stable D,L-amino acid amides are prepared efficiently under mild reaction conditions starting from simple raw materials (Fig. 12.2-1). The reaction of an aldehyde with hydrogen cyanide in ammonia (Strecker reaction) affords the amino nitrile. The amino nitrile is converted in a high yield into the D,L-amino acid amide under alkaline conditions in the presence of catalytic amounts of acetone. The

HCNlNH3

NH2 R&N

1) NH3

3) OH-(pH=13) racemisation 4) H30+

r-

*IPhCH0

ketoneloH R 4 N H 2

I

NH2 DL-amino acid amide

a

0

1) OH-(pH=13) racemisation 2) H30+

L-specific aminopeptidase

(Pseudornonas putida)

L-amino acid

I

D-amino acid amide PhCHO pH=8-11

D-amino acid Figure 12.2-1.

H

DSM’s chemo-enzymatic route for the production of chiral a-H-amino acids.

722

I

72 Hydrolysis and Formation ofC-N Bonds Table 12.2-1.

R

Substrates by Pseudomonas putida cells.

R

R

R

H3C-

H3C-CH2-

resolution step is accomplished with permeabilized whole cells of Pseudornonas putida ATCC 12 633; a stereoselectivityof nearly 100% (E > 100) on hydrolyzing only the r-amino acid amide is combined with a very relaxed substrate specificity (see Table 12.2-1)[27, 2g-311. Not only the smallest optically active amino acid, for example alanine, but also valine, leucine, several (substituted) aromatic amino acids, heterosubstituted amino acids (methionine, homomethionine and thienylglycine) and even an imino acid, proline, are obtainable in both the L- and the D-form. Furthermore, this biocatalyst has recently been reported to hydrolyze azido amino acid amides with high enantioselectivitiesas well (vide in@) [321. No enzymatic side effects are observed and substrate concentrations up to 20 % by mass can be used without affecting the enzyme activity. The biocatalyst is active in a broad pH-range and can be used in soluble form in a batchwise process; thus poorly soluble amino acids can be resolved without technical difficulties. Re-use of the biocatalyst is possible. A very simple and elegant alternative to the use of ion-exchangecolumns or extraction to separate the mixture of D-amino acid amide and the L-amino acid has been elaborated at DSM. Thus addition of one equivalent of benzaldehyde (with respect to the D-amino acid amide) to the enzymatic hydrolysate results surprisingly in the formation of a water insoluble Schiff base with the D-amino acid amide which can be easily separated. Acid hydrolysis (H2S04'HHal, HN03 etc.) results in the formation of the n-amino acid amide, which can be hydrolyzed by cell-preparations of Rhodococcus erythropolis, yielding the D-amino acid. The amidase from this organism lacks stereoselectivity. This option is very useful for amino acids that are highly soluble in the neutralized reaction mixtures obtained after acid hydrolysis of the amide. Process economics dictate the recycling of the unwanted isomer. Path A in Fig. 12.2-1 illustrates that racemization of the D-N-benzylideneamino acid amide is facile and can be carried out under very mild reaction conditions. After removal of

72.2 Formation and Hydrolysis ofAmides I723

N3

+

-

NH2

amino peptidase

from Pputida

N3

-N3

spontaneous racemisation

Scheme 12.2-8.

the benzaldehyde the D,L-amino acid amide can be recycled. This option means that 100% conversion into the L-amino acid is theoretically possible. A suitable method for racemization and recycling of the r-amino acid (path B, Fig. 12.2-1) comprises the conversion of the L-amino acid into the ester in the presence of concentrated acid, followed by addition of ammonia, resulting in the formation of the amide. Addition of benzaldehyde and racemization by base (pH 13) gives the D,L-amino acid amide. In this way 100% conversion into the D-amino acid is possible. For the production of 2-azidophenylaceticacid an even more elegant way of achieving 100% yield of one enantiomer has been reported. Under the conditions used for the resolution a spontaneous racemization of the substrate is achieved, resulting in a dynamic kinetic resolution with a theoretical yield of 100% (Scheme 12.2-8).The distinct advantages of the aminopeptidase process are: The substrate for the enzymatic hydrolysis is a precursor of the amino acid; the number of chemical steps can be kept to a minimum. The use of relatively cheap whole cell biocatalysts contributes to the economical feasibility of the procedure. Both L- and D-amino acids can be prepared with a very high optical purity. The aminopeptidase from Pseudomonas putida ATCC 12 633 has also recently been cloned and overexpressed in E. coli resulting in a highly efficient whole-cell biocatalyst for industrial applications[*'I. The specific activity of this new biocatalyst is substantially increased (25 times) compared with the specific activity of the P. putida wild type cells without changing the other positive characteristics of the aminopeptidase. Even though the aminopeptidase from Pseudomonas putida exhibits the relaxed substrate specificity described above, an a-hydrogen atom in the substrate is an essential structural feature for the enzymatic activity. Therefore this enzyme can not be used for the resolution of higher substituted amino acids. Recently, a new biocatalyst with a broad substrate spectrum of L-specific amidase activity has been identified at DSM. Of 125 microorganisms that were able to use ahydroxy acid amides as the sole nitrogen source, Ochrobactrum anthropi NCIMB 40 321 was selected for its ability to hydrolyze racemic amides with high L-selectivity. The substrate specificity of whole Ochrobactrum anthropi cells is remarkably wide and ranges from a-H-a-amino,a-alkyl-a-amino,and N-hydroxy-a-aminoacid amides to a-hydroxyacid amides[331. After 50% conversion, both the L-acids formed and the

724

I

12 Hydrolysis and Formation ofC-N Bonds

Substrate specificty of Ochrobactrum anthropi cells. L-selective hydrolysis" of amides by Ochrobactrum anthropi.

Table 12.2-2.

Substrate

o,r-a-valine amide D,L-a-methylamide D,L-a-methylleucineamide D,r-tert leucine amide D,L-a-cinnamylalanineamide D,L-phenylglycineamide D,r-a-methylphenylglycineamide D,L-a-ethylphenylglycineamide D,L-a-propylphenylglycine amide D,L-a-allylphenylglycineamide D,L-a-benzylphenylglycine amide D,L-N-hydroxyphenylglycineamide' D,L-mandek acid amide (MAA)

Relative activity ("h)

25 5 15 1

17 lOOb

2

4

1

4

0 25 5

a Activities were measured at pH 8.0 (100mM phosphate buffer) and 40 "C using 3.0 g L-' of amide. b A relative activity of 100 corresponds to 2000 nMol min-' (mg dry mass)-'. c Incubation performed under

anaerobic conditions by flushing with nitrogen.

residual D-amides were present in 99% enantiomeric excess and ammonia accumulated in stoichiometric amounts. The substrate specificity is illustrated in Table 12.2-2. Using this biocatalyst a new route to thiamphenicol, a synthetic analog to the antibiotic chloramphenicol has been developed (Scheme 12.2-9).A precursor of the biologically active (I R,2R)-enantiomer,the (2S,3R)-para-substituted3-phenylserine is obtained by the enzymatic resolution. The residual enantiomer can be efficiently recycled via separation by Schiff base formation with the corresponding para-substituted benzaldehyde and subsequent transformation into the racemic threo-amides[341. A D-aminopeptidasehas been identified at the Sagami Research Institute by Asano et al. 13'1. The group was successful, by using an enrichment culture technique, in selecting a microorganism (Ochrobactriurn anthropi SCRC C1-38)with D-aminopepti-

Scheme 12.2-9.

12.2 Formation and Hydrolysis ofAmides

dase activity from a soil sample. The enzyme, which hydrolyzes D-alanine amide, was purified about 2800 fold. The molecular mass of the native enzyme was approximately 122000 Da, with two identical subunits having a molecular mass of about 59000 Da each. Remarkably, D-valine amide is hydrolyzed very slowly. Generally, the enzyme has higher affinity towards peptide substrates than towards amino acid amides. It does not act on peptides bearing an L-amino acid at the NH2terminus. Thus it exhibits a mode of action typical of aminopeptidases. The optimal pH for activity was 8.0. The immobilized enzyme was also active in organic solvents (benzene, butyl acetate, l,l,l-trichloroethane)[361. ~-Alanine-(3-aminopentyl) amide was quantitatively synthesized in an amination reaction from D-alaninemethylester and 3-aminopentane within 1 h. Asano et al. have also purified a D-stereospecificamino acid amidase from another Ochrobactrum anthropi isolate[37.381. Recently, a new amidase from Comamonas acidovorans has been reported that exhibits a broad substrate specificity and also Damino acid amidase In addition, a D-specific amidase has been identified in Arthrobacter sp. NJ-2G[401. In contrast to the D-selective enzymes of Ochrobactrum sp. and Cornomonas acidovorans, the D-amide hydrolase identified in Arthrobacter sp. NJ-26 was very substrate specific: a good hydrolysis rate was only observed for Dalanine amide.

12.2.3.2.2

I

725

Synthesis of a-Alkyl-a-Amino Acids

Within the pharmaceutical industry a-alkyl-a-aminoacids are regarded as valuable building blocks. An example of this class is ~-a-methyl-3,4-dihydroxyphenylalanine (L-methyl-Dopa),which is used as a drug to treat patients suffering from high blood pressure. More recently, medicinal chemists became interested in bio-active peptides containing a-alkyl-a-amino acids since they tend to freeze specific conformations and slow down enzymatic degradations Nowadays, many a-alkyl-aamino acids have been found in nature. For example, L-isovaline is found in peptaibol antibiotics. Their influence on the conformational behavior of peptides is presently under active investigation. Several routes to enantiomerically pure a-alkyl42, 431. At DSM a Mycobactea-amino acids have been elaborated in recent rium neoaurum biocatalyst has been obtained in a screening, which hydrolyzes a broad range of a-alkyl-a-amino acid amides with high enantioselectivities (Table 12.2-3). The basis of the process leading to the enantiomericallypure acid is essentially the same as that for a-H-a-aminoacids. However, in this case, a ketone is used as the starting material which undergoes a Strecker reaction, followed by hydrolysis of the resulting aminonitrile to form the racemic a-alkyl-a-aminoacid amide. Enzymatic hydrolysis results in the formation of the L-a-alkyl-a-aminoacid (Fig. 12.2-2). Some characteristics of the process are: Using this process both L- and D-a-alkyl-a-aminoacids can be produced. Permeabilized whole cells of Mycobacterium neoaurum ATCC 25795 or crude enzyme preparations can be used.

726

I

12 Hydrolysis and Formation of C-N Bonds

Table 12.2-3.

Substrate specificity of the amidase activity of Mycobacterium neoaurum cells.

Products formed

c recycle

I

D,L-a-alkyl-amino acid amide L-amidase (Mycobacferium neoaurum)

+OH R

E+NH2fiH2 NH2

L-a-alkyl-amino acid

D-a-alkyl-amino acid amide

D-a-alkyl-amino acid Figure 12.2-2.

DSM's chemo-enzymatic route for the preparation of a-alkyl-a-amino acids.

Very high stereoselectivity (> 98% ee) and a remarkably relaxed substrate specificity are observed (table 12-7). The enzyme is active in the pH range from 6.5 to 11,with a broad optimum from pH 8.0-9.5 Recently the enzyme has been purified and thoroughly characterized. It was identified as an amino acid amidase, most probably belonging to the group of

12.2 Formation and Hydrolysis of Amides

I

metallocystein hydrolases [291. In addition to DSM, Ube company reported an analogous biocatalytic route to a-methyl phenylalanine. A Pseudomonas Juorescens ( I F 0 3081) showed the highest conversion (94%) but the stereoselectivity was relatively low (ee 93.4%)[441. 12.2.3.3

Hydrolysis of Cyclic Amides

Cyclic amides can also be hydrolyzed in a highly selective fashion using enzymes. A well known example in this respect is the biocatalytic production of L-lysine from D,La-amino-s-caprolactam(D,L-ACL)[45-471. This process is based on the combination of two enzymatic reactions: the enzymatic enantiospecific hydrolysis of L-a-amino-scaprolactam to 1-lysine and the simultaneous racemization of the residual D-aamino-s-caprolactam (Scheme 12.2-10).

D-ACL

L-ACL hydrolase

L-Lysine Scheme 12.2-10.

In this way L-lysine is produced from D,L-a-amino-s-caprolactam,with a yield of almost loo%, by incubating the racemate with microbial cells of Cryptococcus laurentti, which possess L-a-amino-s-caprolactamase activity, together with cells of Achromobacter obae, which possess a-amino-E-caprolactam racemase activity. The enzymatic hydrolysis of other cyclic amides was also investigated in order to obtain chiral precursors for antibiotics and/or HIV inhibitors. Using isolates capable of growth on a range of N-acyl compounds as the sole carbon and energy source, two strains were selected for the enantioselective hydrolysis of (*)-2-azabicyclo[2.2.l]hept-5-en-3-one(Scheme 12.2-11). Rhocococcus equi NCIMB 40 231 selectively hydrolyzed the (-)-enantiomer, yielding (+)-lactam with > 98% ee (45% yield), whereas, Pseudomonas solanacearum NCIMB 40 249 hydrolyzed the opposite enantiomer with great selectivity yielding (-)-lactamwith > 98% ee (45%yield)f4',491. In

727

72 Hydrolysis and Formation ofC-N Bonds

these biotransformations the relatively low concentration of enzyme (6 g dry mass L-l), the high concentration of substrate (50 g L-'), and the speed of the reaction (3 h) are worth noting. Moreover, mutant strains have been constructed which hyperexpress the amidase activity. Subsequently it was shown that Rhodococcus equi cells can also be applied for the enantioselective hydrolysis of 6-azabicyclo[3.2.0]hept-3-en?'-one, yielding a precursor for the antifungal agent cispentacin f5O]. Evidently, the use of the whole cell biocatalyst enables an efficient biotransformation with high substrate concentrations.

O

D

H

/ \

Scheme 12.2-11.

Recently, BASF has described the enantioselective hydrolysis of substituted lactams. Using strains of Pseudomonas aeruginosa and Rhodococcus erythropolis obtained from soil samples, both enantiomers of 5-vinylpyrrolidinon can be derived ["I. 12.2.4 Selective Cleavage of the C-Terminal Amide Bond

In peptide synthesis selective deprotection of C- or N-terminal groups is common in most methods of chain elongation. The amide groups offer some advantages for Cterminal protection (enhanced chemical stability and increased solubility in water). However, selective cleavage of this amide bond in the C-terminal position was previously impossible. Both chemical and biochemical methods also led to internal peptide bond hydrolysis, giving rise to difficult separation problems. Consequently the amide group has been rather unattractive for C-terminal protection in peptide synthesis. Now, however this situation has changed. Steinke and Kula have isolated an unusual peptide amidase from orange flavedo, which is very selective for the

I hydrolysis of the C-terminal amide bond of peptides[52].The peptide amidase is free 72.2 Formation and Hydrolysis ofAmides

of any proteolytic activity, which would either hydrolyze internal peptide bonds of substrate peptides or side chain amide bonds. The substrate spectrum of this enzyme includes protected and unprotected peptide amides and N-protected amino acid a m i d e ~ ' ~The ~ ] . chain length of the substrate peptide amide, as well as the amino acid composition, including the Cterminal amino acid side chain, are of minor importance. The amidase activity is stereoselective with regard to the C-terminal position, only L-amino acid amides are accepted. Unprotected amino acid amides are not hydrolyzed by this novel enzyme. The broad application of this enzyme is further extended by its broad pH activity range, from 6.5 to 9. Evidently, a new and usehl biocatalyst is now available for selective deaminati~n['~]. Recently the same group has also shown that the reverse reaction, the ammoniolysis of peptides, is catalyzed by this enzyme. Reducing the water activity by carrying out the reaction in acetonitrile containing 5 % of water, ZGly-Phe-NH2has been derived in a thermodynamic ammoniolysis in yields of up to 35 % [55'. 12.2.5

Amidase Catalyzed Hydrolytic and Synthetic Processes in the Production of Semi-synthetic Antibiotics

Since the discovery of the P-lactam antibiotic penicillin G (Fig. 12.2-3)by Fleming in 1929, the use of antibiotics against pathogenic bacteria has increased dramatically. Penicillin G was initially used, which must be applied intravenously because of its instability in the stomach, but now penicillin V, which can be administered orally, has been introduced. However, as a result of the increasing resistance of bacteria, new antibiotics had to be developed. The semi-synthetic antibiotics, which often possess a broad spectrum of antibacterial activity, were produced by altering the side chain of penicillin G through acylation of the amine function of 6-aminopenicillanic acid (6-APA)[561. The first commercial semi-synthetic antibiotic was ampicillin, which was introduced by Beecham in 1961[57].A few years later a new class of antibiotics, the cephalosporins, was marketed. Some of the semi-synthetic cephalosporins are prepared from 7-aminocephalosporanic acid (7-ACA),others from 7-aminodesacetoxycephalosporanic acid (7-ADCA).7-ACA is an intermediate that can be obtained from the fermentation product cephalosporin C; 7-ADCA is an intermediate that was discovered by Morin et al. ["I using chemical ring expansion of the penicillin nucleus (Fig. 12.2-4).The only difference between the two molecules is the absence of an acetoxy moiety in 7-ADCA. Today, the main intermediates for semi-synthetic cephalosporins (SSCs) and penicillins (SSPs), 7-ADCA and 6-APA, respectively, are produced in quantities of many thousands of tons annually in biocatalytic processes using penicillin amidases. The coupling of the side chains to 6-APA and 7-ADCA is still performed chemically. However, in order to obtain improved coupling yields and to overcome the use of toxic and hazardous chemicals and solvents, several leading producers are

729

730

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72 Hydrolysis and Formation of C-N Bonds

@OH

O A O H Penicillin G

Penicillin V

Cephalosporin C Figure 12.2-3.

The three main fermentatively derived fi-lactams.

currently investigating so-called“green routes” again utilizing penicillin amidases to perform this coupling reaction enzymatically. 12.2.5.1

Enzymatic Production o f 6-APA, 7ADCA and 7-ACA Using Amidases: Hydrolytic Processes

Owing to the increasing importance of semi-synthetic antibiotics, commercially feasible routes are of the utmost importance and several methods have been developed. About a decade ago 6-APA and 7-ADCA were mainly produced by chemical deacylation of penicillin G, penicillin V or phenylacetyl 7-ADCA, the last of which was derived from chemical ring expansion of oxidized penicillin G. As a result of the fact that these processes were rather complex and employed hazardous reagents, for example pyridine, phosphorus pentachloride, nitrosyl chloride and dichloromethane, alternative processes have been developed. Penicillin amidases (E. C. 3.5.1.11) catalyze the hydrolysis of the linear amide bond in penicillin molecules producing both the p-lactam nucleus, 6-APA and the corresponding side chain without affecting the p-lactam amide bond in the four-membered ring. Based on their substrate specificity the penicillin amidases are grouped into three classes [’I: 1. Amidases that preferentially hydrolyze penicillin V (phenoxymethylpenicillin). 2. Amidases which are primarily active against penicillin G. 3. Amidases which are most active using ampicillin as the substrate.

12.2 Formation and Hydrolysis ofArnides I 7 3 1 H

H

OAOH

0-OH

Penicillin G

Phenylacetyl7-ADCA

I

I

Penicillin amidase

H2N% O>cH /3

CH3

0-OH 6-APA

OAOH 7-ADCA

Enzymatic production o f 6-APA reaction in which a molecule ofwater is exand 7-ADCA. 6-APA is produced from penicillin cluded. Again, penicillin amidase is used for the G (or V) using the enzyme penicillin amidase. hydrolysis o f phenylacetyl 7-ADCA into 7-ADCA. For the production of 7-ADCA, penicillin C is Both the production o f 6-APA and o f 7-ADCA involve the liberation o f phenylacetic acid, a transformed chemically into phenylacetyl 7-ADCA. This transformation involves oxidation molecule that can be recycled into the fermentation process. of penicillin C followed by a ring expansion Figure 12.2-4.

Penicillin G amidases, in contrast to penicillin V amidases, display a fairly relaxed substrate specificity. Consequentlypenicillin G amidases can also be used for various other applications 611. Major breakthroughs that facilitated enzyme application on an industrial scale were improvements in the area of enzyme isolation, purification and immobilization. Thus, the development of genetically engineered microorganisms accounted for the high yield production of penicillin amidases. Also, the introduction of immobilized enzyme systems, both for whole cell systems and for the isolated and purified amidases [’’)- 62, 631, resulted in prolonged enzyme stability enabling reuse and continuous process modes. As a result of this, the enzymatic routes currently display far better economics for both 6-APA and 7-ADCA production (Fig. 12.2-4) compared with their chemical counterparts. Nowadays, excellent penicillin amidases from various sources are being used on

732

I

72 Hydrolysis and Formation ofC-N Bonds

t

W

6-APA Figure 12.2-5. Schematic representation of industrial production of 6-APA using column packed immobilized penicillin amidase.

an industrial scale for producing either 6-APA or 7-ADCA[’’. 631. These biocatal@c processes are generally performed batchwise at 35-40 “C and pH 7.5-8.5[622. 64,651. Upon formation of 6-APA and 7-ADCA the side chain acid is liberated, which causes a drop in the pH of the reaction mixture. This pH change results in a decrease in the reaction velocity. Since a higher starting pH is not desired because of enzyme deactivation and P-lactam ring hydrolysis, a strict control of the pH is necessary during the process. Generally, pH adjustment occurs separately from the immobilized enzyme, either by packing the immobilized enzyme in columns, as outlined in Fig. 12.2-5, or by cycling the contents of the enzyme reactor through a sieve, retaining the immobilized biocatalyst over a small pH control vessel. Currently, immobilized penicillin amidases can be reused up to 600 times [”]. After completion of the hydrolcc reaction the immobilized biocatalyst is separated from the liquid and the products 6-APA or 7-ADCA are precipitated at their iso-electric points and collected by filtration. After washing and drying an extremly pure product is obtained[65]. In addition to 7-ADCA, 7-ACA is also a very useful intermediate for the production of other SSCs (for example cefazolin, cefotaxime, ceftriaxone and cefuroxime).Until recently, 7-ACA was produced chemically from cephalosporin C using the phosphorus pentachloride process or the “Delft Cleavage”[65].As a result of the good experiences with penicillin amidases and the increasing concern about the amount

12.2 Formation and Hydrolysis of Amides

NH3

0 0

I

Cephalosporin C chemical deamination followed by decarboxylation

I

hydrolysis by glutaryl amidase

7-ACA Figure 12.2-6.

Cherno-enzymatic production of 7-ACA from cephalosporin C.

of waste being produced in chemical side chain cleavage processes, several companies are engaged in the development of enzymatic processes for the production of 7-ACA. Several years ago Asahi commercialized a chemo-enzymatic process in which cephalosporin C is oxidatively deaminated to glutaryl-7-ACA, which is

I

733

734

I

72 Hydrolysis and Formation of C-N Bonds

Cephalosporin C

0,

D-amino acid oxidase;

+

H,O;

0

H

I

I1

HO

0

0

K 0

CH3

2-Ketoadipyl-7-ACA

H

I

K 0

4

CH3

Glutaryl-7-ACA glutaryl-7-ACA amidase;

oy&

glutaric acid HZN

K 0

7-ACA

CH3

Figure 12.2-7.

Two-enzyme bio-

catalytic process for production of 7-ACA from cephalosporin C.

12.2 Formation and Hydrolysis ofAmides

subsequently hydrolyzed enzymatically using a glutaryl amidase from a Pseudornonas species (Fig. 12.2-6)IG5I. Currently,the first step of this process is also carried out enzymatically. Using a D-amino acid oxidase cephalosporin C is oxidized to the corresponding a-ketoadipyl derivative. This latter compound spontaneously decarboxylates to give glutaryl-7-ACA (Fig. 12.2-7)rG5, G71. So far no direct deacylation of cephalosporin C has been commercialized, although enzymes have been identified that can indeed catalyze this one-step hydrolysis[‘*I. After optimization of the production of this biocatalyst, and possibly improvement of its intrinsic properties, it is very likely that a one-step enzymatic hydrolysis of cephalosporin C will be industrialized. 12.2.5.2 A New Fermentation-based Biocatalytic Process for 7-ADCA

With the aid of metabolic pathway engineering a large step forward has now been realized in the production of 7-ADCA by adapting processes within penicillin producing organisms. Thus, the conversion of the five-membered penicillin ring into the six-membered cephalosporin ring can now be performed within the microorganism as outlined in Fig. 12.2-8. By modifymg the responsible gene, the penicillin producing mould can be set to produce a 7-ADCA derivative directly. Thus, several chemical steps from penicillin via penicillin oxide to 7-ADCA can be omitted[G9]. Because of a newly introduced gene, the substrate specificity of the engineered strain changed. Now, dicarboxylic acid is used as the externally added side chain, instead of phenylacetic acid as in penicillin G. Later in the process this side chain is removed enzymatically, using an enzyme quite similar to the glutaryl amidase from Pseudomonas sp. as in the enzymatic production of 7-ACA. For the production of 7-ADCA and the dicarboxylic acid amidase, new plants are currently under construction at DSM in The Netherlands. Compared with the old process for the production of 7-ADCA, the major advantages of this process are higher purity of the end product, much greater energy efficiency and almost complete absence of organic solvents. 12.2.5.3 Enzymatic Formation of Semi-synthetic Antibiotics: Synthetic Processes

The chemical synthesis of semi-syntheticantibiotics (SSAs)from a p-lactam nucleus (such as 6-APA, 7-ACA, or 7-ADCA) and a side chain (such as D-(-)-phenylglycineor an aminothiazoleiminoaceticacid derivative) is difficult to carry out in a single step since both reactants have functional groups that can easily form undesired covalent bonds. In order to obtain the desired product in high yield it is necessary to activate the carboxyl function of the acylating agent, to temporarily protect interfering amino functions, to effect the formation of the amide bond and to remove the protecting groups. Moreover, this condensation should be performed under conditions that will

I

735

736

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72 Hydrolysis and Formation ofC-N Bonds ~

~~

Sugar Fermentation

1

Fermentation

Fermentation

Chemistry

!

1

Penicillin G

Chemistry

1

1

Penicillin G sulfoxide Chemistry

Chemistry

Biocafalysis

Phenylacetyl-7-ADCA Chemistry

Biocatalysis

I

7-ADCA Chemistry

1

Biocatalysis

1

Cefadroxil, Cephalexin, Cephradine

Biocatalysis

1

Figure 12.2-8. Production of 7-ADCA has undergone remarkable changes. In the early days (left-hand side), chemical ring expansion o f penicillin C resulted in the formation of the cephalosporin nucleus. The phenylacetyl moiety was then removed chemically. Later on, this last step was replaced by a biocatalytic step using penicillin arnidase (middle). On the right hand side, a completely new route is presented. Dicarboxyl-7-ADCAis obtained directly by fermentation. A dicarboxyl amidase is used to remove the dicarboxyl group.

preserve stereochemical integrity and leave the fragile four-membered p-lactam ring intact. Today, nearly all SSAs are produced using the methodology described above. However, because of the relative complexity of these chemical processes and the use of toxic reagents and solvents, the application of biocatalysis has promising possibilities here too. Indeed, several biocatalytic processes have been developed and are now being introduced on a production scale. A noteworthy advantage of these processes is that protection of functional groups is not a prerequisite because of the mild reaction conditions and the high selectivity of the enzymes involved. So far, the major focus of research has been directed at enzymatic coupling of D(-)-phenylglycine methylester or D-(-)-phenylglycine amide with either 6-APA or 7-ADCA yielding ampicillin and cephalexin, respectively, and at the coupling of D(-)-4-hydroxyphenylglycine derivatives with either 6-APA or 7-ADCA leading to amoxicillin and cefadroxil, respectively (Fig. 12.2-9) [ 6 3 , 7&751 . During this kinetically controlled condensation, the activated 4-hydroxyphenylglycineforms an acyl-enzyme complex with the penicillin amidase L7'1. Subsequently, this acyl-enzyme complex is deacylated by a nucleophile, the p-lactam nucleus 6-APA or 7-ADCA, or

12.2 Formation and Hydrolysis ofAmides

I

737

OAOH

I

OAOH

I

Penicillin amidase

J.

Ampicillin

f

Cephalexin

Enzymatic conversion o f 6-APA and 7-ADCA into ampicillin and cephalexin using penicillin amidase. The side chain is introduced using an activated form o f D-(-)-phenylglycine, either the amide (R = NHp) or the ester (R = OCH3, OCzHs). Figure 12.2-9.

water. In the first case this leads to product formation, whilst in the second case the activated side chain is hydrolyzed. By carefully tuning reaction conditions and downstream processing sequences for every individual product, yields of up to 90% based on 6-APA or 7-ADCA have been obtained. However, as a result of the competing hydrolysis of the acyl-enzyme complex by water the yield with respect to the ester or amide was quite low (approximately 30%)[72,731. By performing this condensation in the presence of an alcohol, as a result of which the activated phenylglycine is 'recycled' in situ, the yield based on phenylglycine could be improved[75].The latest breakthroughs are the development of immobilized biocatalysts with improved performance, application of low temperatures and using high substrate concentrations L7'1. It is these improvements that make enzymecatalyzed synthesis of SSAs in a purely aqueous environment competitive industrially with traditional chemical synthesis. 12.2.6 Conclusions and Future Prospects

As indicated in the preceding sections, amides and their derivatives are important versatile building blocks for the (agro)chemicaland pharmaceutical industry. Owing to the selectivity of amidases (both regio- and enantioselectivity) and the fact that

738

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12 Hydrolysis and Formation ofC-N Bonds

Reduction of waste volumes resulting from introducing penicillin acylase biocatalysis in antibiotics production.

Table 12.2-4.

Process

Waste reduction factor

Penicillin G + 6-APA Penicillin G --t 7-ADCA 6-APA -t semi-synthetic penicillins 7-ADCA --t semi-synthetic cephalosporins

these conversions can be achieved under very mild conditions, several biocatalytic processes based on amidases have recently been commercialized. The use of these biocatalysts in the chemical industry is expected to increase in importance in the near future as environmental restrictions become more pronounced. The benefits in terms of reduction of waste can be enormous, as can be judged from that already achieved in the production of antibiotics (Table 12.2-4). References M. C. De Zoete, A. C. Kock-van Dalen, F. van Randwijk, R. A. Sheldon, Biocatalysis 1994, 10,307-316. 2 M. C. De Zoete, A. C. Kock-van Dalen, F. van Randwijk, R. A. Sheldon, J. Chem. Soc., Chem. Commun.1993,24,1831-1832. 3 E. M. Anderson, K. M. Larsson, 0. Kirk, Biocatal. Biotransform. 1998, 16, 181-204. 4 B. F. Cravatt, 0. Prospero-Garcia, G. Siuzdak, N. B. Gilula, S. J. Hanriksen, D. L. Boger, R. A. Lerner, Science 1995,268, 1506-1 509. 5 M. J. J. Litjens, A. J.J. Straathof, J. A. Jongejan, J. J. Heijnen, Tetrahedron 1999, 55, 12411-12418. 6 N. W. J. T. Heinsman, S. C. Orrenius, C. L. M. Marcelis, A. De Soma Teixeira, M. C. R. Franssen, A. Van der Padt, J. A. Jongejan, A. De Groot, Biocatal. Biotransform. 1998, IG, 145-162. 7 W. E. Ladner, K. Ditrich, Chimica Oggi 1999, 7/8, 51-54. 8 M. S. de Castro, P. Dominguez, J. V. Sinisterra, Tetrahedron 2000, 56, 1387-1391. 9 R. D. Fallon, K. M. Fried, K. Ingvorsen, W. Jobst, W. J. Linn, B. Yde, B. Stieglitz 1992, W09201062. 10 E. Cerbeland, D. Petre, US Patent 1991, 5,034,329. 11 B. Hirrlinger, A. Stolz, H.-J. Knackmuss, 1. Bacteriol. 1996, 178,3501-3507. 12 U. Joeres, M. Kula, Appl. Microbiol. Biotechnol. 1994, 40, 599-605. 1

U. Joeres, M. Kula, Appl. Microbiol. Biotechnol. 1994, 40, 606-610. 14 K. Yamamoto, K. Otsubo, A. Matsuo, T. Hayashi, I. Fujirnatsu, K. I. Komatsu, Appl. Environ. Microbiol. 1996, 62, 152155. 15 E. Eichhorn, J.-P. Roduit, N. Shaw, K. Heinzmann, A. Kiener, Tetrahedron: Asymm. 1997,8,2533-2536. 16 M. Kobayashi, H. Komeda, T. Nagasawa, M. Nishiyama, S. Horinouchi, T. Beppu, H. Yamada, S. Shimizu, Eur. J. Biochm. 1993, 217,327-336. 17 S. Farnaud, R. Tata, M. K. Sohi, T. Wan, P. R. Brown, B. J. Sutton, Biochem. J . 1999, 340,71-714. 18 R. Hayashi, K. Yamamoto, A. Matsuo, K. Otsubo, S. Muramatsu, A. Matsuda, K. Komatsu, J . Ferment. Bioengineer. 1997, 83, 139-145. 19 S. J. Wu, R. D. Fallon, M. S. Payne, D N A Cell Biol. 1998, 17, 915-920. 20 H. Chebrou, F. Bigey, A. Amaud, P. Galzy, Biochim. Biophys. Acta - Protein Struct. Mol. Enzymol. 1996, 1298,285-293. 21 K. Aida, I. Chibata, K. Nakayama, K. Takinami, H. Yamada, (Eds.),Biotechnology of A m i n o Acid Production: Progress i n Industrial Microbiology, Vol. 24, Elsevier, Amsterdam, 1986. 22 K. Yonaha, K. Soda in: Application ofStereoselectivity of Enzymes: Synthesis of Optically Active Amino Acids and a-Hydroxy-Acids, 13

References and Stereospecific Isotope-Labelingof Amino Acids, Amines and Coenzymes (Ed.: A. Fiechter), Advances in Biochemical Engineering/Biotechnology, Vol. 33, Springer Verlag, Berlin, 1986, pp. 95-130. 23 P. M. Williams, Synthesis ofOptically Active Amino Acids, Pergamon Press, Oxford, 1989. 24 J. Kamphuis, H. F. M. Hermes, J. A. M. van Balken, H. E. Schoemaker, W. H. J. Boesten E. M. Meijer in: Amino Acids: Chemistry, Biology and Medicine (Eds.: G. Lubec, G. A. Rosenthal), ESCOM Science Publishers B.V., 1990, pp. 119-125. 25 J. Kamphuis, W. H. J. Boesten, Q. B. Broxterman, H. F. M. Hermes, J. A. M. van Balken, E. M. Meijer, H. E. Schoemaker in: Advances in Biochemical EngineeringlBiotechnology (Ed.: A. Fiechter), Vol. 42, Springer Verlag, Berlin, 1991, pp. 133-186. 26 J. Kamphuis, E. M. Meijer, W. H. J. Boesten, Q. B. Broxterman, B. Kaptein, T. Sonke, H. F. M. Hermes, H. E. Schoemaker in: Biocatalytic production ofAmino acids and Derivatives (Eds.: D. Rozzell, F. Wagner), Hanser Publ. Munich 1992, pp. 177-206. 27 J. Kamphuis, W. H. J. Boesten, B. Kaptein, H. F. M. Hermes, T. Sonke, Q. B. Broxterman, W. J. J. van den Tweel, H. E. Schoemaker in: Chirality in Industry (Eds.:A. N. Collins, G. N. Scheldrake, J. Crosby), J. Wiley & Sons Ltd., Chichester, UK, 1992, pp. 187-208. 28 A. Taylor, Trends Biotechnol. Sci. 1993, 167-171. 29 T. Sonke, B. Kaptein, W. H. J. Boesten, Q. B. Broxterman, J. Kamphuis, F. Formaggio, C. Toniolo, F. P. J. T. Rutjes, H. E. Schoemaker in: Stereoselective Biocatalysis, (Ed.: R. N. Patel), Marcel Dekker, New York, 2000, pp. 23-58. 30 F. P. J. T. Rutjes, H. E. Schoemaker, Tetrahedron Lett. 1997, 38, 677-680. 31 L. B. Wolf, K. C. M. F. Tjen, F. P. J. T. Rutjes, H. Hiemstra, H. E. Schoemaker, Tetrahedron Lett. 1998, 39, 677-680. 32 C. W. Tornoe, T. Sonke, I. Maes, 13. Schoemaker, M. Morton, Tetrahedron: .4sym. 2000, I I , 1239-1248. 33 W. J. J. van den Tweel, T. J. G. M. van Dooren, P. H. de Jonge, B. Kaptein, A. L. L. Duchateau, J. Kamphuis, 4 p l . Microbiol. Biotechnol. 1993, 39, 296-300. 34 B. Kaptein, T. J. G. M. van Dooren, W. H. J.

Boesten, T. Sonke, A. L. L. Duchateau, Q. B. Broxterman, J. Kamphuis, Org. Process Res. Develop. 1998,2, 10-17. 35 Y. Assano, A. Nakazawa, Y. Kato, K. Kondo, J. Biol. Chem. 1989,264, 14233-14239. 36 Y. Kato, Y. Asano, A. Nakazawa, K. Kondo, Tetrahedron 1989,45,5743-5754. 37 Y. Asano, T. Mori, S. Hanamoto, Y. Kato, A. Nakazawa, Biophys. Res. Commun. 1989, 162,470-474. 38 H. Komeda, Y. Asano, Eur.J. Biochem. 2000, 267, 1-9. 39 T. Hayashi, K. Yamamoto, A. Matsuo, K. Otsubo, S. Muramatsu, A. Komatsu,J. Fern. Bioengineer 1997,83,139-145. 40 A. Ozaki, H. Kawasaki, M. Yagasaki, Y. Hashimoto, Biosci. Biotech. Biochem. 1992, 56, 1980- 1984. 41 C. Toniolo, M. Crisma, F. Formaggio, G. Caricchioni, G. Precigoux, A. Aubry, J. Kamphuis, Biopolymers 1993, 33, 1061-1072. 42 B. Kaptein, W. H. J. Boesten, W. J. J. van den Tweel, Q. B. Broxterman, H. E. Schoemaker, F. Formaggio, M. Crisma, C. Toniolo, J. Kamphuis, Chimica O g i 1996, 14, 9-12. 43 T. Wirth, Angav. Chem., Znt. Ed. Engl. 1997, 36,225-227. 44 Ube, Germany Patent DE 321 7908 1989. 45 T. Fukumura, Agric. Bid. Chem. 1976,40, 1687. 46 T. Fukumura, Agric. Bid. Chem. 1976,40, 1695. 47 T. Fukumura, Agric. Bid. Chem. 1977,41, 1327. 48 S. J. C. Taylor, A. G. Sutherland, C. Lee, R. Wisdom, S. Thomas, S. M. Roberts, C. Evans,]. Chem. SOC.,Chem. Commun. 1990, 1120-1 121. 49 C. Evans, R. McCague, S. M. Roberts, A. G. Sutherland, J. Chem. SOC.,Perkin Trans. 1 1991,656-657. 50 C. Evans, R. McCague, S.M. Roberts, A. G. Sutherland, R. Wisdom, J. Chem. Soc., Perkin Trans. 1 1991,2276-2277. 51 B. Hauer, F. Balkenhohl, W. Ladner, U. Pressler, European Patent Appl. EP 068 7736. 52 D. Steinke, M. R. Kula, Angew. Chem. Int. Ed. Engl. 1990,29,1139-1140 53 D. Kammermeier-Steinke. A. Schwarz, C. Wandrey, M. R. Kula, Enzyme Microb. Technol. 1993, 15, 764-760.

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D. Steinke, M. R. Kula, A. Schwarz, C. Wandrey, G e m a n Patent DE 401 4564. 55 V. Cerovsky, M. R. Kula, Angew. Chem. 1998, 110,19861985). 56 J. Venveij, E. de Vroorn, Red. Trav. Chim. Pays-Bas 1993, 112,66-81. 57 J. H. C. Nayler, TIBS 1991, 16,234-237. 58 R. B. Morin, B. G. Jackson, R. A. Mueller, E. R. Lavagnino, W. B. Scanlon, S. L. Andrews, J . Am. Chem. Soc. 1963,85, 1896-1897. 59 J. G. Shewale, H. Shivaraman, Biochem. 1989,146-154. 60 E. Baldaro, C. Fuganti, S. Servi, A. Tagliani, M. Terreni, The Use of Immobilized Penicillin G Acylase in Organic Synthesis, in: Microbial reagents in organic synthesis (Ed.: S. Servi), Kluwer Academic Publishers, 1992, pp. 175-188. 61 C. Fuganti, P. Crasselli, Tetrahedron Lett. 1986,27,3191-3194. 62 E. J. Vandamrne, Enzyme Microb. Technol. 1983,5,403-416. 63 J. G. Shewale, B. S. Deshpande, V. K. Sudhakaran, s. s. Ambedkar, Process Biochem. 1990, pp. 97-103. 64 E. J. Vandarnrne, Immobilized Biocatalysts and Antibiotic Production: Biochemical, Genetical and Biotechnical Aspects, in: Bioreactors, Immobilized Enzymes and Cells (Ed.: M. Moo-Young),Elsevier, 1988, pp. 261-286. 65 K. Matsumoto, Production of 6-APA, 7-ACA and 7-ADCA by Immobilized Penicillin and Cephalosporin Amidases, in: Industrial Application oflmmobilized Biocatalysts (Eds.: A. Tanaka, T. Tosa, T. Kobayashi),Marcel Dekker Inc., 1992, pp. 67-88. 54

H. W. 0. Weissenburger, M. G. van der Hoeven, Red. Trav. Chim. Pays-Bas 1970, 89, 1081-1084. 67 K. Sauber, Lessonsfiom Industry. in: Stability and stabilization ofenzymes (Eds.: W. van der Tweel, R. Buitelaar, A. Harder), Elsevier Science Publishers, 1993. 68 A. Matsuda, K. Matsuyama, K. Yarnarnoto, S. Ichikawa, K. Kornotsu,j. Bacteriol. 1987, 169, 5815-5823. 69 E. J. A. X. van de Sandt, E. de Vroom, Chimica O& 2000, 18,72-75. 70 R. Okachi, Y. Hashirnoto, M. Kawamori, R. Katsumata, K. Takayama, T. Nora, Enzyme Eng. 1982,6,81-90. 71 A. Bmggink, E. C. Roos, E. de Vroom, Org. Proc. Res. Deu. 1998,2, 128-133. 72 F. Knauseder, N. Palrna, Enzymatic Synthesis of Cephalexin by Iimmobilized Penicillin Acylasefiom E. Coli, ECB-3 Miinchen, Part 1,1984, pp. 431-438. 73 E. M. Baldaro, Efect of Temperature on Enzymatic Synthesis of Cephalosporins in: Bio-organic chemistry in healthcare and technology (Eds.: U. K. Pandit, F. C. Aldenveireldt), Plenum Press, 1991, pp. 237-240. 74 C. K. Hyun, J. H. Kim, Y. J. Kim, Biotechnol. Lett. 1989, 11, 537-540. 75 V. Kasche, Biotechnol. Lett. 1985,7, 877-882. 76 V. Kasche, Enzyme Microb. Technol. 1 9 8 6 8 , 4-15. 77 H. Kakeya, N. Sakai, T. Sugai, H. Ohta, Tetrahedron Lett. 1991, 32, 1343-1346. 78 R. D. Fallon, B. Stieglitz, I. Turner, Jr., Appl. Microbiol. Biotechnol. 1997,47, 156-161. 79 M. Wieser, T. Nagasawa in: Stereoselective Biocatalysis (Eds.: R. N. Patel) Marcel Dekker, New York, 2000, pp. 461-486. 66

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002 72.3 Hydrolysis of N-Acylarnino Acids

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12.3 Hydrolysis of N-Acylamino Acids Andreas 5.Bornrnarius 12.3.1

Introduction

The enzymatic hydrolysis of N-acylamino acids has been known for a century and was first detected in aqueous kidney preparations ['I. Based on the finding that this enzymatic hydrolysis proceeds enantiospecifically[2],Greenstein and coworkers developed a general and very attractive procedure for the resolution of a vast number of racemic N-acylated amino acids to the corresponding L-amino acids catalyzed by aminoacylase (E. C. 3.5.1.14) whereas the N-acetyl-D-amino acid does not reactr31 (Fig. 12.3-1). These initial investigations on a laboratory scale subsequently lead to industrial processes for the production of L-amino acids on a multi-ton scale applied by Tanabe["'I and Deg~ssa['-'~~. The first work on the isolation and characterization of aminoacylases came from Greenstein and coworkers. Fractionation of hog kidney homogenates with ammonium sulfate and acetone revealed that two distinct enzymes were present in the crude preparati~nl'~]. One was found to hydrolyze a large number of N-acetylamino acids and was designated acylase I (E.C. 3.5.1.14) whereas the other was found to hydrolyze preferentially N-acylated L-aspartic acid and was designated acylase I1 (aspartoacylase, E. C. 3.5.1.15). Additionally, a third aminoacylase, which acts preferentially on N-acylated aromatic amino acids, was found in kidney homogenates and was designated acylase 111[14, 151. Besides the LDL-Met

-

HCN + NH3 +

ow + H3C-SH

?;" 0

Racernization Figure 12.3-1. Enantiospecific hydrolysis of N-acetyl-o,L-amino acids catalyzed by aminoacylase I.

742

I

12 Hydrolysis and Formation of C-N Bonds

specific enzymes from kidney preparations, L- as well as D-specific aminoacylases have been isolated from a variety of microorganisms [16-331. Although the physiological function of these enzymes is not known with certainty, it is assumed that they may be involved in the degradation of N-acylated amino acids occupying the N-termini of many proteins and are subsequently formed in the catabolic metabolism of proteins [I7, 34, 351. 12.3.2

Acylase I (N-Acylamino Acid Amidohydrolase, E.C. 3.5.1.14)

Since acylase I has a wide substrate specificity and high enantioselectivity, it is a broadly applicable enzymatic catalyst for the kinetic resolution of most of the natural Table 12.3-1.

Substrate specificity of acylase I

Substrate

N-X-D,L-rnethionine N-X-D,L-ethionine N-X-D,L-norvaline N-X-~,~-aminobutyric acid N-X-D,L-norleucine N-X-D,L-aminoheptanoicacid N-X-D,L-leucine N-X-DJ-alanine N-X-D,L-serine N-X-D,r-glutarnicacid N-X-o,L-arninocaprylicacid N-X-D,L-valine N-X-D,L-arninocyclohexylacetic acid N-X-D ,L-aminophenylaceticacid N-X-glycine N-X-D,L-allothreonine N-X-D,L-threonine N-X-D,L-isoleucine N-X-D,L-arginine N-X-D,r-alloisoleucine N-X-o,L-histidine N-X-D,r-phenylalanine N-X-o,r-diaminopropionic acid N-X-D,r-arninocydohexylpropionic acid N-X-D,L-tyrosine N-X-D,L-ornithine N-X-D,L-lysine N-X-D,L-arninocyclohexylbutyricacid N-X-D,r-S-benzylcysteine N-X-D,L-tryptophan N-X-D,L-asparticacid N-X-D,L-proline N-X-D,r-phenylserine

(E.C.3.5.1.14) f r o m hog kidney["]. Relative activity X = acetyl X = chloroacetyl

looa

413b

64

-

-

-

167 139 126 117 68 48 48 52 32 21 19 19 11 11 3

2 2

-

1.o 0.6

-

-

59

-

22 13

-

13

7

0.6

-

-

1.9' 1.4 1.4 1.3' 0.6' 0.5

0.4

-

< 0.1

< 0.1

< 0.1 < 0.01

< 0.1 < 0.1

-

0

a 400 pmoles x min-' per mg of N at 38 "C. b Dichloroacetyl.c Not determined.

12.3 Hydrolysis ofN-Acyhmino Acids

I

743

Table 12.3-2. Comparison of the kinetic and chemical properties o f pig kidney and Aspergillus oryzae aminoa~yIase[~'1. Property

Molecular mass Subunits Metal ions (Zn2+) SH-groups Inhibition by N-a-ptosyl-r-lysine Chloromethyl ketone HC1 Inhibition by diethylpyrocarbonate Inactivation by metal chelating agents pH-optimum (chloroacetylalanine) 112 Cystine residues Tryptophan residues KM x lo3 mol L-' (chloroacetylalanine) Spec. activity U mg-' (chloroacetylalanine) Peptidase activity CI-Ac-Glu-PABAhydrolysis Heat stability Activation by Co"

Aspergillus oryzae aminoacylase

Pig kidney aminoacylase

73 200 2

85 500 2

6 0

-

+

+

completely reversible 8.5

completely reversible 8.0

4 6 6.3

12 12 6.6 250 (pH 7.8)

319 (pH 8.0) -

+

+

-

60 "C: denaturation

60 "C:denaturation

+

+

as well as unnatural and rarely occurring a-amino acidsl3>361. Thus it is the most important and mostly frequently used aminoacylase in the chemoenzymatic synthesis of L-amino acids from the corresponding racemic N-acetylated precursors (Table 12.3-1). The numerous investigations on acylase I have been reviewed on several occasions [361. Acylase I has been isolated and characterized from kidney preparations [13, 37-39] as well as the fungi Aspergillus oryzae[16. 17, 35* 401 (AA) and Aspergillus melleus (AM).The enzymes from the two Aspergillus species are virtually A comparison of some kinetic and chemical properties of pig kidney acylase (PKA) and the mold enzyme from Aspergillus olyzae (AA) is shown in Table 12.3-2. Both enzymes are dimeric, zinc-containing metalloproteins of roughly the same size and which are strongly activated by external addition of cobalt ions['7, 35, 40, 42-441; Zn2+ is essential for activity[40].The Co2+/acylase-dissociationconstants of PKA and AA are similar with 10-7.5M i4O] und M [431, respectively, the respective constants for Zn*+/acylaseare identical at 10-l' M L40, 451. They differ in the amount of zinc ions bound per subunit and in the number of SH-groups as well as cysteine and tryptophan residues essential for catalyhc activity. The properties of acylase I from Aspergillus oryzue are summarized in Table 12.3-3[461. 12.3.2.1

Genes, Sequences, Structures

The DNA and protein sequences of eight aminoacylases are now known, as of March 2001. Sequences from ~ o m supiens o [47, 481, pig [49, "1, ~ucillusstearothermophilus and Lactococcus lac ti.^[^^] are known for aminoacylase I, sequences from Homo

744

I

12 Hydrolysis and Formation of C-N Bonds Table 12.3-3.

Properties of acylase I from Aspergillus o r y ~ a e ‘ ~ ~ ~ .

~~~

Parameter

Quantity

Reference

Molar mass Da; no. of subunits Specific activity (pure enzyme, U mg-’) T; pH optimum Substrate(s)[KM-value(s), mM] Activators (0.5 mM, in %), pH 6

73 200; 2 (identical) 319 (C1Ac-Ala, pH 8.0) 55 “C; pH 7.5 ClAc-Met [1.5], ClAc-Phe (0.7) CO” (151),Zn2+(loo),Mg” (97), Mn2+(37), Ni2’ (27) Cd”, Cu2+,chelators

WI

Inhibitors Sequences and structure: protein sequence: not known expression system: not known

I351 1151

[35] [35]

1351

DNA sequence: not known 3D crystal structure: not known

sapiensLS3,541 and Mus musculus (house mouse)[”] for aminoacylase I1 and sequences from Achromobacter xylosoxidans A-6 rS61 and Alcaligenes faecalis [”I for Daminoacylase. The DNA sequences of several other acylases, notably r-acylases I, from organisms such as Arabidopsis thaliana, Streptomyces coelicolor, Bacillus subtilis and from two human genome project groups have been annotated as aminoacylases but have not been confirmed to possess aminoacylase activity. Regarding threedimensional structures, as of the beginning of March 2001, no structures of aminoacylases were known or under review according to the Protein Data Bank (PDB). 12.3.2.2

Substrate Specificity

An extensive study of the substrate specificity of both enzymes (AA, PKA), especially for the resolution of unnatural and rarely occurring amino acids has been conducted by Whitesides and (Fig. 12.3-2). Both enzymes have an unusually wide substrate specificity with a preference for hydrophobic substrates. N-acylated aliphatic straight-chain amino acids are the preferred substrates for both enzymes, however, the corresponding aliphatic branched-chain amino acids are also readily accepted, especially by the fungal enzyme[’3, 351. N-acylated amino acids with an aromatic side chain are significantly hydrolyzed only by the fungal enzyme[17](Table 12.3-4).The substrate spectrum of AA was even broader than anticipated[’’]. Sulfur- and selenium analogs react at comparable rates, often even faster than the carbon analogs; four to five atoms are the optimum length of the side chain. S-Methyl-L-cysteine gained significance recently as building block for HIV-protease inhibitors [59, “1, L-selenomethionine was described as part of a suitable treatment for Alzheimer’s disease and Parkinson’s syndrome [61]. Another striking difference is that the renal enzyme hydrolyzes dipeptides whereas the mold enzyme does Acylase I has not only been used for the enantioselective resolution of N-acetyl-D,ramino acids to the corresponding L-amino acids but also for substrate-selective resolution of N-acetyl amino acids using the different activity of the enzymatic

72.3 Hydrolysis of N-Acylamino Acids

R'eCOOH

A

HNYR3 0

K HO

H N ~ CH2'\=~

0

d '

C

NH,O

H ~ ,f H,N

K NH(CH&,'

CH3g

CH3CHzCH(CH3)g

poor, 0.01-1 %

0

NHCH2, H~N(CH~)H

R2, R') are expressed relative to the reactivities of the corresponding substituents of the model substrate, N-acetylmethionine (40 mM racemic: R' = CH'SCH2CH2, R2, RZ = H, R' = CH3. b AA only; reactivity with PKA is fair. c PKA only: reactivity with AA is fair. d AA only: reactivity with PKA is poor. e AA only; no reactivity with PKA. f Data for PKA only. g PKA only: no reactivity with AA.

a Reactivities of substituents (R',

Figure 12.3-2.

Reactivities of substituents of acylase

I

745

R2

746

I

12 Hydrolysis and Formation ofC-N Bonds

Comparison o f the relative activity of Aspergillus and pig kidney aminoacylase with different substrates [351.

Table 12.3-4. Substrate

N-chloroacetyl-r-alanine N-chloroacetyl-L-methionine N-chloroacetyl-D,r-norleucine N-chloroacetyl-r-leucine N-chloroacetyl-L-phenylalanine N-chloroacetyl-L-tryptophan N-acetyl-L-glutamic acid N-acetyl-L-aspartic acid N-acetyl-L-glutamine N-acetyl-L-alanine N-acetyl-r-lysine N-dichloroacetyl-glycine N-dichloroacetyl- L-leucine N-dichloroacetyl-D,L-norleucine N-dichloroacetyl-L-alanine a V,

=

Conc. (rnM1

7.1 7.1 2.1 2.1 3.5 2.1 8.2 8.2 8.2 8.2 8.2 4.1 4.1 4.1 4.1

Aspergillus arninoacylase

100“ 400 207 26 325 125 0 0 13 14 3 0 0 4 0.7

Pig kidney arninoacylase

looa 480 120 96 5 0 21 0 4

7 0 1 3 69 2

4.2 N M s-‘.

catalyst towards different N-acetyl-r-amino acids. Martens and Weigel used kidney acylase for the separation of N-acetyl-L-leucineand N-acetyl-L-isoleucine[631. 12.3.2.3

Stability of Acylases

Acylase I from both sources is very stable as a lyophilized powder. In aqueous solution, the resting stability of acylase from Aspergillus oryzae was found to depend much more on pH than on concentration: while at room temperature (25 “C) and standard pH (7.0) the half-life ~ 1 1 was 2 around GO d for concentrations of between 30 and 120 g crude enzyme L-l, the 5 1 / 2 dropped to 45 d at pH 6.5 and to about 30 d at pH 6.0[46].Also, in solubilized form, the fungal enzyme is fairly stable whereas the pig kidney enzyme is sensitive to auto oxidation and therefore should be kept under Tests for operating stability in repeatednitrogen if stored in a solubilized batch mode L9, 17, 36s 641 reveal that acylase from Aspergillus oryzae again fares much better than the porcine kidney enzyme. Tests for operating stability in a continuous reactor with the acylase from Aspergillus 0ryzae[‘~1again demonstrated: - superior stability of AA (616 U kg-’ L-methionine) over PKA (6000 U kg-I L-met), both measured with [Co”] at S X ~ O -M,~ - tighter binding of Zn” vs. Co2+at 5 x M (308 vs. 616 U kg-’ L-met), - that loss of metal, commonly Zn”, is responsible for activity loss and - possibility of reconstitution over a timescale of several hours, whereas the time

constant of leaching is on the order of 48 h, as well as - the option of pulsing divalent metal addition resulting in 477 U kg-’ L-met at M. [Zn”] of 4 x

W40

20

-

C

-

20-

d

Porcine kidney acylase seems to have a different spectrum of activationrG4]: although Co2+activates PKA most strongly, Ca2+is not far behind whereas Zn2+,just like Mg2+ or Fe", does not seem to exert a strong effect. Both enzymes have moderate therrn~stability~~~] and moderate stability in the presence of organic cosolvents13'1 (Fig. 12.3-3). The behavior of aminoacylase both from porcine kidney and Aspergillus sp. towards a wide range of water-miscible cosolvents was investigated by Iborra et al.["I. They found that enzymatic activity can be correlated with the denaturing capacity of the water-cosolvent system. In 1993,a thermostable aminoacylase from Bacillus stearothermophilus was characterized by Sakanyan et al.rs1].The enzyme hydrolyzes N-acyl derivatives of aromatic amino acids preferentially and even has some dipeptidase activity. Its optimal reaction temperature is 70°C; after incubation for 15 min, 90% of the original activity was retained. The authors write that the similarity ofthe B. stearothermophilus enzyme sequence with that of other enzymes such as aminoacylase I, acetylornithine deacetylase and carboxypeptidase G2 suggests a common origin. The aminoacylase from B. stearothermophilus is well characterized: the gene has been completely G71 and studied for catalytic sequenced[51],cloned into E. coli and overexpres~ed[~~~ and stability properties["]: the intrinsic one Zn2+ion per subunit seems to have a predominantly structural role and activity can be restored to the apo-enzymeby Co2+ and particularly by Cd2+(3-foldactivity!) but not by Zn2+.

748

I

12 Hydrolysis and Formation ofC-N Bonds

Conditions: pH 7.5, J = Table 12.3-5. Thermodynamics o f the N-acetyl amino acid 25 "C; xeq calculated at [So] of0.5 M , K,,-data determined from synthesis and hydrolysis reaction. Amino acid

Keq

Xeq

Glycine

4.5 5.6 5.6 10.5 12.5 3.7

90.8% 92.4% 92.4% 95.6% 96.3% 89.2%

Alanine Aminobutyric acid Norvaline Norleucine Methionine

Another thermostable acylase, aminoacylase SK-1, was reported by the Amano Pharmaceutical Comp. The enzyme is isolated from B. stearothemophilus I F 0 12983. It possesses an optimal temperature for reaction at 60 "C and is stable at 70 "C for at least 30 min. The preferred substrates are dipeptides besides the N-acyl derivatives of Met, Phe and Tyr. K. Soda's group has isolated and characterized a which has many thermostable aminoacylase from Bacillus them~glucosidius~~~] similarities to the Aspergillus enzyme, such as metal content and requirements, activity and specificity profile as well as high stability at elevated temperatures and high content of organic solvents and denaturants. Judged by the identity of the organism used for culturing, of the specificity profile and of some enzyme properties (both are identical dimers with molecular mass of 86000 Da), aminoacylase SK-1 and the aminoacylase from Bacillus therm~glucosidius[~~] seem to be the same enzyme. 12.3.2.4

Thermodynamics and Mechanism of the Acylase-catalyzed Reaction

Equilibrium The hydrolysis reaction of N-acetyl amino acids is equilibrium-limited, however, the equilibrium is well on the side of the hydrolysis so that at low substrate concentrations conversion is almost quantitative. For the case of N-acetyl methionine, Wandrey and Flaschel determined the equilibrium constant K defined as in Eq. (1) K=[

acetate][^-Met] [N-Ac-L-M~~]

and found K = 2.75 M at 37 "C and pH 7"l. Then, at 37 "C and [So] = 100 mM, equilibrium conversion x, is 96 % (basedon N-Ac-L-Met),at [SO]= 500 mM, x, = 86 %. The enthalpy of reaction is 7.9 kJmol-' 191. Data for other substrates are listed in Table 12.3-5. More physicochemical data on the N-acetyl amino acid acylase reaction can be found in ref. [641. pti-Dependence The Michaelis constant for hydrolysis is independent of pH in the pH range 6.0-9.5 whereas the pH-dependence of maximum velocity has a bell-shaped profile with the maximum at pH 7.5 and inflection points at pKa values of 6.7 and 8.9[69].

Table 12.3-6.

Substrate specificityof acylase II (aspartoacylase; E.C. 3.5.1.15) from hog

kidney[”]. Substrate

Relative activitv X = chloroacetyl

X = acetyl

looa

N-X-D,L-asparticacid N-X-D,L-ghtamic acid N-X-D,L-methionine N-X-D ,I-alanine N-X-D,L-leucine N-X-D,L-serine a 0.45 pmolesxmin-’ per

526 22

-b

33

-

-

19 26 11

-

mg of N

at 38 “C.

b Not determined

Mechanism The mechanism of acylase-catalyzed reaction has been studied, particularly for porcine kidney acylase [34, 6g--711 . Wh’ile the mechanism of action was contested for some time between a linear mechanism (random uni-bi)[34, 701 and a doubledisplacement, “ping-pong”,mechanism involving a stable intermediate l7ll, it now seems to have been decided that base-catalyzed attack of the carbonyl carbon by water is the rate-determining step followed by a linear sequence involving an E-P1P2complex[34, 69, 701 E 9. (2): “3

I

Recent work on the non-competitive inhibition of both porcine and fungal aminoacylase by a- and p-fluoro- and -hydroxy acids indicated that the active site of the fungal enzyme should interact with the a-substituent of a substrate via an acidic group while the porcine enzyme has a basic group in the corresponding position with which to recognize substrates 17’1.

Enantiospecijicity Acylase I acts on racemates in a highly enantiospecific way to yield L-amino acids exclusively, with ee values in almost all cases, especially with N-acetyl substrates, exceeding 95 %. According to Cahn-Ingold-Prelogrules, L-amino acids correspond to the (S)-configuration,with the exception of L-cysteine which is in the (R)-configuration owing to first stereochemical priority of the thiomethyl group. In general, the amino acid amide enantiomer bearing the larger substituent in the pro-(S)-position is hydrolyzed preferentially [731. 12.3.3

Acylase II (N-Acyl-L-Aspartate Amidohydrolase, Aspartoacylase, E. C. 3.5.1.1 5)

Apart from acylase I, another aminoacylase was found in kidney preparations by fractionation of hog kidney homogenates with ammonium sulfate and acetone [I3]. Whereas acylase I could be enriched and thus partially purified by this procedure the main activity with N-acylated-L-aspartic acid as the substrate was found in another

750

I

12 Hydrolysis and Formation ofC-N Bonds Figure 12.3-4. Enantioselective hydrolysis of N-acetyl-DL-prolineto L-proline catalyzed by proline a~ylase1~~1.

0 N-Acetyl-D,L-proline

II

L-Proline

0 N-Acetyl-D-proline

Comparison of some kinetic and chemical properties of proline acylases from three different microorganisms~'g-221.

Table 12.3-7.

Property

Enantiospecificity Molecular mass Da No. of subunits Molecular mass of subunits (Da) Isoelectric point pH-optimum pH-stability

Temp. optimum Temp. stability Activation by divalent cations Inhibitors

Reactivation of the apoenzyme by divalent cations

a Not determined.

PS.spec. 1' 1'

Rh. rubraPO]

Corn. Testosteroni"'. *'I

L

L

L

597 000 +/- 12000 10-12 55000 5.0 6.0 7.0-8.0 (30 min, 35 "C) 40 "C (10 min, pH 8.0) none phosphate EDTA o-phenanthroline 2,2-dipyridyl Hg2+> Cu2' > ZnZ+> Fe3+ > Ni2+> Pb2+

560000

Mn" > Ca2+ Pb2+> co2+ Zn" > Ba2+

-a -

6.0 6.0-10.0 50 "C 40 "C

none PCMB

cu2+

380 000

+/-

40000

8 45000 +/- 15000 -

6.8 5.0-10.0 (4weeks, room temp.) 65 "C 70 "C (30 min, pH 7.5) none phosphate 2-mercaptoethanol o-phenanthroline PCMB, PHMB Fe" > Hg2+ > cu2+> Zn2+ > Sn2' > Fe3' co2+> Zn2'

72.3 Hydrolysis of N-Acylamino Acids

fraction of the ammonium sulfate precipitates. To distinguish between these two activities, the former fraction was designated acylase I, and the latter acylase II[13]. In contrast to acylase I, acylase [I has a very narrow substrate specificity. Among the Nacetyl derivatives of the twenty proteinogenic amino acids only N-acetyl-L-aspartic acid is hydrolyzed significantly (Table 12.34). Therefore, acylase I1 from kidney preparations was designated as aspartoacylase or N-acyl-L-aspartateamidohydrolase (E.C. 3.5.1.15) and is the enzyme of choice for the resolution of racemic aspartic acid. Comparison ofthe substrate specificity of proline acylases from four different microorganisms "8-211.

Table 12.3-8.

Substrate

Alc. spec. ['I

N-acetyl-L-Pro N-acetyl-D-Pro N-acetyl-L-Ala N-acetyl-o,r-Ser N-acetyl-L-Val N-acetyl-D,r-Val N-chloroacetyl-L-Pro N-chloroacetyl-r-Met N-chloroacetyl-L-Val N-chloroacetyl-r-Leu N-chloroacetyl-L-Phe N-chloroacetyl-L-Tyr N-chloroacetyl-L-Ile N-acetyl-L-Hyp N-formyl-L-Pro N-propionyl- L-Pro N-butyryl-L-Pro N-valeryl-L-Pro N-caproyl-L-Pro N-capryloyl-L-Pro N-caprinoyl- L-Pro N-myristoyl-L-Pro N-palmitoyl-r-Pro N-benzoyl-L-Pro Gly-L-Pro N-Z-L-Proe N-2-Gly-L-Pro N-2-L-Ala-L-Pro N-2-Gly-L-Ala N-2-Gly-r-Pro N-2-Gly-L-Pro L-Leu-Gly-L-Pro

100"

Relative activity R. spec. [''l Rh. rubra Po]

1OOb 0 0.03 0

100

0 172

-

-

13 24

33 23 59 7 0.6 61 123 24 269 2

100' 0 9 0.2 0.2 -

362 17 14 2 1 1

0.5

10 18 29 14 15 9 -

-

4 0 11 -

1

-

217

-

101

-

a 142 pmoles x min-' x mg-'. b 410 pmoles x min-' x mg-I. c 85 pmoles x min-' x mg-'.

d Not determined. e Z: benzyloxycarbonyl.

Corn. testost. 12'1

I

751

752

I

12 Hydrolysis and Formation ofC-N Bonds

12.3.4

Proline Acylase (N-Acyl-L-ProlineAmidohydrolase)

The acylase-catalyzed resolution of N-acyl-D,L-amino acids has some limitations. Although acylase I from porcine kidney and Aspergillus oryzae has a broad substrate specificity and high enantioselectivity, the enzyme does not accept N-acylated substrates where the hydrogen atom at the amide nitrogen is replaced by an alkyl group. Therefore, N-acylated secondary amines such as N-acetyl-proline and Nacetyl-N-alkyl-aminoacids are not hydrolyzed by this enzyme[13,36, 37, 74, 751 as well as aminoacylases from other This gap in the substrate specificity of aminoacylase I was successfully closed with the isolation of acylases which act specifically on N-acetyl-L-prolineand its derivatives[18-22] (Fig. 12.3-4). The enzyme has been isolated from Alcaligenes sp. [18, 21], Pseudomonas sp. [I9], Rhodotorula rubru(20]and Comamonas testosteroniIZ1.2 2 ] . Some kinetic and chemical properties of proline acylases from three different microorganisms are listed in Table 12.3-7. A comparison of the substrate specificity of proline acylases from four different microorganisms is shown in Table 12.3-8. Proline acylase is a relatively large protein with a molecular mass in the range of 380-600 kDa consisting of 8-12 subunits with a molecular mass of 45-55 kDa. The Substrate specificity of proline acylases from Cornomonas testosteroni towards N-acyl-amino acids [771.

Table 12.3-9.

Substrate

N-acetyl-L-praline N-acetyl-o,L-proline N-acetyl-~-thiazolidine-4-carboxylic acid N-acetyl-~-azetidine-2-carboxylic acid N-acetyl-D,L-pipecolicacid N-chloroacetyl-L-proline N-chloroacetyl-o,L-proline N-chloroacetyl-~-thiazolidine-4-carboxylic acid N-chloroacetyl-~-azetidine-2-carboxylic acid N-chloroacetyl-o,r-pipecolicacid N-chloroacetyl-o,~-indoline-2-carboxylic acid

Relative activity

100 78 255 < 50 58 308 290 357 437 507 0

Conversion (“A)

100 54 99 100 49 100 52 100 100 54 0

Substrate specificity of proline acylases from Comamonas testosteroni towards N-alkyl-amino acids [781.

Table 12.3-10.

Substrate

Relative activity

N-chloroacetyl-L-praline 100 N-chloroacetyl-N-methyl-L-alanine 175 115 N-chloroacetyl-N-methyl-o,L-alanine N-chloroacetyl-N-ethyl-o,L-alanine 82 N-chloroacetyl-N-propyl-o,L-alanine 27 N-ch~oroacetyl-N-methyl-~,~-2-aminobutyric acid 10 2 N-chloroacetyl-N-ethyl-o,~-2-aminobutyric acid a Not determined.

Conversion (“A)

100 100 49 -

-

50 -

12.3 Hydrolysis ofN-Acylamino Acids

hCooH u R'

R2

H20

CH3COOH

-

acylase

HN

KCH3 0

IA

N-acetyldehydroamino acid R'

Z%COOH

I

enarnine N H ~

NH3

H20

_2/

R'

I1

spontaneous

$+ COOH

spontaneous

0 a-keto acid

NH irnine

R1

R F ! C O O H NH2 L-amino acid Figure 12.3-5. Coupled enzymatic reaction of a dehydro amino acid acylase with a amino acid dehydrogenase (from[821).

enzyme is not activated by cobalt ions and has a relatively narrow substrate spectrum. The enzyme from Cornamonas testosteroni preferentially hydrolyzes Nacylated L-proline and the N-acetyl derivatives of other cyclic imino acids [771 (Table 12.3-9)and opens for the first time the route to resolution of racemic N-acylated Nalkyl-aminoacids [781 (Table 12.3-10).Among the proteinogenic amino acids, only the N-acetyl derivatives of r-proline and L-alanine are hydrolyzed to a significant efient[21s22, 771 12.3.5 Dehydroarnino Acid Acylases

A new acylase was found in strains of Breuibacterium sp. by H ~ m m e l [ ~1' ' ' . in 1987, catalyzing the hydrolysis of acetamidocinnamate (ACA) and was named acetamidocinnamate acylase (ACA acylase).A similar, just as enantiounspecific acylase, N-acetyldehydroleucine acylase (ACL acylase), catalyzing N-acyl hydrolysis of branched-chain dehydroamino acids (N-acetyl-dehydrovaline, -1eucine and -isoleucine) was isolated and characterized from Zoogloea ramigera by Kittelmann and Kular8'. 82]. The hydrolysis product in both cases, an enamine, first undergoes

I

753

754

I

12 Hydrolysis and Formation ofC-N Bonds Table 12.3-11. Comparison of the substrate specificityO f D-, and L-aminoacylasein Streptornyces tuirus and Streptomyces olivaceus[28]. Relative activity

5. olivaceus

5. tuirus Substrate

D-

N-acetyl-phenylglycine N-acetyl-leucine N-acetyl-phenylalanine N-acetyl-methionine N-acetyl-tyrosine N-acetyl-valine N-acetyl-tryptophan N-acetyl-alanine N-acetyl-glutamicacid N-acetyl-asparticacid N-acetyl-arginine N-acetyl-proline

100 1130 1004 682 522 314 117 102 69 20 14 0

L-

-

70 8 167 0 29 0 38 0 0 32 0

L-

D-

100 957 723 448 307 261 68 69 38 0

5 0

120 15 208 2 61 0 92 10 21 72 0

a Not determined.

spontaneous rearrangement to the ketimine which is then deaminated spontaneously to the a-keto acid. The dehydroamino acid acylase reaction can be coupled with reductive amination by amino acid dehydrogenases such as PheDH, LeuDH or even AlaDH, respectively, to produce L-amino acids[821 (Fig. 12.3-5). L-Phenylalaninehas been produced continuously from ACA with the help of ACA acylase in an enzyme membrane reactor (EMR) with a space-time-yieldof 277 g L-' d-'[831.With ACL acylase, L-Ieucine was produced at 123-180 g L-' d-' in the same reactor set-up [82].The dehydroamino acid substrates can be prepared conveniently, either from 2-halogencarboxylic acid esters [841,or, specifically in the case of ACA, via the acetamidomalonic ester route by reaction with benzyl halogenides I*']. Apart from the L-specific acylases from kidney and Aspergillus strains it has been shown that similar aminoacylases are widely distributed in microorgani s m ~ [ ~"1. ~ However, - ~ ~ , from the viewpoint of costs, those acylases which are practically employed for large scale industrial purposes, are restricted to the enzyme from Aspergillus oryzae (see Sect. 12.3.7). 12.3.6

D-Specific Aminoacylases

D-Specific aminoacylases have been found in Pseudornonas sp. [33, 87-901 , Streptomyces sp. [28] and Alcaligenes sp. [29-32, 'l]. The first investigations on the use of D-specific aminoacylases for the synthesis of D-amino acids were carried out by Kameda and coworkers. They demonstrated that a strain of the genus Pseudomonas hydrolyzed Nbenzoyl-D-amino acids in addition to N-benzoyl-L-aminoacids [871. The partially purified enzyme was employed to synthesize D-phenylalanine from N-benzoyl-D,Lphenylalanine ["I and D-phenylglycine was synthesized from N-chloroacetyl-D,Lphenylglycine with the crude enzyme preparation Sugie and Suzuki conducted an extensive screening among soil samples as well as 8"

12.3 Hydrolysis of N-AcylarninoAcids

Comparison of the substrate specificityof purified D-aminoacylasesfrom three strains ofAlcaligenes SP.[~’* 30v 321.

Table 12.3-12.

Relative activity of strain DA181

DA1 L-

Substrate

D-

N-acetyl-methionine N-acetyl-phenylalanine N-acetyl-norleucine N-acetyl-leucine N-acetyl-tryptophan N-acetyl-alanine N-acetyl-asparagine N-acetyl-all0isoleucine N-acetyl-valine N-acetyl-phenylglycine N-acetyl-tyrosine N-acetyl-asparticacid N-acetyl-glutamicacid N-acetyl-lysine N-acetyl-arginine N-acetyl-histidine N-acetyl-serine N-acetyl-glycine N-chloroacetyl-phenylalanine N-chloroacetyl-norleucine N-chloroacetyl-isoleucine N-chloroacetyl-alanine N-chloroacetyl-valine N-chloroacetyl-serine N-formyl-methionine N-formyl-phenylalanine N-benzyloxycarbonyl-methionine Glycyl-leucine . .

100 65

52 14 14 8 -

6 3 -

-

33 -

D-

L-

MI-4 L-

D-

100 80 38 (DJ) 17 5 1 0 1 1

0 -

0 (DJ) 0 -

0 (D,L) 0 68 66 (W) 40 (D,L) 38 18 5 (DJ) 56 (DJ) 35 2

20

a Not determined.

among 420 strains of the genus Streptomyces and 16 strains of the genus Streptoverticillium from type culture collections and isolated four Streptomyces strains producing a D-specific aminoacylase suitable for the production of D-phenylglycine[”I. Since the bacteria also produced an t-aminoacylasethe D-aminoacylase had to be separated from the L-specific enzyme by ion exchange chromatography prior to use. Thus, Dphenylglycine could be produced from N-acetyl-D,L-phenylglycinein 99.9 % optical purity. Table 12.3-11lists the substrate specificity of the D- and L-aminoacylases from two Streptomyces species. Microbial D-aminoacylases have also been found in different species and strains of the genus Alcaligenes. The enzyme has been isolated, purified and characterized from Alcaligenes denitii..cans subsp. x y l o s ~ x y d a n s [30, ~ ~321,, Alcaligenes denitnfcans[”] and Alcaligenes f a e ~ a l i s [ ~Several ~]. companies, all of them Japanese, have filed applications for D-aminoacylases The substrate specificity of the Daminoacylases from these strains is shown in Table 12.3-12.

I

755

756

I

12 Hydrolysis and Formation of C-N Bonds Table 12.3-13. Enantioselective deprotection of N-protected D,L-aminoacids by D-aminoacylase from Alcaligenesfaecalis DA-1 Substratea

N-Ac-D,L-methionine N-Ac-D,L-methionine(in 50% DMSO) N-Ac-D,L-leucine N-Ac-D,L-leucine(in 50% DMSO) N-Ac-D,L-phenylalanine N-Ac-glycine N-n-Butyl-D,L-methionine N-Bz-D,L-methionine N-Bz-D, L-leucine N-Bz-D,L-phenylalanine N-Bz-D,L-norleucine N-Bz-~,~-2-amino-n-butyric acid N-Z-D,r-methionine N-Z-D,r-leucine N-Z-D,L-norleucine N-Z-~,~-2-amino-n-butyric acid

Reaction time

Conversion

e e of D-amino

(h)

(W

acid (77)

50.0 53.0 49.3 30.7/48.9 49.9 10 45 47.2 48.1 50 43.9 33.8 32.6 32.6 12.8 15.8

100 30 100 100 100

2

15 2

15 2

2 2

10 10 10 10 10 10 10 10 10

100 89 99 100 53 80 99 100 51 77

a Ac; acetyl; Bz; benzoyl; 2; benzyloxycarbonyl.

As with the D-aminoacylases from Streptomyces sp. the enzymes from Alcaligenes strains have a preference for hydrophobic N-acetyl-aminoacids. In this respect, they are similar to the L-specific acylase I from kidney preparations and Aspergillus sp. The Alcaligenesfaecalis enzyme prefers the N-acyl-D-aminoacid derivatives from Met, Phe and Leu ["I. If a high-affinity substrate residue occupies the hydrophobic side-chain pocket the enzyme even deacylates D-Met methyl esters or N-Ac-D-Met-Xaadipeptide derivatives. Two D-aminoacylases have been described that resemble the L-specific acylase I1 from kidney, which only hydrolyzes the N-acyl derivatives of L-aspartic acid. The Dspecific counterpart of acylase 11, N-acetyl-D-aspartate deacetylase, has been isolated from Alcaligenes xylosoxydans subsp. xylosoxydans L3lI. The same strain produces an aminoacylase which specifically hydrolyzes N-acyl derivatives of D-glutamic acid [311. The latter N-acetyl-D-glutamatedeacetylase has also been found in Pseudomonas sp. (331. All microorganisms producing D-aminoacylases commonly produce L-aminoacylases as well. Therefore, to reach high optical purity of the D-amino acids produced from the respective N-acetyl-D,L-amino acids, the D-aminoacylases have to be separated from the r-aminoacylases (Table 12.3-13). However, this is a disadvantage in view of an industrial application since additional purification steps lead to more expensive enzymes and thus add costs to the whole production process. This is one of several reasons why it is widely accepted today that the production of D-amino acids by enzyme-catalyzedhydrolysis of D,r-hydantoinsseems to be more promising than the ~-aminoacylaseroute via N-acetyl-D,L-aminoacids. The enzyme-catalyzed synthesis of D-amino acids from the respective D,L-hydantoins is described in Chapter 12.4.

12.3 Hydrolysis of N-Acylamino Acids

I

757

Non-proteinogenic amino acids

Proteinogenicamino acids

Y O O H NH2

qCooH a-Aminobutyric acid

Alanine

NH2

WHrooH yCooH Norvaline

Phenylalanine

/

NH2

/tCOOH

TcooH Norleucine

Valine

NH2

NH2

Leucine

~

0 /

Methionine

"

"

" 0-Benzylserine "

NH2

o""'ycooH S-Benzylcysteine

/

NH2

Tryptophan

Q-J-yCOOH

N H

~

NH2

mHYH

Tyrosine

HO

Figure 12.3-6. L-amino acids prepared in bulk quantities by acylase I resolution of N-acetyi-DL-aminoacids.

12.3.7

Acylase Process on a Large Scale

The most established method for enzymatic L-amino acid synthesis is the resolution of racemates of N-acetylaminoacids by acylase I from Aspergillus o r p e fungus. The N-acetyl-r-aminoacid is cleaved to yield L-amino acid whereas the N-acetyl-D-amino acid does not react. After separation of the L-amino acid through ion exchange chromatography or crystallization, the remaining N-acetyl-D-amino acid can be

758

I

12 Hydrolysis and Formation of C-N Bonds

racemized by acetic anhydride in alkaline solution or by adding a racema~e['~] to achieve very high overall conversions into the L-amino acid. N-acetyl-D,L-aminoacids are conveniently accessible on a laboratory as well as an industrial scale through acetylation of w-amino acids with acetyl chloride or acetic anhydride in a SchottenBaumann reaction['8]. As was demonstrated in the synthesis of 13C-~-methionine, the acylase process has a virtually closed material balance because almost 99.5 % of the amino acid components can be retrieved after The acylase is relevant for enzyme reaction engineering along two different lines as follows. With the aminoacylase process, Tanabe Seiyaku commercialized the first immobilized enzyme reactor system ever in 1969 after running the process in batch mode since 19541'. '1. Enzyme from Aspergillus oryzae fungus was immobilized by ionic binding to DEAE-Sephade~[~l. In a fued-bed reactor, the reaction is carried out at elevated temperature to produce r-methionine, L-valine, and L-phenylalanine. Costs are significantly lower than in a batch process with native enzyme. Tanabe started up more fixed-bed reactor processes with immobilized enzymes: L-aspartic acid with aspartase in 1973 and L-malic acid with fumarase one year At Degussa, several enzyme membrane reactor (EMR) set-ups are in operation covering six orders of magnitude from laboratory via pilot stage to full production scale; the process has been scaled up to an annual production level of several 100 tons of enantiomerically pure a-amino acids, mostly L-methionine and 1-valine['*I (Fig. 12.3-6).The enzyme membrane reactor is a recycle reactor operated as a CSTR of up to 200. For both pilot and large-scale with a recycle ratio Frecycle/Finflux operation, the necessary membrane area is configured into polysulfone hollow-fiber modules with a molecular-weightcut-off of 10 kDa resulting in a rejection rate of the 73 kD-acylase far in excess of 99.9%. Aminoacylase has also been immobilized on a nylon membrane ["I. While the half-life as measured by thermal stability, of 161 d is superior to the data for immobilized acylase (65d) 1'1 or soluble enzyme in an EMRr'l, reactor productivity at 0.136 L-valine kg/L-'d-' is lower than that for DEAE-Sephadex-immobilizedacylase (0.5 kg/L-'d-')['] or that for a membrane reactor (0.35 kg/L-'d-l)[']. Results on operational stability of both acylases in a recycle reactor at constant conversionF41 with reaction conditions close to intended large-scale conditions demonstrated much better stability of the Aspergillus enzyme, while renal enzyme is not stable enough for long-term operation [64.651. Moreover, on the process scale achieved today the supply of renal acylase is insufficient, so that fungal acylase is used almost exclusively nowadays, especially since the price per unit is comparable.

References 0. Schmiedeberg, Arch. Exp. Pathol. Pharrnakol. 1881,13, 379-392. 2 I. A. Smorodinzev, 2. Physiol. Chem. 1922, 124, 123. 3 J. P. Greenstein, M. Winitz in: Chemistry of the Amino Acids, Vol. I, John Wiley & Sons 1

Ltd., London, New York, 1961, pp 715-760, and references cited therein. 4 T. Tosa, T. Mori, N. Fuse, I. Chibata, Enzymlogia 1966,31,214-224. 5 T. Tosa, T. Mori, N. Fuse, I. Chibata, Agric. Bid. Chem. 1969,33,1047-1052.

References I 7 5 9

I. Chibata, T. Tosa, T. Sato, T. Mori, Meth. Enzymol. 1976,44,746-759. 7 Tanabe Seiyaku, Ger. Pat. 2828194,1982. 8 Tanabe Seiyaku, U K Pat. Appl. 2082188 1982. 9 C. Wandrey, E. Flaschel, Adu. Biochem. Eng. 1979, 12,147-218. 10 Degussa/GBF, US Pat. 4,304,858,1981. 1 1 W.Leuchtenberger, M. Karrenbauer, U. Plocker, Ann. N. Y Acad. Sci. (Enzyme Eng. 7) 1984,434,78-86. 12 W. Leuchtenberger, U.Plocker, in: Enzymes in Industry (Ed.: W. Gerhartz), VCH, Weinheim, 1990,130-141. 13 S. M. Birnbaum, L. Levintow, R. B. Kingsley, J. P. Greenstein, J . Bid. Chem. 1952, 194, 455-40. 14 Y. Endo, Biochim. Biophys. Acta 1978,523, 207-214. 15 Y. Endo, Biochim. Biophys. Acta 1980,628, 13-18. 16 Amano Pharmaceutical Co. Amano Enzymes, Technical Bulletin 70,1970. 17 I. Gentzen, H.-G. Loffler, F. Schneider in: Metalloproteins. Structure, Molecular Function and Clinical Aspects, A u t u m n Meet. German Biochem. Soc. (Ed.: U. Eser), Thieme, Stuttgart, FRG, 1979, pp. 270-274. 18 Noda Sangyo Kagah Kenkyusho,Jpn. Pat. 55-7015,1980. 19 M. Kikuchi. L. Koshiyama, D. Fukushima, Biochim. Biophys. Acta 1983,744,18&188. 20 Daicel Chemical Industries, Jpn. Pat. 6474987,1989. 21 Degussa AG, US Pat. 5,120,652.1992. 22 U. Groeger, K. Drauz, H. Klenk. Angew. Chem. 1990, 102,428-429; Angav. Chem. Int. Ed. Engl. 1990, 29,417-419. 23 Banyu Pharmaceutical, Eur. Pat. Appl. 020 1039,1986. 24 Amano Pharmaceutical, Jpn. Pat. App/. 184552,1987. 25 Amano Pharmaceutical, Jpn. Pat. Appl. 052732,1988. 26 H.-Y. Cho, K. Tanizawa, H. Tanaka, K. Soda, Agnc. Bid. Chem. 1987,51,2793-2800. 27 H.-Y. Cho, K. Tanizawa, H. Tanaka, K. Soda, J. Biochem. 1988,103,622-628. 28 M. Sugie, H. Suzuki. Agric. Bioi. Chem. 1980,44,1089-1095. 29 Daicel Chemical Industries, /pa. Pat. 645488,1989. 30 M. Moriguchi, K. Ideta, Appl. Enu. Microbiol. 1988,54,2767-2770. 6

31 K. Sakai, K. Imamura, M. Goto, I. Hirashiki, M. Moriguchi, Agnc. Bid. Chem.

1990,54,841-844.

32 K. Sakai, T.Obata, K. Ideta, M. Moriguchi,

J . Fern. Bioeng. 1991,71,79-82.

33 K. Sakai, K. Oshima, M. Moriguchi, Appl.

Enu. Microbiol. 1991, 57, 2540-2543.

34 K. H.Rohm, R. L. van Etten, Eur. J. Biochem.

1986,160,327-332.

35 I. Gentzen, H.-G. Loffler, F. Schneider, 2.

Naturfrsch. 1980, 3%. 544-550.

36 H. K. Chenault, J. Dahmer, G. M. Whitesides, J . Am. Chem. Soc. 1989, 111,

6354-6364.

37 S.-C. Fu, S. M. Bimbaum, J. Am. Chem. Soc.

1953,75,918-920. S. M. Bimbaum, J. P. Greenstein, J. Am. Chem. Soc. 1954,76,6054-6058. 39 H.-G. Lofler, F. Schneider, Bid. Chem. Hoppe-Seyler 1987, 368,481-485. 40 I. Gilles, H.-G. Loffler, F. Schneider, 2. Naturforsch. 1981, 36c, 751-754. 41 M. Bakker, TU Delfi/Netherlands, personal communication 42 R. Marshall, S. M. Birnbaum, J. P. Greenstein, J. Am. Chem. SOC.1956,78, 4636-4642. 43 E. Kumpe, H.-G. Loffler, F. Schneider, 2. Naturforsch. 1981, 36c, 951-955. 44 I. Gilles, H.-G. Loffler, F. Schneider, 2. Naturforsch. 1984,39c, 1017-1020. 45 W. Kordel, F. Schneider, Z. Naturforsch. 1977,326,337-341. 46 A. S. Bommarius, Habilitation thesis, RWTH Aachen, 2000. 47 M. R. Cook, B. J. Burke, D. L. Buchhagen, J. D. Minna, Y. E. Miller, J. Bid. Chem. 1993, 268,17010-17017 48 M. Mitta, I. Kato, S. Tsunasawa, Biochim. Biophys. Acta 1993, 1174, 201-203. 49 M. Jakob, Y. E. Miller, K. H. Rohm, Bid. Chem. Hoppe-Seyler 1992,373, 12271231. 50 M. Mitta, H.Ohnogi, A. Yamamoto, I. Kato, F. Sakiyama, S. Tsunasawa,J. Biochem. 1992,112,737-742. 51 V. Sakanyan, L. Desmarez, C. Legrain, D. Charlier, I. Mett, A. Kochikyan,A. Savchenko, A. Boyen, P. Falmagne, A. Pierard, N. Glansdorff, A p l . Environ. Microbiol. 1993,59,3878-3888. 52 P. Curley, D. van Sinderen, FEMS Microbiol. Lett. 2000, 183, 177-182. 53 R. Kaul, G. P. Gao, K. Balamumgan,

38 S.-C. Fu,

760

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72 Hydrolysis and Formation ofC-N Bonds

R. Matalon, Nature Genetics 1993, 5, 118-123 54 R. Kaul, K. Balamurugan, G. P. Gao, R. Matalon, Genomics 1994, 21, 364-370. 55 M. A. Namboodiri, A. Corigliano-Murphy, G. Jiang, M. Rollag, 1. Provencio, Brain Res. Mol. Brain Res. 2000, 77, 285-289. 56 M. Wakayama, Y. Katsuno, S. Hayashi, Y. Miyamoto, K. Sakai, M. Moriguchi, Biosci. Biotechnol. Bioeng. 1995,59, 2115-2119. 57 C. S. Hsu, W. L. Lai, W. W. Chang, Y. B. Yang, Y. C. Tsai, unpublished work (PDB entry AAK15530). 58 A. S. Bommarius, K. Drauz, K. Giinther, G. Knaup, M. Schwarm, Tetrahedron: Avmm.1997,8,3197-3200. 59 Y. Kiso in: Aspartic Proteinases: Structure, Function, Biology, and Biomedical Implications (Ed.: K. Takahashi), Plenum Press, New York, 1995, p. 413 60 Y. Kiso, Biopolym. (Peptide Science), 1996, 40,235-244 61 J. Birkmayer, Europ. Application EP 0 345 247 A2,1989. 62 W. Kordel, F. Schneider, Hoppe-Seylers 2. Physiol. Chem. 1975, 356,915-920. 63 J. Martens, H. Weigel, Liebigs Ann. Chem. 1983,2052-2054. 64 C. Wandrey, Habilitationschrij, TU Hannover, Germany, 1977. 65 A. S. Bommarius, K. Drauz, H. Klenk, C. Wandrey, Ann. N. Y Acad. Sci. (Enzyme Eng. 11) 1992,929,126-136. 66 J. L. Iborra, J. M. Obon, A. Manjon, M. Canovas, Biotechnol. Appl. Biochem. 1992, 15, 22-30. 67 M. We& G. J. Palm, K.-H. Rohm, Biol. Chem. Hoppe-Seyler, 1995,376,643-649. 68 Amano Pharmaceutical Comp., Japanese patent J P 62044181,1987. 69 I. Y. Galaev, V. K. Svedas, Biochim. Biophys. Acta 1982,701, 389-394. 70 L. Otvos, E. Moravcsik, G. Mady, Biochem. Biophys. Res. Commun.1971, 44, 1056-1064. 71 E. S. Chukrai, D. Lauceniece, A. Arens, 0. M. Poltorak, Vestn. Mosk. Univ.Ser. 2, a i m , 1979,20,118-122. 72 T. Tamura, Y. Oki, A. Yoshida, T. Kuriyama, H. Kawakami, H. Inouye, K. Inagaki, H. Tanaka, Arch. Biochem. Biophys. 2000, 379, 261-266. 73 J. P. Greenstein, M. Winitz in: Chemistry of the Amino Acids, Vol. 2, John Wiley & Sons Ltd., London, New York, 1961, 1763-1767.

J. Kamphuis, W. H. J. Boesten, Q. B. Broxterman, H. F. M. Hermes, J. A. M. van Balken, E. M. Meijer, H. E. Schoemaker, Adv. Biochem. Eng. Biotechnol. 1990, 42, 133-186. 75 S. Kang, Y. Minematsu, Y. Shimohigashi, M. Waki, N. Izumiya, Mem. Fac. Sci., Kyushu Univ., Ser. C 1987, 61-68. 76 A. S. Bommarius, K. Drauz, U. Groeger, C. Wandrey in: Chirality i n Industry (Eds.: A. N. Collins, G. N. Sheldrake, J. Crosby), John Wiley & Sons Ltd., London, New York, 1992, pp. 371-397. 77 K. Drauz, U. Groeger, M. Schilfer, H. Klenk, Chem.-Ztg. 1991, 115,97-101. 78 U. Groeger, K. Drauz, H. Klenk, Angew. Chem. 1992,104,222-224, Angew. Chem. lnt. Ed. Engl. 1992, 31, 196-197. 79 Degussa AG/Ges. F. Biotechnol. Forsch. U S Pat. 4,877,734,1989. 80 W. Hummel, H. Schiitte, E. Schmidt, M.-R. Kula, Appl. Microbiol. Biotechnol. 1987, 27, 283-291. 81 Degussa AG/Ges. f. Biotechnol. Forsch. U S Pat. 5,134,073, 1992. 82 M. Kittelmann, M.-R. Kula, ]. Ferm. Bioeng. 1992,73,99-107. 83 W. Hummel, H. Schiitte, E. Schmidt, C. Wandrey, M.-R. Kula, Appl. Microbiol. Biotechnol. 1987,26, 409-416. 84 F. Effenberger,T. Beisswenger, Angew. Chem. 1982, 94, 210, Angew. Chem., Int. Ed. Engl. 1982, 21, 203. 85 W. Windus, C. S. Marvel,]. Am. Chem. Soc. 1930,52,2575-2578. 86 Y. Yamazaki, W. Hummel, M.-R. Kula. 2. Naturforsch. 1987,426, 1082-1088. 87 Y. Kameda, E. Toyoura, Y. Kimura, H. Yamazoe, Nature 1952, 169, 1016. 88 Y. Kameda, E. Toyoura, Y. Kimura, Nature 1958,181,1225. 89 Y. Kameda, E. Tayoura, Y. Kimura, Pharm. SOC.Jap. 1958, 78, K. Matsui, I. 202. 90 Y.-C. Tsai, C.-P. Tseng. K.-M. Hsiao, L.-Y. Chen, Appl. Environ. Microbiol. 1988, 54, 984-989. 91 Y.-B. Yang, C.3. Lin, C.-P. Tseng,Y.-J.Wang, Y.-C. Tsai, Appl. Environ. Microbiol. 1991, 57, 1259-60. 92 K. Isobe, Y. Hirose, 1999,jpn. Application 09286147, publishedunder CA 11103887 A (Amano). 93 W. Hibino, I. Onishi, S. Abe, K. Yokozeki, 74

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

12.4 Hydrolysis and Formation ofHydantoins

1999,Jpn. Application 092 80073, published under CA 111 1592 A (Ajinomoto). 94 S. Tokuyama, 1999, Eur. Application EP 0 950 706 A2 (Daicel Chemical Industries). 95 H.-P. Chen, S.-H. Wu, K.-T. Wang, Bioorg. Med. Chem. 1994,2,1-5. 96 H.-P. Chen, S.-H. Wu, Y.-C. Tsai, Y.-B. Yang, K.-T. Wang, Bioorg. Med. Chem. Lett. 1992,2, 697-700. 97 Takeda Chemical Industries Ltd., Eur. Pat. Appl. 0,304,021,1989.

0. V. Sonntag, Chem. Rev. 1953,52, 237-416. 99 I. Chibata, T. Tosa, T. Sato, Appl. Biochem. Biotechnol. 1986, 13, 231. 100 I. Chibata, T. Tosa, I. Takata, Trends Biotechnol. 1983, I, 9. 101 I. Takata, T. Tosa, I. Chibata, Appl. Biochem. Biotechnol. 1983,8, 31. 102 A. S. Bommarius, M. Schwarm, K. Drauz, Chimica Oggi 1996, 14(10),61-64. 98 N.

12.4

Hydrolysis and Formation of Hydantoins Markus Pietzsch and Christoph Syldatk 12.4.1 Classification and Natural Occurrence of Hydantoin Cleaving and Related Enzymes

Abbreviations Cit citrulline DTT dithiothreitol EDTA ethylenediaminetetraaceticacid HIC hydrophobic interaction chromatography MTEH methylthioethylhydantoin IEX ion exchange chromatography IMH indolylmethylhydantoin Phg phenylglycine SEC size exclusion chromatography Thienylala thienylalanine 0-Me-Ser 0-methylserine The compound hydantoin was discovered by von Baeyer in 1861 by reduction or hydrogenation of allantoin, which is a naturally occurring cyclic amide in many or plants [l]. The systematic terms for “hydantoin” are “imidazolidine-2,4-dione” “2,4-diketotetrahydroimidazole”.In the literature, a wide spectrum of various 5-mono- and 5,5’-disubstitutedhydantoin derivatives of industrial and pharmacological interest is described, of which 5-monosubstituted hydantoins may be regarded as cyclic ureides of a-amino acids. They are obtained by Strecker synthesis and are important precursors, e. g. in the industrial production of D,L-a-aminoacids. The 5,5’-disubstitutedhydantoin derivatives have been of pharmacological interest since the 1930s,e. g. for the treatment of Parkinson’s disease. Figure 12.4-1gives a survey of the different methods for the chemical synthesis of hydantoins. Detailed reviews on their chemical syntheses and applications are given in references[*]

I

761

762

I

12 Hydrolysis and Formation ofC-N Bonds R-CHO

+ HCN +

(NH,),CO, Strecker Synthesis

R’YcooE: NCO

II

0

Figure 12.4-1.

R-CHO

+

H,N

,f(,

NH,

+ CO

n=O.1 * = Lor D or D.L

Chemical syntheses of hydantoins.

and13],on their structures in solution, and in the solid state in With the increasing interest in new amino acid derivatives, recent investigations on their chemical synthesis concentrates on the development of “one-pot-syntheses”of the corresponding hydantoin derivatives, e. g. by carbonylation of aldehydes in presence of urea derivatives[’I. Many of the hydantoin derivatives are substrates for enzymatic reactions. It has been known since the 1940s that some microorganisms are able to grow on D,L5-monosubstituted hydantoins as the sole C- and/or N-source in a mineral salt medium, often hydrolyzing only one enantiomer of a racemic mixture, and that even enzymes from plant and animal sources are able to hydrolyze and close the hydantoin ring. Various enzymes, so called hydantoinases, facilitate the hydrolysis of the hydantoin ring system in an initial reaction step. The biosynthesis of these enzymes often has to be induced by adding specific compounds during the growth of the microorganisms. The so-formed hydantoinases may have different substrate specificities and in general are selective in forming L- or D-N-carbamoylamino acids (= hydantoic acids). The hydantoinases can often be found in combination with highly stereoselective N-carbamoylamino acid amidohydrolases (N-carbamoylases), which catalyze the further hydrolysis of the hydantoic acids to the free amino acids in an irreversible reaction. In some cases a hydantoin-racemase is involved as a third enzyme. Together, these three enzymes accomplish the total conversion of racemic ~,~-5-monosubstituted hydantoin derivatives into the corresponding enantiomerically pure D- or L-amino acids. This cascade of reactions, whether located in whole cells or carried out using isolated enzymes is called the “hydantoinase-process”.

72.4 Hydrolysis and Formation of Hydantoins

I

763

L-specific

E

a

(L-carbamoyl-aminoacid

I

Il-hydantoin(

@ 4

IPhydantoinI

ID-carbamoyl-aminoacic

R ~ c o o CO2,NHa dH b a

C02,NHs

m o y l a s e l

N"t

[ L-amino acid1

ID-amino aid1

Figure 12.62. Reaction scheme for the enzymatic cleavage of o,L-S-monosubstituted hydantoin derivatives to the corresponding D- or L-amino acids.

Figure 12.4-2 shows the general reaction scheme for the enzymatic cleavage of D,L5-monosubstituted hydantoin derivatives to the corresponding D- or L-amino acids. The great advantages for industrial use of the hydantoinase-process are based on the fact that potentially 100% conversion and a 100% optically pure amino acid can be obtained at the same time if a racemic substrate is used. Until the mid 1990s in most cases, wild type strains, resulting from traditional screening methods (for a review see: reference L6]), were used as whole cell biocatalysts. Detailed reviews on the use of free or immobilized whole cell systems for hydantoin cleavage were given in references [3, '. 1'. More recent activities are summarized in this chapter and concentrate on the use of recombinant free or immobilized enzymes (see Sect. 12.4.2-12.4.G), fusion proteins (see Sect. 12.4.7), specially designed recombinant whole cell biocatalysts (see Sect. 12.4.4) or the optimization of enzyme properties by directed evolution (see Sect. 12.4.7). The hydantoinases belong to the E.C. 3.5.2 group of cyclic amidases"], which is shown in Table 12.4-1. Of this group, four enzymes are original hydantoinases, because their substrates are naturally occurring hydantoin derivatives: carboxymethylhydantoinase (E. C. 3.5.2.4), allantoinase (E. C. 3.5.2.5), l-methylhydantoinase (E. C. 3.5.2.14), and carboxyethylhydantoinase. All other enzymes listed have natural occurring cyclic amides as substrates (e.g. barbiturate, 5,G-dihydrouracil, 5,6-dihydroorotate). From recent investigations on DNA- and amino acid sequences of the different cyclic amidases and subsequent phylogenetic analyses, it is known today that most of these enzymes not only share a number of highly conserved regions and invariant amino acid residues["], but form a protein superfamily and are the product of a divergent evolution["]. Although most of them only share limited sequence homology (identity < 15%) and therefore are only distantly related, it can be shown:

764

I

12 Hydrolysis and Formation ofC-N Bonds Table 12.4-1.

Hydantoinases and cyclic arnidases[’].

Recommended name

Other names

Barbiturase Dihydropyrimidinase

barbiturate amidohydrolase 5,6-dihydropyrimidine amidohydrolase carbamoylaspartic acid ~-5,6-dihydro-orotate amidohydrolase dehydrase ~5-carboxymethylhydantoin amidohydrolase allantoin amidohydrolase penicillin amido-a-lactam p-lactamase, hydrolase Cephalosporinase 4-imidazolone-5-propionate amidohydrolase 5-oxo-~-proline pyroglutamase amidohydrolase creatinine amidohydrolase

Dihydroorotase Carboxymethylhydantoinase Allantoinase Penicillinase

D-hydantoinase

Imidazolone propionase 5-Oxoprolinase (ATP-hydrolyzing) Creatininase r-Lysine-lactamase 6-Aminohexanoate-cyclic dimer hydrolase 2,s-Dioxopiperazine hydrolase 1-Methylhydantoinase (ATP-hydrolyzing) Carboxyethylhydantoinase Indolylmethylhydantoinase

Systematic name

E. C.-number 3.5.2.1 3.5.2.2 3.5.2.3 3.5.2.4 3.5.2.5 3.5.2.6 3.5.2.7 3.5.2.9 3.5.2.10 3.5.2.11 3.5.2.12 3.5.2.13

1-methylhydantoin amidohydrolase L-5-carboxyethylhydantoin amidohydrolase 5-indolylmethylhydantoin amidohydrolase

3.5.2.14

1. that most of them are members of a broad set of amidases with similarities to ureases and build u p into a protein superfamily[’’, ‘*I, whereas

2. the ATP-dependent hydantoinases (see Fig. 12.4-3)are not related, and 3. that they share a metal-binding motif consisting of conserved histidine residues, which seems to have a n important role to play i n structure and activity“’.

“8

131.

The differences i n enantioselectivity, often used for the classification of hydantoinases based o n their biotechnological value, therefore do not reflect the evolutionary relationship of the different hydantoinases, which are forming a more diverse group of enzymes than was assumed earlier (for more details see reviews : referencesIl4I and [131). This protein superfamily probably has its origin i n the prebiotic conditions of the primitive earth, where N-carbamoyl-u-amino acids rather than free a-amino acids are supposed to be the first synthons for prebiotic peptides i n the evolution today[”]. This section will have a detailed look at the occurrence of the different cyclic amides i n nature and their physiological role i n various metabolic pathways. Allantoin is widely distributed in nature and is a n important metabolite in the degradation of purine nucleotides (see Fig. 12.4-4).Allantoin occurs i n all organisms that do not have uric acid as the final product of their purine degradation pathways, and is the substrate for the enzyme allantoinase or 5ureidohydantoinase (E. C.

72.4 Hydrolysis and Formation ofHydantoins

-

Relations based on: Structure homology

A

Superfamily of 'Amidases involved in nucleotide metabolism' Hydantoinase from Arthrobacteraursscens DSM 3145

Dihydropyrimldlnase Allantolnase Dihydroorotase

Sequence homology

Family of: ATP-dependent cyclic amidases'

no relationship

N-Methyihydantoinase L-Oxoprolinase

I

urease others: Adenine deaminase Adenosine deaminase Aminoacylase AMP deaminase Aylphosphatase Cytosine deaminase Chlorohydrolase Formylmethyldehydrogenase imidazolonepropionase Phosphotriesterase

NOsequence information available about:

Carboxyethylhydantoinase

Figure 12.4-3. The evolutionary relationship of hydantoinases derived from sequence and structural similarity. Enzymes i n bold letters are h y d a n t ~ i n a s e s " ~ ] .

3.5.2.5), which can be found in microorganisms, plants and animals, either in combination with an allantoicase (E. C. 3.5.3.4) or an allantoate amidohydrolase (E. C. 3.5.3.9). The latter hydrolyzes allantoin to urea and glyoxylic acid, which are the final products of purine degradation in fishes. A recent paper describes the purification of this enzyme from Bacillusfastidiosus[l6]. In the 1960s, different groups[17.1" described the microbial enzyme as inducible and (+)-specific.Besides allantoin other inducers are compounds with a free ureido group such as N-carbamoyl-L-asparagine,N-carbamoyl-L-aspartate(the corresponding D-compounds were ineffective), hydantoate (i.e. N-carbamoylglycinate) and diureid~methane[~']. Information on the substrate specificity of allantoinases for other hydantoin derivatives is limited but D,L-S-arninohydantoin was shown to be accepted, albeit poorly, as a substrate I2O]. Non-stereoselectiveallantoin hydrolysis and association of the allantoinase with a cofactor-independentallantoin racemase (E. C. 5.1.99.3) has been reported[20,211, so that some microorganisms are also able to use (-)-allantoinas a substrate. An excellent review of these purine as well as pyrimidine degrading enzymes was given by Vogels and van der Drift[22]. The natural function of the carboxymethylhydantoinase(E. C. 3.5.2.2) is postulated to be the hydrolysis of 5-carboxymethylhydantoin,which is described to be the product of a non-enzymatic cyclization of N-carbamoyl-L-aspartic acid [23, 241 and to occur as a side-productin the metabolism of the pyrimidine nucleotide dihydroorotic acid[25].This enzyme often occurs in combination with a ureidosuccinase (E.C. 3.5.1.7)["I, which catalyzes the cleavage of the resulting N-carbamoyl aspartic acid to L-aspartic acid (see Fig. 12.4-5). L-5-Carboxymethylhydantoinwas first isolated after incubating orotic acid, a six-membered cyclic amide, with crude cell extracts of the anaerobic bacterium Clostridium oroticum 12', 26].

I

765

766

I

12 Hydrolysis and Formation ofC-N Bonds

A third naturally occurring hydantoin, ~-S-carboxyethylhydantoin, was first isolated by Brown and Kies[271from the urine of rats, monkeys and humans after being fed '4C-histidine, and it was postulated to be a by-product in the histidine degrada~ ~ ] by induction experiments, tion pathway shown in Fig. 12.4-6. A k a m a t ~ u [proved, that the L-carboxyethylhydantoinasefrom a Bacillus brevis strain, also described by 0

Adenine

Guanine

0

0

bNA" H

Xanthine dehydrogenase

Hypoxanthine

Xanthine

+H201 Xanthine dehydrogenase

2 [HI

Uric acid

Oz+ 2Hz0 Uricase C 0 2 + H,O,

H2N

o""0

-

HNKNH 0 (-)-Allantoin

Racemase Allantoin-

H2N

p)-,fo

HNKNH 0 (+)-Allantoin

Allantoinase

72.4 Hydrolysis and Formation ofHydantoins

I

767

NH3+ CO,

H,O

Hz:”FCOOH

A

1 0

Ureidoglycine A = Allantoate amidohydrolase

Allantoic acid

Hzo

NH3

urea

Allantoicase

S-Ureidoglycolicacid

R-Ureidoglycolicacid

\

R-Ureidoglycolase or Allantoicase

H

OACOOH

I

~

2 NH,

+ CO,

Urease or Allophanate pathway

Figure 12.4-4.

Purine degradation pathway via allantoin i n

Tsugawa et al. 12’] and Hassall and GreenbergL2’I for the formation of L-glutamic acid from ~,~-S-carboxyethylhydantoin, was not able to hydrolyze L-carboxymethylhydantoin and consequently it is not identical to the former enzyme described above. This enzyme has no E. C. number at present. The six-membered ring systems 5,G-dihydropyrimidine, 5,G-dihydrouracil and 5,G-dihydrothyminecan be hydrolyzed by the enzyme dihydropyrimidinase (E. C. 3.5.2.2),which is involved in the degradation of pyrimidine nucleotides. This widely spread, inducible catabolic enzyme is strictly D-selective in contrast to the L-selective dihydroorotase (E. C. 3.5.2.3),which is involved in the opposite anabolic pathway (see above). Another name often used in the literature for the dihydropyrimidinase is Dhydantoinase, because it is also able to hydrolyze ~,~-S-monosubstituted hydantoin derivatives with high activity. Both reactions are shown in Fig. 12.4-7. Natural cyclic amides such as 5,G-dihydrouracil, uracil and 5,G-dihydrothymineas well as hydantoin, 5-methylhydantoin and 5-hydroxymethylhydantoinare effective inducers for enzyme biosynthesis (for a more detailed review on induction experiments see referencef3I).In some cases, the dihydropyrimidinase (D-hydantoinase)is associated with an N-carbamoyl-D-aminoacid amidohydrolase (D-carbamoylase)and a hydantoin racemase [301. The previously proposed identity of the D-N-carbamoylase with the P-ureidopropionase(E. C. 3.5.1.G),which was assumed to be responsible for the hydrolysis of N-carbamoyl-P-alanine (see Fig. 12.4-7)[31-351 is no longer valid since the investigations of Ogawa et al. on different aerobic bacteria showed that the

768

I

12 Hydrolysis and Formation ofC-N Bonds

HN

0,

H 0AOrotic 2 Cacid O O H

1

Methylene blue

f...',.

i

Cytochrome C

I

L-5,6-Dihydroorotic acid

Carboxymethylhydantoinase

Hooc-)---COOH

K

HN

0

Hooc-kf

HZ0

NH2

N-Carbamoyl-L-aspartic acid

'6 Non enzymatic cyclization

HNYNH 0

L-5-Carboxymethylhydantoin

i

Ureidosuccinase

NH,

+ CO, H20

L-Aspartic acid Figure 12.4-5.

Metabolism of orotic acid and dihydroorotic acid[22.241.

12.4 Hydrolysis and Formation of Hydantoins

769

Urocanate

L-Histidine

Hoocw

Hooc-fcooH A H+NH H2O

L-Formiminoglutamicacid

I

HN\//N

L-4-Imidazolone-5-propionicacid

L-5-Carboxyethylhydantoin

J.

HooC-YCooH HNK 0N H z NCarbamoyl-glutamic acid Figure 12.4-6. reaction.

Histidine degradation pathway and carboxyethyl hydantoinase-catalyzed

770

I

12 Hydrolysis and Formation ofC-N Bonds

"w"

""KNH 0

5,6-Dihydrouracil

D,L-5-rnonosubstituted hydantoin

("'

5,6-Dihydropyrirnidinase or " D-Hydantoinase"

P C O O H

RvCOOH

HN 0 KNH2 P-Ureidopropionic acid

0 NCarbarnoyl-D-amino acid

Figure 12.4-7. Analogy between dihydropyrirnidinase- and D-hydantoinasecatalyzed reactions.

L-specific carbamoylase from Pseudomonas putida I F 0 12 996 also hydrolyzes pureidopropionate[14, 3 G ] . The enzyme from Pseudomonas putida IF0 12996 was shown to be strictly L-selective and to be active on L-N-formyl-and also on L-N-acetylalanine13']. In this context it may be of interest that Runser and Meyer described a Dhydantoinase with no dihydropyrimidinase activityL3'] and Ogawa et al. reported on the occurrence of a D-N-carbamoylasewith no relation to a ~-hydantoinase[~*~. Nevertheless, the dihydropyrimidinase seems to be closely related to the barbiturase (E.C. 3.5.2.1), which is able to hydrolyze barbituric acid[39](Fig. 12.4-8). The difference between barbituric acid and the natural compounds uracil and thymine is the presence of a keto-group instead of a methyl- or a hydrogen-group in the 6-position of the ring. Barbiturase was first detected by Hayashi and K ~ r n b e r g r ~ ~ ] in bacteria of the genera Mycobacterium and Corynebacterium and postulated to catalyze a sidereaction in the degradation of pyrimidines. Unfortunately, there are no further data in literature on the substrate specificity and the stereoselectivity of this enzyme, which would allow comparison with the D-hydantoinase, but Kautz and

Barbituric acid Figure 12.4-8.

Barbiturase catalyzed reaction[39].

Malonic acid

Urea

12.4 Hydrolysis and Formation ofHydantoins

I

771

ATP

rCOOH H3CyN).( NH Creatinine

0 I -NMethylhydantoin

1 -WMethylhydantoinase

NH2

0 NCarbamoylsarcosine NCarbamoylsarcosinehydrolase

CO,

+ NH,

rCOOH H3C0N)(NH2

C O H, N ,, H3C

H

NH Creatine

Sarcosine

Figure 12.4-9. 1-Methyl hydantoinase- and N-carbamoylsarcosine-amidohydrolase-catalyzed reactions in creatinine metabolism in bacteria.

Schnackerz were able to show that beef liver dihydropyrimidinase is also able to hydrolyze barbituric acid, although only with low activityIm1. Two other hydantoinases are described in the literature, which have not yet been listed in the Enzyme Nomenclature[’]. Siedel et al.L4l],Yamada et al.[42,431 and Ogawa et al. found a new ATP-dependent 1-methylhydantoinasewith additional nucleoside-triphosphatase activity [451 in different bacteria. This inducible enzyme, which was also shown to act on unsubstituted hydantoin and 5-methylhydantoin14’1, is involved in the degradation of creatinine after its deimination in the 2-position to I-methylhydantoin, resulting in N-carbamoylsarcosine (N-carbamoyl-N-methylglycine) [42,431 (see Fig. 12.4-9).It is associated with a so-called D-N-carbamoylsarcosine hydrolase [431, which eventually hydrolyzes N-carbamoylsarcosine to free sarcosine. Both enzymes can be used for monitoring creatinine levels in blood L4l]. Nishida et al.[46],Syldatk et al.l4’. 481, Yamashiro et al.L4’, ”I, and Yokozeki et al. [51-531 found new L-5-arylalkylhydantoinases and a N-carbamoyl-L-aminoacid amidohydrolases (L-N-carbamoylase),which are involved in the L-selective cleavage of 5-arylalkylhydantoinsand could be most favorably induced by D,L-S-indolylmethylhydantoin or its N-3-methylated derivative(1’. The natural functions of these enzymes are not yet known, while one of the associated N-carbamoyl-L-amino acid amidohydrolases (L-N-carbamoylase)was also shown by Syldatk et al. to be reactive on N-formyl-L-aminoacids [541. In this strain both, hydantoinase and L-N-carbamoylase were shown to occur in combination with a hydantoin racemase[’, 55. 561. Resting cells were used for the industrial production of L-amino acids from D,L5-monosubstituted hydantoin derivatives as shown in Fig. 12.4-2LS71. Concerning their structure, cyclic imides are closely related to dihydropyrimidines and hydantoins. The metabolic transformation pathway for cyclic imides in microorganisms (see Fig. 12.4-10)was studied by Ogawa et al.[”, I’ in Blastobacter sp. and

772

I

72 Hydrolysis and Formation ofC-N Bonds

lrnidase I

nCOOH

HOOC

Succinate

I

TCA cycle

I

Acetyl-CoA

-

COOH Pyruvate

in different aerobic bacteriaL6']. The enzyme involved in this reaction, a so called imidase, was also found to hydrolyze dihydropyrimidines [I4]. Activity for the enzymatic cleavage of disubstituted hydantoins useful in the synthesis of a-,a-disubstituted amino acids was recently detected in crude enzyme extracts from the plant Lens esculenta[6'. 62] and in papain by Rai and Taneja[63]. Of all the enzymes described above, at present only the D-hydantoinase- and the Larylalkylhydantoinase processes are of significance for use in organic synthesis, in particular for the production of natural and non-natural optically pure D- and Lamino acids, and will be discussed in more detail in the following sections.

I

12.4 Hydrolysis and Formation ofHydantoins

773

12.4.2

D-Hydantoinases- Substrate Specificity and Properties

Since the early 1950s it has been known that the inducible catabolic enzyme dihydropyrimidinase (E. C. 3.5.2.2) plays an important role in pyrimidine metabolism[23s31, 33, 39, G4-GG] and is widespread in nature. The natural substrates of this enzyme, which were also reported to be inducers, are 5,G-dihydrouracil and 5,G-dihydrothymine. Both compounds are important intermediates in the degradation of pyrimidine nucleotides. The dihydropyrimidinase-reactionis described to be strictly D-specific and to have a wide substrate specificity (see Fig. 12.4-11). In 1970 and -0

-sw HNYNH HNYNH do 0

H

*

I

W

Q O

0

0

\

O

W

0

h

,"YE

0

HNYNH

""YNH

W

""K""

HNYNH 0

0

C

O

0

%-)+ ""IfE 0

""YNH 0

w ""If"" 0

O \>O

w ""K""

0 2 " W

""YNH 0

0

9;

0$ 0

oA,Xo

""If"" 0

Q

JX0 ox>o H

""If"" 0

Q oA,Xo

O++O

H

/

H

H

Substrates accepted by different D-hydantoinase preparations from mammalian and microbial "1. Figure 12.4-11.

'

""f 0

774

I

72 Hydrolysis and Formation of C-N Bonds

1973, Dudley et al. were the first to publish on the D-selective cleavage of 5-phenylhydantoin to N-carbamoyl-D-phenylglycineby a mammalian enzyme and on the spontaneous in vivo racemization of the residual isomer['^, "1. In 1975, Cecere et al."1' published on the enzymatic production of other N-carbamoyl-D-aminoacids starting from chemically synthesized ~,~-5-monosubstituted hydantoin derivatives using a partially purified fraction of the dihydropyrimidinase from calf liver. They were the first to stress that this enzyme might find an industrial application for the preparation of optically active D-amino acids as the so called "D-hydantoinase" (see Fig. 12.4-7). In 1978, the same group published on the production of various Ncarbamoyl-D-amino acids using an immobilized calf liver dihydropyrimidinase preparation[70r711. Other publications have reported on the occurrence of D-hydantoinases in plant cell cultures[72].Rai and Taneja published on the use of a plant enzyme from Lens esculenta immobilized to DEAE-cellulose for the same purIn other publications, Wallach et al. ["I, Brooks et al. [731 and Kautz and Schnackerz14'] gave detailed reports on the isolation and characterization of the dihydropyrimidinase from beef liver. Table 12.4-2 gives a short overview of the purification procedures and characteristic properties of these mammalian enzymes. The beef liver dihydropyrimidinase consists of four subunits and every active enzyme molecule contains four Zn(")~ations[~~] which are tightly bound (& > 1.33 x lo' M-'). In addition to 5,G-dihydrouracil, glutarimide, thiohydantoin and barbituric acid are also accepted as substrates, but with low reaction rates I4O1. In the late 1970s the group of Yamada et al. in Japan postulated that in microorganisms the reason for the wide spread ability to hydrolyze D-selectively D,L5-monosubstituted hydantoin derivatives was the existence of an enzyme called "Dh y d a n t o i n a ~ e " 751. [ ~ ~With ~ the increasing interest in the production of D-phenylglycine and D-p-OH-phenylglycine,since then several publications have described Dselective hydantoinases isolated from various microorganisms as Pseudomonas ~triata[~'], Pseudomonasfluorescens DSM 84["], Pseudomonas sp. AJ-l1220[35], Arthrobacter crystallopoietes AM2[77],Agrobacterium sp. IP-I 671[37. 781, in anaerobic microorganism~[~'], Pseudomonas sp. KBEL 101["], Agrobacterium turnefaciens["I, thermoPseudomonas d e s m o l y t i ~ u m [ ~Bacillus ~], sp, Cs41, Bacillus philic microorganisms stearothermophilus SD-1 Is', "1 and Bacillus circulans LS71. Runser and co-workers described a D-hydantoinase of an Agrobacterium sp. with remarkably high tem"1. Soong et al. perature and pH stability but no dihydropyrimidinase were recently able to show that D-hydantoinase from Blastobacter sp.Al7p-4 also is able to hydrolyze cyclic imides with bulky substituents to the corresponding halfamides and postulated that this enzyme may also function in cyclic imide metabolism in addition to pyrimidine metabolism ["I. New screening methods for isolation of n-hydantoinase-producing microorganisms were described by Morin et al. using a continuous cultivation systemI,'[ and by LaPointe et al. using a polymerase-chainreaction-amplifiedDNA probe to detect D-hydantoinase-producingmicroorganisms by direct colony hybridization['l]. A survey of the isolation and some characteristic data on some of the bacterial enzymes, which seem to be rather similar to the dihydropyrimidinases from mammalian tissues (Table 12.4-2) and plants, is given in Table 12.4-3.

12.4 Hydrolysis and Formation of Hydantoins

I

775

Table 12.4-2.

Purification and characteristicproperties of D-hvdantoinasefrom animal cells

Source

Reference Purification steps

Yield (%) Purification factor Purity Optimal pH Metal ion requirements

Molecular mass Subunits

Acetone powder from beef liver

Catalase fraction from beef liver

WI

[731

acid and heat hydrophobicchromatreatment, ammonium tography or preparasulfate and acetone tive electrophoresis precipitation

Acetone powder from beef liver ~401

heat treatment, ammonium sulfate precipitation,chromatography on chelating and DEAESepharose

25 200

13 24.2

80 % 8.2 Mn2’ and Mg2+(only when dihydrouracilis the substrate!)

homogeneous no data given Zn2+and Co2+

186 homogeneous 8-10 one Zn” per subunit

226 000 Da 4x56 500 Da

217 000 Da 4 x 5 4 000 Da

44

Figure 12.4-11 gives a survey of the substrates accepted by the different dihydropyrimidinase or n-hydantoinase preparations The differences between the enzyme preparations from mammalian and microbial sources are discussed in more detail in reference L3], but D-hydantoinases or dihydropyrimidinases, respectively, seem to have the following in common: (i) a wide substrate specificity, (ii) metal dependence and (iii) that they are strictly D-specific. Preferably, cyclic amides are hydrolyzed at pH values around 8.5. Furthermore, most of the enzymes are also described to be able to catalyze the hydantoin formation: the optimal pH of this reaction is neutral or weakly acidic. In 1983 the first gene sequence of a D-hydantoinase derived from thermophilic Bacillus sp. LU 1220 and its overproduction in Escherichia coli HB 101 was published[”]. Not until 1994 were cloning, sequencing and expression of a Dhydantoinase gene from Pseudomonas putida DSM 84 in Escherichia coli reported[”], shortly followed by a paper on cloning, sequencing and expression of a thermostable D-hydantoinase from Bacillus stearothermophilus NS 1l22A[”4]. The same was described for the strain Bacillus stearothermophilus SD-1 by Lee et al. in 1997[951.The same group reported that the C-terminal region of the D-hydantoinase was not essential for catalytic activity but affected the oligomeric structure of the In 1998, Chien et al. described the cloning, sequencing and expression of the Dhydantoinase gene from Pseudomonas putida CCRC 12857 in Escherichia coli[”]. Molecular cloning and sequencing of a cDNA encoding dihydropyrimidinase from rat liver was reported by Matsuda et al. [981, and the complete sequencing of a 24.6 kB segment of yeast chromosome XI including homologies to D-hydantoinases by Tzerma et al. “J91. D-Phenylglycine and n-p-OH-phenylglycineare important side chain moieties in the synthesis of semisynthetic penicillins and are produced in several thousand tons per year using the hydantoinase loo].The different methods that this

776

I

72 Hydrolysis and Formation of C-N Bonds

HO

1

D,L-5-pHydroxyphenylhydantoin

1

’

- co,

+ HO ,

Snamprogetti-Process

Kaneka Process

Recordati-Process

- Immobilized dihydropyrimidinase

- Immobilizedresting cells of

- Immobilized resting cells of

from calf liver

Bacillus brevis with D-hydantoinaseactivity

- Reaction conditions: pH 8.0, 30°C

I

- Reaction conditions: pH 9.0, 30°C

Agrobacteriurn radiobacter with D-hydantoinaseand L-Ncarbamoylase activity - Reaction conditions: pH 9.0, 30°C

HO

NGarbamoyl-D-p-hydroxyphenylglycine

D-p-Hydroxyphenylglycine

Industrial production of D-4-hydroxyphenylglycine acids by t h e D-hydantoinase process. Figure 12.4-12.

reaction has been realized in industrial application in recent years can be seen in Fig. 12.4-12. In the 1970s, the company Snamprogetti first reported on the use of the beef liver dihydropyrimidinase immobilized on an ion exchanger for the continuous production of D-phenylglycine[70, 71], while the company Kanekafuchi was reported to use

12.4 Hydrolysis and Formation of Hydantoins

resting cells of a Bacillus sp. containing D-hydantoinaseactivity only[’001.Because of missing D-N-carbamoylase activity or the instability of this enzyme in resting microbial cells, the decarbamoylation of the resulting D-N-carbamoylaminoacid is often performed chemically by treatment with HN02. Because of the high stability of the D-hydantoinase it is possible to use immobilized resting cells, which can be applied repeatedly. With the increasing interest in products other than D-phenylglycine and D-POHphenylglycine, the companies Recordati and Degussa reported on the use of resting cells of an Agrobacterium radiobacter with high activities for both the D-hydantoinase and D-N-carbamoylase[loo,loll. The advantage of this process in comparison with the methods mentioned above is not only the environmental friendly “one pot production’’ of D-amino acids without use of HN02‘ but the possibility of also producing Damino acids, which are unstable against treatment with this acid (e.g. D-tryptophan, D-citrullineor D-pyridylalanine)(for the production of D-citrulline from L-ornithine see Fig. 12.4-13). Nevertheless, the main problem of using resting cells in a “one pot process” still seems to be the stability of the D-N-carbamoylase (see e.g. reference[”]), which is discussed in Sect. 12.4.3.Therefore, a series of papers from the 1990s concentrated on: the optimization of the chemoenzymatic D-hydantoinasecatalyzed production of D-N-carbamoylphenylglycine and ~-N-carbamoyl-4-hydroxy-OH-phenylglycine as the enhanced chemical decarbamoylation of D-N-carbamoylphenylglycine by its interfacial solubilization under micellar conditions; the repeated use of a commercially available covalently immobilized D-hydantoinase at high substrate concentrations [Io2],the repeated use of a thermostable D-hydantoinase from Bacillus stearothermophilus SD-1 immobilized on DEAE-cellulose resin[’03],the mass production of the same enzyme in Escherichia coli using a constitutive expression system[95];the application of numerical modeling for optimization of a complex medium for Dhydantoinase production from Agrobacterium radiobacter NRRL B 11291 [‘041; the modeling, simulation and kinetic analysis of a heterogeneous reaction system for the to the corresponding D-N-carbamoyl conversion of ~,~-4-hydroxy-phenylglycine amino the use of a so called “pressure swing reactor” for the same as well as on the racemization of the remaining substrate enantiomers [lo7I. 12.4.3

D-N-Carbamoylases - Substrate Specificity and Properties

In some cases, D-hydantoinases are described as being associated strictly with Dspecific N-carbamoyl-D-aminoacid amidohydrolases (D-N-carbamoylases).One natural role of these enzymes was discussed as being the P-ureidopropionase (E.C. 3.5.1.6), which catalyzes the decarbamoylation of P-ureido propionic acid in pyrbut with the recent information on its imidine metabolism (see Fig. 12.4-7), stereo~electivity[~~] and its DNA and amino acid sequences, this previously proposed h o m o l ~ g y [is~no ~ longer ~ ~ ~ ]clear. Various D-N-carbamoylases were purified from rat liver as well as from microbial

I

777

1126, 127. 128, 1301

177, 1591

Molecular mass (Da) Subunits (Da)

257 000 4 x 60 000

[761

1 0.53 homogeneous 55 <40 9.0 5.5-8.5 Fe”

10 mol Zn2’per mol of active enzyme, but Mn” and Co2+Enhance the enzymatic activity 200 000 230 000 4x49680 4x60 000

8.8-9.3

77 52.6 homogeneous 50

5-cyanoethyl-hydantoin

I351

11 220

Pseudomonas sp. AJ

190 000

8.0-9.0 6.0-7.0 Fe”, Co”


G 200), crystallization 3 300 homogeneous 45-55

8.0

63 27 crude enzyme 55

protamine sulfate and IEX (DEAE-Toyopearl) ammonium sulfate precipitation, IEX (DEAEcellulose), hydroxyl apatite- and SEC (Sephadex

Hydantoin

1751

Pseudomonasjluorescens Pseudomonas striata

DEAE-Streamline, HIC (Phenylsepharose), HIC (Phenylsepharose), SEC (Sephacryl S-400) Mono Q and preparative electrophoresis

N-3-Methyl-o,~-S-indolylmethylhydantoin

DSM 3745

dihydrouracil, hydantoin and various w-5-monosubstituted hydantoins protamine sulfate and ammonium sulfate precipitation, IEX (DEAEcellulose), HIC (Phenylsepharose), Mono Q , gel

Arthrobacter aurescens

and L-hydantoinases.

poietes DSM 20 117

D-

Arthrobacter crystallo-

Purification and characteristic properties of microbial

filtration Yield (“A) 5 Purification factor 20 homogenous Purity Optimal temperature (“C) 50-60 Temperature stability (“C) 4 0 Optimal pH 8.2-9.2 pH stability 6.5 Zn2’ Metal ion requirements

Purification

Reference Inductor

Source

Table 12.4-3a.

U

>

S

m

2

s9

S

.:3

F

a

Q

-=z

d

t u

0 U

-

250 000 4x62 000

3 30 homogeneous 60 <60 10.0 5.0-10.0

200 000 4x53 000

212 000 4x53 000

1.5 50 homogenous 65 <60 8.0 5.5-11.0 Mn” Mg2+,Mn”, Co”, Ni2’ Ni2’, Mg” 126 000 2 x 54.000

12.4 243 homogeneous 75 <60 8.0-10.0 8.5-9.5 Mnzf. C02+.Ni2+

Yield (%) Purification factor Purity Optimal temperature (“C) Temperature stability (“C) Optimal pH pH stability Metal ion requirements enhance the enzymatic activity enhance the enzymatic activity Molecular mass (Da) Subunits (Da)

137. 881

uracil protamine sulfate and ammonium sulfate precipitation, heat treatment, IEX (DEAE-Sephadexand Trisacyl, HIC (octyl-Sepharose) 9 965 homogeneous 60 < 70 10.0 7.5-10.5

Agrobacterium sp. IP-l 671

methylthioethylhydantoin uracil heat treatment, Sephadex DEAE-Sephacel,HIC G-50, DEAE-cellulose,HIC (phenylsepharose), SEC (phenylsepharose), Fractogel (Sephacryl S-200 HR), Mono Q and superose-12

[89, 1601

hydantoin ammonium sulfate fractionation, Q-Sepharose, heat treatment, HIC (phenylsepharose, preparative gel electrophoresis)

Blastobacter sp. A1 7p-4

Bacillus circulans ~ 7 1

Bacillus stearothermophilusSD-1

and L-hydantoinases.

[85. 861

D-

Reference Inductor Purification

Purification and characteristic properties o f microbial

Source

Table 12.4-3b.

U \o U

$

I -=0 .

P

lu

4

780

I

72 Hydrolysis and Formation ofC-N Bonds

0

L-Ornithine

H2"

""KN" 0

""K"" 0

4

Agrobacterium radiobacter

CO, + NH,

9

D-Citrulline Figure 12.4-13.

Production o f o-citrulline from L-ornithine by means o f

Agrobacterium radiobacter.

cells. In contrast to D-hydantoinases (see above), induction and stability of these enzymes seem to be problemati~[~I. Meyer and Runser reported that both Dhydantoinase and D-N-carbamoylase were found to be highly inducible by the addition of non-metabolizable thiolated hydantoins or pyrimidines to the culture medium of Agrobacterium sp. 1-671[lo']. Rat liver D-N-carbamoylase was isolated by Caravaca and G r i ~ o l i a [ ~ The ~ ] .same authors also found it in the supernatants of liver homogenates of dogs, pigeons and rabbits. In microorganisms, D-N-carbamoylase activity was detected in various strains of Agrobacterium sp.[32, 109-1111 , B1astobacter sp. A17p-4[ll2I,Clostridium u r ~ c i l i c u m [ ~Cornamonas ~1, acid~vorans[~'], Pseudomonas putida 77 [431 and Pseudomonus sp. AJ-l1220[75]. Induction of enzymatic activity during growth was done either by addition of N-carbamoyl amino acids or pyrimidine and hydantoin derivatives. Some of the enzymes were purified and characterized as shown in Table 12.4-4. The enzyme isolated from rat liver and the inducible Clostridium D-N-carbamoylase are both postulated to be involved in the degradation of pyrimidines [331. With only a few compounds having been tested as substrates for these enzymes, they are

34 000

Subunits (Da) 34 285 (calculated)

no details given

68 000

none

65 < 55 "C 7.0 7.0-9.0 no details given

GO < 40 "C 7.0 7.0-9.0

Purity Optimal temperature ("C) Temperature stability Optimal pH pH stability Metal ion requirements Molecular mass (Da)

Purification factor

Yield ("5%)

Purification

30-35 < 45 "C 7.4-7.8

crude enzyme

none none 84 000 (determined by no details given native gel filtration) no details given no details given

7.4-7.6 6.2-9.0

52

clarified crude extract

clarified crude extract

18

MnC12, ammonium sulfate and acetone precipitation, hydroxyl apatite chromatography

clarified crude extract was used for experiments

clarified crude extract

N-carbamoyl-P-alanine

strain was genetically engineered - no data are given on this

11101

1331

urea, N-carbamoyl-phenylglycine, N-carbamoyl-phenylglycine (for the wild strain) Q-Sepharose FF, chelating heat treatment, HIC (PheSepharose, Superose 12 nylsepharose), ammonium (procedure for the recom- sulfate precipitation, DEAE-Sepharose (data for binant enzyme) the recombinant enzyme) 34 (for the recombinant 12.3 (data for recombinant enzyme) enzyme) 20 (for the recombinant 3.9 (data for recombinant enzyme) enzyme) homogenous

11111

1113, 1161

Reference Inductor

N-carbamoylo-phenylglycine

Clostridium uracilicum

Agrobocterium radiobacter Agrobacterium sp. KNK712 Agrobacterium sp. NRRL B11291 (after expression in (BEECHAM-strain) Escherichia colr)

Purification protocols and characteristic properties of microbial D-N-carbamoylases.

Source

Table 12.4-4a.

3 x 4 0 000

111000

none

40 "C < 40 8.0-9.0 6.5-9.5

homogeneous

119

36

ammonium sulfate precipitation, IEX (DEAE-Sephacel, MonoQ), HIC (Phenylsepharose)

N-carbamoyl-0-alanine

1381

Comamonos sp. E 222c

m -4

-

%

4 z 2 2 2. 5

3

2.

on 3

a

0

a

2

-=z 9

A

h,

-

12 Hydrolysis and Formation ofC-N Bonds

Purification protocols and characteristic properties o f microbial o-N-carbamoylases.

Table 12.4-4b.

Source Reference Inductor Purification

Yield (%) Purification factor Purity Optimal temperature ("C) Temperature stability ("C) Optimal pH pH stability Metal ion requirements Molecular mass (Da) Subunits (Da)

Blastobacter sp. A1 7p-4

Pseudomonas sp. AJ-11220

Pseudomonas putida 77

1351

1431

5-cyanoethylhydan- 1-methylhydantoin toin ammonium sulfate pre- IEX (DEAE Toyo- ammonium sulfate precipitation, IEX cipitation, DEAE-Sepha- pearl) cel, HIC (Phenylsephar(DEAE-Cellulose), ose), Sephadex G150, crystallization Mono Q 2.3 37

36 17

63.2 27.4

homogeneous 55 < 50

crude enzyme 55

homogeneous

8.0-9.0 6.0-9.0

7.0

no details given 120 0000 3x40 000

no details given no details given no details given

37 < 40 7.0-8.0 6.0-7.0

no details given 102 000 4 x 2 7 000

obviously different from the other microbial D-N-carbamoylaseslisted in Table 12.4-4. The N-carbamoylsarcosine amidohydrolase from Pseudomonas putida 77 is reported to have its biological function in creatinine metabolism [431. The D-Ncarbamoylases of the various Agrobacterium sp. and the Pseudomonas sp. Aj-11220 are likely to be identical. They have a wide substrate specificity in common, for a survey see Fig. 12.4-14, and hydrolyze only the D-enantiomers of aliphatic and aromatic hydantoic acids r3', '131 . The main problems of this enzyme seem to be (i) its instability and its rapid inactivation in absence of a reducing agent["'] probably caused by an oxidation of an SH-gro~p["~] and (ii) its inhibition by ammonium i ~ n s [ ' ' ~Grifantini ]. et al. were able to prove the role of the cysteine 172 out of five cysteines for enzyme activity by site-directed mutagenesis while Nanba et al. were able to obtain a more thermotolerant D-N-carbamoylase by substitution of Pro 203 by Leu in the gene from Agrobacterium sp. KNK712 before expression in Escherichia c~li[~''I.For stabilization, the same group immobilized the enzyme by glutaraldehyde coupling to Duolite A-568, a macroporous phenol formaldehyde Kim and Kim tried to overcome limitations in the production of D-p-OHphenylglycine with resting cells of Agrobacteriurn sp. 1-671by adsorptive removal of the ammonium ions with a silicate c0rnplex[''~1and proposed the optimized ratio between D-hydantoinaseand D-carbamoylase of about 1 : 3 based on mass for further process optimization('"]. As discussed before, there is a lot of interest in microbial biocatalysts with highly active D-hydantoinase- and D-N-carbamoylase-activity for the direct synthesis of HNOz-sensitiveD-amino acids used as chiral synthons in the production of pharma-

I

12.4 Hydrolysis and formation ofkfydantoins

h

rcooH Y C O O H

\o>COOH

HNyNH2 HNTNH2

W

C

O

O

b C O O H

Y C O O H

clq -0

>COOH

ANTNH2

COOH

HNKNHz 6

HNKNH2 6

I

COOH

COOH

Q

HNyNH2HNyNH2 HNTNHz HNTNH2 COOH

T C O O H ""\COOH

"KNH2

\o&)COOH

HNyNH2HNyNH2 HNTNHz CI

)--GOOH

Ho*COOH

HNyNH2 HNyNHz HNyNHz HNyNHz

-S rCOOH

H

783

O Y C O O H I

HNyNHz

H o q C O O H

\GOOH

COOH

HNyNH2

HNyNH2

HNKNH2

6 HNyNHz HNTNH2 HNyNHz HNyNH2 HNyNHz HNyNHz H

HOOC

\COOH

Ho-$COOH

>COOH

HO R C O O H

COOH

f

h

C

O q HNTNHzHNyNH2HNyNH2 HNyNH2HNTNHz

O

O

H

0

+GOOH

C -:O '"OH

H

~ '\COOH N

~

COOH "-COOH

&COOH

Figure 12.4-14. Substrates accepted by microbial o-N-carbamoylases of Agrobacterium radiobacter and Pseudomonos sp. AJ-11220[35f' 1 3 ] .

ceutical drugs and intermediates. For the synthesis of peptides in particular, a great variety of D-amino acids and derivatives are highly desirable molecules. Recently, cell free extracts of Blastobacter sp. A17p-4 were used for the preparation of optically active D-p-trimethylsilylalanine from the corresponding D,L-carbamoyl amino acid[118]and several biocatalysts (isolated enzymes as well as whole cells) have been compared with respect to stereoselectivityfor the hydrolysis of D,L-S-trimethylsilylhyd a n t ~ i n " ~Cell ~ ] . free extracts of Blastobacter sp. A17p-4 were shown to distinguish stereoisomers of hydantoins not only at the a-carbon but also at the 0-carbon of Ncarbamoyl-a,0-amino acids [120].

HNyNH2

784

I

72 Hydrolysis and Formation of C-N Bonds

12.4.4 L-Hydantoinases - Substrate Specificity and Properties

In the 1960s, Tsugawa et al.[28]were able to isolate strains of Pseudomonas, Micrococcus, Aerobacter, Achromobacter, and Bacillus that were capable of producing Lglutamic acid from ~,~-5-carboxyethylhydantoin by L-5-carboxyethylhydantoinase. Bacillus breuis ATCC 8185 was the first microorganism used for bioconversion of a racemic hydantoin derivative to an L-amino acid in the case of L-glutamic acid with a yield of 90%. In 1988, Yamashiro et al.I4', 501 reported on an L-hydantoinase from Bacillus brevis AJ 12299. This Bacillus L-hydantoinase requires ATP and Mg2+,Mn" or K' as cofactors and acts selectively on r-configured substrates. The optimal reaction conditions for the hydantoin cleavage are pH 8.0 and 50 "C. Only a few substrates have been investigated as shown in Fig. 12.4-15, so it is not clear whether this enzyme may be identical to the L-5-carboxyethylhydantoinase described before (see Sect. 12.4.1). Watabe et al. reported on the cloning and sequencing of genes for an Lhydantoinase deriving from Pseudomonas sp. NS 671 able to convert L-selective D,L5-MTEH, a precursor of methionine['*']. Production of L-methionine from the corresponding hydantoin derivative was also described by Ishikawa et al. for resting cells of Bacillus stearothermophilus NS1122A['221after growth of this strain on a medium containing D,L-5-MTEHas an inducer. The resting cells were described to be stimulated by addition of cobalt and manganese ions, while copper and zinc ions caused a strong inhibition of the enzymatic activities. Wagner et al. described the use of an Arthrobacter sp. DSM 7330 for the production of L-methionine and were able to obtain product concentrations of up to 12Og L-' using a special feed-batch technique for feeding of the hydantoin substrate[123]. From the data available, the three L-hydantoinases from Bacillus breuis and Bacillus stearothermophilus and the enzyme from Pseudornonas mentioned above seem to have a preference for hydantoin derivatives containing aliphatic side chains and therefore differ distinctly from those enzymes found in Arthrobacter sp. by Cotoras et al.

M0

""K"" 0

w

""If"" 0

-sy ""If""

Figure 12.4-15. Substrates accepted by the L-hydantoinase o f Bacillus breuis AJ 12 29914', 501.

0

""If"" 0

12.4 Hydrolysis and Formation ofHydantoins -0

ffo ""Y"" 0

+

""K"" 0

)-w"

""K"" 0

'OW0

""Y"" 0

Figure 12.4-16.

Substrates accepted by the L-hydantoinases of Arthrobader

Yokozeki et a1.[51-53]and Syldatk et al.r7. 1251 as well as in FZavobacteriurn sp. by Nishida et al. [461. These so called "L-5-arylalkylhydantoinases"have comparable substrate specificities and are especially active towards the hydrolysis of hydantoin derivatives with aromatic substituents, as can be seen from Fig. 12.4-16. They could or the corresponding N-3-methyl only be induced by D,L-S-indolylmethylhydantoin derivative of a variety of hydantoins and natural cyclic amides 53* 124* 1251. The L-hydantoinase from Havobacteriurn sp. was reported to be L-selective. Its optimal pH of 9.7 is remarkably high and its optimal temperature is 40 0C[4G]. The enzyme from Arthrobacter aurescens DSM 3745, which has been crystallized and used for initial X-ray analytical studies['26],was described in detail by May et al.['27-'301. The active enzyme is a tetramer consisting of four identical subunits, each with a molecular mass of 49 (570 Da[lz7l,containing 10 mol of zinc per mol of active enzyme, which could be detected by atomic absorption spectrometry and inductive coupled plasma-atomic emission spectrometry['28].By kinetic studies of metal/chelator enzyme inactivation and by identification of specific metal binding ligands, the role of the zinc atoms was found to be in the catalFc activity as well as in A reaction the stabilization of the quaternary structure of the hydant~inase[~~']. mechanism was proposed by Syldatk et al. [131along the lines published for ureases by Jabri et a1.[131]and is shown in Fig. 12.4-17. 1'9

I

785

786

I

72 Hydrolysis and Formation of C-N Bonds Hydrophobic interaction ........

Electrophile (e. g. Zn2*?) Electrophile (e. g. Zn*+?) ,/

.........

,o;-

........

""Y"" 0

-

H

b

w\

.'".

,o,

..........

h2+His62

""YNH

\ Asp274?

('H

0 Nucleophile (e. g. Asp ?)

Electrophile

R t l YCOOH

o",

*

HNKNH2 0 Nucleophile

Proposed reaction mechanism catalyzed by the hydantoinase: after binding o f the substrate, an electrophilic residue (or zinc) stabilizes the negative charge of the carbonyl oxygen. Zinc-bound water is activated and performs a nucleophilic attack on the C4 carbon Figure 12.4-17.

6 K ' I

Electrophile

,../

.......

..'."

2t

/

Zn -His62

HN

H,,,

Asp274?

0

Nucleophile

atom, generating a tetrahedral intermediate. The tetrahedral intermediate undergoes ringopening, assisted by protonation of the ring The proposed residues are derived from a conserved sequence pattern and their respective function in urease['"I.

The enantioselectivity of the enzyme was shown to be strongly dependent on the substrate used[127]:while the enzyme is strictly L-selective for the cleavage of D,L5-IMH, it appears to be D-selective for the hydrolysis of D,L-S-MTEH~~*~]. As part of these investigations, a method based on enzyme activity stain was developed for the detection of hydantoinases with respect to their enanti~selectivity['~'].The isolated enzyme from Arthrobacter aurescens DSM 3745 was recently used for the chemoenzymatic production of optically pure ~-(trimethylsilyl)alanine [l')]. A good stability for the continuous conversion of ~,~-5-indolylmethylhydantoin to N-carbamoyl-L-tryptophan of tIl2 >720 h was first achieved after immobilization of the enzyme by covalent binding to Eupergit CL1321. Further optimization of the immobilization of hydantoin cleaving enzymes has been subsequently carried out [133, 1341. 12.4.5 L-N-Carbamoylases - Substrate Specificity and Properties

In contrast to the D-route, N-carbamoyl-L-aminoacid amidohydrolases (L-N-carbamoylases) were identified in all L-hydantoinase containing microorganisms discussed in Section 12.4.4 (see above). In this section, L-N-carbamoylases from twelve bacterial strains will be discussed with respect to their enzymatic properties and substrate specificities (Table 12.4-5). The biological function of these enzymes is still unknown, with the exception of

72.4 Hydrolysis and Formation ofHydantoins

I

787

the Mn2+/Fe2+-dependent L-selective P-ureidosuccinasefrom Clostridium oroticum (= Zymobacterium oroticum) found by Lieberman and Komberg in 1955 and postulated to play a role in the degradation of orotic acid[26].This hydrolase works best at pH 7.8 to 8.5 and its biological function is postulated to be the conversion of N-carbamoylaspartic acid into L-aspartic acid. It has not been investigated from the biotechnological aspects as yet. The twelve L-N-carbamoylasesderive from seven genera of bacteria: Alcaligenes (1), Arthrobacter (l),Bacillus (4),Blastobacter (l),Clostridium (l),Havobacterium (l),and Pseudomonas (3). Only four of the twelve enzymes have been purified to homogeneity, making a comparison of enzymatic properties difficult. Two of the Bacillus strains have been reported to be thermophilic and the enzymes enriched from these strains have been found to possess optimal temperatures approximately 10 to 20 "C higher than most of the other enzymes (Table 12.4-5).The pH-optima of all L-Ncarbamoylases are between pH 7.5 and 8.5. Whereas hydantoinases are not always strictly L-specific a strictly L-specific carbamoylase, responsible for the optical purity of the amino acid produced with resting cells, has been identified in each strain. The L-N-carbamoylases from Alcaligenes, Arthrobacter, Bacillus brevis A J-12299, Bacillus stearothermophilus NS 1122A and the Pseudomonas putida I F 0 12996 and Pseudomonas sp. NS 671 enzymes have been reported to be (hyper-)activated by one or several of the following heavy metal ions: Mn2+,Co2+,Fe2+Ni2+. In addition to N-carbamoylamino acids some enzymes are able to hydrolyze Nformyl- or N-acetylaminoacids L3', 135-1371. As with the hydantoinases, N-carbamoylases accept N-protected amino acids of unnatural origin. The enzymes of the different genera differ significantly in their substrate specificities. Aliphatic Ncarbamoylaminoacids are preferentially hydrolyzed by the enzymes from the genera Alcaligenes, Bacillus, and Pseudomonas. Only the N-carbamoylasefrom Pseudomonas strain NS 671[13*1 accepts aromatic amino acids as well as aliphatic ones. Aromatic LN-carbamoylamino acids are preferentially hydrolyzed by the enzymes from the genera Arthrobacter and Havobacterium. The substrates hydrolyzed by these enzymes are shown in Fig. 12.4-18. Interestingly, the L-N-carbamoylase from Pseudomonas as a substrate, which is an putida I F 0 12 996 accepts N-carbamoyl-j3-alanine[36] intermediate of the dihydropyrimidine metabolism (see Fig. 12.4-7).In contrast, pureidopropionate is not at all converted by the enzymes from Alcaligenes, Arthrobacter, Bacillus, and Pseudomonas sp. NS 671 and is converted by Havobacterium only, with a very low relative activity. As has been shown by HPLC, whole cells of Alcaligenes xylosoxidans were able to distinguish not only the configuration of the a- but also that of the p-carbon of Ncarbamoyl-o-methylphenylalanine:from the mixture of the four diastereoisomers only threo-L-p-methylphenylalanine was produced [120, 1391. The enzymes from Arthrobacter, Bacillus stearothermophilus NCIB 8224 and NS 1122A, and Pseudomonas sp. NS 671 have been cloned and expressed in E. coli. The enzymes from Bacillus and Pseudomonas share approximately 38 % sequence identity with the Arthrobacter enzyme whereas the 20 amino acids known from the N-termini of the enzymes from Alcaligenes and Pseudomonas putida I F 0 12996 are

7.5

8.5

(‘C) Cloning and Expression Sequenceno identity with Arthrobacfer 1-N-carbamoy. lase (%)

134 000 Da (2 subunits) Optimal Tem- 35 perature (“C) Optimal pH 8.0-8.3

rec. in E. coli

50

44 000 Da (calc. 43993) 93 000 (2 subunits) 50

65000

Pa) .MW natlve (Da)

.MWSDS

homogeneous

homogeneous

partial

1501

11361

11351

Bacillus brevis AJ-12299, Mutant No.102

Reference Purification status

Arthrobacter aurescens DSM 3747 11371

Bacillus steorothermophilus NClB 8224

crude extract ““1

partia1l”‘l

rec. in E. coli 38

38

60

44.000

11121

Blastobocter sp. A17p-I

11531

Bacillus stearothermophilus NS 1122A

rec. in E. coli

60

(calc. 44120)

44 000

whole cells crude extract

1281

Bocillus brevis ATCC 8185

Comparison of L-specific carbamoylases (modified from

Alcaligenes xqlosoxidans

Microorganism

Table 12.4-5.

whole cells

1261

Clostridium oroticum

Flavobacterium

40

parhal

1531

sp. AJ-3912

no

95 000 (2 subunits) 60

45 000

homogeneous

1361

1mqfi

Pseudomonos putido I F 0

crude extract

1351

Pseudomonas sp. AJ-11220

Pseudomonas

37

rec. in E. coli

109 000 (2 subunits) 40

homogeneous (recombinant enzyme) 11381 45 000

11211

sp. NS671

aliphatic C-a-AS: C-r-Met (17)

b

11361

Arthrobacter aurescens DSM 3747

aromatic C-a-AS: C-r-Phe (86). C-L-T,T(45)

aliphatic C-a-AS: C-r-Val (100). C-r-Leu (102), C-r-Ile (84). C-r-Met (73). C-r-Ala (48)

I501

Bacillus brevis A)-12299, Mutant No.102 1281

Bacillus brevis ATCC 8185

C-a-ASC: C-L-Met (97) C-Gly (71) C-DL-Ala (100) C-DL-Val(100) C-r-Leu (94) C-r-lle (55) C-Dr-Ser (86) C-DL.Thr (94) C-L-G~U (56) C-L-ASD(52)

aromatic C-a-ASC: C-D,r-Phe (25) C-i-Trp (trace) C-L-TY(3)

aromatic C-a-AS: C-D,r-Phe (< 0.1) C - L - T(~c~0.1) C-i-Tyr (< 0.1)

aliphatic

C-a-AS: C-r-Met (100) C-o,L-Ala (183) C-L-GIU(112) C-Gly (77) C-r-Leu (28)

1124

aliphatic

11121

Bacillus stear- Blastobacter otherrnophilus sp. A17p-4 NS 1122A

11371

Bacillus stearotherrnophilus NClB 8224 1261

Clortridium oroticum

aromatic C-a-AS: C-~,r-3,4-methylenedioxyPhe (100) C-r-Phe (82

aliphatic C-a-AS: C-r-Met (24) C-D.1-0-Me-Ser (13) C-r-Ser (5) C-Gly (5) C-r-Leu (3) C-i-Ile (2) C-r-Val(2) C-r-Gln (1) C-1-Asn (1) C-r-Ala (0.5)

1531

b

Flauobacterium sp. A)-3912

other: (13) C-r.Tyr (127) C-L-TY (59) Acetyl-Met (38) fOnTlyl-D,LC-r.Trp (55) Acety-Glu (7) Leu (5) other: C-o.r-3,4-dimefOImyl-D,LFormy1-o.~thoxy-Phe (24) Met (5) Trp (98) C-D,L-O-benzylAcetyl-L-Phe serine (15) (0.7) kcetyl-o, Lother: 2-aminohexaP-ureidopronoic acid pionate (3) (0.06) a relative acuvines ~n[%Iare given in brackets () except'. b additional data on substrates not hydrolyzed are given in the cited hterahlre. c isolated yleld m [%] after 24 h ~n brackets ()

aromatic C-a-AS: C-i-Trp (100) C-r-ThienylOther: ala (316) formyl-D,L-Ala C-r-Phe (98)

aliphatic C-a-AS: C-UAla (100) C-Gly (75) C-r.Va1 (28) C-r-Leu (9) C-r-Met (12) C-r-lle (5) C-o,r-2-aminohexanoic acid (24) C-o,r-Ser (19) C-o,r-Thr (9) C-r-Asn (64)

accepted'

aromatic C-a-AS: C-r-Phe (5)

11351

References Substrates

(cont.).

Alcaligenes xylosoxidans

Microorganism

Table 12.4-5.

p-Ureidoisobutyrate (43) formyl-D,L-Ala (75) Acetyl-D,r-Ala (6)

other: J3-ureidopropionate (100) y-ureidobutyrate (290)

aliphatic C-a-AS: C-Gly (16) C-r-Ala (118) C-r-Ser (34) C-or-a-aminobutyrate (31) C-2-aminovalerate (9) C-o.r-Thr (1) C-o,r-Asp (0.1) C-L-GIU(0.3) C-r-Asn (1.6)

134 b

Pseudomonar putida I F 0 12996

aromatic C-a-AS: C-r-Phe (10) C - ~ T y (9) r

aliphatic C-a-AS: C-r-Val (100) C-r-Met (47) C-r-Ala (44) C-r-Leu (98) C-L-GIU(3) C-r-Asn (2)

b

139

Pseudomonar sp. AJ-11220

aromatic C-a-AS: C-D.r-Phe (94) C - L - T(60) ~~

aliphatic C-a-AS: C-r-Met (100) C-o,r-Ala (102) C-D,r-Val (106) C-r-Leu (118) C-r-Ile (97)

b

11211

Pseudomonas 5p. NS671

1

U

W 0

1

9

5

4

3

3

a

D

a

1: -=a

.b .cI

790

I

12 Hydrolysis and Formation ofC-N Bonds

i.;L

\ rCOOH

O Y C O O H

HNKNHz 0 )-COOH

HNKNHz 0

0

-'>COOH

"KNH2 0

\ I

COOH

c

c

l

HNKNH2 0

a

C

O

O

H

h

COOH

HNKNH 0

C

O

O

H

HNKNHz 0

HNKNHz 0 -0.

)-tCOOH

Ho>COOH

HNKNHz 0 5

C

O

O

HNKNHz 0

H

0

F-COOH

HNKNHz 0

HNKNHz 0 H2N+COOH 0

\COOH

HNKNHz 0

HNKNHz 0

+COO,

HNKNHz 0

f\COOH

HNKNH2 0

z'N-@)-

H N 70f N H z

O>COOH

HNKNH

HNKNHz 0 COOH

S

°

HNKNHz 0

C

O

O

H

HNKNH 0

COOH

HNKNH2 0

Figure 12.4-18. Substrates accepted by the L-N-carbarnoylases of Atthrobacter sp.['] and Flauobacterium sp.[46, 51-531.

completely different. In contrast to the D-N-carbamoylases (see Sect. 12.4.3),the L-Ncarbamoylase of Arthrobacter sp. DSM 3747 is induced by N-3-methylated D,L5-indolylmethylhydantoin,which cannot be hydrolyzed by the cells 17]. Resting cell L-hydantoinase processes were first developed for the industrial production of L-tryptophan by the companies Ajinomoto and Tanabe146,51-531. In 1992 the Riittgers company tried to enter the amino acid marked with a resting cell

12.4 Hydrolysis and Formation ofHydantoins

I

791

process for the production of unnatural aromatic L-amino acids using Arthrobacter sp. DSM 3745 or DSM 3747, which both contain an L-hydantoinase, hydantoin racemase and L-N-carbamoylase. However, the productivities obtained (see Fig. 12.419 and for details referenceLs7I)seemed to be too low to fulfill economic requirements. In recent years, new developments have been published, which could overcome these problems: 1. the L-N-carbamoylase from Arthrobacter aurescens DSM 3745 and 3747 could be produced as recombinant enzymes in high cell density culture in Escherichia coli

using an expression system based on the Escherichia coli rha-BAD-promoter[1401, 2. purification of the recombinant L-N-carbamoylases could be optimized by expression of enzymes carrying different tags, making the purification protocols much easier[’41]and, 3. the hydantoin-cleaving enzymes from Arthrobacter aurescens DSM 3747 could be stabilized significantly by immobilization 1341. Reaction rate (“A)

Molar conversion (“A) L-amino acid

after 1 h

after 2 6 h

HNyNH

100

> 90

tryptophan

140 - 160

> 90

phenylalanine

20 - 40

> 70

Obenzylserine

150-200

> 80

170 - 200

> 80

50 - 70

> 80

15-20

> 70

25 - 30

> 70

2 -naphthylalanine

25 - 30

> 80

3,4-dirnethoxy-

0-

-0

I pchloro-phenylalanine

pfluoro-phenylalanine

pnitro-phenylalanine 1‘-naphthylalanine

phenylalanine

QFigure 12.4-19.

170 - 200

> 80

Industrial production of unnatural aromatic L-amino

2’ 4hienylalanine

792

I

72 Hydrolysis and Formation of C-N Bonds

All these developments, together with the directed evolution of the hydantoinase towards a more L-selective enzyme with higher activity['42]will possibly lead to an economically viable production process in future. Additionally, an Escherichia coli whole cell biocatalyst has been constructed containing the genes of hydantoinase, hydantoin racemase and L-N-carbamoylase from Arthrobacter aurescens in optimal proportions, so that during the reaction no LN-carbamoylamino acid occurs as an intermediate product any longer['43]. 12.4.6 Hydantoin Racemases

During enzymatic hydrolysis of 5-monosubstituted hydantoin derivatives in some cases the remaining, non-hydrolyzed enantiomer is racemizing chemically under alkaline reaction conditions. The velocity of this chemical racemization is strongly dependent on electronic factors ofthe substituent in the 5-position (seeTable 12.44). High velocities of racemization are observed particularly for 5-phenyl-and 5-I)-OHphenylhydantoin. From reports in the early literature resting cell bioconversions of hydantoin derivatives, which do not racemize with high velocities, indicated an enzymatic racemization and the presence of a hydantoin racemase. In addition, the chemical and the enzymatic racemization proceed via the keto-enol tautomerism, which is shown in Fig. 12.4-20. Stabilizing effects on the enolate structure such as electronegative substituents are responsible for the velocity of the racemization[2' 1' . Increased racemization rates can be also seen at more alkaline pH-values and with increased temperatures "1. The first hydantoin racemase acting on a cyclic amide substrate reported in the literature was the allantoin racemase (E.C. 5.1.99.3) (Fig. 12.4-4). This enzyme enables several bacteria to use both allantoin enantiomers as substrates [20-22]. Racemic mixtures of allantoin, e. g. from plant materials, can be completely metaboRacemization rate constants k,,, and corresponding half-live times t,,z,rac for various hydantoins at pH 8.5 and 40 "C. Values were calculated from first order rate law: = In 2/krac. In ([al/[a10)= - krac.t; Table 12.4-6.

5-Substituted hydantoin

Substituent: Phenyl Hydroxymethyl Benzyl Methylthioethyl 1'-Hydroxyethyl 3 '-Ureidopropyl 1'-Methylethyl Imidazolylmethyl

Isobutyl Methyl Isopropyl

Correspondingo-amino acid

k,,, (ti')

D-Phg D-Ser D-Phe D-Met D-allo-Thr D-Cit D-allo-Ile D-His D-Leu D-Ala D-Val

2.59 0.43 0.14 0.12 0.11 0.049 0.044 0.043 0.032 0.020 0.012

tbmc

(h)

0.27

1.60 5.00 5.82 6.41 14.26 15.84 16.09 21.42 33.98 55.90

72.4 Hydrolysis and Formation ofHydantoins

R

‘yo

-““K 0 NH

0 Enol

L-Hydantoin

Figure 12.4-20. Ketoenol-tautomerism o f 5-monosubstituted hydantoin derivatives.

0-Hydantoin

lized by various bacteria using a sequence of the L-specific allantoinase and allantoin racemase (see Sect. 12.4.1).Although the natural function of this allantoin racemase is not clear, because allantoin racemizes with high velocities under physiological conditions. The fast and total conversion of r-5-isopropylhydantointo D-valine by resting microbial cells led Battilotti et al.[30]to the suggestion that a hydantoin racemase might be responsible for the racemization of the L-enantiomer.The first hydantoin racemase to be described in detail was a 5-arylalkylhydantoinracemase, which was isolated and purified from Arthrobacter sp. DSM 3747[’’* 1442 14’1. Its substrate specificity is shown in Fig. 12.4-21. As can be seen from Fig. 12.4-21, only some aliphatic and aromatic hydantoin derivativesare accepted by the enzyme out of a variety of substrates. The enzyme was recently cloned and heterologously expressed in Escherichia coli [1461. The gene encoding the hydantoin racemase, designated hyuA, was identified upstream of an LN-carbamoylase gene in the plasmid pAWl6 containing genomic DNA of Arthro-

HN)N fR ..

Substrate R1

R2

Relative activity

Ri

R2

-H

100.0

-H

9.8

-H

20.4

-H

0

HO-

-H

0

HOCC-

-H

0

\f\ I /s-

-H

76.7

-H

62.7

HO-

Relative activity (%)

(%)

w,

Figure 12.4-21. Substrate specificity o f the hydantoin racemase from Arthrobacter sp. DSM 3745 [”. 1441.

I

793

794

I

12 Hydrolysis and Formation of C-N Bonds

bacter aurescens. The matrix assisted laser desorption ionization spectrum (MALDI) of the purified racemase gave a peak at a molecular mass of 25 078.7. This is in good agreement with the calculated value of 25 085 Da for the racemase monomer. On a calibrated column of Superose 12 HR, the relative molecular mass of the native enzyme was estimated to be approximately 170 kDa + 25, so that the native enzyme is suggested to be either a hexamer, heptamer or octamer. The optimal conditions for racemase activity were pH 8.5 and 55 "C with L-5-benzylhydantoinas the substrate. The enzyme was completely inhibited by HgClz and iodoacetamide and stimulated by addition of dithiothreitol, while no effect was seen with EDTA. Kinetic studies revealed substrate inhibition towards the aliphatic substrate L-5-methylthioethylhydantoin. Enzymatic racemization of 0-5-indolylmethylenehydantoinin DzO and NMR analysis showed that the hydrogen at the chiral center of the hydantoin is exchanged for solvent deuterium during the racemization. Comparative analysis of h y u A with various protein databases indicated homology to hydantoin racemases. This hydantoin racemase shared 47.2 % identity in amino acid sequence with the hydantoin racemase of Pseudomonas sp. NSG71 and lower identities to putative hydantoin racemases of Schizosaccharomyces pombe (SwissProt accession no. 409921) and Saccharomyces cerevisiae (SwissProt accession no. P324GO). The multi-alignment of the enzymes showed that the N-terminal region in particular is highly conserved. No significant similarity to the various amino acid racemases or any other racemases deposited was found in the data bases. The hydantoin racemase from Pseudomonas sp. NS 671 is able to racemize both enantiomers of 5-(2-methylthioethyl)hydantoin,5-isopropylhydantoin,S-isobutylhydantoin and 5-ben~ylhydantoin[~~~I. All together, the presence of hydantoin racemases in resting cells used in industrial processes is of importance for a fast and total conversion of hydantoins which racemize chemically very slowly. In future there might be a combination of hydantoin racemases from L-selective microorganisms with D-hydantoinases and D-N-carbamoylaseswhen designing optimal processes leading to D-amino acids. For industrial use, the fast racemization of 5-monosubstituted hydantoin derivatives under mild conditions in the presence of ion exchangers [144, 14'1 could prove more significant, as this procedure also enables fast and total conversion of D,L-S-monsubstitutedhydantoins without enzymatic racemization. 12.4.7

Conclusions

The hydantoinase method has become of significant interest for preparative organic

chemistry: total conversion of racemic hydantoins, synthesized by well-established chemical methods to nearly 100% optically pure products is possible using free or immobilized microbial cells or enzymes. Further, it is possible to prepare a wide range of optically pure D- as well as L-amino acids by this method. Of course there are many factors which influence the competitiveness between enzymatic processes and chemical processes, for example, costs of substrates, costs for production/isolation of enzymes, possible space-time yields and costs for

12.4 Hydrolysis and Formation ofHydantoins

isolation of the products. These factors are strongly dependent on the desired product and therefore there is no single best process for the production of amino acids. For D-p-hydroxyphenylglycine, which is the most important compound produced by the hydantoinase process on an industrial scale (> 1000 tons) at the moment, a first comparison of the feasibility of different methods was given by Tramper and Luyben in the 1 9 8 0 ~ [ ' ~However, ~]. it has already been shown that the hydantoinase process can be employed for the production of many unnatural amino acids which are components of promising pharmaceuticals[l5'1. If these pharmaceuticals reach the market, there will be an augmented demand for these amino acids, which could lead to an increased importance of the hydantoinase process in the future. With the availability of recombinant enzymes, one could expect that the hydantoinase method will also become an important tool in biotransformation of simple precursors to L- and D-amino acids. Some of the current reports on hydantoinase processes focus on isolation and the . Processes at an recombinant expression of thermostable enzymes[84*86* 87, 95* elevated temperature would increase the solubility and racemization rate of hydantoins. Therefore, the increased thermostability of these enzymes is very useful, if the specific activities are still high. Another main advantage of the recombinant expression of the hydantoin cleaving enzymes is to decrease the costs of catalysts, which might contribute to the competitiveness of the hydantoinase processes, which to date do not employ recombinant enzymes. The Kanekafuchi company have published a patent for the production of D-N-carbamoyl-aminoacid from 5-substituted hydantoin, using a recombinant hydantoinase derived from a strain of Pseudomonas, Agrobacterium or Bacillus['52].This might indicate that highly active recombinant Escherichia coli cells could replace the wild-type cells in the near future. Furthermore, the recombinant expression of hydantoinases (and of course carbamoylases[153, 1541) allows enzyme properties such as stability or stereoselectivity to improve by means of protein design. If an X-ray structure was solved, this could be done by a rational protein design[lS5]or, lacking knowledge about a structure, by evolutionary protein design['S6]. May et al. are already able to improve the stereoselectivity of a Lhydantoinase for the conversion of ~,~-S-methylthioethylhydantoin[~~~~, while Kim et al. have shown the possibility of using fusion proteins of D-hydantoinase and D-NlS81. carbamoylase for the production of D-amino Future work will show the impact of these methods on the biotechnological application of hydantoinases. Besides the applied research on hydantoinases for the production of amino acids, the natural functions and genetic organization of distinct hydantoinases, related hydantoin racemases and N-carbamoylasesare still unknown and are of great interest for basic research.

I

795

796

I

72 Hydrolysis and Formation ofC-N Bonds

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798

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12 Hydrolysis and Formation ofC-A! Bonds

S.-G. Lee, L. D.-C., H.4. Kim, Appl. Biochem. Biotechnol. 1997, 62, 251-266. 87 V. Luksa, V. Starkuviene, V. Starkuviene, R. Dagys, Appl. Biochem. Biotechnol. 1997, 62, 219-231. 88 S. Runser, E. Ohleyer, Biotechnol. Lett. 1990, 12,259-264. 89 C. L. Soong, J. Ogawa, M. Honda, S. Shimizu, Appl. Environ. Microbiol. 1999, 65, 1459-1462. 90 A. Morin, N. H. Tran Tmng, G. LaPointe, H. Dubeau, Appl. Microbiol. Biotechnol. 1995,43,259-266. 91 G. LaPointe, D. Leblanc, A. Morin, Appl. Microbiol. Technol. 1995,42, 895-900. 92 E. Jacob, K. Henco, S. Marcinowski, G. Schenk, G e m . Pat. DE 353 5987,1987. 93 G. LaPointe, S. Viau, D. Leblanc, N. Robert, A. Morin, Appl. Environ. Microbiol. 1994, 60, 888-895. 94 Y. Mukohara, T. Ishikawa, K. Watabe, H. Nakamura, Biosci. Biotech. Biochem. 1994, 58,1621-1626. 95 D.-C. Lee, G.-J. Kim, Y.-K. Cha, C.-Y. Lee, H.3. Kim, Biotechnol. Bioeng. 1997,56, 449-455. 96 G.-J.Kim, H . 3 . Kim, Biochem. Biophys. Res. Commun.1998,243,96100. 97 H. R. Chien, Y.-L. Jih, W.-Y. Yang, W.-H. Hsu, Biochim. Biophys. Ada 1998, 1395, 68-77. 98 K. Matsuda, S. Sakata, M. Kaneko, N. Hamajima, M. Nonaka, M. Sasaki, N. Tamaki, Biochim. Biophys. Acta 1996, 1307, 140-144. 99 M. Tzermia, 0.Horaitis, D. Alexandraki, Yeast 1994, 10,663-679. 100 E. M. Baldaro, P h a m a c . Man$ Int. 1993, 137-139. 101 K. Drauz, M. Kottenhahn, K. Makryaleas, H. Klenk, M. Bemd, Angew. Chem. 1991, 103,704-706. 102 C.-K. Lee, K.-C. Lin, Enzyme Microb. Technol. 1996, 19, 623-627. 103 D.-C. Lee, S.-G. Lee, H.3. Kim, Enzyme Microb. Technol. 1996, 18, 35-40. 104 A. Achary, K. A. Hariharan, S. Bandhyopadhyaya, R. Ramachandran, K. Jayaraman, Biotechnol Bioeng. 1997, 55, 148-154. 105 D.-C. Lee, J.-H.Park, G.-J. Kim, H . 4 . Kim, Biotechnol. Bioeng. 1999, 64, 272-283. 106 C.-K. Lee, C.-H. Fan, Bioproc. Eng. 1999, 21, 3412-3347. 107 C.-K. Lee, C.-H. Fan, Enzyme Microb. Technol. 1999, 24, 659-666. 86

108 P. Meyer, S. Runser, FEMS Microbiol. Lett.

1993, 109,67-74.

109 A. Buson, A. Negro, L. Grassato, M. Ta-

gliaro, M. Basaglia, C. Grandi, A. Fontana, M. P. Nuti, FEMS Microbiol. Lett. 1996, 145, 55-62. 110 A. Louwrier, C. J. Knowles, Biotechnol. Appl. Biochem. 1997,25, 143-149. 111 H. Nanba, Y. Ikenaka, Y. Yamada, K. Yajima, M. Takano, S. Takahashi, Biosci. Biotechnol. Biochem. 1998,62,875-881. 112 J. Ogawa, M. C.-M. Chung, S. Hida, H. Yamada, S. Shimizu, J . Biotechnol. 1994, 38, 11-19. 113 R. Olivieri, E. Fascetti, L. Angelini, L. Degen, Enzyme Microb. Technol. 1979, 1 , 201-204. 114 H. Nanba, Y. Ikenaka, Y. Yamada, K. Yajima, M. Takano, K. Ohkubo, Y. Hiraishi, K. Yamada, S. Takahashi, Biosci. Biotechnol. Biochem. 1998,62,1839-1844. 115 G.-J.Kim, H.-S. Kim, Biotechnol. Lett. 1994, 16,17-22. 116 R. Grifantini, C. Pratesi, G. Galli, G. Grandi, J . Biol. Chem. 1996, 271, 9326-9331. 117 G.-J. Kim, H.-S. Kim, Enzyme Microb. Technol. 1995, 17, 63-67. 118 H. Yamanaka, T. Kawamoto, A. Tanaka,J. Ferment. Bioeng. 1997, 84, 181-184. 119 M.Pietzsch, T. Waniek, R. J. Smith, S. Bratovanov, s. Bienz, C. Syldatk, Chem. Month. 2000, 131,645-653. 120 J. Ogawa, A. Ryouno, S.-X. Xie, V. M. Rakesh, R. Indrati, H. Miyakawa,T. Ueno, Y. Ikenaka, H. Nanba, S. Takahashi, S. Shimizu, Appl. Microbiol. Biotechnol., 1999, 55, 797-801. 121 K. Watabe, T. Ishikawa, Y. Mukohara, H. Nakamura,J. Bacteriol. 1992, 174,962969. 122 T. Ishikawa, Y. Mukohara, K. Watabe, S. Kobayashi, H. Nakamura, Biosci. Biotech. Biochem. 1994,58,265-270. 123 T. Wagner, B. Hantke, F. Wagner,J. Biotechnol. 1996, 46, 63-68. 124 D. Cotoras, Dissertation, 1985, Technische Universitat Braunschweig, Germany. 125 C. Syldatk, F. Wagner,/. Food Biotechnol. 1990,4,87-95. 126 0.May, M. Siemann, C. Syldatk, K. Niefind, D. Schomburg, Acta Crystallogr. 1996, D52, 1209-1210. 127 0.May, M. Siemann, M. Pietzsch, M. Kiess,

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R. Mattes, C. Syldatk,]. Biotechnol. 1998,61, 1-13. 128 0.May, M. Siemann, M. G. Siemann, C. Syldatk,J. Mol. Catal. B: Enzymatic 1998, 5, 367-370. 129 0.May, M. Siemann, C. Syldatk, Biotechnol. Techn. 1998, 12, 309-312. 130 0. May, M. Siemann, C. Syldatk,J. Mol. Cat. B: Enzymatic 1998,4,211-218. 131 E. Jabri, M. B. Carr, R. P. Hausinger, P. A. Karplus, Science 1995,268,9981004. 132 G. Wohlfahrt, R. Miiller, C. Syldatk, DEC H E M A Biotechnology ConJ Ed. 1992, Vol. y pp. 45-49. 133 M. Pietzsch, H.Oberreuter, B. Petrovska, K. Ragnitz, C. Syldatk, in: Stability and Stabilization ofBiocatalysts (Eds.: A. Ballesteros, F. J. Plou, J. L. Iborra, P. Halling), Vol. 15, Elsevier Science, Amsterdam, 1998, p. 517-522. 134 K. Ragnitz, M. Pietzsch, C. Syldatk,]. Biotechnol. 2001, 92, 179-186. 135 J. Ogawa, H. Miyake, S. Shimizu, Appl. Microbiol. Biotechnol. 1995,43, 1039-1043. 136 B. Wilms, A. Wiese, C. Syldatk, R. Mattes, J. Altenbuchner, M. Pietzsch, J. Biotechnol. 1999,68,101-113. 137 N. Batisse, P.Weigel, M. Lecocq, V. Sakanyan, Appl. Environ. Microbiol. 1997, 63, 763-766. 138 T. Ishikawa, K. Watabe, Y. Mukohara, H. Nakamura, Biosci. Biotech. Biochem. 1996, 60,612-615. 139 J. Ogawa, A. Ryouno, S.-X. Xie, V. M. Rakesh, R. Indrati, H. Miyakawa, T. Ueno, S. Shimi2u.J. Mol. Cat. B: Enzymatic 2001, 12, 71-75. 140 B. Wilms, A. Hauck, M. Reuss, C. Syldatk, R. Mattes, M. Siemann, J. Altenbuchner, Biotechnol. Bioeng., 73, 95-103. 141 M. Pietzsch, A. Wiese, K. Ragnitz, B. Wilms, J. Altenbuchner, R. Mattes, C. Syldatk, J. Chromatop. B 2000,737,179-186. 142 0.May, P.T. Nguyen, F. H. Arnold, Nat. Biotechnol. 2000, 18, 317-320.

B. Wilms, A.Wiese, C. Syldatk, R. Mattes, J. Altenbuchner, J. Biotechnol. 2001, 86, 19-30. 14.4 M. Pietzsch, Dissertation, 1992, Technische Universitat Braunschweig, Germany. 145 M. Pietzsch, D E C H E M A Biotechnology Conferences Ed. 1990, Vol. I V , p. 259. 146 A. Wiese, M. Pietzsch, C. Syldatk, R. Mattes, J. Altenbuchner, 1.Biotechnol. 2000, 80,217-230. 147 K. Watabe, T.Ishikawa, Y. Mukohara, H. Nakamura, J. Bacterial. 1992, 174, 3461-3466. 148 F. Wagner, M. Pietzsch, C. Syldatk, US Pat. 5,449,7861995. 149 J.Tramper, K. C. A. M. Luyben, PVProcestechniek 1984, 39, 61-67. 150 K.-H. Drauz, Chimia 1997,51. 310-314. 151 G.-J. Kim, J.-H. Park, D.-C. Lee, H.3. Ro, H.4. Kim, Mol. Gen. Genet. 1997, 225, 152-156. 152 Y. Ikenaka, H. Nanba, S. Takahashi, M. Takano, K. Yajima, Y. Yamada, Eur. Patent 801 131 A l , 1997. 153 Y. Mukohara, T. Ishikawa, K. Watabe, H. Nakamura, Biosci. Biotech. Biochem. 1993, 57,1935-1937. 154 R. J.Neal, A.M. Griffin, M. Scott, A. R. Shatzman, H. C. Gorham, world patent WO 94 00577,1994. 155 2. Shao, F. H. Arnold, Cum. Opin. Stnrct. Bid. 1996, 6, 513-518. 156 F. H. Arnold, J. C. Moore, Adv. Biochem. Eng. 1997,58,1-14. 157 G. J. Kim, Y. H. Cheon, H. S. Kim, Biotechnol. Bioeng. 2000,68, 211-217. 158 G. J. Kim, D. E. Lee, H. S. Kim, Appl. Enuiron. Microbiol. 2000, 66, 2133-2138. 159 M. Siemann, A. Alvarado-Marin,M. Pietzsch, C. Syldatk,J. Mol. Catal. B: Enzymatic 1999, 6, 387-397. 160 J. Ogawa, M. Honda, C. L. Soong, S. Shimizu, BiosGi. Biotech. Biochem. 1995, 59, 1960-1962. 143

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

800

I

12 Hydrolysis and Formation ofC-N Bonds

12.5

Hydrolysis and Formation of Peptides Hans-DieterJakubke 12.5.1

Introduction

Peptides and proteins play a fundamental role in the formation and maintenance of structure and function of living systems. Peptides comprise a variety of biologically active linear and cyclic compounds with diverse functions. The different classes of peptides include, for instance, hormones and other signalling or regulatory factors, antibiotics, alkaloids, toxins, enzyme inhibitors, and sweeteners. There is permanently great interest in pharmaceutically active peptides and proteins since they have many applications and great potential in medicine, such as in cardiovascular diseases, mental illness, connective tissue diseases, the therapy of cancer, regulation of fertility and growth, and the control of pain. The demand for peptides and proteins is enormous, and rising all the time. In a peptide chain amino acids are linked together by bonds between the carboxyl group of one and the amino group of another amino acid, known as peptide bonds. This amide or peptide bond has some characteristics of a double bond: it does not rotate freely and is shorter than other C - N bonds. Nature provides a wide range of special enzymes, the proteol$c enzymes or correctly designated as peptidases, which can cleave these bonds in peptide and protein substrates. In contrast, for catalyzing the formation of peptide bonds the number of efficient enzymes is rather low. Peptidases catalyze a single reaction, the hydrolysis of a peptide bond. The ubiquitous distribution among all life forms and their enormous diversity of function makes the peptidases one of the most fascinating families of enzymes. As a result of complete analysis of several genomes it has been shown that about 2 % of all gene products are proteolytic enzymes. In biological and biochemical research proteolytic enzymes play a contrary role: some researchers either love them or other hate them. In the first case, the only good peptidase is a dead one, no longer capable of degrading the desired protein during isolation and purification. Irreversible inhibition of any contaminating proteolytic enzyme is the best way to solve this problem. However, for most purposes proteolytic enzymes are of great importance. Owing to the special physiological functions, some proteolytic enzymes are active in degrading proteins for digestive and nutritional purposes. These enzymes act both extracellulary (e.g. in the intestine of animals) and intracellulary (in the hydrolytic subcellular organelles, preferentially in liver and kidney cells). Other peptidases are responsible for controling processes, e. g. they can act to cause limited proteolysis of peptide and protein substrates. In limited proteolytic processes a single susceptible peptide bond may be cleaved followed by a dramatic change in the biological activity of the products formed. Physiological functions are a result of proteolytic conversion of inactive precursors into biologically active proteins, e. g. in blood coagulation, prohormone or proenzyme activation. Pancreatic peptidases frequently exist as

72.5 Hydrolysis and Formation of Peptides

zymogens, a special inactive proenzyme arrangement that ensures that the pancreas does not digest itself. These enzymes have their function outside cells and will be activated by another peptidase at the place of action. The number of peptidases within the cell are more numerous but much more difficult to investigate in comparison with the extracellular enzymes"]. A much smaller group are the cellsurface peptidases which are specialized in the hydrolysis of relatively simple peptides rather than proteins. This group of peptidases does not need activation. Usually the biological function is the inactivation of signalling peptides in order to terminate a hormonal or neuropeptide signal but sometimes they activate peptide substrates, e. g. the conversion of angiotensin I to angiotensin 11F2, 1' . Contrary to the well-known native function of peptidases the reverse reaction, the peptidase-catalyzed peptide bond formation, can only be successfully carried out by manipulating the reaction conditions, the enzyme or the substrate. Besides enzymatic techniques, classical chemical synthesis in solution, solid-phase synthesis and recombinant techniques belong to the most important methods of peptide synthesis. The main aim of this chapter is to give an overview of the present importance of proteases in the technology of peptide synthesis. 12.5.2 Hydrolysis of Peptides 12.5.2.1 Peptide-CleavingEnzymes 12.5.2.1.1

Introduction and Terminology

More than 500 proteolytic enzymes are known and, in a general sense, they all catalyze the same reaction: hydrolysis of peptide bonds. An excellent handbook c41 provides a ready reference to the approximately 500 proteolytic enzymes known up to the end of the 1990s. These enzymes are classified as peptidases or proteases. In the past there has been widespread uncertainty about the exact meaning of the terms proteases, peptidases and proteinases, as well as proteolytic enzymes. There is no doubt that proteolytic enzymes was the most generally understood term in the current usage. However, this is ambiguous since many of the enzymes which are capable of hydrolyzing peptide bonds do not accept proteins as substrates. The Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (NCIUBMB) recommends the term peptidase as the general term for all peptide bondhydrolyzing enzymes. The E. C. List can be found in its revised version on the World Wide Web (www) at http://www.chem.qmw.ac.uk/iubmb/enzyme/index.html. The acceptable terms for the major types of peptidases are shown in Fig. 12.5-1. The meanings of the words below are described by the italicized semi-systematic terms. The terms in bold type are preferred, whereas the terms in parentheses have historical precedence and are satisfactory when used in the correct context. Most of the peptidases fall into one of two categories, depending on the positional specificity of the peptide bond cleavage process. An enzyme is said to be an endopeptidase when

I

802

I

72 Hydrolysis and Formation ofC-N Bonds

Peptide bond hydrolase Peptidase (=Protease)

I I Endo-acting peptide bond hydrolase Endopeptidase (=Proteinase) Figure 12.5-1.

Exo-actingpeptide bond hydrolase Exopeptidase (=Carboxy- and Aminopeptidases)

Proposed terms for the major types of peptidases.

L3 I T

Site of endopeptidase action

Y

9’

T5

H,N-CH-CO-NH-CH-CO-NH-CH-CO-NH-CH-CO-NH-CH-COOH

t

aminopeptidases

t

A

carboxypeptidases

L S i t e of exopeptidase action

Figure 12.5-2.

Scheme o f the action o f endopeptidases and exopeptidases.

the susceptible peptide bond is an internal one in a peptide or protein. In contrast, an enzyme is termed an exopeptidase when the susceptible peptide linkage is at the carboxyl terminus or at the amino terminus of the substrate. In the E. C. List there are also terms for subtypes of exopeptidases and endopeptidases. Exopeptidases acting at the free N-terminus liberating a single amino acid residue (aminopeptidases) or a dipeptide or a tripeptide (dipeptidyl-peptidases and tripeptidyl-pepidas), whereas those acting at the free C-terminus liberate a single residue (carboxypeptidases) or a dipeptide (pepidyl-dipeptidases). Furthermore, other exopeptidases are specific for dipeptides (dipeptidases)or remove terminal residues which are substituted, cyclized or linked by isopeptide bonds (omegapeptidases). Endopeptidases act on bonds in the middle of the peptide chain (see Fig. 12.5-2). The term oligopeptidase is used to refer to endopeptidases that act optimally on oligopeptide substrates rather than on proteins. peptidases differ in the specificities that they display in a hydrolysis reactions. It is somewhat simplistic to designate a peptidase on the basis of a single amino acid residue at the active site. Near the active site of the peptidase is a “pocket” in the surface of the enzyme molecule which is specific for amino acid side chains of the substrate. Owing to different interactions in this region there are great differences in the so-called primary specificity of the peptidases. Trypsin, for example, cleaves only those peptide bonds adjacent to the amino acids lysine or arginine which carry a positive charge and are hydrophilic. In the binding pocket of trypsin a negatively charged aspartic acid unit is at the back, holding the positively charged lysine or arginine side chain in the pocket by electrostatic forces. Despite the fact that this pocket for specific side chains is obviously important for binding, it is not the only binding site. It has been followed from kinetic studies that the binding of substrates (and inhibitors) involved interactions at a number of subsites on either side of the

Figure 12.5-3. Simplified representation of the DeDtidase sDecificiW

Protease

HN ,

s3 s,

s,

s;

s;

s;

ger[2551.The amino acid residues of

pz

pi

p;

p;

p;

COOH thecorrespondingsand S'subsites

p3

pair of residues containing the peptide bond to be hydrolyzed. The enzyme and substrate must be fixed at several points, so that the susceptible bond is oriented at the active site in optimal configuration. In 1967, a system of nomenclature to describe the interaction of peptidases and their substrates was introduced by Schechter and Berger[2551. According to this system the binding site for a peptide substrate in the active site of a peptidase is envisioned as a series of subsites S which interact with the amino acid building blocks P ofthe peptide or protein substrate (see Fig. 12.5-3).The amino acid residues of the substrate are denoted by P and P', respectively, which interact with the corresponding S and S' subsites within the active site of the peptidase. The sites are numbered from the catalytic site, S1....S, towards the N-terminus of the peptide substrate, and S1'. ...S', towards the C-terminus. In analogy, the residues which they accommodate are numbered PI.. .. P,, and PI'....P,,', respectively. The arrow indicates the site of enzymatic cleavage of the substrate between the residues PI -PI'. With the increasing knowledge of the amino acid sequences of peptidases and particularly when the three-dimensional protein structure began to emerge, a functional division of peptidases became possible. Detailed mapping of the active sites has provided a better understanding of the interaction of substrate and peptidase and has permitted both the design and synthesis of highly specific inhibitors as well as a useful prediction of the outcome of the reverse peptidase action in peptide synthesis (see Sect. 12.5.3.3). The general stoichiometry for the hydrolysis of a peptide bond is shown in Fig. 12.5-4. Water attacks the electron-deficient carbonyl atom targetting first a tetrahedral adduct, which then eliminates the amine fragment and produces the acid. The process is characterizedby transferring the aminoacyl moiety of the peptide to water. In this type of group-transfer reaction the nucleophilic co-substrate is water; 55.5 M water is the most nearly ubiquitous weak nucleophile in degradative enzymatic

->

R4-NH-R' '?I

H,O

peptide Figure 12.5-4.

+-

[

R ~ ~ H . 1

tetrahedral adduct

+

R?

It'

+

HZNR'

OH

acid

amine fragment

The general mechanism for the hydrolysis o f a peptide bond.

804

I

72 Hydrolysis and Formation ofC-N Bonds

processes in the cell. Under physiological conditions the hydrolysis of peptide bonds will proceed in the absence of peptidases, but only at an exceedingly low rate. The reactants only rarely attain the high internal energy required for the hydrolysis process. In contrast, enzymes allow the reaction to follow a different pathway from the substrate to the products, and, therefore, reduce the energy barriers. In the course of the reaction new intermediate states of highest energy appear, with energy lower the internal energy barriers, e. g. the high-energy transitions between one intermediate and the following one. Proteolysis is functionallyirreversible,since energy is liberated in the hydrolysis of peptide bonds. From the overall change in energy it follows that the ionized hydrolysis products are thermodynamically more stable. On the other hand, aminoacyl-grouptransfer is involved in protein biosynthesis. As a result of the ionized state of amino acids at physiological pH, the attack by the amino group of another amino acid to form a peptide bond would involve formal expulsion of 0 z 2 - . This species is very instable and, therefore, would not proceed to any reasonable extent. In protein biosynthesis the carboxylate must be chemically modified so that an oxygen atom can be eliminated with a low energy activation. The key concept in protein biosynthesis is that the aminoacyl group from an activated intermediate is transferred to the specific nitrogen of the amino group catalyzed by the ribosomal peptidyltransferase. The reaction takes place via the transfer of a peptidyl residue from peptidyl-tRNAin the ribosomal P site to the amino group of the aminoacyl-tRNA in the A site. Despite many years of intensive research, the nature and the basic mechanism of the ribosomal peptidyltransferase reaction is still largely unknown. Recently, Zhang and Cech[’] demonstrated that an in vitro-selected ribozyme can catalyze the same type of peptide bond formation as a ribosome. The ribozyme resembles the ribosome in such a way that a very specific RNA structure is necessary for substrate binding and catalysis, and both amino acids to be coupled are attached to nucleotides. Despite the presence of many different possible peptidyltransferase ribozymes, one of these must be strikingly similar in sequence and secondary structure to the “helicalwheel” portion of 2 3 s rRNA implicated in the activity of the ribosomal peptidyltransferase.These results from Cechs group demonstrate that a ribozyme is capable of catalyzing peptide bond formation analogous to the action of the ribosome, providing evidence that RNA itself can make peptides and support the “RNA world” hypothesis in biological evolution. Since the ribosomal peptidyltransferaseactivity is not suitable for practical use as a simple C - N ligase and, in addition, the multienzyme complexes involved in bacterial peptide synthesisr6Ido not seem to possess a general applicability,only the reverse catalFc potential of peptidases can be considered as valuable supplement to chemical coupling methods (cf. Sect. 12.5.3). In addition, peptidases have been used successfully for enzymatic manipulation of protecting groups in peptide synthesis 17-91.

72.5 Hydrolysis and Formation ofpeptides

12.5.2.1.2

Catalytic Mechanism[”*

The overall process of peptide bond scission is identical in all classes of peptidases and differences between the catalyhc mechanisms are rather subtle. The attack on the carbonyl group of the peptide bond requires a nucleophilic agent, either oxygen or sulfur, in order to approach the slightly electrophilic carbonyl carbon atom. To remove a proton from the attacking nucleophile, general base catalysis will assist this process. Furthermore, some type of electrophilic action on the carbonyl oxygen increases the polarization of the C - 0-bond. Generally, the four classes of peptidases (serine, cysteine, aspartic and metallopeptidases) differ in the groups that perform nucleophilic attack, general base catalysis, and electrophilic assistance. Also, different groups are involved in the breakdown of the tetrahedral intermediate which is formed in the initial nucleophilic attack, requiring general acid catalysis to promote the departure of the amine fragment. The four types of peptidases are based on the different catalytic mechanisms, which were first recognized by the use of some group-specificinhibitors. The reactive serine residue in the active site of serine peptidases (but also in other serine hydrolases, such as acetylcholine esterase) react in an irreversible step with organophosphate compounds, e. g. diisopropyl phosphofluoridate (DFP or DipF) resulting in the death of the appropriate enzyme. Owing to the high toxicity of DFP other reagents, e. g. phenylmethylsulphonylfluoride (PMSF) and 3,4-dichloroisocoumarin (3,4-DCl)have been used in its place. The reactive cysteine residue of cysteine peptidases is susceptible to oxidation and can react with various reagents: iodoacetate, N-ethyl-maleimide,heavy metals (for example Hg) and with the highly selective inhibitor N-[~-3-tr~~~-carboxy0xiran-2-carbonyl-~-leucyl-amido(4-guanidino)butane] (E-64).The highly acidic pH optima led to the first recognition of aspartic peptidases. Later, with pepstatin A from a strain of Streptomyces, a specific inhibitor was found. Chelating agents, e. g. EDTA and 1,lO-phenanthroline are prone to inhibit metallopeptidases. Serine PeptidasesI’21 These form the most studied class of peptidases. They have a reactive serine residue, e. g. the hydrolysis of a peptide substrate involves an acylenzyme intermediate in which the hydroxyl group of Ser19’ (from the chymotrypsin numbering system) is acylated by the acyl moiety of the substrate, releasing the amine fragment of the substrate as the first product. The formation of the acylenzyme is the slow step in peptide bond hydrolysis, but the acylenzyme often accumulates in the hydrolysis of ester substrates. The acylenzyme thus formed will be the same for a series of substrates which differ in their leaving group. The catalytic mechanism of serine peptidases will be given in terms of chymotrypsin (Fig. 12.5-5). After chymotrypsin has bound the substrate to form the Michaelis complex, the attack of Ser”’ on the peptide bond of the substrate forms a high energy tetrahedral intermediate. At the same time the proton of the serine hydroxyl group is transferred to the nearby His”, the serine hydroxy group forms a covalentbond with the carbonyl atom of the peptide bond to be cleaved. The liberated proton is taken by the imidazole ring of Hiss7thereby forming an imidazolium ion

806

I

12 Hydrolysis and Formation ofC-N Bonds

/His

Substrate polypeptide

lN-?\

H

Lo

Tetrahedral intermediate

Michaelis complex

I2

q<:

Ser

0-. -

0

Acyl-enzyme intermediate

(His

(His

4

1:F

-.O-C-R

Tetrahedral intermediate

+

IR

P-c,

H

O

Active enzyme

Figure 12.5-5. Scheme o f the catalytic mechanism o f serine proteases (chymotrypsin numbering).

(general base catalysis). This process is supported by the polarizing effect of the unsolvated carboxylate ion of Asp"' which is hydrogen bonded to Hiss7 in the sense of electrostatic catalysis. Mutagenic replacement of Asp'04 by Asn in trypsin, for example, did not changed the K M substantially at neutral pH. On the other hand, kcat

72.5 Hydrolysis and formation ofpeptides I807

was reduced to < 0.05% of its wild-type value. Furthermore, neutron diffraction studies have shown that Asp104 remains as a carboxylate ion rather than a proton being abstracted, as from the imidazolium ion of HisS7 to form an uncharged carboxylic moiety. The active site of serine peptidases is complementary in structure to the transition state of the reaction, a structure which is very close to the tetrahedral adduct of Ser19’ and the carbonyl carbon of the peptide substrate. Indeed, transition state binding catalysis provides the catalytic power of the appropriate serine peptidase. In the course of the formation of the tetrahedral intermediate a conformational distortion causes the carbonyl oxygen of the scissile peptide bond to move deeper into the active site to occupy the oxyanion hole. The resulting oxyanion is hydrogenbonded to the backbone of NH groups of G l ~ l ’and ~ Ser’”, whereas the NH group of the peptide bond preceding the scissile bond forms a hydrogen bond to the backbone carbonyl of Glyl”. The decomposition of the tetrahedral intermediate forming the acylenzyme intermediate and the amine product occurs under the driving force of proton donation from the N3-atom of Hiss7 through general acid catalysis. The Nterminal part of the cleaved peptide chain (amine product) will be released in the next step and replaced by a water molecule forming a second tetrahedral intermediate. The latter decomposes to the reaction’s carboxyl product (C-terminal portion of the cleaved peptide chain) and the active enzyme. Generally, all the serine peptidases employ the same catalytx three amino acid units to hydrolyze peptide bonds. The diversity of serine peptidases results entirely from the way they accommodate their specific substrates. Cysteine Peptidases l1 1’ Other terms for cysteine peptidases are cysteine-type peptidases, thiol peptidases or sulfhydryl peptidases. They are peptidases in which the attacking nucleophile is the ~ papain numbering system). The sulfhydryl group of a cysteine residue ( C Y Sin~ the mechanism of catalysis is similar to that of serine peptidases because a covalent intermediate is formed. Beside the cysteine nucleophile a proton donorlgeneral base is required, which in the majority of cysteine peptidases is a His residue (His”’). Despite the fact that in some families of cysteine peptidases a third amino acid residue is required to orientate the imidazolium ring of the histidine moiety in the course of the catalytic process, in general, only a catalytic dyad is necessary. The archetypal cysteine peptidase is papain which was isolated from the latex of the tropical papaya fruit (Carica papaya)[’3.141. It is a single protein of 212 amino acid residues containing three disulfide bonds and the three-dimensional structure is known with 1.65 8, resol~tion[~’]. The catalytic amino acid residues have been identified as Cys2’, His”’ and Asn”’, whereas Gin’’ helps to stabilize the oxyanion hole. A second group of cysteine peptidases which is very diverse in sequence is the “papain-like’’endopeptidasesof RNAviruses containing only the catalytic dyad Cys/His without any additional residues being involved in the catalytic mechanism. The same is true for caspases, a group of ten cytosolic endopeptidases with strict specificity for cleavage of aspartyl bonds. Clostripain from the anaerobic bacterium Clostridium histolyticum is a heterodimeric protein of 526 amino acid residues. The

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12 Hydrolysis and Formation ofC-N Bonds

heavy chain ( M ,- 43000 Da) and the light chain ( M , - 15398 Da) are held together by strong noncovalent forces rather than by disulfide bridges. Cys4’ of the heavy chain was identified as the catalytic residue of the active site. This peptidase is well known for the selective cleavage of arginyl bonds, whereas lysyl bonds are hydrolyzed at a lower rate. The catalytic mechanism of the adenovirus endopeptidase is similar to that of papain, the difference being that four amino acids His, Glu (or Asp) Gln and Cys are involved in it. Last but not least, the caspases with a strict specificity for cleavage of aspartyl bonds should be mentioned as the last family of cysteine peptidases. Members of this family transmit the events leading to apoptosis of animal cells.

Aspartic Peptidases The aspartic peptidases comprise peptidases which catalyze the hydrolysis of peptide bonds without the use of nucleophilic attack by a functional group of the enzyme. The nucleophile attacking the scissile peptide bond in this case is an activated water molecule and no covalent intermediate will be formed between the enzyme and a fragment of the substrate. The name of this group of peptidases is based on the catalytic domain which consists of two aspartic acid side chains (Asp32and Asp215of the porcine pepsin numbering system) activating the water molecule directly. These two side chain carboxyl groups are close enough to share a hydrogen bond between two of their oxygens holding the water in place. However, there are not two Asp residues in the catalytic dyad in all members of aspartic peptidases. An endopeptidase from nodavirus has an Asp and an Asn as catalytic residues, and in a related tetravirus endopeptidase the Asp residue is replaced by Glu. It is interesting to note that all the enzymes so far described are endopeptidases. Metallopeptidases As with the aspartic peptidases, metallopeptidases do not form covalent intermediates and the nucleophilic attack on the peptide bond to be cleaved is mediated by a water molecule. The latter is activated by a divalent metal cation, usually Zn2+ but sometimes also Co2+or Mg2+.In order to assist in attack of a water molecule the metal ion provides a strong electrophilic “pull”. The metallopeptidase has a water molecule coordinated to the fourth tetrahedral site. Beside the metal ion the other ligands are two histidine building blocks and a glutaminic acid residue in thermolysin and carboxpeptidase A. The enzymes of this family can be divided in two groups depending on the number of metal ions necessary for catalysis. In many cases only one zinc ion is required, but often two metal ions act cocatalytically.All the enzymes which contain cobalt or manganese require two metal ions, but zinc-dependent enzymes are also known in which two zinc ions act in a cocatalytic manner. Enzymes known to date containing cocatalytical metal ions are exopeptidases, whereas those with one catalyhc metal ion belong to exopeptidases or endopeptidases. His, Glu, Asp or Lys are known metal ligands in metallopeptidases. Together with the metal ligand very often a Glu residue is engaged in the catalytic process. In the leucyl aminopeptidase Lys or Arg fulfill this function.

12.5 Hydrolysis and Formation ofpeptides

12.5.2.1.3

E.C. Classification

As shown above, based on the chemical groups that are responsible for their catalytic activity, peptidases have been classified into four distinct groups. Recommended by the International Union of Biochemistry and Molecular Biology (1992) [16] all hydrolases are designated as E.C. 3., and the peptidases as E.C. 3.4. defining the main classes of peptidases by a third numeral (11to 24) as indicated in Table 12.5-1. The sub-subclasses are not further divided. The enzymes are listed in arbitrary order within each of them. Unfortunately, the molecular structures and evolutionary relationships are not taken into account in the E. C. classification. In this E. C. list the exopeptidases are mainly classified based of their action. Generally, only peptides with an unblocked terminus are attacked. The only exception are so-called omega peptidases which comprise a very small number that are capable of releasing certain modified terminal residues. To this group belong, for example, acylaminoacyl peptidases which release acetyl or formyl moieties from the N-terminus, and the pyroglutamyl peptidase, capable of releasing the cyclic residue. An isopeptide bond will be cleavaged by the P-aspartylpeptidase. Other omega peptidases are directed to the substituted C-terminus, e. g. the peptidyl glycinamidase releasing a C-terminal glycine amide, and the y-glutamyl carboxypeptidase which splits a C-terminal glutamic acid linked by an isopeptide bond.

12.5.2.1.4

Peptidase Families and Clans

Starting with the earlier work of Rawlings and Barret1l7]and improved in the handbook[4]another level of sophistification to the classification of peptidases has been developed. Evolutionary considerations can be taken into account due to the relative ease by which cDNA-derived sequences can now be obtained. According to this principle of classification a family of peptidases is defined as a group in which Table 12.5-1. Principles of peptidase classification according to the Enzyme Commission (E. C.) of the international Union of Biochemistry and Molecular Biologyl’61. E.C. Number

Exopeptidases 3.4.1 1.3.4.14.3.4.14.3.4.15.3.4.16.3.4.17.3.4.18.3.4.19.Endopeptidases 3.4.21.3.4.22.3.4.23.3.4.24.3.4.99.-

Type of peptidase

Type of cleavage

Aminopeptidase Dipeptidase Dipeptidyl peptidase Tripeptidyl peptidase Peptidyl dipeptidase Carboxypeptidase (serine) Carboxypeptidase (metallo) Carboxypeptidase(cysteine) Omega peptidase

N-terminal residue Dipeptides only N-terminal dipeptide N-terminal tripeptide C-terminal dipeptide C-terminal residue C-terminal residue C-terminal residue Terminal modified residue

Serine endopeptidase Cysteine endopeptidase Aspartic endopeptidase Metalloendopeptidase Endopeptidase with unknown mechanism

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72 Hydrolysis and Formation ofC-N Bonds

every member indicates a statistically significant relationship in the amino acid sequence to at least one other member of the family in the part of molecule which is responsible for peptidase activity. Applying strict statistical criteria implies confidence that any two peptidases that are placed in the same family have evolved from a common ancestor and thus are homologous according to the definition of Reeck et al. ['*I. Each peptidase family is named with a letter that denotes the catalpc type (S, T, C, A, M or U, for serine, threonine, cysteine, aspartic acid, metallo- or unknown), followed by an arbitrarily assigned number ( see Table 12.5-2). The term clan is used for defining a group of families the members of which have evolved from a single ancestral protein, but have diverged so far that their relationship can no longer proved by comparison of the primary structures. Clan-level relationships between families can at best be made evident by similarities in three-dimensional structures. The name of the clan is formed from the letter for the catalytic type (in analogy to families) followed by an arbitrary second capital letter. About 40 families of serine- and threonine-type peptidases can be distinguished on the basis of sequence comparison. However, only a few known families of threonine-dependent peptidases are included therein. By comparing the tertiary structures and the order of the catalytic residues in the sequence most of these families can be grouped into seven clans (cf. Table 12.5-2). The serine peptidases and their clans can be used to demonstrate this type of classification in more details. In clan SA with the order of the catalytic triad His, Asp, Ser the tertiary structure is characterized by a p sheet-based two-domain structure. Each domain contains a p barrel and between the domains the active site cleft is located. The largest family S 1 of trypsin consists of more than 70 sequenced proteins. Well-known members of the family S2 are, e. g. streptogrisin A, glutamyl endopeptidase, and lysyl endopeptidase (Achromobacter). Togavirin (S3), IgAl-specific serine-type prolyl endopeptidase (SG), flavivirin (S7), hepatitis C polyprotein peptidase (S29), helper component proteinase (S30), pestivirus NS2-3/NS3 serine peptidase and arterivirus serine endopeptidase (S32) complete the families of clan SA. The order of the catalytic triad of clan S B is Asp, His, Ser and the tertiary structure contains both p sheets and a helices. This clan contains only the subtilisin family (S8) including peptidases from archaea, bacteria and eukaryotes. Clan SC contains peptidases with the alp hydrolase fold bearing the catalytic triad in the order Ser, Asp, His. This clan includes the families (characteristic member in parentheses) S9 (prolyl oligopeptidase), S10 (carboxypeptidase C), S15 (Xaa-Pro dipeptidyl -peptidase), S28 (lysosomal Pro-Xaa carboxypeptidase), S33 (prolyl aminopeptidase), and S37 (StreptomycesPS-10 peptidase). The characteristic catalytic dyad Ser, Lys of clan SE is represented by the motif Ser-Xaa-Xbb-Lys,and the fold consists of helices and an a+p sandwich. The families of this clan S 1 1 (penicillin-binding protein 5), S12 (Streptomyces RG1 D-Ala-D-Ala carboxypeptidase), S13 (penicillinbinding protein 4) are involved in the biosynthesis, turnover and lysis of bacterial cell walls. The catalytic residues in clan S F (catalyticdyad Ser, Lys or Ser, His) are more widely spaced in comparison with clan SE. The families of this clan include only endopeptidases from bacteriophages, bacteria, archaea and eukaryotes with the members: S24

12.5 Hydrolysis and Formation ofpeptides Table 12.5-2. Evolutionary classification o f peptidases into families and clans based on primary and teriary structure. Class (E.C. list)

Serine (E. C. 3.4.21.)

Cysteine (E.C. 3.4.22.)

Aspartic (E.C. 3.4.23) Metallo (E.C. 3.4.24)

Families

Clans (families)

Catalytic residues

SA (Sl-3,6,7,29-32,35,43)

His, Asp, Ser

SB (S8) SC (S9,10,15,28.33,37) SE (Sll-13) SF (S24,26,41,44) SH (S21) TA (S42) SX (14,16,18,19,34,38,39,43) CA (C1,2,10,12,19)

Asp, His, Ser Ser, Asp, His Ser, Lys Ser, Lys, (His) His, Ser, His Thr

CB (C3,4,24,30,37,38) CC (C69,1G,21,23,27-29,31-36, 41-43) CD (C14) CE (C5) CX (C11,13,15,22,25,2G,39,40) AA (A1-3,9,10-18)

His, Cys Cys, His

AB (AG, 21) MA (M1,2,4,5,9,13,30,36,48)

Asp, Asn His, Glu, His (HEXXH) His, His/Asp (HGXXHXXGXXH/D) His, Glu, His (HXXE/H) His, His, Asp (HMYGHAAD) His, Glu, His (HXXEH) LYS, Asps, Glu (NTDAEGRL) Aspz, His, Gluz His, Asp,, Glu

MB (M6-8,10-12) MC (M14) MD (M15) ME (M16,44) MF (M17) MG (M24) MH (M18,20,25,28,40,42) MX (M3,19,22,23,26,27,29,32,34-38, 41,43,45,47)

Cys, His, Asp (Asn)

His, Cys His, Glu(Asp),Gln, Cys Asp, Asp

(Lex A repressor), S2G (signal peptidase I), S41 (TSP protease), and S44 (tricorn protease). All known members of clan SH (catalytic triad: His, Ser, His) are endopeptidases from DNA viruses which are involved in virus prohead assembly. The clan includes only the family S21 (Cytomegalouirusassemblin). Clan TA with the catalyhc residue Thr, Ser or Cys, and an a,p,a,p sandwich fold includes a number of peptidases whose only proteolytic activity is self-activation.Important families of this clan are T1 (proteasome),and S42 (y-glutamyltranspeptidase). Other families (clan SX) of serine peptidases including S14 (endopeptidase Clp), S1G (endopeptidase La), S18 (omptin), S19 (cell wall-associated endopeptidase of

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12 Hydrolysis and Formation ofC-N Bonds

Trichophyton), S34 (HflA endopeptidase), S38 (Treponema chymotrypsin-like endopeptidase), S39 (cocksfoot mottle virus endopeptidase), S43 (porin) cannot yet be assigned to clans, since neither the tertiary structure nor the order of catalytic residues are known. The cysteine peptidases comprise the clans CA, CB, CC, CD, CE and CX. The last includes a number of other families of cysteine peptidases for which tertiary structures are unknown and virtually nothing is known about the specificity of the catalytic machinery. The clan CA contains papain and its relatives. Papain was the first clearly studied cysteine peptidase. From the crystal structure of papain and a few closely related peptidases of the family C1, it could be concluded that the catalybc residues are Cys, His and Asn in that order of sequence. Further members of C1 are the cathepsine B, H, K, L and 0, the dipeptidyl peptidase I, and glycyl endopeptidase. The C2 family contains various calpains, whereas streptopain belongs, to C10 ubiquitin C-terminal hydrolyse PGP 9,s to C12, and the isopeptidase T to C19. Clan CB contains viral “chymotrypsin-like” cysteine peptidases that process the viral polyproteins, and in clan CC are listed viral “papain-like’’endopeptidases. The only family of clan CD (C14) comprises a number of cytosolic endopeptidases which cleave aspartyl bonds with high specificity. This family of caspases consists of ten members from which caspase-1 and caspase-3 are best known. The mature caspase1, processed from a single-chain precursor by presumably autocatalytic cleavage of four aspartyl bonds, is a heterodimer of a 22 kDa heavy chain and a10 kDa light chain [I9]. This peptidase was formerly known as interleukin 1P-converting enzyme (ICE) since it mediates, among other things, the processing of interleukin l p at aspartyl bonds. Human caspase-3 is also a heterodimer consisting of the subunit p12 (11896 Da) and the subunit p17 (16617 Da) with a tertiary and quarternay structure similar to caspase-1 L20]. This peptidase appears to function in order to proteolytically inactive proteins which are involved in cellular repair and homeostasis during the effector phase of apoptosis. Clan CE contains only the adenovirus endopeptidase[”]. A catch-all clan CX comprises all other families of cysteine peptidases which could not been classified up to now due to the lack of necessary data of structure and catalytic maschinery. Aspartic peptidases have so far been described for all endopeptidases. Unfortunately, the tertiary structure has only been elucidated for four families. Endopeptidases of the family A 1 consist of two lobes, with the active site between them. One lobe has been derived from the other by gene duplication. In the active site each lobe, with very similar three-dimensional structures, bears one Asp residue of the catalytic dyad. It is interesting to note that the crystal structure of retropepsin from family A2 of clan AA showed a single lobe with one catalytic Asp residue with structural similarity to one lobe of the pepsin from family Al. Retropepsin is only active as a homodimer forming the catalytic site between the two monomeric molecules. There is evidence that the peptidases of families A1 and A2 have evolved from a common ancestor. Unfortunately, a number of other families could not yet been assigned to any clan. Metallopeptidases are allocated to eight clans. A couple of families could not yet be

12.5 Hydrolysis and Formation of Peptides

assigned to these clans since, in particular, the metal ligands have not been biochemically characterized. Zinc-dependent metallopeptidases,both exopeptidases and endopeptidases, with the HEXXH motif are listed in the clan MA. The family M4 contains along with thermolysin, and elastase (Staphylococcus) well-known peptidases. The tertiary structure has been determined for members of this family showing a two-domain structure with the active site between the domains. The Nterminal domain contains the HEXXH motif and includes both a helices and /3 sheets as dominating structure elements and shows some similarities to the domain structure of clan MB. In the C-terminal domain are five helices in a closed bundle. This characteristic fold is typical of thermolysin-like peptidases. Clan MC contains metallocarboxypeptidases which belong to only one family (M14) which is divided into the subfamilies A, B and C. Typical for this clan is that one zinc ion is tetrahedrally coordinatedby a water molecule, two histidine and one glutamate residues. Clan MF includes aminopeptidases that require cocatalyhc zinc ions for their enzymatic activity. The well-known leucyl aminopeptidase has a two-domain structure bearing the active site in the C-terminal domain. Whereas exopeptidases of clan MG require cocatalytic ions of cobalt or manganese, clan MH contains the third group of metallopeptidases that also require cocatalytic metal ions, but here these are all zinc ions. The third clan in which cocatalytic metal ions are necessary is clan MF with zinc or manganese. Only one catalytic zinc ion is required for peptidases of clans MA, MB, MC, MD and ME. 12.5.2.2 Importance of Proteolysis

Historically, enzymatic proteolysis has generally been associated with protein digestion. Therefore, the digestive peptidases of the pancreatic and gastric secretions are among the best characterized peptidases and much of the current knowledge of structure and function has been derived from investigations of those proteolytic enzymes. Activation of the pancreatic digestive enzymes is initiated by enterokinase, an enzyme secreted by the mucous membrane of the stomach. It converts some trypsinogen into active trypsin, which then activates all the proenzymes, including more trypsinogen. The function of the digestive proteases is merely to breakdown all the proteins they encounter. Later, it became evident that peptidases play regulatory roles in a great variety of physiological processes [22* 231. These include processing and molecular assembly of nascent polypeptide chains, and the processing of protein hormones, developing enzyme precursors to mature enzymes, fertilization, many other proteolytic processes important for cellular functions, and the regulation of the programmed cell death (apoptosis).The last is a mechanism that regulates cell number and is vital throughout the life of all animals. Apart from various biochemical events involved in apoptosis, the most fundamental one is the participation of members of the caspase family in both the initiation and execution phases of cell death. The mechanism of activation of the caspases constituting the different apoptosis-signaling complexes

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12 Hydrolysis and Formation ofC-N Bonds

can be explained by an unusual capability of the caspase zymogen to autopress to an active Proteolytic processing occurs in many different ways and is triggered by different proteases. Limited proteolysis is the key to this selectivity which depends on the accessibility of the scissile peptide bond to the acting peptidase and on its specificity. In this cases proteolysis is directed and limited to the cleavage of specific bonds in the target protein. A wide variety of prokaryotic and enkaryotic proteins are synthesized as larger pre- or pre-proforms. Some of these are biologically inactive and become activated upon limited proteolysis. Lysomal enzymes, mitochondria1 proteins, membrane proteins, secreted proteins etc. undergo intracellular proteolytic maturation. Various viruses code for specialized peptidases which are essential for virus A couple of viral peptidases are interesting therapeutic targets. An extremely large number of publications have been dedicated to the aspartic peptidases, especially to the enzyme of the human immuno deficiency virus (HIV),which is a key target in the treatment of AIDS. HIV-1 protease (HIV-1 PR), more exactly named as human immunodeficiency virus 1 retropepsin (HIV-1retropepsin; E. C. 3.4.23.16), has become the most thoroughly investigated system in the history of peptidases. The biological function of the retroviral peptidase is to cleave the polyprotein precursor into its constituent functional units such as the matrix, capsid, and nucleocapsid structural proteins of HIV to permit assembly. For this reason, the great interest in HIV-1 retropepsin has centered on the development of compounds that selectively inhibit the viral enzyme and not the related human aspartic peptidases. Useful principles of inhibition have been combined by several companies to produce antiviral compounds that have achieved approval from the Food Despite the development and Drug Administration (FDA) in the USA (cf.review[25]). of extremely strong and selective inhibitors which have been demonstrated to be effective in human trials one major problem remains: the extremely rapid development of forms of the virus that are resistant to the drugs containing the inhibitors. Secretory proteins are usually synthesized as precursors bearing an aminoterminal extension. The signal peptide is removed co-translationallyby signal peptidases during translocation across the membrane. In the next step precursors of protein hormones, growth factors and certain polycistronic precursor proteins are processed by specific enzymes. In contrast to consecutive zymogen activation consecutive prepro-cleavage reactions are regulated independently. The pathway of processing of many pre-proteins is known, but many of the maturation peptidases can not yet be characterized. For this reason, the application of molecular cloning techniques will be helpful in the near future for the sequence elucidation of pro-proteins as well as the cDNA and genomic sequences for maturation enzymes. The structure changes range from the relatively simple alterations in zymogen activation to more complex processing events in multidomain peptidase precursors, such as prothrombin or plasminogen. Generally, proteolpc processing induces intramolecular rearrangements required for the expression of biological response. Like blood coagulation,the complementary system is triggered by a signal that activates several consecutive zymogen activation reactions. This later system takes part in the immune reaction

72.5 Hydrolysis and Formation ofpeptides

directed against foreign organisms of tissues. Several components of the complementary system are serine peptidases. Peptidases as integral components of cells have only been partly explored, e. g. lysosomal peptidases, granulocyte serine peptidases, membrane-bound peptidases, and enzymes of specialized tissues, such as the reproductive tract, skins, lens, muscle, pituitary, adrenals etc. Various ATP-dependent peptidases have been isolated. The proteasome is a large multifunctional protease complex that degrades intracellular proteins. The name is derived from protease (“protea-”) and large particle (“-some”) This complex is an exception among peptidases as regards the nucleophilic residue and the general structure. Both in eukaryotes and archea, the proteasome is a multisubunit complex comprising four stacked rings each containing seven subunits (M, -20-30 kDa). In eukaryotes the 20s proteasome (E.C. 3.4.99.46; also named multicatalytic proteinase, macropain and prosome) has the form of a hollow cylinder (length 148 A, diameter 113 A). It shows several different catalytic activities and contains 14 different but homologous subunits, whereas in archea there are just two different kinds of subunits, and the enzyme complex possesses only one catalytic activity. In bacteria the proteasome is built up of two rings of six subunits. One of the two different subunits is related to the eukaryote and archaen proteasome subunits, the other is an ATPase. The 26s protea~ome[’~,”1 ( M , - 2100 kDa) consists of the 20s proteasome and at least one other multisubunit regulatory protein known as PA700, 19s cap, p-particle, ball, and ATPase complex. It was first found in extracts of rabbit reticulocytes by its capabilityto degrade ubiquitinated proteins in an ATP-dependent manner. Since this complex can also degrade various nonubiquitinated proteins the older designation ubiquitin-conjugate degrading enzyme (UCDEN) is probably inappropriate. The 20s proteasome subcomplex of the 26s proteasome containing multiple catalytic sites with distinc specificities is responsible for the whole proteolytic activity. In addition, the PA700 regulatory complex displays further enzymatic activities, such as ATPase activity, isopeptidase activity and seems to contain a substrate protein unfolding activity. The ATPase activity is necessary for assembly of the 26s proteasome from the 20s proteasome and PA700 subcomplexes and also for the degradation process. Since peptide bond hydrolysis is not energy dependent, the hydrolysis of ATP might be required for unfolding protein substrates and/or for translocation of the unfolded peptide substrate into the central channel of the proteasome. The proteasome is responsible for turnover of most cellular proteins in mammalian cells and the selective degradation of proteins with abnormal structures. Last but not least, the proteasome is involved in the production of antigenic peptides for presentation by MHC class I complexes. The generation of antigenic peptides seems to be performed by a specific subpopulation of proteasomes containing two or three subunits encoded in the major histocompatibility complex. Considerable attention has been paid to a group of intracellular serine peptidases associated with granulocytes as well as leukocytes and mast cells as mentioned above. These peptidases are stored in granulas and released in response to inflammatory or allergic stimuli. Many of the peptidases are relevant to human health

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12 Hydrolysis and Formation of C-N Bonds

and disease r2’1, some as natural components of the human body, and others because they are important in species which provide us with food, or cause diseases. In oder to understand proteolytic activity in biological processes, knowledge of the contribution of the natural peptidase inhibitors to the regulation of the activity is Inhibitors are as diversified as the proteases themselves. Generally, they can be divided into two main classes: (a) active site-specific low-molecular-mass inhibitors, and (b) naturally occuring protein peptidase inhibitors. Examples of the first group are the serine peptidase inhibitors diisopropyl phosphofluoridate (DFP) and phenylmethanesulphonyl fluoride (PMSF).Both react with the active site serine. Many of the naturally occuring peptidase inhibitors, isolated from animal, plant and bacterial organisms, behave as pseudosubstrates. They combine essentially irreversibly with the active site and are converted into a modified form in which a peptide bond, related to the primary substrate specificity of the peptidase, is cleaved. Of special physiological interest are inhibitors which react with mammalian plasma serine peptidases, especially those involved in blood coagulation. In principle, such inhibitors have both protective and regulatory functions. Approximately 10% of the nearly 200 proteins in blood serum are peptidase inhibitors. ?lie alproteinase inhibitor secreted by the liver, for example, inhibits leukocyte elastase which is thought to be part of the inflammatory process. Furthermore, special variants of this inhibitor with reduced inhibiting power are associated with pulmonary emphysema. The latter is a degenerative disease of the lungs which results from the hydrolysis of its elastic fibers. Interestingly, certain plants release peptidase inhibitors in response to insect bites in order to inactivate the digestive enzymes of the attacking insect. Peptidases are valuable tools in the study of the primary and higher-order structure of proteins r3l]. Proteolysis of proteins for sequence analysis and peptide mapping can be carried out according to different Based on the extent of proteolytic reaction, it is allowed to reach completion or it is prevented from reaching completion. In the first case the products constitute an equimolar set of peptides whose composition will not be influenced by further digestion with same enzyme. Depending on the restriction imposed by the primary specificity of the peptidase used, a protein will be fragmented to varying degrees. The fragments can subsequently be separated and sequenced. Combining these data with sequence data of other overlapping sets which are generated with different peptidases allows the reconstruction of the sequence. If proteolysis is prevented from reaching completion a different set of data is obtained. Inhibition or removal of the peptidase are desirable interventions to determine the initial cleavage products. Furthermore, peptidases are also structural probes of conformation of soluble and two-dimensionalN M R L3’1 are the proteins [331. Although X-ray crystallography[34] methods of choice for the determination of the three-dimensional structure of globular proteins, some weaknesses of these techniques demand alternative methods even if these will provide structural information at a lower level of resolution. For example, limited proteolysis can be used to probe the structure and the dynamics of proteins in solution, which provide experimental data that are easy to obtain and complement well those results derived from the techniques mentioned above. The

72.5 Hydrolysis and Formation ofpeptides

goal of investigations are soluble proteins in their native or near-native states. Limited proteolysis occurs in this case at the level of only one or very few peptide bonds which leads to the formation of “nicked proteins. This species of proteins consists of rather large fragments which remain associated in a stable and often also functional complex. Usually, a nicked protein is much more labile than the native form. Therefore, the unfolding leads to a suitable substrate for an extensive proteolytic degradation to small peptides, and further proteolysis is much faster in comparison with the initial peptide bond cleavage at the level of the native protein. Consequently, in this case, during proteolysis the intact protein and small proteins are present in the incubation mixture, without intermediate sized products. In the case where nicked proteins are sufficiently stable, they may resist further extensive proteolytic degradation and can be isolated and characterized. It is assumed that the limited proteolysis phenomenon derives from the fact that a specific polypeptide chain segment of the compact, folded protein substrate is exposed and flexible so that it can fit the active site of the appropriate peptidase for an efficient and selective limited hydrolysis. There is no doubt that enhanced chain flexibility or segment mobility is the key feature of the site of peptide bond hydrolysis demonstrated by a clear-cutcorrelation between sites of proteolytic attack and sites of enhanced chain flexibility. The present availability of automatic, efficient and sensitive techniques of protein sequencing and, particularly, the recent dramatic advances of mass spectrometry[36]in the analysis of peptides and proteins, allows a more systematic use of the limited proteolysis approach as a simple first step in the elucidation of structure-dynamics-function relationships for novel proteins which are only available in minute amounts. Since a growing number of newly discovered peptidases are specifically expressed in single tissues, especially, at low expression levels or often only at certain development stages, it is very complicated to isolate the enzymes in sufficient quantities using classical biochemical procedures. Therefore, the only alternative is the cloning and expression of these peptidases. In addition, recombinant techniques allow directed structural alterations in order to program mechanistic or functional features. Peptidases can be expressed in most of the developed expression systems (yeast, viral, bacterial, insect cells and mammalian). It is not usually easy to predict which expression system is the method of choice. For functional expression of recombinant peptidases various examples have been presented [37]. Last but not least, it should be mentioned that a couple of peptidases have industrial importance. In particular, since subtilisins have a broad substrate specificity and are highly stable at neutral and alkaline pH they are of considerable industrial interest as protein-degrading additives to detergents. These reasons combined with their large data base make subtilisins attractive for protein engineering. Extensive engineering studies have been carried out on the Bacillus subtilins and more than 500 site-directed mutants have been produced to alter specific enzyme properties, such as pH profile, thermal stability or substrate specificity (see e.g. referenceSL37-391 ).

I

817

818

I

72 Hydrolysis and Formation of C-N Bonds

12.5.3 Formation of Peptides 12.5.3.1

Tools for Peptide Synthesis

Although the origins of peptide chemistry are usually traced back to the early 20th century when Emil Fischer obtained the simplest dipeptide glycyl-glycineby cleavage of the appropriate diketopiperazine, the first peptide bond in a chemical laboratory was synthesized by the young Theodor Curtius in the laboratory of Hermann Kolbe at Leipzig University in 1881. Despite the fact that Emil Fischer with co-workers in Berlin made basic contributions to peptide synthesis, the productive epoch of peptide chemistry began some decades later in the1950s. Wieland and Bodan~zky[~'] have written an excellent account of the history of peptide synthesis. Peptides belong to an increasingly important class of bioactive molecules in physiology, biochemistry, medicinal chemistry and pharmacology. They act as hormones, neurotransmitters, cytokines, growth factors etc. However, it is not only naturally occuring physiologically relevant peptides that are the subjects of interest. Peptide analogs possessing agonist or antagonist activity are also useful tools in investigations when searching for suitable drugs. Radiolabeled analogs and molecules bearing affinity labels have been applied for the characterization and isolation of receptors. Furthermore, peptides are useful as substrates of peptidases, kinases, phosphatases and special transferases in investigations on enzyme kinetics, and mechanisms of action. In the preparation of polyclonal and monoclonal antibodies peptides play an important role as synthetic antigens, and epitope mapping using synthetic peptides has been developed as a valuable approach for the identificationof specific antigenic peptides for the preparation of synthetic vaccines, and also for the determination of protein sequence regions which are important for biological function. In addition, the design of small peptide mimetics of protein function or structure, and the development of various peptidomimetics in drug development are further goals in peptide chemistry. In particular, in the last ten years the number of known peptides has doubled and besides the development of efficient chemicals for peptide synthesis methods, the field of peptide and protein chemistny has been opened up to molecular biology and genetic engineering. The classical chemical peptide synthesis is a synthesis in a homogeneous s o l ~ t i o n [ ~ ~Even - ~ ~ in ] . the 1950s this approach had started to gain industrial importance followed by the solid-phasetechnique in the early 19GOs, invented by the Nobel laureate Bruce Merrifield147-501. The most fundamental time-consuming operations in chemical peptide synthesis (sometimes not free from undesirable side reactions) are the selective protection, and after synthesis the deprotection of the aamino function, the carboxyl group and the various side chain functionalities of trifunctional amino acids. Despite the development of numerous efficient protection methods based on chemical techniques, the whole process is rather slow as all intermediate products have to be purified and characterized after each reaction step. The formation of each peptide bond requires the activation of the carboxylic acid function of the carboxyl moiety.

72.5 Hydrolysis and Formation of Peptides

An important point in selecting a coupling method is its degree of safety from racemization, since all synthetic operations carried out at a center of chirality have this permanent risk. Therefore, the synthesis of peptides with a multitude of chiral centers continues to be a formidable chemical effort. The existence of more than 150 chemical variations for peptide bond formation indicates that an ideal coupling method does not exist, e.g. a fast procedure without racemization or other side reactions to realize quantitative coupling of equimolar amounts of the carboxyl and amine components. There is no doubt that the use of well known strategies and the application of activation methods with well established safety steps to protect from racemization in simple model systems does not assure the loss of optical purity during a multitude of coupling steps in the synthesis of medium-sized and long peptides. For the chemical peptide synthesis in homogeneous solution, which still plays an important role in the production of large quantities of peptides for pharmaceutical use, highly skilled personel are required. In this manner multikilogram quantities or even tons of peptides consisting of the range of 2-30 amino acid residues can be produced. Since peptides for research purposes are usually required in only mg tog can be used. amounts, the time-saving solid-phase peptide synthesis The strategy is in principle similar to that in solution, with the difference that there is no need for isolation of the intermediate products. As the growing peptide chain is synthesized on a suitable resin the whole procedure lends itself to automation. The drawback is that every reaction step at the resin has to be forced to give an almost 100% yield. In practice, this cannot be accomplished,with the consequence that the desired product must be isolated from a mixture of side-products by the final, normally HPLC, purification procedure which is sometimes difficult to perform and also expensive. Peptides of up to - 50 amino acid residues are now readily accessible using stepwise solid-phaseprocedures C5O1. An alternative for the preparation of larger polypeptides and proteins is the biotechnological production (geneticengineering, recombinant DNA technology,) in bacteria, yeast, or cultured mammalian cells [51-531. In principle, this is an economic way to produce peptides of more than 50 amino acid residues and even small proteins with complicated glycosyl or other groups attached to amino acid side chains. Compared with the problems connected to the chemical synthesis strategies, recombinant techniques provide quite a different set of problems. Whereas the principle of the expression of a gene in host cells through the normal biosynthetic and genetic machinery of the host cell using a suitable expression vector is relatively simple, putting this technique into practice poses some problems: Selection of the appropriate expression strategy, and the host cell system as well as the optimal vector system, the control of the stability of mRNA and also of the translated protein, isolation and purification of the product, scale-up,downstream processing etc. The development of cloning vectors which propagate in eukaryotic hosts, e.g. yeast or cultured animal cells, has in particular eliminated many of the problems associates with the synthesis of eukaryotic proteins. It should be noted that posttranslational processing may also vary among different eukaryontes. It is an advantage that shuttle vectors are available that are capable of propagating in both

820

I

72 Hydrolysis and Formation ofC-N Bonds Figure 12.5-6. Principle of native chemical ligation according to Dawson

-/(o>s->-] SR Step 1

H2N

et al.[551.

1

Trans thioesterification

0 Peptide 2 H2N Step 2

i

S to N Acyl transfer

yeast and E. coli and thus transfer genes between these two cell types. Recombinant protein production is of great medical, agricultural, and industrial importance [541. For example, human insulin, human growth factor, erythropoietin, various types of colony-stimulating factors, blood clotting factors are typical examples of recombinant proteins which are in routine clinical use. Despite the fact that heterologous expression of recombinantly cloned genes is by far the most commonly employed method of to engineering proteins this approach is only applicable to naturally occurring amino acids. This limitation is in principal overcome by unnatural amino acid mutagenesis [s41 and some other chemistrydriven approaches. Among the various chemical ligation methods the so-called “native chemical has proved to be a useful route to fully synthetic proteins 155, 56-60] . A s shown in Fig. 12.5-6 this procedure relies on the reaction that occurs between a peptide fragment possessing an essential N-terminal cysteine residue (peptide 2; Fig. 12.5-6), which can be expressed in principal using recombinant DNA procedures, and a second peptide fragment possessing an athioester group (peptide 1; Fig. 12.5-6).In an initial intermolecular, chemoselective reaction a thioester-linked intermediate is formed (step 1) which spontaneously rearranges via S+ N acyl transfer to the final amide-linked product (step 2). The rearrangement step corresponds mechanistically to an intramolecular S+ N acyl transfer reaction described by Wieland et al. r61] in 1953. Pulling together protein splicing (for a review see reference[”]) and native chemical ligation led to “expressed protein ligation” (EPL) [631 or also termed “inteinmediated protein ligation (IPL) [641. As shown in Fig. 12.5-7 the protein fragment of interest is expressed in E. coli as an intein-CBD (chitin binding domain) fusion protein. The chitin binding domain allows protein affinitity purification using chitin

72.5 Hydrolysis and Formation of Peptides

Expressed protein ligation Clone gene into intein vector

Figure 12.5-7. Principle of expressed protein ligation according t o Muir et a1.[63].

Express in

E. coli

0

N 1 Recomb. protein

HSJ

~--J--NH-C~~-/GH-

k

Affinity purification

Contaminants

HSj

0

N+ Recomb. protein P N H - C y s

4

N to S Acyl transfer

0 N

Recomb. protein

Synthetic peptide (both in large excess)

Native chemical ligation

QUICK

Semi-synthetic protein

beads. The necessary expression vector is commercially available. The N-t S acyl transfer results in a thioester-linked intermediate. In the next step a large excess of a suitable thiol agent (for example thiophenol) generates, by trans-thioesterification in situ, the protein a-thioester which reacts quickly with the simultaneously added synthetic amine component. The latter has to bear an N-terminal cysteine residue. Customized peptides containing N-terminal cysteine residues are available from a variety of sources. There is no doubt that the extension of the native chemical ligation to EPL led to significant progress in protein semi-synthesi~[~~I, despite the remaining requirement of an N-terminal cysteine residue in the amine fragment to

I

822

I

12 Hydrolysis and Formation of C-N Bonds

be coupled. Apart from these advantages it must keep in mind that direct reaction of a thiophenyl ester with the amine component could result in partial epimerization of the C-terminal amino acid residue of the protein a-thioester. 12.5.3.2 Choice of the Ideal Enzyme

Enzymes have become valuable tools in medium to large-scale synthetic organic Owing to the fact that hydrolases possess a wide substrate spectrum and do not usually need cofactors for their catalpc function, they are at present the enzymes most widely used as biocatalysts in preparative organic chemistry. Among the hydrolases the huge family of peptidases plays an important role in various processes of proteolysis as shown above. Unfortunately, a universal C - N ligase with a high catalytic efficiency for all possible combinations of the 21 proteinogenic amino acids both as C- and Nterminal amino acid residues, respectively, in fragments to be coupled could not be developed during evolution. Such heavy demands on specificity could not even be solved by nature. Therefore, protein biosynthesis has been developed as a step-wise strategy starting with the N-terminus of the growing peptide chain and catalyzed by the ribosomal peptidyltransferase. Limited proteolysis of the biosynthesis precursor molecules and posttranslational modifications provide the bioactive peptides and proteins. In nature the peptide bond formation is accomplished on the ribosome and takes place via the transfer of a peptidyl residue from the peptidyl-tRNA in the ribosomal P site to the amino group of aminoacyl-tRNA in the A site. Despite intensive investigations in recent years, the nature and the basic mechanism of the peptidyltransferase reaction within the ribosome is largely unknown. According to recent studies from the Nobel laureate Thomas R. Cech and coworkers[5]an in uitro selected ribozyme is capable of catalyzing the same type of peptide bond formation as a ribosome, e.g. its sequence and secondary structure seems to be strikingly similar to the “helical wheel” portion of 23s rRNA implicated in the activity of the ribosomal peptidyltransferase. These results provide evidence for the feasibility of the “RNA world” hypothesis by demonstrating that RNA itself is capable of catalyzing peptide bond formation. Furthermore, from these findings the idea that the rRNA has a catalpc function in the ribosomal peptide bond formation is supported. It can be assumed that the selection of the individual aminoacyl-tRNAfor the A site is mostly attributed to the specificity of the appropriate aminoacyl-tRNA synthetase together with the specific codon-anticodon interactions, whereas the 2 3 S rRNA participates in catalyzing peptide bond formation but without side-chain specificity for the amino acid esterified to the tRNA’s 3’-terminalnucleoside. In comparison with the prerequisites for specificity of peptidases the peptidyltransferase seems to be an old unspecific ribozyme in accordance with its function in evolution as precursor to extant life. It is of interest to note that in a recent paper a possible mechanism for peptide bond formation on ribosome without the mediation of peptidyltransferase has been These authors assume, by analysis of the energetics using a semi-

72.5 Hydrolysis and formation ofpeptides

I

823

empirical method for the formation of a cyclic intermediate, that the peptide bond formation through the tetrahedral intermediate in an S-configurationmay not need assistance from an enzyme or ribozyme. From the tetrahedral intermediate a cyclic intermediate will be formed, where the 2’-OHof the ribose sugar of the P-site tRNA is a member of the ring, which produces a free tRNA and a tRNA attached to a planar peptide unit. Since the free 2’-OH group of the peptidyl-tRNA was proposed to be involved in peptide bond formation, it has been argued that the appropriate tRNA may be acting as a biocatalyst (enzyme or ribozyme). Even in the case it should be possible to separate ribozyme activity from the ribosome or to isolate an in vitro selected ribozyme that can catalyze the same type of peptide bond formation as a ribosome, however such a biocatalyst seem does not to be suitable for simple practical use rather than using a chemical coupling reagent. In principle, this conclusion is also valid for the nonribosomal poly- or multienzymes which are involved in the biosynthesis of peptide antibiotics[721. Up to now, they have only found application in the synthesis field of cyclosporin, gramicidin S, special plactam antibiotics and analogs. At the end of this short assessment only those enzymes that usually act as hydrolases catalyzing the cleavage of peptide bonds remain to be discussed. The fundamental suitability of peptidases for catalyzing the formation of peptide bonds is based on the principle of microscopic reversibility that was predicted by van? Hoff On the last page of his contribution he had proposed the basic idea of in 1898[731. peptidase-catalyzedformation of the peptide bond, as follows: “Die Frage ist berechtigt, ob . ... auch nicht das Trypsin imstande ist, unter Umstanden, durch die Gleichgewichtslage gegeben, Eiweiss zu bilden aus den Spaltprodukten, die es selber bildet”. The concept of van’t Hoff of the equilibrium constant of a reversible chemical reaction, along with the function of a catalyst (including biocatalysts) for accelerated achievment of the equilibrium according to O ~ t w a l d [ ~is~ Ithe , theoretical background of enzyme-catalyzed peptide synthesis. However, about 40 years elapsed before the first experimental proof of van’t Hoff‘s prediction became evident through the first clear-cut peptidase-catalyzed synthesis of an amide bond carried out by Bergmann and Fraenkel-Conrat[”].Before this approach gained any industrial importance another 40 years had elapsed, and in recent decades considerable efforts have been made to find the optimum conditions for peptidase-catalyzed peptide synthesis as can be seen in various reviews [7G-981. 12.5.3.3 Principles of Enzymatic Synthesis

As shown in Fig. 12.5-8the equilibrium of a peptidase-catalyzedreaction is normally shifted to the thermodynamically more stable cleavage products. In contrast to proteolysis, the peptide bond formation is a two-substrate reaction and requires not only a specificity-dependentinsertion of the carboxyl component into the S-subsites of the active site, but also an optimal binding of the amine component in the S’ region. To shift the equilibrium in favor of fragment product formation various

824

I

12 Hydrolysis and Formation ofC-N Bonds

Responsiblefor cleavage specificity [Primary specificity]

R'

R2

R3

I

I

?

R1'

I

R2'

I

73'

I

-NH-CH-CO-NH-CH-CO-NH-CH-CO-NH-CH-CO-NH-CH-CO-NH-CH-COp3

p2

T3

R2

p3

p2

I

t R1

-NH-CH-CO-NH-CH-CO-NH-CH-C<

t

+ O

Responsiblefor the primary specificity in synthesis Figure 12.5-8.

P'l Protease R"

oe

I

PI

1

( 8 1

P'2

P'3

R2'

R3'

I

I

P'2

P'3

H3N-C H-CO-N H-C H-CO-N H-C H-COP'l

t

Important for the nucleophile efficiency in kinetically controlled peptide bond formation

Peptidases function in vivo as hydrolases rather than as ligases.

manipulations are necessary which also differ mechanistically. The approaches to peptidase-catalyzedpeptide bond formation are generally classified into basic strategies (see below) according to the type of carboxyl component used. In the equilibrium-controlled approach the carboxyl component bears a free carboxyl group as shown in Fig. 12.5-9, p. 826, whereas in the kinetically controlled approach the carboxyl component is employed in a slightly activated form, mainly as an alkyl ester. Both strategies are fundamentally different due to the energy required for the conversion of the starting components into the peptide products. Before interpreting the two mechanisms in more detail some general considerations of reversing proteolysis must be discussed.

12.5.3.3.1

General Manipulations in Favoring Synthesis

Looking at the equilibrium for the reversal of proteolysis, under normal conditions the equilibrium is shifted towards the hydrolysis products. For example, a synthesis of a dipeptide from its constituent free amino acids is, from the energetic point of view a very unfavorable process because of considerable increase in the free enthalpy involved. Under these circumstances it is not possible to accomplish peptide bond formation by simple reversal of hydrolysis, even using high concentrations of the starting amino acid zwitterions. Energetically more favorable is the reaction of an anion and a cation using an P-protected amino acid as a carboxyl component and a P-blocked amino acid as an amine component, respectively. According to the underlying thermodynamic principles, the outcome of peptide synthesis in aqueous solution depends on (a) the value of the equilibrium constant, (b) the ionization constants of the selectively protected starting compounds and (c) the initial concentrations of the ionized and nonionized forms of the carboxyl and amine component.

72.5 Hydrolysis and Formation ofpeptides

The thermodynamic parameters only allow statements relating to the free enthalpy change between the start and the end of the reaction, e. g. the equilibrium of the reaction. Only the velocity with which the equilibrium is reached depends on the catalytic action of the enzyme used. According to the law of mass action the product yield is proportional to the starting component concentration. Using the least expensive starting component in excess, manipulations described in the following make it possible to transform the other starting component almost quantitatively into product. The formation of insoluble products is a useful way of shifting the equilibrium towards synthesis.The reaction medium must be designed so that both starting components on the left-hand side of the equation are soluble in the medium while the peptide product on the right-hand side is insoluble. Under these conditions the product is continuously removed from the reaction medium by precipitation and sometimes an almost quantitative product yield can be obtained. A second way of reversing the proteolysis reaction can be performed by product extraction, a concept quite close to the solubility-controlled process of precipitation. The reaction is carried out in a biphasic system where the product is much more soluble in the organic phase and is continuously removed from the aqueous phase where the starting components and the enzyme are soluble. In both approaches to product removal the benefits of the appropriate organic solvent must be taken into consideration, which will be discussed later. Last but not least, in special cases the formed product can be separated from the equilibrium by molecular traps, where the desired product will removed by specific complex formation, as demonstrated, for example, in the course of clostripain-catalyzed fragment condensation of the ribonuclease (RNase)fragments 1-10 with 11-15 using RNase S(21-124) as a trap["].

12.5.3.3.2

Equilibrium-controlledSynthesis

This equilibrium-controlledor thermodynamic approach (see below) represents the direct reversal of proteolysis. Consequently, all peptidases, independent of their mechanisms, can be used. Apart from this advantage the necessary high enzyme requirement and the low reaction velocity are drawbacks of this approach. Preceding the conversion, determined by Ken, is an ionization equilibrium Kion: +

KLmm

RCOO- + H3NR' + RCOOH + HzNR'

IL."

RCO- NHR' + HzO

(1)

If the water concentration is taken into the equilibrium constant Eq. (2) is obtained: K s p = Kion . &on

+ = [RCO-NHR']([RCOO-][H3NR'])-'

(2)

The reaction medium, especially the pH, determines the constants for a given pair of reactants. To obtain an equilibrium that is shifted in favor of peptide product formation the ionization equilibrium must be manipulated. One efficient method is the addition of water-miscible organic solvents to the aqueous reaction mixture thereby lowering the dielectric constant of the medium, reducing the acidity of the carboxyl group, and to a lesser extent the basicity of the amino group of the nucleophilic amine component [loo,l o l l . The use of biphasic systems (for a review

I

825

826

I

12 Hydrolysis and Formation ofC-N Bonds

see reference[102]),e.g. solvent systems consisting of an aqueous phase and a nonmiscible phase (nonpolar organic solvents)does not damage the enzyme since it is localized in the aqueous phase. Under ideal conditions the reactants diffuse from the organic phase into the aqueous phase and after peptide bond forming step the product diffuses back into the organic phase. Only the insufficient solubility of the reactants in nonpolar organic solvents limits the general application of the biphasic approach, particularly for the condensation of longer segments. For the direct reversal of catalybc hydrolysis of peptides, discussed in this chapter, the term equilibrium-controlled approach should be preferred. Because of the thermodynamic control of both equilibria in Eq. (1)the reversal of proteolysis is often denoted as a thermodynamic approach. In order to increase the product yield of this endergonic process various manipulations are required. In addition to those mentioned above, reverse micelles I1O3], anhydrous media containing minimal water concentrations [lo41,water mimics [*''I, and reaction conditions promoting product precipitation as discussed in first part of this chapter are often employed.

Kinetically Controlled Synthesis

12.5.3.3.3

In contrast to the equilibrium-controlled approach the peptidase-catalyzed kinetneeds much less ically controlled peptide synthesis (for a review see enzyme, the reaction time to reach maximal product yield is significantly shorter, and the product yield depends both on the properties of the enzyme used and the substrate specificity. Kinetic control means that the product appearing with the highest rate and disappearing with the lowest velocity would accumulate. Whereas the equilibrium-controlled approach ends with a true equilibrium, in the kinetic approach the concentration of the product formed goes through a maximum before the slower hydrolysis of the product becomes important. The product will be hydrolyzed if the reaction is not stopped after the acyl donor ester is consumed and true equilibrium is allowed to be reached. In Fig. 12.5-9 both approaches are compared schematically. The kinetic approach (b)requires the use of an acyl donor ester as a carboxyl component and is limited to a

R-COOH + H2N-R'

41

R-CO-NH-R'

t -

+ H,O

b

R-CO-X

+ H,N-R'

1

R-CO-NH-R'

+ HX

t -

Comparison of the equilibrium (a) and the kinetically controlled approach (b) of peptidase-catalyzed peptide synthesis.

Figure 12.5-9.

12.5 Hydrolysis and Formation of Peptides

I

827

Ac-E..HN

EH + Ac-N

Kinetics of peptidase-catalyzed acyltransfer reaction. EH =enzyme, Ac-X = acyl donor ester (carboxyl component), H N = nucleophile (amine component), H X = leaving group o f t h e acyl donor ester, Ac-OH = hydrolysis product, Ac-N = peptide product; E..Ac-X = enzyme-substrate complex (Michaelis complex), Ac-N..HN = acylenzyme-nucleophile complex. Figure 12.5-10.

peptidases which rapidly form an acylenzyme intermediate, e. g. serine and cysteine peptidases. The peptidase acts as a transferase catalyzing the transfer of the acyl moiety to the amino acid- or peptide-derived amine component. Specifically, the acylenzyme reacts, in competition with water, with the nucleophilic amine component to form the peptide bond. The ratio of formation of aminolysis and hydrolysis products is of decisive importance for successful preparative peptide synthesis. Figure 12.5-10 describes the kinetics of the peptidase-catalyzedacyl transfer reaction. First, the acylenzyme is formed via the Michaelis-Menten complex, which binds the amine component to the acylenzyme. The resulting acylenzyme-nucleophile complex can undergo aminolysis as well as hydrolysis. The acyl transfer efficiency of the peptidase for the corresponding substrates is determined by the ratio of the aminolysis and hydrolysis product formed, which is also denoted as selectivity.

12.5.3.3.4

Prediction of Synthesis by S’ Subsite Mapping.

Serine and cysteine peptidases are not perfect acyltransferases.Therefore, it is useful to have a method for the prediction of the outcome of the kinetically controlled peptide synthesis. In order to get a simple efficiency parameter we decided to introduce the partition value p1106]analogous with the definition of the Michaelis constant according to Eq. (3), where Pz = Ac-OH, P3 = Ac-N, and N = HN.

The p value corresponds to the nucleophile concentration at which hydrolysis and aminolysis of the acylenzyme proceeds with the same velocity. The advantage of p is that the definition is not based on a particular kinetic scheme. Furthermore, p allows a rapid estimation of the yield of any acyl transfer reaction. A concentration of the >> p is necessary for peptide formation in high nucleophilic amine component yield. Assuming an equilibrium between the acylenzyme and the acylenzyme-

[w

828

I

12 Hydrolysis and Formation ofC-A! Bonds

nucleophile complex, Eq. (4) and (5) can be derived from Fig. 12.5-10 for the velocities of hydrolysis and aminolysis of the acylenzyme, where E = EH, EA = Ac-E, A = Ac-X, and EAN = Ac-E.. HN. V H = [EA]k3

VA =

k + [EAN] KN

k4 [EAN] KN

Eq. (6) results from combination Eq. (4) and (5).

It follows from Eq. (6)that a linear correlation between the partition value p and the nucleophile concentration is obtained. The quotient ks/k4 corresponds to the ratio of hydrolysis and aminolysis of the EAN complex whereas the term kNk3/k4 is a measure of the nucleophile efficiency. ~ ' ~the . The partition value p can be determined by differnt m e t h ~ d s [ ' ~ ~ - 'In presence of a large excess of nucleophile ([W >> [Ale) the decrease in the nucleophile concentration during the reaction course can be ignored. Under these conditions vH/ V A = [P2]/[P3]. The determination ofp can be established out from the product ratio obtained by HPLC analysis according to Eq. (7).

In the preparative application of acyl transfer reactions, however, a large excess of the nucleophile is not useful because a complete turnover of both reactants is desired. For this reason, we developed the determination of p from the integrated rate eq~ation[~~'1 according to Eq. (8).

A plot of [Pz]/[P3]versus ln([Nlo/([Nlo- [P3]))/[P3] gives a straight line with the slope KN k3/k4 and an intercept with the y axis at kslk4. Since this method permits the determination of p under the conditions employed in preparative peptide synthesis it should be useful for the optimization of the reaction conditions. An understanding of the molecular interactions between the acylenzyme and the attacking nucleophilic amine component allows an optimization of the acyl transfer efficiency. The efficiency of the nucleophilic attack of the amine component depends essentially on an optimal binding within the active site by S ' - P' interactions (Fig. 12.5-11). Consequently, more information on the specificity ofthe S ' subsites of serine and cysteine peptidases are useful, which can be obtained by systematic acyl transfer studies using libraries of nucleophilic amine components. According to the definition of the p value (see above) small values of p indicate high S' subsite specificity for the appropriate amine component in peptidase-catalyzed acyl transfer reactions.

72.5 Hydrolysis and Formation ofpeptides

I

829

Figure 12.5-11. Schematic representation o f subsite-substrate interactions in the course o f t h e acyl transfer from the acylenzyme t o the nucleophilic amine component catalyzed by a serine peptidase.

We have studied a couple of different serine peptidases (for a review see and clostripain['", 'I2], respecreference ["I), the cysteine peptidases papain [I' and tively, and the prolyl endopeptidase from Hauobacterium meningosept~m[~'~I, have determined p values for various series of nucleophilic amine components. Apart from clostripain none of the enzymes under investigation catalyzed acyl transfer to nucleophilic amine components with P'1 = Pro or D-amino acids. The efficiency of chymotrypsin-catalyzedacyltransfer decreases in the order of positively charged > aliphatic > aromatic > nagatively charged PI1 side chains. The specificity of chymotrypsin for P'1= Arg and Lys is attributed to electrostatic interactions between these side chain moieties and Asp3' and Asp36in the active site. A statistical analysis of proteolysis data confirmed that chymotrypsin possesses a specificity for peptide bonds bearing Arg or Lys at the Po1position, whereas Leu-Asp bonds of proteins were cleaved by this enzyme considerably less frequently than one expects from the frequency of occurence of this peptide bond['l4]. Our results confirm this statistical evaluation exactly. Furthermore, remarkably chymotrypsin prefers arginine residues at the P'1 and PI3 positions, which offers an interesting option for using chymotrypsin in the sense of a restriction peptidase for peptide-catalyzed processing of recombinant proteins (cf. Fig. 12.5-27 ). The selectivity of the S' subsites of different peptidases is reflected by the broad range of data obtained as shown for simple amino acid amides in Table 12.5-3. The values demonstrate the preference of basic and hydrophobic P'' residues for chymotrypsin and also for papain. In the case of chymotrypsinthe strongly basic side chain of arginine amide gives rise to a higher efficiency than all other nucleophiles. Despite the difficulties in catalyzing Xaa-Pro bonds, we have studied the clostripaincatalyzed acyltransfer using a large number of proline-containing peptides as well as

830

I

12 Hydrolysis and Formation ofC-N Bonds Table 12.5-3. Comparison ofp values of selected amino acid amides H-Xaa-NH2 i n acyltransfer reactions catalyzed by various serine and cysteine peptidases according t o Schellenberger and Jakub ke Ig51. ~

~~

P

Enzyme Xaa

EndoproteinaseGlu-C V8 Endoproteinase Glu/Asp-C Chymotrypsin Tiypsin Elastase Papain

Arg

Leu

Val

Met

> 500

16 132 4.2 72 62 0.41

117 n. d. 6.7 130 69 3.9

64 382 3.3 12 34 1.5

30 0.11 66 16 1.3

Ala-Xaa dipeptides and amino acid amidesL"" 'I2] . The efficiency of clostripaincatalyzed acyltransfer, using Bz-ArgOEt as the acyl donor to amino acid amides decreases in the order Leu > Lys > Gly > Arg > Gln > Ser > Pro > Thr > Ala > Asn > Asp > Glu. S' subsite mapping using an Ala-Xaa library led to the result that clostripain prefers PIz residues with positively-charged side chains, followed by proline, whereas negatively-charged side chains of Asp and Glu are weak nucleophilic acceptors. In the pentapeptide series, containing only one proline residue, the efficiency decreases in the order Pro-Pt3> Pro-P'2 > Pro-Ptl.Surprisingly, PAPAG, PPAAG and PF-NH2 act as very weak nucleophilic acceptors. The variety of different conformations of proline-containing peptides should be the reason for the extreme differences in enzyme-nucleophile interactions.

12.5.3.3.5

What Approach Should be Preferred?

As mentioned above, the equilibrium-controlledapproach has the advantage that all peptidases can be used. However, the high enzyme requirement and the low reaction velocity are serious drawbacks. Owing to the endergonic process the reaction conditions must be manipulated in order to increase the product yield. The addition of high concentrations of water-miscibleorganic solvents to decrease the pK value of the carboxyl component very often decreases the catalytic activity of the peptidases. Furthermore, by carrying out equilibrium-controlled synthesis in aqueous media using reactants with unprotected side chain functions, the specificity-determining amino acid residue should again not occur in the segments to be coupled. In the kinetic approach, the serine or cysteine peptidase rapidly reacts with a suitable acyl donor ester to form the acylenzyme intermediate, which can be deacylated competitivelyby the added nucleophilic amine component and water. The ratio between aminolysis and hydrolysis of the acyl donor ester is of great importance for the outcome of the synthesis route.This selectivity is essentially determined by the S' subsite specificity of the enzyme as shown above. To establish an optimum synthesis strategy, it is useful to know the basic kinetic parameters for the reaction course, in particular those obtained by S' subsite mapping are of great importance for planning and optimization of the enzymatic synthesis. Depending on the specificity of the peptidase used, pH and solvent conditions, the

12.5 Hydrolysis and Formation ofpeptides

peptide product formed in the kinetic approach is quite stable since the amidase activity of most enzymes is lower than the esterase activity. In addition, the esterase activity can be positively manipulated by varying the type of leaving group, as shown later. For preparative peptide synthesis such a manipulation is very important as it allows complete conversion of the acyl donor ester before the product is hydrolyzed. There is no doubt that the course of kinetically controlled protease-catalyzedpeptide synthesis can be influenced more efficiently than the equilibrium approach. Although the kinetic approach should be preferable, the decision must depend on the overriding total synthesis concept. The largest industrial scale application of the equilibrium approach is probably the enzymatic synthesis of Z-Asp-Phe-OMe,the precursor of the peptide sweetener The best known use of transpeptidation technology is the large scale conversion of porcine insulin into human insulin by trypsin['lG1or Achrornobacter lyticus protease [ll'l. 12.5.3.4

Manipulations to Suppress Competitive Reactions

The most important factors which limit the widespread routine application of peptidases in kinetically controlled peptide synthesis are undesired hydrolysis of the acyl donor ester and proteolysis of both the starting segments to be coupled and the final peptide product, respectively (Fig. 12.5-12). An elimination or minimization of these undesired reactions can be performed by various manipulations concerning the reaction medium, the enzyme and the substrate as well as on mechanistic features of the process. In particular, an efficient leaving group of the acyl donor ester can provide high reaction rates in combination with a decreasing danger of possible proteolysis of the starting segments and the final product.

12.5.3.4.1

Medium Engineering With Organic Solvents

In peptidase-catalyzed peptide synthesis the solubility of the starting components dramatically influences the course of the synthesis. From the ideal medium, water, the spectrum of solvents ranges from water-miscibleorganic solvents and aqueousorganic biphasic systems to monophasic organic solvents with trace amounts of Avoidance of hydrolysis Y-NH-C-OH Cleavable

Hydrolysis

H,Of

J :

Y-NH*C-(S,O)R

T

Leaving group

Enzyme

?

$Y-NH-ll-C-Enzyme H(S,O)R

NH,UR'

\?

---)-Y - N H I I ) C - N H D R '

NH,UR'

(Thio-)acyl enzyme Arninolysis Peptide product

Figure 12.5-12. General course of the kinetic approach to fragment condensation catalyzed by serine or cysteine peptidases.

I

831

832

I

12 Hydrolysis and Formation ofC-N Bonds

Reaction medium

Advantages

Drawbacks

Alternatives

ideal medium for enzymes

poor solubility for partially protected reactants kinetic approach only promotion of hydrolysis

use of solubilizing protecting groups

increased reactant solubility

reduzed enzyme activity

use of chemically or genetically modified enzymes

promoting equilibrium-controlled approach

difficult product isolation

Water

optimal ecological conditions

WaterNaterMiscible Organic Solvents

WatermaterNonmiscible Organic Solvents

Monophasic Organic Solvents

[Biphasic Systems] prevention of enzyme activity

higher enzyme reauirement

easy product isolation

limitation of reactant solubility lowering of velocity reduced enzyme activity

prevention of hydrolysis no solubility problems of partially protected reactants adjusting media between chemical and enzymatic strategies

use of chemically or genetically modified enzymes

use of chemically or genetically modified enzymes

change of stereoand regiospecificity higher enzyme requirement

-

Figure 12.5-13.

Influence of the reaction medium on peptidase-catalyzedpeptide synthesis

water necessary for the catalytic activity of the enzyme (Fig. 12.5-13). Not only for ecological reasons, but water should be the preferred reaction medium for enzymatic processes, since it is i n vivo the medium of choice for enzymes anyway.

Solubilizing Protecting Groups These are the only alternative way of bypassing the poor solubility of most amino acid-derived starting components, and synthesis of peptides can only be performed if one or both reactants bears such a solubility-promoting group. A successful synthesis of kyotorphin (Tyr-Arg)in a continuous large scale procedure using highly solubilizing P-protecting groups was carried out by Fischer et a1.[1181.They used maleyl (Mal-, 3-carboxyacryloyl-),a group which increases both the solubility of the tyrosine ethyl ester as well as the activity of chymotrypsin. This procedure was performed with concentrations of Mal-Tyr-OEt of up to 1.5 mol L-' and an equimolar

72.5 Hydrolysis and Formation ofpeptides

concentration of H-Arg-OEt.A 72.7 mol procedure resulted in 12 kg of the diacetate of Tyr-Arg which corresponds to an overall yield of 50.4% including protecting group removal, purification by ion exchange chromatography, and final product isolation by spray drying. Further large scale procedures using solubilizing protecting groups were carried out by Florsheiner et al. and Hermann et al. [120]. It was also reported that carboxypeptidasesare capable of coupling N-terminally unprotected amino acid esters (50 mM) to unprotected amino acids as well as amino acid derivatives (0.2-1.5 M) in one step at room temperature in aqueous solution[12'1.This synthesis principle is more generally applicable to other esterolpc endopeptidases or lipases [122-1241. The reduced stereoselectivityallows synthesis of D,L-dipeptides in higher yields than the corresponding L,L-dipeptides [12'1. The chymotrypsin-catalyzed coupling of HPhe-OMe with nucleophilic amine components in a frozen aqueous starting from lower acyl donor/nucleophile ratios should be mentioned as an interesting alternative, and enzyme-catalyzed synthesis in frozen-aqueous systems will discussed later in more detail (see Sect. 12.5.3.4.2). Water/Water-Miscible Organic Solvent Systems Such systems promote the solubility of partially protected starting compounds and increase the pKvalue of the carboy1 component in equilibrium-controlledprocesses thereby promoting this synthesis course. However, reduced enzyme activity in the presence of high portions of organic solvents and difficulties in product isolation are sometimes serious drawbacks. Despite these limitations such media with a small organic solvent content are preferred in enzymochemical peptide synthesis. The application of more stable immobilized enzymes as well as chemically or genetically engineered enzymes offers advantages in cases of high contents of organic solvents, as will discussed below. Biphasic Aqueous/Organic 12*1 These have been developed as an alternative to water/water-miscible organic solvents systems. This approach leads to preservation of enzyme activity and allows simple product separation, an advantage which is counteracted by prolonged reaction times where additional partition equilibria are most likely to be the ratedeterming steps. The general use of biphasic systems is mostly limited by the solubility of the starting components in the nonpolar organic phase. This alternative to the use of water-miscibleorganic solvents has been used with various peptidases and good yields were obtained using no more than two equivalents of the nucleophilic amine component (for a review see Synthesis in Reversed Micelle~('~'~ l3O] This is principially very similar to the approach discussed above. After adding small amounts of water and a surfactant to a hydrocarbon, the polar ends of the surfactant form a sphere which contains the water. Since the lipophilic group of the surfactant is facing outside into the surrounding hydrocarbon, the reverse structure of a normal micelle is formed. Liposome-assisted selective polycondensation of amino acid and peptides shows an interesting continuation along this line [l3'1.

I

833

834

I

72 Hydrolysis and Formation of C-N Bonds

Monophasic Organic Solvents['32] The ultimate way of preventing undesired hydrolpc side reactions in the course of peptide synthesis is offered by these solvents. Trace amounts of water between approximately 0.3 to about 1% are necessary to maintain the catalytic activity of the enzyme. Although it has been generally assumed that higher concentrations of water-miscible organic solvents significantly reduce the catalytic activity of the peptidases, few papers have demonstrated successful enzymatic peptide synthesis performed in some hydrophilic organic solvents, such as aliphatic alcohols and . Generally, enzyme specificities change dramatically in organic a~etonitrile['~~-'~~1 solvents. Higher enzyme requirement and reduced rates should be noted. It is interesting to mention that peptidases also catalyze esterification and transesterification reactions in organic solvents when the appropriate alcohol is added. Chemically or Genetically Modijied Peptidases They provide a useful alternative for peptide synthesis in high concentrations of organic solvents since they are more stable than the native enzymes. Various possibilities for modification are known. Immobilized Enzymes Such enzymes can be used in a very simple way for enzymatic peptide synthesis as first reported by Jakubke and coworkers['*', 137-1391 at the beginning of the 1980s. The effort involved in immobilizing an enzyme is mostly compensated for by the possibility of its repeated use. Immobilized biocatalysts have almost the same efficiency as the native enzymes. The peptidase is covalently linked or adsorbed to an insoluble gel or resin. The water content in these systems plays an important role in modulating the catalytic properties of the immobilized peptidase. The presence of water molecules on the enzyme is required in order to retain the catalytic activity. The measurement and control of the thermodynamic water activity is necessary to quantify the water effect on enzyme activity and the intrinsic influence of other variables such as support, solvent and e d u ~ t s [ 1411. ' ~ ~The ~ advantage of these systems have been demonstrated in the synthesis of various biologically active 1421 .The effect of water-miscible aprotic solvents on kyotorphin synthepeptides sis catalyzed by immobilized chymotrypsin was studied by Lozano et a1.['43] Of special technical interest are the continuous synthesis of the aspartame precursor ZAsp-Phe-OMewith thermolysin immobilized on amberlite XAD-7 in a plug flow and the conversion of porcine insulin into human insulin catalyzed type reactor by Achromobacter lyticus protease I immobilized on SiOz-polyglutamicacid['45]. Solvent-Modijkd Enzymes These are named as enzymes which are modified, for example, with polyethylene glycol (PEG) allowing synthesis in monophasic organic solvents as described, e. g. for chymotrypsin[146r 14'1, papain [1481 and thermolysin [14')1. Using PEG-modified enzymes in monophasic organic solvents undesired proteolytic reactions can be almost completely eliminated. However, owing to the solubility properties the use of hydrophobic organic solvents makes the application for the synthesis of longer

72.5 Hydrolysis and Formation ofpeptides

I

835

peptides very complicated and often impossible. Insoluble cross-linked chymotrypsin can be obtained using glutaraldehyde concentrations several times higher in contrast to the procedure for soluble polymeric preparations of chymotrypsin[”’I. Insoluble cross-linked chymotrypsin was used in a medium with GO% (v/v) dimethylformamide (DMF) for successful synthesis of short peptides. High amounts of powdered suspensions of peptidases in DMF have been used for peptide synthesis [1521. An very interesting synthesis approach has been described using crosslinked enzyme crystals (CLECs)[153* 1541.

Chemically Modijed Enzymes Enzymes are often prepared with the aim of reducing the peptidase activity with some of the esterase activity remaining, thus preventing the hydrolyhc cleavage of peptide bonds [*‘I. Methyl-chymotrypsin(MeCT) obtained by N-methylation of His57 shows a significant change in the enzymatic catalysis. MeCT is less active than native chymotrypsin by a factor lo4 to lo5 but it is virtually without any peptidase activity[155].Owing to the low activity more activated cyanomethyl ester is used instead of methyl ester. Subtilisin can also be changed to an acyltransferase via modification of the active site serine to cysteine (thiol subtilisin with low amidase activity[’5G1) or seleno subti1isin[’’’1. Successful synthesis of various L,D-dipeptides using [Met(0)”2]chymotrypsin[’58~ were carried out as well as the synthesis of AcTyr-OEt from Ac-Tyr-OH and ethanol catalyzed by hexyl-chymotrypsinin a biphasic system[’’’]. Genetically Engineered Enzymes They have elevated solvent tolerance and also owing to the lowering amidase activity have been successfully used for synthetic purposes [lG01. Enzyme engineering describes a range of techniques from deliberate chemical modification as shown above to remodeling a wild-type enzyme by gene technology. The aim of engineering peptidases to generate peptide ligases by conversion of serine and cysteine peptidases via site-directed mutagenesis, is to make enzymes more stable and favor aminolysis rather than hydrolysis. Using multiple site-directed mutagenesis subtilisin can be converted into a mutant which allows kinetically controlled synthesis to be performed in the presence of high concentrations of DMF. An ingenious combination of chemical and enzymatic steps should promote the progress in peptide and protein synthesis as was demonstrated with subtiligase,a double mutant of subtilisin BNP’. This variant was prepared by protein design and used in a further total synthesis of Ribonuclease A (RNase A) [‘“I by combining solid-phase synthesis for fragment synthesis and enzymatic coupling of these fragment to form the protein (cf. 12.5.3.7.2, p. 856). The selection for improved subtiligases by phage display results in the identification of two new mutants that increased the activity of subtiligase[IG2].

836

I

12 Hydrolysis and Formation of C-N Bonds

Enzyme engineering

Enzyme engineering

1

T

Promoting equilibrium approach (reducted enzyme activity)

Media adjusting between chemical and enzymatic strategies Preventionof enzyme activity (limited application)

Organic-aqueoussolvent mixtures

Biphasic systems

f

f

(reduced enzyme activity and changed specificity) Micro-aquousmonophasis organic solvents

t

Addition of organic solvents

Advantages: * Ideal medium for enzymes optimal ecological conditions

Disadvantages: promotion of hydrolysis * bad reactant solubility

tI

-

*

Water [Solubilizing protecting groups]

Reducing of water concentration

I

Solvent-free micro-aqueoussystems (Diffusion-controlledsynthesis; using sonification and fluidization, resp.)

Freezingthe reaction mixture (Frozen-stateenzyme-catalyzed peptide synthesis)

Figure 12.5-14. Extended approaches to medium engineering in enzymatic peptide synthesis1961.

12.5.3.4.2 Medium Engineering by Reducing Water Content Competitive reactions in enzymatic peptide synthesis are, as mentioned above, mainly undesired hydrolysis of the acyl donor ester in the kinetic approach, and undesired proteolytic side reactions in both the starting components in fragment condensation as well as the final product. It can be demonstrated that side reactions of these types can be largely, but not completely, avoided by synthesis in organic solvents of controlled water activity. However, since the main drawbacks caused by organic solvents are enzyme deactivation and changes in specificity, which can only partly be improved by enzyme engineering, new strategies (Fig. 12.5-14)in reducing the water concentration without substitution by organic solvents have been described (for a review see reference["]).

Enzymatic Peptide Synthesis in Frozen Aqueous Systems This is based on observations by Grant and Alburn['"l that trypsin-catalyzed hydroxylaminolysis of amino acid esters was favored over hydrolysis in frozen reaction mixtures (for a review see Hansler and Jakubke('"1). In 1990 Schuster et

72.5 Hydrolysis and Formation of Peptides

I

837

al. [lG5] first reported on the influence of freezing on peptidase-catalyzed kinetically controlled peptide synthesis. The peptidase is added to the reactants precooled to 0 "C in a polypropylene tube and immediately inserted into liquid nitrogen. After 20 s the tube is transferred into a cryostat at - 15 "C or similar temperature. Amino components that are considered to be inefficient nucleophiles in enzymatic synthesis at room temperature gave substantially higher yields in frozen reaction mixtures. Later these results could be explained on the basis of the so-called freeze-concentration model[16G] and were confirmed by other investigators(1G71. In frozen aqueous systems the endopeptidase chymotrypsin is capable of acting as a reverse carboxypeptidase catalyzing coupling of free amino acids as amino components (168]. Various amino acids were acylated under catalysis of chymotrypsin starting from 2 mM Mal-Phe-OMeand 50 mM (50% as free base) of the appropriate amino acids at - 25 "C in unexpectedly high yields (% given in parentheses): Met (75), Val (58),Ser (52), Ile ( 3 5 ) ,Thr (30),Asn (29),Leu (2G), Lys (GO). Tougu et a1.[169] described similar results on coupling Mal-Tyr-OEt with free amino acids. The surprising catalytic behavior of chymotrypsin under frozen state conditions is demonstrated in Table 12.5-4. N"-unprotected amino acid esters as well as dipeptide esters, even containing unusual amino acids, can be coupled in frozen aqueous systems in high yields indicating both reverse aminopeptidase and dipeptidylpeptidase activities. Furthermore, cysteine proteases, with the exception of clostripain, were capable of catalyzing peptide synthesis in high yields using amine components with low efficiency at room temperature in frozen reaction mixtures. The specific properties of peptide synthesis in frozen solutions such as changes in specificity observed in serine and cysteine peptidase-catalyzed reactions strongly suggest that factors other than concentration of the reactants are probably involved in yieldenhancement by freezing. This assumption is supported by investigations reported by Jakubke et a1.[171]who determined the amount of unfrozen water in frozen samples at - 15 "C using the 'H-NMR-relaxation time technique and obtained an apparent concentration factor of 50. Synthesis experiments carried out under these concentration conditions at room temperature gave substantially lower yields com-

Comparative model peptide synthesis catalyzed by chyrnotrypsin in frozen aqueous systems and at room temperature.

Table 12.5-4.

Acyl donor

Amino component

Peptide Ice

Mal-Tyr-OMe Mal-Tyr-OMe Mal-Phe-OMe H-Tyr-OEt H-Phe-OMe H-4-fluoro-PheOMe H-2-naphtyl-Ala-OMe H-Leu-Phe-OMe H-Asp-Phe-OMe H-Gly-Phe-OMe

H-p-Ala-Gly-OH H-D-Leu-NHZ H-Lys-OH H-Lys-OH H-Leu-NH2 H-Leu-NH2 H-Leu-NH2 H-Ala-Ala-OH H-Ala-Ile-OH H-Ala-Ile-OH

79 73 60 71 94 90 93 88 91 85

Yield ("A) 25 "C

Reference

838

I

12 Hydrolysis and Formation ofC-N Bonds

pared with frozen reaction mixtures and, therefore, could not simulate the reaction conditions in ice. In addition to the freeze-concentration effect, a catalytic role for ice crystals, a favorable orientation of substrate and biocatalyst, the markedly lower dielectric constant of ice compared with water, and the high proton mobility in ice, have been discussed as further factors that possibly influence reactions in frozen systems. In summary, the reverse action of hydrolases provides an attractive alternative to the chemical synthesis of peptides but this approach could also be verified for the synthesis of oligosaccharides and oligonucleotides using glycosidases and ribonucleases,

Peptidase-catalyzed Synthesis in Solvent-jee Micro-aqueous Systems Such systems show a second route to reducing water concentration without substitution by organic solvents. This interesting development allows the application of reaction systems with partly undissolved reactants and is based on an extensive theoretical treatment of the equilibrium position described by Halling et al. [1721. The principle of a solid-to-solidconversion is illustrated graphically in Fig. 12.5-15 and selected examples of its experimental implementation are illustrated in Table 12.5-5. The application of solid phase substrate pools combines the equimolar (or nearly equimolar) supply of starting compoments with high obtainable yields, easy work-up procedures and compatibility with chemical standard procedures. The key parameter for obtaining high product yields via acyltransfer reactions is the ratio of aminolysis and hydrolysis favored by high nucleophile concentrations. In combination with solid phase acyl donor pools, this approach allows an equimolar supply of starting materials without any addition of organic solvents. The synthetic potential of systems with partly unsolved reactants was proven by pilot scale synthesis of Z-HisPhe-OMe and the low calorie sweetener precursor of Z-Aspartame in the thermodynamic approach [1731 and by kinetically controlled synthesis of enkephalin derivatives Furthermore, Halling and co-workers have studied the effect of water and A

B

n n before

substrates

A + n

____,

B

n

P

e

in equilibrium

substrates

m

product

General principle of application o f “equilibrium shift” towards the product by solid-phase substrate pools (botton) compared with synthesis starting from Figure 12.5-15.

72.5 Hydrolysis and Formation ofpeptides

Table 12.5-5.

Selected examples for peptide synthesis in water-based solid-liquid systems according to Eichhorn et aI."73]and Jakubke et al.[g'l. Carboxyl component

Amine component

Peptide yield (%)

Time (h)

Enzyme

2-Ala-OH 2-Asp-OH 2-Gln-OH 2-Phg-OH 2-Ser-OH 2-His-OH Z-Phe-OH Ac-Tyr-0E t Ac-Tyr-OEt 2-His-Phe-OBzl 2-Ser-OCam 2-Gly-His-ONb 2-Arg-His-ONb

H-Leu-NHz H-Phe-OMe H-Leu-NHZ H-Leu-NHz H-Leu-NH2 H-Leu-NH2 H-Met-NHz H-Arg-NH2 H-Gly-Gly-OH H-Arg-Trp-NHz H-His-ONb H-Lys-NHZ H-Gly-NH2

95 90

0.5

94

4

89 89 95 88 90

2 2.5 3

Thermolysin Thermolysin Thermolysin Thermolysin Thermolysin Thermolysin Thermolysin Chymotrypsin Chymotrypsin Chymotrypsin Papain Chymotrypsin Subtilisin

7

24

1 2 2.5 2.5 1.5

63

95 85 90 55

6

enzyme concentration of thermolysin-catalyzed solid-to-solid peptide synthesis in and reviewed the recently developed approach to enzymatic synthesis with mainly undissolved substrates at very high concentrations

12.5.3.4.3

Substrate Engineering

In the case where undesired subsequent reactions occur during kinetically controlled synthesis it is of minor importance which bond is cleaved by the enzyme. These side reactions underline the issue that the specificity of the enzyme for the acyl donor ester does not lie sufficiently above its specificity for the peptide product. Since the sequence of the starting components cannot be changed, the only practical alternative to suppress such competitive reactions is to use a highly specific leaving group of the acyl donor ester. As a simple model peptide with a highly sensitive cleavage site for chymotrypsin Schellenberger et al. [1771 used the chromogenic chymotrypsin substrate Mal-Leu-Phe-pNA (Ma1 = maleyl), which is formed by chymotrypsin-catalyzed coupling of Mal-Leu-OY with H-Phe-pNA. Table 12.5-6 shows that the leaving group moiety Y is of major influence on the reaction course. When Mal-Leu-OMe is employed as the carboxyl component, the velocity of the product cleavage reaches the rate of its formation after a short time. By using the Table 12.5-6.

Influence of the specificityconstants of acyl donor esters on the yield ofthe chymotrypsin-catalyzedsynthesis of Mal-Leu-Phe-pNA"starting from Mal-Leu-OYwith varying leaving groups Y and H-Phe-pNA according to Schellenberger et al.[1771, Leavinggroup Y

Me(Methy1) Bzl(Benzy1) Nb~~~inobenrv~~

KM

kcat

kcat/KM

(mM)

(s-7

(M-l S-l)

Reaction time (min)

Yield ("A)

120 f 30 1.7 f 0.4 0.38 f 0.1

5.6 i 0.4 5.4 f 0.4 5.9 f 0.4

4.7x 10 3.21 ~0' 1.5x lo4

10

-3

5 5

65 80.5

a k,,,/K, of Mal-Leu-Phe-pNA1 . 2 10 ~ M-'s-'

(Mal; maleyl).

I

839

840

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72 Hydrolysis and Formation ofC-N Bonds

more specific acyl donor esters (higher specificity constants kcat/KM) a clear product accummulation is attained. With Mal-Leu-ONbor ester of similar or higher specificity, starting components containing highly protease-labile cleavage sites can be coupled even in a homogeneous phase in high yields. According to this finding Bongers et al. published a two-step enzymatic semisynthesis of the superpotent analog of human growth hormone releasing factor [deNHzTyr',D-Ala2, Alal'] GFR(1-29)-NH2 from the amine component [Ala1']GRF(4-29)-NH2 and the carboxyl component deNH2Tyr-D-Ala-Asp-OY(Y = Et or 4-NOzBz1, respectively) catalyzed by V8 protease and Glu/Asp-specific endopeptidase (GSE) from Bacillus lichenijomis, respectively. Using the 4-nitrobenzyl leaving group compared with the ethyl moiety results in a higher yield without the undesired proteolytic side reactions. The state-of-the-artof substrate engineering is without doubt the substrate mimetic-mediated C - N ligation strategy which allows irreversible peptide bond formation and will be presented separately (see Sect. 12.5.3.6). 12.5.3.5 Approaches to Irreversible Formation of Peptide Bond

Despite the development of various possibilities to suppress competitive reaction, as shown in the preceding section, an absolute avoidance of proteolytic cleavage of the peptide bond formed cannot be guaranteed. The only alternative seems to be the use of biocatalysts that do not have the catalytic potential to hydrolyze peptide bonds.

12.5.3.5.1

Use of Nonpeptidases

Nonpeptidases are supposed to possess favorable prerequisites for the formation of peptide bonds because undesired proteolytic cleavages in the starting components and the product can be ruled out. Enzymes involved in protein synthesis also possess potential, e. g. aminoacyltRNA-synthetase-aminoadenylate complex['7g~l, the arginyl-tRNA: protein arginyltransferase1'80], and nonribosomal poly- or multienzyme complexes [721 which require ATP or GTP to activate the carboxyl group of an amino acid, and seem to accept various amino acid nucleophiles for peptide bond formation. However, the application ofthese enzyme systems for generally practical peptide bond formation is rather limited. Furthermore, lipases ['", lx2I containing the catalytic triade typical for serine peptidases have been used for peptide synthesis as well as pig liver esterase[ls3. 1841. These enzymes accept both D- and L-amino acid derivatives as weak acyl donors or nucleophilic acceptors, but concerning the practical importance the situation is similar to the enzyme systems reported previously. Particularly promising from the theoretical point of view seems to be the developments in catalytic antibodies. It could be established that antibodies raised against suitable transition state analogs are capable of catalyzing the formation of peptide bonds['85, lS6l. At present the practical importance is rather low, but the development of more tailor-made catalytic antibodies for peptide bond formation could change the situation in the future.

12.5 Hydrolysis and Formation of Peptides

12.5.3.5.2 Use of Proteolytic Inactive Zymogens

In 1994, it was firstly established that zymogens, the catalytically inactive precursors of various peptidases, can be used as biocatalysts for practically irreversible peptide bond formation[91,96, 97, lS71. The capability of reacting slowly with site-specific reagents indicated that such reactions proceed via the formation of an acyl-zymogen intermediate [lS8.lS9l. Although, the second-order rate of ester hydrolysis is 106-107 times slower than by the appropriate active enzyme, the deacylation rates of zymogen and active enzyme do not differ significantly.Therefore, it can concluded that the conversion of a zymogen to an enzyme should not be the activation of an inert zymogen, but the potentiation of catalytic activity intrinsic to the zymogen. Based on this feature Jakubke’s group has used the zymogens of the well studied serine peptidases trypsin and chymotrypsin, respectively, in peptide synthesis experiments, and has surprisingly observed catalysis of peptide bond formation by the zymogens trypsinogen and chymotrypsinogen. In several cases S’ subsite mapping studies showed significant differences in the deacylation of the acyl enzymes compared with the corresponding acyl zymogens, based on acyl transfer to various peptide derivatives.Although the zymogens possess the same catalytic triad, which is necessary for the formation of the appropriate covalent acyl intermediate, the non-optimal formed substrate binding cleft prevents proteolysis. In particular, Glylg3is distorted and is not capable of forming a hydrogen bond to the carbonyl oxygen of the substrate which is necessary for the stabilization of the oxyanionh ~ l e ~ ”However, ~]. because of the high flexibility in this region, a principle oxyanion stabilization takes place, although not in an ideal manner. To confirm true zymogen catalysis it was essential to prove that the zymogen preparations were not contaminated with traces of the appropriate active enzyme. Based on the significantly different affinity of both enzyme and zymogen to the basic pancreatic trypsin inhibitor (BPTI) it was possible to analyze the esterase activity of the zymogen, which is an efficiency parameter used in estimating their peptide bond forming potential. Since the differences in L t / K cover ~ a range of about 5 orders of magnitude, for general use of zymogen catalysis it is essential to improve the acylation rate. The application of zymogens for irreversible fragment condensations was studied by coupling a synthetic tetrapeptide methyl ester with a recombinant 24-peptide according to the procedure of Cerovsky et al.L9’]. A comparison of the coupling reactions (Fig. 12.5-16)was carried by dropping the acyl donor ester 1into a solution of the amine component 2 and, alternatively,under batch conditions. The first way was chosen in order to minimize the undesired ester hydrolysis of 1and, in addition, to manipulate a large excess of the amine component 2. In this case 5.4 mg (0.002 mmol) of 2 was coupled with 4 mg (0.008 mmol) of 1 in the presence of 0.5 mg of chymotrypsinogen and resulted, after 400 min, in the complete conversion of the acyl donor ester in GO % yield to the desired product 3. The batch procedure led to a product yield of 52 %. In order to avoid any undesired zymogen activation by limited proteolysis, e. g. of the L y ~ * ~ - I peptide l e ’ ~ bond in the case of trypsinogen, it would be useful to prevent this reaction by chemical means. The guanylation of trypsinogen by l-guanyl3,s-dimethylpyrazolecauses a stable zymogen because of the conversion of all lysine

I

841

842

I

72 Hydrolysis and Formation ofC-N Bonds

Z-AGGF-OMe

+

1

1

H2N-G’ KLSQELHKL”QTYPRTDVGA2’ GTPA-OH chymotrypsinogen

2

Z-A’ GGFGKLSQE’’ LHKLQTYPRT” DVGAGTPA-OH

3 (4+24)-Fragrnent condensation catalyzed by chymotrypsinogen according t o Cerovsky et al. (cf. reference[98]). Figure 12.5-16.

residues into homoarginine (Har), including the crucial Lys”. Peptide synthesis with the guanylated zymogens led to very surprising results. Using dipeptides with a free carboxyl group as the amine components they are much more effectively accepted by the the guanylated species Lgll. From molecular modelling studies it can be concluded that there is an interaction between the carboxyl group of the dipeptide with the only lysine within the active site (LysG’). The conversion of LysG1 to homoarginine increases the pK of the side chain and therefore the basic character. 12.5.3.6

Irreversible C-N Ligations by Mimicking Enzyme Spe~ificity”~’]

The synthetic importance of peptidases as biocatalysts for peptide synthesis is undisputed due to a couple of advantages over pure chemical coupling methods. The mild reaction conditions and the high degree of regio- and stereospecificity guarantees both freedom from partial epimerization and that there is no need for temporary protection of side-chain functions. On the other hand, there are some serious drawbacks of the classical peptidase approach which has been discussed below in detail. Most important is the fact that the formed peptide bond formed can be cleaved in the course of the catalytic process by the same enzyme. There are no differences in the requirements of the specificity in both the peptide bond forming step and cleavage step, respectively. Since the specificity is manifested by the sidechain of the PI amino acid residue, e.g. Arg or Lys in the case of trypsin, an irreversible peptide bond formation seems not to be possible according to the classical concept of reversal of proteolysis. In ribosomal peptide bond formation the mechanism is based on an acyl transfer of the acyl moiety from the peptidyl-tRNA (or Met-tRNA at the start of the prokaryotic biosynthesis) located at the P site of the ribosome to the amino group of the aminoacyl-tRNA in the A site catalyzed by the side-chain unspecific ribozyme peptidyltransferase. Learning from nature our philosophy was that mimicking specificity is the only way to make the peptidase-catalyzed peptide bond forming step irre~ersibleI”~~ Ig3, lg71. Since from the mechanistic point of view the kinetic approach with serine and cysteine peptidases is also an acyl transfer process, the idea arose of to transfering the specificity moiety of the PI amino acid side chain to the leaving group of the acyl donor ester. In this manner the enzyme should recognize the acyl donor ester. However, after the acylation of the enzyme the leaving group

12.5 Hydrolysis and Formation of Peptides

with the specificity determinant is released from the enzyme with the consequence that the peptide bond formed cannot be cleaved by the enzyme due to the lack of specificity for recognizing this bond. In 1991 we were able to confirm this assumption by model peptide synthesis catalyzed by trypsin using various nonspecific W protected amino acid 4-guanidinophenyl esters (OGp) as acyl donors and various amino acid and peptide derivatives as nucleophilic acyl acceptors [192, 1931, and later extended by further examples from another 19’1. At that time this type of acyl donor ester was named an inverse substrate according to time-dependent irreversible inhibitors of trypsin and trypsin-likepeptidases, such as 4-amidino- and 4-guanidinophenyl esters which were found to be hydrolyzed by these peptidases 19’1. Although this fact was first virtually idependently of their acyl published in 1973 by Wagner and Horn[’96],very little was known about the basic mechanism of the hydrolysis of these inverse esters. In 1997 an extension of this new approach to irreversible peptide segment condensation with other peptidases was described and the term substrate mimeticswas introduced by Bordusa et al.

12.5.3.6.1

Mechanism o f Substrate Mimetic Hydrolysis

The most striking structural differences of W-protected amino acid or peptide 4-guanidinophenyl esters compared with common peptide substrates are the nonspecific acyl residue and the highly specific leaving group. It was established by Bordusa’s group [19’1 that all 4-guanidinophenyl esters, independently of structure and chirality of the acyl moiety, are hydrolyzed despite the lack of trypsin-specificacyl moieties, with the exception of the lysine derivatives (Table 12.5-7).This behavior is in contrast to common trypsin substrates. According to the familiar model, conventional trypsin substrates bind with their acyl residue to the S-binding site of the enzyme having the leaving group at the S’-subsite and the scissile bond between attacked by Ser”’. Table 12.5-7. Steady-state kinetic parameters for the hydrolysis of Boc-Xaa-OCp by trypsin” according to Thormann et a1.[’98].

r-Ala D-Ala GlY L-Leu

o-Leu L-Gln o-Gln

L-Phe o-Phe L - G ~ D-GIu L-LYS D-LYS

0.206 0.161 0.087 0.146 0.035 0.239 0.071 0.211 0.249 0.071 0.039 0.107 0.314

32.4 0.61 23.5 38.8 0.85 35.2 0.68 66.1 9.0 5.5 0.43 270 15.7

1.6 x 3.8 x 2.7 x 2.7 x 2.5 x

105 10’ 105

1.5 x 9.6 x 3.1 x 3.6 x 7.6 x 1.1 x 2.5 x

105

5.0 x

105

lo4

lo3 lo5

lo4 104

lo4 10‘

lo4

a Conditions: 25 m M Mops, pH 7.6,100 m M NaC1.5 m M CaC12,25 “C: errors less than 15%.

I

843

844

I

72 Hydrolysis and Formation ofC-N Bonds

cleavage site Protease S3

S2

: m ~

S1

%

$ S'q

S>

S;

Figure 12.5-17. Schematic comparison of the binding of a peptide 4-guanidinophenyl ester and a common trypsin substrate t o the active site o f the enzyme according t o the conventional binding model.

As shown schematically in Fig. 12.5-17 applying the same binding principles for the acyl moiety of the substrate mimetics leads to a catalytically unproductive binding. The acyl residue binds at the S-subsite of trypsin, but the scissile bond would be far away from the active site and, therefore, and cannot be attacked by Ser"5. However, docking calculations show that the specificity-bearing OGp group binds to the S1-bindingpocket like the side chain of L-arginineof commom peptide substrates. Surprisingly, this holds even for the substrates Boc-L-Arg-OGpand BocL-Lys-OGpdespite the presence of the S1 specific arginine and lysine residues, thus indicating a higher S1-specificityfor the 4-guanidinophenylmoiety. Indeed, all L- and D-substrate mimetics realize an arrangement in such a way that the scissile bond is very close to the hydroxyl group of the active Ser195.Furthermore, the carbonyl group of the scissile ester bond of the appropriate substrate mimetic is located at exactly the same position as the carbonyl group of the scissile peptide bond between P1-Lysl5 and PI'-Ala16in the trypsinogen-BPTI complex. This implies a possible attack by trypsin, which was confirmed by the hydrolysis studies. How does it work from the mechanistic point of view? Contrary to common trypsin substrates, the acyl residues of these enzyme-substrate mimetic arrangements bind to the s'-subsiteof trypsin (Fig. 12.5-18). For this reason, all binding sites beyond S1 are only of minor importance for the substrate mimetics. Furthermore, the acyl residues of the substrate mimetics do not reflect the specificity of the S-

Ac-X

- 0

Hz0 HX

W C O O H Ac-OH

Figure 12.5-18. Schematic representation o f the new extended kinetic model o f peptidasecatalyzed hydrolysis o f substrate mimetics according t o Thormann et EH, free enzyme; Ac-X, substrate (substrate mimetic); [E..Ac-XI, Michaelis-Menten complex; HX, leaving group; E-Ac, acyl enzyme intermediate located i n S'-region; Ac-E, acyl enzyme intermediate located i n S-region; KR, rearrangement equilibrium constant; Ac-OH, hydrolysis product.

12.5 Hydrolysis and Formation of Peptides

binding site of the enzyme. Since the direction of the peptide backbone chain is reversed, the S’-subsitespecificity is also not reflected. Therefore, substrate mimetics show a unique specificity behavior. The deacylation step, however, requires an unoccupied S’-subsitesince water can only attack the acyl enzyme from this site without hindrance. Hence, the flipping acyl moiety acts like a “sliding window” within the active-site, spanning the primed and unprimed subsite regions. The extended kinetic model requires a rearrangement step between the two arrangements (E-Ac and Ac-E) of the acyl enzyme described by the equilibrium constant KR (Fig. 12.5-18).From the experimental data of Table 12.5-7it follows that D-configured substrates exhibit lower k,,, values, which might be related to lower KR values. Exploring the dynamic behavior by molecular dynamics simulations of Boc-L-Ala-trypsinand Boc-D-Ala-trypsinindicated that the flip of the D-Ala complex to the S-subsite takes about 1.5 ns, much more than in the L-Ala complex (300 ps). For an experimental study of the S’-subsite accessibility, S’ mapping studies (cf. Sect. 12.5.3.3.4) are suitable. By their specific S-binding capacity, peptide nucleophiles should be capable of pushing aside the acyl moiety from the S’ region more efficiently than water. Therefore, the aminolysis of acyl enzymes bearing the acyl moiety in S’ should proceed at higher rates compared with their hydrolysis. Indeed, from the mapping studies it follows that the p-values for the deacylation of Bz-D-Alatrypsin are dramatically lower than for Bz-L-Ala-trypsin. Consequently, the experimental data of aminolysis also support this unique catalysis mechanism for substrate mimetics. 12.5.3.6.2 Cationic Substrate Mimetics

The Na-protected amino acid 4-guanidinophenyl ester was the first example of substrate mimetics for Arg-specific peptidases used for irreversible peptide bond formation[192,1931. Apart from the guanidino group linked at various aromatic and aliphatic spacers, also the amidino moiety is also suitable as a specificity-determing residue in the leaving group of cationic substrate mimetics 1192-1951 . After the basic studies with trypsin we could also establish that other Arg-specific peptidases such as thrombin and clostripain are suitable enzymes for peptide synthesis using cationic substrate mimetics [1971. Im particular, clostripain has been very useful in substrate mimetic-mediated fragment condensation. As shown in Fig. 12.5-19 the (3 + 5) fragment condensation provided a product yield of over 90% within a few minutes and the product formed remains unchanged after 72 h. The course of this synthesis clearly proves the irreversibility of this model C - N ligation. For synthesis planning, clostripain has an additional decisive advantage due to the extremely low PI’ specificity for the N-terminal amino acid residue of the amine component. Firstly, Bordusa and co-workers[19’1 demonstrated impressively the capability of the cysteine peptidase clostripain as a biocatalyst for the synthesis of peptide isosteres. These authors have investigated the function of clostripain for acylating aliphatic noncyclic and cyclic amines varying in chain length and ring size using the trypsin standard acyl donor ester Bz-Arg-OEt. Furthermore, using a model

I

845

846

I

72 Hydrolysis and Formation ofC-N Bonds

BOC-Phe-Gly-Gly-OGp 122 mg (0.174 mM)

1

+

H-Ala-Phe-Ala-Ala-Gly-OH 157 rng (0.286 mM) Tos x H,O

2

Clostripain

3 Boc-Phe-Gly-Gly-Ala-Phe-Ala-Ala-Gly-OH 177 mg (91% yield); 3 x 2 5 TFA x 2 H,O MALDI-TOF. m/z calc for [M+Na+]= 819.36, found. 819.29 Figure 12.5-19. Clostripain-catalyzed (3 +5) fragment condensation of Boc-Phe-Cly-ClyOCp and H-Ala-Phe-Ala-Ala-Cly-OH['"I. Conditions: 50 mM HEPES-buffer, pH 8, 100 mM NaCI, 10 mM CaC12, 25OC, [Clostripain]: 1.6 p ~ .

substrate mimetic, clostripain was capable to catalyze the reaction with noncoded and non-amino acid-derived amines. The results of these investigations indicate that the substrate mimetic approach may extend outside of peptide synthesis. In a recent paper Bordusa's group presented a novel enzymatic approach to the synthesis of carboxylic acid amides using substrate mimetics and clostripain as a biocatalyst [200]. This unexpected peptidase-mediated approach to the coupling of non-coded and non-amino-acid-derivedamines with pure organic esters could only be realized by the combination of the substrate mimetic strategy with the use of clostripain that possesses a broad tolerance towards amines. Selected examples of the clostripain-catalyzedcoupling of Bz- P-Ala-OGp and the 4-guanidinophenyl ester of 4-phenylbutyricacid (Pbu-OGp)with various amino acid amides and peptides are summarized in Table 12.5-8. Furthermore, the broad tolerance of clostripain toward non-coded amino acids and even simple amines, such as aliphatic, aromatic, or substituted amines including unnatural amino acids, and diamines as acyl acceptors is demonstrated by the results of appropriate syntheses compiled in Table 12.5-9. The substrate mimetic approach has opened a new range of synthesis applications beyond peptide synthesis offering efficient and selective organic amide bond formation under extraordinarily mild reaction conditions. Clostripain-catalyzed coupling of 4-guanidinophenyl esters o f 4-phenylbutyric acid (Pbu-OCp) and benzoyl-P-alanine (Bz-P-Ala-OCp),respectively, with various amino acid amides and peptides according to Cunther et al.[1991.

Table 12.5-8.

Acyl donor ester

Acyl acceptor

Product

Yield ("96)

Pbu-OGp Pbu-OGp Pbu-OGp Pbu-OGp Bz-P-Ala-OGp Bz-P-Ala-OGp Bz-P-Ala-OGp Bz-P-Ala-OGp

H-Leu-NH2 H-LYS-NH~ H-Ala-Pro-OH H-AFAAG-OH H-Leu-NH2 H-Lys-NHZ H-Ala-Pro-OH H-AFAAG-OH

Pbu-Leu-NHz Pbu-Lys-NHz Pbu-Ala-Pro-OH Pbu-AFAAG-OH Bz-0-Ala-Leu-NHz Bz-0-Ala-Lys-NHz Bz-P-Ala-Ala-Pro-OH Bz-P-Ala-AFAAG-OH

98 96 93 92 98 93 91 93

12.5 Hydrolysis and Formation ofpeptides

I

847

Clostripain-catalyzed coupling of non-amino acid-derived carboxyl and amine componentsa according t o Gunther and 6 0 r d u s a ' ~ ~ ~ . Table 12.5-9.

Product

Yield ("h)

Pbu-OGp

PbuNH-

81

Pbu-OGp

WUNH-J"

80

Acyl donor

Acyl acceptor

Pbu-OGp

H2N

P~UNH-

53

Pbu-OGp

H2N-OH

mu N H - O ~

65

Pbu-OGp

wuNH'-"OH

78

Pbu-OGp

P~UNH-"'OH

70

NHTH

Pbu-OGp

92

Pbu-OGp

2

95

Pbu NH

Pbu-OGp

wu-o-

Bz-OGp

BZ-NH-+J

82

Bz-OGp

BZ NH--"'

76

Bz-OGp

BZ NH-

56

Bz-OGp

BZ-N

Bz-OGp

Bz-NH-OH

84

BZNH-OH

70

Bz-OGp

H2N-OH

H-O

n. s.

57

NHTH 82

Bz-OGp Bz

Bz-OGp

2

94

Bz-NH

Bz-OGp

-

Bz-0-

n. s?

a Conditions: 0.2 M HEPES-buffer (pH 8.0),0.1 M NaCl, 0.01 M CaCL 5 % DMF, 25 "C, (acyl donor): 2 m M , (acyl acceptor): 12 m M ; b n. s., no synthesis.

12.5.3.6.3

Anionic Substrate Mimetics

Owing to the general validity of the concept of substrate mimetics Giinter and Bordusa[*'*]have expanded this strategy to anionic leaving groups in the appropriate mimetic structures based on the specificity determinants of Glu-specific endopeptidases. Since the leaving group moiety of a substrate mimetic binds in place of the specificity-determining amino acid side chain, for the strong Glu-preferred V8 protease from Staphylococcus aurens a carboxylate function linked with a suitable spacer was chosen as the ester leaving group. Unfortunately, the so far unknown 3D structure of this enzyme allows only the design of suitable mimetic structures by

848

I Z-Pro-Leu-Gly-SCm

12 Hydrolysis and Formation of C-N Bonds

1

+ H-Leu-Ala-Phe-Ala-Lys-Ala-AspAla-Phe-Gly-OH

2 mM

2

10 mM

i

V8 protease

Z-Pro-Leu-Gly-Leu-Ala-Phe-Ala-Lys-Ala-Asn-Ala-Phe-Gly-OH

3

Yield 55% (determined by analytical RP-HPLC) MALDI-TOF m/z calculated for [M+Na+] = 1433 71, found 1434 17 Figure 12.5-20. V8 protease-catalyzed (3 +lo) fragment condensation o f Z-Pro-Leu-Cly-LeuAla-Phe-Ala-Lys-Ala-Asp-Ala-Phe-Cly-OH[*"I. Conditions: 0.2 M HEPES-buffer, p H 8.0, 37 "C, [V8 protease] = 4 9 FM.

empirical structure-function relationship studies. Apart from other structures, the carboxymethyl thioester moiety in particular was selected as a potentially suitable leaving group for imitating the Glu residue in PI' position. The capability of the carboxymethyl thioester group to act as an artificial recognition site for the V8 protease was initially studied by steady-state hydrolysis kinetic studies. As a general result it can be summarized that the carboxymethyl thioester moiety was found to mediate specific hydrolysis of all carboxymethyl thioester substrate mimetics independently of the PI' amino acid residue. This also holds for Pro and even for D-Ala, which causes only a slight decrease in specificity compared with the L-enantiomer. Generally, a one to four orders of magnitude lower specificity compared with the 0 ~ s-l) was found. common substrate Z-Glu-SMe ( k c a t / K ~= 1 . 1 2 ~ 1 M-' In contrast to the non-specificity of the V8 protease for the acyl part, the negative charge of the leaving group is essential to mimick substrates. Lacking this charge in Z-Phe-Scam ( -S-CH2-CONH2instead -S-CH2-COOH)a complete loss of specificity results since no hydrolysis of Z-Phe-Scam could be observed. The utility of carboxymethyl thioester for V8 protease-catalyzed peptide synthesis could be demonstrated both by model acyl transfer reactions using amino acid and dipeptides as acyl acceptors and fragment condensation, respectively. Fig. 12.5-20 indicates a semipreparative (3 + 5) model fragment condensation. After 2 h the enzymatically nonoptimized coupling reaction of 1with an excess of 2 in HEPES-buffer containing 5 % DMSO was stopped by the addition of diluted trifluoroacetic acid and resulted in a yield of 55 %. has investigated In addition to carboxymethyl thioesters, Bordusa's further types of thioesters and phenylester bearing the carboxyl group, e. g. carboxyethyl thioester, 2-carboxyphenyl thioester, 3- and 4-carboxyphenyl ester, which also mediate acceptance by V8 protease. It is surprising to note that despite the lower degree of structural similarities, the aromatic part of the leaving group led to even higher specificity constants than found for the aliphatic counterparts. In addition, these studies have been expanded to the use of the not so expensive but equally Gluspecific endopeptidase from Bacillus licheni$obmis (BL-GSE),which can easily be purified from alcalase in good yields.

72.5 Hydrolysis and Formation of Peptides

12.5.3.6.4

I

849

Hydrophobic Substrate Mimetics

In addition to the enzymes mentioned above with a high specificity for positively and negatively charged P1’ amino acid residues, a third important class of enzymes are represented by peptidases with specificity for aromatic and hydrophobic functionalities. Well-known representatives of this family are the serine peptidases chymotrypsin and subtilisin, which have application in classical enzymatic peptide synthesis and, therefore, they should also be interesting biocatalysts for the substrate mimetic approach. Both enzymes primarly prefer bulky hydrophobic and aromatic P1‘ amino acid residues. In addition, the S1 binding pocket of subtilisin contains a carboxylic acid moiety (G~U’’~) which causes additional activity towards Arg and Lys r2031. For this reason, aromatic leaving groups with additional positively charged substitutions, e. g. 4-guanidinophenyl ester should fit the natural specificity of these peptidases. Parallel to an empirical design of specific mimetic structures, the well-known 3D structures of the two enzymes allow the use of rational approaches such as the computer-assisted protein-ligand docking approach. Using the latter to predict the function of the 4-guanidinophenyl ester functionality, Bordusa and co-worker selected Boc-Ala-OGp as a model ligand and docked it towards the enzyme[”’]. Fig. 12.5-21shows the arrangement of the ligand Boc-Ala-OGp at the active site of chymotrypsin in the lowest energy complex (A) in comparison with that found for trypsin (B) [lg81. In analogy to the natural specificity of chymotrypsin, hydrophobic contacts between the phenyl moiety of the ester group and the residues Cys”’ and Val213of the enzyme predominante. Interestingly, the guanidino functionality favors this binding mode by formation of additional hydrogen bonds with three serine residues which are located at the bottom of the S1 binding pocket. This specific binding pattern, specifically, the orientation of the carbonyl oxygen to Glylg3

Asp 1 A 191

--+ Asp189

Figure 12.5-21. Arrangements o f Boc-Ala-OCp at the active site o f chymotrypsin (a) and trypsin (b), respectively according to Cunther, Thust, Hofmann and Bordusa (see e. g. r e f e r e n ~ e ” ~ ’ ~ ) .

850

I

12 Hydrolysis and Formation ofC-N Bonds

80

10

Figure 12.5-22. Chymotrypsin-catalyzed peptide synthesis using 4-guanidinophenyl esters o f various non-specific and non-coded acyl moie-

H-Ala-Ala-NH -Gly-Leu-NHp

(oxyanionhole), the distance between the carbonyl C-atom of the scissile ester bond and the active Ser'", and the reversed binding of the acyl moiety fulfill the conditions for the binding and catalytic mechanism of substrate mimetics. Indeed, acyl 4-guanidinophenyl ester was hydrolyzed by chymotrypsin, and also peptide bond formation using various 4-panidinophenyl esters with nonspecific coded and non-coded acyl residues could be successfully performed as shown in Fig. 12.522[191]. The yields obtained are in the same range as the yield obtained using the normal-type acyl donor Bz-Phe-OMe. Furthermore, phenyl ester are also suitable substrate mimetics for chymotrypsincatalyzed peptide synthesis, as was established by Bordusa's group and will demonstrated by sophisticated fragment condensations in Sect. 12.5.3.7.

12.5.3.6.5

Enzymochemical Substrate Mimetic Approach

In order to synthesize longer polypeptides and proteins the condensation of the initial fragments is an essential prerequisite. Despite different chemical ligation techniques in the field of protein semisynthesis (cf. 12.5.3.1,p. 820-821) enzymatic C - N ligation seems to be the only way to avoid partial epimerization, which cannot be completely eliminated in the course of a chemical fragment coupling reaction. Consequently, the application of the substrate mimetic strategy for the peptidasemediated condensation of peptide fragments indisputably needs to be combined with the solid-phase peptide synthesis approach. Since a peptide ester can be 2051 Cerovsky and Bordusa[2061 achieved using of the oxime resin ~tartegy[~'~, developed a procedure for the synthesis of peptide fragments in the form of substrate mimetics esterified as 4-guanidinophenyl-,phenyl- and mercaptopropionic acid esters. The synthesis protocol involves covalent attachement of the first N"-Bocprotected amino acid to the oxime resin, blocking free hydroxylic groups by acetic

72.5 Hydrolysis and Formation of Peptides a)

b)

+ Protease

00000

a protecting group

-A

Figure 12.5-23. General approach t o fragment substrate mimetics via the oxime resin strategy (a) and substrate mimetic-supported peptide fragment condensation (b) catalyzed by specific peptidases according t o Cerovsky and Bordusa [2061.

0individual amino acid A specific leaving group

anhydride, deprotection of the N*-amino group of the attached amino acid, followed by successive chain elongation according to the well-known SPPS methodology.The generation of the peptide fragment in the form of the substrate mimetic can be performed by aminolysis of the oxime ester linkage between the peptide and resin, with the appropriate free amino acid substrate mimetic ester as shown schematically in the upper part (a) of Fig. 12.5-23. After deprotection of side-chain functions of the amino acid residues, and if necessary also those of the ester leaving group, the only Nu-protected peptide ester can be coupled with an amine component using the suitable peptidase (b).Some examples of model fragment condensations using this approach with catalysis from three different peptidases are given in Fig. 12.5-24. The coupling reactions were performed on a preparative scale using 1 : 2 ratios of acyl donor ester to the nucleophilic acyl acceptors (in the case of trypsin 1 : 2.5) resulting in product yields between GO-70 %. 12.5.3.7

Planning and Process Development of Enzymatic Peptide Synthesis

The high enantio- and diastereoselectivity in peptidase-catalyzed peptide synthesis allows, in constrat to most chemical coupling methods, the formation of peptide bonds without partial epimerization in the C-terminal amino acid residue of the carboxyl component. Furthermore, owing to the regiospecificity of the enzymes, tedious protection/deprotection steps are not problems in the enzymatic approach. Using serine and cysteine peptidases a further point needs to be decided; namely, should the carboxyl component be used as the acylamino acid or should an ester be used in order to favor acylation of the enzyme. Enzymatic synthesis using peptide esters or amino acid esters as substrates has the clear advantage of proceeding at a high rate, thereby demanding a low concentration of enzyme and, furthermore, being completely independent of the solubility of the starting materials and product. Although the kinetically controlled synthesis would be preferable, the decision should depend on the total synthetic concept. An unfavorable nucleophile specificity may be better taken care of in an equilibrium-controlledreaction with the necessary manipulations of conditions. In spite of some limitations the equilibrium-controlled approach has proved to be worthwhile in the trypsin-catalyzed semisynthesis of

852

I

72 Hydrolysis and Formation of C-N Bonds

a)

Boc-Tyr(Bz1)-Pro-Ser(Bz1)-Ala-Leu-0-P + H-Ala-OGp(Z)z

Boc-Tyr(Bzl)-Pro-Ser(Bzl)-Ala-Leu-Ala-OGp(Z)2 1 HZlPd

1 + I

Boc-Tyr-Pro-Ser-Ala-Leu-Ala-OGp H-Met-Ala-Ala-Ala-GIy-OH 2 I 3 trypsin Boc-Tyr-Pro-Ser-Ala-Leu-Ala-Met-Ala-Ala-Ala-Gly-OH 4

b)

I

Boc-Trp-He-He-Leu-0-P + H-Gly-SCe

Boc-Trp-He-He-Leu-Gly-SCe 5

+

1

H-Leu-Ala-Ala-Ala-GIy-OH 6 V8 protease

Boc-Trp-lle-lle-Leu-Gly-Leu-Ala-Ala-Ala-Gly-OH 7

c)

Boc-Leu-Asn-Lys(Z)-Ile-0-P

+

H-Val-OPh

i

I

Boc-Leu-Asn-Lys(Z)-He-Val-OPh 8 HdPd Boc-Leu-Asn-Lys-lle-Val-OPh + H-Arg-Ala-Ala-Ala-Gly-OH 10 9 chymotrypsin

1

Boc-Leu-Asn-Lys-He-Val-Arg-Ala-Ala-Ala-Gly-OH 11 Combination of solid-phase peptide synthesis and sub. strate mimetic-supported segment condensations with different peptidases and substrate mimetics according t o Cerovskyy and Bordusa[2061. Figure 12.5-24.

human insulin as well as the industrial aspartame synthesis using therrnolysin. In order to overcome poor solubility of the starting components the introduction of solubilizing protecting groups is frequently necessary.

12.5 Hydrolysis and Formation ofpeptides

12.5.3.7.1

I

Stepwise Chain Elongation

Contrary to chemical synthesis, enzymatic stepwise chain building may start either from the N-terminus or from the C-terminus. In chemical synthesis, incremental chain lengthening from the N-terminus, as performed in ribosomal protein synthesis, is normally not recommended under preparative conditions, since the efforts needed to avoid the permanent risk of partial epimerization outweigh the potential gain. Despite these principal limitations, investigations on solid-phase peptide synthesis in an N-to-Cdirection, called inverse synthesis, has been performed using HOBt salts of the amino acid 9-fluorenylmethyl esters L2O71. Unfortunately, the racemization problem could not be excluded. Furthermore, Mitin and Ryadnov[208] have described inverse peptide synthesis in order to exclude deprotection reactions at every solution synthesis stage. This could be realized using the high solubility of free amino acids in dimethylformamide containing Ba(C104)2, Ca(C104)2 or Ca(N03)2. An attempt to solve the extensive exclusion of racemization was tried using copper(n)ions (CuC12)during activation of the carboxyl group with ethyldimethylaminopropylcarbodiimide (EDC) as the coupling reagent in the presence of HOBt [2091. In a general sense exopeptidases should be the enzymes of choice for stepwise chain assembly since once formed the internal peptide bonds of the growing chain can no longer be proteolytically cleaved from this type of peptidase. Carboxypeptidase exhibit superior properties for the stepwise synthesis, especially, carboxor other serine peptidases of this type. In principle, ypeptidase Y (CPD-Y)[2101 aminopeptidases can also be used starting from the C-terminus. Because under these conditions not only the carboxyl component but also the amine component has a free a-amino function, product isolation is more difficult, particularly, if one component is used in excess. Otherwise, stepwise synthesis from the C-terminus is not a problem in chemical peptide synthesis. A classical example for a kinetically controlled synthesis starting from the Nterminus and using CPD-Y as an enzyme for all coupling steps was described by Bz-Arg-OEt was couWidmer et al. f211] for [Metlenkephalin(Tyr-Gly-Gly-Phe-Met). pled with H-Tyr-NH2 at pH 9.6 giving the Bz-dipeptide amide in 85% yield. The CPD-Y-catalyzeddeamidation at pH 9.6 provided Bz-Arg-Tyr-OHin 90 % yield. After chemical esterification with EtOH/HCl, the resulting Bz-Arg-Tyr-OEtwas coupled with H-Gly-OEt at pH 9 to give the protected tripeptide derivative (yield: GO%), followed by the successive addition of the other amino acid derivatives in the same manner. Amino acid amides were preferred as the amine components, since free amino acids (except Met) only give low yields and amino acid esters give rise to side reactions that are difficult to control. Finally, the protecting group for the P-amino function of Tyr, the Bz-Arg moiety, was easily removed with trypsin. The disadvantage of this synthesis strategy seems to be the complicated route of selectively removing the C-terminal amide grouping by means of CDP-Y. This step followed by chemical esterification of the peptide had to be resorted to before it was possible to use the intermediate in the next coupling reaction as the carboxyl component. A second step-by-steppeptide synthesis from the N- to C-terminus was described

853

OEt

SBzl H 1

Z

a) Yield: 89%

Z

Z H

OEt H

3

OPr

b)

Yield: 84%

5

OPr H

OBZ'

C)

Yield: 83%

4

7

Yield: 100%

8

Tyr-Arg-Ser-OH from N- t o C-terminus using clostripain and chymotrypsin, respectively, as biocatalysts according t o Bordusa et al.[208].a) and c): Clostripain; b): chymotrypsin; d): catalytic hydrogenation using 10% Pd/C; -OPr, propyl ester; -SBzl, thiobenzyl ester.

OBzl

OH

by Bordusa et al. 1212] for the model tetrapeptide H-Lys-Tyr-Arg-Ser-OHbut using the endopeptidases clostripain and chymotrypsin as biocatalysts (Fig. 12.5-25). The synthesis could be performed without side chain protection for all trifunctional building blocks and the only nonenzymatic reaction was the final catalytic hydrogenation for cleavage the terminal blocking groups. As a rule, peptidases can only make a meaningful contribution to a synthesis strategy if the full advantagee of the enzymatic reactions can be utilized. An a priori completely unrealistic position is the comparison of a stepwise peptidase-catalyzed assembley of a peptide chain with the automatic solid-phase technique. On the other hand, selected di- and tripeptides can be synthesized enzymatically using solubilizing protecting groups on a large scale, even in a continuous process[118-120] (cf. 12.5.3.4.1, p. 832-833). In addition, the solid-to-solidconversion has proven to be a very useful method for the synthesis of selected short peptides which fulfil the requirements for this special synthetic procedure (cf. 12.5.3.4.2, p. 838-839).

12.5.3.7.2

Fragment Condensation

This approach has some advantages over the stepwise strategy. Firstly, if small fragments are combined to make one which is larger its isolation is more easily facilitated in contrast to a stepwise synthesis, and, secondly, the fragment condensation approach offers the possibility of synthesizing a set of related analoges with variabel sequences in a region. In principle it is possible to synthesize peptides using enzymes both for protection/deprotection procedures as well as for the formation of the peptide bonds. Fig. 12.5-26 shows the fully enzymatic synthesis of the tert.-butyl ester of Leu-enkephalin12131 using both equilibrium and kinetically controlled coupling steps. In order to obtain the unprotected Leu-enkephalin, the C-terminal protecting group must be split off by chemical means. Although, in principal it is possible to perform totally enzymatic synthesis of peptides, in practice combined chemical and enzymatic steps are preferred. For the classical enzymochemical synthesis of polypeptides and even small proteins, the optimum approach is usually synthesis of fragments using the SPPS methodology

Hl

72.5 Hydrolysis and Formation of Peptides

1

G~Y p

n PhAc .;iH

OMe

Phe h

i

PhAc

Figure 12.5-26. Fully enzymatic synthesis of ~ ~ ~ [Leulenkephalin ~ ~ tert-butyl OBU' PA.. DeniciIIin . acylase; CT, chymotrypsin; ~ OBU' P, P papain; PhAc, h phenyla~

A

~

OBu'

OMe

~

I

855

Leu

~

cevl.

P PhAC

1

I

I

I

for enzymatic conjunction in a overall divergent strategy. In a given synthesis project initially it is necessary to separate the whole sequence into segments containing favorable combinations of amino acids which permit peptidase-catalyzed segment coupling according to the eluciated S'-subsite specificity. Since the kinetic parameters of the enzymatic synthesis course are often not available, they can be estimated from the data for similar substrates and nucleophilic amine components. Based on such estimates an optimum synthesis strategy can be established. In Table 12.5-10selected examples of enzymatically synthesized peptides are compiled. During the last decade in particular, remarkable efforts have been made to find optimum conditions for peptidase-catalyzedpeptide synthesis including the development of new reaction conditions and new biocatalyts. Once the optimal synthesis conditions have been recognized, kg amounts of biologically active peptides can be produced. The synthetic biotransformations can normally be achieved with commercially available enzymes which are easy to handle. In addition, owing to the application in only catalytic amounts the higher costs of the enzymes used are usually insignificantly in comparison with highly sophisticated chemical coupling reagents plus the financial expense of the reagents necessary for protection/ deprotection procedures in chemical synthesis. As a model system for peptidase-catalyzed modification of peptides produced by recombinant DNA technology Schellenberger et al. [243, 244] developed a new approach to the production of peptides based on chemical synthesis and peptidasecatalyzed processing (Fig. 12.5-27). First, they produced an artificial substance P precursor as a P-galactosidase (1-459) fusion protein containing nine copies of the sequence H-Arg-Leu-Arg-Argl-Pro-Lys-Pro-Gln-Gln-Phe7-OH. The sequence of the peptide precursor was designed to meet the specific requirements of chymotrypsin and papain, respectively, used in conversion reactions as the complete amino acid sequence should be regenerated by addition of the appropriate dipeptide derivatives. After isolation and purification of the fusion protein, which was accumulated in E. coli as inclusion bodies, the dodecapeptide ester H-Arg-Leu-Arg-Arg'-Pro-Lys-Pro-

~

~

12 Hydrolysis and Formation ofC-N Bonds Table 12.5-10.

Selected examples of enzymaticallysynthesized peptides.

Peptide/Protein

Synthesis route*

References

Angiotensin I1 (analog) Aspartame Calcitonin (salmon) Calcitonin (dicarba analogs) Caerulein Caerulein (analog) Cholecystokinin-8 Cholecystokinin-8 (analogs) Delta sleep inducing peptide (DSIP) Dynorphin-(1-8) EGF (3-14,21-31, 33-42) EGF (29-44) Eledoisin (611) Eledoisin [Met/Leu]Enkephalin [MetIEnkephalin [LeuIEnkephalin [LeuIEnkephalinderivatives Growth hormone releasing factor (human) analog Hepatitis B S antigen (122-137) Ht31(493-515) peptide Insulin (human) Kyotorphin LH-RH [D-Phe6]LH-RH MSH (5-8,9-12,13-16) Oxytocin (1-9) Ribonuclease A Somatostatin Substance P (6-11,7-11) Vasopressin (1-6) * E, equilibrium approach: K, kinetic approach; total, totally enzymatic coupling; part, partly enzymatic

coupling

Gln-Gln-Phe-Gly'-OMe was formed by chymotrypsin-catalyzed transpeptidation in the presence of H-Phe-Gly-OMe. In a papain-catalyzed acyl transfer reaction and subsequent tryptic cleavage, the resulting dodecapeptide ester was converted into substance P. These results indicate that peptides can be readily produced by a combination of recombinant DNA technology and peptidase-catalyzed conversion with the advantage of possible incorporation of groups other than coded amino acids into the recombinant product. The chemoenzymatic synthesis of RNase A['"] using a mutant of subtilisin BNP', called subtiligase, underlines the progress of enzyme-catalyzed fragment condensations in the course of the synthesis of a small protein. The fragments (98-124, 77-97,64-76,52-63,21-51 and 1-20) were synthesized by standard SPPS methodology. The choice of the fragments was solved in such a way that the C-terminal residues of the appropriate fragments (Tyr", Tyr", Val63and Alazo)were the closest

12.5 Hydrolysis and Formation ofPeptides

I

857

Linker Sequence

\

Helper Protein

c

<Substance P-(l-7)] 00000 Arg-Leu-Arg-Arg-Pro-Lys-Pro-Gln-Gln-Phe-Arg-Leu-Arg-[

SP1-7],00000

+ ff-Phe-Gly-OMe Chymotrypsin-catalyzedtranspeptidation 00000 Arg-Leu-Arg-Arg-Pro-Lys-Pro-Gfn-Gln-Gln-Phe-Phe-Gly-OMe

1

+ /+-/-eu-Met-Nff2

Papain-catalyzedacyltransfer 00000 Arg-Leu-Arg-Arg-Pro-Lys-Pro-Gln-Gln-Phe-Phe-Gly-Leu-Met-NH~

1

Trypsin-catalyzedcleavage n H-Arg-Pro-Lys-Pro-GIn-Gln-Phe-Phe-Gly-Leu-Met-NH2 Substance P Figure 12.5-27. Peptidase-catalyzed modification of an artificial substance P precursor protein according to Schellenberger et a/. [2431.

to matching the substrate specificity of the subtilisin mutant. Using a considerable excess of the fragments bearing a Phe-NH2-modified carboxamido methyl ester ensured that most of the side reactions could be suppressed. Starting with the Cterminal fragment (98-124) the total yield after five fragment condensations was 15%, and after folding the final protein could be obtained in 8% yield. In a similar manner the three analogs of RNase A were synthesised in which the two residues His” and His’19 of the active center were exchanged individually and simultaneously for L-4-fluorohistidine. Despite this impressive example of five successful enzyme-catalyzed fragment condensations with average yields of roughly 75 % in the course of the synthesis of RNase A all the peptide bond forming steps could not be performed irreversibly. Even though the new C - N ligation strategy based on the substrate mimetic concept (cf. Sect. 12.5.3.6)has not as yet been proved for the synthesis of a similar protein target, it guarantees the irreversibility of the enzymatic coupling reaction, as can be demonstrated by the chymotrypsin-catalyzed (8 + 16) fragment condensation of the Ht 31(493-515) peptide derived from the human protein kinase A anchoring protein (sequence 493-5F1)[~~~]. The synthesis of the 24-peptide was accomplished by the chymotrypsin-catalyzed fragment condensation at a nonspecific Ser-Arg peptide bond via the substrate mimetics strategy (Fig. 12.5-28). The fully protected carboxyl component 1 was synthesized on Kaiser’s oxime resin and was released from the support by aminolysis with H-Ser(Bz1)-OPaccording to the procedure described on p. 850-851. After side-chain deprotection by catalytic hydrogenation, 2 was coupled with the unprotected amine segment 3 catalyzed by chymotrypsin, leading to the complete conversion of both peptide segments. Finally, the N-terminal Boc group was cleaved by TFA giving the desired Ht 31 (493-515) peptide 5.

858

I

12 Hydrolysis and Formation ofC-N Bonds

resin

Boc-Asp(OBzl)-Leu-lle-Glu-(OBzl)-Glu(OBzl)-Ala-Ala-O-oxime

I I

TFA.Ser(Bzl)-OPh

Boc-Asp(OBzl)-Leu-lie-Glu-(O6zl)-Glu(OBzl)-Ala-Ala-Ser(Bzl)-OPh

1

H,IPd

Boc-AspLeu-Ile-Glu-Glu-Ala-Ala-Ser-OPh

2

H-Arg-lle-Val-AspAla-Val-lle-Glu-Gln-Val-Lys-Ala-AIa-Gly-Ala-Tyr-OH

I

I

3

1

Chyrnotrypsin

Boc-Asp-Leu-lle-Glu-Glu-Ala-Ala-Ser-Arg-lle-Val-Asp -Ala-Val-lle-Glu-Gln-Val-Lys-Ala-Ala-Gly-Ala-Tyr-OH

4

1

TFA

H-Asp-Leu-lle-Glu-GIu-Ala-Ala-Ser-Arg-ile-Val-Asp-Ala-Val-lle-Glu-Gln-Val-Lys-Ala-Aia-dy-Ala-Tyr-OH

5

Figure 12.5-28. Chymotrypsin-catalyzed (8+16) segment synthesis of the Ht 31 (493-51 5) peptide via substrate mimetic strategy[233].

12.5.4 Conclusion and Outlook

Despite the fact that chemical methods are popular for the synthesis of peptides a huge number of papers has been published in recent decades dealing both with enzymatic formation of peptide bonds and enzymatic manipulation of protecting groups. Enzymatic methods have several advantages over chemical procedures but at present more peptides are synthesized by chemical synthesis than in peptidasecatalyzed processes. The use of peptide synthesizers, in addition to recent new developments in the field of chemical ligation procedures, still favor chemical methods compared with the enzymatic approach. However, there is no doubt that enzymatic methods have advantages, including the prevention of racemization, no need for time-consuming and expensive protection/deprotection procedures of sidechain functions, the reduced use of problematic (toxic) solvents and reagents and possible reuse of the biocatalysts. The question should not be whether to use a chemical or an enzymatic approach in peptide synthesis; an ingenious combination of chemical and enzymatic steps should promote the general progress in peptide synthesis. It could be demonstrated that after establishing the optimal synthesis conditions, kg amounts of biologically active peptides and analogs can be obtained using enzymatic coupling methods. The semisynthetic synthesis of human insulin and the

References I859

production of aspartame in a ton-scale underline the industrial importance of the enzymatic approach. However, the enzymatic approach does not have the versatility of chemical synthesis methods and suffers from some limitations. The main reason seems to be the lack of a universal enzyme which is capable of catalyzing peptide bond formation for all possible combinations of the 21 proteinogenic amino acid residues located both as C- and N-terminal building blocks in peptide fragments to be coupled. Such an enzyme could not be developed during evolution due to the extremely high specificity requirements. In ribosomal protein synthesis nature prefers the stepwise synthesis from N- to C-terminus followed by maturation procedures based on limited proteolysis and further modifications. The only biocatalyst involved in ribosomal synthesis, the peptidyl transferase, seems to be an old ribozyme without any specificity for the PI side chain functions of the amino acids, only catalyzing the acyl transfer reaction of the selected aminoacyl-tRNAs. Since such a biocatalyst has no practical importance in peptide synthesis in a peptide laboratory, the only alternative for this purpose is the reverse catalytic hydrolysis potential of proteases. The advantages and drawbacks of peptidases used for catalyzing peptide bond formation have been demonstrated in this contribution. An ingenious combination of chemical and enzymatic strategies as demonstrated in a new synthesis of RNase A should be the state-of-the-artin this field at present. Furthermore, using the new C N ligation strategy based on the substrate mimetic concept, irreversible peptide bond formations catalyzed by high specific peptidases can be performed for the first time. In combination with peptidase mutants which lack amidase activity, this new C - N ligation approach will contribute to significant progress in enzymatic peptide synthesis, especially in clear-cut fragment condensations using recombinant polypeptide thioester as the substrate mimetics with chemically synthesized or recombinant fragments. This specific programming of enzyme specificity by molecular mimicry corresponds in practice to a conversion of a peptidase into a C - N ligase, a biocatalyst which could not developed by nature during evolution.

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862

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72 Hydrolysis and Formation of C-N Bonds

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51,83-128. 211 F. Widmer, K. Breddam, 1. T. johansen in: Peptides 1980 (Ed.: K. Bmnfeldt) Scriptor, Copenhagen, 1981, pp. 46-55. 212 F. Bordusa, D. Ullmann, H.-D. Jakubke, Angm. Chem., Int. Ed. Engl. 1997,36, 1099-1101. 213 R. J. Didziapeptris, B. Drabnig, V. Schellenberger, H.-D. Jakubke, V. Svedas, FEES Lett. 1991,287,31-33. 214 Y. Isowa, M. Ohmori, M. Sato, K. Mori, Bull. Chem. SOC.Jpn. 1977,50,2766-2772. 215 K. Oyama, S. Nishimura, Y. Sonaka, K. Kihara, T. Hashimoto, J. Org. Chem. 1981,46, 5242-5244. 216 M. Condo, H. Yamashita, K. Sakakibara, Y. Isowa in: Peptide Chemistry 1981 (Ed.: T. Shiori), Protein Research Foundation, Osaka, 1982, pp. 93-98. 217 V. Cerovsky, E. Wiinsch, 1. Brass, Eur. J. Biochem. 1997,247,231-237. 218 H. Takai, K. Sakato, N. Nakamizo, Y. Isowa in: Peptide Chemistry 1980 (Ed.: K. Okawa), Protein Research Foundation, Osaka, 1981, pp. 213-218. 219 W. Kullmann, Proc. Natl. Acad. Sci. USA 1982,79,2840-2844. 220 V. Cerovsky, 1. Hlavacek, 1. Slaninova, K. Jost, Coll. Czech. Chem. Commun. 1980, 53,2766-2772. 221 V. Cerovsky, J. Pirkova, P. Majer, J. Slaninova, 1. Hlavacek in: Peptides 1988 (Eds.: G. Jung, E. Bayer), de GruyterBerlin, New York, 1989, pp. 265-267. 222 M. Capellas, G. Caminal, G. Gonzalez, J. Lopezsantin, P. Clapes, Biotechnol. Bioeng. 1997,56,456-463. 223 K. Sakina, K. Kawazura, K. Morihara, Int. J . Peptide Prot. Res. 1988, 31, 245-251. 224 W. Kullmann,/. Org. Chem. 1982,47, 5300-5303. 22s F. Widmer, S . Bayne, G. Houen, B. A. Moss, R. D. Rigby, R. G. Whittaker, J. P. Johansen in: Peptides 1984 (Ed.: U. Ragnarson), Alqvist Wiksell, Stockholm, 1984, pp. 193-200. 226 F. Widmer, S. Bayne, G. Houen, B. A. Moss, R. D. Rigby, R. G. Whittaker, J. T. Johansen in: Forum Peptides Le Cap d'Agde 1984 (Eds.: J. B. Castro, J. Martinez), Groupe Francais de Peptides, 1985. 227 H.-D. Jakubke, P. Kuhl, A. Konnecke, G.

Doring, 1. Walpuski, A. Wilsdorf, N. P. Zapevalova in: Peptides 1982 (Eds.: K.Blaha, P. Malon), de Gruyter, Berlin, New York, 1983, pp. 43-45. 228 P. Kuhl, G. Doring, K. Neubert, H.-D. lakubke, Monatsh. Chem. 1984,115,423-430. 229 P. Bjorup, 1. L. Torres, P. Adlercreutz, P. Clapes, Bioorgan. Med. Chem. 1998,6, 891-901. 230 W. Kullmann, Biochem. /. 1984, 220, 405-416. 231 Y. H. Ye, G. L. Tian, D. C. Dai, G. Chen, C. X. Li, Tetrahedron 1998,54, 12585-12596. 232 S.Aasmul-Olsen, A.J. Andersen, P. Thorbek, F. Widmer in: SthInt. Con& on Tetanus (Eds.: G. Nistica, B. Bizzini, B. Butchenko, R. Trian), Pythagora Press, Rome, Milan, 1989, pp. 191-208. 233 V. Cerovsky, J. Kockskaemper, H. G. Glitsch, F. Bordusa, ChemBioChem. 2000, 126-129 234 R. Obermeier, G.Seipke, Process Biochem. 1984,29-34. 235 A. J. Andersen, F. Widmer, J. T. Johansen in: Peptides 1986 (Ed.:D. Theodoropoulos), de Gruyter, Berlin, New York, 1987, pp. 183-188. 236 M. Schuster, A. Aaviksaar, H.-D. Jakubke, Tetrahedron Lett. 1992, 33, 2799-2802. 237 V. Schellenberger, U. Schellenberger, H.-D. Jakubke, A. Hansicke, M. Bienert, E. Krause, Tetrahedron Lett. 1990, 31, 7305-7308. 238 W. Kullmann,]. Prot. Chem. 1983,2, 289-301. 239 P. Thorbek, 1. Lauridsen, F. Widmer in: Peptides: Chemistry and Biology, Proc. loth Am. Pept. Symp. (Ed.: G.R. Marshall), ESCOM, Leiden, 1988, pp. 279-281. 240 V. Bille, C.Ripak, I. van Assche, L. Forni, J. Degelaen, A. Scarso in: eptides 1990 (Eds.: E. Giralt, D. Andreu), ESCOM, Leiden, 1991, pp. 253-254. 241 P. Kuhl, G.Doring. K. Neubert, H.-D. Jakubke, Pharmazie 1984,39,814-816. 242 V. Cerovsky, Coil. Czech. Chem. Commun. 1986,51,1352-1360. 243 V. Schellenberger, W. Tegge, R. Frank, Int. 1. Pept. Protein Res. 1992, 39,472-476. 244 V. Schellenberger, M. Pompejus, H.-J. Fritz, lnt. J. Pep. Protein Res. 1993,41, 326332. 255 I. Schechter, A. Berger, Biochem. Biophys. Res. Commun. 1967,27,157-162.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

866

I

72 Hydrolysis and Formation ofC-N Bonds

12.6 Addition of Amines to C = C Bonds

Marcel Wubbolts

The ammonia lyases (E. C. 4.3.1.x),which catalyze the addition of amines to carboncarbon double bonds, belong to the class of carbon-nitrogen lyases. The reactions catalyzed by ammonia lyases are in equilibrium and depending on reaction conditions the reaction can be directed either towards ammonia addition or in the direction of elimination of ammonia. Ammonia lyases in their natural role are involved in the metabolism of amino acids and also play a role in, for instance, the degradation of amino sugars, but only a limited amount of these enzymes have been characterized biochemically. Application of a broad range of different ammonia and lyases in organic chemical synthesis on an industrial scale has thus far not occurred, which is due to both their limited commercial availability and their lack of stability under process conditions. Exceptions are the commercially applied aspartase, which is an ammonia lyase that is utilized for the synthesis of 1-aspartic acid from fumaric acid, and phenylalanine lyase. The latter is an example of a commercial application of an ammonia lyase in a process for the production of L-phenylalanineand more importantly L-phenylalanine derivatives. 12.6.1

Addition of Ammonia to Produce Amino Acids 12.6.1.1

Aspartic Acid

L-aspartic acid ammonia lyase, or aspartase (E.C. 4.3.1.1) is used on a commercial scale by Kyowa Hakko, Mitsubishi, Tanabe and DSM to produce L-aspartic acid, which is used as a building block for the sweetener Aspartame, as a general acidulant and as a chiral building block for synthesis of active ingrediants [ll. The reaction is performed with enzyme preparations from E. coli, Breuibacterium Jauum or other coryneform bacteria either as permeabilized whole cells or as isolated, immobilized enzymes. The process is carried out under an excess of ammonia to drive the reaction equilibrium from fumaric acid (1)in the direction of r-aspartic acid (L-2) (see Scheme 12.6-1) and results in a product of excellent quality (over 99.9% e. e.) at a yield of practically 100%.The process is carried out on a multi-thousand ton scale by the diverse producers of L-aspartic acid. Site directed mutagenesis of aspartase from E. coli by introduction of a Cys430Trp mutation has resulted in significant activation and stabilization of the Since maleic acid is a cheaper starting material than fumaric acid, the process that is probably the most economical makes use of both a maleate isomerase (E.C. 5.2.1.1) and aspartase (E. C. 4.3.1.1), Scheme 12.6-1. Mitsubishi has succeeded in

12.G Addition ofAmines t o

C = C Bonds

I

867

Aspartase

+COOH H

O

O

C

NH:,

L-2

Maleate lsomerase

COOH L C O O H

3

Aspartase

’

COOH H

O

O L-2

Aspartase

COOH H

O

O

C

-

T

T

C

T

NH2

Fumarate hydratase

-

+-----H OO C , - HOOC , - ,

+ I.c-- H

-

NHZ

COOH O

O

Aspartase

&COOH HOOC 1

___L

4 -

4

Aspartate-p decarboxylase

L-2

-

T

OH

NHz D-2

rac 2

C

Y C O O H NH2

Scheme 12.6-1.

combining both activities in a Brevibacteriumjavumrecombinant for the large-scale production of L-aspartic acid13]. Mitsubishi has also developed a process for production of D-asparticacid (D-2)and L-malic acid ( 4 ) by incubation of racemic aspartic acid with the exclusively L-selective aspartase in combination with fumarase, thereby preventing the reaction going backwards by conversion of the generated fumaric acid into L-malic acidI4]. The combined utilization in a single reactor of both aspartase from Brevibactevium flavum and aspartate-P-decarboxylase from Pseudomonas dacunhae, thereby catalyzing the reaction from fumaric acid via L-aspartic acid to L-alanine (S),has also been developed by Mitsubishi[’I. Another combination reaction is the biocatalytic production of the herbicide phosphinotricin [ ~-2-amino-4-(hydroxymethylphosphinyl)butyric acid, (7)in Scheme 12.6-21 by the company Meiji Seika, whereby an amino-transferase that acts on 4-(hydroxymethylphosphinyl)-2-oxo-butyric acid and that utilizes aspartic acid as the amino donor was used in combination with aspartase to generate the amino donor from fumaric acid and ammonia[‘].

868

I

72 Hydrolysis and Formation ofC-N Bonds 0

OH

Aspartase Amino transferase

0

6

OH

7

Scheme 12.6-2.

12.6.1.2

Aspartic Acid Derivatives

The enzyme methylaspartate ammonia lyase (P-methylaspartase, E. C. 4.3.1.2) is involved in the metabolism of branched pentanoic acids. The enzyme catalyzes the addition of ammonia to mesaconic acid (8) to yield ~-threo-3-methylaspartate(9) as depicted in Scheme 12.63. The enzyme has been shown to be induced under anaerobic conditions in facultative anaerobes such as Citrobacter, Proteus, Escherichia coli and Enteroba~ter[~. 81 and has been applied for the synthesis of 3-substituted (S)aspartic acid derivatives, such as (2S,3S)-3-methylasparticacid (9), (2S,3S)-3-ethylaspartic acid (ll),and (2R,3S)-3-chloroasparticacid (13)"1. In addition, a process for the preparation of dialkyL(2S,3S)-3-ethylaspartatesusing methylaspartate ammonia lyase has been developed by Merck['I]. Bear et al. have been using methylaspartate ammonia-lyase from Clostridium tetanomorphum to produce optically active pure precursors [3-methyl-,3-ethyl and

-

Methylaspartate Ammonia Lyase

HOOCL

C

O

O

H

O

O

H

O

O

H

7

a

HOOCL

C

___)

-c--.---

10

HOOCL

C

____)

7

13

12

HOOCL

C

14

HOOC+

O

O

H

____)

P

NH2

12.6 Addition ofAmines to C = C Bonds

I

869

Rp rcoo Phenylalanine

P _____) Lyase Ammonia

RT

NH2

19

e0/ C;3.iUr

R = NOz, CI. NH2. OH, CH3 at o. rn and p position

0

Phenylalanine Ammonia Lyase

=

q-cooH y 3,4-Dihydroxyphenylalanine Ammonia Lyase

COOH

=

HO

23

HO

OH

OH

Scheme 12.6-4.

3-iso-propylaspartic acids, (15)] for the synthesis of benzyl 3-alkylmalolactonates, which are suitable building blocks for semi-crystallinepolyesters ("1. 12.6.1.3

Histidine Ammonia Lyase

Histidine ammonia lyase (HAL, histidinase, histidine-a-deaminase, E. C. 4.3.1.3) is capable of abstracting ammonia from L-histidine (17),resulting in the formation of urocanoic acid [Scheme 12.6-4, (G)], an intermediate in the metabolism of Lhistidine("]. HAL has also been identified as a key enzyme in the synthesis of secondary metabolites such as Nikkomycin in Streptomyces teradae['21. The mechanism of the enzyme has been investigated and seems to proceed via the carbanion intermediate [l', 131. Synthetic applications of HAL are difficult to achieve, particularly as the enzyme is sensitive to oxygen[13].The utility of HAL is limited to niche applications such as the synthesis of radiolabeled urocanic acids as tracers of histidine metabolism [ll].

870

I

12 Hydrolysis and Formation of C-N Bonds L-Serine Deaminase

0

NH2

L-Threonine Deaminase

[ "r:cooH] /\ljCOOH

NH2

0

Scheme 12.6-5.

12.6.1.4

Phenylalanine, Tyrosin and L-DOPA

Phenylalanine ammonia lyase (PAL, E. C. 4.3.1.5)is an enzyme of relaxed substrate (18),R = H) and p specificity that accepts both trans-cinnamic acid (Scheme 12.6-4: coumaric acid [(19),R = OH] as substrates and thus results in the formation of the natural amino acids L-phenylalanineand L-tyrosine.The enzyme plays an important role in the synthesis of alkaloids, flavenoids and lignin in plants. The reaction has been exploited by Mitsui[14,"1, Great Lakes/NSC[lGland others to implement synthetic routes for non-natural substituted derivatives of L-phenylalanine starting from trans-cinnamic acids, for instance using the PAL enzymes from Rhodotorula rubra, Rhodotorula glutinis or Rhodosporidium toruloides. The PAL mediated synthesis of a variety of L-phenylalanine derivatives, carrying aromatic ring substituents such as nitro-, chloro-, amino-, hydroxy- and methyl groups at the 2, 3 and 4 position have Also, the synthesis of N-heterocyclicmolecules, derived thus been described116181. The direct synthesis of from phenylalanine by PAL has been shown['7, 19, I.' (21)] as a building block for aspartame, phenylalanine methyl ester [Scheme 12.6-4, from trans-cinnamyl methyl ester (20) by PAL from Rhodotorula glutinis further illustrates the synthetic versatility of PAL'", 1'. Radioactive tracers derived from Lphenylalanine have also been made with the aid of PAL[23s241s An enzyme that is related to PAL, dihydroxy-L-phenylalanine ammonia lyase (E. C. 4.3.1.11), is capable of synthesizing L-DOPA (23) from 3,4-dihydroxy-trans-caffeic acid (22), but this starting material is not as readily available as catechol, pyruvate, and ammonia are. As a result, the tyrosine phenol-lyase (TPL, E.C. 4.1.99.2) of Envinia herbicola is the enzyme of choice for biocatalytic L-DOPAproduction[25,"1, particularly as productivity has been increased since the TPL encoding gene from Enuinia herbicola was cloned and has been overexpressed successfully[25].

12.G Addition ofAmines t o

C = C Bonds

12.6.1.5

Serine and Threonine Deaminases

Both the L- and D-serine deaminase catalyze the elimination of the amino functionality of both L- and D-serine,but the mechanism proceeds via the initial elimination of water and these enzymes are thus classified as hydrolyases (L- and D-serine dehydratases E. C. 4.2.1.13 and E. C. 4.2.1.14, respectively)[27, *‘I. The aminoacrylate generated is unstable and subsequent elimination of the amine results in the formation of pyruvate. Similarly, threonine deaminase is in effect a dehydratase that converts L-threonine into 2-oxobuturate, water and ammonia (E. C. 4.2.1.16) (Scheme 12.6-1). 12.6.1.6 Ornithine Cyclodeaminase

Ornithine cyclodeaminase (E.C. 4.3.1.12) is an ammonia lyase that is not ubiquitously present but which has been identified in genera such as Rhizobium,Agrobactehum, Pseudomonas, Rhodobacter and Clo~tridium[~”~~]. Ornithine cyclodeaminase, which contains NAD that is tightly bound to the enzyme, catalyzes the conversion of L-ornithine, an intermediate in the metabolism of L-arginine, into L-proline. The reaction is peculiar among the ammonia lyases in that it involves a deamination of the amino group at the a-position followed by attack of the &amino group to give 2-0x0-5-aminopentanoicacid to form proli line[^^]. Conversions other than that from L-ornithine to L-proline have not been described. 12.6.2 Ammonia Lyases that Act on Other Amines 12.6.2.1

Elimination of Ammonia from Ethanolamine

The elimination of ammonia from ethanolamine to give acetaldehyde, which involves vitamin BIZ and which has been demonstrated to proceed via a radical anionL34,351, is catalyzed by ethanolamine ammonia lyase (EAL, E.C. 4.3.1.7). Genetic and biochemical analysis of the ethanolamine ammonia lyase isolated from Salmonella tphimurium and Rhodococcus sp. have been carried 371 and the enzyme appears to belong to a class of BIZ dependent enzymes that catalyze similar rearrangements, such as diol dehydratase and methylmalonyl-CoA mutase [351. Ethanolamine ammonia lyases are induced under anaerobic conditions, which is required since the radical reaction intermediates are highly reactive with dioxygen[38].Despite the interesting chemistry, we did not come across synthetic applications of ethanolamine ammonia lyases, other than the observation that the enzyme of Acetobacterium catalyzes the elimination of ammonia from triethanolamine in addition to ethanolamine r3’1.

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72 Hydrolysis and Formation of C-N Bonds

References 1 A. Liese, K. Seelbach, C. Wandrey, Industrial

Biotransformations, Wiley-VCH Verlag GmbH, Weinheim, 2000. 2 S. Murase, J. S. Takagi, Y. Higashi, H. Imaishi, N. Yumoto, M. Tokushige, Biochem. Biophys. Res. Commun. 1991, 177,414-419. 3 K. Hatakeyama, M. Goto, M. Terasawa, H. Yukawa, Fermentative Manufacture ofr-Aspartic Acid from Maleic Acid and Ammonia, Mitsubishi Chemical Industries Ltd., 1997, J P 103 13889. 4 M. Terasawa, S. Nara, H. Yamagata, H. Yugawa, Manufacture of D-Aspartic Acid and/or r-Malic Acid with Aspartase and Fumarase, Mitsubishi Petrochemical Co., 1991, JP06014787. 5 M. Gotoh, T.Nara, M. Terasawa, and H. Yukawa, Single Reactor Microbial Manufacture ofr-Alanine, Mitsubishi Petrochemical Co., 1990, EP 386476. 6 S. Imai, M. Takahashi, S. Fukatsu, Y. Ogawa, New Processfor the Production ofr2-Amino-4(hydroxymethyl-phospinyl) butyric Acid, Meiji Seika Kaisha, 1989, EP 367 145. 7 Y. Asano, Y. Kato, Biosci., Biotechnol., Biochem. 1994,58,223-224. 8 Y. Kato, Y. Asano, Arch.Microbiol. 1997, 168, 457-463. 9 U.Heywang, H. Schwartz, M. Casutt, Preparation ofN-protected dialkyl (2S,3S)3-ethylaspartates, Merck, 1990, DE 4007038. 10 M. M. Bear, V. Langlois, M. Masure, P. Guerin, Macromol. Symp. 1998, 132, 3 37-348. 1 1 T. Furuta, H. Takahashi, H. Shibasaki, Y. Kasuya, ]. Bid. Chem.,1992, 267, 12 600- 12605. 12 U.Roos, S. Mattern, H. Schrempf, C. Bormann, FEMS Microbiol. Lett. 1992, 97, 185-190. 13 J. D.Galpin, B. E. Ellis, M. E. Tanner,J. Am.

Chem. Soc. 1999,121,10840-10841.

14 N. Naito, R. Taneda, M. Koito, H. Ito, N.

Fukuhara, Manufacture of L-Phenylalanine with Ammonia Lyase, Mitsui Toatsu Chemicals, 1991, JP 051 68487. 15 N. Naito, D. Ura, M. Koito, N. Fukuhara. Separation of L-Phenylalaninefrom Cinnamic Acid, Mitsui Toatsu Chemicals, 1990, JP 04069370.

16 W. Liu, Synthesis ofoptically Active Dhenyl-

alanine Analogs using Rhodotorula graminis, Great Lakes Chemical Corp., 1997, US 598 1239. 17 J. S. Zhao, S. K. Yang, Huaxue Xuebao 1997, 55,196-201. 18 J . S. Zhao, J. Q. Cao, S. K. Yang, YaoxueXuebao 1995,30,466-470. 19 J. Zhao, S. Yang, Y. Jiang, Youji Huawue 1993, 13,486-489. 20 M. Yanaka, D.Ura, A. Takahashi, N. Fukuhara, Manufacture ofa-Substituted Alanines with L-PhenylalanineAmmonia-Lyase,Mitsui Toatsu Chemicals, 1992, J P 061 13870. 21 G. B. D’Cunha, V. Satyanarayan, P. M. Nair, Enzyme Microb.Techno1. 1996, 19,421-427. 22 G. B. D’Cunha, V. Satyanarayan, P. M. Nair, Enzyme Microb.Technol. 1994, 16, 318-322. 23 J. L. Coquoz, A. Buchala, J. P. Metraux, Plant Physiol. 1998, 117, 1095-1101. 24 J . Jemielity,M. Kanska, R. Kanski, Isot. Environ. Health Stud. 1998, 34, 335-339. 25 F. Foor, N. Morin, K. A. Bostian, Appl. Environ. Microbiol. 1993, 59, 3070-3075. 26 T. Katayama, H.Suzuki, T. Koyanagi, H. Kumagai, Appl. Environ. Microbiol. 2000, 66 4764-4771. 27 A. E. M. Hofmeister, S. Berger, W. Buckel, Eur.]. Biochem. 1992,205,743-749. 28 K. D. Schnackerz, C. H. Tai, R. K. W. Potsch, P. F. Cook,J. Bid. Chem. 1999,274, 36 935-36943. 29 M. J. Soto, P. van Dillewijn, J. Olivares, N. Toro, FEMS Microbiol. Lett. 1994, 119, 209-214. 30 M. I. Igeno, C. Gonzalez del Moral, F. J. Caballero, F. Castillo, FEMS Microbiol. Lett. 1993,114,333-337. 31 C. Tricot, V. Stalon, C. Legrain,]. Gen. Microbiol. 1991, 137, 2911-2918. 32 W. L. Muth, R. N. Costilow,]. Biol. Chem. 1974,249,7457-7462. 33 W. L. Muth, R. N. Costilow,]. Bid. Chem. 1974,249,7463-7467. 34 T. T. Harkins, C. B. Grissom, Science 1994, 263,958-966. 35 J . Retey, Angew. Chem. 1990, 102, 373-379. 36 L. P. Faust, B. M. Babior, Arch. Biochem. Biophys. 1992, 294, 50-54. 37 R. De Mot, I. Nagy, G. Schoofs, J. Vanderleyden, Can. J. Microbiol. 1994, 40,403-407. 38 D. M. Roof, J. R. Roth,J. Bacteriol. 1989, 171,3316-3323.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

12.7 Transaminations

12.7 Transaminations

J. David Rozzell and Andreas 5.Bomrnarius 12.7.1 Introduction

Given their critical role in biological systems, it is not surprising that numerous applications for amino acids have developed, particularly in the pharmaceutical industry. Fourteen of the twenty common proteinogenic L-amino acids are essential in human diets, which has led to the development of a significant market for these as components in intravenous feeding solutions. L-Glutamic acid is used as a flavor enhancer in foods with annual sales estimated at greater than one billion dollars. LLysine, D,L-methionine,and L-threonine have already become established as largevolume additives to animal feeds that require enrichment in certain deficient amino acids, and L-tryptophan is developing a similar application. L-Phenylalanineand Laspartic acid have very important markets as key components in the manufacture of the high-intensity sweetener aspartame. A competitive product in development, alitame, is synthesized from D-alanine. The importance of non-naturally occurring amino acids can be seen from the increasing number of pharmaceutical products that incorporate one or more such compounds as intermediates. Numerous chiral drug candidates are synthesized from various natural and non-natural amino acid building blocks and have been submitted for biological testing. Inevitably, applications for amino acids, both naturally-occuringand non-natural, will result from this activity. There are already numerous examples. The synthesis of two thrombin inhibitors, Tirofiban from Merck & Co. and Inogatran from Astra-Zeneca,is based on analogs of L-tyrosine and D-cyclohexylalanine, respectively. D-2-Aminoadipicacid is one of the amino acids found in the tripeptide that is converted biologically into the p-lactam nucleus, and its use as a precursor for producing semi-synthetic penicillins and cephalosporins has been suggested. The L-antipode is also a common component of combinatorial synthesis approaches that incorporate non-naturally occurring amino acids. Fluorine substitution is also becoming increasingly common in the preparation of peptide analogs. In particular, p-fluoro-L-phenylalanineis a good choice as a non-naturally occurring amino acid for such work because it is almost isosteric with L-phenylalanine, but contains a strongly electron withdrawing fluorine atom to modify its dipole moment. In particular, the non-naturally occurring amino acid r-tert-leucine has received significant attention due to several pharmaceutically active compounds into which it is incorporated[']. HIV-protease inhibitors developed by Novartis and Abbott are based on L-tert-leucine[ 2 , 1' . Roche has developed the anti-arthritic compound Ro 31-9790 based on its potent inhibition of collagenase14]and a key component in the synthesis of Ro 31-9790 is the methylamide of L-tert-leucine. Boehringer Ingelheim developed a series of compounds that inhibit the ribonucleotide reductase of Herpes

I

873

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12 Hydrolysis and Formation of C-N Bonds

simplex virus; several of the most active structures contained ~-tert-leucine[~]. As a result, a market for L-tert-leucine as a pharmaceutical intermediate is developing. In addition, the derivative L-tert-leucinolis widely used as a chiral auxiliary[']. The markets for non-naturally occurring amino acids can be substantial. Inhibitors of angiotensin-converting enzyme, or the so-called ACE inhibitors, have developed strong markets as anti-hypertensivedrugs. One of the most successful is the product Enalapril, which has achieved sales of more than $1 billion annually. Marketed by Merck, Sharpe and Dohme, Enalapril is due to go off patent soon, leading to the emergence of generic competitors. Other similar ACE inhibitors include Ramipril, Benazapril, Lisinopril, Zestril, Trandolopril, and Quinipril. A key acid, or Lcomponent in all of these compounds is L-4-phenyl-2-amino-n-butanoic homophenylalanine. As price competition for generic ACE inhibitors intensifies, Lhomophenylalanine will probably become an important non-naturally occurring amino acid product. Other large volume products include D-phenylglycine and D-phydroxyphenylglycine,key intermediates in the synthesis of ampicillin and amoxicillin, respectively, D-penicillamine, a chelator used to treat cystinuria and severe arthritis, D-valine, a building block for the synthetic pyrethroid Fenvalerate, and phosphinothricin, an important herbicide marketed by AgrEvo. Additional commercial opportunities exist for the production of isotopically labeled amino acids, particularly 15N, 15N/13C, and 15N/13C/2H amino acids for use in medical research, with a larger potential market in magnetic resonance imaging. Various methods have been developed for the production of amino acids. Most naturally-occurring, proteinogenic amino acids can be produced by fermentation, although chemical synthesis, isolation from hydrolyzed proteins, and enzymatic conversion are used in a few instances. For the production of non-proteinogenic or non-natural amino acids for which no metabolic pathways exist, traditional fermentation methods cannot be used without re-engineering of the metabolic pathways in the cell. For these types of amino acids, various chemical and enzymatic synthetic methods have become increasingly common. Among the various enzymes capable of producing optically-active amino acids, transamination reactions, catalyzed by enzymes known as aminotransferases or transaminases, have broad potential for the synthesis of a wide variety of enantiomerically pure (R)-and (S)-compounds containing amine groups. Indeed, various examples of the use of aminotransferases for the production of D- and L-amino acids, both naturally-occurring and non-natural, have been published "I.[' In addition, certain aminotransferases have been found to act on amines, and methods for the production of enantiomerically pure amines by transamination have been deThis method allows for yields of up to 100% whereas routes based on scribed[1G21]. hydrolases require external racemization to reach such yield levels. In this section we will focus on the application of aminotransferases.

12.7 Transaminations

12.7.2 Description of Transarninases 12.7.2.1

Homology and Evolutionary Subgroups o f Aminotransferases

About one third of all known sequences of vitamin BG-dependent enzymes belong to aminotransferases which in turn can be divided into four subgroups based on sequence homology: the most common species such as aspartate, tyrosine, or phenylalanine aminotransferase belong to subgroup I, subgroup I1 takes (acetyl)ornithine, o-amino acid and y-aminobutyrate aminotransferases, subgroup 111 comprises the D-amino acid transferases, and subgroup IV the (phospho)serine aminotransferases[221.Only 4 of the about 400 amino acid residues proved to be invariant among all aminotransferase sequences: Gly 197, Asp/Glu 222, Lys 258, and Arg 386. Apparently, aminotransferases form a group of homologous proteins, the chemistry of which already existed very early in evolution. 12.7.2.2 Mechanism of Transamination

Aminotransferases are key enzymes in a number of metabolic pathways, and as a result, enzymes from this class are widely distributed in nature. The first evidence for the presence of an enzyme catalyzing a transamination reaction was published by Needham and Szent-Gyorgyi and co-workerswho noticed a relationship between the L-glutamic acid, L-aspartic acid, and oxaloacetic acid levels in pigeon breast mus~le[~ Banga ~ ] . and Szent-Gyorgyi demonstrated the reversibility of glutamic-pyruvic transaminase (E. C. 2.6.1.2, alanine aminotransferase) by chemically isolating the that time, a large amino acid products L-glutamate and ~ - a l a n i n e [251.~ ~Since . number of different aminotransferases have been discovered and characterized, including aminotransferases, capable of catalyzing the transamination of all naturally-occurring amino acids. There are now more than 2500 sequences of aminoAs of the middle of transferases known, compared with 51 sequences in 1993L221. February 2001, the Entrez databank contained 121 3D-structures of 9 aminotransferases from 13 organisms. The mechanism of the reaction is well understood as a result of the detailed studies of Meister [26,271. Aminotransferases catalyze the transfer or an amino group from an amino acid donor to a 2-ketoacid acceptor (Fig. 12.7-1).This amino group transfer is mediated by the cofactor pyridoxal phosphate, which is reversibly bound to the enzyme through a Schiff-baselinkage to the epsilon-amino group of an activesite lysine. Mechanistically, the reaction catalyzed by an aminotransferase can be thought of as the result of two discrete steps. The first step is the transfer of an amino group from the amino group donor to pyridoxal phosphate, generating a 2-ketoacid byproduct that dissociates from the enzyme and an enzyme-bound pyridoxamine phosphate intermediate. The second step involves the transfer of the amino group from the enzyme-bound pyridoxamine phosphate to the 2-ketoacid acceptor, and

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I

72 Hydrolysis and Formation ofC-N Bonds R1\

,COOH C

R’\

0

H,N

II

2-Ketoacid Acceptor

H,N

rr OCOOH

2%H

General reaction catalyzed by arninotransferases. Figure 12.7-1.

L-Amino acid

4%

H

L-Amino acid Donor

2-Ketoacid Coproduct

4%

H 2-Ketoacid Acceptor

+

D-Amino acid Donor

NH,

D-Amino acid

-

L

2-Ketoacid Coproduct

producing the corresponding amino acid product and regenerating the pyridoxal phosphate cofactor for another catalyhc cycle. As a result, aminotransferases characteristically exhibit ping-pong kinetics r2’]. 12.7.2.3

Protein Engineering and Directed Evolution with Aminotransferases

Aminotransferase (AAT), the enzyme catalyzing the reversible transformation of aspartate and glutamate into the respective 0x0 acids, has been studied most among the vitamin B6-dependentenzymes. An X-ray crystal structure is now known for the aspartic-glutamic aminotransferase from E. coli r2’1. Active site residues have been identified, laying the groundwork for further detailed mechanistic studies and modification of the enzyme by specific mutagenesis. Several workers have been successful at changing the relative activity of aminotransferase towards different groups of substrates or even different reactions through structure-based protein engineering and directed evolution.

12.7 Transaminations

12.7.2.3.1

I

877

Structure-based Protein Engineering

Multiple active-site site-specific mutations of AAT led to an increase in P-decarboxylase activity with the double mutant Y225R/R386A (1380-fold)f3O]. Coupled with a decreased transaminase activity by a factor of 500 in the single mutant R292KL3lI, workers found a combined 20 000-fold decrease in the rate of transamination in the triple mutant Y225R/R292K/R386A [321. In fact, the triple mutant catalyzed pdecarboxylation %fold faster than transamination, a change of ratio from the wildtype enzyme by a factor of 25 million. The observed changes in substrate specificity were rarely additive however, because triple mutants containing R292X, i. e. mutations to amino acids other than lysine, were mostly completely inactive towards pdecarboxylation even though they contained the double mutant Y225RJR386A eliciting p-decarboxylase activity. Previously, AAT had been transformed into an L-tyrosine aminotransferase (TAT) by site-specificmutation of up to six amino acid residues lining the active site of wildtype TAT. The hexuple AAT-mutant achieved kinetic data towards the transamination of aromatic substrates such as L-phenylalaninewithin an order of magnitude of wildtype TAT[33].

12.7.2.3.2

Directed Evolution o f Aminotransferases

Meanwhile, directed evolution methods that combine mutagenesis of genes with high-throughput screening of functional gene products have developed rapidly. In a selection strategy based on the substrate 1-phenyl-n-propylamine(PPA)as the sole source of nitrogen in a chemostat, a recombinant Pseudornonas putida strain carrying the R-transaminase gene, a single amino acid change, Y112F, presumably at or near the active site, improved enantioselectivity of the reaction of racemic and propiophenone to 37.8 % 1-phenyl-n-propylamineto (S)-1-phenyl-n-propylamine e. e. from 6.5 % e. e. in the wildty~e[~’]. Further site-directedmutagenesis of position 112 yielded 99.4% e.e. in the mutant Y112L. In a related example, a single mutant T51S, generated by error-prone PCR in about 10000 samples, both improved tolerance of (R)-transaminasetowards the reaction product, a substituted I-phenyl2-propylamine (an amphetamine), from 85 to 105 mM as well as reaction rate[”I. Lastly, a p-tetralone was converted into the corresponding (S)-aminein 65 % e. e. by Random mutation followed by activity wild-type (S)-transaminase (Fig. 12.7-2)[‘’I. screening for the colored ketone starting from the enantiomerically pure amine, produced a number of single mutants such as M245V, P247L, and F407L with higher enantioselectivity,up to 84% e. e., at similar level of activity. It was further found that combination of advantageous mutants through site-directed mutagenesis around

+wo

*-(S)-Transaminase

2-aminobutane Figure 12.7-2.

.-mNH

methylethylketone

Conversion oftetralone-2 to 2-arninotetraline by (S)-transarninase.

878

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12 Hydrolysis and Formation ofC-N Bonds

sensitive sites such as 245-247 and 405-407 improved enantioselectivity further, up to 94%. Efforts to evolve aminotransferases with improved activity on new ketoacid substrates have been initiated with encouraging results [341. Using directed evolution, the substrate specificityof AAT has been changed to one favoring P-branched amino acids and their respective oxoacids, effectively converting AAT into a branched-chain aminotransferase (BCAT). By employing an E. coli auxotroph deficient in the branched-chain aminotransferase (BCAT) gene, ilvE, the authors set up a stringent selection system which provided a powerful advantage for cell growth to the mutated AAT systems[35,3Gl. The resulting evolved aminotransferases had 13 [351 and 1713'] amino acid substitutions and showed 10S-fold and 2x10G-foldimprovement in catalytic efficiency (kcat/&), respectively, towards the unnatural substrate, valine, and between 10- and 100-fold decrease towards the natural substrate, L-aspartate, compared with the wild-type. A high degree of conserved amino acid substitutions was found in most active mutants. Interestingly, only one mutated amino acid residue in each case is located at a distance to the substrate that would allow interactions, the remainder were mutated far away from the active site. This work demonstrates that 1OG-foldshifts in substrate specificity can be achieved when employing directed evolution methods, that combinatorial or evolutionary methods are probably superior to rational design methods when changing substrate specificity, and most importantly, that remote residues and their interactions with the active site environment are important determinants of enzyme activity and specificity. Such remote residues act cumulatively, possibly by remodelling the active site, by altering the subunit interfaces, or by shifting different enzyme domains. 12.7.3 Use o f Arninotransferases in Biocatalytic Reactions 12.7.3.1

Synthesis o f a-L-Amino Acids

Aminotransferases (transaminases) have been studied as potentially useful biocatalysts for the production of a wide range of different amino acids. The general reaction catalyzed by aminotransferases is shown in Fig. 12.7-1.An amino group is transferred from a donor amino acid to a 2-keto acid acceptor. As described earlier, a cofactor, most commonly pyridoxal phosphate, is involved in the catalysis. The cofactor, which is only required in concentrations of 50-100 PM,is reversibly bound to the enzyme through a Schiff-baselinkage to the epsilon-amino group of active-site lysine[2628].Using an aminotransferase, a desired amino acid can be produced from a given 2-keto acid precursor using an inexpensive L-amino acid as the amino group donor. As a co-product of the reaction, a second 2-keto acid corresponding to the amino acid donor is produced along with the desired amino acid product in equimolar amounts. Among the advantages of transaminases as biocatalysts for the production of optically pure amino acids are as follows:

12.7 Transaminations

Aminotransferases have high stereoselectivity for a given enantiomer. Optically active L- or D-amino acids are produced stereoselectively; the process is a chiral synthesis, not a resolution. The catalytic rates of these enzyme-catalyzed reactions are generally relatively rapid. Capital costs for such a biocatalytic process are low; in contrast to the situation with fermentations, already existing chemical process equipment can be used for performing the enzyme-catalyzed reaction. A large number of the required 2-keto acid precursors are accessible through chemical synthesis, expanding the range of potential products. Aminotransferases are potentially applicable to the production of a wide range of amino acids, because enzymes are available for D- and L-amino acids. In addition, a wide range of aminotransferases with side-chain specificity are known, including enzymes for the production of amino acids with aromatic side chains, acidic side chains, branched alkyl side chains, etc. In some cases, the 2-keto acid by-products may also have significant value. For example, important markets exist for pyruvic acid, 2-ketoglutaric acid, and other similar compounds. One of the simplest examples of an efficient transamination process is the production of L-alanine and 2-ketoglutarate from the precursors L-glutamate and pyruvic acid (Fig. 12.7-3). Porcine glutamic-pyruvic transaminase is available commercially, and this enzyme was used as a model system for studying the transamination on a preparative scale. The equilibrium constant was measured for this reaction and found to be 1.86, slightly favoring the formation of r-alanine and 2-ketoglutarate. Glutamic-pyruvic transaminase was immobilized on porous aminopropyl glass using water-soluble carbodiimide as a coupling agentL71. At a loading of 20 mg of total protein bound per gram of glass, the activity of the biocatalyst when assayed or the production of L-alanine was 400 units per gram of biocatalyst. The enzymatic activity retained after immobilization was 40%, and the immobilized enzyme was used for the continuous production of L-alanine and 2-ketoglutarate from pyruvate

Pyruvic acid

+

L-Glutamic acid Figure 12.7-3.

L-Alanine

Glutamate-pyruvate aminotransferase

-

L

+

2-Ketoglutarate

Transamination using glutarnic-pyruvic arninotransferase.

I

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880

I

12 Hydrolysis and Formation ofC-N Bonds

@

+ L-alanine

-

(S)-Transaminase

acetophenone Figure 12.7-4.

-

@

+ pyruvate

(S)-phenylethylamine Enantiomerically pure (S)-amines via w-transaminases.

and L-glutamate over a three-month period with less than 40% loss in activity. Volumetric productivity was 200 gL-lh-l of L-alanine. 12.7.3.2

Synthesis of Enantiomerically Pure Amines

While most methods for the synthesis of enantiomerically pure amines have employed kinetic resolution with the help of lipases or esterases, a method independent of kinetic resolution has been developed using the transamination of ketones catalyzed by o-transaminases (w-TA), shown in Fig. 12.7-4 with acetophenone as an example[20*211. The w-transaminases can be employed in two ways to produce both enantiomers in a pure form [l81: a racemic mixture can be separated, by kinetic resolution, into the corresponding ketone and the remaining amine enantiomer, which is typically obtained in high enantiomeric excess, the ketone can be recycled as a starting material for the racemic amine; the same o-transaminase can be employed to synthesize the enantiomer of the opposite configuration straight from the ketone. Both ( R ) -and (S)-aminotransferasehave been employed at Celgene for the synthesis of enantiomericallypure amines from racemic amines. Degrees of conversion were at or close to 50 % for resolutions and enantioselectivitiesnormaly reached > 99 % e. e. for the amine product from both resolutions or syntheses from ketones [18, 19]. The donor for resolutions of amine racemates was usually pyruvate whereas either isopropylamine or 2-aminobutane served as donors for reduction of ketones. The amine products ranged from phenylethylamine and tetramines with the amine group at activated benzylic sites or in a cyclic structure, to phenylisoproylamines (amphetamines) or phenoxyisopropylamines where the amine group is hardly or not activated at all. A selection of products synthesized with o-aminotransferase technology is shown in Fig. 12.7-5.The Celgene process has been scaled up to the 500 kg level [I9]. The (S)-w-TA from Vibrio fluvialis was found to catalyze the reduction of acetophenone to (S)-a-methylbenzylaminewith the concomitant oxidation of L-alanine to pyruvate. The enantiomeric excess was always > 99% e.e. As thermodynamic equilibrium strongly favors the reverse reaction, however, high yields were achieved only when an excess of acetophenone was added and upon removal of pyruvate:

&

+WOH

/

MeO

R

RO

i"'

/

I

72.7 Transaminations 881

UOJ

R=H.Me

oy

/

NHz

I

R = H, Me. CI, Br, NOp

List o f amines produced by o-arninotransferase technology (both enantiorners produced i n each case).

Figure 12.7-5.

yields of z 90 % were achieved with acetophenone and benzylacetone in the presence of a 10-fold excess of L-alanine if pyruvate was removed by using whole cells. The reaction suffers from strong inhibition by both products, pyruvate and (S)-amethylbenzylamineI2O]. Interestingly,the authors found a linear correlation between the reactivities of amino acceptors and the inverse reactivity of amino donors[2']. 12.7.3.3 Other Preparative Applications o f Arninotransferases 12.7.3.3.1

Preparative Applications: L-Phosphinothricin

L-Phosphinothricin, the active ingredient of the broad-spectrum herbicide Basta (AgrEvo),can be obtained through enzymatic transamination of the corresponding oxoacid, 2-0x0-4-[(hydroxy)(methyl)phosphinoyl]butyric acid, in a coupled system with aspartate aminotransferase (AAT) and 4-aminobutyrate:2-ketoglutaratetransaminase (E.C. 2.6.1.19) from E. coli (Fig. 12.7-G)[371.In solutions containing 10% substrate, 85 % conversion was reached with only i3 % amino acid by-products. For

882 pyruvate + CO

I

12 Hydrolysis and Formation ofC-N Bonds

t

L-phosphinothricin

oxaloacetate glutarnateoxaloacetate transarninase (GOT)

P-ketoglutarate4-arninobutyfate transaminas6

L-AsP glutarate

Figure 12.7-6. Coupled process for the herbicide ingredient L-phosphinothricin with transaminases.

this process, a new AAT from B. stearothermophilus has been screened and characterized (Topt= 95 "C, pH,,, = 8.0) before being cloned and overexpressed in E. coli.

12.7.3.3.2

Synthesis of an Omapatrilat Building Block with L-Lysine

E-am hotransferase &-0x0-L-norleucineacetal is a key intermediate for the synthesis of Omapatrilat (BMS-186716), a novel dual-action vasopeptidase inhibitor under development at Bristol-Myers-Squibb(BMS). The BMS researchers developed a novel synthesis of a key building block of Omapatrilat, the bicyclic compound BMS-199541-01, by oxidation of the &-groupof L-lysine in the N-protected dipeptide N-Cbz-L-homo-cys-Llys with a newly found L-lysine ~-aminotransferase[~~]. The enzyme was isolated from Sphingornonas paucimobilis and was cloned and overexpressed in E. coli (2 kUL-I). It is an 81 kDa homodimer with a specific activity towards the product BMS199541-01 of 1.68 Umg-' of protein; the enzyme requires a-ketoglutarate as a cosubstrate which is recycled back into the process after oxidation of the L-glutamate back to a-ketoglutarate by glutamate oxidase (isolated from Streptomyces noursei). LLysine E-aminotransferasewas found to be one of the most important rate-limiting enzymes in cephalosporin biosynthesis 13'1. The process scheme (Fig. 12.7-7) starts from the N-protected dipeptide dimer [Llys-~-homocys]~ disulfide which, after reduction of the S - S bond, is oxidized enzymatically to N-Cbz-L-homo-cys-L-lys-&-aldehyde. Under acidic conditions, the aldehyde group is present as a gem-diol, attacks the a-N and closes the ring to the aminol. After nucleophilic attack of the S - H group, the hydroxyl group acts as a leaving group and affords closure of the 1,3-thiazepinering. In the biotransformation process to BMS-199541-01, yields of 65-70 mole-%were achieved without recycling of the L-glutamate resulting from the reduction of aketoglutarate yields were substantially lower. L-Lysine &-aminotransferasealso catalyzes the oxidation of N-a-protectedL-lysines as well as L-lysine peptides such as Nprotected L-met-L-lys.

72.7 Transaminations

I

883

Dithiothreitolor Tributylphosphine

c

SH

Dipeptide Monomer

a-keto glutarate Dipeptide Dimer BMS-201391-01

L-Lysine-E-aminotransferase

Glutamatt Oxidase

from Sphingomonas paucimobilis or rec E. coli L-Glutamate

PGHN Acid

SH

cb-

PGH%

(protecting group)

BMS-199541-01 Figure 12.7-7. BMS process to the bicyclic intermediate BMS-199541-01 via L-lysine ~-aminotransferase[~~’.

884

I

72 Hydrolysis and Formation ofC-N Bonds

12.7.4 Driving the Reaction to Completion

There is one major disadvantage to most of the transamination technology as presented above: because the transamination reaction involves an amino acid reacting with a 2-keto acid to generate products which consist of a 2-keto acid and an amino acid, the equilibrium constant is often close to unity. As a result, the net conversion of substrates to products is thermodynamically limited. The key to the development of an efficient transamination technology lies in overcoming the problem of incomplete conversion of the 2-keto acid precursor to the desired amino acid product. One approach to this problem is the coupling of the transamination reaction to a second reaction that consumes the keto acid by product in an essentially irreversible step; this drives the transamination reaction to completion. By using an aminotransferase that can utilize aspartic acid efficiently as the amino group donor (instead of glutamic acid), the corresponding 2-keto acid by product is oxaloacetate (rather than 2-ketoglutarate).Oxaloacetate is a P-ketoacid and can be easily decarboxylated to pyruvate. This decarboxylation occurs spontaneously in aqueous solution, catalyzed

RKcooH 0

H,N

,COOH C

4%

H

L-Amino acid

2-Ketoacid

-

+

R,

L

+

H O O C VCOOH

0 Oxaloacetic acid

L-Aspartic acid

Pyruvic acid

Acetolactate synthase (ALS)

\R-

- co, 3---

0

acetoin Figure 12.7-8.

Driving the transamination reaction to completion.

12.7 Transaminations

I

Time course for the transamination o f phenylpyruvate t o L-phenylalanine i n the presence and absence o f oxaloacetate decarboxylase.

Table 12.7-1.

Reaction time (min) 0 10 20 45 80

Transaminase alone Phenylpyruvate (mM)

Transaminase & Oxaloacetate decarboxylase Phenylpyruvate(mM)

200 184

200 140

166

116

124

44 6

116

by various metal ions and amines, and can be accelerated chemically, as shown in Fig. 12.7-8, or enzymatically using the enzyme oxaloacetate decarboxylase. The important feature of the process is that the essentially irreversible decarboxylationof oxaloacetate to pyruvate drives the entire process to completion, allowing the transamination of 2-keto acids to amino acids in yields approaching 100% of the 12, 131. I mportantly, this method of driving the reaction to completion may be used for the production of either D-amino acids or L-amino acids. The decarboxylation reaction catalyzed by the enzyme oxaloacetate decarboxylase has been examined using enzymes from four different sources: Pseudornonas putida, Micrococcus luteus, and two strains of Azotobacter vinelandii. The highest rates were obtained with the oxaloacetate decarboxylase isolated from Pseudornonas, a Mg2+requiring enzyme[7. l21. The effectiveness of decarboxylation in driving the reaction to completion was demonstrated in a coupled enzymatic process by using phenylpymvate as the starting 2-keto acid. In this experiment, phenylpyruvate sodium salt and L-aspartate were incubated with E. coli broad-range transaminase at room temperature and pH 7.5 in both the presence and absence of oxaloacetate decarboxylase from Ps. putida. Magnesium ion, which is cofactor for the decarboxylase, was also present in both reaction mixtures at a concentration of G mM. The transamination reaction was monitored by following the disappearance of phenylpyruvate. The results are summarized in Table 12.7-1. As demonstrated by the data, when oxaloacetate decarboxylase was included in the mixture the reaction proceeded to completion much more rapidly than in the case when the decarboxylase was omitted [I2]. Other methods can also be used for driving the transamination reaction to produce amino acids in high yields. For example, if L-lysine or L-ornithineare used as the donor in the two-enzyme process shown in Fig. 12.7-9, the cyclization of the aldehyde is strongly favored, creating an essentially irreversible reaction that can lead to high yields of a desired amino acid from the corresponding 2-ketoacid[lO,40, 411 8'

12.7.5 Production of L-Amino Acids Using Immobilized Transaminases

Continuous decarboxylationof oxaloacetate as it is formed is an important part of an efficient, high-yielding transamination process. This decarboxylation occurs readily

885

886

I

72 Hydrolysis and Formation ofC-N Bonds

HOOcTcOO HooC7fCooH 0

NH,

2-Ketoglutarate

L-Glutamic acid

+

-

Transaminase

-

+ R,

RKcooH 0

HN ,

,COOH

4%

H

HOOcTcO Hooc7fcooH NH,

L-Glutamic acid

0

-

2-Ketoglutarate

Lysine

+

Aminotransferase

H L-Lysine

Cyclized lmine Coproduct

Figure 12.7-9. Coupled reactions using L-lysine for driving the transamination of 2-ketoacids to amino acids.

in aqueous solution, and may be accelerated enzymatically as described above, or chemically using a metal ion such as Mg2*in sufficient concentration. Immobilization of the enzyme allows reuse of the enzyme or continuous production of amino acid in a flow reactor system. Immobilization of the E. coli broad-range transaminase has been accomplishedby covalent attachment using glutaraldehyde PVC-silica support matrix that had been activated with a polyamine[13].In the example described in Table 12.7-1,4 L of cell lysate containing 61.6 g of enzyme (activity of 5.2 million international units) were clarified by centrifugation at 13 000 g for 30 min and recirculated through a preactivated support matrix for 1.5 h. After washing, 57 g or 93% of the enzyme remained bound to the support. Bound activity was 4.2 million units. The retained activity of the enzyme after immobilization was approximately 89 %. The pH-rate profile for the reaction catalyzed by the E. coli broad-range transaminase was determined using the immobilized transaminase with p-fluorophe-

I

12.7 Transaminations 887

Concentrations of reactants for the production of L-p-fluorophenylalanineby transamination ofp-fluorophenylpyruvate. Table 12.7-2.

Reactant

Concentration (mM)

Sodium p-fluorophenylpyruvate r-Aspartate Pyridoxal phosphate MgCl2

100 110 0.1

50

nylpyruvate as the keto acid and L-aspartate as the amino group donor. The transamination reaction displayed a fairly broad useful pH range; the immobilized transaminase had a pH optimum of approximately 7.5, but retained activity in the range of pH 6.0-9.5. At pH 5.0 and 10.0, activity fell to less than 20% of that measured at pH 7. For continuous production of L-p-fluorophenylalanine,a typical set of operating conditions is shown in Table 12.7-2.L-Aspartate is used at a 10% molar excess to the starting 2-ketoacid. The cofactor pyridoxal phosphate is added to the reaction mixture to achieve a final concentration of 0.1 mM. The initial pH of the feed solution is 7.2. Mg2+ion was used to accelerate the decarboxylationof oxaloacetate to pyruvate. The reaction was maintained with a temperature range of 37-40 "C. Under these conditions using an immobilized broad-range aminotransferase, the volumetric productivity of the reactor for the production of L-phenylalanine at 85% conversion was 20 gL-lh-'. One of the main advantages of the transamination system is its applicability to a range of other L-amino acids, including non-naturally occurring amino acids. For example, broad-range aminotransferase (encoded by the aspC gene) will efficiently transaminate the 2-keto acids corresponding to L-phenylalanine,p-fluoro-L-phenylL-tryptophan,L-methionine,~-1iorrioalanine, L-tyrosine, rn-hydroxy-L-phenylalanine, phenylalanine, L-2-aminoadipicacid and a number of others. Using other aminotransferases, the transamination of other 2-ketoacids to the corresponding amino Table 12.7-3.

Amino acids produced by transamination

Amino Acid

Aminotransferase

r-Phenylalanine r-Tyrosine L-Tryptophan L-p-fluorophenylalanine L-meta-tyrosine r-Homophenylalanine L-2-Aminoadipicacid L-2-Aminopimelicacid r-Valine r-Leucine r-tert-leucine D-Alanine D-Leucine D-Tyrosine D- Phenylalanine

Broad-range, aromatic Broad-range, aromatic Broad-range, aromatic Broad-range Broad-range Broad-range, aromatic Broad-range Broad-range Branched-chain Branched-chain Branched-chain D-broad-range D-broad-range D-broad-range D-broad-range

888

I

12 Hydrolysis and Formation ofC-N Bonds

HOOc-fcOOH Hooc7fCooH NH,

0

2-Ketoglutarate

L-Glutamic acid

-

+

Branched-chain L

Transarninase

H3C. 22,I CH,

,COOH C

II

0

Trirnethylpyruvate

L-tert-Leucine

HOOc-TfcOOH HOOcTc 0

NHZ

2-Ketoglutarate

L-Glutarnic acid Broad-Range

+

+

L I

Transarninase HOOC/YcooH

co, COOH H O O C V

0

NHZ L-Aspartic acid

Net: L-Aspartate + Trirnethylpyruvate Figure 12.7-10.

0

Oxaloacetic acid

-

H3c7fC02H Pyruvic acid

L-tert-Leucine+ Pyruvate + CO,

Coupled arninotransferases for the production o f L-tert-leucine.

acids can be carried out. A list of amino acids that have been produced by transamination is shown in Table 12.7-3. The broad-range aminotransferase has low catalytic activity for the group of branched-chain amino acids, including L-leucine, L-isoleucine and L-valine. To enable production of this group of L-amino acids, another transaminase, the socalled branched-chain amino acid transaminase (BCAT), has been used. This enzyme has also been shown to catalyze the transamination of trimethylpyruvate to produce the commercially interesting unnatural amino acid L-tert-leucine,although the rate of the reaction is significantly less than that for L-valine. Unlike the broadrange transaminase, the branched-chain aminotransferase is not active with L-

I

12.7 Transaminations 889

aspartate as the amino donor. L-Glutamate is used for efficient transamination using this enzyme. To drive this reaction, a coupled transamination reaction was established with both the broad-range and branched-chain aminotransferases acting together as shown in Fig. 12.7-8 for the production of L-tert-leucine.In the first reaction, the branchedchain aminotransferase catalyzes the reaction of L-glutamate with trimethylpyruvate to produce L-tert-leucineand 2-ketoglutarate.The second reaction catalyzed by broadrange aminotransferase converts L-aspartate and 2-ketoglutarate into oxaloacetate and L-glutamate.The donor L-aspartate is present in stoichiometric amounts relative to 2-ketoisovalerateand is used to continuously recycle the 2-ketoglutarateformed in the first step to L-glutamate as the reaction proceeds. Oxaloacetate is decarboxylated to pyruvate in an essentially irreversible reaction, driving the entire sequence of reactions to completion. The net reaction is the transamination of trimethylpyruvate to L-tert-leucine with L-aspartate using 2-ketoglutarate as an intermediary amino transfer agent. This sequence of reactions has also been used to produce L-leucine and L-valine in the laboratory (Fig. 12.7-10). In laboratory-scaleexperiments, solutions containing 200-GOO mM keto acid were transaminated to the corresponding branched-chain L-amino acid, with a concentration of L-glutamate between 50 mM and 100 mM and a 1.1 molar excess of Laspartate. Yields obtained for the branched-chain amino acids have typically been in the range of 80-90% based on starting with a 2-keto acid['']. Another example of a coupled enzyme reaction demonstrates the versatility of the transaminase system in biocatalysis. Using a racemic D,L-amino acid mixture as the starting material, the enzyme D-amino acid oxidase from Trigonopsis variabilis will convert the D-amino acid in the mixture selectively into the corresponding 2-keto acid. The 1-amino acid of the D,L- pair is neither a substrate nor an inhibitor of Damino acid oxidase. If a transaminase is present in the same reaction mixture, the 2-keto acid can be transaminated in the presence of L-aspartate to the corresponding L-amino acid. The entire reaction can be driven to completion as described previously by decarboxylation of the oxaloacetate. Thus, in a single pot, racemic D,Lamino acids can be convened directly into optically active L-amino acids (Fig. 12.711). 12.7.6

D-Amino Acid Transferases

The aminotransferase reaction can be utilized for the synthesis of D-amino acids as well as the better-known route to L-amino acids (Fig. 12.7-11).Regarding sequence similarity, D-aminotransferases form a distinct subgroup among the transferases, however, it has been found, with the help of crystal structures[42-431that some striking similarities exist between L-amino acid aminotransferases with respect to active site structure and to branched-chain aminotransferase (BCAT) with respect to sequence. D-Aminotransferases utilize the same PLP chemistry as L-aminotransferases to effect tran~amination[~~l. Mutagenesis of a distant interdomain loop of Darninotransferase to produce enhanced conformational flexibility (proll9-argl20-

890

I

12 Hydrolysis and Formation of C-N Bonds

RYcooH

RYCooH

NH,

NH,

L-Amino acid

+

D-Amino Acid Oxidase

Figure 12.7-11. Conversion of racemic amino acids into L-amino acids with D-amino acid oxidase and an L-aminotransferase.

+

m

Racemic Amino acid

2-Ketoacid

RKCooH 0

,R

H,N

,COOH

4%

H

L-Amino acid

+ L-Aspartic acid

L-Transaminase

m

+ Pyruvic acid

+ CO,

pro121 to gly-gly-gly)resulted in higher catalytic constants towards most D-amino acid substrates [&I. D-Amino acids can also be produced directly by transamination using a Daminotransferase. Since these enzymes require a D-amino acid donor, we developed a coupled enzymatic reaction with aspartate racemase to generate D-aspartic acid in situ from inexpensive L-aspartic acid. The reaction scheme is shown in Fig. 12.7-12. Aspartate racemase, cloned from Streptococcus thermophilus and expressed in E. coli, is used in conjunction with a D-aminotransferaseto produce D-amino acids from corresponding 2-ketoacidsin a reaction that is analogous to that for the production of L-amino acids. Oxaloacetate,produced from D-aspartateduring the transamination, is decarboxylated to pyruvate, driving the reaction to completion as with the Ltransamination. Significant amounts of D-alanine are produced using the D-aminotransferase cloned from Pseudomonas sphaericus ATCC 10208 as it has activity toward pyruvate. Directed evolution efforts are in progress to develop an enzyme having reduced D-alanine production, resulting in a cleaner product mixture. For the synthesis of D-glutamate,a two-enzyme system consisting of of glutamate racemase and D-aminotransferasehas been found in B. s p h a e r i c u ~ [ ~ ~ ] .

I

12.7 Transaminations 891

R

RIYcooH -0

+

~ Figure ~ 12.7-12.~

Production ~ of D-amino acids by transamination

GH,

2-Ketoacid Acceptor

D-Amino acid

R2Yc02H 0

NH, D-Amino acid Donor

2-Ketoacid Coproduct

12.7.7

Synthesis of Labeled Amino Acids

Isotopically labelled amino acids are particularly amenable to production by transamination. Because the reaction catalyzed by aminotransferases transfers a specific amino group from the donor, amino acids highly enriched in isotopes such as 15N can be produced. For example, 15N L-tyrosinehas been produced in greater than 90 % yield from "N L-aspartate and p-hydroxyphenylpyruvate using the broad-range aminotransferase from E. coli. The reaction is shown schematically in Fig. 12.7-13. Analysis of "N-isotope incorporation was carried out by mass spectrometry by Cambridge Isotope Laboratories. The samples showed incorporation of 98.4 % "N, which was almost identical to the isotopic purity of the starting L-aspartic acid. Only the L-isomer of tyrosine was detectable by chiral HPLC. This result established the feasibility of the production of 15N amino acids by transamination by meeting three important criteria for success: there was no detectable loss of isotopic purity in the transfer of the amino group from 15N aspartic acid to the 2-ketoacid,

mooH

COOH

HO

HO

p-Hydroxyphenylpyruvic acid

+ A ,COOH

HOOC

c ,3H 15NH,

15NL-Aspartic acid Figure 12.7-13.

15N L-Tyrosine Transaminase

*

+

H3CKC00H 0

Pyruvic acid

Production of "N-labeled amino acids by transamination.

+

co2

892

I

12 Hydrolysis and Formation of C-N Bonds

the stereochemical fidelity of the transamination reaction was perfect within detection limits, and The yield of conversion of the 15N aspartic acid (the most costly starting material in this reaction) was high (> 90%). 12.7.8

Availability of Enzyme

In order to facilitate the production of adequate amounts of transaminase at low cost, the genes encoding aminotransferases have been cloned and overexpressedin E. coli. Two examples are the aspC and ilvE genes from E. coli. The expression of these genes has been described previously, with the levels of aminotransferase enzyme reaching approximately 30-40% of the total cell protein['*! More recently, the genes encoding other aminotransferase genes have been c l ~ n e d [leading ~ ~ ~to~the ~ ,availability of a broader group of aminotransferase enzymes for evaluation. Given the high reaction rates observed and the potential for wide applicability for the production of amino acids, both D and L, natural or unnatural, transamination reactions should prove to be useful method for the chemist.

References

For a review on tert-leucine, see: A. S. Bommarius, M. Schwarm, K. Stingl, M. Kottenhahn, K. Huthmacher, K. Drauz, Tetrahedron: Asymmetry 1995, 6, 2851-2888. 2 P. Ettmayer, M. Hubner, A. Billich, B. Rosenwirth, H. Gstach, Bioorg. Med. Chem. Lett. 1994, 4 , 2851-2856. 3 D. J. Kempf, L. M. Codacovi, D. W. Norbeck, J. J. Plattner, H. Sham, S. J. Wittenberger, C. Zhao, 1992, Eur. Patent Applic. EP 486948. 4 P. A. Brown, W. H. Johnson, G. Lawton, 1992, Eur. Patent Applic. EP 0497192. 5 R. Deziel, N. Moss, R. Plante, 1993, Eur. Patent Applic. EP 0560274. 6 N. J. Turner, J. R. Winterman, R. McCague, J. S. Parratt, S. J. C. Taylor, Tetrahedron Lett. 1995,36,1113-1116. 7 J. D. Rozzell, Methods Enzymol. 1987, 236, 479-497. 8 J. D. Rozzell, Production of Amino Acids by Transamination, 1985, U . S . Patent 4,518,692. 9 J. D. Rozzell, Production of ~-4-Phenyl-2-Aminobutanoic Acid by Transamination, 1985, U. S . Patent 4,525,454. 10 J. D. Rozzell, Alpha Amino Acidsfrom Alpha Ketoacids Using Coupled Transaminase Enzymes, 1989, U. S . Patent 4,518,692. 1

J. D. Rozzell, Production of Amino Acids Using Coupled Enzyme Systems, 1989, U . S. Patent 4,880,738. 12 S. P. Crump, J. S. Heier, J. David Rozzell in: Biocatalysis (Ed.: D. A. Abramowicz), Van Nostrand Reinhold, New York, 1990, pp. 155-133. 13 S. P. Crump, J. David Rozzell in: Biocatalytic Production of Amino Acids and Derivatives: New Developments and Process Considerations (Eds.: J. D. Rozzell, F. Wagner), Hanser Publishers, Munich, 1992, pp. 43-58. 14 D. J. Ager, S. C. Laneman, I. G. Fotheringham, P. P. Taylor, D. P. Pantaleone, Proc. Chiral Europe '97, 1997, 33-36. 15 D. P. Pantaleone, P. P. Taylor, R. F. Senkpeil, I . G. Fotheringham, TIBTECH 1998, 16(10), 412-418. 16 D. I. Stirling, A. L. Zeitlin, and G. W. Matcham, Enantiomeric Enrichment and S Stereoselective Synthesis of Chiral Amines, 1990, U . S. Patent 4,950,606. 17 D. I. Stirling, A. L. Zeitlin, G. W. Matcham, J. D. Rozzell, Jr. Enantiomeric Enrichment and Stereoselective Synthesis of Chiral Amines, 1992, U. S. Patent 5,169,780. 18 D. I. Stirling in: Chirality in Industry (Eds.: A. N. Collins, G. N. Sheldrake, J. Crosby), 11

References I893 Wiley, New York, 1992, Chap. 9,209222. 19 G. W. Matcham, A. R. St. G. Bowen, Chimica Oggi 1996 (6), 20-24. 20 J . 4 . Shin, B.-G. Kim, Biotech. Bioeng. 1998, 60(5), 534-540. 21 J.-S. Shin, B.-G. Kim, Biotechnol. Bioeng. 1999,65,206-211. 22 P. K. Mehta, T. I. Hale, P. Christen, Eur. ]. Biochem. 1993, 214(2), 549-61. 23 D. M. Needham, Biochem. J. 1930,24,208. 24 E. Annau, I. Banga, A. Blazo, V. Bruckner, K. Laki, F. B. Staub, A. Szent-Gyorgi,2. Physiol. Chem. 1936, 224, 105. 25 I. Banga, A. Szent-Gyorgi,2. Physiol. Chem. 1937,248, 118. 26 A. Meister. Adv. Enzymol. 1955, 16, 185-246. 27 A. Meister, Annu. Rev. Biochem. 1956,25, 29-56. 28 P. Christen, D. E. Metzler, Transaminases 1985, John Wiley & Sons, New York. 29 D L. Smith, D. Ringe, W. L. Finlayson, J. F. Kirsch,]. Mol. Biol., 1986, 191, 301-302. 30 R. Graber, P. Kasper, V. N. Malashkevich, E. Sandmeier, P. Berger, H. Gehring, J. N. Jansonius P. Christen, Eur. ]. Biochem. 1995, 232,686-690. 31 R. A. Vacca, S. Giannattasio, R. Graber, E. Sandmeier, E. Marra, P. Christen, ]. Biol. Chem. 1997,272(35),21 932-7. 32 R. Graber, P. Kasper, V. N. Malashkevich, P. Strop, H. Gehring, J.N. Jansonius, P. Christen, J. Biol. Chem. 1999,274(44),31 203-8. 33 J. J. Onuffer, J. F. Kirsch, Protein Sc. 1995,4, 1750-1757. 34 J. D. Rozzell, Methodsfor Producing A m i n o Acids by Transamination, 1999, U.S. Patent Applic. 09/334,821. 35 T. Yano, S. Oue, H. Kagamiyama, Proc. Natl. Acad. Sci. U S A , 1998, 95(10), 5511-5. 36 S. Oue, A. Okamoto, T. Yano, H. Kaga-

miyama,]. B i d . Chem. 1999, 274(4), 2344-9. 37 K. Bartsch, R. Schneider, A. Schulz, Appl. Environ. Microbiol. 1996, 62(10), 3794-3799. 38 R. N. Patel, A. Banerjee, V. B. Nanduri, S. L. Goldberg, R. M. Johnston, R. L. Hanson, C. G. McNamee, D. B. Brzozowski, T. P. Tully, R. Y. KO, T. P. LaPorte, D. L. Cazzulino, S. Swaminathan, C.-K. Chen, L. W. Parker, J. J. Venit, Enzyme Microb. Technol. 2000,27(6), 376-389. 39 L.-H. Malmberg, W.4. Hu, D. H. Sherman, Appl. Microbiol. Biotechnol., 1995,44, 198-205. 40 I. G. Fotheringham, D. P. Pantaleone, P. F. Taylor, Chimica OggilChemistry Today, 1997, Sept.-Oct., 33-36. 41 K. Soda, Biochemistry 1968,7,4102-4109. 42 S. Sugio, G. A. Petsko, J. A. Manning, K. Soda, D. Ringe, Biochemistry 1995, 34, 9661-9669. 43 D. Peisach, D. M. Chipman, P. W. Van Ophem, J. M. Manning, D. Ringe, Biochemistry 1998, 37(14), 4958-4967. 44 A. Gutierrez, T. Yoshimura, Y. Fuchikami, K. Soda, N. Esaki, Protein Eng. 1998, 11(1), 53-58. 45 I. G. Fotheringham, S. A. Bledig, P. P. Taylor, /. Bacteriol. 1998, 180(16), 4319-23. 46 J. D. Rozzell, unpublished results. 47 P. V. Warren, R. V. Swanson, Transaminases and Aminotransferases, 1998, U. S. Patent 5,814,473. 48 P. V. Warren, R. V. Swanson, Transaminases and Aminotransferases 1999, U. S. Patent 5,962,283. 49 P. V. Warren, R. V. Swanson, Transaminases and Aminotransferases 2000, U. S. Patent 6,013,509. 50 K. Nakata, T. Narita, H. Tsunekawa, T. Yoshioka, Processfor Producing L-2-Aminoadipic Acid, 1999, U. S. Patent 5, 906, 927.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

I895

13 Formation and Cleavage o f P - 0 Bonds George M. Whitesides

13.1 Introduction

The use of isolated enzymes to form or cleave P - 0 bonds is an important application of biocatalysts. Restriction endonucleases, (deoxy)ribonucleases,DNA/ RNA-ligases, DNA-RNA-polymerases, reverse transcriptases etc. are central to modern molecular biology[']. Enzyme catalyzed phosphoryl transfer reactions have also found important applications in synthetic organic chemistry. In particular, the development of convenient cofactor regeneration systems has made possible the practical scale synthesis of carbohydrates, nucleoside phosphates, nucleoside phosphate sugars and other natural products and their analogs. This chapter gives an overview of this field of research. Hundreds of potentially useful enzymes are available in nature. It is often worthwhile to survey enzymes for applicability in the synthesis of a specific compound, but how to find the best enzyme? Enzymes have been reviewed and classified by many schemes [2-41. Enzymes involved in reactions at phosphoryl groups are, unfortunately for the synthetic chemist, spread almost over all classes. Without a good knowledge of enzymology, it is not easy to find the enzyme classes of interest for a particular transformation. This review links the compound classes and enzyme classification systems in Section 13.1.1 to help overcome this barrier. Most synthetically useful phosphorylating enzymes require nucleoside triphosphates as cofactors. The central importance of cofactor regeneration, and the most used regeneration methods for these cofactors, are discussed in Section 13.2.1. The end of Chapter 13 includes tabular surveys of the most important applications, classified in compound or structural classes (see Sections 13.2.2 and 13.3.3), to facilitate the search for relevant enzymes and procedures.

896

I

73 Formation and Cleavage of P - 0 Bonds

13.1.1

Enzymes Forming or Cleaving Phosphorous-Oxygen Bonds

Phosphoesters are ubiquitous in biochemistry and serve several functions 1'1. Genetic information is stored in DNA and RNA. In cellular control mechanisms, phosphorylation of proteins is an important mechanism for regulating protein activitiesr6]. Phosphorylation can activate metabolites or change solubility properties. Enzymecatalyzed formation and cleavage of P - 0 bonds are central to the cellular energy balance l71, Biosynthesis depends heavily on phosphorylated intermediates. A useful classification for enzymes involved in phosphoryl transfers was introduced by Knowles[*](see Fig. 13-1).This classification, based on enzyme functions and mechanisms, differentiates primarily between two groups of enzymes. The first group contains only enzymes that accept phosphoric monoesters as substrates (type A and B). The second group includes all enzymes catalyzing reactions at phosphoryl groups of phosphodiesters (type C-E). Table 13-la and 13-lb link Knowles' classification and the enzyme classification recommended by the International Union of Biochemistry (IUB; compare Chapter 1)121.The IUB classes give a direct access to the specific enzymes in reference works and to the CA registry numbers necessary for an efficient literature searchI2*1'. Tables 13-la and 13-1b list only the most important categories of enzyme classes (E. C.'s). Some enzymes that are involved in reactions at phosphorus are hidden in other classes. For example glyceraldehyde-3-phosphatedehydrogenase,which catalyses the oxidative phosphorylation of glyceraldehyde-3-phosphateto 1,3-diphosphoglycerate, is classified under E.C. 1.2.1.12 and 1.2.1.13. Neither the name of the enzyme nor its IUB-classification,gives information about the phosphorylating step. Identifying enzymes potentially useful in synthesis that have been ambiguously classified is difficult for those outside of biochemistry because no complete reference is available connecting enzymatic activity with synthetic applicability. A second important point is that many enzyme catalyzed reactions are reversible. Some hydrolytic enzymes can be used in enzyme catalyzed phosphorylation reac-

?-

R-*Of[-0

to A

-

4- ?-

4-

-O-P-'j-O-ZfOfL-Nu

"I

B

OC t

1"

D

4-

R-O-$[-O-R'

to E

Classes o f enzymes involved in reaction at phosphorus. A and B represent enzyme types that handle phosphoric monoesters and related compounds ("0 may be an oxygen o f a hydroxyl, carboxyl, or phosphoryl group, or the nitrogen o f a guanidine group. For simplicity, displacements at t h e y phosphoryl groups o f nucleosides triphosphates were classified with these reaction). C , D and E represent the enzymes that catalyze transformations o f phosphoric diesters (displacements at a or fi phosphorous groups o f nucleoside triphosphates and transfer of pyrophosphates were classified with the reactions of phosphoric diesters). Figure 13-1.

13. I Introduction Table 13-la.

Enzymes accepting phosphoric monoesters as substrates.

Enzyme Functional classb type"

Fundion'

IUB classes with titles, containing

such types of enzymesd

Phosphomutases Phosphoryl group transfer, 2.7.5. for which the acceptor is 5.4.2. another functional group on the donor molecule.

Phosphomutases Intramolecular phospho transferases

Phosphorylases

Formation of a P - 0 bond under phosphorolytic cleavage of a C-Heteroatom bond.

Hexosyltransferases Pentosyltransferases

Nucleotidases

Phosphoryltransfer from 3.1.3. Phosphoric ester hydrolases a nucleotide to water as an (3.1.4 Phosphoric diester hydroacceptor molecule. lases) (Nucleotides are cleaved hydrolytically).

Phosphatases

Phosphoryl group transfer 3.1.3. Phosphoric ester hydrolases from a phosphoric mono- 3.6.1. Hydrolases acting on acid ester to water as an accepanhydrides in phosphoroustor molecule. (Phosphoric containing anhydrides monoesters are cleaved hydrolytically).

Phosphokinases

2.4.1. 2.4.2.

Phosphoryl group transfer: 2.7.1. Nucleoside triphosphate is and the donor and some other 2.7.2. molecules than HzO are Phosphotransfera- the acceptors. Compounds 2.7.4. ses different than nucleoside triphosphates are the donor and some other molecules than HzO are the acceptors. ATPases

Phosphatases which are responsible for the coupling of ATP cleavage to other metabolic processes.

Phosphotransferaseswith an alcohol group as acceptor Phosphotransferaseswith a carboxyl group as acceptor Phosphotransferaseswith a phosphate group as acceptor

3.6.1.3 ATPases

a See figure 13-1; b functional classes bases on ref."'; c see ref."' and 14'; d see ref."'

tions. Alkaline phosphatase (E.C. 3.1.3.1), for example, was used in enzymecatalyzed phosphorylation of glycerol with inorganic phosphate (1' . In some cases enzymes may catalyze unexpected reactions with unnatural substrates: aminoacyl tRNA synthetases (ARS) were used to synthesize p',p4-di(adenosine 5'-)tetraphosphate ( A p d ; l),a natural inhibitor of human platelet aggregation["] (Fig. 13-2). Here, in the first step an amino acid (AA) reacts reversibly with ATP and ARS and forms an aminoacyl-AMP-ARS complex and PPi; the back reaction of this intermediate with ATP leads to the desired product A P ~ [ " - ' ~ ] .

I

897

898

I

73 Formation and Cleavage ofP-0 Bonds Table 13-lb.

Enzymes accepting phosphoric diesters as substrates.

Enzyme Functional type" classb

Function'

IUB classes with titles, containing such types of enzymesd

2.7.6. Diphosphotransferases

C

Pyrophosphokin- Pyrophosphate group ases transfer from ATP to an acceptor molecule other than water.

D

Nucleotidyl transferases

Transfer of nucleotidyl moieties

2.7.7. Nucleotidyltransferases

D

Nucleotidyl cyclases

Nucleoside triphosphate cyclisation under formation of pyrophosphate

4.6.1. Phosphorous oxygen lyases

E

3.1.5. Triphosphoric monoester Triphosphohydro- Triphosphate transfer hydrolases from a nucleoside triphoslases phate to water as an acceptor molecule.

E

Polynucleotide synthetases

Responsible for the linkage of two poly- or oligonucleotide moieties to form polynucleotide chains

6.5.1. Ligases forming phosphoric ester bonds

E

Phospholipases

Hydrolytic cleavage of phosphoglycerides (essentiallyphospholipase C and D)

3.1.4. Phosphoric diester hydrolases

E

Nucleases

Phosphonucleotide trans- 3.1.4. Phosphoric diester hydrolases fer from a polynucleotide 3.1. Endo- and exonucleases to water as an acceptor molecule. (Polynucleotides are cleaved hydrolytically).

E

Phosphodiesterases

Phosphomonoester trans- 2.7.8. Transferases for other fer from a phosphodiester substituted phosphate groups other than polynucleotide 3.1.4. Phosphoric diester hydrolases to water as an acceptor molecule. (Phosphodiesters are cleaved hydrolytically).

a See figure

13-1;b functional classes bases on ref.? c see ref."' and r4'; d see ref.['].

One of the most important criteria in the evaluation of a new process is the availability of an (see Chapter 20: Tabular Survey of Commercially Available Enzymes). If the enzymes are not commercially available, their isolation and purification can be expensive and time consuming (see Chapter 2: Production and Isolation of Enzymes). The importance of the product to be synthesized may sometimes justify the additional effort. Mechanistic aspects of L O bond formations and cleavages have been reviewed["] and are outside the scope of this work. The use of enzymes catalyzing the formation

I

13.7 Introduction 899

t ARS

AA

+

AT?

ATP

AA-AMP-ARS AA

pi

HO OH

+ ARS

HO OH 1

Figure 13-2. Enzymatic synthesis of p’,p4-di(adenosine 5’-)tetraphosphate (Ap4A 1) with aminoacyl tRNA synthetases (ARS). AA can be leucine, for example, and ARS leucyl t-RNA synthetase[”].

of P-N bonds - for example, phosphorylations of amino acids (E.C. 2.7.3) - are discussed only briefly. Enzymes dealing with the formation of aminoacyl tRNA (E. C. 6.1.1),acyl-CoA derivatives (E. C. 6.2.1) or peptides (E. C. 6.3.2) are also not covered, even if cleavages of nucleoside phosphates are involved. 13.1.2

Biological Phosphorylating Agents

To compare the ability of different compounds to transfer a phosphoryl group, phosphorylation of water was chosen as a standard reaction[’7].The free energy of hydrolysis of a phosphorus compound (AG2ydr) is called its phosphorylating potential. Table 13-2 summarizes the phosphorylating potentials of the most important biological compounds (Fig. 13-3)having phosphoryl donor abilities. By far the most important strong biological phosphorylating agent is adenosine 5’-triphosphate (ATP, 8). ATP is ubiquitous and plays a central role as cofactor in anabolic and catabolic processes. Moreover, many enzymes involved in the formation of P-0 bonds are ATP dependent. The biologically active form of ATP is, in most cases, the magnesium salt MgATP2-[221. Other nucleoside triphosphates have similar phosphorylating potentials but they are rarely used as phosphoryl group donors[23,241; usually GTP, CTP and UTP act as nucleoside or nucleoside phosphate donors (see Section 13.2.2.2). Creatine- and arginine phosphate (7 and 9) play important roles in the storage of phosphorylating potential in vertebrates and invertebrates, respectively[25* 26]. In living cells, these N-phosphoguanidine derivatives are formed by phosphoryl group transfer from ATP, and in the reverse reaction ADP is the only acceptor for 7 and 9. 1,3-Diphosphoglycerate(5) and phosphoenolpyruvate (2) are important phosphorylating agents of ADP in the glycolytic pathway. P~lyphosphate[~~], phosphoramidate 12’] and pyrophosphate[”] are involved in the biochemical phosphorylation

900

I

73 Formation and Cleavage of P - 0 Bonds Table 13-2.

Free energies of hydrolysis of some important biological phosphorus "1.

Compound (R-OPOs2-)

PH

[kcal/mol]

[kJ/mol]

Phosphoenolpyruvate (2) Methoxycarbonylphosphateb (3) Carbamyl phosphate (4) 1,3-Diphosphoglycerate ( 5 ) Acetyl phosphate (6) Phosphocreatine (7) ATP (8) (+ ADP + Pi)' ATP (8) (+ AMP + PPi) Arginine phosphate (9) Pyrophosphate' (PPi) Glucose 1-phosphate (10) Glucose 6-phosphate (11)

7.0 7.0 9.5 6.9 7.0 7.0 7.4 7.0

12.8 12.4 12.3 11.8 10.3 10.3 7.3-9.6 7.7 7.7 4.5-8.0

53.5 51.8 51.4 49.3 43.1 43.1 30.5-40.1 32.2 32.2 18.8-33.4

7.0

3.3 5.0

20'9 13.8

Glycerol-1-phosphate (12)

8.5

2.2

9.2

8.0

7.0

strong phosphorylating agents

1

rl?:phorylating agents

The standard free energies are bases on a standard state of IM total stoichiometric concentration of reactants and products, except hydrogen ion, and on an activity of pure water of 1.0: see ref."8]; Hydrolysis of ATP and PI', depend strongly on the concentration of Mg2+in solution and on pH[1s211, a

2

3

4

I

8

5

6

9

/OP

HO

OP OH 10

Figure 13-3.

Ho*HO OH

OH 11

P O L O H 12

Structures of the most important biological phosphorylating agents. P =

phosphate.

of D-glucose, hexoses and L-serine respectively in some organisms. Carbamylphosphate (4) and acetylphosphate (6) have high phosphorylating potentials (see Table 13-2), but nature uses them mainly as donors of ~arbamyl[~'] or acetyl groups[31]. Only in a few cases do they act as phosphoryl donors[30,321. Phosphorylations with low-potential phosphorylating agents are thermodynamically not favorable. In biological systems, these processes are made possible by

13.2 Phosphorylation

coupling them to a thermodynamically more favorable process. Examples of weak phosphorylating agents are sugar phosphates such as glucose- and ribose phosphates, which can transfer their phosphate group to other sugars[32]or to nucleosides like riboflavin [331. Phosphate sugars are formed when polysaccharides are cleaved with a phosphorylase and inorganic phosphate[34].

13.2

Phosphorylation

Chemical phosphorylations usually involve many protection and deprotection steps. Enzymatic phosphorylations can make synthesis more efficient by eliminating many of these steps. In addition, enzyme-catalyzedintroduction of phosphoryl groups can be diastereo-[351 or enantiospecific[36, 371. One of the major challenges in enzyme-catalyzedphosphorylation reactions is, as mentioned above, the choice of the most convenient enzyme. The other major difficulty is the availability of the coenzymes. Cofactors act as biological phosphoryl donors and in enzyme-catalyzed synthesis, they have to be added in stoichiometric amounts or coupled to an efficient regeneration system. 13.2.1 Regeneration of Nucleoside Triphosphates

In enzyme-catalyzed synthesis, adenosine 5'-triphosphate (8) is the cofactor most often used as phosphoryl group donor. Other nucleoside phosphates, UTP, or CTP are used principally as donors of a nucleoside phosphate moiety to form activated intermediates in biological pathways (see Section 13.2.2.2). For example: UTP precedes the activated from a glucose, UDP-glucose, in the Leloir synthesis of polysaccharides, CTP precedes CDP-choline in the synthesis of phospholipids and CMP-NeuAc in the formation of glycosides of sialic acids (see Chapter 11.3). The costs for a mole CTP, GTP or UTP vary from $ 32000 to 90000 (as research biochemicals)[381. The high price of these cofactors precludes their large-scale use in stoichiometric quantities and makes cofactor regeneration necessary. Even with ATP, one of the least expensive cofactors used in organic synthesis[3', 381 and available through mole scale synthesis from RNA r4OI regeneration remains of central importance. The use of a cofactor regeneration system not only eliminates the need for stoichiometric quantities of cofactor but it can also favorably influence the position of the reaction equilibrium and prevent the accumulation of cofactor byproducts that may inhibit the forward process. Product isolation is simplified as well. A nucleoside phosphate regeneration system must meet several specifications to be practical. To be economical, a regeneration method must be capable of recycling the cofactor 102-106 times [391. All materials should be readily available, inexpensive, easily handled, stable under reaction conditions and compatible with the rest of the reaction system. The transfer of phosphate should be thermodynamically and

I

902

I

13 Formation and Cleavage of P - 0 Bonds

kinetically favorable and it should be regioselective in forming a high-energy P - 0 bond. 13.2.1.1

Regeneration ofATP from ADP and AMP

At the scale required for synthesis of fine chemicals, the major problems of ATP regeneration have been solved13'. 41, 421 . Three strategies have been applied: chemical synthesis; biological methods including whole cells, organelles, and fermentation processes: and cell-free enzymatic catalysis. Chemical methods often lack the necessary specificity and are not compatible with biochemical transformations. Biological and enzymatic systems provide the most efficient ATP regenerating systems 13'1. The use of cell-free enzymes requires a greater initial effort or expense than do the biological methods, but are more specific than biological systems and often generate fewer by-products (see ref. r3'1 and references cited therein). a) From ADP. Several procedures for the large-scaleregeneration of ATP from ADP using isolated enzymes as catalysts are 391. These methods have in common the characteristic that phosphoryl groups are transferred from a highenergy phosphoryl donor to ADP (compare Section 13.1.2). The advantages and disadvantages of these methods are summarized in Table 13-3. In practice, for most synthetic applications, either acetyl phosphate/acetate kinase or phosphoenolpyruvate/pyruvatekinase are used to regenerate ATP. Because of the ease of preparing AcP, AcP/AcK is the most economical method for large-scalework. Its application is, however, limited to fast phosphorylation reactions where the hydrolysis of AcP is not important. The PEP/pyruvate kinase system is used in instances where the requirement for a strong, stable phosphorylating reagent outweighs the relative inconvenience of preparation of PEP.

Phosphoenolpyruvate/pyruvate kinase. Phosphoenolpyruvate(PEP; Z)/pyruvatekinase (PK; E.C. 2.7.1.40) is the most efficient system for the regeneration of ATP from ADP. The phosphorylating agent PEP can be prepared in a mole Starting from crude pyruvic acid, the crystalline monopotassium salt PEP-K' is synthesized in a three-step procedure. For transformations on a scale <1 mol, PEP can be prepared from commercially available 3-phosphoglyceric acid in an enzyme-catalyzed reaction["]. This method is more expensive than the chemical preparation, but is more convenient because it requires less time and produces less organic waste (see Section 13.2.1.2; Fig. 13-5). Pyruvate kinases are commercially available from multiple sources F3*1. The enzyme generally used in ATP regeneration - from rabbit muscle - has high specific activity (-500 U per mg of protein), is inexpensive (Is 2-4/1000 U), and is stable when immobilized[Gs5-G7]. This regeneration system can be used in membrane enclosed enzyme catalysis (MEECtechnique) kG8, "1 as well. The stability of PEP in solution and its strength as a phosphoryl donor (Table 13-2) make PEP particularly convenient for use in slow and thermodynamically un-

product inhibition/ KL[mMlb

costs [$/loo0 U], (so~rce)[~~l

-

- 3.0; - 12.6 21

- lo3

+++I51

0.4b

5.5; - 23.0

+I471

[481

0.41501

++++e.

400-1200

acetate 400, NC

thermophilusj

-

CP/CK

0.3

2.2

- 5.0; - 20.9

+++[52J

++b

5.1; - 21.3

0.1'

O.OSC

+I301

400-900

C

10 (Streptococcusfaecalisj

l.Gb

-

+++f,

2 140b

HCOj500, NC

thermophilusj

MCP/AcK

378 ( B . stearo-

AcP/AcK

378 (B. stearo-

-

1491

- 1o2I

3.0; - 12.6

+WI

Sf, b

0.05[25]

+++f.

150-250

[*I

-

2 - 0.7; - 2.9*

+++++[S4J

0.003[441

0.17[441

++++e,

15&250[44]

-

muscle) creatinelZsJ 6-40d, NC

Pn/PnK

isolated from E. coii[44~

CrP/CrK

2.6 (rabbit

Pi/glycolysis

- 16; - 67.5k

-

+a. f

8h

-

sources)a

66 (from diff.

a calculated from 1000 U of each enzyme; see ref.l4'I and 1381: b see ref."*l C = competitive, NC = non-competitive; c carbamate kinase kinetics 1s complex, inhibition plays an important rolel4'I; d K,depends strongly on the anions present in ~ o l u t i o n ~e~immobilized '~; enzyme($ fvalue(s) for the free enzyme(s);g calculation based on the values from Table 13-2 (AG(P-transfer)= AG$, (P-donor)- A d , , , (ATP); AG;,,, (ATP)= - 7.3 kcal/mol; 30.5 kJ/mol);h calculation based on the sum of all enzymes present; i based on Table 13-2,values for PP,; k the driving force of this process is the transformation of glucose to 2 equivalents of lactate (AG = - 197 kJ/mol)lz2~; I calculated from data in ref.ls51.

half life for hydrolysis (25 "C, pH 7)b

P-donors AG (P-transfer)g properties [kcal;kJ/mol]

ease of preparation/ availability

O.lb

0.07b

K, (P-donor) [ m ~ ]

1471

Km (ADP)'[mM]

++++e.

30&500

pyruvate 10, c

muscle)

2.5 (rabbit

PEP/PK

Properties o f ATP-regeneration systems.

Enzyme spec. activity properties [u/mg pr~tein][~'J stability

Table 13-3.

-5-

I!

-

904

I

13 Formation and Cleavage of P - 0 Bonds

favorable phosphorylation reactions. It is also the method of choice for the regeneration of ATP at low concentrations of ADP, since the Michaelis constant for PK is smaller (K,(MgADP) = 0.1 mM)ll'l than for acetate kinase (K,(MgADP) = 0.4 mM) [SO]. The PEP/PK regeneration method has two minor disadvantages. First, the synthesis of PEP [64. 471 requires more effort and expense than does the synthesis of AcP rS71. Second, pymvate is a strong inhibitor of PK (see Table 13-3). The reactions are therefore carried out in dilute solutions to keep the pyruvate concentration low, and pyruvate is either removed from the reaction solution or PEP is used at high concentrations to minimize the effects of inhibition.

Acetyl phosphatelacetyl kinase. Acetyl phosphate (AcP; G)/acetyl kinase (AcK; EC 2.7.2.1) is the most widely used large scale ATP-regeneration system. AcP is modestly stable in aqueous solutions and is a phosphoryl donor of intermediate strength (Table 13-2and 13-3).Diammonium acetyl phosphate can be prepared from ketene and anhydrous phosphoric or, more easily, from acetic anhydride and anhydrous H3P04[57].However, the use of the diammonium salt in ATP regeneration has three disadvantages. First, NH; reacts with acetyl phosphate in solution. Second, it forms an insoluble precipitate with Mg2+ under reaction conditions. Third, its preparation involves several steps that require careful experimental control and that are difficult to carry out at large scale. Preparation of aqueous solutions of acetyl phosphate as its sodium r5l] or potassium salt [l'] circumvents these drawbacks. Two types of commercially available 13'1 acetate kinases, from Escherichia coli ["I and from Bacillus ste~rothe~ophilus~~'1, have been used in ATP regeneration. The latter kinase is more expensive but it is preferred for synthetic use because it is thermostable and it is stable to auto~idation[~'1.Both enzymes have acceptable specific activities (150-300 and 400-1200 U respectively per mg protein) and can be stabilized by imm~bilization[~', ", G662]. Acetate is a weak inhibitor of AcK, but product inhibition is not a serious problem (see Table 13-3)unless reaction solutions have acetate concentrations greater than 1 M [ ~ ~The ] . relative instability of AcP in solution compared with PEP is the major disadvantage of the AcP/AcK system. The contribution of the enzymes to the total cost of the process in generally low when they are recycled, making the slightly higher cost of AcK compared with PK a minor disadvantage [391. Polymer bound ATP was regenerated in a membrane reactor with AcP/AK Ib31.

Methoxycarbonyl phosphate/acetate kinase. Methoxycarbonyl phosphate (MCP; 3) was designed to replace AcP as phosphoryl donor['']. It is comparable to PEP in its high phosphorylating strength (see Table 13-2),but resembles acetyl phosphate in its ease of synthesis. Aqueous solutions of MCP are prepared from aqueous phosphate and methyl chloroformate and used in ATP regeneration without purification. The reaction product after phosphoryl transfer is methyl carbonate, which hydrolyses rapidly to form C 0 2 and MeOH. Product isolation is simple and bicarbonate inhibition can be avoided by purging the reaction mixture.

13.2 Phosphoryhtion

MCP is accepted as an unnatural substrate by AcK (E. C. 2.7.2.1) and CK (E. C. 2.7.2.2) but not by PKL1'l.The principal disadvantage of methoxycarbonyl phosphate as phosphorylating agent in ATP regeneration is MCP's rapid spontaneous decomposition. The half-life of MCP in aqueous solution is only 0.3 h (25 "C, pH 7)'''). Because of this short half-life, MCP is only used in a few cases where high phosphorylating potentials are required to push the phosphorylation reaction to the product side (see Table 13-4, entry 20). Others. ATP has been regenerated from ADP with propionylphosphate/acetate kinase, but propionylphosphate is a poorer substrate than AcP[*']. Carbamoyl phosphate (CP; 4)lcarbamyl kinase (CK E. C. 2.7.2.2) was described as a regeneration method for ATP in 1973 but it has seldom been used[70].CP is a very strong phosphorylating agent (see Table 13-2) and can easily be prepared from aqueous potassium cyanate and KH2P04[52]. Rapid decomposition of carbamyl phosphate generates ammonium ions and magnesium ammonium phosphate complex is formed as a gelatinous precipitate. This precipitation lowers activity, both by precipitating the magnesium(I1) required for activity of the kinases, and because it occludes enzymes. ADP phosphorylation with polyphosphate (P,) and polyphosphate kinase (P,K E. C. 2.7.4.1) has also been demonstrated[44].The cheap, stable polyphosphates and the stability of P,K are highly attractive. Unfortunately, P, has a low phosphorylating potential and P,K is not readily available. These facts may explain why this regeneration system has not found any practical application. Another very interesting but little-used regeneration method is based on phosphocreatine (PC; 7) and creatine kinase (CrK E. C. 2.7.3.2)r7l].PC is comparable in its phosphorylating potential to AcP, but it is more stable in aqueous solutions (Table 13-3). CrK is inexpensive and fairly stable. The current lack of an efficient and simple laboratory scale synthesis for PC seems to have limited the applications of this method to a few syntheses of sugars r7lI and nucleosides [721. Recently, a promising regeneration system based on a multienzyme system from the glycolytic pathway was described[45](Fig. 13-4). Glucose and inorganic phosphate were used as low-energy phosphorylating agents. Eleven enzymes are used to convert glucose to lactate. Two equivalents of inorganic phosphate are consumed and two equivalents of ATP are formed. The overall process of this regeneration system has a favorable free energy (see Table 13-3), which can be useful in the synthesis of compounds with high phosphorylating potentials (see Table 13-4, entry 20). The main backdraw of this method is the complexity of this multienzyme system and the poor stability of some of the enzymes used. CHpOH

Figure 13-4. ATP regeneration via the glycolytic catalyze the conversion of glucose t o lactate.

Eleven enzymes are used t o

I

905

906

I

73 Formation and Cleavage of P - 0 Bonds

b) From A M P . In biochemical processes, ATP may be converted to either ADP or AMP. The regeneration of ATP from AMP is slightly more complicated than from ADP. Methods coupling one of the above mentioned regeneration systems and adenylate kinase (AdK; E. C. 2.7.4.3)have been used extensively in the production of ATP[48s431. AdK catalyzes the formation of 2 ADP from ATP and AMP. AcP/AcK and PEP/PK are the methods most often used to convert ADP to ATP (vide infra). 13.2.1.2

Regeneration o f Other Nucleoside Triphosphates

The preparation of nucleoside triphosphates (NTP) from nucleoside diphosphates (NDP) follows the same regeneration systems described above (Section 13.2.1.1). ~, AcK and PK have broad substrate specificities and recognize all of the N D P s [ ~731. The efficient generation of NDP from nucleoside monophosphates (NMP)has been solved on preparative scale[74].Adenylate kinase was used in the preparation of cpdine S'-triphosphate (CTP) and uridine 5'-triphosphate (UTP) from the corresponding nucleoside monophosphates. Nucleoside monophosphate kinase (NMP; E. C. 2.7.1.4)was used in the synthesis of UTP. Guanosine 5'-triphosphate (GTP)was prepared with guanylate kinase as catalyst, coupled to a conventional ATP regeneration system. Best results were achieved when 3-phosphoglyceric acid served as the ultimate phosphorylating agent, and a multienzyme system was used as transfer catalyst (Fig. 13-5). XMP

li ii

OP

A 0 0 0

AooEnzymatic synthesis of nucleoside triphosphates. i) Phosphoglycerate mutase (E.C. 2.7.5.3); ii) enolase (E.C. 4.2.1.1 1); iii) pyruvate kinase (E.C. 2.7.1.40); iv) adenylate kinase (E.C. 2.7.4.3, X = A , C, U), guanylate kinase (E.C. 2.7.4.8, X = C) or nucleoside monophosphate kinase (E.C. 2.7.4.4, X = U). P = phosphate. See Figure 13-5.

13.2 Phosphorylation

I

907

13.2.2

Applications 13.2.2.1

Phosphorylationswith ATP as a Cofactor

The most widely used and best developed enzyme-catalyzed phosphorylations are the ones that are coupled to ATP regeneration systems. Sugar phosphates, nucleoside phosphates and glycerides are the major classes of compounds prepared with these methods. Kinases are the enzymes most often used for the phosphorylation of saccharides (Table 13-4,entries 7-19).For example, glucose-6-phosphate(ll),a useful reagent for the regeneration of nicotinamide cofactors (see Chapter 15), was prepared from glucose in a one-step reaction by phosphorylationwith ATP 17'1. ATP was regenerated with AcP/AcK and the phosphoryl transfer was catalyzed with hexokinase, (HK;E. C. 2.7.1.1), and enzyme with broad substrate specificity. Both enzymes were immobilized and reused after product isolation. Alternatively, fluorinated hexopyranose phosphates and glucose phosphate analogs, with sulfur or nitrogen in the ring, were prepared with PEP/PK and HK[761 (Table 13-4,entry 9). The synthesis of 11 starting from polysaccharides or from fructose 1,6-diphosphate[771 was demonstrated, but the former method is less convenient and the latter is more expensive than the procedure starting from D-glucose. 5-phospho-D-ribosyla-1-pyrophosphate (PRPP; 13) is a key intermediate in the biosynthesis of various nucleotides and other natural products. An interesting application of an ATP/AMP regeneration system is demonstrated in the synthesis of PRPP from D-ribose in a multienzyme (Table 13-4,entries 13 and 14). In the first step, ribose was phosphorylated with ATP using ribokinase ( R K E.C. 2.7.1.17) as catalyst. In the second step, PRPP-synthetase(E. C. 2.7.6.1) catalyzed the transfer of a pyrophosphate group from ATP to ribose 5-phosphate (Fig. 13-6). The preparations of ATP, GTP, CTP and UTP have been discussed in Section 13.2.1. Kinases are the enzymes most popular for the synthesis and regeneration of nucleoside triphosphates from their mono- and diphosphates. Nucleoside phosphate analogs have been synthesized using the same enzymes. For example, ribavarin triphosphate (RTP; 14), a compound with anti-viral properties, was prepared from ribavarin monophosphate with adenylate kinase (E. C. 2.7.4.3) and pyruvate kinase (E. C. 2.7.1.40) as catalysts with PEP as ultimate phosphorylating agent (Fig. 13-7)[481. Here PEP/PK has proved to be more useful as regeneration system for ATP than AcP/AcK (see Section 13.2.1.1) in a typical example of a kinetically unfavored reaction. RMP is one of the rare unnatural substrates accepted by adenylate kinase. Other nucleotide analogs - for example ATP-a-S and ATP-)I-S- have also been synthesized using kinases (Table 13-5, entries 4 and 5) 17', 791. Glycerol kinase (GK; E. C. 2.7.1.30) catalyzes the enantiospecific phosphorylation of glycerol to form sn-glycerol-3-phosphate, an important intermediate for the synthesis of phospholipids. The enzyme is inexpensive and stable when immobilized. Studies with enzymes from a variety of microbial sources have shown

908

I

i-203pw

73 Formation and Cleavage of P - 0 Bonds

HO

*OH

i

HO OH

pyruvate

HO OH

1

ATP

AMP + ATP

PYmate

Fw

13

Coupled enzymatic synthesis o f PRPP from ~ - r i b o s e [ ' ~ i) ]ribokinase; ii) pyruvate kinase; iii) PRPP-synthetase; iv) adenylate kinase. Figure 13-6.

RMP

RTP

RDP

H

pymvate

p- p- p'0-P-0-P-0-P-0

HO OH

PEP 14

Enzymatic synthesis of ribavarin 5'-triphosphate (RTP; 14) from ribavarin 5'-monophosphate (RMP) 1481. i) adenylate kinase (EC 2.7.4.3); ii) pyruvate kinase (EC 2.7.1.40). Figure 13-7.

that glycerol kinase accepts a wide range of glycerol analogs as substrate (Table 13-4, entries 3 - 5 ) [361. sn-Glycerol-3-phosphate(12) and analogs were synthesized in gram scales, using glycerol kinase as catalyst and PEP/PK or AcP/AcK as ATP regeneration system. The phosphorylation of racemic mixtures produced chiral organic phosphates with enantiomeric excess (ee) >90-95% and yields of 75-95%. The unphosphorylated enantiomers of the chiral substrates were recovered in yields of 30-40% (80-90% ee). Alkaline phosphatase was used to hydrolyze the phosphorylated enantiomer and to provide enantiomerically pure unphosphorylated mateFor example, ~-3-Chloropropane-l,2-diol (15) was prepared from a racemic mixture in a two step procedure with a 53 % overall yield (97% ee) (Fig. 13-8). Another application of glycerol kinase is the monophosphorylation of dihydroxy-

13.2 Phosphorylation

HO H

CI &OP03'ATP

j

ADP

C

ii

HO H I A O

H

15 Enzyme catalyzed separation of a racemic mixture of ~,~-3-chloropropane1,2-diol. i) glycerol kinase; ii) alkaline phosphatase. Figure 13-8.

Dihydroxyacetone phosphate (DHAP),an important intermediate in the aldolase catalyzed synthesis of monosaccharides (see Chapter 14), was prepared in a 0.4 mol scale using AcP/AcK for the regeneration of ATP (Table 13-4; entry 6 ) . Guanidine derivatives were phosphorylated with ATP as well. The syntheses of arginine- and creatine phosphate (7 and 9), two relatively strong biological phosphorylating agents, are not economical, but they demonstrate the potentials of different ATP regeneration systems (see Table 13-4, entries 20 and 21). Further applications of ATP regeneration systems are given in Table 13-4 and 13-5. 13.2.2.2

P - 0 Bond Formation with Other Nucleoside Triphosphates than ATP

The use of stoichiometric quantities of nucleoside triphosphates or their regeneration from the corresponding mono- or diphosphates have found important applications in the synthesis of activated natural products; for instance, nucleoside phosphate sugars are important biological intermediates in the synthesis of complex carbohydrates, glycoproteins, glycolipids and proteoglycans. All of the eight nucleoside phosphate sugars, used in vivo by mammalian glycosyltransferases in the Leloir pathway, are accessible today by practical enzymatic or chemoenzymatic approaches (see reviews ref.[''] and [81]). The use of these activated monosaccharides in the enzymatic preparation of oligosaccharides and glycoconjugates is discussed in Section 11.3. The enzymatic synthesis of sucrose is, however, discussed here, to illustrate the efficient in situ generation of UDP-glucose from UDP in a complex multienzyme reaction (Fig. 13-9). In a typical example, a nucleoside triphosphate is recycled to regenerate a nucleoside phosphate sugar The synthesis, starting from glucose-1-phosphateand fmctose, used sucrose synthetase (E. C. 2.4.1.13), pyruvate kinase (E. C. 2.7.1.40) and UDP-glucose pyrophosphorylase (E. C. 2.7.7.9). Inorganic pyrophosphatase (E. C. 2.6.1.1) was used to keep the pyrophosphate concentration in the reaction mixture low and to drive the equilibrium to the product side.

I

909

910

I

13 Formation and Cleavage of P - 0 Bonds

HO HO

OH

D-fructose

UDP-glucose

UDP

4

a-D-glucose 1-phosphate General scheme for an enzyme-catalyzed synthesis o f sucrose with UDPglucose[821.i) sucrose synthetase; ii) pyruvate kinase; iii) UDP-glucose pyrophosphorylase; iv) inorganic pyrophosphatase. Figure 13-9.

13.2.2.3

Other Phosphorylating Agents

Agents other than ATP are rare in enzyme-catalyzed formation of P - 0 bonds. Inorganic phosphate, pyrophosphate and polyphosphates were used to prepare and phenols rS41. The yields phosphorylated monosaccharides, alcohols, polyols were poor and the reactions lack specificity (Table 13-4, entries 1, 2 and 4). Glycerol1-phosphate was prepared from glycerol, for instance, using inorganic phosphate and alkaline phosphatase (E.C. 3.1.3.1): 75 g of the product was isolated in a 41% yieldL9I. The reaction was regio- but not stereospecific. Phosphorylases were used with isotopically labeled phosphate or with inorganic thiophosphate to prepare (thi0)phosphorylated monosaccharides from oligo- or polysaccharides (see Table 13-4, entries 18 and 15). 32P-labelingof phospholipase with [32P]Piwas used to study the biological function of this enzyme complex[85].p-Nitrophenyl phosphate was employed as phosphoryl group donor in the synthesis of allo-uridine, using a phosphotransferase as catalyst (see Table 13-5, entry 6 ) .

R=H

?R

R = PO: (45 %)

Phosphorylated products

(rat)

X = 0, NH, H; Y OH, H, CH20H, F, SH, NHz; Z = H, CH3, CH2CH3; Y=Z=O.

CH2XH

Y Z

5

R

X=OH;Y=O

X = C1, SH, OCH3, CHIOH, Br, CHzCH3, OH; Y = 0 or NH

J/YH

4

3

+ non

(45-96 %)

RxCN2XPO:-

Y Z

(rac)-glycerolphosphate

phosphorylated enantiomer (29-95 %)

xJ,um:-

Glycerol, GlycerolAnalogs, Dihydroxyacetone

2

Aliphatic and Aromatic Alcohols 1 Monosaccharides, alcohols, polyols

Product

P - 0 bond formation at non-nucleoside compounds.

Entry Starting material

Table 13-4.

(E.C. 2.7.1.30)

Alkaline Phosphatase (E.C. 3.1.3.1) Glycerol Kinase

Glycerol Kinase (E.C. 2.7.1.30)

Alkaline Phosphatase (E.C. 3.1.3.1)

Alkaline Phosphatase (E.C. 3.1.3.1)

Enzyme

ATP

Pi, PPi

ATP

Pi

PPi

P-Source

~~

PEP/PK

PEP/PK or AcP/AcK

Cofactor regeneration

~~

References

13'1; see also "[I

~ 3 1 [91 .

~

D

a

d

2

3

2.

i

s-

-Q

3-

h,

w

d

Starting material

(cont.).

OH

OH

OH

D O HO

D-arabinose

HO

H

(and other glucose analogs)

ibid

HO

Monosaccharides

Entry

Table 13-4.

(63%)

(86%)

11 (80%)

HO

OH

11 (65%)

OH

PEP/PK

PEP/PK

ATP

Hexokinase (E.C. 2.7.1.1)

ATP

AcP/AcK

AcP/Ac

PEP/PK or AcP/AcK

Cofactor regeneration

ATP

ATP

ATP

P-Source

y-Glutamyl-Cysteine Synthetase (E.C. 6.3.2.2)

Hexokinase (E.C. 2.7.1.1)

GK (E.C. 2.7.1.30)

0 HOAOP0:-

DHAP (83%)

Enzyme

Product

I7'j1; see

also

I131, IS91. [771

1631; see also

[431; see also [")I

References

N

2

B

3

m

0

P

9

%

6

9 f!

SL

Q

3

s.

3

F

(u

-

-

(cont.).

16

15

14

13

12

11

OH

0PO;-

HO

OH

OH

Om;-

om:-

OPO,”

OH

(74%)

0

0 0-7-0-7-0 OH 0- 0-

D-ribose 5-phosphate

HO

Ribokinase (E.C. 2.7.1.17)

2-

PRPP Synthase (E.C. 2.7.6.1)

Phosphorylase a

[=PIPi

ATP

ATP

HO

0PO:-

Fructokinase (E.C. 2.7.1.11) and Phosphoglucoisomerase (E.C. 5.3.1.9)

03psPoH

[35P]glucose1-phosphate

Glycogen

OH

PRPP (75%)

D-ribose 5-phosphate

HO

2-03polp&

ATP

ATP

ATP

Fucose Kinase (E.C. 2.7.1.52)

Phosphoribulokinase (E.C. 2.7.1.19)

P-Source

Enzyme

2-03p0T2-

ribulose 1,s-biphosphate

Ho &OH

(80%)

HO OH

H 3 c F o H

Product

2-03p0w

D-ribose

HO

ribulose 5-phosphate

0

Ho &OH

HO OH

H3C77-&?

Entry Starting material

Table 13-4.

none

none

AcP/AcK

PEPIPK

PEP/PK (82%) or AcP/AcK (6G %)

PEP/PK

Cofactor regeneration 1911

References

L

3

P

a -.,

(cont.).

OH

r-Arginine

creatine

H2NKTVoCH3 0

NH;

Enzyme 22 Phospholipase C

21

20

Guanidine derivatives

R=H

Sucrose

18

19

Galactose

17

material

Entry Starting

Table 13-4.

NH;

Phosphorylated Phospholipase

Phospho-arginine

phosphocreatine

CH, 0

bo$'HNKT/Ko-

R = PO: (18%)

Glucose I-thiophosphate (55 %)

Galactose 1-thiophosphate (25%)

Product

Acid Phosphatase (E.C. 3.1.3.2)

Arginine Kinase (E.C. 2.7.3.3)

Creatine Kinase (E.C. 2.7.3.2)

Hexokinase (E.C. 2.7.1.1)

Galactokinase (E.C. 2.7.1.6) Sucrose Phosphorylase (E.C. 2.4.1.7)

Enzyme

Pi

ATP

ATP

ATP

NasSP03

ATWS)

P-Source

AcP/AcK (24%) MCP/AcK (55%) Glucose, Pi + Mutienzyme system PEP/PK (67 %) or AcP/AcK (31%)

PEP/PK

none

none

regeneration

Cofactor

1851

11001

[451

[I81

LSSI

1991

[971; see also ["I

1971

References

B

B

2

g

5

n

n

3

2.

$

bJ

1

OH

0

R = PO:- (31%)

ATP-.I-S (80%)

ATP-a-S (53%)

HO

AdK (E.C. 2.7.4.3)lPK (E.C. 2.7.1.40)

RTP (14) (93%)

RMP

(AMPS)

Adenylate Kinase (E.C. 2.7.4.3)

ATP, GTP, CTP, UTP

NMP (RNA)

Phosphotransferase (Malt Sprouts)

PhosphoglycerateKinase (E.C. 2.7.2.3)

NMP-Kinase (E.C. 2.7.4.4)

dATP (67%) Adenylate Kinase (E.C. 2.7.4.3)

Enzyme

dAMP (DNA)

Nudeotides and Analogs

Product

P - 0 bond formation at nucleosides.

Entry Starting Material

Table 13-5.

PEP/PK

PEP/PK

Cofactor regeneration

see

1781;

1941

References

1-(Thiophosph0)- Dihydroxyacetone, [791 3-phosph0NaZHSP03, glycerate Multienzyme System [103]. none p-Nitrophenyl phosphate see a l ~ ~ [ ~ ~ ~ l

ATP

ATP

ATP

ATP

P-Source

Starting Material

(cont.). Product

ATP

O-

O-

OH

HO

OH

P-Source

UDP-Glucose Pyrophosphorylase (E.C. 2.7.7.9)

Leucyl t-RNA Synthetase (E.C. 61.1.4)

ATP

cHz-C;l*

\

P+

\ o

OH

UDP-Glucose (95%)

A p A (98%)

(-90%)

-0'

0

Phosphorylase (E.C. 2.7.7.8)

Polynucleotide

DNA Polymerase 1 (E.C. 2.7.7.7) and T4 DNA Ligase (E.C. 6.5.1.1)

Enzyme

WA pppoYYA

HO

A=Adenine

t

I

-O-P-O-P-CH,-CHz

F1F1

Nucleoside Phosphate Sugars 10 UTP + Glucose 1-Phosphate

9

8

Oligonucleotidesand Analogs 7 Single Stranded DNA+ Phosphorothioate + Mixture of Deoxynucleoside Analogs of DNA 5 ' - 0 - (1-thiotriphosphate) phosphorothioates

Entry

Table 13-5. Cofactor regeneration

11051

References

-

B

3

m

P 0

4

%

z

;

SL

Q

6'

4

3

F

(u

Uridine-5'-0-(2thiodiphosphogalactose) (UDP@S)-Galactose (50%)

Galactose 1-thiophosphate and UDP-glucose

11

NAD+ (> 90%)

NADP+ (quant)

ATP + Nicotinamide Mononucleotide

NAD+

12

13

NAD(P)+

Product

Starting Material

(cont.).

Entry

Table 13-5.

NAD Pyrophosphorylase (E.C. 2.7.7.1) NAD Kinase (E.C. 2.7.1.23)

Galactose-1-Phosphate Uridyl Transferase (E.C. 2.7.7.12)

Enzyme

ATP

ATP

P-Source

AcP/AcK

AcP/AcK AdK

Cofactor regeneration

[I081

[lo81

[971

References

918

I

73 Formation and Cleavage of P - 0 Bonds

13.2.3

Tables Containing Typical Examples Ordered According to the Classes of Compounds

Sugars, nucleosides and their analogs are the classes of compounds most often involved in enzyme catalyzed phosphorylation. Typical carbohydrate phosphorylations are included in Table 13-4, together with the phosphorylation of other nonnucleosidic compounds. Table 13-5 gives an overview of the enzyme catalyzed phosphorylation reactions of nucleosides and their analogs. A few representative examples of nucleoside sugars are listed, for more detailed information consult the review. refs [74, 'l].

13.3

Cleavage of P - 0 Bonds

In vivo, cleavage of P - 0 bonds are performed by enzymes such as phosphatases, phosphodiesterases, phosphohydrolases, nucleases, DNases and RNases (see Section 13.1.1). In vitro, cleavage of a P - 0 bond is often a trivial synthetic step. Even for an easy step, enzymes attract increasing attention. The enzymatic reactions are preferred when regio- or stereoselectivity is required, and when the substrates are temperature or pH sensitive. Many phosphate analogs have been tested as substrates of enzymes that hydrolyze phosphoryl groups. These analogs are often accepted as substrates for the enzymes, and such reactions could be synthetically valuable. Typical examples are presented in the tables. Table 13-6.

-

Hydrolysis of phosphate and pyrophosphate monoester.

R-OH R-O-PO:R-O-P(OP-)-PO;- --t R-OH Entry

1

R-O

Enzyme

References

Polyprenol (phosphates and pyrophosphates)

Acid Phosphatase (E.C. 3.1.3.2) or Alkaline Phosphatase (E.C. 3.1.3.1)

1110-1161

2

Acid Phosphatase (E.C. 3.1.3.2) or Alkaline Phosphatase (E.C. 3.1.3.1)

[l1'1

Acid Phosphatase (E.C. 3.1.3.2)

[1181.see a ~ s 0 [ 1 1 9 - w

KDO 8-Phosphate Phosphatase

[1231

q0 OH

4

HO

H

13.3 Cleavage ofP-0 Bonds

I

919

(cont.).

Table 13-6. Entry

R-O

5

Enzyme

References

5'-Ribonucleotide phosphohydrolase

11241

(E.C. 3.1.3.5)

I

HO

I

OH

Alkaline Phosphatase (E.C. 3.1.3.1) or Acid Phosphatase (E.C. 3.1.3.2)

[1031

Alkaline Phosphatase (E.C. 3.1.3.1)

I1041

Alkaline Phosphatase (E.C. 3.1.3.1) or Acid Phosphatase (2-Phases System) (E.C. 3.1.3.2) Alkaline Phosphatase (E.C. 3.1.3.1)

[ I 2 5 1261

(1271

0 in 3 or 6

10

9

0-7-00-

Inorganic Pyrophosphatase (E.C. 3.6.1.1)

PSI

13.3.1 Hydrolysis of Phosphate and Pyrophosphate Monoesters

Both acid and alkaline phosphatases have been used to cleave aliphatic and aromatic phosphate monoesters. Table 13-6 shows typical examples ordered according to the substrate class. This table includes an example where the enzymatic reaction was run with a sensitive substrate (entry l), and examples where regio- or a stereoselectivity was required (entries 2 and 5, respectively). Polyprenyl phosphates and pyrophosphates have been hydrolyzed by acid and alkaline phosphatases (Table 13-6, entry 1). For this hydrolysis, classical chemical methods are inadequate as the reaction products decompose under acid conditi~n~[~~~]].

920

I

13 Formation and Cleavage ofP-0 Bonds

A regioselective dephosphorylation was used in the synthesis of 2'-carboxy-~arabinitol 1-phosphate (Table 13-6, entry 2), a natural inhibitor of ribulose 1,sbisphosphate carboxylase. Either acid or alkaline phosphatases can be used for the 1,sselective hydrolysis of the 1-phosphoryl group of 2'-carboxyl-~-arabinitol bisphosphate. With acid phosphatase, the conversion was essentially quantitative yielding exclusively the 1-phosphate derivative (cleavage of the 5-phosphoryl group). On the other hand, hydrolysis with alkaline phosphatase gave a 4 : 1mixture of the 1and 5-phosphate derivatives. Many natural and unnatural monosaccharides have been prepared by aldolase catalyzed condensation. The synthesized sugars were often dephosphorylated in situ by an acid phosphatase (Table 13-6,entry 3). These reactions illustrate multienzyme synthesis. In this case, no isolation of the phosphate intermediate is required: both enzymatic reactions are run in the same pot after adjustment of the pH value. One of the best examples of an enzymatic dephosphorylation for a synthetic purpose is shown in the entry 5 ofTable 13-6.A 5'-ribonucleotidephosphohydrolase was used in the synthesis of (-)-aristeromycin, a carbocyclic analog of adenosine. The (-)-enantiomer of aristeromycin shows some cytostatic and antiviral activity, while the (+)-enantiomeris inactive. The racemate (*)-5'-phosphorylatedaristeromycin was resolved by selective hydrolysis of the (-)-enantiomer with the hydrolase. The (-)-alcohol and the (+)-S'-phosphatederivative were separated easily on a silica gel column. Hydrolysis of the (+)-enantiomer with calf intestinal phosphatase yielded pure (+)-alcohol. Phosphorylated p-nitrophenol was hydrolyzed with an alkaline phosphatase['29]l. This hydrolysis was also performed in a two-phase system with an acid phosphatase [lzsI. The naphtol derivative, Table 13-6, entry 9, is dephosphorylated by an alkaline phosphatase. The resulting naphtol decomposes with chemiluminescent emission and can be used in bioassays to generate a chemiluminescence signal proportional to the concentration of an alkaline phosphatase label. Inorganic pyrophosphate may be considered as a particular case of a phosphate monoester. The enzymatic decomposition of pyrophosphate by inorganic pyrophosphatase (Table 13-6, entry 10) can be used to drive a multienzyme synthesis (see[351). 13.3.2

Hydrolysis of 5- and N-substituted Phosphate Monoester Analogs

Enzymatic hydrolysis of oligonucleotide-analogs containing modified phosphoryl moieties have been examined extensively to study their resistance to the enzymatic hydrolysis. Thiophosphates (Table 13-7) were subjected to hydrolysis with both acid and alkaline phosphatases. Most authors claimed that these compounds are substrates for alkaline phosphatases, but the reaction rate is much lower than with the N e ~ m a n n [ ~ ~however, '], reported that these corresponding phosphates [12'. same S-substituted analogs are resistant to alkaline phosphatases but hydrolyzed by acid phosphatases.

73.3 Cleavage of P - 0 Bonds

-

Hydrolysis ofthiophosphates.

Table 13-7.

R-O-PS0;-

R-OH

Entry

R-O

Enzyme

References

1

H,C-0

Alkaline Phosphatase (E.C. 3.1.3.1)

[12'.

Alkaline Phosphatase (E.C. 3.1.3.1) or Acid Phosphatase (E.C. 3.1.3.2)

(126, 1291

Alkaline Phosphatase (E.C. 3.1.3.1)

[1281

2

D

N

=

N

e

O

1291

Only the alkaline phoshatases have been used with phosphorothioates (Table 13-8).The presence of sulhr between the phosphoryl moiety and the residue does affect the enzymatic reaction with alkaline phosphatases. Imidodiphosphates are also potential substrates for phosphoryl hydrolyzing enzymes (see Table 13-8, entry 7). They have been used less often than the S-substituted phosphate analogs. Another goal of these studies involving analogs with modified phosphoryl groups or isotopicallylabeled nucleotides was mechanistic elucidation of the stereochemical course of the r e a c t i ~ n I *13',~ ~1331. Table 13-8.

-

Hydrolysis of phosphorothioates and irnidodiphosphates.

R-S-POf R-SH R-NH2 R-NH-PO:--+ Entry

R-S or R-NH

Enzyme

References

1

HZN-CH,-CH2-S

Alkaline Phosphatase (E.C. 3.1.3.1)

[12g1

Alkaline Phosphatase (E.C. 3.1.3.1)

[lZ9l

Alkaline Phosphatase (E.C. 3.1.3.1)

(1291

Alkaline Phosphatase (E.C. 3.1.3.1)

[1301

2

?

H3C-C-NH-CH,-CH,-S -0OC-CHZ-CH2-S

3

4

5

1

co,

=to;

Alkaline Phosphatase (E.C. 3.1.3.1) or Pyruvate Kinase (E.C. 2.7.1.40)

[1301

Alkaline Phosphatase (E.C. 3.1.3.1)

[961

Alkaline Phosphatase (E.C. 3.1.3.1)

[l3lI

S

6

fS OH

7

R a

Ae 0-p-0-p-NH 0-

HO

OH

0-

I

921

922

I

13 Formation and Cleavage of P - 0 Bonds

13.3.3 Hydrolysis of Phosphate and Phosphonate Diesters 13.3.3.1

Nucleic Acids and their Analogs

Endo- and exonucleases have been used successfully with nucleic acids and their analogs for organic synthetic purposes. For example, ATP was synthesized from AMP for use in cofactor recycling (Table 13-9, entry 1).The AMP was obtained from yeast RNA by cleavage with the nuclease P1 yielding a mixture of nucleoside monophosphates [lo*].In another report[73],nucleoside diphosphates were obtained by hydrolysis of RNA with nuclease PI and a polynucleotide phosphorylase (the diphosphates are preferred because the diphosphates were more easily transformed to the nucleoside triphosphates than the monophosphates). Similarly, dATP was synthesized from dAMP, obtained by cleaving herring sperm DNA with DNase I and nuclease PI (Table 13-9, entry 2). Selective phosphorylation was obtained with adenylate kinase in the presence of pyruvate kinase and phosphoenol pyruvate. Synthetic oligonucleotide analogs are interesting in applications in which they suppress translation of mRNAs by hybridization (antisense technology). A good antisense agent would be resistant to nucleases, and able to maintain its biological activity for substantial periods in living organisms [13'1. Oligonucleotide analogs modified at the phosphodiester linkage with a phosphorothioate group are the subject of numerous papers (see Table 13-9). Other oligonucleotide analogs have been tested as substrates for endo- and exonucleases. The natural substrates were modified at either the base residues (Table 13-9, entry 4) or at the sugar moieties (Table 13-9, entries 5, G and 7). The tetraphosphate Ap& and its analogs are other examples of a cleavage of a phosphodiester (Table 13-9, entry 8). 13.3.3.2 Other Phosphate and Phosphonate Diesters

Enzymes have often been used as mild catalysts to hydrolyze phosphate and phosphonate diesters. Cyclic phosphate diesters can be hydrolyzed selectively with RNases and phosphodiesterases to give the corresponding phosphate monoesters (Table 13-10,entries 1 and 2). Phosphodiesterases have been used to deprotect phosphonate diesters (Table 13-10, entries 3-5). This method is especially useful for sensitive compounds (see Table 13-10, entry 6: a P - 0 bond could be cleaved selectively in the presence of a P N bond).

13.3 Cleavage o f P - 0 Bonds Table 13-9.

Hydrolysis of nucleic acids, nucleosides and their analogs.

Entry Starting material

Product

Enzyme

References

RNA

Nucleoside Monophosphates or Nucleoside Diphosphates

[lol,731

denatured DNA

Deoxy Nucleoside Monophosphates

Nuclease P 1 (E.C. 3.1.30.1) or Nuclease P1 (E.C. 3.1.30.1) and Polynucleotide Phosphorylase (E.C. 2.7.7.8) DNase I (E.C. 3.1.21.1) Nuclease P1 (E.C. 3.1.30.1)

P41

Phosphorothioate Substituted Nucleic Acids

Endo- (E.C. 3.1.30.1)and Exonudeases (E.C. 3.1.4.1) (E.C. 3.1.16.1)

[lo5,

Nucleotide Analogs Containing Modified Bases

Restriction Endonucleases

[14'1

Nucleic Acid Analogs Containing r-Ribose

Exonucleases

11421

cytidine + allo-uridine RNase A (E.C. 3.1.27.5) 6'-phosphate RNase T2 (E.C. 3.1.27.1) Nuclease S 1 (E.C. 3.1.30.1)

0

OH

l1O31

?

Lo-

HO

13+1401

OH

Nucleotide Analogs Containing Acyclic Sugar Analogs

Nucleases Phosphodiesterases (E.C. 3.1.16.1)

1143. 1441

Thiophosphate Analogs of APPPPA

Ap4 Hydrolases (E.C. 3.6.1.17) (E.C. 3.6.1.41)Ap.+A Phosphorylase (E.C. 2.7.7.53)

[145, 1071

13.3.4

Other P - 0 Bond Cleavages

Phosphate and phosphonate esters can also be cleaved enzymatically to give products different from those obtained by enzymatic hydrolysis. The formal migration of a phosphoryl group between the CG and the C1 of glucose is catalyzed by phosphoglucomutase. Mechanistic studies were performed with the

I

923

924

I

13 Formation and Cleavage of P - 0 Bonds

Hydrolysis of phosphate and phosphonatediesters.

Table 13-10.

R-O-PO,Ri--* R-CRi-PO3R2 Entry

-

R-O-PO3H R' R-CRi-PO3Hz or R-CRi-PO3HR" ~

Starting Material

Product

Enzyme

Acoa

1

~

~~

References

RNase TI (E.C. 3.1.27.3) and RNase Tz (E.C. 3.1.27.1)

OH

o=p-00-

2

HzF-YH-CHzB

H,F-FH-CH,B

9P

OH

A

RNases or Phosphodiesterase

11471

o=y-o-

0 0-

0-

B = Base 3

0

II

R = R' = H

Phosphodiesterase I (E.C. 3.1.4.1)

R =H

Phosphodiesterase I (E.C. 3.1.4.1)

R= H

Phosphodiesterase I (E.C. 3.1.4.1)

R=H

Phosphodiesterase I (E.C. 3.1.4.1)

(EtO),HC-CH=CH--CH,-P-OR'

l4'I

I

OR

R = R' = Et 4

TFA-Ala.AspNH. CHCOOEt I

CHFH CHzP03R,

R = Et

5 R2°3P

L

(1481

4 '

11501

N PN7P03R, 0

R = Et [1481

R = Et

thiophosphate analog of glucose 6-phosphate 16. In the presence of phosphoglucomutase, this analog yields 6-thioglucose 1-phosphate 17; albeit at a slower rate than the natural substrate (Figs. 13-10). Aminolysis of phosphonate diester derivatives have been used to form organophosphorus analogs of peptides (18) with phosphatases and phosphodiestera~~

ses[151,

1521

The equilibrium between phosphoenolpyruvate and phosphonopyruvate (19, Fig. 13-10)is catalyzed by a phosphomutase. The mechanism of the transformation of a phosphoryl into a phosphonoyl group has been studied with labeled and Ssubstituted analogs of the natural substrate [153-1581.

13.3 Cleavage o f P - 0 Bonds

&

s ~ o ~ ~ Phosphoglucomutase

-

HO &OH OH

HO

17

16

YOOEt AcNH-FH ?i3 H2C-CHzr- OEt 0

HO op032-

Phosphodiesterase 4

OP032A C O O

YOOEt

AcNH-FH

cH3

H,C-CH~~-N~H

qH3

CH~-CH2~r-NHCHzC0OEt 0 18

9

Phosphomutase *

o -'3pJ ,-(

coo19

Figure 13-10. hydrolysis.

P - 0 bond cleavages with hydrolytic enzymes, not leading t o the products o f

Figure 13-11. Phosphorylase catalyzed formation of polysaccharides and modified polysaccharides. i) phosphorylase.

Numerous analogs of carbohydrate polymers (i.e., amylose, glycogen) have been prepared from modified monosaccharide 1-phosphateswith phosphorylase (Fig. 1311 shows the natural substrates) 1159-1621. Abbreviations AcK: acetate kinase; AcP: acetyl phosphate; AdK adenylate kinase; AP,,A: pl,pndi(adenosine 5'-) n-phosphate; ARS: aminoacyl tRNA synthetase; ATP, ADP, AMP: adenosine 5'-tri-, di-, monophosphate; ATP-a-S: (&)-adenosine 5 ' - 0 - ( I-thiotriphosphate), ATP-y-S: adenosine 5'-0-(3-thiotriphosphate); CK carbamyl kinase; CP: carbamyl phosphate; CrK: creatine kinase; CTP, CDP, CMP: cytidine Sl-tri-, di-, monophosphate; dATP, dAMP: deoxyadenosine S'-tri-,monophosphate; DNA: deoxyribonucleic acid; AG: change in free energy; GK glycerol kinase; GTP, GDP, GMP: guanosine 5'-tri-, di-, monophosphate; HK: hexokinase; IUB: International Union of Biochemistry; MCP: methoxycarbonyl phosphate; NTP, NDP, NMP: nucleoside Sl-tri-, di-, monophosphate; PC: phosphocreatine; PEP: phosphoenol pyruvate; Pi: orthophosphate; PK: pyruvate kinase; P,: polyphosphate; P,K poly-

I

925

926

I

13 Formation and Cleavage o f f - 0 Bonds

phosphate kinase; PPi: pyrophosphate; PRPP: S-phospho-~-ribosyla-l-pyrophosphate; RNA: ribonucleic acid; tRNA: transfer RNA; RK: ribokinase; RTP, RMP: ribavarin tri-, monophosphate; U: one unit: the amount of enzyme that catalyzes the formation of 1 pmol/minute; UTP, UDP, UMP: uridine Sl-tri-, di-, monophosphate.

References

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13 Formation and Cleavage of P - 0 Bonds

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30,4978-4987.

100 J. Bolte, G. M. Whitesides, Bioorg. Chem.

1984, 12,170-175. H.J. Leuchs, J.M. Lewis, V.M. Rios-Mercadillo, G. M. Whitesides, J . Am. Chem. Soc. 1979,101,5829. 102 P.A. Frey, Stud. Org. Chem. (Amsterdam) 1985,20,465-480. 103 A. Billich, U. Stockhove, H. Witzel, Nucleic Acids Res. 1983, 11, 7611-7624. 104 R. Stolarski, J. M. Ciesla, D. Shugar, 2. Natuforsch. C: Biosci 1990,45, 293-299. 105 B.V. L. Potter, F. Eckstein, J . Biol. Chem. 2984,259,14243-14248. 106 R. R. Breaker, G. R. Gough, P.T. Gilham, Nucleic Acids Res. 1990, 18, 3085-3086. 107 A. Guranowski, G. Just, E. Holler, H. Jakubowski, Biochemistry 1988,27, 2959-2964. 108 D.R. Walt, M.A. Fineis, V. M. Rios-Mercadillo, J. Auge, G. M. Whitesides, J . Am. Chem. Soc. 1984, 106,234-237. 109 H. G. Davies, R. H. Green, D. R. Kelly, S. M. Roberts, Biotransformations in Preparative Organic Chemistry, Academic Press, London, 1989, p. 75-86. 110 H.Fujii, T. Koyama, K. Ogura, Biochim. Biophys. A d a 1982,712,716-718. 111 J. F. Wedgwood, J. L. Strominger,]. Bid. chem. 1980,255, 1120-1123. 112 G. S. Adrian, R. W. Keenan, Biochim. Biophys. Acta 1979, 575, 431-438. 113 G.R. Daleo, R. P. Lezica, FEBS Lett. 1977, 74,247-250. 114 E. Willoughby, Y. Highasi, J. L. Strominger, J. Biol. Chem. 1972,247, 5113-5115. 115 T.Koyama, H. Inoue, S. Ohnuma, K. 0gura, Tetrahedron Lett. 1990, 31, 4189-4190. 116 S. Ohnuma, T. Koyarna, K. Ogura, Tetrahedron Lett. 1991, 32, 241-242. 117 S. Gutteridge, G.S. Reddy, G. Lorimer, Bio6hem.J. 1989, 260,711-716. 118 M.D.Bednarski, E.S. Simon, N. Bischofberger, W.-D. Fesner, M.-J. Kim, W. Lees, T. Saito, H. Waldmann, G. M. Whitesides, J. Am. Chem. Soc. 1989,111,627-635. 119 K.K.-C. Liu, T.Kajimoto, L. Chen, 2. Zhong, Y. Ichikawa, C. H. Wong,J. Org. Chem. 1991,56,6280-6289. 120 S.T. Allen, G. R. Heintzelman, E. J. Toone, J. Org. Chem. 1992,57,426-427. 121 A. Straub, F. Effenberger, P. FischerJ. Org. Chem. 1990,55, 3926-3932. 101

References I929 122

E. J. Toone, G. M. Whitesides, A C S Symp.

Ser. 1991,466, 1-22. 123 T. Baasov, A. Jakob,J . Am. Chem. SOC.1990, 112,4972-4974. 124 P. Herdewijn, J. Balzarini, E. De Clercq, H. Vanderhaeghe,J. Med. Chem. 1985,28, 1385-1386. 125 M. Cantarella, L. Cantarella, F. Alfani, Enzyme Microbl. Technol. 1991, 13, 547553. 126 R. Breslow, I. Katz,]. Am. Chem. SOC.1968, 90,7376-7377. 127 B. Edwards, A. Sparks, J. C. Voyta, R. Strong, 0. Murphy, I. Bronstein, ]. Org. Chem. 1990,55,6225-6229. 128 M. Mushak, J.E. Coleman, Biochemistry 1972, 1 I , 201-205. 129 H. Neumann, ]. Biol. Chem. 1968,243, 4671-4676. 130 K. D. Sikkema, M. H. O’Leary, Biochemistry 1988,27,1342-1347. 131 R. G. Yount, D. Babcock, W. Ballantyne, D. Ojala, Biochemistry 1971, 10, 2484-2489. 132 J. B. Jones, Tetrahedron 1986, 42, 3351-3403. 133 S. Mehdi, J.A. Gerlt, Biochemistry 1984,23, 4844-4852. 134 A. W. Nicholson, K. R. Niebling, P. L. McOsker, H. D. Robertson, Nucleic Acids RCS. 1988, 16,1577-1591. 135 K. Shinozuka, T. Morita, Y. Hirota, H. Sawai, chem. Lett. 1991,1941-1944. 136 K. Kariko, S. W. Li, W. J. Sobol, L. Suhadolnik, N. L. Reichenbach, R. J. Suhadolnik, R. Charubala, W. Pfeiderer, Nucleosides Nucleotides 1987, 6, 173-184. 137 P. M. J. Burgers, F. Eckstein, Biochemistry 1979,18,592-596. 138 M. Koizumi, E. Ohtsuka, Biochemistry 1991, 30, 5145-5150. 139 H. P. Vosberg, F. Eckstein, ]. Biol. Chem. 1982,257,6595-6599. 140 S. Spitzer, F. Eckstein, Nucleic Acids Res. 1988, 16 11691-11704. 141 J. W. Bodnar, W. Zempsky, D. Warder, C. Bergson, D. C. Ward, ]. Biol. Chem. 1983, 258,15206-15213. 142 M. J. Damha, P. a. Giannaris, P. Marfey, L. S. Reid, Tetrahedron Lett. 1991, 32, 2573-2576.

K. Augustyns, A. Van Aerschot, A. Van Schepdael, C. Urbanke, P. Herdewijn, Nucleic Acids Res. 1991, 19, 2587-2593. 144 A. Holy in Phosphorus Chem. Directed Biol., Lect. rnt. Symp., Meeting Date 1979 (Ed.: S. J. Wojcieck), Pergamon Press, Oxford, 1980, 53-64. 145 D. Lazewska, A. Guranowski, Nucleid Acids Res. 1990, 18,6083-6088. 146 A. Holy, Collect. Czech. Chem. Commun. 1970,35,3686-3711. 147 A. Holy, G. S. Ivanova, Nucleic Acids Res. 1974, I, 19-34. 148 I.A. Nachev, Tetrahedron 1988,44, 1511-1522. 149 I.A. Nachev, Phosphorus Sulfir 1988, 37, 133-141. 150 I.A. Nachev, Synthesis 1987,1079-1084. 151 I. Nachev, Tetrahedron 1991,47, 12391248. 152 I. Nachev, Bull. Chem. Soc.]pn. 1988, 61, 3711-3715. 153 H. Nakashita, A. Shimazu, H. Seto, Agnc. Biol. Chem. 1991,55, 2825-2829. 154 P. M. Cullis, R. Misra, ]. Am. Chem. SOC. 1991, 113,9679-9680. 155 M. S. McQueney, S. L. Lee, E. Bowman, P. S. Mariano, M. D. Dunaway,]. Am. Chem. SOC. 1989, 11 1,6885-6887. 156 S. Freeman, H. M. Seidel, C. H. Schwalbe, J. R. Knowles, ]. Am. Chem. SOC.1989, 111, 9233-9234. 157 M. S. McQueney, S. L. Lee, W. H. Swartz, H. L. Ammon, P. S. Mariano, D. DunawayMariano,]. Org. Chem. 1991,56, 7121-7130. 158 E. Bowman, M. McQueney, R. J. Barry, D. Dunaway-Mariano,]. Am. Chem. SOC. 1988, 110,5575-5576. 159 C. Niemann, R. Nuck, B. Pfannemiiller, W. Saenger, Carbohydr. Res. 1990, 197, 187-196. 160 S.G. Withers, Carbohydr. Res. 1990, 197, 61-73. 161 J. Zemek, S. Kucar, J. Zamocky, Eur.]. Biochem. 1978,89,291-295. 162 M. Kitaoka, T. Sasaki, H. Taniguchi, Agric. Biol. Chem. 1991,55, 1431-1432. 143

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

I

14 Formation of C-C Bonds Chi-Huey Wong

14.1 Aldol Reactions

The aldol reaction is one of the most powerful methods for carbon-carbon bond formation, and its catalytic asymmetric variants have great potential in contemporary organic synthesis [‘I. Aldolases are enzymes which catalyze reversible and irreversible asymmetric aldol condensations in r ~ a t u r e [ ~ via - ~ ]one , of two distinct reaction mechanisms 1‘1. Type I aldolases activate the donor/nucleophilic substrate via Schiff base formation with an active-sitelysine residue. These enzymes are predominantly found in animals and higher plants, and do not require metal cofactors. Type I1 aldolases activate both donor and acceptor substrates via chelation to an active-site Zn”, and are found mainly in microorganisms. Aldolases can be conveniently classified into groups according to their natural donor substrates, i. e. dihydroxyacetone phosphate (DHAP), pyruvate/phosphoenol pyruvate (PEP), glycine, acetaldehyde, and a small number of other molecules. The ability of aldolases to accept a variety of unnatural acceptor substrates, and to generate new stereocenters of known absolute and relative stereochemistry reliably, has made them powerful tools for asymmetric synthesis. 14.1.1

DHAP-Utilizing Aldolases 14.1.1.1

Fructose 1,dDiphosphate (FDP) Aldolase (E.C. 4.1.2.13)

FDP aldolase catalyzes the reversible aldol addition reaction of DHAP and Dglyceraldehyde3-phosphate (D-G~Y 3-P) to form D-FDP(Fig. 14.1-1).The equilibrium constant for this reaction has a value of - lo4 M-’ in favor of FDP formation. The enzyme has been isolated from a variety of eukaryotic and prokaryotic sources, both in type I and type I1 forms [7-211. Generally, the type I FDP aldolases exist as tetramers (M. W. - 160 KDa), while the type I1 enzymes are dimers (M. W. - 80 KDa). For the

931

932

I

74 Formation of C-CBonds 0 PO&OH

+

H+op

FDPaldolase c _

OH DHAP (PO = phosphate)

Figure 14.1-1.

D-G’y3-p

OH OH D-FDP

Aldol addition reaction catalyzed in uivo by FDP aldolase.

type I enzymes there is a high degree of sequence homology (<SO%), with the active site residues being highly conserved through evolution[12-22]. However, significant differences identified in the C-terminal regions may control substrate specificity[221. No sequence homology between type I and type I1 aldolases, or between different type 11 enzymes, has been identified. Mechanistic studies have mainly been carried out on FDP aldolases from rabbit muscle (RAMA)[231 and yeast[241,and the X-ray structures of the enzymes from rabbit muscle (2.7 A and human muscle (3 A resolution) [251 have been determined. Some of the type I aldolases are commercially available, inexpensive, and have useful specific activity (-GO U mg-I). These enzymes are not particularly air-sensitive,though there is an active site thiol group. The free enzyme has a half-life of - 2 days in aqueous solution at pH 7.0[26r 271, but this is lengthened by immobilization or enclosure in a dialysis membrane. The type I aldolase from rabbit muscle has been cloned and expressed in E. c ~ l i [ ~The l . equivalent enzyme from Staphylococcus carnosus is much more stable The type I1 aldolases from several microbial sources have for synthesis recently been cloned and overexpressed[”. 27s 29. 301 . D espite the small degree of homology in primary sequence between the enzymes from E. coli and rabbit muscle, studies have shown that they possess almost the same substrate To date, FDP aldolase, especially the commercially available RAMA, is the most widely-used aldolase in organic synthesis. A few studies which compare the stability and lunetic parameters of RAMA vs. bacterial fmctose-1,G-bisphosphate aldolases 331, and FDP aldolase from spinach leaves has also been have been employed for synthesis purposes. RAMA accepts a wide range of aldehyde acceptor substrates, with DHAP as the donor, to generate vicinal diols with D-threo stereochemistry reliably L5. 26, 27, 35-441. Suitable acceptors include unhindered aliphatic and a-heteroatom substituted monosaccharides, and derivatives thereof 1441. Aromatic, sterically hindered aliphatic, and a, p-unsaturated aldehydes are generally not substrates [261. The specificity for the donor substrate is much more stringent. Initially, only three DHAP analogues were shown to be substrates, but they were so weak (- 10% cf. DHAP),that their general use in organic synthesis was 451. However, recently, a DHAP phosphonate analog has been shown to be a good substrate for FDP aldolases from rabbit and S. carnosus, as well as Rha 1-P aldolase from E. c 0 l i [ ~ ~ 1 . FDP aldolase exhibits kinetic diastereoselectivity with unnatural chiral aldehyde acceptor substrates. However, even though there is significant discrimination (- 20 : 1)between the D- and L-enantiomersof the natural substrate Gly 3-P[261, this is usually not the case with unnatural aldehydes. In fact, resolutions of racemic aldehydes are normally only successful if carried out under thermodynamic control. Often the aldol products can cyclize via formation of a hemiketal, leading to

74.7 A h / Reactions 1.0,

NHAc

2.reductlon

933

I

1 FDP A. DHAP 2 Pase 3 Hz,PdC

0

,LNHA, HO NHAc

i 3

P

h

v

N

H

2

N3

PhvNHAc HbNHAc 1. FDP A, DHAP 2. Pase

2. reduction

3. Hz, Pd/C

N3

Figure 14.1-2.

*

N3

NHAc

Preparation o f optically active aldehyde acceptors for FDP aldolase.

significant energy differences between the two diastereomeric products, and ultimately favoring one product after equilibration. For example, with racemic phydroxybutyraldehyde[26, 371 as a substrate, only a single diastereomer was obtained, with the methyl group in the more stable equatorial position. Synthetically, FDP aldolase has been employed in the production of I3C-labeled r3’, 46, 471, nitrogen-containingr27* 38-40], deoxy-[3’-371, fluoro-136, 481, and high35, 37, 431. Most of these syntheses require the preparation of the carbon sugarsL342 aldehyde acceptor. In cases where the aldehyde is optically active, this necessitates either asymmetric synthesis of the required enantiomer, or use of a racemic aldehyde, with subsequent separation of diastereomeric products. In general, ozonolysis of a terminal olefin (Fig. 14.1-2)r4’1 and acid-catalyzedacetal deprotection are convenient routes to the acceptor aldehydes. a-Chiral aldehydes have also been prepared by ring opening of readily-available ( R ) -and (S)-glycidaldehydeacetal, or the corresponding thirane and aziridine, by appropriate nucleophiles c4’1. Both enantiomers of glycidaldehyde acetal may be prepared by lipase-catalyzedresolution of 3-chloro-2-hydroxypropanal diethyl acetal r4’1. Alternatively, tandem use of Sharpless asymmetric dihydroxylation (AD) and aldolase-catalyzed condensation allows quick and facile synthesis of carbohydrates with complete stereocontrol (Fig. 14.13 ) [SO]. A 1 : 4 mixture of deoxynojirimycinand deoxymannojirimycin was obtained when was used as a substrate for RAMA [38, 391, indicatracemic 3-azido-2-hydroxypropanal ing that the D-aldehyde is a better substrate for the enzyme. A similar result was obtained with FDP aldolase from E. c0lir~~1.Since both deoxynojirimycin and ., .o.

Sharpless

EtO

N,

1. FDP A, DHAP 2. Pase

HOfiOR epoxidalion ~0-0~ 3.Hz. Pd/C ~

-

: H

HO

OH

OH

.o. .~

HO

Porcine Pancreas Lipase

.o. ..

. nOH i

HO A

O

C

O

R

-

LoH

EtO El0

1.FDPA.DHAP

I

OH

R = protecting group, R‘ = H, Bn. or OBn

Figure 14.1-3.

Chernoenzyrnatic stereo-controlled synthesis o f azasugars.

2.Pase 3. H, Pd/C

*

HO

0%

&&NR HO

934

I

74 Formation of C-CBonds

+

CH3

co2

HO HO

OH DAHP

H HO (+)-exo-brevicomin

a

o

H

6H - . cyclilol

H%&

OH

1-deoxynojirimycin

+H r HO

OH

aza-suaar analoa of ManNAc

C-glycosides

Figure 14.1-4.

Various classes o f m o l e c u l e s synthesized u s i n g FDP aldolase.

deoxymannojirimycin are potent glycosidase inhibitors, each compound was also prepared in an optically pure form from the respective optically pure azidoaldewere obtained via LP-80 catahydes L2'1. Both (R)-and (S)-azido-2-hydroxypropanal lyzed resolution of the racemic acetal precursor .]'41 Similar strategies have been employed to prepare the P-glycosidase inhibitors P-1-homonojirimycin,P-1-homomannojirimycin and the azasugars corresponding to N-acetylglucosamine and N-acetylmannosamine(Fig. 14.1-4)[491. Similarly, employing 2-azidoaldehydes as RAMA substrates allowed the preparation of polyhydroxylated pyrrolidines (Fig. 14.1-5)[38, 52, 531. 1,4-Dideoxy-l,4-iminoD-arabinitol was synthesized from azidoacetaldehyde, and both (2R,5R)- [491 and (2S,5R)-bis(hydroxymethyl)-(3R,4R)-dihydro~~olidine were synthesized from racemic 2-azido-3-hydroxypropana1, respectively. In the latter case, the kinetic product of the aldol addition was transformed into the (ZR,SR)-stereoisomerof the pyrrolidine, while the thermodynamic product gave the (2S,SR)-stereoisomer.Furthermore, pyrrolidines structurally related to GlcNAc have been prepared stereoselectively by a similar transformation from lipase-resolved aldehyde precursors [541. H

2. Pase

OH thermodynamic product

kinetic product

Figure 14.1-5.

Synthesis of polyhydroxylated pyrrolidines u s i n g RAMA.

HO

O 'H

1. AcSK AcSH

'u

1. DHAP, RAMA

___)

EtO EtO&

2. HCI

'0

*

Et,SiH, BF3*Etz0

P

AcO OAc

Figure 14.1-6.

OH

A,..& AcO

HO OAc

2. Pase

14.1 Aldol Reactions

..JGJS" OH

0

-..

-..

OH

OH

I

935

tl ~

HO HO OH

Preparation of deoxy-thio sugars.

The 6-deoxyazasugars and their analogs can also be easily prepared by direct reductive amination of the aldol products prior to removal of the phosphate group rS51. Studies using glucose 6-phosphate (Glc 6-P) indicate that the phosphate group is probably reductively cleaved from the imine 6-phosphate rather than the azasugar 6-phosphate. Use of 3-azido-4-hydroxy aldehydes results in the formation of homoaza sugars[51,"I. The optically pure aldehydes can be obtained either by Sharpless epoxidation of the olefins[51]or enzymatic resolution of the epoxides[57]. The lipase-resolvedmaterial was also used to prepare another class of glycosyl cation mimics, the tetrahydropyrimidineslsS,591. These compounds exist in equilibrium with their guanadinotetraose forms which predominate at low pH. The tetrahydropyrimidines are potent inhibitors of a-galactosidase,due to their close resemblance to the transition state half-chair conformation of the enzymatic reaction. Interestingly, an inhibitor with an OBn group attached to the nitrogen has a much lower pK, and inhibits a-galactosidasein the region of a physiological pH ["I. Similar to the synthesis of azasugars, a series of deoxy-thiosugars was prepared by aldol condensation of thioaldehydes with DHAP followed by reduction of the Regioselective ring opening of the (S)-glyciresulting thioketoses (Fig. 14.1-6)I.'[ daldehyde diethyl acetal with potassium thioacetate introduced the thio function. RAMA-catalyzed aldol condensation followed by dephosphorylation gave the corresponding thioketose[''], which was then acetylated and reduced to the 1-deoxy-5-thioAlso, in a similar manner, l-deoxy-5-thio-~-manD-glucopyranose peracetate I.'[ nopyranose was obtained from the other aldehyde enantiomer, while a Fuc-1-P aldolase-catalyzed reaction provided 1-deoxy-5-thio-galactopyranoseand l-deoxy5-thio-altropyranose, and Rha-1-P aldolase catalyzed reaction produced l-deoxyS-thio-~-mannopyranose I.'[ With other aldolases in place of FDP aldolase, a wide range of other polyhydroxylated piperidines and pyrrolidines have been synthesized (vide in@) LS31. Aldolase-catalyzed condensation followed by reductive amination has become a cyclic imine general strategy for the synthesis of 5-["], 6- LG2, G31, and 7-membered[G4] sugars. The resulting compounds have become the gold standard template for glycosidase inhibitor design. Use of racemic methyl N-acetylaspartateP-semialdehydeas a substrate for RAMA provides a precursor to 3-deoxy-~-arabino-heptulosonic acid 7-phosphate (DAHP,

936

I

74 Formation of

C-CBonds

Fig. 14.1-4) L4l]. This compound is an important intermediate in the shikimate pathway for the biosynthesis of aromatic amino acids in plants. The RAMA reaction produced the desired D-threo stereochemistry, and chemical reduction of the keto group gave the desired (GR)-stereoisomerin GO% diastereomeric excess. Other analogs of DAHP are also potentially available by this route, due to the broad substrate specificity of RAMA. The use of pentose and hexose phosphates as RAMA substrates provides a route to high-carbon sugars, including analogs of sialic acid and KDO[44,65, 66]. 0ther carbohydrate derivatives prepared by RAMA include unsaturated C8-C9 sugars [671, phosphonic acid derivatives["], fluorescently-labeled fructose derivatives["], perfluoroalkylated sugars L7O], and those protected by thioacetals L7'1. Furthermore, the S. carnosus enzyme has been employed for the synthesis of bicyclic sugars [721 and disaccharide mimetics [731. Complex xylulose structures can also be synthesized by RAMA[741. Employing a one-pot, three-enzyme system with RAMA, triose phosphate isomerase, and 1-deoxy-D-xylulose-5-phosphate synthase, l-deoxy-~-xylulose-5-phosphate could be obtained in 47% Furthermore, a four-enzyme, one-pot system employing [761. FDP-aldolasefrom S. carnosus furnished 5-deoxy-5-ethyl-~-xylulose The synthesis of (+)-exo-brevicomin(Fig. 14.1-4) was the first example ofthe use of RAMA to synthesize a non-carbohydrate RAMA was employed to catalyze the key aldol addition step, in which the two chiral centers of the target molecule were established. RAMA has also been employed for the synthesis of a key ~ I ,for that of acyclic polyols [791. Single aldol condensafragment of (+) a s p i ~ i l i n [ ~and tion on remote dialdehydes has also been achievedI.'[ Other molecules synthesized by FDP aldolase include C-glycosides[43, 1' and cyclitols (Fig. 14.1-4) LS31. Cyclitols are an interesting class of bio-active compounds, and the use of aldolases provides a chemo-enzymatic strategy towards their synthesis. An example is the synthesis of nitrocyclitolswhich was accomplished by an FDP aldolase catalyzed reaction with nitroaldehyde, followed by a non-enzymatic intra[83a1. A one-pot synthesis of cyclitols has molecular nitro-aldol reaction (Fig. 14.1-7) been reported, involving an FDP aldolase-catalyzedreaction between a phosphonate aldehyde and DHAP. The aldol product cyclized in situ via an intramolecular Horner-Wadsworth-Emmonsolefination to give the polyhydroxylated cyclopentane 1. FDP A,

O

DHAP 2. Pase

z

AcO

"2'

*

OH

-$- + ozNq-Oo;c

OzN

OAc

OAc

OAc

(1:l)

Figure 14.1-7.

*

OH

HO

c]

Ac20, BF3-Etz0

V

*

OH

;02N--f5

N

Preparation o f nitro-cyclitols.

I

14.7 Aldol Reactions 937

/

NC Figure 14.1-8. 0

NC

One-pot synthesis of cyclitols. 0

OEt

OH OEt

0

OH OH H

* E HO&~ l.IDH,NADH o t POAOH + H U O E t 1. RAMA “U s ; 2. Pase 2. H+ ~

DHAP

I

OH

OH

L-xylose NaBH(0Ac)s

OH OH OEt H

o

a O OH

E

1. IDH, NADH

t

2.H+

-

OH OH H O U O R ; OH

H

2-deoxy-Darabino-hexose

Figure 14.1-9. Use of the “inversion strategy” t o synthesize L-xylose and 2-deoxy-D-arabino-hexose.

(Fig. 14.1-8)[83b1. Using this approach, different functionalized cyclitols may become easily accessible. FDP aldolase is a useful catalyst for the direct synthesis ofketose monosaccharides and their analogs (vide suprcz). However, a number of the important naturallyoccurring carbohydrates are aldoses. Various FDP aldolase products can be isomerized to a mixture of the ketose and aldose, and subsequently separated with Ca2+ or Ba2+treated cation exchange resins. Another strategy involves the use of glucose isomerase (GI)LS41, which catalyzes the isomerization of fructose (Fru) to glucose (Glc),and is used in the food industry for the production of high fructose corn syrup. GI also accepts analogs of Fm with modifications at positions 3, 5 and 6 as substrates L3‘1. Aldose analogs including 6-deoxy, 6-fluoro, 6-0-methyl and 6-azido371. glucose have been synthesized using this FDP aldolase/GI However, not all FDP aldolase products are substrates for GI, and in the case of 5-deoxy-~-fructose,the equilibrium lies completely in the favor of the ketose. Furthermore, in the inversion strategy (Fig. 14.1-9)[421, monoprotected dialdehydes are used as substrates for FDP aldolase, generating protected aldehyde ketoses. The ketone group is then chemically or enzymatically stereoselectivelyreduced, and the aldehyde subsequently deprotected to produce the aldose. The strategy also places the vicinal diol produced in the aldol reaction in a position other than C3/C4. One enzyme suitable for the reduction is the NADH-dependant iditol dehydrogenase

938

I

14 Formation ofC-C Bonds Products prepared from FDP aldolase-catalyzed reactions with DHAP.

Table 14.1-1.

[a1

[c] R=H, [d] R = P032-

[bl

O -H PO

0

OH OH

-

. .

-

PO O -H

0

PO O -R

-

OH OH

OH OH

OH OH

[f,g] R=H, [h-k] R = P03'.

[el 0 PO&OH

OH OH

0

. . .OH OH

[h] 0

OH OH

P O + o R

OH

0

OH OH

O H +' '

OH OH

OH OH

p o + O P

poO *P OH OH OH

OH OH OH

If1

OH OH OH

[h] R=H,OH,NH2 0

0

OH OH OP

[hl

OH OH OH

PO&

R2

0

OP

[bl 0

OH

OH

OH

PO-

OH OH OH

PO

[ml 0

OH OH OH

0

OP

PO

[I1

[hl

OH

If1

[h-kl 0

OH OH

P O + o p

OH OH OH

If1

OH OH

0

OH OH

OH R2

OH

P0-R'

P

0 0A

R OH 3

~. OH R'

OH R

R'

R2

H H H H H H H H H H

THPO BzO CHJCHZ CH3 (Et0)zPO Ph

Ref.

R'

R2

OH

OH R

Ref.

R'

R2

Ref

R3

OHC(CHz), HOCH2 CbzNH CbzNHCH2

J. Gorin, J. K. N. Jones,/. Chem. Soc. 1953, 1537. b J. K. N. lones. N. K. Matheson, Can./. Chem. 1959, 37, 1754. c B. L. Horecker, P. 2. Smymiotis,j . Am. Chem. Soc. 1952,74,2123. d C. E. Ballou, H. 0. L. Fischer, D. L. MacDonald,/. a P. A.

Am. Chem. Soc. 1955.77,5967: H. A. Lardy, V. D.

Wiebelhaus, K. M. Mann,/. Biol. Chem. 1950, 187, 325. e K. N. Jones, R. B. Kelly, Can. /. Chem. 1956,34,95. f J. K. N. Jones, H. H. Sephton, Can./. Chem. 1960, 38, 753. g G . Haustveit, Carbohydr. Res. 1976,47, 164.

74. I Aldol ReaGtions r J. R. Dumachter, C:H. W0ng.J. Org. Chem. 1988, h M. D. Bednarski, H. J. Waldmann, G. M. White53,4175. sides, Tetrahedron Lett. 1986, 27, 5807. s N. J. Turner, G. M. Whitesides, J. Am. Chem. SOC. i F. P. Franke, M. Kapuscinski, J. K. MacLeod, J. F. 1989,111,624. Williams, Carbohydr. Res. 1984, 125, 177. j A. H. Mehler, M. E. Cusic Jr. Science 1967,155, 1101. t L. Hough, J. K. N. Jones,J. Chem. SOC.1952,4052. u P. Huang, 0. N. Miller,J. B i d . Chem. 1958, 330,805. k M. Kapuscinsci, F. P. Franke, 1. Flanigan, J. K. Mav C.-H. Wong, G. M. Whitesides,J. Org. Chem. 1983, cleod, J. F. Williams, Carbohydr. Res. 1985, 140, 65. 48, 3199. I F. E. Charalampous,J. B i d . Chem. 1954,211, 249. w C.-H. Wong, F. P. Mazenod, G. M. Whitesides,J. m P. A. J. Corin, L. Hough, J. K. N. Jones,J. Chem. Org. Chem. 1983,48,3493. SOC. 1953,2140. x R. L. Pederson, M. J. Kim, C.-H. Wong, Tetrahedron n M. D. Bednarski, E. S. Simon, N. Bischofberger,W: Lett. 1988,29,4645. D. Fessner, M. J. Kim, W. Lees, T. Saito, H. J. Waldmann, G. M. Whitesides, J . Am. Chem. SOC.1989, I 1 I, y T. Ziegler,A. Straub, F. Effenberger,Angew. Chem. Int. Ed. Engl. 1988,27, 716. 627. o F. Effenberger, A. Straub, Tetrahedron Lett. 1987,28, L C. H. von der Osten, A. J. Sinskey, C. F. Barbas Ill, R. L. Pederson, Y.-F. Wang, C.-H. Wong, J. Am. Chem. 1641. p A. L. Lehninger, J. Sice, J. Am. Chem. SOC. 1955,77, SOC.1989,111,3924.

I

5343.

q J. R. Dumachter, D. G. Drueckhammer, K. Nozaki, H. M. Sweers, C.-H. Wong,]. Am. Chem. SOC. 1986, 108,7812.

(IDH) from Candida utilis, also known as sorbitol or polyol dehydrogenaseL3’>851. Reduction of the ketone occurs to give the alcohol with (S)-stereochemistry.The corresponding (R)-alcohol was obtained by non-stereoselective reduction of the ketone with NaBH(OAc)3‘ and the (S)-epimer was selectively removed by IDHwere synthesized by each catalyzed oxidation. L-Xyloseand 2-deoxy-~-arabino-hexose of these two processes, respectively. Other aldose/ketose isomerases with different substrate specificity have been cloned and overexpressed[86], including Fuc isomerase (Fuc I, EC 5.3.1.3) and Rha isomerase (Rha I, EC 5.3.1.14). Fuc isomerase, in combination with Fuc 1-P aldolase or Rha 1-P aldolase, has been used to prepare L-glucose, L-galactose, L-fucose, and derivatives from the corresponding L-glyceraldehydederivatives and DHAP iS71. RAMA has been the most popular synthetic aldolase, due to its commercial availability. Notably, no significant differences in substrate specificity or stereoselectivity between FDP aldolases from different sources have been observed[88]. However, it is still important to verify this, especially for the type I1 aldolases which operate by a different mechanism. In fact, the type I1 aldolase from E. coli, which has been subcloned and o~erexpressed[~~], has the potential to supplant RAMA as the FDP aldolase of choice for synthesis. It has enhanced stability compared with RAMA (vide supra),and is available from a microbial as opposed to an animal source. Table 14.1-1 illustrates products prepared from FDP aldolase-catalyzed reactions with DHAP. 14.1.1.2

Fuculose 1-Phosphate (Fuc 1-P) Aldolase (E.C. 4.1.2.17), Rhamnulose 1-Phosphate (Rha 1-P) Aldolase (E. C. 4.1.2.1 9) and Ragatose 1,C-Diphosphate (TDP) Aldolase

Fuc 1-P aldolase, Rha 1-P aldolase, and TDP aldolase also use DHAP as the donor substrate in aldol condensation. Fuc 1-P aldolase catalyzes the reversible condensa-

939

940

I

14 Formation of C-C Bonds

0 PO&OH

+

-

0 ,,kCH3

0

FUC 1-p aldolase,

OH DHAP

OH OH

L-lactaldehyde

Figure 14.1-10.

FUC1-P

Aldol addition reaction catalyzed in vivo by Fuc 1-P aldolase.

DHAP

L-lactaldehyde

Figure 14.1-11.

Rha 1-P

Aldol addition reaction catalyzed in vivo by Rha 1-P aldolase.

0 & ‘‘OH

+

H

G

O

P

-

0

DHAP

OH

TDP aldolase *

O ‘+OP

OH D-GIY3-P

Figure 14.1-12.

OH

PO*CH3

OH OH D-TDP

Aldol addition reaction catalyzed in vivo by TDP aldolase.

tion of DHAP and L-lactaldehyde to provide L-FUC1-P (Fig. 14.1-10). With the same substrates, Rha 1-P aldolase produces L-Rha 1-P (Fig. 14.1-11). Both of these enzymes are type I1 aldolases and are found in many microorganisms rS91. Both E. coli enzymes have been cloned, overexpressed in E. coli, and purifiedI”O-’*I. TDP aldolase, a type I aldolase involved in the galactose metabolism of cocci, catalyzes the reversible condensation of D-G~Y 3-P with DHAP to give D-TDP(Fig. 14.1-12), and has also been cloned and overe~pressed[’~l. Both Fuc 1-P and Rha I-P aldolase show specificity with regard to the aldehyde component, generating vicinal diol units of D-erythro and L-threo configurations, respectively (Fig. 14.1-13) [90-931. While the stereospecificity for the absolute (3R)configuration is mechanism-based, the configuration at C4 is somewhat substrate dependent. However, these two aldolases also show significant kinetic preference for the L-enantiomer of 2-hydroxyaldehydes (c95 : 5), facilitating resolution of racemic mixtures of these compounds (Fig. 14.1-14) LS6]. Both enzymes have been used in the synthesis of rare ketose l-phosphates[86], azasugars, and deoxyazasugars[54.64, 951. Rha 1-P has also been employed in the synthesis of bicyclic carbohydrate structures L9‘l. Fuc 1-P and Rha 1-P aldolases have also been utilized in whole cell systems with DHA and catalyhc inorganic aresenate [971. With L-lactaldehydeas the substrate in the Rha 1-P aldolase reaction, the aldol product L-rhamnulose was subsequently isomerized to L-rhamnose, catalyzed by rhamnose isomerase. No such isomerization was observed with L-xylulose, the corresponding aldol product using glycolaldehyde as the substrate. Recent studies have since shown that both rhamnose and fucose isomerase require fixed stereochemistry only up to C3 for aldohexose substrates; 943

14.7 Aldol Reactions

Aldehyde substrate

Product

L-lactaldehyde c

<3 : 97

Fuc 1-P

glycolaldehyde

.

Diastereomeric Ratio trans ; cis

Aldolase

3

D-Giy

Rha 1-P

OH OH

Rha I - P

H

H

L-Gly

OH

<3 : >97

Fuc I - P

0

H

>97 : 3

a o OH OH

H

OOH GOH o

H

>97 ; <3

Fuc I - P Rha 1-P

43 100 100

28

. <3

42

c3 : 297

Fuc I - P

38

<3 : > 9 7 >97

>97 : <3

Rha I - P

Relative Rate (%)

17

41

OH 0 3-hydroxyproplonaidehyde

Fuc I - P

HO

<3 : >97 >97 : <3

Rha 1-P

OH

,,AoH

Formaldehyde

OH

Fuc I-P

44

Rha 1-P

22

.

Fuc I - P

5

95

14

OH

Rha 1-P

69

31

32

Fuc I - P

30

70

20

Rha 1-P

97

3

22

H3C=OH

acetaldehyde

11

29

ebutyraldehyde

Figure 14.1-13. Acceptor substrate specificity and diastereoselectivity o f Fuc 1-P and Rha 1-P aldolase.

FUC1-P aldolase/ HO CHO OH

+

O / H O A O P

(L)

*OH

DHAP

(L) Figure 14.1-14. Kinetic resolution of 2-hydroxyaldehydes using Fuc 1-P and Rha 1-P aldolase.

(2R,3R) for rhamnose isomerase and (2S,3R) for fucose isomerase186].A sequential application of Fuc 1-P aldolase and fucose isomerase was employed for the preparation of L-fucose analogs [981. TDP aldolase has been isolated from several sources [991. The enzyme from E. coli has a narrow pH profile with an optimum at pH 7.5, but still displays acceptable

I

941

942

I

14 Formation ofC-C Bonds

o

+

0 HO&OP

TDP aldolase

DHAP

w

OH

OH

O

L-erythro

D-threo


>go%

protection

H

1protection

J

OH

\ AD-mix-a

H

OH ,

;

"

O

0

0

OH

OH

OH

sL;H *o OH

$

OH ,..-Rs W RL

OH

O $ *H

)

bH

bH

OH

OH

+ R+oP

Diastereoselectivity of TDP aldolase.

Figure 14.1-15.

Rs R .,,, L

OH 0

OH 0

R +OP

'H

OH

OH

:

OH

6H

OH

6H

0

, OH

OH

OH

OH

/

OH

\

; ;++o H

OH

Figure 14.1-16.

0

OH

OH

OH

OH

Product stereochemistries generated by the four complementary

DHAP aldolases.

activity within pH 6.5-7. TDP aldolase accepts a variety of substrates, including glycoaldehyde, D- and L-glyceraldehyde, acetaldehyde, and isobutyraldehyde['OO]. However, a diastereomeric mixture of products is generally formed. Also, only with the natural D-substrate does the major product (D-TDP)have the tagatose configuration (Fig. 14.1-15). Owing to this lack of stereoselectivity, TDP aldolase is not as synthetically useful as the other FDP aldolases. However, with suitable protein engineering, this may change in the future. The four DHAP-utilizingaldolases generate all four stereochemical permutations of the vicinal diol at C3/C4 of the ketose product, and can be used to generate all four stereoisomers of a desired product (Fig. 14.1-16). In this manner, these enzymes have been utilized for the synthesis of sLeXmimetic side chains[81],among other targets.

I

74.7 Aldol Reactions 943

14.1.1.3

Synthesis of DihydroxyacetonePhosphate (DHAP)

All four aldolases described previously use DHAP as the donor substrate, and several approaches have been taken towards its synthesi~1'~'-~~~]. Enzymatic in situ generation of DHAP from FDP can be accomplished using FDP aldolase and triosephosphate isomerase (TI)[341. FDP aldolase catalyzes the retro-aldol reaction of FDP to y into DHAP. give ~ - G l 3-P y and DHAP, and TI catalyzes the conversion of ~ - G l 3-P However, the overall reaction may not go to completion, depending on the thermodynamic stability of the product compared with FDP. The presence of FDP may also complicate product isolation. Another enzymatic method involves glycerol kinasecatalyzed phosphorylation of dihydroxyacetone (DHA) using ATP, with in situ regeneration of ATP [34* 1"' . This procedure generates DHAP directly in high yield, but may be expensive for large-scale synthesis. Another method involves the generation of DHAP from phosphatidyl choline and DHA using phospholipases C and DI1OG]. DHAP and the corresponding phosphoramidate and phosphorothioate have been generated in situ enzymatically from the reduced form and used as substrates [104b]. DHA dimer can be phosphitylated chemically with (PhCH20)2PNEt2'and subsequently oxidized to the phosphate with H202 (Fig. 14.1-17)['021,or it can be phosphorylated directly with either POC13[lo3]or (PhO)2POC1['04a].Alternative efficient chemical syntheses of DHAP have also been These phosphate derivatives are all subsequently transformed into the stable dimer precursor of DHAP, which can easily be converted into DHAP by acid hydrolysis. The main drawback to preparing DHAP chemically is the lengthy synthetic procedure. Alternatively, DHAP can be replaced by a mixture of DHA and inorganic M-'s-') a r ~ e n a t e [971.~ ~DHA * reacts reversibly with inorganic arsenate (k - 2.4 x to form dihydroxyacetone arsenate, an analog of DHAP and a donor substrate for aldolases. In fact, in the presence of RAMA, triose phosphate isomerase, and inorganic arsenate, DHA was converted into D - F ~ uin almost quantitative yield["]. However, reaction rates tend to be very slow, and low yields of aldol products are often obtained. Inorganic vanadate and DHA has also been investigated,but cannot be successfully utilized as a DHAP analog[97]. OH

,H ,O "J

0

HO

OH EtoH'

H+

EtO$$OEt 0 HO

dihydroxyacetone dimer Figure 14.1-17.

Chemical synthesis of DHAP.

1. (PhCH20)2PNEt2, then H202

2.HzlPd

EtO$$Ilt PO 0

H+ P

O 0 A O H DHAP

944

I

14 Formation ofC-C Bonds 14.1.2

Pyruvate/Phosphoenolpyruvate-UtilizingAldolases 14.1.2.1 N-Acetylneurarninate (NeuAc) Aldolase (E.C. 4.1.3.3) and NeuAc Synthetase (E.C. 4.1.3.19)

NeuAc aldolase, or sialic acid aldolase, catalyzes the reversible condensation of pyruvate with N-acetylmannosamine(ManNAc)to form NeuAc (sialicacid; N-acetyl~-am~no-3,~-d~deoxy-~-g~ycero-~-ga~acto-2-nonu~opyranoson~c acid) (Fig. 14.118)[108, 1091.

The a-anomer of NeuAc serves as the aldolase substrate, even though the panomer predominates in solution. The initial products of aldol cleavage are aManNAc and pyruvate["o]. In uiuo, the enzyme has a catabolic function, with an equilibrium constant for the retro-aldol reaction of 12.7 M-'. However, for synthetic purposes, the production of the aldol product can be achieved using excess pyruvate['O'I. NeuAc aldolase is a Schiff base-forming type I aldolase and has been isolated from both bacteria and animals. The optimum pH for activity is 7.5, although it retains activity between pH 6 and 9, and is stable under oxygen["'. 'l']. The enzymes from Clostridia and E. coli are now commercially available (Toyobo), and that from E. coli has been cloned and overexpressed in E. c01i["~].It can be used free in solution, in immobilized f ~ r r n [ ~ ' ~,-or ~'~ I enclosed in a dialysis membrane['l']. Good conversion (-90%) of ManNAc to NeuAc has been achieved using the isolated enzyme, and purification can be achieved by decomposing excess pyruvate or acid-catalyzedesterification of the products [ll'l. with pyruvate decarboxylase['"I, The need for excess pyruvate and purification of NeuAc can be circumvented by coupling the synthesis of NeuAc to a more thermodynamically stable process. For example,the NeuAc aldolase reaction can be coupled to a sialyltransferasereaction to produce sialyloligosaccharides[I2']. Another variant of this process used a mixture of ManNAc and GlcNAc, whereby the GlcNAc was epimerized to ManNAc chemically[121.1221 or enzymatically['23.1241. Extensive substrate specificity studies have indicated that only pyruvate is acceptable as the NeuAc aldolase donor substrate"'']. However, this enzyme has broad acceptor specificity, and over s x t y aldoses have been characterized as substrates. Substitutions at C2,4 and 6 of ManNAc are allowed,with only a slight preference for absolute stereochemistry at C4, 5 and 6[11&120s125-1301 . Some pentoses and their analogs are also substrates, although 2 and 3 carbon molecules are not accepted. 0

"s OH

HO OH

NeuAc aldolase

D-ManNAc

Figure 14.1-18.

H6

*

H

OH

a-NeuAc Aldol addition reaction catalyzed in uivo

o

G

C

H6

OH

P-NeuAc

by

NeuAc aldolase.

0

2

H

14. I Aldol Reactions 0 OH

OH NeuAc aldolase

OH

OH

0

-

H C O -;

OH

OH

D-arabinose

Figure 14.1-19.

945

H?

0

HO+H

I

HO

OH

OH

KDO

Synthesis of KDO using NeuAc aldolase. n

NeuAc aldolase, pyruvate

-

0 subtilisin BPN'

*

HO OH

HO OH

1. HCI

0

2. dansyl-CI,

B

O

c

-

H

N

~

o

~ Na2C0, w

AcHN

C02H

HO OH

HO OH

/ Figure 14.1-20. Synthesis of 9-0-acetylNeuAc by combined use of subtilisin and NeuAc aldolase.

The stereoselectivity of NeuAc aldolase is unusual, as it is thermodynamically controlled. With the natural substrate D-ManNAc, attack occurs exclusively on the si face of the carbonyl group. However, a mixture of KDO and 4-epi-KDOwas obtained when D-arabinose was used as a substrate[1281, and a complete reversal of diastereoselectivity was observed with L-mannoseand 6-deoxy-~-mannose[~~'. '19, 13'1 . W'lth these latter aldehydes, exclusive attack on the re face of each carbonyl group gives the more thermodynamically favored products. NeuAc aldolase has been investigated as a catalyst for the condensation of various pentoses and hexoses, and enzyme stereoselectivity characterized['32].Furthermore, KDN has been produced on a 100 gram scale from pyruvate and D-mannose in a crystallized yield of 75 %[1331. NeuAc, derivatives thereof, and polysialic acids play important roles in cell-cell 13'1 . Th e adhesion and communication in bacterial and mammalian wide substrate specificity and ready availability of NeuAc aldolase provide the opportunity for the synthesis of many sialic acid derivatives. Azasugar analogs of NeuAc were synthesized using the 3-deoxy-3-azidoanalogs of ManNAc, mannose, and glucosamine as acceptor substrates Similarly, KDO was produced using Darabinose as the acceptor (Fig. 14.1-19). A facile synthesis of 9-0-acetyl-NeuAc has been accomplished via regioselective irreversible acetylation of ManNAc catalyzed by subtilisin, followed by NeuAc aldolase-catalyzed condensation of the resulting 6-0-acetyl-ManNAcwith pyruvateI"'1. A 9-0-glycyl-NeuAcderivative was prepared in a similar fashion, and was further converted into a fluorescent derivative (Fig. 14.1-20)[13'1. A large number of NeuAc derivatives modified at the 5-amino the C7l 4 O ] have been reported. Many other compounds or the C9-p0sition[~~",

946

I

74 Formation of C-C Bonds

R'

AcNH

Figure 14.1-21.

R2

H

R3

OH

R4 H

R5

Ref.

CHzOH, CH~OAC, CHzN,, CHzF, CH,OMe, CHzOCOCHOHCH,

5,11

12.3,

OH

H

OH

H

C W H , CHZOAC,H

7,4,10

OH

H

H

H, F

CHzOH, CHpF

83

H

H,OH

OH

H

CHzOH

10,8

Ph

H

OH

H

CHzOH

10

N,

H

OH

H

CH2OH

8

NeuAc analogs synthesized using NeuAc aldolase.

have been prepared using this aldolase (Fig. 14.1-21)[47.141-1441, including an amethyl ketoside of 5-amino-NeuA~['~~1, and polyacrylamides bearing pendant asialoside groups [1431 or C-linked sialosides ll#]. The latter strongly inhibit agglutination of erythrocytes by the influenza virus. The synthesis of NeuAc in vivo is accomplished using NeuAc synthetase["ll. This aldolase catalyzes the irreversible condensation of PEP and N-acetylmannosamine. Although this enzyme has not yet been isolated and characterized, it may prove synthetically useful in the future. 14.1.2.2

3-Deoxy-~-manno-2-ocu~osonate Aldolase (E.C. 4.1.2.23) and 3-Deoxy-~-manno-2-octulosonate 8-Phosphate Synthetase (E. C. 4.1.2.16)

3-Deoxy-~-manno-2-octulosonatealdolase, also known as 2-keto-3-deoxyoctanoate (KDO) aldolase, catalyzes the reversible condensation of pyruvate with D-arabinose to form KDO (Fig. 14.1-22). KDO and its activated form CMP-KDO are key intermediates in the synthesis of the outer membrane lipopolysaccharide (LPS) of Gramnegative bacteria[145].Inhibitors of LPS biosynthesis or LPS binding KDO aldolase has a catabolic functherefore serve as antimicrobial agents [147-1511. tion, with an equilibrium constant for degradation of 0.077 M. It has been isolated and purified from E. coli [1521 and Aerobacter cloacae [1531. Preliminary investigations on this enzyme showed high specificity for KDO in the direction of cleavage, whereas the condensation reaction proceeded with some flexibility. Several unnatural substrates, including D-ribose, D-xylose, D-lyxose, L-arabinose, D-arabinOSe 5-phosphate and N-acetylmannosaminewere reported to be weak substrates (relative Studies on the substrate specificity of KDO aldolase rate >5% cf. ~-arabinose['~~]). from Aureobacterium barkerei strain KDO-37-2, have indicated that this enzyme OH 0

OH OH 0

KDO aldolase

OH OH D-arabinose Figure 14.1-22.

H

O

W C OH OH

pyruvate

Aldol addition reaction catalyzed in vivo by KDO aldolase.

O KDO

Y

14.7 Aldol Reactions

A 0

0

-0pc

+

OH OH

R

RHO

OH

OH

H

b

o

R = OH, 57% R = H, 47%

-

KDO aldolase

H

-

O

0

OH D-GIY

Figure 14.1-23.

&

OH W O

H

- -

11%

KDO aldolase-catalyzed synthesis of carbohydrates.

I

"

OH

KDO 8-P

synthetase

o-arabinose 5-P

Figure 14.1-24.

H 0 H q c 0 2 -

OH

OP

I

4c0

-

-02C-OH

OH

947

HO,,,.

R = OH, D-ribose R = H, 2-deoxy-~-ribose

+

-o,cJ---

0

KDO aldolase

O +H ,

R

pyruvate

-

OH

I

p?

.-

.

:

OH

I

C

KDP 8-P

Go;

Aldol addition reaction catalyzed in uiuo by KDO 8-Psynthetase.

widely accepts trioses, tetroses, pentoses and hexoses as substrates['54].The best substrates have (R)-configurationat C3, with the substituent at C2 having little effect. Several aldol addition reactions have been conducted on a preparative scale, including the synthesis of KDO itself, which was obtained in 67% yield (Fig. 14.123). In each case, attack of the pyruvate took place on the re face of the carbonyl group of the acceptor substrate. Excess pyruvate can be decomposed with pyruvate decarboxylase to simplify the i~olationI'~~1. 3-Deoxy-~-manno-2-octulosonate 8-phosphate synthetase, also known as phospho2-keto-3-deoxyoctanoate(KDO 8-P) synthetase, catalyzes the irreversible aldol reaction of PEP and D-arabinose 5-phosphate to give KDO 8-P (Fig. 14.1-24)[155]. The and the enzyme has been isolated from E. coli B[15'1 and Pseudomonas aer~ginosa('~~1, E. coli enzyme has been cloned and overexpressed in E. coli and Salmonella typhirnuri~m['~~]. It has been used in the synthesis of KDO 8-P, using D-arabinose 5-phosphate generated either by hexokinase-catalyzed phosphorylation of arabiStudies nose 11521, or an isomerase-catalyzed reaction of D-ribose 5-phosphate indicate KDO-8-P is very specific for its natural substrates, although some KDO analogs may be accessible. 14.1.2.3 3-Deoxy-~-arabino-2-heptulosonicAcid 7-Phosphate (DAHP) Synthetase (E.C. 4.1.2.1 5)

In vivo, DAHP synthetase, also known as phospho-2-keto-3-deoxyheptanoate synthetase, catalyzes the synthesis of DAHP from PEP and D-erythrose 4-pho~phate['~~]]. DAHP is a key intermediate in the shikimate pathway for the biosynthesis of

OH

948

I

74 Formation of C-C Bonds

0

OP

Ao2-*-

Aco2ADP

ATP

PEP 3

D-fructose

upo&OH

OH OH 0

OH 0

p

1

OH OH 0-fructose 6-P

o

M

C

OH

OH

D-erythrose 4-P

DAHP

0

2

-

1. Hexokinase,2. Pyruvate kinase, 3. Transketolase + D-ribose 5-P, 4. DAHP synthetase

Multi-enzyme synthesis of DAHP.

Figure 14.1-25.

aromatic amino acids in plants['60].The enzyme has been cloned['"] and used to synthesize DAHP (Fig. 14.1-25) In this synthesis, D-erythrose4-phosphate was generated in situ from Fru 6-P, catalyzed by transketolase in the presence of D-ribose 5-phosphate. Fru 6-P was generated from D-FWand ATP, catalyzed by hexokinase in the presence of an ATP regeneration system. In general, it is more efficient and economical to use whole cells containing a DAHP synthetase plasmid['"]. Such a system also provides the necessary enzymes for the synthesis of DHAP. Recently, DAHP synthase purified and overexpressed in E. coli has been characterized with respect to substrate specificity, and catalyzes the condensation of PEP with ribose5-phosphate, deoxyribose-5-phoshate,and arabinose-5-phosphate This enzyme has also been employed as a component of a biocatalytic process for large-scale production of vanillin from glucose [1651. 14.1.2.4

2-Keto-4-hydroxyglutarate (KHC) Aldolase (E. C. 4.1.2.31)

In viuo, KHG aldolase catalyzes the reversible condensation of pyruvate and glyoxylate to form KHG (Fig. 14.1-26)W6* 1671. This enzyme participates in the terminal step of mammalian catabolism of L-hydroxyproline The enzymes isolated and purified from bovine liver and E. coli are both type I aldolases. Limited substrate

'

-0pc

pyruvate

+

0

KHG aldolase

HKC02.

-

glyoxylate

-

0

OH

-02c KHG

Other pyruvate analogs which are donor substrates for KHG aldolase

-02cLR

Hk

Et02C

0 R = CH3, CHpCO;,

Br, OH, SH, Ph, imidazole, PhOH, CO;

Figure 14.1-26. Aldol addition reaction catalyzed in vivo by KHC aldolase and the donor substrate specificity of this enzyme.

74. I Aldol Reactions

I

949

specificity studies on KHG aldolase from bovine liver indicate that it accepts both 2-ketoenantiomers of KHG equally well, and also cleaves 2-keto-3-deoxyglucarate, 4,5-dihydroxyvalerate, and oxaloacetate [1671. In the condensation direction, this enzyme is relatively specific for glyoxylate, although it does accept other pyruvate derivatives[l6'I. The enzyme from E. coli prefers the natural substrate [KHG with (S)configuration]and also cleaves 2-keto-4-hydroxybutyrate and oxaloacetate [1691. Using the E. coli enzyme, both L- and D-4-hydroxy-2-ketoglutarate have been prepared on a millimole scale[170].In the condensation reaction, glyoxylate can be replaced with glyoxaldehyde, formaldehyde, acetaldehyde, and formic acid, while pyruvate can be substituted by a-ketobutyrate and bromopyruvate. 14.1.2.5

2-Keto-3-deoxy-dphosphogluconate (KDPC) Aldolase (E. C. 4.1.2.14)

In vivo, KDPG aldolase catalyzes the reversible condensation of pyruvate with D-G~Y 3-P to form KDPG (Fig. 14.1-27).The equilibrium constant lies in favor of the aldol addition (K- lo3 M - ~ ) .KDPG aldolase accepts a number of unnatural acceptor aldehydes, although at rates much lower than the natural Various sources of KDPG aldolase have been investigated as C - C bond forming catalysts in organic synthesis [17*1, such as for the synthesis of non-carbohydrate components of the nikkomycin natural products The related enzyme KDPGal aldolase has also been utilized for similar purposes sp[1741. Unlike other aldolases, simple aliphatic aldehydes are not KDPG aldolase substrates. However, other than the presence of polar functionality at C2 or C3, there appears to be no other structural requirement for the acceptor aldehyde. These studies also demonstrate that KDPG aldolase stereospecifically generates the new stereocenter at C4 with (S)-configuration. Furthermore, by using the technique of directed evolution, KDPG aldolase has been altered with respect to its acceptor enantioselectivity and phosphate requirement to accept non-phosphorylatedenantiomeric aldehydes [1751. 0

PO

~

0

+

ACO; -

KDPG aldolase

OH

D-GIY3-P

-

OH 0 PO--,J-%Oi OH

pyruvate

KDPG

Other acceptor substrates of KDPG aldolase Acceptor

VEI

Acceptor

Vrd

nitropropanal chloroacetaldehyde D-glyceraldehyde D-lactaldehyde ribose 5-P

200

erythrose glycoaldehyde benzaldehyde butyraldehyde ribose

1.5 1.5

120 100 27 5

0 0

0

Figure 14.1-27. Aldol addition reaction catalyzed in vivo by KDPC aldolase and the acceptor substrate specificity of this enzyme.

950

I

74 Formation of C-C Bonds

14.1.2.6 2-Keto-3-deoxy-~-glucarate (KDC) Aldolase (E. C. 4.1.2.20)

In vivo, KDG aldolase catalyzes the reversible reaction of pyruvate and tartronic acid semialdehyde to form KDG (Fig. 14.1-28).This aldolase has been found in various bacteria and the enzyme from E. coli has been isolated and purified['76]. KDG aldolase accepts several other aldehyde acceptor substrates, including glycoaldehyde, glyoxylate, and D- and L-glyceraldehyde. It has been used to synthesize 2-keto3-deoxy-~-gluconate on a preparative scale[1771. 14.1.3

2-Deoxyribose 5-phosphate Aldolase (DERA) (E.C. 4.1.2.4)

DERA[1781is unique among the aldolases, in that the donor of the aldol reaction is an aldehyde, rather than a ketone. In vivo,the enzyme catalyzes the reversible condensation of acetaldehyde and ~ - G l y3-P to form D-2-deoxyribose5-phosphate, with an equilibrium constant in the cleavage direction of 2 x M (Fig. 14.1-29).It is a type I aldolase, and has been isolated from animal tissues [17q1 and microorganisms [180]. The E. coli gene encoding DERA has been sequencedIls1I, subcloned, and the enzyme overexpressed in E. c01i[182-184].At 25 "C and pH 7.5, DERA is fairly stable (70% activity retained after 10 days). A number of unnatural substrates are accepted by DERA (Fig. 14.1-29), and it ~ ~ ~ 1 various generates (R)-configuredchiral centers. DERA from L. p l a n t a r ~ r n [accepts acceptor substrates including L-G~Y 3-P, D-erythrose 4-phosphate, glycoaldehyde phosphate, D-ribose 5-phosphate, D,L-glyceraldehyde, D-erythrose,and D-threose [lS6]. Only propionaldehyde can weakly replace acetaldehyde as the donor. The E. coli enzyme[lS2]accepts acetaldehyde, propionaldehyde,acetone, fluoroacetone, aliphatic aldehydes, sugars, and sugar phosphates as acceptor substrates. However, the rates of the aldol reactions are very slow (0.4-1 % CJ the natural substrates). More recently, DERA has been used to obtain key intermediates in the synthesis of the epothilone class of natural products['88].Several syntheses of azasugars conducted using DERA are illustrated in Fig. 14.1-30. When acetaldehyde is used as the donor, the products from the DERA-catalyzed reaction are aldehydes, capable of being acceptor substrates for a second aldol condensation (Fig. 14.1-31)[lS71. For example, when a-substituted acetaldehydes were employed as substrates, products of the first aldol condensation could not cyclize to a hemiacetal, and the products reacted with a second molecule of acetaldehyde to form 2,4-dideoxyhexoses. These products could then cyclize to stable

-02cL +

PYrUVate Figure 14.1-28.

H&coi

OH tartronic acid semialdehyde

- KDG aldolase -

0 OH -02c+co2

OH KDG

Aldol addition reaction catalyzed in vivo by KDC aldolase.

14.7 Aldol Reactions

acceptor

donor

I

951

product

OH R = H, F, CI, Br, OH, or CH,

1 OH HO R = CH,. CH,OH, or Ph

Hoa P

N3

OH

OH

P

OH

OH

HoTsT-oH OH

OH

OH I

9 Figure 14.1-29. Aldol addition reaction catalyzed in vivo by DERA, and reactions with unnatural substrates.

952

I

74 Formation ofC-C Bonds Figure 14.1-30. Syntheses of azasugars using DERA.

/J

-

N3

:

P

OH

donor

product

Hz,Pd/C

azasugar

OERA

N3v0H

9

OH

OH

P

"

'

POH O CH3 H

OH

A

HO OH

DERA R

I]

OH

0

DERA

R

R = CH3, MeOCH,. MOMOCH,, CICHz. N3CHn

'm

CH,CH,COOH, CH,0HCH20P

OH

Figure 14.1-31.

~

I'

OH

0

R

t

Br,/HZO

BaCO,

72%

'Tor -

OH

OH

Sequential aldol reactions catalyzed by DERA.

hemiacetals, thus stopping the polymerization after two sequential aldol reactions. Conversion to chiral lactone derivatives of mevinic acids, which are active as cholesterol-loweringagents, could then be accomplished. The best substrate for the DERA-catalyzed sequential reaction appeared to be succinic semialdehyde (R = CH2CH2COOH) in which the carboxylic acid mimics the Gly 3-P phosphate group [1841. One-pot sequential aldol reactions were performed by combining DERA with FDP

74.7 Aldol Reactions

OH HO

R = MeOCH,, MOMOCH,, CICH,,

HO OH

DERA

R

P

R = CH, MeOCH,, MOMOCH,, CICH,. NsCHz, CHZCHZCOOH, CHzOHCH2OP

OH

OH

R&

N&H,

87-

Figure 14.1-32. One-pot aldol reaction employing RAMA and DERA.

0

OH

1\1111 OH

DERA

P

R

OH

R

0

Figure14.1-33. Tandem use of DERA and NeuAc

aldolase.

Rr R,oroH ~

Br2/H,O

BaCO,

OH

72%

v

OH

aldolase (Fig. 14.1-32)[18’), ”)‘I . The products of these reactions are 5-deoxy ketoses with three substituents in axial positions. Owing to the formation ofthese thermodynamically unfavored products at long reaction times, some inversion of the usual stereochemistry of both DERA and FDP aldolase was observed. Combination of DERA and NeuAc-aldolase catalysis gave sialic acid derivatives (Fig. 14.1-33)[1891. In this case, however, one-pot synthesis was not possible, due to the incompatibility of the reaction conditions for the two aldolases. Glycine-dependentAldolases The glycine-dependent aldolases, including serine hydroxymethyltransferases (SHMT) and threonine aldolases, are pyridoxal 5-phosphate-dependent enzymes which catalyze the reversible aldol reaction of glycine with an aldehyde acceptor to In vivo SHMT (EC 2.1.2.1) catalyzes the conform a p-hydroxy-a-amino densation of glycine and formaldehyde to give L-serine, and requires the cofactor tetrahydr~folate[”’~].SHMT has been used for the resolution of racemic erythro phydroxy a-amino acids, the large-scale synthesis of ~-serineI’”>1931, and the production of 2-amino-3-hydroxy-l,6-hexanedicarboxylic acid [1941. Although SHMT is selective for the L-configuration at the a-center, it generally displays poor erythro-threo discrimination, resulting in product mixtures [195,1961. Threonine aldolases catalyze the reversible aldol reaction between glycine and acetaldehyde to give threonine (Fig. 14.1-34),and both D- and L-Thr aldolases have been reported. The substrates for the L-threonine aldolases (E.C. 4.1.2.5) are also substrates for L-SHMT (vide supra). Many threonine aldolases also accept allo-

I

953

954

I

,P

C-CBonds

poH- -

74 Formation of

OH

L-threonine aldolase

+

0

NHz erythro Yield (%)

threo Ratio (elythro:threo)

CH,

-

38

93 : 7

Ph

-

87

60 : 40

45-75

70 : 30 to 100 : 0

N&Hz

BnOCH, BnOACH2 BnO-0,

CHp

PhS-CHZ-

Figure 14.1-34. substrates.

0

R

NHp

R

OH

-

78

92 : 8

53

53 : 47

45

92 : 8

80

50 : 50

Reaction catalyzed in vivo by L-Thr aldolase, and unnatural

threonine derivatives as substrates, sometimes preferably over compounds with the "'l. threo configuration Threonine aldolases have been used extensively for the resolution of racemic phydroxy a-amino acids. For example, with a L-threonine aldolase isolated from Streptornyces arnakusaensis, several racemic mixtures of 3-@-substituted-phenyl)serines were resolved to give the enantiomers with the D-threo stereochemistry in >95 % ee['99*2001 . Recently, both D-[~''] and L-Thr aldolases ['01* 2021 have been used in the preparation of novel P-hydroxy-a-aminoacids. In addition, D-threonine aldolase has been utilized to prepare a small molecule that acts as a gelator of organic solvents [2031. L-Threonine aldolase has been employed in the synthesis of fragments of the mycestericin class of natural as well as peptidic RNA mime t i c ~ [ ~L-Threonine ~~]. aldolase (E. C. 4.1.2.5) from Candida hurnicola has been and has been investigated for use in condensation reactions['"]. The enzyme accepted a broad range of aldehydes, but in general mixtures of L-erythro and L-threo products were obtained, with the L-erythro configuration being the preferred one (Fig. 14.1-34). When hydroxyaldehydes are employed as L-Thr aldolase substrates, complex product mixtures result. Protection of the hydroxyl groups prevents this, and allowes the preparation of CCprotected L-threonineand L-allothreoninederivatives.Acceptor aldehydes with an oxygen functionality at the a-position gave high erythro/threo

14. I Aldol Reactions

I

955

& ‘OH

.!.

OH

Figure 14.1-35.

Use of L-Thr aldolase in the preparation of sLe” mimetics.

ratios, a ratio which was reduced when the oxygen was in the S-position.Although a,S-unsaturated aldehydes did not serve as substrates, several thiophenol derived aldehydes were accepted, providing a route toward unsaturated amino acids. One LThr aldolase product, the 4-hydroxy-~-allothreonine derivative,has been used as a key C2O71. intermediate in the synthesis of potent sialyl Le” mimetics (Fig. 14.1-35) Other known aldolases whose substrate specificity remains to be examined are summarized in Table 14.1-2. Catalytic Antibodies

In recent years, catalytic antibody technology has provided methods for developing new protein catalysts[208]. Monoclonal antibodies (mAbs)elicited against “transitionstate” haptens catalyze reactions with remarkable rate accelerations. By appropriate antigen design, functional groups that perform general acid/base catalysis, nucleophilic/electrophilic catalysis, and catalysis by strain or proximity effects can be induced into the binding site of an antibody. Even reactions which are unfavorable or otherwise unattainable have been achieved using the catalyhc antibody approach. Aldolase catalytic antibodies developed recently have the ability to match the efficiency of the natural aldolases while accepting a more diverse range of substrates. Initial catalytic antibodies were developed to bind a primary amine cofactor as a mimic of the type I aldolases. The hapten designed mimicked the transition state the iminium ion, resulting in the production of an antibody that catalyzed the aldol [2091. Even though no condensation of acetone and aldehyde acceptors (Fig. 14.1-36) stereochemical information was built into the transition-state mimic, the antibody catalyzed stereoselective addition to the si face of the aldehyde. The subsequent development phase, namely reactive immunization L2l0], involved

OH 0

OH 0

acetone, pH 9.0

AcHN

AcHN > 95% de

Ar

Figure 14.1-36. Aldol reaction catalyzed by catalytic antibody 72D4, and a transition-state hapten.

1

:

2.8

65% de

956

I

14 Formation ofC-C Bonds Other aldolases and the reactions they catalyze in uiuo.

Table 14.1-2.

0 P O L O H

ketotetmse pimphate & O 'H "

+

aldolase (EC 4.1 2.29) [a]

DHAP 0

'

'02C pyruvate

+

phospho-5-keto-Zdeoxy-

HIccoz.

PO & JO i.z

gluconate aldolase (EC 4.1.2.29) [b]

OH

'bop

2-keto-3.deoxy-S-phospho-

+

galactonate aldolase (EC 4.1.2.21)

OH

[i

0

OH

-02c+oP 6H

4hydroxy-2-keto4rnethyl

+

'OZC

glutarate aldolase (EC 4.1.3.17) [c]

H K O H

+

Hj C O H

+

-

2-keto-3deoxy-c-pentanoate aldolase (EC 4.1.2.28) [c]

0

2-keto9deoxy-~-pentanoate

0

OH OH

WOH

Nacetylneurarninate(NeuAc) -0zc

synthetase (EC 4.1.3.19)

PEP

OH

'0,C &OH

aldolase (EC 4.1.2.18) [d]

-0zc

OH

ACHN

serine hydmxymelhyl

-0,c R

transferase (EC 2.1.2.1) [el

O

OH

H

OH dlhydroneopledn

HoT)$lNH2 + HKoH

K

PO

+

H

b

O

p

+

phosphoketolase

HzO

0

H

(EC 4.12.9) [g]

OH

+

OH

fructose-6-phosphate

+H20

H

OH

@

17a-hydmmrwestemne +

pH

aldolase (EC 4.1 2.30)[i]

O 0 A OHO

P

+ Pi

OH OH

-

+ Pi

phosphoketolase(EC 4.1.2.22) [h]

OH

NHZ

HoG:;7Jl

aldolase (EC 4.1.2.25) [fj

0

OH

0

-0 0 -ozc+

+

a Isolated from rat liver, see: F. C. Charalampous,

Methods Enzymol. 1962, 5, 283. Acetaldehyde, glycoaldehyde or glyceraldehydecannot replace formaldehyde.

ketopantoaldolase (EC 4.1.2.12) 1

0 .OzC?OH

b W. A. Andeson, B. Magasanik, J. Bid. Chem. 1971, 246, 5662. c W. A. Wood in: The Enzymes (Ed.: P. D. Boyer),Academic Press, New York, 1970;Vol. VII, p. 281.

14.1 Aldol Reactions with acetaldehyde to give L-allothreonine.originally d This enzyme also catalyzes the aldol addition of thought to be catalyzed by I-allothreonine aldolase pyruvate with formaldehydeto give 4-hydroxy-2-oxobutyrate, originally thought to be catalyzed by hy(E.C. 4.1.2.6). droxyoxobutyrate aldolase (E. C. 4.2.1.1). Phenylpyru- f I. B. Mathis, G. M. Brown,]. Bid. Chem. 1970,245, 3015. The reaction requires thiamine pyrophosphate vate is also a donor substrate, while acetaldehyde, and favors cleavage. benzaldehyde and crotonaldehydeare not acceptor g E. C. Heath, J. Hunvitz, B. L. Horecker, A. Ginsberg, substrates, see: H. Hift, H. R. Mahler,]. B i d . Chem. /. B i d . Chem. 1958,231, 1009. The reaction favors the 1952,198,901. e L. Schirch, Adv. Enzymol. 1982,53,83. A multicopy cleavage of ~-xylulose-5-phosphate.The enzyme from Leuconostoc msenteroides also accepts fructose-6-phosplasmid containing the E. coli serine hydroxymethyl transferase was introduced to Klebsiella aerogenes for phate, hydroxypyruvate and glycoaldehyde as suboverexpression of the enzyme. The enzyme requires strates. tetrahydrofolate (THF) and pyridoxal phosphate. THF h E. Racker, Methods Enzymol. 1992,5,276. The reaction favors degradation. first reacts nonenzymaticallywith formaldehydeto form NS,NlO-methyleneTHF which is then accepted i D. E. lohnston, Y:B. Chiao, 1. S. Gavaler, D. H. Van by the enzyme to form serine, see: B. K. Hamilton, Thiel, Biochem. Pharm. 1981,30, 1827. H. Y. Hsiao, W. E. Swanm, D. M. Anderson, J. Delej W. K. Maas, H. J. Vogel,]. Bacterial. 1953,65,388; E. N. McIntosh, M. Purko, W. A. Wood,]. B i d . Chem. nte,]. Trends Biotechnology, 1985, 3,64. This enzyme 1957,228,499. also catalyzes the reversible aldol reaction of glycine

raising antibodies against a 0-diketone “chemical trap” to imprint the lysinedependent type I aldolase mechanism in the active site (Fig. 14.1-37)[211!The Eamino group of a lysine side chain reacts with the 0-diketoneto give a 0-ketoimine, which tautomerizes to the stable vinylogous amide. By using this method, two catalyix antibodies with aldolase selectivity, 38C2 and 33F12, were identified and subsequently shown to have remarkable scope[212]. The structure of 33F12 has been determined and shown to have the Schiff base forming Lys residue buried in a hydrophobic pocket at the base of the binding site[211]. Unlike natural aldolases, catalytic antibodies accept a wide range of ketone donor substrates (Fig. 14.1-38A). Small aliphatic ketones are well tolerated, but mixtures of products result with unsymmetrical ketones, due to reaction at both a-positions. aHeteroatom-substituted ketones show much higher levels of regioselectivity, with reaction occurring almost exclusively at the carbon atom bearing the heteroatom. Interestingly, the regiochemistry of the reaction of fluoroacetone is opposite to that observed with the natural aldolase DERA, thus providing a complementary approach. A wide variety of aldehydes serve as acceptors (Fig. 14.1-38B),including those that

Figure 14.1-37.

Reactive immunization strategy.

H

I

957

958

A.

I

14 Formation ofC-C Bonds

A it

1 ,

,),OeM

FJ

OH, OMe

0

I

R = NHAc, NO2

AH

AcHN

J H n = 3, 4, 5

Figure 14.1-38. A, Catalytic antibody ketone donor substrates. B, Catalytic antibody aldehyde acceptor substrates.

resemble the hapten, and simple aliphatic aldehydes. Polyhydroxylated aldehydes, such as glyceraldehyde, glucose, and ribose, are not substrates, most likely because of the hydrophobic nature of the active site. In contrast to the natural aldolases, aromatic and a$-unsaturated aldehydes are excellent substrates. The stereochemistry of the addition is donor dependent. When acetone is used as the donor substrate, addition occurs from the si face of the carbonyl group; with hydroxyacetone, addition occurs from the re face. The stereoselectivity is generally quite high, with ee values greater than 99 % commonly observed. As a general rule, high enantioselectivity is observed with acceptors having an sp2 center in the aposition, and lower enantioselectivitiesare observed for a-position sp3 centers. The utility of catalytic antibodies was demonstrated with the antibody-catalyzed aldolase approach to the brevicominssp[213]and the epothilones l2l4l (Fig. 14.1-39). Antibody 38C2 is commercially available and has recently been used as a catalyst to activate prodrugs i2l5].Generic, drug-masking groups can be selectively removed by sequential retro-aldol and retro-Michael reactions catalyzed by 38C2 (Fig. 14.1-40). The antibody was also used in the enantioselective retro-aldol reaction of tertiary aldols containing heteroatom-substituted quaternary carbon centers F21G]. This gave enantiomerically enriched tertiary aldols, most with ee values greater than 95 %. Synthesis of enantiomerically pure tertiary aldols using the catalytic asymmetric aldol reaction with ketone acceptors represents a significant challenge. Compounds prepared in this study have been used in the synthesis of (+)-frontalin,the side chain of saframycin H, and mevalonolactone. In order to increase the repertoire and efficiency of the aldol reaction further, and

$

74. I Aldol Reactions

OH

40%,96%ee

Ar

Ar

(*)

+;flcHo 51%, 75%ee Figure 14.1-39.

0 O *H /

Use of catalytic antibody 38C2 for the preparation o f epothilone intermediates. OH

0

/

"'OH

3*c2 *

Me0

0

pro-drug Figure 14.1-40.

OH

0

retroaldol

Me0

0

OH

0

drug Retro-aldol reaction catalyzed by Ab 38C2 for the unmasking o f pro-drugs.

to develop antibodies with complementary enantioselectivity, a P-diketone sulfone was employed as the h a ~ t e n [ ~ (Fig. ' ~ I 14.1-41). The tetrahedral geometry of the sulfone moeity in this hapten mimics the rate-determining tetrahedral transition state of the C-C bond forming reaction. It is thus expected to facilitate nucleophilic attack of the enaminone intermediate on the acceptor aldehyde. It was indeed demonstrated that catalytic antibodies with broad reaction scope can be generated using this approach. In addition, antibody 93F3 was more efficient (k,,, - 3min-l) than and enantiocomplementary to 38C2, providing the unreacted (S)-aldolwith >96% ee. The mechanism-based approach to eliciting catalytic antibodies combined with the rapid, immune-selection process as illustrated in these studies provides a new and exciting direction for catalyst design and development.

I

959

960

I

14 Formation of C-C Bonds

5-

antibody * +[>7&N-Ab R

6'

R'

transition state

R o s 2

+

H~N/\/\/~~

transition-state analog reactive immunization

H

hapten, R = - O 2 C V N 0

1 CH3

Figure 14.1-41.

fi-Diketone sulfone as hapten for reactive immunization.

14.2 Ketol and Aldol Transfer Reactions

14.2.1 Transketolase (TK)(E.C. 2.2.1 .l)

TK is one of the enzymes involved in the oxidative pentose phosphate pathway, and requires the cofactors thiamine pyrophosphate (TPP)[21'1and Mg2+[2181.It reversibly transfers the C1-C2 ketol unit from D-xylulose 5-phosphate to D-ribose 5-phosphate, and generates D-sedoheptulose7-phosphate and D-GIY3-P. D-Erythrose 4-phosphate also functions as an acceptor of the ketol unit from D-xylulose 5-phosphate, to 3-P (Fig. 14.2-1).TK from baker's yeast is commercially produce Fru 6-P and ~ - G l y available, and the enzyme can also be isolated from spinach[220.221]. TK from E. coli has been overexpressed and prepared on a large scale[222]. In ketol transfer reactions, OH 0 $ ,

)

0

OH

+ H*op

, OP OH D-XylUlOSe 5-P

OH 0

OH D-xylulose 5-P

OH OH TK

-

P

O

0

OH

OH D-erythrose 4-P

A

H +Ho>\+op

0

OH OH D-ribose 5-P

+ H+oP

H o, - ,k- .O P +

-

D-GIY3-P

-

0

OH TK

-

OH OH

D-sedoheptulose 7-P

OH OH

+ Ho++.OP

PO+H

0 D-GIY3-P

0

OH

D-Fru 6-P

Figure 14.2-1. Ketol transfer reactions in the oxidative pentose phosphate pathway catalyzed by TK.

14.2 Ketol and Aldol Transfer Reactions

I

961

0 HL

+

HOJLoz HPA

O

TK

H

*

H

O

L

o

H

OH

R = CHZOH, CH3, N3, CHzCH3

0

0

OH OH

TK

H+OH

+

HOO *H

. .

. .

OH OH OH

OH OH 0

OH OH

0

OH

OH

OH

OH OH

0

OH

TK

H&

H

-*

OH

O -. . L\ OH OH OH 0

TK

*

OH 0 Figure 14.2-2.

HOQH OH OBn

Acceptor substrate specificity o f TK.

the enzyme isolated from yeast shows a higher diastereoselectivity(- 100%)[221] than that from spinach (- 95 %), with the newly-formed hydroxymethine chiral center always possessing an (S)-configuration. TK also accepts p-hydroxypyruvic acid (HPA) as a ketol donorlzz3],and an efficient multi-enzyme synthesis of D-xylulose 5-phosphateemploying FDP aldolase and E. coli transketolase has been reported[224]. The ketol unit is transferred to an aldose acceptor with an activity of 4 % compared with D-xylulose 5-pho~phate[~~']. This has been an invaluable discovery for the use of TK in synthesis, as the decarboxylation of HPA and subsequent loss of carbon dioxide, render the overall condensation reaction irreversible. A wide range of aldehydes are ketol acceptors, including aliphatic, a,P-unsaturated, aromatic, and heterocyclic aldehydes, although some are relatively poor substrates (Fig. 14.22)[225, 2261. The presence of a hydroxyl or an oxygen atom at C2 and/or C3 has a positive effect on the rate, while steric hindrance near the aldehyde exerts a negative effect. P-D-Hydroxy aldehydes (and not L-) are substrates, producing vicinal diol products of D-threo configurationLzz5, 2271. This allows efficient resolution of aldehydes epimeric at C2 by transketolase. The enzyme appears to have no preference for configuration beyond C2.

962

P

OH O 4 H

I

14 Formation of C-C Bonds

OH OH

+H

o 0

0

D-GIY3-P

~

~

TAo =======

OH OH

0

D-sedoheptulose7-P Figure 14.2-3.

0

OH OH H o p~ , ; r / O P

+

OH

H+OP

OH

OH

D-Fru 6-P

D-erythrose4-P

Aldol transfer reaction in the oxidative pentose phosphate pathway

catalyzed by TA.

Starch phosphorylasea

__j__j______

D-Glc ,-p

phosphate

Figure 14.2-4.

phosphoglucomutase

D-Glc 6-p

phosphoglucose isomerase

-

-

D-Fru 6-P

7~7; TA

u

D-G~Y ~~~

D-Fru

D-GIY3-P

3-phosphoglycerate phosphatase

Multi-enzyme synthesis of D-Fru from starch.

TK has been used to catalyze the key step in the synthesis of the naturally occurring beetle pheromone (+)-exo-brevicomin[228] and the azasugar 1,4-dideoxy1,4-imino-~-arabinitol [391. Both syntheses involve the condensation of H PA with racemic 2-hydroxyaldehydes,whereby the ketol unit is diasteroselectivelytransferred to only the D-enantiomer of the aldehyde. In addition, transketolase has been employed in the synthesis of complex heptuloses [2291, fructose analogs [2301, and other sugars [231j. Erythrulose has been continuously produced through transketolase-catalysisin a membrane reactor [2321. 14.2.2 Transaldolase (TA) (E.C. 2.2.1.2)

TA is also an enzyme ofthe oxidative pentose phosphate It catalyzes the transfer of the C1-C3 aldol unit from D-sedoheptulose7-phosphate to ~ - G l 3-P, y and produces D-FI-U6-P and D-erythrose 4-phosphate (Fig. 14.2-3).TA forms a Schiff base intermediate and does not require any co-factors. This enzyme is commercially available, and was used in a multi-enzyme synthesis of D - F ~ ufrom starch (Fig. 14.24)[2331.Here, it accomplished transfer of an aldol moiety from Fru 6-P to Dglyceraldehyde,and formed ~ - G l 3-P y and D - F ~ .

14.3

Acyloin Condensation

Acyloin condensation catalyzed by yeast was first observed in the early part of the twentieth century[234,2351 . Yeast-catalyzed acyloin condensations between acetaldehyde and benzaldehyde derivatives have since been reported, giving products with a (R)-configurationin all cases (Fig. 14.3-1)[ 2 3 G , 2371. The acyloin formed from benzaldehyde alone has been used in the industrial manufacture of It is

$fi

74.4 C-CBond Forming Reactions lnvolving AcetylCoA

0

FyH JH 7-

+

I

963

R’

AcyloinYeast condensation-

R2 acetaldehyde benzaldehyde derivative

R2

Products obtained from yeast-catalyzed acyloin condensation CI

Me0

Me0

OH

HO

Acyloin condensation between acetaldehyde and benzaldehyde derivatives catalyzed by yeast. Figure 14.3-1.

probably the enzyme a-carboxylase (E. C. 4.1.1.1)that is responsible for catalyzing the acyloin reactions, as the carboxylase-catalyzed reaction of pyruvate and benzaldehyde in the presence of the cofactor thiamine pyrophosphate gives the corresponding acyloin product [2391. Pyruvate decarboxylase in highly purified [2401 or partially purified catalyzes acyloin condensation to give products of the ( R ) configuration.

14.4

C-C Bond Forming Reactions InvolvingAcetylCoA

Enzymatic reactions which utilize coenzyme A thioesters as substrates are involved in the biosynthesis of steroids, terpenoids, macrolides, fatty acids, and other natural products. Owing to the high cost of CoA, these enzymes can only be practically used in organic synthesis if the CoA thioesters can be recycled. AcetylCoA can be efficiently regenerated by using one of several enzymatic systems [242-2441 . Phosphotransacetylase (E. C. 2.3.1.8)/acetylphosphate, carnitine acetyltransferase (E. C. 2.3.1.7)/acetylcarnitine,and acetylCoA synthetase (E. C. 6.2.1.1)/ATPhave all been employed for this purpose. These enzymatic recycling systems have been coupled to the synthesis of citric acid catalyzed by citrate synthetase. An interesting non-enzymatic regeneration of acetylCoA utilizes phase transfer catalysts in a twophase aqueous-organic system (Fig. 14.4-1) L2451. Citric acid was efficiently prepared using this procedure, and this method also offers the potential to prepare many different acylCoA derivatives for use as substrates of CoA-dependent enzymes. AcetylCoA is also involved in the biosynthesis of poly-p-hydroxybutyrate(Fig. 14.42, x = 0). Many whole cell systems have been used to synthesize this polymer and other interesting materials in this class [2461. For example, copolymers consisting of x = 0 and 1, respec(R)-3-hydroxybutyland (R)-3-hydroxyvalerylunits (Fig. 14.4-2, tively) were prepared by feeding propionate to whole cells of A. e ~ t r o p h u s [ ~ ~ ~ ] .

964

I

14 Formation of C-C Bonds

aqueousphase organic phase

acetylCoA H02C

i.

Figure 14.4-1.

0 RASCOA

thiolase acetylCoA-

acylCoA R = CHS(CH~),

Chemical regeneration of acetylCoA using a phase transfer catalyst.

0

0

RuSCoA

P-ketoacylCoA

OH 0

reductase RuSCoA

p-hydroxyacylCoA

synthetase

poly-p-hydroxyester n = 500-15,000

X = 0-7

Figure 14.4-2. Enzyme-catalyzed reactions involved in the whole-cell synthesis o f poly-p-hydroxyesters.

AcetoacetylCoA thiolase (E. C. 2.3.1.9),acetoacetylCoA reductase (E. C. 1.1.1.36),and polyhydroxybutyrate ~ y n t h e t a s e I ~ are ~ ~the 1 enzymes involved in polyester synthesis. AcetoacetylCoA thiolase catalyzes the head-to-tail Claisen condensation of two acetylCoA molecules. In this reaction, the active site cysteine attacks acetylCoA to form a thioester enzyme intermediate, which then reacts with the enolate derived from enzymatic deprotonation of the other acetylCoA. Mechanistic studies have been performed on this enzyme from Zooglea ramigera, which has been cloned and overexpres~ed[~~~I. It has been established that the thiolase will form acyl enzyme intermediates with a number of acylCoA substrates, but will only accept acetylCoA as the nucleophile. After subsequent reduction, this results in all polymer units possessing a fi-hydroxy group. These polymers are also useful sources of (R)-fihydroxy acids [2481.

14.5 lsoprenoid and Steroid Synthesis

I

965

14.5 lsoprenoid and Steroid Synthesis

Enzymes involved in the biosynthesis of isoprenoids and steroids have been used in organic synthesis [2491. 2,3-Oxidosqualenelanosterol cyclase was used to synthesize a number of lanosterol analogs (Fig. 14.5-1)[2sG2531. When using an enzyme suspension from baker’s yeast containing this cyclase, ultrasonic irradiation proved very . property of lanosterol cyclase effective in promoting catalysisr2”, 2521 . An interesting was utilized during the synthesis of C30 functionalized lanosterols, whereby the enzyme rearranged a vinyl group rather than the usual hydrogen or methyl group L2”1. This product was subsequently converted into (+)-3O-hydroxylanosterol and the corresponding aldehyde. These compounds are natural receptor-mediated feedback inhibitors of HMG-CoA reductase, and therefore are of interest in the design of hypocholesteremic drugs [2s41. Both enantiomers of 4-methyldihomofarnesol were synthesized using farnesyl diphosphate synthetase from pig liver, the (S)-enantiomer being a precursor of juvenile hormone (Fig. 14.5-2)[2s51. Alkyl group homologs of isopentenyl diphosphate have also been examined as substrates for farnesyl diphosphate synthase [2sGl.

&R3 R2

O

(from ? ~ ~ ~ ~ ~ baker’s $ ‘ , ” , ” , ” _ yeast)

\

I

R’

Figure 14.5-1.

\

HO R’ H H OH H

R2 H H H OH

R3 CH3 CO2CH3 CH3 CH,

.

% ...

R’

R~

lanosterol analogs

Synthesis of lanosterol analogues using 2,3-oxidosqualene lanosterol cyclase.

uopp (Sj-4-methyldihomofarnesol

farnesyl diphosphate

qthetase

(R)-4-rnethyldihomofarnesol

Figure 14.5-2. Synthesis o f both enantiomers o f 4-methyldihomofarnesol using farnesyl diphosphate synthetase.

juvenile hormone

966

I

74 Formation ofC-C Bonds

YH2 c 1 4 c o g

tryptophan synthase, RH

YH2 *

c'4co,

-0ZCCHzSH

*coz

0-substituted a-amino acids

0-chloroalanine

YH2

R

RH = PhCHZSH, CH3(CH&SH,

tyrosine phenol base. RH ~.

-

0-chloroalanine

H

NHZ

R*C-,__ 0-substituted a-amino acids

RH =

R' =OH, CI, alkyl

R'

Synthesis o f fi-substituted a-amino acids from fi-chloroalanine using tryptophan synthase and tyrosine phenol lyase.

Figure 14.6-1.

14.6 6-Replacement o f Chloroalanine

Methods have been developed for the synthesis of unnatural amino acids using pyridoxal phosphate-dependent enzymes 12571. These enzymes usually catalyze transaminations, a$-eliminations, a,y-eliminations, and decarboxylations of amino acids. However, using fbchloroalanine as a substrate, unusual amino acids are produced by P-replacement. Tryptophan synthase (E. C. 4.2.1.20) from E. coli catalyzes the formation of tryptophan and analogs. This enzyme has been employed to incorporate various heteroatoms into tryptophan, such as selenium [258J, ~ulfurI~~'1, chloride[261],and Notably, tryptophan synthase could be used to catalyze exchange of the a-proton from Asn, Glu, Ser, Ala, Phe, and Met as well as that of Trp[262]. Tyrosine phenol lyase (E.C. 4.1.99.2) (Fig. 14.61)has been utilized to synthesize tyrosine, DOPA, and rneth~latedL~~~1, fluorinated[264],and azido-tyrosineanalogs [2651.

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a) C.-H. Wong in: Enzymes as Catalysts in Organic Synthesis (Ed.: M. P. Schneider), D. Reidel Publishing Co., Dordrecht, 1986; p. 199; b) H. J. M. Gijsen, W. Fitz, L. Qiao, C.-H. Wong, Chem. Rev. 1996,96,443;c) W.-D. Fessner, CUT. Opin. Chem. Biol. 1998, 2, 85. 4 C.-H. Wong in: Biocatalysis (Abrarnowics), von Nostrand Reinhold, New York, 1990, p. 319. 5 E. J. Toone, E. S. Simon, M. D. Bednarski, G. M. Whitesides, Tetrahedron 1989,45, 53G5. 3

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C. Demuynck, J. Bolte, I>. Hecquet, V. Dalmas, Tetrahedron Lett. 1991,32, 5085. 227 F. Effenberger, V. Null, T. Ziegler, Tetrahedron Lett. 1992,33, 5157. 228 D. C. Myles, P. J. Andrulis 111, G. M. Whitesides, Tetrahedron Lett. 1991, 32,4835. 229 a) V. Dalmas, C. Demuynck, Tetrahedron: Asymm. 1993,4, 1169; b) C. Andre, C. Demuynck, T. Gefflaut, C. Guerard, L. Hecquet, M. Lemaire, J. Bolte, J. Mol. Catal. B: Enzym. 1998,5, 113; c) C. Andre, C. Guerard, L. Hecquet, C. Demuynck, I. Bolte, J. Mol. Catal. B: Enzym. 1998, 5, 459. 230 a) L. Hecquet, J. Bolte, C. Demuynck, Tetrahedron 1994,50, 8677; b) C. Guerard, V. Alphand, A. Archelas, C. Demuynck, L. Hecquet, R. Furstoss, J. Bolte, Eur. J. Org. Chem. 1999,12,3399. 231 a) A. J. Humphrey, N. J. Turner, R. McCague, S . J. C. Taylor, J. Chem. Soc., Chem. Commun. 1995,24,2475;b) K. G. Moms, M. E. B. Smith, N. J. Turner, M. D. Lilly, R. K. Mitra, J . M. Woodley, Tetrahedron: Asymm. 1996,7,2185. 232 J. Bongs, D. Hahn, U. Schoerken, G. A. Sprenger, U. Kragl, C. Wandrey, Biotechnol. Lett. 1997, 19, 213. 233 A. Moradian, S. A. Benner, J. Am. Chem. SOC.1992, 114,6980. 234 C. Neuberg, J. Hirsch, Biochem. 2.1921, 1IS, 282. 235 H. Beevarova, 0. Hane, K. Mauk, Folia Microbiol. 1963, 8, 165. 236 a) M. Behrens, N. N. Iwaneff, Biochem. Z. 1921, 121, 311; b) C. Fuganti, P. Grasselli, Chem. Ind. 1977,983. 237 G. Seoane, Cur. Org. Chem. 2000,4, 283. 238 A. H. Rose, Industrial Microbiology, Butterworths, Washington, 1961, p. 264. 239 G. Grue-Sorensen, I. D. Spenser, J. Am. Chem. SOC.1988,110, 3714. 240 V. Kren, D. H. G. Crout, H. Dalton, D. W. Hutchinson, W. Koenig, M. M. Turner, G. Dean, N. Thomson, J. Chem. SOC.,Chem Commun. 1993,4, 341. 241 Z.Guo, A. Goswami, K. D. Mirfakhrae, R. N. Patel, Tetrahedron: Asymm. 1999, 10, 4667. 242 U.-M. Billhardt, P. Stein, G. M. Whitesides, Bioorg. Chem. 1989, 17, 1. 243 E. Rieke, S. Barry, K. Mosbach, Eur. J. Biochem. 1979,100,203. 244 a) S. S. Patel, H. D. Conlon, D. R. Walt,]. 226

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

974

I

74 Formation ofC-C Bonds

Org. Chem. 1986,51, 2842; b) S. S. Patel, D. R. Walt,]. Bid. Chem. 1987, 262, 7132. 245 T. Ouyang, D. R. Walt, S. S. Patel, Bioorg. Chem. 1990,18,131. 246 a) D. Byrom, TIBECH 1987,5,246; b) T. Suzuki, T. Yamane, S. Shimizu, Appl. Microb. Biotech. 1986, 24, 370. 247 a) 0. P. Peoples, S. Masamune, C. T. Walsh, A. J. Sinskey,J. Bid. Chem. 1987, 262, 97; b) S. Masamune, M. A. J. Palmer, R. Gamboni, S. Thompson, J. T. Davis, S. F. Williams, 0. P. Peoples, A. J. Sinskey, C. T. Walsh, J. Am. Chem. Soc. 1989, 111, 1879. 248 a) D. Seebach, M. F. Zuger, Tetrahedron Lett. 1984,25,2747; b) D. Seebach, M. F. Zuger, Helv. Chirn. Acta 1982, 65,495. 249 K. S. Kyler, M. J. Novak, NATO A S 1 Ser., Ser. C 1992, 381, 3. 250 E. E. van Tamelen, E. C. Leopold, S. A. Marson, H. R. Waespe,]. Am. Chem. SOC.1982, 104,6479. 251 a) J. Bujons, R. Guajardo, K. S. Kyler,]. Am. Chem. Soc. 1988, 110,604; b) J. C. Medina, K. S. Kyler, J . Am.Chem. Soc. 1988, 110, 4818. 252 J. C. Medina, R. Guarjardo, K. S. Kyler,]. Am. Chem. Soc. 1989, 111, 2310. 253 B. J. Robustell, I. Abe, G. D. Prestwich, Etrahedron Lett. 1998, 39,9385. 254 S. M. Grundy, Nav Eng.]. Med. 1988,319, 24 and references therein.

255

a) T. Koyama, M. Matsubara, K. Ogura,

/. Biochem. 1985,98,457;b) T. Koyama,

256

257 258 259

260 261 262 263 264

265

K. Ogura, F. C. Baker, G. C. Jamieson, D. A. Schooley,]. Am. Chem. Soc. 1987, 109, 2853. a) M. Nagaki, H. Kannari, J. Ishibashi, Y. Maki, T. Nishino, K. Ogura, T. Koyama, Bioorg. Med. Chem. Lett. 1998,8, 2549; b) T. Gotoh, K. Ogura,]. Biochem. 1992, 112, 20. T. Nagasawa, H. Yamada, Appl. Biochem. Biotech. 1986, 13, 147. M. Welch, R. S. Phillips, Bioorg. Med. Chem. Lett. 1999, 9, 637. R. S. Phillips, L. A. Cohen, U.Annby, D. Wensbo, S. Gronowitz, Bioorg. Med. Chem. Lett. 1995, 5, 1133. M. J. Sloan, R. S. Phillips, Bioorg. Med. Chem. Lett. 1992, 2, 1053. M. Lee, R. S. Phillips, Bioorg. Med. Chem. Lett. 1992, 2, 1563. J. J. Milne, J. P. G. Malthouse, Biochem. SOC. Trans. 1996, 24, 133s. K. Kim, P. A. Cole, Bioorg. Med. Chem. Lett. 1999, 9, 1205. R. S. Phillips, R. L. Von Tersch, J. G. Fletcher, A. H. Lai, Amino Acids: Chem. Biol. Med. 1990, 166. D. Hebel, D. C. Furlano, R. S. Phillips, S. Koushik, C. R. Creverling, K. L. Kirk, Bioorg. Med. Chem. Lett. 1992, 2, 41.

14.7

Enzymatic Synthesis of Cyanohydrins

Martin H . Fechter and Hefried Criengl

In the last decade, optically pure cyanohydrins (a-hydroxynitriles) have become a versatile source for the synthesis of a variety of chiral building blocks. Diverse methods for the enantioselective synthesis of cyanohydrins have been published and reviewed Ill. Besides enzyme catalyzed methods, hydrocyanation or silylcyanation of aldehydes or ketones controlled by chiral metal complexes or cyclic dipeptides, as well as diastereoselective hydrocyanation of chiral carbonyl compounds, have been applied with moderate success. However, the most advantageous preparations of optically active cyanohydrins, with respect to the obtained enantioselectivities, are the enzymatically controlled approaches discussed in the present chapter. Two common enzyme systems are described and reviewed['-16]: firstly, esterases or lipases, which have been employed

14.7 Enzymatic Synthesis ofcyanohydrins

R’\

C=O + H C N

R2’

A

PH R’/f\CN R

Figure 14.7-1. Cyanohydrin forrnation: R’ = alkyl, cycloalkyl, aryl, heteroaryl; RZ = H, alkyl.

for the resolution of racemic cyanohydrins or alkoxynitriles, and secondly, oxynitrilases - also known as hydroxynitrile lyases (HNLs),which catalyze the reversible formation of cyanohydrins (Fig. 14.7-1),using HCN and aldehydes or ketones. About 3000 plant species are known to release HCN from their tissues, a process which is known as cyanogenesis[”. “1. Storage compounds are cyanohydrins where the hydroxy function is glycosylated to a carbohydrate or protected as a fatty acid ester. The plant defence mechanism in the case of sugar compounds is a two-step reaction. Initially a glycosidase liberates the cyanohydrin moiety, which is cleaved either spontaneously by base catalysis or enzymatically by the action of oxynitrilases to release the corresponding carbonyl compound and HCN [I9]. The application of an HNL was the subject of one of the earliest reports in the field of biocatalysis, namely the synthesis of mandelonitrile from benzaldehyde and hydrocyanic acid using a crude enzyme preparation obtained from almonds (termed “emulsin”)f2O1. However, little attention was paid to this d i s c o ~ e r y [ ~until ~ - ~ ~the I 1960s, when this enzyme (E. C. 4.1.2.10) was isolated, characterized12”261,and used for the preparation of enantiomerically enriched (R)-cyanohydrinsfrom aromatic and aliphatic aldehydes [27-291. The first examination of an (S)-oxynitrilasein millet revealed that this enzyme only seedlings (Sorghum bicolor, E. C. 4.1.2.11) [3s331 accepts aromatic substrates. At this time, the best enantiomeric excess obtained was 87% for the formation of (R)-mandelonitrile; other aldehydes gave even lower enantiomeric ratios. 14.7.1

The Oxynitrilases Commonly Used for PreparativeApplication

At present, the oxynitrilases from eleven cyanogenic plants (from six plant families) have been purified and characterisedL9. lo1.Theproperties of a selection of these are outlined in Table 14.7-1.The oxynitrilases E. C. 4.1.2.10 from Rosaceae (e.g. Prunus sp.) contain the cofactor FAD. However, the latter is not involved in redox reactions. Instead, it seems to have a structure-stabilizing effect, and its presence might be Some of these enzymes are glycosylated explained on evolutionary grounds [34-3G1. and most of them are constructed from several Recently the crystal structure of the oxynitrilase from Hevea brasiliensis (E.C. 4.1.2.39) was r e p ~ r t e d [ ~ ’ -The ~ ~ ]enzyme . was found to contain a large j3-sheet which is surrounded by a-helices and a cap region on both sides. The active site is deeply buried inside the protein and connected to the surface by a narrow channel. Similar discoveries were published very recently for the (S)-HNLfrom Manihot esculenta (E.C. 4.1.2.39)[42.431. A big step forward, toward further applications of the Prunus amygdalus HNL, was achieved by the Kratky group by elucidating the crystal structure of this

I

975

976

I

14 Formation ofC-C Bonds Table 14.7-1.

Oxynitrilases available for organic synthesis.

Plant

Enzyme availability

Pnrnus amygdalus Almonds

Natural substrate

Substrate acceptance for syntheses

Stereoselectivity

Linum usitatissimum

All R’ and R2 Aliphatic aldehydes and ketones

(4

Flax seedlings overexpression

Sorghum bicolor

Millet seedlings (S)-4-Hydroxymandel- Aromatic aldehydes onitrile

(S)

Hevea brasiliensis

Rubber tree leaves overexpression

Manihot esculenta Manioc leaves overexpression

(R)-Mandelonitrile Acetone cyanohydrin (R)-2-Butanonecyanohydrin

(R)

Acetone cyanohydrin

All R’ and R2

(S)

Acetone cyanohydrin

All R’ and R2

(S)

Until quite recently, all HNLs had to be isolated from natural sources. To supply the industrial demand, enzymes from Hevea b r a ~ i l i e n s i s [461~ ~ , Manihot escuhave been successfully l e n t ~ ~ [and ~ ~ Linum - ~ ~ ] usitatissimum (E. C. 4.1.2.37) overexpressed in several microorganisms. Presently, the (S)-cyanohydrinof 3-phenoxybenzaldehyde is used as an intermediate for various pyrethroid type insecticides; this reaction is catalyzed by overexpressed (S)-HNLfrom H. brasiliensis and the cyanohydrin is produced on the hundred ton per year scale[53]. In contrast to the HNLs from H. brasiliensis and M.esculenta, where aliphatic and aromatic aldehydes or ketones function as substrates, the HNL from Sorghum bicolor only catalyzes the formation and cleavage of aromatic (S)-cyanohydrins[54-591. The most convenient natural sources of enzymes yielding products with (R)-stereochemistry are almonds (Prunus amygdalus)I ‘ ‘ [ and almond meal r6’]. In addition to Linum usitatissimum[621,other sources of (R)-HNL have also recently been reported[63,641. Concerning the substrate spectrum, the P.amygdalus HNL catalyzes the HCN addition to aliphatic and aromatic carbonyl moieties, the L.usitatissimum oxynitrilase accepts only aliphatic ketones or aldehydes I6’1. 14.7.2 Oxynitrilase Catalyzed Addition of HCN to Aldehydes

(R)-Hydroxynitrile lyases. For preparative applications, (R)-HNLfrom almonds has been extensively investigated. Brussee et al. [“, 671 showed that without enzyme purification a crude extract from almond meal in aqueous methanol using in situ HCN generation from a solution of KCN in an acetate buffer affords cyanohydrins in up to 93 % ee. Apple meal, in the form of unpurified enzyme preparations, accepts sterically hindered aldehydes (e.g. pivalaldehyde) as substrates, leading to ( R ) cyanohydrins with high enantiomeric purity (usually ee > 90 %) [63, “1. A purified enzyme from Prunus amygdalus supported on cellulose using nonaqueous systems was employed for the first time by Effenberger and co-workersLG9]. Optimal results were obtained by almost completely suppressing the non-enzymatic HCN addition

74.7 Enzymatic Synthesis of Cyanohydrins

using ethyl acetate as solvent. In this manner enantiomeric purity could be improved. Besides crystalline cellulose (Avicel), other hydrophobic enzyme immobilization systems such as Celite were used[70,711. Utilizing the natural support, unpurified almond meal in organic solvents with small amounts of aqueous phase (4%), provides products with ees of up to 99%rG1, 72-751. Similar results were achieved with so-called "microaqueous systems" In order to reduce the amount of racemic cyanohydrin produced by chemical conversion, low concentrations of HCN were used by employing a relatively safe and convenient source of this reagent: Kanerva has developed a method where HCN acetone c y a n ~ h y d r i n [73, ~ ~77-791. , diffuses into the reaction mixture from a second flask[74].Wandrey used an enzyme membrane reactor for the continuous production of product employing an (R)-HNL. In a production run the volumetric yield was increased to 2400 g (R)-mandelonitrilel L x day with a residence time of just 3.8 min. The enzyme consumption was 17000 U/kg product["]. Applying a biphasic system a second industrial scale procedure was Based on these findings, four parameters (pH, concentration of HCN and benzaldehyde, temperature) were optimized to obtain a throughput of 6700 g (R)-mandelonitrile/Lx day. A novel synthesis of (R)-cyanohydrinswas described based on the use of cross-linked and subsequently polyvinyl alcohol-entrapped (R)oxynitrilases. These immobilized lens-shaped biocatalysts have a well-defined macroscopic size in the mm range, show no catalyst leaching and can also be efficiently recycled. Furthermore, this immobilization method is cheap, and the entrapped ( R ) oxynitrilases gave similar results to those using free enzymes. Accordingly, ( R ) cyanohydrins were obtained in good yields and with high enantioselectivitiesof up to "3

ee > 99 % [821.

Some substrates, e. g. acrolein, gave only low optical purity with the P. amygdalus HNL. The catalytic capability of (R)-specific HNL from L. usitatissimum for the preparation of aliphatic cyanohydrins was investigated L50, 51, 651 and gave encouraging results (ee of up to 99 %). (S)-Hydroxynitrile lyases. As already mentioned, the (S)-hydroxynitrilelyase from Sorghum bicolor adds HCN only to aromatic and heteroaromatic aldehydes. Initial investigationswere performed on the natural substrate 4-hydroxybenzaldehyde,and rather promising results concerning the enantiomeric excess were found[83].These results were confirmed and extended using a suspension of enzyme immobilized on "1 or etiolated shoots of S.bicolor[861 in diisopropyl ether. The Avicel ~ellulose['~~ Sorghum enzyme was one of the first recombinant hydroxynitrile lyases[871,overexpressed in Escherichia coli. In parallel to this work the H . brasiliensis HNL was also o~erexpressed[~~], giving access to sufficient quantities of this enzyme both on a preparative scale and for industrial use. To date only a few preparative applications for Sorghum HNL[74]are known because of the narrow substrate range. A similarly broad substrate range to that for the (R)-HNLfrom Prunus amygdalus is revealed by the (S)-HNLs from Manihot esculenta and Hevea brasiliensis (E.C. 4.1.2.39). Detailed sequence studies have revealed high homologies between both enzymes (M. e s ~ u l e n t a ["I;~ ~H , . brasilien~is[~~~ 451). This result was confirmed by the crystal structures. The latter was solved for H. brasiliensis in Grazc3'] and for the M. esculenta enzyme in S t ~ t t g a r t [ ~Expectations ~]. that these enzymes would be similar

I

977

978

I with respect to substrate specificitywere established by experimental data from both 74 Formation of C-C Bonds

groups. The cyanoglycoside Linamarin was found in 1965 in the seeds of the rubber tree (Hevea brasiliensis) [891.Tw0decades later the corresponding hydroxynitrile lyase was described[”. 911. Studies regarding the synthetic potential of this enzyme with respect to the preparation of optically pure cyanohydrins started with the wild type113s92-941 . A s already mentioned groundbreaking results were obtained with the synthesis of the (S)-cyanohydrinof 3-phenoxybenzaldehyde,this being a precursor for some important synthetic pyrethroids 95-971. HNL from Manihot esculenta Crantz (termed E.C. 4.1.2.37 at this time because E. C. 4.1.2.39 was not created earlier than 1999[”])was purified to homogeneity from young leaves of the cyanogenic tropical crop plant cassava in 1994[471.First experiments demonstrated a broad substrate range, but only unsatisfactory optical purities were obtained[”]. The overexpression of the cloned M.esculenta HNL gene in E. coli increased the accessibility and specific activity of the biocatalyst as well as the ee of produced cyanohydrins [871. A selection of substrates with typical enantioselectivitiesof the obtained cyanohydrins, from the respective HNLs, is shown in table 14.7-2. 14.7.3 HNL-Catalyzed Addition of Hydrogen Cyanide to Ketones

Preparative elaboration of (R)-cyanohydrinsof ketones employing oxynitrilase from Prunus amygdalus was first investigated in organic solvents l1O71. Alkyl methyl ketones were obtained in moderate yields and in high optical purity, whereas with alkyl ethyl ketones the chemical and optical yields were reported to be lower[lo8].The alteration, working with almond meal instead of purified enzyme, resulted in an astonishingly high enantiomeric excess [‘*I. Similar results with 98 % ee for the (R)-cyanohydrinof butyl methyl ketone, were obtained[lo9I. (R)-Oxynitrilasefrom Linum usitatissimumhas been used for the synthesis of ( R ) butan-Zone cyanohydrin on a preparative scale I”]. Concerning (S)-ketonecyanohydrins, impressive results were gained on aliphatic or aromatic ketones, e. g. acetophenone cyanohydrin. The latter was obtained using the oxynitrilase from H . brasiliensis. (40% conversion, 99 % ee)I‘[ or M . esculenta HNL (87% conversion, 98% ee) [‘lo]. In table 14.7.3.the results gained by HNL-catalyzed conversions of selected methyl ketones to the corresponding cyanohydrins are shown. 14.7.4 Transhydrocyanation

The transcyanation (exactly termed transhydrocyanation ) of aromatic and aliphatic aldehydes with acetone cyanohydrin, catalyzed by (R)-oxynitrilaseto give cyanohydrins (see Fig. 14.7-2.), was first performed in This innovative method avoids the use of free HCN as the cyanide source and is mostly accompanied

14.7 Enzymatic Synthesis ofcyanohydrins Table 14.7-2.

Aldehydes R-CHO as substrates for oxynitrilase-catalyzed cyanohydrin formation.

R

HNL

Ph

R

(El-PhCH=CH 3-PhO(CsH4)

Source

Conversion 1%1

ee I%]

P. a.

99 97 99 98

S S S

S. b.

H. b. M. e.

100 97 96 100

R S

P. a. H. b.

54 93

87 98

R

P. a. S. b. H. b.

99 93 99

98 96 99

S

S PhCH20CH2

S

H. b.

92

12

PhCHz

R

83 44

88 99

10 88

10 93

S

P. a. H. b.

PhCHzCH2

R S

L. u. H . b.

2-CH30(C&)

R

P. a. H. b.

65 61

96 77

S S

P. a. S. b. H . b.

85 93 80

98 89 99

R

P. a.

S S S

S. b.

47 54 49 82

99 71 95 98

96 95

99 (S)" 80(R)" 98(R)"

96 88 98 98

99 87 98 92

P. a. S. b.

71 64 98 85

99 (S)" 91(R)" 99(R)" 96(R)*

R

P. a.

S S

S. b. H. b.

S R

M. e.

95 95 49 98

99 98 99 98

100 92 70

74 98 56

99 80 100

98 86 92

96

99

S

3-CH@(C&)

4-CH3

0 (c6H4)

2-fury1

R

R S S

3-fuvl

2-thienyl

R S S S R S S

S 3-thienyl

CH4H

S S

(E)-CHjCH=CH

R S

S

(E)-CH,(CH2)4CH=CH S

H. b. M.e. P. a. S. b. H. b.

P. a. S. b.

H . b. M. e.

H. b. M . e.

L. u.

H. b. M.e. P. a.

H . b. M. e. H . b.

80

Reference

I

979

14 Formation ofC-C Bonds Table 14.7-2.

(cont.).

HNL

Source

(E)-CH3(CH2)2CH=CH

S S

H . b. M . e.

R

Conversion ["h]

ee ["h]

46 82

95 97

(Z)-CH,(CHz)*CH=CH

S

H . b.

35

80

CH~(CH~)~CGC

S

H . b.

88

80

3-cyclohexenyl

R

P. a. H . b.

86 87

99

P. a. H.b. M . e.

90 95 100

99 99 92

P. a. H. b.

82 35

96 85

P. a. H.b.

72 81

97 96

P. a. L. u.

M . e.

H . b.

99 91 80 70

98 98 80 88

P. a. L. u. H . b. M . e.

99 100 80 91

83 93 81 95

P. a.

58

L. u. H.b. M. e.

100 80 80

92 89 67 94

S

Reference

55

-L Change of product configuration owing to a priority replacement according CIP rules Abbreviations: HNL, hydroxynitrile lyase; P. a., Prunus amygdalus; S. b., Sorghum bicolor; H . b., Hevea brasiliensis; M. e., Manihot esculenta; L. u., Linum usitatissimum.

Figure 14.7-2. Principle o f transhydrocyanation: R' aryl, heteroaryl; R2 = H, alkyl.

= alkyl,

cycloalkyl,

by a slight decrease in ee compared to standard conditions. It was optimized in Turku [721 by comparing the feasibility of powdered almond meal as a catalyst to that of a purified enzyme preparation in an organic solvent. as the cyanide donor The attempt to use racemic 2-methyl-2-hydroxyhexanenitrile was rewarded by obtaining aliphatic o-bromo cyanohydrins from the corresponding aldehydes in 90-97% ee[781. As a biocatalyst, (R)-oxynitrilasewas used.

14.7 Enzymatic Synthesis ofcyanohydrins

I

981

Table 14.7-3. formation.

R

Methyl ketones R-CO-Meas substrate for oxynitrilase-catalyzedcyanohydrin

I"/.]

Source

Conversion

R R

P. a.

L. u. M . e.

80 100 91

76 95 18

P. a. L. u. H . b. M . e.

70 100 99 36

97 93 74 69

P. a.

73 59 58

99 99 80

P. a.

H.b.

54 99

90 98

P. a. H . b. M.e.

57 86 69

98 99 91

H. b. M.e.

49 81

78 28

S S

P. a. H. b. M . e.

14 40 87

90 99 98

S

H. b.

74

95

S

R

R S

S

R S S

R S

R S

S S

S

R

H.b. M . e.

ee

rh]

HNL

Reference

.

.

Abbreviations: HNL, hydroxynitrile lyase; P. a,, Pnrnus amygdalus; L. u., Linum usitatissiwum: H. b., Heuea brasiliensis; M. e., Manihot esculenta.

14.7.5

Experimental Techniques for HNL-Catalyzed Biotransforrnations

HNL catalysis in aqueous medium. Reaction in aqueous solution is performed with an appropriate acidic component and alkali cyanide for in situ development of the required HCN. The following procedure is a typical e~arnple['~I. To a stirred solution of 1 mmol aldehyde in 1.7 mL of 0.1 mol/L sodium citrate buffer (pH 4.0),1 mL of a crude cytosolic extract of (S)-HNLfrom Heuea brasiliensis (100IU/mL) was added and the mixture was cooled down to 0 "C. Subsequently, 2 mmol of potassium cyanide adjusted to pH 4.0 with cold 0.1 moljL citric acid (17 mL) were added in one portion. After stirring for 1 h at 0-5 "C, the reaction mixture was extracted with methylene chloride (3 x 50 mL). The combined organic layers were dried over anhydrous sodium sulfate and the solvent was removed to give the crude cyanohydrin. This was then purified by column chromatography on silica gel using petroleum ether / ethyl acetate acidified with trace amounts of anhydrous HCl as the eluent. H N L catalysis in organic medium. A significant advancement in cyanohydrin production was made by performing the transformation in organic solvents immiscible with water. It has been observed that there is virtually no spontaneous

982

I chemical addition of HCN to the carbonyl moiety[48, 74 Formation ofC-C Bonds

A representative protocol for cyanohydrin formation in organic solvents with immobilized oxynitrilase is the A suspension of Avicel cellulose (0.5 g) in 0.05 mmol/L phosphate buffer (pH 4.5, 10 mL) containing ammonium sulfate (4.72 g) was stirred for 1 h, and a solution of (S)-HNLfrom Sorghum bicolor (50 pL, 1000 IU/mL, specific activity 70 IU/mg) was added. The mixture was stirred at room temperature for 10 min and filtered, and the immobilized enzyme was suspended in diisopropyl ether (10 mL). After addition of aldehyde (2 mmol) and dry liquid HCN (300 pL, 7.5 mmol), the mixture was stirred until all aldehyde had reacted. After removal of the immobilized enzyme, the filtrate was concentrated to yield the crude cyanohydrin. H N L catalysis in biphasic medium. Biphasic solvent mixtures were reported 11'1 as well as (S)-HNLfrom Hevea b r a ~ i l i e n s i s ~ "*I.l ~ ~ employing (R)-oxynitrilase[812 A typical procedure is as follows Is']. Freshly distilled benzaldehyde (37.1 g, 0.35 mmol), HCN (12.2 g, 0.45 mmol) and (R)-oxynitrilase(78 mg) were dissolved in 225 mL of methyl t-butyl ether (MTBE) and 250 mL of citrate buffer (50 mmol/L, pH 5.5) at 22 "C. After stirring for 20 min the MTBE layer was separated and the aqueous layer was extracted once with 25 mL of MTBE. The combined organic layers were dried over MgS04, filtered and concentrated under reduced pressure. Yield: 45.2 g (97%), purity 98%, ee 98%. The aqueous layer was reused in a series of four consecutive experiments using the same amounts of reagents in the organic phase. A total of 185.5 g of benzaldehyde was converted into 226 g of (R)-mandelonitrileusing 78 mg of (R)-oxynitrilase (0.035 69-71, ",

lo'.

lo7, 111-1141.

wt%).

Transhydrocyanation for HCN generation. An alternative method of employing organic solvents that allows the safe use of HCN is transhydrocyanation L7', 73, 77-79, 'I6, 'I7]. An example of cyanohydrin formation using acetone cyanohydrin as the cyanide source is given in the following procedure[77]. To a solution of 120 mg (1 mmol) of phenylacetaldehyde and 110 mg (1.3 mmol) of acetone cyanohydrin in 11 mL of diethyl ether at 23 "C, 0.5 mL of (R)-oxynitrilase buffer solution (10mg/mL, 0.4 mol/L acetate buffer, pH 5.0) was added. The mixture was stirred for 18 h at 23 "C and diluted with 50 mL of ether. The aqueous phase was extracted with 2 x 10 mL of ether and the combined organic phases were dried over anhydrous magnesium sulfate. Evaporation of solvent gave a pale amber liquid which was chromatographed on a flash silica gel column in 1 : 30 : 50 ethyl acetate / benzene / dichloromethane to afford 122 mg (83%) of cyanohydrin, ee 88%. 14.7.6

Resolution of Racemates

Oxynitrilase as catalyst. It is possible to treat a racemic cyanohydrin with a (R)-or (S)HNL to decompose selectively one enantiomer of this mixture (exemplified in Fig. 14.7-3.).The (R)-HNLfrom Prunus amygdalus was used for the resolution of racemic cyanohydrins. Employing a biphasic system, namely citrate buffer/diisopropyl ether (40:l) at 39"C, catalytic amounts of PhNH2 and semicarbazide were added for

14.7 Enzymatic Synthesis ofCyanohydrins I983

Figure 14.7-3. Enantioselective HNL catalyzed decomposition of racemic cyanohydrins: R’ = alkyl, cycloalkyl, aryl, heteroaryl; R2 = H, alkyl.

Figure 14.7-4. Lipase-catalyzed formation of optically enriched cyanohydrins: R’ cycloalkyl, aryl, heteroaryl; R2 = acyl; R’ = H, acyl.

= alkyl,

aldehyde capture. In this manner the (S)-cyanohydrinof 3-phenoxybenzaldehyde was obtained with 91 % ee at 50% conversion[”’]. Recently, almond meal was used for the resolution of rac-2-hydroxy-2-phenylpropanenitrile. Under the optimized conditions, (S)-2-hydroxy-2-phenylpropanenitrile, as the less reactive enantiomer, was obtained in 98-99% ee at approximately 50% conversion[’18].In a similar way the (S)-cyanohydrinwas afforded from racemic 2-methyl-2-hydroxyhexanenitrilewith P. amygdalus HNL in more than 90% ee~73, 781

Esterase or lipase as catalyst. Application of hydrolytic enzymes is realized in three different systems: enzymatic hydrolysis or transesterification of racemic cyanohydrin esters (see Figure 14.7-4.)as well as enzymatic acylation of racemic cyanohy&ins [W 1201 A series of cyanohydrin acetates with an e.e. up to 98% has been prepared by enzymatic hydrolysis of their racemic acetates in the presence of an esterase from Pseudomonas sp. [1371. Lipoprotein lipase from Pseudomonas sp. catalysed irreversible transesterification using enol esters was applied to the resolution of different aromatic cyanohydrins[13’, 139]. The enantioselectivehydrolysis of the racemic acetate by Arthrobacter lipase gave the optically pure (S)-3-phenoxybenzaldehyde cyanohydrin. The unhydrolysed (R)acetate was reracemised by heating with triethylamine and submitted again to enzymic hydrolysis[l4O]. In addition, the resolution of the racemic acetate ester of the cyanohydrin of 3-phenoxybenzaldehydeusing a highly enantioselective lipase from Pseudomonas sp. was carried out recently with an e.e. of >9G%[1411.Both the cyanohydrin esters and the free cyanohydrins (which are prone to racemization) can be isolated as enantiomers with high optical purity (ee 97%) on a preparative scale by the hydrolysis of the racemic butyrates with Candida cylindracea lipase and Pseudomonas sp. lipase11211. A one-pot synthesis of optically active cyanohydrin acetates from aldehydes has been accomplished by lipase-catalyzed kinetic resolution coupled with in situ formation and racemization of cyanohydrins in an organic solvent. Racemic cyanohydrins, generated from aldehydes and acetone cyanohydrin in diisopropyl ether under the catalysis of basic anion-exchangeresin, were acetylated stereoselectivelyby a lipase from Pseudomonas cepacia (Amano)with isopropenyl acetate as an acylating

984

I

14 Formation ofC-C Bonds

HH R'

H

H o ~ NH,

Y A CN H

grs

PZR2

HoR

j y

XYMS

1.c"

lq

-

HH

TBDMSOR

P

CN

- g r

CHO

R&

NH2

\I

- CR2 K k

CN

A7Y H

Y

\

x;

g th % \i

COOR2

X

&R CN

NHR3

H COOH

to

T COOR' H

CN

CN

Figure 14.7-5. Follow-up reactions of optically pure cyanohydrins: a) TBDMSCl/imidazole[66,671; b) R2MgX/ether,NaBH4, H30+[84f1251. , c) CH3Mgl/ether,H J O + [ ~d) ~ ]R2CH2Mgl/ether, ; MeOH, R3NH2,NaBH4[126];e) IAH[84];f) H30+['O71;g) R20H/CHC13/wolfatite[127];h), R3CI/Nal/ k) CHsCN/pyr/O 0C[1271;i) DIBALH/hexane/ -78 "C, conc. HCI/MeOH [1271; j) R2S02Cl/pyrL1021; KN3/DMF, (inversion) [12']; I) LiAIH4/Et20/-80 "C, phosphate buffer pH 7.0/-70 "C['28];m) potassium phtalimide/DMF, (inversion) ["'I; n) KOAc/DMF/r.t., (inversion)[lo2.12', l2'I; 0)conc. HCl/r.t. or lipase['02a12', lZ91; p) Me3SiCl/pyr/Et20/0to +25 0C[1301;q) diethylaminosulfurtrifluoride (DAST)/CH2CI2/-80to +25 "C['301;r) DIBALH/CHzC12/ -78 "C, 1 N H2S04"3'1.

reagent. The (S)-cyanohydrinwas preferentially acetylated by the lipase, while the unreacted (R)-isomer was continuously racemized through reversible transhydrocyanation catalyzed by the resin. These processes consequently led to a one-pot conversion with up to 94% ee in 63-100% conversion yields[122,1231. The Pseudomonas aeruginosa lipase (immobilised on hyflo Super-Cel) catalysed kinetic resolution of (~ac)-2-(acetyloxy)-2-(pentafluorophenyl)acetonitnle gave enantiomerically pure cyanohydrin and its antipodal ester [142-1441.

14.7 Enzymatic Synthesis ofCyanohydrins

14.7.7 Follow-up Chemistry of Enantiopure Cyanohydrins

',

Optically pure cyanohydrins are important synthetic building blocks[', 2 . 4. Is* 1241, as can be seen from Fig. 14.7-5. in selected examples. Both functional groups, the hydroxy and the cyanide moiety, can be easily converted into a large range of other chiral intermediates such as a-hydroxy acids and esters, ahydroxy aldehydes and ketones, p-amino alcohols and a-fluorocyanides. These structural moieties are present in a large number of industrially valuable products such as drugs, agrochemicals,flavorings and fragrances. 9.

14.7.8 Safe Handling of Cyanides

Hydrogen cyanide smells like bitter almonds, although many people cannot smell it at all[132]. Cyanide is a fast acting poison in the human body; it affects the ability of all cells to breathe. Severe breathing difficulties develop very rapidly when cyanide is swallowed, inhaled, or absorbed through the skin. Cyanide poisoning symptoms in the early stages indude: general weakness, breathing difficulty, headache, nausea, giddiness, vomiting, the victims breath smelling like bitter almonds, and irritation of the nose, mouth, and throat. Hydrogen cyanide is liberated by the addition of acid to cyanide compounds. ~ ] . limits The TLV (threshold limit value) for HCN is 11 mg/m3 or 10 ~ p m [ ' ~These include the potential contribution of skin absorption to the overall exposure. Proper gloves should be worn when handling dry sodium cyanide. Rubber gloves and splash-proof goggles should also be worn when substantial amounts of sodium cyanide solution are used. All reaction equipment in which cyanides are used or produced should be placed in well-ventilated hoods, and it should be determined immediately whether anyone has been exposed to cyanide vapors or liquid splashVapor-detector tubes sensitive to 1 ppm of HCN are available commercially. The presence of free cyanide ion in aqueous solution may be detected by treating an aliquot of the sample with ferrous sulfate and an excess of sulfuric acid. A precipitate of Prussian blue indicates that free cyanide ion is present. More sophisticated for continuous warning is the use of electrochemical sensors for HCN detection. Waste solutions containing cyanides treated with sodium hypochlorite are converted to harmless cyanate, which can be further processed to ammonia and carbon dioxide by addition of diluted sulfuric acid to pH 7. Surplus HCN gas can be neutralized by aqueous sodium hydroxide and then oxidized. Caution has to be advised with liquid hydrogen cyanide because bases including sodium hydroxide and sodium cyanide may initiate a violent polymerization[133]. Explosive hazards can occur on exposure of HCN to air in the presence of sources of ignition (flammable limits in air: 5.640% v/v) including heat (polymerizes explosively at 5 0 4 0 "C)and when HCN is stored for long periods of time.

I

985

986

I

74 Formation of C-C Bonds

14.7.9 Conclusions and Outlook

The enzymatic synthesis of enantiopure cyanohydrins has been brought to a high stage of development. Both (R)- and (S)-cyanohydrins are accessible for a broad variety of substrates in as a rule excellent yield and enantiopurity. Following recent progress in overexpression, HNLs are also available in quantities needed for industrial production. The procedures for safe handling of cyanides are well established so that they do not restrict the exploitation of HNLs.

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B: Enzym. 2000,11, 37-43.

M.Inagaki, J. Hiratake, T. Nishioka, I. Oda.

Bull. Inst. Chem. Res., Kyoto Univ. 1989. 67, 132-135. 120 M.Inagaki, J.Hiratake, T. Nishioka, J. Oda, / . A m . Chem. Soc. 1991, 113,936&9361. 121 F. Effenberger, B. Gutterer, T. Ziegler, E. Eckhardt, R. Aichholz, Liebigs Ann. Chem. 1991,47-54. 122 M. Inagaki, J.Hiratake. T. Nishioka, J. Oda, /. Org. Chem. 1992,57, 5643-5G49. 123 M. Inagaki, A. Hatanaka, M. Mimura, J. Hiratake, T. Nishioka, J. Oda, Bull. Chem. Soc./pn. 1992,65, 111-120. 124 C.G.Kruse, H. W. G e l d , G. J. M. van Scharrenburg, Chim. Om'1992,10,59-63. 125 F. Effenberger, B. Gutterer, J. Syed, Tetrahedron: Asymmetry 1995,6,2933-2943. 126 J. Brussee, A. van der Gen, Recl. Trav. Chim. Pays-Bas 1991,110,25-26. 127 F. Effenberger, M. Hopf, T. Ziegler, J. Hudelmayer, Chem. Ber. 1991,124,1651-1659. 128 F. Effenberger, U. Stelzer, Angau. Chem. 1991,103,866867:Angau. Chem., Int. Ed. End., 1991,30,873-874. 129 F. Effenberger, U. Stelzer, Chem. Ber. 1993, 126,779-786. 130 U. Stelzer, F. Effenberger, Tetrahedron: Asymmetry1993,4,1G1-164. 131 W. R. Jackson, H. A. Jacobs, G. S. Jayatilake, B. R. Matthews, K. G. Watson, Aust. /. Chem. 1990,43,2045-2062.

132 Anon., Dangerous Prop. h d . Mater. Rep.

1992, 12,116-130.

133 National Research Council, Prudent Prac-

tices in the Laboratory: Handling and Disposal ofchemicals, National Academic Press, Washington, D. C., 1995. 134 R.J. Lewis, Sax's Dangerous Properties of Industrial Materials, 10th Edition, Wiley, New York, N.Y., 1999. 135 R. L. Somerville, Chem. Eng. Prog. 1990,86, 64-68. 136 R. Baxter, Ind. Finish. (Wheaton, 111) 1977, 53,38-41. 137 A. Van Almsick, J. Buddms, P. HoenickeSchmidt, K. Laumen, M. P. Schneider, 1. Chem. Soc., Chem. Commun. 1989, 1391-1393. 138 S . H. Hsu, S. S. Wu, Y. F. Wang, C. H. Wong, Tetrahedron Lett. 1990, 31, 6403-6406. 139 Y.F. Wang, S.T. Chen, K. K.C. Liu, C. H. Wong, Tetrahedron Lett. 1989, 30, 1917-1920. 140 S. Mitsuda, H. Yamamoto, T. Umemura, H. Hirohara, S. Nabeshima, Agnc. Bid. Chem. 1990,54,2907-2912. 141 A. Fishman, M. Zviely, Tetrahedron: Asymmetry 1998,9,107-118. 142 T.Sakai, Y. Miki, M. Tsuboi, H. Takeuchi, T. Ema, K. Uneyama, M. Utaka, /. Org. Chem. 2000,65, 2740-2747. 143 T.Sakai, Y. Miki, M. Nakatani, T. Ema, K. Uneyama, M. Utaka, Tetrahedron Lett. 1998, 39, 5233-5236. 144 T. Sakai, 'I: Takayama, T. Ohkawa, 0. Yoshio, T. Ema, M. Utaka, Tetrahedron Lett. 1997,38,1987-1990.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyriqht 0Wiley-VCH Verlaq GmbH, Weinheim 2002

I991

15 Reduction Reactions 15.1

Reduction of Ketones Kaoru Nakamura and Jomoko Matsuda 15.1.1

Introduction 15.1.1.1

Enzyme Classfication and Reaction Mechanism

Research on the asymmetric reduction of ketones by biocatalysis is expanding, and its practical applications to organic chemistry have resulted in success in the enantioselective synthesis of pharmaceuticals, agrochemicals and natural products [1-41. It is attracting increasing attention because of the following advantages: - providing a green and sustainable process (natural catalysis), - high enantio-, regio- and chemo-selectivity compared with most man-made

reagents and catalysts, - achiral ketones can be transformed into the corresponding alcohols with 100%

-

yield and 100% ee theoretically, whereas kinetic resolution of racemic substrates by hydrolytic enzymes such as lipases yields only 50% of products to achieve 100% ee. the resulting alcohol functionality can be easily transformed, without racemization, into other useful functional groups such as halides, thiols, amines, azides, etc.

Dehydrogenases, classified under E.C.l.l., are enzymes that catalyze reduction and oxidation of carbonyl groups and alcohols, respecti~ely[~]. The natural substrates of the enzymes are alcohols such as ethanol, lactate, glycerol, etc. and the corresponding carbonyl compounds, but unnatural ketones can also be reduced enantioselectively. To exhibit catalytic activities, the enzymes require a coenzyme; most of the dehydrogenases use NADH or NADPH, and a few use flavin, pyrroloquinoline quinone, etc. The reaction mechanism of the dehydrogenase reduction is as follows:

992

I

15 Reduction Reactions

Step 1 A holoenzyme (an enzyme with its coenzyme) binds a ketone. Step 2 A hydride on the coenzyme is transferred to the ketone to produce an alcohol. (With concurrent oxidation of the coenzyme) Step 3 The enzyme releases the product alcohol. Step 4 The oxidized coenzyme is transformed back into the reduced form. (With concurrent oxidation of an auxiliary substrate) There are four stereochemical patterns in the transfer of the hydride from the coenzyme, NAD(P)H,to the substrate (Step 2) as shown in Fig. 15-1.With E l and E2 enzymes, the hydride attacks the si-face of the carbonyl group, whereas with E3 and E4 enzymes, the hydride attacks the re-face, which results in the formation of R and S alcohols, respectively. On the other hand, E l and E3 enzymes transfer the pro-R hydride of the coenzymes, and E2 and E4 enzymes use the p r o 3 hydride. Examples of the El-E3 enzymes are as follows: E l : Pseudornonas sp. alcohol dehydrogenaseL6] Lactobacillus kefir alcohol dehydrogenase171 E2: Geotrichurn candidurn glycerol dehydrogenase[8-101 Mucorjavanicus dihydroxyacetonereductase [11] E3: Yeast alcohol dehydrogenase[l2] Horse liver alcohol dehydrogenase[l3-l61 Moraxella sp. alcohol dehydrogenase[I' E2

-ry ADPR

Figure 15-1. Stereochemistry ofthe hydride transfer from NAD(P)H to the carbonyl carbon on the substrate (S is a small group and L is a large group).

sifac re face

I

ADPR

15.1.1.2

Coenzyme Regeneration

Reduction of the substrate accompanies the oxidation of the coenzyme (Step 2). Before the next cycle of the reduction of the main substrate can occur, the coenzyme has to be reduced (Step 4). Many methods for the regeneration of the reduced form of coenzyme [NAD(P)H]have been developed, so that only a catalytic amount of the coenzyme is required for the reaction. The coenzyme regeneration methods can be classified into two types: - two-enzmye system: different enzymes reduce the substrate and NAD(P)+, - one-enzyme system: the substrate and NAD(P)+ are both reduced by the same

enzyme.

IS. 1 (a)

(b)

0

Enzvme 1

u

main substrate for asymmetric reduction NADH

COZ

OH

Enzyme 1

for asymmetric reduction NAD(P)H

NAD'

Enzyme 2

0

Reduction ofKetones

NAD(P)*

A

A

HCOzH auxiliary substrate

U

Enzyme2

l

x auxiliary substrate

Figure 15-2. Regeneration o f NAD(P)H: (a) Two-enzyme system using a formate dehydrogenase as an auxiliary enzyme and formic acid as an auxiliary substrate; Enzyme 1 = Enzyme for the reduction o f t h e main substrate Enzyme 2 = Formate dehydrogenase (b) one-enzyme system using 2-propanol as an auxiliary substrate. Enzyme 1 = Enzyme 2 For example alcohol dehydrogenase from Thermoanaerobium brockiil"* "1, Pseudomonas sp. 1'1, Lactobacillus kefirlq, and Ceotrichum candidum I2O, "1.

One of the examples of the two-enzymesystem uses a formate dehydrogenase for the recycling of coenzyme [Fig. 15-2(a)]['*3, 22-24] . It catalyzes oxidation of HC02H to C02 in order to drive the reduction of NAD' to NADH. The system is one of the most widely used due to the advantages such as: 1)the enzyme is commercially available, 2) COZ can be easily removed from the reaction, 3) formate is strongly reducing, therefore no back reaction occurs, and 4) C02 and HCOzH are innocuous to enzymes. For example, the reduction of ethyl 4-chloro-3-oxobutanoateby a carbonyl reductase from Rhodococcus eruthropolis uses NAD'/formate dehydrogenase as shown in Fig. 15-3["I. As exemplified, the system is very useful for the recycling of NADH. However, it does not accept NADP', so it cannot be used for the direct reduction of NADP+. For the reduction of NADP' using formate dehydrogenase, catalytic amounts of NAD' and NAD(P)' transhydrogenase are required. Changing the coenzyme specificity of a formate dehydrogenase using genetic methods is discussed in Sect. 15.1.3.7. Two-enzyme systems using glucose dehydrogenase or glucose-6-phosphatedehydrogenase (commercially available enzymes) have also been widely employed [2G-311. Carbonyl reductase from Rhodococcus erythropolis

0

cIdk/cozEt

*

NAD+ HC02H formate dehydrogenase

Conv. 100%. ee >99% (R)

Carbonyl reductase from Rhodococcuseiythropolis COpEt

*

NAD' HCOpH formate dehydrogenase

OH CI &COzEt

OH A C O z E t

Conv. 49%, de 95%, ee>95% (2R. 3s)

Figure 15-3. Examples of reduction using the formate/formate dehydrogenase N A D H recycling system[251.

I

993

994

I

15 Reduction Reactions

0 CI&COpEt

HLADH

t

NAD' Glucose Glucose dehydrogenase from Baci//uscereus

OH C I A C O Z E t

yield '*%

ee "%(5)

Figure 15-4. Example o f reduction using glucose/glucose dehydrogenase NADH recycling

F

Figure 15-5.

F Photosynthetic microorganism Synechococcussp. PCC 7942

yield 90% ee >99% (S) F

Utilization o f light energy for an efficient

They oxidize glucose or glucose-6-phosphate to form gluconolactone or gluconolactone-6-phosphate, respectively, which is spontaneously hydrolyzed to give gluconic acid. Both NAD' and NADP' act as substrates for these enzymes. For example, a thermostable glucose dehydrogenase form Bacillus cereus was used to recycle NADH in the asymmetric reduction of ethyl 4-chloro-3-oxobutanoate by horse liver alcohol dehydrogenase (HLADH)as shown in Fig. 15-4[261. Another example of a two-enzyme system involves molecular hydrogen and a hydrogenase[lI.Hydrogenases catalyze the reduction of NAD' or other redox dyes by dihydrogen. The system is attractive because dihydrogen is inexpensive, strongly reducing and innocuous to enzymes and NAD(H), and no by-product is formed. However, a drawback is the extreme sensitivity of the hydrogenase enzymes to inactivation by dioxygen, preventing this system from being widely used. To provide an environmentally friendly system, photochemical methods have been developed, which utilize light energy for the regeneration of NAD(P)H['. 32, 331. Recently, the use of cyanobacterium, a photosynthetic biocatalyst, for the reduction was reported where the effective reduction occurred under illumination (Fig. 155) [321. When a photosynthetic organisms is omitted, the addition of a photosensitizer is necessary. The methods utilize light energy to promote the transfer of an electron from a photosensitizer via an electron transport reagent to NAD(P)+[']. One-enzyme recycling systems are also well developed. One of the most frequently utilized is the alcohol-alcohol dehydrogenase system as shown in Fig. 15-2(b).The system does not need an auxiliary enzyme, but an auxiliary substrate is necessary. Ethanol or 2-propanol is frequently used as an auxiliary substrate. For example, HLADH uses ethanol as shown in Fig. 15-G[13-16]and Thermoanaerobium brockii"'. *'I, Pseudomonas sp. 16], Lactobacillus kefid71, and Geotrichum candidum[20.21] alcohol dehydrogenases recycle NAD(P)H by employing an excess of 2-propanol.A detailed investigation of the type and amount of the auxiliary substrate needed by G. candidum revealed that it can use 2-alkanols from 2-propanol to 2-octanol (and cyclopentanol as well), and 15-20 equivalents of the supplementary alcohol are necessary to shift the equilibrium (between the oxidation and reduction) towards the reduction of the main substrate. Because a much higher concentration

15.1 Reduction of Ketones

I

995

0

%, Yield 29% Yield 35% ee 100% ee36%

Yield 11% ee 100%

H

Yield 26% ee >97%

Figure 15-6. Reduction of heterocyclic ketones by HIADH using ethanol as an auxiliary

s u b ~ t r a t e [ l6]. ’~~

H

Yield 48% ee 60%

of the auxiliary substrate to that of the main substrate is required, 2-propanol is deemed most suitable for synthetic purposes due to its high volatility. Electrochemical regeneration of NAD(P)H represents another interesting The system involves electron transfer from the electrode to the method [34-361. electron mediator such as methyl viologen or acetophenone etc., then to the NAD(P)+ (which is catalyzed by an electrocatalyst such as ferredoxin-NADP’ reductase or alcohol dehydrogenase, etc.) [34]. Other methods involve the direct reduction of NAD’ on the Both one-enzyme systems and two-enzyme systems have been reported. 15.1.1.3 Form ofthe Biocatalysts: Isolated Enzyme vs. Whole Cell

Enzymes in a pure form, in a partially purified form, and in the whole cell can be used for organic synthesis, and each has advantages and disadvantage^[^]. The proper choice of the form of the biocatalyst is important because it affects the enantio-, regio- and chemo-selectivities,the requirement (or not) of a coenzyme and an auxiliary enzyme, the ease of catalyst preparation and work up procedures, etc. as shown in Table 15-1. The most widely used whole cell biocatalyst is bakers’ yeast. Since it has many different kinds of enzymes, many kinds of substrate can thus be reduced, and various types of the reactions are expected. For example, 0-keto esters, aromatic, aliphatic, cyclic and acyclic ketones can be reduced with high yield[’. 37-391. Therefore, it is a versatile “all-round”reagent. However, since bakers’ yeast contains many kinds of dehydrogenases, some of them may be S selective, while others are R selective, so that the enantioselectivities can be low to high depending on the substrate structure. Further degradation of the product may also be a problem, again associated with the fact that there are many kinds of enzymes in the cell. Not only the enzymes but also the cellular components such as coenzymes and carbohydrates are conserved in the cell, which makes the whole cell processes favorable. For example, the addition of an expensive coenzyme and an auxiliary enzyme for coenzyme regeneration is not necessary, which makes the system simple and economical when comparing with the equivalent isolated enzyme process.

996

I

75 Reduction Reactions Table 15-1.

The form of biocatalyst: whole cell vs. isolated enzyme

Parameter

Whole Cell

Isolated Enzyme

Kinds of enzymes Kinds of reactions Regio- and enantioselectivity Coenzyme Catalyst preparation Work up Example

Many Many Low to high Unnecessary Easy Difficult Bakers' yeast

One One High Necessary Difficult Easy Horse liver alcohol dehydrogenase

However, the product isolation may be complicated due to large amounts of biomass and metabolites. On the other hand, isolated enzyme processes also have many advantages. The problem associated with the product isolation and overmetabolism can be avoided using an isolated enzyme. More importantly, chemo-, regio-, and enantioselectivities of isolated enzyme systems are usually higher than that of whole cell processes because two competing enzymes with different stereoselectivities are not present. One of the most widely used isolated enzymes is horse liver alcohol dehydrogenase (HLADH) which reduces, for example, S-heterocyclic ketones to give the corresponding tetrahydrothiopyran-4-01with 100% ee [Fig. lS-G(a)]['I. However, when the selectivity is so high, the substrate specificity is not wide; thus HLADH can reduce cyclic ketones with excellent enantioselectivity but cannot reduce acyclic ketones. Another advantage of the isolated enzyme system is that the reaction pathway can be understood and predictions made. For example, for HLADH, the crystal structure[4s421 and the active site (diamond lattice) model[13,141 are available to understand the reduction, whereas, in a whole cell process, even the catalytic species itself may not be clear. In summary, whole cell and isolated enzyme biocatalysts both have various advantages and disadvantages. Using a recombinant yeast having the gene of a requisite enzyme is the way to access a single predominant enzyme in a microorganisms, a strategy which will be further discussed in Sect. 15.1.3.2. 15.1.1.4

Origin of Enzymes

Enzymes from various sources have been used for asymmetric reductions in organic synthesis. Microorganisms are the most important sources. There are a huge number of species (mostly in soil), containing a variety of enzymes. Commercially available microbial dehydrogenases are alcohol dehydrogenases from yeast, Thermoanaerobium brockii (TBADH),and the hydroxysteroid dehydrogenase from Pseudomonas testosteroni. One of the most attractive kinds of microorganisms for organic synthesis is a thermophilic microorganism such as Themoanaerobium brockii['8, l91, or T h e m o anaerobacter ethanolicus, etc. (43-491. The thermostability of the dehydrogenase en-

15.1 Reduction of Ketones

I

997

zymes from these microorganisms is very high: TBADH is stable even at 86 "C [I8* and an alcohol dehydrogenase from Thermoanaerobacter ethanolicus can be used at 50-60 'T[",471. Since the enzymes with high thermostability usually have a high tolerance to organic solvent or substrates, the enzymes from thermophilic microorganisms are most suitable for organic synthesis. Another interesting class of biocatalyst encompasses the photosynthetic micro"]. Owing do the high growth rate, a large amount of the organisms, the algaeL3** biomass for use as the biocatalyst is available. Importantly, such organisms can use light energy as power for coenzyme recycling as described in Sect. 15.1.1.2, so an environmentally friendly system can be constructed using them. The second most widely studied source of enzymes are mammalian enzymes as exemplified by horse liver alcohol dehydrogenase (HLADH).Detailed investigations on this enzyme have been reviewed elsewhere[13,1'. The third and least studies source is from plant cell cultures, which have only recently been used in biocatalysis[51-571. Although the number of species available are much less than microorganisms, plants possess a much larger gene. More importantly since plants can effect photosynthesis, different types of enzymes exist in plants to those of microorganisms. Therefore, different enzymes which catalyze unique reactions with man-made substrates may be expected. Despite the strong possibility of the discovery of interesting enzymes, plant cell cultures have not been fully investigated for use in biocatalysis due to their relatively slow growth rate. 15.1.2

StereochemicalControl 15.1.2.1

Enantioselectivity of Reduction Reactions

The synthesis of enantiomerically pure compounds is becoming increasingly important for research and development in chemistry and especially in the pharmaceutical industry, as chiral drugs now represent close to one-third of all pharmaceutical sales world wide["]. In most of the cases, one enantiomer is more effective as a drug than the other. The influence on the environment is also different between the enantiomers; different enantiomers of chiral pollutants in soils are preferentially degraded by microorganisms in various environments [Go]. Therefore, synthetic methods exhibiting extremely high enantioselectivitiesare necessary. The enzymatic reactions occurring in Nature involving natural substrates usually show very high enantioselectivities. On the other hand, with man-made substrates the enantioselectivity can also be high (> 99% ee) but this is not always the case as shown in Fig. 15-7. Low enantioselectivity results when the catalyst is a low selectivity enzyme [Fig. 15-7 (C)] and/or when there are more than two competing enzymes with different enantioselectivities [Fig. 15-7 (D)]. In case (C), either an enzyme or substrate has to be changed. On the other hand, in case (D), a change in a microorganism or substrate as well as a change in reaction conditions may be effective in improving the enantioselectivity. In case (D), by choosing the proper

998

I

75 Reduction Reactions OH

&R+

(high]: Figure 15-7.

QH

RAW

(3

OH

Enantioselectivity of the product and improvement methods.

Inhibitor of R-enzyme or activator of S-enzyme

-

OH

OH

RAR’

0

Inhibitor of S-enzyme or activator of R-enzyme

OH

Figure 15-8. Synthesis of both enantiomers using one microorganism by choosing appropriate conditions.

conditions, both enantiomers can be synthesized by using only one microorganism; when a selective inhibitor for an S-directing enzyme or on R-directing enzyme is added to the reaction mixture, the (R)-alcohol or (S)-alcohol will be enantioselectively produced, respectively, as shown in Fig. 15-8. 15.1.2.2 Modification of the Substrate: Use of an “Enantiocontrolling” Group

The enantioselectivity of a biocatalytic reduction can be controlled by modifying the substrate because the enantioselectivity of the reduction reaction is profoundly affected by the structure of substrates. For example, in the reduction of 4-chloro3-oxobutanoate by bakers’ yeast, the ester moiety can be used to control the stereochemical course of the reduction 161-631. When the ester moiety was smaller than a butyl group, then (S)-alcoholswere obtained, and when it was larger than a pentyl group then (R)-alcoholswere obtained as shown in Fig. 15-9. After the reduction, the ester moiety can be exchanged easily without racemization, so both enantiomers of an equivalent synthetic building block are obtained using the same reaction system by changing an “enantiocontrolling”group, the ester moiety. The “enantiocontrolling”group can also be introduced into the keto esters at the a- or a‘-positions. For example, sulfur functionalities such as methyl- and

15.1 Reduction ofKetones

S

R

yeast cell

n=1-4

-100

I

999

n=5-12

I , , , , , ,

0

2

4

6

8 1012

n Figure 15-9. Stereochemical control on yeast-catalyzed reduction by changing the ester group[61-631.

0

-

Bakers' yeast

/ICC02Me

;

j

%%%

' e*

Bakers'yeast

b

0 Ph02S&C02Me

QH

hCozM

f

+CO2Me

SR

t

ee ,96%

OH

0

ee87%

SR

Bakers' yeast

OH

OH PhOzS&C02Me

_c

ee 98% M eO C, , - ) 2

Figure 15-10. Stereochemical control on yeast-catalyzed reduction by introducing sulfur function aIities "1. lS43

Bakers' yeast ....*

-i

ee69% ee96%

~0 l B a k e r s ' y e a s t OH ~

OH

p

I

Figure 15-11. Improvement of enantioselectivity by substituting iodide at the para position; yeast reduction followed by dehalogenation (dh) [651.

phenylthiorG4land phenyl~ulfonyl[~~1 groups can be used to improve the enantioselectivities as shown in Fig. 15-10. Other types of ketones can also be modified to improve the enantioselectivities, and various functionalities can be used to modify the substrate to produce the corresponding alcohol with higher enantioselectivities.For example,the reduction of acetophenone by yeast results in the formation of phenylethanol in 69 % ee, whereas the reduction of p-iodoacetophenone followed by the dehalogenation results in a product of96% ee (Fig. 15-11)[G51. As shown above, the substrate modification and "de"modification steps can be used to improve the enantioselectivity, although on the negative side the strategy may introduce extra steps into a synthetic route.

IS

Reduction Reactions

Table 15-2.

Screening for the synthesis of important chiral building blocks. Microorganisms Result screened

Reactions

Candida magnoliae

H o- go (D'

0

C02Et

-

OH -CO2Et

(2R,3s)

90 g/L, 96.6% ee (99% ee after heat treatment)

Reference

67

bacteria 191actinomycetes 59 45 mg/mL stoichiometric yield Rhodoforula minufa IF0 0920:86% ee 230 yeasts Candida parapsilosis IF0 0708:87% ee 68 81 molds 42 basidiomycetes Aspergillus niger IF0 4415:87% ee

450 bacteria

Klebsiella pneumoniae IF0 3319 99% de, >99% ee, 99% yield (2Kg in 200 L fermentor)

70

15.1.2.3

Screening of Microorganisms

Screening for a novel enzyme is a classical method and one of the most powerful tools available to find the system to convert a selected ketone into a desired alcoh01[~"~~1. It is possible to discover a suitable enzyme or microorganisms by the application of the newest screening and selection technologies that allows rapid identification of enzyme activities from diverse sources 166]. Enzyme sources for screening can be soil samples, commercial enzymes, culture sources, a clone bank, etc. From these sources, enzymes which are regularly expressed and enzymes which are not expressed in the original host can be tested to establish whether they are suitable for the transformation of certain substrates [661. For example, 400 yeasts were screened for the reduction of ethyl 4-chloro-3-oxobutanoate, and Candida magnoliae was found to be the best one as shown in Table 15-2[", 7 2 , 731. For the reduction of ketopantoyl lactone, various kinds of microorganisms were screened, and several microorganisms which produce D-pan pantoy lactone stoichiometrically at a concentration of 45 mg mL-' with high enantioselectivity were found['*]. For the reduction of ethyl 2-methyl-3-oxobutanoate,out of 450 bacteria, Klebsiella pneumoniae I F 0 3319 and 4 other strains were found to give the corresponding (2R, 3S)-hydroxyesterswith more than 98 % de and > 99 % ee["]. Screening techniques have also been applied for the purpose of drug synthesis. For example, a key intermediate in the synthesis of the anti-asthma drug, Montelukast, was prepared from the ketone 1 by microbial transformation as shown in Fig. 15-12[711. The biotransforming organism, Microbacterium campoquemadoensis (MB5614),was discovered as a result of an extensive screening programme.

15. I Reduction of Ketones

1

-

Montelukast (Singulair)

Figure 15-12. Reduction of a ketone by Microbacterium carnpoquemadoensis (MB5614) in a synthesis of the anti-asthma drug, M o n t e l ~ k a s t [ ~ ’ ] .

Table 15-3.

Control on diastereoselectivity by heat treatment74.

QH COzEt

Yeast

/\/COZEt ee , 9 5 2 1

Yeast cell

Svn (%)

OH

+ ee >95% Anti (%)

No heat treatment 50 OC, 30 min

heat + inhibitor

15.1.2.4

Treatment of the Cell: Heat Treatment

Treatment of the cell before the reaction is sometimes an effective method of controlling the selectivity of some biocatalysts. When reducing with a whole cell and the selectivity is not as is desired due to the presence of plural enzymes with different selectivities, heat treatment of the cell to selectively deactivate one or more enzymes can change the selectivity of the reduction. For example, the diastereoselectivity in the yeast reduction of 2-allyl-3-oxobutanoatewas changed from anti-selectivityto synselectivity by pre-treatment of the yeast before the reaction as shown in Table 15-3[741. In this case, the diastereoselectivityis further improved to 96 : 4 by using an enzyme inhibitor. Another example is the use of heat treatment as a supplement to the screening process. The enantioselectivity of the reduction of ethyl 4-chloro-3-oxobutanoateby Candida magnoliae was improved from 96.6% ee (S) using untreated cells to 99% ee (S) with heat treated cells rG71. 15.1.2.5

Treatment of the Cell: Aging

When a whole cell system is used for a reduction, the substrate is usually added to the cultivation medium after a certain growth period, or to the mixture of the

I

’Ool

1002

I

75 Reduction Reactions

medium and freshly harvested cells. However, when the mycelium of a local strain of Geotrichum candidum was not used immediately after growth, but filtered and preincubated by shaking in deionized water for 24 hours at 27 "C ("agedmycelium"), then used for the reduction of ethyl 3-oxobutanoate, the stereochemistry of the product alcohol was different from that obtained from the reduction using fresh mycelium [75-781. When fresh mycelium was used, the enantioselectivity and the absolute configuration of the product shifted from S (26% ee) to R (58% ee) on raising the substrate concentration from 1 to 20 g L-'. When aged mycelium was used, the absolute configuration was always R and showed constant enantioselectivity (ca. 50% ee) regardless of the substrate concentration, although the reduction proceeded at a slightly slower rate. In the aging process, an S-forming activity, was lost, leaving unaffected either one low-specificityreducing enzyme with major R-forming activity, or several enzymes having opposite enantioselectivitiesbut similar KM values. 15.1.2.6 Treatment of the Cell: High Pressure Homogenization

High pressure homogenization is a new technology in food processing. It was found that the same technology can be applied to effect the microbial reduction of chemical compounds[791.The cell culture with substrate (such as acetophenone, 5-hexenz-one, etc.) was poured into the high pressure homogenizer, and then it was incubated for 48 h and the enantioselectivity of the product was evaluated. During the process the reaction mixture was forced under pressure through a narrow gap where it was subjected to rapid acceleration [l (blank experiment), 500, 1000, 1500 bar] after which it undergoes an extreme drop in pressure. Various strains of Saccharomyces cerevisiae and Yarrowia lipolytica are utilized in the reduction processes and higher enantioselectivitieswere generally achieved albeit in lower yields than the standard process. 15.1.2.7 Treatment of the Cell: Acetone Dehydration

A dried cell mass is often used as a biocatalyst for a reduction, since it can be stored

for a long time and can be used whenever needed, without cultivation. One of the useful methods to dry the cell mass is acetone dehydration["]. For example, the cells of Geotrichum candidum I F 0 4597 were mixed with cold acetone (-20 "C) and the cells were collected by filtration[20.21]. The procedure was repeated five times and then the cells were dried under reduced pressure. The dried cells (acetone powder of G. candidum I F 0 4597: APG4) were obtained; they can be stored for a long time in the freezer. The drying of the cell not only aids the preservation of the cell but also contributes to the stereochemical control as shown in Table 15-4.The reduction of acetophenone catalyzed by G. candidum I F 0 4597 resulted in poor enantioselectivity [28% ee(R)]. When the form of the catalyst was changed from wet whole-cell to dried powdered-

75.7 Reduction of Ketones

Acetone treatment o f Ceotrichum candidum for the improvement of enantioselectivity", ".

Table 15-4.

Acetone dried cell (APG4)

Untreated whole cell P'h

*

NAD' or NADP'

Ph

*

oH j\Ph

2-propanol or cyclopentanol >99% ee (S)

28% ee (RJ

Catalyst

Coenzyme

Additive

Yield ("h)

Untreated whole cell Acetone dried cell (APG4) Acetone dried cell (APG4) Acetone dried cell (APG4) Acetone dried cell (APG4)

none none NAD' NAD' NADP'

none none 2-propanol cyclopentanol cyclopentanol

52 0 89 97 8G

ee (%)

28(R) -

>99(S) >99(S) >99(S)

cell (APG4), no reduction was observed, which would indicate the loss of the necessary coenzyme(s) and/or coenzyme regeneration system@)during the treatment of the cells with acetone. Addition of coenzyme, NAD', did not have a significant effect on the yield. Addition of 2-propanol resulted in only a small increase in the yield, but a significant improvement in the enantioselectivity was observed. Surprisingly, addition of both NAD' and 2-propanolprofoundly enhanced both chemical yield and enantiomeric excess. Addition of NADH, NADP' or NADPH instead of NAD' and addition of cyclopentanol instead of 2-propanol also gave enantiomericallypure alcohol in high yield. The improvement in the enantioselectivity from 28 % (R) to > 99 % (S) was due to the suppression of every enzyme which reduces the substrate, followed by the stimulation of an S-directingenzyme by the addition of the coenzyme and an excess amount of 2-propanol, agents which push the equilibrium towards the reduction of the substrate. It was confirmed, by separating the enzymes in the powder, that many S- and Rdirecting enzymes exist in the biocatalyst. The addition of coenzyme and cyclopentano1 stimulates only one particula S-enzymebut not other S-enzymesand R-enzymes because the spec& S-enzyme can oxidize cyclopentanol [concomitantly reducing NAD(P)'], while other S- or R-enzymes cannot use cyclopentanol as effectively["]. This is a very interesting case where the reduction with a cell initially having both Sand R-directing enzymes was modified and resulted in excellent S enantioselectivity. 15.1.2.8

Cultivation Conditions of the Cell

The dependence of enantioselectivity in microbial transformations on the cultivation conditions of the microorganisms has also been investigated[82-8G1.The enzymes induced during the growth phase and during starvation are certainly different, therefore the enantioselectivity of the product may be different when two competing enzymes with different enantioselectivitiescatalyze the reduction. Since the enzyme

1004

I

75 Reduction Reactions

reducing the non-natural substrate is not usually known, cultivation conditions which induce the desired enzyme have to be found by trial and error. For example, the effect of cultivation time and different carbon sources on the enantioselectivity of the reduction of sulcatone by some anaerobic bacteria has been Another example is the investigation on the effect of the medium concentrations for cultivation of Geotrichum candidurn I F 0 4597 on the enantioselectivity of the reduction of acetophenone derivatives. The yield of R-alcohol (the minor enantiomer) increased with the medium concentration; therefore, the medium concentration was kept low, optimally to produce the S-enantiomer[**]. The effect of the aeration during cultivation on the enantioselectivity of bakers' yeast production of 3-hydroxyesters has also been reported["]. Inducers such as a substrate analog may also induce the desired enzyme to improve the enantioselectivity. 15.1.2.9

Modification of Reaction Conditions: Incorporation o f an Inhibitor

In the case of the observation of poor overall enantioselectivity due to the presence of two competing enzymes with different enantioselectivities,one of the most straightforward methods to improve the enantioselectivity is the use of the inhibitor of the unnecessary enzyme(s). Ethyl chloroacetate, methyl vinyl ketone, allyl alcohol, allyl bromide, sulfur compounds, Mg2+,Ca2+,etc. have been reported as inhibitors of enzymes in yeast [87-971. For example, the low enantioselectivity in the yeast reduction of P-keto ester was improved by addition of ethyl chloroacetate or methyl vinyl ketone as described in Fig. 15-13. The enzymes inhibited and those not inhibited were identified by enzymatic studies using purified enzymes["]. The mechanism of the inhibition is reported to be non-competitive. These inhibitors were also used to improve the enenatioselective reduction of inhibitor

0

inhibitor

a

L-enzyme-l

without any inhibitor

i OH R A C H 2 C O 2 R ' low ee

Figure 15-13. enzymes187.

Improvement o f t h e enantioselectivity by using an inhibitor of undesired 88. 971

15.1 Reduction of Ketones

uR r: UR Allyl alcohol or methyl vinyl ketone

OH

F3c

Bakers’ yeast

OH 0

F3C

Allyl bromide Figure 15-14.

F,C

aR (R)

Stereochemical control using an inhibitor[89].

fluorinated diketones (Fig. 15-14). By applying a suitable inhibitor, both enantiomers of the alcohol can be obtained using only one kind of microorganism, namely bakers’ 15.1.2.10

Modification of Reaction Conditions: Organic Solvent

Organic solvents have been used widely for esterifications and transesterifications using hydrolytic enzymes to shift the equilibrium towards esterification by avoiding hydrolysis. Organic solvents can also be used for reductions using dehydrogenases [98-1091 . They can be used to control the overall enantioselectivity of the reduction, when there are more than two competing enzymes with different enantioselectivities,K M and V., Enzymatic reactions follow the Michaelis-Menten equation, therefore, the rate of the enzyme catalyzed reaction depends on the substrate concentration. When an organic solvent is introduced, most organic substrates usually dissolve in the organic phase, and the effective substrate concentration in the aqueous phase around the enzyme decreases. The change in substrate concentration by the addition of the organic phase causes the change in the enzyme species catalyzing the reduction. For example, as shown in Fig. 15-15, if the K M for an S-directingenzyme is much smaller than that for an R-directing enzyme, and V, for the S-directing enzyme is much smaller than that for the R-directing enzyme, then when the substrate concentration is low, the S-enzymewill dominate, whereas at high substrate concentration, the Renzyme will dominate the biotransformation. In fact, when the yeast reduction of ethyl 2-oxohexanoatewas conducted in water,

’

R-enzyme

velocity €enzyme

IS1

Substrate concentration At low substrate conc. the Senzyme predominates.

At high substrate conc. the R-enzyme predominates.

Figure 15-15. Effect of the substrate concentration on enantioselectivity of the reduction with the system having both an S-enzyme with small KM and small V, and an R-enzyme with large KM and large V,.

I

1005

1006

I

15 Reduction Reactions

(9 Further,, decomposition

BuKCO,Et yeast

\

in benzene

Figure 15-16.

OH

B U L C o 2 E t (4

Stereochemical control by using an organic solvent.

Table 15-5.

Mechanism of stereochernical control using benzene: kinetic parameters of yeast a-keto ester reductases (YKERs)'". 'lo.

(s-')

Enzyme

Enantioseledivity

KM (mM)

t a t

YKER-I YKER-IV YKER-V YKER-VI YKER-VII

R R

8.40 0.142 5.72 1.03 27.3

1.53 4.59 27.8 2.10 127

s R S

VmaX(U kg-' yeast)

37.7 41 649 1774 501

both (R)- and (S)-alcoholswere produced and the (S)-alcohol was obtained as the major product as a result of the further enantioselective decomposition of the (R)enantiomer (Fig. 1S-16)['00slo9]. H owever, when the biotransformation was conducted in benzene, then the (R)-alcoholwas formed selectively in high yield. KM and V,, for all enzymes existing in yeast and catalyzing the reduction were determined and it was found that an R-enzyme, YKER-IV, has a KMwhich is smaller than other enzymes by an order of magnitude (Table 15-S), and, therefore, predominantly catalyzes the reduction in benzene"". llo, lll]. 15.1.2.1 1

Modification of Reaction Conditions: Use of a Supercritical Solvent

Supercritical fluids, materials above their critical pressure and critical temperature (Fig. 15-17),have been attracting attention as solvents with the advantages of gas-like low viscosities and high diffusivities coupled with their liquid-like solubilizing power r112]. Supercritical carbon dioxide (SCCOZ) has the added benefit of an environmentally benign nature, nonflammability, low toxicity, ready availability, and ambient critical temperature (T, = 31.0 "C) that is suitable for biotransformations. The attraction of combining natural catalysts with a "natural" solvent has been the driving force behind a growing body of literature on the stability, activity and specificity of enzymes in SCCOZ.The first report on biotransformations in superand the benefit of using supercritical fluids for critical fluids was in 1985[113-115], biotransformations has been demonstrated, e. g. through improved reaction rates, etc.[116.

1171

Recently the alcohol dehydrogenase from Geotrichum candidum was found to

15.1 Reduction of Ketones Figure 15-17.

Phase diagram o f carbon dioxide.

_______------

-100

0

100

Temperature (OC)

I:

Immobilized Geotrichum candidurn cell R ~ R ' Supercritical CO?

R = CH, CH,F, etc. R' = Ph, 0- , rn- or pfluorophenyl, Ph-(CH,),-

etc.

-

OH R-R'

-

Yield 11 96% ee 96 - 299%

Figure 15-18. Reduction of fluoroketones by Ceotrichum candidum I F 0 5767 in supercritical C O Z " ' ~ ~ .

catalyze the reduction of fluoroacetophenones etc. in scCO2 at around 100 atm and 35 "C (Fig. 15-18)[''81. The enantioselectivity obtained was equivalent to the system using an organic solvent. 15.1.2.12 Modification of Reaction Conditions: Cyclodextrin

Cyclodextrin has also been used to control the enantioselectivity of bioreduction~['~'-'~'1.When added to a reaction mixture, the substrate can reside in the cyclodextrin, which decreases the effective substrate concentration around the enzyme and results in the domination of reactions involving enzymes with low KM. The effect can be demonstrated by the reduction of ketopantoyl lactone by yeast. The enantioselectivity was improved from 7 3 % to 9 3 % by adding P-cyclodextrin to the reaction mixture. The improvement in enantioselectivity of the reduction in the presence of enzymes with different enantioselectivitiesand KMvalues by decreasing the substrate concentration was confirmed by the ineffectiveness of a-cyclodextrin which is too small to include the substrate. It was also confirmed by dilution of the reaction mixture, which improved the enantioselectivity in the absence of cyclodextrin. 15.1.2.1 3 Modification o f Reaction Conditions: Hydrophobic Polymer XAD

A decrease in the effective substrate concentration around the enzyme but not in the bulk can also be achieved using hydrophobic polymer XAD instead of using cyclodextrin or an organic solvent['22-'26].For example, the technique was used in the reduction of methyl benzyl ketone by Zygosaccharomyes rouxii for the synthesis of LY300164, a noncompetitive antagonist of the AMPA subtype of excitatory amino acid The adsorption properties of the resin on both substrate and

I

1007

1008

I

15 Reduction Reactions

Zygosaccharomyces rouxii

a ketone loading

5K2,3-benzodiazepine (LY300164) Ydrl.7

I'

,,,+

-

" croui?$ 2 g/L of product alcohol

Adsorbed species: Initial: 80 g/L of ketone final: 75 glL of product alcohol 0

Y

I

*OH 0

Y

Figure 15-19. Decrease in the effective substrate concentration around the enzyme by using hydrophobic polymer XAD[12z].

product allowed a ketone loading of 80 g L-l, while limiting the effective solution concentration of both substrate and product to sublethal concentrations of 2 g L-' (Fig. 15-19). The hydrophobic resin has also been used for the purpose of controlling selectivity[123, 1241. E~ antioselectivity, chemoselectivity and space-time yields of the yeast reduction of a,P-unsaturated carbonyl compounds were impressively enhanced. The distribution of substrates and products between the resin and the water phase showed that the improved selectivity could be attributed to the control of substrate concentration. The powerful influence of the hydrophobic resin was also demonstrates in the Geotrichum candidum catalyzed reduction of simple aliphatic and aromatic ketones [126]. For example, the enantioselectivity of the reduction of 6-methylhept-5-en2-one was improved from 27 % ee ( R ) to 98 % ee (S). 15.1.2.14

Modification o f Reaction Conditions: Reaction Temperature

Reaction temperature is one of the parameters that affects the enantioselectivity of a r e a ~ t i o n [ ~ ~For - ~the ~ ] oxidation ]. of an alcohol, the values of k,,,/KM were determined for the ( R ) -and (S)-stereodefining enantiomers; E is the ratio between them. From the transition state theory, the free energy difference at the transition state between (R)-and (S)-enantiomers can be calculated from E [Eq. (2)], and AAG is in turn the temperature function [Eq. (3)) The racemic temperature (T,) can be calculated as shown in Eq. (4).With these equations, T, for 2-butanol and 2-pentanol of the Thertnoanaerobacter ethanolicus alcohol dehydrogenase was determined to be 26 "C and 77 "C, respectively.

Moraxella sp. TAEl23 alcohol dehydrogenase 0°C

NADH

-OH

ee >99% (SJ

75.7 Reduction of Ketones

Figure 15-20. Reduction of 2-butanone by the alcohol dehydrogenase from Moraxella sp. TAE123 a t 0 0C[171.

E = (kcat / KM)R/ ( k a t / from transition state theory - RTln(E) = AAGI AAGI = AAHI - TAASI When AAGI = 0, T, = AAHI 1 AASI

(4)

Since the transition state for alcohol oxidation and ketone reduction must be identical, the product distribution (under kinetic control) for reduction of 2-butanone and 2-pentanone is also predictable. Thus, one would expect to isolate (R)2-butanol if the temperature of the reaction was above 26 "C. On the contrary, if the temperature is less than 26 "C, (S)-2-butanolshould result. In fact, the reduction of 2-butanone and 2-pentanone at 37 "C resulted in 28 % ee (R)- and 44% ee (S)-alcohol, respectively, as expected [431. The temperature range that can be used for a biocatalytic reduction is very wide because alcohol dehydrogenases from various types of microorganisms (thermophilic and psychrophilic) are available. The extremely high stability of enzymes from thermophilic microorganisms are discussed in Sect. 15.1.1.4. On the other hand, conducting reactions at temperatures as low as 0 "C is also possible using an Antarctic psychrophile [171. For example, the reduction of 2-butanone, which is an extremely challenging substrate for enantioselective reduction, with alcohol dehydrogenase from Moraxella sp. TAE123, at 0 "C afforded (S)-2-butanol in > 99% ee (Fig. 15-20). 15.1.2.15

Modification of Reaction Conditions: Reaction Pressure

The effect of high hydrostatic pressure (400 bar) on microbial reductions of the ketones such as acetophenone, etc. has been examined using various strains of Saccharomyces cerevisiae and Yarrowia lipolytica. Higher enantioselectivitiesare generally achieved together with lower yields compared with the results obtained at atmospheric pressure as in the case of treatment of cells with high pressure homogenation [791. Although the enantioselectivity obtained here is not as high as > 99% ee, this finding added pressure as an adjustable parameter to control the enantioselectivity of the bioreduction.

1010

I

75 Reduction Reactions

15.1.3

Improvement of Dehydrogenasesfor use in Reduction Reactions by Genetic Methods 15.1.3.1

Overexpression o f the Alcohol Dehydrogenase

Recent developments in molecular biology have contributed to the development of useful biocatalysts. Overexpression as well as rational and random mutations of many alcohol dehydrogenases have improved the function of enzymes so that they can be useful in organic synthesis[22,127-1341 . E xamples of overexpressed enzymes are introduced here, and Sect. 15.1.3.2-15.1.3.8 will describe the improvement of catalytic functions achieved by using genetic methods. Although the non-genetic chemical modifications of enzymes can also be important in order to improve a bio~atalyst['~~], they are not mentioned here. Example 1: The Thermoanaerobacter ethanolicus 39E adhB gene encoding the secondary alcohol dehydrogenase was overexpressed in Escherichia coli to form more than 10% to total protein[136].The recombinant enzyme was purified by heat treatment and precipitation with aqueous (NH4)2S04and isolated in 67 % yield. Enzymes with mutation@) around the active site residues were also created to examine the catalytically important zinc binding motif in the proteins. Example 2: The gene encoding a phenylacetaldehyde reductase with a unique and wide substrate range was cloned from the genomic DNA of the styrene-assimilating Corynebacteriurn strain ST10[137-139J . The enzyme was expressed in recombinant E. coli cells in sufficient quantity for practical use and purified to homogeneity by three column chromatography steps [140]. The amino acid residues assumed to be three catalytic and four structural zinc-binding ligands were characterized by site-directed mutagenesis of two zinc-binding centers within the enzyme

Besides these examples, many other important enzymes for biocatalytic reductions, ~~I, such as the NADPH-dependent carbonyl reductase from Candida r n a g n ~ l i a e [ ~the ketoreductase from Zygosaccharornyces rouxii and the aldehyde reductase from Sporobolomyces salmonicolor AKU4429 [1441, etc. have also been expressed in E. coli etc. and shown to be active. The availability of sufficient quantities of enzymes for crystallization studies has led to the crystal structures been obtained for several dehydrogenases. For example, two tetrameric NADP+-dependentbacterial secondary alcohol dehydrogenases from the mesophilic bacterium Clostridiurn belj'erinckii and the thermophilic bacterium Thermoanaerobium brockii have been crystallized in the apo- and the holo-enzyme forms, and their structures are available in the Protein Data Bank[145]. The crystal structure of the alcohol dehydrogenase from horse liver is also available [40-421.

75.1 Reduction of Ketones

15.1.3.2 Access to a Single Enzyme Within a Whole Cell: Use of Recombinant Cells

The advantages and disadvantages of using whole cell and isolated enzymes are described in Sect. 15.1.1.3. Here, genetic methods are used to build the systems with the advantages of both whole cells and isolated enzymes; the technology enables one to access essentially a single enzyme within a whole cell[127]. For example, to improve a low enantioselectivity due to the presence of plural enzymes in a cell with overlapping substrate specificities but different enantioselectivities, a recombinant cell with only the enzyme possessing the desired enantioselectivity was used (Fig. 15-21). Isolation of the enzyme, of course, improves the enantioselectivity.However, the requirement of a laborious enzyme isolation process and expensive cofactor with its associated regeneration enzyme (if necessary) have limited the practical utility of isolated enzyme processes. However, once the gene encoding the enzyme with high enantioselectivity has been overexpressed in E. coli, then the essentially single enzyme system can be accessed within the whole cell. Since it is a whole cell system, it can be cultivated to supply an appropriate amount without involving a laborious process for the isolation of an enzyme. The fact that there is no coenzyme requirement is also a merit for the system. Because it has only one enzyme which transforms the substrate, the problems of overmetabolism or low selectivity are also resolved. Using E. coli expressing Gcylp and E. coli expressing Gre3p, various 0-keto esters and a-alkyl-0-ketoesters were reduced with excellent enantio- (up to > 98% ee) and diastereo-selectivities (> 98% de) [*'*I.

I

Yeast catalyzed blotransfonnations ~

WPH

catalyst=

- ......

Isolated enzyme catalyzed biotransformation

.- - -

:S-enqmel - ----

r-------,

Isolation of an enzyme I

I

Sometimes result in low selectivity Enzymes with overlapping substrate specificities but ditferent enantioselectiiitiespresent

A I

i

Coenzyme will be necessary Limited supply due to the

Creation of engineered E. coli strains expressing an enzyme

I

Recombinant cell expressing the enzyme of interest from yeast catalyzed biotransformation

I

Single catalytic species (high enantio- and chemoselectivity, no overmetabolism) No coenzyme necessary No laborious process for the isolation of an enzyme Figure 15-21. Advantages and disadvantages of whole cell, isolated enzymes and recombinant cell as biocatalysts.

I

1012

I

I5 Reduction Reactions Figure 15-22. Use o f FAS deficient yeast t o improve the diastereoselectivity of a

cis (3R,4S) Commercial yeast

48

FAS deficient yeast

36

trans (3R,4R) :

38 3

15.1.3.3. Use o f a Cell Deficient in an Undesired Enzyme

This is a similar approach to that described above. Use of a yeast strain deficient in fatty acid synthase (FAS) suppressed formation of the undesired trans-diastereomer of a 0-lactam as shown in Fig. 15-22['*']. 15.1.3.4 Point Mutation for the Improvement of Enantioselectivity

Point mutation of enzymes has played an important role in determining those amino acid residues involved in catalytic activities. It has also been used to improve the enantioselectivity of dehydrogenases. For example, even a single point mutation of a secondary alcohol dehydrogenase from Themoanaerobacter ethanolicus can change substantially the enantioselectivity for the reduction of 2-butanone and 2-pentanone as shown in Table 15-6[451. 15.1.3.5 Broadening the Substrate Specificity of Dehydrogenase by Mutations

Developments in molecular biology enable us to change the substrate specificity of enzymes; the enzymes can be engineered to be more suitable for the requisite substrate. For example, variations have been made to the structure of the NAD' dependent L-lactate dehydrogenase from Bacillus stearothemophilus (LDH)[1301. Two regions of LDH that border the active site (but are not involved in the catalytic Table 15-6. Control o f enantioselectivity by a single mutation (serine-39 t o threonine) of the secondary alcohol dehydrogenase from Thermoanaerobacter e t h a n ~ l i c u s ~ ~ .

Parameter

Wild type

Mutant (S39T)

3.1 x i 0 5 1.1 x105 0 . 8 7 ~105 1.3 xi05

2.8 x10' 0.29~ lo5 3.5 x10' 2.1 X l O S

kat/K~ ( M ' s-') for oxidation at 55 "C of: (R)-2-butanol (S)-2-butanol (R)-2-pentanol (S)-2-pentanol

Ee of the reduction at 55 "C of: 2-butanone 2-pentanone

75.7 Reduction of Ketones

I

1013

Table 15-7.

Broadening the substrate specifity o f L-lactate dehydrogenase from Bacillus stearothermophilus by rational protein engineerit~g'~'.

Wild Type

CH3 CHiGH(CH3)z

250 0.33

0.06 6.7

4200000

102-'oSGlnLysPro MetValSer

CH3 CH2CH(CH3)2

66 0.67

0.16 1.9

410000 353

23GZ37AlaAla+ GlyGly

CH3 CH2CH(CH3)2

167 1.74

4 15.4

42 000

'02c'05GlnLysPro + MetValSer /236237AlaAla GlyGly

CH3 CH;?CH(CH3)2

32 18.5

4 14.3

8000 1300

-

+

50

110

reaction) were altered in order to accommodate substrates with hydrophobic side chains larger than that of the naturally preferred substrate, pyruvate. The muta--* MetValSer and [23G-2371AlaAla -+ GlyGly were made to tions [102-'051GlnLy~Pr~ increase to tolerance for large hydrophobic substrate side chains as shown in Table 15-7.The five changes together produced a broader substrate specificity LDH, with a 55 fold improved k,,, for a-keto isocaproate [R = CH2CH(CH+]. The substrate specificity of isocitrate dehydrogenase (IDH) has also been redesigned by genetic methods [13'1. Despite the structural similarities between isocitrate (ISO) and isopropylmalate (IPM),wild type isocitrate dehydrogenase (IDH) exhibits a strong preference for its natural substrate (ISO). The substrate specificity of IDH was changed to that of isopropylmalate dehydrogenase (IPMDH) using a combination of rational and random mutagenesis. Three amino acids of IDH (S113, N115, V l l b ) were changed and the chimeric enzyme ETV (S113E, N114T, V116V) showed Table 15-8.

0

Redesigning the substrate specificity of isocitrate dehydr~genase'~'. OH

~~z~Xr,.~ HOzc&co2H

OH

OH

isocitrate (ISO)

Enzyme

Wild Type IPMDH Wild Type ID H EVG

ENA ETV

isopropylmalate (IPM)

IDH position 113 115

E S E E E

L N V N T

kcat/b

IPM

(tW1S-')

(W' S-')

kcat/Khn

IS0

L V

1.4~10 1.7x10-" 1.1~10-~ 1 . 5 lo-' ~ 1.8~10-~

0 1.6~10 1.1~10-~ 5.9~ 3.9x10-'

-

Lt/KM

G

A V

IPM

IS0

116

kcat/KM

1.0~10-~ 1.0 2.5 4.6

1014

I

15 Reduction Reactions Table 15-9.

Elimination of the cofactor requirement by “blind” directed e v o l ~ t i o n ’ ~ ~ .

Bacillus stearothermophillus lactate dehydrogenase

Wild Wild Mutated (R118C, 203L, N307S) Mutated (R118C, 203L, N307S)

Cofactor (Fructose 1,dbisphosphate)

Kt.APYruvate

+

0.05

-

5 0.05 0.07

+ -

(mM )

a preferred substrate specificity for IPM over ISO; [kcat/K~IPM] / [kCat/K~ISO] of ETV was 4.6 while that of wild type IDH was 1.0 x 15.1.3.6 Production of an Activated Form of an Enzyme by Directed Evolution

One of the drawbacks of using alcohol dehydrogenases as catalysts for organic synthesis (comparing them with hydrolytic enzymes) is the cofactor requirement For example, Bacillus stearothermophillus lactate dehydrogenase is activated in the presence of fructose l,G-bi~phosphate[~~*]. The activator is expensive and representative of the sort of cofactor complications that are undesirable in industrial processes. Three rounds of random mutagenesis and screening produced a mutant which is almost fully activated in the absence of fructose 1,G-bisphosphate as shown in Table 15-9. 15.1.3.7 Change in the Coenzyme Specificity by Genetic Methods: NADP(H) Specific Formate Dehydrogenase

Formate/formate dehydrogenase is one of the most useful coenzyme regeneration systems as has been described in the Sect. 15.1.1.2. However, the known wild type formate dehydrogenases only accept NAD+; NADP’ is not the substrate. Multipoint site-directed mutagenesis was used to create a formate dehydrogenase which was able to accept NADP+.This mutant enzyme was then coupled to the reduction using the alcohol dehydrogenase from Lactobacillus sp as shown in Fig. 15-23t2*l. The activity of the NADP(H)-specific mutant (with NADP’ as substrate) is about GO% of the activity ofwild type formate dehydrogenase (with NAD’ as substrate). 15.1.3.8 Use of a Mutant Dehydrogenase for the Synthesis of 4-Amino-2-HydroxyAcids

The usefulness of a mutant dehydrogenase was demonstrated in a practical synthesis of 4-amino-2-hydroxy acids, which themselves are valuable as y-turn mimics for investigations into the secondary structure of peptides [1461. Chemoenzymatic synthesis of these compounds were achieved by lipase catalyzed hydrolysis of a a-keto esters to the corresponding a-keto acids followed by reduction employing a lactate dehydrogenase in one pot. Wild type lactate dehydrogenase from either Bacillus

15. I Reduction ofKetones 0

)cph

Lactobacillus sp. alcohol dehydrogenase ~

NADPH

NADP’

Figure 15-23. Recycling o f NADPH with protein engineered formate dehydrogenase[”].

protein engineered formate dehydrogenase from Pseudomonas sp.101

Table 15-10.

I

1015

- APh OH

The use o f a mutant dehydrogenase for the synthesis of 4-amino-2-hydroxyacids’46.

Dehydrogenase

R

Reaction Time

Wild type Staphylococcusegidemidis lactate dehydrogenase

a:CH3 b:CH(CH3)2 C: CH2CH(CH3)2 d: CHzPh

4 days no reaction no reaction no reaction

H205Q mutant of Lactobacillus delbrueckii bulgaricus D-hydroxyisocaproatedehydrogenase

a:CH3 b: CH(CH3)z c: CHzCH(CH3)z d: CHzPh

4h 5h 4h 5h

Yield (“A)

67

-

85 90 78 85

stearothermophilus (BS-LDH)or Staphylococcus epidermidis (SE-LDH)could be used specifically to reduce the ketone of the alanine derived a-keto acid, 2a, giving the (S)and (R)-2-hydroxyacids, respectively, in good yields. However, more bulky a-keto acids 2 b 2 d were not substrates for these enzymes. In contrast, the genetically engineered H205Q mutant of Lactobacillus delbrueckii bulgaricus D-hydroxyisocaproate dehydrogenase proved to be an ideal catalyst for the reduction of all the a-keto acids 2a-2d, giving excellent yields of the CBZ-protected (2R, 4S)-4-amino-2-hydroxyacid as a single diastereomer (Table 15-10).This genetically engineered oxidoreductase has great potential value in synthesis, not only due to its broad substrate specificity but also due to the high catalytic activity. For example, reduction of 1mmol of 2a took just 4 h with the H205Q mutant, whereas with SE-LDH the reaction required 4 days. 15.1.3.9

Catalytic Antibody

Nakayama and Schultz have developed antibodies to carry out the catalyhc enantioselective reduction of an a-keto amide using NaBH3CN as the r e d ~ c t a n t “ ~Mono~]. clonal antibodies raised to phosphonate 3 were prepared (Fig. 15-24), and one antibody showed activity for the enantioselective reduction of a chiral keto amide 4.

1016

I

I5 Reduction Reactions

hapten 3

‘“W;t 0

antibody NaBH3CN

~

0zNq;h+

CH3

4

Figure 15-24.

(2s)

0

EH3

Reduction of a ketone by a catalytic antibody”47!

Reduction with the antibody gave the 2s product with a diastereomeric excess greater than 99 % (oppositeto the stereoselectivityof the uncatalyzed reaction which afforded the 2 R product). 15.1.4 Reduction Systems with Wide Substrate Specificity 15.1.4.1 Bakers’ Yeast

Many methods for asymmetric reduction have been developed and some of these are used for the synthesis of optically active alcohols on a preparative scale. Bakers’ yeast is one of the most widely used microorganisms due to its commercial availability and its wide substrate specificity, which enables the non-expert in biochemistry to use the biocatalyst as a reagent for organic synthesis. Detailed reactions will not be described in this text since there are many reviews and original reports on this subject[’. 37-39, 148-1621 . H owever, one of the most important and useful reactions using yeast, the reduction of a hydroxymethyl ketone, is featured here due to the excellent enantioselectivity obtained even on a large scale (Fig. 15-25).1163-1661 . For example, 1-hydroxy-2-heptanone(50 g) was reduced to the corresponding (R)-diol in an Another example [Fig. 15-25(c)] optically pure form in 56% yield [Fig. 15-25 (b)][1641. is the reduction of a sulphenyl hydroxyketonewith yeast in the synthesis of a natural product [166]. Products isolated from the mandibular glands of the oriental hornet were synthesized using yeast reduction of an S-substituted hydroxyketone. 15.1.4.2 Rodococcus erythropolis

A carbonyl reductase isolated form Rhodococcus erythropolis accepts a broad range of substrates, including a variety of compounds useful for synthetic chemistry, as shown in Table 15-1112s1.Reduction of all the carbonyl compounds tested yielded

(S)-configuredhydroxyl compounds with high enantioselectivities.

15.1 Reduction of Ketones OAc

(a)

0

Yield 97% ee >95%

OH

yeast

Yield 56%(isolated)

large scale (509)

(c)

0

yeast

H O A S P h

A

OH

H O A S P h

- 0

-

&SPh

Yield 90% ee 78% Yield 63% ee 100% (after recrystallization)

Figure 15-25.

Reduction of hydroxyketones by bakers’ yeast[’63, 164, 1661.

Table 15-11.

Kinetic constants of the R. erythropolis carbonyl reductase2’.

Substrate

A L An

L

V, Pmg-1

KM (mM)

3.5

330

3.5

260

4.8

59

7.7

0.46

& M0/

1.4

2.6

0

3.8

0

C

5.5

0

0

10.4

A

0

u

0.59

V, (U mg-’1

Substrate

I

&04

A

KM (mM)

18 7.3 16 3.1

-

0

O

9.9 -

4.2

10.3

0.42

10.8

0.34

10.6

11.1

0.54

1.7

7.6

8.3 0.039 3.8

15.1.4.3 Pseudomonas sp. Strain PED and Lactobacillus kefir

The substrate specificities of the alcohol dehydrogenases from Pseudomonas sp. strain PED and Lactobacillus ke$r have been investigated. It was reported that they reduce wide varieties of ketones [6, ’1. Both reactions use 2-propanolfor the regeneration of coenzyme and produce (R)-alcoholsas depicted in Table 15-12. However, they require different coenzymes. The alcohol dehydrogenase from the Pseudomonas sp. uses NADH and transfers to pro-R hydride of NADH to the si-face of carbonyl compounds as shown in Sect. 15.1.1.1. The mechanism is ordered bi-bi with the coenzyme binding first and released last. On the other hand, the enzyme from

I

1017

1018

I

I5 Reduction Reactions Table 15-12. Enantioselectivities of the alcohol dehydrogenases from Pseudornonas sp. strain PED and Lactobacillus kefiP, '.

ee (%) Pseudomonas Lactobacillus sp. strain PED kefr

Product

OH

0

PhACF,

92

> 99

OH

94

-

ee

Product

\o&cl

OH

(%%I

Pseudomonas Lactobacillus sp. strain PED kefr

98

-

97

> 99

93

> 97

Phi

OH

P h / K

OH

OH

86

OH

0

Ph+',

CI4

45 98

0

27

Lactobacillus kejr uses NADPH and transfers the pro-R hydride from the cofactor to the si-face of carbonyl compounds. 15.1.4.4

Thermoanaerobium brockii

The alcohol dehydrogenase from Themoanaerobium brockii is very suitable for the reduction of aliphatic 191. Even very simple aliphatic ketones can be reduced enantioselectively.An interesting substrate size-induced reversal of enantioselectivity was observed. The smaller substrates (methyl ethyl, methyl isopropyl or methyl cyclopropyl ketones) were reduced to the (R)-alcohols, whereas higher ketones produced the (S)-enantiomers. This example and the next one (Sect. 15.1.4.5) using G. candidurn show that the biocatalytic reduction system is very beneficial for the reduction of aliphatic ketones over a non-enzymatic system where no report on highly enantioselective (> 99% ee) reduction of unfunctionalized dialkyl ketones can be found, to the best of our knowledge.

75.7 Reduction of Ketones

-

Table 15-13. Asymmetric reduction o f aliphatic ketones with the alcohol dehydrogenase from Therrnoanaerobiurn brockii". Product

Relative rate ee VO)Config.

12.0

48

R

3.0

8G

R

OH

Product OH

0.8

44

oH

R

/+/.v-/

OH

3.3

79

s

1.0

96

S

OH

0.3

95

s

OH

0.1

81

2S,3R

OH L

0.9

97

0.9

99

s

0.2

95

s

0.G

97

s

0.3

99

s

0.3

98

S

0.1

99

s

1.5

98

S

OH

w

XLl

Relative rate ee (%) Config.

C

l

s

15.1.4.5 Ceotrichum candidum

Reductions using an acetone powder of G. candidum (APG4), NAD' and 2-propanol exhibit one of the widest substrate specificities together with very high enantioselectivities (Table 15-14) 21].Various ketones such as acetophenone derivatives can be reduced with APG4 with excellent enantioselectivities(> 99% ee). The nature and electronegativityof substituents on the phenyl ring did not affect the enantioselectivity although the yield was slightly lower for para derivatives than for the corresponding ortho and meta derivatives. Reduction by APG4 of several aromatic ketones having different length alkyl chains demonstrated the scope and limitations of the substrate specificity. The phenyl moiety of acetophenone can be replaced by a benzyl or even by a 2-phenylethyl group with slightly better results in terms of chemical yield without any decrease in enantioselectivity.However, when the methyl moiety of actophenone was replaced by an ethyl, isopropyl or methoxymethyl group, the yield decreased dramatically, although the enantioselectivity remained high (z 99% ee). When the alkyl chain was elongated to a propyl or enlarged to a t-butyl group, the reaction was observed scarcely to proceed. The versatility of the APG4 reduction system is further exemplified by the use of pketo esters as substrates. 3-Oxobutyrates involving methyl, ethyl, t-butyl, or neopentyl esters are reduced to the (S)-hydroxyesterswith > 99% ee and in quantitative yield. Moreover, simple aliphatic ketones from 2-octanoneto 2-undecanone, as well

1020

I

75 Reduction Reactions Table 15-14. Reduction ofvarious ketones by the acetone powder of C. candidum, 2-propanoIz0.21.

Product

Yield ("A)

X=H

ee ("A)

89

O-F > 99 OH

m-F

95 74 0-CI >99 m-CI 95 p-C1 62 o-Br 97 m-Br 92 p-Br 95 o-Me 96 m-Me 86 p-Me 78 o-Me0 84 m-Me0 90 p-Me0 29 o-CF~ 6 m-CF3 96 p-CF, 73 1',2',3',4',5'-Fs 62

pF

X

>99(S) > 99 (S) > 9 9 (S) >99(S) >99 (S) 99 (S) > 9 9 (S) >99 (S) >99(S) > 9 9 (S) > 99 (S) >99(S) >99(S) >99(S) > 99 (S) > 9 9 (S) 97 (S) >99(S) >99 (S) > 99 (S)

Yield ("A)

Product

R = Et Pr i-Pr t-Bu CHzOMe CHzCl

OH

RAPh

R=Me Et t-Bu neo-Pentyl

ee ("A)

41 > 9 9 (S) 0 12 99(S) 1 8 >99(R) 80 98(R) >99 >99 >99 > 99

>99(S) >99 (S) > 99 (S) > 99 (S)

72

>99 (S)

87 87 85 60

>99 >99 >99 >99

OH 0

*o.., OH W

R

R=me Et Pr Bu

(S) (S) (S) (S)

90

99 (S)

92

99 (S)

OH

OH

&Ph

NAD' and

96

> 99 (S)

93

>99 ( S )

OH

*Ph

as 6-rnethyl-S-heptene-2-one and 5-chloro-2-pentanoneare also reduced by the APG4 system to the corresponding (S)-2 alkanols giving high yields with 99% ee. In summary, a detailed investigation of substrate specificity for the acetone powder of a G. candidum system reveals that as long as there is a methyl group at the aposition of the carbonyl group, high yield and enantioselectivity can be obtained regardless of the substituent on the other side of the ketone moiety. Apart from acetone-dried G. candidum I F 0 4597, intact whole cells of various strains of G. candidum have been found to be useful for asymmetric reductions[75-78, 101. 126, 1G7-1711 . For example, methyl 2-acetylbenzoatewas reduced by G.

candidum ATCC 34614, I F 0 5767 or I F 0 4597 as well as by other microorganisms such as Mucor javanicus, Mucor heimalis, Endomyces magnusii, Endomyces reessii and bakers' yeast to afford phthalide derivatives (Fig. 15-26)which have various pharmacological profiles such as relaxant, antiproliferative or antiplatelet effects, e t ~ . [ ' ~ ~ ] .

75.7 Reduction ofKetones

I

lo2'

Figure 15-26. Asymmetric reduction by G. candidurn ATCC 34614 for the synthesis of a bioactive phthalide d e r i ~ a t i v e " ~ ' ] .

15.1.5

Reduction of Various Ketones 15.1 S.1

Reduction of Fluoroketones

The biocatalytic reduction of fluoroketones is useful in order to gain an insight into the enzyme recognition of fluorinated groups, and is also very important due to the high synthetic values of the products, optically active fluorinated alcohols [lc.O, 172-1851. Sometimes the monofluorinated substrate can be a straightforward mimic of the unsubstituted counterpart, but with difluorinated and trifluorinated substrates, different recognition patterns compared with unfluorinated or monofluorinated substrates and with each other are often observed. For example, the enantioselectivity of yeast reduction is definitely affected by the fluorination pattern on the One of the most prominent effects of the fluorination of a substrate is seen in the reduction of acetophenone derivatives by the acetone powder of Geotrichum candidurn (APG4) as shown in Fig. 15-27[173,1741. Reduction of methyl ketones afforded (S)-alcohols in excellent ee, whereas the reduction of trifluoromethyl ketones gave the corresponding alcohols of the opposite configuration, also in excellent ee. Monofluoroacetophenone and difluoroacetophenone were also reduced under the same conditions. The reduction proceeded quantitatively for both substrates. As expected, the stereoselectivity shifted from the acetophenone type to the trifluoroacetophenone type according to the number of fluorine substituents at the a-position as shown in Fig. 15-28. The replacement of the methyl moiety with a trifluoromethyl group alters the bulkiness and electronic properties: the effect on the enantioselectivity has been examined. No inversion in stereochemistry was observed for the reduction of hindered ketones such as isopropyl ketone, while the stereoselectivity was inverted for the reduction of ketones with electron-withdrawingatoms such as chlorine. The mechanism for the inversion in stereochemistry was investigated in further studies. Several enzymes with different enantioselectivities were isolated; one of them OH H3CAph (s)

Yield 90% ee >99%

-

0

-

X3CKPh acetone powder G. candidurn NADP+ Cyclopentanol X=HorF

OH

(9 Yield ,99% ee 98%

Figure 15-27. Reduction of acetophenone and trifluoroacetophenone by an acetone powder of Geotrichum candidurn, NADP' and cyclopentano11' 73. 1741.

1022

I

15 Reduction Reactions

Configuration =

OH

100

X3CAPh

50

ee of product (%)

0

-50

OH

Configuration =

-1 00

X3CA Ph

I

I

JPh

FJPh

b

I

F$Ph F

L$Ph F

Substrates Figure 15-28. Effect o f introducing a fluorine atom or atoms at the a-position of acetophenone on the stereoselectivity in the reduction by C. candidurn acetone

&cx3

&

cx3

X=HorF

Figure 15-29. Substrates used for the examination o f t h e stereodirecting effects o f trifluoromethyl and methyl

catalyzed the reduction of methyl ketones, and another, with the opposite enantioselectivity, catalyzed the reduction of trifluoromethyl ketones. The differing abilities of trifluoromethyl and methyl groups to direct enantioselection in the reduction of carbonyl substrates has also been analyzed using various other microorganisms including different strains of G. candidum, Hansenula anomala, Saccharomyces cervisiae, Streptomyces, e t ~ . [ ’ ~The ~ ] . reduction of the cyclic ketone and enones shown in Fig. 15-29was investigated. The differences in the electronic and steric properties of the trifluoromethyl and methyl residues resulted in different chemo- and enantioselectivities in the reduction of the phenylbutenones, while the cyclohexanones showed similar enantioselectivities. Many synthetically valuable reactions involving reductions of fluoroketones have Various monofluoroketones are been reported as shown in Fig. 15-30[17G-1781. reduced with yeast; some of them proceeded with high diastereoselectivity. Chiral trifluoromethyl benzyl alcohols are useful synthons for ferroelectric liquid crystals. Therefore, Fujisawa et al. investigated the asymmetric reduction of the corresponding ketones using bakers’ yeast [179. l8’]. The enantioselectivity of the bakers’ yeast reduction of trifluoroacetylbenzene derivatives was improved by the introduction of some functional groups at the para-position to give the corresponding (R)-trifluoromethyl substituted benzylic alcohols in high chemical and optical yields as shown in Fig. 15-31.The “enantio-controlling”functional group at the paraposition was then used in further transformations. Yeast and G. candidurn acetone powder (APG4) are complementary to each other in the reduction of various trifluoromethyl biphenyl ketones. Yeast reduction affords the (R)-alcohol, whereas G. candidurn reduction affords the (S)-alcohol (Fig. 1532) [181].

15.1 Reduction of Ketones

OH yeast

Diastereomeric ratio up to 72/28 ee up to 86%

RA.y Ri...y F F R = Me, Et, Pr, Bu

R, = Me, Et, Pr, Bu R, = Me, Et

yeast

LR -

F3C R = Ph, Pr, Bu

F3C

Figure 15-30.

Reduction of fluorinated ketones by y e a ~ t [ ’ ~ ~ - ’ ~ ~ ~ .

F

h

-

OH

yeast

3

C

R

F

3

C

m

upto 92% ee

E -

ferroelectric liquid crystals

R

R = C02H, CO Me, NH, NHBz. NHTs, N H A ~OH, . o d e , OAC, O T ~OBZ ,

Figure 15-31. Asymmetric reduction of trifluoroacetylbenzene derivatives by bakers’ yeast[’79’ 180]. OH yeast

*-3“‘

up to 96% ee OH

’ R

G. candidurn R = H. Br. OMe, OH,

C02H, C0,Me.

NH2

acetone powder F3c%

up to 99% ee ’ R

Figure 15-32. Reduction of trifluoromethyl biphenyl ketones: bakers’ yeast vs C. candidurn acetone

Moreover, various optically pure fluorinated alcohols are produced by employing G. candidurn reductions as shown in Table 15-15(174].Monofluoroacetophenoneand difluoroacetophenoneare reduced to (R)-alcoholsby the acetone powder, NAD’ and

I

1023

1024

I

15 Reduction Reactions Table 15-15. Synthesis o f chiral fluorinated alcohols by the reduction with acetone powder and isolated enzymes of Ceotrichurn candidurn I FO 4597’74. Product

X=H

x=c1

F3C$

X=Br

Yield (“A)

ee (“A)

Product

84 81 80

98 (S) >99(s) >99(S)

‘ d p h

X

Yield (“A)

OH

F x p h

F)“

a The isolated enzyme was used

ee (“A)

93

> 99 (R)

91

> 9 9 (S)”

for the reduction.

2-propanol, and to (S)-alcohols by a constituent enzyme previously separated by anion-exchange chromatography and using glucose-6-phosphate/glucose-6-phosphate dehydrogenase as the cofactor recycling system. Both enantiomers of monofluorophenylethanol can be obtained with excellent ee using only one microorganism. 15.1 S . 2

Reduction of Fluoroketones Containing Sulfur Functionalities

As the demand for optically active fluorinated compounds increases, the importance of the development of asymmetric synthetic methods for fluorinated building blocks grows. On the other hand, sulfur functionalities such as phenylthio and dithianyl groups have been used as useful reactive units for a variety of chemical transformations. Therefore, various trifluoromethyl ketones containing a sulfur functionality have been reduced with various microorganisms [182-1851. For example, several microorganisms have been employed for the reduction of a,a,a,-trifluoromethyl a’-sulphenyl ketones (Fig. 15-33). Some of them produce the corresponding alcohols in high diastereo- and enantioselectivities;the high converSPh PhScF3

Candida sake CBSl59

(2&

Phk.,\CF,

0

de 94%. ee 84%

Rq C F 3

0

-

_____z_________L Candida lypolytica CBS 2074

R

TCF3

OH (2R.3R) R = Ph de 92% ee >96% R = CH2CH2Ph de >96% ee >96% Figure 15-33.

tion[’821.

Reduction o f sulphenyl ketones followed by epoxide forrna-

75.7 Reduction $Ketones 1) KHMDS, BnBr n-Bu,NI, THF

G. candidurn acetone powder

* F3c%

2) EtOH Raney Ni (W-2;

0

G. candidurn acetone powder

OH

CHzCH2CH2SPh

78

F3CAR 3-Thienyl 1,J-Dithian-P-yl

F3c ee >99%

Yield 88% (4.16 g) ee >99% (Rj

42

(3

>99 (9

D99 >99

Figure 15-34. Asymmetric reduction o f trifluorornethyl ketones containing a sulfur functionality by the acetone powder o f C. ~ a n d i d u r n ~ ’ ~ ~ ] .

sion into a single enantiomer is secured by the racemization of starting ketones under the biotransformation conditions. Transformation of the resulting sulphenyl trifluoromethyl alcohols into trifluoromethyl epoxides was also The acetone powder of G. candidurn (APG4)has also been used for the reduction of sulfur containing trifluoromethyl ketones (Fig. 15-34) [lS3].This reaction can be scaled up easily without the loss of enantioselectivity. For example, the reduction of trifluoro(2-thieny1)ethanoneon the gram scale proceeded quantitatively and yielded the optically pure (R)-alcohol in 88% yield after purification (4.16g, ee > 99%). The thienyl alcohol can be further transformed into a fluorinated aliphatic alcohol without racemization. 15.1.5.3 Reduction o f Chloroketones

The reduction of chloroketones has been widely investigated since it can produce versatile chiral intermediates. For example, reduction of an a-chloroketone results in the formation of a chlorohydrin, which can easily be transformed into an epoxide on treatment with a base. On recently published example involves the reduction of 3,4-dichlorophenacylchlorideby Rhodotorula mucillaginosa CBS 2378 or Geotrichum candidurn CBS233.76to give the (R)- or (S)-chlorohydrinwith > 99% ee and > 98% ee, respectively, as shown in Fig. 15-35[18G1. The (S)-enantiomer was transformed into the corresponding epoxide and then into a dichlorophenylbutanolide, an intermediate in the synthesis of (+)-cis-lS,4S-sertraline,which is an antidepressant drug of the selective serotonin reuptake inhibitor (SSRI) type. There are also many other examples of the reduction of a-halomethyl ketones as shown in Table 15-16[187-1891 . Vanous . microorganisms are able to reduce fluoro-, chloro- and bromoketones [161, 19s192’. However, reduction of iodoacetophenone usually results in a poor yield, producing, mainly, acetophenone or phenylethanol. Another example of the reduction of a-chloroketone involves dynamic kinetic resolution. The reduction of an a-chloroketo ester by M. racernosus and R. glutinis resulted in optically active syn- and anti-chlorohydrin, respectively, as shown in

I

1025

1026

I

IS Reduction Reactions OH

Rhodotoru/a muci//aginosa CBS 2378

OH

Geotrichum

cI

CBS233.76

CI

*

XAD-1180

CI

Figure 15-35. Reduction of a chloroketone followed by epoxidation for the synthesis o f sertraIine”861. Table 15-16.

x+

Reduction o f a-halogenated acetophenones Biocatalyst

- xv OH

Catalyst

Cryptococcus macerans

X

Yield” (%)

ee (“A)

Reference

c1

80 95

100 93

187 187

67 37 9

97 90 97

188

55 6 (40) 0 (15)

35 68

189 189 189

65

75 87.4 94

Br

F c1 Br

188 188

Bakers’ yeast

F c1 Br

F C1

Geotrichum candidum sp. 38

86 15 (25)

Br

OH M racemosus

/

0

0

CI

-A-

189

RCONH

ph&C02H

R = fert-butoxy side chain of taxotere

K C O 2 E t Ph ~

Cl Figure 15-36.

189 189

OH R = Ph side chain of taxol or Diltiazein3

Ph%COpEt CI

-

~

p.\\C02Et 0 Ph

Enantio- and diastereo-selective reduction o f a c h l o r ~ k e t o n e [ ’lg4]. ~~~

Fig. 15-36[1931. The syn-isomer was transformed into the corresponding epoxide, followed by conversion into the side chain of taxol and taxotere[’”]. One of the most studies a-chloroketones is ethyl 4-chloro-3-oxobutanoate. ( R ) -and (S)-enantiomers of the corresponding alcohol were produced by various micro-

75.1 Reduction of Ketones

I

1027

Table 15-17. Comparison of various microorganisms for the reduction of ethyl 4-chloro-3-oxobutanoate.

0 C I A C O 2 E t

-

Microorganism

OH Cl&C02Et (s)

Microorganism

Yield (“A)

ee (“A)

Reference

Geotrichum candidum

98 100

96 90 55 100 100

170 90 61 195 142

ee (%)

Reference

Bakers’ Yeast Bakers’ Yeast

Lactobacillus kefr Candida magnoliae

100 88

(recombinant and overexpressed in Escherichia coli)

0 CI A C O 2 E t

Microorganism

OH Cl*CO,Et (R)

Yield (“A)

Microorganism

Dancus carota Sporobolomycessalmonicolor Lactobacillusfermentum Saccharomyces cerevisiae

52 86 98 16

42 95 70 55

196 197 195 63

(FAS (P-keto reductase) negative)

organisms as shown in Table 15-17. The (R)-enantiomer is a promising chiral building block for the synthesis of L-carnitine,an essential factor for the P-oxidation of fatty acids in mitochondria. As shown in Fig. 15-37, a chiral intermediate for a human immunodeficiency vims protease inhibitor (HIVPI) was also synthesized by the reduction of an achloroketone with a Streptomyces strain [1981. Another example of the reduction of chloroketone is the reduction of 5-chloro2-pentanone by TBADH as shown in Fig. 15-38[19].Using this biotransformation in the synthetic pathway, a naturally occurring heterocycle isolated from the glandular secretion of the civet cat (Viverru civettu), was prepared.

COEN H

cI

StreptomycesnodosusSC 13149 *

O

H

OH

0

EMS-186318(Antiviralagent)

Figure 15-37. Synthe. sis of a chiral intermediate for an HIVpi 11981.

1028

I

75 Reduction Reactions 0

TBADH

A

OH

C

I

-a

m

Figure 15-38. Reduction o f 5-chloro-2-pentanone by TBADH for natural product

Yield 65% de 88%

sex attractant of the pine saw-fly

ee>96% (2R3.5~

Figure 15-39. Reduction o f ketones containing sulfur or nitrogen f u n ~ t i o n a t i t y ~ *191. '~~. 15.1.5.4

Reduction of Ketones Containing Nitrogen, Oxygen, Phosphorus and Sulfur Functionalities

Ketones with useful heteroatomic functional groups containing n i t r ~ g e n [ ' ~ ~ - ~ ' ~ ] , 213-2171 phosphoms 12181 and sU]fUr1154s 184, 219-2271 h ave been reduced by biocatalysts. For example, an intermediate in the synthesis of P-lactam antibiotics was obtained by microbial reduction of a P-keto ester as shown in Fig. 5-39(a)['"], while yeast reduction of a 0-keto dithioester afforded an easily separable mixture of P-hydroxy-dithioesters, the major component of which was converted enantioselectively into a sex attractant of the pine saw-fly as shown in Fig. 15-39(b)r2191. oxygen[lG3,

15.1.5.5

Reduction of Diketones

Regio- and enantioselective reduction of diketones can be achieved readily by using a b i o ~ a t a l y s t [. ~A ~ - ~ ~ ~ optically ~ s a~ result, active hydroxyketones and diols have been synthesized successfully. For the reduction of a-diketones, the selectivity between the reduction to diol and to hydroxyketone can be controlled using a diacetyl reductase from Bacillus stearothemophilus (Fig. 15-40)[233]. When a one-enzyme system was used for the coen( S ) , both carbonyl groups zyme recycling using endo-bicyclo[3.2.0]hept-2-en-G-ol were reduced selectively to produce a diol. On the other hand, a-hydroxyketones were obtained using a two-enzyme system glucose 6-phosphate/glucose 6-phosphate dehydrogenase for coenzyme recycling. The synthetic potential of both systems has been illustrated by the synthesis of the male sex pheromone of the grape borer Xylotrechus pyrrhoderus, identified as a two-component mixture of the reduction products, G and 7.

15. I Reduction ofKetones

I One-enzvme svsteml Bacillus sfearafherrnophilus diacetyl reductase

oH

"%"' 0

NADt

I

1029

[Two-enzymesystem]

Bacillus sfearofherrnophilus diacetyl reductase

glucose 6-phosphate/ glucose 6-phosphate dehydrogenase

OH

0

NADH

Bacillus stearofherrnophilus

I

Product

OH

Yield (%) R=Me Pr Ph Pentyl(6)

eR 6~ OH

40

92 80

82

80

ee (%)

1

>98(S,S) >98(S,S) >98(S,S) >98(S,S)

95(S,S)

OH

Figure 15-40.

Reduction o f a-diketones by diacetyl reductase from Bacillus s t e a r ~ t h e r m o p h i l u s ~ ~ ~ ~ ~ .

Regio- and enantioselective reduction of P-diketones may be carried out using biocatalysts. For example, a diketo ester 8 was reduced by the alcohol dehydrogenase from Lactobacillus brevis, to provide the corresponding hydroxyketo ester with 99.4 % ee in 78% yield; this was used as an intermediate for the synthesis of dimeric metabolite vioxanthin of Penicillium citreo-viride in order to develop an assay system to monitor phenol oxidative coupling in lignan formation [Fig. 15-41(a)][228]. Yeast reduction also proceeds regio- and enantioselectively with aliphatic diketones producing hydroxyketones with perfect selectivities as shown in Fig. 15-41(b)[2321. The yeast reduction also proceeds satisfactorily with 2,2-disubstitutedcycloalkanediones, producing hydroxyketones with excellent enantio- and diastereoselectivities as shown in Fig. 1 5 - 4 1 ( ~ ) [ ~ ~ ~ ! 15.1S . 6 Reduction o f Diary1 Ketones

Bulky ketones such as diaryl ketones can be also reduced by biocatalysts. For example, a rice plant growth regulator, (S)-N-isonicotinoyl-2-amino-5-chlorobenzhydrol, was prepared by microbial reduction of 2-amino-5-chlorobenzophenone with Rhodosporidium toruloides followed by isonicotinoylation as shown in Fig. 1542(a)[2431. A phosphodiesterase 4 inhibitor was also prepared by microbial reduction of a diaryl ketone 9 with Rhodotorula pilimanae, which was found by the screening of 310 microbial strains [Fig. 15-42(b)][244].

1030

I

75 Reduction Reactions alcohol dehydrogenase from Lactobacillus brevis

OH

8

Dimeric metabolite vioxanthin of PeniciNium citreo-viride

(b)

,y ,. , ) ,0

-

0

yeast

OH o

Yield 42% ee>99%

( 2 s 3s)

Figure 15-41.

Regio- and enantioselective reduction of

diketones[228.

231. 2321

0

(a)

NH2

Rhodosporidium toruloides

,

(J"f$ -

OH

QH NH2

/

CI Yield 60% ee 99%(s)

CI

N H C O ~ N

/

CI Rice plant growth regulator

- a a

)fo&

(b)

Rhodorola pilimanae 9

/

/ OMe Yield 10% ee96%(S)

OMe P

N

Figure 15-42. Reduction of diary1 ketones for the synthesis of bioactive compounds[243,2441.

15.1.5.7

Diastereoslective Reductions (Dynamic Resolution)

Enantio and diastereoselective reduction (dynamic resolution) of keto esters and ketones can be achieved using yeast and other microorganisms[55* 70r 74, 245-2531 . As shown in Fig. 15-43, when the racemization rate of the keto ester is faster than that for the yeast reduction, and the product hydroxyester is not racemized under the reaction conditions, then the yeast reduction may proceed enantioselectively and

15.1 Reduction ofKetones I1031

&C02R

2s

yeast

Rk

OH

Figure 15-43.

/YCOzR'

Diastereo-

c o p ~ ' selective reduction. +

R h

R h

2s, 3s

2S, 3R

2R,35

2R, 3R

faster racemization rate than reduction rate

2R

Rhizopus arrhizus

8

O

E

Kloekera rnagna or Cunninghamella t echinulata

- okoEt

gOEt (yo&(1 s, 25)

Mucor racernosus

Rhodotorula glufinis

(1 s, 25)

Figure 15-44.

QH 0

(1 s, 2R)

_OH 0 ....&oEt

0

(1s, 2R)

Diastereoselective reduction of cyclic keto esters[245].

diastereoselectively;thus only one stereoisomer out of the four possible ones can be obtained in one step. Actually, when bakers' yeast was used for the reduction of neopentyl2-methyl-3-oxobutanoate(R = Me, R = neopentyl), then the ratio of (2R, 3s) : (2S, 3R) : (2S, 3s) : (2R, 3R) products was found to be 96 : c 1 : 4 : c 1[2471. When an enzyme was isolated from the yeast, then the diastereoselectivitywas improved to > 99 : 1, and only a single isomer was obtained[248]. Another example is the large scale reduction of ethyl 2-methyl-3-oxobutanoate by Klebsiella pneurnoniae I F 0 3319I7']. On a 200 L scale, 2 Kg of the substrate were converted into the (2R, 3S)hydroxyester with 99 % de, > 99 % ee, and 99 % chemical yield as shown in Table 15-2.

Enantio- and diastereoselective reduction of cyclic keto esters are also achieved using various microorganisms (Fig. 15-44)[2451. By selecting a suitable organism, synand anti-hydroxyestersmay be synthesized enantio- and diastereoselectively. 15.1.5.8

Chemo-enzymaticSynthesis of Bioactive Compounds

Ketones with various functionalitis, containing F, C1, N, S , 0, etc., have been shown to be reduced by a biocatalyst, and by using the biocatalytic reduction as a key step, the chemoenzymatic synthesis of many bioactive compounds have been re-

75 Reduction Reactions

* Figure 15-45. Synthesis o f all four isomers o f t h e western corn rootworm sex pheromone [2341.

Figure 15-46.

Synthesis o f natural products from a key intermediate obtained by yeast reduction.

15.2 Reduction ofvarious Functionalities

I

ported['229 '2% '99. 228-230. 7-34! 235, 243, 254-2741 For example, 2,8-nonandione can be reduced enantioselectivelyby TBADH to furnish the corresponding diol, from which all four isomers of 8-methyldec-2-ylpropanoate, the western corn rootworm sex pheromone, were prepared (Fig. 15-45)[2341. One of the most versatile key intermediates discovered to date is the hydroxyketone 10 which is synthesized by the yeast reduction of the corresponding diketone [229, 2301. Starting with 10, many terpenes have been enantioselectively synthesized by Mori et al., as shown in Fig. 15-46.

15.2 Reduction of Various Functionalities

Kaoru Nakamura and Tomoko Matsuda 15.2.1

Reduction of Aldehydes

Many aldehyde reductases transform both aldehydes and ketones 275, 2761. For example, phenylacetaldehyde reductase from a styrene-assimilating Corynebacteriurn strain, ST-10, reduces aldehydes and ketones as shown in Table 15-18[138]. Other aldehyde reductases such as one from Sporobolornyces salrnonicolor also reduce aldehydes as well as ketones['&, 2751. Organometallicaldehydes can be reduced enantioselectivelywith dehydrogenases. For example, optically active organometallic compounds having planar chiralities were obtained by biocatalytic reduction of racemic aldehydes with yeast [277, 2781 or H L A D H [ 2 7 9 1 as shown in Fig. 15-47. The dynamic resolution of an aldehyde is also possible as shown in Fig. 15-48[280j. The racemization of the starting aldehyde and enantioselective reduction of a carbonyl group by bakers' yeast resulted in the formation of tertiary chiral carbon centers. The ee of the product was improved from 19% to 90 % by changing the ester moiety from the isopropyl group to the neopentyl group.

Examples of substrates of phenylacetaldehyde reductase from Corynebacteriurn strain, ~ ~ - 1 0 ' ~ ~ .

Table 15-18.

Substrate (mM) (aldehyde)

Relative activity

Substrate (mM)

Relative activity

(%I

(ketone)

(W

Acetaldehyde (3) n-Valeraldehyde (3) n-Hexyl aldehyde (3) Phenylacetaldehyde (3) 3-Phenylpropionaldehyde (1)

0 181 1220 100 364

Acetone (3) 2-Hexanone (3) 2-Heptanone (3) Acetophenone (3) 4-Phenyl-2-butanone (3)

0 207 760 35 29

1033

15.2 Reduction ofvarious Functionalities

I

ported['229 '2% '99. 228-230. 7-34! 235, 243, 254-2741 For example, 2,8-nonandione can be reduced enantioselectivelyby TBADH to furnish the corresponding diol, from which all four isomers of 8-methyldec-2-ylpropanoate, the western corn rootworm sex pheromone, were prepared (Fig. 15-45)[2341. One of the most versatile key intermediates discovered to date is the hydroxyketone 10 which is synthesized by the yeast reduction of the corresponding diketone [229, 2301. Starting with 10, many terpenes have been enantioselectively synthesized by Mori et al., as shown in Fig. 15-46.

15.2 Reduction of Various Functionalities

Kaoru Nakamura and Tomoko Matsuda 15.2.1

Reduction of Aldehydes

Many aldehyde reductases transform both aldehydes and ketones 275, 2761. For example, phenylacetaldehyde reductase from a styrene-assimilating Corynebacteriurn strain, ST-10, reduces aldehydes and ketones as shown in Table 15-18[138]. Other aldehyde reductases such as one from Sporobolornyces salrnonicolor also reduce aldehydes as well as ketones['&, 2751. Organometallicaldehydes can be reduced enantioselectivelywith dehydrogenases. For example, optically active organometallic compounds having planar chiralities were obtained by biocatalytic reduction of racemic aldehydes with yeast [277, 2781 or H L A D H [ 2 7 9 1 as shown in Fig. 15-47. The dynamic resolution of an aldehyde is also possible as shown in Fig. 15-48[280j. The racemization of the starting aldehyde and enantioselective reduction of a carbonyl group by bakers' yeast resulted in the formation of tertiary chiral carbon centers. The ee of the product was improved from 19% to 90 % by changing the ester moiety from the isopropyl group to the neopentyl group.

Examples of substrates of phenylacetaldehyde reductase from Corynebacteriurn strain, ~ ~ - 1 0 ' ~ ~ .

Table 15-18.

Substrate (mM) (aldehyde)

Relative activity

Substrate (mM)

Relative activity

(%I

(ketone)

(W

Acetaldehyde (3) n-Valeraldehyde (3) n-Hexyl aldehyde (3) Phenylacetaldehyde (3) 3-Phenylpropionaldehyde (1)

0 181 1220 100 364

Acetone (3) 2-Hexanone (3) 2-Heptanone (3) Acetophenone (3) 4-Phenyl-2-butanone (3)

0 207 760 35 29

1033

1034

I

75 Reduction Reactions

Yield 53% ee78% (s)

Yield 32% ee >99% (R) CHO I

Yield 33% ee 91%(s) Yield 51% ee 81%(R) Figure 15-47. Reduction of organometallic aldehydes to produce alcohols with planar chiralities [277-2791.

OHC-C0zR

yeast

ti =//

OHCYC02R

Figure 15-48.

c

HOH$2,COzR ee (%)

HOHzCyC02R

I

-CH2C(CH3)3

90

Reduction o f aldehyde with dynamic resolution1280].

15.2.2

Reduction of Peroxides to Alcohols

Horseradish peroxidase has been used for the reduction of peroxide to alcohol [2*1-*84l . The enzyme selectively recognizes sterically uncumbered (R)-alkyl aryl hydrogenperoxides,which allows kinetic resolution to provide (Rj-alcohol and (S)peroxide. However, poor enzyme recognition is observed with hydroperoxides possessing larger R2 groups such as a propyl or an isopropyl moiety as shown in Fig. 15-49. This reaction can be performed on a preparative scale conveniently to provide optically pure hydroperoxides. 15.2.3

Reduction of Sulfoxides to Sulfides

Asymmetric synthesis of sulfoxides can also be achieved by biocatalytic reduction. One example is the reduction of alkyl aryl sulfoxides by intact cells of Rhodobacter sphaeroides f sp. denitrijcan~[~~~I. In the reduction of methyl p-substituted phenyl sulfoxides, (S)-enantiomers were exclusively deoxygenated while enantiomerically pure (R)-isomers were recovered in good yield. For poor substrates such as ethyl phenyl sulfoxide, the repetition of the incubation after removing the toxic product was effective in enhancing the ee of recovered (R)-enantiomers to 100% as shown in Table 15-19.

15.2 Reduction of Various Fundonahies

I

1035

QOH Horseradish peroxidas: Guaiacol

R2%

-2"

+

\

Ri R1

ee (Oh) (-)-(S)-ROOH (+)-(Rj-ROH

'2

OH

%% \

R1

(9

R1

(Rj

5

-

OOH SiMezPh Horseradish peroxidase M if,-e2Ph+ Guaiacol

SiMe2Ph

E = 14.2

QOH

R2

OOH

+

+R1

OH

Horseradish peroxidase

*Rl

R2

Guaiacol

OH OOH

I Figure 15-49.

Table 15-19.

?

Me

Me

2

Reduction o f peroxides t o alcohols [281-2841.

Reduction o f sulfoxide to obtain optically pure ( R ) - s u l f o ~ i d e ~ ~ ~ .

4 9'. - R's\Ar

Rhodobactersphaeroidesfsp. denitrificans

R"\A~

R

Ar

Me Ph Me pMe-C6H4 Me pBr-C6H4 Me pMeO-C&, PhCH2 Me Ph Et Ph &Pr

Yield (%) ee (%) 46 40 43 47 41 41 54

+

*{

-<<-

R's'.Ar

(R)-sulfoxide

100 100 100

>99 90 100

21

15.2.4

Reduction ofAzide and Nitro Compounds to Amines

Bakers' yeast catalyzes the reduction of azides and nitro compounds to amiFor example, it catalyzes chemoselective reduction of azidoarenes to nes [2862911. arenamines as shown in Fig. 15-50[286*2871. Excellent yields are obtained for various aromatic compounds on reaction at room temperature. Aromatic nitro compounds

1036

I

75 Reduction Reactions Bakers' yeast R-N3

-

R-NHz

Figure 15-50.

x = H, MeCO, MeO, CI. Br, I, Me, NO,,

Reduction of azide to amine

by bakers!yeast[286.

Yield U P to 90%

2871

OH, etc

containing 0-,rn-, or p-electron withdrawing groups, such as carbonyl, halogen and nitro, were also selectively and rapidly reduced to their corresponding amino derivatives in good yields using bakers' yeast in basic solution[288]. N-oxides can also be reduced. For example, the microbial deoxygenation of a series of aromatic and heteroaromatic N-oxide compounds, including quinoline N-oxides, isoquinoline N-oxides, 2-aryl-2H-benzotriazole1-oxides, benzo[c]cinnoline N-oxide and azoxybenzene, has performed with bakers' yeast-NaOH to afford quinolines and pyridines [2911. 15.2.5

Reduction of Carbon-Carbon Double Bonds

The reduction of carbon-carbon double bonds to single bonds has been studied with various substrates ['l, 124, 292-3061 . For example, Ohta et al. demonstrated that the reduction of a number of 1-nitro-1-alkenesby fermenting bakers' yeast was enantioselective, resulting in the formation of optically active 1-nitroalkanes as shown in 2951 . 0n the other hand, Fuganti et al. reduced a,P-unsaturated-& Fig. 15-51(a)[294s lactones to produce enantiomerically pure (+)-(R)-goniothalamins [Fig. 15-51(b)], which show CNS activity. They also performed the kinetic resolution of the corresponding embryotoxic epoxide with yeast [2961. One of the most studied enzymes for the reduction of carbon-carbon double R2

,

(a)

,JL

R1

NO2

Yeast or old yellow enzyme N R O ,2-? .l,

etc

ee 99% (+)-(R)-goniothalamins Figure 15-51.

olefins

1294-2961

ee

H 1 Ph 1 Ph Me 98 pCI-Ph Me 89 pBr-Ph Me 94 Ph Et 97 Ph n-Pr 89 Hexyl no reaction Ph Hexyl Me (E) 83 Me Hexyl (Z) 66 2-Thienyl H

ee 77%

Examples of substrate specificity of yeast reduction of

6

15.2 Reduction of Various functionalities

I

1037

0, +,o

o;,+,oold yellow enzyme

old yellow enzyme

Figure 15-52. Mechanism of the reduction of nitro olefin by “old yellow enzyme” from y e a ~ t [ ~ ’ ~ - ~ ’ ~ ] .

&R

S

-

Nicotiana tabacum p90 reductase

Q

R

Nicotiana tabacum p44 reductase

11

R R

ee

n-Pr 95 Figure 15-53. Reduction o f carbon-carbon double bonds by reductases from plant cell

bonds is the “old yellow enzyme’’ from y e a ~ t [ ~ ’ ~ -which ~ ’ ~ ] has been shown efficiently to catalyze the NADPH-linked reduction of nitro olefins. The reduction of the nitro-olefin proceeds in a stepwise fashion (Fig. 15-52). The first step involves hydride transfer from the enzyme-reduced flavin to the P-carbon of the nitro-olefin which forms a nitronate intermediate that is freely dissociable from the enzyme. The second step, protonation of the nitronate at the a-carbon to form the final nitroalkane product, is also catalyzed by the enzyme. Photosyntheticmicroorganisms and plant cell cultures are very important sources of enzymes for the reduction of ole fin^['^, 2981. For example, Hirata et al. found that reduction of enone 11 with Nicotiana tabacum p90 reductase and Nicotiuna tabacum p44 reductase affords (S)- and (R)-alkylcyclohexanones,respectively, with excellent enantioselectivities as shown in Fig. 15-53. They also found two enone reductases from Astasia longa, a nonchlorophyllous cell line classified in Euglenales, and studied the mechanism. Both catalyzed enantiospecific trans-addition of hydrogen atoms to carvone from the si-face at the a-position and from the re-face at the P-position. 15.2.6 Transformation o f a-Keto Acid to Amine

A dehydrogenase can also be used for the transformation of an a-keto acid to an amine (Fig. 15-54). The chiral intermediate for an antihypertensive drug was prepared by reduction of an a-keto acid with glutamate dehydrogenase from beef liver. The cofactor NADH was regenerated using glucose dehydrogenase from Bacillus sp. [3071.

1038

I

15 Reduction Reactions Glutamate dehydrogenase from beef liver NADH NH3 Glucose I glucose dehydrogenase

HO dco2H

Figure 15-54.

-

NHp

HOC -OpH yield 92% ee>99%

Antihypertensive drug Reduction of a-keto acid to amine[307].

15.2.7 Reduction of Carbon Dioxide 15.2.7.1 Reduction of C 0 2 to Methanol

Syntheses using C 0 2 as a carbon source are attracting growing interest. The development of environmentally benign methods to utilize C02 is very important due to the abundance of COz. For this purpose, dehydrogenases have been successfully utilized. Formate, formaldehyde and alcohol dehydrogenases are used for the reduction of C 0 2 to methanol as shown in Fig. 15-55[333 308-3111. For the efficient production of methanol, electrochemical methods have been used (Fig. 15-55)[33,308, 310, 3111. Electrochemically, C02 was converted into formate by formate dehydrogenase with the aid of methyl viologen or pyrroloquinolinequinone as a mediator. Methanol dehydrogenase was used to reduce formate to formaldehyde and methanol with the same system[308,310. 3111. An approach for the conversion of COz into formic acid which combines a semiconductor photoelectrode with formate dehydrogenase is very interesting [331. Electrons in the semiconductor can be produced with light of wavelengths shorter than 900 nm. Then, the photogenerated electrons were transferred to C02 through methyl viologen to produce formic acid as shown in Fig. 15-5G[331. Another highly efficient process involves the immobilization of three enzymes in a silica ~ol-gel[~~’]. Since the process consists of a sequential reaction of in situ generated substrates with three different enzymes, the confinement of the system in a porous matrix resulted in an enhanced probability of the reactions as shown in Fig. 15-57 due to an overall increase in local concentration of reactants within the nanopores of the sol-gel processed glasses [3091.

Formate dehydrogenase

COZ

* HCOpH

Formaldehyde dehydrogenase

-

Alcohol dehydrogenase

HCHO

*

CH3OH

dehydrogenase dehydrogenase e- I electron mediator Figure 15-55.

Reduction of COZto methanol with dehydrogenases.

75.2 Reduction of Various Functionalhies

I

1039

MV2+'

a

'ght

M e - N w N - M e

Me-N-N-Me Photoelectrochemical pumping of enzymatic COz reduction[33].

Figure 15-56.

Sol-gel

Methanol (pmol)

Solution

10

0

'

0

50

100 150 200

NADH (pmol)

0

+

Malic enzyme

cop

* FCOzH NADP', NADP' reductase CO,H methyl viologen e-

AC02H

HOpCJ C O p H Figure 15-58.

Figure 15-57. Effect of the confinement of the three enzymes in a porous matrix on methanol productionISo9I.

+

OH lsocitrate dehydrogenase * HO,C+CO,H methyl viologen COpH e'

3131. Electrochemical reductive fixation o f COzPi2~

15.2.7.2

Reductive fixation of COZ

Reductive fixation is an another important process. Malic enzyme and isocitrate dehydrogenase catalyze both the reduction of the carbonyl group in an a-keto acid and fixation of C02 at the a-position with the aid of an electric power source and an electron mediator (Fig. 15-58)1312. 3131. Uniquely, the reaction using isocitrate dehydrogenase does not require the use of NADP+. When C02 is reductively fixed in an organic molecule, the enzyme is oxidized;the oxidized enzyme is ultimately reduced back to its original form by methyl viologen cation radicals[312!

1040

I

75 Reduction Reactions

References 1

2

3

4

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G. Fantin, M. Fogagnolo, A. Medici, P. Pedrini, S. Poli, F. Gardini, M. E. Guerzoni, Tetrehedron: Asymm.1991,2, 243-246. 227 R. Hayakawa, M. Shimizu, T. Fujisawa, Tetrahedron Lett. 1996, 37, 7533-7536. 228 D. Drochner, M. Miiller, Eur. ]. Org. Chem. 2001,211-215. 229 K. Mori, H. Mori, Org. Synth. 1993, Coll. VO~. 8, 312-315. 230 K. Mori, S. Takayama, S. Yoshimura, Liebigs Ann. Chem. 1993,91-95. 231 D. W. Brooks, H. Mazdiyasni, P.G. Grothaus,]. Org. Chem. 1987,52,3223-3232. 232 H. Ohta, K. Ozaki, G. Tuchihashi, Agric. Bid. Chem. 1986,50,2499-2502. 233 0. Bortolini, G. Fantin, M. Fogagnolo, P. P. Giovannini, A. Guerrini, A. Midici,J . Org. Chem. 1997,62,1854-1856. 234 E. Keinan, S. C. Sinha, A. Sinha-Bagchi, J. Org. Chem. 1992,57,3631-3636. 235 K. Fuhshuku, N. Funa, T. Akeboshi, H. Ohta, H. Hosomi, S. Ohba, T. Sugai, 1.Org. Chem. 2000, 65, 129-135. 236 R. Bel-Rhlid, A. Fauve, M. F. Renard, H. Veschambre, Biocatalysis 1992, G, 319-337. 237 H. Ikeda, E. Sato, T. Sugai. H. Ohta, Tetrahedron 1996, 52,8113-8122. 238 Y.-Y. Zhu, D. J.Burnell, Tetrahedron: Asymm. 1996,7,3295-3304. 239 S . 4 . Lee, J.-L. Yan, K. C. Wang, Tetrahedron: Asymm. 1997,8,3051-3058. 240 P. Besse, J.Bolte, A. Fauve, H. Veschambre, Bioorg. Chem. 1993, 21, 342-353. 241 K. Nakamura, S. Kondo, Y. Kawai, K. Hida, K. Kitano, A. Ohno, Tetrahedron: Asymm. 1996,7,409-412. 242 M. Sakakibara, A. Ogawa-Uchida, Biosci. Biotech. Biochem.1995,59, 1300-1303. 243 M. Kato, K. Sasahara, K. Ochi, H. Akita, T. Oishi, Chem. Pharm. Bull. 1991, 39, 2498-2501. 244 M. Chartrain, J.Lynch, W.-B. Choi, H. Churchill, S. Patel, S. Yamazaki, R. Volante, R. Greasham, J . Mol. Catal, B: Enzymatic 2000,8,285-288. 245 S. Danchet, C. Bigot, D. Buisson, R. Azerad, Tetrahedron: Asymm. 1997,8,1735-1739. 246 K. Nakamura, T. Miyai, K. Nozaki, K. Ushio, S. Oka, A. Ohno, Tetrahedron Lett. 1986, 27, 3155-3156. 247 K. Nakamura, T. Miyai, A. Nagar, S. Oka, A. Ohno, Bull. Chem. SOC.Jpn. 1989,62, 1179-1187. 248 K. Nakamura, Y. Kawai, N. Nakajima, 226

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I

15 Reduction Reactions

T. Miyai, S. Honda, A. Ohno, Bull. Chem. SOC.Jpn. 1991,64,1467-1470. 249 C. Abalain, D. Buisson, R. Azerad, Tetrahedron: Asymm. 1996,7,2983-2996. 250 G. Fantin, M. Fogagnolo, P. Giovannini, A. Medici, E. Pagnotta, P. Pedrini, A. Trincone, Tetrahedron: Asymm.1994, 5,1631-1634. 251 T. Kuramoto, K. Iwamoto, M. Izumi, M. Kirihata, F. Yoshizako, Biosci. Biotech. Biochem. 1999, 63, 598-601. 252 K. Nakamura, T. Miyai, Y. Kawai, N. Nakajima, A. Ohno, Tetrahedron Lett. 1990, 31, 1159-1 160. 253 K. Nakamura, Y. Kawai, T. Miyai, A. Ohno, Tetrahedron Lett. 1990, 31, 3631-3632. 254 G. Gibbs, M. J. Hateley, L. McLaren, M. Welham, C. L. Willis, Tetrahedron Lett. 1999,40, 1069-1072. 255 B. Das, P. Madhusudhan, A. Kashinatham, Bioorg. Med. Chem. Lett. 1998, 8, 14031406. 256 C. Gonzales-Bello,M. K. Manthey, J.H. Harris, A. R. Hawkins, J. R. Coggins, C. Abell,J. Org. Chem. 1998,63,1591-1597. 257 R. N. Patel, R. L. Hanson, A. Banerjee, L. J. Szarka,J. Am. Oil Chem. SOC.1997,74, 1345-1360. 258 N. W. Fadnavis, S. K. Vadivel, M. Sharfuddin, U. T. Bhalerao, Tetrahedron: Asymm. 1997,8,4003-4006. 259 M. Amat, M.-D. Coll, J. Bosch, E. Espinosa, E. Molins, Tetrahedron: Asymm.1997,8, 935-948. 260 A. Sutherland, C. L. Willis, Tetrahedron Lett. 1997, 38,1837-1840. 261 C. Fuganti, P. Grasselli, M. Mendozza, S. Servi, G. Zucchi, Tetrahedron 1997,53, 2617-2624. 262 N. M. Kelly, R. G. Reid, C. L. Willis, P. L. Winton, Tetrahedron Lett. 1996, 37, 1517-1520. 263 H. Watanabe, T. Watanabe, K. Mori, Tetrahedron 1996,52, 13939-13950. 264 M. Miyazawa, K. Tsuruno, H. Kameoka, Tetrahedron: Asymm. 1995,6,2121-2122. 265 T. Sugai, 0. Katoh, H. Ohta, Tetrahedron 1995,51,11987-11998. 266 M. Zarehcka, M. Rejzek, 2. Wimmer, D. Saman, L. Strcinz, Tetrahedron 1993,49, 5305-5314. 267 S. Robin, F. Huet, A. Fauve, H. Veschambre, Tetrahedron: Asymm.1993, 4, 239-246. 268 Y.J. Surh, S. S. Lee, Biochem. Int. 1992, 27, 179- 187.

269

J.-X. Gu, 2.-Y. Li, G.-Q. Lin, Tetrahedron: Asymm. 1992,3,1523-1524.

G. Fronza, C. Fuganti, P. Grasselli, G. Pedrocchi-Fantoni, S. Servi, Tetrahedron Lett. 1992,33,5625-5628. 271 E. Santaniello, P. Ferraboschi, P. Gnsenti, F. Aragozzini, E. Maconi,J. Chem. Soc., Perkin Trans I , 1991,601-605. 272 E. Keinan, S. C. Sinha, A. Sinha-Bagchi, J. Chem. SOC., Perkin Trans. 1,1991, 3333-3339. 273 K. Pabsch, M. Petersen, N. N. Rao, A. W. Alfermann, C. Wandrey, Red. Trav. Chim. Pays-Bas, 1991, 110, 199-205. 274 A. Kumar, D. H. Ner, S. Y. Dike, Tetrahedron Lett. 1991, 32, 1901-1904. 275 S. Shimizu, M. Kataoka in: Biotransformations (Ed.: K. Faber), Springer-Verlag, Berlin, 2000, p. 109-123. 276 A. Muheim, R. Waldner, D. Sanglard, J. Reiser, H. E. Schoemaker, M.S.A. Leisola, Eur. /. Biochem. 1991, 195,369-375. 277 J.A.S. Howell, M. G. Palin, G. Jaouen, S. Top, H. E. Hafa, J. M. Cense, Tetrahedron: Asymm. 1993,4,1241-1252. 278 J.A.S. Howell, M. G. Palin, H. E. Hafa, S. Top, G. Jaouen, Tetrahedron: Asymm. 1992,3,1355-1356. 279 C. Baldoli, P. D. Buttero, S. Maiorana, G. Ottolina, S. Riva, Tetrahedron: Asymm. 1998,9,1497-1504. 280 K. Nakamura, T. Miyai, K. Ushio, S. Oka, A. Ohno, Bull. Chem. Soc./pn. 1988, 61, 2089-2093. 281 W. Adam, U. Hoch, M. Lazarus, C. R. SahaMoller, P. Schreier,J. Am. Chem. SOC.1995, 117, 11898-11901. 282 W. Adam, C. Mock-Knoblauch,C. R. SahaMoller, Tetrahedron: Asymm. 1997, 8, 1947-1950. 283 W. Adam, M. Lazarus, U. Hoch, M.N. Korb, C. R. Saha-Moller , P. Schreier,/. Org. Chem. 1998,63,6123-6127. 284 W. Adam, B. Boss, D. Harmsen, 2. Lukacs, C. R. Saha-Moller,P. Schreier,J. Org. Chem. 1998,63,7598-7599. 285 M. Abo, A. Okubo, S. Yamazaki, Tetrahedron: Asymm. 1997,8, 345-348. 286 M. Baruah, A. Boruah, D. Prajapati, J. S. Sandhu, Synlett. 1996, 1193-1194. 287 A. Kamal, B. Laxminarayana, N. L. Gayatri, Tetrahedron Lett. 1997, 38, 6871-6874. 288 W. Baik, J. L. Han, K. C. Lee, B. H. Kim, J.-T. 270

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyriqht 0Wiley-VCH Verlaq GmbH, Weinheim 2002

75.3 Reduction ofC=N bonds

Hahn, Tetrahedron Lett. 1994, 35, 3965-3966. 289 W. Baik, D. 1. Kim, H. J . Lee, W.-J. Chung, B. H. Kim, S.W. Lee, Tetrahedron Lett. 1997, 38,4579-4580. 290 J . A . Bladcie, N. J. Turner, A. S. Wells, Tetrahedron Lett. 1997,38,3043-3046. 291 W. Baik, D.I. Kim, S. Koo, J. U.Rhee, S. H. Shin, B. H. Kim, Tetrahedron Lett. 1997, 38, 845-848. 292 K. Stott, K. Saito, D. J. Thiele, V. Massey, J. Bid. Chem. 1993,268,6097-6106. 293 A.D.N. Vaz, S. Chakraborty, V. Massey, Biochemistry 1995, 34,4246-4256. 294 Y. Meah, V. Massey, Proc. Natl. Acad. Sci. USA 2000,97,10733-10738. 295 H. Ohta, N. Kobayashi, K. Ozaki, J . Org. Chem. 1989,54,1802-1804. 296 C. Fuganti, G. Pedrocchi-Fantoni, A. Sarra, S. Servi, Tetrahedron: Asymm. 1994,5, 1135-1 138. 297 M. S. v. Dyk, E. v. Rendsburg, I.P.B. Rensburg, N. Moleleki,]. Mol. Catal, B: Enzymatic 1998, 5, 149-154. 298 T. Hirata, K. Shimoda, T. Gondai, Chem. Lett. 2000,850-851. 299 G. Fronza, C. Fuganti, M. Mendozza, R. S . Rallo, G. Ottolina, D. Joulain, Tetrahedron 1996,52,4041-4052. 300 K. Takabe, M. Tanaka, M. Sugimoto, T. Yamada, H. Yoda, Tetrahedron: Asymm. 1992,3,1385-1386.

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301

15.3 Reduction of C=N bonds

Andreas 5.Bommarius 15.3.1

Introduction

Enantiospecific reduction of C=N bonds is of interest for the synthesis of a-amino acids and derivatives such as amines. While nonenzymatic reductive amination has been known since 1927['],only recently have enzymatic procedures to L-amino acids became established. The reduction can be achieved by different enzymes following different mechanisms, e. g. by pyridoxalphosphate (PLP)-dependenttransaminases (E.C. 2.6.1, discussed in Chapter 12.7)or by amino acid dehydrogenases (E.C. 1.4.1) using NADH or NADPH as the cofactor. The synthetic usefulness of the transaminase reaction is diminished by the location of the equilibrium (&, often is close

I

1047

15 Reduction Reactions

to one), so that complex mixtures result, which are often laborious to separate (for solutions to this problem, see Chapter 12.7). For this reason, this chapter focuses on the reduction of C=N bonds by reductive amination with amino acid dehydrogenases, AADHs. Reductive amination of a-keto acids to a-amino acids is similar to the reduction of C = 0 bonds to the corresponding a-hydroxy acids. In an equilibrium reaction, aketo acids can be reductively aminated to a-amino acids or, vice versa, a-amino acids can be oxidatively deaminated:

RyCooH+ NAD+ + H20 NH2 A very promising process route is the reductive amination of prochiral a-keto acids to a-amino acids with AADHs and the cofactor NADH and its regeneration by cooxidation of formate to COz by formate dehydrogenase (Fig. 15.3-1). This asymmetric synthesis route possesses a number of advantages rendering it attractive in today's context of seeking environmentally benign processes:

compact synthesis of a-keto acid substrates, formation of harmless and easily separable COZ as the only co-product, extreme enantioselectivity of amino acid dehydrogenases, and yields of up to 100% with respect to a-keto acid, resulting in no undesirable enantiomers and other by-products.

Ry CooH-aGGiGq

RKCooH

+h

OH

[ L-dehydrogenase I

or

RyCooH NHZ

[NADH +

pziGcq

H@I FDH formate dehydrogenase

co2 Figure 15.3-1. regeneration.

HCOOH or HCOONH4 Schematic o f enzymatic reductive amination with cofactor

75.3 Reduction of C=N bonds

Table 15.3-1.

List of NAD(P)+-dependent amino acid dehydrogenase~(~1.

I

~~~~

E.C. Number Enzyme

Coenzyme

Sourcea

1.4.1.1 1.4.1.2 1.4.1.3 1.4.1.4 1.4.1.7 1.4.1.8 1.4.1.9 1.4.1.10 1.4.1.11 1.4.1.12 1.4.1.15 1.4.1.16 1.4.1.20 1.4.1.-

NAD+ NAD+ NAD(P)' NAD' NAD' NAD(P)+ NAD' NAD' NAD' NAD+ NAD' NADP' NAD' NAD(P)+

B (Bacillus, Streptomyces, Halobacterium) B, F, Y,P A, F, Etrahymena B, F, Y,Chlorella P B (Alcaligenes,Streptomyces),P B (Bacillus,Clostridium) B (Mycobacterium) B (Clostridium) B (Clostridium) Human, B (Agrobacterium) B (Bacillus, Colynebacteriurn) B (Brevibacterium,Bacillus, Rhodococcus) P

Alanine DH Glutamate DH Glutamate DH Glutamate DH Serine DH Valine DH ucine DH Glycine DH 3,5-Diaminohexanoate DH 2,4-Diaminopentanoate DH Lysine DH Diaminopimelate DH Phenylalanine D H Tryptophan DH

a Abbreviations: B: bacterium; F: fungi; Y: yeast; A: animal; P: plant; DH: dehydrogenase

With three exceptions (AlaDH from Phormidium lapideum, L-lysine-&-dehydrogenase and meso-a,&-diaminopimelateDH) all of the AADHs (Table 15.3-1) catalyze reduction of prochiral keto acids to the L-amino acids [(S)-configuration].The natural function of L-AADHsis not known. The D-AADHsthat have been found appear to be iron-sulfur membrane-associated flavoenzymes which seem to catalyze the oxidative reaction from keto acids to amino acids only; artificial dyes and the coenzyme Q analog serve as electron acceptors but not AADHs have been screened from a variety of organisms (Table 15.3-l),the most important enzymes for synthesis are alanine dehydrogenase (AlaDH, E. C. 1.4.1.1), phenylalanine dehydrogenase (PheDH, E. C. 1.4.1.20), and particularly leucine dehydrogenase (LeuDH, E. C. 1.4.1.9). The ubiquitous glutamate dehydrogenase (GluDH, E. C. 1.4.1.2.-4), however, is still the most studied member of the group. Reviews on AADHs: Apart from early review articles on individual amino acid dehydrogenases by Schiitte et al. (1985; LeuDH from B. cereus)I6],Ohshima et al. (1985a; LeuDH from B. species)17]and Hummel et al. (1987; PheDH from Rh. rhodocrous) [8],comprehensive reviews have been published by Hummel and Kula (1989)['], Ohshima and Soda (1989 and 1990)[5~'o~1'] and by Brunhuber and Blanchard (1994)['21. 15.3.2 Structural Features of Amino Acid Dehydrogenases (AADHs)

Most of the AADHs possess hexameric structure, although octamers, tetramers, dimers and even monomers have been found. The subunits are usually of similar size: for instance, most bacterial AADHs are hexamers with a molecular weight of around 49 000 per subunit.

1049

1050

I

75 Reduction Reactions Identities o f protein sequences of different amino acid dehydrogenases (in per cent) [221. The data were calculated via BLAST search i n the database 'Swissprot'[231.

Table 15.3-2.

Protein

LeuDH, B. stearothemophilus LeuDH, B. cereus LeuDH, B. sphaericus PheDH, Rh. rhodocrous PheDH, 'I: intemedius

LeuDH, LeuDH, B. B. cereus sphaericus

PheDH, Rh. rhodocrous

PheDH, Th. intermedius

CIuDH, C. symbiosum

82.5

32.0 31.5 31.7

45.6 44.5 41.8 26.4

12.6 13.4 14.0 12.4 14.2

-

79.9 76.9

-

-

-

15.3.2.1 Sequences and Structures

Several amino acid dehydrogenases have been screened from a variety of microorganisms, the preparatively most important are phenylalanine dehydrogenase (PheDH, from Rhodococcus sp. M4) and leucine dehydrogenase (LeuDH, from Bacillus stearothemophilus and Bacillus cereus). As of the end of February 2001, more than 20 gene and protein sequences for AADHs except GluDH (which more than triples the number) and 3D crystal structures from five different AADHs have been ~], from B. sphaeri~us['~I, deposited (GluDH from Clostridium s y m b i ~ s u r n [ ~LeuDH AlaDH from Phomidium lapideum[l5I, PheDH from Nocardia sp 239L''l and PheDH from Rhodococcus sp. M4[178'81). Sequence homologies and similarities of 3D structures of the members of several organisms are so high that amino acid dehydrogenases can be termed a single superfamily, generated through divergent evolution [19-211 (Table 15.3-2). Remarkable, on one hand, is the high degree of identity of the three leucine dehydrogenases, and on the other hand the sequence of glutamate dehydrogenase, which bears no homology to the other dehydrogenases. Although overall sequence homology varies from around 20 % up to 80 %, the residues essential for the threedimensional structure of a subunit, for nicotinamide cofactor binding, and for catalysis have been conserved[*']. While a complex between NAD' and GluDH from Clostridium symbiosum left the overall conformation unaltered [241, a drastic conformational change (hinge movement) was observed on binding of the gl~tamateI'~1. 15.3.3 Thermodynamics and Mechanism of Enzymatic Reductive Amination 15.3.3.1 Thermodynamics

For reductive amination, basically no thermodynamic limitation exists: for the equals 9x1012[251, for phenylalanine/ leucine/ketoleucine reaction at pH 11.0, ~ been thus, the maximum phenylpyruvate at pH 7.95 a k& of 2 . 5 ~ 1 0has degree of conversion is very close to 100%. Coupling of the reductive amination reaction with cofactor regeneration via the FDH/formate reaction, which is irreversible, further helps to pull the equilibrium towards the amino acid product.

75.3 Reduction ofC=N bonds

I

1051

15.3.3.2 Mechanism, Kinetics

As will be elucidated below, the mechanism of reductive amination and the geometry of the active center[13,18, *'* 271 cause the (S)-configuredamino acid products of the reaction to be completely enantiomerically pure, an important criterion for a large-scaleapplication. The catalytic mechanism of AADHs has been studied most thoroughly with GluDH from C. s y r n b i o ~ u r n [ ~and ~ , ~with ~ I PheDH from Rhodococcus M4["1. The mechanism was found to be remarkably similar in both cases so that the prediction by Stillman et al.[131seems to have been borne out. In Fig. 15.3-2, the study on PheDH is illustrated[18]: Following the scheme in Fig. 15.3-2, which depicts oxidative deamination, in a clockwise fashion starting from the top left, the a-N-protonated L-Phe molecule is stabilized by the &-groupof LysGG at the carbonyl group as well as by the &-groupof Lys78 via a water molecule, the carbonyl group of Pro117 and the P-carboxyl group of Asp118 at the a-amino group. The first intermediate is the protonated imine after steps (2) and (3)in which Lys78 picks up the proton from the a-N-group of L-Phe and delivers a hydrogen to the Si face of the cofactor NAD' with deprotonation of Lys78. Accompanied by another Lys78 protonation, the water molecule adds to the imine 3''

-K78

H'

Figure 15.3-2.

Proposed mechanism for amino acid dehydrogenases (with PheDH as an example) [la].

H '

0

1052

I

75 Reduction Reactions

carbon to form the carbinolamine, the second intermediate [step (4)].The Lys78 proton is picked up by Asp118 [step (S)] and in turn by the amino group [step (G)] of the substrate to liberate NH3 and with the formation of phenylpyruvate. The keto group is stabilized by the protonated c-sidechain of Lys78 as well as a by a proton from Gly40. The positioning of Lys78 and Gly40 also prevents the oxidation of phenyllactate, so that PheDH cannot act as a HicDH. A similar mechanism had already been proposed for GluDH from C. syrnbiothe only major difference seems to be the attribution of the initial deprotonation of the amino acid molecule to Asp165 (which corresponds to Asp117 on PheDH) instead of Lys125 (Lys78 in PheDH). The Lys125 in GluDH is known to have a low pK value[28],which causes this residue to act as a proton shuttle more easily. The optimum degree of protonation and catalybcally important amino acid residues can be determined from a log V,,-pH on the acidic and alkaline side of the optimum pH, log V,, decreases nearly linearly with pH, the two slopes intersect at the optimum degree of protonation, which is also the optimum point of activity. The experimentally observed optimum pH value of 9.2-9.3 for LeuDH l3O], corresponding to two pK values of around 8.7 and 10.0 for amino acid residues participating in the catalytic step, can be linked to lysine residues, corroborating the results of Rife and Cleland (1980)rZ6] and Sekimoto et al. (1993)["I for the case of GluDH. Brunhuber et al. in their study of PheDH assigned their pKas values of8.1 and 9.4 to Asp118 and Lys78, respectively["]. The influence ofpH on reductive aminations with AADHs can also be explained by the dissociation equilibrium of ammonia (pK, value 9.25). Only an uncharged ammonia molecule can be accepted by LeuDHIz6,301 so that a minimal pH of around 7.5 has to be kept throughout the reaction. 15.3.4 Individual Amino Acid Dehydrogenases 15.3.4.1

Leucine Dehydrogenase (LeuDH, E. C. 1.4.1.9)

Isolation and characterization of LeuDH has been pioneered by Hummel et al.131] (from B. sphaericus), Schiitte[6](from B. cereus), and by Ohshima and Soda (from mesophilic Bacillus sphaericus and from moderately thermophilic Bacillus stearothemophilus['O, 321). The biochemical data for the last two enzymes, however, do not differ much, as Table 15.3-3reveals. The LeuDH from B. stearothemophilus as compared with the B. sphaericus enzyme has an extended pH range of activity (5.5-10 vs. 6.5-8.5), a higher heat stability (70 vs. 50 "C after a heat treatment of 5 min), a longer half-life (several months vs. six days at pH 7.2 and G "C), and much greater stability against organic solvents and denaturants [I'. LeuDH from B. stearothemophilus had already been cloned and overexpressed['" 331 during early studies. Recently, the production of recombinant enzyme from B. cereus even on a large scale has been demonstrated[34.351. 8"

15.3 Reduction ofC=N bonds Table 15.3-3.

Properties of LeuDH from Bacillus sphaericus and Bacillus stearothermophilus['O1. ~

~~~

Source

B. sphaericus

B. stearothermophilus

M, (kDa)

245 000 41 000

300 000 49 000

hexamer

hexamer 11.0

Subunit (M,) Optimum pH: deamination amination Coenzyme Substrate specificity (in % of L-leucine) Deamination: L-leucine L-valine L-isoleucine L-norvaline L-a-aminobutyrate L-norvaline D-leucine Amination: a-ketoisocaproate a-ketoisovalerate a-ketovalerate a-ketobutyrate a-ketocaproate

10.7 9.0-9.5

I

9.0-9.5

NAD (KM 0.39 mM) NAD (KM 0.49 mM) 100 (KM 1.0 mM) 74 (1.7) 58 (1.8) 41 (3.5) 14 (10) 10 (6.3) 0 100 (0.31) 126 (1.4) 76 (1.7) 57 (1.7) 46 (7.0)

100 (KM4.4 mM) 98 (3.9) 73 (1.4)

-

0 100 167 86 45

The substrate specificity of LeuDHs, catalyzing mainly branched-chain a-keto acids to the a-amino acids, has been investigated by Zink and Sanwal(1962)[361 and subsequently by Schiitte et al. (1985: B. cereus)161,Ohshima and Soda (1989; Bacillus stearothemophilus and Bacillus sphaericus) f51, Nagata et al. (1990; Bacillus DSM 7330)[371, Misono et al. (1990; Corynebacterium pseudodiphtheriticum) 13*1 and by Bommarius et al. (1994; Bacillus stearothemophilus) i3')1. In addition to the proteinogenic amino acids valine, leucine, and isoleucine, unnatural amino acids such as tertle~cine[~'] or L- P-hydroxy-valinek4l1 can be synthesized. The kinetic parameters of several leucine dehydrogenases show a similar pHprofile. The opposite tendency of V, and KM for all substrates is remarkable: the dimethyl-substituted substrates show values above 10 mM (Vm, values are between 0.2 and 30% of the reactivity of 2-0x0-4-methyl-pentanoicacid, the base case), whereas KM values below 1 mM are typical for good substrates (V,, = 100% compared with the base case, 2-0x0-4-methyl-pentanoic acid). 15.3.4.2

Alanine Dehydrogenase (AlaDH, E.C. 1.4.1.1)

AlaDH has been isolated and characterized from both mesophilic (B. subtilis and B. sphericus) [421 and thermophilic (B. stearothemophilus) [431 organisms. For cloning and purification of AlaDH, see ref.[*]. The narrow substrate specificity of AlaDH[421 renders the enzyme useful for synthesis of L-alanine and analogs only, such as [15N]L-alanine[451, 3-fluoro-L-alanine1461, and 3-chloro-~-alanine [471.

1053

1054

I

15 Reduction Reactions

NADH

Figure 15.3-3.

NAD’

Synthesis of 6-hydroxy-~-norleucine with CluDH/glucose DH(’’1.

15.3.4.3 Clutamate Dehydrogenase (CIuDH, E. C. 1.4.1.2-4)

GluDH has been investigated by the groups of Engel and Rice since the 1980s so that more is known about GluDH, especially from C. syrnbiosurn, than about any other AADH. Although there is no sequence identity to other AADHs beyond random similarity (Table 15.3-2),site-directed mutagenesis of two amino acids residues, K89L and S380V, led to similar activity levels towards glutamate, norleucine and methionine and demonstrated the importance especially of the K89L mutation[48, 491. Stud’ ies on GluDH from the same source define the knowledge base regarding conformational change of the enzyme upon binding of the substrate but not upon the preceding binding of the cofactor. These conformational changes also seem to be responsible in part for substrate spe~ificity[’~1. Just as with other AADHs, GluDH has potential as a catalyst in synthesis: beef liver GluDH was the best catalyst for the reductive amination of 2-keto-6-hydroxyhexanoic acid Na salt to 6-hydroxy-~-norleucine,a potentially important building The reaction of 95 mM block for the vasopeptidase Vanlev (BMS) (Fig. 15.3-3)[511. substrate (2: 1 mixture of 2-keto-6-hydroxy-hexanoicacid Na salt in equilibrium with 2-hydroxy-tetrahydropyran-2-carboxylic acid) was complete in 3 h, resulting in an amino acid product of 89-92 % chemical yield and >99% optical purity. As the keto acid substrate is very cumbersome to synthesize, an alternative way of providing the keto acid substrate was the separation of D,L-6-hydroxynorleucine, which can be prepared easily from 4-hydroxybutylhydantoir1,by D-amino acid oxidase to L-amino acid and keto acid where the latter in turn was reduced by G~uDH/NADH[~*]. Both FDH/formate and glucose DH/glucose were employed for cofactor regeneration. 15.3.4.4 Phenylalanine Dehydrogenase (PheDH, E. C. 1.4.1.20)

An enzyme catalyzing the reductive amination of phenylpyruvate to the desired LPhenylalanine was first found by Hummel et al.[52]in a strain of Brevibacteriurn and later in Rhodococcus sp. ,’[ 531. Table 15.3-4 summarizes the microbiological and kinetic data [’)I.

15.3 Reduction of C=N bonds Table 15.3-4.

Comparison of PheDH from Brevibacteriurn and Rhodococcusspecies[9].

Parameter

Microbiological data: enzyme yield (U L-I) after addition of 1% of L-phenylalanine L-histidine L-phenylalaninamide L-isoleucine D-phenylalanine DL-phenylalanine Enzymological data: pH optimum reductive amination oxidative deamination

KM (mM) phenylpyruvate phydroxypyruvate indolepyruvate 2-0x0-4-methylmercaptobutyrate V,

(relative to phenylpyruvate) phenylpyruvate p-hydroxypyruvate indolepyruvate 2-0x0-4-methylmercaptobutyrate

KM (pM) NADH KM (mM) N H 2 Stability: stored at 4 "C (tip) deactivation ("3 d-I) under operation Reference

Table 15.3-5.

Brevibacteriurn

Rhodococcus

210 120 0 204 214 9.0

15200 1800 3500 0 0 0 9.25

10 0.11

10 0.16

0.24 8.0 3.0 100

2.4 7.7 2.1 100

96 24 59 47

5 3 33 130

431 4-8h

387 10 d

26 8

5 53

Substrate specifity of different P ~ ~ D H S [ ~ ' ] .

Substrate'

Ketoisocaproate Keto-methionineb Phenylpyruvate p-OH-phenylpyruvateb Indolepyruvateb Keto-4-phenylbutyrate Keto-5-phenylvalerate

Rhodococcus rhodocrous VmaX(U m L-') KM (mM) 50 150 7.5 4.5 96 46

2.1 0.16 2.4 7.7 0.01 0.65

Rel. activity (%)

4.2 33 = 100 5 3 64 30

B. sphaericus Rel. activity (%)

6.0

= 100

138

n. d.' 1.9 1.5

a Conditions: pH 8.0, T = 25 "C, 1.31= 0.1 M; comparison: LeuDH from 5.cereus: 2-0x0-4-methyl-pentanoic

acid = loo%, 2-0x0-4-phenylbutyrate= 0.2% B. sphaericus data fromis5] b As in a except for a pH of 8.5''' c Not determined

I

1055

1056

I

15 Reduction Reactions

w PheDH

NADH

cop Figure 15.3-4.

NAD+

[:mCo NH,

L-allysine ethylene

acetal

NH,HCOO

Synthesis o f allysine ethylene acetal with PheDH/FDH1581.

Apart from L-phenylalanine, the homolog L-homophenylalanine (L-Hph), important as a component in ACE inhibitors, can be obtained from 2-keto-4-phenylbutyrate with PheDH [541. The substrate specificity of PheDH from Bacillus sphaericus has been investigated by Asano et al.rS51.Table 15.3-5compares the activities of two PheDH from Rhodococcus rhodocrous['I and Bacillus s p h ~ e r i c u sfor ~ ~ the ~ ] transformation of aromatic and aliphatic keto acids. Sequencing, cloning, and heterologous expression of PheDH from Rhodococcus was first described by Brunhuber et al.[sGl.A double mutation G124A/L307V was created by site-directed mutagenesis of PheDH from Bacillus sphaericus to change the substrate specificityfrom a PheDH closer to a LeuDH. This led to a mutant with decreased activity towards L-phenylalanineand enhanced activity towards almost all aliphatic amino acid substrates, thus confirming the predictions made from molecular modeling [57J. PheDH from Trtermoactinornyces intemedius ATCC 33 205 was utilized recently to synthesize allysine ethylene acetal [( S)-2-amino-5-(1,3-dioxolan-2-yl)-pentanoic acid (2)] from the corresponding keto acid with regeneration of NAD' cofactor by FDH/ f ~ r m a t e [ ~(Fig. * ] 15.3-4);the specific activity towards the keto acid was 16% compared to the standard substrate phenylpyruvate. The system was used in three different configurations: (i) the system with heatdried cells from Trt. intermedius (PheDH) and C. boidinii (FDH) yielded on average only 84 M% and could not be scaled up owing to lysis of the 7%. intemedius cells; (ii) a similar system with recombinant PheDH from E. coli improved the yield to 91 M%; (iii) heat-dried Pichia pastoris containing endogeneous FDH and expressing recombinant PheDH from Th. intemedius yielded 98 M% with an optical purity of >98%. Altogether, more than 200 kg of allysine ethylene acetal have been produced. 15.3.5 Summary of Substrate Specificities

The most comprehensive investigation of substrate specificity of LeuDH and PheDH has been conducted by Krix et al. (1997)[30!Table 15.34 lists the relative rates of various substrates.

15.3 Reduction of C=N bonds I1057 Table 15.3-6.

Relative V,,, values of keto acid substrates ofvarious LeuDHs and PheDHP’].

Keto acid

Specific activity (u mg-‘ of protein) 2-Oxobutyricacid 2-0x0-3-methylbutyricacid 2-0~0-3,3-dimethylbutyric acid 2-Oxopentanoic acid 2-0x0-3-methylpentanoicacid 2-0x0-4methylpentanoicacid” 2-0~0-3,3-dimethylpentanoic acid 2-0~0-4,4-dimethylpentanoic acid 2-Oxohexanoicacid 2-0x0-4-methylhexaoicacid 2-0x0-4-ethylhexanoicacid 2-0xo-4,4-dimethylhexanoic acid 2-0xo-5,5-dimethylhexanoic acid 2-0x0-3-cyclohexylpropanoic acid 2-Oxooctanoicacid 2-Oxo-3-(l-adamantyl)propanoic acid

B. stearo-

B. cereus

thermophilus LeuDH

LeuDH

B. sphaericus Rhodococcus LeuDH Rhodocrous PheDH

120 48 113 31 63 110 = 100 2 7 15 22 1 0.5 0.8 0.8 0.2 0

15.9 74 152 74 81 114 = 100 11 14 63 19 11 1.2 0.3 0.1 n. d.b n. d?

3.3 6G 205 51 102 88 = 100 5 11 75 n. d? n. d? 0.2 n. d? 0.3 n. d.b n. d?

54.8 72 96 8 157 193 = 100 4 54 250 296 79 146 257 140 n. d.b 16

V, values refer to 2-0x0-4-methylpentanoic acid (= loo%),pH 8.5, T = 30 “C. Absolute activity of LeuDHs with 2-0x0-4-methyl-pentanoic acid (ketoisocaproic acid) were 120 U mg-’ (B. stearothemophilus), 15.9 U mg-’ (B. cereus) and 3.3 U mg-’ (B. sphaericus) as well as 54.8 U mg” with PheDH (Rh.rhodocrous). b Not determined

a All

LeuDHs from B. cereus, B. sphaericus and B. stearothermophilus display a remarkably similar substrate spectrum: LeuDHs accept 2-oxoacids with hydrophobic, aliphatic, branched and unbranched carbon side chains of up to six C atoms as well as some alicyclic keto acids as substrates, however, not the adamantyl group, where the geometric limit seems to be reached. 2-0x0-3-methylpentanoicacid is the preferred substrate, the preferred chain length is C5. The keto acid substrate should have at least four C atoms; pyruvate is only converted at less than 3 % of standard. Short-chainketo acids with branching at the C3 position are only preferred by the enzyme from B. sphaericus. The different amino acid dehydrogenases differentiate substrate side chains mainly based on steric parameters in the C3 and C4 position of branched ketoacids. Functionalized keto acids such as ketoglutarate are not accepted (activity < O . l % of the base case). Phenylpyruvate as a model compound of an aromatic substrate was inert[’’]. A correlation of LeuDH activity with van-der-Waals volumes [601 or hydrophobicities[“] for different C atom configuration of side chains only yielded a moderate

correlation [39, ‘*I. PheDH differs markedly from all LeuDHs, as it can convert not only aromatic substrates but also the aliphatic substrates typical for LeuDHs. Owing to the high

1058

I

15 Reduction Reactions

intrinsic specific activity of PheDH from Rhodococcus, in many cases the enzyme actually registers higher specific activity with many sterically demanding a-keto acid substrates than LeuDH. The substrate specifity of PheDH from Rhodococcus rhodocrous and Bacillus sphaericus seems to vary more between the two PheDHs than the specificity between the different LeuDH species. PheDH from B. sphaericus mainly converts (substituted) phenylpyruvates whereas the enzyme from Rhodococcus sp. displays a fairly high degree of activity in the presence of a phenylalkyl group in the substrate. 15.3.6 Process Technology: Cofactor Regeneration and Enzyme Membrane Reactor (EMR) 15.3.6.1 Regeneration of NAD(P)(H) Cofactors

Enzymatic reductive amination with NADH as the cofactor can only be operated on a large scale if the cofactor is regenerated. Wandrey and Kula have developed a regeneration scheme using formate as the reductant of NAD' generated upon reductive amination (Fig. 15.3-1). The formate is oxidized irreversibly to COz by formate dehydrogenase (FDH, E.C. 1.2.1.2) [62]. For soluble reactants and products, enzymes are preferentially immobilized in an enzyme-membrane reactor (EMR). To prevent the cofactor from penetrating through the membrane, it can be enlarged with polyethyleneglycol (PEG)[631. L-leucine was produced in an EMR with LeuDH from both B. ~phaericus[~'] and B. stearothermophil~s[~~1. LeuDH has also been employed successfully for the synthesis of L-tert-leucine in batch pr0cesses[~~1 and in its continuous version[40b* 651. L-tertleucine is an important building block for several novel pharma developments["* 671 as well as being on intermediate for templates for asymmetric L-Phe was produced in an EMR with PheDH starting from phenylpyruvate [Fig. 15.3-5, (i)]["]. Owing to the instability and high cost of this compound, two additional processes were devised generating phenylpyruvate in situ (Fig. 15.3-5): (ii) intermittent oxidation of DL-phenyllactate with D- and L-hydroxyisocaproate DH (HicDH)[691, or (iii) hydrolysis of acetamidocinnamic acid (ACA) with ACA acylase["]. For productivities of all processes, see Table 15.3-7. Another regeneration scheme for NADH from NAD' utilizes glucose which is oxidized to gluconic acid with the help of glucose dehydrogenase (see Fig. 15.3-3for an example)[51].Regeneration to NADPH from NADP' can be afforded by glucose721; the 6-phosphate dehydrogenase with glucose-6-phosphate as the system, however, has not found widespread use yet, probably owing to the higher price of NADP' vs. NAD' and the cost associated with the generation of glucose6-phosphate from glucose. With the advantage of the potentially quantitative use of a keto acid substrate and with suitable processes of cofactor regeneration, reductive amination of keto acids is an interesting route to a-amino acids worthy of consideration in comparison with more established routes.

15.3 Reduction of C=N bonds

a

mCo0" r \ W H f i " " "

PheDH

Phenylpyruvate

u NAD@

NADH+H@

co2

m H C o o H

HCOO@N@

FDH

L-HicDH D-HicDH+

D,L-Phenyllactate

COOH , NAD@

NADH+H@

t

I

0 Acetamido-cinnamate COOH

PheDH

I

f

Enzymatic routes to L-phenylalaninevia phenylpyruvate[''. (i) Reductive amination o f phenylpyruvate by PheDH with simultaneous NADH regeneration using FDH. (ii) Oxidation o f DL-phenyllactate with D- and L-2-hydroxy-4-methylpentanoate (HicDH) and simultaneous reductive amination of the phenylpyruvate formed in situ with PheDH. NADH is "substrate-coupled" regenerated from phenyllactate. (iii) In situ formation of phenylpyruvate by enzymatic deacetylation o f N-acetamidoocinnamic acid by the respective acylase followed by simultaneous reductive amination with PheDH. Figure 15.3-5.

I

1059

1060

I

75 Reduction Reactions

Continuous production of L-amino acids with the aid of dehydrogenases in an enzyme membrane reactor(g].

Table 15.3-7. AADH

Regeneration Precursor enzyme(s)

LeuDH

FDH

LeuDH

D-HmpDH L-HmpDH

LeuDH

D-HmpDH L-HmpDH

LeuDH

FDH

AlaDH

D-LDH L-LDH FDH

PheDH PheDH PheDH

+ ACA

D-HmpDH L-HmpDH FDH

Product Product Degree s.t.y. conc. ofcon- g/(Lxd) version

oxomethyl- L-leu pentanoate DL-OHL-leu methyl-pentanoate

80

80

250

70

70

72

D,LL-met OH-methionine trimethylL-tle pyruvate D,L-lactate L-ala

240

60

143

425

85

640

184

46

134

114

95

456

22

43

28

70

88

277

acylase

Ref.

(U kg-7

(mM)

phenyl-pyru- L-phe vate D,L-phenyl- L-phe lactate acetamid- L-phe ocinnamate

Enzyme consumptions

300/300 40c (LeuDH,FDH) 730/350/650 40a (LeuDH, D-HmpDH, L-HmpDH) 40c

1000/2000 40c (LeuDH, FDH) 4700/2600 40a (LeuDH, FDH) 1500/150 68 (PheDH, FDH) 69 1170/1770/ 70 400 (acylase, PheDH, FDH)

HmpDH: 2-hydroxy-4-methylpentanoate-DH;LDH: lactate dehydrogenase; L-tle: L-tert-leucine

15.3.6.2

Summary of Processing to Amino Acids

The production of L-tert leucine on a multi-100 kg scale and of L-neopentylglycine on a 30 kg scale with LeuDH from B. stearothemophilus demonstrates the suitability of enzymatic reductive amination on a large scale and even for slow substrates. The economics of the process is influenced decisively by the retention and regeneration of both production (AADH) and regeneration enzyme (FDH). If yields of less than 100% are acceptable enzyme consumption can be lowered by running the process in a continuous Owing to the broad substrate specificity of AADHs, reductive amination can be utilized especially for the synthesis of hydrophobic amino acids. LeuDH and PheDH feature complementary specificities for aliphatic and aromatic L-amino acids. Both enzymes are enantioselectiveto the highest degree and stable in a coupled process with FDH. As non-enzymatic processes of reductive amination often lead to low yields and enantioselectivities[73-761, enzymatic schemes are superior to chemical ones. Additionally, enzymatic reductive aminations are conducted solely in water so that organic solvents can be avoided.

References I1061

References 1

F. h o o p and H. Oesterlin, Hoppe Seylers Z.

Physiol. Chem. 1927, 170,186-211. 2 P. J.Olsiewski, G. J. Kaczorowski and C. Walsh, J. Bid. Chem. 1980,255,4487-94. 3 S. Nagata, N. Esaki, K. Tanizawa, H. Tanaka, K. Soda, Agnc. Bid. Chem. 1985,49, 1134-41. 4 M. Lobocka, J. Hennig, J. Wild, T. Klopotowski,]. Bacteriol. 1994,176,1500-10. 5 T. Ohshima, K. Soda, Znt. Znd. Biotech. 1989, 9, 5-11. 6 H. Schiitte, W. Hummel, H. Tsai, M.-R. Kula, Appl. Microbiol. Biotechnol. 1985.22, 306317. 7 T. Ohshima, S. Nagata, K. Soda, Arch. Microbiol. 1985, 141,407-411. 8 W. Hummel, H. Schiitte, E. Schmidt, C. Wandrey, M.-R. Kula, Appl. Microbiol. Biotechnol. 1987, 26,409-416. 9 W. Hummel, M.-R. Kula, Eur. /. Biochem. 1989, 184,l-13. 10 T. Ohshima, K. Soda, TIBTECH 1989,7, 210-214. 11 T. Ohshima, K. Soda, Adv. Biochem. Eng./ Biotech. 1990,42, 187-209. 12 N. M. W. Brunhuber, J. S. Blanchard, Crit. Rev. Biochem. Mol. Bid. 1994, 29(6), 415-467. 13 T. J. Stillman, P. J. Baker, K. L. Britton, D. W. Rice, 1.Mol. Bid. 1993, 234, 1131-1139. 14 P. J.Baker, A. P. Turnbull, S. E. Sedelnikova, T. J. Stillman, D. W. Rice, Structure 1995,3,693-705. 15 P. J.Baker, Y. Sawa, H. Shibata, S. E. Sedelnikova, D. w. Rice, Nature Struct. Bid. 1998,5,561-567. 16 A. Pasquo, K. L. Britton, P. J. Baker, G. Brearley, R. J. Hinton, A. J. G. Moir, T. J . Stillman, D. W. Rice, Acta Crystallop. 1998, 054,269-272. 17 J.L. Vanhooke, J. B. Thoden, N. M. W. Brunhuber, J. S. Blanchard, H. M. Holden, Biochemistry 1999,38,2326-2339. 18 N. M. W. Brunhuber, J. B. Thoden, J. S. Blanchard, J. L. Vanhooke, Biochemistry 2000,39(31), 9174-9187. 19 K. L. Britton, P. J. Baker, P. C. Engel, D. W. Rice, T. J. Stillman, J. Mol. Biol. 1993, 234, 938-45. 20 S. Nagata, K. Tanisawa, N. Esaki, Y. Saka-

moto, T. Ohshima, H. Tanaka, K. Soda, Biochemistry 1988,27,9056-62. 21 H. Takada, T. Yoshimura, T. Ohshima, N. Esaki, K. Soda,J. Biochem. 1991, 109, 371-6. 22 A. S. Bommarius, Habilitation thesis, RWTH Aachen, 2000. 23 a) M. C. Peitsch, Bio/Technology 1995, 13, 658-660; b) M. C. Peitsch, Biochem. Soc. Trans. 1996,24, 274-279. 24 P.J.Baker, K. L. Britton, P. C. Engel, G. W. Farrants, K. S. Lilley, D. W. Rice, T. J. Stillman, Proteins 1992, 12,75-86. 25 B. D. Sanwal, M. W. Zink, Arch. Biochem. Biophys. 1961, 94,430-435. 26 J. E. Rife, W. W. Cleland, Biochemistry 1980, 19,2328-33. 27 T. Sekimoto, T. Matsuyama, T. Fukui, K. Tanizawa,J. Bid. Chem. 1993,268, 27 039-27045. 28 D. Piszkiewicz, E. L. Smith, Biochemistry 1971,10,4538-44. 29 W. W. Cleland, Enzyme Kinetics as a Toolfor Determination of Enzyme Mechanisms. Investigation of Rates and Mechanisms of Reactions, 4th ed., John Wiley & Sons, New York, 1986, pp. 791-870. 30 G.Krix, A. S. Bommarius, K. Drauz, M. Kottenhahn, M. Schwarm, M.-R. Kula,]. Biotechnol. 1997, 53, 29-39. 31 W. Hummel, H. Schiitte, M.-R. Kula, Eur. J. Appl. Micobiol. Biotechnol. 1981, 12, 22-27. 32 T. Ohshima, S. Nagata, K. Soda, Arch. Microbiol. 1985, 141,407-411. 33 M. Oka, Y.4. Yang, S . Nagata, N. Esaki, H. Tanaka, K. Soda, Biotechology and Applied Biochem. 1989, 11,307-311. 34 M. B. Ansorge, M.-R. Kula, Biotechnol. Bioeng. 2000,68,557-562. 35 M. B. Ansorge, M.-R. Kula, 4 p l . Microbiol. Biotechnol. 2000, 53(6), 668-673. 36 M. W. Zink, B. D. Sanwal, Arch. Biochem. Biophys. 1962, 99, 72-7. 37 S . Nagata, H. Misono, S. Nagasaki, N. Esaki, H. Tanaka, K. Soda,/. Ferment. Biotechnol. 1990, 69, 199-203. 38 H. Misono, K. Sugihara, Y. Kuwamoto, S. Nagata, S. Nagasaki, Agnc. Biol. Chem. 1990,54,1491-1498. 39 A. S. Bommarius, K. Drauz, W. Hummel,

1062

I

15 Reduction Reactions

M.-R. Kula, C. Wandrey, Biocatalysis 1994, 20,37-47. 40 a) C. Wandrey, B. Bossow, Biotechnol. Bioind. 1986, 3, 8-13; b) U. Kragl, D. VasicRacki, C. Wandrey, Chem. Ing. Tech. 1992, 64,499-509; c) C. Wandrey in: Enzymes as Catalysts i n Organic Synthesis" (Ed.: M. Schneider), D. Reidel, Dordrecht, 1986, pp. 263-284. 41 R. L. Hanson, J. Singh, T. P. Kissik, R. N. Patel, L. J. Szarka, R. H. Mueller, Bioorg. Chem. 1990, 28,116-30. 42 a) T. Ohshima, K. Soda, Eur. J . Biochem. 1979, ZOO,29-39; b) H. Pommb, D. Vancea, L. Muresan, J . Biol. Chem. 1987,262, 4610-4615; c) I. Vancurova, A. Vancura, J. Vole, Arch. Microbid. 1988, 250, 438-440. 43 T. Ohshima, C. Wandrey, M. Sugiura, K. Soda, Biotechnof. Lett. 1985, 7 , 871-876. 4.4 a) S. Kuroda, K. Tanizawa, H. Tanaka, K. Soda, Biochemistry, 1990,29,1009-1015; b) K. Soda, S. Nagata, H. Tanaka, JP 60 180 580,1985; c) K. Soda, S. Nagata, H. Tanaka, JP 60 180 590,1985. 45 A. Mocanu, G. Niac, A. Ivanof, V. Gorun, N. Palibroda, E. Vargha, M. Bologa, 0. Barzu, FEBS Lett. 1982, 143,153-156. 46 T. Ohshima, C. Wandrey, D. Conrad, Biotechnol. Bioeng. 1989, 34, 394-397. 47 Y. Kato, K. Fukumoto, Y. Asano, Appl. Microbiol. Biotechnol. 1993, 39, 301-4. 48 X.-G. Wang, K. L. Britton, P. J. Baker, S. Martin, D. W. Rice, P. C. Engel, Protein Eng. 1995,8(2), 147-152. 49 X.-G. Wang, L. K. Britton, P. J. Baker, D. W. Rice, P. C. Engel, Biochem. SOC.Trans. 1996, 24(2), 126s. 50 T. J. Stillman, A. M. B. Migueis, X.-G Wang, P. J. Baker, L. Britton, P. C. Engel, D. W. Rice, /. Mol. Bid. 1999, 285, 875-885. 51 R. L. Hanson, M. D. S., A. Banejee, D. B. Brzozowski, B.-C. Chen, B. P. Patel, C. G. McNamee, G. A. Kodersha, D. R. Kronenthal, R. N. Patel, L. J. Szarka, Bioorg Med. Chem. 1999,7,2247-2252. 52 W. Hummel, N. We% M.-R. Kula, Arch. Microbid. 1984, 237, 47-52. 53 W. Hummel, E. Schmidt, C. Wandrey, M.-R. Kula, Appl. Microbiol. Biotechnol. 1986, 25, 175-185. 54 C. W. Bradshaw, C.-H. Wong, W. Hummel, M.-R. Kula, Bioorg. Chem. 1991, 19, 29-39. 5 5 Y. Asano, A. Yamada, Y. Kato, K. Yama-

guchi, Y. Hibino, K. Hirai, K. Kondo,/. Org. Chem. 1990,55,5567-71. 56 N. M. W. Bmnhuber, A. Banejee, W. R. Jacobs Jr., J. S. Blanchard, /. Bid. Chem. 1994, 269(23), 16 203-16211. 57 S. Y. Seah, K. L. Britton, P. J. Baker, D. W. Rice, Y. Asano, P. C. Engel, FEBS Lett. 1995, 370(1-2), 93-6. 58 R. L. Hanson, J. M. Howell, T. L. LaPorte, M. J. Donovan, D. L. Cazzulino, V. Zannella, M. A. Montana, V. B. Nanduri, S. R. Schwarz, R. F. Eiring, S. C. Durand, J. M. Waslyk, W. L. Parker, M. S. Liu, F. J. Okuniewicz, Bang-Chi Chen, J. C. Harris, K. J. Natalie Jr., K. Ramig, S. Swaminathan, V. W. Rosso, S. K. Pack, B. T. Lotz, P. J. Bernot, A. Rusowicz, D. A. Lust, K. S. Tse, J. J. Venit, L. J. Szarka, R. N. Patel, Enzyme Microb. Technof.2000, 26(5-6), 348-358. 59 T. Ohshima, C. Wandrey, M. Sugiura, K. Soda, Biotechnol. Lett. 1985, 7 (22), 871-876. 60 A. Bondi,J. Phys. Chem. 1964,68 (3), 441-451. 61 J.-L. Fauchere, Q S A Rojoligopeptides (Amino-acid Side Chain Parameters and Some Specgc Q S A R Studies), QSAR Des. Bioact. Compd., Barcelona, Spanien 1984. 62 H. Schiitte, J. Flossdorf, H. Sahm,M.-R. Kula, Europ.]. Biochem. 1976, 62, 151-60. 63 M.-R. Kula, C. Wandrey, Methods Enzymol. 1988, 136, 34-45. 64 T. Ohshima, C. Wandrey, M.-R. Kula, K. Soda, Biotechnol Bioeng. 1985, 27, 1616-1618. 65 R. Wichmann, C. Wandrey, A. F. Buckmann, M.-R. Kula, Biotechnol. Bioeng. 1981, 23,2789-2802. 66 A. S. Bommarius, M. Schwarm, K. Stingl, M. Kottenhahn, K. Huthmacher, K. Drauz, 1995, Tetrahedron: Asymm. 6, 2851-2888. 67 M. Whittaker, C. D. Floyd, P. Brown, A.J.H. Gearing, Chem. Rev. 1999,99,2735-2776. 68 W. Hummel, H. Schutte, E. Schmidt, M.-R. Kula, Appl. Microbid. Biotechnol. 1987, 27, 283-291. 69 E. Schmidt, D. Vasic-Racki,C. Wandrey, Appl. Microbid. Biotechnol. 1987, 26, 42-48. 70 E. Schmidt, W. Hummel, C. Wandrey, Proc. 4th Eur. Conp. Biotechnol. 1987, 189-191. 71 C.-H. Wong, G. M. Whitesides, j . Am. Chem. SOC. 1981, 103,4890-99. 72 B. L. Hirschbein, G. M. Whitesides, /. Am. Chem. SOC.1982, 104,4458-GO. 73 U.Groth, T. Huhn, B. Porsch, C. Schmeck,

References I1063

U . Schollkopf, Liebigs Ann. Chem. 1993, 71 5-7 19. 74 U. Groth, C. Schmeck, U. Schollkopf, Liebigs Ann. Chem. 1993, 321-323. 75 2. X. Shen, J. F. Qian, W. J. Qiang, Y.W.

Zhang, Chin. Chem. Lett. 1992,3 (4), 237-238. 76 G.C. Cox and L.M.Hanvood, Tetrahedron: Aqmm. 1994,5,1669-1672.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyriqht 0Wiley-VCH Verlaq GmbH, Weinheim 2002

16 Oxidation Reactions 16.1

Oxygenation of C-H and C=C Bonds

Sabine Nitsch, Cideon Grogan and D. Ashcroft 16.1.1 Introduction

Reactions catalyzed by oxygenase enzymes (mono or dioxygenases) are interesting for applications in organic synthesis. There are numerous examples of such reactions in biological systems, yet there are few chemical reagents or catalysts that can compete with biocatalysts. Examples of monoxygenases catalyzed biotransformations are shown in Figure 16.1-1. These include heteroatom oxygenation, aromatic hydroxylation, Bayer-Villiger oxidation, double-bond epoxidation and hydroxylation of nonactivated hydrocarbon atoms. The latter can occur with regio-, stereo-,

I

I

Monoxygenases

Figure 16.1-1. Some examples of monooxygenase-catalyzed biotransformations.

1066

I

76 Oxidation Reactions

and in some cases enantioselectivity that is difficult to achieve using conventional chemistry. There is a large body of literature describing the exploitation of these oxygenase enzymes for synthetic applications, and the current chapter will only give a few representative examples of what has been done. There are few reports on biotransformations using isolated oxygenase enzymes because of several problems with cell free enzymes. Many of the oxygenases are membrane-bound, and require a complex set of co-factors and co-proteins. Despite the fact that a large number of their genes have been identified, the enzymes themselves are difficult to isolate and quite unstable. Thus, oxidative bioconversions, especially on an industrial scale, generally use whole-cellbioconversion techniques, which makes them less accessible for use by organic chemists in organic synthesis laboratories. However, very recently new techniques have been developed for the isolation, cloning and over-expression of oxygenases in heterologous expression systems, and the number of reports using isolated systems is increasing. These new developments will be discussed at the end of the chapter, and point to the possibility of overcoming technical difficulties that have hampered the application of oxygenase systems in biocatalysis in the past. A number of excellent reviews with comprehensive coverage on the literature of biooxidations have appeared in journals and books['-*]. In this chapter we will only try to highlight some of these biotransformation reactions, in particular hydroxylation of non-activatedcarbon atoms and double-bond epoxidation reactions. 16.1.2

Hydroxylating Enzymes

Hydroxylation reactions in nature are generally catalyzed by monooxygenase, a subclass of the oxidoreductase enzyme group. These enzymes are very important and ubiquitous proteins found in almost all living cells, ranging from bacterial to mammalian. One of the most important groups of this type of enzyme is the cytochrome P450 family. These are heme-dependent monooxygenases whose essential role, among other functions, is to ensure detoxificationof exogenous compounds by rendering these very often lipophilic molecules water soluble, thus facilitating their excretion. Because of this essential function, mammalian monooxygenases have been thoroughly studied in the context of drug metabolism[1, 9-141. The majority of the cytochrome P450 systems reported to date are multicomponent, requiring the involvement of additional proteins for transport of reducing equivalents from NAD(P)H to the terminal cytochrome P450 component. Increasing attention is given to microbial monooxygenases, in particular in their application for biotransformations. One of the earliest and most important industrial applications of microbiologicallymediated bioconversions is the 11-a-hydroxylation of progesterone using Rhizopus arrhizus cells (Figure 16.1-2) [''I. Microbial monooxygenases tend to be soluble enzymes that can be purified fom cell free extracts, and a number of crystal structures of microbial monooxygenases are now available[", 171. The first X-ray structure for P450 was that of cytochrome P450,,,, which was isolated from Pseudomonas putida, and catalyzes the 5-exo hydroxylation of its natural substrate D-camphor to 5-exo hydroxycamphor as shown in Figure 16.1-2.

16.7 Oxygenation of C-H and C=C Bonds

Progesterone

11-a-hydroxyprogesterone

Pseudomonas putida

Camphor

11067

*

O k 0 ” 5-exo-hydroxycamphor

Figure 16.1-2. Regio- and stereoselective microbiological hydroxylation of progesterone and D-camphor.

The P45OC,, enzyme has served as a model system for general studies of cytochrome P450 enzymes in terms of structure, function and mechanism[16,‘*I. The exquisite regio- and stereoselectivitycan be explained by the active site geometry of the P45OC,,, which shows several van der Waals interactions with hydrophobic side chains and a key hydrogen bond between tyrosine 96 and the carbonyl oxygen of the substrate. Removal of the tyrosine-96hydroxyl group by site-directedmutagenesis or removal of the carbonyl oxygen by using camphane results in loss of selectivity[”J. There are also an increasing number of non-P450 type biohydroxylases. Examples are the n-octane o-hydroxylaseof Pseudomonas oleovorans and the n-decane hydroxylase of Pseudomonas denitriicans, which have been shown to be also responsible for epoxidation of 1-octeneand for O-demethylation of heptyl methyl Similarly, the progesterone 9-a-hydroxlase from Norcardia sp., one of the first microbial hydroxlases obtained in crude form, was shown not to be a cytochrome P450 Interestingly, this enzyme allows for the functionalization of the steroid skeleton, thus opening the way to production of the C-11-oxygenated corticosteroids[26] as shown in Figure 16.1-3. Other very important examples of non-cytochrome P450 enzymes are methane monooxygenases. These appear to be more reactive than P450 oxygenases and are able to catalyze the conversion of methane to methanol, chemically one of the most difficult steps [271. These enzymes have been shown to reductively activate dioxygen for incorporation into a wide variety of hydrocarbon substrates, including alkanes, alkenes and alicyclic or aromatic hydrocarbons. The enzyme harbors a hydroxybridged dinuclear iron cluster in its active site, and its structure has been determined by X-ray crystallographyL2*].

1068

I

do

IG

Oxidation Reactions

Nocardia sp.

/

0

Progesterone

o-&o

OH

/

I

Figure 16.1-3. Use of9-a-hydroxylation of progesterone as a way to corticosteroids.

16.1.2

Hydroxylating Enzymes

The active site of P450 monooxygenases contains an iron-heme center that is directly involved in the oxidation process by activating molecular oxygen. The catalpc cycle by which cytochrome P450-mediated alkane hydroxylation occurs is by now well studied[*,181. The reaction cycle of cytochrome P450,,, is outlined in Figure 16.1-4. This mechanism involves (i) reversible substrate binding which converts the sixcoordinate, low spin form of the protein to the penta-coordinate high-spin form, (ii) electron reduction of the ferric substrate-enzyme complex by flavoprotein NADPH-

I Fe3'ROH

k

2H+l Figure 16.1-4. The catalytic cycle of cytochrome P450 enzymes.

16. I Oxygenation ofC-H and C=C Bonds

cytochrome P450 reductase leading to the ferrous enzyme, (iii)binding of molecular oxygen to give the six-coordinateiron-dioxygen intermediate, (iv) + (v) reduction of this species with a second electron and addition of two protons, thus leading to an activated oxygen intermediate, (vi) insertion of an oxygen atom into the substrate, and (vii)release of the iron atom in its original ferrous state. Apart from the products of steps (iv) and (v), all these intermediates have now been investigated by crystallography using trapping and cryocrystallographymethods [lS1. The exact mechanistic details of oxygen insertion into the C-H bond are still the subject of intense discussion. One of the most popular proposals appears to be that of the so-called “rebound mechanism”, which proceeds by an initial hydrogen abstraction from the alkane (RH) by the active oxygen intermediate to form a radical R and a hydroxo-iron species as intermediates. The radical then rebounds on the hydroxy group and generates the enzyme-product species.I’[ Alternative proposals involve cationic intermediates f3O1 or two-state reactivity with multiple electromer species for epoxidations L3l]. It should be noted that the same cytochrome P450 enzyme is able to achieve reactions as different as double-bond epoxidation or heteroatom demethylation. Thus it appears clear that the chemo-, regio- and stereoselectivity of the reaction is a function of the nature and the fit of the substrate, or, more properly, of its transition state with the protein, rather than being governed by enzyme reaction specificity. It is obvious that all these very powerful enzymes are of high synthetic value for the organic chemist since they prove to be able to achieve, at normal temperature and in aqueous media, reactions which are very difficult, if not impossible, to perform using conventional chemistry. One additional bonus offered by these biological tools is their generally high selectivity. It is thus understandable that a variety of oxygenative biotransformations have been explored using numerous substrates. We will focus in the following pages on two such particularly interesting reaction types, namely the hydroxylation of non-activated carbon atoms and the stereospecificepoxidation of “isolated double bonds. 16.1.4 Hydroxylation of Non-Activated Carbon Atoms 16.1.4.1

Hydroxylation of Monoterpenes

Because of their involvement in the flavor and fragrance industry, monoterpenes are one type of natural compounds which have been considered as interesting substrates for biohydroxylation studies. For instance, geraniol, nerol and linalool were studied by different groups and were shown to lead to the 8-hydroxylatedproducts with the fungus Aspergillus r ~ i g e r [ ~as~ ]well as with four strains of Botrytis ~ i n e r e a [ ~ ~ ] . Interestingly, the same 8-hydroxylated products were found starting from the corresponding acetates, which were shown to be hydrolyzed to the starting alcohol by the fungus Aspergillus niger prior to hydroxylation [341. This C-8 regioselectivity has also been observed in hydroxylations of geraniol and nerol with reconstituted

1070

I

76 Oxidation Reactions

-I_OH

P450cath

'H

Figure 16.1-5. Retention of configuration during the hydroxylation at C-8 o f isotopically labeled geraniol.

'''[3H]

(R)-( 1 -'Hl)[l -3Hl]

+OH

$H

+OH

"H

+,OH

[%I

'H

(S)-(1-'Hl)[l -3Hl]

hydroxylating enzyme systems from rabbit l i ~ e r [ ~ ~and - ~ from ' ] plant cells like Vinca rosea as well as Catharanthus roseus (L.) G. In this last case, incubation of different I3C- and 2H-labeledgeraniols revealed that hydrogen abstraction is completely regioselective in favor of the CH3 group trans to the chain at C-6, i. e. at position C-8. An intramolecular isotope effect of KH/KD 8.0 was determined, suggesting that the hydrogen abstraction is one of the major rate determining steps. Furthermore, Fretz and Woggon [411 have studied incubation of the (R)(8-'H1) (8-3H1)and (S)(8-2H1)(8-3H1)geraniols (Fig. 16.1-5).This resulted in the formation of the chiral 8-hydroxy products, thus indicating clearly retention of configuration during the allylic hydroxylation process. Interestingly, in neither of these cases has an allylic radical rearrangement (migration of the double bond) been observed. Bioconversions of (+)-limonene,the major constituent of citrus essential oils, have also been studied in recent years in order to afford biotechnological routes to interesting products of natural source valuable for the perfume and/or flavor industries. For instance, (+)-limonenewas shown to be transformed by Pseudornonas gladioli to (+)-a-terpineol(one of the most commonly used products in fragrances and flavors i. e. lemon, nutmeg, orange, ginger, peach and spices), which is resistant to further degradation by the bacterium, and to (+)-perillicacid, which is further The corresponding levorotatory limonene antipode is known as being the primary olefinic constituent of the volatile oils of immature Mentha piperita (peppermint), Mentha spicata (spearmint) and Perrilla f i t e s c e n s leaves, whereas (-)-menthone, (-)-camoneand (-)-perilly1aldehyde, respectively, are the major oxygenated compounds. The enzymatic hydroxylation of (-)-limoneneat C-3, C-6 and C7 to the corresponding derivatives has been studied using light membrane preparations from leaves of each of these plants. It has thus been shown that they lead to the

1 6 1 Oxygenation of C-H and C=C Bonds

c:

Pseudornonas gladioli

A

T

(+)-lirnonene

O

COOH

A

H

(+)-a-terpinol

(+)-perillic acid

ro

7

(OH

(-)-carvone

(-)-trans-carveol

(-)-lirnonene

(-)-PeWl alcohol

(-)-perilly1 aldehyde

M I : Mentha spicata M2: Perilla frutescens M3: Mentha piperita

(-)-trans-isopiperitenol

A.

+

cellulosae

19%

Figure 16.1-6.

(-)-rnenthone

5%

12%

21%

Bioconversion of limonene using various biocatalysts.

corresponding oxygenated compounds in a mutually exclusive manner in each species. This suggests very strongly that different forms of cytochrome P450 are present in each type of plant, each one showing an exclusive regiochemistry of as shown in Fig. 16.1-6. oxygen Another terpene interesting for flavor chemistry is 0-ionone (Fig. 16.1-7). Its microbiologically mediated transformation has been explored in order to afford a mixture of derivatives that is utilized as an essential oil of tobacco, used for tobacco 451. One of the best microorganisms capable of flavoring at the ppm converting 0-ionone to the desired mixture of its useful derivativeswas an Aspergillus niger strain. This process has been recently improved using bioconversion in the presence of organic solvents and immobilization techniques. Thus, this fungus

1072

I

1G Oxidation Reactions

go k0k0 bH

p-ionone

(4R)

(25)

45%

M : A . niger or

30% 11Yo

46%

A. awamo

$o

Qo /

(R)-(+)-pulegone

58%

OH 4%

10%

OH 6%

+

Sfreptomyces albus

OH sulfone of

HO

24%

20%

rnyrcene Figure 16.1-7.

Biohydroxylations o f various monocyclic terpenes

could be repeatedly used for microbial conversion of p-ionone in the presence of isooctane for more than 480 hr4'1. Another Aspergillus strain, A. awarnori, has also been shown recently to achieve hydroxylation of p-ionone. This led to a mixture of two alcohols, the major product being a building block usable for further synthesis of abscisic acid (an important phytohormone) analogs 1471. Other monocyclic terpenes like for instance R-(+)-pulegone (a mint-like odor monoterpene ketone which constitutes the main component of Mentha pulegium essential oil)L41' menthols, terpinolene and carvotanacetone r4'] have been investigated. All these substrates proved to be transformed by various Aspergillus strains, including A. nigec leading essentially to monohydroxylation. Also, monocyclic

1 6 I Oxygenation ofC-H and C=C Bonds

I

A. niger

30°C,pH 7.2 (+)-6-endo-hydroxyfenchone

(+)-fenchone

Acetobacter mefhanolicus

(+)-fransverbenol

(+)-a-pinene

(+)-verbenone

(+)-transSobrerol

OH

60%

(+)-camphor

7%

v

9%

Y Bacillus cereus

OH 1,&cineole

Figure 16.1-8.

74%

Biohydroxylation of various bridged bicyclic terpenes.

sulfoxide derivatives of the linear terpenes myrcene and ocimene were shown to be good substrates for several bacteria and fungi, whereas they were themselves only very poorly Finally, some bicyclic monoterpenes have also been recently described to be subject to microbiological hydroxylation (Fig. 16.1-8).For instance (+)-fenchonewas transformed to (+)-6-endo-hydroxyfenchoneby A. niger[’l], and a-pinene led to the predominant metabolites (+)-trans-verbenol,(+) -verbenone and (+)-trans-sobrerolby the action of several strains of the methylotrophic species Acetobacter methanoliC U S [ ~ ~(+)-Camphor ]. is transformed to the major metabolite 6-endo-hydroxy-camphor by cytochrome P450,,, enriched intact cells of Streptomycesgri~eus1~~1, and 1,S-cineole (the major component of the oil from leaves of Eucalyptus radiata var.) is hydroxylated to 6-(R)-exo-hydroxy-l,&cineoleby the bacterium Bacillus cereus following a high yielding (74%) and stereospecific route [541. Interestingly, this same bacterium had been previously described to be able to achieve hydroxylation of 1,4-cineole yielding good yields of essentially pure 2-(R)-endoand 2 4 R)-exo-hydroxy-l,4-cineole[551.

1073

1074

I

IG

Oxidation Reactions

16.1.4.1

Hydroxylation of Monoterpenes

Similarly to monoterpenes, several studies aimed at the microbiological hydroxylation of sesquiterpenes have been described, and a review has summarized the most interesting results obtained in this area Since then, additional examples have been published. Thus, three germacrone-type sesquiterpenoids, (+)-germacrone4,5-epoxide, germacrone and (+)-curdione,were described as being transformed by Aspergillus niger[57].The interesting feature of these results is the fact that they essentially led to hydroxylated guaiane-type sesquiterpenoids (together with allylic alcohols and spirolactone) which arise from transannular cyclization of the carbon

ReHo+ q +

HO

1

(+)-gerrnacrone 4,5-epoxide

3%

4%

3%

4%

__c

nger

/

HO

'

(+)-germacrone

R1 =OH,R2 = H 9% R1 = H, R2 = O H 10%

(+)-curdione

;

H

Beauvaria sulfurescens

H'

\

OCONHPh Phenylcarbarnoyl derivative of dihydroarternisinin Figure 16.1-9.

Biohydroxylation of various higher terpenes.

H" \ OCONHPh 15%

1 6 1 Oxygenation of C-H and C=C Bonds I1075

skeleton as shown in Fig. 16.1-9. The biohydroxylation of an N-phenylcarbamoyl derivative of dihydroartemisinine by the hngus Beauveria sulfirescens has been described[58].This allowed the preparation of new and novel derivatives of artemisinine, a drug known to be active against Plasmodiumfalciparum, the strain responsible for malaria, which claims more than one million lives a year. The C-1O-Nphenylcarbamoyl derivative of dihydroartemisinine, a highly oxygenated sesquiterpene, is thus converted into its 1Chydroxymethyl derivative in 15% yield. Although this yield can be considered as rather modest, this biohydroxylation is interesting since it allows us to prepare derivatives which retain the peroxide group required for biological activity of these drugs. Also, it emphasizes the possibility to favor biohydroxylation by introducing an amide or urethane group into a substrate, as already observed on other model substrates (Fig. 16.1-9)[59* 60]. A series of biotransformations of GP-santonin and of some of its derivatives, achieved by Cuwularia lunata and Rhizopus nigricans cultures, have also been described (Fig. lG.l-lO)[G1]. Depending on the strain used and on the starting substrate, several metabolites were obtained, including products resulting from hydroxylation as well as double bond and/or carbonyl reduction. The same group also described biotransformations of several 1,G-difunctionalizedeudesmanes leading to 12-hydroxyderivatives which are interesting intermediates for the synthesis of G,l2-e~desmanolides[~~]. Similarly, starting from GP-acetoxyeudesmanone,biohy-

‘0 6P-santonine

19%

p& ’

12%

6p-acetoxy-l p-4pdihydroxyeudesmane

p& ”

deans-

OAc

6p-acetoxyeudesmanone

Figure 16.1-10.

1 10%

0

0

HO

OAc

h0;

HO

OAc

72%

$J),,Jy

HO

OAc

15%

Further examples of biohydroxylationsof bicyclic natural compounds.

1076

I

7G Oxidation Reactions

Gibberellin 20 -19-lactone

Figure 16.1-11.

R'=OH, R2=H, R3=CH,0H R'=H, R'=OH, R3=CH20H R'=H, R2=OH, R3=CH0

Some examples o f diterpenes hydroxylation by Cibberellafujiikuroi

droxylation was achieved at C-11 by the fungus Rhizopus nigricans, thus opening another way to the synthesis of the same lactone targetsFG3]. Several diterpenes have also been described recently to be subject to microbiological hydroxylations. Thus, as shown in Fig. 16.1-11, a compound prepared from gibberellin A13 was transformed by the fungus Gibberella fijikuroi, affording three metabolites, two of these arising from hydroxylation at its 3a- and 19-p0sitions[~~]. Similarly, the same fungus was shown to transform isoatisene derivatives into rearranged isoatisagibberellin derivatives[G51. Sclareol, a natural product first isolated from the essential oil of Salvia sclarea L. (Labiatae) in 1931, is used for diverse applications in the perfumery and flavoring industries and in folk medicine. This diterpene has been described recently to be hydroxylated by three strains, i. e. Cunninghamella sp., Septomyxa afinis[661and Mucor p l u m b e ~ s [68], ~ ~leading , essentially to hydroxylation reactions on the A ring of this compound (Fig. 16.1-12).Some of these metabolites could be used for further synthesis of some biologically active targets or as mammalian metabolism models. Amazingly, as in the case of sciareol, the presence of an oxygenated function on the C ring position of grindelic acid again orients the hydroxylation process towards the A ring, since mainly 3P-hydroxylation is observed [691. This is to be compared with the results obtained in studying the behavior of Rhizopus and Aspergillus strains as potential hydroxylating species for kaurene sesquiterpenes 1"' (Fig. 16.1-13).Inter-

IG. 1 Oxygenation of C-H and C=C Bonds

Cunninghamella

sp. NRRL 5695

R' = OH, R2 = CH3 R' = H, R2 = CHzOH

36% 50%

OH Septomyxa affinis

R' = H, R2 = R3 = 0 49% R' = R3 = H, R' = OH 3%

Scalerol

R'=OH,R2=R3=H

27%

Mucor plumbeus

k3

R1 = OH, R2 = R3 = H 84% R1 = R3 = H, R 2 = OH 6% R1 = R 2 = H, R3 = OH 6% Figure 16.1-12. Hydroxylation of natural sclareol using different microorganisms.

estingly, the fact that the starting substrate bears an oxygenated function on the A ring now orients the oxidations catalysed by R. nigricans toward the C ring (and in particular to the C-13 position). This nicely resembles the results previously observed in the steroid family by Jones and and once more emphasizes the role of a preexisting oxygenated function which operates as an anchoring and thus a sitedirecting entity inside the hydroxylating active site. By modifylng the location of this function on the starting material, it should therefore be possible to orient the hydroxylation locus differently. This is nicely exemplified by hydroxylation of stemodine, a diterpene bearing a preexistent OH group at position 2 of the A ring. Hydroxylation by C. elegans and by Polyangium cellulosum then orients the hydroxylation process toward positions 17 and 19[721.

I

1077

1078

I

16 Oxidation Reactions

t

nigsr

H

O

mixture

W

Grindelic acid

+ mixture

:ans

I

OAc

I

\

OAc

Kaurene derivative

22%

Stemodin

M: C. elegans

M: I? celldosum

I

R' = R 3 = R 4 = H, R'= a-OH R' = R3 = R 4 = H, R'= P-OH R' = R2 = R 4 = H, R 3 = O H R2 = R 3 = R 4 = H, R' = O H R2 = R 3 = H, R' = R 4 = OH

Figure 16.1-13. Effect of a preexisting oxygenated function on the orientation of a biohydroxylation process.

16.1.4.3

Hydroxylation of Steroids

Because of their utmost importance as bioactive molecules, steroids have been the most thoroughly studied family as far as microbiological hydroxjlations are concerned. The most important features and references have been put together by Holland in his important monograph'73].At the present time, one could presumably almost consider that one or even several strains are known which are able to introduce a hydroxyl group at every carbon atom of the steroidal framework. Obviously, however, further work will have to be achieved in order to improve the selectivities and yields

16.7 Oxygenation ofC-H and C=C Bonds

I

Curvularia lunata o#oH

&OH

fl op

0

Cortexolone

C E CHH3

0

ZEZH3

Curvularia lunata

/

Norethisterone acetate Figure 16.1-14.

Examples of steroid hydroxylations by Curvularia lunata.

of these bioconversions. Thus, the course of the Ilp-hydroxylation of cortexolone by Curuularia lunata, a hydroxylation of considerable commercial importance, has been more recently reexamined. The work described by Chen and Wey[741focused on the improvement of this process by studying the characteristics of mycelial growth as well as the role of substrate addition time and dissolved oxygen tension. These studies have provided some more insight into the fundamental aspects of this biotransformation. The 110-hydroxylation of norethisterone acetate by the same microbial strain has also been described[75](Fig. 16.1-14). 16.1.4.4

Miscellaneous Compounds

Although being initially the major part of literature concerning microbiological hydroxylations,natural compounds of the terpene or steroid families have not been the only ones to be studied in this context. Indeed, more recently studies have focused on using biohydroxylationsto provide interesting synthetic intermediates, in particular chiral intermediates. Linear or branched-chain alkanes have been shown previously to undergo hydroxylation by various microorganisms, and can lead for instance to fatty acids, hydroxyacids or a-dicarboxylicacids of commercial importance. Pseudornonas oleovoram is one of the strains capable of achieving such transformations, but this process suffers from the fact that the monocarboxylic acids formed in the first step are submitted to P-oxidation and thus used as a source of energy and carbon. Such problems can now be overcome using heterologous expression of P450 genes in microorganism, which will be discussed later on. An interesting aspect of biohydroxylations is that the biocatalyst can in principle generate one single enantiomer starting from a prochiral substrate - a reaction which could be defined as “enantiogenic”1“‘. This enantiogenic hydroxylation can lead to enantiopure compounds by stereospecific attack of one single enantiotopic

1079

1080

I

76 Oxidation Reactions

n=l n = 2, 3, 4

yield = 26%, ee = 20% yield > 65%, ee > 98%

Montierellc isabellina

chroman Figure 16.1-15.

yield = 1070,ee > 98%

Examples of regio- and stereoselective benzylic hydroxylation.

face of the starting substrate. Biohydroxylation reaction can therefore be used to prepare high value chiral synthetic intermediates from low value prochiral starting materials, and there are now a number of reports of such reactions. Thus, benzocycloalkenes have been described to undergo bacterial hydroxylation by the Pseudomonas putida strain UV4. As shown in Fig. 16.1-15, this yielded exclusively hydroxylation at the benzylic position, and also one single enantiomer, i. e. the (R)-alcohol.The biotransformation of benzocyclobutene proved, however, to be different from that observed for higher benzocycloalkenes, presumably because A similar result has been observed by Holland of its particular chemical and coworkers in the course of chroman biotransformation by the fungus Mortierella isabellina [781, which leads, although in low yield, to the benzylic (R)-alcohol. Another interesting example of asymmetric synthesis from a prochiral substrate is the preparation of (S)-naproxen,a non-steroidal anti-inflammatory drug. It has been shown that several strains are able to regioselectively oxidize one of the enantiotopic methyl groups of the isopropyl moiety. This allows the preparation of the corresponding acid, which is obtained with high enantiomeric purity (Fig. 16.1-16). Next to Aspergillus niger the fungus Beauueria bassiana (previously classified as Sporotrichum sulfirescens and B. sulfirescens) is one of the most frequently used fungal biocatalystl2). In particular, hydroxylations of piperidine and pyrrolidine derivatives have been studied by several groups, and interesting regio- and sterpro R

CH3

Organism Cordyceps rnihfaris Graphinium fructicola Exophiala rnansonni Exophialajeanselmei

ee (%) 99 96 68 66

Figure 16.1-16. Selective hydroxylation o f one enantiotopic methyl group as an approach t o optically pure Naproxene.

1 6 1 Oxygenation ofC-H and C=C Bonds

Beauveria sulfurescens

-

&

HO COPh

83%

Beauveria sulforescens * \

COPh

0

Figure 16.1-17. Hydroxylation versus epoxidation of two spiro-bicyclic amides.

COPh

\

74%

CoPh

eoselectivities have been rep0rted[~'-'~1.It should be noted that the ring nitrogen generally needs to be protected for a successful biohydroxylation.This can be used to advantage since the choice of protecting group can influence the regio- and stereochemistry of the hydroxylati~n[~~]. Some examples of hydroxylations of spirobicyclic amides are shown in Fig. 16.1-17.Similarly to previous results described by "I, these led to good Fonken and coworkers[". 861 and by Furstoss and coworkers[87* yields of hydroxylated products. In all these cases, the regioselectivity of the reaction is partly or even exclusively oriented toward the C-9 carbon atom, a result which could have been predicted on the basis of the previously described results. Interestingly, a similar substrate bearing a double bond at carbon C-9 led to the corresponding epoxide. Because they constitute partial structures of various higher terpenes and/or steroids, enantiomers of different substituted hexahydronaphthalenones are pivotal intermediates in the total synthesis of these target compounds. Therefore, several differently substituted octalone derivatives have been studied for microbiological hydroxylations. These substrates were prepared in optically active form by chemical ketone and were submitted for screening synthesis from (S)-(+)-Wieland-Miescher's with nine strains known to hydroxylate polyterpenic or steroidal substrates. Thus, submitted to a culture of Rhizopus arrhizus, these substrates led to allylic hydroxylation at the B ring, as shown in Fig. 16.1-18f8',"1. Similar results were obtained by Azerad and coworkers ['*I in the same series, starting from differently substituted octalones. These authors have investigated the biotransformation of their substrates with a variety of fungal strains. For most of these strains, the (R)-enantiomer of hydronaphthalenone led to the 8-hydroxyenone as the main product, i.e. again a product of allylic hydroxylation, which is quite disappointing since this product is easily accessible by (e1ectro)chemicaloxidation. However, the fungus Mucor plumbeus produced another hydroxylated metabolite, the 6a-hydroxyl derivative. Interestingly, the S-enantiomer ofthe starting substrate only led to the 8-hydroxyenonein this last case. Introduction of an additional methyl group on the carbon framework of the starting octaenone also led to different regioselectivities of the hydroxylation. Hydrindane derivatives, which bear a five-membered B ring (instead of a sixmembered ring in the decalones derivatives) have also been examined for bio~ Ithe . hydroxylations observed hydroxylation by the fungus Rhizpous a r r h i ~ u s [ ~All now occur at position 3, to the a$-unsaturated ketone. This can be considered as

I

1081

42% 55%

R’=OH, RLH R’=H, $=OH

27% 9% 0

0

@

0

Rhizopus

+

arrhizus

* HO@OH

HO

HO@

63%

9%

54%

16%

n

n’/>(j -

403

0

Mucor plumbeus

~

0

22%

bH

aotBu ofio +

0

58% OH

Figure 16.1-18.

,-,\OH 6

+

;a

66%

Rhizopus arrhizus

-

8%

bH

Microbiological hydroxylation of differently substituted octalones.

being formally analogous regioselectivity as compared to the results obtained on decalones. In this case, however, reactivity is identical in both antipodal series, and led almost quantitatively but with moderate or low stereoselectivityto the formation of the epimeric alcohols. Interestingly, these biohydroxylations prove to be complementary to lead tetraacetate oxidation of these substrates, which affords the 6-acetyl substituted products. Stmctually much more complex molecules have also been submitted to regioselective enzymatic hydroxylation. Two such examples have been described involving milbemycin, a sixteen-membered macrolide which exhibits broad-spectrum insecticidal and acaricidal activity, and monensin, a carboxylic polyether antibi~tic”~. 941. Milbemycin (Fig. 16.1-19) was thus regioselectivelyhydroxylated at the 130 position (followed eventually by a C-29 hydroxylation) to afford the 138,29-

1 6 1 Oxygenation of C-H and C=C Bonds

H

H

Monensin Sebekia

/

benih/

<

Figure 16.1-19.

Regioselective hydroxylation o f structurally complex substrates.

hydroxylated product by a strain isolated from solonized brown Mallee oil (collected in Adelaide in Australia) and identified as Streptomyces cavourensis. It is interesting to emphasize here the high regioselectivity observed for this hydroxylation of a rather complex and multifunctional compound. Even more complex is the structure of monensin, a compound which has been extensively used as an anticoccidial agent for poultry and shown to improve the efficiency of feed utilization in ruminant animals. When submitted to a culture of Sebekia bevihana, monensin was first quantitatively converted by enzymatic reduction of the 6-hydroxy-ketone(which is in equilibrium with its hemiketal tautomeric form) and was regioselectively further hydroxylated at the C-29 methyl group as well as at the nearby ethyl group ~ubstituent[”~]. All the previously described examples exemplify the ability of various monooxygenase enzymes to achieve, often with good to reasonable yields and interesting

I

1083

1084

7G Oxidation Reactions

I regioselectivities, the hydroxylation of non-activated carbon atoms which are inaccessible using conventional chemistry. This thus allows one-step syntheses of these metabolites, which can in certain cases be of high enantiomeric purity. Another type of oxygenation reaction which is of interest is the stereoselective epoxidation of double bonds, the essential aim being in this matter the access to epoxides of high enantiomeric purity. This will be the subject of the following part of the discussion. 16.1.5

Epoxidation of Olefins

As discussed previously, monooxygenases provide highly activated oxygen intermediates that can oxidize a wide range of functional groups. One of the most studied among these has been the epoxidation of olefins [95-971. This epoxidation is particularly interesting when applied to prochiral double bonds. Spectacular success has been obtained in the field of asymmetric chemical epoxidation, notably using Sharpless epoxidation catalysts for ally1 alcohols and Jacobsen catalysts for aryl olefins, which has made epoxides key intermediates in the synthesis of chiral compounds. However, these chemical catalysts often have a limited “substrate” range, and biocatalysts can provide access to complementary structural motifs. Without attempting to be exhaustive, we will try in this chapter to focus on results allowing us to directly oxidize olefins to their corresponding epoxide, using microbial cells. Other sources of monooxygenases, such as mammalian cells (microsomes) or plant cells, have been studied in this respect. However, these will not be considered in this review. 16.1.5.1

Epoxidation of Straight-Chain Terminal Olefins

One of the earliest observations implicating the formation of epoxides during microbial olefin metabolism was the report by Bmyn in 1954 that Candida lipolytica grown on I-hexadecene produced 1-hexadecanediol (about 5 % of the hydrocarbon consumed was accounted for as the diol)[98].Molecular “ 0 was shown to be incorporated into this diol, and the 1,2-epoxide was identified as one of the bySeveral further reports confirmed that enzyproducts of this metabolism[”. “1‘. matic systems are able to achieve epoxidations. For instance, Van der Linden showed in 1963 that Pseudomonas aeruginosa grown on n-heptane and resuspended in a buffer solution produced the epoxide from 1-octene (Fig. 16.1-20)110’1. This led the authors to conclude that this epoxide was formed by enzymes already present in the alkane-growncells and that epoxidation might be catalyzed by the same hydroxylases that would normally oxidize alkanes. A similar conclusion was reached by Maynert and coworkers [lo2],who demonstrated that epoxides are obligatory intermediates in the metabolism of simple olefins in rat liver microsomes. However, the real breakthrough in the study of enzymatic epoxidations is due to Abbot and coworkers I1O3] and to May and coworkers [lo41,who established unequivo-

1G. 1 Oxygenation ofC-H and C=C Bonds M *

W& ’ o % + H O -

/

H 1-octene

(R)

ally1 benzene M: Pseudomonas oleovorans Figure 16.1-20.

I

Stereospecific epoxidation of straight-chain terminal olefins.

cally that epoxides are formed from terminal olefins by the bacterial strain Pseudomonas oleovorans (Fig. 16.1-20).They showed that 1-octene is epoxidized to 1,2-epoxyoctane of (R)-configuration(ee 70%) or hydroxylated to 7-octen-1-01.The 1,7-dieneis exclusively epoxidized, affording 7,8-epoxy-l-octene,which can be further processed to the corresponding diep~xide[’~~]. It was shown later that this monoepoxidation was stereospecific, leading to the R(+)-7-epoxideshowing an ee of about 80%. Furthermore, the diepoxide was shown to be essentially of (R,R)configuration. This interestingly indicates that the configuration of the monoepoxide formed at one end of the molecule profoundly affects the stereochemicalcourse of the reaction. Indeed, the authors showed that when starting from racemic monoepoxide, the diepoxide was essentially formed from the (R)-monoepoxide.Interestingly it was observed that olefins bearing an allylic (or homoallylic) hydroxyl were not epoxidized, but were converted instead to the corresponding saturated ketones. One of the most useful characteristics of this work is the fact that these epoxides could be routinely produced at yields approaching (at best) 1 g L-’ after simple overnight shaking using whole-cell or even crude cell-free systems. Thus, these results clearly opened the way to a new type of biotransformation which should be very useful for organic synthesis. The enzymatic system involved in hydroxylation reactions of long-chain alkanes had been previously studied by Coon and coworkers, who isolated an enzyme system from P. oleovorans that catalyzes co-hydroxylation of alkenes and fatty ac106-1151. This was resolved into three protein components: rubredoxin (an iron-sulfur protein of molecular weight 19 000), an NADH-rubredoxin reductase (a flavoprotein of molecular weight 55 000) and an “o-hydroxylase”(characterized as being a non-heme iron protein, with one iron atom and one cysteine per polypeptide

1085

1086

I chain). Interestingly,it was shown that this same enzyme system is responsible for 16 Oxidation Reactions

the conversion of terminal olefins to their corresponding 1,2-epoxides[lo4].This leads to a competition between the two types of biotransformations, which results in a specific pattern for each type of substrate. Thus, further investigation demonstrated that this monooxygenase can produce epoxyalkanes with from six to twelve carbon atoms containing terminal alkenes. As a result of the influence of carbon chain length on epoxidation versus hydroxylation it was shown that hydroxylation predominates for the “short” substrates propylene and 1-butene, but that epoxidation activity falls off much less readily than hydroxylation for “long”substrates. For the “medium” length substrates, like for instance 1-octene, both reactions do occur. Thus, this substrate is epoxidized to 1,2-epoxyoctaneor hydroxylated to 7-octen-1-01,while for 1-decene epoxidation largely predominates. Interestingly, the epoxidation reaction exhibits a specificity far different from that expected for chemical reactivity. Indeed, terminal olefins are epoxidized exclusively even in the presence of more highly substituted (electron-rich)double bonds. Thus, cyclic and internal olefins were not epoxidized. This indicated that the substrate specificity pattern observed severely moderates the inherent reactivity of the activated oxygen species involved in these transformations. Methyl imidoesters as well as sodium cyanide were found to be inhibitors of enzymatic epoxidation, and the potency of a homologous series of imidoester inhibitors was examined. In the reaction with dienes, 1,s-hexadiene to 1,ll-dodecadiene were epoxidized while dienes with a smaller number of carbon atoms were hydroxylated to the corresponding unsaturated alcohols[116]. The reactivity was shown to be maximal for octadiene (leadingto 0.3 to 0.4 g of diepoxyoctaneper liter) and falls off rapidly as the carbon chain is shortened, but decreases only slightly as the chain is lengthened. In a further study, it was shown that a very efficient conversion of 1,7-octadiene to 7,8-epoxy-l-octeneand 1,2-7,8-diepoxyoctane could be obtained by incorporating a high concentration of cyclohexane into the conventional fermentation medium. Thus, a 90% yield of product was achieved within 72 h, instead of a 18.5 % yield in the absence of cyclohexane, when a 20% (v/v) amount of cyclohexane was used. Clearly, this is an early example of the use of organic solvents applied to microbial transformations l1”1. A similar result was obtained later on using the 1-octene substrate itself as the organic phase (20% v/v), leading to comparable results (70% ee) [118]. Interestingly it also has been shown in the course of this work that, when n-hexadecane (which is not metabolized by the cells) is used as a solvent, racemic epoxide is enantioselectively degraded by the “a-hydroxylation” enzymatic system of P. oleovorans, leading to an enrichment in (S)-1,2-epoxyoctane. Further work by Wynberg and coworkers was aimed at even increasing the yield of 1,2-epoxyoctaneusing an optimized two-phase system and a cell renewal proced~re‘’~’]. Thus, yields up to 150 mg 1,2-epoxyoctaneper mL I-octene and up to 20-25 mg 1,2-epoxyoctaneper mL culture was obtained. Some other substrates were tested in this optimized system. Of these, 1-decenewas converted into (R)-1,2-epoxydecane (GO % 0. p.), while allylbenzene was converted to the corresponding epoxide. However, no effort was made to determine the absolute configuration and the optical purity of this product.

16.1 Oxygenation ofC-H and C=C Bonds

lM

I

H

Chemical

R. equi NClB 12035

95.4

f! putida NClB 9571

98

/? oleovorans AT CC 29347

98.4

f! aeruginosa NClB 8704

98.8

I

1087

R = CH2CH20CH3: Metropolol R = CH2CONH2 : Atenolol (M: f! oleovorans 0.p. = 97%) Figure 16.1-21. Microbiological epoxidation as a way t o optically

pure fl-blocker drugs.

All these results led to an interesting application for asymmetric organic synthesis. Thus, P. oleovorans has been used, among some other microorganisms, for stereospecific epoxidation of some arylallylethers into (+)-arylglycidylethers (Fig. 16.121). These intermediates were chemically converted into (S)-(-)-3-substituted-1-alkylamino-2-propanols, which are the physiologically active components of the padrenergic receptor blocking drugs. This method has been used to synthesize (S)(-)-Metoprolo1 and (S)-(-)-Atenolo1with enantiomeric purities of 95.4-98.8 % and 97 % respectively['201.These applications are of great industrial interest, since it has been shown that (S)-(-)-Metoprolo1is 270-380 times more active than its antipode[121]. Microorganisms screened for epoxidation activity were selected from bacteria belonging to the genera Rhodococcus, Mycobacterium, Nocardia and Pseudomonas. Species of Pseudomonas gave the best activities, but there were variations between the individual members, and P. oleovorans was the most active organism. The activity was further enhanced by carrying out the transformation in the presence of a cosubstrate such as glucose. This pioneering work on microbial expoxidationof straight-chain terminal olefins has triggered several further studies aimed at preparing enantiopure epoxides via biotransformations. Thus, a number of alkene-utilizing microorganisms have been described in the literature. In the context of aliphatic substrates these efforts have been developed essentially along two lines: epoxidation of long-chain olefins and epoxidation of short (CI-C,) chain compounds. Thus, for instance, it was shown that Corynebacteriurn equi ( I F 0 3730) grown on n-octane is able to oxidize 1-hexadeceneto give the corresponding optically pure (R)-(+)-epoxide(41% yield based on consumed substrate) 123J. This strain also assimilated other terminal olefins and produced

1088

I

IG Oxidation Reactions

the corresponding epoxides from substrates which have a carbon chain longer than fourteen, although in very low yields (less than 1%). Production of 7,8-epoxy1-octenefrom 1,7-octadieneby non-growing Pseudomonas putida species using twophase transformation has also been achieved [1241. Similarly, a gaseous hydrocarbonassimilating microorganism Nocardia c o r a l h a B-276 grown on I-alkenes (C3,C4 and c13-c18) was described as being able to produce the corresponding 1,2-epoxyalkanes. One of the products, 1,2-epoxytetradecane,was shown to be optically active. Glucose-grown cells could also transform styrene and C2.C18 1-alkenes to their epoxyalkanes[12'1. Similarly, production of epoxides from c&10 I-alkenes and styrene was shown to be enhanced by using n-hexadecane as an additional solvent, while this led to a decreased rate for epoxidation of longer chain l-alkenes1126]. Epoxidation of unsaturated fatty acids such as palmitoleic acid by Bacillus rnegateriurn has also been reported [1271. Here again, experiments indicated that epoxidation and hydroxylation were catalyzed by the same soluble cytochrome P450dependent enzymatic system. 16.1.5.2 Short-Chain Alkenes

Short-chain alkenes are another type of substrates which have been studied for microbiological epoxidation during the last thirty years. In this context, an extensive study has been conducted by De Bont and coworkers in order to prepare epoxides from gaseous olefins. Thus, a Mycobacteriurn sp. (E 20) was isolated from soil and shown to excrete ethylene oxide when grown on ethylene['28, 1291 . Stud'ies carried out using ' * 0 2 showed that a monooxygenase was involved in these epoxidations, as proved by incorporation of only one l80into the product. Another Mycobacterium (Py 1) was also shown to achieve this reaction. Experiments were performed in a gassolid reactor to prevent accumulation of the toxic ethylene oxide in the immediate vicinity of the biocatalyst[1301. An experimental set-up, allowing for automatic gas chromatography analysis of circulation gas in a batch-reactor system, was also described allowing on-line monitoring of the microbial oxidation of the gaseous alkenes propene and 1-butene (Fig. 16.1-22)[1311. Optimization was achieved by studying the influence of various organic solvents on the retention of immobilized cell a~tivityIl~~1. High activity retention was favored by a low polarity in combination with a high molecular weight. Using chiral gas chromatography (at that time recently described by Schurig and Biirkle [1331), eleven strains of alkene-utilizingbacteria were screened with respect to the stereospecific epoxidation of propene, 1-butene and 3-chloro-1-propene. The results obtained showed that seven of these bacteria strongly resembled each other, in that they all produced 1,2-epoxypropane and 1,2-epoxybutanemainly in the (R)-form(93 and 85 % ee respectively).Several of these thus strains were also able to epoxidize stereoselectively l-chloro-2,3-epoxypropane, leading to the synthetically very useful (S)-epichlorohydrin (ee > 95 %). Stereoselective epoxidation of 4-bromo-1-butene and of 3-buten-1-01was similarly studied using three strains. The results showed that the epoxides were again obtained predominantly in the (R)-formbut that their enantiomeric purity depended on both

16.1 Oxygenationof C-H and C=C Bonds Figure 16.1-22. epoxidation.

X = Br, CI

M: Mycobacteria

Nocardia corallina

n=34

Short-chain alkene

&O CnH2n+1

e.e. = 76-90%

(-)-fosfomycin (90%)

the strain used and on the substrate studied(87].Inactivation of the alkene oxidation enzymatic system by the produced epoxide was also investigatedin view of setting up a biotechnological procedure for producing these epoxides Modeling the effects of mass transfer on the kinetics of propene epoxidation was also achieved by the same authors[135,13'1, and they showed that product inhibition can be reduced by absorbing the epoxide in the gas phase in cold di-n-octylphthalate['37]. In addition to the Mycobacterium species, several other strains have been reported to achieve epoxidation of olefins. Thus, three distinct types of methane-grown methylotrophic bacteria (Methylosinus trichosporium, Methylobacterium capsulatus and Methylobacterium organophilum) were shown by Hou and to be able to oxidize terminal C2 to Cq n-alkenes to their corresponding 1,2-epoxides,which accumulated extracellularly. Results from inhibition studies indicated, as in the case of the previously discussed o-hydroxylation system of P. oleovorans, that the same monooxygenase enzyme was responsible for the hydroxylation of methane and the epoxidation of alkenes. Further work achieved by the same group showed that whole cells of Methylosinus sp. CRL 31, immobilized by adsorption on glass beads, were able to convert propylene to propylene oxide for several hours until the reduced NAD cofactor was depleted. This could be regenerated by periodic addition of methanol. These authors also observed that attempts to immobilize the cells by covalent binding or entrapment in polyacrylamide gel led to complete loss of propylene

1090

I

76 Oxidation Reactions

epoxidation activity(139]. However, no mention is made in this work of the enantiomeric purities of the obtained epoxides. Further studies carried out by Subramanian1'401 revealed that these were nearly racemic compounds, and also that the major problem of these biotransformations was again product (epoxide)inhibition. More information about the reaction mechanism of the epoxidation achieved with whole-cell M. trichosporium was gained by Okura and coworkers1l4l],who showed that the configuration of the double bond was retained during the epoxidation of cis2-butene. This result was further confirmed by studies of the epoxidation of 1,2-deuterated-cis-propene. A concerted insertion of oxygen was postulated to account for this result Oxidation of propylene to propylene oxide by Methylococcus capsulatus (Bath) was studied in order to optimize the biotransfomation for a possible industrial production. However, the high rates obtained could only be sustained for 3-4 min before loss of biocatalytic activity 0ccurred[l~~1. Similar results were obtained by Wyngard and in the course of a study aimed at exploring how immobilization of the whole cells on solid supports would influence the rate and duration of the epoxidation of propylene by the strain Nocardia corallina B-27G initially isolated by Furuhashi and coworkers [I2'. 12'] . Here again the results suggested that entrapment in a hydrophobic matrix might be a favorable system, but that loss of activity was quite rapid with time. The same Nocardia strain has been shown to be able to epoxidize branched chain terminal olefins in an asymmetric manner leading to (R)-epoxidesshowing optical purities of 7G-90% depending on the chain length. These epoxides were used as chirons for further synthesis of prostaglandin a-chains. The same strain was shown by these authors to be also able to epoxidize trifluoromethylethylene (75% ee) [1451. Some newly isolated Xanthobacter sp. were recently shown to be able to accumulate 1,2-epoxyethanefrom ethene or, when grown on propene, to accumulate 2,3-epoxybutane from cis- or trans-2-butene but with apparently low yields [14'1. Similarly, Rhodococcus rhodochrous, a propane-oxidizingstrain, was shown to produce 1,2-epoxyalkanes from short-chain terminal alkenes. Interestingly, its oxygenase enzyme appeared to be capable of tolerating high levels of product without inhibiti~n"~']. Finally, a very useful and industrially interesting epoxidation which deserves special attention is the stereospecificepoxidation of cis-propenylphosphonate.Eighteen species of Penicillium, one of Oidium and one of Paecilomyces were found to effect this reaction, which affords directly (-)-fosfomycin,a broad-spectrum antibiotic. Using the strain Penicillium spinulosum MB 2843 at optimum culture conditions, a 90% efficiency (based on olefin charged, 0.5 g L-l) was obtained after G days, leading to a product claimed to be optically pure [1481. 16.1.5.3 Terpenes

Besides the extensive studies aimed at preparing optically active epoxides starting from short or long straight-chain alkenes, another area of investigation has been the microbiological epoxidation of various natural substrates, essentially in the terpene and steroid area. Interestingly enough, it appears that terminal olefins (and only

76.7 Oxygenation ofC-H and C=C Bonds

"

9H

I

1091

i +

Methyl geranate

w1-m e.e

S.

P

albus

HO Linalool

- 100%

OH

+(&

HO D. gossypina

7

bH (-)-Linalool

Figure 16.1-23.

(SR,GR)-Linalool oxide

Some examples of olefinic terpene epoxidation.

these) are epoxidized almost exclusively by bacteria, and lead to accumulation of the corresponding epoxide in the culture. On the other hand, more substituted double bonds are often preferentially oxidized by higher organisms like fungi. The product is generally the corresponding vicinal diol arising from further metabolism (hydrolysis) of the primarily formed epoxide. Numerous publications describe microbial transformations of various terpenes [14', lS0l. However, there are few cases of an accumulation of intermediates in sufficient amounts for further use in synthesis [w. One of the first examples of such a transformation has been described by Marumo and coworkers (Fig. 16.1-23)I1'*l. Their investigations, aimed at preparing optically active insect juvenile hormone, showed that methylgeranate was metabolized by the fungus Colletotrichurn nicotianae, leading to 19.6% of S(-)-methyl-6,7-epoxygeranate and to 15.6% of R(+)methyl-6,7-dihydroxygeranate after 9 h incubation. Longer incubation times (24 h) produced only the optically pure glycol with an isolated yield as high as 85%, showing that the first epoxidation step had to be stereospecific. Unfortunately, this analpcal study was not pursued on a preparative scale, and no accurate results concerning the stereochemical and kinetical aspects of these interesting biotransformations have been described. A similar microbial oxidation of the isoprene double bond has been studied by

1092

I Veschambre and coworkers starting from l i n a l o 0 1 [ ~Thus ~ ~ ~ . Streptomyces albus, a I6 Oxidation Reactions

strain which synthesized nigericine, transforms each enantiomer of linalool, as well as the racemic compound, into a mixture (10-20% yield) of two diastereoisomeric linalool oxides. In this case, the epoxide formed primarily is trapped by an intramolecular cyclization. Based on the reported proportions of these products, one can deduce that the ee of the formed epoxide was about 35 %. Further work achieved using several other microorganisms showed that Beauveria sulfirescens gave similar yields (15-20 % analytical) of an equimolar mixture of linalool oxides [lS4].Botyris cinerea, a fungus which participates in the formation of flavors in sweet wines, was also checked for linalool biotransformation. This led to several metabolites including linalool oxides, presumably arising from prior epoxidation of the olefinic bond [1551. Interestingly, these products were also detected in the Carica papaya fruit flavor, together with the diastereoisomeric epoxides [1s61. It was also observed by Abraham and coworkers that (-)-linalool is processed by Diplodia gossypina exclusively to a mixture of trans-(3R,G R)-linalool oxide and to the corresponding tetrahydropyran. These were proposed to arise by intramolecular cyclization of the intermediate G(S)epoxide. Some other similar substrates have been studied in the course of this study, but they generally led to low yield mixtures of products. Comparable results were obtained from linalool using the strain Streptomyces cinnamonensis [15*l. Similar transformations were observed starting from 2-methyl-2-heptene-6-one[1s91. However, because of the number of metabolites formed and the low yields obtained, these biotransformations cannot be usefully employed for organic synthesis. Myrcene and trans-nerolidol were also shown by Abraham and Stumpf to be transformed by two fungi (Diplodia gossypina and Corynespora cassiicola respectively) into a mixture of several products including vicinal diols arising from oxidation of the isoprenyl double bond. These were shown to be further degraded, presumably via an acyloin-splittingmechanism [l6O1. During the course of the fermentation, the diol occurred at first in the culture medium followed by the nordiols and the trialcohols. So, the formation of these compounds from diols seemed to be very likely. Some other related substrates were also studied in the same context, and it was shown that both strains revealed a pronounced and almost opposite substrate selectivity. Much more impressive is the result obtained by the same groupL'"1,who conducted a broad screen of 800 various microorganisms using both the ( S )(-)- and the (R)(+)-limoneneenantiomers as a starting substrate, as well as some other terpenes which were tested with the best suited strains (Fig. 16.1-24).The most interesting results were observed with Diplodia gossypina (ATCC 1093G), which afforded 380 mg of a diol which was found to be the ( l R , 2R, 4S)-8-p-menthen1.2-diol from 1 g of (S)(-)-limonene. Similarly, Corynespora cassiicola (DSM 62474) from 1.8g of a-terpene. was described to yield l.lg of (lR, 2R)-3-p-menthen-l,2-diol (R)(+)-limonenewas shown to afford (lS, 2S, 4R)-p-8-menthene-1,2-diol. Because of the interest of these products in flavor chemistry, the preparative-scale transformation of this enantiomer by the fungus Diplodia gosvpina has been undertaken: thus 1300 g were transformed, yielding 900 g of the (IS, 2s) diol showing high optical purity[1621. Interestingly, these strains convert the substrates fast with only negligible amounts of side products. Also, it is noteworthy that the

2 y2 gossypina Diplodia

(S)-(-)-Limonene

\

-1

-

,

a-Terpinene Figure 16.1-24.

30%

( I R , 2R, 4S)-8-p-Menthene-l,2-diol ,,,\OH

Diplodia gossypina

(R)-(+)-Limonene \

1 6 1 Oxygenation of C-H and C=C Bonds

corynespora

cassiicola

0.6%

55%

OH ,,,,\OH

/

49%

+

f' /

2%

pJ OH

+

/

1Yo

Stereoselective oxidation of rnonocyclic terpenes.

obtained diols are almost exclusively of trans configuration. No indication is provided concerning the determination and the values of the obtained products' optical purities. It was suggested that these trans-diols were formed via an intermediate epoxide, which could be further cleaved enzymatically to the obtained diols. Surprisingly, both these microorganisms were shown not to attack 3,3,5,5-tetramethyllimonene[1631.However, geranylacetone, nerylacetone, trans-nerolidol, ciswere transformed by these nerolidol, farnesol and 2,5-dimethyl-1,3-hexadiene strains to the corresponding glycols in yields of up to 70%[1643 16'1 and interesting optical purities of up to 98%. Using (+)-trans-nerolidolas a substrate, the strain Nocardia a k a DSM 43 130 was shown to be lacking an epoxide hydrolase, thus leading to a 27 % yield of the corresponding (S)-epoxidewhich accumulates in the culture medium [lS71. Also, the ability of the monensin-producing organism Streptomyces cinnamonensis to convert the cis and trans isomers of nerolidol has been investigated['58].However, here again this led to a low-yield mixture of several products. Much more useful in that sense are the results obtained by Furstoss and coworkers in the course of their study of biooxygenation of geraniol derivatives (Fig. 16.1-25). Indeed, it has been described in a first paper that, if the N-phenylcarbamate of geraniol is used instead of geraniol itself, its transformation by the fungus Aspergillus niger leads to a 49 % isolated yield of the 6,7-dihydroxylatedproduct. Moreover, this diol proved to be of (6s) absolute configuration and was shown to possess an This diol, which is a very versatile enantiomeric excess of about 95%[76, substrate for further organic synthesis, can thus be obtained without problem in

I

1093

1094

I

76 Oxidation Reactions

A. niger 18

02

R = CONHPh

(6R) ee = 95% Figure 16.1-25.

Stereoselective pH-dependent oxidation of geraniol N-phenyl

carbamate.

gram-scale quantities (1g substrate treated for 36 h in 1 L culture afforded 550 mg pure diol). Further work aimed at exploring the influence of the culture conditions showed that a unique stereochemical control could be achieved simply by modulating the pH of the medium. Thus, although when the culture was at pH 2 the diol of (S)-configurationwas obtained, at pH 6-7 the diol of opposite ( R )absolute configuration was isolated in similar yields and with an ee again as high as 95%. This interestingly showed that the fungus A. niger not only is able to convert the substrate across the pH 2-7 range, but that the (6s)-epoxide must be the primarily formed metabolite. This can then be further hydrolyzed in acidic medium (following the classical acid-catalysismechanism) to afford the (6s)-diolor, at pH 6, be hydrolyzed enzymatically to the (6R)-diol by attack on the less substituted oxirane carbon Experiments conducted in the presence of " 0 confirmed this hypothesis. When the incubation was carried out at pH 2, the distribution of the " 0 label in the obtained diol was 95 % on C-6 and 5% on C-7.This ratio was inverted at pH 7. These results show clearly that, whatever the pH, molecular oxygen is involved in these oxygenations but only one labeled oxygen atom is incorporated into the diol, leading to an epoxide which is differently hydrolyzed, depending on the pH of the medium. Very interestingly as far as organic synthesis is concerned, these biooxygenations can be conveniently performed on a scale of several grams (5 g), thus allowing easy preparation of either enantiopure diol. These can be conveniently used as "chirons" for the synthesis of various natural or non-natural products. For instance they can be cyclized to the optically pure linalool oxides [16'] or the corresponding tetrahydropyranols [1691. Biooxygenation of some other similar compounds, i. e. 7-geranyloxycoumarin, citronellyl N-phenylcarbamate and sulcatol N-phenylcarbamate were studied . The reaction was shown to be operative in all these cases, (Fig. 1G.1-26)['70-'721 leading, for instance to either enantiomer of marmin (a member of the umbelliferone family). Moreover, this result opens the way to an easy preparation of either

:I ,<\OH

7 6 7 Oxygenation of C-H and C=C Bonds I1095

A. niger

A.niger PH 6

PH 2 60%

43%

OH

OH

(-)-marmine

7-geranyloxycournarin

(+)-marmine

0

0

>,

+

1.: p ,:::o

(G‘R),i”epoxyaurapten

*

cR

(G’S),i”epoxyaurapten

AP? 85%

(3RS) ee=90%

$

A.niger

PH 2 73%

Sulcatyl N-phenylcarbamate

A60% P E .

(3R)-citronellyl N-phenylcarbamate

~

T

R

\,\OH

(2R 5s)

+

(3R96R) ee=92%

$R

,\\OH

P S , 55)

Figure 16.1-26. Application of the pH-dependent oxidation of geranyl derivatives to the synthesis of some natural products.

enantiomer of 6‘,7’-epoxyauraptenand of 3’,6‘-epoxyaurapten,both these compounds being natural products isolated from various sources. Similar results were obtained from both commercially available citronellol enantiomers, leading to the corresponding diols showing ee’s as high as 90 and 92 %. Bioconversions conducted at pH 2 on racemic sulcatol N-phenylcarbamate led to a 73 % yield of a 1/1mixture ofthe two expected diastereoisomeric diols, which can be readily separated by flash chromatography. They both show ee’s > 95 %, indicating that the first (epoxidation) step again occurred in a highly stereospecific manner. Interestingly in this case, it was also possible to avoid hydrolysis of the intermediate epoxide by changing the preculture conditions and performing the reaction at neutral pH. This intermediate can thus be obtained directly with high enantiomeric purity. Using this chiron allows the four-step synthesis of optically pure pityol, a

1096

I

IG

Oxidation Reactions

(25) Figure 16.1-27. steps.

(2S,5S)

(2R,SS)-pityol

A four-step synthesis o f (ZR,SS)-pityol using microbiologically mediated

male-specific attractant of the bark beetle Pityophtorus pityographus. Thus, prochiral 6-methyl-hept-5-en-2-onewas reduced with baker’s yeast to the corresponding alcohol (60% yield, 98.5% ee). This was converted to its N-phenylcarbamate,which was subsequently subjected to epoxidation using A. niger, thus affording a 50% preparative yield of the corresponding enantiopure epoxide. In a final step, treatment of the epoxycarbarnate with an alcoholic NaOH solution led to the natural (2RS.S)pityol(7.5% overall yield, 100% ee, 98% de) (Fig. 16.1-27). 16.1 S.4

Cyclic Sesquiterpenes

Various cyclic sesquiterpenes have also been studied in order to explore the possibility of achieving their microbiological transformations. Very often these were shown to lead to epoxidation processes when one (or several) double bonds were present in the starting substrate (Fig. 16.1-28). Thus germacrone, which is thought to be the precursor of a variety of bicarbocyclic sesquiterpenoids, was shown to be transformed by the fungus Cunninghamella blakesleena. This led primarily to regio- and stereoselective epoxidation of one of the intracyclic double bonds of this prochiral triene, thus affording two epoxides. The third product isolated from this experiment was due to subsequent epoxidation of the remaining intracyclic double bond. Interestingly, the exocyclic olefinic bond conjugated to the carbonyl function appeared resistant to oxidation [1731, Valencene, another olefinic sesquiterpene, has been studied in the same context using microorganisms isolated from It was observed that these biotransformations led in reasonable yields to a mixture of three main metabolites, including an epoxide and nootkatone, an interesting flavoring compound. The microbial transformation of humulene, a substrate showing a structure similar to that of germacrone, was studied by Abraham and Stumpf using a screen of about 300 strains[175]. This led the authors to select the fungi Diplodia gossypina and Chaetonium cochlioides for preparative scale experiments. It was thus observed that the main reaction path starts with the epoxidation of the 1,2-doublebond, as shown by direct biotransformation of this monoepoxide obtained by chemical synthesis. This is then further oxidized to yield a multitude of products including diepoxides and hydroxy-epoxides (Fig. 16.1-28). Comparable results were obtained from caryophyllene, a compound similar to humulene. Again, the biotransformation of this substrate with cultures of Chaeton-

76.7 Oxygenation of C-H and C=C Bonds

p+C.b

Germacrone

\

my-

+

Soil

- Valencene

(D. gossypina C.cochlioides

Nookatone 12%

............

* mixture

D. gossypina

C. cochlioides

mixture

Carophyllene Figure 16.1-28.

Epoxidation steps i n the course of sesquiterpene biotransforrnations

i u m cochlioides as well as of Diplodia gossypina give a broad spectrum of products, resulting from an initial epoxidation of the 1-2 double bond followed by additional epoxidation or hydroxylation processes (Fig. 16.1-28)[176, 1771. 16.1.6 Conclusions, Current and Future Trends

This review has illustrated the very broad range of biohydroxylations and epoxidations that can be achieved using monoxygenase enzymes. In fact, one can propose

I

1097

1098

l that almost all organic compounds are potential substrates for these enzymes. Since 16 Oxidation Reactions

each substrate can lead to many different oxidized products, the range of compounds that can be generated is clearly enormous. Finding new enzymes with novel substrate specificities and selectivities of reaction has in the past been achieved by screening organisms and substrates and has very much been down to good luck. Current and future work is focused on finding methods to make this process faster and more rational and predictable. This is now possible because of new technologies in genetics, molecular biology and structural biology, of which a few highlights are discussed below. More and more P450 monoxygenases have been sequenced and cloned into heterologous expression systems. This can have the advantage of higher turnover yields because of higher expression of the enzyme in the host or because higher cell mass can be obtained when using easy growing organisms such as E. coli as hosts[’78,1791. Heterologous expression can also overcome problems of loss of product because of further metabolic degradation[l8’I as in the case of the alk gene of P oleovorans. Such expression systems also allow the facile generation of chimeric enzymes and mutants with more desirable biocatalytic properties, such as increased activity towards a particular substrate[lS1,182]. Some of the popular organisms for biohydroxylationssuch as Beauvaria bassiana also might contain several endogenous P450 enzymes that can interfer with selectivity of one enzyme and make predictions of reactions very difficult [1831. The rapid emergence of whole genome sequences has made a major impact on the study of P450 monooxygenases [lS41, since they are often easily identifiable by small conserved consensus sequences, in particular around the heme binding site. We now know that Mycobacterium tuberculosis contains probably twenty different P450 monoxygenases; Bacillus subtilis contains seven. The a priori prediction of substrate specificity and selectivity from gene sequence is at the moment impossible and presents a great challenge to the researcher. However, there has been some success in prediction of substrate specificity by “in silico screening” based on available threedimensional structures of P450-monooxygenases[1851. Thus, substrate docking algorithms were used to predict substrate suitability for P450cam and its L244A mutant from a library of commercially available compounds. The most practical way of using P450-based biocatalysts is still in whole-cell systems, because of cofactor requirements and problems with enzyme stability. However, some P450 monooxygenases, such as the P450cam, can be isolated in sufficient quantities and reconstituted for cell-free preparative scale biotransformations[1821.This might be particularly useful for substrates that cannot penetrate cell walls, are toxic to the organism or are unstable in the organism. One solution for overcoming co-factor requirements might be the use of electrochemical methods, and is has indeed been shown that P450cam can be immobilized on an electrode and can take up electrons from the electrode[186]. Another novel area of intense research is the application of mutagenesis (random and directed) to obtain desired changes in substrate specificity. Thus P45Ocam, which is highly selective for camphor and closely related analogs, was subjected to site-specific mutagenesis, changing the tyrosine in position 96 to a phenylala-

1G. 1 Oxygenation of C-H and C=C Bonds

I which resulted in about a 20-fold increase in the reactivity towards naphthalene. The P450 monooxygenase was independently subjected to random mutagenesis by Arnold and co-workers[18’1, and mutants were screened for increased activity towards naphthalene. Similar improvements to those observed by specific mutagensis were obtained. However, interestingly, the mutations that were found to be responsible for improved activity were not at position 96, but were distant from the active site of the enzyme. Such a “directed evolution” approach has great promise in quickly generating desired biohydroxylationcatalysts, provided that a suitable screening system for the product can be found. The method has also been recently used by the same group on P450BM3 [188]. In conclusion, the application of biocatalysts in biohydroxylations and epoxidations is rapidly expanding in terms of practicality, substrate range and selectivity. A vast diversity of P450 genes is generated by genomics programmes and mutagenesis. Methods for screening such oxidation catalysts are becoming more rapid, and one can forsee a future where designer biooxidation catalysts, tailored for a specific substrate and even for selectivity of reaction, can be generated within short time spans using a combination of rational and screening methods. 16.1.7

Cis Hydroxylation of Aromatic Double Bonds 16.1.7.1

Introduction

The microbial dioxygenation of aromatic compounds 1 has been known for over thirty years through the pioneering efforts of D. Gibson et al., who characterized the In lower organisms, the chiral cis metabolic pathway of toluene degradati~n[’~’]. glycol intermediates 2 are rapidly oxidized by dihydrodiol dehydrogenase, involving rearomatisation to the diol3, which is further oxidized by ring cleavage dioxygenase to give dicarboxylic acid 4, which can be channeled into the organism’s normal metabolic pathways (Scheme 16.1-1)[190-1921. dioxygenase 02

aoH

R

R 1

~ ‘ C O ~ H /C02H

R

4

NADP+ \I

OH

2

dihydrodiol dehydrogenase

ring cleavage dioxygenase

Oxidative degradation of aromatic compounds by microorganisms.

Scheme 16.1-1.

OH

3

1099

1100

I

7 G Oxidation Reactions

The use of certain strains of Pseudomonas putida, most notably the mutant 39 D with blocked dehydrogenase allows accumulation of the chiral glycols in the fermentation medium associated with high stereospecificity while the substrate tolerance remains high with respect to ring substituents. The enzymology of dioxygenases has been and refinement of the mechanistic details of the dioxygenases c o n t i n ~ e s [ ~ 'but ~ J , only those enzymes and applications of relevance to the preparative biotransformations will be considered here. 16.1.7.2

Preparation o f cis Dihydrodiols

An impressive number of substituted aromatic compounds 5 have been converted by mutant strains of Pseudomonas putida into the corresponding chiral cis glycols G with often excellent stereosele~tivity[~~~~ 1961 . The remarkable substrate range and selectivity of this dioxygenase system for the aromatic ring have been demonstrated by the conversion of a series of substituted benzenes and of alkenyl benzenes with the side chain double bond being left intact (Scheme 16.1.2) 191 ' . An analogous product was obtained from para-fluorotoluene, but the dihydrodiols from parachloro- and para-bromotoluenes were found to be racemic [1991. Unlike the substrates shown in Scheme 16.1-2,benzoic acid, toluic acid, and their halogenated analogs, for example 7, undergo enzymatic dioxygenation by Alcaligenes eutrophus B 9 and two strains of Pseudomonas, for example JT 103, mainly at the 1,2-position (Scheme 16.1-3)[200]. However, with other strains of Pseudomonas putida, for example JT 106, enantiospecific cis 2,3-dihydroxylationis possible, too [201]. The structure of the substrates is not necessarily restricted to monocyclic aromatic compounds such as those shown in Scheme 16.1-2. The dioxygenase activity of Pseudomonas putida and Belj'erinckia species has been used exclusively for the synthesis of cis dihydrodiols from polycyclic [202] and heterocyclic[2031 derivatives, Such products have been obtained from naphthalene, anthracene, phenanthrene, benz[a]pyrene, benz[a]anthracene, and methylsubstituted benz[a]anthracenes, and

?

R Pseudomonas putida

JTI 03

6

5

OH

R = H, Me, Et, nPr, nBu, BU, EtO, nPrO, halogen, CF,, Ph, PhCH, PhCO, CH(OH)CH,, COCH,, CHz=CH, CHz=CHCHz, CH$H=CH CH3C=CHCH3, HC-C, CF,, CN, COP!+ SiMe,

HOZC,

Scheme 16.1-2. Synthesis of cis diols by Pseudomonas putida.

OH

Pseudomonas putida JTI 03 F

F

7

F

Scheme 16.1-3. Cis-hydroxylation of aromatic carboxylic acids by Pseudomonas putida JT 103.

a \

1 G. 7 Oxygenation of C-H and C=C Bonds Scheme 16.1-4.

I

'lol

Beijerinckia 0 8/36

monas putida.

x=o.s

Pseudomonas putida

33%

a

Ho""

03

Pseudomonas putida \

\

N

47%

9

HON Q OH

Pseudomonas putida

55% 10

Pseudomonas putida

45%

many of the enzymes responsible have been identified and ~haracterized[~'~]. Benz[a]anthracene,for example, is converted to three cis dihydrodiol regioisomers by Beijeierinckia B 8/36['04]. This organism has also been reported to produce dihydrodiols from dibenzofuran [2051 and dibenzothiophene (Scheme 16.1-4)['06]. The ability of a Pseudomonas putida mutant to metabolize heteroaromatic compounds is demonstrated by the bioconversion of quinoline 8, isoquinoline 9, quinazoline 10, and quinoxaline 11 L2O7]. Attack occurred exclusively in the carbocyclic ring (Scheme 16.1-4). The impact of the genetic revolution has been greater in the area of dioxygenasecatalyzed reactions than in many other areas of bioconversion, largely because of the bacterial origin of the enzymes concerned. The bacterial oxidation of aromatic double bonds to cis diols in an enantiospecific manner leads to highly interesting synthons for organic chemistry. For example, the diene may be subjected to DielsAlder reactions, as well as Michael-type addition reactions. Alternatively, oxidative cleavage of the cyclohexadienering leads to open chain products, which further react to yield cyclopentanoids. The large synthetic potential of chiral cis glycols is illustrated in Scheme 16.1-5. T. Hudlicky et al. efficiently synthesized the prostaglandin PGE2 12 through an the vinyl oxidative ring cleavage of the methyl-substituted diol G (R = CHS)[208],

1102

I

I6 Oxidation Reactions

OH

HO

21

1‘

HO“’

//

R=CH3

\

.

.

-OH

15

OH 19

16

HO

18

.

HO’

Et6 17

Scheme 16.1-5.

Syntheses of natural products from substituted cyclohexadienediols.

derivative G (R = CH = CHI) was used for the construction of the plant metabolite (-)-zeylena13r209), and the chloro-substituted diol for the synthesis of the alkaloid trihydroxyheliotridane 14[2101 and the carbohydrates L-ribonolactone 15i 2 1 1 ] and Derythrose 16[’12].Hudlicky et al. also prepared the sesquiterpene specionin 17L2l3],an antifeedant to the spruce budworm, and the narcissus alkaloid lycoricidine 18i214]in only nine steps. D-myo-inositol ZO[217],conduritol Biologically active polyols like pinitol 19L2”, CF218]and conduritol E r2”)1 were obtained from diol6 in both enantiomeric forms in only a few steps using this approach. Futhermore C. R. Johnson et al. synthesized (-)-shikimic acid 21, the biosynthetic precursor of the benzene moiety of aromatic amino acids [220]. In the case of the cyclohexadienediols 6, the current development promises to complement the traditional and rather arduous use of carbohydrates as starting materials from the chiral pool. The popularity of diol-based methods will, therefore, be directly proportional to their ready commercial availability and to the operational

References

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I

1103

1104

I

7G Oxidation Reactions

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219

16.2 Oxidation o f Alcohols

Andreas Schmid, Frank Hollmann, and Bruno Bijhler 16.2.1

Introduction

The enzymatic oxidation of alcohols is catalyzed by different oxidoreductases. Here, examples of dehydrogenases, oxidases, and peroxidases are discussed. Single enzymes were selected based on representative or demanding reactions that are catalyzed, or because of interesting reaction engineering solutions applied. Reactions catalyzed by whole microbial cells are described in a separate chapter. A focus is put on presenting or introducing enzyme catalysts and their substrate spectra in order to give the reader a basis for designing his or her own, new reactions with sterically or electronically similar compounds or with such compounds which are compatible with a certain reaction mechanism. Biocatalysis usually exploits advantageous features of enzymes such as chemoselectivity, regioselectivity, enantioselecivity and substrate spectrum of a certain broadness as depicted in Fig. 16.2-1. These points are addressed in examples in the following chapters. 16.2.2

Dehydrogenasesas Catalysts 16.2.2.1

Regeneration of Oxidized NicotinamideCoenzymes

Regeneration of NAD(P)' from NAD(P)H is a redox reaction involving the transfer of two electrons and a proton (successivelyor at once as hydride ion H') to a suitable acceptor. Most commonly these acceptors are carbonyl functions, molecular oxygen or the anode. Apart from a few exceptions the direct hydride transfer is slow or disadvantageous so that catalytic procedures have to be applied. Here we selected representative examples to give an overview. Excellent review articles are available, tooil-3.

101

16.2 Oxidation ofAlcohols

A H

R-0

t

G

B

PH -.A-

C .COOH

D E

Figure 16.2-1. Enzyme-catalyzed oxidations o f alcohols. Reactions are grouped according t o the feature mainly exploited in the preparative application. A-C: Chemoselectivity (e. g. Sects. 16.2.2.3, 16.2.2.6, and 16.2.2.11); C, D: Regioselectivity (e.g. Sects. 16.2.2.9, 16.2.2.10, and

16.2.3.4); E, F: Enantioselectivity (e. g. Sects. 16.2.5.2 and 16.2.6.4); C: Non-natural substrates (e.g. Sect. 16.2.2.3); H: Complex structures from simple starting materials (e.g. Sect. 16.2.2.3).

16.2.2.2

Dehydrogenases as Regeneration Enzymes

Today, the utilization of a dehydrogenase-catalyzed reduction reaction is still the most widespread approach for the regeneration of oxidized NAD(P)+.Its principle is displayed in Fig. 16.2-2. Most commonly, alcohol dehydrogenase (E. C. 1.1.1.1) from yeast (YADH), horse liver (HLADH), or Themzoanaerobiurn brockii (TBADH) as well as glutamate dehydrogenase (E.C. 1.4.1.2.) or lactate dehydrogenase (E.C. 1.1.1.27) are used for NAD(P)+regeneration (Table 16.2-1).Thus, the reduction equivalents are transferred to an aldehyde or ketone as terminal electron acceptor yielding the corresponding alcohols. The drawbacks of this approach result from the necessity to use a second enzyme, whose optimal reaction conditions may differ significantly from those of the actual production enzyme, and the presence of cosubstrates and coproducts. Furthermore,

I

1109

1110

I

7 G Oxidation Reactions

NAD(P)+

red.

NAD(P)H

regeneration enzyme

OX.

Enzymatic regeneration of oxidized NAD(P)+.

Figure 16.2-2.

Comparison of commonly used dehydrogenases for NAD(P)' regeneration.

Table 16.2-1. Regeneration

Cosubstratel

Specific Activity

enzyme

coproduct

[U mg-'I

Stability

Coenzyme

E'o M vs.

NHE" [11

YADH TBADH Glutamate DH Lactate DH

Acetaldehyde/ ethanol Acetone/ isopropanol a-Ketoglutaratel glutamate Pymvate/lactate

NAD'

- 0.199

30-90

Low, sensitive to 0 2 Thermostable

NADP'

- 0.286

40

High

- 0.121

1000

High

NAD' and NADP' NAD'

300

- 0.185

a NHE: normal hydrogen electrode.

1 H. K. Chenault, G. M. Whitesides, Appl. Biochem. Biotech. 1987, 14, 147-197.

the thermodynamical driving force is low because the formal redox potential of the cosubstrate/coproduct couple is often close to that of the NADH/NAD' couple. Some of these problems can be addressed using the following regeneration concepts:

16.2.2.2.1

Enzyme-Coupled Regeneration

Since dehydrogenase catalysis is reversible, the production enzyme can be used to perform the regeneration reaction of NAD(P)+using a suitable cosubstrate as electron acceptor. In this case, the regeneration enzyme in Fig. 16.2-2 is identical with the production dehydrogenase. However, conversion rates in this set-up tend to be low because of a given reaction equilibrium, which requires an efficient method to withdraw the products and coproducts.

1 6.2.2.2.2

lntrasequential Regeneration

One elegant way of in situ product removal is to use the product of a first dehydrogenase reaction as substrate for a subsequent enzymatic reaction, thus recycling the oxidized nicotinamide coenzyme (Fig. 16.2-3). Various NAD(P)-dependent enzymes can be applied as regeneration enzymes in this cascade reaction.

16.2 Oxidation ofAlcohols

dehydrogenase

substrate

NAD(P)+

product

intermediate

I

regeneration enzyme lntrasequential regeneration of NAD(P)'. The strategy applied is the synthetic coupling of a dehydrogenase-catalyzed oxidation and a regeneration reaction yielding the final product and NAD(P) regeneration. Figure 16.2-3.

If the regeneration enzyme is a second dehydrogenase, an overall redoxisomerization takes place. But also monooxygenases are reported as regeneration enzymes thus yielding an overall double oxidation of the substrate (see Sect. 16.2.2.6.1). 16.2.2.3

Molecular Oxygen as Terminal Acceptor

The application of molecular oxygen as oxidant is favorable for several reasons. It is cheap and easily applicable. Furthermore, the high redox potentials of the 02/H20 or 0 2 / H 2 0 2couples (in acidic solution + 1.23 V and + 0.682 V, respectively) result in a strong thermodynamic driving force for the regeneration reaction. Since direct oxidation of NAD(P)H by molecular oxygen is very the electron transfer has to be accelerated via enzymatic or chemical techniques. NADH oxidases (NADH dehydrogenases, E. C. 1.6.99.x) from several organisms have been characterized in recent years[']. Two types of NADH oxidases can be distinguished, namely those reducing molecular oxygen to water and those performing the reduction to hydrogen peroxide. Interestingly, few examples are found in literature employing NADH oxidases for the regeneration of NAD', probably because of stability reasons. However, an NADH oxidase from Themus aquaticus was reported to be stable at 80 "C for at least 1 hL6],which might allow small scale applications. FMN reductase (NAD(P)H dehydrogenase (FMN), E. C. 1.6.8.1) catalyzes the transhydrogenation from NAD(P)H to FMN r71, yielding the oxidized nicotinamide coenzyme and FMNH2, which reacts spontaneously with molecular oxygen (Fig. 16.2-4). The reaction might be coupled to the catalase reaction in order to decrease the degree of enzyme inactivation over longer reaction times. Compared to the non-catalyzed hydride transfer from NAD(P)H to FMN 1'1, up to 1000-fold increases in the transhydrogenation rate are reported, which is not very high when applied synthetically. In this respect also the operational stability of FMN reductase has to be optimized. Besides the native substrate, cheaper alloxazine-based analogs are also accepted"].

I

1111

1112

I

76 Oxidation Reactions

Figure 16.2-4.

Transhydrogenation catalyzed by F M N reductase.

Among the chemical mediator systems especially o-pinones are capable of accepting the hydride equivalent from reduced nicotinamides. The oxidized mediators are regenerated by molecular oxygen. Since these mediators can also be recycled electrochemically,they are discussed in the following chapter. 16.2.2.4

Electrochemical Regeneration

A very elegant method to regenerate NAD(P)+from NAD(P)His to use the anode as terminal electron acceptor. The most common approaches are summarized in Figure 16.2-5.

Direct electrochemical N A D ( P J H oxidation (Fig. 16.2-5 A) The easiest way to oxidize NAD(P)H is to withdraw the excess electrons anodically. Although the formal potential of the NADH/NAD' couple is - 320 mV [- 324 mV for NADP] vs NHE["], overpotentials as large as 1 V are required to achieve significant oxidation rates at bare electrodes[", '*I. The number of enzymes, substrates, and

NAD(P)H

Figure 16.2-5. Electrochemical regeneration of NAD(P)'. A: direct anodic oxidation; 8:indirect electrochemical oxidation; C: diaphorase-accelerated indirect electrochemical oxidation.

IG.2 Oxidation ofAlcohols

products that can withstand this oxidizing power is limited. In addition, direct oxidation is often accompanied by electrode fouling, which is attributed to the formation of NAD dimers or stable adducts [12,131.

Indirect electrochemical NAD(P)H oxidation (Fig. 16.2-5 B,C) The high overpotentials needed for NAD(P)Hoxidation can be considerably lowered by the use of redox mediators. Organic compound (such as ortho- and purasubstituted quinones [I4lSl, diimines I2O1, and organic dyes 121-221) undergoing twoelectron transfer processes were found to be ideal fpr NAD(P)' regeneration. [23, 241 are probably the most potent Amongst these, l,lO-phenanthroline-5,6-diones mediators. Furthermore, quinoid mediators can be generated in the surface of carbon electrodes by oxidative pretreatment ["I. Besides these hydride acceptors, single-electron-transfermediators (e.g. transition metal complexesr2', 22], viologene derivatives12'* 271, ferrocenes 12*1, heteropolyanions[2g1, conducting polymers[3o]or ABTS C3l1 are also capable of oxidizing NAD(P)H. Examples for one- and two-electron acceptors are listed in Table 16.2-2. These mediators have been applied mostly freely diffusing but also immobilized at the electrode surface. A great variety of immobilization techniques have been used for the preparation of these modified electrodes - the mediator molecules have, for example, been directly adsorbed onto electrode surfaces, incorporated into conducting polymers or covalently linked to functional groups on electrode surfaces. Often the electron transfer between the reduced nicotinamide coenzyme and the mediator is rather slow because of kinetic limitations. In many of these cases electron transfer catalyzed by diuphoruse (E. C. 1.6.99.x) results in a drastic enhancement of the reaction rate (Fig. 16.2-5C). Diaphorase-catalyzedNAD(P)+regeneration was reported for example with methylene blue[32],PQQ[33](under aerobic conditions), ferrocene 12'1, N-methyl-p-aminophenol1341, N,N-dimethylindoaniline, 2,G-dichlorophenol indophenol (DCIP), [ F e ( c N ) ~ ] ~ -viologenes [~~l, or several quinoid structures f3'1. Many of the quinone-based mediators react in their reduced states with molecular oxygen. This aerobic regeneration has the advantage that no additional electrochemical equipment is necessary to perform NAD(P)+regeneration. On the other hand, reactive oxygen species are generated, which might inactivate enzymes and which therefore need to be removed from the reaction mixture. It should be mentioned at this point that most of the mediators described here were developed for analytical purposes. Only a few systems were applied to electrochemically driven dehydrogenase-catalyzed oxidations. This is partially because some systems exhibit moderate half-life times. In conclusion it can be said that, for each individual case, a mediator with a good performance and stability under the given production conditions has to be found.

1114

I

7 6 Oxidation Reactions Table 16.2-2. NAD(P)'.

Selection of frequently used mediators for indirect electrochemical regeneration of One-electron acceptors

SO,NH,

2,2'-azino-bis-(3-ethylbenzothiazoline-6sulfonic acid)-diammonium salt (ABTS)

[Os(bpy)z(PVI)ioCl]'

GD R = Me: methyl viologene R = Bz: benzyl viologene

Ferrocenes Two-electron acceptors

orthoquinone (and various derivatives)

para-quinone (and various derivatives)

16.2.2.5

Photochemical Regeneration

Various methods for photosensitized oxidation of NAD(P)Hhave been developed[37]. Photochemical methods are based either on the light-induced excitation of a mediator enabling it to oxidize NAD(P)H (reductive quenching mechanism) or on the light-induced excitation of the already reduced mediator, thus facilitating its reoxidation (oxidative quenching mechanism) (Fig. 16.2-6). For reductive quenching, photosensitizers such as tin porphyrins r3'1, methylene blue[3'], and other dyes L4O] are reported (Fig. 16.2-7).Ruthenium(11) tris bipyridine complexes in combination with viologenes are used for oxidative quenching. After

7 6 2 Oxidation ofAlcohols

NAD(P)H

Figure 16.2-6. Electron transfer from NAD(P)H to acceptors (A) via photosensitizers (S) facilitated by photochemical activation.

*-

NAD(P)+

reductive quenching

0

oxidative quenching

I

R z CH, rnethylene blue 5

-N+/

/

\

H

thionine

/

c) I

O CI + b -

/ \0 -

Sn(ll)-rneso-tetrarnethylpyridiniurnporphyrin 2,&dichlorophenol indophenol (DCP/P) Figure 16.2-7. Photosensitizers used for photochemical regeneration of NAD(P)'from NAD(P)H.

the oxidation of NAD(P)H,the reduced Ru complex is excited by light. The resulting powerful reduction agent transforms methyl viologene into the radical cation. The electrons from NAD(P)Hare usually transferred to molecular oxygen, protons or the 402 411

Next to soluble photosensitizers, semi-conductors were reported for NAD' regenerationI4*I.The advantage of these photochemical systems is that some of them utilize visible light, pointing towards the possibility of using sunlight for driving organic reactions. Disadvantageous, however, are the still low performances (TTN and TF of the photosensitizers and coenzymes) and the fact that photoexcitation results in the formation of strong oxidizing agents and the formation of free reactive radicals. Therefore, photochemical regeneration has not become one of the standard procedures, yet[37]. 16.2.2.6

Oxidations Catalyzed by Alcohol Dehydrogenase from Horse Liver (HLADH)

HLADH is certainly one of the most prominent and widely used oxidoreductases. The NAD-dependentenzyme is a dimer consisting of two almost identical subunits,

1116

I

7G Oxidation Reactions

which both contain two zinc atoms[43, The 3-dimensional structure was elucidated via X-ray analysis c4'. 461. HLADH exhibits a unique combination of a very broad tolerance for primary and secondary alcohols (or aldehydes and ketones in the reductive direction) with an 481. HLADH exhibits tolerance almost invariable and predictable stereospe~ificity[~~, to many organic solvents[49]and is active even in water-saturated organic solvents[", 421. Even though HLADH exhibits a rather poor specific activity in the range of 1-2 U mg-', it is commercially available at reasonable prices ($ 570/1000 U, Sigma 2001) and, more importantly, is fairly stable even in oxygen-containing mediaf4'].Also because of that, HLADH has been studied extensively during the last few decades.

16.2.2.6.1

Regeneration of NAD' in HLADH-catalyzed Reactions

Various concepts for the enzymatic regeneration of NAD' in combination with isolated HLADH have been reported, ranging from a second dehydrogenase such as glutamate dehydrogenase['I, 521 to enzyme-coupledor intrasequential approaches. A Baeyer-Villiger monooxygenase was applied to oxidize cyclic ketones produced in situ by HLADH with concomitant regeneration of NAD' (Fig. 1G.2-8)[531. Even though yields and enantiomeric excesses are moderate, this concept has synthetic significance and should be optimized in future. A very elegant reaction sequence was reported by Tanaka and coworkers[54]. HLADH was used for the kinetic resolution of a series of racemic 0-hydroxysilanes yielding one enantiomer in ee values ranging from 20 to 97 % in reasonable yields and the corresponding P-ketosilane.This P-ketosilane hydrolyzes spontaneously and drives the regeneration of NAD' catalyzed by HLADH (Fig. 16.2-9). Other NAD' regeneration approaches are based on the transfer of hydride either to PQQ (catalyzedby diaphorase) [331, directly to flavins [55-571, or to flavins via FMN reductase catalysis["]. Direct hydride transfer to flavins has the advantage that the alloxazine acceptor can be chosen freely, e. g. cheap riboflavin instead of FAD. On the HO,,, HLADH

H2O

0 2

lntrasequential regeneration o f N A D with HLADH and a Baeyer-Villiger monooxygenase (BVO) from Acinetobacter calcoaceticus.

Figure 16.2-8.

16.2 Oxidation ofAlcohols

SiMe,R

HLADH

+

NAD+ RMe,Si-OH

HLADH lntrasequential NAD' regeneration for HLADH-driven kinetic racemate resolution of P-hydroxysilanes.

Figure 16.2-9.

other hand, the spontaneous hydride transfer suffers from sluggish kinetics (k = 0.2 M-' s-'; turnover rates ranging between 0.06 and 1.8 h-I) which is app. 1000-fold slower than the values reported for enzymatic regeneration. For this reason, high excesses of the acceptor have to be applied in order to achieve acceptable regeneration rates. Introduction of FMN reductase accelerates this reaction remarkably. Electrochemical methods utilizing quinoid mediators [23, 241 or ferrocenes["I as well as photo~hemical[~~1 methods have also been applied to regenerate NAD' in combination with HLADH. Especially the electrochemical variants utilizing quinoid shuttle systems proved to be very efficient, with mediator performances as high as 130 catalytic cycles per hour and quantitative yields.

substrate

product

NAP

NADH

HlADH-catalyzed oxidations i n two-liquid phase systems (in the case o f buffer-saturated organic solvents, the aqueous phase is limited t o a layer around HLADH).

Figure 16.2-10.

1118

I

715 Oxidation Reactions Table 16.2-3.

Synthetic application o f HLADH in organic solvents.

Substrate(s)

Product(s)

o^""" o^"" Geraniol

Solvent

Remarks/Ref.

Hexane

Plugged-flowreactor for continuous production

Geranial

c::: ao"" Cinnamylalcohol

&& Racemic

Isopropyl ether

[31

Hexane

(41

Cinnamylaldehyde

&+A

2 R. Lortie, I. Villaume, M. D. Legoy, D. Thomas, Biotech. Bioeng. 1989, 33, 229-232. 3 T. Kawamoto, A. Aoki, K. Sonomoto, A. Tanaka, J. Fern. Bioeng. 1989, 67, 361-362. 4 J. R. Matos, C.-H. Wong. J. Org. Chem. 1986, 51, 2388-2389.

Ethyl acetate, chloroform, Isopropyl ether, butyl acetate

glass beads [51

Hexane

HLADH in polyacrylamide particles

HLADH immobilized on

[GI

5 J. Grundwald, B. Wirz, M. P. Scollar, A. M. Klibanov, J. Am. Chem. SOC. 1986,108.6732-6734. 6 C. Gorrebeck, M. Spanghoe, G. Lanens, G. L. Lerniere, R. A. Dornrnisse, J. A. Lepoivret, F. C. Adlerweireldt, Rec. Trav. Chim. Pays-Bas 1991, 110,231-235.

16.2.2.6.2 HLADH in Organic Media

Several applications of HLADH in organic/aqueous media have been reported (Table 16.2-3).The concept of these two liquid-phase reaction systems is shown schematically in Fig. 16.2-10.This approach is especially suitable for substrates and products with low solubility in aqueous media. Furthermore, the organic phase serves as a sink for products, thus decreasing problems resulting from product inhibition or back reactions.

16.2.2.6.3

Kinetic Resolution of Alcohols using HLADH

Because of its high enantioselectivity,H LADH has found widespread applications in the kinetic resolution of racemic alcohols and a-amino alcohols. Total turnovers of up to 10' for HLADH and 800 for NAD were reported with 90% residual activity, yielding the corresponding aldehydes in enantiomeric excesses up to 96%. The ahydroxy aldehydes were metabolized in situ by an aldehyde dehydrogenase to the corresponding a-hydroxy acids (Fig. 16.2-11)["I. Examples of further kinetic resolutions of racemates via regioselective oxidation using HLADH are given in Fig. 16.2-12.

162 Oxidation ofAlcohols

RE

O

I

H

I

NAD’

~

NADH

-NA<

NAD‘

NADH

f” HLADH

Figure 16.2-11. HLADH as enantioselective catalyst in the kinetic resolution of uic-diols (A) and a-amino alcohols (6). R = CHzOH, CHzF, CHzCI, CHZBr, CH,, CH=CH2,

C ~ H SCH2NH2, , (CH3)z.

k

OH

HLADH

k

+

OH

‘ 7 OH

HLADH

Figure 16.2-12.

Chemo- and stereoselective oxidations of sec-alcohols.

I

1119

1120

I

7G Oxidation Reactions

16.2.2.6.4

HLADH for the Oxidation o f meso-Compounds

Probably the most prominent application of HLADH is the oxidation of rneso-diols to homochiral lactones. Both 1,4-and 1,s-diolsare accepted as substrates (Table 16.2-4). The overall 4-electron oxidations proceed via two successive steps (tandem oxidation). The enantiomeric excesses often exceed 97 %. 16.2.2.7 Alcohol Dehydrogenase from Yeast (YADH)

Even though the primary sequences differ significantly, YADH exhibits almost the same quaternary structure as HLADH I['. Nevertheless, far fewer applications in biocatalflc processes are known for YADH than for HLADH. In part this is due to its low overall stability and its low resistance towards organic solvents[61].Furthermore the substrate spectrum of YADH is limited to primary alcohols and 2-hydroxyalkanes["]. It has been used in a few oxidative applicati~ns''~,G41. On account of its high specific activity (about 300 U mg-') together with its very low price (less than 1.2 $/lo00 U, Sigma, 2001), YADH has been used as a regeneration enzyme for NADHrG51.In this approach it is a problem that both ethanol and acetaldehyde as cosubstrate and coproduct of the regeneration reaction inactivate YADH and also other enzymes at low concentrations. This problem can be addressed by elegant techniques such as the use of gas membranes. Only volatile compounds such as ethanol or acetaldehyde can pass into the gas phase. This concept has been applied for lactate dehydrogenase (Fig. 16.2-13) 1"- G71. Hazardous acetaldehyde is removed and even recycled to form ethanol by treatment with sodium borohydride in the gas phase. Cycle numbers of over 10 000 are reported. 16.2.2.8 Alcohol Dehydrogenase from Thermoanaerobium brockii (TBADH)

TBADH is a NADP-dependent dehydrogenase with remarkable thermostability up to 65 oC[G8]. Neither HLADH nor YADH are able to convert linear secondary alco-

gaseous interphase Figure 16.2-13. Regeneration of N A D H with YADH. Acetaldehyde diffuses through the gaseous interphase into the second liquid phase where it is regenerated chemically to ethanol.

16.2 Oxidation ofAlcohols Table 16.24.

Examples of H LADH-catalyzed enantioselective oxidations of meso-diols. HLADH

HO * n HO

Meso-diol

'ri

HO

OH

8

H

O

0 OH

Lactone

Yield [%]

e e I"'%]

References

0

99

> 97

171

90

95

181

NDa

> 97

191

68

> 97

90

> 97

P

55

99

(2

95

"100"

> 99

> 99

70

99

fiOH

HO

=======

0

0

0

0

0

w 0

0

ko 0

a ND:not determined

1122

I

7G Oxidation Reactions 7 G.Hilt, B. Lewall, G. Montero, I. H. P. Utley, E. Steckhan, Liebigs Ann.lRecueil1997, 2289-229G. 8 T. Osa, Y.Kashiwagi, Y. Yanagisawa, Chem. Lett. 1994,3G7-370. 9 K. Mori,M. Amaike, J. E. Oliver, Liebigs Ann. Cham. 1992,1179. 10 Y Yamazaki, K. Hosono, Tetrahedron Lett. 1989, 30, 5313-5314.

11 M.-E. Gourdel-Martin, C. Cornoy, F. Huet, Tetra-

hedron: Asym. 1999, 10, 403-404. 12 R. N. Patel, M. Liu, A. Banejee, S. L., Ind.]. Chem. 1992,318,832-836. 13 Y. Yamazaki, K. Hosono, Tetrahedron Lett. 1988, 29, 57G9-5770.

hols. TBADH fills this gap: its activity is highest for secondary alcohols,being low for primary alcohols[48].Because of this rather narrow substrate spectrum, TBADH is mostly used for the regeneration of NADPH. Only a few synthetic applications are reported[24,"1. Figure 16.2-14gives one example where YADH was used simultaneously as an oxidizing enzyme and a NADPH regeneration enzyme (intrasequential cofactor regeneration). 16.2.2.9

Glycerol Dehydrogenase (GDH, E.C. 1.1.1.6)

GDH was isolated from various bacterial strains, especially from Schizosaccharomyces pombeC7'. 711 and Cellulomonas SP.['~,731. It displays a somewhat complementary substrate specificity to HLADH. While HLADH oxidizes meso-diols with secondary hydroxyl groups rather badly, they are readily oxidized by GDH to the corresponding (S)-a-hydroxyketones1'1. Furthermore, the natural substrate glycerol is transformed to achiral dihydroxy acetone by GDH while HLADH produces optically active (S)-glyceraldehyde.In many cases GDH seems to prefer secondary hydroxyl groups (Table 16.2-S), although this rule of thumb has some exceptions. In aqueous buffers GDH exhibits only low enantioselectivity, e. g. for the kinetic resolution of l-phenyl-l,2-ethanediol (which is most probably due to spontaneous racemization via enolization)[74]; furthermore, it suffers from pronounced product inhibition, accounting for low yields. Both problems (product inhibition and TBADH

0

Figure 16.2-14. lntrasequential regeneration of NADP with TBADH and a Baeyer-Villiger monooxygenase (BVO) from Acinetobacter calcoacetiGUS.

7 6.2 Oxidation ofAlcohols

Alcohol oxidations catalyzed by glycerol dehydrogena~e['~].

Table 16.2-5.

Product

Substrate

-

HOTOH HOTOH OH H O T O '

0 H

O

O

G

' HO 0

OH

OH

H 14

O

0

0

OH

H

T

. 0

T

OH

0

J. H.Marshall, I. W.May, J.

Sloan,J. Gen. Microbiol. 1985,131, 1581-1588.

++ /

& /

a

NAu in situ extraction into hexane via hollow fibre module

+COOH OH -

LDH

I I OC O O H

Deracernization of roc 1-phenyl-1,Z-ethandiol coupled t o in situ product extraction via a hollow fiber module. Figure 16.2-15.

1124

I

IG

Oxidation Reactions

K13CN+HCH0

-

d

YH H0
2-

- -

R

GPDH

HO
2-

2-

phosphatase

H07y0p03

“’OH

HO””

“’OH OH

OH

Figure 16.2-16. Synthesis of”C-labeled sugars in a tandem reaction of CPDH, aldolase, and phosphatase.

Figure 16.2-17. Pired anodic regeneration of NAD’ coupled to cathodic reduction ofpyruvate.

racemization) can be solved by in situ extraction into a second (organic) phase (Fig. 1G.2-15)[741. This biphasic system yielded higher ee values (99% instead of 58%) at maximal theoretical conversions (50% instead of 38 %) in significantly shorter reaction times (GO h instead of 170 h) compared to the solely aqueous system. Since GDH contains autooxidizable thiol groups, it is necessary to perform such reactions in media essentially free from oxygen. 16.2.2.10 Glycerol-3-phosphate Dehydrogenase (GPDH, E. C. 1.1.1.8)

GPDH has been isolated from various organisms. The enzyme from rabbit muscle is commercially available. Its synthetic applications are limited because of its very

7 6.2 Oxidation ofAlcohols

narrow substrate spectrum it almost exclusively accepts ~-glycerol-3-phosphate [75, “1. The product 3-hydroxyacetone phosphate, however, is an essential substrate of aldolases and therefore can serve as a building block in the enzymatic synthesis of non-native sugars and polyols. Although the redox equilibrium of GPDH favors the reduced substrates even more than in the case of GDH, it has been employed in the synthesis of radioactively labeled carbohydrates starting from K13CN and formaldehyde (Fig. 16.2-16)[77, 781. Depending on the substrates, singleor double-labeled glucose, fructose or sorbose are available by the sequence outlined in Fig. 16.2-16. 16.2.2.1 1

Lactate Dehydrogenase (LDH, E.C. 1.1.1.27)

LDH was used to catalyze the deracemization of lactate in a very elegant electrochemical approach. The driving force of the endergonic reaction was supplied by anodic regeneration of NAD’ and cathodic reduction of pyruvate (Fig. 16.217)17’), 80]. Thus, both LDH products were removed efficiently, avoiding product inhibition. The electrochemical reduction of pyruvate leads to racemic lactate, producing 50% of the desired product and 50% of “new” substrate for LDH. An

0

A o o -

e-

Figure 16.2-18. Electrical wiring of lactate dehydrogenase (LDH).

1126

I

7G Oxidation Reactions

interesting approach to direct “electricalwiring” of LDH to an electrode was reported recently (Fig. 16.2-18)[*‘I. NAD was covalently linked via a PQQ spacer to a gold electrode. This modified electrode is capable of binding LDH over the exposed nicotinamide groups. Upon oxidation of lactate to pyruvate the excess electrons tunnel from NADH in the active site to PQQ and eventually to the anode. Thus, a kind of electrical linkage between the enzyme and the electrode is established. The enzymes were crosslinked, as LDH is a homotetramer and might dissociate during the reaction. This approach is not only useful for electrochemical biosensors but might be transferred to other oxidoreductase reactions. 16.2.2.12 Carbohydrate Dehydrogenases

Many so-called polyol dehydrogenases have been reported in literature, for example various glucose dehydrogenases, mannitol dehydrogenase, fructose dehydrogenase, and uridine-5‘-diphosphoglucose dehydrogenase. Glucose dehydrogenase (E. C. 1.1.1.47)was applied for the production of D-gluconic acid in a plug-flowreactor with direct electrochemical regeneration of NAD+[82].Glucose-6-phosphate dehydrogenase (E.C. 1.1.1.49) is a common regeneration enzyme for NADPHIG9]. Most polyol dehydrogenases are not specific for their native substrate, but also catalyze the oxidoreduction of various carbohydrates. Thus, they can be applied for the production of (non-)naturalsugars which are especially valuable in the sweetener industry. Yet their applications are limited compared to the polyol oxidases (see Sect. 16.2.3)

COOH

0

“‘OH

Figure 16.2-19.

HO’”

“OH

Regioselective oxidation o f cholic acid by hydroxysteroid dehydrogenases.

16.2 Oxidation ofAlcohols

HO'"

NAD+

NADH

NADH

NAD'

Figure 16.2-20. 3a- and 3fi-hydroxysteroid dehydrogenase (HSDH) catalyzed stereoinversion in steroids.

16.2.2.13 Hydroxysteroid Dehydrogenases (HSDH)

The hydroxysteroid dehydrogenases comprise another group of synthetically interesting dehydrogenases. For many hydroxylated positions of the steroid backbone, individual NAD(P)' dependent dehydrogenases exist, which selectively oxidize the respective residue. For example, the three hydroxy groups of cholic acid in the 3-,7-,and 12-positions can all be oxidized regioselectively(Fig. 16.2-19)[83-851. In addition to the regioselective oxidation of the hydroxy groups in virtually every position, a discrimination of the absolute stereochemistry can be achieved by various a- or P-selective HSDHs. Thus, the stereoinversion of various steroids was achieved by successive oxidation at position 3 with 3a-HSDH and subsequent reduction with 3P-HSDH (Fig. 16.2-20) Hydroxy functions in other positions were not modified, and the products at the end of the sequence were essentially pure. Because of the low solubility of the reactants, biphasic systems with ethyl (butyl) acetate as organic solvents were used as reaction media. 16.2.2.14 Other Dehydrogenases

In addition to the alcohol dehydrogenases mentioned above, ADHs from various other sources were examined, especiallywith respect to increased stability,resistance to organic solvents, and catalpc properties. A NAD+dependent ADH isolated from Sulfolobus solfataricus was found to exhibit better thermostability than HLADH [tl/z (GO "C) = 20 h] together with a distinctive preference for (S)-alcohols(complementaryto HLADH) ["I. The enzyme has a broad substrate specificity that includes linear and branched primary alcohols and linear and cyclic secondary alcohols[48].The highly purified enzyme exhibits a specific activity of 4 U mg-' (for benzyl alcohol at 65 "C)18', 1'. To date, this enzyme is not commercially available. Hummel et al. established a new route to enantiomerically pure alcohols by the

1128

I

76 Oxidation Reactions

R. erythopolis

NAD(P)t

NAD(P)H

NADH-oxidase

HZOZ

* )/

Hzo

catalase

A

SY"

OH

Deracemization of 1-phenyl1-ethanol. The ADHs from R. erythropolis and L. kefir exhibit complementary steroespecificity. Combination o f both in an oxidation-reduction sequence yields the desired enantiopure alcohol. Figure 16.2-21.

0 2

NADH

AOH o o -u

&R'

+

L , R

OH

OH

48-50 (RR) % >99 % ee

3-20 YO (9 299% ee

Aoo o -

LDH

++ OH

OH

QoH OH

&OH OH

Kinetic resolution o f racemic syn-diols by Bacillus stearothermophilus diacetyl reductase (BSDR). A: reaction with LDH-catalyzed regeneration of NAD+; B: selection of syn-diols applied. Figure 16.2-22.

0

7G.2 Oxidation ofAlcohols I1129

combination of a (R)-specific,NADP-dependent ADH from Lactobacillus keJr and a (S)-specific,NAD-dependentADH from Rhodococcus erythropoli~[~~~. In a first step, a kinetic resolution yielded 50 % of the desired alcohol. Subsequently the ketone was reduced with the suitable ADH, finally yielding the desired optically pure enantiomer in 100% yield (Fig. 16.2-21). Recently, diacetyl reductase (Acetoin reductase, E. C. 1.1.1.5) from Bacillus stearothemophilus (BSDR) was reported to be a powerful catalyst in the oxidative kinetic resolution of vic-diols (Fig. 16.2-22)[901. All syn-diols tested yielded the enantiopure (R,R) diols in almost maximum theoretical yields, a-hydroxy ketones were largely further oxidized to the corresponding diketones. Oxidation of vic-anti diols only gave ee values in the range of 62-76 %. 16.2.3 Oxidases as Catalysts

16.2.3.1 General Remarks

Oxidases utilize molecular oxygen as terminal electron acceptor. This can be considered as aerobic regeneration of the prosthetic group of the oxidase. At first glance, this seems to offer a simpler enzymatic oxidation procedure compared to the coenzyme-dependent dehydrogenases or monooxygenases. However, with few exceptions such as cytochrome c oxida~e”~1, some NADH oxidases”’] or laccases[”], which reduce molecular oxygen directly to water in an overall four-electron transfer step, O2reduction generally leads to hydrogen peroxide (transfer of two electrons) or to the superoxide radical anion (transfer of one electron) as primary reduction products. 16.2.3.2 Methods to Diminish/Avoid H202formation

Autoregeneration of oxidases with concomitant catalase-catalyzed disproportionation of hydrogen peroxide is a simple and effective regeneration method (Fig. 16.223); it is quite commonly used with oxidase reactions.

oxidase (E.C. 1 .x.3.y)

catalase (E.C. 1.11.1.6)

Figure 16.2-23. Coupling o f oxidase autoregeneration and catalase for dismutation of hydrogen peroxide.

1130

I

71 Oxidation Reactions

Figure 16.2-24.

Indirect electrochemical regeneration of an oxidase.

Hydrogen peroxide, however, is highly reactive and irreversibly inhibits enzyme activity (also catalase) even in low concentrations. Hydrogen peroxide can be avoided if excess electrons are transferred to the anode. However, direct electron transfer between enzymes and solid electrodes is usually very slow because the enzymatic active sites are often deeply buried within the protein shell and therefore inaccessiblefor the electrode (the tunneling probability of electrons is a function of distance). In order to accelerate the electron transfer, low molecular weight redox active substances can be used to shuttle the electrons between the enzyme and the electrode. This indirect electrochemical enzyme regeneration is represented schematically in Figure 16.2-24. For the anaerobic electrochemical regeneration of a given oxidase, a suitable mediator can be chosen from various organometallic complexes, especially ferrobut also bipyridine/phenanthroline, terpyridine, or hexacyano comcenes [94'021, Also, quinoid salts such as TTF/TCNQ (tetrathiofulvalene/tetraplexes [Io3, cyanoquinodimethane)['05. 1"' as well as benzoquinones [lo7] and redox dyes such as phenazine and phenothiazine derivatives (MPMS, thionin, azure A, and azure C) ['08] proved to be useful redox agents for indirect electron transfer. Even incorporation of oxidases into conducting polymers made of polypyrrole or polythiophene derivatives proved to function for electrochemical regeneration ['091. It should be mentioned at this point that most of the research in the field of electrochemical oxidase regeneration concentrates on analccal applications, inspired by the search for electrochemical biosensors ['lo]. However, it was demonstrated that indirect electrochemical methods are suitable for prolonging oxidase operational stability. In a particular example, glucose oxidase (E.C. 1.1.3.4) was immobilized on a carbon felt anode and regenerated with the benzoquinone/hydroquinone redox couple (Fig. 16.2-25)[lo7].Thus, the operational stability of glucose oxidase could be increased at least 50 times compared to the use of molecular oxygen as oxidant. Productivities as high as 1OOg h-' L-' were reached. One disadvantage of the electrochemical methods is the need for rather elaborate equipment. Recently, Baminger et al. proposed a novel concept of enzymatic regeneration of a range of redox mediators including quinones and various redox dyes [931. Instead of reoxidizing these mediators via the anode, laccases are employed. Laccases (E. C. 1.10.3.2)are multi-copper oxidases['"] that are found in various trees and fungi["2. '131 . Laccases catalyze the oxidation of various structurally diverse

n I-g$ glucose oxidase

D-glucose

0

Figure 16.2-25.

"OH

H20

I

1131

JoHO f;!

OH gluconic acid

OH

Indirect electro-enzymatic oxidation of glucose using glucose oxidase.

flavoprotein

Figure 16.2-26.

HO'"'

162 Oxidation of Alcohols

laccase

Laccase-based regeneration concept for oxidized flavoproteins

(oxidases).

substances with concomitant reduction of molecular oxygen to water['14], thus avoiding the generation of hazardous hydrogen peroxide (Fig. 16.2-26). This regeneration concept was tested with pyranose oxidase (P20, E.C. 1.1.3.10)~933]. Interestingly, it was found that P20 shows higher affinity for some mediators than for 0 2 (KM value for 1,4-benzoquinone is 120 m M compared to 650 mM for 02)with otherwise comparable activities yielding a 6 times higher kCat/KM value. Preparative scale biotransformations could be performed with two-fold volumetric productivities. The TTNs were 1.1x lo6 for P20, with a residual activity of 85 %, and 800 for 1,4-benzoquinone. Similar results were obtained with the enzyme cellobiose-dehydrogenase (E. C. 1.1.99.18),which is incapable of autoregeneration, in combination with ABTS or DCIP and laccase.

1132

I

7G Oxidation Reactions

r~

R H HO o

R

S

O

P20

H

OH

HoHO

0,

R = CH,OH H

HzO, Figure 16.2-27.

OH

0

catalase

-

o,

+

H,O

Oxidation of carbohydrates specifically at C-2 by pyranose oxidase (P20).

16.2.3.3

Pyranose Oxidase (P20, E.C. 1.1.3.10)

P20 is common among wood-degrading basidiomycetes [ll5].It has been isolated and characterized from various microorganisms Although the substrate specificity vanes to some extent among the P20s isolated from different fungi, P20s have some properties in common, such as the homotetrameric structure with covalently bound FAD. The main metabolic role of P 2 0 appears to be as a constituent of the fungal ligninolytic system that provides the lignin-degrading lignin peroxidase and manganese peroxidase with hydrogen peroxide l1 171. Natural substrates of P 2 0 are probably D-glucose, D-galactose, and D-xylose,which are abundant in lignocellulose and which are oxidized to the corresponding 2-keto sugars. In addition, P20 exhibits significant activity with a number of other carbohydratesIll8]. During such oxidations, electrons are transferred to molecular oxygen, yielding hydrogen peroxide. In addition, benzoquinones, 2,G-dichloroindophenol, as well as ABTS were reported to function as electron acceptors[’3, ll’l. Interestingly, up to 11-fold increased reactivity (compared to molecular oxygen as electron acceptor) was found. P20 is currently used in various analytical applications, e. g., in clinical chemistry for the determination of 1,S-anhydro-D-ghcitol,an important marker for glycemic control in diabetes patients or in amperometric biosensors for the detection of monosaccharides [120, ‘’1 . For the last two decades, P20 has received increased attention as the key catalyst in several biotechnological applications. Only a few can be mentioned here. The essential structural requirements of substrates for P20 are the six-membered ring of pyranoid saccharides and an equatorially orientated 2-OH group [1221. In some cases regioselective oxidation at C-3 was observed[”81. The general reaction scheme is given in Fig. 16.2-27. Table 16.2-6 gives a selection of preparative oxidations reported with P20. The “Cetus process” P 2 0 is involved in the so-called “Cetus process”, in which D-fructose is produced

from cheap D-glucose (Fig. 16.2-28).

7 6.2 Oxidation ofAlcohols

I

1133

Table 16.2-6.

Substrates and oxidation products of pyranose oxidase.

Substrate

Yield p4"

Product

Activity [%la

Oxidation of monosaccharides

100

100

94

40

70

8

5

very low

100

96

o-Glucosone

D-Glucose

HO"'

"OH

OH

OH

D-Allose

~-Ribo-hexos-2-ulose

H O T : I I I l HO

HO OH

OH

D-Galactose

~-Lyxo-hexos-2-ulose

HO"'

HO"

.

OH

OH

~-Erythro-pentos-2-ulose

D-Ribose

HO"

" OH

3d-~-Glucose

HO"

3d-~-Erythro-hexos-2-ulose

HO'"

100

92

100

75

OH

HO"

HO"' OH

1,5-Anhydro-~-glucitoI

OH

References

1134

I

16 Oxidation Reactions Table 16.2-6.

(cont.).

[%r Activity [%I" ~~

Substrate

Product

Yield

~

~

~~~~

References

Oxidation o f Disaccharides HO

HO

Hobob: HO

6H HO

OH

Allolactose

100

N D ~

P I

100

N D ~

(161

OH

Allolactulose

HO

HO

OH OH

Meliobiose

Meliobiulose

OH

OH

Gentiobiulose

Gentiobiose HO

6H

OH

Isomaltose

Palatinose

a expressed as percentage ofyield and activity of o-glucose oxidation: b ND: not determined

15 S . Freimund, A. Huwig, F. Giffhom, S . Kopper, Chem. Eur. J. 1998,4,2442-2455.

16 C. Leitner, P. Mayr, S. Riva, J. Volc, K. D. Kulbe, B. Nidetzky, D. Haltrich,J. Mol. Cat. B: Enzymatic 2001,11,407-414.

Hydrogen peroxide is not merely dismutated by catalase, but used as substrate in a second enzyme cascade reaction producing propylene oxide['23-125] . In an alternative process 1126] the reduction step was performed enzymatically using aldose reductase and formate dehydrogenase for NADH regeneration. Thus, essentially glucose free D-fructose was obtained.

16.2 Oxidation ofAlcohols

I

1135

H A

HCI

‘I-

i.

fi

haloperoxidase

halohydrine epoxidase

Figure 16.2-28. lsornerization of D-glucose to o-fructose with pyranose oxidase (P20) and coupling o f hydrogen peroxide t o a synthetic reaction (Cetus process). Table 16.2-7. Kinetic resolution of some racernic 2-hydroxy acids t o the (R)-2-hydroxy acids and the corresponding 2-keto acids[”1.

Substrate

Yield [“/.I

e e I%]

49-50

> 98

50

> 99

47

86

9H *OOH

1,2.4,7

OH

&/trans -0-COOH

c

OH 17 W. Adam, M. Lazarus, 9. Boss, C. R. Saha-Moller,H.-U. Humpf, P. Schreier, /. 0%.Chem. 1997,62, 7841-7843.

16.2.3.4

Glycolate Oxidase (E. C. 1.1.3.1 5) Glycolate oxidase is a peroxisomal enzyme that is found in the leaves of many green plants and in the liver of mammalians. The enzyme isolated and for economic reasons only partially purified from spinach (Spinacia oleracea) was applied to the enantioselective oxidation of various 2-hydroxy acids yielding the corresponding Enantiopure 2-hydroxy acids are 2-keto acid and the remaining ( R ) valuable building blocks in the synthesis of glycols [‘’I, haloesters [12’1 or epUnless the steric demand of the substituents close to the alcohol function is too big, the oxidation proceeds smoothly to the full theoretical conversion with enantiomeric excesses of the alcohols usually in the range of 98-99% (Table 16.2-7).

1136

I

7G Oxidation

Reactions

Rq~~

Figure 16.2-29.

glycolate oxidase

. q O H

0

0

('

0 2

Deracemization o facids racemic 2-hydroxy in a combination of glycolate oxidase and lactate dehydrogenase (LDH).

H*O NAOH

NAD'

co2

HCO,

FDH Table 16.2-8.

Conversion of racemic 2-hydroxy acids into (R)-2-hydroxy acids by the combined action of glycolate oxidase and D-lactate dehydrogenase~'81.

Substrate

Oxidase [U]

Dehydrogenase[U]

Reaction time [h]

Yield ["/.I ~

2

450

e e pi] ~~

66

100

> 99

210

100

94

0

One unit (U) is defined as the amount of enzyme which converts 1 pmol of substrate per minute.

18 W. Adam, M. Lazarus, C . R. Saha-Moller, P. Schreier, Tetrahedron Asymmetry 1998.9,351-355.

Kinetic resolutions have a maximum yield of only 50%. Therefore, a second enzymatic process was added after completion of the glycolate oxidase-catalyzed kinetic resolution['31! By addition of D-lactate dehydrogenase (E. C. 1.1.1.28) together with formate dehydrogenase for NADH regeneration, enantiospecific reduction of the 2-keto acid was achieved. Overall, a quantitative transformation (deracemization) of the racemic 2-hydroxy acid into the corresponding (R)-2-hydroxyacid was achieved (Fig. 16.2-29). Unfortunately, this process cannot be performed in a more elegant and more efficient one-pot synthesis. On the one hand, the pH optima for the three enzymes are not compatible with each other, and on the other, lactate dehydrogenase is air sensitive. In addition to this, glycolate oxidase also catalyzes the reverse reaction under aerobic conditions, thus lowering the ee-value.Therefore, the reaction mixture is filtered (glycolate oxidase can be reused) and, after pH adjustment, the second enzymatic transformation is performed. Table 16.2-8 shows some results of this procedure. Glycolate oxidase has been studied thoroughly not only for specific oxidation of

762 Oxidation ofAlcohols

HO-OH

m

ethyleneglycol

Homo

b

H O T o

OH

glycolaldehyde

glycolic acid

I

0-0

D

OH glyoxylic acid

glyoxal Figure 16.2-30.

Sequential oxidation of ethylene glycol t o glycolic acid.

glyoxylic acid

Vf4

glycolate oxidase

1

CO,

+ HCO,H + OH

Figure 16.2-31. Synthesis o f glyoxylic acid by glycolate oxidase. The undesired sidereactions (A) with hydrogen peroxide and (B) overoxidation by glycolate oxidase are prevented by in situ forrnation o f an irnine.

(S)-2-hydroxypropionic acid (lactate)[1321 and for the kinetic resolution of racemic 2-hydroxy acids[127,l3l],but also for selective oxidations of 1,Zdiols such as ethylene glycol (Fig. 16.2-30). Reports on the specific conversion of glycolic acid into glyoxylic acid are numerous. Isobe et al. Introduced an in vivo system utilizing Alculigenes sp. isolated from media containing 1,2-propanediol.By carefully adjusting the pH, a yield of 95 % was obtained f 1331. DiCosimo and coworkers optimized the in vitro production of glyoxylic acid from glycolic acid with glycolate oxidase from spinachll3'1. Improvements in operational stability as well as in productivity were achieved by enzyme immobilization either onto a solid or in permeabilized, metabolically inactive cells of Pichia pastoris or Hansenula polymorphu, containing overexpressed glycolate oxidase from spinach together with catalase. The undesired oxidation of glyoxylic acid by hydrogen

I

1137

1138

I

16 Oxidation Reactions

Figure 16.2-32. Non-natural substrates for nucleoside oxidase from Pseudomonas sp. These compounds are converted selectively t o their corresponding 5'-carboxylic acids.

peroxide (yielding formate and carbon dioxide) and further metabolization by glycolate oxidase could be prevented by trapping the aldehyde function of glyoxylic acid as imine (Fig. 16.2-31)[13'1. 16.2.3.5

Nucleoside Oxidase (E.C. 1.1.3.28)

Nucleoside oxidase is produced by Pseudomonas species and related Gram negative bacteria [1371. The hetero-tetramer with covalently bound FAD oxidizes the 5'-hydroxyl group of purine and pyrimidine nucleosides to the corresponding carboxylic acids. It has found application in the analytical determination of nucleosides (e.g. in assessing food freshness)[1381. At Glaxo Wellcome R&D it found attention as key step in the production of anti-inflammatory compounds [139-1411. Several non-natural substrates were selectively converted on multi-gram scale into their 5'-carboxylic acids (Fig. 16.2-32). The operational stability of the enzyme was improved by immobilization onto a solid matrix and especially by substitution of molecular oxygen as the primary electron acceptor by stoichiometric amounts of hydroquinone. 16.2.3.6 Glucose Oxidase (E. C. 1.1.3.4)

The most prominent of the alcohol oxidases is glucose oxidase. The dimeric flavoenzyme catalyzes the oxidation of P-D-glucoseto D-glucono-&lactone,a reaction that has attracted the attention of generations of analytical chemists because of its

I62 Oxidation ofAlcohols

possible applicability in glucose sensors for diabetes The reaction of the stoichiometrically formed hydrogen peroxide with various dyes can be used as the analpcal More elegant variants (that at the same time avoid the formation of hazardous hydrogen peroxide) utilize anaerobic, electrochemical regeneration with a suitable mediator. Thus, the catalytic current becomes the analpcal signal. Several approaches have been reported, e. g. the utilization of freely diffusible quinones [Io7], the incorporation of glucose oxidase in a conducting polymer (produced from 1,4-hydroquinones and soybean peroxidase), or the immobilization of several mediators in the vicinity of the prosthetic redox center['*. 991 Because of the high substrate specificity of glucose oxidase, which almost exclusively accepts glucose (other substrates such as D-maltose, D-xylose, or Lsorbose are converted with less than 6% of the activity on glucose[lU*14'1), this oxidase has not found any synthetic application, but it is frequently used in the food industry to remove traces of molecular oxygen from vacuum sealed products. Immobilized glucose oxidase is also used for the deoxygenation of juices and beer [1461. 16.2.3.7

Alcohol Oxidase (E. C. 1.1.3.1 3)

The aliphatic alcohol oxidase, a FAD-dependent enzyme, catalyzes the oxidation of primary short-chain alcohols to the corresponding aldehydes. Dioxygen can be replaced by synthetic acceptors such as dichlorophenolindophenol or phenazine methosulfate [14'1. By utilizing an alcohol oxidase from Pichia pastoris or Candida sp.[l4'I, almost complete conversion of ethylene glycol into glyoxal (Fig. 16.2-30) was observed. These enzymatic routes were shown to be superior in terms of reaction conditions and yields compared to the chemical variants that make use of metal catalysts or even nitric acid for the oxidation of ethylene glycol. Recently, aliphatic alcohol oxidase was applied as dehydrated enzyme in a gas-solid bioreactor an excess amount of catalase was added to prevent oxidase inactivation.

GAOX

HO

"'0H

HO

OH Figure 16.2-33. Galactose oxidase (GAOX) catalyzed oxidation of a-o-galactose t o meso-galactohexodialose.

OH

1139

I

I

16 Oxidation Reactions Table 16.2-9.

Substrates and products o f galactose oxidase. Product

Substrate

References

OH

rneso-Galactohexodialdose

OHOH

OH

OHOH

UDP-[14C]-Galactose

OH

UDP-[14C]-Galacturonicacid

HO{OH

HO

D,L-Threitol

D-Threose + L-Threitol

HO

) "03-OH HO

HO

Xylitol

Hot OH

OH

"$OH HO

811

HO

HO

OH

OH OH

OH

L-Glucose + D-Glucitol

L-Galactose + D-Galactit01

OH

OH

7 G.2 Oxidation ofAlcohols I1141 Table 16.2-9.

(cont.).

Substrate

Product

References

OH

HOA

O

HO-0

H

L(-)Glyceraldehyde

H O L C I (S)-Halodiol + (R)-Aldehyde 19 S. S. Basu, G. D. Dotson, C. R. H. Raetz, Anal. Biochem. 2000,280,173-177. 20 D. G . Drueckhammer, W. J. Hennen, R. L. Pederson, D. F. Barbas, C. M. Gautheron, T.Krach, C. H. Wong, Synthesis 1991,7, 499-7525.

21 A.M. Klibanov, B. N. Alberti, M. A. Marletta, Biochem. Biophys. Res. Commun. 1982,1982, 108.

Table 16.2-10. Substrates and products in the kinetic resolution of allylic alcohols with cholesterol oxidaselZ21. Substrate (R = H, OH)

Product

d l

HO

HO

J35 +-'F

0

No product detected

HO 22 S . Dieth, D. Tritsch, J:F.

Biellmann, Tetrahedron Lett. 1995,36,2243-2246.

16.2.3.8 Galactose Oxidase (GAOX, E. C. 1.1.3.9)

Galactose oxidases belong to the group of copper-dependent oxidases. For the GAOX from Dactylium dendroides the existence of covalently bound pyrroloquinoline quinone (PQQ)could be shown['45].It catalyzesthe specific oxidation of the hydroxyl group in position 6 of galactose (Fig. 16.2-33)[l5O]. The enzyme regeneration can be performed aerobically or utilizing mediators

1142

I

7 B Oxidation Reactions Figure 16.2-34. Ferric protoporphyrin IX as prosthetic group in most peroxidases.

such as ferrocene["'], tetracyano-iron-1,lO-phenanthroline, or cobalt tert-pyridine complexes[lo31. GAOX stereospecifically oxidizes a broad range of substrates (Table 16.2-9). In synthetic applications, the oxidation of racemic or meso-polyolssuch as D,L-threitol or xylitol to the non-native sugars are of special interest['51, lS2]. In addition to the monosaccharides represented in Table 16.2-9, GAOX also converts di- or oligosaccharides[lS31. 16.2.3.9 Cholesterol Oxidase (ChOX, E. C. 1.1.3.6)

ChOX from Rhodococcus erythropoliswas applied for the kinetic resolution of racemic mono- and bicyclic ally1 alcohols (Table 16.2-10)(lS41.Although the substrates tested were much smaller than the native substrate cholest-4-en-3P-01,reasonable enantioselectivities (E) in the range of 7-20 were found for the (S) alcohols. Both enantiomers of the alcohol (entry 1)were oxidized with moderate enantioselectivities ( E = 7) for the (S) enantiomer. For bicyclic alcohols, the position of the hydroxyl group with respect to the methyl group is essential. Only at a relative trans configuration of both substituents significant oxidation occurred. By utilizing organic redox dyes as primary electron acceptors and concomitant reoxidation at a glassy carbon electrode, amperometric biosensors for cholesterol based on cholesterol oxidase were developed[108]. 16.2.4

Peroxidases as Catalysts 16.2.4.1

Introduction

Peroxidases (E. C. 1.11.1.7) are ubiquitously found in plants, microorganisms, and animals. Most peroxidases studied so far contain ferric protoporphyrin IX (protoheme, Fig. 16.2-34) as the prosthetic group[1ss].However, some peroxidases also contain selenium (glutathione peroxidase)(1561, vanadium (bromoperoxidase)[1571,

76.2 Oxidation ofAlcohols

A

D E

2 H++ 0,

Figure 16.2-35. Methods of generating appropriate hydrogen peroxide concentrations for chloroperoxidase reactions, (A) enzymatically with glucose oxidase and (B) electrochemically by cathodic reduction of molecular oxygen.

manganese (manganese peroxidase)[1581,and flavin (flavoperoxidase)[1591 as prosthetic groups. Most peroxidases accept a variety of peroxides, such as hydrogen peroxide or alkyl hydroperoxides, as oxidizing agents. The mechanism includes the activation of oxygen in a high valence iron-oxo specie^['^^^ "'1. 16.2.4.2

Methods to Generate HZOZ

At a first glance, utilization of cheap hydrogen peroxide as electron acceptor seems appealing. The major drawback, however, is the sometimes rapid inactivation of peroxidases by their substrate. For example, chloroperoxidase (CPO, E. C. 1.11.1.10) exhibits a half-life time of 38 min even at an H202 concentration of 50 pM [1611. Several approaches to controlling hydrogen peroxide at a constant low concentration have been reported. In aqueous/organic emulsions, the use of tert-butyl hydroperoxide is beneficial. On the one hand, the peroxide concentration is limited according to the partition coefficient, and on the other hand, tert-butanolwas shown to exert a stabilizing effect on CPO['"]. The slow continuous addition of hydrogen peroxide results in better CPO performance [lG3], which can be even further improved by sensor-controlledaddition of H202[1G2], increasing the CPO total turnover number for indole oxidation more than 20-fold to ca. 860 000.

I

1143

1144

I

IG

Oxidation Reactions

Table 16.2-11.

Chloroperoxidase-catalyzed oxidation of some alcohols to the corresponding

aldehydes. Substrate

Yield I"/.]

Remarks and reference

94

H 2 0 2 or

95

H20z or tert-butyl hydroperoxide as oxidants [231

92

H202

0"""

quantitative

O -H

81

/\/=\/OH

95

O -H

99 OH

or tert-butyl hydroperoxide as oxidants

3 times higher activity with tea-butyl hydroperoxide in biphasic systems compared to H202 in b~ffer['~I

Production in gram-scale; low, non-enzymatic cis/trans isomerization observed (*'I

97

0 &OH

tert-butyl hydroperoxide as oxidants

50 (40% ee)

Production in gram-scale, low yield with cis-isornerl2'1

46 (45% ee)

1251

92

74

+ O .H \

Quantitative conversion; significant amounts of acid as the product of overoxidation were found[z61

25 E. Kiljunen, L. T. Kanerva,J. Mol. Cat. B: Enzy23 S. Hu, L. P. Hager, Biochem. Biophys. Res. Commatic 2000, 9, 163-172. mun. 1998,253, 544-546. 24 B. K. Samra, M. Anderson, P. Adlercreutz, Biocat. 26 M. P. J. van Deurzen, F. van Rantwijk, R. A. Sheldon,J. Carbohydr. Chem. 1997, 16,299-309. Biotran$l999, 17, 381-391.

7 6.2 Oxidation ofAlcohols

I

1145

COOH COOH

2e- 2 H +

~

H

o

o

c

~

C

O

O

0

HO

0

OH

Figure 16.2-36. Pyrroloquinoline quinone (PQQ) in its oxidized and reduced form as prosthetic group for most quinoprotein dehydrogenases.

However, external HzOz addition still has the disadvantage that locally high concentrations occur at the entry points, resulting in CPO inactivation at these hot spots. This can be circumvented via in situ generation of hydrogen peroxide. Two promising approaches have been reported so far: (i) another enzymatic reaction producing HzOz e. g. with glucose ~ x i d a s e [ and ~ ~ ~(ii) ] , electrochemicalreduction of molecular oxygen (Fig. 16.2-35)["l, "'1. In both approaches, drastic increases of the number of CPO catalytic cycles up to 1.1 x 10' were achieved. 16.2.4.3 Chloroperoxidase(CPO,

E. C. 1.1 1.1.lo)

Publications on CPO-catalyzed oxidations of alcohols are rare. However, some selective oxidations of aliphatic, allylic, propagylic and benzylic alcohols to the aldehyde stage have been reported (Table 16.2-11). 16.2.4.4 Catalase (E. C. 1.11.1.6)

Most commonly, catalase is applied for the dismutation of hydrogen peroxide[16G]. On reaction of catalase with one molecule of hydrogen peroxide, the intermediate high valence iron-oxo species is generated. This species, however, is a potent oxidant and readily reacts not only with a second molecule of hydrogen peroxide (yielding water and molecular oxygen) but has been reported to oxidize various other compounds such as methanol or nitrite [166]. Klibanov and coworkers enlarged the substrate spectrum by including a variety of alcohols that were oxidized to the corresponding aldehydes. Depending on the substrate and the reaction medium, high enantioselectivitiesare reported The generation of reactive catalase in its oxidized stage can also be achieved by direct electrochemical oxidation (transfer of electrons from ferric protoporphyrin IX to the electrode). Thus, catalase immobilized on graphite electrodes has been used for the hydrogen peroxide-free oxidation of phenol [lG81.

H

1146

I

71 Oxidation Reactions QHDH

glycidol

QHDH C. testosteroni

'>( O

q

+

_____)

O

H

'>( O W 0


COOH

solketal Resolution o f alcohols by enantioselective oxidation using quinohemoprotein dehydrogenases (QHDH) from different microorganisms. Figure 16.2-37.

16.2.5

Quinoprotein Dehydrogenases (QDH) 16.2.5.1

General Remarks

Quinoproteins constitute a class of dehydrogenases distinct from the nicotinamideand flavin-dependent oxidoreductases They use different quinone cofactors to convert a vast variety of alcohols and amines into their corresponding carbonyl products [1701. Proteins containing the cofactor pyrroloquinoline quinone (PQQ) (Fig. 16.2-36)form the largest and best-characterizedsub-group. QDHs are independent from classical coenzymes like NAD(P)+.The substrate electrons are preferentially transferred to organic acceptors (quinones) and nonnative redox mediators such as phenazine derivatives, DCPIP, Wursters blue[171], 1731, osmium complexes or direct contact to an ferrocene[lO1],ferricyanide electrode[17'1. One advantage of the PQQ-dependent dehydrogenases over the NAD(P)-dependent dehydrogenases is the more positive redox potential of the PQQJPQQH, couple (+ 90 mV/pH 7[1761compared to - 320 mV[177,17811. Similarly to the flavin-dependent reactions, several mechanisms have been discussed, including covalent substrate-PQQ intermediates or hydride transfer [179-1811. The most important QDHs are methanol (alcohol) dehydrogenase (E. C. 1.1.99.8) and glucose dehydrogenase (E. C. 1.1.99.17), which will be discussed briefly.

1 G.2 Oxidation of Alcohols

I

1147

EP

Figure 16.2-38. A'-dehydrogenation of 6-a-methyl-hydrocortisone-21-acetate with polyurethane-entrappedArthrobacter simplex cells in buffer-saturated 1-decanol. The dehydrogenase (DH) activity is largely increased on addition o f quinoid electron acceptors (EA).

16.2.5.2

Methanol Dehydrogenase (E. C. 1.1.99.8) In addition to PQQ, the methanol dehydrogenases from Comamonas testosteroni and Gluconobacter suboxydans contain a heme group, which is indicated in their synonym quinohemoprotein dehydrogenase. The regeneration of these enzymes has been achieved by anodic reoxidation of ferricyanide [1731, 0s-modified anodes [1741, or even direct contact to the anode [17'1. Quinohemoprotein dehydrogenases (from Comamonas testosteroni and Gluconobacter suboxydans) have been reported to oxidize the alcohols solketal and glycidol (Fig. 16.2-37)enantio~electively['~~~. Alcohol oxidases from various strains, and especially NAD(P) dependent dehydrogenases (except HLADH together with thio-NAD+[1821), were found to be extremely inefficient for the oxidations in Figure 16.2-37,a fact, which is attributed to the significantly lower redox potential of the NAD(P)+/NAD(P)Hredox system[172]. The QDH from C. testosteroni was further characterized[1831. It oxidizes stereospecifically the (R) enantiomer of secondary alcohols. Both, kcat/KM and E increased with the substrate chain length. In vitro, ferricyanidewas used as sacrificial electron acceptor. In uivo, the excess electrons are most probably transferred to molecular

1148

I

IG

Oxidation Reactions

NAD+

NADH

R. eryfhropolis metabolism Figure 16.2-39. Enantiospecific oxidation of racemic carveol t o (-)-cawone and (-)-cis carveol using whole cells of Rhodococcus erythropolis.

oxygen via the respirator chain. This process is considerably accelerated (by a factor of 12) upon addition of external quinoid electron acceptors such as vitamin K (that are capable of autoregeneration) (Fig. 16.2-38)[ls41. 16.2.5.3 Glucose Dehydrogenase (E.C. 1.1.99.17)

So far, a m e m b r a n e - b o ~ n d and [ ~ ~ a~ soluble ] glucose dehydrogenase[l’G]have been identified. The latter oxidizes a wide range of mono- and disaccharides[’8G].In addition to cytochrome b562, regeneration with artificial acceptors such as DCPIP or ferr~cene[”~, 188] is effective and unproblematic, as no autoregeneration with molecular oxygen (producing reactive 0-species) is possible. It has commercial interest as a component of glucose test strips for diabetes c0ntrol[~”1. 16.2.6 Whole-Cell Oxidations 16.2.6.1

Stereoselective Oxidation of (-)-Carve01 to (-)-Carvone[’901

By using whole cells of Rhodococcus erythropolis DCL14, a racemic mixture of (-)-carve01was converted to (-)-carvone and (-)-cis-carveol(Fig. 16.2-39).The system was optimized using the two-liquid concept, in which a second organic phase serves as substrate and product reservoir. (-)-Carvone is an important flavor compound. The enzyme responsible for this bioconversion, catalyzed by wild-type cells of A high enantioseRhodococcus erythropolis DCL14, is carveol dehydrogena~e[~”I. lectivity and no further conversion of (-)-carvone was obtained. Carveol dehydrogenase has a broad substrate specificity and prefers substituted cyclohexanols as substrate^["^]. The regeneration of the cofactor NAD’ was accomplished by the use of living cells.

762 Oxidation ofAlcohols

$1

$ CH,OH

CH,OH

Enterobacter agglomerans

-

Figure 16.2-40. Production of the low calorie sweetener tagatose from D-galactitol by whole cells of Enterobacter agglomerans.

CH,OH

CH,OH

Gluconobacter oxydans

OH

LOH 1-amino-D-sorbitol

I

6-amino-L-sorbose

Figure 16.2-41. Oxidation of N-protected 1-amino-D-sorbitol to 6-amino-L-sorbose using Gluconobacter oxydans.

The use of a two-liquid phase system consisting of a 1:1 mixture of phosphate buffer and dodecane resulted in an increase of the initial (-)-trans-carveolconversion rate by 70% (to 26 nmol per minute and per mg protein). The production was increased from 4.3 to 208 pmol (-)-carvone formed per mg protein as compared to the aqueous system. A simple downstream process consisting of phase separation, methanol extraction, evaporation, and separation of (-)-cis-carveol and (-)-carvone over a silica gel column, was developed. In another study, Rhodococcus globendus PWD8 was found to oxidize D-limonene regio- and enantioselectivelyvia (+)-trans-carveolto (+)-carvone[”*]. 16.2.6.2 Sugar DehydrogenasesApplied in Whole Cells

Cofactor regeneration by the cell metabolism is the main advantage of whole cells in polyalcohol oxidations. The induction of whole-cell biocatalyst activity is dependent on the nature of the growth substrate. An example is the production of the low calorie carbohydrate sweetener tagatose from D-galactitol (Fig. 16.2-40). As biocatalysts, wild-type strains of Enterobacter agglomerans and Gluconobacter oxydans DSM 2343, in which sugar dehydrogenases catalyze the reaction of interest, were described 1193, 1941 In the case of Enterobacter agglomerans, cells growing on 1% glycerol plus 1% erythritol resulted in the best biocatalcc performance. In 30 h, galactitol (50 g/L) was converted with a tagatose yield of 86 %. Immobilization and storage at - 20 “C are possible. With Gluconobacter oxydans, growing cells were found to be more effective than resting cells. Furthermore, galactitol adaptation gave a notable increase in tagatose yield. Another example is the oxidation of 1-amino-D-sorbitol(N-protected)to 6-amino-~sorbose (Fig. 16.2-41)[’95! This reaction was published as a step in the synthesis of

1149

1150

I

IG Oxidation Reactions

fl-0

1

organic phase buffer

Figure 16.2-42. Selective oxidation of linear and branched aliphatic alcohols t o the corresponding aldehydes using P. pastoris i n aqueous/organic reaction mixtures.


6 /

Figure 16.2-43. Oxidation of benzylic alcohol t o benzaldehyde using whole cells o f P. pastoris in organic/aqueous emulsions or with purified alcohol oxidase. In vitro hydrogen peroxide was removed by catalase.

1-desoxynojirimycin.Derivatives of 1-desoxynojirimycinare pharmaceuticals for the treatment of carbohydrate metabolism disorders (e.g. diabetes mellitus). Suspended whole cells of Gluconobacter oxydans were used as the biocatalyst, in which D-sorbitol dehydrogenase is responsible for this biotransformation. To prevent undesired follow-upreactions of G-amino-L-sorbosein water, the amino group has to be protected by, for example, a benzyloxycarbonyl group (R).Cells are produced by fermentation on sorbitol and used for the bioconversion step as resting cells in water without added nutrients. The biotransformation is carried out by Bayer in a 10 000 L reactor with 90 % yield. 16.2.6.3

Oxidation of Aromatic and Aliphatic Alcohols to Corresponding Aldehydes and Acids

Havin-containing alcohol oxidase combined with catalase in peroxisomes of Pichia pastoris naturally catalyzes the oxidation of methanol to formaldehyde. In vivo and in vitro applications are possible. The alcohol oxidase has a broader substrate specificity than the subsequent enzymes of the methanol degradation pathway. Therefore,

16.2 Oxidation ofAfcohols

I

1151

Figure 16.2-44. Oxidation of alcohols by whole cells ofAcinetobacter. In aqueous media the oxidation proceeds until the acid stage, whereas the aldehyde is accumulated in the presence of organic solvents.

NAD+

NADH

Figure 16.2-45. Preparation of isovaleraldehyde using an alcohol dehydrogenase (ADH) in whole cells o f Cfuconobacter oxydans.

products other than formaldehyde are not degraded further. The spectrum of alcohols oxidized by whole cells of Pichia pastoris includes aliphatic Cl-Cs alcohols (saturated, unsaturated or branched). In biphasic media, Pichia pastoris also oxidizes c&1 alcohols, phenylethyl alcohol and 3-phenyl-1-propanol[1961. Up to 70 g/L acetaldehyde was produced from e t h a n ~ l [ ~ ” ~ Here, ” ~ ] . competitive product inhibition was partially overcome by high Tris buffer concentrations. Tris is able to bind acetaldehyde and markedly improve reaction yields. In a biphasic system consisting of 97 % hexane and 3 % aqueous phase, hexanol(l1 g/L) was converted to hexanal (Fig. 16.2-42)within 24 h at a yield of 9G%[1961. In another example, benzyl alcohol was oxidized by whole cells of Rchia pastoris and purified alcohol oxidase (Fig. 16.2-43)[2001.For this reaction the importance of solute partitioning in the biphasic reaction system was studied[201].With immobilized cells in organic (xylene)/aqueous media, benzaldehyde concentrations up to 30 g/L were reached in the organic phase[”*]. With purified alcohol oxidase, up to 45 g/L benzaldehyde was produced within 8 h and with an enzyme concentration of 0.94 g/L. Dehydrogenases of Acinetobacter and Gluconobacter strains catalyze the oxidation of various alcohols to corresponding aldehydes and acids in uivo. Substrates tested

NAD+

NADH

Figure 16.2-46. Preparation o f 2-phenyl acetaldehyde with Acinetobacter sp. in organic/aqueous emulsions.

1152

I

7 G Oxidation Reactions Table 16.2-12.

Oxidations catalyzed by Acinetobacter sp. in aqueous and biphasic medial27]. Water/isooctane (vol/vol l / l )

Water Substrate

T

O

Acid yield ["A]

Time [h]

Aldehyde yield

> 97

3

74

1

> 97

3

90

1

> 97

3

87

1

> 97

24

72

4

25

24

<5

24

H

)+=fOH

0"""

rO" > 97

90

45 min

> 97

93

45 min

> 97

77

45 min

<5

24

40 ((S)-alcohol: 95% ee)

3

r/.1 Time [h]

24

racemic 27 R. Gandolfi, N. Ferrara, F. Molinari, Tetrahedron Lett. 2001, 42, 513-514.

include ethanol, propanol, butanol, 2-methyl-1-butanol,3-methyl-1-butanol,l-pentanol, 1-hexanol, geraniol, 2-phenylethanol, 2-phenylthioethanol, cinnamyl alcohol, benzyl alcohol and (R,S)-2-phenyl-l-propanol (Tables 16.2-12 and 16.2-13)[202. 2031. The molecular structure of substrates and products as well as physicochemical conditions significantly influence bioconversions of short-chain aliphatic and aromatic alcohols into acids[204.2051 . Y. ields depend on the toxicity of the alcohol (different inhibitory concentrations for different alcohols), since product inhibition is often the major limiting factor L2O4]. Furthermore, dissolved oxygen concentrations and pH conditions are important factors for improving such bioconversions. Depending on strain and substrate (specificity of dehydrogenases), the reaction is directed to aldehyde or acid accumulation (Fig. 16.2-44).In principle, acid accumulation is favored in aqueous media, whereas aldehydes preferentially accumulate in biphasic media[203].

7G.2 Oxidation ofAlcohols

I

1153

Table 16.2-13.

Oxidations catalyzed by Gluconobacterasaii in aqueous and biphasic systems1271.

Water Substrate

T

O

Acid yield [%]

Time [h]

Aldehyde yield [%]

Time [h]

> 97

4

93

45 min

> 97

4

90

1

> 97

3

91

45 min

16

24

29

5

<5

24

<5

24

> 97

5

85

2

> 97

5

96

1

20

24

24

4

33

24

<5

24

H

>-3-'OH

0""" T

Water/isooctane (vol/vol 1/1)

O

H

racemic 27 R. Gandolfi, N. Ferrara, F. Molinari, Tetrahedron Lett. 2001,42, 513-514.

OH

OH

0 (3-2-phenylpropanoic acid

Figure 16.2-47.

Resolution of racemic (R,S)-2-phenylpropionic alcohol with whole cells of Gluconobacter oxydans yielding (S)-2-phenylpropanoic acid and (R)-2-phenylpropionic alcohol.

An example of aldehyde formation is the production of isovaleraldehyde by Gluconobacter oxydans R (Fig. 16.2-45) [202, 2061. Glycerol-grown Gluconobacter oxyduns slowly oxidizes 3-methyl-1-butanolto isovaleraldehyde,with yields of over 90%. The product was recovered by bisulphite trapping or cold Extractive bioconversion in a hollow-fiber membrane bioreactor allowed continuous produc-

1154

I

1G Oxidation Reactions

/

C.parapsilosis IF0 1396

P”,,H

(@-I ,3-butanediol

oxidative

\

:11267

-OH (S)-l,3-butanediol

Figure 16.2-48. Preparation of both enantiomers of 1,3-butanediol with whole cells of K. /actis and C.parapsilosis either by enantioselective oxidation o f 1,3-butanediol (oxidative) or enantioselective reduction of 4-hydroxybutanone (reductive).

tion of isovaleraldehyde at overall productivities of 2-3 g L-l h-’ 1206]. Yields between 72 and 90 % were reached. Another example of a synthesis is the production of phenylacetaldehyde using Acinetobacter strains (Fig. 16.2-46) 20sl. Different two-liquid phase systems were tested for their ability to remove the aldehyde into the organic phase before its further conversion to acid. In an optimized two-liquid-phase process, in which isooctane (at a volume fraction of 50%) was used as the organic carrier solvent, product concentrations of 9 g/L were reached in 4 h of reaction, corresponding to a yield of 90 %[208].The production strain Acinetobacter sp. ALEG showed satisfactory long-term stability, being able to perform the transformation with 80 % of the original activity after 3 days of contact with the solvent. Besides the multigram-scale production of different aliphatic carboxylic acids by biocatalytic alcohol oxidation, especially the enantioselective oxidation of racemic 2-phenyl-1-propanol to (S)-2-phenylpropanoic acid with Gluconobacter oxydans (Fig. 16.2-47)is another good example of acid production from alcohols~209~. After optimization of the parameters temperature, pH, substrate concentration, and agitation speed using a simplex sequential method, the resolution involving two oxidation steps yielded 45 % product with an ee of 98 %. 16.2.6.4 Enantiospecific Reactions

Two ways of producing optically pure 1,3-butanediol via microbial resolution have been reported: the oxidation of a racemic mixture of 1,3-butanediolyielding one enantio-

16.2 Oxidation ofAlcohols

n

OH

ADH

ethyl-4-chloro-3-oxobutanoate NADH

I

0

CI

0(R)-ethyl-4-chloro-3-hydroxybutanoate

NAD+

Asymmetric reduction of ethyl-4-chloro-3-oxobutanoate catalyzed by an alcohol dehydrogenase (ADH) in recombinant E. coli. The necessary reduction equivalents were derived from the oxidation o f isopropanol with the same enzyme. Figure 16.249.

K

Rhodococcus erythropolis

~

OH


COOH

Figure 16.2-50. Enantioselective oxidation of isopropylideneglycerol utilizing Rhodococcus erthyropolis.

mer and 4-hydroxy-2-butanone,and the reduction of the 4-hydroxy-2-butanone yielding one enantiomer of 1,3-butanediol (Fig. 16.2-48). (R)-1,3-butanediolis an important chiral synthon for the synthesis of various optically active compounds such as azetidinone derivatives, which are intermediates in the production of antibiotics, pheromones, fragrances, and insecticides. From a screening procedure, Kluyveromyces lactis I F 0 1903 and Candida parapsilosis I F 0 1396 were found to be effective in the enantioselectiveoxidation of (R)1,3-butanedioland (S)-1,3-butanediol,respectively, and in the asymmetric reduction of 4-hydroxy-2-butanone to (R)-1,3-butanediol and (S)-1,3-butanediol, respectively[2101. The equilibria between ketones and alcohols are catalyzed by secondary alcohol dehydrogenases. The secondary alcohol dehydrogenase of C. parapsilosis I F 0 1396

OH

-

0

enantiospecific oxidation

OH

+

RIAR2

racemic alcohol

RlAR2

desired enantiomer asymmetric reduction

R1

1

R2

desired enantiomer

Figure 16.2-51.

Stereoinversion catalyzed by two different alcohol dehydrogenases via enantiospecific oxidation followed by an asymmetric reduction.

1155

1156

I

7G Oxidation Reactions

was purified and characterized as an NAD’-dependent dehydrogenase with a broad substrate specificity (secondary alcohols > primary alcohols)r2111. The alcohol dehydrogenase gene of C. purupsilosis was cloned and expressed in recombinant E. coli JMlO9, which showed more than twofold higher specific alcohol dehydrogenase activity than C. purupsilosis[212]. Resting cells of C. parupsilosis were used for the large-scale (2000 L) production of (R)-1,3-butanediol(94% ee) from racemic 1,3-butanediol.After down-stream processing 3.1 kg product was isolated (overall yield: 15.5%), and a chemical purity of Table 16.2-14.

Biocatalytic stereoinversions with Ceotrichum candidum [**l.

With allyl alcohol (33 mM)

Without allyl alcohol Substrate

d

28

O

H

Yield [“A]

e e [“A]

Configuration

Yield [“A]

e e [“h]

Configuration

96

99

fR)

94

98

fR)

K.Nakamura, Y. Inoue, T. Matsuda, A. Ohno, Tetrahedron Lett. 1995,36.

6

16.2 Oxidation ofAlcohols

enantioselective oxidation

&j$ /

OH

c

& +

OH

OH

/

OH

0

OH

asymmetric reduction Figure 16.2-52.

Synthetic application of the stereoinversion concept using Candida sp.

and Pichia sp.

98.8 %was reached[213]. Resting cells of recombinant E. coli were reported to produce (R)-1,3-butanediol(93.5% ee, 94.7 % yield) from the racemate without any additive to regenerate NAD’ from NADHL2”I. In another application, recombinant E. coli produced 36.6 g/L ethyl-(R)-4-chloror2l0].Here, 3-hydroxybutanoate (99% ee) from 40 g/L ethyl-4-chloro-3-oxo-butanoate the secondary alcohol dehydrogenase served as both synthetic (asymmetric reduction) and regenerating (NADH-regeneration via isopropanol oxidation) enzyme (Fig. 16.2-49). Enzymatic resolution of (R/S) isopr~pylideneglycerol[~’~~ 2151 Whole cells of Rhodococcuserythropolis were used for the selective oxidation of the (S)enantiomer of isopropylideneglycerol (Fig. 16.2-50).With a 50 % conversion of the racemate, an ee value of over 98% was reached for (R)-isopropylideneglyceroland of over 90 % for (R)-isopropylideneglycericacid. (R)-Isopropylideneglycerolis a useful C3-synthon in the synthesis of (S)-P-blockers; e. g. (S)-metoprolol.(R)-Isopropylideneglycericacid can also be used as starting material for the synthesis of biologically active compounds. 16.2.6.5

Stereoinversions using Microbial Redox Reactionsp14

Racemic mixtures of secondary alcohols can be resolved completely by enantiospecific enzyme-catalyzed oxidation resulting in one enantiomer of the alcohol and the ketone followed by asymmetric enzyme-catalyzedreduction of the ketone (Fig. 16.251). For oxidation and reduction, two separate microorganisms [217-2191 or two may be used. different enzymes in a single microorganism[22&222] An example of a suitable biocatalyst is Geotrichum candidum, harboring both an oxidizing and a reducing enzyme activity. Table 16.2-14shows the catalFc performance of Geotrichum candidum towards different substrates [220] when the biocatalyst is incubated for 24 h with 27 mM substrate. Ally1 alcohol effectively shifts the stereoselectivityof the reduction. It is presumed to inhibit enzyme@)that reduce aryl

I

1157

1158

I

1G Oxidation Reactions

mo x

m-

OH

1

/

"OH

MO

chemical

Chemoenzymatic synthesis of 2-hydroxy-1-indanone.The racemic syn and anti diols were prepared by chemical dihydroxylation of indane. Asymmetric induction was achieved by microbial oxidation (MO) of these diols. Figure 16.2-53.

Table 16.2-15.

Substrate spedificity ofA\rthrobacter and Pseudomonas strains[29].

Taxonomy

Substrate specificity (no substrates)

Arthrobacter sp. strain 1HB

cis-(lS,2R)-diol > trans-(lS, 2s)-diol >> (cis-(1R, 2s)-diol, trans-(1R, 2R)-diol) cis-(lS,2R)-diol>>trans-(lS,2S)-diol>> (cis-(lR,2s)-diol, trans-(1R, 2R)-diol) trans-(IR, 2R)-diol>cis-(lS,ZR)-diol> cis-(IR, 2S)-diol>>(trans-(lS,2S)-diol)

Arthrobacter sp. strain 1HE Pseudomonas aeruginosa strain I N

29 Y. Kato, Y.Asano, /. Mol. Cut. B: Enzymatic 2001, 13, 27-36,

Table 16.2-16.

Microbial stereoselective oxidation of cis- and trans-l,2-indandiol~[~~1.

rh]

Strain

Substrate

Product

Reaction time [h]

Yield

Arthrobacter sp. 1HB

Cis Trans Cis Trans Cis Trans

R

4 12 4 24 5 24

46 35 47 8

> 99.9 > 99.9 > 99.9

7

82.5 > 99.9

Arthrobacter sp. 1HB

P. aeruginosa IN

S R S R R

40

e e [%]

> 99.9

29 Y. Kato, Y. Asano, /. Mol. Cut. B: Enzymatic 2001, 13, 27-36,

methyl ketone to (S)-1-arylethanol.The inhibition of yeast reductases by ally1 alcohols has been reported[223]. Another example is the deracemization of (RS)-1-{2',3'-dihydrobenzo[b]furan4'-yl}-ethane-1,2-diolby biocatalytic stereoinversion (Fig. 16.2-52)[2241. In order to find an appropriate biocatalyst to accomplish such a deracemization, different microorganisms were screened. Several microorganisms belonging to the genera Candida and Pichia allowed yields of 60-70% with 90-100% enantiomeric excess. Substrate dissolved in DMF was added to the biotransformation mixture consisting of resting cells suspended in phosphate buffer (pH 7). The presence of glucose generally increased the yield but lowered the enantiomeric excess. Different microorganisms can be suitable for a given stereoinversion and the optimal biocatalyst should be chosen by screening.

16.2 Oxidation ofAlcohols

I

1159

nu

cholesterol

oxidations androst-2-en-3,i 7-dione

asymmetric reduction

I

testosterone Selective oxidation o f cholesterol to testosterone by whole cells o f Mycobacteriurn sp NRRL 8-3805.

Figure 16.2-54.

Stereoselective oxidation of racemic 1,2-indandiol~[*~~1 Kato et al. described the stereoselective microbial synthesis of both enantiomers of 2-hydroxy-l-indanone,selecting cis- or trans-diol as the substrate (Fig. 16.2-53).Cis1-amino-2-indanolis an important synthon in organic chemistry (for example in the synthesis of the leading HIV protease inhibitor Crixivan) and can easily be synthesized from optically active 2-hydroxy-1-indanone[2261. Microorganisms degrading indane derivatives were screened for stereoselective oxidation of racemic cis- or trans-l,2-indandiol. Three promising strains specifically oxidizing the benzylic hydroxyl group were found (see Table 16.2-15). All strains produced inducible enzymes responsible for the oxidation reaction, recognizing the stereochemistry of the 1-or 2-positionsof the diol regardless of their cis and trans geometry. By using the resting cells of the strains, both enantiomers of 2-hydroxy-1-indanonewere synthesized in enantiomerically pure form simply by selecting cis- or trans-1,2-indandiol as the substrate. Growth conditions were optiactivity. mized to promote cell growth and the formation of 1,2-indanediol-oxidizing The biocatalyst activity was optimally induced with 0.05 % indanol. Carefully choosing appropriate carbon and nitrogen sources is crucial for optimal biocatalyst activity and cell growth. Table 16.2-16 shows the stereoselective oxidation of racemic cis-diol or trans-diol into optically active 2-hydroxy-1-indanoneat a 2 mL scale with 50 mg dry cells per ml.

1160

I

16 Oxidation Reactions

/

A

I

Figure 16.2-55. Regioselective three-step oxidation of ebastine (A) t o carebastine using Cunninghamella blakesleeana.

(B)

Production of testosteronefrom cholesterol using Mycobacterium sp. [2271 In this multistep reaction the microbial degradation of sterol side chains combined with the reduction of an intermediate thereof is used to accumulate testosterone from cholesterol. A cholesterol-assimilatingand androst-2-en-3,17-dione-accumulating mutant of Mycobacteriurn sp. NRRL B-3805 oxidizes cholesterol through multiple steps of the sterol side chain degradation pathway, also involving alcohol oxidations, to androst-2-en-3,17-dione(Fig. 16.2-54).This multistep oxidation is followed by the reduction of androst-2-en-3,17-dioneto testosterone by the NADH requiring activity of 17P-hydroxysteroiddehydrogenase. This activity is dependent on the presence of glucose as the carbon source. After the glucose in the fermentation culture is completely consumed, most testosterone is oxidized to androst-2-en-3,17-dione. Adding a larger amount of glucose prevents this oxidation. On a 2.5 L scale a yield of 51 %was reached in 120 h of cultivation. Here, the initial substrate concentration amounted to 0.1 % (w/v). Microbial oxidation of ebastine [228] Ebastine is a new generation antihistaminic drug with fewer side-effects. The microbial three-step oxidation of ebastine, using whole cells of the mold Cunninghamella blakesleeana as biocatalysts, involves an alcohol and an aldehyde oxidation step and results in the formation of carebastine, which is the pharmacologically active The initial step in the oxidation of ebastine is hydroxylation by a cytochrome P-450-dependent monooxygenase to the corresponding alcohol. The two consecutive oxidations are catalyzed by oxidoreductases,which are not further characterized, and lead via the aldehyde to the corresponding carboxylic acid carebastine (Figure 16.2-55). Growth in a complex medium containing soybean-peptone and yeast extract is necessary for biocatalyst activity. A component of soybean-peptone, genistein, is thought to act as an inducer of cytochrome P-450 enzymes. Growing cells provide a higher yield than resting cells. Addition of 1% poly(viny1 alcohol) was found to prevent pellet formation and thereby to guarantee constant mass transfer rates. From a 3 L batch fermentation, 270 mg carebastine was isolated (yield: 45%).

16.2 Oxidation ofAlcohols

\f-

co2

NADH

-

FDH

Q ~

Diaphorase

Enzymatic three-step oxidation o f methanol t o carbon dioxide in the anodic compartment o f a biofuel cell.

Figure 16.2-56.

N

A

HO'"

"OH

-P0s=

E B Figure 16.2-57. Mediated electron transfer steps in the electroenzymatic oxidation o f glucose (A) and reduction o f 02.

I

P1

1162

I

IG

Oxidation Reactions

Therefore, after 24 h of cultivation, GOO mg ebastine was added and the incubation was continued for 68 h. . 16.2.7 Miscellaneous 16.2.7.1 Biofuel Cells

In recent years, biofuel cells have gained tremendous attention. The use of methanol instead of dihydrogen as the oxidizable substance offers special advantages as it is readily available and easy to store and handle. At the same time, the theoretical cell voltage of an MeOH/02 cell (1.19 V) is near that of H2/02 (1.23V). Whitesides and coworkers recently developed a biofuel cell based on the step-wise In the anodic enzymatic oxidation of methanol to carbon dioxide (Fig. 16.2-56)[2301. compartment of the biofuel cell, methanol is oxidized to carbon dioxide in three steps: by an alcohol dehydrogenase, an aldehyde dehydrogenase, and ultimately formate dehydrogenase. In each of these enzymatic steps, one equivalent of NADH is produced. NADH itself transfers its electrons via diaphorase to viologene and in the end to the anode. The redox potential of the reducedloxidized viologene couple (- 0.55 V) is only slightly less negative than MeOH/C02 (- 0.64 V) and NADH/ NAD' (-0.59 V). Thus, the loss in cell potential was minimized. The catholyte consisted of platinum gauze in an 02-saturated buffer ( 0 2 + 4e-+4Hf 2H20). An open-circuit potential of 0.8 V and a maximum power output of 0.67 mW cm-2 was achieved. Another biofuel cell concept is based on the oxidation of glucose to gluconolactone catalyzed by glucose oxidase (Fig. 16.2-57)[231. 2321. Because of the slow kinetics of the electron transfer to 02,dioxygen is usually reduced at a potential several hundred millivolts more negative than its formal potential, thus lowering the power density of a fuel cell. Utilizing laccase to catalyze this reaction can circumvent that. ABTS is a suitable mediator between the electrode and laccase because of its quite positive redox Wiring laccase reduction to the electrode via an osmiummodified electrode also facilitates the electroreduction of molecular oxygen. The same modification serves as the conductor between glucose oxidase and the anode. +

-PEG

-V Figure 16.2-58.

OH

NAD modified with polyethylene glycol (PEG).

OR

16.2 Oxidation ofAlcohols Table 16.2-17.

Kinetic constants of different dehydrogenases for NAD(P)' and PEG-NAD(P)+. Native cofador

NAD'-dependent enzymes

KM

FDH YADH HLADH LDH 3a-HSDH Glucose DH

15 175 154 62 182 29 9G

NADP+-dependentenzymes

KM [@I

Glutamate DH Malic enzyme

160 5

Glutamate DH

13

TBADH a 100% correspond to

V,

PEG-bound cofador

KM [@I

VmaX[as % o f NAD']a

82

57

444

53

1310 1150

64 72 21 66 3

142 647 2030 KM

425 12 28

bM1

V,,,

[as % o f N A D V ]

96 86 84

values of the dehydrogenases determined with native coenzymes.

A miniaturized cell was constructed which exhibited a power output of 0.137 mW cm-2. After 72 h of operation, 75 % of the initial power output was still present. Even though biofuel cells are generally considered to be in their infancy[234], their potential, which is based on non-hazardous, easy-to-handlesubstrates and electrolytes (especiallythe moderate temperatures compared to those of conventional fuel cells: 80-1000 "C) cannot be neglected. Even photosynthetic biofuel cells (converting light energy into electrical energy) have been shown to work in principle[235]. 16.2.7.2

Biomimetic Analogs to NicotinamideCoenzymes

For large-scale applications of NAD(P)-dependent enzymes, continuous-flow reactors with ultrafiltration membranes have been proposed[236]. In order to retain low molecular weight nicotinamide cofactors in the reactor, charged membranes have been used, retarding the overall negatively charged nicotinamide coenzymes by electrostatic repulsion[237,2381. Retention rates of approx. 99% and TTNs (NAD) of up to 10 000 were reported. Another approach makes use of polymer-modified NAD [modification with polyethylene glycol (PEG; MW = ZOOOO)], thus retaining it on account of its The polymer modification usually drastically increased size (Fig. 16.2-58)[239-2411. leads to a drastically increased KM value, whereas the V,, value is generally over 50% of that of low molecular weight NAD(P) (Table 16.2-17). Another area of research deals with synthetic analogs of NAD(P) coenzymes. Besides the lower costs, these analogs may offer better stability or easier regeneration and may add new functionalities to known enzyme systems (e.g. thio-NAD together with HLADH [ls2I). Some artificial redox coenzymes were developed mim244]. Activity icking the "shape" of native nicotinamide coenzymes (Fig. 16.2-59) with various NAD-dependent enzymes was found, even though the activity was only

1164

I

7G Oxidation Reactions Figure 16.2-59.

Synthetic analogs

of NAD.

so3CL4

blue N-3

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S.J. B. Duff, W. D. Murray, Biotech. Bioeng. 1988, 3 1 , 6 4 9 . 198 S. J. B. Duff, W. D. Murray, Biotech. Bioeng. 1988,31,790-795. 199 S. J. B. Duff, W. D. Murray, R. P. Overend, Enz. Microb. Tech. 1989, 11, 770-775. 200 S. J. B. Duff, W. D. Murray, Biotech. Bioeng. 1989,34,153-159. 201 K. Kawakami, T. Nakahara, Biotech. Bioeng. 1994,43,91&924. 202 F. Molinari, R. Villa, M. Manzoni, F. Aragozzini, Appl. Microbiol. Biotech. 1995, 43, 989-994. 203 R. Gandolfi, N. Ferrara, F. Molinari, Tetrahedron Lett. 2001,42, 513-514. 204 J. Svitel, P. Kutnik, Lett. Appl. Microbiol. 1995,20,365-368. 205 J. Svitel, E. Sturdik, Enz. Microb. Tech. 1995, 17,546-550. 206 F. Molinari, F. Aragozzini, J. M. S. Cabral, D. M. F. Prazeres, Enz. Microb. Tech. 1997, 20,604-611. 207 M. Manzoni, F. Molinari, A. Tirelli, F. Aragozzini, Biotech. Lett. 1993, 15, 341-346. 208 F. Molinari, R. Villa, F. Aragozzini, R. Leon, D. M. F. Prazeres, Tetrahedron Asym. 1999, 10,3003-3009. 209 F. Molinari, R. Gandolfi, F. Aragozzini, R. Leon, D. M. F. Prazeres, Enz. Microb. Tech. 1999,25,729-735. 210 A. Matsuyama, H. Yamamoto, N. Kawada, Y. Kobayashi,J. Mol. Cat. B: Enzymatic 2001, 11,513-521. 211 H. Yamamoto, N. Kawada, A. Matsuyama, Y. Kobayashi, Biosci. Biotech. Biochem. 1995, 59,1769-1770. 212 H. Yamamoto, N. Kawada, A. Matsuyama, Y. Kobayashi, Biosci. Biotech. Biochem. 1999, 63,1051-1055. 213 A. Matsuyama, Y. Kobayashi, Biosci. Biotech. Biochem. 1994,58, 1148-1149. 214 M. A. Bertola, H. S. Koger, G. T. Phillips, A. F. Man, V. P. Claassen. A process for the preparation of (R)- and (S)-2,2-R1,R21,3-dioxolane-4-methanol: EP, 1987. 215 A. Liese, K. Seelbach, C. Wandrey, Oxidase of Rhodococcus erythropolis. In Industrial Biotransformations. A. Liese, K. Seelbach, C. Wandrey (eds),Wiley-VCH,Weinheim, 2000, pp. 163-164. 216 A. J.Camell, Adv. Biochem. Eng. Biotech. 1999,63, 57-72. 217 G. Fantin, M. Fogagnolo, P. P. Giovannini, 197

A. Medici, P. Pedrini, Tetrahedron Asym. 1995,6,3047-3053. 218 E. Takahashi, K. Nakamichi, M. Furui,J. Fern. Bioeng. 1995,80,247-250. 219 S . Shimizu, S. Hatori, H. Hata, H. Yamada, Enz. Microb. Tech. 1987, 9, 411-416. 220 K. Nakamura, Y. Inoue, T. Matsuda, A. Ohno, Tetrahedron Lett. 1995,36. 221 M. Takemoto, K.Achiwa, Tetrahedron Asym. 1995,6,2925-2958. 222 J. Hasegawa, M. Ogura, S. Tsuda, S. I. Maemoto, H. Kutsuki, T. Ohashi, Agrc. Bid. Chem. 1990,54,1819-1828. 223 K. Nakamura, K. Inoue, K. Ushio, S. Oka, A. Ohno, Chem. Lett. 1987,4,679-682. 224 A. Goswami, K. D. Mirfakhrae, R. N. Patel, Tetrahedron Asym. 1999, 10,4239-4244, 225 Y. Kato, Y. Asano, J . Mol. Cat. B: Enzymatic 2001, 13,27-36. 226 H.Kajiro, S. Mitamura, A. Mori, T. Hiyama, Bull. Chem. Soc.]pn. 1999,72, 1093-1100. 227 W.-H. Liu, C.-K. Lo,J. Ind. Microbiol. Biotechnol. 1997, 19, 269-272. 228 H. Schwartz, A. Liebig-Weber, H. Hochstatter, H. Bottcher, Appl. Microbiol. Biotech. 1996,44,731-735. 229 M. Matsuda, M. Sakashita, Y. Mitsuki, T. Yamaguchi, T. Fujii, Y. Sekine, ArzneimittelForschung 1994,44, 55-59. 230 G.T.R. Palmore, H. Bertschy, S. H. Bergens, G. M. Whitesides, J. Electroanal. Chem. 1998,443,155-161. 231 S . C.Barton, H.-H. Kim, G. Binyamin, Y. Zhang, A. Heller,J. Am. Chem. SOC.2001, 123,5802-5803. 232 T. Chen, S.C. Barton, G. Binyamin, 2. Gao, Y. Zhang, H.-H. Kim, A. Heller,J. Am. Chem. Soc. 2001,123,8630-8631. 233 G. Tayhas, R. Palmore, H.-H. Kim, J. Electroanal. Chem. 1999,464,110-117. 234 J. St-Pierre, D. P. Wilkinson, AIChEJ. 2001, 47,1482-1486. 235 S.Tsujimura, A. Wadano, K. Kano, T. Ikeda, Enz. Microb. Tech. 2001, 29, 225-231. 236 A. F. Biickmann, G. Carrea, Adv. Biochem. Eng. Biotechnol. 1989, 39, 97-152. 237 B. Nidetzky, W. Neuhauser, D. Haltrich, K. Kulbe, Biotech. Bioeng. 1996, 52, 387396. 238 J.M. Obon, M. J. Almagro, A. Manjon, J. L. Iborra, J . Biotechnol. 1996, 50, 27-36. 239 E. Steckhan, S. Herrmann, R. Ruppert, J. Thoemmes, C. Wandrey, A n g m . Chem. Int. Ed. Engl. 1990, 29, 388-390.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

1170

I

240

1G Oxidation Reactions

P. Pasta, G. Carrea, N. Gaggero, G. Grogan,

A. Willetts, Biotech. Lett. 1996, 18, 1123-1128. 241 G. Ottolina, G. Carrea, S. Riva, Enz. Microb. Tech. 1990, 12, 596-602. 242

R. 7. Ansell, C. R. Lowe, Appl. Microbiol. Biotech. 1999,51, 703-710.

Dilmaghanian, C. V. Stead, R. J. Ansell, C. R. Lowe, Enz. Microb. Tech. 1997,20,

243 S.

165-173. 244

R. J. Ansell, D. A. P. Small, C. R. Lowe, J. Mol. Cat. B: Enzymatic 1999, 6 , 111-123.

16.3 Oxidation of Phenols Andreas Schmid, Frank Hollmann, and Bruno Buhler 16.3.1

Introduction

Several classes of oxidoreductases accept phenols and their derivatives as substrates for oxidation reactions. A broad range of products can be obtained depending on the substrates and enzymes applied (Fig. 16.3-1). Several monooxygenases catalyze the hydroxylation of the aromatic ring specifically ortho or para to the existing phenolic alcohol function (Fig. 16.3-1 A). Oxidases can be used to catalyze the stereospecific benzylic hydroxylation of aliphatic side chains to (R) or (S) alcohols and the further oxidation of benzylic alcohols to corresponding ketones or aldehydes; furthermore, elimination to (2)or (E') alkenes can be obtained if desired (Fig. 16.3-1 B). Laccases and peroxidases generate phenoxy radicals which - depending on the reaction conditions - can react further with phenols to structurally complex dimers or conducting polymers (Fig. 16.3-1 C). Even nitration reactions are reported (Fig. 16.31 D). Thus, enzymatic modification opens up new possibilities for synthetic chemistry with aromatic compounds under mild and non-toxic conditions. 16.3.2

Oxidases 16.3.2.1

Vanillyl-alcohol oxidase

(E.C. 1.1.3.38)

The enzyme vanillyl-alcohol oxidase (VAO, E.C. 1.1.3.38) was examined in detail with respect to mechanism, structural properties, and biotechnological applications by van Berkel and coworkers, giving an excellent example of how detailed biochemical studies provide a basis for preparative biocatalytic applications (for recent reviews see['. '1). The homooctamer with a monomer mass of 65 kDa was isolated and purified from Penicillium simplicissimum. The catalytic mechanism of VAOcatalyzed oxidation of para-alkyl phenols was studied in detail r3-'1. After initial hydride abstraction from the Ca atom, a binary complex of the intermediate paraquinone methide and reduced FAD reacts with molecular oxygen, regenerating the

76.3 Oxidation of Phenols

I

1171

I--\

HO

/

OMe

R

R

Figure 16.3-1. Enzyme-catalyzed oxidations of phenols. A ortho- and para-hydroxylations catalyzed by monooxygenases (Sects. 16.3.3.2and 16.3.6.2);B: oxidation at the benzylic position catalyzed by oxidases (Sects. 16.3.2.1and 16.3.5);C:coupling reactions catalyzed by peroxidases and laccases (Sects. 16.3.4.1and 16.3.2.2);D: nitration reactions catalyzed by peroxidases (Sect. 16.3.4.3).

oxidized prosthetic group. Depending on the nature of the aliphatic side chain, the para-quinone methide is hydroxylated to (chiral) benzylic alcohols (short aliphatic side chains) or rearranges yielding benzylic alkenes (long aliphatic side chains) (Fig. 16.3-2).Table 16.3-1 shows a selection of reactions catalyzed by VAO as well as the kinetic constants thereofl3, 1'. OH

- r" rR

HOq

-

R

0, + H,O

R = short chain

0

HO

VAO-FAD

VAO-FADH

% 0

HzO2

/o^'"

HO

cis+trans Figure 16.3-2.

Reaction mechanism ofvanillyl oxidase (VAO).

R = medium chain

1172

I

16 Oxidation Reactions Table 16.3-1.

Substrate spectrum and kinetic constants of vanillyl oxidase.

Substrate

KM [ W ]

kat

9

2.5

280

4

4.2

1050

6

4.9

820

16

1.3

81

72

0.5

7

2

1.2

600

100%alkene

8

0.3

38

100% alkene

42

< 0.001

< 0.02

65

1.4

21

77

0.5

7

HO

16% alcohol 60%ketone 24 % alkene

94

0.7

7

HO

4% alcohol 2 % ketone 94% alkene

Product(s)'

[S-'1

k,t/K~

76 % alcohol

/o^

HO

24% alkene 68 % alcohol

Xy-

HO

32 % alkene 90 % alcohol

HO

10% alkene

dd OMe

20 % alcohol 80 % alkene

HO

26 % alcohol 74 % alkene

HO

1 % alcohol

-o^"

r

HO

99 % alkene

HO

10^"/"

HO

40% HO HO

[W3][s-' M-'1

16.3 Oxidation ofphenols Table 16.3-1.

(cont.)

Substrate

Product(s)"

K, [pu] kcat[sd]

kcot/Khn [lo-'] [s-' M-'j

100 % ketone

222

0.7

3

100 % ketone

4.9

13.0

2700

4.8

6.5

1400

290

5.4

19

240

1.3

5.4

5.3

82

QH

/o^

HO

q-

HO

bMe

/o^"

HO

HO&OH

qJ- -9""

HO

HO

OMe

OMe

HO

-0""

HO

/o"""'

HO

bMe

OMe

$yo

HO

65

a Beside s the structure shown the products formed include benzylic alcohols, benzylic alkenes and benzylic ketones.

VAO exhibits a remarkable activity towards 4-alkylphenols, bearing aliphatic side chains of up to seven carbon atoms. The maximum chain-length of 7 is in accordance with structural data obtained from X-ray crystallography"1. Short-chain 4-alkylphenols are mainly hydroxylated at the Ca position, whereas medium-chain 4-alkylphenols are dehydrogenated to 1-(4'-hydroxyphenyl)alkenes (Fig. 16.3-2)['I. The hydroxylation reaction is highly stereospecific, producing the (R)-enantiomer with ee values of up to 94 % 1'1. Furthermore, VAO also catalyzes the further oxidation of the alcohols to the corresponding ketones. Here, the VAO-catalyzed oxidation of (S)-alcoholsis far more efficient than the oxidation of (R)-alcohols,promoting a possible application in kinetic resolution reactions. Substrates with more spaceconsuming alkyl side chains are dehydrogenated by the action of VAO. With paramethyl phenols (e.g. cresol), a very low conversion rate is found which is due to the formation of a stable intermediate formed through a nucleophilic attack of the reduced FAD on the para-quinone methide, yielding a covalent bond[']. Since the rate-limiting hydrolysis of this intermediate is acid-catalyzed,the pH optimum of the reaction shifts from alkaline to acidic values. The formation of such a covalent

I

1173

1174

I

7G Oxidation Reactions

liver microsmomes

HO

/

OMe capsaicin

qNH2 VAO

HO

-c

HO

OMe

OMe vanillin

Figure 16.3-3. Potential biotechnological production route to vanillin from natural components with vanillyl oxidase.

intermediate is supposed to be more unlikely with increasing length of the aliphatic side chain, because of increasing steric hindrance. Much attention has been paid to the shift from hydroxylation to dehydrogenation with increasing length of the side chain. The product ratio between alcohols and alkenes is strongly influenced by the extent of hydratation of the intermediate, paraquinone methide. Thus, by using organic media with a low water content the overall alkene yield could be significantly increased. The same is true for monovalent anions such as CP, Br-, or SCN-, which bind to the active site, thereby decreasing the water concentration at the active site[']. By enzyme engineering based on the threedimensional structure [71, the ratio between hydroxylation products and dehydrogenation products could be shifted either in favor of the alcohols, when Asp170 was exchanged with Glu, or in favor of the alkenes, when Asp170 was exchanged with Ser["I. Double mutants of VAO (D170S/T457E and D170A/T457E) were produced based on the same rational approach, thus inverting the stereospecificity of the VAOcatalyzed hydroxylation of 4-ethyl phenol from (R) to (S) (ee = 80%) [I1]. The VAO-catalyzed production of vanillin is of special synthetic interest. In particular, a route starting from capsaicin that is readily available from red hot pepper has some biotechnological potential. Here, vanillylamine is obtained by hydrolysis of capsaicin using rat liver microsomes and further oxidized by VAO (Fig. 16.3-3). Furthermore, a one-pot synthesis using carboxylesterase for capsaicin hydrolysis is proposed [''I. 16.3.2.2

Laccase (E. C. 1.10.3.2)

Recently, laccases found some interest for synthetic application. Laccases are widely distributed in plants and fungi['3].The copper-containing enzymes are some of the few oxidases so far reported to reduce molecular oxygen to water (aside from cytochrome c oxidase and others). This ability was recently exploited in a novel regeneration concept for flavin-dependentenzymes (see Chapter 16.2) [I4]. Purified laccase oxidizes various phenolic compounds via hydrogen abstraction. The resulting phenoxy radical undergoes various dimerization and oligomerization reactions. Even though the synthetic potential of such reactions has to be considered as moderate, in some cases interesting products (such as complex coumaran type compounds) can be obtained in reasonable yields from simple phenols["1. Laccases alone are not able to oxidize benzyl alcohols. Bourbonnais and Paice["]

16.3 Oxidation ofPhenols

I

1175

Table 16.3-2.

Laccase/ABTS-catalyzed oxidations to corresponding aldehydes.

Catalned reaction

P\ O / "

-

Po \ /

O

H

-

Po

CI

94

PI

92

PI

92

PI

89

[21

CI

\ /

*OH-

Literature

Me0

Me0

P

Yield I"?]

\ /

-0

-0

p-p0 CI

CI

1 A. Potthast, T. Rosenau, C. L. Chen, I. S . Gratzl, I . Mol. Cat. A.: Chemical 1996,108, 5-9.

2 A. Potthast, T. Rosenau, C. L. Chen, J. S. Gratzl,

I. Org. Chem. 1995,60,432&4321.

were the first to report that laccase in the presence of a specific compound, usually called a "mediator", is able to catalyze the oxidation of benzyl alcohols. Mostly ABTS (2,2'-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid), HOBT (l-hydroxybenzotriazole) [l71,and NHAA (N-hydroxyacetanilide)[18] have been used as mediators so far. The actual role of the mediator is not yet fully understood, although Potthast et al. recently found evidence that laccase produces reactive radical species of ABTS and

1176

I

7G Oxidation Reactions

* R

creolase activity

-OH

0 2

II I RwoH catecholase activity n un ” I,

Oxidation of phenols catalyzed by tyrosinase displaying so-called creolase and catecholase activities. Figure 16.3-4.

HOBT, which perform the actual oxidations [”]. Nevertheless, some preparative oxidations of various benzylic alcohols are reported (Table 16.3-2). It should be pointed out here that the laccase-mediator system still is far from being economically feasible. 16.3.3 Monooxygenases 16.3.3.1 Tyrosinase (E.C. 1.10.3.1)

Tyrosinases (synonyms: phenol oxidases, poly-phenolases or polyphenol oxidases) are copper-containing monooxygenases, which catalyze two consecutive reactions with molecular oxygen as cosubstrate, namely the ortho-hydroxylation of phenols and the oxidation of the resulting catechols to ortho-quinones (Fig. 16.3-4). The initial (phenol-hydroxylating)activity is usually referred to as creolase activity, whereas the second (catechol-oxidizing)activity is most commonly called catecholase activity[”]. The classification of tyrosinases (polyphenol oxidases) is somewhat ambiguous; enzymes exhibiting monophenol oxidase activity are classified as E. C. 1.14.18.1., but those with catechol oxidase activity as E. C. 1.10.3.2. However, many enzymes exhibit both activities, and a more appropriate classification of all twoelectron-acceptingcopper monooxygenases as E. C. 1.14.18.1was proposed[”]. In animals, tyrosinase is involved in the formation of melamines, and in plants, tyrosinase leads to the well-known browning of open surfaces of Much attention has been paid to the mechanism[20,22]. In the active site, two copper(1) ions bind molecular oxygen. Upon binding of the phenolic substrate, the ortho-position is attacked electrophilically by one of the activated oxygen atoms. The resulting copper-bound catechol serves as an internal electron donor and leaves the active site as ortho-quinone.Figure 16.3-5 illustrates this mechanism. In order to prevent rapid quinone polymerization in aqueous media, the quinones are usually reduced to the catechols (most commonly by ascorbic acid) (Fig. 16.3-6). Several tyrosinase-catalyzed oxidations of phenols have been reported; some of these are presented in Table 16.3-3. Tyrosinase was reported to hydroxylate and oxidize tyrosine residues in proteins [23], which is important in the production of moisture-resistant adhesives. In fact, tyrosinase has been used for the production of synthetic glues with similar compositions to those of naturally occurring adhesives such as mussel glue [24]. An interesting cascade reaction was reported by Waldmann et al.[252 2 G ] . Tyr-

16.3 Oxidation ofPhenols

I

1177

0 2

,cut

-

+/

\

Q

cu

\

0

0

\ /2+

f

,C",

0 \

:2

o/cy

Figure 16.3-5. Reaction mechanism for the oxidation o f phenols by tyrosinase.

OH

HO*

o HO

Ho*o

-

0

OH

1

PH

L

0

Figure 16.3-6. Ascorbic aciddriven reduction o f quinones.

R

A

Figure 16.3-7. Chemoenzymatic Diels-Alder reactions. Ortho-quinones (dienes), derived from phenols by oxidation with tyrosinase, spontaneously react with dienophils.

osinase, immobilized on glass beads, was used to oxidize several phenols in chloroform as the organic medium. The products of the enzymatic oxidation step, the ortho-quinones, served in situ as dienes in a Diels-Alder reaction (Fig. 16.3-7). Table 16.3-4 summarizes some phenols (dienes after enzymatic oxidation) and dienophiles with which such a reaction cascade was observed.

1178

I

16 Oxidation Reactions Table 16.3-3.

Oxidations of phenols catalyzed by tyrosinase.

Substrate

Product

References and remarks

Electron-rich phenols are preferred131

---b R

R

R = OCH3, OCzHs, CH3, C(CH3)3,Halogen, etc.

-

OH

,O/\bNHz HO

(4, 51 HO

R = H, CH3

L-DOPA production[6] HO

HO

NH, H

°

F

O

H

Possible agent in melanoma treatment 1'. 8l

HO

HO

F

F

OH

Coumestans L91

OH

S . Passi, M. Nazzaro-Porro, Brit. /. Dematol. 1981, 7 M. E. Rice, B. Moghaddam, C. R. Creveling, K.R. Kirk, Anal. Chem. 1987,59, 1534. 104,659. 8 R. S . Phillips, 1.G. Fletscher, R. L. Von Tersch, M. Jimenez, F. Garcia-Carmona, F. Garcia-CanoK. L. Kirk, Arch. Biochem. Biophys. 1990,276, vas, 1. L. Iborra, ].A. Lozano, F. Martinez, Arch. Biochem. Biophys. 1984,235,438. 65. M. Jimenez, F. Garcia-Carmona, F. Garcia-Cano9 U.T.Bhalearo, C. Muralikrishna, G. Pandey, vas, J. L. Iborra, ].A. Lozano, Int. J. Biochem. 1985, Synth. Commun. 1989,19,1303. 17,891. 10 0. Toussaint, K. Lerch, Biochem. 1987,26, 8567. G . M. Carvalho, T. L. M. Alves, D. M. G. Freire, Appl. Biochem. Biotech. 2000,84-86,791-800.

By this reaction sequence, highly functionalized bicyclo-[2.2.2]-octenescan be obtained from simple phenols and alkenes as starting materials. The overall yields reported are usually satisfactory (> 70%). The Diels-Alder products are racemic, probably because the Diels-Alder reaction proceeds in the bulk organic phase without involvement of tyrosinase.

16.3 Oxidation ofphenols Table 16.3-4. Substrates for the reaction cascade including tyrosinase catalyzed oxidation of ohenols and a Diels-Alder-reaction[ ' l , '*I.

Phenols

Dienophiles

Via a tyrosinase catalyzed reaction the phenols are transformed to dienes, which subsequently react with the dienophiles in a Diels-Alder-reactionas shown in Figure 16.3-7. 11 G. H . Miiller, H. Waldmann, Tetrahedron Lett. 1996,37,3833-3836.

12 G. H . Miiller, A. Lang, D. R. Seithel, H. Wald-

mann, Chem. Eur.]. 1998,4,2513-2522.

16.3.3.2 2-Hydroxybiphenyl-3-monooxygenase(HbpA, E. C. 1.14.13.44)

The flavin-dependent,homotetrameric HbpA is the first enzyme in the biodegradation pathway of 2-hydroxybiphenylin Pseudomonas azelaica HBPl L2'1. HbpA catalyzes the selective ortho-hydroxylation of a broad range of phenols to the corresponding catechols, utilizing NADH as cofactor (Fig. 16.3-8and Table 16.3-5). Compared to the chemical synthesis of ortho-substitutedcatechols (ortho-hydroxysuch an enzymatic approach is superior lation and aromatization procedures)[28-311, with respect to the number of steps involved as well as simplicity, selectivity, and yield. The resulting ortho-substitutedcatechols are valuable building blocks [321. HbpA is an excellent example of in vivo as well as in uitro biocatalysis. Since the desired catechols are rapidly degraded via the D. azelaica rneta-cleavage pathway by two catechol-2,3-dioxygenases, the gene coding for HbpA was expressed in E. coli JM109, which served as a biocatalyst accumulating the desired products [331. Drawbacks such as inhibition by substrate and product can be overcome by continuous substrate feeding and in situ recovery of the catechol products with solid adsorbents 0 2

NADH

NAD'

R = Ph, 2-OH-Ph, 2,3-(OH),Ph, F, CI, Br, Me, Et, Pr, iPr, But Figure 16.3-8. Reaction scheme for the ortho-hydroxylation o f phenol derivatives catalyzed by 2-hydroxybiphenyl-3-monooxygenase (HbpA).

I

1179

1180

I

IG Oxidation Reactions Table 16.3-5.

Substrates and relative activities o f 2-hydroxybiphenyl-3-monooxygenase

(HbpA) [131. Substrate

Product

dp HO

69

Relative activity I"?]"

HO

100

HO

34 HO

OH

HO

OH

49 Native substrate OH

O "&

24

OH

3G OH

10 OH

Ho$cl

&

20

HO

33

a Relative activities were determined polarographicallywith whole cells of recombinant E. coli containing

HbpA. 100%corresponds to the HbpA-dependent specific oxygen uptake rate of whole cells incubated with 2,2'-dihydroxybiphenyl.

13 A. Schmid, H:P. E. Kohler, K.-H. Engesser,]. Mol. Cat. B: Enzymatic 1998,5,311-316

in such a way that substrate and product concentrations can be kept below toxic levels [321. Thus, several 3-substituted catechols were produced in gram amounts with satisfactory to high yields (Table 16.3-6). The in uivo processes are based on a recombinant E. coli as catalyst[33].Optimized space-time yields of up to 0.39 g L-' h-' for the formation of 3-phenyl catechol from 2-phenyl phenol can be reached[34]. The enzyme itself was purified and characterized in 351. Based on this knowledge and via directed evolution, HbpA characteristics were modified (Meyer, Schmid and Witholt, unpublished results) yielding HbpA variants with improved

7 G.3 Oxidation of Phenols Table 16.3-6. Preparative-scale production of 3-substituted catechols using E. coli J M l O l containing 2-hydroxybiphenyl-3-rnonooxygena~e~’~]. Product

Product recovered 191

Molar yield

Ho&

8.1

94

2.1

95

0.6

71

2.2

77

1.7

85

0.9

71

2.1

71

&

HO

HO

OH

HO,

,OH

HO

OH

M

14 M. Held, W. Suske, A. Schmid, K. Engesser, H. Kohler, B. Witholt, M. Wubbolts, j.Mol. Cut. B: Enzymatic

1998,5,87-93.

catalytic properties and changed substrate spectrum. For example, a new mutant with drastically decreased unproductive NADH oxidation and concomitant formation of hydrogen peroxide was developed. This so-calleduncoupling reaction is quite common amongst flavin-dependent monooxygenases, and represents the major mechanism of autoregeneration amongst oxidases. Furthermore, the activity toward several substrates that are poorly converted by native HbpA, such as 2-sec-butylpheno1 (30 % activity increase), 2-tert-butylphenol (fivefold activity increase) or guaiacol (more than eightfold increase in KM/kCat), could be The HbpA substrate spectrum could be enlarged even more via directed evolution. Recently, an HbpA mutant was found that initiated the production of indigo stating from indole. It is assumed that HbpA converts indole into the 2,3-epoxide,which spontaneously dimerizes to indigo (Fig. 16.3-9)13’1. In vitro application of HbpA (and monooxygenases in general) offers some advantages over whole-cell biotransformations. For example, toxic effects on cell metabolism and further metabolization of the desired product can be avoided, and experimentally demanding in vivo set-ups are not necessary (beneficial for organic chemists). The major challenge in in vitro biotransformations is the efficient

I

1181

IG Oxidation Reactions 0 2

H indole NADH

I

NAD'

Figure 16.3-9. Proposed reaction sequence catalyzed by 2-hydroxybiphenyl-3-monooxygenase (HbpA) for the formation o f indigo from

0

indole.

H indigo

-

--

m S I ~ Usubstrate JPCKJ

NADH

A

B

C0,-

\ J FDH

12+

HC0,H cathode

Figure 16.3-10. Formation o f 3-phenylcatechol from 2-phenylphenol catalyzed by partially purified 2-hydroxybiphenyl-3-monooxygenase(HbpA) in organic aqueous emulsions. Regeneration o f N A D H was achieved in situ with formate dehydrogenase (FDH) (A) or indirectly electrochemically with [Cp*Rh(bpy)(H20)I2+ (B).

76.3 Oxidation of Phenols Substrates and products of peroxidase - catalyzed oxidative di- and oligomerizations of phenols.

Table 16.3-7. Substrate

Products

References and applications

Alkaloid synthesis [ I 5 ]

Me0

OMe OH

Me0

# OH

J$

HO

OH

OH

/

Alkaloid synthesis [I'

OMe

.:WlH

Antimicrobial compounds ( I 6 ]

HO@ HO

"3 \

'

'

COOCH,

o

'0H

@ /

Phytoalexin activity~"1

/

C A COOCH,

regeneration of reduced nicotinamide coenzymes. The general strategies are described in Chapter 7. Furthermore, the production enzyme must be easily available in large amounts. HbpA was obtained in gram amounts from recombinant E. coli in a one-step operation via expanded bed adsorption Limitations

I

1183

1184

I

7G Oxidation Reactions Table 16.3-7.

(cont.).

Substrate

Products

References and aDdications

HO

Melanin synthesis [l9I HO

ti

HO

aoH

Racemic ['I

%OH, ' , ' OH

woH

a

Racemic I2O1

OH

RO

RO

Quest Int. Naarden, The Netherlands, R = arrabinoxylan, carbohydrate gel which retains water

0 Me

15 A. R. Krawczyk, E. Lipkowska, J.T. Wrobel, Coll. 18 A. E. Goodbody, T. Endo, J. Vukovic, J. P. Kutney, Czech. Chem. Commun. 1991,561147. L. S. L. Choi, M. Misawa, Planta Med. 1988,136. 16 A. Kobayashi, Y. Koguchi, H. Kanzaki, S.I. Ka19 M.dlschia, A. Napolitano, K. Tsiakas, G . Prota, jiyama, K. Kawazu, Biosci. Biotech. Biochem. 1994, Tetrahedron 1990,46,5789. 58, 133. 20 M. M. Schmitt, E. Schiiler, M. Braun, D. Haring, 17 D. M.X. Donelly, F. G. Murphy, J. Polonski, P. Schreier, Tetrahedron Lett. 1998,39,2945-2946. T. Prang6, J. Chern. Soc. Perkin Trans. 1 1987,2719.

due to low solubility of substrates and products can be overcome in biphasic reaction systems (Fig. 16.3-10).HbpA exhibits significant activity in the presence of various organic solvents such as 1-decanol,hexadecane or heptaner3'1. Thus, the synthetic in vitro application of HbpA was done via an emulsion process. Several regeneration strategies for NADH were reported (Fig. 16.3-10). In the emulsion process, a high 3-phenylcatechol concentration in the organic phase and the same or higher productivities (up to 0.45 g L-' h-') as in the in vivo process were achievedL4']. Here, formate dehydrogenase and formate served as the coenzyme regeneration system (Fig. 16.3-10 A). The benefits of this regeneration

16.3 Oxidation ofphenols

I

1185

Hooclf;oH HO

HooI'IOOH

@OH R

Figure 16.3-11. Hydroxylation of phenols t o catechols catalyzed by horseradish peroxidase (HRP).

system are described in Chapter 16.6. Even electrical power could be used as a source of reduction equivalents (Fig. 16.3-10B) l4l]. 16.3.4

Peroxidases 16.3.4.1 Oxidative Coupling Reactions

Phenols are typical substrates for peroxidases. Quite similarly to the laccasemechanism (described earlier in this chapter), peroxidases catalyze phenol oxidations via hydrogen abstraction. The radicals thus generated leave the active site and Table 16.3-8.

Suberate

Selected hydroxylation reactions of phenols catalyzed by horseradish peroxidase. Product

"&OH HO

Tyrosine

L-Dopa

OH OH

Adrenaline

21 A.M. Klibanov, 2. Berman, B.N. AlbertiJ. Am. Chem. SOC.1981,103,6263-6364.

Literature

1186

I

7 G Oxidation Reactions Selected nitration reactions of phenols catalyzed by soybean peroxidase.

Table 16.3-9.

Product(s), Yield I"/.]

Substrate

nara

ortho

O *H

/

NO2

O Z N a o H

OH

58

I

OH

0

27

OH

OH

22

25

0

D

O

H

o&oH41 20

yl

0

OH

VNoz 25

react with other aromatic compounds (depending on the reaction conditions) to form dimeric and polymeric products[42].A selection of dimeric products is presented in Table 16.3-7. Recently, peroxidases, especially horseradish (HRP) and soybean peroxidase, found increasing interest in resin manufacturing. The peroxidase-catalyzed coupling of phenols [431, catechols [441, hydroquinones C4'1, or anilines [46, 471 is a potential substitute for the conventional production of phenolic resins using toxic formaldehyde [481. The resins find applications as conductive polymers [45, 4gl. 16.3.4.2 Hydroxylationof Phenols

As early as 1961, Mason and coworkers reported that HRP, in the presence of dihydrofumaric acid as cofactor, catalyzes the hydroxylation of arenes (Fig. 16.311)'501.

Also lignin peroxidase was found to catalyze the oxidation of phenol, cresol, and tyrosine ['ll.

76.3 Oxidation ofPhenols Table 16.3-10.

Oxidation reactions o f arylamines catalyzed by peroxidases.

Substrate y

Product

References and remarks

2

Bromoperoxidase1")

Chloroperoxidase [23] CI

CI

Aminopyrrolonitrin

R

0

Pyrrolonitrin

R

0

22 N. Itoh, N. Morinaga, T. Kouzai, Biochem. Mol. Bid. 1993, 29,785-791. 23 S. Kirner, K.-H. van Pee, Angew. Chem. Int. Ed. 1994, 33, 352.

Chloroperoxidase R = 0-, m-,pC1; pCH3; p-COOH 24 V.N. Burd, K:H. van Pee, Bioorg. Khim. 1998,24, 462-464.

16.3.4.3 Nitration of Phenols

Khmelnitsky and coworkers recently reported a rather unusual application of soybean peroxidase. In the presence of nitrite and hydrogen peroxide, phenols are nitrated. The nitration of tyrosine has been reported earlierrS2*s31. The substrate spectrum was enlarged by various phenolic compounds (Table 16.3-9).Thus, such an enzymatic nitration represents an alternative to chemical nitration (especially for acid-labile phenols, which cannot by nitrated chemically). Other peroxidases such as HRP or CPO were also able to perform such reactions. Another approach to the production of nitroarenes with peroxidases is based on the CPO (or bromoperoxidase)-catalyzedoxidation of arylamines. Table 16.3-10 gives a selection of peroxidase-catalyzedconversions of aniline derivatives to corresponding nitroarenes. For example, aniline was converted into nitrobenzene by a bromoperoxidase from Pseudornonas p ~ t i d a ( ' ~ 1and , aminopyrrolonitrin was converted into the antibiotic pyrrolonitrin by a CPO from P. p y r r ~ c i n i a [ ~ ~ ] .

I

1187

1188

I

7G Oxidation Reactions

Table 16.3-11. Substrate

Substrates and products of 4-cresol-o~idoreductase~~~~ 26].

Product

Substrate

6 bH OH

$o /

OH

/

6- 4 OH

0 ’

@

OH

OH

Product

OH

6 OH

0 ’

OH

OH 25 W. Mclntire, D. J. Hopper, T. P. Singer, Biochem.j. 1985,228,325-335.

26 W. Mclntire, D.J. Hopper, J. C. Craig, E.T. Everhart, E.V. Webster, M. J. Causer, T. P. Singer, Biochem.J 1984,224,617-621.

16.3.5 Other Oxidoredudases

16.3.5.1 4-Cresol-oxidoreductase (PCMH,

E. C. 1.17.99.1)

This enzyme shares structural and mechanistic properties with VAO[’ll. In contrast to VAO it is not an oxidase as regeneration of the covalently bound FAD with molecular oxygen is not possible. It is a flavocytochrome enzyme. The reduction equivalents from the substrate are transferred to a type c cyto~hrome[’~, 571. In

7 6.3 Oxidation of Phenols Oxidations of 4-alkylphenols catalyzed by 4-ethylphenol 0xidoreductase1*~].

Table 16.3-12.

Substrate

Relative conversion rate

[%r

44

p-Cresol 4-Ethylphenol

100

4-Propylphenol

112

4-Butylphenol

114

4-Pentylphenol

116

4-Heptylphenol

52

4-Nonylphenol

14

a 100% corresponds to the 4-ethylphenol conversion rate.

27 C. D. Reeve, M.A. Carver, D. J. Hopper, Biochem.J. 1990,269,815-819

addition to a cytochrome c / cytochrome c oxidase regeneration sy~tem['~1, chemical reoxidation agents such as phenazine methosulfate, dichlorophenol indophenol[57], and ferrocenes[6G621 have been used. The reaction mechanism is quite similar to the one of VAO and also includes an intermediate, the para-quinone methide. Like VAO, 4-cresol-oxidoreductase also exhibits a high enantioselectivity for (S)-l-(4'-hydroxyphenyl)alkylalcohols ["I. This enzyme accepts a broad range of substrates; para-methylphenols are preferably oxidized to the corresponding aldehydes, whereas the oxidation of para-alkylphenols results in the formation of significant amounts of (S)-alcohols (Table 16.3-11)

631.

16.3.5.2 4-Ethylphenol Oxidoreductase

4-Ethylphenol oxidoreductase from Pseudomonas putida JD1 is structurally almost identical to 4-cresol oxidoreductase,but catalyzes the hydroxylation of para-alkylphenols with longer aliphatic chains (Table 16.3-12). The hydroxylation reactions enantioselectively produce (R)-alcohols[64, "1. The regeneration properties of this enzyme are quite similar to 4-cresol oxidoreductaselG11.

1190

I

IG Oxidation Reactions

2-aminotetralines

9-hydroxy N-(n-propyl) hexahydronaphthoxazine

Figure 16.3-12. Substrates for phenol oxidase from Mucuna pruriens. 5-, 6-, or 7-Hydroxylated 2-aminotetralins with R = H or C,H7 and 9-hydroxy-N-(n-propyl)-hexahydronaphthoxazine are substrates for the phenol oxidase.

Figure 16.3-13. Formation o f 7,g-dihydroxy N-(di-n-propyl)-2-aminotetralin with Mucuna-phenoloxidase. Quinone formation is prevented in situ with ascorbate as reductant.

16.3.6 In vivo Oxidations 16.3.6.1 Phenoloxidase of Mucuna pruriens

Like other phenoloxidases, this enzyme has a low substrate specificity and is able to ortho-hydroxylate a whole range of para-substituted monocyclic phenols. The catechols produced belong to groups of fine chemicals and pharmaceuticals["]. Furthermore, also bi- and tri-cyclic phenols were converted into catechols (Figurel6.312) [G71. 2-Aminotetralines, on the basis of their dopaminergic properties, are compounds of pharmaceutical interest. Phenoloxidase (monophenol monooxygenase, E. C. 1.14.18.1) introduces one atom of molecular oxygen into the substrate and was used in alginate-entrappedcells or in partially purified form. The pharmaceutical 7,8-dihydroxy-N-(di-n-propyl)2-aminotetralin was produced continuously using a phenol oxidase suspension in dialysis tubing in an airlift fermenter coupled to an aluminium oxide column for selective product isolation (Figure 16.3-13)["j. A product concentration of 130 mg/L and a yield of 25 % were reached.

16.3 Oxidation of Phenols

aoycooH J3 “y 0, I Beauveria bassiana

(R)-2-phenoxypropionic acid

HO

(R)-2-(4-hydroxyphenoxy)propionic acid

Regioseledive para-hydroxylation o f (R)-2-phenoxypropionic acid catalyzed by Beauveria bassiana (HPOPS process). Figure 16.3-14.

16.3.6.2

Monohydroxylation of (R)-2-PhenoxypropionicAcid and Similar Substrates[69.701

The product is a frequently used intermediate for the synthesis of enantiomerically pure aryloxyphenoxypropionic acid type herbicides. The enzyme catalyzing the hydroxylation of the phenolether is an oxidase, which is not further characterized. The biocatalyst Beauvena bassiana was found by an extensive screening of microorganisms for regioselective hydroxylation of (R)-2-phenoxypropionicacid and for substrate tolerance. This fungal strain was improved by random mutagenesis and screening, which resulted in strain LU 700. The hydroxylation is not growthassociated and the ee is increased during oxidation from 96% for the substrate to 98% for the product. After process optimization, a productivity of 7 g L-l d-’ was reached. The biotransformation is carried out in a 120 000 L reactor at BASF in Germany. The biocatalyst has a broad substrate spectrum. A compound needs the structural elements of a carboxylic acid and an aromatic ring system to be a substrate for the oxidase. Hydroxylation primarily takes place at the para position if it is free. If an alkyl group is in the para position, only the side chain is oxidized. In systems with more than one ring, the most electron-rich ring is hydroxylated. 16.3.6.3

Biotransformation of Eugenol to Vanillin L7’]

The biotechnological production of vanillin is of interest because there is a large demand for vanillin originating from so called “natural” sources. Possible strategies for the biotechnological production of vanillin are reviewed by Priefert et al. [72]. One synthetically interesting strategy is the production of vanillin from eugenol. Here, a part of a catabolic pathway is used to accumulate an intermediate of this pathway. This was achieved by the knock-out of the enzyme catalyzing the further conversion of the putative product. For the accumulation of vanillin from eugenol, the catabolism of eugenol in Pseudornonas sp. Strain HR199 (DSM7063) was used. In order to prevent further degradation of vanillin, the gene enconding vanillin dehydrogenase, responsible for the oxidation of vanillin to vanillic acid, was inactivated by insertion mutagenesis. In a non-optimized biotransformation using growing cells in an aqueous mineral salts medium containing gluconate as a source of carbon and energy and 6.5 mM eugenol, vanillin accumulated up to a concentration of 2.9 mM, corresponding to a

I

1191

1192

I

FoMe qoHqo

11 Oxidation Reactions

~$$;Yla~

hydroxylas: eugenol

L

/

OMe OH

eugenol

coniferyl alcohol

0

I

enovl-CoAhydratase/aldolase F 0

M

OH

vanillin

;

T

acetyl-CoA

-

r

I

coniferyl aldehyde coniferyl aldehyde dehydrogenase

%

OMe

OH

OH

o

A

feruloyCCoAsynthetase

' OMe

qo OMe OH

OH

feruloyl-CoA

ferulic acid

Multistep biotransformation of eugenol to vanillin catalyzed by whole cells o f Pseudomonas sp. HR 199. Figure 16.3-15.

molar yield of 44.6%. The major drawback of the process is the degradation of vanillin by the action of coniferyl aldehyde dehydrogenase when coniferyl aldehyde is depleted from the medium.

References

R.H. H. van den Heuvel, M. W. Fraaije, A. Mattevi, C. Laane, W. J. H. van Berkel, J. Mol. Cat. B: Enzymatic 2001, 11,185-188 2 R. H. H. van den Heuvel, C. Laane, W. J. H. van Berkel, Adv. Synth. Cat. 2001, 343, 515-520. 3 M. W. Fraaije, C. Veeger, W. J. H. van Berkel, Eur. J. Biochem. 1995,234, 271-277. 4 M. W. Fraaije, W. J. H. van Berkel,J. Bid. Chem. 1997,272,18111-18116. 5 M. W. Fraaije, R. H. van den Heuvel, J. C. Roelofs, W. J. H. van Berkel, Eur. J. Biochem. 1998,253,712-719. 6 R. H. H. van den Heuvel, M. W. Fraaije, C. Laane, W. J. H. van Berkel,J. Bacterial. 1998, 180,5646-5651. 7 A. Mattevi, F. M. W., A. Mozzarelli, A. Olivi, W. J. H. van Berkel, Structure 1997, 5, 907-920. 8 F. P. Drijfhout, M. W. Fraaije, H. Jongejan, W. J. H. van Berkel, M. C. R. Franssen, Biotech. Bioeng. 1998,59, 171-177. 1

R. H. H. van den Heuvel, J. Partridge, C. Laane, P. J. Halling, W. J. H. van Berkel, FEBS Lett. 2001,503,213-216. 10 R. H. H. van den Heuvel, F. M. W., W. J. H. van Berkel, FEBS LETT 2000,481, 109- 112. 11 R. H. H. van den Heuvel, F. M.W., M. Ferrer, A. Mattevi, W. J. H. van Berkel, PNAS 2001,97,9455-9460. 12 R. H. H. van den Heuvel, M. W. Fraaije, C. Laane, W. J. H. van Berkel, j.Argi. Food Chem. 2001,49,29542958. 13 H. P. Call, I. Mucke,J. Biotech. 1997,53, 163-202. 14 U.L. R. Baminger, C. Galhaup, C. Leitner, K. D. Kulbe, D. Haltrich, J. Mol. Cat. B: Enzymatic 2001, 1 I , 541-550. 15 T.Shiba, X. Ling, T. M., C.-L. Chen,]. Mol. Cat. B: Enzymatic 2000, 10,605-615. 16 R. Bourbonnais, M. G. Paice, Febs Lett. 1990,267,99-102. 9

References 17 A. Potthast, T. Rosenau, K. Fischer, H o b

forschung 2001,55,47-56.

18 M. Amann.; Proceedings of the Intema-

tional Symposium an Wood and Pulping Chemistry, 1997, Montreal, Canada. 19 D. Kertesz, D. Zito, Biochim. Biophys. Acta 1965,96,447. 20 S. G. Burton, Catalysis Today 1994,22, 459-487. 21 D. Strack, W. Schliemann, Angew. Chem. 2001,113,3907-3911. 22 P. Capdeville, M. Maumy, Tetrahedron Lett. 1982,23,1573-1576. 23 K. Mammo, J.H. Waite, Biochim. Biophys. Acta 1986,872,98. 24 H. Yamamoto, H. Tanisho, S. Ohara, A. Nishida, rnt. I . Bid. Macromol. 1992, 14,66. 25 G. H. Miiller, H. Waldmann, Tetrahedron Lett. 199637, 3833-3836. 26 G. H. Mtiller, A. Lang, D. R. Seithel, H. Waldmann, Chem. Eur. J. 1998,4, 2513-2522. 27 W. A. Suske, M. Held, A. Schmid, T. Fleischmann, M. G. Wubbolts, H.-P. E. Kohler, ]. Bid. Chem. 1997, 272, 24 257-24265. 28 F. Chioccara, P.Gennaro, G. la Monica, R. Sebastino, B. Rindone, Tetrahedron 1991, 47,4429-4434. 29 D. H. R. Barton, D. M. X. Donnelly, P. J. Guiry, J.-P. Finet, J. Chem. Soc. Perkin Trans. r 1994,2921~. 30 A. Feigenbaum, J.-P. Pete, A. Poquet-Dhimane, Tetrahedron Lett. 1988, 29,73-74. 31 K. A. Parker, K. K. A.,Joumal ofOrganic Chemistry 1987,52,674-676. 32 M. Held, W. Suske, A. Schmid, K. Engesser, H. Kohler, B. Witholt, M. Wubbolts,J. Mol. Cat. B: Enzymatic 1998,5,87-93. 33 A. Schmid, H.-P. E. Kohler, K.-H. Engesser, ]. Mol. Cat. B: Enzymatic 1998, 5, 311-316. 34 M. Held, A. Schmid, H.-P. E. Kohler, W. A. Suske, B. Witholt, M. G. Wubbolts, Biotech. Bioeng. 1999,62,641-648. 35 W. A. Suske, W. J. H. van Berkel, H.-P. E. Kohler,J. B i d . Chem. 1999, 274, 33355-33365. 36 A. Meyer, A. Schmid, M. Held, A. H. Westphal, M. Rothlisberber, H.-P. E. Kohler, W. J. H. van Berkel, B. Witholt, 2001, submitted. 37 A. Schmid, 2001. 38 J. Lutz, B. Krummenacher, B. Witholt, A. Schmid. "2-Hydroxybiphenyl3-Monooxygenase: Large Scale Preparation and Cell Free

Application in Emulsions"; BioTrans 2001, 2001, Darmstadt, Germany. 39 A. Schmid, J.Lutz, V. V. Mozhaev, L. Khmelnitsky, B. Witholt, J . Mol. Cat. B: Enzymatic, submitted. 40 A. Schmid, I. Vereyken, M. Held, B. Witholt, J. Mol. Catal. B: Enzymatic 2001, 11, 455-462. 41 F. Hollmann, A. Schmid, E. Steckhan, Angew. Chem. 2001,113,190-193. 42 W. Adam, M. Lazarus, C. R. Saha-Moller, 0. Weichold, U. Hoch, D. Haring, P. Schreier, Biotransformations with Peroxidases. In K. Faber (ed), Adv. Biochem. Eng. Biotech., Springer, Berlin, Heidelberg, 1999, Vol. 63, pp. 74-104. 43 H. Kurioka, H. Uyama, S. Kobayashi, PolymerJ. 1998,30,526-529. 66 S. Dubey, D. Singh, R. A. Misra, Enz. Microb. Tech. 1998, 23. 45 P. Wang, S. Amarasinghe, J. Leddy, M. Arnold, J. S. Dordick, Polymer 1998, 39, 123- 127. 46 J. A. Akkara, P. Salapu, D. L. Kaplan, 1nd.J. Chem. 1992,31B, 855-858. 47 j. Y. Shan, S. K. Cao, Polym. Adv. Technol. 2000,l I , 288-293. 48 P. W. Kopf, Encyclopedia of Polymer Science and Engineering; Wiley, New York, 1986, VO~.11; pp. 45-95. 49 S. Kobayashi, I. Kaneko, H. Uyama, Chem. Lett. 1992, 393. 50 D. R. Buhler, H. S. Mason, Arch. Biochem. Biophys. 19Gl,2, 224. 51 M. W. Schmall, L. S. Gorman, J. S. Dordick, Biochim. Biophys. Acta 1989,999, 267. 52 H. Shibata, Y. Kono, S. Yamashita, Y. Sawa, H. Ochiai, K. Tanaka, Biochim. Biophys. Acta 1995, 1230,45-50. 53 A. van der Vliet, J. P. Eiserich, B. Halliwell, C. E. Cross, J. Bid. Chem. 1997,272, 7617-7625. 54 N. Itoh, N. Morinaga, T. Kouzai, Biochem. Mol. Bid. 1993, 29, 785-791. 55 S. Kimer, K.-H. van Pee, Angew. Chem. Int. Ed. Engl. 1994, 33, 352. 56 W. McIntire, D.E. Edmondson, T. P. Singer, D. J. Hopper,]. Biol. Chem. 1980, 255, 6553-6555. 57 A. L. Bhattacharyya, G. Tollin, W. McIntire, T. P. Singer, Biochem.]. 1985, 228, 337-345. 58 W. McIntire, C. Bohmont. In de Gruyter, Havins and Havoproteins, Berlin, 1987, pp. 677-686.

I

1193

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

1194

I

1G Oxidation Reactions

W. McIntire, D. J. Hopper, J. C. Craig, E. T. Everhart, E.V. Webster, M. J. Causer, T. P. Singer, Biochem.J. 1984,224,G17-621. 60 H. A. 0. Hill, B. N. Oliver, D. J.Page, D. J. Hopper, J. Chem. SOC.,Chem. Commun. 1985,1469-1471. 61 B. Brielbeck, M. Frede, E. Steckhan, Biocatalysis 1994,10,49-G4. 62 E. Steckhan, Electroenzymatic Synthesis. In Top. Curr. Chem.; Springer-Verlag:Berlin; Heidelberg, 1994;Vol. 170;pp. 84-111. 63 W. McIntire, D. J. Hopper, T. P. Singer, Biochem.]. 1985,228,325-335. 64 C. D. Reeve, M. A. Carver, D. J. Hopper, Biochem. J. 1989,263,431-437. 65 C. D. Reeve, M . A. Carver, D. J. Hopper, Biochem. J . 1990,269,815-819. 66 N. Pras, H. J. Wichers, A. P. Bruins, T. M. Malingre, Plant Cell, Tissue and Organ Culture 1988,13,15-26. 59

N. Pras, G. E. Booi, D. Dijkstra, A. S. Horn, T. M. Malingre, Plant Cell, Tissue and Organ Culture 1990,21,9-15. 68 N. Pras, S. Batterman, D. Dijkstra, A. S. Horn, T. M. Malingre, Plant Cell, Tissue and Organ Culture 1990,23,209-215. 69 C. Dingler, W. Ladner, G. A. fiei, B. Cooper, B. Hauer, Pesticide Science 1996,4G,3335. 70 B. Cooper, W. Ladner, B. Hauer, H. Siegel. Verfahren zur fermentativen Herstellung von 2-(4-hydroxyphenoxy-)propionsaure, 1992,EP0465494Bl. 71 J.Overhage, H. Priefert, J. Rabenhorst, A. Steinbiichel, Appl. Microbiol. Biotech. 1996, 52,820-828 72 H. Priefert, J. Rabenhorst, A. Steinbiichel, Appl. Microbiol. Biotech. 2001,56, 296-314. 67

16.4 Oxidation of Aldehydes

Andreas Schmid, Frank Hollmann, and Bruno Buhler 16.4.1 Introduction

To date, few reports on synthetic enzymatic oxidations of aldehydes have been published. Preparative applications reported include bioconversions of natural products such as retinal (Fig. 16.4-1 A) and various aliphatic and unsaturated aldehydes (Fig. 16.4-1 B). A broad range of aromatic acids can be obtained from their corresponding aldehydes (Fig. 16.4-1 C). Another reported reaction type is the production of olefins from aldehydes by oxidative removal of formic acid from the substrate (Fig. 16.4-1 D). 16.4.2 Alcohol Dehydrogenases

Alcohol dehydrogenases are generally applied for the interconversion of alcohols and aldehydes. Yet, these enzymes have also attracted interest due to their ability to oxidize aldehydes[lI. HLADH was shown to oxidize butanal12]. This reaction, however, shows no potential for synthetic application unless a very efElcient NAD' regeneration system is applied (Fig. 16.4-2).The catalyhc activity of HLADH for the reduction of the aldehyde is more than 100 times higher than that for aldehyde oxidation (examined for benzaldehyde)f31. As a result, the initially formed NADH is

76.4 Oxidation ofAldehydes 11195

RAOH

Figure 16.4-1. Selected enzymatic oxidations o f aldehydes. A oxidation o f complex natural products such as retinal; 6:oxidation of aliphatic and a$-unsaturated aldehydes; C: oxidation o f (hetero)arylic aldehydes; D: oxidative cleavage of the aldehyde-carbon atom yielding terminal alkenes.

fi-

NAD'

-OH

NADH

O '

* NADH

NAD+

Figure 16.4-2. Oxidation activity for aldehydes exhibited by horse liver alcohol dehydrogenase (HLADH). Only minor amounts of acid are produced because of the higher HLADH activity for aldehyde reduction.

-0

+

HO ,

JOH

0-

TBADH * NAD+

TBADH * NADH

/\OH Figure 16.4-3. Aldehyde dismutase acitivity of Thermoanaerobium brockii alcohol dehydrogenase (TBADH). A high affinity o f the TBADH-NAD' complex for hydrated acetaldehyde is proposed, explaining the stochiometric acetaldehyde dismutation.

1196

I

7G

Oxidation Reactions

used for aldehyde reduction, yielding a dynamic equilibrium between alcohol and aldehyde. TBADH also exhibits the so-called aldehyde dismutase activity[41.In contrast to HLADH, stochiometric dismutation of acetaldehyde into one equivalent of ethanol and acetic acid has been reported. A gem-diol mechanism was proposed for this reaction (Fig. 16.4-3). 16.4.3

Aldehyde Dehydrogenases

Several aldehyde dehydrogenases have been reported for biocatalytic applications. Recently, aldehyde dehydrogenase (E. C. 1.2.1.5) from yeast was applied to oxidize (Z,Z)-nona-2,4-diena1[’1. Recycling of NAD’ was achieved in situ by addition of an alcohol dehydrogenase, reducing (Z,Z)-nona-2,4-dienalto the corresponding alcohol. Since both reactions are stochiometrically linked via NAD, this corresponds to an overall dismutation of the aldehyde (Fig. 16.4-4).This concept was extended to industrially relevant metabolites of linoleic acid (detergents and polymer buildingblocks) (Fig. 16.4-5). No isomerization of the double bonds and yields up to 90% were reported[’]. Enzymatic transformation of (Z,Z)-nona-2,4-dienal t o the corresponding alcohol and acid catalyzed by an alcohol and an aldehyd e d ehyd rogenase from yeast. Figure 16.4-4.

A

/AD?

\

NAhidDH

NADH

OH

I

H y & &

/-

lipoxygenase

I

hydroperoxide lyase

J

O\

A

AldDH

C o*Oo*

\“.‘Am

H C > o O O H

Figure 16.4-5. Enzymatic cleavage o f linoleic acid t o o-hydroxy and dicarboxylic acids.

16.4 Oxidation of Aldehydes Table 16.41. Kinetic constants o f bovine kidney aldehyde dehydrogenase for different substrates 1’1. Substrate

Vm,

[%I”

KM

bM1

100

9.1

758

1

855

1.5

1960

30

1683

33.9

3026

8.2

A H

-0 0

a The

V,

values are relative to retinal as substrate.

1 P. V. P. Bhat, L., Wang, X. L., Biochem. Cell Bid. 1996.74,695-700. R

Mechanism proposed for light emission in the course o f the luciferase reaction. Figure 16.4-6.

Another NAD’-dependent aldehyde dehydrogenase (from bovine kidney) was characterized with respect to its activity toward retinal and other aldehydes (Table 16.4-1) [GI.

I

1197

1198

I

76 Oxidation Reactions Oxidation of aldehydes t o corresponding carboxylic acids catalyzed by P450 rnonooxygenases.

Table 16.4-2.

0

R H'

*-

RKOH

0,, NAD(P)H

H,O, NAD(P)*

Substrate

Reference

Aliphatic aldehydes

~

RA

H

3

1

[31

&

0

\

Losartan

171

~~~

2 Y. Terelius, C. Norsten-Hoog, T. Cronholm. M. Ingelman-Sundberg, Biochem. Biophys. k s . Commun. 1991, 179,689-694. 3 K. Watanabe, T. Matsunaga, S. Narimatsu, 1. Yamamoto, H. Yoshimura, Biochem. Biophys. Res. Commun. 1992,188, 114-119. 4 S. Tomita, M. Tsujita, Y. Matsuo, T. Yubisui, Y. Chikawa, 1nt.J. Biochem. 1993,25,1775-1754.

5 K. Watanabe, T. Matsunaga, I. Yamamoto, H. Yashimura, Drug. Metab. Dispos. 1995, 23, 261-265. G K. Watanabe, S . Narimatsu, T Matsunaga, I. Yamamoto, H. Yoshura, Biochem. Qhamacol. 1993,46, 405-41 1.

7 R. A. Steams, P. K. Chakravarty, R. Chen, S.-H. L. Chiu, Drug. Metab. Dispos. 1995, 23, 207-215.

16.4.4 Monooxygenases 16.4.4.1

Luciferase (E.C. 1.14.14.3)

Probably the most prominent oxidation reaction of aldehydes is the well-known luciferase reaction. The flavin-dependentluciferase is present in a number of marine and terrestrial species[" 1'. Light of about 490 nm (blue-green) is emitted as a by-

16.4 Oxidation ofAldehydes

I

1199

Table 16.43. Oxidations and subsequent decarboxylations of aldehydes catalyzed by P450 monooxygenases.

NAD(P)H. 0,

Substrate

8 E. S. Roberts, A. D. N. Vaz, M . J. Coon, Proc. Natl. Acad. Sci USA 1991,88,8963-8966. 9 A. D. N. Vaz, E. S. Roberts, M. 1. Coon,]. Am. Chem. Soc. 1991,113, 5886-5887.

NAD(P)+,H,O

Reference

10 A. D. N. Vaz, K. J. Kessel, M. J. Coon, Biochern 1994,33,13651-13661.

product of the oxidation of aliphatic aldehydes. Excited flavin species are discussed as emitters (Fig. 16.4-6)1'- lo]. 16.4.4.2

Cytochrome P 4 5 0 ~ ~ . 3

The oxidation of an aldehyde to the corresponding carboxylic acid with P450 systems is reported for various substrates (Table 16.4-2).In some cases oxidative decarboxylation is observed yielding formic acid and an olefin, one carbon atom shorter than the substrate (Table 16.4-3). Several o-0x0 fatty acids are transformed to the corresponding a,w -dicarboxylic acids, whereas o-formylesters of fatty acids are decarboxylated to the o-hydroxy fatty acids and carbon dioxide["]. For several w-0x0 fatty acids turnover frequencies (measured as O2consumption) between 1.8 to 25 s-l were found. Many P450 systems are multi-component enzymes with small protein cofactors such as putidaredoxin performing the electron mediation between NAD(P)H and the active site of the enzyme. Vilker and coworkers recently were able to show that NADPH can be omitted from the catalyhc cycle by direct electrochemical reduction of putidar-

1200

I

7G

Oxidation Reactions

Table 16.4-4.

Substrate

A

H

J

Kinetic constants ofxanthine oxidase"'].

KM [mM]

vmax

141.5

22.2

130

100

430

23.3

142

2.4

0.34

3.4

0.046

2.7

1.7

4.2

1.03

7.7

0.068

15.7

0.085

1.8

1

1

2

0.1

Is-']

H

0

11 F.

F. Morpeth, Biochim. Biophys. Acta 1983,744,328-334.

edo~in['~-'~I, thus oxidizing styrene or camphor. Other approaches utilize Co sepulchrate as reducing agent, which can be regenerated either chemically (via Zn) ['I or electrochemically['G.l71.

References I1201 16.4.5 Oxidases 16.4.5.1 Xanthine Oxidase (E.C. 1.1.3.22)

Xanthine oxidase was examined for its catalyhc applicability for the oxidation of aldehydes as early as 196711*].In addition to 02,xanthine oxidase was reported to accept e. g. methylene blue, PMS or ferricyanide[”I as electron acceptors. Table 16.4-4gives kinetic data for some substrates L2O]. 16.4.6 Oxidations with Intact Microbial Cells[*’]

Burkholderia cepacia was reported to transform aromatic aldehydes into the corresponding acids. Vanillin, para-hydroxybenzaldehyde,and syringaldehyde were converted to corresponding acids with high yields of 94%, 92 %, and 72 %, respectively (Fig. 16.4-7)[22]. The acid produced is not further metabolized as long as the aldehyde still is accessible to the cells. The enzyme responsible for aldehyde oxidation in Burkholderia cepacia was not further characterized. However, the gene of an NADdependent vanillin dehydrogenase of Pseudomonas sp. strain HR199 was cloned and characterized[23].Recombinant E. coli containing this vanillin dehydrogenase transformed vanillin to vanillate at a clearly higher rate than Burkholderia cepacia.

R

Burkholderia cepacia *

Figure 16.4-7. Oxidation of aromatic aldehydes by Barkholderia cepacia TM1.

References 1

L. P. Olson, J. Luo, 0. Almarsson, T. C. Bruice, Biochemistry 1996, 35, 9782-9791.

G. T. M. Henehan, N. J. Oppenheimer, Biochemistry 1993, 32, 735-738. 3 G. L. Shearer, K. Kim, K. M. Lee, C. K. Wang, B. V. Plapp, Biochemistry 1993, 32, 2

1118611194.

S. Trivic, V. Leskova, G. W. Winston, Biotech. Lett. 1999, 21, 231-234. 5 A. Nunez, T. A. Foglia, G. J. Piazza, Biotechnol. Appl. Biochem. 1999, 29, 207-212. 6 P. V. Bhat, L. Poissant, X. L. Wang, Biochem. Cell Bid. 1996,74,695-700. 7 T. 0. Baldwin, M. M. Ziegler. In Chemistry and Biochemistry of Flavoenzymes, CRC 4

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

1202

I

76 Oxidation Reactions

Press, Boca Raton, 1992, Vol. 111, pp. 467-530. 8 A. Palfey, V. Massey, Flavin-Dependent Enzymes. I n ComprehensiveBiological Catalysis. M. Sinnott (ed),Academic Press, San Diego, London, 1998, Vol. 111, pp. 83-154. 9 C. T. Walsh, Y.C.J. Chen, Angav. Chem. 1988,100,342-352. 10 P. Macheroux. S. Gishla, Nachr. Chem. Tech. Lab. 1985, 33, 785. 11 S. C. Davis, 2. Sui, J. A. Peterson, P. R. Ortiz de Montellano, Arch. Biochem. Biophys. 1996,328, 35-42. 12 M. P. Mayhew, V. Reipa, M. J. Holden, V. L. Vilker, Biotechnol. Prog. 2000, 16, 610-616. 13 V. Reipa, M. Mayhew, V. L. Vilker, PNAS 1997,94, 13554-13558. 14 V. L. R. Vilker, Vytas; Mayhew, Martin; Holden, Marcia J.,/. Am. Oil Chem. SOC.1999, 76,1283-1289. 15 U. Schwaneberg, D. Appel, J. Schmitt, R. D. Schmid,/. Biotech. 2000,84, 249-257.

R. W. Estabrook, K. M. Faulkner, M. Shet, C. W. Fisher, Application of Electrochemistry for P450-CatalyzedReactions. I n Methods in Enzymology, Academic Press. San Diego, London, Boston, New York, Sydney, Tokyo, Toronto, 1996, Vol. 272, pp. 44-51. 17 K. M. Faulkner, M. S. Shet, C. W. Fisher, R. W. Estabrook, PNAS 1995,92, 7705-7709. 18 F. Dastoli, S. Price, Arch. Biochem. Biophys. 1967, 118,163-165. 19 G. Pelsey, A. M. Klibanov, Biochim. Biophys. Acta 1983,742,352-357. 20 F. F. Morpeth, Biochim. Biophys. Acta 1983, 744, 328-334. 21 M. Tanaka, Y. Hirokane, /. Biosci. Bioeng. 2000,90, 341-343. 22 S. Adachi, M. Tanimoto, M. Tanaka, R. Matsuno, Chem. Eng./. 1992,49, B17-B21. 23 H. Driefert, J. Rabenhorst, A. Steinbiichel, 1. Bacteriol. 1997, 179, 2595-2607. 16

16.5 Baeyer-VilligerOxidations

Sabine Flitsch and Cideon Crogan 16.5.1 Introduction

The enzymatic Baeyer-Villiger oxidation continues to receive attention from synthetic organic chemists a s it offers advantages of regio- and enantioselectivity still rarely exhibited by reagents such a s meta-chloroperbenzoic acid (m-CPBA). S o m e recent advances have resulted in abiotic catalytic reagents capable of i n d u c i n g modest enantioselectivity in the Baeyer-Villiger reaction but these reactions are outside the scope of this section. The most encouraging examples of enantioselective Baeyer-Villiger reactions a r e still those catalyzed by microorganisms and enzymes and the extensive research in this area over the last decade has been covered in a number of recent reviews[”’). 16.5.1.1

Steroidal Substrates I t h a d been known for m a n y years t h a t Baeyer-Villiger-type processes occur during the catabolic transformations of natural compounds. In 1953, it w a s described that the C17 side chain of steroids c a n be cleaved by several microorganisms including

16.5 Baeyer-Vdliger Oxidations

Fusarium, Penicillium, Cylindrocarpon, Aspergillus and Gliocladium speciesf8-'']. One example reported was the conversion of progesterone into A'*4-androstadien-3,17-dione in 84% yield as illustrated in Fig. 16.5-1 181. Since these reports, many others describing the microbiological Baeyer-Villiger oxidation of various steroids have been publi~hed[~~-'~I. Interestingly, it has been shown that depending on the microbial strain used, further oxidation may occur leading to incorporation (of an oxygen atom into the D-ring, thus affording the corresponding lactone. In general, these oxidations are restricted to this ring. This selectivity may be due to the fact that the A-ring bears an a, P-unsaturated ketone moiety, which appears to display a different reactivity compared with the other carbonyl functions [l51. Introduction of a A' double bond also often occurs during these processes. Other eicamples involving oxidation of the A ring have been described with a Glomerellujkaroides strain[lGland with Gymnoascus r e e ~ i i l ~Thus, ~]. eburicoic acid affords a 30% yield of A-secoacid whereas the steroidal alkaloid tomatidine leads to the corresponding ketone as the major product, but a smaller amount of A-seco acid is also obtained. This could well be due to hydrolysis of the lactone which would be formed from Baeyer-Villiger oxidation of the parent ketone Fig. 16.5-2. The mechanism of these reactions has been studied by several groups. Fonken and coworkers[18]first showed using 21-14C labelled progesterone, that the testosterone acetate formed during degradation of progesterone by Cladosporium resinae is not an artefact but is indeed an intermediate in the degradation pathway. Further work by Prairie and Talalay['9]using the strain Penicillium liliacinum established the involvement of two enzymes, a 6.l-dehydrogenaseand an NADPH-dependent oxygenase. They also showed that I8O2 molecular oxygen is incorporated as the ring oxygen atom of testololactone. Rahim and Sih[20]succeeded in showing that an oxygenase (requiring the presence of oxygen) as well as an esterase were involved in the degradation of the progesterone side-chain. In other studies using the 17a-labelled substrate, Singh and Rahkit[21]showed that retention of the deuterium label at the C17 position occurs and that the molecular oxygen is incorporated into the product (Fig. 16.5-3). More recently, a gene from Rhodococcus rhodochrous has been cloned and expressed[22],which encodes for a steroid monooxygenase that inserts an atom of oxygen between the C15 and C20 carbons of progesterone, forming testosterone acetate.

Fusarium sp.

progesterone Figure 16.5-1.

A1~4-androstadien-3,17-One

Biotransformation of progesterone using Fusarium spp.

1204

I

76 Oxidation Reactions

Glornerella fusaroides

HO

* 30%

eburicoic acid

Gyrnnoascus reesii H

~

0

2

C

e

HO Figure 16.5-2.

A-ring cleavage by Glornerellafusaroides and Cyrnnoascus reesii.

Figure 16.5-3. Retention of the deuterium label and oxygen incorporation during the side-chain degradation of progesterone.

All these results led to the conclusion that a process similar to the Baeyer-Villiger oxidation must occur during these degradations. The general scheme for the formation of testololactone from progesterone can thus be described, as shown in Fig. 16.5-4. It involves four successive steps; first a Baeyer-Villiger oxidation of the steroid sidechain leading to a testosterone acetate, secondly an esterase hydrolysis, thirdly oxidation of the C17 hydroxyl leading to the corresponding 3,17-dione and finally a second Baeyer-Villiger oxidation of this diketone at the D-ring leading to the corresponding 8-lactone. It has been shown in the fungus Cylindrocarpon radicicola that one bifunctional enzyme is involved in these transformations, which is able to catalyze oxygenative esterification of 20-ketosteroids as well as oxygenative lactonisation of 17-ketosteroid~[~~~ 241. It is noteworthy that all the above investigations into steroid substrates for lactonization were conducted on single enantiomers and thus, no reference to the enantioselectivity of the processes had been recorded.

I

16.5 Baeyer-ViUiger Oxidations 1205

L

O

I

PAC

4 -& 0

0

progesterone

androstenedione

testosterone acetate I

testosterone

testololactone Figure 16.5-4. Mechanism of the biotransformation of progesterone into testololactone.

16.5.1.2

Aliphatic Substrates

Baeyer-Villiger oxidation has also been reported for aliphatic ketones. Several strains able to grow on various aliphatic or alicyclic substrates have been isolated, and it has been shown that their degradation often involves a Baeyer-Villiger oxidation. For example, it has beeen observed that Pseudomonas multivorans, Pseudomonas aeruginosa, Pseudomonas cepacia and Nocardia sp. are able to grow on tridecan2-one P-281. Forney and Markovetz isolated undecyl acetate directly from growing cultures of Pseudomonas aeruginosa. They showed that all early intermediates in the pathway arise biologically and sequentiallyfrom their precursors, indicating involvement of a Baeyer-Villiger type oxidation. In a further study they also showed that cell-free

1206

I

16 Oxidation Reactions

tridecanone

0

J. undecyl acetate

-0

undecanol

O H-

undecanoic acid

-

0

undecyl undecanoate

Figure 16.5-5. Degradation of tridecan-2-one with a crude cell-free preparation from a Pseudomonas aeruginosa strain.

preparations obtained from methylketone grown Pseudomonas aeruginosa, when supplemented with NADH or NADPH in the presence of 0 2 , carry out a reaction sequence visualized in Fig. 16.5-5. Using Pseudomonas cepacia grown on tridecan-&one, Markovetz and coworkers[28] later showed that experiments conducted with I8O2 led to 84% incorporation of into the C - 0 - C linkage, rather than into the carbonyl function, indicating the occurrence of a Baeyer-Villiger type process. They also observed that the undecyl esterase involved in the degradation process is able to hydrolyze both aliphatic and aromatic acetate esters. They also reported that this enzyme is strongly inhibited by organophosphates such as tetraethylpyrophosphate (TEPP), as well as by other esterase inhibitors like p-chloromercuribenzoate1271. A similar degradation pathway was described for oxidation of tetradecane and 1-tetradecene with Penicillium sp. L2’I. Similar mechanisms were proposed for the degradation of other aliphatic substrates such as butan-2-one[281, acetol l3O1, acetophenone 13’1 and l-phenylethan~lr~~]. Interestingly,cell extracts of Nocardia sp. LSU 169 grown on butan-2-onewere also shown to be capable of oxidizing tridecan-2-one. Generally,the Baeyer-Villiger reaction was followed by an esterase catalyzed hydrolysis [331.

16.5 Baeyer-Villiger Oxidations Figure 16.5-6.

w,,

-&

Pseudomonas sp.

Degradation of Z-heptylcyclopentanone by a Pseudomonas sp.

C7H,,

5%

16.5.1.3

Alicyclic Substrates

Baeyer-Villiger oxidation is also a common feature during the catabolic degradation of a variety of other compounds, including monocyclic, bicyclic or polycyclic molecules. For monocyclic compounds, one of the first reports describing formation of a lactone from racemic a-substituted cyclopentanone by various Pseudomonas sp. was by This could be regarded as the first indication that these reactions were to prove of interest for asymmetric synthesis since the lactone product displayed some optical activity (Fig. 16.5-6). Further studies showed that other substrates such as cyclopentanolr3’1, cyclohexane [36391, cyclohexanol [40-421, cyclohexan-l,2-diol[43-451, cycloheptanone [461 and, more recently, cycl~dodecane[~~] were degraded via analogous pathways. These were studied using bacterial strains including Pseudomonas sp. NCIMB 9872 L3’, *I, Nocardia globerula CL1 14’1, Acinetobacter TD 63 [431, Acinetobacter calcoaceticus NCIMB 9871L3’1, Xunthobacter sp. L3’1 and Rhodococcus All these degradation pathways were shown to involve a Baeyer-Villiger oxidation of a cycloalkanone that led to formation of the corresponding lactone. Further degradation then occured via hydrolysis of this lactone by a lactone hydrolase which has, in some cases, been isolated. As an example, the reaction sequence for the degradation of cyclopentanol by Pseudomonas sp. NCIMB 987213’] is shown in Fig. 16.5-7. A pathway for the degradation of (-)-menthol and menthane-3,4-diol by a bacterium classified as a Rhodococcus sp. was proposed by Shukla and coworkers. Again, the proposed scheme involves formation of the corresponding lactone by a Baeyer-Villiger process[49].Interestingly, an identical process has been shown to occur in the degradative pathway of menthol and menthone in peppermint (Mentha Rhodococcus erythropolis DCL 14[”] has also been reported to piperita) rhizomes .I‘’[ degrade menthone in addition to 1-hydroxy-2-0x0-limonene and dihydrocarvone via an enzymatic Baeyer-Villiger reaction. Some other monocyclic compounds bearing ketonic side chains have also been shown to undergo degradation processes involving Baeyer-Villiger type oxidation. For example, oxidation of p-ionone by Lasioplodiu the~bromae[’~] affords, among other products, the alcohols shown in Fig. 16.5-8.In this case, the loss of two carbons from the sidechain has been attributed to a contribution of Baeyer-Villiger oxidation followed by ester hydrolysis and reduction. Similar results were described by Nespiak and coworkers in the course of their study of cyclopentyl ketones by Acremonium roseum (Fig. 16.5-9).When R = CH3 or

1208

I

y:

71 Oxidation Reactions

NA<

cyclopentanol dehydrogenase

NADPH cyclopentanone oxygenase NADP’

valerolactone hydrolase

?OH

4 T

CH,OH

H-0 L

hydroxyvalerate dehydrogenase and oxovalerate dehydrogenase NADPH (CH)--CozH 3

\

~

~

2

~

-

Acetyl-CoA

Figure 16.5-7. Reaction sequence for the oxidation o f cyclopentanol by Pseudomonas sp. NCIMB 9872.

C2H5, the alcohol formed via Baeyer-Villiger oxidation and ester hydrolysis was the only product isolated after 2 days. However, higher esters (R = n-or i-C3H7, R = nbutyl) have also, if not predominantly, some amount of allylic oxidation product. In this study, it was shown that the (S)-enantiomer of the substrate methyl ester was oxidized more rapidly than the (R)-isomer and that the reaction proceeded with retention of configuration at the chiral center. Thus, by using short incubation times (2 days) the racemic substrate led to the (S)-alcohol,but the optical purity was low (around 20 %). Although interesting, this apparent enantioselectivity could also be due eventually to an enantioselectivehydrolysis of the intermediate ester or to some other catabolic pathway. However, the butyl ketone led to the (R)-alcoholshowing 100% optical purity. An extensive study by Fuganti and coworkers [54-571 showed that the metabolism of 4-(4-hydroxyphenyl)butan-2-one (“raspberryketone”) by the fungus Beauveria bassiana unexpectedly yielded tyrosol, through insertion of oxygen via a Baeyer-Villiger reaction and subsequent acetate (Fig. 16.5-10).Only a narrow range of

1 6 5 Baeyer-Villiger Oxidations 11209

S-ionone esterase

koH reductase

--&?

Figure 16.5-8.

Biotransformation o f 6-ionone by Lasiodiploida theobromae.

Acremonium roseum

-

R Me Bu Figure 16.5-9.

alcohol

(+)-s o.p.= (-)-R

O.P.

20%

= 100%

Transformation of cyclopentyl ketones by Acremonium roseum.

substrates was converted in this manner, The authors were able to show via deuterium incorporation experiments, that the configuration of the migrating carbon-carbon bond was retainedIS61,this being a defining characteristic of the peracid-catalyzedBaeyer-Villiger process. Camphor and its analogs are the most studied bicyclic substrates for the biological It has Baeyer-Villiger r e a c t i ~ n [ ~ ~ ~ been ~ ] . shown by Gunsalus and coworkers that, in the early steps of D-(+)-camphoroxidation by Pseudomonas putida C1, both alicyclic rings are cleaved by lactonization reactions: Thus, the conversion of (+)-camphorto 5-keto-1,Zcampholide involves three reactions; hydroxylation, oxidation and lactonization. Using a different Pseudomonas strain, the non-hydroxylated campholide has been isolated, suggesting that in this case lactonization occurs prior to hydroxylat i ~ n [ ~ *ItI was . also shown, by analysis of extracted metabolites, that an analogous, enantiocomplementary pathway existed for the metabolism of L-(-)-camphor. Several further studies have been devoted to clarifylng these steps. Interestingly, it has been shown that P. putida does not express lactone hydrolases that are active towards

1210

I

7G Oxidation Reactions

Beauveria bassiana ATCC 71 59

HO raspberry ketone

I

tyrosol Figure 16.5-10. Biotransformation of raspberry ketone to tyrosol by Beauveria bassiana ATCC 7159.

the lactone intermediate. The intermediate bicyclic lactone is unstable under reaction conditions, and spontaneously opens to a cyclopentenone. This is then again oxidized via a Baeyer-Villiger reaction to the corresponding lactone. The degradation ofthe enantiomers of camphor is shown in Fig. 16.5-11.Three enzymes catalyzing the Baeyer-Villiger reaction, i. e. 2,s-diketocamphane 1,2-monooxygenase [which forms the bicyclic lactone analog of (+)-camphor][641, 3,G-diketocamphane 1,G-monooxygenase[which forms the bicyclic lactone analog of (-)-camphor]16'1 and 2-0x0-A3-4,5,5-trimethylcyclopentenylacetyl-Co-A monooxygenase (which catalyzes the lactonization of the monocyclic intermediate) [631 from Pseudornonas putida ATCC 17453 have been purified to homogeneity and thoroughly characterized. Enzymatic Baeyer-Villiger oxygenations are not restricted to microbial cells. It has been shown that (+)-camphor,a major constituent of the volatile oil of immature sage (Salvia oficinalis L.) leaves, is converted into a water soluble metabolite via enzymatic lactonization to 1,2-campholide, followed by conversion into the 8-Dglucoside-6-0-glucoseester of the corresponding hydroxy acid[", 691. The oxidation of racemic fenchone by a Corynebacteriurn SP.[~'] (reclassified as Mycobacteriurn rhodochrous), an organism which grows at the expense of either (+)- or (-)- camphor, has also been reported. This was shown to lead, in a 45% yield, to a 90/10 mixture of 1,2 and 2,3-fencholides, as shown in Fig. 16.5-12. This result contrasts with the chemical oxidation of fenchone with peracetic acid, where 2,3-fencholide is the major product in a 40/60 mixture. Accumulation of these lactones is a priori surprising as compared with the total degradation of the structurally similar camphor substrate. However this may simply be due to the fact that this lactone, unlike that formed from camphor, is chemically stable in the medium. Of course, one has also to assume that, here again, the strain is devoid of any lactone hydrolase. This bioconversion was the first gram-scalepreparative report

7 6.5 Baeyer-Villiger Oxidations

(+)-camphor

4

(-)-camphor

Pseudomonas putida C1

so

J.

& 'OH

HO

$.

+

diketocamphane monooxygenases

P

J

0

\COSCoA

2-A3-4,5,5-trimethylcyclopentenylacetate Co-A ester monooxygenase

& --

0

I

COSCoA

Figure 16.5-11. Metabolism of both enantiomers of camphor by Pseudomonas putida C1 (= NCIMB 10007=ATCC 17453).

of a non-steroidal product, yet no indication of any enantioselectivity for this reaction was presented. Similarly, it has been shown that 1,8-cineoleand 6-0x0-cineoleare degraded via the scheme shown in Fig. 16.5-1317*1.As in the case of camphor, the first step involves a

I

1211

1212

I

16 Oxidation Reactions

9 fenchone

1

1,2-fencholide

Figure 16.5-12.

2,3-fencholide

Oxidation of racemic fenchone to the corresponding fencholides.

hydroxylation, followed by oxidation of the alcohol to form 6-oxocineole.This is then processed via a Baeyer-Villiger reaction leading to a lactone which is spontaneously opened to the hydroxy acid. 16.5.1.4

Polycyclic Molecules

Enzymatic Baeyer-Villiger reactions have also been described in the metabolic processing of larger, polycyclic non-steroidal molecules (Fig. 16.5-14).This is the case for the biosynthesis of aflatoxin B 1 where it has been demonstrated that formation of versiconal acetate intermediate from avemfin occurs via such a Similarly, aflatoxin G1 was shown to be formed from aflatoxin B[731 whereas degradation of the anthraquinone questin to desmethylsulochrin was shown to imply a Baeyer-Villiger process [741. Furthermore, biological Baeyer-Villiger reactions have been reported in the biosynthesis of polyketides such as DTX-417’] and the aureolic acid antibiotics such as mithramycin 1762 771. The oxygenase MtmOIV from Streptomyces argillaceus responsible for cleavage of the fourth ring of premithramycin B is unique amongst those responsible for biological Baeyer-Villiger reactions, in that it displays sequence homology not with other “Baeyer-Villiger monooxygenases” (vide infa), but with flavin-type hydroxylases encoded in polyketide synthase gene clusters from other Streptomyces spp. [761.

1

6-0x0 cineole

*

c

o

2

y

&cozH

6-0x0-cineole Figure 16.5-13.by Degradation a Rhodococcuso f

HO SP.

w

HO

\

/

0

7 G. 5 Baeyer-Villiger Oxidations

1213

averufin

0

Aspergillus parasiticus

*:

versiconal acetate

HO

0

4P

HO

questin

/

\

0

1

Aspergillus terreus

desmethylsulochrin HO Figure 16.5-14.

I

CO,H Involvement of enzymatic Baeyer-Villiger processes in the degradation

of non-steroidal polycyclic compounds.

16.5.2

Baeyer-Villiger Monooxygenases

The reactions described above illustrate that there are numerous metabolic routes wherein biological Baeyer-Villiger reactions have been implicated. The synthetic potential of the enzymatic Baeyer-Villiger reaction has dictated that intensive

1214

I

76 Oxidation Reactions Figure 16.5-15. Riboflavin derivatives: the coenzymically active forms o f flavoprotein.

HO

R =H

riboflavin

research efforts have been devoted to studying the nature of the enzymes that catalyze these reactions. Enzymes that catalyze the Baeyer-Villiger reaction are a subset of the flavin monooxygenases. In the mechanism of oxidation catalyzed by such enzymes one atom of molecular oxygen is incorporated into the substrate, whereas the other is reduced to HzO.Two cofactors are required for catalyticactivity. The first is a reduced flavin (FAD or FMN) bound non-covalently in the active site. The riboflavin moiety of flavin monooxygenase holoproteins is shown in Fig. 16.5-15; the second is a reduced nicotinamide cofactor (NADPH or NADH),which is required to furnish the enzyme with electrons to reduce the flavin. Several Baeyer-Villiger monooxygenases (BVMOs) have been purified and in rare cases, the relevant genesm cloned and expressed. Some of these are listed in Table 16.5-1. There appear to be two types of BVMOs. Type 1are homogeneous; both flavin reduction and substrate oxygenation are carried out on a single polypeptide, these are most usually FAD and NADPH dependent. Type 2 are heterogeneous, a substrate oxygenating subunit appears to require a separate flavin reductase/NADH dehydrogenase in order to generate reduced flavin. Type 2 BVMOs are usually FMN and NADH dependent. 16.5.2.1 Type 1 BVMOs

Cyclopentanone monooxygenase, which catalyzes the conversion of cyclopentanone to valerolactone, has been isolated from Pseudomonas sp. NCIMB 9872[46,781. This has been shown to be made up of three identical subunits, each using one FAD equivalent, and to be NADPH dependent. Cyclohexanone monooxygenases have been purified from Acinetobacter calcoaceticus NCIMB 9871 and Nocardia globerula CL1 r4'1 and Rhodococcus copr~philus[~']. These enzymes were shown to be single polypeptides and to be FAD and NADPH dependent. Tridecanone monooxygenase from Pseudomonas cepacia is a dimer of two identical subunits, however, but is also FAD plus NADPH dependent["]. A cyclohexanone monooxygenase from Xanthobacter sp. is unusual in that it is dependent on FMN, but NADPH as a nicotinamide cofactor[82].Steroid monooxygenase from Rhodococcus rhodochrous [831 and monocyclic monoterpene ketone monooxygenase from Rhodococcus erythropolis DCL 14[511

123 (55 each)

NADPH

2 identical subunits

1

Cyclohexanonemonooxygenase Xanthobacter sp. Monocyclic monoterpene ketone monooxygenase Rhodococcus erythropolis DCL 14 Cyclohexanonemonooxygenase Rhodococcus coprophilus Steroid monooxygenase Rhodococcus rhodochrous

FAD FAD

58

60

NADPH NADPH

single polypeptide single polypeptide

1

1

9.0

FAD

60 NADPH

single polypeptide

1

8.8

FMN 50

NADPH

single polypeptide

1

7.8

7.2

9.0

7.8-8.0

1 FAD per subunit

115 (56 each)

NADPH

1 FMN per subunit

2 identical subunits

72 (36 each)

NADH

2 identical substrate oxygenating subunits + NADH dehydrogenase

2

3,6-Diketocamphane1,6monooxygenase Pseudomonas putida ATCC 17453

1 FMN per subunit

1 FAD

1 FAD

1

78 (39 each)

NADH

2 identical substrate oxygenating subunits + NADH dehydrogenase

2

2,s-Diketocamphane 1,2monooxygenase Pseudomonas putida ATCC 17453

Steroid monooxygenase Cylindrocarpon radicicola ATCC 11011

106

NADPH

2 identical subunits

1

2-0x0-A3-4,5,5-trimethylcyclopentenyI acetyl Co-A monooxygenase Pseudomonas putida ATCC 17453

53

NADPH

single polypeptide

1

8.4

9.0

1 FAD

59

NADPH

single polypepetide

1

Cydohexanone monooxygenase Acinetobacter calcoaceticus NCIMB 9871 Cyclohexanonemonooxygenase Nocardia globurela CL 1 2-Tridecanonemonooxygenase Pseudomonas cepacia 1 FAD

7.7

1 FAD per subunit

200 (54-58 each)

NADPH

3-4 identical subunits

1

Optimum pH

Mole offlavin/ mole of protein

Native molecular mass x 1000 Da

Cofictor specificity

Subunit structure

Cyclopentanone monooxygenase Pseudomonas NCIMB 9872

Number o f proteins

Characteristicsof various Baeyer-Villigermonooxygenases.

Enzyme and source

Table 16.5-1.

1641

~ 3 1

1801

1811

1811

1481

Reference

1216

I are also Type

16 Oxidation Reactions

1 BVMOs as is the 2-oxo-A3-4,5,5-trimethylcyclopentenyl acetyl Co-A monooxygenase from Pseudomonas putida ATCC 17453 [631. Cyclohexanone monooxygenase (CHMO, E.C. 1.14.13.X) is by far the most studied Type 1 BVMO and has been used extensively for as a model for mechanistic studies and as a catalyst in synthesis (vide inpa). CHMO was purified from Acinetobacter calcoaceticus NCIMB 9871 grown on cyclohexanol as the sole carbon source, by Tmdgill and coworkers["]. It was found to be active as a monomer and to contain one non-covalently bound FAD molecule per monomer. The gene was cloned and the protein expressed in Escherichia c0li['~1 and more recently in Saccharomyces cerivi~iae[~~I. Each subunit is a polypeptide of 542 amino acids and, although no definitive structure of a BVMO has yet been published, a potential flavin binding site at the N-terminus was identified, in addition to a potential NADP binding site. Analysis of the sequence reveals that the N-terminus of the enzyme bears strong homology with the FAD binding domain of other flavoproteins such as glutathione reductase from Escherichia coli. 16.5.2.2 Type 2 BVMOs

The diketocamphane monooxygenases (DKCMOs) from Pseudomonas putida ATCC 17453 involved in camphor degradation, are FMN plus NADH dependent and are heterogeneous, consisting of two identical substrate oxidizing polypeptides and an NADH dehydrogenase. The enzymes have been purified, extensively characterized["* 671 and their N-terminal amino acid sequences determined[86].These data showed the oxygenating subunits of the DKCMOs to have homology with the NADH an enzyme which catalyzes the plus FMN dependent luciferase of Vibrio ha~eyi['~], Baeyer-Villiger oxidation of dodecanal to dodecanoic acid with the release of a photon of light. The application of the DKCMOs enzymes to synthesis has also been investigated (vide inja) and, whilst the genes encoding these proteins have not been identified, preliminary X-ray crystallographic data on 3,G-diketocamphane-l,Gmonooxygenase has been reported["]. 16.5.2.3 Mechanism of the Enzymatic Baeyer-Villiger Reaction

The mechanism of the enzymatic Baeyer-Villiger oxidation, with reference to CHMO, has been studied by the group of W a l ~ h I"1~ ~who . proposed the scheme shown in the top cycle in Fig. 1G.5-lG.The tricyclic isoalloxazine ring is the center of catalysis. Initially, the exogenous reductant NAD(P)H acts as the electron donor to afford the reduced flavin. This can be readily reoxidized by both one-electron or two electron processes in the presence of 0 2 to yield a 4-a-hydroperoxyflavin.This intermediate undergoes an 0-0 bond fission upon nucleophilic attack on an electrophilic ketone substrate, a mechanism similar to the chemical Baeyer-Villiger oxidation of ketones by peracids. This initially affords the 4-a-hydroxyflavinwhich, by loss of H20 regenerates the starting FAD for a subsequent catalytic cycle.

7G.S Baeyer-Villiger Oxidations I1217

U Figure 16.5-16. Proposed mechanisms for the enzymatic Baeyer-Villigeroxidation of cyclohexanone.

1218

I

16 Oxidation Reactions

However, the FAD-4-a-OOHcan also break down directly via liberation of H202. A variation on this model has recently been proposed by Kelly et al.l7] who suggested that the hydroxy group of the Criegee intermediate could not be immobilized in such a mechanism, and that unreasonable steric constraints would be imposed for many of the substrates transformed reported for these enzymes. A new tautomer of the the flavin hydroperoxide was proposed as part of an alternative scheme (lower cycle, Fig. 16.5-16) in which an intermediate trioxane decomposes to yield the lactone and flavin hydrate. In addition to ketone substrates, the 4-a-hydroperoxyflavin can also react by nucleophilic attack on other molecules. Thus, boronic acid substrates were transformed into the corresponding alcohols via the intermediate borate esters as hydrolytically labile initial enzyme products r90* 911. However, the 4-hydroperoxyflavin,acting in these cases as an electrophile, is also able to oxidize other nucleophilic substrates and in particular heteroatoms such as and phosphorous 1911. Indeed, CHMO oxygensulfur[911,selenium1"~ 921,

NADPH, 0, enz-FAD

0

Ca

(5

Q

0 NADPH, 0,

NADPH, 0,

____)

enz-FAD

NADPH, 0, _____)

enz-FAD

NADPH, 0, -

enz-FAD

f

enz-FAD

$1

(5 S

0-s

S

f

f

O"-s

I

NADPH, 0, ____)

enz-FAD I

I

Figure 16.5-17. Studies on heteroatom oxidation using purified cyclohexanone mono oxygenase

76.5 Baeyer-VilligerOxidations

ates trimethyl phosphite to trimethyl phosphate, sulfides to sulfoxides (one equivalent) or sulfones (two equivalents). If 3- or 4-thiocyclohexanones were used as substrates, these were converted exclusively into the lactone products, showing that Baeyer-Villiger oxidation is preferred in these cases (Fig. 16.5-17).The synthetic applications of heteroatom, notably sulfur, oxidation by BVMOs have been thoroughly explored and and reviewed L94. These results illustrate that reactions performed by BVMOs are similar to those of peroxide containing reagents (hydrogen peroxide, alkyl hydroperoxides or peracids), which are able to deliver either a formally nucleophilic or a formally electrophilic oxygen atom to a substrate. Indeed, whereas Baeyer-Villiger oxidation or boronic acid oxygenation involve initial attack of a nucleophilic oxygen, the sulfide, selenide or phosphite ester oxygenations require the transfer of an electrophilic oxygen to a nudeophilic electron pair of the substrate. Interestingly, no epoxidation of olefinic double bonds by BVMOs have been reported The substrate selectivity of CHMO was first explored by Trudgill and coworkers [70, 81],who demonstrated that the enzyme processes C4-C8 cyclic ketones. The migratory aptitude of the enzymatic oxygen insertion process was probed initially with two types of substrates. First, in an attempt to explore the stereochemical mode of these reactions, Schwab et al. studied the Baeyer-Villiger oxidation of (2R)deuterated cyclohexanone. Detailed NMR multinuclear spectroscopic studies led to the conclusion that CHMO catalyzes the conversion of cyclohexanone to E-caprolactone with complete retention of configuration at the migrating carbon center ["I, a result identical to the chemical route (Fig. 16.5-18).To eliminate the possibility of an was also enolization and/or rearrangement route, 2,2,6,6-tetradeuterocyclohexanone incubated with the enzyme. The fact that no loss of deuterium was observed by GC again militates in favor of a mechanism similar to that proposed for chemical Baeyer-Villiger oxidation. In an elegant further study["^, these authors confirmed their preliminary proposal of the (R)-absolute configuration of the starting 2-deuterocyclohexanone as well as the occurrence of a total retention of configuration of the CHMO catalyzed Baeyer-

(EnzFAD-0-\OH

Enz-FAD-OH

0.

Figure 16.5-18.

Stereochemical studies using deuterated cyclohexanone.

?

D

-

<

1220 IG Oxidation Reactions

I

1-0xepanone + H,O

cyclohexanone + 0,

cyclohexanone oxygenase

NADPH

NADP’

G-6-PDH

6-phosphogluconolactone Figure 16.5-19.

glucose-6-phosphate

NADPH recycling in the course of CHMO catalyzed oxidation.

Villiger reaction, Interestingly, they described for the first time that the efficient conversion of ketone into lactone could be brought about in the presence of a catalytic amount of NADPH, with cofactor recycling accomplished by the glucose6-phosphate dehydrogenase as shown in Fig. 16.5-19. In order to test enzyme regio- and enantioselectivity rigorously, Schwab and coworkers also studied the asymmetric substrate 2-methylcyclohexanone. A “virtual racemate” made up of equivalent quantities of (2R)-2-[methyl-2H3]methylcyclohexanone and of (2S)-[methyl-13C]methylcyclohexanone (each one prepared by different methods) was studied, using a multinuclear NMR technique. The conclusions from is this experiment were two fold. First, they confirmed that 6-methyl-~-caprolactone the only reaction product, thus indicating a total regioselectivityof oxygen insertion into the “more substituted carbon-carbon bond. Second, these results showed for the first time a two-fold rate difference between transformation of the two enantiomers of cyclohexanone. This was an interesting result that suggested that CHMO could show far greater discrimination toward enantiomers of a substrate that bore a far bulkier C2 substituent. Finally, the measurement of the reaction kinetics for each one of the substrate enantiomers showed that, after about 50 % reaction, there is a progressive decrease in both the degree of enantioselectivity as well as the absolute rate of lactonization of the two substrate enantiomers. However, the reasons for these diminishing rates of reaction were not clear. Further work exploring the migratory aptitude of different substituents has been described[”]. Phenylacetone is converted into benzyl acetate, showing exclusive benzyl migration in accordance with the chemical reaction achieved with trifluoroacetic acid. Different results were observed with phenacetaldehyde, where an inverted preference is seen as compared with the peracidic reactions. Using purified CHMO from A. calcoaceticus NCIMB 9871, Taschner and coworkers r9‘, 971 showed that several prochiral substrates including some 4-substituted cyclohexanoneswere efficiently converted into their corresponding lactones, each of them showing very high enantiomeric purities (Fig. 16.5-20).Thus, CHMO prove to

76.5 Baeyer-Villiger Oxidations

Substrate

M

e

Product

0

% yield ('77 e.e.)

80 (>98)

0

73 (>98)

27 (>98)

d

Me0

25 (>98)

76 (75)

88 (~98)

*oLoH 0

73 (9.6)

Figure 16.5-20.

CHMO catalyzed oxidation o f various prochiral substrates.

be extremely effective at discriminating between the two sides of the carbonyl function of such prochiral substrates. However, the presence of an alcohol or methyl ether function at position 4 leads unexpectedly to products of lower ee values.

I

1221

1222

I

IG

Oxidation Reactions

16.5.3

Synthetic Applications

With the exception of steroid type substrates, the results described up to now have dealt with small-scaleanalytlcal studies. However, in view of the potential of BVMOs for regio- and even enantioselective transformations of various substrates, studies into the scale-up of these transformations began in earnest soon after these earlier investigations, followed by considerations of their application in chiral organic synthesis. In this context, Abril et al.[”] examined a variety of readily available ketones in order to determine the substrate selectivity, regioselectivityand enantioselectivity of CHMO immobilized in a polyacryalamidegel. They also used the NADPH recycling system previously desribed by Schwab for in situ regeneration of this cofactor. These experiments showed that 2-norbornanone, L- and D-fenchone, (+)-camphor and (+)-dihydrocarvoneare processed by CHMO. In a typical experiment, 10.2 g of were obtained from 11.4 g of 2-norbornaracemic 2-oxabicyclo[3.2.l]octan-3-one none, using 1.7 g of NADP cofactor. The authors concluded that the enzyme did not display a useful degree of enantioselectivity,therefore offering no major advantages over chemical oxidation. One major drawback of employing CHMO as a catalyst is the necessity to regenerate the expensive nicotinamide cofactor NADPH. One strategy for circumventing this problem is use of whole-cellpreparations of microorganisms for BaeyerVilliger oxidations. One early example of this technique involved the oxidation of 2,2,5,5-tetramethyl-1,4-cyclohexanedione to the optically pure (S)-ketolby Cuwularia lunata described by Azerad and coworkers[’9]. They showed that during the fungal reaction of the dione, as shown in Fig. 16.5-21, the already formed (S)-ketolwas isomerized to its five-membered isomer. Moreover, when submitted to appropriate culture conditions, the racemic ketol afforded the (S)-lactone(81% ee) as well as the unchanged (R)-lactol of 97% ee The remaining substrate could then be further treated by rn-chloroperbenzoic acid to afford the (R)-hydroxylactoneenantiomer. Extensive studies have been performed on the microbial Baeyer-Villiger oxidation of bicyclic [3.2.0] ketones and analogues. These studies were prompted by the important findings of Furstoss and coworkers[’00],who determined that the oxygenation of bicyclo[3.2.0]hept-2-en-G-one using Acinetobacter sp. TD 63 led to two regioisomeric lactones in equal quantities and almost quantitative yield. The first arises from the “normal” oxygen insertion mode into the more substituted carboncarbon bond, whereas the second is the result of an oxygen insertion into the less substitiuted bond leading to the so-called “abnormal” lactone. Moreover, both these lactones were of high optical purity i. e. showing a 98% ee for the (-)-(l S, 5R) isomer and a 95% enantiomeric excess for the (-)-(1R. 5s) enantiomer. These results appeared to suggest that biological Baeyer-Villiger oxidations could indeed be used for the large-scale preparation of optically active lactones. In the case of bicyclo[3.2.0]hept-2-en-G-one, each one of the substrate enantiomers reacts with a different and divergent regioselectivity for the oxygen atom insertion. This result is noteworthy since it describes for the first time such an almost perfect

76.5 Baeyer-Villiger Oxidations

4Y

Figure 16.5-21. ’ Baeyer-Villiger oxidation o f 2,2,5,5-tetramethyl-1,4-cyclohexane-dione by Curvularia lunata.

0

Curvularia lunata

$ OH

4 OH

regio- vs. enantioselectivity for the Baeyer-Villiger oxygenation. A more complete study[lol],aimed at exploring the synthetic potential of these reactions, confirmed that this enantiodivergent selectivity is not restricted to one particular substrate but is a general phenomenon within a series of similar compounds. Two strains of bacteria, Acinetobacter sp. TD 63 and A. calcoaceticus NCIMB 9871 were used throughout this study and led to almost identical results. In most cases, both “normal” and “abnormal” lactones were obtained in approximately 1: 1 ratios and with almost quantitative yields. Also, it was observed as shown in Fig. 16.5-22 that the “abnormal” lactone, which is not accessible using conventional Baeyer-Villiger oxidation, always shows very high ee values, whereas the enantiomeric purity of the “normal” lactone is somewhat lower for the substrate bearing a saturated sixmembered ring. Both of these lactones are interesting chiral synthons; the “normal” one being an important chiron for prostaglandin synthesis. It is noteworthy that all lactones of a particular type are formed from the same enantiomer of the starting ketone: thus, the substrate enantiomer bearing an (S)-configurationat the bridgehead carbon atom a to the carbonyl group leads to the “normal” lactones, whereas the (R)-configuration affords the “abnormal” ones. Similar results were obtained in the course of a study conducted on bicyclic

I

1223

1224

I

16 Oxidation Reactions

A. calcoaceticus NCIMB 9871

,ao

44%

> 95% e.e.

36%

> 95% e.e.

43%

> 95% e.e.

41yo 86% e.e.

52%

60% e.e. Figure 16.5-22.

42%

> 95% e.e.

31yo

> 95% e.e.

37%

> 95% e.e.

36%

> 95% e.e.

28%

> 95% e.e.

Oxidation of various In.2.01 bicyclic ketones with Acinetobacter calcoaceticus

NCIMB 9871.

substrates bearing an oxygen atom in the five or six-membered ring[’02](Fig. 16.523). Here again, equivalent ratios as well as high ee values were obtained for both the ‘normal’and “abnormal”lactones. Since the lactones are unreported in the literature in their optically inactive form, detailed studies using circular dichroism were conducted in order to attribute the absolute configuration of the products. Whilst the whole-cell approach has proved invaluable, the associated problems of overmetabolism and side reactions can be encountered. Another way to counter the problems of high cost in using isolated BVMOs is to use an NADH dependent enzyme, as NADH retails at approximately one tenth of the cost of NADPH. The Type 2 DKCMOs from Pseudornonas putida ATCC 17453 (= NCIMB 10007) are NADH dependent, and Grogan et al. were successful in applying a complement of these enzymes, termed M 0 1 , to the transformation of bicyclo[3.2.0]hept-2-en-6-one, to yield another enantiodivergent mix of lactones enantiomeric to those obtained

76.5 Baeyer-Vihger Oxidations

A. calcoaceticus NCIMB 9871

*

("3::::Fo 35%

91% e.e.

35%

99% e.e.

34%

98% e.e.

33%

+

32%

> 99% e.e.

35%

97% e.e.

42% > 99% e.e.

33%

72% e.e.

97% e.e.

60%

18% > 99% e.e.

35% e.e.

Figure 16.5-23. Oxidation of various 0x0-[n.2.0] bicyclic ketones with Acinetobacter calcoaceticus NCIMB 9871.

with A. calcoaceticus NCIMB 9871/TD 63. The use of NADH dependent enzymes is also important in this context, as it allows use of the NAD dependent formate dehydrogenase/sodium formate recycling strategy for cofactor regeneration[lo3I, reducing costs still further. Interestingly, the separated isoenzymes, 2,s-diketocamphane 1,2-monooxygenase and 3,G-diketocamphane 1,G-monooxygenase were shown to have different selectivities for this transformation, compromising the result obtained with M 0 1 [lo41(Fig. 16.524). Further transformations of this ketone by luminescent bacteria containing NADH dependent luciferases (also Type 2 BVMOs) have also been reported [lo5],although characterization of cell-free systems employing these enzymes has not been investigated further. The biotransformation of bicycl0[3.2.0]hept-2-en-G-one using whole cell suspensions of the fungus Cylindrocarpon destructans gave not only different ratios of both lactones depending on the degree of conversion, but also no enantioselectivity was

1226

I

7G Oxidation Reactions

0

NADH-dependent BVMOs 0 from Pseudomonas putida

NADH

a

co*

NAD’ /

Na+O,CH formate dehydrogenase

‘M01‘

63%, 60% e.e.

37%, 95% e.e.

2,5-DKCMO

57%, 82% e.e.

43%, 100% e.e.

3,6-DKCM0

17%, 10% e.e.

13%, 72% e.e.

Biotransformation of bicyclo[3.2.0]hept-2-en-6-one by NADH dependent BVMOs from camphor grown Pseudomonas putida ATCC 17453.

Figure 16.5-24.

observed [‘06]. Further fungal biotransformations described by Carnell and Willetts showed that a series of dematiaceous fungi were also able to lactonize the same substrate [‘071. These included various Cuwularia and Dreschlera species. Some of these fungi produced both regioisomeric lactones with a high degree of stereoselectivity, whilst others produced mostly the 3-oxa lactone. The test strains of Curvularia lunata and Dreschlera australiensis gave lactones with equal and almost opposite degrees of regio- and stereoselectivity. Importantly, the biotransformation of bicyclo[3.2.0]hept-2-en-G-one by another fungus, Cunninghamella echinulata NRRL 3655, is unique in that it results in a resolution of the parent substrate to yield only the “abnormal” (-)-(lR, 5S)-3-oxalactone in 30% yield and 95% ee[108].This chiral synthetic intermediate has been used to synthesize both single enantiomer cyclosarkomycin[’08]and the marine brown algae pheremones (+)-multifidene and (+)-viridiene[lo’](Fig. 16.5-25). Further reports by Furstoss and coworkers concerned Baeyer-Villiger oxidation of a-substituted cyclopentanones[’lo].Using the same two Acinetobacter strains used previously, this study aimed to explore the possibility of synthesising optically active 6-lactonesbearing aliphatic chains, these compounds being of particular interest as chiral synthons. This study showed that various lactones of (S) configuration can be obtained in fair yields with moderate to excellent ee values depending on the chain length and on the conversion ratio. Using Acinetobacter calcoaceticus NCIMB 9871 it was, however, necessary to run these biotransformations in the presence of tetraethylpyrophosphate (TEPP), a well known inhibitor of hydrolases. This was necessary in order to avoid hydrol$c degradation of the 6-lactonesformed. The use of this inhibitor was, however, unnecessary when using the Acinetobacter sp. TD 63 strain which is known to lack a lactone hydrolase. One interesting application of this study was the preparative two-step synthesis of both enantiomers of 5-hexadecanolide, a

7 6.5 Baeyer-Villiger Oxidations

Cunninghamella echinulata

03-

I

1227

(-)-(1 R, 5S)-cyclosarkomycin

steps

35% 95%

(+)-(3R,4S)-viridiene steps

(+)-(3S, 4S)-multifidene Figure 16.5-25. Biotransformation of bicyclo[3.2.0]hept-2-en-6-one by Cunninghamella echinulata NRRL 3655 and synthetic targets.

CllH23

A. calcoaceticus

> lh

ko

I

A. calcoaceticus Figure 16.5-26.

mCPBA

* $c~,Ha

Baeyer-ViIIiger oxidation of a-undecylcyclopentanone: synthesis o f either enantiomer of hexadecanolide.

pheromone isolated from the oriental hornet Vespa orientalis. As shown in Fig. 16.526, Baeyer-Villiger oxidation of racemic undecylcyclopentanone with A. calcoaceticus NCIMB 9871 led to a 25% isolated yield of (S)-5-hexadecanolideshowing an ee of 74%. Interestingly, a 30 % yield of remaining (R)-2-~ndecylcyclopentanone of 95 % optical purity can also be isolated using a longer incubation time, thus allowing direct access, via chemical Baeyer-Villiger oxidation, to the (R)-(+)-5-hexadecanolide known to be the sole bioactive enantiomer. The biotransformation of a-substituted cycloalkanones using the BVMOs from camphor grown Pseudomonas putida has also been investigated in depth. Whilst the NADPH dependent activity corresponding to 2-0x0-A3-4,5,5-trimethylcyclopentenylacetyl-Co-A monooxygenase (and termed M 0 2 ) resolved a series of a-alkyl cyclopentanones with good selectivity, poorer resolution of these compounds was per-

1228

I

IG Oxidation Reactions

R

M01 or M02 from camphor

'VR

cl

grown Pseudomonas putida ATCC 17453

M01 R

Yield ketone

e.e. ketone

Yield lactone

e.e. lactone

C4H9

14

9

16

58

&HI3

48

48

34

74

%HI7

35

22

11

90

e.e. ketone

Yield lactone

e.e. lactone

40

95

R

Yield ketone

C4H9

26

&HI3

51

75

35

92

%HI7

44

59

29

95

M02 from camphor grown Pseudomonas putida ATCC 17453

+

b

R

+

bR

M02

R

Yield ketone

e.e. ketone

Yield lactone

e.e. lactone-

%HI3

30

65

36

72

c8H17

49

61

34

77

CHZCQEt

43

89

30

93

13

75

34

83

CH~CH~OAC

Figure 16.5-27. Biotransformation o f 2-substituted monocyclic ketones by BVMOs from camphor grown Pseudomonas putida ATCC 17453.

formed by the NADH dependent M 0 1 complement[104](Fig. 16.5-27). An extension to this study revealed that M 0 2 could be used to resolve a series of a-substituted cyclohexanones wherein the subsituents consisted of esters, acetates and common protecting groups [''I. This led to the development of a chemoenzymatic synthesis of (R)-(+)-lipoicacid incorporating a BVMO catalyzed resolution as the key step (Fig. 16.5-28). Interestingly, the preferred selectivity of cyclopentanone monooxygenase from Pseudomonas sp. NCIMB 9872, is opposite to that of M 0 2 , and in a

I 6 5 Baeyer-Villiger oxidations

Q----

I

1229

0

M 0 2 from camphor grown Pseudomonas putida ATCC 17453

*

JJ

0

steps, including Mitsunobu inversion of chiral centre

II

M

e

o

S-S

w

Figure 16.5-28. Chemoenzymatic synthesis of (+)-lipoic acid incorporating a BVMO catalysed resolution as the key step.

separate investigation,it was suggested that this enzyme be used in the place of M 0 2 to eliminate the need for the Mitsonobu inversion in the chemoenzymatic synthesis [1121. The biological Baeyer-Villiger oxidation has also been applied, in a variety of forms, to the production of optically active lactones from prochiral 3-substituted cyclobutanones. A series of cyclobutanones was subjected to oxidation by Acinetobacter sp. and to the M 0 1 and M 0 2 enzyme preparations derived from camphorgrown Pseudornonas putida ATCC 17453['13]. The results are summarized in Fig. 16.5-29.In general, the reactions performed with Acinetobacter sp. displayed better enantioselectivities, but the value of a multi-biocatalyst approach was illustrated by the fact that certain BVMOs from P. putida displayed opposite enantioselectivity. A further series of cyclobutanone substrates was oxidized by Acinetobacter sp. and by the fungus Cunninghamella echin~lata[l'~I(Fig. 16.5-30). The lactonization of 3-(4'-chlorobenzyl)-cyclobutanone was performed by this fungus to yield (R)lactone of 99 % ee in 30 % yield, which was used in a chemoenzymatic synthesis of baclofen [lls1, a lipophilic derivative of y-aminobutyric acid. The Cunninghamella strain was also used to oxidize 3-(benzyloxymethyl)-cyclobutanoneto the optically pure (R)-(-)-y-butyrolactone, which was used in enantiodivergent chemoenzymatic syntheses of (R)-and (S)-proline["'I . The oxidation of either enantiomer of menthone and dihydrocarvone by Acinetobacter sp. were also reported['l71. (-)-Menthone is not metabolized but (+)-menthone leads to the expected lactone, whereas both enantiomers of dihydrocarvone are oxidized. Thus (-)-dihydrocarvone leads to the expected lactone, whereas (+)-dihydrocarvone afforded the unexpected 'abnormal' lactone product (Fig. 16.5-31).Both enantiomers of dihydrocarvone are also transformed by MMKMO ["I from Rhodococcus erythropolis DCL 14,which in contrast to Acinetobacter sp., also transforms both enantiomers of menthone. Taschner and coworkers described the oxidation of cis-3,5-dimethylcyclohexanone by whole-cell preparations of A. calcoaceticus NCIMB 9871 [118], which led directly to

1230

I

76 Oxidation Reactions

Baeyer-Villiger monoxygenase or whole cell catalyst R

R

R Bu Bu' CHpPh

Conversion 95 98 100 100

Yield lactone

100

89

(q-, 55

Conversion 100 78 58 48

Yield lactone nd nd 40 38

e.e. lactone (R-,69 (R)-,91 (q-, 15 (q-37

68

56 57 83

e.e. lactone (s)-,17% (R)-, 84% (6-, 82% (R)-, 95%

I Mnl R Bu Bul CHnPh

Figure 16.5-29.

Biotransformation o f prochiral 3-substituted cyclobutanones using BVMOs.

the corresponding optically active lactone and thence to the hydroxyacid, which was converted into the methylester by reaction with diazomethane. This methylester, which was shown to be optically active, is a key intermediate in the synthesis of the polyether antibiotic ionomycin. In addition, several bridged bicyclic compounds have been examined as potential substrates (Fig. 16.5-32). In contrast to the regiodivergent behaviour of the [n.2.0] bicyclic compounds, in these cases, only one lactone product is usually obtained. This high selectivity compares favorably with the chemical Baeyer-Villiger oxidation of compounds of this type, which often afford regiomixtures[119].In addition, the

7 13.5 Baeyer-Villiger Oxidations

I

1231

go

Organism

R

~

1, R = Ph 2, R = pFC,H, 3, R =p-CLC,H, 4, R =p-MeGH, 5, R = C $ c

do

'

R

o )

0

6, CH,C,H,-p-OMe 7, CH,OCH,Ph 8, CH,Ot-Bu

8

AcinetobacterTD63 C. echinulata A. calcoaceticus AcinetobacterTD63

90 25

43 15

(R)-,25

98 89 88

obtained bridgehead lactones are often described to be of high optical purity. The benzyloxy derivative is known to be an important intermediate for prostaglandin synthesis. The residual fluorinated bicyclic ketone of high enantiomeric excess was used to synthesize an antiviral carbocyclic nucleoside['201. In this last case, detailed studies showed that the first formed product is the corresponding alcohol (about 80% conversion) and that over the next 3 h period, the alcohol concentration decreased, the amount of ketone rose and the production of lactone This observation led to an elegant closed-loop recycling procedure, as shown in Fig. 16.533, where the alcohol dehydrogenase from Thermoanaerobium brockii was used in conjunction with the purified monooxygenase from A. calcoaceticus NCIMB 9871. In

1232

I

16 Oxidation Reactions

A. calcoaceticus or Acinetobacter TD 63

*

n rac-rnenthone

(-)-menthone

A. calcoaceticus or Acinetobacter TD 63

*

A racdi hydrocarvone Figure 16.5-31. Oxidation o f dihydrocarvone enantiomers with Acinetobacter calcoaceticus NCIMB 9871 and Acinetobacter sp. TD63.

this case, the substrate alcohol also serves as a co-substrate for the NADPH recycling reaction. Thus, endo-bicyclo[2.2.1]heptan-2-01was transformed using catalytic amounts of NADP. An analogous recycling loop was set up using the NAD dependent alcohol dehydrogenase from Pseudomonas sp. NCIMB 9872 and the NADH dependent M 0 1 isozyme complement from Pseudomonas putida ATCC 17453, for the oxidation of 7- endo-methylbicyclo[3.2.0]hept-2-en-6-ol[1221. A further series of prochiral bicyclic [2.2.1] substrates have also been studied by Taschner and coworkers and lead generally to lactones of high enantiomeric purity. One of these is a valuable precursor for chorismic acid synthesis [971. The transformation of a series of norbornanone derivatives (Fig. 16.5-34) was studied by Roberts and coworkers who determined that both the M 0 1 complement of NADH dependent BVMOs from Pseudomonas putida ATCC 17453 and the NADPH dependent fraction M 0 2 were successful in the resolution of hydroxy, acetoxy and benzyloxy norbornanones [1231. Interestingly 25DKCMO and 36DKCMO when separate, displayed notably different reactivity toward the hydroxy and acetoxy derivative, again emphasizing their complementary nature as potential individual biocatalysts. The benzyloxy lactone is an intermediate in the synthesis of the insect antifeedant azadirachtin. Further studies also been performed on the bicyclo[3.2.0]heptan-6-oneseries of compounds [124* 12’1. These results are summarised in Fig. 16.5-35.Oxidation of this ketone with Pseudomonas NCIMB 9872 gave the (lS, 5R)-lactoneoflow optical purity (23% ee) with only small amounts (5%) of the isomeric lactone, whereas its oxidation with an Acinetobacter sp. gave these lactones in a 9: 1 ratio and a modest yield, a result quite different from the one described previously. However, oxidation using either Pseudomonas sp. or Acineof 7-endo-methylbicyclo[3.2.0]hept-2-en-6-one tobacter sp. produced optically pure (ee > 96 %) of both lactones in equal quantities

1 6 5 Baeyer-Villiger Oxidations

I

1233

Pseudomonas sp.

NCIMB 9872

:

38

4

1

0

Cylindrocarpon destructans

0

0% e.e.

F Acinetobacter

&Lo+F $ Y o

*

NCIMB 9871

A d

A d

OAc 11%

*ao

BzO-Acinetobacter

h

NCIMB 9871

O H &:

‘0

26%, 95% e.e.. F

F

Acinetobacter

NCIMB 9871

*

0

Figure 16.5-32.

&Ao+ F$7° Br

36%, 95% ex.

Baeyer-Villiger oxidation of various [2.2.1] bicyclic substrates.

dehydrogenase

NADP+

NADPH + H+

monooxygenase

0

Figure 16.5-33. Closed-loop recycling procedure for NADPH recycling using the substrate alcohol as the reducing agent.

1234

I

1 G Oxidation Reactions

NADH-dependent BVMO

* *R

l,R=H

R&o

0 (1 S,5S, 6R)-

2, R = O H 3,R = OAC 4, R = OBn

Enzyme

Substrate

Conversion (%)

25DKCMO

1

20

Lactone e.e. (%)

60

36DKCMO

1

48

>90

25DKCMO

2

0

36DKCMO

2

33

>95

25DKCMO

3

35

>95

36DKCMO

3

0

‘M01’

4

39

>95

Figure 16.5-34. Biotransformation of norbornanone derivatives using NADH dependent BVMOs from camphor grown Pseudomonas putida ATCC 17453.

(combined yields 50-55 %). Surprisingly, 7,7-dimethylbicyclo[3.2.0]hept-2-en-6-one was oxidized by the Acinetobacter strain to give exclusively one lactone of 29% ee, a very low enantioselectivity. The bromohydrin obtained from this substrate led to similar results, yielding the same type of oxidation. This can be considered as being the “normal” lactone since substitution with two methyl groups makes this carboncarbon bond the more substituted one. Again, the M 0 1 isozymic complement from Pseudomonas putida was successful in generating the complementary enantiomers from endo-methyland dimethyl derivatives with good enantiomeric excess [*031. 16.5.4

Models for the Action of Baeyer-Villiger Monooxygenases

The results of biological Baeyer-Villiger oxidations have been, in some cases unpredictable and surprising, and, in the continued absence of a structure of one of these enzymes, several groups have attempted to explain the various observations of selectivity with an increasingly complex series of models. Initially, some workers proposed that enantiodivergent biotransformations of the type witnessed in the oxygenation of bicyclo[3.2.0]hept-2-en-6-one by, for instance CHMO and 25DKCMO could be due to the presence in either of these preparations of two separate enzymatic activities. Whilst this was once and indeed still is, a reasonable assumption in the light of results obtained with whole-cell preparations, the use of highly purified preparations of the two named enzymes to effect this biotransformati~n[’~~* 1‘’‘ have eliminated this possibility in these cases. The phenomenon of enantiodivergence has therefore been addressed with respect to one enzyme active site.

I

76.5 Baeyer-Villiger Oxidations 1235

Pseudomonas

*

0:::::Fo +

NCIMB 9872

75%, 23% e.e.

ofo

Acinetobacter

/

NCIMB 9871

5%

0::::yo +

96% e.e.

96% e.e. Acinetobacter

0

*

NCIMB 9871

+ 29% e.e.

Ho,q,,,qo Acinetobacter D

-“111

0

NCIMB 9871

Br 98% e.e. Figure 16.5-35.

Baeyer-Villiger oxidation of various [n.2.0] bicyclic compounds.

The first model was proposed by Furstoss and coworkers, based on steric and stereoelectronic considerations. In this model, shown in Fig. 16.5-36, the 4-ahydroxyperflavin is considered as being the oxygen transfer agent, according to the hypothesis of Walsh and coworkers[841. The enantioselectivity of the reaction would be due to a different positioning of each intermediate in the active site. It is supposed, primarily, that the attack of the hydroperoxyflavin should take place on the least hindered face of the ketone. On the other hand, the migrating C-C bond of the peroxidic intermediate should be antiperiplanar to the peroxidic bond and to a nonbonded electron pair of the hydroxide group, as suggested for chemical BaeyerVilliger oxidations. Thus, the cycloalkyl part of the (S,S)-enantiomer of the ketone (the one leading to the “normal”lactone) could be accommodated in only one region of the active site (position 1).Position 2 would never be adopted due to some steric hindrance with the active site (dotted cube). Similarly, in the case of the (R,R)enantiomer, position 4 would be favored over position 3 leading to the “abnormal” lactone. This model was augmented by further work by the inclusion of results obtained with both monocyclic monterpene 3-substituted cyclobutanone substrates [1131 and a-substituted cyclohexanones

1236

I 1 G Oxidation Reactions

Position 1

;

i

::

I

.

Position 2

Position 3

Position 4

Figure 16.5-36. Furstoss model for the active site o f cyclohexanone monooxygenase from Acinetobacter calcoaceticus NClMB 9871.

Taschner and coworkers proposed a similar model based on two other flavoenzymes; the human and E. coli glutathione reductase. The FAD binding domain of glutathione reductase and p-hydroxybenzoate hydroxylase have been shown to resemble each other closely via comparison of their respective X-ray crystal structures. Extrapolating this information to CHMO leads to the proposal that the hydroperoxide is attached to the re-face of the isoalloxazine ring and that the ketone substrates approach the hydroperoxide from the direction of the dimethylbenzene Further stereochemical and stereoelectronic considerations lead to a hypothesis explaining the observed stereoselectivities. In the model of Furstoss and coworkers, stereoselectivityof CHMO is determined by the differentiation of groups of different sizes in the active site. A different model, proposed by Kelly and coworker^[^^^-^^^^, extends Taschner’s idea that the source of stereoselectivity might be the flavin cofactor itself. It was suggested that the stereoselectivityof oxygen insertion arises solely as a result of the flavin face, re- or si-, from which the hydroperoxide attacks. This would lead to two distinct Criegee intermediates of opposing absolute configuration (Fig. 16.5-37). Hence it was

16.5 Baeyer-Villiger Oxidations

S or si-

Ror re-

Non-migratinggroup Migrating group

CHMO

I

1237

Non-migratinggroup RI-o\O~!*A R21 0 Migrating group

25DKCMO or 36DKCMO

Figure 16.5-37. Schematic representation o f enantiomeric Criegee intermediates for the enzymatic Baeyer-Villiger reaction.

CHMO, NADPH Sor si

Ror re 25DKCMO or 36DKCMO NADH Figure 16.5-38. Enantioselective Baeyer-Villiger oxidation of a tricyclic ketone by Type 1 and Type 2 BVMOs.

demonstrated that for the tricyclic ketone shown in Fig. 16.5-38 for which attack from only the exo-face is possible, pure preparations of BVMOs always resulted in lactones of >95 % ee Interestingly, all Type 1, FAD plus NADPH dependent BVMOs yield lactone from the (R)-configurationof the intermediate, and all Type 2, NADH plus FMN dependent BVMOs yield lactone from the (R)-intermediate. Substrate interaction with the topology of the active site must also be considered however, as the enantiocomplementary DKCMOs, both proposed to catalyze oxygen insertion via (R)-Criegee intermediates, catalyze complementary resolutions of racemic camphor [671. This additional dependence on active site topology for selectivity in CHMO was carefully considered by Ottolina et al. who developed a sophisticated cubic space model for the active site of CHMO (Fig. 16.5-39).This group was able to show that, for example, for the biotransformation of 7-endo-methylbicydo[3.2.0]hept-2-en6-one, of the eight possible intermediates in oxidation, the only two “allowed by the model were the two which led to the lactones observed by experiment. The model was successfully applied to a series of other ketones and also predicts the stereoselectivity of sulfur oxidation by this The group of Colonna established in a series of reports that CHMO was able to catalyze the oxidation of a range of alkylaryl sulfides, benzyl alkyl sulfides, functionalized sulfides and 1,3-dithioacetals with absolute configuration and enantiomeric excesses being highly dependent

1238

I

1 G Oxidation Reactions

Side

\

Front

Top

nP

I Lj

Side HS

Figure 16.5-39. Cubic space filling model o f the active site o f cyclohexanone monooxygenase from Acinetobacter calmaceticus NClMB 9871, based on the results o f the oxidations o f a series o f bicyclic ketones. The catalytic oxygen is circled. The main (M) hydrophobic large (HL)and hydrophobic small (Hs) pockets are depicted. The correct arrangements o f the Criegee intermediate are also shown.

on the structure of the substrate['3]. This group has also recently reported the first asymmetric oxidation of tertiary amines using CHM0['331. The ability of BVMOs to oxidize sulfur was also exploited by Beecher and Willetts in order to construct space filling cubic models of the active site of the DKCMO enzymes from Pseudomonas putida ATCC 17453 (Fig. 16.5-40). They note that the more relaxed enantiospecificity of 3GDKCM0, at least in terms of sulfoxidation, appears to be due to an overall larger 3D cubic space available in the active 3GDKCMO appears to be the best candidate for a first X-ray structure of a BVMO, as preliminary crystal data have been 16.5.5 Conclusion and Outlook

It is apparent from the many application of BVMOs in synthesis, that these enzymes currently represent the most valuable method of effecting the enantioselective

I G.5 Eaeyer-Villiger Oxidations

I

1239

Figure 16.5-40. Cubic space filling models o f active sites o f : right, 3,6-diketocamphane 1,6-monooxygenase; and left, 2,S-di ketocamphane, 1,2-monooxygenase based on results o f sulfoxidations o f a series of sulfide substrates.

‘engineered’ Saccharomyces cerivisiae expressing CHMO

+

>

+ R

R

R

b

1

R

Ratio 1:2

2

e.e. lactone 1 (%)

e.e. lactone 2 (%)

9

36

80:20

33

19

83:17

33

60

Combined yield lactones Me

13:87 95%

Et

80%

n-Pr

44%

n-Bu

99:1

38

99:1

16

34% moct

19%

Figure 16.5-41. Biotransformation o f 3-alkylcyclopentanones by “engineered” Saccharomyces cerivisiae expressing CH MO.

Baeyer-Villiger reaction. The primary sources of BVMO enzymes carry associated disadvantages that must now be addressed, although recent biotechnological advances suggest that BVMOs will be more accessible to the synthetic organic chemist in the future.

CHMO Rhodococcus coprophilus

CHMO Acinefobacter NCIMB 9871

CPMO Pseudomonas NClMB 9872

Steroid monooxygenase Rhodococcusrhodochrous

2

3

4

5

3,6-DKCMO Pseudomonas putida

7

M-S-Q-L-M-D-F-D-A-I-V-I-G-G-G-F-G-G-L-Y-A-V-K-K-

A-Q-T-I-H-G-V-D-A-V-V-I-G-A-G-F-G-G-I-Y-A-V-H-K-

A-E-W-A-E-E-F-D-V-L-V-V-G-A-G-A-G-G-

1

1

1

A-M-E-T-G-L-I-F-H-P-Y-M-Y-P-G-K-S-A-A-Q-

-M-Q-A-G-F-F-G-T-P-Y-D-L-P-T-R-T-A-R-Q-M-

M-N-G-Q-H-P-R-V-V-V-A-A-P-D-A

1 4 -N-S-V-N-D-K-L-D-V-L-L-I-G-A-G-F-

1

2

2

Figure 16.5-42. N-terminal amino acid sequence alignment ofType 1 BVMOs (1-5) and Type 2 BVMOs (6 and 7). Conserved residues are marked in bold.

2,5-DKCMO Pseudomonas putida

6

Type 2 BVMOs

Steroid monooxygenase Cylindrocarpon radicicola

1

Type 1 BVMOs

2

8'

2

P

% 5.

B

m

-

References I1241

this problem has been the cloning and expression of the gene encoding CHMO in Saccharomyces ceri~isiae[*~1. In a series of reports by Stewart and coworkers[135-1371, the “designer yeast” was shown to catalyze many of the reactions which had previously been shown to be catalyzed by either whole cells of Acinetobacter sp. or CHMO in addition to some new ones (Fig. 16.5-41). Recently, a similar strategy has seen whole-cell preparations of Escherichia coli expressing recombinant CHMO for the same purpose[138].It remains to be seen whether constraints on the use of genetically engineered microorganisms of this type will render these strains as “difficult”to manipulate as the wild-type strains. The use of purified enzyme would circumvent the need for whole-cell containment procedures, and indeed, amounts of CHMO are now available from Fl~ka[~~’]]. However, the attendant costs associated with cofactor recycling must be addressed if this approach is to prove viable. The recent production of a formate dehydrogenase suitable for use in NADP/NADPH recycling s y ~ t e m s I ~ should ~ ~ 1 prove attractive in this regard, as should the further investigation of NADH dependent enzymes. The practicalities associated with the industrial scale up of biological Baeyer-Villiger reactions are currently being investigated [14’1. New sources of enzyme will also become important and with the advent of genomic science, paralogs of genes that encode CHMO-like proteins are being identified amongst whole bacterial genomes, most recently those of Pseudomonas aerugin~sa[’~~] and Mycobacterium tubercul~sis~’~~]. The availability of gene and amino acid sequence data for BVMOs will prove useful in identifylng more new activities in this manner. BVMOs of the same Type (1 or 2) exhibit sequence homology within their N-terminal amino acid sequences although homology between types is not (Fig. 16.5-42). In the hture, the “tailoring”of enzyme characteristics by either rational redesign or so-called “directed” evolution approaches could also doubtless be applied to BVMOs. Fundamental to these studies would be the development of an efficient, rapid screen for BVMO activity. Rational redesign would require more knowledge of the 3D structure of these enzymes. This is one reason why the acquisition of a complete X-ray crystal structure of a BVMO must be considered of fundamental importance to the ongoing development of this area.

References A. Gusso, C. Baccin, F. Pinna, G. Struhl, Organomtallics 1994, 13, 342-3451, 2 C. Paneghetti, R. Gavagnin, F. Pinna, G. 1

Strukul, Organornetallics 1999, 18, 5057-5065. 3 C. Bolm, G. Schlingoff, F. Bienewald,I. Mol. Catal. A 1997, 117, 347-350. 4 A. J.Willetts, Trends. Biotechnol. 1997, 15, 55-62. 5 S. M. Roberts, P. H. W. Wan, J . Mol. Catal. B. Enz. 1998,4,111-136.

R. Kelly, P. Wan, J. Tang in: Biotransforrnations, Vol. 8a in the series Biotechnology. (volume Ed. D. R. Kelly; series Ed. H.-J. Rehm, G. Reed, A. Puhler, P. J. W. Stadler), Wiley VCH, Weinheirn, 1998, pp. 535-588. 7 D. R. Kelly, Chern. Oggi. 2000,18, 33-39 and 6 D.

52-56. 8

E. Vischer, A. Wettstein, Experientia 1953, 9, 371-372.

9

J. Fried, R. W. Thoma, A. Klingsberg,J . Am. Chem. SOC.1953,75, 5764-5765.

1242

I

76 Oxidation Reactions

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I

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S. M. Roberts, H. Sandey, N. F. Shipston, J .

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Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

7 G. G Oxidation of Acids 140

141

PEDANT http://pedant.mips.biochem. rnpg.de/ 143 C. A. Rivera-Mamero,M. A. Burroughs, R. A. Masse, F. 0. Vannberg, D. L. Leimbach, J. Roman, J. J. Murtagh, Microb. Pathogenesis 1998, 25,307-316.

142

K. Seelbach, B. Riebel, W. Hummel, M. R. Kda, V. I. Tishkov, A . M . Egorov, C. Wandrey, U. Kragl, Tetrahedron. Lett. 1996,9,1377-1380. M. C.Hogan, J. M. Woodley, Chem. Eng. Sci. 2000,55,2001-2008.

16.6

Oxidation of Acids

Andreas Schmid, Frank Hollmann, Bruno Buhler 16.6.1

Introduction

At a first glance, synthetically relevant oxidations of carboxylic acids, except for oxidations at positions other than the carboxylate group, can hardly be found in literature. However, some preparative applications in whole cell catalysis were reported and will be discussed in the following (Fig. 16.6-1A,B,C). In vitro, the high thermodynamic driving force for the oxidation of fonnate and pyruvate [l?(formatel COZ)= - 0.42 V"]; l? (pyruvate/(acetate, COZ)) = - 0.70 V[']] are used for the regeneration ofcoenzymes such as NAD(P)H or, indirectly, ATP (Fig. 16.6-1 D,E). NAD(P)H regeneration

t

D

a C O O H

-C

6""

Hoot,,

OH

- R-COOH

-

-

E

ATP regeneration

C

COOH

\

COOH

Figure 16.6-1. Synthetic and preparative applications of oxidations of acids. A, 6:Oxidations of benzoic acid initiated by dihydroxylation (Sects. 16.6.4.2 and 16.6.4.3); C: oxidative decarboxylation (Sect. 16.6.4.1); D,E : energy coupling for the regeneration of coenzymes (Sects. 16.6.2, 16.6.3).

I

1245

1246

I

76 Oxidation Reactions

H A

0 2

Figure 16.6-2. (PYOX).

Oxidative phosphorylation o f pyruvate by pyruvate oxidase

16.6.2 Pyruvate Oxidase (PYOx, E. C. 1.2.3.3)

PYOx from Lactobacillus plantarum L3, 4l or Streptococcus s a n g u i ~ [catalyzes ~] the decarboxylative phosphorylation of pyruvate to acetylphosphate, or the homologous arsenylation (Fig. 16.6-2). Acetylphosphate is an important substrate for the enzyme acetate kinase (E.C. 2.7.2.1), which catalyzes the phosphorylation of various nucleotide diphosphates such as ADP, GDP, TDP, IDP, or UDP to the activated triphosphates["'I. This reaction can be applied to regenerate ATP in ATP-dependent enzymatic in vitro reactions (Fig. 16.6-3). In a recent example, PYOx-catalyzed regeneration of ATP was coupled to in vitro protein biosynthesis (e.g. for human lymphotoxin)['I. Under aerobic conditions, no external regeneration system for PYOx has to be applied; catalase however has to be added in order to destroy harmful hydrogen peroxide. An alternative to this autoregeneration approach (Fig. 16.6-3A) was reported by Steckhan and coworkers for cases where hydrogen peroxide formation has to be prevented (Fig. 16.6-3B) [lo]. 0

n \ L

~ow.2-

.AtOOH +

PYOX

PYOX rerl

0,

?

W a Fa. GF

* D n T t o H

\

Br/AJ w -R

FB

0

A

acetate krnasa

H A

Eatarass

further enzymetic reactions

e-

Figure 16.6-3. Decarboxylative phosphorylation o f pyruvate by pyruvate oxidase as driving force for the regeneration of ATP; A: aerobic regeneration; B: indirect electrochemical regeneration.

NAD'

f"T

HCOO

1 G. G Oxidation of Acids

I

1247

Figure 16.6-4.

"I-I

Regeneration of NADH using the formate

dehydrogenase (FDH) reaction.

CO,

Here, the anode, together with the mediation by ferrocene, removes excess electrons from the PYOx active site. Another possible application of the PYOx-catalyzed production of acetylphosphate lies within the in uitro regeneration of acetyl-CoA [ll]. 16.6.3

Formate Dehydrogenase (FDH, E. C. 1.2.1.2)

Probably the most prominent oxidation of a carboxylic acid is catalyzed by the enzyme formate dehydrogenase (FDH, E. C. 1.2.1.2). FDH was isolated from various bacteria, yeasts, and plants, where its physiological role is the regeneration of NADH I1*l. FDH catalyzes the oxidation of formate to carbon dioxide, concomitant with the reduction of NAD' to NADH (Fig. 16.6-4). Because ofthe favorable thermodynamic equilibrium of the reaction and the volatility of the reaction product, the enzyme is commonly applied for in situ regeneration of NADH during asymmetric synthesis of chiral compounds [I3]. FDH from Cundida boidinii is mostly used as regeneration enzyme. It found industrial application at Degussa-Huls AG in a leucine dehydrogenase-catalyzed reductive amination of 2-keto acids yielding various amino acids (e.g. tert-leuNative FDH is very selective for NAD'. Recently a new FDH was developed by site-directed mutagenesis that shows all advantages of the NAD+dependent enzymes and additionally accepts NADP' as substrate [I7]. The activity of the mutant with NADP' is about GO% of the wild-type FDH with NAD+['8]. 16.6.4 Oxidations with Intact Microbial Cells 16.6.4.1

Production of Benzaldehydefrom Benzoyl Formate or Mandelic Acid

Benzaldehyde can be produced from benzoyl formate with whole cells of Pseudomonas putidu ATCC 12633 as biocataly~tI'~~ 201 (Fig. 16.6-5). Alternatively, but less effectively, mandelic acid can be used as starting material. A pH of 5.4 was found to be optimal for benzaldehyde accumulation. At this proton concentration, partial inactivation of the benzaldehyde dehydrogenase isoenzymes and activation of the benzoyl formate decarboxylase are reported. Fed-batch cultivation prevented substrate inhibition. In situ product removal is necessary to prevent product inhibition.

1248

I

16 Oxidation Reactions mandelate racemase COOH

mandelate dehydrogenase COOH

benzoyl formate decarboxylase

COOH

benzaldehyde dehydrogenase isoenzymes

__c

Figure 16.5-5. Degradation of tridecan-2-one with a crude cell-free preparation from a Pseudomonas aeruginosa strain.

Activated charcoal served as a solid-phase adsorption device [201. Thus, benzaldehyde and thiophene-2-carboxaldehydewere obtained from benzoyl formic acid and thiophene-2-glyoxylicacid respectively, in final concentrations of up to 4.8 g L-’ and molar yields exceeding 85 %. 16.6.4.2

Microbial Production of cis,cis-Muconic Acid from Benzoic Acid

Significant effort was put into the oxidation of benzoic acid to cis,cis-muconic acid via a multi-step reaction catalyzed by whole microbial cells [21-241. Cis,cis-muconic acid is used as raw material for the synthesis of resins and polymers (precursor of adipic acid). Furthermore, it is widely used as building block in the synthesis of pharmaceuticals and agrochemicals. As biocatalyst, growing cells of a mutant Arthrobacter strain (lacking cis&muconate derivatization activity) was used. The reaction cascade (Fig. 16.6-6) is initiated by a dioxygenation of the benzylic ring followed by decarboxylationyielding catechol, which is transformed to the product via dioxygenase-catalyzedring cleavage. benzoate 1,2-oxidoreductase

0

dehydrogenase

&HoH OH

NADH

NAD+

~

NAD+

p” aoH NADH

OH _. .

+

02

catechol 1,2dioxygenase

C

1COOH \

COOH

Figure 16.6-6. Sequential oxidation of benzoate to (cis,cis)-muconic acid catalyzed by Arthrobocter sp.

References I1249 Figure 16.6-7. Dioxygenation of benzoate to corresponding cis-l,2-diols.

COOH

O,+NADH

NAD+

Benzoic acid was fed continuously to the fermentation medium. The space-time yield of the process including downstream processing amounts to 70 g L-’ d-l. 16.6.4.3

Biotransformationof Substituted Benzoatesto the Corresponding cis-Diols

Enantiopure 1,2-cis-dihydroxycyclohexa-3 ,S-diene carboxylic acids have considerable synthetic potential as building blocks in chiral synthesis. Such cis-diols can be produced from benzoic acid derivatives by the action of toluate-1,2-dioxygenaseof Pseudomonasputida mt-2F2’]or homologous enzymes of a different origin (Fig. 1G.G7). Growing cells or recombinant Pseudomonas oleovorans GPol2 containing toluate1,2-dioxygenaseefficiently transform a whole range of meta- and para-substituted benzoates to the corresponding cis-diols, which are not further degraded by the Pseudomonas host. In the ortho position only hydrogen and fluorine were accepted as substituents. Toluate-l,2-dioxygenaseactivity is induced by ortho-toluate or the substrates themselves. Similar reactions were reported for the broad-substrate-specificbenzoate dioxygeRecombinant E. coli containing this enzyme nase of Rhodococcussp. strain 19070[2Gl. transform benzoate and anthranilate to catechol and 2-hydro-1&dihydroxybenzoate, respectively.

References 1 2 3

4

5 6

7

8

D. D. Woods, Biochem. J. 1936,30,515. K. Burton, Ergeb. Physiol. 1957,49, 275. B. Sedewitz, K. H. Schleifer, F. Gotz,J. Bacteriol. 1984, 160, 273-278. B. Sedewitz, K. H. Schleifer, F. Gotz,]. Bact e n d . 1984, 160, 462-465. J. Carlsson, U. Kujala, FEMS Microbiol. Lett. 1985,25,53-56. H. Vigenschow, H.-M. Schwarm, K. Knobloch, Bid. Chem. 1986,367,951-956. K. Suzuki, H. Nakajima, K. Imahori, Methods Enzmol. 1982,90,179-185. J. S. Nishimura, M. J. Griffith, Methods Enzmol. 1981,71,311-316.

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D.-M. Kim, J. R. Swartz, Biotech. Bioeng.

1999,66,180-188. 10 E. Steckhan. Kontinuierliche enzymatische

Synthesen enantiomerenreiner organischer Zwischenprodukte durch elektrochemische Aktiviemng von Redoxenzymen in elektrochemischen Enzymmembranreaktoren Final report for the period 01.03.1998 to 31.08.2000 on the Research Project 11 556 N/1; AiF: Bonn, 2000. 11 U. M. Billhardt, P. Stein, G. M. Whitesides, Bioorg. Chem. 1989, 17, 1-12. 12 V. 0. Popov, V. S . Lamzin, Biochem. J . 1994, 301,625-643.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

1250

I

IG Oxidation Reactions

M.-R. Kula, U. Kragl, Dehydrogenases in synthesis of chiral compounds. In Stereoselective Biocatalysis. R. N. Patel, (ed) Marcel Dekker, New York, 1999, pp. 839-866. 14 A. Liese, K. Seelbach, C. Wandrey. In Industrial Biotransfomations,WileyVCH,Weinheim, 2000, pp. 125-128. 15 A. S. Bommarius, M. Schwarm, K. Drauz,J. Mol. Cat. B: Enzymatic 1998, 5, 1-11. 16 U. Kragl, D. Vasic-Racki,C. Wandrey, Bioproc. Eng. 1996,14,291-297. 17 K. Seelbach, B. Riebel, W. Hummel, M.-R. Kula, V. I. Tishkov, A.M. Egorov, C. Wandrey, U. Kragl, Tetrahedron Lett. 1996, 37, 1377-1 380. 18 V. I. Tishkov, A. G. Galkin, G. N. Marchenko, Y. D. Tsyganov, H. M. Egorov, Biotech. Appl. Biochem. 1993, 18, 201-207. 19 J. Simmonds, G. K. Robinson, Enz. Microb. Tech. 1997,21, 367-374. 20 J. Simmonds, G. K. Robinson, Appl. Microbiol. Biotech. 1998, 50, 353-358. 13

Mizuno, N. Yoshikawa, M. Seki, T. Mikawa, Y. Iamada, Appl. Microbiol. Biotech. 1988, 28, 20-25. 22 N. Yoshikawa, S. Mizuno, K. Ohta, M. Suzuki,]. Biotech. 1990, 14, 203-210. 23 N. Yoshikawa, 0. Ohta, S. Mizuno, H. Ohkishi, Production of cis,cis-muconic acid from benzoic acid. In Industrial Application oflmmobilized Biocatalysts. A. Tanaka, T. Tosa, T. Kobayashi (eds), Marcel Dekker, New York, 1993, pp. 131-147. 24 A. Liese, K. Seelbach, C. Wandrey, Oxygenase of Arthrobactersp. In Industrial Biotransfmations. A. Liese, K. Seelbach, C. Wandrey (eds), Wiley-VCH,Weinheim, 2000, pp. 137-138. 25 M. G. Wubbolts, K. N. Timmis, Appl. Environ. Microbiol. 1990, 56, 569-571. 26 S. Haddad, D. M. Eby, E. L. Neidle, Appl. Environ. Microbiol. 2001, 67, 2507-2514. 21 S.

16.7

Oxidation of C-N Bonds

Andreas Schmid, Frank Hollmann, and Bruno Buhler 16.7.1

Introduction

Enzymatic oxidations of carbon-nitrogen bonds are as diverse as the substances containing this structural element. Mainly amine and amino acid oxidases are reported for the oxidation of C-N bonds. The steroespecificity of amine-oxidizing enzymes can be exploited to perform resolutions and even deracemizations or stereoinversions (Fig. 16.7-1 A). Analogous to the oxidation of alcohols, primary amines are oxidized to the corresponding imines, which can hydrolyze and react with unreacted amines (Fig. 16.7-1 B). In contrast to ethers, internal C-N bonds are readily oxidized, yielding substituted imines. This can be exploited for the production of substituted pyridines (Fig. 16.7-1 C). Furthermore, pyridines can be oxidized not only to N-oxides but also to a-hydroxylatedproducts (Fig. 16.7-1 D).

A

16.7.2.1 and 16.7.3.1); B: preparation of aldehydes (and subsequent formation o f imines) by oxidation of primary

R

R

16.7.3.2); D: hydroxyla-

16.7.2

Oxidations Catalyzed by Dehydrogenases 16.7.2.1

L-Alanine Dehydrogenase (L-Ala-DH, E.C. 1.4.1.1)

L-Alanine dehydrogenase (1-Ala-DH,E. C. 1.4.1.1) catalyzes the specific deaminative oxidation of L-alanine and thus can potentially be exploited for the resolution of racernic alanine (e.g. derived from the Strecker-synthesis).However, the oxidation of secondary alcohols and amines is thermodynamically unfavorable ['I, so that the equilibrium of the reversible dehydrogenase reaction is on the substrate side. Therefore, an additional thermodynamic driving force has to be introduced into the system in order to drive the desired reaction towards completion. Moiroux and coworkers recently introduced such a system (Fig. 16.7-2)[-'I. The general philosophy of their approach is the utilization of electrical power to remove the dehydrogenase products NADH and pyruvate (which is in situ transformed into the corresponding irnine), thus driving the equilibrium reaction towards completion. The electrochemical oxidation and reduction reactions produce NAD' and racemic alanine, respectively, as substrates for the dehydrogenase reaction. Using this procedure, not only a racemate resolution (with maximum 50 % yield) but a deracemization (100% yield) is achieved. The overall rate-limiting step is the slow, non-enzymatic formation of the imine. Consequently, the process is very slow (at best, the complete conversion of a 10 mM solution of r-alanine required 140 h).

1252

I

1G Oxidation Reactions

TI

L-Ala-DH

Stereoinversion o f L-alanine t o o-alanine catalyzed by L-alanine dehydrogenase (L-Ala-DH) in an electrochemical reactor.

Figure 16.7-2.

16.7.2.2

Nicotinic Acid Dehydrogenase (Hydroxylase) (E.C. 1.5.1.13)

The membrane-bound molybdoenzyme[G1 nicotinic acid dehydrogenase catalyzes the first step in the microbial degradation of nicotinic acid by inserting a hydroxyl function a to the nitrogen atom (Fig. 16.7-3).A possible mechanism for this reaction is given in Fig. 16.7-417]. The inserted hydroxyl function originates from water, which was confirmed by H2180 experiments [,' 1' . While nicotinic acid dehydrogenase does not accept NAD' as electron acceptor, artificial mediators such as benzyl viologene and 2,3,5-triphenyltetrazolium dyes can replace NADP+[']. Various bacterial strains have been reported to convert a broad range of nicotinic acid derivatives (Table 16.7-1)[lo,12]. An industrial process (according to the first entry in Table 16.7-1) was set up by

QCOOH Figure 16.7-3.

*

HOQCOOH

c ~

Microbial mineralization o f nicotinic acid.

Figure 16.7-4. Proposed mechanism for enzymatic hydroxylation o f nicotinic acid (A = acceptor). The reaction scheme is based on the so-called arine mechanism.

c

citric acid cycle

16.7 Oxidation ofC-N Bonds I1253 Table 16.7-1.

Microbial a-hydroxylationof substituted pyridines.

Reactions catalyzed by whole cells

Final product concentration [g L-'1

Enzymes and reference

74

Dehydrogenase [11

191

Dehydrogenase ['I

301

Dehydrogenase 131

6.4

Dehydrogenase 14]

98

Dehydrogenase ['I

N R"

Dehydrogenase and decarboxylaseIG]

NRa

Dehydrogenase [71

45

DehydrogenaseI['

40

Nitrilase and Dehydrogenase [91

55

Nitrilase and Dehydrogenase1'

40

Nitrilase and Dehydrogenase [lo]

8

Dehydrogenase [''I

Akdgenesop UK21

coon

Z O O C O O H

Rhmbiurn Sp LA17 COOH

oCN

coon

a N R not reported.

1 H. Kulla, Chimia 1991,45,81-85. 2 T. Nagasawa, B. Hurh, T. Yamane, Biosci. Biotech. Biochem. 1994,58,665-668. 3 B. H u h , M.Ohshima, T. Yamane, T. Nasagawa,]. Fern. Bioeng. 1994,77,382-385. 4 M. Ueda, R. Sashida,]. Mol. Cat. B: Enzymatic 1998,4,199-204. 5 A. Kiener, R. Glockler, K. Heinzmann, /. Chem. Soc. Perkin Trans. I1993,1201-1202. 6 T. Yoshida, A. Uchida, T. Nagasawa. "Regiospecific ..."; Annu. Meet. SOC. Biosci. Bioeng., 1998, Japan.

7 T. Yoshida, T. Nagasawa, Biosci. Biotech. Biochem. 2000,89,111-118. 8 M. Yasuda, T. Sakamoto, R. Sashida, M. Ueda, Y.Morimoto, Biosci. Biotech. Biochem. 1995,59, 572. 9 A. Kiener. USP5266469 (1993). 10 M. Wieser, K. Heinzmann, A. Kiener, Appl. Microbid. Biotechnol. 1997,48,174. 11 A. Kiener, Y. van Gameren, M. Bokel.; USP 5,284,767, 1994.

1254

I

IG

Oxidation Reactions

-

QCOOH

___)

HO Figure 16.7-5.

xNoz

H./ i"-y-(/ CI

N

6-Hydroxynicotinic acid as synthon for the pesticide

Imidachloprid.

Lonza AG, Switzerland. 6-Hydroxynicotinicacid is precipitated from the fermentation broth as magnesium salt in the so-called pseudocrystal process, thus enabling not only easy downstream processing but also continuous fermentation [I3]. 6-Hydroxynicotinic acid is the key building block in the synthesis of Imidachloprid (Fig. 16.7-S), an effective pesticide against hemipterans and other sucking insects [10, 111 16.7.3

Oxidations Catalyzed by Oxidases 16.7.3.1

Amino Acid Oxidases

Among the enzymes catalyzing oxidations of carbon nitrogen bonds, the amino acid oxidases (AAO, E. C. 1.4.3.x) are the most interesting for synthetic applications. Compared to some specific amino acid oxidases such as aspartate oxidase or glutamate oxidase, the two D- and L-amino acid oxidases (E. C. 1.4.3.2 for L-AAOand E.C. 1.4.3.3 for D-AAO) are advantageous on account of their broad substrate NH2 R ~ C O O H

L-AA

\

NH

HO ,,

RA k O H

0

+ NH, +

R ACOOH

L-AAO

\

NH R AO : OH

D-AA Resolution of racemic amino acids (AA) catalyzed by and (L)-specific amino acid oxidases (AAO).

Figure 16.7-6. (D)-

7 G. 7 Oxidation

HowNHz

of C-N Bonds

HO, P

O

O

H

+

NWNH

YH,


I 0,

HO ,,

1

+ NH,

bleomycine Figure 16.7-7. Resolution of D,L-erythro-P-hydroxyhistidine as the enantiospecific step i n bleomycine synthesis.

t

R = CH,CH,SCH, methionine CH,CH(CH,), leucine

L-amino acid

FDH

Enzymatic deracemization of amino acids catalyzed by D-amino acid oxidase (D-AAO). Leucine dehydrogenase (LeuDH) transforms the oxidation product o f the undesired amino acid enantiomer in situ into the racemic amino acid. Regeneration o f N A D H is performed by formate dehydrogenase (FH D). Figure 16.7-8.

spectrum and their strict stereo~pecificity~'~. 'I. Therefore, AAOs are most commonly used for the resolution or deracemization of racemic amino acid mixtures (Fig. 16.7-6). The approach outlined in Fig. 16.7-6 was used for example to remove traces of Dmethionine from 99% pure t-methionine[16,"1 or to transform racemic phenylalanine quantitatively into D-phenylalanineand phenylpyruvic acid ["I. Coimmobilization with catalase on a solid matrix (EupergitB) resulted in largely increased DAAO stability. In an enzyme-membrane-reactor,space-time-yieldsas high as 90 g L-l d-' were reached. In another example, a racemic mixture of D,L-erythro-P-hydroxy[191. The histidine was converted into the ketoacid and L-erythro-j3-hydroxyhistidine

1256

I

IG Oxidation Reactions

L-pipecolic acid

One-pot chemo-enzymatic deracemisation of pip pipe colic acid catalyzed by D-amino acid oxidase (D-AAO).Utilization of catalase was not reported.

Figure 16.7-9.

latter compound is a key intermediate in the synthesis of the anti-tumor agent bleomycine (Fig. 16.7-7) Simple racemate resolutions have a maximal yield of 50% for the desired compound. Furthermore, additional (potentially laborious) separation steps are necessary. As a consequence, alternativeprocesses that involve the stereoinversion of the undesired enantiomer are gaining increasing interest[*']. One approach for these so-called deracemization processes is to reconvert the oxidation product either enzymatically (Fig. 16.7-8)or chemically (Fig. 16.7-9)to the racemic substrate. The enzymatic variant of this concept was reported for the deracemization of D,Lmethionine or D,L-leucine (Fig. 16.7-8)[I7]. Soda and coworkers developed a chemoenzymatic racemization procedure utilizing boron hydrides for non-enantioselective reduction of the undesired D-AAO product (Fig. 16.7-9)[213 221. Using the same procedure, the authors achieved conversion of D-proline into L-proline["I. Furthermore, D,L-lactate and 2-hydroxy butyric acid were deracemized by utilizing Llactate oxidase[231. The D-AAOcatalyzed oxidative deamination of cephalosporin C found industrial application (Hoechst Marion Roussel, Germany) as the first step in the so-called 7-aminocephalosporanicacid (7-ACA)process (Fig. 16.7-10)[24-261. Using this process, this application of heavy metals and chlorinated hydrocarbons can be avoided, and the volumes of waste-gas as well as of mother liquors are drastically reduced L2'1. 16.7.3.2

Amine Oxidases

16.7.3.2.1 Monoamine Oxidase (MAO, E.C. 1.4.3.4)

The flavoenzymes monoamine oxidase A and B (MAO-A, MAO-B)[271 catalyze the oxidative deamination of various primary and secondary amines and the oxidation of tertiary amines. Their physiological role, as the various synonyms such as epineph-

16.7 Oxidation of C-N Bonds

Figure 16.7-10.

Enzymatic reaction sequence for the production o f 7-ACA from cephalosporin C.

I

1257

YgP&oy

K' 0

cephalosporin C

D-MO

COOH

0

H202

NH3

""""miY N&ou 0

K:

COOH

spontaneous

0

H O O C - - T i ~ &0 oy

COOH

0

glutaryl amidase E.C. 3.1.1.41 K~~~CvCOOH

b $ L o ,

0

COOH

0

7-aminocephalosporanicacid (7-ACA)

(i'

INH2 /

serotonine

phenetylamine

milacemide

, . +O H

HO noradrenaline

HO

adrenaline

Figure 16.7-11. Various neurotransmitters as substrates for mitochondria1 monoamine oxidase (MAO).

1258

I

7G Oxidation Reactions U

MA0 (HAT or SET)

R I

FAD

FADH

FADH '

HP,

0,

FADH,

Proposed mechanisms for the oxidation of primary and secondary amines by monoamine oxidase (MAO). Figure 16.7-12.

MA0

I

I

I

Oxidation of l-methyl-4-aryl(heteroaryl)-l,2,3,6-tetrahydropyridines catalyzed by rnonoamine oxidase (MAO). Figure 16.7-13.

rine oxidase, serotonin oxidase, tyramine oxidase, or adrenaline oxidase suggest, is the transformation of neurotransmitters via oxidative deamination (Fig. 16.7-11) as well as the detoxification of xenobiotics[28-301. The mechanism of M A 0 is still a topic of hydrogen atom transfer (HAT)[321 or single electron transfer (SET)[331 are discussed as initial oxidation steps in the overall mechanism (Fig. 16.7-12). Various substrates have been specified for M A 0 with respect to synthetical, mechanistical and biochemical purposes. Castagnoli and coworkers elucidated structural requirements of MAO-B with various substituted l-methyLl,2,3,G-tetra-

MA0 HO

HO

dopamine

HO spontaneous Picet-Spengler condensation HO HO

norlaudanosine

Preparation of norlaudanosine initiated by the oxidation of dopamine by monoamine oxidase (MAO) (A). The oxidation product reacts spontaneously in a Picet-Spengler condensation with unreacted dopamine (B).

Figure 16.7-14.

167 Oxidation ofC-N Bonds Table 16.7-2. v/v water) ["I.

Oxidation o f various amines catalyzed by monoamine oxidase in n-octane (0.5 %

Substrate

0""'

Product

0"""' P"

6

Q

Yield I"/.] 99

99

69

14

12 7. C. G. Woo, X. Wang, R. B. Silverman,J. Org. Chem. 1995,60,6235-6236.

hydropyridines to produce dihydropyridines that are further oxidized to pyridinium structures (Fig. 16.7-13)[31* 341. M A 0 was used in viuo and in uitro as a catalyst for the production of norlaudanosine from dopamine (Fig. 16.7-14)r3'1. Norlaudanosine is an important synthon for benzylisoquinoline alkaloids, providing the upper isoquinoline portion of the morphinan skeleton. In uitro and in viuo yields were in the range of 20%. MAO-B was also tested in low water content organic media such as ether, tetrachloromethane, octane, benzene and cyclohexane. Under optimized conditions quantitative conversions of various substrates were achieved (Table 16.7-2)f3'1. 16.7.3.2.2 Diarnine Oxidase (E.C. 1.4.3.6)

The copper-containing amine oxidases (copper amine oxidases, diamine oxidases) possess either a topaquinone or a 6-hydroxydopamine cofactor (Fig. 16.7-15), generally integrated in the oxidase primary structure. Tyrosine residues of the enzyme backbone in the active site are discussed as precursors for the prosthetic group[37]. As the name suggests, diamine oxidase catalyzes the oxidative deamination of diamines. Preferably a,w-diamines such as putrescine (1,4-diaminobutane) or cadaverine (1,s-diaminopentane) (the names already suggest their smell), but also various derivatives are readily converted. Quite often cyclic imines are obtained via internal nucleophilic attack by the unreacted amino function (Fig. 16.7-16)[38-401.

I

1259

1260

I

7G Oxidation Reactions

o$

H_

2e-2H+

-

H

o

COOH d

Figure 16.7-15. Topaquinone and 6-hydroxy-dopamine as prosthetic groups of diamine oxidases.

OH OH topaquinone

H2N n = 1,2,3

OH 6-hydroxydopamine

n diarnine oxidase

(-3b) N

Figure 16.7-16. Application of diamine oxidase in the synthesis of different azaheterocycles.

X = 0, S , CH(CH,)

In the presence of suitable nucleophiles (such as benzoyl acetic acid) the primary imines can be spontaneously further modified in situ. A convenient approach to obtain phenacyl-derivatives,building blocks in the synthesis of certain alkaloids, was reported r3’]. In some cases, diamine oxidases exhibit activities complementary to monoamine oxidases. For example vanillylamine is far more efficiently converted into vanillin by a diamine oxidase from Aspergillus niger than by the monoamine oxidase from E. coli[”]. Even enantioselectiveoxidations of some alkyl-,benzyl-, or phenylethyl- (arylethyl) amines were reported with diamine oxidase from pea settlings[41].Porcine kidney diamine oxidase was used for the oxidative transformation of Nitruria alkaloids such as na~lininI~~1. For the conversion of poorly water-soluble amines (and to avoid product inhibition), diamine oxidase can also be applied in non-aqueous media[43]. 16.7.4 Oxidations Catalyzed by Transaminases

Transaminases are generally not considered to be enzymes catalyzing redox reactions, which is obvious considering the meaning of the E. C. code for transferases (E. C. 2.6.1.x = transferring amino groups). Nevertheless, the exchange of an amino functionality between an amino acid and an a-keto acid implies the oxidation of the amino acid. Transaminases are described elsewhere in this book (Chapter 12).

References References L. G. Lee, G. M. Whitesides, J . Am. Chem. SOC.1985, 107,6999-7008. 2 J. M. Laval, J. Moiroux, C. Bourdillon, Biotech. Bioeng. 1991, 38,788-796. 3 A. Anne, C. Bourdillon, S. Daninos, J . Moiroux, Biotech. Bioeng. 1999, 64, 101-107. 4 A.-E. Biade, C. Bourdillon, J.-M. Laval, G. Mairesse, J. Moiroux, J . Am. Chem. Soc. 1992,114,893-897. 5 A. Fassouane, J.-M. Laval, J. Moiroux, C. Bourdillon, Biotech. Bioeng. 1990,35, 935-939. 6 M. Nagel, J. R. Andreesen, Arch. Microbiol. 1990,154,605-613. 7 G. Wittig, Angew. Chem. 1957,69,245-251. 8 D. E. Hughes, Biochem. J. 1955,60, 303-310. 9 J. S. Holcenberg, E. R. Stadtman,]. Bid. Chem. 1969,244,1194-1203. 10 M. Petersen, A. Kiener, Green Chem. 1999, 99-106. 11 T. Yoshida,T. Nagasawa, Biosci. Biotech. Biochem. 2000,89,111-118. 12 A. Tinschert, A. Tschech, K. Heinzmann, A. Kiener, Appl. Microbiol. Biotech. 2000, 53, 185-195. 13 P. Lehky, H. Kulla, S. Mischler. Verfahren zur Herstellung von 6-Hydroxynikotinsaure; EP 015 2948 A2: Lonza, Switzerland, 1995. 14 V. W. Rodwell, Methods Enzmol. 1971, 17B, 174-188. 15 P. Wikstrom, E. Szwajcer, P. Brodelius, K. Nilsson, K. Mosbach, Biotechnol. Lett. 1982, 4,153-158. 16 K. Parkin, H. 0. Hultin, Biotech. Bioeng. 1979,939-953. 17 N. Nakajima, D. Conrad, H. Sumi, N. Esaki. C. Wandrey, K. Soda, Ferment. Bioeng. 1990, 70, 322-325. 18 R. Fernandez-Lafuente, V. Rodriguez, J. Guisan, Enz. Microb. Tech. 1998, 23. 19 S. M. Hecht, K. M. Rupprecht, P. M. Jacobs, J. Am. Chem. Soc. 1979, 101,3982-3983. 20 W. Kroutil, K. Faber, Tetrahedron: Asym. 1998,9,2901-2913. 21 J. W. Huh, K. Yokoigawa, N. Esaki, K. Soda, J . Ferment. Bioeng. 1992,74, 189-190. 22 J. W. Huh, K. Yokoigawa, N. Esaki, K. Soda, Biosci. Biotech. Biochem. 1992, 56, 2081-2082. 1

K. Soda, T. Oikawa, K. Yokoigawa,J. Mol. Cat. B: Enzymatic 2001, 11, 149-153. 24 J. Verweij, E. D. Vroom, Rec. Trav. Chim. P ~ ~ s - 1993, B ~ s 112,6681. 25 F. Alfani, M. Cantarella, A. Gallifuoco, Biocat. Biotransf: 1998, 16, 395-409. 26 A. Liese, K. Seelbach, C. Wandrey. In Industrial BiotranSformations, Wiley-VCH, Weinheim, 2000, pp. 129-130 and 225230. 27 R. B. Silverman, Biochem. SOC.Trans. 1991, 19,201-206. 28 W. Weyler, Y.-P. P. Hsu, X. 0. Breakefield, Pharmacol. Ther. 1990,47, 391-417. 29 J. F. Powell, Biochem. SOC.Trans. 1991, 19, 199-214. 30 P. Dostert, M. Strolin Benedetti, K. F. Tipton, Med. Res. Rev. 1989,9,45-89. 31 S. K. Nimkar, S. Mabic, A. H. Anderson, S. L. Palmer, T. H. Graham, M. de Jonge, L. Hazelwood, S. J. Hislop, N. Castagnoli, J. Med. Chem. 1999,42,182&1835. 32 A. Anderson, S. Kuttab, N. J. Castagnoli, Biochem. 1996,35, 3335-3340. 33 B. Y. Zhong, R. B. Silverman,J . Am. Chem. SOC.1997, 119,6690-6691. 34 J. Yu, N. Castagnoli. Bioinorg. Med. Chem. 1999, 7. 35 L. K. Hoover, M. Moo-Young,R. L. Legge, Biotech. Bioeng. 1991, 38, 1029-1033. 36 J. C. G. Woo, X. Wang, R. B. Silverman,J. Org. Chem. 1995,60,6235-6236. 37 N. K. Williams, J. P. Klinman, J. Mol. Cat. B: Enzymatic 2000,8,95-101. 38 J. E. Cragg, R. B. Herbert, M. M. Kgaphola, Tetrahedron Lett. 1990, 31, 6907-6910. 39 A.M. Equi, A. M. Brown, A. Copper, S. K. Ner, A. B. Watson, D. J. Robins, Tetrahedron 1991,47. 40 E. Santaniello, A. Manzocchi, P. A. Biondi, C. Secchi, T.Simonic, J. Chem. Soc., Chem. Commun.1984,803-804. 41 A. R. Battersby, J. Staunton, M. C. Summers, J . Chem. Soc., Chem. Commun.1974, 465,548-549. 42 E. Cheng, J. Botzem, M. J. Wanner, B. E. B u m , G.-J. Koomen, Tetrahedron 1996, 52, 5725-6732. 43 J. A. Chaplin, C. L. Budde, Y. L. Khmelnitsky,J. Mol. Cat. B: Enzymatic 2001, 13, 69-75. 23

11261

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

1262

I

lG Oxidation Reactions 16.8

Oxidation at Sulfur Karl-Heinz van Pee 16.8.1

Enzymes Oxidizing at Sulfur and their Sources

The oxidation at sulfur is catalyzed by a number of different enzymes produced by a variety of organisms. They have been isolated from a fungus"], soybeanr2],rat, pig and rabbit liver L3-'1, horseradish r61, bacteria ['-I, milk[10],and human white blood cells ~ ' 1 . The enzymes catalyzing oxidation reactions at sulfur belong to two different classes of enzymes: monooxygenases, including cytochrome P-450monooxygenases and FAD-containing monooxygenases, and heme-containing peroxidases (Figs. 16.8-1and 16.8-2, Table 16.8-1). Some of these enzymes such as chloroperoxidase from Caldariomyces $mago, horseradish peroxidase, lactoperoxidase from bovine milk, and myeloperoxidase from human white blood cells are commercially available. Others such as pig liver microsomal FAD-containing monooxygenase have to be isolated from tissue with very low yieldsI41or like hydrocarbon monooxygenase from Pseudomonas oleovorans [12-131 require several protein components and cofactors, substantially limiting the use of these enzymes for the production of oxidized sulfur compounds.

S-CH,

monooxygenase

SOCH,

H A (R)

or

(S)

Figure 16.8-1. Oxidation of methyl p-tolyl sulfide to methyl p-tolyl sulfoxide by a monooxygenase. The product can either be ofthe R- or S-configuration depending on the monooxygenase used.

peroxidase/haloperoxidase

SOCH,

HZOZ (R)

or

(S)

Figure 16.8-2. Oxidation of methyl p-tolyl sulfide to methyl p-tolyl sulfoxide by a peroxidase or haloperoxidase in the presence of hydrogen peroxide. The product can either be predominantly of the R- or S-configuration depending on the peroxidase or haloperoxidase used.

168 Oxidation a t Sulfur Table 16.8-1.

Classification o f enzymes oxidizing at sulfur and their sources.

Enzyme class

Source

Reference

Monooxygenases

pig liver microsomes rat liver microsomes rabbit liver microsomes bovine adrenals Pseudomonas oleovorans Acinetobacter sp. soybean horseradish Caldariomyces&mago Ascophyllumnodosum Corallina ojicinalis human white blood cells bovine milk

14 3 16 17 12,13 15 2 23 1 27 28 30 25

Peroxidases Haloperoxidases

16.8.2

Oxidation of Sulfides 16.8.2.1

Oxidation of Sulfides by Monooxygenasesand by Whole Organsims

Fujimori et al. [I4] used pig liver microsomal FAD-containing monooxygenase and phenobarbital-induced rabbit liver microsomal cytochrome P-450 to catalyze the oxidation of unsymmetrical sulfides to the corresponding optically active sulfoxides with varying degrees of enantiomeric excess (12-96%). Comparison of the oxygenation of racemic 2-methyl-2,3-dihydrobenzo[b] thiophene showed that the enantiotopic, diastereotopic, and enantiomeric differentiating abilities of the FADcontaining monooxygenase are higher than those of the cytochrome P-450 monooxygenase. They found that the oxygenation with the FAD-containing monooxygenase is sterically much more highly controlled than that with cytochrome I?-450. Whereas higher ee-values are observed in the oxygenation of smaller sulfides with the FAD-containing monooxygenase, the oxygenation of large sulfides by the cytochrome P-450 monooxygenase results in higher ee values than those of sulfides bearing small substituents. Hydrocarbon monooxygenase from Pseudomonas oleo~orans[~-'*I also catalyzes the stereoselective sulfoxidation of methyl thioether substrates [131 with up to 80% ee. The products obtained with this enzyme are probably ofthe R-configiration. The (S)-(-)-sulfoxideis predominantly produced (82 % S, 18 % R) from p-tolyl ethyl sulfide when cyclohexanone monooxygenase from Acinetobacter sp. NCIB 9871[I' was used, whereas the the FAD-containing monooxygenase from hog liver microsomes oxidizes p-tolyl ethyl sulfide to yield the (R)-(+)-sulfoxideenantiomer as the major product (95 % R, 5 % S) Llsl. The enzymatic oxidation of various diaryl, dialkyl, and aryl alkyl sulfides by cytochrome P-450 from rabbit liver resulted predominantly in the formation of the sulfoxides with the R-configuration[16]. '9

I

1263

1264

I

7G Oxidation Reactions

The S-(-)configuration was predominantly obtained when two cytochrome P-450 isoenzymes from rat liver were used for the oxidation of p-tolyl ethyl Oxidation of phenyl2-aminoethyl sulfide by dopamine b-hydroxylase from bovine adrenals in the presence of ascorbate as the electron donor resulted in the formation of phenyl 2-aminoethyl sulfoxide. The product was probably of the S-configuration[l7I. Holland et al. [18] obtained the (R)-sulfoxidesfrom various para-substituted phenyl 3-chloropropyl and phenyl 3-hydroxypropyl sulfides by biotransformation with the fungus Mortierella isabellina with an enantiomeric excess of 82-88%. The (S)sulfoxides were produced using the fungus Helrninthosporiurn sp. and the bacterium Acinetobacter calcoaceticuswith ee values of > 95 % and 94%, respectively. 16.8.2.2 Oxidation of Sulfides by Peroxidases and Haloperoxidases

A number of peroxidases were investigated for their use in oxidizing organic sulfides. p-Substituted thioanisols were oxidized by partially purified soybean sulfoxidase using 13(S)-hydroperoxylinoleicacid as the peroxide. Methyl p-tolyl sulfide gave the (S)-sulfoxidewith about 90 % ee 12]. The sulfoxidation of organic sulfides by chloroperoxidase from Caldariomyces firnago was investigated by different groups [19-261. Colonna et al. ['I compared the oxidation of sulfides by this enzymes with that catalyzed by horseradish peroxidase. Chloroperoxidase catalyzed the formation of sulfoxides with tert-butyl and other peroxides with an R absolute configuration in up to 92% ee, whereas horseradish peroxidase gave racemic products. When sterically hindered oxidants such as cumyl hydroperoxides and chloroperoxidase were used, racemic or almost racemic products were obtained. tert-Butyl hydroperoxide also had the advantage of giving higher yields and higher ee. Using vanadium bromoperoxidases from marine algae the (S)- or (R)-sulfoxides can be obtained from methyl phenyl sulfide derivatives, respectively, depending on the source of the enzyme. While bromoperoxidase from Ascophgllum nodosum produces the (R)-sulfoxidewith 91% ee1271,the (S)-enantiomer is obtained with bromoperoxidases from CoralZina o#cinalis and C. piluli&ra[281. When investigating the substrate selectivity using a series of aryl, alkyl, dialkyl, and heterocyclic sulfides, it was found that p-substitution led to higher enatioselectivity and higher chemical yields with respect to o-substitution[*'].A similar influence of the p-substitution was found for sulfoxidation catalyzed by bromoperoxidase from the marine alga ~scophyZLum~ o ~ o s u ~ ~ ~ I . Benzyl methylsulfide, thioanisol, and thiobenzamide were oxidized by chloroperoxidase, lactoperoxidase, and horseradish peroxidase to the respective sulfoxides. Whereas lactoperoxidase and horseradish peroxidase had low activities towards benzyl methylsulfide, thiobenzamide was efficiently oxidized by lactoperoxidase. Chloroperoxidase had high activity in halide-independent reactions towards all three substrates['']. This enzyme was also used for the asymmetric sulfoxidation of a series of cyclic sulfides. In all cases the (R)-sulfoxideswere obtained. In the case of

S

Ethyl p-tolyl

Methyl alkyl Diaryl, dialkyl, aryl alkyl Phenyl2-aminoethyl Phenyl3-chloropropyl Phenyl3-chloropropyl 2,3-Dihydrobenzo[b]thiophene

S

Methyl p-tolyl

S

R R

Predominant configuration of sulfoxide obtained

chloroperoxidase vanadium bromoperoxidase vanadium bromoperoxidase soybean hydroperoxidedependent oxygenase rat liver cytochrome P-450 cyclohexanone monooxygenase FAD-containingmonooxygenase alkane monooxygenase rabbit liver cytochrome P-450 dopamine P-hydroxylase Mortierella isabellina Helminthosporiurn sp. chloroperoxidase

Enzyme or organism

Products and absolute configuration obtained in the oxidation of various sulfides by different enzymes.

Methyl phenyl

Sulfide

Table 16.8-2.

3 15 15 13 16 17 18 18 29

19-21 27 28 2

Reference

Q

1266

I

76 Oxidation Reactions

2,3-dihydrobenzo[b]thiophenethe yield was 99.5 % with an ee of 99%[*'1. Table 16.8-2shows some examples of sulfides oxidized to sulfoxides by different enzymes and the absolute configuration of the products. When using peroxidases, care has to be taken, as the peroxidase-catalyzed oxidation is in competition with the spontaneous oxidation of the sulfides by the oxidant. Depending on the enzyme used for oxidation of organic sulfides, sulfoxides with S- or R-configuration can be obtained with high ee, whereas at present there is only one chemical oxidation method which leads to high ee in alkyl aryl sulfoxides. This method uses chiral titanium complexes and cumene hydroperoxide for the oxidation of organic sulfides L2'].

References 1

2

D. R. Morris, L. P. Hager,]. B i d . Chem. 1966,241,1763-1768.

E. Blee, F. Schuber, Biochemistry 1989, 28,

4962-4967.

Kim, T. Iyanagi, S. Oae, Bull. Chem. Soc. Jpn. 1983,56,2300-2310. 17 S. W. May, R. S. Phillips, J . Am. Chem. SOC. 1980,102,5983-5984.

3 D. J. Waxrnan, D. R. Light, C. Walsh, Bio-

18 H. L. Holland, J.-X. Gu, A. Kerridge, A.

4

19 S. Colonna, N.Gaggero, A. Manfredi, L.

chemistry 1982,21,2499-2507. D. M. Ziegler, L. L. Poulsen, Methods Enzymol. 1978, Vol. 52, 142-151. 5 Y. Irnai, R. Sato, Biochem. Biophys. Res. Comm u n . 1973, 60, &14. 6 L. M. Shannon, E. Kay, J. Y. Lew, J. Biol. Chem. 1966,241,2166-2171. 7 A. G. Katopodis, K. Wimalasena, J. Lee, S. W. May,]. Am. Chem. SOC.1984, 106, 792&7935. 8 S. W. May, L. G. Lee, A. G. Katopodis, J.Y. Kuo, K. Wirnalasena, J. R. Thowsen, Biochemistry 1984,23,2187-2192. 9 N. A. Donoghue, D. B. Norris, P. W. Trudgill, Eur. J. Biochem. 1976, 63, 175-192. 10 C. Dumontet, B. Rousset,]. Biol. Chem. 1983,258,14166-14172. 11 J. Schultz, Reticuloendothel. Syst. 1980, 2, 231-254. 12 S. W. May, A. G. Katopodis, Enzyme Microb. Technol. 1986,8, 17-21. 13 A. G.Katopodis, H. A. Smith, Jr., S. W. May, /.Am. Chem. SOC.1988, 110,897-899. 14 K. Fujirnori, T. Matsuura, A. Mikami, Y. Watanabe, S. Oae, T. Iyanagi, /. Chem. Soc. Perkin Trans. 1 1990, 1435-1440. 15 D. R. Light, D. J. Waxrnan, C. Walsh, Biochemistry 1982, 21, 2490-2498. 16 T. Takata, M. Yarnazaki, K. Fujirnori, Y. H.

Willetts, Biocat. Biotrans. 1999, 17, 305-317.

Casella, M. Gullotti, /. Chem. Soc., Chem. Commun. 1988,1451-1452. 20 S . Colonna, N. Gaggero, A. Manfredi, L. Casella, M. Gullotti, G. Carrea, P.Pasta, Biochemistry 1990, 29, 10465-1048. 21 S. Colonna, N. Gaggero, L. Casella, G. Carrea, P. Pasta, Tetrahedron: Asymmetry 1992, 3,95-106. 22 L. Casella, S. Colonna, G. Carrea, Biochemistry 1992,31,9451-9459. 23 S. Kobayashi, M. Nakano, T. Goto, T. Kirnura, A. P. Schaap, Biochem. Biophys. Res. Commun. 1986, 135, 166-171. 24 S. Kobayashi, M.Nakano, T. Kimura, A. P. Schaap, Biochemistry 1987,26, 5019-5022. 25 D. R. Doerge, Arch. Biochem. Biophys. 1986, 244,678-685. 26 D. R. Doerge, N. M. Cooray, M. E. Brewster, Biochemistry 1991,30,8960-8964. 27 H. B. ten Brink, H. L. Holland, H. E. Shoemaker, H. van Lingen, R. Wever, Tetrahedron: Asymmetry 1999, 10,4563-4572. 28 M. A. Anderson, S. G. Allenmark, Tetrahedron 1998,54,15293-15304. 29 S . G. Allenmark, M. A. Anderson, Chirality 1998, 10,246-252. 30 M.-F. Tsan,/. Cell.Physiol. 1982, I l l , 49-54.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002

76.9 Halogenation

I

1267

16.9 Halogenation Karl-Heinz van Pee 16.9.1 Classification of Halogenating Enzymes and their Reaction Mechanisms 16.9.1.1

Haloperoxidases and Perhydrolases

The only type of halogenating enzymes known until 1997 were peroxidases and perhydrolases which catalyze the formation of carbon halogen bonds using halide ions, hydrogen peroxide and an organic substrate activated for electrophilic attack. According to the halide ions they can utilize they are arranged into three groups: iodoperoxidases, bromoperoxidasesand chloroperoxidases. Iodoperoxidases catalyze the formation of carbon-iodine bonds, whereas bromoperoxidases catalyze iodination and bromination reactions, and chloroperoxidases catalyze the iodination, bromination, and chlorination of organic substrates. As haloperoxidases are oxidoreductases using hydrogen peroxidase as the oxidant for the oxidation of halide ions producing hypohalogenic acids, the existence of fluoroperoxidases can be ruled out. The overall reactions catalyzed by haloperoxidases and perhydrolases are shown in Fig. 16.9-1. All haloperoxidases isolated until 1984 were heme-containing enzymes[’]. The first non-heme haloperoxidase was isolated by Vilterr2]. Instead of heme, vanadium is responsible for the halogenating activity of this algal enzyme 41. Non-heme and non-metal “haloperoxidases”were isolated from bacteria however, elucidation of the three-dimensional structure and the reaction 139

‘-+

a) heme-haloperoxidase

’)

H’

b) vanadium haloperoxidase

2) R-H + HOX

1) CH,COOH

*

+ HO ,,

2) CH,COOOH

3)HOX + R-H

+ X-

c) perhydrolase

HOX

R-X+ H,O

-

CH,COOOH

* CH,COOH + HOX *

R-X+H,O

X- = CI-, Br, IFigure 16.9-1. Overall reaction catalyzed by (a) heme-IS’]and (b) vanadiumcontaining haloperoxidases[5’’ and (c) perhydrolases []’‘.

1268

I

7G Oxidation Reactions

mechansim of this type of halogenase showed that they are not real haloperoxidases. They are actually perhydrolases which produce hypohalogenicacids via the oxidation of halide ions by enzymatically formed peracetic acid[10-121.Thus, in addition to grouping the haloperoxidases according to the range of halide ions oxidized, they can be classified according to their prosthetic group into heme type and non-heme type haloperoxidases L1 '1. The heme type haloperoxidases are inactivated during the halogenation reaction, because the heme group of these enzymes is attacked by the hypohalous acids produced by the enzymes [l].Thus, heme type haloperoxidases have the disadvantage that the reaction velocity slows down considerably during the course of the reacti0n['~1.With non-heme type haloperoxidases this does not seem to be the case. They are not inactivated during the halogenation reaction and are very stable under reaction conditions [l41. However, the disadvantage of inactivation is partly compensated for by the fact that some of the heme type haloperoxidases have much higher specific activities than non-heme type haloperoxidases. Some of the non-heme haloperoxidases are very stable with respect to organic solvents[15]which is of great importance when the substrates that are to be halogenated are not very soluble in water. In these cases water missible organic solvents can be added to the reaction mixture or a two phase-system can be used. 16.9.1.2 FADH2-dependent Halogenases

In 1997 the existence of a novel class of halogenating enzymes was reported[161. These halogenases showed no relationship to any of the known haloperoxidases[l7.1" and did not require hydrogen peroxide for halogenating activity. Initially these new halogenases were thought to require NADH [16],but more detailed studies showed that they actually require FADHZ['~, 1" which is produced by NADHdependent flavin reductases. Figure 16.9-2 shows the hypothetical reaction mechanism of FADH2-dependent halogenases. 16.9.2 Sources and Production of Enzymes 16.9.2.1 FADH2-dependent Halogenases

Although FADH2-dependent halogenases seem to be present in many bacteria producing halometabolites[19z21, 22], only one example of this new class of halogenases has been isolated to homogeneity until now. This enzyme, tryptophan 7-halogenase, is produced by several Pseudomonas strains producing the antibiotic pyrrolnitrin such as Pseudomonas fluorescens and P. aureofaciens and by Myxococcus f i l ~ u s [ ~Monodechloroaminopyrrolnitrin-3-halogenase, ~]. another FADH2-dependent halogenase from pyrrolnitrin-producing Pseudomonas strains has so far only been purified partially [ I 6 . 2ol.

1G. 9 Halogenation

FAD + NADH + H+

flavin reductase

halogenase FADH,, 0,

QTC C & O :OH

FADH,

~

+ NAD+

07

C !{;iOOH

0

H

CI-, H+

-

PTC&:COOH

halogenase - H,O

halogenase

QTC C & O :OH HO

CI

CI

Hypothetical reaction mechanism of FADH2-dependent tryptophan 7-halogenase as an example of FADH2-dependent halogenases[201.

Figure 16.9-2.

From biosynthetic and hybridization studies it is known that FADH2-dependent halogenases are involved in the biosynthesis of many halometabolites produced by 221 and it can be expected that other FADH2-dependent halogenases bacteria['', will be purified and characterized in the near future. "9

16.9.2.2

Haloperoxidases and Perhydrolases

Iodoperoxidases such as horseradish peroxidase [241 and thyroid peroxidase LZ51 can be isolated from many different organisms. Bromoperoxidases have been obtained in a pure form from mammals (lactoperoxidase)[26], sea urchin (ovoperoxidase)[I', marine algae 128-301, fungi (lignin peroxidase)[321 and bacteria [33, 341. Chloroperoxidases have been found in mammals (myeloperoxida~e[~~1 and eosinophil peroxidase [36j), a marine worm [371, and fungi [3s401. Several perhydrolases have been isolated from bacteria L5-', 4Ll. Chloroperoxidase can be produced in batch culture at concentrations of 280 mg L-1[421 and 20 mg of lactoperoxidase can be isolated from 1 L of bovine milk[Z6].The sources for these two enzymes, bovine milk and culture broth of Caldariomyces

+ FAD + H,O

1270

IG

I filmago,

Oxidation Reactions

are easily obtained. Chloroperoxidase can also be obtained in larger quantities from the fungus Curvularia i n a e q u ~ l i s [ ~Thus, ~ ] . a number of different haloperoxidases from various sources are available in quantities necessary for the enzymatic halogenation of organic compounds.

tryptophan

tryptamine

3-methylindole

rnonodechloroaminopyrrolnitrin

indole-3-acetonitrile

5-methylindole

aminopyrrolnitrin

Figure 16.9-3. Substrates accepted by tryptophan 7-halogenase: (a) indole derivatives, (b) phenylpyrrole derivatives; the positions of chlorination are indicated by

1G. 9 Halogenation

I

1271

16.9.3 Substrates for Halogenating Enzymes and Reaction Products 16.9.3.1 Halogenation of Aromatic compounds

The recently detected FADHz-dependenthalogenases are substrate specific. Tryptophan 7-halogenase catalyzes the chlorination and bromination of D- and L-tryptophan to 7-chloro-or 7-bromotryptophan, This enzyme also accepts a number of other indole derivatives such as tryptamine, indole-3-acetonitrile,3-methylindole and 5-methylindoleas substrates (Fig. 16.9-3a)[431. In addition to indoles, aminophenylpyrrole derivatives are also chlorinated by tryptophan 7-halogenase (Fig. 16.9-3b)[431. Monodechloroaminopyrrolnitrin 3-halogenase catalyzes the regioselective chlorination of the aminophenylpyrrole derivative monodechloroaminopyrrolnitrin to aminopyrrolnitrin ["l, however, nothing is known about the substrate specificity of this enzyme. In contrast to FADH2-dependent halogenases, haloperoxidases have no substrate specificity. The enzymatic iodination, bromination, and chlorination of a number of different aromatic compounds by haloperoxidases have been reported in the last few years. All aromatic substrates halogenated successfully by haloperoxidases are aromatic compounds activated for electrophilic substitution (Table 16.9-1). Phenols and phenol ethers are very good substrates for haloperoxidases. The first aromatic substrate to be used in enzymatic iodination was tyrosine. This substrate was iodinated using chloroperoxidase from Caldariomyces fimago[441and thyroid pero~idase[~'l. Horseradish peroxidase and lactoperoxidase have been used to lable proteins with radioactive isotopes of iodide 1'1 and bromoperoxidase from Penicillus capitatus has been employed to lable human serum albumin with the radioactive isotope of bromine 14'4. Phenolsulfonephthalein (Phenol Red) is brominated to 3,3',5,5'-tetrabromophenolsulfonephthalein (Bromophenol Blue) by many haloperoxidases fl, "1. This reaction has been used for the detection of halogenating enzymes by different groups 19, 471. Corbett et al.[481obtained 2,6-dibromo-4-chloroaniline or 2,4,6-trichloroaniline when they incubated 4-chloroaniline with chloroperoxidase in the presence of hydrogen peroxide and bromide or chloride, respectively. Several obscurolides, secondary metabolites produced by Streptomyces viridochromogenes T7 [491, were brominated using perhydrolase from Streptomyces aureofaciens Tu24[501.The obscurolides were monobrominated in the 2-position and dibrominated in the 2,4-positions of the aromatic ring system of the obscurolides (Fig. 16.9-4). In the case of dibromination, the hydroxymethyl group was replaced by bromine. No bromination of the olefinic double bond could be detected. A number of aromatic heterocycliccompounds have been halogenated by different haloperoxidases. Franssen et al. Is']used chloroperoxidase from Cddariomycesfimago to chlorinate

Substrate (halide)

indole derivatives (Cl-, Br-) phenylpyrroles (Cl-, Br-) monodechloroaminopyrrolnitrin (Cl-, Br-) phenol ether (Cl-, Br-) phenols (Cl-, Br-, I-) anilines (Cl-, Br-) pyrazoles, pyridines (Cl-) nucleic bases (Cl-, Br-, I-) phenols (I-) estrone (I-) phenols (I-) phenols (Cl-, Br-) phenols (Br-, I-) phenol red (Br-) nudeic bases (Br-, I-) phenols (Br-) nikkomycin (Br-) phenylpyrroles (Cl-, Br-) obscurolide (Br-) phenol red (Br-) indole (Cl-, Br-) phenylpyrroles (Cl-, Br-)

Source

pyrrolnitrin-producing Pseudomonads pyrrolnitrin-producing Pseudomonads Caldariornycesfumago

bovine milk thyroid glands Notomastus lobatus Asophyllum nodosum Corallina pilulijera Streptomyces aureofaciens

Pseudomonas pyrrocinia

Tryptophan 7-halogenase (FADHz-dependent) Monodechloroaminopyrrolnitrin 3halogenase (FADHz-dependent) Chloroperoxidase (heme)

Lactoperoxidase(heme)

Thyroid peroxidase (heme) Chloroperoxidase (heme-flavin) Bromoperoxidase (non-heme)

Bromoperoxidase (non-heme)

Perhydrolase (non-heme, non-metal)

Perhydrolase (non-heme, non-metal)

20 67-68 44 48 51 14 72 73 45 37 74 15 14 75 52 56 50 9 29 7, 54, 56

20,43 43

Reference

FADHz-dependenthalogenases, haloperoxidases and perhydrolases used for biotransformation of aromatic compounds and their sources.

Enzyme (type)

Table 16.9-1.

4

48' a

P

% 8'

3.

0

m

d

N

-

c:

7G. 9 Halogenation

I

1273

Figure 16.9-4. Bromination of obscurolide A3 by perhydrolase from Streptornyces aureofaciens Tu24 [501.

pyrazole, 1-methylpyrazole and 3-methylpyrazole to their corresponding 4-chloroderivatives. The same enzyme was used to produce 5,7-dibromo-8-hydroxyquinoline from 8-hydroxyquinoline.2-Aminopyridinewas regiospecifically chlorinated to 2-amino-3-chloropyridine. The chlorination, bromination, and iodination of various nitrogen-containing heterocycles catalyzed by chloroperoxidase from Caldariomyces &mago and bromoperoxidase from Corallina pilulijira were compared by Itoh et al. [141. The nucleoside antibiotic nikkomycin Z was brominated using the perhydrolase from Streptomyces aureofaciens Ti24[521.Bromination occurred at the 6-position and at the 4,G-positions of the pyridine system of nikkomycin Z. The antifungal antibiotic pyrrolnitrin [3-chloro-4-(2-nitro-3-chlorophenyl)pyrrole] was brominated at the 2-position of the pyrrole moiety by bromoperoxidase from Streptomyces phaeochromogenes ts31. Pyrrolnitrin was chlorinated at the 2-position and at the 2,5-positions of the pyrrole system by perhydrolases from Pseudomonas pyrrocinia and Streptomyces aureofaciens. The corresponding bromo-derivatives were also obtained with these enzymes [541. Another phenylpyrrole compound, 2-(3,5-dibromo-2-methoxyphenyl)pyrrole was brominated to 2-bromo-,2,3-dibromo-,3,4-dibromo-,2,3,4-tribromo-5-(3,5-dibromo2-methoxypheny1)pyrroleby perhydrolase from Streptomyces aureofaciens Tu24[”1. When the same substrate was chlorinated using the perhydrolases from Pseudomonas pyrrocinia and Streptomyces aureofaciens Tu24, 2-chloro-, 3-chloro-, 4-chloro-, 2,3-dichloro-, 2,4-dichloro, and 3,4-dichloro-5-(3,5-dibromo-2-methoxyphenyl)pyrrole could be isolated (Fig. 16.9-5)[56]. 16.9.3.2

Halogenation of Aliphatic Compounds

Haloperoxidases catalyze the halogenation of a wide range of alkene substrates. Ethylene was iodinated, brominated, and chlorinated to the corresponding 2-haloethanol by chloroperoxidase from Caldariomycesfimago. Using the same enzyme and propylene as the substrate the 1-halo-2-propanolsand the 2-halo-1-propanolswere obtained. 1,3-Butadienewas converted into 1-bromo-3-butene-2-01,2-bromo-3-buby lactoperoxidase (Fig. 16.9-6). Brominatene-1-01and 1,4-dibrom0-2,3-butanediol tion of allene by chloroperoxidase from Caldariomyces f i m a g o resulted in 2-bromo2-propen-1-01~~~1. Propylene, allyl chloride and allyl alcohol were halogenated to yield halohydrins and dihalogenated products [’*I. When several halide ions were present in the reaction mixture, heterogeneous dihalides were obtained[’”].The chlorination

1274

I

7G Oxidation Reactions

B

r

p

H,O,,ACOH,CIperhydrolase

'' " " c v c c l + $+$",Brv r v cc ' + B l r

/

/

Br

/

pe

/

Br br

Br

Br 8r

Br

Br

Br

Figure 16.9-5. Chlorination o f 2-(3,5-dibromo-2-methoxyphenyl)pyrrole by perhydrolase from Streptornyces aureofaciens Tu24 [561.

/ chloroperoxidase w H,O, B r Figure 16.9-6.

H3C

A

&+++Rr

~

OH

Br

Br

OH

Bromination o f 1J-butadiene by lactoperoxidase from bovine

chloroperoxidase

HO ,,

CI-

CI

OH

Figure 16.9-7. Chlorination o f methyl cyclopropane by chloroperoxidase from Caldariornycesfirnago 16'].

bromoperoxidase

H,O,

Br

Bromination o f monochlorodimedone, the substrate used for the search for halopereoxidasesl']. Figure 16.9-8.

of propenylphosphonic acid resulted in the formation of 1-chloro-2-hydroxypropylphosphonic acid[60].Phenyl acetylene was brominated to a-brornoacetophenone and a-dibromoacetophenone by lactoperoxidase. Chloroperoxidase from Culdariomyces &mugo was used to chlorinate methyl cyclopropane to 4-chloro-2-hydroxy-butane (Fig. 16.9-7)l6I1. Monochlorodirnedone, the substrate used for the detection and isolation of haloperoxidases and perhydrolases (Fig. 16.9-8), and other p-diketones such as barbituric acid[''] is brominated at the 2-position by all known haloperoxidases and perhydrolases. Oxooctanoic acid and other p-ketoacids form mono- and dihalogenated ketones and carbon dioxide["]. When p-alanine and taurine were used as substrates for myeloperoxidase the corresponding N-chloroamines could be detected [G6G51. As can be seen from the number of substrates halogenated by the different

1G.9 Halogenation

I

1275

haloperoxidases and perhydrolases, these enzymes show no substrate specificity. Examples of aliphatic substrates halogenated by haloperoxidases and perhydrolases are shown in Table 16.9-2. 16.9.4

Regioselectivityand Stereospecificityof Enzymatic Halogenation Reactions 16.9.4.1

FADH2-dependent Halogenases

FADHz-dependenttryptophan 7-halogenase shows regioselctivity which is dependent on the substrate used. With tryptophan, the enzyme is highly regioselective and catalyzes only halogenation at position 7 of the indole ring. However, with other indole derivatives halogenation occurs at positions 2 and 3 of the indole ring (Fig. 16.9-3a).Chlorination of aminophenylpyrrole derivatives by tryptophan 7-halogenase also proceeds with relaxed regioselectivity (Fig. 16-3b)[431. Nothing is know about the regioselectivity of monodechloroaminopyrrolnitrin 3-halogenaseand other FADH2-dependenthalogenases, but biosynthetic investigations suggest that many of these halogenases catalyze halogenation reactions 'j'j]. regio~e1ectively['~~ So far no investigations on the stereospecificity of this type of halogenating enzymes have been reported. Haloperoxidases show very poor regioselectivity. There are only very few reports on regioselective reactions catalyzed by haloperoxidases. Franssen et al. [ ' j 2 ] reported the regioselective chlorination of 2-aminopyridine to 2-amino-3-chloroaminopyridine by chloroperoxidase from Caldariomycesfimago. When anisole was brominated using chloroperoxidase from Caldariomyces fimago Walter and Ballschmitter ['j71 found a para-preference for the bromination reaction with a para :ortho ratio of 16 compared with 9 for the normal electrophilic bromination. The para: ortho ratio for the chlorination of anisole with the same enzyme obtained by Brown and Hager ['j8] was 1.9. This discrepancy could be due to the different reaction conditions used. Walter and Ballschmitter ['j71 used a 50 times higher anisole concentration and only about half the amount of chloroperoxidase compared with Brown and HagerL"]. If one takes into consideration that anisole could reach the active site of the enzyme and is present at a high concentration, a relatively large part of the substrate could be chlorinated at the active site with a certain orientation and only a smaller part would be chlorinated by enzymatically produced hypochlorous acid. This effect could be amplified by smaller amounts of the enzyme and thus lower concentrations of hypohalous acid produced. However, this would mean that halogenation occurring at the active site showed a higher degree of regioselectivity, even without a specific binding site for the organic substrate. Similar results were obtained by Itoh et al. [I4, 'j91 for the halogenation of different substrates using bromoperoxidase from Corallina pilulifera and chloroperoxidase from Caldariomycesfimago. In addition to poor regioselectivity, haloperoxidases also show poor stereospeci-

Caldariomycesfumago

bovine milk

mammals Penicillus capitatus Bonnemasoinia harngera Ascophyllum nodosum

Chloroperoxidase (heme)

Lactoperoxidase (heme)

Myeloperoxidase(heme)

Bromoperoxidase (heme)

Bromoperoxidase (non-heme)

Source

barbituric acid (Br-)

2-hydroxymethylene testosterone (Br-) 2-hydroxymethylene-17p-hydroxyandrostan-3-one (Br-) glycals (Cl-, Br-, I-) alkynes (Cl-, Br-, I-) cyclopropanes (Cl-,Br-) p-diketones (C1-, Br-) alkenes (Br-, I-) alkynes (Br-) cyclopropanes (Br-) alanine, taurine (Cl-) 0-alanine (Cl-) a-amino acids, peptides (Br-)

alkenes (Cl-, Br-, I-) 9(11)-dehydroprogesterone(Br-)

Substrate (halide)

Haloperoxidases used for biotransformations of aliphatic compounds and their sources.

Haloperoxidase (type)

Table 16.9-2.

62

71 61 61 44,70,79-81 57,59,82-84 61 61 65 64 85

78

77

Reference 57-60,70,75 76

B

8' ,3

3

2 a. ?iF:

m

-

U

m -

N

4

lG.9 Halogenation

I

1277

ficity. Kollonitsch et al. Ib0] obtained optically inactive erythro-dimethyl l-chloro2-hydroxypropylphosphonatefrom trans-propenylphosphonicacid using chloroperoxidase from Caldariomyces@mago. Ramakrishnan et al.I'[ investigated the bromiacid to the nation of racemic 2-e~o-methylbicyclo-[2.2.l]hept-5-ene-2-endo-carboxylic to the 2-exo-bromo-3-endo-hyS-lactone and racemic bicyclo-[3.2.0]hept-2-en-G-one droxybromohydrin. The products were obtained in racemic form. 2-Methyl-4-propylcyclopentane-1,3-dionewas chlorinated to 2-chloro-2-methyl-4-propylcyclopentane-1,3-dione. Here the product was obtained as a 40: GO ratio of the racemic diastereomers. From these findings they concluded that active site chlorination by chloroperoxidase from Caldariomyces &mago proceeds without appreciable stereoselectivity. On the other hand, Liu and W ~ n g ( ~ described *] the stereoselective bromohydrations of D-galactal and L-fucal to 2-bromo-2-deoxy-~-ga~actose (P/a= 3) and 2-bromo2-deoxy-~-fucose(p/a = 2), respectively. They also obtained the corresponding chlorinated products, however, in much lower yields. 16.9.5

Comparison of Chemical with Enzymatic Halogenation

NADH2-dependenttryptophan 7-halogenasecatalyzes the incoporation of a chloride atom into the indole ring at a position were direct chemical chlorination is not possible. The structures of metabolites containing halogenated indole rings suggest that similar halogenases exist which catalyze halogenation reactions at positions 2-7 of the indole ring. These enzymes are certainly very promising candidates as tools in organic synthesis, especially as they catalyze the incorporation of the halide atoms as nucleophiles, which allows regioselective and possibly stereoselective halogenation reactions. Enzymatic halogenation catalyzed by haloperoxidases and perhydrolases involves the oxidation of halide ions to a halonium ion species which leads to the formation of hypohalous acids (Fig. 16.9-1). The products obtained by enzymatic halogenation with these enzymes are the same as the products obtained by chemical electrophilic halogenation with hypohalous acids. The differences in the para :ortho ratios in the halogenation of some aromatic compounds could be due to a mixture of halogenation at or near the active site and in solution. The major advantage of enzymatic halogenation using haloperoxidases and perhydrolases is that the enzymes have a very low substrate specificity and that no free halogen is needed which makes halogenation catalyzed by these enzymes less hazardous than chemical halogenation. Some of the non-heme haloperoxidases and perhydrolases are very stable, even against organic solvents, and easy to use as they do not need any cofactors. However, care has to be taken not to use too high concentrations of hydrogen peroxide, as this could lead to oxidation of the substrate.

1278

I

16 Oxidation Reactions

References

S. L. Neidleman, J. Geigert, Biohalogenation: Principles, Basic Roles and Applications, Ellis Honvood Ltd., Chichester, UK. 1986. 2 H. Vilter, LeJol. Bot. Mar., 1983, 26, 451-455. 3 H. Vilter, Phytochemistry, 1984, 23, 387-1390. 4 R. Wever, H. Plat, E. De Boer, Biochim. Biophys. Acta 1985,830, 181-186. 5 K.-H. van Pee, G. Sury, F. Lingens, Biol. Chew. Hoppe-Seyler 1987,368,1225-1232. 6 B. E. Krenn, H. Plat, R. Wever, Biochim. Biophys Acta 1988,952, 255-260. 7 W. Wiesner, K.-H. van Pee, F. Lingens, J . E d . Chem. 1988,263,13725-13732. 8 R. Zeiner, K.-H. van P,e, F. Lingens, /. Gen. Microbiol. 1988, 134, 3141-3149. 9 M. Weng, 0. Pfeifer, S. Krauss, F. Lingens, K.-H. van Pee, J . Gen. Microbiol. 1991, 137, 2539-2546. 10 H. J. Hecht, H. Sobek. T. Haag, 0. Pfeifer, K.-H. van Pee, Nature Struct. Bid. 1994, 1, 532-537. 11 M. Picard, 7. Gross, E. Lubbert, S. Tolzer, S. Krauss, K.-H. van Pee, A. Berkessel, Angew. Chem. Int. Ed. Engl. 1997, 36, 1196-1199. 12 B. Hofmann, S. Tolzer, I. Pelletier, 7. Altenbuchner, K.-H. van Pee, H. J. Hecht, J . Mol. B i d . 1998, 279, 889-900. 13 N. Itoh, Y. Izumi, H. Yamada,/. Bid. Chem. 1986,261, 5194-5200. 14 N. Itoh, Y. Izumi, H. Yamada, Biochemistry 1987,26,282-289. 15 E. De Boer, H. Plat, M. G. M. Tromp, R. Wever, M. C. R. Franssen, H. C. van der Plas, E. M. Meijer, H. E. Schoemaker, Biotechnol. Bioeng. 1987, 30, 607-610. 16 K. Hohaus, A. Altmann, W. Burd, I. Fischer, P. E. Hammer, D. S. Hill, J. M. Ligon, K.-H. van Pee, Angew. Chem. Int. Ed. Engl. 1997, 36,2012-2013. 17 P. E. Hammer, D. S. Hill, S. T. Lam, K.-H. van Pee, 7. M. Ligon, Appl. Environ. Microbiol. 1997, 63, 2147-2154. 18 S. Kirner, P. E. Hammer, D. S. Hill, A. Altmann, I. Fischer, L. J. Weislo, M. Lanahan, K.-H. van Pee, J. M. Ligon,J. Bacteriol. 1998, 180,1939-1943. 19 K.-H. van Pee, S. Keller, T. Wage, I. Wynands, H. Schnerr, S. Zehner, Biol. Chem. 2000,381,l-5. 1

S. Keller, T. Wage, K. Hohaus, E. Eichhorn, K.-H. van Pee, Angav. Chem. Int. Ed. Engl. 2000,39,2300-2302. 21 S. Pelzer, R. SuBmuth, D. Heckmann, J. Recktenwald, P. Huber, G. Jung, W. Wohlleben, Antimicrob. Agents Chemother. 1999,43, 1565-1573. 22 B. Nowak-Thompson, N. Chaney, J. S. Wing, S. J. Gould, J. E. Loper, J . Bacteriol. 1999, 181,2166-2174. 23 P. E. Hammer, W. Burd, D. S. Hill, J. M. Ligon, K.-H. van Pee, FEMS Microbiol. Lett. 1999, 180, 39-44. 24 L. M. Shannon, E. Kay, J. Y. Lew, J . Biol. Chem., 1966,241,2166-2172. 25 N. M. Alexander,J. B i d . Chem. 1959,234, 1530-1533. 26 C. Dumontet, B. Rousset, J . Bid. Chem. 1983,258,14166-14172. 27 T. Deits, M. Farrance, E. S. Kay, L. Medill, E. E. Turner, P. J. Weidman, B. M. Shapiro, /. Bid. Chem. 1984,259,13 525-13533. 28 D. G. Baden, M. D. Corbett, Biochem. J. 1980, 187,205-211. 29 7. A. Manthey, L. P. Hager,]. Biol. Chem. 1981,256,11232-11238. 30 N. Itoh, Y. Izumi, H. Yamada, Biochem. Biophys Res. Commun. 1985, 131,42&435. 31 H. Plat, B. E. Krenn, R. Wever, Biochem. /. 1987,248,1123-1131. 32 V. Renagathan, K. Miki, M. H. Gold. Biochemistry 1987,26,5127-5132. 33 K.-H. van Pee, F. Lingens,J. Gen. Microbiol. 1985, 131,1911-1916. 34 M. Knoch, K.-H. van Pee, L. C. Vining, F. Lingens, J. Gen. Microbiol. 1989, 135, 2493-2502. 35 J. Schultz, Reticuloendothel. Syst. 1980, 2, 231-254. 36 J. T. Archer, G. Air, M. Jackas, D. B. Morell, Biochim. Biophys. Acta 1965, 99, 9 6 1 0 1 . 37 Y. P. Chen, D. E. Lincoln, S. A. Woodin, C. R. Lovel1,J. Bid. Chem. 1991, 266, 23 909-23915. 38 D. R. Morris, L. P. Hager, J . B i d . Chem. 1966,241,1763-1768. 39 T.-N. E. Liu, T. M'Timkulu, J. Geigert, B. Wolf, S. L. Neidleman, D. Silva, J. C. Hunter-Cevera, Biochem. Biophys. Res. Commun. 1987, 142,329-333. 40 B. H. Simons, P. Barnett, E. G. M. Vollen20

References I1279

broek, H. L. Dekker, A. 0. Muijsers, A. Messerschmidt, R. Wever, Eur. J. Biochem. 1995, 229,566-574. 41 W. Wiesner, K.-H. van Pee, F. Lingens, FEBS Lett., 1986,209, 321-324. 42 R. D. Carmichael, M. A. Pickard, Appl. Environ. Microbiol. 1989, 55, 17-20. 43 K.-H. van Pee, M. Holzer, in Tryptophan, Serotonin and Melatonin - Basic Aspects and Application. Advances i n Tryptophan Research (Eds: G. Huether, W. Koch, T. J. Simat, H. Steinhardt, Plenum Press, New York 1999. 44 L. P. Hager, D. R. Morris, F. S. Brown, H. Ebenvein, 1.Biol. Chem. 1966,24 1 , 1769-1777. 45 J. Nunez, Methods Enzymol. 1984, 107, 476-488. 46 J. A. Manthey, L. P. Hager, K. D. McElvany, Methods Enzymol. 1984, 107,439--445. 47 j. C. Hunter-Cevera, L. Sotos, Microb. Ecol. 1986,12,121-127. 48 M. D. Corbett, B. R. Chipko, A. 0. Batchelor, Biochem.]. 1980, 187,893-903. 49 H. Hoff, H. Drautz, H.-P. Fiedler, H. Zahner, j. E. Schultz, W. Keller-Schierlein,S. Philipps, M. Ritzau, A. Zeek,]. Antibiot. 1992,45, 1096-11107. 50 F. Thiermann, G. Bongs, K.-H. van Pee, D. Braun, H.-J.Cullmann, H.-P. Fiedler, H. Zahner, 10. DECHEMA-Jahrestagung der Biotechnologen, Karlsruhe, G e m a n y , 1992. 51 M. C. R. Franssen, H. G. van Boven, H. C. van der Plas,]. Heteroqcl. Chem. 1987, 24, 1313-1316. 52 H. Decker, U. Pfefferle, C. Bormann, H. Zahner, H.-P. Fiedler, K.-H. van Pbe, M. Rieck, W. A. Konig,]. Antibiot. 1991,44, 626-634. 53 K.-H. van Pee, F. Lingens, FEBS Lett. 1984, 173, 5-8. 54 G. Bongs, K.-H. van Pee, Enzyme Microb. Technol. 1994, IG, 53-GO. 55 H. Laatsch, H. Pudleiner, B. Pelizaeus, K.H. van Pee, Liebigs Ann. Chem. 1994, 65-71. 56 V. N. Burd, K.-H. van Pee, F. Lingens, A. Voskoboev, Bioorg. Khim 1992,18, 1002-1006. 57 J. Geigert, S. L. Neidleman, D. j. Dalietos, S. K. DeWitt, Appl. Environ. Microbiol. 1983, 45, 366-374. 58 J. Geigert, S. L. Neidleman, D. J. Dalietos, S. K. DeWitt, Appl. Environ. Microbiol. 1983, 45,1575-1581.

S. L. Neidleman, j. Geigert, Trends Biotechnol. 1983, 1, 1-5. 60 j. Kollonitsch, S . Marburg, L. M. Perkins, ]. Am.Chem. SOC.1970,92,4489-4490. 61 j. Geigert, S. L. Neidleman, D. j. Dalietos,]. Biol. Chem. 1983,258,2273-2277. 62 M. C. R. Franssen, J. D. jansma, H. C. van der Plas, E. de Boer, R. Wever, Bioorg. Chem. 1988,16,352-363. 63 R. F. Theiler, J. F. Siuda, L. P. Hager, Science 1978,202,1094-1096. 64 R. J. Selvaraj, J. M. Zgliczynski, B. B. Paul, A. J. Sbarra,]. Infect. Dis. 1978, 137, 481-485. 65 M. B. Grisham, M. M. Jefferson, D. F. Metton, E. L. Thomas,]. Biol. Chem. 1984,259, 10 404-10413. 66 K.-H. van Pee, Annu. Rev. Microbiol. 1996, 50,375-399. 67 B. Walter, K. Ballschmitter, Chemosphere 1991,22, 557-567. 68 F. S . Brown, L. P. Hager,J. Am. Chem. SOC. 1967,89,719-720. 69 N. Itoh, A. K. M. Q. Hasan, Y. Izumi, H. Yamada, Eur.]. Biochem. 1988, 172, 477-484. 70 K. Ramakrishnan, M. E. Oppenhuizen, S. Saunders, j. Fisher, Biochemistry 1983,22, 3271-3277. 71 K. K.C. Liu, C.-H. Wong,]. Org. Chem. 1992,40, 3748-3750. 72 R. E. Huber, L. A. Edwards, T. j. Carne, J . Biol. Chem. 1989,264,1381-1386. 73 B. Matkovics, 2. Rakonczay, S. E. Rajki, L. Balaspiri, Steroidologia 1971, 2, 77-79. 74 H. Vilter, Lejol. Bot. Mar. 1983, 26, 429-435. 75 H. Yamada, N. Itoh, S. Murakami, Y. Izumi, Agnc. Biol. Chem. 1985,49, 2961-2967. 76 S. L. Neidleman, S. D. Levine, Tetrahedron Lett. 1968, 46, 4057-4059. 77 S. L. Neidleman, M. A. Oberc,]. Bacteriol. 1968,95,2424-2425. 78 S. D. Levine, S. L. Neidleman, M. Oberc, Tetrahedron 1968,24, 2979-2984. 79 R. P. Martyn, S. C. Branzer, G. T. Sperl, Bios 1981,52,8-12. 80 S. L. Neidleman, P. A. Diassi, B. Junta, R. M. Palmere, S. C. Pan, Tetrahedron Lett. 1966,44,5337-5342. 81 M. C. R. Franssen, H. C. van der Plas, Bioorg. Chem. 1987,15, 59-70. 82 J. M. Boeynaems, D. Reagan, W. C. Hubbard, Lipids 1981, IG, 246-249. 59

1280

I

IG Oxidation Reactions W. C. Hubbard, Biochim. Biophys. Acta 1983,

83 J. M. Boeynaems, J. T. Watson, J. A. Oates,

W. C. Hubbard, Lipids 1981,16, 323-

327. 84 J. Turk, W. R. Henderson, S. L. Klebanoff,

85

751,189-200. M. Nieder, L. P. Hager, Arch. Biochem. Biophys. 1985,240,121-127.

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyriqht 0Wiley-VCH Verlaq GmbH, Weinheim 2002

I

lZ8’

17 lsomerizations Nobuyoshi Esaki, T. Kurihara and K. Soda

17.1

Introduction

Isomerases catalyze the isomerization of substrates, and are classified into five groups as follows: Racemases and epimerases (E. C. class 5.1) They are defined as enzymes that catalyze the isomerization of a substrate through stereochemical reverse rearrangement of a substituent bound to a chiral center (usually a chiral carbon) in the substrate molecule. Racemases act on molecules containing only the asymmetric center concerned in the reaction. Epimerases act on substrates containing one or more asymmetric centers in addition to the reactive chiral center. cis-trans-Isomerases (E. C. class 5.2) They catalyze the interconversion of cis-trans geometrical isomers. Sugar isomerases, tautomerases, A-isomerases, etc. (E. C. class 5.3) Sugar isomerases catalyze the interconversion between aldose and ketose. Tautomerases catalyze a keto-enol tautomerization. A-Isomerases catalyze the shift of a double bond. The reactions catalyzed by these enzymes proceed through intramolecular oxidation and reduction. Mutases (E. C. class 5.4) They catalyze the transfer of a substituent to produce a structural isomer. Cycloisornerases (E. C. class 5.5) They catalyze the ring formation through an intramolecular lyase reaction.

Isomerizations catalyzed by most of these enzymes proceed through 1,1-,1,2-, or 1,3-hydrogen shifts (Table 17-1), while mutases catalyze exchange of a hydrogen

1282

I

77 lsomerizations Table 17-1. Type

1,l-Shifts

Enzyme-catalyzedisornerizations classified as hydrogen shifts. Examples Rl

R,+H R,

ir

F=O

1,2-Shifts

H-C-OH

I

__

Category Rl

H+R*

Epimerases, Racemases

R3

H I

H-:-OH C=O

Aldose-ketoseisomerases

I

1,3-Shifts

atom with particular functional groups such as amino, hydroxy, and a-amino-acarboxymethyl groups attached at neighboring carbon atoms of the substrates through homolytic cleavage. Here we describe enzymological properties of representative racemases, epimerases, and isomerases, and their application to production of various optically active compounds.

17.2

Racernizations and Epirnerizations

Since the discovery of enzymatic racemization of lactate by lactic acid bacteria '1, Clostridium acetobutyri~um[~], and CE. butyri~um1~1, a variety of racemases and epimerases have been demonstrated, and they are classified into the four groups as follows: Amino acid racemases and epimerases catalyzing racemization and epimerization at the chiral center containing an NH2 or NH group (E. C. class 5.1.1); Mandelate racemase, lactate racemase, and others acting at the chiral center containing an OH group (E.C. class 5.1.2); Various carbohydrate epimerases such as UDP-~-glucose-4'-epimerase (E. C. class 5.1.3); Methylmalonyl CoA epimerase and some others, in whose substrates a CH3 group is bound to the chiral centers (E. C. class 5.1.99). Racemases and epimerases have been used for production of various optically active compounds from cheaply-availableracemic substrates by combination of enzymes that act specifically on one of the isomers of the racemates to catalyze hydrolysis, oxidation, reduction, elimination, replacement, and other reactions. The racemases and epimerases used act exclusively on the substrates, but not on the products of the

77.2 Racemizations and Epimerizations

reaction. Thus, total conversion of the racemic substrates into the desired opticallyactive compounds is achieved. Here we describe enzymological characteristics of the representative racemases and epimerases, and their application to production of optically active compounds. 17.2.1

Pyridoxal 5'-phosphate-dependentAmino Acid Racemases and Epimerases 17.2.1.1

Alanine Racemase (E.C. 5.1.1.1)

Alanine racemase is a bacterial enzyme that catalyzes racemization of L- and Dalanine, and requires pyridoxal 5'-phosphate (PLP) as a cofactor. The enzyme plays an important role in the bacterial growth by providing D-alanine,a central molecule in the peptidoglycan assembly and cross-linking,and has been purified from various The enzyme has been used for the production of stereospecifically deuterated NADH and various D-amino acids by combination of L-alanine dehydrogenase (E. C. 1.4.1.1),D-amino acid aminotransferase (E. C. 2.6.1.21), and formate dehydrogenase (E. C. 1.2.1.2) [I7, "1.

17.2.1.1.1

Gene Cloning and Primary Structure

Two distinct alanine racemase genes were cloned from the Salmonella typhimurium chromosome. One mapped at minute 37 on the chromosome is termed dadB, and the other mapped at minute 91 is termed the alr The dadB alanine racemase is formed inducibly and functions in the catabolism of L-alanine: the alr enzyme is synthesized constitutively, and functions in the anabolic assembly of peptidoglycan Alanine racemase genes were also cloned from Bacillus stearothemophilus['OI, Bacillus subtilis [''I, Bacillus p~ychrosaccharolyticus['~~, and Aquijkx pyrophiUS['^]. Two distinct alanine racemase genes were assigned in the genome sequences of Escherichia coli["], B. subtilis, Pseudomonas aeruginosa, and Vibriocholerae, but only a single one occurs in the other bacterial genomes whose complete nucleotide sequences were determined as shown at internet sites such as http://www.geno me.ad.jp/kegg/catalog/org-list.htm1. Uo et al. [16]have found that fission yeast, Schtzosaccharomyces pombe, has also the alanine racemase gene, which is involved in the catabolism of D-alanine in S. pombe in the same manner as dadB of S. typhimurium. The yeast enzyme only shows any high degree of similarityto the alanine racemases of y-proteobacteria(gram-negative phylum). Therefore, the gene of S. pombe has possibly been acquired from yproteobacteria through some events of horizontal gene transfer such as conjugation: S. pombe is known to be a recipient of the genes from E. coli through direct conjugation. D-Alanine occurs in various natural compounds produced by fungus. For example, cyclosporin A contains D-alanine as a component and is produced by Tolypocladium niveum[121. Alanine racemase is involved in the biosynthesis of D-alanine in this fungus.

1284

I

I7 lsomerizations 17.2.1.1.2

Stability

The native dadB and alr racemases from Salmonella typhimurium are readily inactivated by digestion with a-chymotrypsin, trypsin, and subtilisin [221. However, the Bacillus stearothermophilus enzyme is stable even after fragmentation into two pieces [232 241. A. pyrophilus, a hyperthermophilic bacterium, produces extremely stable alanine racemase ["I. It maintains catalytic activity in the presence of organic solvents as well. On the other hand, Bacillus psychrosaccharolyticus, a psychrophyilic bacterium, produces a thermo-labile enzyme[14].However, it shows high catal9c activity at low temperatures, such as at 0 ° C . Similar cold activity and thermal instability was found in the enzyme from a psychrophile isolated from raw milk, Pseudomonasfluorescens I"].

17.2.1.1.3

Reaction Mechanism

Reaction of alanine racemase proceeds through the steps shown in Fig. 17-1. PLP bound with the active-sitelysyl residue (A) reacts with a substrate to form an external Schiff base (B) through transaldimination. The subsequent a-hydrogen abstraction

Figure 17-1. Mechanism o f the alanine racemase reaction. A, An internal aldirnine o f PLP with a lysyl residue; 6, an external aldimine o f PLP with o-alanine; C, a quinonoid intermediate formed after removal o f a hydrogen from alanyl external aldimines B or D; D, an external aldimine o f PLP with L-alanine. Reprinted from Watanabe et al.'"'.

17.2 Racemizations and Epimerizations

results in the formation of a resonance-stable deprotonated intermediate (C). If reprotonation occurs at the a-carbon of the substrate moiety on the opposite face of the planar intermediate (C), then an antipodal aldimine (D) is formed. The &-amino group of the lysine residue is substituted for the isomerized amino acid through transaldimination, and the internal aldimine (A) is regenerated. According to D ~ n a t h a n [ ~the * ] , Ca-H - bond to be broken is positioned perpendicularly to the plane of the conjugated x-system of the external Schiff base intermediate, in order to achieve maximum orbital overlap with the 7c electron system of the complex, resulting in a substantial rate enhancement for the cleavage of that bond. The racemization reaction proceeds via either a one-base[2G]or tw~-base[~'] mechanism. The one base mechanism is characterized by the retention of the substrate-derived proton in the product (internal return) [2G1. By this criterion, reactions catalyzed by a-amino-&-caprolactamracemase 12*] and amino acid racemase with low substrate specificity (E. C. 5.1.1.10) L2]' have been considered to proceed through the one-base mechanism. However, such internal returns were not observed Bacillus stearin the reactions catalyzed by alanine racemases from E. c0li[~~1], and Salmonella typhimurium (dadB and alr) [291. The internal return otherm~philus[~'], is not expected to occur in the reactions of two-base mechanism. In fact, kinetic analyses 1301indicated that the alanine racemase reaction proceeds through a twobase mechanism: proton donors and proton acceptors are situated on both sides of the planar intermediate (Fig. 17-1, C) and accomplish removal and return of the ahydrogen of the substrate amino acid. X-ray crystallographic studies [31* 321 suggested that Lys 39 and Tyr 265 of alanine racemase from B. stearothemophilus serve as the bases (Fig. 17-2).Watanabe et al. [331 showed that the lysyl residue binding PLP in the racemase (Lys 39) acts as the base catalyst specific to the D-enantiomer of alanine. The crystal structure of the enzyme complex with R-1-aminoethylphosphonic acid[32],a tight-bind inhibitor of the enzyme[341,demonstrated that the phenolic oxygen of Tyr 265 is appropriatelyaligned for proton abstraction from an L-isomer in the active site of the structure: Tyr 265 is the second base specifically acting on the Lalanyl-PLP aldimine.

Tyr 265'

Lys 39

Tyr 265'

Lys 39

Figure 17-2. Stereodiagram o f the aldimine formed from l-aminoethylphosphonate and PLP viewed perpendicular to the plane o f the PLP ring. The catalytic residues Tyr 265' and Lys 39 are shown. Reprinted from Stamper et aI.'32J.

1286

I

7 7 lsornerizations

A D-[tx-'H] Ala D/L-Ala AlaDH L-[a-'H] Ala

[u-'H]-L-Aia

@roR)

7[4R-2H]NAD2H+ Pyruvate + NH3

[C4-' HINAD' Oxidized Form of Substrate

[2H] Reduced Form of Substrate

Oxidized Form of Substrate

['HI Reduced Form of Substrate

Figure 17-3. A, Preparation of [4S-'H]-NADH by coupling of alanine racernase and L-alanine dehydrogenase. B, In situ determination o f stereospecificity of H-transfer by 'H-NMR. AlaR, AlaDH, and D H represent alanine racernase, L-alanine dehydrogenase, and dehydrogenase, respectively.

17.2.1.1.4

Production of Stereospecifically Deuterated NADH

NAD-linked dehydrogenases show either pro-S or pro-R-stereospecificityfor hydrogen removal from the C4 position of the nicotinamide moiety of the reduced coenzymes. The stereospecificity of hydrogen transfer is examined by means of stereospecifically C4-deuterated NADH, which is prepared enzymatically from NAD' and deuterated substrates by tedious procedures. Esaki et al. developed a simple method to produce the stereospecificallydeuterated NADH by an NAD'-dependent dehydrogenase by combination with amino acid racemase 13'1. L-Alanine dehydrogenase transfers deuterium of [2-'H]-~-alanineto NAD' to produce [4R-2H]-NADH[3G1.Alanine racemase catalyzes the C2-deuteration of D and L-alanine in 2H20110], and [4R-2H]-NADH was produced from D-alanine and NAD' by coupling of the reactions catalyzed by alanine racemase and r-alanine dehydrogenase in 2H20 (Fig. 17-3A).Furthermore, this finding led to development of a simple procedure for the in situ analysis of stereospecificity of hydrogen transfer of NADH by an NAD-dependent dehydrogenase by means of 'H-NMR (Fig. 173B)C3'1.

17.2.1.1.5

Production of D-Amino Acids

Considerable attention has been paid to multi-enzyme reaction systems as a means to the stereospecific production of L-amino Wichmann et al. have developed a continuously operated membrane reactor for production of L-leucine from aketoisocaproate13'1. The system is also applicable to the production of several other aliphatic L-amino acids such as L-valine, 1-tert-leucine and ['SN]-~-leucine.The

yDH xATA 17.2 Racemizations and Epimerizations

Formatex

NAD’

T====== D-Alanine

L-Alanine aR :

Keto acid

FDH

COP

NADH +H+

\ Pyruvate

D-Amino acid

NH3

Figure 17-4. Synthesis o f o-amino acids from a-keto acid, formate, NAD+, D-alanine, and ammonia by coupling of L-alanine dehydrogenase (AlaDH), formate dehydrogenase (FDH), alanine racemase (AlaR), and D-amino acid aminotransferase (D-ATA).

process has been successfully scaled-up for industrial production of these L-amino acids [391. A similar system has been developed for production of L-phenylala411 and ~-P-chloroalanine[~~~~1. However, little attention has been paid to the stereospecific production of D-amino acids by means of multi-enzyme reaction systems, although D-amino acids have been paid considerable attention [&I. For example, substantial amounts of D-serine,D-aspartateand other D-amino acids occur in mammalian l ~ r a i n [ ~ ’ -and ~ ~ ]13N-labeled , D-amino acids are expected to be useful for the study of their metabolism in brain[481. A simple procedure was established for the synthesis of various D-amino acids by means of four types of thermostable enzymes: alanine racemase, D-amino acid aminotransferase [49* ”1, L-alanine dehydrogenase [’I, and formate dehydrogenase (Fig. 17-4)[171. The commercial preparation of formate dehydrogenase from Cundida boidinii used by Wichmanri et a1.19 is not sufficiently stable. However, Galkin et al.[52]cloned and expressed the gene of thermostable formate dehydrogenase in E. coli. D-Phenylalanine and D-tyrosine, which are the poor substrates for D-amino acid aminotransferase, were synthesized in an optical purity of essentially 100 %, but with yields of lower than 50 %. However, the yields were increased by addition of excess amounts of the D-amino acid aminotransferase (Table 17-2)[171. Selenium is an essential micronutrient for mammals, fish and several bacteria, although it is toxic at 531. D-Selenomethionine was produced in an 80% yield a high c~ncentration[’~, based on 2-0~0-4-methylselenobutyrate[’~]. Norvaline, valine, and a-aminobutyrate were also produced with high yields. However, a-aminobutyrate was synthesized as a racemic mixture. D-Norvaline was obtained at an enantiomeric excess of only 30%. The low optical purity is probably due to the action of L-alanine dehydrogenase: aketobutyrate and a-ketovalerate are reduced by t-alanine dehydrogenase at rates of 79 and 6.6% relative to that of pyruvate, respectively[”]. Moreover, alanine racemase also racemizes a-aminobutyrate and norvaline, though very sl0wly[~~1. Thus, this method is not applicable to the stereospecificproduction of D-a-aminobutyrateand D-norvaline. The preparations of D-valine, D-methionine and D-norleucine also suffered contamination by the antipodes at concentrations of 4, 3, and 1%, respectively, due to the action of L-alanine dehydrogenase on the a-keto analogs of these amino acids [’ll. However, D-glutamate, D-phenylalanine and D-tyrosine were efficiently produced in the system. The final concentration of D-glutamateproduced

1288

I

7 7 lsornerizations Table 17-2.

Synthesis of o-amino acids from a-keto acids by combination offour purified enzymes: alanine racemase, L-alanine dehydrogenase, formate dehydrogenase, and o-amino acid aminotransferase.

Substrate

Product

Yield (“h)”

ee (“A)

a-Ketoglutarate

D-glutarnate D-leucine D-norleucine D-rnethionine D-valine n-norvaline a-aminobutyrate D-phenylalanine D-tyrosine

98 80 82 95 90 92 93 72b 70b

100 >99 98 94 92 30 0 100 100

a-Ketoisocaproate a-Ketocaproate a-Keto-y-thiornethylbutyrate a-Ketoisovalerate a-Ketovalerate a-Ketobutyrate Phenylpymvate Hydroxyphenylpyruvate

a The yields were determined after an 8 h

incubation.

b The amount of 0-amino acid aminotransferase used (30 units) was 10-foldhigher than that in other systems

was only around 0.3 M, limited because of the equilibrium of the D-amino acid aminotransferase reaction. The method is most suitable for stereospecific conversion of a-keto acids into the corresponding D-amino acids, in particular labeled compounds, for example with 13N by means of 13N-NH3. The industrial use of the above-mentionedsystems depends predominantly on the cost of the enzymes, although the intact cells of microorganisms containing the enzymes can be used as catalysts in order to decrease costs[56].In most cases, however, additional genetic improvements through metabolic engineering are required, thereby new functional combinations are made by the rational transfer of pathways from one organism to another[”]. The transfer of the ethanol pathway from Zymomonas mobilis to other enteric bacteria represents an example of this approach[58].In the above-mentioned system, various D-amino acids can be produced from the corresponding a-keto acids, if four functional genes are introduced into one microorganism. The simultaneous expression of all enzymes in a single cell Synthesis of D-amino acids from a-keto acids by E. coli cells harboring pFADA which codes for four enzyme genes: alanine racemase, L-alanine dehydrogenase, formate dehydrogenase, and D-amino acid aminotransferase.

Table 17-3.

Substrate

Product

Yield (“A)”

ee (“A)

a-Ketoglutarate a-Ketoisocaproate a-Ketocaproate a-Keto-y-thiornethylbutyrate a-Ketoisovalerate a-Ketovalerate a-Ketobutyrate Phenylpymvate H ydroxyphenylpyruvate

D-glutamate o-leucine D-norleucine D-methionine D-valine D-norvaline a-arninobutyrate D-phenylalanine D-tyrosine

85 76 70 80 85 90 95 15 5

100 >99b 88 90 92 35 0 ND‘ ND

a The yields were determined after a 12 h

incubation

b The optical purity determined by HPLC is >99.9% c ND, not determined.

17.2 Racernizations and Epirnerizations

4

Hindlll BamHl

plac

ptac

sm

ATG

FDH

T G A ~ S D ATG ~

AlaDH

T G A S~ D ATG ~

HSD~ I

Pstl I Nsil

Bgfll

Sphl

DAAT

TAA

ATG

AlaR T A A ~ ~ F A D A

Figure 17-5. Construction of the plasmid used for the production o f o-amino acids by expression in E. coli cells; formate dehydrogenase (FDH), L-alanine dehydrogenase (AlaDH), alanine

racernase (AlaR), and D-amino acid aminotransferase (DAAT).

- IE

g 0,2

Ec

C

0

0

f _ 1

-

0,l-

D-Glutamate

I a-Ketoglutarate

I

EcoRl

Figure 17-6. Time course for the production o f D-glutarnate with E. coli cells containing pFADA. a-Ketoglutarate was added after 4 and 10 h o f incubation at

provides additional benefit for industrial applications:the intracellular pool of NAD’ (supplied by the cell itself) could be used for NADH regeneration without any additional supplies. Galkin et al. I‘[ constructed plasmids containing, in addition to the thermostable formate dehydrogenase gene, all three genes required for the synthesis of D-amino acids (Fig. 17-5). D-Enantiomers of glutamate and leucine were produced at high optical purity and high conversion rates with the recombinant E. coli cells harboring the plasmid for coding of the four heterologous genes (Table 17-3). a-Keto acids, particularly branched-chain and long-chain a-keto acids, are toxic, inhibiting the growth of E. coli when added at concentrations of only 15-30 mM. Therefore, Galkin et al. used the resting cells of the recombinant E. coli instead of growing ones. Moreover, the isolation of products in the resting-cell system is much easier than when using growth media containing complex ingredients such as yeast extracts. The final concentration of D-glutamateproduced was around 0.3 M (Fig. 17-6). 17.2.1.2

Amino Acid Racemase with Low Substrate Specificity (E.C. 5.1.1.10)

An amino acid racemase which shows very broad substrate specificity was discovered in Pseudoinonas striata (= Ps. putida), purified, and characterized[”1. The enzyme catalyzes racemization of various amino acids except aromatic and acidic

1290

I

7 7 lsomerizations

amino acids. A similar enzyme also occurs in Aerornonas punctata[60].Arginine racemase, which also shows a broad substrate specificity, has been demonstrated in Pseudomonas graveolens (= Pseudornonas taetrolens) ['*I. These amino acid racemases do not act on threonine, valine and their analogs, whose P-methylene group is substituted. Recently, Lim et al.L621 found, in Ps. putida ATCC 17642, a new amino acid racemase catalyzing not only racemization of various amino acids but also epimerization of D- and L-threonineby stereoconversionat the a-position: it catalyzes epimerization of L-to D-allo-and also of D- to L-allo-threonine. Amino acid racemase with low substrate specificity catalyzes racemization of leucine and various other amino acids, which are also a-deuterated in 2 H z 0during their ra~emization[~']. Therefore, [4S2H]-NADHwas produced in the same manner as described above with the racemase and L-leucine dehydrogenase (E. C. 1.4.1.9), which is pro$ Amino acid racemase with low substrate specificityof Ps. putida ATCC 17642 does not racemize aromatic and acidic amino acids. However, phenylalanine and phenylglycine undergo a-hydrogen exchange with deuterium from the solvent when incubated with the racemase in 2H20. Lim et al. [641 found that each enantiomer of both a-deuterated phenylalanine and phenylglycine are produced stereospecifically with retention of the C2 configuration. This a-hydrogen exchange reaction is applicable to the production of a-deuterated phenylalanine and phenylglycine. Makiguchi and coworkers established a method to synthesize L-tryptophan from D,L-serine and indole by means of tryptophan synthase (E. C. 4.2.1.20) from E. coli and the amino acid racemase with low substrate specificity of Ps. striata (= Ps. putida) [651. Both D,L-serine and indole are cheaply available by chemical synthesis. Tryptophan synthase catalyzes the p-replacement reaction of L-serine with indole to produce L-tryptophan, and the amino acid racemase with low substrate specificity converts unreacted D-serine into L-serine. Because the racemase does not act on tryptophan, almost all D,L-serine is converted into optically pure L-tryptophan. Makiguchi et al.["] succeeded in producing L-tryptophan in a 200 L reactor using intact cells of E. coli and Ps. put id^[^^]. Under the optimal conditions established, 110 g L-l of L-tryptophan was formed in molar yields of 91 and 100% for added D,Lserine and indole, respectively, after 24 h of incubation with intermittent indole feeding. Continuous production of L-tryptophan was also achieved using immobilized cells of E. coli and ps. putida. The maximum concentration of L-tryptophan formed was 5.2 g L-' (99% molar yield for indole). S-Adenosyl-L-methionine is the important methyl donor in biological transmethylation to form S-adenosyl-L-homocysteine,which is hydrolyzed to adenosine and homocysteine by S-adenosyl-L-homocysteinehydrolase (E. C. 3.3.1.1) in vivo. However, equilibrium of the S-adenosyl-L-homocysteinehydrolase reaction favors the direction toward synthesis of S-adenosyl-L-homocysteine.Shimizu et al. developed a simple and efficient method for the high yield preparation of S-adenosyl-Lhomocysteine with S-adenosyl-L-homocysteinehydrolase of Alcaligenes faecalis, in which the cellular content of S-adenosyl-L-homocysteinehydrolase was about 2.5 % of the total soluble protein. S-Adenosyl-L-homocysteinewas produced at a concentraHowever, when racemic tion of about 80 g L-' with a yield of nearly

17.2 Racernizations and Epimerizations

I

1291

H

HO

1

OH OH

2

OH OH

OH

7

OH OH

8

OH OH

YH2

9

OH OH

Structures of adenosine and related nucleosides which serve as substrates for S-adenosyl-L-homocysteinehydrolase. 1, Adenosine; 2, formycin A 3, neburalin; 4, adenosine "-oxide; 5 , 2-chloroadenosine; 6, tubercidine; 7, N6methyladenosine; 8, inosine; 9, 1-methyladenosine. Figure 17-7.

homocysteine was used, the D-enantiomerremained unreacted. When Ps. striata (= Ps. putida) cells were used as the catalyst, D-homocysteine was converted into Sadenosyl-r-homocysteine:the amino acid racemase with low substrate specificity acts on homocysteine, but not on S-adenosylhomocysteine.A. faecalis is better than Ps. striata in showing higher S-adenosyl-r-homocysteinehydrolase and lower adenosine deaminase activities than those of Ps. striata. Therefore, a mixture of both bacterial cells was used to produce 70 g L-l of S-adenosyl-r-homocysteinefrom D,Lhomocysteine and adenosine with a molar yield of nearly 100%~G6]. S-Adenosyl-Lhomocysteine hydrolase acts on various adenosine analogs, and the corresponding S-nucleotidyl-L-homocysteines(Fig. 17-7) were synthesized from the analogs and D,L-homocysteineby means of both bacterial cells LG7].

1292

I

7 7 lsornerizations

17.2.1.3 a-Amino-E-caprolactamRacemase

a-Amino-E-caprolactam (ACL) is a chiral heterocyclic compound synthesized from cyclohexene, which is a by-product in the industrial production of nylon. Fukumura168-701established an enzymatic method to produce L-lysine from D,L-ACL. The process is composed of two enzyme reactions: the selective hydrolysis of L-ACL to Llysine, and the racemization of ACL (Fig. 17-8).The L-ACL-hydrolyzing enzyme (aamino-e-caprolactam hydrolase (E. C. class 3.5.2) is distributed in the cells of Cryptococcus laurentii and other yeasts (68-701, and its synthesis is induced by D,L-ACL. The enzyme purified to homogeneity from a cell extract of C. laurentii has a molecular weight of about 185000, and is activated by MnC12 and MgC12[71].L-ACL is the only substrate of the hydrolase: D-ACL and e-caprolactam are not hydrolyzed. ACL racemase has been found in the cells of Achromobacter obae and other bacterial7*],and is a unique enzyme among racemases in acting exclusively on cyclic amides derived from a,o-diamino acids. Ahmed et al.[731purified the enzyme to homogeneity from the cell extract of A. obae, and characterized it. The enzyme is composed of a single polypeptide chain whose molecular weight is about 50 000, and contains 1 mol of PLP per mol of enzyme as a coenzyme. In addition to both isomers of ACL, D- and L-a-amino-6-valerolactamalso serve as effective substrates [741. The enzyme catalyzes the exchange of the a-hydrogen of the substrate with deuterium or tritium during racemization in deuterium oxide or tritium By tritiumincorporation experiments, the enzyme was shown to catalyze both inversion and retention of configuration of the substrate with a similar probability in each hrnover. When [a-2H]-~-ACL and unlabeled D-ACLwere converted into the L-isomer by ACL racemase in water and in deuterium oxide, respectively, in the presence of excess LACL hydrolase, a-hydrogen (or a-deuterium) was retained significantly in the product[28].Therefore, a single base mechanism has been proposed for the racemization catalyzed by ACL racemase. The ACL racemase gene has been cloned from the chromosomal DNA of A. obae, and its complete nucleotide sequence determined, which revealed that the enzyme consists of 435 amino acids and that its molecular weight is 45 568[751. H

:"4

""NH;!

-

ACL Racemase

D-ACL

I

L-ACL A C L Hydrolase

H

z

N

~

L-Lysine

c NHZ

o

o Figure H 17-8. Total conversion of racemic ACL into L-lysine by coupling of ACL racemase and ACL hydrolase reactions.

7 7.2 Racemizations and Epirnerizations 11293

17.2.2 Cofactor-independent Racemases and Epimerases Acting on Amino Acids 17.2.2.1 Glutamate Racemase (E.C. 5.1.1.3)

D-Glutamateas well as D-alanineis an important component of the peptidoglycan of bacterial cell walls [761, and is produced by glutamate r a ~ e m a s e c1'.~ ~ ,Lactic acid bacteria show high activity of the enzyme[7'], and glutamate racemase was first purified from Pediococcuspentosaceus 811.

17.2.2.1.1 Gene Cloning

Nakajima et al. cloned the glutamate racemase gene of Pediococcus pentosaceus["! Glutamate racemase genes have been also cloned from various other sources: Lactobacillus fermenti 831, Lactobacillus bre~is['~],E. ~oli['~], Bacillus purnilu~[~~1, Aquijix pyrophilu~[~~I, and Bacillus subtilis[88*891.

17.2.2.1.2 Enzymological Properties

The glutamate racemase gene from Pediococcuspentosaceus was over-expressedin the recombinant cells, but formed an inclusion body[81]. However, the enzyme was solubilized with G M urea, renatured by dialysis to remove urea, and purified to homogeneity with a high overall yield["]. The amount of enzyme produced by the clone cells corresponded to about 38 % of the total insoluble proteins. However, the glutamate racemase gene was solubilized in vivo in an active form when it was coexpressed with the gene of chaperonin GroESL["]. Choi et al. isolated the active enzyme and purified it The enzyme is composed of a subunit with a molecular mass of about 29 kDa. The enzyme acts specifically on glutamate with KM values of 14 and 10 mM for D- and L-glutamates,respectively. None of other amino acids occurring in proteins including aspartate, asparagine, and glutamine are racemized. Other glutamate analogs (homocysteate, a-aminoadipate, glutamate ymethyl ester, N-acetylglutamate,a-hydroxyglutarate, and cysteine sulfinate) are also inert. However, L-homocysteine sulfinate, a y-sulfinate analogue of glutamate, is racemized at a rate of about 10% of that of L-glutamate. Amino acid racemases generally require PLP as a cofactor, but glutamate racemase is dependent on neither PLP nor on any other cofactor[80,'l]. Proline racemase (E. C. 5.1.1.4) [921, diaminopimelate epimerase (E. C. 5.1.1.7) and hydroxyproline epimerase (E. C. 5.1.1.8) also require no coenzyme. Glutamate racemase from E. coli is unique because it is activated about 100 fold in the presence of UDP-N-acetylmuramoyl-L-alanine (UDP-MurNAc-L-Ala),the precursor of peptidogly~an~~~]. UDP-MurNAc-L-Alais ligated to D-glutamate,a product of the glutamate racemase reaction, by the catalysis of UDP-N-acetylmuramoy1-Lalanyl-D-glutamate synthetase (E. C. 6.3.2.9). Thus, the activation of the E. coli glutamate racemase by UDP-MurNAc-L-Ala has a physiological importance in the

1294

I

I 7 lsornerizations

regulation of peptidoglycan biosynthesis[951. In contrast, glutamate racemases of Gram-positive bacteria such as Lactobacillus fernenti, Lactobacillus breuis, Bacillus pumilus are not activated by UDP-MurNAc-L-Ala,though these enzymes show about 30 % sequence similarities to the E. coli enzyme. The predominant difference between the E. coli enzyme and the glutamate racemases of the Gram-positive bacteria is that the former has a 21-amino acid extension at the N-terminus as compared with the latter enzymes: the N-terminal region is responsible for the activationL9'1. Glutamate racemase produced in cell extracts of Bacillus subtilis, an abundant producer of poly-y-glutamate,is a monomer with a molecular mass of about 30 kDa containing no cofactor[881.It almost exclusively catalyzes racemization of glutamate and is mainly concerned in D-glutamate synthesis for poly-y-glutamate production. B. subtilis produces another isozyme of glutamate racemase encoded by the YrpC Ashiuchi et al. cloned both enzyme genes and compared their enzymological properties [88, 891. Enzymological properties of YrpC, such as the substrate specificity and optimum pH, are similar to those of the other glutamate racemase (Glr).The thermostability of YrpC, however, is considerably lower than that of Glr. In addition, YrpC shows higher affinity and lower catalytic efficiency for L-glutamate than Glr [891.

17.2.2.1.3 Structure and Mechanism

Glutamate racemase contains one essential cysteine residue per mol of enzyme, whose chemical modification results in complete inactivation c9l1. Choi et al. determined the amount of tritium incorporated into the substrate and product enantiomer during incubation with the enzyme in tritium water, and found that tritium is exclusively incorporated into the product enantiomer regardless of the configuration of the substrate used["l. This is compatible with a model in which two different bases participate in abstraction and return of a-hydrogen of the substrate. One of the two bases involved in catalysis is suggested to be the essential cysteine residue: a thiolate from one of the cysteines abstracts the a-proton, and the other cysteine thiol delivers a proton to the opposite face of the resulting carbanionic intermediate["]. Kim et al.[871cloned the glutamate racemase gene from Aqufifex pyrophilus, a hyperthermophilic bacterium, and expressed it in E. coli. The enzyme shows strong thermostability in the presence of phosphate ion, and it retains more than half of its original activity after incubation at 85 "C for 90 min. Hwang et al.["] crystallized the glutamate racemase of A. pyrophilus and determined the tertiary structure of the enzyme by X-ray crystallography.The enzyme is composed of two identical subunits, and each monomer consists of two alp fold domains. Hwang et al. has also proposed a mechanism in which two cysteine residues are involved in the catalysis (Fig. 179) [9G].

Glavas and Tanner replaced the two cysteine residues, Cys 73 and Cys 184, by serine, and analyzed the reactions catalyzed by the mutant enzymes: the elimination of water from a substrate analog, N-hydroxyglutamate,through a one-base requiring reaction[97].The C73S mutant was a much poorer catalyst than the wild-typeenzyme

17.2 Racemizations and Epimerizations

I

1295

fyHiS I8O

Ser 8 - 0 H H

H

o F G I u 147

HO CYS70-S-

H-S-Cys D-Glu

-

Ser 8-0 I H

180

H

HO

L

178

(""' (YHiS

Cys 70-S-H

\

L-Glu

I

\\

OA0-

0

i Asp73 :

Cys70-SH

o F G l ~ 147 HO HS-CYS 178

Figure 17-9. Mechanism o f glutamate racemase reaction. Cys 70 and Cysl78 serve as the bases t o abstract an a-proton from the substrate, and a carbanion intermediate is formed. Alternatively, the racemization may proceed through a concerted mechanism. Reprinted from Hwang et a1.(95].

toward D-N-hydroxyglutamate, whereas the C184S mutant was better than the wildtype. When L-N-hydroxyglutamate was used as a substrate, C73S was better but C184S was poorer than the wild-type.Thus, Glavas and Tanner concluded that Cys73 is responsible for the deprotonation of D-glutamate and Cys 184 is responsible for the deprotonation of L-glutamate

17.2.2.1.4

Synthesis of D-AminoAcids with Clutamate Racemase

Nakajima et al. [981 have developed an efficient method for the synthesis of various Damino acids from the corresponding a-keto acids and ammonia by coupling of four enzyme reactions catalyzed by D-amino acid aminotransferase ["I, glutamate racernase 17'), 911, glutamate dehydrogenase and formate dehydrogenase (Fig. 17-10). Various D-amino acids are produced by this method. Under the optimum conditions established by Nakajima et al. ("1, D-enantiomersof valine, alanine, a-aminobutyrate,

1296

I

1

I 7 lsomerizations

Formate

NAD+ xuL~~utamate

GluR

======= D-GlutamateXA;;Keto

acid

FDH

co2

NADH

+ H+

a-Ketoglutarate + NH3

-

D-Amino acid

Figure 17-10. Enzymatic synthesis o f D-amino acids by combination o f glutamate racemase, glutamate dehydrogenase, D-amino acid aminotransferase and formate dehydrogenase reactions.

Table 17-4. Production ofvarious D-amino acids by means o f f o u r purified enzymes: glutamate racemase, D-amino acid aminotransferase, glutamate dehydrogenase, and formate dehydrogenase". DAmino acids

D-Valine D-Alanine o-a-Aminobutyrate D- Aspartate u-Leucine D-Methionine D-Serine D-Histidine D-Phenylalanine D-Tyrosine a

Molar yield ("9)

100 100

100 100 84

80 50

36 28 13

Reprinted from N. Nakajima et al."981.

leucine, methionine and aspartate are synthesized from their a-keto analogs with a molar yield higher than 80% under the conditions used (Table 17-4)1981.D-Histidine and a few other D-amino acids, which are poor substrates of D-amino acid aminotransferaseI"1, are produced in a yield lower than 40% under the same conditions. However, Bae et al.I1OOlestablished an efficient method for production of Dphenylalanine and D-tyrosine by feeding a-keto acid intermittently in order to keep its concentration at less than 50 mM, above which the productivity decreased greatly (Fig. 17-11). By running the multi-enzyme system for 35 h, 48 g L-' of D-phenylalanine and 60 g L-' of D-tyrosine were produced with 100% of optical purity from the equimolar amounts of phenylpyruvate and hydroxyphenylpyruvate,respectively. An enzyme-membrane reactor system containing polyethyleneglycol-NAD' developed by Wandrey and associatesI"'1 is probably applicable to this system. The production level of D-aminoacids are mainly dependent on the stability of glutamate racemase. Therefore, thermostable glutamate racemases produced by A. pyrophi1 ~ 4 ~ and 1 ~ ~B. 1 subtilis[881are probably usehl as catalyst of this multi-enzyme system. Yagasaki et al. (lo21 developed a new method for the synthesis of D-glutamatefrom L-glutamate by means of E. coli recombinant cells harboring a plasmid containing glutamate racemase gene from L. brevis ATCC 8287. r-Glutamate was first racemized to D,L-glutamate at pH 8.5, and L-glutamate was then decarboxylated at pH 4.2 by glutamate decarboxylase, which was inherently produced by the E. coli host cells.

17.2 Racemizations and Epimerizations Figure 17-11. Production o f D-phenylalanine by successive feeding o f phenylpyruvate. Phenylpyruvate (U)was added intermittently. The dotted line indicates the expected producon tivity o f D-phenylalanine )(. the basis o f the initial production rate. Reprinted from Bae et aI.["'O]. K

0 C

8

0 '

5

10

15

20

Time (h)

25

30

35

Starting from 100 g L-' of L-glutamate, they obtained 50 g L-' of D-glutamate in a 15 h reaction. D-Glutamate can be produced successively from L-glutamate with L. breuis ATCC 8287 cells because this strain produces both glutamate racemase and glutamate decarboxylase simultaneously. Thus, 50 g L-' of optically pure D-glutamate was produced from 100 g L-' of glutam am ate['^^]. Oikawa et a1.['O4I replaced glutamate decarboxylase by glutamate oxidase because the oxidase has optimum pH values similar to that of glutamate racemase. They developed a bioreactor consisting of two columns sequentially connected and containing immobilized glutamate racemase from B. subtilis and L-glutamate oxidase from Streptomyces sp. X119-6: Lglutamate was racemized by the glutamate racemase column, and then L-glutamate was oxidized by the L-glutamate oxidase column. D-Glutamate was produced in about 90 % of the theoretical yield I1O41. 17.2.2.2

Aspartate Racemase (E.C. 5.1.1.13)

D-Aspartate occurs in the peptidoglycan layer of bacterial cell walls, and is produced from L-aspartate through an aspartate racemase (E. C. 5.1.1.13) reacti0n1~~'l.The enzyme has been demonstrated as being present in various Lactobacillus and Streptococcus strains [*06] such as Lactobacillus &rmenti['051 and Streptococcus faecalis1'"I. Recently, archaea such as Desulfirococcus strain SY [lo'] and Themococcus strains["'] were shown to produce aspartate racemase. It is interesting to note that various other archaea such as Pyrobaculum islandicum, Methanosarcina barkeri and Halobacterium salinarium produce D-amino acids, although their function is not yet known ["I'. Okada et al. purified the enzyme to homogeneity from the cell extract of S. thermophilus, the specific activity of the crude extract of which was elevated 3400-f01d[~~~]. The gene encoding aspartate racemase was cloned from S. thermophilus, and overexpressed in E. coli['ll]. The amount of the enzyme produced reached

I

1297

1298

I

17 lsomerizations

about 20% of the total soluble proteins of the E. coli clone cells. Thus, the enzyme was efficiently purified to homogeneity from the clone cells['*'! The enzyme is a homodimer of a subunit with a molecular weight of about 28000. In addition to aspartate, cysteate and cysteine sulfinate are the only substrates of the enzyme: they are racemized at a rate of 88 and 51 %, respectively, of that of L-aspartate['''I. The presence of the acidic group at the fi-carbon is essential; none of asparagine, cysteine, serine, and alanine are the substrates. Both isomers of glutamate are also inert. The KM values for L- and D-aspartate are 35 and 8.7 mM, respectively. Aspartate racemase requires no cofactors and contains an essential cysteine residue in the same manner as glutamate racemase[80].When L- or maspartate was incubated with aspartate racemase in tritiated water, tritium was incorporated preferentially into the product enantiomer. This is consistent with the results of glutamate racemase as described above["]. Yamauchi et al.['12] concluded that aspartate racemase also uses two bases to remove and return the a-proton of the substrate. Aspartate racemase contains three cysteine residues: Cys 84, Cys 190 and Cys 197, and only Cys 84 is essential for the enzyme activity. The alkylation of one cysteine residue/dimer with 2-nitro-5-thiocyanobenzoic acid results in a complete loss of activity. Therefore, the enzyme shows a ha1f-of-the-sites-reactivity[l1'1.Yamauchi et al. suggested that the enzyme has a composite active site formed at the interface of two identical subunits in the same manner as proposed for proline ra~emase['~]. Kumagai and c o w ~ r k e r s [ ' ~developed ~l an enzymatic procedure to produce Dalanine from fumarate by means of aspartase (E.C. 4.3.1.1), aspartate racemase, and D-amino acid aminotransferase (Fig. 17-12). Aspartase catalyzes conversion of fumarate into L-aspartate, which is racemized to form D-aspartate. D-Amino acid aminotransferase catalyzes transamination between D-aspartate and pyruvate to produce D-alanine and oxalacetate. This 2-0x0 acid is easily decarboxylated spontaneously to form pymvate in the presence of metals. Thus, the transamination proceeds exclusively toward the direction of D-alanine synthesis, and total conversion of fumarate into D-alanine was achieved.

Fumarate

J

Aspartase

L-Aspartate

1

ASPR

D-Aspartate

D-Alanine

z

A 1

\f

Pyruvate

Oxalacetate D-ATA

/--co2 Figure 17-12. Enzymatic production of D-alanine by combination of aspartase, aspartate racemase, and D-amino acid aminotransferase reactions.

7 7.2 Racemizations and Epimerizations

I

1 7.2.2.3

Diarninopirnelate Epirnerase (E. C. 5.1.1.7)

meso-a,&-Diaminopimelateis the direct precursor of L-lysine, and is an essential component of the cell wall peptidoglycans in Gram negative bacteria. meso-a,&Diaminopimelate is formed from L-a,&-diaminopimelateby diaminopimelate epimerase. The enzyme gene (dupF) was mapped at 85 min on the E. coli chromos0me['~~1. Richaud et al. isolated an E. coli mutant lacking diaminopimelate epimerase activity by insertional mutagenesis, and showed that the mutant does not require meso-a,&-diaminopimelate in a minimal medium l1l4]. Thus, meso-a,E-diaminopimelate epimerase encoded by the dapF gene is not essential for E. coli, but meso-a,&diaminopimelate still occurs in the mutant cells. Richaud et al. proposed that E. coli has another enzyme with diaminopimelate epimerase The diaminopimelate epimerase gene (dupF) was cloned from E. c0li['~~1, and the amino acid sequence of the enzyme was deduced from the nucleotide The enzyme was purified to homogeneity from the wild-type E. coli cells and the recombinant E. coli cells carrying a plasmid coding for dapF gene[116].The enzyme is composed of two identical subunits with a molecular weight of about 32 000. The enzyme is independent of PLP or of any other cofactors. The enzyme shows a V, value of 132 pmol min-' per mg of protein, and a I& value of 0.24 mM for L-a,Ediaminopimelate. The thiol group of Cys 73 of the enzyme is specifically labeled by a mechanism-based inactivator, 2- (4-amino-4-carboxybutyl)-2-aziridine carboxylic acid. Higgins et al. discovered an interesting similarity in amino acid sequences around the catalytically essential cysteine residue of proline racemase 19*1, hydroxyproline epimera~e['~], and diaminopimelate epimerase (Cys 73), and proposed that PLPindependent racemases/epimerases derive from a common evolutionary origin (l16]. However, no significant similarity in the entire amino acid sequence was found between diaminopimelate epimerase and glutamate racemase, and also between diaminopimelate epimerase and aspartate racemase. Cirilli et al. [11'1 cloned the gene of diaminopimelate epimerase from Huemophilus injuenzae, and purified and crystallized the enzyme. The enzyme is monomeric and has a unique protein fold, in which the amino terminal and carboxyl terminal halves of the molecule fold into structurally homologous and superimposable domains (Fig. 17-13).Cys 73 of the amino terminal domain is found in the disulfide linkage, at the domain interface, with Cys 217 of the carboxy terminal domain["']. Thus, it is most conceivable that these two cysteine residues stay in reduced form in the active enzyme and function as the acid and base in the mechanism. Koo and Blanchard['l8I explored a number of kinetic and isotope approaches to clarify the mechanism of the enzyme. However, which of the two cysteine residues is responsible for proton abstraction from the two enantiomeric Ca-H bonds is not yet known.

1299

1300

I I7 lsomerizations

,----(208

130

Figure 17-13. Top: Ribbon diagram of diaminopimelate epimerase from Haernophilus influenzae. The disulfide bridge between Cys 73 and Cys 21 7 connects domain I (residues 1-1 17 and 263-274) and domain II (residues 118-262).

,-I

312

201

BB

14b'

Bottom: Topology of the secondary structural elements of diaminopirnelate epimerase. The position of pseudo-2-fold symmetry axis is indicated by the black dot between p-strands B7 and B8. Reprinted from M. Cirilli et

7 7.2 Racernizations and Epirnerizations

I

1301

17.2.2.4

Proline Racernase (E. C. 5.1 .1.4)

Proline racemase occurs in Clostridium sticklandii, which produces 6-aminovalerate from L-proline. Proline racemase and D-proline reductase are responsible for the conversion: L-proline is racemized by proline racemase to form D-proline,which is converted into F-aminovalerate by D-proline reductase (E. C. 1.4.4.1). Rudnick and Abeles purified proline racemase to 95 % homogeneity from Clostridium sticklandii, and characterized The enzyme is composed of two identical subunits with a molecular weight of about 38000, and is independent of any cofactors or metals. Most amino acid racemases require pyridoxal 5'-phosphate, which labilizes the bond between the a-hydrogen and the chiral center by aldimine formation with the a-amino group of the substrate. However, PLP is not involved in the reaction of proline racemase acting on an a-imino acid. The enzyme also acts on 2-hydroxy-~-prolineand 2-allo-hydroxy-~-proline although slowly: they are epimerized at a rate of 2 and 5 % of the rate of r-proline racemization, respectively. L-Proline and D-proline showed KM values of 2.9 and 2.5 mM, respectively["'l. Pyrrole-2-carboxylate is a competitive inhibitor of proline racemase, and stoichiometrically binds with the enzyme (1mol per dimer). Thiol groups of the enzyme are alkylated by iodoacetate at a stoichiometry of 1 mol of cysteine residue per mol subunit. However, the enzyme is inactivated completely by modification of only one cysteine residue per dimer. Thus, Rudnick and Abeles proposed a reaction scheme in which the active site is located at the interface of two identical subunits, each of which furnishes one of the two active site thiol groups positioning appropriately at the composit active site: a thiolate anion derived from one thiol group abstracts the aproton from the substrate, and another thiol group protonates the intermediate derived from the substrate from the opposite face["]. They proposed occurrence of two forms of free proline racemase: one binds with D-proline and the other binds with L-proline. According to their proposed mechanism, the product enantiomer is released much faster than the release of the substrate-derived proton. The proton release also proceeds much faster than the interconversion of the two forms of the enzyme. Knowles and coworkers defined the energetics and delineated the complete free energy profile for the proline racemase reaction [119-1251. Yagasaki and Ozaki[1261developed a method for production of D-proline from Lproline using the recombinant proline racemase of Clostridiurn sticklandii. L-Proline was degraded by Candida sp. PRD-234, and optically pure D-proline was obtained. 17.2.3 Other Racernases and Epirnerases Acting on Amino Acid Derivatives 17.2.3.1

2-Arninod2-thiazoline-4-carboxylateRacernase

Sano et al. [12'1 have found several bacterial strains that are capable of producing Lcysteine from ~,~-2-amino-2-thiazoline-4-carboxylate (ATC),an intermediate in the

1302

I

7 7 lsomerizotions

0

. . hydrolase I

),sT~~~~

H2N

H2Ng & C O O H

L-2-amino-A2-thiazoline4-carboxylate

li

hydrolase II

HS C (--ooH

NH2 S-carbamoyl-L-cysteine

NH2 L-cysteine

racemase

L>

H2N

"'/C0OH

D-2-amino-A2-thiazoline4-carboxylate

Figure 17-14.

Enzymatic synthesis of L-cysteine from ~,~-2-amino-A*-thiazoline-4-carboxylate.

chemical synthesis of D,L-cysteine. These include several Pseudornonas species isolated from soil and other strains belonging to different genera such as E. coli, Bacillus breuis, and Micrococcus s~denensis['~~]. Three enzymes are probably involved in this pathway: L-ATChydrolase, S-carbamoyl-L-cysteinehydrolase and ATC racemase (Fig. 17-14). Pseudornonas thiazolinophilurn isolated from soil was shown to have the highest activity of the enzymes that produce L-cysteine from D,L-ATC.The enzymes are inducibly formed in the bacterial cells by addition of D,L-ATCto the growth medium. Degradation of L-cysteine by cysteine desulfhydrase or other PLP enzymes present in the cells was successfully prevented by addition of hydroxylamine or semicarbazide to the incubation mixture. A mutant strain of Ps. thiazolinophilum lacking cysteine desulfhydrase was isolated and used to produce L-cysteine from D,L-ATCin a molar yield of 95% and at a product concentration of 31.4 g L-'['28]. Pseudornonas desrnolytica AJ 3872, one of the L-cysteine producers isolated was found to lack the ability to convert D-ATC into L-cysteine: it is an ATC racemase-deficient strain['*']. However, little is known about the enzymological properties and function of the racemase. Among the three enzymes participating in L-cysteine production, L-ATChydrolase was found to be the least stable['30].However, the stability of L-ATChydrolase was sharply enhanced as water activity decreased from 0.93 to 0.80. In the absence of sorbitol, the stability of L-ATChydrolase increased in proportion to ionic strength. Thus, Ryu et al. succeeded in enhancing the half life of L-ATChydrolase by 10-foldto 20-fold in sorbitol-saltmixtures [1301.

7 7.2 Racemizations and Epimerizations

I

17.2.3.2 Hydantoin Racemase

5-Substitutedhydantoin derivatives have been used as precursors for D- and L-amino acids in chemical synthesis. However, they are hydrolyzed enantioselectively by the enzymes named hydantoinases: some act specifically on D-5-substitutedhydantoins, and others on the r-isomers. N-Carbamoyl amino acids formed are also hydrolyzed enantiospecifically by N-carbamoyl amino acid amidohydrolases to produce D- or Lamino acids (Fig. 17-15). Since the Kanegafuchi Chemical Industry, Japan, commercialized an enzymatic procedure for the production of D-p-hydroxyphenylglycine, which is a building block for the semisynthetic j3-lactam antibiotic amoxycillin, various processes for amino acid production by means of hydantoinases have been devel~ped[~~~-'~~]. Subsequent to the discovery that hydantoin is hydrolyzed by extracts of mammalian livers [1341 and plant seeds [1351,various microorganisms have been shown to utilize D- and L-5-substituted hydantoins as a sole carbon or nitrogen source by means of D- as well as L-specific hydantoinases inducibly formed [131-1331. Distribution of D-hydantoinase in microorganisms has been shown by Yamada and coworker^^^^^]. The enzyme is identical to dihydropyrimidinase (E. C. 3.5.2.2), and is widely distributed in bacteria, in particular in Klebsiella, Corynebacterium, Agrobacteriurn, Pseudornonas, and Bacillus, and also in actinomycetes such as Streptornyces and Actinoplanes. The enzyme activity occurs also in eukaryotes: yeasts, molds, plants and mammals. Pseudomonas putida was found to be the best strain, which produced D-hydantoinasemost abundantly and inducibly by addition of 5-methylhydantoin. Most of D-hydantoinase producers form N-carbamoyl D-amino acids from the corresponding 5-substituted hydantoins. Accordingly, to obtain free D-amino acids, N-carbamoyl amino acids need to be isolated and hydrolyzed chemically or enzymatically. However, a few bacterial strains produce N-carbamoyl D-amino acid amidohydrolase in addition to D-hydantoinase. Thus, optically pure D-amino acids were produced from D-hydantoinswith these bacterial cells. Olivieri et al. [1371 found that Agrobacteriurn turnefaciens cells grown on uracil as a sole nitrogen source catalyze the complete conversion of racemic hydantoins into D-amino acids. Hartley et a1.[138] obtained a mutant strain which expresses both the hydantoinase and Ncarbamoylamino acid amidohydrolase in the absence of an inducer. In contrast, other bacterial strains belonging to the genera of Flavobacteri~m['~'],Arthro-

R H O

Hydantoinase H20

HNKNH 0 R = aryl, alkyl

*

R y c o o H

Chemical or enzymatic hydrolysis

HNKNH2 H20 0

*

RyCooH + + COz NH2

* LOrD

Figure 17-15. Enzymatic synthesis of D- or L-amino acids from 5-substituted D,L-hydantoinsthrough N-carbamoyl-D- or L-amino acids.

NH3

1303

1304

I b a ~ t e r [ ' ~Pseud~monas['~'~ ~], 7 7 lsomerizations

1421, and B a ~ i l l u s [ ' ~convert ~ - ~ ~ ~whole ] racemic 5-substituted hydantoins into the corresponding L-amino acids. In these bacteria, 5-substituted hydantoins are hydrolyzed by L-hydantoinase to form N-carbamoyl L-amino acids, which are hydrolyzed further to L-amino acids by N-carbamoyl L-amino acid amidohydrolase in the same manner as described above except that the enzymes involved show opposite stereospecificity. 5-Mono-substituted hydantoins can racemize spontaneously under weakly alkaline conditions, and this chemical racemization participates at least partly in the total conversion of the racemic hydantoins into free L- or D-amino acids. However, if chemical racemization proceeds only a hydantoin racemase was suggested to occur and participate in the total conversion[146, 1471. Watabe et a1.[148]isolated a plasmid which is responsible for the conversion of 5-substituted hydantoins into the corresponding L-amino acids from a soil bacterium, Pseudomonas sp. NS 671, which is able to convert racemic 5-substituted hydantoins into the corresponding L-amino acids. The genes involved in the conversion were cloned from the Pseudomonas plasmid into E. coli, and functions of four genes were identified and named hyuA, hyuB, hyuC and hyuE. Both hyuA and hyuB are required for the conversion of D- and L-5-substitutedhydantoins into the corresponding N-carbamoyl-D- and N-carbamoyl-L-amino acids, respectively, although the individual reactions catalyzed by the gene products have not yet been identified. HyuC codes for an N-carbamoyl-L-aminoacid amidohydrolase, while hyuE is a hydantoin racemase gene[l4'I. Significant nucleotide sequence similarity was found between hyuA and hyuC (43%), and also between hyuB and hyuC (46%). Watabe et al. suggested that these genes have evolved from a common ancestor by gene However, no proteins registered in NBRF and SWISS protein data bases showed similarity with the deduced amino acid sequences of the four genes. Wagner and associates purified hydantoin racemase from Arthrobacter aurescens DSM 3747 and characterized Watabe et al. [14'1 also purified the enzyme from E. coli clone cells harboring a plasmid coding for the enzyme gene derived from Pseudomonas sp. NS 671. The Pseudomonas enzyme is a hexamer composed of a subunit with a molecular weight of about 32000, which is consistent with the value deduced from the amino acid sequence. The D- and L-isomers of 5-(2-methylthioethy1)hydantoin and 5-isobutyrylhydantoinare racemized effectively. ~-5-(2-Methylthioethy1)hydantoinis racemized at a V,, value (79 pmol min-' mg-') which is about 2.5 times higher than that for the L-isomer. Wiese et al.['501 cloned the hydantoin racemase gene from Arthrobacter aurescens DSM 3747 and purified the enzyme to homogeneity. The Arthrobacter enzyme has a molecular mass of 25.1 kDa['50] and acts on aromatic and aliphatic hydantoin derivatives such as 5-indolylmethylhydantoin, 5-benzylhydantoin, 5-(p-hydroxybenzyl)hydantoin,5-(2-methylthioethyl)hydantoin, and 5-i~obutylhydantoin['~~l, although hydantoins with arylalkyl side chains are preferred substrates [1591. Free amino acids, amino acid esters and amides are inert, but the enzyme suffers from inhibition by aliphatic The hydrogen at the chiral center substrates such as L-5-methylthioethylhydantoin. is exchanged with solvent deuterium of a substrate, ~-5-indolylmethylenehydantoin,

17.2 Racemizations and Epimerizations

I

1305

during racemizati~n['~~]. Pietzsch et al. established a method for the synthesis of in optically pure ~-3-trimethylsilylalaninefrom ~,~-5-trimethylsilylmethylhydantoin 88 % yield and 95 % enantiomeric excess with whole resting cells of Agrobacterium sp. IP I 671, immobilized in a Ca-alginatematrix. On the other hand, L-3-trimethylsilylalanine was also prepared from the racemic substrate by enantiomer-specific hydrolysis of the L-form in the presence of L-N-carbamoylase from Arthrobacter aurescens DSM 3747[1521. Watabe et al. found that the Pseudornonas enzyme is inactivated by a substrate, L5-methylhydantoin,during ra~emization~'~']. However, the enzyme was not affected by the D-isomer. Both enantiomers of 5-isopropylhydantoininactivated the enzyme to the same extent. Interestingly, divalent sulfur-containing compounds such as methionine, cysteine, glutathione, and biotin protected the enzyme effectively from inactivation. E. coli cells expressing the racemase are capable of racemizing all of these hydantoin derivatives: the enzyme is protected from inactivation by divalent sulfur compounds occurring in the cells. Watabe et al. concluded that the protective effect by the divalent sulfur-compounds is not due to their reducing Both Pseudom~nas['~~] and Arthr~bacter['~~] enzymes are inhibited strongly by Cu2+.The Arthrobacter enzyme is completely inhibited by HgCl2 and iodoacetamide, and stimulated by addition of dithiothreitol [1501. Therefore, the enzyme may contain essential cysteine residues, which are possibly modified by some activated intermediate derived from the particular substrates leading to the enzyme inactivation. E. coli cells canying a plasmid coding for hyuA, hyuB, hyuC, and hyuE convert only ~ - 5 2methylthioethyl)hydantoin -( into L-methionine. On the other hand, E. coli cells harboring a plasmid coding for only hyuA, hyuB, and hyuC first convert the Lhydantoin, then the D-isomer is hydrolyzed slowly when the L-isomer is depleted. is only conTherefore, Watabe et al. believe that ~-5-(2-methylthioethyl)hydantoin verted into L-methionine in the presence of the hydantoin racema~e['~'].The mechanism of stereospecific conversion of D,L-S-substituted hydantoins to the corresponding L-amino acids by Pseudomonas sp. strain NS 671 has been clarified by Ishikawa et al. ~,~-S-substituted hydantoins are converted exclusively into the Lforms of the corresponding N-carbamoylamino acids by the hydantoinase in combination with hydantoin racemase, and then the N-carbamoyl-t-amino acids are converted into L-amino acids by N-carbamoyl-t-aminoacid amidohydrolase (Fig. 1716). By directed evolution May et al.

succeeded in inverting the enantioselectivity of D-hydantoinase from Arthrobacter sp. DSM 9771 into an L-selective enzyme. The improved hydantoinase also acquired a five-fold increase in activity. The recombinant E. coli cells expressing three heterologous genes (i. e. the evolved L-hydantoinase, L-N-carbamoylase, and hydantoin racemase) were found to produce 91 mM Lmethionine from 100 mM ~-5-(2-methylthioethyl)hydantoin in less than 2 h[1541.

1306

I

7 7 lsomerizations

“yCooH HNKNH = HNyNHz 0 ATP ADP

0 L-5-Substituted Hydantoin

II Hydantoin Racemase

Acid KCarbamyl-L-Amino* Amidohydrolase

0

/I

N-CarbamylL-Amino Acid

RYCooH NH2 L-Amino Acid

U

L-preferential Hydantoinase

D-5-Substituted Hydantoin

N-CarbamylD-Amino Acid

Figure 17-16. Stereospecific conversion of o,~-5-substitutedhydantoins into the corresponding L-amino acids by Pseudomonos sp. NS 671. Reprinted from lshikawa et al.11531.

17.2.3.3 N-Acylamino Acid Racemase

~-Aminoacylases(E. C. 3.5.1.14) catalyze the hydrolysis of the amide bond of various N-acyl-L-amino acids, such as N-acetyl-, N-chloroacetyl- and N-propionyl- amino acids [1551, and is widely distributed in animals [155-1571, plants [lS8, and microorganisms [lGo, l6l1. Greenstein[”’] first studied the reactivity of pig kidney enzyme, and showed its application to the optical resolution of racemic amino acids. Chibata et al. [IG2] found that L-aminoacylase is produced abundantly by fungal species belonging to the genera Aspergillus and Penicillium. L-Aminoacylaseswere purified from pig kidney and A. oryzae, and their reaction mechanism and physiological function were studied[l“. 163-1Gs1 . Cho et a]. [‘“I showed that various thermophilic Bacillus strains produce thermostable L-aminoacylase, and purified it to homogeneity from Bacillus themoglucosidius DSM 2542, which produces the enzyme most abundantly. L-Aminoacylases of pig kidney, Aspergillus oryzae and B. thermoglucosidius share many features with each other: they contain Zn” as a prosthetic metal, are strongly activated by Co2+,and have a pH optimum in the range of 8.0-8.5. Sugie and S u ~ u k i [ ~ ”demonstrated ] the occurrence of D-aminoacylase, which specifically hydrolyzes the amide bond of N-acyl-D-aminoacids, in actinomycetes, and applied the enzyme to the production of ~-phenylglycine.Recently, a new Daminopeptidase was found in Alcaligenes denitnicans, and shown to act on various Nacyl-~-amino acids including N-acetyl-D-methionine[“*. lG9].

17.2 Racemizations and Epimerizations

I

1307

N-Acylamino acids are usually racemized much more readily than the corresponding free amino acids. Therefore, by combination of chemical racemization and enantioselective hydrolysis of N-acylamino acids, racemates of N-acylamino acids can be fully converted into the desired enantiomer of the free amino acids according to the stereospecificity of the aminoacylases used. For example, L-tryptophan is produced industrially by combination of chemical racemization of N-acetyltryptophan and enantiospecific hydrolysis of its L-isomer with the Aspergillus L-aminoacylase, which shows high reactivity towards N-acyl derivatives of aromatic L-amino acids. When N-acetyl-D,L-tryptophanis incubated with the fungal enzyme, N-acetylL-tryptophan is selectively hydrolyzed to L-tryptophan, which is then crystallized from the solution. N-Acetyl-D-tryptophan in the mother liquor is racemized with acetic anhydride, and the racemate is again used as a starting material. In principle, D- and L-amino acids can be produced from their corresponding N-acyl derivatives in the same manner, provided that N-acyl derivativesof the desired amino acids serve as the substrates of the available aminoacylases,and are racemized chemically without any major loss by decomposition. However, the chemical racemization can be achieved only under extreme conditions in order for the aminoacylases to be inactivated, and the enzymes are usually required to be saved for the subsequent cycles for reasons of economy. Therefore, the antipode of the substrate is separated from the enzyme and preferably from the product in order to avoid its possible racemization. Tosa et al. have developed a continuous method to produce Ltryptophan, which is now utilized in industry, by means of the Aspergillus Laminoacylase immobilized on DEAE-Sephadex[17']. Takahashi and Hatano of Takeda Chemical Industries, Japan, succeeded in finding a racemase that acts on N-acylaminoacids, but not on the corresponding free amino acids, and named it acylamino acid racema~e["~]. They have established a method of producing optically active a-amino acids from the corresponding D,L-N-acylamino acids by means of the acylamino acid racemase and aminoacylases. Acylamino acid racemase occurs widely in various actinomycete strains belonging to the genera of Streptomyces, Actinomadura, Actinomyces, Iensenia, and Amycolato psi^["^]. The enzyme was purified to homogeneity from Streptomyces atratus Y-53, which shows the highest enzyme activity among the strains tested['73].The enzyme is composed of G subunits with identical molecular masses (about 41000), and shows a molecular mass of 244000 in the native state. Tokuyama and Hatan0['~'1 purified thermostable N-acylamino acid racemase from Amycolatopsis sp. TS-1-60 and purified it to homogeneity. The molecular masses of the native enzyme and the subunit are 300000 and 40000, respectively. The enzyme is stable at 55 "C for 30 min. The enzyme catalyzes the racemization of N-acylaminoacids such as N-acetylL- or D-methionine, N-acetyl-L-valine,N-acetyl-L-tyrosineand N-chloroacetyl-L-valine (Table 17-5). In addition, the enzyme also catalyzes racemization of dipeptide Lalanyl-L-methionine.By contrast, N-alkylamino acids and methyl and ethyl esters of N-acetyl-D- and L-methionine are not racemized. The apparent KM values for Nacetyl-L-methionine and N-acetyl-D-methionine are 18.5 mM and 11.3 mM, respectively. The enzyme activity is markedly enhanced by the addition of divalent metal ions such as Co2+,Mn2+ and Fe2' and inhibited by addition of EDTA and p

1308

I

17 lsornerizations Table 17-5.

Substrate specificity of acylamino acid racemasea.

Substrate

Relative activity

N-Acetyl-o-methionine N-Acetyl-r-methionine N-Formyl-D-methionine N-Formyl-L-methionine N-Acetyl-D-alanine N-Acetyl-L-alanine N-Benzoyl-o-alanine N-Acetyl-D-leucine N-Acetyl-L-leucine N-Acetyl-n-phenylalanine N-Acetyl-L-phenylalanine N-Chloroacetyl-D-phenylalanine N-Chloroacetyl-L-phenylalanine N-Acetyl-D-tryptophan N-Acetyl-L-tryptophan N-Acetyl-D-vahe N-Acetyl-r-valine N-Chloroacetyl-D-vahe N-Chloroacetyl-L-valine N-Acetyl-D-alloi s o h c i n e

100 100 40 63 33 21 14 37 74 64 84

90 112 10 8

35 19 80 105 33

a Inert: D- and r-methionine, D- and r-alanine, D- and L-leucine, wand L-phenylalanine. D-

and L-tryptophan,D- and L-valine.

chloromercuribenzoate. The gene of N-acylamino acid racemase was cloned from Amycolatopsis sp. TS-1-60[1751, and overexpressed in E. coli host cells with T7 promoter[’76].The gene codes for a protein of 368 amino acids with a molecular mass of 39411 Da. Palmer et al.[177]found that N-acylamino acid racemase of Amycolaptosis sp. TS-1-60 is similar to an unidentified protein encoded by the Bacillus subtilis genome. N-Acylamino acid racemase efficiently catalyzes an 0succinylbenzoate synthase reaction, which is responsible for menaquinone biosynthesis. Tokuyama et al. [1721 found that most of acylamino acid racemase-producing strains produce not only acylamino acid racemase but also aminoacylases; one of either D- or r-aminoacylase or both of them. Moreover, acylamino acid racemase shows the optimum pH at around 8.0, which is close to that of aminoacylases. Therefore, Nacylamino acid can be converted as a whole into L- or D-amino acids in one step by means of microbial cells of appropriate strains producing either L- or D-aminoacylase in addition to acylamino acid racemase. 17.2.3.4 lsopenicillin N Epimerase

Isopenicillin N is a precursor of penicillin, and synthesized from 6-(L-aminoadipoy1)Isopenicillin N is then conL-cysteinyl-D-valineby isopenicillin N synthetase verted into penicillin N by isopenicillin N epimerase. Penicillin N is ring-expandedto deacetoxycepharosporin C by penicillin N expandase. The latter compound is

17.2 Racemizations and Epimerizations

sCOOH Hs$H isopenicillin N epimerase

isopenicillin N synthetase

7-T02

O

H2N

i

H20

ICOOH

H2N

COOH

isopenicillin N

&(L-a-aminoadipoyl)L-cysteinyl-0-valine

penicillin N

S%COOH

SI$COOH -N

I-N

penicillin N expandase 02

deacetoxycepharosporin C hydroxylase

Ir

02 a-ketoglutarate

a-ketoglutarate

deacetoxycepharosporin C Figure 17-17.

deacetylcepharosporin C

Biosynthetic pathway for cepharosporin C.

hydroxylated to form deacetylcepharosporinC by deacetoxycepharosponnC hydroxylase. These reactions proceed sequentially in the biosynthesis of cepharosporin C in 180] (Fig. 17Streptomyces clavuligerus, a producer of various p-lactam antibiotics 17). However, in Cepharosporium acremonium, conversion of penicillin N into deacetoxycepharosponn C is catalyzed by a bifunctional enzyme, penicillin N expandaseldeacetoxycepharosporin C hydroxylase in Cepharosporium acremonium11811. Isopenicillin N epimerase activity, demonstrated in the extract of Cepharosporiurn acremonium protoplasts was found to be very unstabIe['821. Usui and Yu11831, however, succeeded in purifying the enzyme to homogeneity after development of a simple assay procedure of the enzyme. They studied its enzymological properties[183].The enzyme has a monomeric structure with a molecular mass of 47000. The enzyme contains 1 mol of PLP per mol of protein. The enzyme shows a V,, value of 3.93 pmol min-' per mg and a KM of 0.30 mM for isopenicillin N, whereas it of 9.47 pmol min-' per mg and a KM of 0.78 mM for penicillin N. The shows a V,, Gqvalue for the conversion between isopenicillin N and penicillin N is 1.09, which is in good agreement with the theoretical value. In addition to isopenicillin N and penicillin N, deacetoxycepharosporin C was epimerized only slowly: the rate relative

1310

I

I7 lsomerizations

to isopenicillin N is about 1%. However, the following penicillin derivativesare inert: deacetylcepharosporin C, ceparosporin C, &(L-a-aminoadiopoyl)-L-cysteinyl-D-valine, L-a-aminoadipate,and D-a-aminoadipate.The enzyme is inhibited strongly by thiol reagents such as p-chloromercuribenzoate[1831. 17.2.4

Racemization and Epimerization at Hydroxyl Carbons

Various epimerases acting on carbohydratederivatives and acyl-CoA derivativeswere demonstrated, purified, and characterized as reviewed previously['84].Lactate racemase (E.C. 5.1.2.1) is the first racemase to he The mechanism of lactate racemase reaction was studied with the enzyme preparations partially purified from Clostridium b u t y r i c ~ r n [ ' ~ ~Hiyama ]. et al. [186] highly purified the enzyme from Lactobacillus sake, but little is known about its enzymological properties. In contrast, mandelate racemase (E. C. 5.1.2.2) is the enzyme best characterized among various racemases and epimerases: its tertiary structure and functional groups that participate directly in catalysis has been clarified. 17.2.4.1

Mandelate Racemase (E.C. 5.1.2.2)

Mandelate racemase catalyzes the racemization of mandelate, which is the first step of the mandelate assimilation pathway in Pseudomonas putida. Although the mandelate pathway occurs widely in various bacteria, fungi and yeasts, most of them utilize one enantiomer or the other of mandelate in a benzoate-forming pathway. A few strains such as Acinetobacter calcoacetic~s[~~~] and Aspergillus nigar[188]are capable of using both enantiomers with two complementary dehydrogenases with different stereospecificities. However, a single strain of Pseudomonas putida producing mandelate racemase can utilize both enantiomers [1891. In Pseudomonas putida, D-mandelate is converted into L-mandelate by mandelate racemase, then oxidized to benzoylformate by mandelate dehydrogenase (Fig. 1718). Benzoylformate decarboxylase is the second enzyme of the pathway and catalyzes decarboxylationof benzoylformateto form benzaldehyde,which is oxidized to benzoate by NAD- and NADP-linked benzaldehyde dehydrogenases. The genes encoding these five enzymes constitute an operon that is induced by either enantiomer of mandelate [l9O]. Stecher et al. ["I' established large-scale production of mandelate racemase by Pseudomonas putida ATCC12633 by optimization of enzyme induction: both glucose and mandelate were added to the culture right from the start as the carbon source. Thus, about 300-fold enhancement in the enzyme production was achieved. Strauss et al. [1921 showed that immobilized mandelate racemase is an efficient biocatalyst used for repeated batch reactions to produce (R)-mandelatefrom (S)-mandelateunder mild conditions. Kenyon and coworkers purified mandelate racemase to homogeneity, and characterized it[ls9].Divalent metal ions such as Mg2+,Mn2+,Co2+,and Ni" were required for the catalysis. In addition to mandelate, p-hydroxymandelate and p-(bromome-

7 7.2 Racemizations and Epirnerizations

H : hOH

Hoh:o*

mandelate racemase \

-..

Figure 17-18.

1311

dehydrogenase

L

6

O benzoylformate dehydrogenase*

I

O6;OoH

mandelate

P

\

H

benzaldehyd dehydrogenase,

\

d ‘OH

p-keto adipate pathway

acetylCoA + succinate

Mandelate assimilation pathway in Pseudomonas putida.

thy1)mandelate serve as the substrates. p(Bromomethy1)mandelate is decomposed to p-(methy1)benzoylformateand bromide by action of the enzyme. The KM values for D- and L-mandelateare 0.23 and 0.26 mM, respectively. Ransom et a1.[193]cloned the gene for mandelate racemase from Pseudomonas putida in Pseudomonas aeruginosa on the basis of the inability of the latter strain to grow on D-mandelateas a sole carbon source. The amino acid sequence was deduced from the nucleotide sequence, and the predicted molecular mass of the enzyme was 38750[1931.The enzyme is composed of eight identical subunits. The crystal structure of mandelate racemase has been solved and refined at 2.5 A re~olution[”~1. The secondary, tertiary and quaternary structures of mandelate racemase are quite similar to those of muconate lactonizing enzyme[”’, 1961 . Mandelate racemase is composed of two major structural domains and a small C-terminal domain. The Nterminal domain has an a + p structure, and the central domain has an a/P-barrel topology. The C-terminal domain consists of an L-shaped loop. Divalent metal ions, which are essential catalykally, are ligated by three distal carboxyl groups of Asp 195, Glu 221, and Glu 247, all of which occur at the central domain[194].The active site location was determined by analysis of a complex between mandelate racemase and p-iodomandelate, whose iodine atom has high electron density and contributes greatly to the analysis. The active site of the enzyme is located between the two major domains. The ionizable groups of Lys 166 and His 297 are located at the positions interacting with the chiral center of the substrate (Fig. 17-19).Neidhart et al. proposed that they participate in general acid/base catalysis: Lys 166 abstracts the a-proton of r-mandelate, and His 297 abstracts the aproton from D-mandelate. Landro et al. [19’] then replaced His 297 by asparagine, analyzed the crystal structure of the H297N mutant enzyme at 2.2 A resolution, and studied the mechanism of catalysis of the mutant enzyme. Although the mutant enzyme has no mandelate racemase activity, it catalyzes the stereospecific elimination of bromide from p-(bromomethy1)-L-mandelateat a rate equivalent to that catalyzed by the wild-typeenzyme. Moreover, the mutant enzyme catalyzes exchange of the a-hydrogen of L- but not D-mandelatewith deuterium in deuterium oxide at a rate 3.3 times less than that of the wild-type enzyme. Thus, Landro et al.[”’, ”)‘I concluded that the mandelate racemase reaction proceeds through a two-base

1312

I

17 lsomerizations

139SEA

f\

GLU

GLU

GLU

Models o f the mandelate racemase active site with complexed substrate, p-iodomandelate. Reprinted from Neidhart et al.[194]. Figure 17-19.

mechanism in which Lys 166 abstracts the a-proton from L-mandelateand His 297 abstracts the a-proton from D-mandelate (Fig. 17-20). In fact, the X-ray crystal studies of mandelate racemase inactivated by (R)-a-phenylglycidaterevealed that the E-amino group of Lys 166 is covalently bound to the distal carbon of the epoxide ring[”’]. KlGGR mutant enzyme catalyzes the stereospecific elimination of bromide ion from (R)-p-(bromomethy1)mandelate to form p(methy1)benzoylformateat a rate similar to that catalyzed by the wild-typeenzyme[200], while H297N acts stereospecif[2011. This is compatible with the mechanism ically on (S)-p-(bromomethy1)mandelate that Lys 166 and His 297 participate as the (S)- and (R)-specificcatalyst, respectively. Bearne and Wolfenden[2021 proposed that the complementary nature of the structures of mandelate racemase and its substrate is optimized in the transition state otherwise the general acid-generalbase catalysis will not become an efficient mode of catalysis.

17.3

lsomerizations

We describe here the enzymological characteristics and application of isomerases, especially D-xylose (glucose) isomerase, phosphoglucose isomerase, triose phosphate isomerase, L-rhamnose isomerase, L-fucose isomerase, maleate cis-trans isomerase, and unsaturated fatty acid cis-trans isomerase. i%ketyl-D-glucosamine 2-epimerase is not an isomerase, but for convenience we will also describe the characteristics and use of the enzyme because this section deals with sugarmetabolizing enzymes.

17.3 lsomerizations

o y G l 317 ~

I

LYs

166-NH3

H%...H-O

Figure 17-20. . I .

N H i-==/Nt

HO. ~

. ,,\O---HaN-Lys 164 'Mi2+

His 297

Mechanism o f t h e

reaction catalyzed by mandelate racemase with concerted general acidgeneral base through an enolic intermediate. Reprinted from Mitra et aI. [' 981.

17.3.1

D-Xylose (Glucose) lsomerase (E. C. 5.3.1.5)

D-Xylose isomerase catalyzes the interconversion between D-xylose and D-xylulose (Fig. 17-21). Since this enzyme acts on D-glucose to produce D-fructose, it is often referred to as glucose isomerase (Fig. 17-21). The isomerization of glucose to fructose by this enzyme is a very important process for the industrial production of high fructose corn syrup. This enzyme is also applicable to the synthesis of many aldoses and ketoses because of its wide substrate specificity. The enzyme gene has been cloned from various microorganisms, and the enzyme has been overexpressed, purified, and characterized. Their three dimensional structures have also been determined [203-20G1. 17.3.1.1

Properties

Xylose isomerases have been purified from various microorganisms, such as Lactobacillus brevis, Streptomyces sp., Bacillus stearothemophilus, and Actinoplanes

I

1313

1314

I

77 lsomerizations

CHO

YHO H-C-OH I HO-C-H

YH2OH

c=o I

___)L

H-+-OH CH2OH D-Xylose Figure 17-21.

YH20H

c=o

H-?-OH HO-C-H

HO-C-H I

H-?-OH CH20H

D-Xylulose

H-C-OH H-C-OH I

CH2OH

D-Glucose

d

HO-I;-H I H-C-OH

~ - 6 - 0 ~ I

CH20H

D-Fructose

Reactions catalyzed by D-xylose isomerase.

rnissouriensis[207-2101. They consist of four identical subunits whose molecular mass are in the range 42 000-51 000. The optimum pH usually ranges from 7.0 to 9.0. The cDNA for barley (Hordeurn uulgare) enzyme gene has been cloned, and the recombinant enzyme characterized[211]. It is unique because it is a dimer composed of a subunit with a molecular mass of 53620, which is much larger than those of microbial enzymes. Thermostable xylose isomerases were purified and characterized from many thermophilic bacteria f204, 205, 212-2221 . The enzyme isolated from Themotoga neapolitana is extremely thermostable, with the optimal activity being above 95 oC[21Gl. The catalytic efficiency (kcat/ht) of the enzyme is essentially constant between GO and 90 "C, and decreases between 90 and 98 "C primarily because of a large increase in KM. Xylose isomerase requires divalent metal cations, usually Mg2', Mn2+,or Co2+for the maximum activity and thermal stability. The enzyme has a wide substrate specificity[223]: glucose and fructose derivatives modified at the 3-, 5- or 6-position are isomerized by the enzyme as will be described later. 17.3.1.2

Reaction Mechanism

The reaction mechanism of xylose isomerase was proposed based on X-ray cryst a l l ~ g r a p h y [and ~ ~ molecular ~] mechanical and molecular orbital studies [2251. The a-pyranose form of the substrate binds to the active site of the enzyme, and the reaction is initiated by ring-opening involving hydrogen transfer from the first hydroxyl group to 0 5 (Fig. 17-22).After extension of the substrate, a water molecule abstracts the proton from the hydroxyl group at 0 2 of xylose and transfers it to Asp 257 in the second step. The following hydride shift causes isomerization. The 01 atom of the ketose is negatively charged and most probably abstracts a proton from Asp 257. The stable cyclic conformation is then formed. This hydride shift reaction mechanism is quite different from the base-catalyzed enolization mechanism proposed for phospho sugar isomerases such as triosephosphate isomerase which generally do not require a metal ion for activity[226].

17.3 lsomerizations

B Glu 217

-

.N

. ’ -..-o ‘M$T?F .\ --_ - F A s p 2 5 5 *’

t

H

,Mg*i

,‘

N=r\ His54

bf

O\\

1315

H (yHis220

),

P

I

,I

‘.

\

“0

’0

I

Asp 57

OH

OH

H

+

H N 7 NH

His 54

Asp 57 Asp 257

GIU217,

H

H

His 54

Asp 57 Asp 257

Reaction mechanism for xylose-xylulose conversion by o-xylose isomerase through ring opening (A) and hydride shift (B). Reprinted from Fuxreiter et al. [2251. Figure 17-22.

1316

I

77 /sorner;zations

17.3.1.3

Production o f Fructose

Xylose isomerase derived from various microorganisms, such as Actinoplanes missouriensis, Streptomyces griseofiscus, Havobacterium arborescens, Streptomyces phaechromogenes, Bacillus coagulans, Streptomyces murinus, Streptomyces rubiginosus, and Streptomyces oliuochromogenes, is utilized in the annual conversion of 3 million tons of glucose into fructose for use as high fructose corn syrup. The enzyme is immobilized by glutaraldehyde cross-linking or adsorption on an insoluble resin for the fixed bed isomerization process [2271. The isomerization is reversible, and the final fructose content depends on the reaction temperature. The reaction is usually carried out in the region of 60-65 "C. However, a higher temperature gives a higher fructose content. It is reported that the degree of conversion is raised from 42 %, which is the normal fructose content of the syrup, to 55 % by isomerization with xylose isomerase at about 95 0C[2271.Therefore, the thermostability of the enzyme is an important issue. Recently, several thermo205. 212-222]. It is also stable xylose isomerases were found and reported that the thermostability of the enzyme is enhanced by site-directed mutagenesis [22sl. a-Amylasesand xylose isomerases with low optimum pH values are expected to be useful for fructose production from cornstarch because raw cornstarch solutions have an acidic pH of around 4.5 and the glucoamylase reaction, the second step in the process, prefers an acidic pH. Fructose can be produced from cornstarch without pH adjustment throughout the process at acidic pH values by means of such acidophilic a-amylases and xylose isomerases. Takasaki et al. [2291 found an acidophilic a-amylase in a Bacillus licheni&mis strain isolated from soil, and showed that the enzyme is suitable for digestion of cornstarch at an acidic pH of 4.5-5.0. Acidophilic xylose isomerases have been demonstrated in Thermoanaerobacterium sp. JW/SL-YS[2171 and Streptomyces sp. SK[22'], and purified and characterized. Both ofthese have optimum pH values around G.5, but are highly active at acidic pHs such as 5.0. Since they are highly thermostable, they are expected to be useful for fructose production. 17.3.1.4

Production o f Unusual Sugar Derivatives

Xylose isomerase has a wide substrate specificity, and 3-, 5-, or 6-substituted glucose and fructose are isomerized by this enzyme. Since this enzyme requires the 4-OH group for hexoses to be substrates, phosphoglucose isomerase instead of xylose isomerase is used for the synthesis of kubstituted fructose as described below.

17.3.1.4.1

Preparation of Glucose Derivatives Modified at Position 3 or 6

Bock and coworkers [2301 showed that D-glucose derivatives bearing modifications at the C 3 or CG position are converted by xylose isomerase from Streptomyces sp.

17.3 lsomerizations

A

I

OH X=F X = N3

I

OH

OH

I

1317

Figure 17-23. Conversion by xylose isomerase of (ZR,3R)-configuredaldotetrose modified at C5 into open-chain 2-ketoses (A), and L-erythrose into L-erythrulose (B). Reprinted from Ebner and S t u t ~ [ ~ ~ ~ ] .

B

However, epimers of D-glucose are inert as substrates of the enzyme: D-mannose, Dallose, and D-galactose. Various 5-modified D-glucofuranoses are quantitatively converted into the corresponding D-fructopyranoseswith the enzyme [2311. Ebner and S t i i t ~ [ ~showed ~ * ] that various (2R,3R)-configuredaldofuranoses such as D-erythrose and CS-modified D-ribose derivatives serve as substrates of the enzyme: D-erythrose is quantitatively converted into D-glycero-tetrulose,with D-ribofuranoses being the corresponding open-chain 2-ketoses (Fig. 17-23).L-Erythrose, the enantiomer of Derythrose, is also isomerized quantitatively by the enzyme to L-erythrulose(L-glycerotetrulose) (Fig. 17-23).Fructose bisphosphate aldolase catalyzes a stereospecific aldol condensation between dihydroxyacetone phosphate and a number of aldehydes to form hexoketose 1-phosphates, the phosphate groups of which are removed by hydrolysis. The resultant hexoketoses are converted stereospecifically into hexoaldose derivatives by xylose isomerase. Thus, unusual hexoaldose derivatives such as 3-deoxy-~-glucose, 6-deoxy-~-glucose, 6-O-methyl-~-glucose and 6-deoxy-6-fluoroD-glucose were prepared by this method[223,2331.

17.3.1.4.2

Preparation o f Fructose and Sorbose Derivatives Modified at Position 5

Xylose isomerase converts a wide range of D-glucose as well as L-idose derivatives modified at position 5 into the corresponding ketose. 5-Deoxy-5-fluoro-D-xylulose and a variety of 5,6-dimodified open-chain analogs of D-fructose, namely the 5,6-diazido-S,G-dideoxy, 6-azido-S,6-dideoxy, 6-azido-5,6-dideoxy-5-fluoro, 5,6-diderivatives were deoxy-5-fluoro, 5.6-dideoxy-6-fluoro and 5,6-dideoxy-5,6-difluoro prepared with glucose isomerase (Fig. 17-24)[234, 23s1. 17.3.1.4.3

Preparation of Sucrose Derivatives with Modified Fructose Moieties

Xylose isomerase is also used for the synthesis of modified sucroses, which is important in the study of the topographical aspects of the binding of sucrose to a sucrose carrier protein [23G1. 6-Deoxy- and 6-deoxy-6-fluoroglucosechemically synthesized are isomerized to the corresponding 6-substituted fructose by xylose isomerase. The resultant substrates are subsequently condensed with UDP-glucose by sucrose synthase. Although the equilibrium of the first step lies towards the glucose

1318

I

77 fsomerizations

F

A

w

OH

o

H

h

F

+

o

0

H OH

OH

OH 0 B

OH

Y + O H

OH

X

OH

X = H, Y = F X = H, Y = N3 X=Y=F X = F, Y = N3 X = F, Y = H X=Y=N3 X=Y=H Figure 17-24. Production of 5-deoxy-5-fluoro-o-xylulose and 5,6-dimo. dified open-chain analogs o f D-fructose with xylose isomerase. Reprinted from Hadwiger et al.[235].

derivatives, this problem is overcome by coupling the isomerization reaction with the sucrose formation, which is irreversible. The second reaction completely drives the isomerization reaction almost to completion. Incubation of 6-deoxy- or 6-deoxy6-fluoroglucose and UDP-glucose with both the xylose isomerase and sucrose synthase afforded 6'-deoxy- and 6'-deoxy-6'-fluorosucrose in 73 and 53 % isolated yield, respectively. 17.3.2 Phosphoglucose Isomerase (E.C. 5.3.1.9)

Phosphoglucose isomerase catalyzesthe interconversion of glucose 6-phosphate and fructose 6-phosphate. This enzyme is involved in the gluconeogenesis, glycolytic pathway, and pentose phosphate cycle. Since thermostable enzymes are generally useful for industrial application, thermostable phosphoglucose isomerase was purified from Bacillus stearothem~philus[~~'] and Bacillus ~ a l d o t e n a d ~B. ~ ~stearl. othemophilus produces two isozymes of phosphoglucose isomerase, and they were overexpressed in E. coli, purified to homogeneity, crystallized[239], and the X-ray structure of the enzyme was 2411 . The structure of the rabbit muscle enzyme complexed with a competitive inhibitor D-gluconate 6-phosphate was also 2431. The enzyme is a dimer with two a/Pdetermined by X-ray crystallography[242, sandwich domains in each subunit. Lys 518 and His 388 are located at the active center and are probably involved in the catalytic mechanism. Since gluconate 6-phosphate occurs predominantly in its cyclic form, phosphoglucose isomerase probably catalyze the opening of the hexose ring to give initially its straight chain form with Lys 518 and His 388. Then the enzyme undergoes isomerization of the

I

17.3 lsornerizations 1319

Arg 272

Arg 272

OH

H

/"

phosphate

I

I

I

glucose-6-

Arg 272

O V 0

I

His 3 8 8 - x y N H

O Y O Glu 357

I

Glu 357

Figure 17-25. Mechanism o f phosphoglucose isomerase reaction. His 388 and Clu 216 catalyze the ring opening. The side-chain o f Glu357 abstracts a proton from the C2 position of the open chain form ofthe substrate, and the cis-

Glu 357

fructose-6-phosphate

enediol is formed. Then, a proton is transferred from the protonated Clu 357 to the C1 position ofthe intermediate. Reprinted from Jefferyet ai. [2421.

substrate through formation of a cis-enediol intermediate with the double bond between C 1 and C2 (Fig. 17-25). Glu 357 transfers the proton from the C2 of glucose 6-phosphate to its C1 position. The side chain of Arg 272 stabilizes the negative charge of the intermediate (Fig. 17-25). Xylose isomerase requires the 4-OH group for glucose derivatives to be substrates [2301. On the other hand, phosphoglucose isomerase can act on 4-substituted phosphoglucose. Therefore the latter enzyme is applicable to the preparation of glucose or fructose derivatives modified at position 4. For example, 4-deoxy4-fluorofructose was prepared from 4-deoxy-4-fluoroglucosewith phosphoglucose isomerase because xylose isomerase cannot isomerize 4-deoxy-4-fluoroglucose[23G1. 4-Deoxy-4-fluorofructosewas then converted into 4'-deoxy-4'-fluorosucrose, which is useful for the analysis of the interaction between sucrose and a sucrose carrier protein, with fructose-6-phosphatekinase [2361. Fructose 1,6-bisphosphatehas attracted attention due to its important applications in the field of medicine, and is produced from glucose in three step by enzymatic reactions catalyzed by glucokinase, phosphoglucose isomerase, and phosphofructokinase. ATP is regenerated by acetate kinase (Fig. 17-26). Ishikawa and coworkers established an efficient method for production of fructose 1,G-bisphosphate in a Glucose + ATP PGI G6P -2F6P + ATF'

GK

Glucose-6-phosphate (G6P)

Fructose-6-phosphate (F6P)

PFK

FDP + ADP

AK

ADP + Acetyl phosphate

ATP + Acetic acid

+

ADP

Figure 17-26. Synthesis o f fructose 1,6bisphosphate from glucose by combination o f glucokinase (CK), phosphoglucose isomerase (PCI), phosphofructokinase (PFK), and acetate kinase (AK) reactions.

1320

I

17 lsomerizations

0

O H ,k,P O -.O .~

-

Triosephosphate isomerase L

Ix,

QH !

opog-

D-Glyceraldehyde 3-phosphate

Dihydroxyacetone phosphate Glu 165

o,

Figure 17-27. Reaction catalyzed by triosephosphate

isomerase.

Glu 165

$

-0

Glu 165

I

Glu 165

i

0

\o

Glu 165

x I

c? ' 0

Figure 17-28. Triosephosphate isomerase reaction through a cis-enediol intermediate. The pro-R proton is removed from C1 o f dihydroxyacetone phosphate by the side chain of Clu 165, and the carbonyl group o f t h e substrate is polarized by the side chain of His 95. Reprinted from Harris et a~.[*49~.

batch reactor system using the purified enzymes[244] and the crude extract of Bacillus The yield of fructose 1,G-bisphosphatedepended on the stearothermophilus cells[245]. activity of glucokinase in the reactor[246]. 17.3.3 Triosephosphate lsomerase (E.C. 5.3.1.1)

Triosephosphate isomerase is involved in the glycolybc pathway, and catalyzes the interconversion of dihydroxyacetone phosphate and D-glyceraldehyde phosphate (Fig. 17-27).The refined three-dimensional structures of chicken, yeast, and trypano-

I

17.3 lsornerizations

1321

OH

OH

Figure 17-29. Synthesis of [3',4'-''Cz]-thymidine from [2',3'-'3Cz]-dihydroxyacetone phosphate with triosephosphate isomerase (TPI) and D-2-deoxyribose-5-phosphate(DHAP). Asterisks indicate the positions selectively labeled with "C. Other positions that can be isotopically substi. tuted are marked with ', 4and 0.Reprinted from Ouwerkerk et al.[2511.

soma1enzymes have been elucidated[247]. The reaction is thought to proceed through a cis-enediolintermediate with Glu 165 and His 95 as acid and base catalysts (Fig. 1728)[248,24ql. The side chain of Glu 165 removes the pro-R proton from the C1 of dihydroxyacetonephosphate, and that of neutral His 95 polarizes the carbonyl group of the substrate. Fructose 1,6-bisphosphate, a precursor molecule for sugar synthesis, can be prepared from dihydroxyacetone phosphate with this enzyme and ald0lase1~~~1. Triosephosphate isomerase has been used for various other purposes. For example, [3',4'-'3C2]-thymidinehas been prepared from [13C2]-aceticacid through [2',3'-'3C2]-dihydroxyacetonephosphate and ~-[3',4'-'~C2]-2-deoxyribose5-phosphate with triosephosphate isomerase and D-2-deoxyribose-5-phosphate aldolase (E.C. 4.2.1.2) (Fig. 17-29)12511. 17.3.4 L-Rharnnose Isomerase (E.C. 5.3.1.14)

L-Rhamnose is an important component ofbacterial cell walls, and is metabolized in E. coli through a pathway similar to that of glucose 6-phosphate in glycolysis. Rhamnose isomerase catalyzes the first reaction in the pathway to produce Lrhamnulose from t-rhamnose (Fig. 17-30).The enzyme gene was cloned from E. coli and o~erexpressed[~~*], and the enzyme was purified and Rhamnose isomerase is composed of four identical subunits with a molecular mass of about 47 kDa. It has the maximum activity around 7.6, and requires Mn2' to provide the highest activity. The enzyme shows no significant sequence similarity to any other ketol isomerases including xylose isomerase. However, rhamnose isomerase was found, by X-ray crystallography, to be most similar to xylose isoof rhamnose isomerase is composed of (P/a)s-barrels, m e r a ~ e l ~The ~ ~monomer ], and the structure and arrangement of the barrel are very similar to those of xylose isomerase. However, each of them has an additional a-helical domain, which is involved in subunit assembly and differs from each other only in its structure. The

1322

I

17 lsornerizations

L-R hamnose (6-Deoxy-L-mannose)

L-Rhamnulose (6-Deoxy-L-fructose)

Figure 17-30. Reaction catalyzed by rhamnose isomerase. Since both substrate and product occur in cyclic forms, L-rhamnose isomerase catalyzes ring opening before isomerization. Reprinted from Korndorfer et aI.[2521.

Figure 17-31. Superposition o f the metal binding sites o f rhamnose isomerase (residues named and drawn with thick bonds) and zylose isomerase (thin bonds). Reprinted from Korndorfer et al. [2521.

residues surrounding the catalyix Mn2+ site (Asp 302, Asp 304 and His 270) are conserved in the two structures (Fig. 17-31). Therefore, the reaction catalyzed by rhamnose isomerase is thought to proceed through a metal-mediated hydride-shift mechanism in the same manner as xylose isomerase [2521. Bhuiyan et al. 12531 immobilized L-rhamnose isomerase from Pseudomonas sp. LL172 on chitopearl beads, and used it to produce L-mannose from L-fructose. The immobilized enzyme was found to be stable: it retained about 90% of the initial activity after five repeated batch reactions. The concentration of L-mannose relative to L-fructose was about 3:7 at equilibrium. D-Allose was also produced from Dpsicose with the immobilized L-rhamnose isomerase. Since D-psicose is readily produced from D-fructose with D-tagatose 3-epimerase, D-allose can be produced from D-fructose by combination of the two enzymes immobilized on chitopearl beads. Bhuiyan et al. [2541 found that the reaction progresses steadily until 40% of the D-psicose is converted into D-allose. The immobilized D-tagatose 3-epimerase was also stable even after repeated uses, and D-allose was produced efficiently in the system.

77.3 lsornerizations

I

1323

17.3.5 L-Fucose Isomerase (E. C. 5.3.1.3)

Fucosylated oligosaccharides are important components of glycoproteins and glycolipids which are useful for cancer diagnosis and immunotyping. Therefore, efficient production methods for L-fucose and its analogs would be useful. L-Fucose isomerase acts on D-arabinose, which was known as D-arabinose isomerase in earlier literatures. L-Fucose is metabolized through a pathway similar to that of D-glucose in glycolysis, and L-fucose isomerase corresponds to glucose 6-phosphate isomerase. However, none of the aldose-ketose isomerases including glucose 6-phosphate isomerase shows sequence similarity to L-fucose isomerase. LFucose isomerase shares the common characteristics with other aldose-ketose isomerases acting on unphosphorylated substrates: the requirement of metal ions such as Mn2+for L-fucose isomerase. Aldose-ketose isomerases acting on phosphorylated substrates generally require no metal ions with the exception of phosphomannose isomerase (E. C. 5.3.1.8) which requires Zn" for its activity. Seemann and Schulz [2551 determined the three-dimensional structure of L-fucose isomerase from E. coli, a hexamer from a subunit with a molecular mass of 64 976 Da. The enzyme shows no structural similarity to any other aldose-ketoseisomerases analyzed thus far. However, Seemann and Schulz, on the basis of the tertiary structure, suggested that the r-fucose isomerase reaction proceeds through an enediol intermediate [2551. Fessner et a1.[2561 developed an efficient method for the synthesis of L-fucose analogs modified at the nonpolar terminus by means of L-fucose isomerase and Lfuculose 1-phosphatealdolase from E. coli. Various L-fucose analogs bearing linear or branched aliphatic side chains were prepared in about 30% overall yield with hydroxyaldehyde precursors and dihydroxyacetonephosphate as the starting materials (Fig. 17-32). 0

OH

R'&o

3

+ HO&OPOi:-

R2

RL

op0:-

R2 R'

OH

-

R2

Fucl

%OH

R'

OH

OH

R2

R@ ' "' OH HO OH

Figure 17-32. Enzymatic synthesis o f L-fucose analogs with L-fucose 1-phosphate aldolase (FucA), phosphatase (P'ase), and L-fucose isomerase (Fucl). Reprinted from Fessner et al.[2561.

R'

R2

CH3 CH2-CH3 CH=CHz CECH

H H H H

CH3 CF3

CH3 H

OH

1324

I

77 lsomerizations 0

ACOOH

\

J"""

HO

Neu5Ac

2-epimerase

Ho& HO

AcNH OH GicNAc Synthesis o f N-acetylneuraminate (Neu5Ac) from N-acetylD-glucosamine (ClcNAc) and pyruvate through N-acetyl-D-mannosamine (ManNAc) with N-acetylneuraminate and N-acetyl-D-glucosamine2-epimerase. Reprinted from Maru et aI.[259]. Figure 17-33.

17.3.6 N-Acetybglucosamine 2-Epimerase

N-Acetylneuraminate is a sialic acid with various biological functions that is widely distributed in animals. It has been prepared only from natural resources such as colominic acid, edible birds nests, milk or eggs. Alternatively, it has been prepared enzymatically from N-acetyl-D-mannosamine and pyruvate with N-acetylneuraminate lyase as the catalyst[257,25s1.However, N-acetyl-D-mannosamine is expensive, and the method is not suitable for large-scale production of N-acetylneuraminate. Maru et a1.[2591developed an elegant method for the enzymatic production of Nacetylneuraminate from the inexpensive N-acetyl-D-glucosamine and pyruvate by means of N-acetylneuraminate lyase and N-acyl-D-glucosamine2-epimerase, whose genes were cloned from E. c0li[~"1 and pig kidney[261],respectively (Fig. 17-33). Simultaneous use of these enzymes and feeding of appropriate amounts of pyruvate to the reaction mixture enabled production of N-acetylneuraminate from N-acetyl-Dglucosamine with a 77% conversion rate, and 29 kg of N-acetylneuraminate were obtained from 27 kg of N-acetyl-D-glucosamine. 17.3.7 Maleate cis-trans lsomerase (E. C. 5.2.1.1)

Maleate cis-trans isomerase catalyzes the conversion of maleate into fumarate. This enzyme is applicable to the production of L-aspartateby coupling with the aspartase reaction as shown in Fig. 17-3412", 2631.First, maleate is isomerized to fumarate by

17.3 lsornerizations

Maleate cis-trans isomerase

OOCHCOO

coo

I

1325

Aspartase O O C ~ c o o

H

H

-OOCHH

Maleate Figure 17-34.

NH4

Fumarate

_NH3 L-Aspartate

Synthesis of L-aspartate using maleate cis-trans isomerase and aspartase.

cis-trans isomerase, and then the fumarate formed is aminated to L-aspartate by aspartase. In this procedure, the resting cells of Alcaligenesfaecalis containing both enzymes can be used as a catalyst. Thermostable maleate cis-trans isomerase was purified from Bacillus stearothermophilus MI-102 and characterized, and the enzyme gene was cloned and Two cysteine residues, Cys 80 and Cys 198, among the three conserved cysteines were found by site-directed mutagenesis studies to be catalytically important, although their catalytic roles are not yet known. 17.3.8 Unsaturated Fatty Acid cis-trans lsomerase

trans-Unsaturated fatty acids occur in membrane phospholipids of some bacterial genera such as Pseudomonas and Vibri~[~"].They are produced by cis-trans isomerase from cis-unsaturated fatty acids in response to environmental stresses such as elevated temperatures, increased salt concentrations, and the presence of organic The structural gene for the cis-transisomerase was solvents such as toluene [2662691. cloned from Pseudomonas putida P8[270].The E. coli recombinant cells carrying the gene were shown to produce trans-unsaturated fatty acids in response to the organic solvent, although E. coli has no inherent ability to produce these fatty acids [2701. Okuyama et a1.1271]purified the cis-trans isomerase from Pseudomonas sp. E-3 and characterized the enzyme catalyzing cis-trans isomerization toward 9-hexadecenoate. It catalyzes the cis-to-transconversion of a double bond of cis-mono-unsaturatedfatty acids with carbon chain lengths of 14, 15, 16, and 17 at positions 9,10, or 11,but not at 6 or 7: the enzyme shows a strict specificity for both the position of the double bond and the chain length of the fatty acid. A similar enzyme was also discovered by Witholt and coworkers, which was purified from the periplasmic fraction of Pseudomonas oleovorans[2721. Not only 9-cis-hexadecenoatebut also 1l-cis-octadecenoate were found to serve as substrates of the enzyme. Moreover, the enzyme acted only on free unsaturated fatty acids and not on esterified fatty acids in contrast to the enzyme from Pseudomonas sp. E-3.Therefore, the Pseudomonas oleovorans enzyme differs from the enzyme of Pseudomonas sp. E-3 in substrate specificity, although both are monomeric enzymes with a molecular mass of about 80 kDa. The cis-trans isomerases are expected to be useful for biotransformation of unsaturated fatty acids.

1326

I

77 lsornerizations

17.4

Conclusion

Total conversion of racemic starting materials into a particular stereoisomer of a desired compound is very useful in the chemical industry. Half or more of the starting materials can be saved and steps for the laborious separation of the products from the starting material remaining reduced. Thus, racemases and epimerases are very useful in the chemical industry, when their reactions are coupled with some stereospecific reactions. Isomerases are also powerful catalysts for the production of particular enantiomers or diastereomers of interest from cheaply-availablestarting materials especially in the field of carbohydrate chemistry. Various new racemases and isomerases useful for industrial applications will no doubt be discovered from microorganisms at some point. However, established and well-known enzymes can be remodeled in order to expand their uses by various protein engineering technologies such as directed evolution. A good example for this is L-specific hydantoinase derived from D-specific hydantoina~e('~~1. The engineered enzymes can be incorporated into metabolic engineering studies in order to develop powerful microbial cells. References H. Katagiri, K. Kitahara,J. Agr. Chem. Soc. Jpn. 1936,12,844. 2 H. Katagiri, K. Kitahara, Biochem. J. 1937, 1

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66, 245. 191 192

Y.-C. Tsai, C.-P. Tseng, K.-M. Hsiao, L.-Y. Chen, Appl. Environ. Microbiol. 1988, 54,

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Tokuyama, K. Hatano, Appl. Microbiol. Biotechnol. 1995, 42, 853. S. Tokuyama, K. Hatano, Appl. Microbiol. Biotechnol. 1995, 42, 884. S. Tokuyama, K. Hatano, Appl. Microbiol. BiotechnoL 1996, 44, 774. D. R. J. Palmer, J. B. Garrett, V. Sharma, R. Meganathan, P. C. Babbitt, J. A. Gerlt, Biochemistry 1999,38,4252.

174 S. 175 176 177

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1997, 56, 33.

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U. T. Strauss, A. Kandelbauer, K. Faber, Biotechnol. Lett. 2000, 22, 515. S. C. Ransom, J. A. Gehlt, V. M. Powers, G. L. Kenyon, Biochemistry 1988,27, 540. D. J. Neidhart, P. L. Howell, G. A. Petsko, V. M. Powers, R. Li, G. L. Kenyon, J. A. Gerlt, Biochemistry 1991, 30, 9264. D. J. Neidhart, G. L. Kenyon, J. A. Gerlt, G. A. Petsko, Nature 1990,347,692. P. C. Babbitt, G . T. Mrachko, M. S. Hasson, G. W. Huisman, R. Kolter, D. Ringe, G. A. Petsko, G . L. Kenyon, J. A. Gerlt, Science 1995,267,1159.

J. A. Landro, A. T. Kallarakal, S. C. Ransom, J. A. Gerlt, J. W. Kozarich, D. J. Neidhart, G. L. Kenyon, Biochemistry 1991,30,927. 198 B. Mitra, A. T. Kallarakal, J. W. Kozarich, J. A. Gerlt, 1. G. Clifton, G. A. Petsko, G. L. Kenyon, Biochemistry 1995, 34, 2777. 199 J. A. Landro, J. A. Gerlt, J. W. Kozarich, C. W. Koo, V. J. Shah, G. L. Kenyon, D. J. Neidhart, S . Fujita, G. A. Petsko, Biochemistry 1994, 33, 635. 200 A. T. Kallarakal, B. Mitra, J. W. Kozarich, J. A. Gerlt, G. L. Kenyon, J. G. Clifton, G . A. 197

References I1331 Petsko, G. L. Kenyon, Biochemistry 1995, 34, 2788. 201 S. L. Schafer, W. C. Barrett, A. T. Kallarakal, B. Mitra, J. W. Kozarich, J.A. Gerlt, J. G. Clifton, G. A. Petsko, G. L. Kenyon, Biochemistry 1996,35,5662. 202 S. L. Bearne, R. Wolfenden, Biochemistry 1997,36,1646. 203 K. A. Briggs. W. E. Lancashire, B. S. Hartley, EMBO]. 1984,3,611. 204 K. Dekker, H. Yamagata. K. Sakaguchi, S. Udaka, Agnc. Bid. Chem. 1991,55, 221. 205 K. Dekker, A. Sugiura. H. Yarnagata, K. Sakaguchi, S. Udaka, AppI. Microbiol. Biotechnol. 1992, 36, 727. 206 S. D. Feldmann, H. Sahrn, G. A. Sprenger, Mol. Gen. Genet. 1992, 234, 201. 207 K. Yamanaka, Biochim. Biophys. Actu 1968, 151, 670. 208 Y. Takasalu, Y. Kosugi, A. Kanbayas, Agnc. Biol. Chem. 1969,33,1527. 209 N. Murarnatsu, Y. Nosoh, Arch. Biochem. Biophys. 1971, 144,245. 210 C. S. Gong, L. F. Chen, G. T. Tsao, Biotechnol. Bioeng. 1980, 22, 833. 211 P. Kristo, R. Saarelainen, R. Fagerstrorn, S. Aho, M. Korhola, Eur. ]. Biochem. 1996,237, 240. 212 S. H. Brown, C. Sjoholm, R. M. Kelly, Bitechnol. Bioeng. 1993,41, 878. 213 C. Lee, J.G. Zeikus, Biochem.]. 1991,273, 565. 214 K. Dekker, H. Yamagata, K. Sakaguchi, S. Udaka, ]. Bacteriol. 1991, 173, 3078. 215 J.Chauthaiwale, M. Rao, Appl. Environ. Microbiol. 1994, 60, 4495. 216 C. Vieille, J. M. Hess, R. M. Kelly, J. G. Zeikus, Appl. Environ. Microbiol. 1995, 61, 1867. 217 S. Liu, J. Wiegel, F. C. Gherardini,]. Bacteriol. 1996, 178, 5938. 218 S. S. Deshrnukh, V. Shankar, Biotechnol. Appl. Biochem. 1996, 24, 65. 219 C. J. Moes, I. 4. Pretorius, W. H. Van-Zyl, Biotechnol. Lett. 1996, 18, 269. 220 B. C. Park, S. Koh, C. Chang, S. W. Shu, D. S. Lee, S. hd. Byun, Appl. Biochem. Biotechnol. 1997, 62, 15. 221 B. K. Srih, S. Bejar, Biotechnol Lett. 1998, 20, 553. 222 C. Chang, H. K. Song, B. C. Park, D. S. Lee, S. W. Suh, Acta Crystallogr. Sect D Bid. Crystallogr. 1999, 55,294.

R. Dumvachter, H. M. Sweers, K. Nozaki, C. H. Wong, Tetrahedron Lett. 1986,27, 1261. 224 C. A. Collyer, K. Henrick, D. M. Blow, ]. Mol. Bid. 1990, 212, 211. 225 M. Fuxreiter, 8.Farkas, G. Niray-Szabo, Protein Eng. 1995, 925. 226 I. A. Rose, Philos. Trans. R. Soc. London, Ser. B 1981,293,131. 227 V. J.Jensen, S. Rugh, Methods Enzymol. 1987,136,356. 228 M. Meng, M. Bagdasarian, J. G. Zeikus, Biotechnology 1993, 11, 1157. 229 Y. Takasaki, S. Furutani, S. Hayashi, K. Irnada, ]. Ferment. Bioeng. 1994,77,94. 230 K. Bock, M. Meldal, B. Meyer, L. Wiebe, Acta Chem. Scand., Ser. B 1983, 37,101. 231 A. Berger, A. de Raadt, G. Gradnig, M. Grasser, H. Low, A. E. Stiitz, Tetrahedron Lett. 1992, 33, 7125. 232 M. Ebner, A. E. Stiitz, Carbohydr. Res. 1998, 305, 331. 233 J. R. Dunwachter, D. G. Drueckharnrner, K. Nozaki, H. M. Sweers, C. H. Wong,]. Am. Chem. SOC.1986,108,7812. 234 A. Berger, A. de Raadt, G. Gradnig, M. Grasser, H. Low, A. E. Stutz, Tetrahedron Lett. 1992, 33, 7125. 235 P. Hadwiger, P. May, B. Nidetzky, A. E. Stutz, A. Tauss, Tetrahedron: Asymmetry 2000, 1 1 , 607. 236 P. J. Card, W. D. Hitz, K. G. Ripp,]. Am. Chem. Soc. 1986, 108, 158. 237 N. Murarnatsu, Y. Nosoh, Arch. Biochem. Biophys. 1971, 144, 245. 238 M. Takama, Y. Nosoh, ]. Biochem. 1980,87, 1821. 239 C. D. Hsiao, C. C. Chou, Y. Y. Hsiao, Y. J. Sun, M. Meng,J. Structural Bid. 1997, 120, 196. 240 Y. J. Sun, C. C. Chou, W. S. Chen, R. T. Wu, M. Meng, C. D. Hsiao, Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 5412. 241 C. C. Chou, Y. J.Sun, M. Meng, C. D. Hsiao, ]. Biol. Chem. 2000,275,23 154. 242 C. J. Jeffery, B. J. Bahnson, W. Chien, D. Ringe, G. Petsko, Biochemistry 2000, 39, 955. 243 C. J. Jeffery, R. Hardre, L. Salmon, Biochemistry 2001,276, 1560. 244 A. Widjaja, M. Shirishima, M. Yasuda, H. Ogino, H. Nakajirna, H. Ishikawa,]. Biosci. Bioeng. 1999, 87, 611. 245 A. Widjaja, M. Yasuda, H. Ogino, H. Naka223 J.

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I. Maru, J. Ohnishi, Y. Ohta, Y. Tsukada, Curbohydr. Res. 1998,306,575. 260 Y. Ohta, Y.Tsukada, T. Sugimori, K. Murata, A. Kimura, Agric. B i d . Chem. 1989,53, 477. 261 I. Maru, Y. Ohta, K. Murata, Y. Tsukada,]. Biol. Chem. 1996,271,16294, 262 Y. Takamura, I. Kitamura, M. Iikura, K. Kono, A. Ozaki, Agric. Bid. Chem. 1966, 30, 338. 263 Y. Takamura, I. Kitamura, M. Iikura, K. Kono, A. Ozaki, Agric. Bid. Chem. 1966,30, 345. 264 K. Hatakeyama, M. Goto, Y. Uchida, M. Kobayashi, M. Terasawa, H. Yukawa, Biosci. Biotechnol. Biochem. 2000, 64, 569. 265 H. Keweloh, H. J. Heipieper, Lipids 1996, 31, 129. 266 N. Morita, A. Shibahara, K. Yamamoto, K. Shinkai, G. Kajimoto, H. Okuyama,]. Bucteriol. 1993, 175, 916. 267 R. Diefenbach, H. Keweloh, Arch. Microbiol. 1994,162,120. 268 Q. Chen, D. B. Janssen, B. Withoh,/. Bacteriol. 1995, 177, 6894. 269 H. J. Heipieper, G. Meulenbeld, Q. van Oirschot, J. A. M. de Bont, Appl. Environ. Microbiol. 1996, 62, 2773. 270 R. Holtwick, F. Meinhardt, H. Keweloh, Appl. Environ. Microbiol. 1997, 63, 4292. 271 H. Okuyama, A. Ueno, D. Enari, N. Morita, T. Kusano, Arch. Microbiol. 1998, 169, 29. 272 V. Pedrotta, B. Witholt,]. Bacterial. 1999, 181, 3256. 259

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyriqht 0Wiley-VCH Verlaq GmbH, Weinheim 2002

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18 Introduction and Removal of Protecting Groups Dieter Kadereit, Reinhard Reents, Duraiswamy A.Jeyaraj and Herbert Waldmann

18.1 Introduction

The proper introduction and removal of protecting groups is one of the most important and widely carried out synthetic transformation in preparative organic chemistry. In particular, in the highly selective construction of complex, polyfunctional molecules, e. g. oligonucleotides, oligosaccharides, peptides and conjugates thereof, and in the synthesis of alkaloids, macrolides, polyether antibiotics, prostaglandins and other natural products, regularly the problem arises that a given functional group has to be protected or deprotected selectively under the mildest conditions and in the presence of functionalities of similar reactivity, as well as in the presence of structures that are sensitive to acids, bases, oxidation and reduction. Numerous classical chemical methods have been developed for the manipulation of protecting groups [1-31. Nevertheless, severe problems still remain caused by the need to introduce or remove selectively specific blocking functions which can not, or only with great difficulties,be solved by using classical chemical tools only. However, the arsenal of the available protecting group techniques has been substantially enriched by the application of biocatalysts. In addition to their stereodiscriminating properties, enzymes offer the opportunity to carry out highly chemo- and regioselective transformations. They often operate at neutral, weakly acidic or weakly basic pH values and in many cases combine a high selectivity for the reactions they catalyze and the structures they recognize with a broad substrate tolerance. Therefore, the application of these biocatalysts to effect the introduction and/or removal of suitable protecting groups offers viable alternatives to classical chemical methods ["I.

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18 Introduction and Removal of Protecting Groups

18.2 Protection of Amino Croups16121 18.2.1

N-Terminal Protection of Peptides

The selective protection and liberation of the a-amino function, the carboxy group and the various side chain functionalities of polyfunctional amino acids constitute some of the most fundamental problems in peptide chemistry. Consequently, numerous efficient protective functions based on chemical techniques have been 13, 141 However, since the mid-1970s,a developed to a high level of practicability. systematic search for blocking groups being removable with a biocatalyst has been carried In addition to the mild deprotection conditions they promise, protecting groups of this type are expected to be particularly useful for the construction and manipulation of larger peptide units, i. e. for transformations which, for solubility reasons, in general have to be carried out in aqueous systems. Also applications in the reprocessing of peptides obtained by recombinant DNA technology are foreseen (for an interesting appropriate example see Chapter 12.5). Initial attempts to introduce an enzyme-labile amino protecting group involved the use of chymotrypsin for the removal of N-benzoylphenylalanine(Bz-Phe)from the tripeptide Bz-Phe-Leu-Leu-OH("1. The desired dipeptide H-Leu-Leu-OH was obtained in 80% yield under mild conditions (pH 7.3, room temperature). Chymotrypsin, however, is an endopeptidase with a rather broad substrate tolerance, catalyzing the hydrolysis of peptide bonds on the carboxy groups of hydrophobic and of aromatic amino acid residues. Since such amino acids appear widely in peptides, and since no method is available to protect them against attack by the enzyme during the attempted deprotection, the use of chymotrypsin is problematic. Its use is therefore limited to special cases [16] in which no danger of competitive cleavage at undesired sites has to be feared. A protease of much narrower specificity is trypsin which catalyzes the hydrolysis of peptide bonds at the carboxylic group of lysine and arginine. These amino acids carry polar, chemically reactive side chain functional 141. The high specificity of groups which can be protected by various techniques trypsin together with the possibility of hiding the critical amino acids which function as primary points of tryptic cleavage allowed for the development of a broadly applicable system for the protection of the a-amino group of peptides [12, l7-l91. I n several studies the application of trypsin-labileprotecting groups, along with suitable blocking functions for the side chains of arginine and lysine were d e ~ c r i b e d [ l ~ - ~ ~ ] . Thus, for instance Z-Arg-OH served as the enzymatically removable protecting group in a stepwise synthesis of deamino-oxytocin 1 (Fig. 18-1)[18, 191. Starting with a pentapeptide the amino acid chain was elongated with Z-Argprotected amino acid p-nitrophenyl esters. The N-terminal Z-Arg protecting group was successively removed in moderate to high yield and without attack on the other peptide bonds by treatment with trypsin. Unfortunately, the preparation of the protected arginine p-nitrophenyl esters is difficult,thus preventing this method from becoming generally useful for the stepwise assembly of larger peptides. The trypsin-

18.2 Protection ofArnino Groups

I

1335

H-Asn-Cys(Acm)-Pro-Leu-Gly-NH2

I

1) 2-Arg-AA-ONp 2) trypsin

ONp =

t

(iterate)

I Mpr -fffxf%tAsn-Cys-Pro-Leu-Gly-NH2

-s-s1

deamino-oxytocin

Bz-GIy-His-He-Glu BLeu-AspmTyr-Thr-Cys(Acm)-NHEt 2

21-31 fragment of murine epidermal growth factor

n=

N-terminally deprotected by enzymatic removal of Z-Arg (1) or Bz-Arg (2) with trypsin

Construction of oligopeptides via removal of N-terminal arginine residues with trypsin. Figure 18-1.

labile blocking groups have, however, proven to be very useful for the construction of oligo- and polypeptides via condensation of preformed peptide fragments. An illustrative example consists of a chemoenzymatic construction of the 21-31 fragment 2 of murine epidermal growth factor (Fig. 18-1). In the course of this synthesis the deblocking by trypsin was applied twice[1G]. The enzyme first liberated the N-terminus of a tetrapeptide and subsequently of a heptapeptide. In a synthesis [241 of human p-lipotropin an Ac-Arg-residue was introduced by a solid-phase technique at the N-terminus of the 29 C-terminal amino acids of the desired polypeptide. After cleavage from the resin and protection of the side chain functionalities, the arginine moiety was removed with trypsin, leaving the peptide chain intact. Finally, coupling of this 61-89 fragment to a partially protected 1-GO segment, and subsequent deprotection delivered P-lipotropin. Further examples are found in syntheses of oxypressin [I2], Met-enkephalin[251 and Glu4-oxytocin In addition to chymotrypsin and trypsin, the collagenase from Clostridiurn histolyticum has been proposed as a catalyst for the removal of N-terminally attached dummy amino acids from peptides [26]. The enzyme recognizes the tetrapeptides Pro-X-GlyPro and cleaves the X-Gly bond. The use of this biocatalyst permitted the construction of des-pyroglutamyl-[15-leucine]humanlittle gastrin I by selective hydrolysis of the dipeptide Pz-Pro-Leu (Pz = 4-phenylazobenzyloxycarbonyl)from the N-terminus of the octadecapeptide Pz-Pro-Leu-Gly-Pro-Trp-Leu-(Glu)s-Ala-Tyr-Gly-Trp-Leu-AspPhe-NH2. Transformations of this type are analogous to the naturally occuring conversion of prohormones into hormones and may prove to be useful for the processing of peptide factors produced by recombinant DNA technology.

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78 lntroduction and Removal offrotecting Croups

Despite the impressive syntheses that have been made possible using proteases, the use of these enzymes is always accompanied by the danger of a competitive (and sometimes unexpected and unforeseeable) cleavage of the peptide backbone at an undesired site. At a minimum, complex protecting group schemes may become necessary if the amino acid which serves as the recognition structure for the protease occurs several times in the peptide chain to be constructed. This disadvantage can be overcome if a biocatalyst devoid of peptidase activity is used for the liberation of the N-terminal amino group. This principle has been illustrated by the application of in industry for the large scale synthesis of penicillin G acylase from E. c0li[~~-@1 semisynthetic penicillins and by using a phthalyl imidase from Xanthobacter agilis[45-473(vide infiu).Penicillin G acylase attacks phenylacetic acid (PhAc)amides and esters but does not hydrolyze peptide bonds. The acylase accepts a broad range of protected peptides as substrates and selectively liberates the N-terminal amino group under almost neutral conditions (pH 7-8, room temperature) leaving the amide bonds as well as the C-terminal methyl, allyl, benzyl and tert-butyl esters unaff e ~ t e d [ ~ ~381.- ~ The ' , PhAc group is easily introduced into amino acids by chemicall4'I or enzymatic[49]methods and is stable during the removal of the C-terminal protecting groups employed [29-321. Recently, it has been shown that a phthalyl amidase isolated from Xanthobacter agilis is able to deprotect a variety of phthalimido substrates once the substrates are partially hydrolyzed to their monoacids (Fig. 18-2)[45-471. The phthalyl group is commonly used for amine protection, because it completely blocks this functionality by double acylation[2s '1. The enzymatic phthalyl removal proceeds via a two step process ofweakly basic hydrolysis to yield the monoacid 4 and subsequent treatment with the phthalyl amidase (Fig. 18-2).Because the hydrolysis of the phthalimide 3 to the corresponding monoacid 4 can be catalyzed by imidases such as the rat liver imidase,I'[ this procedure in particular represents a powerful alternative to the classical phthalyl deprotection which requires relatively drastic conditions and toxic reagents. However, the general applicability of the enzymatic phthalyl removal is yet to be investigated. If the construction of PhAc- or phthalyl-peptides is carried out by chemical activation of the PhAc-amino acids, the application of the non-urethane blocking 301. However, this disadvantage can be group results in ca. 6% racernizati~n[~', overcome by forming the peptide bonds enzymatically,e. g. with tryp~in['~I, chymotryp~in['~] or carboxypeptidase Y [39, "1, or by using urethane-type protecting groups (vide infa). For such condensation reactions and the subsequent enzymatic removal ofthe PhAc group, a continuous process was developed which has the potential to be transferable to a larger scale [391.

~1

@N-(R2

3

0

Figure 18-2.

-@&;Base

0

CH&N/H20

or 0.2 M buffer (PH 8.0)

Phthalyl arnidase

C02H

C02H +

R'

O

R' HWR2

y 2

R

4

Enzymatic removal of the phthalyl group.

5

6

78.2 Protection of Amino Croups

s-s PhAC-Gly

I

PhAc-Phe

n=

I

?S

?S

OH Lys-OH

I

7

(PhAc),insulin

8

leucine enkephalin

PhAcHN

N-terminally deprotected using penicillin G acylase

Is-sl

Mpr-Tyr-Phe-Glu-Asn-Cys-Pro-Lys-Gly-NH2

mN j /

9

H 1-deamino-Lyse-vasopressin

penicillin G acylase. 74% pH 7,37"C,

Figure 18-3. Application ofthe phenylacetamido (PhAc) group as an enzymatically removable amino protecting group.

The applicability of the penicillin acylase-catalyzed deprotection for the constmction of larger peptides has been demonstrated by the complete deprotection of the presumably at the Nporcine insulin derivative 7 carrying three PhAc terminal glycine of the A-chain, the N-terminal phenylalanine of the B-chain and the side chain of the lysine in position 29 of the B-chain (Fig. 18-3). The enzymatic hydrolysis proceeded to completeness and the peptide backbone was not attacked. A further interesting example is given by a recent biocatalyzed synthesis of leucine in which all critical steps are performed by enzymes, enkephalin tert-butyl ester 8[381 two of them through the agency of penicillin G acylase: i) phenylacetates are introduced as N-terminal protecting groups of the amino acid esters by using penicillin G acylase, ii) the elongation of the peptide chain is carried out with papain or a-chymottypsin, iii) the deprotection of the N-terminal amino group is achieved again by means of penicillin G acylase. These examples and also the application of this technique for aspartame syntheses[28*40, 41), as well as the deprotection of glutathione derivatives[351 demonstrate that penicillin G acylase can be used advantageously for the N-terminal unmasking of peptides. In addition, the enzyme has

I

1337

1338

I

78 Introduction and Removal of Protecting Croups

been used for the liberation of the side chain functionalities of lysine and cysteine, as well as in p-lactam, nucleoside and carbohydrate chemistry (vide infia). 18.2.2 Enzyme-labile Urethane Protecting Groups

The enzyme-labile N-protecting functions described so far are simple acyl groups which typify the danger of razemization during chemical peptide syntheses. This problem can, in general, be overcome by the use of urethane blocking functions. However, so far only few examples of a biocatalytic removal of classical urethane protecting groups such as the Z- and Boc-group are known['*]. Apparently, the enzymatic attack on the urethane carbonyl group, which would initiate the cleavage process, is too inefficient to be useful for synthetic purposes. To overcome this problem, two different strategies were developed. Both concepts have in common the fact, that the enzyme-labile bond is no longer part of the urethane. However, the first approach includes the introduction of a spacer (the AcOZ- and PhAcOZ groups), while the second strategy relies on the cleavage of a glycosidic C - 0-bond of a glycoside urethane by the respective biocatalyst, e.g. a glucosidase (the BGloc group). Through the introduction of a spacer between the group which is recognized by the enzyme and the urethane, the substrate is kept at a distance from the enzyme during the reaction (Fig. 18-4). Therefore, any steric effects caused by the bulk of certain amino acids are expected to be minimal and, as the amino acid sequence does not influence the reactivity, this concept should be generally applicable to the synthesis of peptides and peptide conjugates. An additional advantage of the introduction of the spacer is the option to choose the group that is recognized by the enzyme and thus the enzyme itself. This concept was first realized by using p-hydroxybenzyl alcohol as a spacer in the p-(acetoxy)-benzyloxycarbonyl (AcOZ) group which encorporates an acetic acid ester as the enzyme-labile bond (Fig. 18-4).Accordingly, the AcOZ group can be removed under conditions typical for acetyl ester hydrolysis, for instance by treatment with lipases or esterases [53-551. As lipases display a broad specificity, other esters present in the substrate molecule might be hydrolyzed during the AcOZ removal. Thus, the p-(phenylacety1)benzyloxycarbonyl (PhAcOZ) group was developed, which takes advantage of the high selectivity of penicillin G acylase for the phenylacetyl group (Fig. 18-4). The versatiliy of this enzyme-labile urethane protecting group was demonstrated by the synthesis of phosphorylated [56-601, glycosylated [56-601 and lipidated['l] peptides. A second approach takes advantage of a characteristic property of glycosidases. It is well known that glycosidases hydrolyze their substrates by cleaving the glycosidic bond via nucleophilic attack at the anomeric carbon atom. Therefore, a carbohydratederived urethane protecting group would provide the desired enzyme-lability. In additional, such sugar derivatives have increased solubility in aqueous solutions, a necessary requirement for all biotransformations. This concept was successfully realized by using glucose and galactose as the carbohydrate component

18.2 Protection of Amino Groups Figure 18-4. Principle of the spacerbased protecting groups AcOZ and PhAcOZ.

group which /s recognized by the enzyme

J-

I

1339

0

enzyme-la bile linkage gmup which undergoes spontaneous fragmentation upon cleavage of the enzyme-labile linkage

Jenzyrnatic cleavage

1

fragmentation

0

0 + GO2+ H2N-Peptids

0

PhAcOZ =

(Fig. 18-5)['*, G31. During the synthesis the carbohydrate hydroxy functions are blocked by either benzyl ethers in the tetra-0-benzyl-D-glucopyranosyloxycarbonyl (BGloc) group or acetyl groups in the tetra-0-acetyl-D-glucopyranosyloxycarbonyl (AGloc) or the tetra-0-acetyl-0-D-galactopyranosyloxycarbonyl (AGaloc) protecting groups. The removal of these carbohydrate-basedprotecting groups proceeds via a two step process by removing the hydroxy blocking function in a first step followed by treatment with a glucosidase (AGloc, BGloc) or galactosidase (AGaloc), respectively. I n the case of the acetyl derivatives AGloc and AGaloc a sequential two step process as well as a one-pot procedure were developed for the deprotection reaction, allowing for a convenient deprotection protocol as demonstrated for dipeptide 11 (Fig. 18-5)LG2].

1340

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78 introduction and Removal ofprotecting Groups

I

AGloc

BGEoc

AGaloc

Galactose X=O-PG, Y=H Glucose X=H, Y=O-PG 0 PG=Bn, Ac PG’

Peptide or Peptide

Conjugate

Galactose X=OM, Y=H Glucose X=H, Y=QH Enzyme-labile ) Glycosidic Bond

I

Glycosidase

HHO O S , ,

,OAc

H OAc Q 10

1) Lipase WG, 5% MeOH 0.07 M phosphate buffer pH 6.0,37 ‘C, 16 h 2) alp-glucosidase, 24 h

OH

11 64% over two steps

Figure 18-5.

Carbohydrate-based urethane protecting groups.

18.2 Protection ofArnino Groups

18.2.3 Protection ofthe Side Chain Amino Group of Lysine

During chemical peptide syntheses and if trypsin is used for the construction of the peptide bonds or N-terminal deprotection, the side chain amino group of lysine generally has to be protected to prevent side reactions[13*141. This goal can be achieved enzymatically by applying the penicillin G acylase-catalyzed removal of the PhAc group (vide supru)[G4]. Thus, the first application of the PhAc group in peptide chemistry was a synthesis of l-deamino-Lys'-vasopressin from the protected congener 9, during which the lysine side chain was masked as the phenylacetamide (Fig. 18-3).After the peptide chain had been assembled and the disulfide bond was formed by oxidative cyclization, the PhAc group could be removed enzymatically in 74% yield without side reaction. A further interesting example which demonstrates that this technique can be applied advantageously to the synthesis of even larger peptides is found in the complete deprotection of (PhA~)~porcine insuline (vide supru, Fig. 18-3)LZ71 and modified insuline fragments I['. Since penicillin acylase is commercially available and devoid of peptidase activity["], this method appears to be generally useful for the construction of lysine-containingoligopeptides. In addition to the PhAc group, pyroglutamyl amides (Glp) were proposed as enzymatically removable blocking functions for the lysine side chain LZ31. Their removal was achieved with pyroglutamate aminopeptidase from calf liver. Thus, all N-protecting groups were split off from the protected RNAse 1-10 fragment GlpLys(Glp)-Glu-Thr-Ala-Ala-Ala-Lys(Glp)-Phe-Glu-Arg-OH and from a model dipeptide. The general usefulness of this method remains to be demonstrated, however. 18.2.4

Protection of Amino Groups in fi-Lactam Chemistry

The enzymatic removal of acyl groups plays an important role in the industrial production of semisynthetic penicillins and cephalosporins.To this end, penicillin G 12 (R = CH2-Ph) and penicillin V 12 (R = CH2-0-Ph),or the respective cephalosporins are first deacylated by means of penicillin acylases (Fig. 18-6)[", 68]. The 6-aminopenicillanic acid and the 7-aminocephalosporanic acid thus obtained are subsequently acylated by non-enzymatic or enzymatic methods to give the semisynthetic antibiotics 13. The manu.facture of therapeutically important cephalosporins from penicillin G and V includes a chemical ring expansion of the thiazolidine ring to a dihydrothiazine. In the course of this sequence the amino group remains protected as phenylacetyl or phenoxyacetyl amide, which is finally removed using penicillin G or V acylase. Of particular importance is the choice of a suitable protecting function for the COOH group. It must be stable during the ring expansion but removable without damaging the ceph-3-em nucleus. As an alternative to chemical methods, the use of the phenylacetoxymethyleneester was suggested for this purpose[41, It is easily introduced and is stable during the construction of the cephalosporin framework (Fig. 18-6).Together with the phenylacetamidethe ester can eventually be

I

1341

1342

I

18 introduction and Removal of Protecting Groups

'"m

R = Ph-CH2penicillin G acylase

R = Ph-0-CH2penicillin V 12

bHNm

non-enzymatic or enzymatic R* methods 0

0

acylase

13

I COOH

'COOH semisynthetic penicillins

semisynthetic cephalosporins

kHNX& -0

Ph

ring

0

0

expansion

0&o-()J$

Ph

acylase penicillin G

14

H

2

N

E

Ph

+

70-90%

0 COOH

1) cholesterol esterase, PH 7

OTBDMS

2) Jones oxidation

16

0 Figure 18-6.

Enzymatic deprotection of amino- and carboxy groups in B-lactam chemistry.

removed in high yield from penicillin G and the cephalosporins 14 by penicillin G acylase. The formaldehyde formed in the deprotection is not harmful to the enzyme.

18.3 Protection ofThiol Croups

In a new approach to the well known versatile 0-lactam building blocks, an enzymatic deprotection of an acylated methylol amide was applied with advantages (Fig. 18-6)[70j. Thus, the dibenzoate 15 was regioselectively saponified by cholesterol esterase at pH 7 giving rise to a monoacylated aminal. After Jones oxidation and subsequent loss of formaldehyde, the azetidinone 16 was obtained, which can be transformed into various enantiomerically pure penem and carbapenem building blocks. As an alternative to the well established phenylacetyl group in p-lactam chemisq, recently a biocatalyzed procedure for the removal of phthalyl irnide has been described (Fig. 18-2)F4', 711. Its general usefulness remains to be demonstrated, however. 18.2.5

Protection o f Amino Groups o f Nucleobases

In general, the amino groups of the nucleobases adenine, guanine and cytosine in general must be protected during oligonucleotide synthesis to prevent undesired side reactions. To this end, they usually are converted into amides which are finally hydrolyzed under fairly basic conditions. If the amino functions are, however, masked as phenylacetamides, the protecting functions can be cleaved off by again employing penicillin G acylase (Fig. 18-7)[72-781. The enzyme, for instance, selectively liberates the amino groups of the deoxynucleosides 17 without attacking the acetates in the carbohydrate parts and without damage to the acid-labile N-glycosidic bonds. The biocatalyzed phenylacetylremoval can be carried out using both solubilized or immobilized substrates [771. The latter methodology has been developed using controlled pore glass (CPG) as a solid support (Fig. 18-7).

18.3

Protection ofThiol Groups14-6, *'

12]

18.3.1

Protection o f the Side Chain Thiol Group o f Cysteine

The liberation of the P-mercapto group of cysteine was also achieved by means of the penicillin G acylase mediated hydrolysis of phenylacetamides[33-351. To this end, the SH group was masked with the phenylacetamidomethyl(PhAcm)blocking function (Fig. 18-7).After penicillin acylase-catalyzed hydrolysis of the amide incorporated in the acylated thioaminal (see, e.g. 18),a labile S-aminomethyl compound is formed which immediately liberates the desired thiol. This technique was for instance applied in a synthesis of glutathione which was isolated as the disulfide 19. In a related glutathione synthesis the method was used for the simultaneous liberation of ~~* the SH- and the N-terminal amino function of g l ~ t a m i n e [351.

I

1343

1344

I

18 fntrodudion a n d Removal ofProtecting Croups

AcowBphAc 17

2'-deoxyguanosine

2'-deoxyadenosine

2'-deoxycytidine

O-3,d(TGPhAcGPhAc G PhAc G )PhAc 5 '

1) conc. NH3

3'd(TGGGG)5'

WN&O-3'd(TGGGG)5' H

0

Boc-GlU-OtBU

I

a

CYS-Gly-OH

I fH2

s,

\

N H

J

1) CFjCOOH 2) penicillin G acylase, PH 8 31H202

PhAcm 19 glutathione

18

77%

Figure 18-7. Enzymatic deprotection o f t h e amino groups o f nucleobases and the mercapto group ofcysteine by means o f penicillin G acylase. The shaded balls represent controlled pore glass (CPG).

18.4

Protection of Carboxy

croup^[^^*

12,

791

18.4.1 C-Terminal Protection o f Peptides

As in the enzymatic liberation of the N-terminus of peptides, initial attempts to achieve an enzyme-catalyzed deprotection of the corresponding carboxyl groups

78.4 Protection of Carboxy Groups

I

1345

concentrated on the use of the endopeptidases chymotrypsin[8&821, trypsin[81983s 841 and thermolysin PSI, a protease obtained from Bacillus themoproteolyticus which hydrolyzes peptide bonds on the amino side of hydrophobic amino acid residues (e.g. leucine, isoleucine, valine, phenylalanine). This latter biocatalyst enables the cleavage of the “supporting” tripeptide ester H-Leu-Gly-Gly-OEt from a protected undecapeptide to take place (pH 7, room temperature). The octapeptide thereby obtained was composed exclusively of hydrophilic amino acids. Owing to the broad substrate specificity of thermolysin and the resulting possibility of unspecific peptide hydrolysis this method can not be regarded as being generally applicable. The exploitation of the esterase activities of chymotrypsin and trypsin opened routes to the hydrolysis of several peptide methyl, ethyl and tert-butylesters at pH 6.4 to 8 and room temperature[80,81]. The transformations are not only successful with peptides carrying the respective enzyme-specificamino acids at the C-terminus, but in several cases different amino acids were also tolerated at this position. However, severe drawbacks of this methodology are that numerous peptides are poor substrates or are not accepted at all. Moreover, a competitive cleavage of the peptide bonds occurs if the peptides contain trypsin- or chymotrypsin-labile sequences. Therefore, these proteases appear not to be generally useful for a safe C-terminal deprotection as well. The disadvantages of using by the endopeptidases can be overcome by using carboxypeptidase Y from baker’s yeast [25, 8G, 871. This serine-exopeptidase also has esterase activity and is characterized by quite different pH-optima for the peptidase and the esterase activity (pH >8.5). Even in the presence of various organic cosolvents the enzyme selectively removes the carboxy protecting groups from a variety of differently protected di- and oligopeptide methyl and ethyl esters[25,871 without attacking the peptide bonds. An additional attractive feature is, that its esterase activity is restricted to a-esters, consequently j3-and y-esters of aspartic and glutamic acid, respectively, are not attacked. Carboxypeptidase Y was used advantageously for the stepwise C-terminal elongation of the peptide chain in aqueous solution employing a solubilizing poly(ethy1ene glycol) derived polymeric support as the N-terminal blocking In a further remarkable synthesis which did not include the use of a polymeric N-protecting group, Met-enkephalin 20 was built up employing carboxypeptidase Y for C-terminal deprotection of intermediary generated peptide amides as well as for the formation of the peptide bonds (Fig. 188) (251.

The additional opportunity to hydrolyze selectively C-terminal peptide amides with carboxypeptidase Y is of particular interest if, as is demonstrated in the above mentioned example, enzymatic methods are applied to the formation of the peptide bonds, because amino acid amides are often the nucleophiles of choice in these biocatalyzed processes. For this purpose a peptide amidase from the flavedo of oranges shows very promising p r o p e r t i e ~ 1 ~ The ~ ~ ~enzyme 1. is equipped with a broad substrate specificity and accepts Boc-, Trt-, Z- and Bz-protected and Nterminally unprotected peptide amides (Fig. 18-8). The C-terminal amides are saponified in high yields at pH 7.5 and 30°C without affecting the N-terminal blocking groups or the peptide bonds. A noticeable advantage of this biocatalyst is

1346

I

18 Introduction and Removal of Protecting Groups

H m G l y - G l y *Met-OH 20

methionine enkephalin

[7=C-terminally deprotected by enzymatic saponification of the peptide arnide with carboxypeptidase Y;Tyr was N-terminally deprotected by removal or Bz-Arg with trypsin

PG-peptide-NH2

amidase from the flavedo of oranges pH 7.5,30°C

Tyr-Ser Leu-Val Gly-Leu-Val Gly-Gly-Leu

PG-peptide-OH

100

Figure 18-8. C-terminal deprotection o f peptide amides by carboxypeptidase Y and an amidase from the flavedo of oranges.

that N-deprotected amino acid amides, in contrast to the respective peptide amides, do not belong to its substrates. They can, therefore, be used as nucleophiles in peptide syntheses catalyzed by this enzyme, i. e. the formation of the peptide bond together with the subsequent C-terminal deprotection is achieved in a single step. A further possibility for the enzymatic removal of C-terminal blocking groups is opened up by the application of enzymes which generally display a high esterase/ protease ratio. Such a biocatalyst is the alkaline protease from Bacillus subtilis DY which shows similarities to Subtilisin Carlsberg. For this enzyme the ratio of esterase to protease activity is >lo5. It selectively removes methyl, ethyl and benzyl esters from a variety of Tit-, Z- and Boc-protected di- and tripeptides and a pentapeptide at pH 8 and 37 "C (Fig. 18-9)L9l1. The N-terminal urethanes and the peptide linkages are left intact. A further protease which fulfills the requirements for a successful1 application in peptide chemistry is alcalase, a serine endopeptidase from Bacillus lichenijomis whose major It can advantageously be component is subtilisin A (Subtilisin Carlsberg)[92-941. employed with advantage to selectively saponify peptide methyl and benzyl esters (Fig. 18-9).In a solvent system consisting of 90% tert-butanol and 10% buffer (pH 8.2) even highly hydrophobic and in aqueous solution insoluble Fmoc peptides were accepted as substrates and deprotected at the C-terminus without any disturbing side reactions. A selective classical alkaline saponification of methyl esters would be impossible due to the base-sensitivity of the Fmoc group.

78.4 Protection of Carboxy Groups

PG-peptide-OR

Boc

alkaline protease from Bacillus subtilis DY pH 8,37"C

PG-peptide-OH

w

Tyr(tBu)-Glu-Leu Leu-Glu-Val Ala-Glu-Asp-Leu-Glu

PG-peptide-OR

Bzl Bzl

alcalase, pH 8.2.35°C

85 80

PG-peptide-OH

t

90 vol% tert-butanol, 10 vol% buffer PG Fmoc Fmoc Boc Z

peptide Ala-Val-lle Asn-Phe Met-Leu-Phe Met-Asp(0Me)-Phe

R Me

Bzl Me Me

yield [%.I

85 90 80 90

C-terminal deprotection of peptide esters by the alkaline protease from Bacillus subtilis DY and alcalase. Figure 18-9.

A very promising and unusually stable biocatalyst is thermitase, a thermostable extracellular serine protease from the thermophilic microorganism ntermoactinornyces vulgaris whose esteraselprotease ratio amounts to >lo00 : 1. The enzyme shows a broad amino acid side chain specificity and cleaves methyl, ethyl, benzyl, methoxybenzyl and tert-butyl esters from a variety of Nps-, Boc-, Bpoc- and Zprotected di- and oligopeptides in high yields at pH 8 and 35-55 "C (Fig. 1810)[33, 34, 95-971. In addition, it is specific for the a-carboxygroups of Asp and Glu. To enhance the solubility of the substrates, furthermore, up to 50 vol% of organic cosolvents such as DMF and DMSO may be added which also serve to reduce the remaining peptidase activity to a negligible amount [34. 971. In the discussion of the protease-catalyzed cleavage of the N-terminal protecting groups it has already been pointed out that the use of biocatalysts belonging to this class of enzymes in general, i. e. also for the C-terminal deblocking, may lead to an undesired hydrolysis of peptide bonds. In particular, this has to be expected if the respective ester or amide to be hydrolyzed turns out to be only a poor substrate, which is only attacked slowly, an experience not uncommon if unnatural substrates are subjected to enzyme mediated transformations. This undesired possibility would, however, be overcome if enzymes were used which were not able to split amides at all. This principle has been realized in the development of the heptyl

I

1347

1348

I

18 Introduction and Removal of Protecting Groups

PG-peptide-OR

thermitase, pH 8, 55°C

*

PG-peptide-OH

10-60 vol% organic cosolvent PG

peptide

R

yield [%I

Z Boc Bpoc NPS

Leu-VaCGlu(tBu)-Ala Pro-Gly Tyr(tBu)-Glu-Leu Ser(Bzl)-His(Dnp)-LeuVal-Glu(tBu)-Ala

Me Me Me Me

92 73 55 90

lipase from Rhizopus niveus PG-peptide-OR 21 R = (CHZ)&H3 22 R = (CH&Br

+

pH 7,37"C

PG

peptide

R

Boc Z Aloc 2 Boc

Ser-Thr Thr-Ala Met-Gly Ser-Phe Val-Ala

Hep Hep Hep EtBr EtBr

F moc -Met

PG-peptide-OH

yield

[%I

95 85 90 84 95

tG'y#xtPro-

23 C-terminal pentapeptide of the N-Ras protein

n=

C-terminally deprotected by employing lipase from Rhizopus niveus

Figure 18-11. C-terminal

deprotection of peptide esters by lipase from Rhizopus niveus.

(Hep),[+,' 31, 32, 98-1001 the 2-bromoethyl (EtBr)IG6, 3 1 s 32s "1' and the p-nitrobenzyl (PNB) esters[lo21as carboxy protecting groups for peptide synthesis which can be enzymatically removed by means of lipases or esterases, respectively (Fig. 18-11). The Hep-esters proved to be chemically stable during the removal of the Nterminal Z-, Boc- and the Aloc-group from the dipeptides 21. The selective removal of the Hep-esters was achieved by a lipase-catalyzed hydrolysis. From several enzymes investigated, a biocatalyst isolated from the fungus Rhizopus niveus was superior to the others with respect to substrate tolerance and reaction rate. The enzyme accepts a variety of Boc-, Z- and Aloc-protected dipeptide Hep-esters as substrates and hydrolyzes the ester functions in high yields at pH 7 and 37 "C

18.4 Protection of Carboy Croups

without damaging the urethane protecting groups and the amide bonds (Fig. 1811)[98* 991. Z- and Boc-dipeptide-2-bromoethyl esters 22 are also attacked, at a comparable or in some cases even higher rate. In the presence of either one of the enzyme-labile protecting groups the N-and C-terminal amino acid can be varied considerably. With increasing steric bulk and lipophilicity of the amino acids, in particular the C-terminal one, the rate of the enzymatic reactions decreases. If the Cterminal amino acid is proline, the enzymatic reaction does not take place. The lipase-mediateddeprotection of peptides was for instance successfully applied in the construction of the C-terminal pentapeptide methyl ester 23 of the N-Ras-protein, which is localized in the plasma membrane and which plays a vital role in cellular signal transduction (Fig. 18-11)[lo3]. The use of lipases for the removal of protecting groups from peptides in addition to the absence of protease activity has several advantages. Various enzymes belong ing to this class and stemming from different natural sources (including mammals, bacteria, fungi and thermophilic organisms) are commercially available and fairly inexpensive, This variety provides the opportunity of replacing a chosen biocatalyst by a better one if a particular substrate is only attacked slowly (videinf;a). The lipases are not specific for L-amino acids but also tolerate the presence of the D-enantiomer['041. A noticeable feature is that, in contrast to proteases and esterases, they operate at the interface between water and organic solvents[105]. This is particularly important if longer peptides, which are composed of hydrophobic amino acids and/ or carrying side chain protecting groups, and that do not dissolve well in the aqueous systems, have to be constructed. The full capacity of the lipase mediated technique for C-terminal deprotection was demonstrated by the synthesis of complex base-labile phosphopeptideslaI and 0glycopeptides, which are sensitive to both acids and bases [loG, lo7]. To this end, e. g. the serine glycoside 24 was selectively deprotected at the C-terminus by lipase from the fungus Mucorjavanicus (Fig. 18-12). The carboxylic acid 25 liberated thereby was then coupled with an N-terminally deprotected glycodipeptide and after subsequent enzyme-mediated deprotection the glycotripeptide carboxylic acid 26 was obtained in high yield. This compound was finally condensed with a tripeptide to give the complex diglycohexapeptide27, which carries the characteristic linkage region of a tumor-associated glycoprotein antigen found on the surface of human breast cancer cells. In the course of these enzymatic transformations, the N-terminal urethanes, the peptide bonds, the acid- and baselabile glycosidic linkages and the acetyl protecting groups, being sensitive to bases, were not attacked. In these cases lipase from Rhizopus niveus which was the enzyme of choice for simple peptides only attacked the substrates slowly, so that a different biocatalyst had to be used. This demonstrates the above mentioned advantage of being able to apply several catalytic proteins of comparable activity but different substrate tolerance for the solution of a given synthetic problem. The viability and the wide applicability of the principle of using enzymes for the removal of individual protecting groups from complex multifunctional compounds such as lipo- and glycopeptides is furthermore proven by the finding that proteases can also be used for this purpose. Thus, by means of thermitase-catalysis the C-

I

1349

1350

I

78 Introduction and Removal ofprotecting Groups

H

O

H

z' N&-

&

0=

AcO

zSN?OH 0 '

lipase from Mucor javanicus 88%

AcO OAc

O

A

c

O

~

I A::q

AcO OAc

24

25

1) chain elongation 2) lipase from Mucor javanicus 76%

AqGalNHAc Z-ker-Thr-Ala-Pro-Pro-Ala-OHep I AqGalNHAc

27

chain elongation

H d HO o AcHN

** @ 3o

o

g

HO

AcO AcO

AcHN

H

therrnitase, pH 7.5,45"C, 20% DMF, 86%

Boc-Asnm

OH 29

e

r

HO

26

m

papain, pH 6.6, quant.

HO&-peptide+OMe HO

papain? pH 6.6, 96%

Z-Ser-Thr-Ala-OH AcHN 0

Ace@ AcO OAc

characteristic linkage region of a tumor associated antigen

Teoc-Ser-A l a I K ]

AcHN

I

31: peptide = Ser-Gly subtilisin, pH 7, 68% 32:peptide = Gly-Ser subtilisin pH 7, 65%

Figure 18-12. Construction of acid- and base labile glycopeptides via enzyme-mediated C-terminal deprotection.

terminal tert-butyl ester was removed from the glycopeptide28 (Fig. 18-12)[34s"'I. In a different study, this enzyme was also used for the cleavage of methyl and p nitrobenzyl esters[lo9I.From the serine glycoside 29["', 'l1I and from the asparagine conjugate 30['121the methyl esters could be cleaved offwithout disturbing side reactions by using papain as the biocatalyst. Similarly, the liberation of the Cterminal carboxy group of the glycosylated dipeptides 31 and 32 was achieved by means of subtilisin-catalyzedhydrolysis[1131. However, in these cases papain could not be used since this protease preferably cleaved the peptide bonds. This example again highlights the danger associated with the use of a protease for the removal of protecting groups from peptides.

78.4 Protection ofcarboxy Croups

I

1351

A problem arising regularly in the enzymatic deprotection is the poor solubility of the fully blocked peptides in the required aqueous media, resulting in a limited accessibility of the substrates to the enzymes. To overcome this difficulty, in many cases solubilizing organic cosolvents are added, however, a more general and viable approach consists of the introduction of solubilizing protecting groups, e. g. in the enzyme-mediated formation of peptide bonds (see Chapter B 2.5) [l14]. An enzymatically removable solubilizing ester protecting group could be found in the ethylene glycol derived esters such as the methoxyethyl (ME) estersL7**'151, and the methoxlipase PG-peptide-0

PG-peptide-OH

pH 7,37"C

n=1: methoxyethyl (ME) n=2: methoxyethoxyethyl(MEE)

33

-+

Boc-peptide-0 34

NMe3 B r-

butyrylcholine esterase from horse serum pH 6.5, r.t.

*

Boc-peptide-OH

Cho

o=l;.,. H-Ser-GIy-Asp(0H)-OH

HO HO ,HE\

H-Thr-Gln-Thr-Ser-Ser-Ser-Gly-OH OH adenovirus 2 nucleoprotein

o k w HI,

j

serum response factor (SRF)

Aloc-Cys-Met-Gly-Leu-Pro-Cys-OMe SJ

O\

G,-,-proten i

N-Ras protein

Boc-Phe-Cys-Asp-Phe-OH

'

I

0

human Y, receptor

Figure 18-13. Use o f hydrophilic esters as solubilizing enzymatically removable protecting groups for the synthesis o f characteristic protein fragments.

1352

I

78 htroduction and Removal ofprotecting Groups

H

'0

PG-peptide-NMN

H

35

peroxidase or tyrosinase pH 7.37 "C

r

i

PG-peptide-N"No spontaneous fragmentation

PG-peptide-OH Figure 18-14.

+

N2

+

Phenylhydrazide as a carboxy protecting group.

yethoxyethyl (MEE)

13)[58,59, 76, 78, 118-1211 . ~he

0 1

115-1171 and in the choline esters (Fig. 18ME and MEE esters serve both as hydrophilic analogues

of the heptyl esters discussed above and can therefore be removed by the same biocatalysts such as the lipase from Mucor javanicus. Their increased solubility in aqueous media has been used successfully in the synthesis of small peptides and peptide conjugates including glyco-[115-1171 and nucleopeptides F7'1. Similarly, the respective dipeptide choline esters 34 are readily soluble in purely aqueous media (i.e. without added cosolvent) and are converted into the corresponding carboxylic acids under the mildest conditions, and without side attack on the peptide bonds and the N-terminalurethanes, by means of the commercially available butyrylcholine esterase from horse serum. The increased hydrophilicity of peptide choline esters was used advantageously used for the synthesis of peptides and very sensitive peptide conjugates such as lipidated peptides [118-121], phosphorylated and glycosylated peptides Is', "1 and nucleopeptides (Fig. 18-13) [76, 781. Recently, phenylhydrazide has been introduced as an enzyme-labile carboxy protecting group[122,1231 . Th'is protecting group can be removed by mild enzymatic oxidation using a peroxidase[122.1231 or mushroom t y r ~ s i n a s e [ ' (Fig. ~ ~ ] 18-14). 18.4.2

Protection ofthe Side Chain Groups of Glutamic and Aspartic Acid

The stepwise removal of arginine methyl ester by proteases has been investigated as a possibility for the enzymatic deprotection of the side chain carboxylate groups of the aminodicarboxylic acids aspartic acid (Asp)and glutamic acid (Glu).To this end, Z-Asp(ArgOMe)-NHzand Z-Glu(ArgOMe)-NHzwere converted into Z-Asp(0H)NH2 and Z-Glu(OH)-NH2by subsequent treatment with trypsin, which hydrolyzes the arginine methyl esters, and with porcine pancreatic carboxypeptidase B, which splits off the arginines[125].Since the second step is slow and requires high concentrations of the carboxypeptidase, this method can, most probably, not be applied routinely in peptide synthesis because it introduces too much of a danger of competitive side reactions. However, enzymatic transformations have proved to be useful for the synthesis of selectively functionalized aspartic and glutamic acid derivatives. For instance,

18.5 Protection of Hydroxy Croups

I

alcalase selectively hydrolyzes the a-benzyl esters of H-Asp(Bz1)-OBzl and HGlu(Bz1)-OBzl in 82% and 85% yield, respectively, on a decagramm scale[’261. Similarly, aspartyl- and glutamylpeptides can be deprotected selectively at the Cterminus by this enzyme, however, in these cases an undesirable attack on the peptide bonds may occur[’27].In addition, Z-Asp(OAl1)-OAll is converted into ZAsp(OAl1)-OH in quantitative yield by Also a lipase from Candida cylindracea is able to differentiate between the two carboxylic acid groups of glutamic acid. From the respective di-cyclopentylester it preferably (ratio 20 : 1)removes the y-ester in 90% yield[12’]. In addition, the enzyme thermitase and the alkaline protease from Bacillus subtilis (vide supra) also have great potential for the selective manipulation of dicarboxylic amino acids. The examples given in Sections 18.2 to 18.4 demonstrate that the selective deprotection of peptides can be achieved advantageouslyby making use of enzymatic reactions. In the light of the increasing number of available biocatalysts it appears that in the near future a host of new and superior enzymatically removable blocking groups for the synthesis of peptides will be developed. However, these techniques will definitely not be used for the preparation of simple small peptides in the laboratory. Most probably they will be applied to the synthesis of sensitive polyfunctional compounds and long oligopeptides, the construction of which is cumbersome by standard chemical methods. Furthermore, they offer significant advantages if a technical process for the manufacturing of a given peptide has to be developed. Finally, together with the recently developed methods for the biocatalyzed formation of peptide bonds (see Chapter 12.5) (l3Ol, enzymatic protecting group techniques could prove to be the tools of choice for the construction of peptides in aqueous solution, the practical development of which has been tried for several decades [131,1321

18.5 Protection of Hydroxy Groups

[4-93

’

33-1

361

Mono- and oligosaccharides,alkyl- and arylglycosides and various other glycoconjugates generally include a multitude of hydroxyl groups of comparable chemical reactivity. Also, the synthesis of oligonucleotides and nucleosides, B-lactams, alkaloids, steroids and peptides often requires the selective protection of one or more alcoholic functions. Consequently, for the directed construction of polyhydroxy compounds these functional groups have to be manipulated selectively, in general making cumbersome protection and deprotection steps necessary. Although numerous chemical techniques are available to mask or to liberate hydroxyl groups, the development of enzymatic methods for this purpose has been progressing steadily and appears to complement the arsenal of classical tools. In addition, the enzymatic protection of hydroxy goups (and vice versa of carboxy groups) in racemic compounds as well as their enzyme-catalyzeddeprotection has been used extensively for the separation of enantiomeric alcohols and carboxylic acids (see Chapter 11).

1353

1354

I

18 fntroduction and Removal of Protecting Croups

18.5.1 Protection of Monosaccharides[133f 1371

The selective protection and deprotection of carbohydrates can be achieved with various classical chemical techniques 38-1401. In addition, however, owing to the synthetic challenge the multifunctional carbohydrates pose, enzymatic techniques for the introduction of blocking groups into sugars and/or their subsequent removal offer further, different opportunities. The enzymatic acylation of sugars in aqueous solution has been reported but gives low yields as the equilibrium for the reaction favors hydrolysis. However, enzymatic acylation in dry organic solvents has shown substantial success. While direct enzymatic esterification of alcohols with acids is often not practical, good to excellent yields have been obtained using transesterification techniques (Table 18-1).The displacement of the equilibrium toward products has been accomplished by using an excess of the acyl donor and by using activated, irreversible acyl donors such as trihaloethyl esters [l4l], enol esters [1421, acid anhydrides or oxime esters [134, l3'1. In particular,the enol esters have the advantage that the liberated enol tautomerizes to a ketone or an aldehyde, thereby shifting the equilibrium toward the desired products and consequently giving higher yields. This technology, however, is not restricted to carboxylic acid derivatives being the acyl donor. Organic carbonates [1431, either activated as the or, even better, as an 0xime[~~'1 derivative, allow for the enzyme-catalyzed synthesis of carbonates such as the methoxycarbonyl, the benzyloxycarbonyl (2) and the allyloxycarbonyl (Aloc) carbonate. The last two examples can later be removed by non-enzymatic means. The high polarity of sugars and their derivatives requires that polar solvents be used to dissolve them. Solvents found to be suitable include pyridine, DMSO, DMF and dimethylacetamide. However, these solvents also often inactivate enzymes, although some enzymes, for instance the lipases from the porcine pancreas (PPL), from Candida antarctica (CAL), from Candida GyEindracea (CCL, later renamed Candida rugosa) and the lipase from Pseudomonas cepacia (PSL) as well as the proteases subtilisin and proleather, maintain their inherent acitvity [14G]. A less polar solvent such as THF allows the use of a broader variety of lipases, but does not dissolve unmodified pyranoses. Nevertheless, it should be noted that even glucose suspended in THF has been successfully acylated by using lipase of Candida antar~tica[~~']. To remain active in an organic solvent, the enzyme must contain a small amount of water which is required for maintaining the correct protein structure. In the absence of this essential water, highly polar compounds such as carbohydrates form excessively tight enzyme-product complexes. This inhibits association and dissociation of substrates and products from the active site and thus slows down the reaction. Accordingly, the addition of drying agents such as zeolite CaA not only influences activity of the the biocatalyst but also its selectivity. For instance, the acylation of 1-0methyl 0-D-glycopyranoside49 catalyzed by lipase SP 435 (an immobilized lipase from Candida antarctica) in ethyl butanote as the solvent and acyl donor led to 1491. If zeolite CaA was added, a acylation predominantly in the G-po~ition['~~,

'

18.5 Protection ofHydroxy Groups

I

1355

mixture of 2,6- and 3,6-bisacylatedpyranosides (95 : 5) was formed. In the presence of zeolite CaA and tert-butanolas a cosolvent, again monoacylation in the 6-position was observed. Alternatively, precipitation of the enzyme from aqueous solution at its optimum pH prior to its use in an organic solvent has also been reported to increase the enzyme’s activity greatly. The results of enzymatic acylation of several pyranose and furanose sugars are shown in Table 18-1. Other lipophilic carbohydrate derivatives such as alkyl glycosides also display a higher solubility in less polar organic solvents, in which most lipases tend to be more stable than in polar solvents. A further interesting finding is that heat stable lipases are capable of transferring long-chain fatty acids to the 6-hydroxy group of ethyl glucoside on a kilogram-scale, utilizing the molten fatty acids themselves as solvent^^'^^]. On a somewhat smaller scale, the acylation of glucose has also been carried out using only a minute amount of solvent or in supercritical CoZ 1741. The regioselectivity observed in the acylation of underivatized pyranoses in principle parallels that recorded for the classical chemical introduction of acyl groups into carbohydrates. However, if the 6-OH groups are protected first or deoxygenated, in the corresponding enzymatic reactions selectivities are observed which can not be realized with classical chemical methods. By careful choice of solvent and lipase, it is possible to rnodifiy selectively a number of C6 protected pyranoses at the secondary hydroxy groups (Table 18-2). By combination of enzymatic with non-enzymatic protection group chemishy, carbohydrates can be selectively modified in the primary and secondary hydroxy positions. To demonstrate this versatility, the straightforward synthesis of differently mono-acylated glucose derivatives is described in Fig. 18-15. For instance, 6-0butyrylated glucose GGa (R = n-butanoyl; prepared enzymatically, see Table 18-1)is converted into the 3,6-dibutanoate 93 by lipase from Chromobacterium uiscosum (CVL) or from Aspergillus niger (ANL). The 2,6-dibutanoate 94 can conveniently be built up with the lipase from porcine pancreas (PPL; Fig. 18-15)[1641.Similar observationswere reported for n-octylglucoside,but for the corresponding galactoseand mannose 6-esters the selectivity was lower. In contrast, the chemical butyrylation of glucose derivative GGa with the acid anhydride in pyridine gave a complex mixture of various diesters without any significant regiodiscrimination. The enzymatic approach was also used to convert the 6-0-tritylglucose GGb (R = Trt) into the 3-butanoate 95 by a chemoenzymatic approach with lipase from Chromobacterium glucose GGc (R = TBDPS) could uiscosum (CVL), and the 6-tert-butyl-diphenylsilylated be acylated exclusively at the 2-position when employing lipase from Candida cylindracea (CCL) From the disubstituted glucoses obtained by the enzymecatalyzed reactions, the protecting functions in the 6-position could be split off chemically or enzymatically, thus making the glucose esters 95 and 96 carrying a single acyl group in the 2- or the 3-position available in a convenient way (Fig. 1815).

The monoacylated saccharides used in these studies dissolve in several organic solvents, of which tetrahydrofuran and methylenedichloride were found to be

40

39

38

36 Hi-

H

OH OH

Solvent

pyridine dioxane TH F pyndine pyridine pyridine DMF pyridine pyridine pyridine pyridine dioxane

pyridine benzene/pyridine 2:1 pyridine dioxane DMF benzene/pyridine 2:1 DMF DMF 97% DMF

pyridine dioxane

Enzyme"

PPL CAL CAL PS L PSL proleather subtilisin subtilisin optimase M-440 PPL PSL CAL

PPL CCL PSL CAL protease N CCL protease N subtilisin 8399 subtilisin BNP'

CAL PSL

Selective acylation of the primary hydroxy group in monosaccharides.

Compound No. Structure

Table 18-1.

RC02N=CMe2 RC02N=CMe2

MeC02C (Me)=CH2 MeC02C(Me)=CH2 MeC02CH=CH2 BOC-Gly-OCHzCN

MeCOlCH2CC13 MeC02CH=CH2 RC02N=CMe2 ROC02N=CMe2 MeC02C(Me)=CH2

MeC02CH2CCl3 RC02N=CMe2 ROC02N=CMe2

RC02CH2CC13 ROC02N=CMe2 RC02CH=CH2 MeCOzCH2CC13 EtC02CH2CC13 PhC02CH2CC13 PrC02CH2CC13 PrC02CH2CCl3 Boc-l'he-0CH2CF~

Acyl Donor

6 6

6 6 6 6

6 6 6 6 6

6 6 6

6 6 6 6 6 6 6 6 6

Position

45-83 50-72

73 92 65

65-80 44-53 40

36

57 70-85 43-68

79 29 33 60 64

19-35 15-72

Yield ("7)

[1561 11561

11421 P531 P541 (1551

WI

~411 ~421 ~521 [I451

~411 ~521 P451

11511

11501

~411 [I451 [I471 ~461 ~461 ~461 11501

Ref.

Y

-z

u

5

a0

3' og

'OI

n

%a

0

5

a

a

2

g.

2

a

0

18.5 Protection ofHydroxy Croups EZ In

z

ziz In* 66

In

d

3 .

. . IIn

6

13

m

6

In

d

el vl

c

-

B I

cl

c c

0

r

0 I 0

F d' d

d N

D;

I

tI l

I

d

P

rn d

d u

I

1357

(cont.).

51

H & ;OC8H ,i

49 Hi%

47b %; H

0

OH OCEHi7

OH

OH OMe

OH

Compound No. Structure

Table 18-1.

CAL

THF PrCOzEt/tBuOH (1:l) tBuOH THF THF

CAL CAL CAL CVL ANL

MeC02CH=CH2 PrC02Et Ph(CHz),COzH PrC02CH2CC13 PrC02CH2CC13

CHz=CHCOZEt/ tBuOH (l:l)CH2=CHCOzEt THF/pyridine (41) MeCOzCH=CH2

MeCOzCH=CH2 MeCOZCH=CHz PrCOzEt

CAL CAL

CCL CAL CAL

benzene/pyridine 2:1 THF/pyndine (4:l) PrCOzEt/tBuOH (1:l)

CiiHz3C02H

acetone/pyridine 3:l

CAL

CAL

Acyl Donor

Solvent

Enzyme"

67

51

6

6

6

6 6 52 6 6;3,6 (1:l) 6;3,6(1O:l)

6 6

6 3,6 6

Yield ("A)

Position

Ref.

-

w

4

-2

00

-

00 U

78.5 Protection of Hydroxy Croups

I

1359

5 a 3

I

m 0

m rn

in h

a

I N

el

el

a

m

m fi

ln N

ln d

el

a

m

ln W

(cont.).

G1

OH

OMe

PPL

THF

pyridine

MeC02CH=CH2

MeC02CH=CHz/THF

PSL

PPL

Acyl Donor

Solvent

Enzyme”

OH

PPL

PPL

TH F

TH F

“VMe

HO

HO OH

HowoMe voMe

Hi=3

0

‘cog

GO HO

59

58

57

Compound No. Structure

Table 18-1.

5

5

5

6

6

Position

84

77

77

5

[I681

~

[W

94

81

Ref.

Yield (“h)

1

4

2

a

oq

5. CI

0,

3P

D

;

B

sl.

n

$s

a

.b

s

0

0

-

W

m

(cont.).

THF

PPL

Acyl Donor

CCL

CCL

kOH

HO

9

THF

MeCOzCH=CH*

6

6

5 3

Position

EtOAc

EtOAc

MeC02CH2CF3

HvMe

Solvent

Enzyme'

60

93

90

39 17

Yield (%)

~701

Ref.

a Many enzymes were usually screened for activity, only the best results are listed. CAL Candida antarctica lipase; CCL lipase from Candida cylindracea (later renamed Candida rugosa): PPL porcine pancreas lipase: PSL Pseudomonas cepacia lipase.

63

62

Compound No. Structure

Table 18-1.

3

g.

F

u

Po m D 2

MeCOzCH=CH2

MeC02CH=CH2 MeCOzCH=CHz

MeC02CH=CH2

MeC02CH=CH2 MeC02CH=CH2

lipase from Mucor miehei

PFL lipase from ft2Mucor miehei

69

70

OH

PrC02CH2CC13

TH F

O M ?-€;

PrC02CHzCC13

TH F

CVL

ANL CVL PPL CVL PFL CCL

CVL

0

a: R=butyryl b: R=trityl C: R=TBDPS

PrC02CH2CC13 PrC02CH2CC13 PrC02CH2CC13 PrC02CH2CC13 MeC02CH=CH2 PrC02CH2CC13

THF THF THF THF MeC02CH=CH2 CHzClz

68

67

66

A q l Donor

Solvent

Enzymea

Selective acylation o f secondary hydroxy groups in monosaccharides.

Compound No. Structure

Table 18-2.

3 2

2

2 3

2 3

3 (GGa) 3 (GGa) 2 (GGa) 3 (6Gb) 2 (GGb) 2 (GGc)

Position

52

13

20 31

45

80 51 88

Yield (“h)

11641

11641 11641 ~641 ~ 4 1 F751 [I641

Ref.

2

$

?

9

0

2 8’

5

00

N

--

-

m

W

(cont.).

\

a: R=OMe OH b: R=SEt

-

a: R=OMe R b: R=SR c: R=OPh

HO

a: R=butyryl b: R=trityl c: R=benzyl

71 phT%

Compound No. Structure

Table 18-2.

MeC02CH=CH2

PFL

3

2 (74a) 2 (74a) 2 (74a) 2 (74b) 2 (744

THF/pyridine (41) THF/pyridine (4:l) CHzClz/pyridine (4:l) MeC02CH=CH2 MeC02CH=CH2

PPL PFL CCL PFL PFL

3 (72a) 3 (72a) 3 (724 3 (72b)

80

81

84

93

86 86 86

98 94 73 76

2 (71a) 2 (71a) 2 (71a) 2 (71a) 2 (71b) 2 (71c) 3 (72a)

Yield (“h)

Position

2

MeC02CH=CH2

Acyl Donor

THF/pyridine (41)

Solvent

PPL

PSL PFL PFL

PSL

PSL PFL PFL

PSL

Enzyme”

Ref.

8

W

d

(cont.).

OH

a: R=OAII b: R=SEt

h

bMe

R PFL

PSL

PPL

OH

PFL

.:WoMe

Pr

PSL

79

o<

PFL

80

MeCOzCH=CH2

MeCOzCH=CH*

MeC02CH=CH2/THF

MeCOzCH=CH2

Solvent

4 4

PrC02CHzCF3 THF

3

3

68

65

92

90

10

3 (7Gb)

2

91

Yield (“h)

3 (7Ga)

Position

PrC02CHzCF3

RCOzCH=CH2

MeCOzCH=CH*

MeCOzCH=CHz

MeCOZCH=CHz

MeCOzCH=CH2

Acyl Donor

THF

RC02CH=CHz


O HO

phTo

Enzymea

78

77

76

Compound No. Structure

Table 18-2.

11771

11751

[180]

11771

Ref.

4

-

9

9

%

?

$

D 4

s

2

2

a

3,

1

00

5 -

18.5 Protection of Hydroxy Groups

z7

Ln

d

4 Ln

8

m

m

N N

el

44

fi

a h

-1

m fi

-

00

00 N

a

m

*

0

N

&r&

& &

r&

00 d

I 00 n

-1

fi

I

1365

(cont.).

90

OH

Hexane

PSL

CAL

6H

PSL

MeC02CH=CH2

PrC02Et

dioxane dioxane MeC02CH=CH2 PrCO2EtltBuOH

MeCN

PSL

CAL CAL PSL CAL

Solvent

Enzymea

p:H

OH

m0

89 HO

88

87 H R S O OH C * H l 1

Compound No. Structure

Table 18-2.

MeCOzCH=CHz

PrCOzEt

RC02N=CMe2 MeOCOzN=CMe2 MeC02CH=CH* PrCOzEt

MeCOzCH=CH2

MeC02CH=CH2

Acyl Donor

4

4

4 4 4; diester

70-72 42

28

4

70

3,4

Yield (“h)

2,4

3,4

Position

1187,1891

Ref.

a a

-1

d 00

-

18.5 Protection ofHydroxy Groups

I

1367

1368

I

18 Introduction and Removal ofprotecting Croups

R=Trt 1) CVL, THF, 2, H+ 88% TCE-But

R = n-But HHO

O

q

94

TCE-But = 0 ~ O ^ C C I ,

R = n-But

THF OH TCE-But 80% 66a:R = n-But 66b:R = Trt 66c:R = TBDPS

CCL 85%

OBut

HEo%

~

OH TCE-But 51%

ButO

95

HO

OH 93

R = TBDPS 2) Bu~NF/AcOH I) CCL -5O"C, THF CHZCIZ 75% TCE-But 75%

96 Figure 18-15.

Selective enzymatic introduction of protecting groups into partially acylated

hexoses.

particularly suitable for the enzymatic reactions. This was also observed in the lipasemediated acylation of the methyl glycosides of both D- and L-fucose and -rhamnose, Using lipase from Pseudomonasfluorescence (PFL), both D-carbohydrates were converted into the 2-monobutanoates with high regioselectivity. The naturally occurring L-enantiomers of these 6-deoxysugars, however, were esterified preferably at the 4-hydroxy groups. These results contrast favorably with chemical derivatizations, since the 4-hydroxy groups of the 6-deoxy-L-carbohydrateshave only slight reactivity toward chemical acylating reagents. In addition, methyl-L-fucoside can be converted into the 3-butanoate with lipase from Candida cylindracea. The introduction of an acyl-substituent into the 6-positions of the D-fucoside and the Lrhamnoside does not influence the regioselectivity of the enzymatic acylation Finally, it should be mentioned, that some attempts were made to differentiate between the hydroxy groups of fructose by enzymatic methods, however, with lipases as well as with subtilisin, only mixtures of the 1- and 6-isomers were ob-

78.5 Protection of Hydroxy Croups

tained [141, 150, 1921. Regioselectively monosubstituted fmctoses can, however, be obtained by an enzymatic approach from sucrose (vide infu). 18.5.2

Deprotection of Monosaccharides['33.l37I

Initial attempts to apply lipases for the enzymatic removal of acyl groups from glucose pentaacetate only resulted in low levels of selectivity[1932 'O4I. However, later was found to hydrolyze exclusively the on lipase from porcine pancreas (PPL) anomeric acetate from peracetylated pyranoses while the esterase from Rhodosporium tomloides (RTE)F1O5I releases the primary hydroxy group in preferance (Table 18-4).On the other hand, if the anomeric center is derivatized as a methyl glycoside, the regioselective enzymatic liberation of the 6-OH group becomes feasible with a number of hydrolytic enzymes [168,195-1991 . Thus, from methyl a-D-glucose tetraoctanoate 97a and the corresponding tetrapentanoate 97b, lipase from Cundidu cylindruceu (CCL) removes only the primary ester group in yields of ca. 75%. Similarly, the a-D-gdactoside103,as well as the corresponding mannoside 104b and the 2-acetaniido-2-deoxy-mannoside105 were converted into the 6-deprotected pyranosides in 29-50 % yield (Table 18-3), but the 2-acetamido-2-deoxy-glucoside was only a poor substrate. In the latter cases the regioselectivitywas less pronounced and the 4,G-dideoxyderivatives were also formed in ca. 20 % yield. In addition to this class of compounds, lipases also accept hexopyranosides carrying several different functionalities (e.g. acetals [1971, enol ethers [160, I'*' and, in particular, 1,G-anhydropyranoses as substrates (Tables 18-3and 18-4).In all cases the reaction conditions are so mild that the acid sensitive structures of these compounds remain unaffected. Particularly remarkable is the regioselectivity displayed by lipase from Pseudomonas cepucia (PSL:)in the deprotection of the glycall31 [16', 2001. The biocatalyst exclusively attacks the 3-acetate and leaves the primary ester intact. The enzymatic deprotection strategy can also be used to synthesize carbohydrates carrying a single acyl group in selected positions. Thus, 3,G-dibutyryl glucose 93 (prepared by enzymatic acylation of glucose) was converted into the 3-butanoate 95 by lipase mediated hydrolysis of the 6-ester (Fig. 8-15)[1641. The principles and the enzymes mentioned above which allow the regio- and chemoselective protection and deprotection of the various pyranoses to be carried out were also successfully applied to the enzymatic manipulation of acyl groups in furanoses. Of particular interest in this context is the finding that the five-membered rings can also be handled by the biocatalysts with a pronounced regioselectivity, although furanoses can adopt more flexible conformations with similar energies in solution. The cleavage of the primary acetyl groups from the furanosides 106-111 could be carried out in high yields with lipase from Cundida cylindruceu (Table 18-3)[lG8]. For and the a- and the P-xylo-compoundsthe hydrolysis the 2-deoxy-a-~-ribofuranoside was less selective. From the peracetylated furanoses 125 and 126 the anomeric acyl group was removed with total selectivity by means of lipase from Aspergillus niger (Table 18-4). 1,G-Anhydropyranosesserve as convenient starting materials for various synthetic

I

1369

101

99

98

AcO

AcO

) 0( o g

0

OMe

R=octanoyl

PPL

PPL

PPL

CCL

RTE

c: R=acetyl

OR

CRL

a: R=octanoyl b: R=pentanoyl

RoOMe

CCL CCL CCL CCL PEG-modifiedCCL

97

R;;g

Enzyme"

6 6

0.1 M phosphate buffer 0.1 M phosphate buffer 0.1 M phosphate buffer 0.1 M phosphate buffer, BuzO (10%) Cl,CCH,

G

0.1 M phosphate buffer

0.1 M phosphate buffer, acetone 1O:l

0.1 M phosphate buffer, acetone 10:1

6

6

6

6

citrate buffer

0.1 M phosphate buffer, acetone 1O:l

G

0.1 M phosphate buffer

6 6 4,6

G

Position

Solvent

Selective deacylation of primary hydroxy groups in monosaccharides.

Compound No. Structure

Table 18-3.

75

82

90

77

78 (97a) 75 (9%) 90 ( 9 7 4 (974 27 (97c) 48 (97c) 91 (97c) 77 (97c)

Yield ("A)

[I991 [I951

[1681 [16531 ~961 [I971 [198]

Ref.

a

a a

0

9

s%

f

a

2. %

-5

oa

0 U

-

W

(cont.).

AcO

OAc

a: k a c e t y l OMe b: R=pentanoyl

106 AcoQoMe

104

Ro OMe a: R=acetyl b: R=pentanoyl

103 RO

6

85

6

0.1 M phosphate buffer

CCL

0.1 M phosphate buffer, 10% DMF

[199] [195] [168] 94 (104a) 70 (104a) 33 (104b) 6 6 6

0.1 M phosphate buffer citrate buffer 0.1 M phosphate buffer

CRL RTE CCL

CCL

[168]

[195] 29 (103b)

85 (103a)

~641

Ref.

6

6

85

Yield ("h)

0.1 M phosphate buffer

citrate buffer

6

Position

CCL

RTE

Tris.HC1

0.1

CCL M

Solvent

Enzyme"

Roe

Compound No. Structure

Table 18-3.

(cont.).

o

Acb

c

~

bAc

~

CCL

CCL

e CCL

0.1 M phosphate buffer, 10% DMF

0.1 M phosphate buffer, 10% DMF

0.1 M phosphate buffer, 10% DMF

0.1 M phosphate buffer, 10% DMF

Solvent

AcO

CCL

5

5 3

5 3

5

Position

63

50

40

50 30

98

Yield YO)

Ref.

Candida rugosa;CRL); PPL: porcine pancreas lipase; RTE: Rhodosporiurn toruloides esterase.

0.1 M phosphate buffer, 10% DMF

AcowoMe

M

Enzyme"

a Many enzymes were normally screened for activity, only the best results are listed. ANL Aspergillus niger lipase: CCL lipase from Candida cylindracea (later renamed

ll1

OAc

109 A C 0 q g O M e

108 A

AcO

Compound No. Structure

Table 18-3.

. %

P

-

0

-

-

W

N U

4

18.5 Protection of Hydroxy Croups

I

1373

m w

3

*- 00 0

r.w

Ink

I 00 n

2"

IL

elw &I-

4

a F&

N

3 3

m

3 3

f3

& &

I n 3 3

ua 3 3

4

2

(cont.). Solvent

PPL RTE

0.05 M phosphate buffer, 10% DMF citrate buffer

hog kidney acylase phosphate buffer, DMF (1O:l) Aspergillus niger pecti- phosphate buffer, DMF (1O:l) nase ANL phosphate buffer, DMF (1O:l)

Enzyme”

‘ACO

OAc

1

4 4

1

tAmyl-OH Cl3CCH3

0.05 M phosphate buffer, 10% DMF

PSL PEG-modifiedCCL

PPL

3

0.05 M phosphate buffer, 10% DMF

0.1 M phosphate buffer, 10% acetone

54

84 82

88

61

11

93 27

4

Yield (“h)

Position

2 3

PPL

ANL

;aoMe

6Ac

AcOO B 0OAc . e

120 A

118

Compound No. Structure

Table 18-4.

Ref.

11681

~071 ~981

P681

12061

[2061 PO61

a

4

0 3

S

9.

a 2

d

co

P U

-a

w

d

(cont.).

Enzyme‘

OAc

128 OAc

OAc

F7

a: R=acetyl b: R=butyvl

127 OR

F7

OR

AcO

~~

0.1 M phosphate buffer 0.1 M phosphate buffer 0.05 M citrate-phosphate buffer 0.1 M phosphate buffer 0.1 M phosphate buffer

WGL PLE PPL CVL CCL

alcalase

0.1 M phosphate buffer 0.1 M phosphate buffer

0.1 M phosphate buffer

RJL

CCL

0.1 M phosphate buffer, 10% DMF

0.1 M phosphate buffer, 10% DMF

0.05 M phosphate buffer, 10% DMF

Solvent

ANL

ANL

Aco?a’”c

126 Aco*oAC

125

Compound No. Structure

Table 18-4.

2

4

2.4

2 4 3 4 4 4

1

Position

85-90 82

15 (127a) 67 (127a) 69 (127a) 42 (127a) 91 (12%) 77 (127b)

47 (127a)

50

Yield (“h)

[210] [210]

[209]

[208] [208]

[208]

Ref.

W Ul U

d

% -=I Q.

3

9.

P

a

D

u

cn

Po

(cont.).

OAc

OAc

OBut

PGA

3

3 3,4 3,4,6

2

80-85

90 24 22

60

90 16 19 65

Yield ("A)

(74,2131

Ref.

liver esterase; PPL: porcine pancreas lipase; PSL Pseudomonas cepacia lipase; RJL Rhizopusjaponicus lipase; RTE: Rhodosporium tomloides esterase; WGL wheat germ lipase.

0.1 M phosphate buffer

PSL 0.25 M phosphate buffer acetyl esterase from 0.15 M NaCl buffer the flavedo of oranges

phosphate buffer, DMF (1O:l)

2 2 2 4 2,4

0.1 M phosphate buffer 0.1 M phosphate buffer 0.1 M phosphate buffer

CCL CCL PPL

WGL

Position

Solvent

Enzyme"

a Many enzymes were normally screened for activity, only the best results are listed. ANL Aspergillus niger lipase; CAL Candida antarctica lipase; CCL lipase from Candida cylindracea (later renamed Candida mgosa; CRL); PGA: penicillin-G-acylase; PLE: porcine

OPhAc

12g But0

Compound No. Structure

mo

Table 18-4.

4

5

0

--

U

W

m -

18.5 Protection of Hydroxy Groups

HO OR 133a R = acetyl 133b R = butyryl R = acetyl PPL or PLE

63-69%

R = n-butyryl lipase from Chromobacterium viscosum (CVL), Pseudomonas sp. or Mucor miehei (MML) 91%

R = acetyl chymotrypsin RO

OR

:;Jgk

CCL 77%

R = n-butyryl CCL, 47%

OR

RO

HO

127a R = acetyl 127b R = n-butyryl

OH 134

R = acetyl, RJL, 47%

7

0 135

RO Figure 18-16.

OH

Selective enzymatic removal o f protecting groups from 1,6-anhydropyranoses.

purposes in carbohydrate chemistry. Therefore, the directed manipulation of their hydroxy groups is of particular interest. Each of the three OH-groups in 1,G-anhydroglucopyranose can be liberated selectively making use of enzymatic reactions (Fig. 18-16, Table 18-4)[208-210s 2121. Thus, the 4-protecting group was split off from the triacetate 127a using lipase from porcine pancreas (PPL)[2091or pig liver esterase (PLE)[208, 209]. The acetate in the 3-position could be attacked preferentially using c h y m ~ t r y p s i n [or ~ ~lipase ~ I from wheat germ (WGL)[zos],and the 3,4-diacetate 135 was obtained by hydrolysis with lipase from Rhizopusjavanicus (RJL)[208! In each case, however, other derivatives were formed as undesired by products. High yields could be obtained from the tri-n-butanoate 12%. It was converted into the 2,3-dibutanoate 133b in 91 % yield by means of several lipases, but the enzyme from Candida cylindracea (CCL) removed two acyl groups successivelyto yield the monobutanoate 134. Similarly, the analogous 3-azido-1,G-anhydropyranose 128 is regioselectively deacylated at 0 2 and 0 4 by means of lipase OF from Candida cylindracea and

I

1377

1378

I

18 Introduction and Removal of Protecting Groups

alcalase, respectively[2111. Of particular importance is the stereochemistry at C4 of the bicyclic substrates. If the alcohol at this position is equatorial, as for instance in the corresponding 1,6-anhydrogalactopyranose129 and the analogous lactone 130, several enzymes act only in a random fashion or not at However, the acyl group in the 2-position seems to be preferred (Table 18-4).The results obtained from these studies indicate that the reactivity of acyl protecting groups in 1,6-anhydropyranoses toward hydrolysis by lipases decreases in the order C4, > C2, > C3, >> c4q. The above mentioned investigations revealed that the lipase-mediated hydrolysis proceeds at higher reaction rate and, in many cases with better selectivity, if butanoates or pentanoates are employed as substrates instead of acetates. However, the use of enzymatic deacylations is by no means restricted to simple alkanoates. An illustrative and impressive example is found in the hydrolysis of generally basestable carbohydrate pivaloylates using an esterase from rabbit serum (ERS) [214-2171. For instance, the biocatalyst selectively splits off the 6-pivaloyl group from a-methyl 3,4,6-tripivaloyl-2-acetamido-2-deoxy-glucoside. On prolonged incubation the complete removal of pivaloylates from carbohydrates is also possible. Of particular significance is, that the enzyme does not have to be purified, but that crude serum preparations are sufficient for the preparative purposes. A further enzyme which allows the chemo- and regioselective unmasking of different carbohydrate derivatives to be carried out is acetyl esterase from the flavedo of oranges, a biocatalyst 218]. It can be applied for the which preferably hydrolyzes acetic acid synthesis of selectively deacylated pyranoses. Thus, from pentaacetylglucose 112 the 2,3,4,6-tetraacetateis obtained by means of the regioselective saponification of the 1-acetate.If the hydrolysis is allowed to proceed further, the 6-acetate is also cleaved and the 2,3,4-triacetatebecomes available in ca. 40% yield. If tri-0-acetyl-glucall31 is subjected to the enzymatic hydrolysis, at 40% conversion the 6-acetate is the main product. By introducing acyl groups which are specifically recognized by certain enzymes into carbohydrates, not only the regioselectivitybut also the chemoselectivity of the biocatalysts can be exploited. This can, for instance, be achieved by the selective saponification of phenylacetates catalyzed by penicillin G acylase [3G321. The enzyme glucose without affectliberates the 2-OH group of 1,3,4,G-tetraacetyl-2-phenylacetyl ing the acetic acid esters. In this case, moreover, an ester of a secondary hydroxy function is chemoselectively hydrolyzed in the presence of the chemically more reactive acetates at the 6-position and at the anomeric center. This approach was also adopted for the enzymatic deprotection of the glucal 132. Thus, its 3-OH group was liberated without cleaving the acetates that were present [1901. 18.5.3

Di- and Oligosaccharides[’37]

For enzymatic protecting group manipulations on di- and oligosaccharides in particular the use of subtilisin together with dimethylformamide as the solvent is advantageous.As has already been pointed out, the use of DMF is often critical, since

18.5 Protection of Hydroxy Croups

I

1379

its dissolving ability is high enough to solubilize even highly polar polyhydroxy compounds (comparable experiments with pyridine as the solvent generally failed)[l4l].Only a few reports about the successful use of other solvents such as pyridine L2l91 or tert-butanol[220] have been published. subtilisin accepts several disaccharides as substrates and transfers butyric acid from ethyl or trichloroethyl butanoate to the primary G'-hydroxy functions of the nonreducing monosaccharide of the p-(l-3)-linked cellobiose 136 and the respective 220]. For lactose the regioselectivity was less pronounced, maltobiose (Fig. 18-17) however, methyl and benzyl p-D-lactoside 137 were converted into the 6'-butanoates in 71-73% Rutinose in which the primary hydroxy group of the glucose moiety is blocked (see also 149, Fig. 18-19),is selectively substituted in the 3-position [2221. In addition, higher maltooligomers could also be acylated in the 6-position of the terminal nonreducing carbohydrate.For instance, 6"-0-butyrylmaltotriosewas isolated in 29 % yield, but also the corresponding tetra-, penta-, hexa- and heptamer were substrates for the biocatalyst. These enzymatic esterifications open a route to discriminating between the primary hydroxy groups in di- and oligosaccharides in a convenient and straightfonvard way. Classical chemical one step methods of compal4O], and multistep sequences rable selectivity are not available for this purpose[139* usually have to be carried out if the selective protection of a specific primary hydroxy group in a di- or oligosaccharide is desired. Owing to its great commercial importance as a renewable resource, sucrose 138 has been subjected to several enzymatic hydroxy group manipulations. This nonreducing disaccharide turned out to be a substrate for subtilisin In contrast to chemical acylations in which the most reactive OH-groups are found in the 6- and the G'-position, the enzyme selectively transfers various acyl functions to the 192, 2 2 3 ] . This acylation was usually carried out in DMF as a 1'-alcohol(Fig. 18-17)[lSoS solvent, but the use of anhydrous pyridine gave similar results[219].The monoacylated disaccharides 139 thereby obtained could then be further transformed enzymatically. On the one hand, with the lipase from Chromobacteriurn viscosum (CVL) the free primary 6-OH group was acylated in 31 % yield. On the other hand, the 1'-esters 139 are substrates for yeast a-glucosidase which hydrolyzes the glycosidic bond and thus makes the 1-0-acylfiuctoses 140, potentially useful as chiral synthons, available [1921. Alternatively, the 6'-OH-group in sucrose 138 can be selectively acylated,if the carbohydrateis converted into the 2,1':4,6-bisacetalprior to the treatment with a lipase (NovozymTM435) [2241. On considering hydrolysis, several enzymes were investigated[225-2291. Depending on the biocatalyst used, acetyl groups from different positions of octaacetyl sucrose 141 could be removed selectively in usehl yields. For instance, alcalase and protease N preferably attack the acetate on Ol'[226s2301, the lipase from Candida cylindracea preferably liberates the OH-group on C4'of the furanoid ring[225,2301 and wheat germ lipase preferentially liberates the 1'-, 4'- and 6'-OH-groups (Fig. 1817)[223* 2311. The deacylation of the octaacetates of cellobiose, lactose, maltose and melibiose with Aspergillus niger lipase leads to the formation of the respective carbohydrate heptaacetates with a free anomeric OH-group at C1 in high yield[230,2321 . Wlth

1380

I

18 Introduction and Removal of Protecting Groups

HHOo

eHOo

*

HO

HO H

OH

oHO @HO o

~

OR

HO 137 lactosides 71-73% R = Me, Bzl

136 cellobiose 47% subtilisin, trichloroethyl butyrate, DMF

0

subtilisin, DMF

K

R

138

OCH,CX, 12-64% X = CI, F

L

I

a-glucosidase

HO O HO V alcalase or protease N 65-74%

TOH

140

54%

AcO AcO

OAc

141 Figure 18-17.

Selective enzymatic protection and deprotection of disaccharides.

prolonged reaction times, the acetates at C1 and C2 are hydrolyzed from cellobiose and lactose octaacetate in 51 % or 42% yield, respectively. 18.5.4

Nucleosides1’35,2331

The directed protection of nucleoside functional groups is a fundamental problem in nucleoside and nucleotide chemistry. Although several chemical methods are available for the regioselective acylation of the nucleoside carbohydrates, enzymatic

18.5 Protection of Hydroxy Croups

methods offer significant advantages with respect to yield, regioselectivity and the number of synthetic steps which have to be carried out. Earlier studies focussed on the use of the dihydrocinnamoyl group as an enzymelabile nucleoside protecting function which can be removed through the agency of a~ h y m o t r y p s i n [2351. ~ ~ ~Although . the enzyme shows an interesting tendency to attack preferably the S'-position, this technique was not exploited further. Highly regiodiscriminating biocatalyzed acyl transfer reactions to the carbohydrate parts of various nucleosides could be carried out again employing the protease subtilisin together with dimethylformamide as solvent. In particular, a mutant of this enzyme, obtained via site specific mutations appears to display advantageous properties. It transfers the acetyl group from isopropenyl acetate to the primary hydroxy functions of various purine and pyrimidine nucleosides and 2'-deoxynucleosides142 in high yields (Fig. 18-18)[2361. Commercially available subtilisin (protease N from Amano) provided the same compounds with identical yields and selectivities, however, five times more enzyme is required for this purpose. In addition, in the transfer of butyric acid from trichloroethyl butanoate to adenosine and uridine, carried out earlier I'[, this biocatalyst showed inferior properties with respect to regioselectivity and yields. The selective introduction of protecting groups into the hydroxy functions of different nucleosides can also be achieved by means of lipases. Thus, unprotected pyrimidine and purine 2'-deoxynucleosides143 (X = H) are selectively converted into the 3'-O-aqlated derivatives 144 in 6 6 8 2 % yield making use of lipase from Pseudornonas cepacia (PSL) and employing oxime carbonates as acyl donors (Fig. 1818)[237-2391. Similarly, by applying oxime esters or acid anhydrides, different ester functions can be selectively introduced into the 3I-position of nucleotides by using the lipases from Candida cylindracea (CCL),porcine pancreas (PPL) or Pseudomonas cepacia (PSJd)[240-244].If lipase from Candida antarctica (CAL) is used, however, the esters and carbonates are predominantly generated at the primary 5'-OH group of (deoxy)nucleotides[238, 239, 241* 242, 244-2471. Furthermore, in the case of ribonucleotides, complete regioselectivity can be achieved by using the same methodology[241]. The regioselectivity of the CAL-catalyzed alkoxycarbonylation is profoundly influenced significantly by the structure of the starting oxime In the alkoxycarbonylation of thymidine the use of the phenyl derivative leads to almost exclusive formation of the 5' carbonate, while the corresponding ally1 carbonate is introduced without any regioselectivity. An investigation of the enzyme-catalyzed acylation of a-, xylo-, anhydro-, and arabino-nucleosidesshowed that in these cases the primary 5'-hydroxygroup can be selectively acylated using lipase from Candida antarctica (CAL)[249-2511. A selective derivatization of the 3'-OH-group,however, was unsuccessful. When acylations of nucleosides with acid anhydrides in the presence of lipase from Pseudomonaspuorescence (PFL) in DMF or DMSO as the solvent first proceeded, the regioselectivity was However, this lipase together with subtilisin can be utilized to effect highly specific deacylations of various pyrimidine nucleosides 145 (Fig. 18-18)[2531. Thus, lipase from Pseudomonas Jluorescence (PFL) preferably attacks the hexanoyl group on the secondary hydroxy function of the N-

1382

I

18 Introduction and Removal of Protecting Croups

subtilisin 8350 or protease N

JOk, DMF

HO R 142

I R

HO

R

H

OH

H

OH

H

OH

quant.

90

80

80

80

65

I

I

yield [%I

X=H,OH

base

x = H, PSL, THF 60°C

oxime carbonaL carbonate HO

HO

X

0 144 64-82%

143

45-68%

0

base = A, U,T R = alkyl. vinyl, ally1

oxime carbonate = R O K O / N

0

su btilisin 12-31%

X

b

3

'

X

= H, Br, F, CH3

Pseudomonas fluorescence 58-74% 145 Figure 18-18.

nucleosides.

Selective enzymatic protection and deprotection of the carbohydrate parts o f

18.5 Protection of Hydroxy Croups

I

1383

glycosides, giving rise to the 5-esters in good yields. On the other hand, subtilisin gives rise to the $esters with moderate results. It should be noted, however, that in both cases from considerable to large amounts (6-71%) of the completely deprotected nucleosides were also formed. Subtilisin in phosphate buffer also selectively hydrolyzes the 5'-acetate of purine and pyrimidine triacetylated esters to give the A similar prefercorresponding 2',3'-diacetylribonucleosidesin 40-92 % ence was observed for the lipase from porcine pancreas, but with poorer selectivity and a slower reaction rate. This enzyme, however, deacetylated the deoxynucleoside 3',5'-di-O-acetylthymidine at the 5'-position in almost quantitative yield[255! In contrast, if lipase from Candida cylindracea (CCL) was used in the catalysis, the 3'-ester of this diacetate was preferentially hydrolyzed [2551. Using acetyl esterase of the flavedo of oranges, bisacylated purine deoxynucleotides can be selectively deprotected at the 3'-hydroxy group in 31-40 % yield [741. Interestingly, by introducing a phenylacetyl group for amino protection in the purine moiety the regioselectivity of the acetyl removal is reversed. Now the primary acetate is hydrolyzed by acetyl esterase in 22-52 % yield. In addition, the complete hydrolysis of an anomeric mixture of peracetylated 2'-deoxynucleosidesby wheat germ lipase or porcine liver esterase has been used to synthesize the pure p-anomer of e.g. thymidine, this being the only completely deprotected product [2561. The alcoholysis peractylated uridines catalyzed by Candida antarctica lipase leads to the formation of the completely deprotected nucleotide[2571. Although this reaction can be stopped after removal of the first acetyl group, no regioselectivity was observed for the formation of di-0-acetyluridine. 18.5.5

Further Aglycon Clycosides

In addition to nucleosides, several other naturally occurring carbohydratederivatives can be selectively protected/deprotected by means of enzymatic techniques. For instance, salicin 146, a wood component that contains a primary hydroxy group located in a glucose moiety and a second one in a benzylic position, was butyrylated exclusively at the 6-OH of the monosaccharide in 35 % yield by applying subtilisin Under the same conditions, in and trichloroethyl butanoate in DMF (Fig. 18-19)I'[. riboflavin (vitamin B2) 147 only the primary alcohol was esterified in 25 % yield["'], and colchicoside 148a as well as a thio analog 148b were converted into the 6'-butanoates by treatment with trichloroethyl butanoate in pyridine in the presence of subtilisin [2581. The corresponding 6'-acetatesof 148a,bwere obtained by treatment with vinyl acetate in the presence of Candida antarctica lipase as the biocatalyst (Fig. 18-19) Similarly, the carbohydrate parts of flavonoid disaccharides were regioselectively functionalized. Thus, for instance in the disaccharide rutin 149 and the related hesperidin only the 3"-OH group of the glucose moiety was esterified upon treatment with trifluoroethyl butanoate and subtilisin in 53 % yield (Fig. 1819)[2221. In the presence of lipase from Candida antarctica, however, both the 3"- and the 4"'-positions were acetylated['621. If only the glucose moiety is present in the molecule, as in the related isoquercitrin 150, the regioselectivty in the subtilisin-

1384

I

18 lntroduction and Removal of Protecting Groups

HHOO

G o HO

R

fq

R = Me 0

HAc

Me0

X=O: 148a X=S: 148b

subtilisin, pyridine, 86% 0 -OCH2CF3

.

or CAL, vinyl acetate f-amyl alcohol

0

XMe subtilisin, pyridine, 53 % 0

0-

-OCH2CF3 W H + O R HO Y HO

or CAL, vinyl acetate f-amvl alcohol. 91 %

149 rutin

H

o/ e HO

0

:

:

CAL, f-amylvinyl alcohol, acetate 79 %

B or CAL, vinyl cinnemate acetone, 68 %

150 isoquercitrin Figure 18-19.

Selective enzymatic acylation of aglycon glycosides.

catalyzed reaction was less pronounced[259].However, in the presence of lipase from Candida antarctica the 3",6"-bisacylatedproduct is formed if vinyl acetate is used as the acyl donor['62]. Interestingly, by using vinyl cinnamate as the acyl donor, this biocatalyst only acylates the primary 6"-hydroxy group[260].Naringine 151 was converted into the 6-glucosyl ester in the presence of subtilisin (Fig. 18-20). In all cases the rhamnose and the phenolic hydroxyls remained unattacked (for the protection of phenolic hydroxy groups in flavonoids see Sect. 18.5.8). The steroidal glucoside ginsensoside Rgl 152 can be selectively monoacylated in high yields at the 6'-position using Candida antarctica lipase as the biocatalyst1261,2G2]. In this case, similar results were obtained with different acyl donors such as vinyl acetate, dibenzyl malonate and bis(trichloethy1)malonate (Fig. 18-20). Two impressive examples of selective enzymatic deacylations of complex sub-

78.5 Protection ofHydroxy Groups

I

1385

I

subtilisin, pyridine, 49 % 0 40CH2CF3

151 naringin

CAL, f-arnyl alcohol vinyl acetate (87 %) dibenzyl rnalonate (85 %) bis(2,2,2-trichloroethyl)rnalonate (71 %)

OH 152 ginsenoside Rgl Figure 18-20.

Selective enzymatic acylation of aglycon glycosides.

strates consist in the removal of all acetates from the peracetylated fi-D-glucopyrand of the gibberellinicacid derivative 154[264], anosyl ester 153 of abscisinic acid[263] containing one glucose tetraacetate glycosidically bound and a second one attached as an ester (Fig. 18-21).In both cases the removal of the acetyl groups by chemical methods in particular was complicated by an undesired cleavage of the ester linkages to the glucoses. However, the four acetyl groups present in 153 could be hydrolyzed chemoselectively by means of helicase, an enzyme occurring in the seeds of Helianthus annus, whereby the unprotected glucose ester was formed in 82 % yield without destroying the ester bond between abscisinic acid and glucose. Similarly, the biocatalyst removed all acetates from 154.In this case the yield reached only 8%, it should, however, be kept in mind that ten acetic acid esters had to be cleaved in the enzymatic process and that the aglycon is rather complex. In conclusion, the various enzyme-mediated protecting group manipulations carried out on numerous carbohydrate derivatives indicate that biocatalysts can be used advantageously in the protecting group chemistry of carbohydrates. In particular, subtilisin and several lipases from different sources (from porcine pancreas, from Candida cylindracea, Aspergillus niger, Chromobacterium viscosum, Mucor javanicus, Pseudomonas fluorescence and from wheat germ) allow the chemo- and regioselective acylation and deprotection of various saccharides, the structures of

1386

I

18 Introduction and Removal of Protecting Groups

"+"'/=
w. 0

AcO

0 -0

H

AcO

--0

H&

0

AcO

153 helicase 82%

154

OAc

8%

helicase

O

Figure 18-21.

Enzymatic deprotection of complex glucosyl esters.

which differ widely, to be carried out. A general principle that emerges from these studies is that the enzymes exhibit a predominant preference toward primary hydroxy groups. If these functionalities are not present or protected, the biocatalysts are capable of selectively manipulating secondary hydroxy groups or the esters thereof. In the introduction and removal of acyl groups, the regioselectivitydisplayed by the enzymes often parallels the findings recorded for classical chemical transformations, although it is significantly higher in many cases. Furthermore, in several cases regioselectivities were observed in the biocatalyzed processes which can not or only slightly be achieved by means of chemical methods. Finally, it should be realized that subtilisin and the lipases are capable of introducing specific acyl groups into the carbohydrates which can later be removed selectively by different enzymatic or chemical methods. 18.5.6 Polyhydroxylated Alkaloids

The plant alkaloid castanospermine 155 and the related piperidine alkaloid l-deoxynojirimicin 160,like several other polyhydroxylated octahydroindolizidines,piperidines and pyrrolidines, are potent glycosidase inhibitors. These nitrogen bases are of considerable interest for the study of biosynthetic processes and, in addition, castanospermine and some of its derivatives may be of clinical value as antineoplastic agents and as drugs in the treatment of AIDS. In the light of the analogy between the structures of these alkaloids and glucose, some of the above mentioned enzymatic methods for the selective functionalization of carbohydrates were applied to prepare several acyl derivatives of 155 and 160. Thus, subtilisin transfers the acyl moieties from several activated esters to the 1-OH group of the bicyclic base in moderate to high yields (Fig. 18-22)[2G52 266] . A gain,

I

18.5 Protection ofHydroxy Croups

R = CH3 = CHTPh

} R = CH=CH2

R = Ac-L-Phe R = (CH2hCI Ac-D-Ala

157 R = n-C3H7 CVL ~ O C H THF 72%

OH 161

56%

1.5 equiv. trichloroethyl butyrate

~

C

C

~

~

-

subtilisinl HO HO

subtilisinl pyridine

HO OH 160

6 equiv. trichloroethyl butyrate

HO HO

162 Figure 18-22.

1387

77%

Selective enzymatic protection of polyhydroxylated alkaloids.

pyridine had to be used as the solvent for the polyhydroxy compound. The monoesters 156 obtained by this technique, like the monoesters of hexoses could subsequently be dissolved in THF and were further acylated by means of different enzymes, e. g. to the 6-butanoate 157 and the 1,7-dibutanoate158. Finally, the 1-ester was removed from 158 by subtilisin in aqueous solution to deliver the 7-butanoate 159 in 64% yield. In contrast to castanospermine, 1-deoxynojirimicine 160 contains a primary hydroxy group as well as a much more nucleophilic amino function. If a small excess of trifluoroethyl butanoate is employed, subtilisin converts this alkaloid preferably into the 6-monoester 161 (Fig. 18-22)I2"]. However, with 6 equiv. of the acylating agent, the 2,G-diester 162 is formed in 77% yield. This diester 162 may be subsequently deacylated regioselectively at the 6-position by means of several different enzymes.

1388

I

18 lntroduction and Removal offrotecting Croups

It should be noted that under the conditions of the enzymatic acylation the amino group is not derivatized, an observation which has also been made in related cases [266. 2671, e. g. N-terminally deprotected serine-peptides. 18.5.7

Steroids

Enzymatic acyl transfer reactions are also practical processes for the acylation of hydroxy groups in steroids. The lipase from Chrornobacteriurn viscosurn (CVL) for instance selectively transfers butyric acid from trifluoroethyl butanoate to equatorial (B) C3-alcoholic functions that are present in a variety of sterols, e. g. 163 and the respective 5,6-didehydrocompound (Fig. 18-23)[268].Axially oriented alcohols at C3 and secondary alcohols at C17 or in the sterol side chains are not derivatized. In addition to the equatorial alcohols, the compounds being accepted as substrates by the lipase must have the A/B-ring fusion in the trans configuration. In the B-ring a double bond is tolerated, in the A-ring, however, it is not. Similarly, lipase from Candida antarctica acylates the 3-hydroxy group in steroids such as 163 and its 5,6-didehydro derivative[269].Interestingly, acylation in this position is preferred regardless of the orientation of the hydroxy group. For instance, treatment of 164 with vinyl acetate in the presence of Candida antarctica lipase leads to the formation of corresponding 3-acetylated derivative in 82 % yield. In contrast, subtilisin does not recognize the hydroxy group at C3 of the steroid nucleus, but rather transfers the acyl moiety to alcoholic groups in the 17-position or in the side chains (Fig. 18-23). Changes in the A- or in the B-ring do not dramatically influence the selective mode of action of this biocatalyst. This behavior is the same as that determined for the lipase of Pseudomonas cepacia, which was recently used for the regio- and stereoselective acylation of steroids r2’O]. Thus, using these enzymes, the completely regioselective protection of either alcoholic group in several steroid diols is possible. This feature opened a route to a new chemoenzymatic process for the oxidation of selected positions of the steroid framework via an enzymatic protection/oxidation/ deprotection sequence. Chemoenzymatic approaches of this type are expected to provide attractive alternatives to the currently utilized enzymatic oxidation of steroids by hydroxysteroid dehydrogenases. A further biocatalyst comes into play when bile acids serve as starting materials, e. g. deoxycholic acid methyl ester 165[271].The cis-configuration of the A/B-ring fusion prevents the application of lipase from Chromobacteriurn viscosurn (CVL) and the aliphatic chain hinders the esterification of the C12a hydroxy group by subtilisin. The lipase from Candida cylindracea (CCL) has proved to be the most suitable enzyme for the enzymatic acylation of bile acids. In hydrophobic solvents, i.e. hexane, toluene, butyl ether, benzene, etc. (except acetone) and employing trichloroethyl butanoate as the acyl donor, the 3a-0-butanoyldeoxycholic acid methyl ester 166 is formed in 80% yield without any by-products, suggesting that the enzyme is ineffective towards 12a-OH. In addition, the 7a-OH and the 7p-OH, present in 167 and 168 are not esterified by the enzyme. In both cases, the 3-butanoate is also formed (Fig. 18-23).

18.5 Protection of Hydroxy Croups

I

1389

163 3-monoburyrate 83% CVL: CAL: 3-monoburyrate subtilisin: q7-monobutyrate 60%

CCL, trichloroethyl butyrate, hydrophobic solvent 164 R = R' = R2 = OH, R3 = OH 165R = R1 = RZ = H, R3 = OH 80% 166 R = But. R1= R2 = H, R3 = OH 167 R = But. R1 = H. R2 = OH. R3 = H 168 R = But, R1 = OH, R2 = H, R3 = H

171 CCL: 3.1 7a-dihydroxyestradiol 60% 3-hydroxy-I 7a-acetoxyestradiol 25%

CCL: 169 R = 3a-OAc, R = 17P-OAc no reaction 170 R = ~P-OAC, R " = 17p-OAc -+ SP-OH, R = 17P-OAc 79%

172 R1=R3=OAc,R2=(0), R4=R5=H 173 R'=R5=OAc, R2=(0), R3=R4=H 174 R1=R2=R3=OAc,R4=R5=H Figure 18-23.

175 R ~ R~=OAC. = R~=(o) 176 R1=OAc, R2 =C(0)CH3, R3=H

Selective enzymatic protection of steroids.

Saponification of steroid esters can also be steered with Candida cylindracea lipase (CCL)[272. 2731. This process occurs in the presence of octanol in organic solvents and is characterized by a pronounced stereospecificity and regioselectivity. Thus, the 3a-

1390

I

18 Introduction and Removal ofprotecting Croups

esters of 3a,17P-diacetoxysteroid 169 resisted liberation, whereas the 3g-isomer 170 is transformed into the corresponding alcohol in 79% yield. The 17a-acetate of 3,17a-diacetoxyestradioll71 is also saponified, but at a slower rate than the 3-acetate (Fig. 18-23). In the case of the androstane derivatives 172 and 175 different Thus, the selectivities of Candida antarctica lipase (CAL) and CCL were alcoholysis of 172 in the presence of CAL afforded the C3 deprotected product in 75% yield whereas CCL led to the removal of the acetate at C1G in 66% yield. Treatment of 173,174 and 176 with CCL led to the cleavage of the C3 acetate in 79%, 2741. 87% and 83 % yield, 18.5.8

Phenolic Hydroxy Groups

Polyphenolic compounds occur widely distributed in nature and may possess a variety of interesting biological properties, e. g. antibiotic, antiviral and antitumor activity. The synthesis and further elaboration of these compounds often requires the selective protection or deprotection of specific phenolic hydroxy groups. To achieve this goal, the methods highlighted above for the various aliphatic polyols can also be applied successfully. For example, for the the enzyme-catalyzed acetylation of phenols six different 2761. Out of these, only the lipase from lipases was initially screened for activity[275. Chromobacterium viscosum (CVL) showed significant activity. In a subsequent study, the lipase from Pseudomonas cepacia (PSL) turned out to be a more efficient biocatalyst, which was succesfully used for the regioselective acylation of various aromatic dihydroxycarbonyl compounds [2771, and (+)-catechin[2781. Thus, by using PSL as the biocatalyst the dihydroxy aldehydes and ketones 177, 178 and related compounds were selectively acetylated in conversions ranging from 20 to 97 % using vinyl acetate as the acyl donor (Fig. 18-24)[2771. (+)-Catechin179 was also subjected to irreversible acyl transfer conditions. In this case, both the 5- and 7-monoacetates Interestingly,the inability of were obtained in 40% and 32% yield, respectively[278]. the lipase from Aspergillus niger to acylate aromatic hydroxy groups has consequently been used for the selective acylation of primary aliphatic hydroxy functions in molecules containing both aromatic and aliphatic OH-groups [2791. In fact, even PSL preferentially acylates primary aliphatic hydroxy groups if they are present in the compound [280]. In the deprotection of peracetylated polyphenolic compounds a somewhat different scheme has emerged. In this area, a broader spectrum of lipases has been used successfully. For example, the pentaacetyl derivative of catechine 179 was treated with PSL under alcoholysis conditions (THF, n-butanol) to give the 3,3',4'-trisacetate in 50% yield after 12 On longer exposure to the biocatalyst, the 3-monoacetylderivative was isolated in 95 % yield. Thus, the coumarine 180, the chromanone 181, the chalcone 182, the flavanone 183 as well as several flavones, e.g. 183 and 185 were regioselectivelydeacylated by employing different lipases in organic solvents (Fig. 18-24). Porcine pancreatic lipase (PPL) predominantly attacks one of the phenolic acetates present in 180-183 with

18.5 Protection of Hydroxy Croups

I

1391

0 &R HO

OH

' OH

OH

177a R=H (78 Yo) 178a R=H (20%) 177b R=CH3 (97 Yo) 17Bb R=CH3 (20 %) 1 7 7 RzCHZCH3 ~ (93 Yo) 178C R=CH,CH3 (22%)

wo

(only conversion given)

OH

179 (+)-catechin

(only conversion given)

AcO

\

AcO

CH3

180 PPL, 65%

183 PPL,55%

185 PPL, 78%

0

181 PPL, 73%

AcO

\

181 PPL,50%

0

184 PCL,55%

186 PCL, 95%

187 PPL, 75%

Figure 18-24. Selective enzymatic protection and deprotection o f polyphenolic compounds.

good to high regioselectivity and produces the respective selectively protected compounds available in good yields [281-2831 . The flavone acetates 184 and 186 can be partially deacylated with high regioselectivityby transesterification using lipase from Pseudomonas cepacia (PSL) and n-butanol in THF. [284s2851 However, in other cases the positional specificity displayed by the enzyme was less pronounced. This technique has allowed for an efficient construction of a selectively 0-methylated flavonoid [2841. In addition, aryl alkyl ketones which are important starting materials for the synthesis of polyphenolic natural products may be manipulated selectively by making use of an enzymatic saponification[283,285-2871 . 1n general, in these cases the sterically better accessible ester groups are cleaved, as for instance in 185[2851.All of these examples have in common the fact that a carbonyl group is either directly or vinylogously attached to the aryl moiety. Without such a function present in the

1392

I

78 introduction and Removal of Protecting Groups

molecule, the biocatalysts failed to differentiate the ester groups or completely deacylated the substrates. However, by using the lipases from porcine pancreas (PPL)or Candida cylindracea (CCL)immobilized on microemulsion-basedgels it was possible to monodeacylate resorcinol and related diesters such as 187 in high yields [lSs1.Alternatively, by using tert-butyl methyl ether saturated with water as the solvent, it was possible to monodeacetylate diacetoxynaphthalenes selectively~288~. The influence of the solvent was exemplified by charging the solvent system to acetone/buffer: under such conditions only completely deacylated products were obtained.

18.6

Biocatalysis in Polymer Supported Synthesis: Enzyme-labile Linker Groups

Combination chemistry and parallel synthesis of compound libraries on polymeric supports are efficient methods for the generation of new substances with a predetermined profile of propertie~[~*~-~’~1. The anchoring of one reactant to a polymeric support has the advantage that an excess of reagent may be used, while purification is kept manageable. This is particularly important if the reaction is to be carried out with several reactants in the same reaction vessel. Solid phase synthesis involves the use of linkers between the compounds to be varied combinationallyand the solid supports which are stable during the reactions. These linkers have to be cleavable as desired, usually at the end of the synthetic sequence, with high selectivity and in good yield, without affecting the structure(s) of the product(s) that are released from the polymeric supports. Linkers have previously usually been cleaved by classical chemical methods, for instance using strong acids. Such conditions often restrict the application of the linkers, i. e. acid-sensitivelinkers are not suitable for acid-labile compounds, such as carbohydrates. Specific linkers have therefore been developed for acid-labile compounds, such as silylether linkages, thioether linkages [2921, and ester linkages [2931. Although such linkers may be cleaved in the presence of acid-labile groups, they have the disadvantage that they are themselves quite labile to common chemical reagents that one might want to employ on the solid phase. For example, esters and silylethers are unstable to bases and thioethers are unstable in the presence of oxidants, such as rn-chloroperbenzoic acid, and to electrophilic reagents, such as alkylating agents. In principle linker groups are polymer-enlarged versions of blocking functions used in regular solution phase chemistry. Therefore, enzymatic transformations that may be employed for the removal of protecting groups in solution in principle may also open up alternative opportunities for releasing compounds from polymeric supports. The linkers developed so far can be divided into exo- and endo-linkers (Fig. 18-25) cleavable by exo- endo-enzymes, respectively, as proposed by Flitsch et al. [2941. Exo-linkers are composed of three units: (i)a group providing the site for enzyme catalyzed hydrolysis (R1); (ii) a site for attachment of the target molecule (R3); and (iii) a site for attachment to a further optional spacer (R2).

18.6 Biocatalysis in Polymer Supported Synthesis: Enzyme-labile Linker Groups

Exo-linkersd e a d by exo-enzymes

Enddinkers cleaved by endhemymes

R’: group providingthe site for enzyme catalyzed hydrolysis, R‘: optional intermediatelinked to a solid support, R3: residue to be synthesized and varied in the course of a synthesis on the support, X: 0, N(H). N(R”). C(O)O, S, C(O)N(H) or C(O)N(R), R” is a noninterfering substituent. Y: 0 or NH.

Figure 18-25.

Graphical representation of exo- and endo-linkers.

Endo-linkers are linkers in which the target molecule (R3), the group, which provides a site for enzyme catalyzed hydrolysis (R’) and a further optional spacer (R’) are attached to the polymeric support in a linear arrangement. By means of enzyme mediated dissection, the target molecule, in many cases tagged with the functional group recognized by the enzyme, is released. Examples of endo-cleavablelinkers have been reported (Table 18-5).However, in many cases the product is tagged with part of the linker. For instance, the endopeptidase chymotrypsin cleaves endo-linkers towards the middle of a peptide-chain or “internally”.Not only does this limit the methodology to a very small number of enzymes, but it may also restrict the structure of molecules that can be generated. For instance, this method will typically (but not necessarily, see Figs. 18-26 and 18-27)generate compounds containing C-terminal aromatic amino acids, which are necessary for recognition by chymotrypsin. By contrast, exo-linkers do not restrict the structure of the reactant and can be cleaved by more readily available exoenzymes, which act at the end of a chain or “externally”(Table 18-5). Furthermore exo-cleavable linkers yield untagged products upon cleavage from the solid support. 18.6.1

Endo-linkers

For a better overview, examples of endo-linkers and the enzymes used for the cleavage of the product from the solid phase which have been described in the literature so far are given in Table 18-5. Wong and coworkers[’”I introduced a silica-based solid support with a specific enzymatically cleavable linker for the synthesis of glycopeptides and oligosaccharides. They found that styrene- and sugar-based polymers tend to swell which leads to a low coupling yield. Their choice of solid support is aminopropyl silica based on the

I

1393

1394

I

18 Introduction and Removal of Protecting Croups Table 18-5.

Examples o f endo-linkers and the appropriate cleavage enzymes.

Linker

Enzyme

Examples

Ref.

a-Chymotrypsin

Oligosaccharide synthesis

[297-2981

Ceramide glycanase

Oligosaccharide synthesis

[299]

Peptide synthesis

[301]

0

R

O

oAo;,oa

Phosphodiesterase

0

HO 0

facts that: (a) it is compatible with both aqueous and organic solvents, (b) it has a large surface area accessible to biomolecules, and (c) it has sufficient density of functional groups. A hexaglycine spacer was attached to the solid support to give a substitution of 0.2 mmol g-' of dry silica and the excess amino groups were then capped using acetic anhydride. In the next step a selectively cleavable, a-chymotrypsin sensitive, phenylalanine ester 189 was implemented for the release of the products from the solid support under mild conditions. Then it was transformed to 190 followed by reactions with glycosyl transferases to yield 191. Finally, the desired glycopeptidewas cleaved from the solid support in high yield by treatment of 191 with a-chymotrypsin (Fig. 18-26). Nishimura and coworkers1296-2971 described a novel method for the enzymatic synthesis of oligosaccharide derivatives employing an a-chymotrypsin sensitive linker. The synthesis of the water soluble GlcNAc-polymer 197, sensitive to achymotrypsin, is shown in Fig. 18-27. Oxazoline derivative 193 was coupled with 6-(N-benzyloxycarbonyl-~-phenylalanyl)-amino-hexanol-l (194) followed by N-deprotection of the phenylalanine and subsequent condensation with 6-acrylamidocaproic acid 195. De-0-acetylationgave the polymerizable GlcNAc derivative 196. Finally, copolymerization of acrylamide and monomer 196 in the presence of ammoniumpersulphate (APS) and N,N,N,N-tetramethyl ethylene diamine (TMEDA) gave the

( 1 ) 25% TFA (CH2C12) (2) Boc-Gly-OH ((7 eq), BOPIHOBt,DIEA (3) 25% TFA (CH2C12) (4) Boc-Asn(G1cNAcp)-OH

0

-

190

glactosyl transferase, sialyl transferase

PH (1) a-chymotrypsin, H20. pH 7.0

(3) a-l.3-fucosyltransferase, GDP-Fuc (2.5 eq), 0.1 M HEPES (pH 7.0)

.

!I

0

192

1396

I

18 Introduction and Removal ofprotecting Croups Figure 18-27. Oligosaccharide synthesis and a-chymotrypsin catalyzed release from the solid support.

(1) Z-Phe-NH-(CH2)6-OH(194) CSA, (CHCI&, 70" C (2) H, Pd/C, MeOH. 50" C (3) CH2=CHCONH(CH,),COOH (195) EtOH-C,H, (4) MeONa (cat.), MeOH/THF

CH,=CHCONH, TMEDA. APS

Ig7 X : Y = 1 : 4 Galactosyl transferase Sialyl transferase

OH

:v

/-

I

198

C6H5

a-chyrnotrypsin Tris-HCI buffer PH 7 . a , 4 v c

199

0 (cH HZN 2)5N3:

X:Y=1:4

18.1 Biocatalysis in Polymer Supported Synthesis: Enzyme-labile Linker Groups

A;i&+Co.o+ OAc

OAc

nu

OAc

bH OH

I

4

-N H

NHCOOCHpC6H, (1) Pd/C, MeOH (2) 195, EEDQ, EtOH-C& (3) MeONa

0 CH*=CHCONH2 TMEDA. APS DMSO-H20,50° C

0

,

HO OH OH

0

202

.OH

HO

1

bH

X:Y=1:5 CMP-NeuAc a -2,3-sialyltransferase BSA, MnCI2,ClAP 50 nM sodium cacodylate buffer, pH 7.49

OH ceramide

1 H N K0( C H 2 ) 5 '

ceramide glycanase Triton CF-54 sodium citrate buffer pH 6.0, 37" C

(GM3) Figure 18-28.

,,

204

Ceramide glycanase mediated release by transglycosylation.

0

X:Y=1 :5

I

1397

1398

I

18 Introduction and Removal ofProtecting Croups

polymer 197 in high yield. The polymer 197 was then subjected to galactosylation and subsequent sialylation with the corresponding glycosyl transferases to yield 198. The final product 199 was cleaved from the water-soluble support by treatment with a-chymotrypsin at 40 "C for 24 h in 72 % overall yield from 197. Nishimura and Yamada [29s1introduced a water-solublepolymeric support having a linker recognized by ceramide glycanase for a synthesis of ganglioside GM3 (204). Synthesis of the polymerizable lactose derivative 201 with a ceramide glycanase sensitive linker is shown in Fig. 18-28. The lactosyl ceramide (LacCer) mimetic glycopolymer 202 is obtained from the monomeric precursor 201 by co-polymerization with acrylamide. This solid support 202 was converted into the intermediate product 203 by sialylation using PGa1l-t 3/4GlcNAc a-2,3-sialyltransferase.Finally, the polymeric support was cleaved by transglycosylation with leech ceramide glycanase in the presence of excess ceramide as the acceptor to give the desired product 204 in high yield (Fig. 18-28).An advantage of the water-solublepolymer is that the transfonnation can be monitored by NMR spectroscopy during the enzymatic glycosylation steps. Arrays of up to 1000 peptide nucleic acid (PNA) oligomers of different sequence were synthesized by Jensen et al. on polymer membranes (Fig. 18-29)[2991. The PNA chain was linked to the peptide spacer glutamic acid-(y-tee-butylester)-(c-aminohexanoic acid)-(c-aminohexanoic acid) (Glu[OtBu]-~Ahx-cAhx) via an enzymatically cleavable Glu-Lys handle. The Glu[OtBu]-~Ahx-~Ahx spacer was coupled to the amino-functionalized membrane by standard Fmoc-Chemistry. Then the membranes were mounted in an ASP 222 Automated SPOT Robot and a grid o€ the desired format was dispensed at each position. The free amino groups outside the spotted areas were capped and further chain elongation was performed with Fmocprotected PNA monomers to synthesize the desired PNA oligomers. After completion of the synthesis, the PNA oligomers were cleaved from the solid support by incubation with bovine trypsin solution in ammonium bicarbonate at 37 "C for 3 h. One of the very first papers concerning endo-linkers was published by Elmore et al. (Fig. 18-30)[3001. They described a new linker containing a phosphodiester group for solid phase peptide synthesis using a Pepsyn K (polyacrylamide) resin. After completion of coupling and deprotection cycles, the phosphodiester 207 was cleaved with a phosphodiesterase. In this way p-casomorphin, Leu-enkephalin and a col-

*05 Figure 18-29.

AOOH

\

trypsin

Trypsin mediated cleavage of a peptide bond in PNA oligomer synthesis.

78.G Biocatalysis in Polymer Supported Synthesis: Enzyme-labile Linker Groups

I

206

I

1399

Synthesis via symmetrical anhydrides of N-Fmoc amino acids

Fmoc-Ala-Pro-Gly-Leu-Ala-Gly-0 T

L

O

\

II

C

0

*07

1

O

A

/qzbHN-@

N H

1

Phosphodiesterase,pH = 5.7,7d (83%)

Fmoc-Ala-Pro-Gly-Leu-Ala-Gly-OH 208 Figure 18-30.

Synthesis of a collagenase substrate on a phosphodiesterase-scissile linker.

lagenase substrate were synthesised in high yields. In the context of enzymatic cleavage of linkers on polymeric supports particular attention was paid to the general question of whether enzymatic transformations on resins are viable and high yielding. An in-depth treatment of this problem is beyond the scope of this review. However, a few examples for the application of biocatalyzed transformations on solid supports will serve to illustrate that such transformations can indeed be employed advantageously for various purposes. Meldal et al. described the proteolytic cleavage of the alanine-tyrosine bond in a resin-bound decapeptide by treatment with the 27 kDa protease subtilisin BNP’ to demonstrate the accessibility of the interior of the newly designed S P 0 C C - r e ~ i n [ ~ ~ ~ ] to enzymes L3021. Furthermore, enzymatic hydrolysis of model isopeptides F-oligo(L-methiony1)-Llysine from Bio-beads L3O3] by pepsin, chymotrypsin, cathepsin C (dipeptidyl peptidase IV) and intestinal aminopeptidase N was investigated using high-performance liquid chromatography to identify and quantify the hydrolysis products L3O4]. Larsen et al. reported the enzymatic cleavage of a desB30 insulin B-chain from a presequence (LyS(B0C))a. This spacer shifts the conformation of the growing peptide chain from a p-structure to a random coil conformation and reduces peptide-chain aggregation, which otherwise causes serious synthetic problems. Novasyn KA[3051 was employed as a solid support, but unfortunately, no information about the enzyme used was reported L3O61. Barany et al. were the first to exploit the different enzyme accessibilities of surface and interior areas of a given bead and the resulting differentiated bead was used to synthesize a peptide library on the surface and the code for this on the interior simultaneously[307]. This clever strategy is illustrated in Fig. 18-31. Selective cleavage of short Na-protectedpeptide substrates with chymotrypsin from the surface area of

1400

I

78 introduction and Removal ofProtecting Groups

*<-e Cleavage of the peptide bonds on the surface

r

P’

210

209

1

Attachment of Fmoc -S

1. Boc-cleavage 2.Coupling of Boc-C

Fm0c-S-

:*

FITDGS-P ’

‘

1

--

211

212

I.Fmoc- and Boc-cIeavage 2. Receptor sereeninu

a

Bead Selection

Hits

a

Code Analysis

Structure

Figure 18-31. Peptide encoded combinatorial peptide libraries via enzyme-mediated spatial segregation. P-P’: substrate with a scissile bond between P and P’; S : terminal residue o f t h e screening structure, C: terminal residue o f t h e coding structure.

TentaGel-AM-beads209 leaves the majority of the peptide attachment sites in the interior uncleaved to afford 210 (“shaving”methology).The first residue is attached using orthogonal FMOC-chemistryto provide 211. Coding is done by using standard BOC-chemistry on the interior of the bead to yield 212. Repetition of this process furnishes a surface peptide, which is encoded internally (213). This generation of two structures on the same bead allowed the investigation of the synthesized peptide library (1x lo5 members) with different receptors (anti-pendorphin antibody, streptavidin and thrombin). After the staining procedure had been carried out, the beads that showed a color were selected for sequencing and the coding peptides present within the bead were used to deduce the binding structures. This screening led to the discovery of a new thrombin ligand, which binds with an affinity one order of magnitude higher than the natural motif.

78.G Biocatalysis in Polymer Supported Synthesis: Enzyme-labile Linker Groups

OH

2. Precipitation

214 OH

HO

3. Filtration

OH

NP = o-Nitrophenyl

1401

1. P-Galactosidase

+

HO

I

, . o*O H HO OH 215

Lo,

I

- GIU OH

*+&I

H o & - o . ,

HO

OH

217b

i

OH 216b

OH

OH

OH

OH

Q + & . +

HO

OHHOHO

218a

Cleavage of the linker

+&/+&I

HO

OH

____)

Figure 18-32.

HO

HO

OH

218b

OH

General strategy for the liquid-phase synthesis o f disaccharides using glycosidases.

1402

I

18 fntroduction and Removal ofProtecting Groups

Fernandez-Mayoralas and coworkersL3081 used the high substrate specifity of enzymes in their synthesis of galactose-glucose-disaccharides(218) on an MPEGsupport[3o9].After galactosylation of glucose immobilized on the soluble support (215)using B-galactosidase,the unreacted monosaccharide glucose was removed by the combined use of a- and j3-glucosidases to obtain only MPEG-bound disaccharides (216, Fig. 18-32).Finally the disaccharides 218 obtained were released from the support by ethanolysis. Schmitz and Reetz described the solid phase enzymatic synthesis of oligonucleotides on Kieselguhr-PDMA-resinsvia T4 RNA ligase. Concomitantly,they found that RNase A selectively cleaves the last bound nucleotide at the ribose sugar leaving a 3 ' 5 ' - diphosphorylated oligomer behind on the resin, but application in actual synthesis has not yet been undertaken[31o]. 18.6.2

Exo-linkers

An exo-linkeraccordingto Fig. 18-25must contain an enzyme labile group R1, which is recognized and attacked by the biocatalyst. Possible combinations could be: phenylacetamidelpenicillin amidase, ester/esterase, monosaccharide/glycosidase, Table 18-6.

Examples o f exo-linkers and the appropriate cleavage enzymes.

Linker

R

X

y

O

G

Z

0

Enzyme

Examples

Ref.

Lipase

Picet-Spengler reaction, nucleoside immobilization

[313]

Penicillin acylase

Palladium cat. [318, 3191 C-C-couplings, Mitsunobu- and Diels-Alder reactions, 1,3-dipolar cycloadditions

Penicillin acylase

Immobilization of alcohols (e. g. Fmoc protected serine methyl ester, glycosides) and amines (e.g. phenylalanine)

m

0

H RX H

0

0

[312, 3131

18.6 6ioc:atalysis in Polymer Supported Synthesis: Enzyme-labile Linker Groups

I

1403

phosphate/phosphatase, su.lfate/sulfatase and peptides/peptidases L3l1].Up till now only the following systems have been worked out (Table 18-6). In independent and simultaneous investigations Flitsch and coworkers r311, 3121 and Waldmann and coworkers[313, 314] developed a selectively cleavable exolinker, which can be cleaved with penicillin G acylase, a commercially available and widely used enzyme[77]. Penicillin acylase catalyzes the hydrolysis of phenylacetamides and has been used in peptide synthesis for the cleavage of protecting 3151. In linker 219 developed by Flitsch and c ~ w o r k e r s [3121 ~ ~(Fig. * ~ 18-33) -XR represents the alcohol or amine group of the target molecule. Hydrolysis of the phenylacetamide moiety generates the hemiaminal221 which readily fragments in an aqueous medium and thereby releases the desired products, RXH. The thioethyl group present in the anchor group of 219 was activated by treatment with N-iodosuccinimide (NIS) followed by displacement with a variety of alcohols (223-225). To prove the possible application of this linker i n solid phase carbohydrate synthesis protected glycosides 226 and 227 were coupled to linker 219 and released enzymatically. Flitsch et al. also described the immobilization and enzymatic cleavage on a variety of a r n i n e ~ [ ~ l ~ ] . Nevertheless, the application of this enzyme-labile linker group in multi-step syntheses on the solid phase and subsequent enzyme-initiated release from the polymeric support has not been described yet. Waldmann and coworkers described designed exo-linker 228 r313, 3141 . Theanchor group comprises a 4-acy:loxy-3-carbo~benzyloxy group, which is recognized and attacked by the biocatalyst, so that a spontaneously fragmenting intermediate is generated, thereby releasing the desired compound (Fig. 18-34)[53, 54, 571. The linker 228 is attached as an amide to the solid phase. Cleavage of the acyl group by a lipase generated a phenolate 229, which fragments to give a quinone methide 230 and releases the product 231. The quinone methide remains on the solid phase and is trapped by water or an additional nucleophile. Following on from this cleavage principle, amines (bound as urethanes), alcohols (bound as carbonates), arid carboxylic acids (bound as esters) can be detached from the polymeric carrier. The substrate specificity of the enzyme guarantees that only the intended ester is cleaved. TentaGelS-NH2was chosen as the polymeric support, i. e. a polystyrene resin equipped with terminally NH2-functionalized oligoethyleneglycol units. It has a polar surface and swells in aqueous solutions allowing the biocatalyst access to the polymer The applicability of the enzyme-labile anchor group was demonstrated by the synthesis of tetrahydro.+carbolins 237 employing the Pictet-Spengler reaction (Figure 18-35).The benzylic alcohol group of the linker 232 was first esterified with Boc-L-tryptophan, and after its N-terminal deprotection the support-bound tryptophan 233 was reacted with aliphatic and aromatic aldehydes to give imines 234, which cyclized immediately in reasonable to high yields to the tetrahydro-0-carbolins 235. Lipase RB 001-05 :selectively attacked the acetate incorporated into the linker and generated the corresponding phenolate 236, which then fragmented spontaneously. Following these multistep transformations the desired tetrahydro-p-carbolins 237 were obtained in 70-80 % yield.

1404

I

18 Introduction and Removal ofprotecting Groups

Penicillin acylase

f+:-;b 1

I

222

FrnocHN

223 225

Figure 18-33.

Loading and cleavage of a penicillin acylase scissile linker.

18.6 Biocatalysis in Polymer Supported Synthesis: Enzyme-labile Linker Groups

0

Lipase

* N H y 2 3 0

Nu

e

1

HO

X-R

-RC02H

X = NH: amines (-COP) X = CR2: 0: alcohols carboxylic (-COP) acids

231

Figure 18-34. Principle for the development of the enzyme-labile 4-acyloxy-benzyloxy linker group.

Waldmann and coworklers developed a second exo-linker following a new approach[317,3181 which makes use of a safety-catchlinker. It is based on the enzymatic cleavage of a functional group embodied in the linker. In this way an intermediate is generated, which subsequently cyclizes intramolecularly according to the principle of assisted removal L2, 319-3221 and thereby releases the desired target compounds (Fig. 18-36). The linker group is immobilized as a urethane on the amino-functionalized carrier 238. It facilit,atesthe attachment of a variety of molecules such as alkyl halides, alcohols or amines bound as carboxylic acid esters and amides. According to the safety-catchprinciple, the separation of the desired products proceeds in a twostep process. First, penicillin G acylase hydrolyzes the phenylacetamide with complete chemo- and regioselectivity, under exceptionally mild conditions (pH 7.0, room temperature or 37 "C) [30, 72*741 . Then the activated intermediate generated,

I

1405

1406

I 18

introduction and Removal of Protecting Groups

1) Boc-trypothan DIC, DMAP

2) CF3COOH, r.t. 68 232

\OH 233

1

R-CHO, molecular sieves, CHZC12, 50°C OAc

vow '7

234

I

lipase RB 001-05, 50 mM MES buffer/ CH30H (60/40), pH 5.8, 30°C

R

R = Ph, 4-N02C6H4,i-Pr 55-85 %

p6::--] 0

0

H N

237

R

70-80 %

Figure 18-35. Solid phase synthesis of tetrahydro-6-carbolins and subsequent detachment by enzyme initiated fragmentation of the anchor group.

i. e. benzylamine 239, cyclizes to polymer-bound lactam 240 and releases the desired target molecule 241. POE 6000 was used as the polymeric support, a soluble polyethyleneglycol derivative functionalized at both termini with an amino group and with an average molecular mass of 6000 Da [323-3241. After completion of the homogeneous reactions

18.G Biocatalysis in Polymer Supported Synthesis: Enzyme-labile Linker Groups

X=O,NH,NR

-

pmH 1 Penicillin Gacylase

239

Figure 18-36. Principle of the enzyme-labile safety catch linker.

it can be precipitated, filtered off, and washed with diethyl ether, thereby facilitating the separation of surplus reagents and the side products. Furthermore it allows for NMR spectroscopic monitoring of the reactions [3251. Most importantly, it is soluble in aqueous solutions, thereby allowing efficient access of the enzyme to the polymerfixed linker group. The suitability of the polymer-linker conjugate was examined for a variety of transformations, in particular Pdo-catalyzedreactions. For instance, the polymerbound aryl iodide 242 was transformed quantitatively in a Heck reaction to a cinnamic acid ester 243 and to biphenyl 245 in a Suzuki reaction. It gave an alkine 244 in a Sonogashirareaction (Fig. 18-37).The desired benzyl alcohols 246248 were released by incubation of the corresponding polymer conjugates 243-245 with penicillin G acylase at pH 7 and 37 "C in high yields and isolated with a purity of >95 % by simple extraction with diethyl ether. Furthermore, the applicability in a Mitsunobu esterification reaction and a DielsAlder reaction was proven (Fig. 18-38).The polymer-bound benzyl alcohol 249 was

I

1407

1408

I

78 Introduction and Removal ofProtecting Groups

242

OMe

I

245

Penicillin G acylase pH 7.0, 10 % MeOH,

/ /

Penicillin G acylase pH 7.0, 10 % MeOH,

/ \

OtBU 247

246

Figure 18-37.

Pdo-catalyzed reactions on enzyme labile linker-conjugates.

reacted with 4-acetamidophenol in the presence of the Mitsunobu reagent to give phenyl ether 250 in quantitative yield. It was released from the polymeric support in high yield. For the Diels-Alder reaction, polymer-bound acrylic acid ester 252 was treated with cyclopentadiene. The cycloaddition product 253 was formed with an endolexo ratio of 2.5 : 1 and with quantitative conversion. The subsequent enzymatic release delivered the corresponding alcohol (251,254) in high yield and purity.

18.7 Outlook

During recent decades substantial progress was achieved in the development of enzymatic protecting group techniques. In particular, it was demonstrated that these methods offer viable alternatives to classical chemical approaches. Not only do the biocatalyzed transformations complement the arsenal of chemically removable protecting groups, but in many cases they additionally offer the opportunity to carry out useful functional group interconversions with selectivities which can not or only barely be matched by chemical techniques. However, the overwhelming majority of the investigations carried out in this area has restricted themselves to the study of the protection and deprotection of model compounds. Complex synthetic schemes were nearly always avoided. Whereas this appears to be particularly true in the area of carbohydrates, noticeable examples which demonstrate the capacity of these biocatalyzed processes were recorded in peptide and peptide conjugate chemistry, i. e. in

References I1409

249

0

H

nNr

DEAD, PPh3

, THF, 60°C

HO

H

I

250

+

Nr

Figure 18-38.

o

a

251 o

t

H

I H

penicillin G acylase, pH 7.0,37"C, 48h (75 %)

penicillin G acylase, pH 7.0,37"C, 48h (81 %)

D

HOo0 254

Mitsunobu and Diels-Alder reaction on enzyme labile linker-conjugates.

the synthesis of lipo-, glyco and nucleopeptides. The data and observations highlighted above, however, provide a solid basis for the application of biocatalysts in the handling of protecting group problems in complex multistep syntheses. On the other hand, the use of biocatalystsin protecting group chemistry in the sense of a general method deserves and is certainly awaiting further intensive development. Numerous applications of the known enzymes appear to be possible in all areas of preparative chemistry. In addition, the use of catalpc proteins which have not yet been applied to carry out protecting group manipulations and of biocatalysts unknown today or which will be developed in the future, e.g. by evolutionary approaches, will create new opportunities for improved organic syntheses. References 1

2

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3

P. J. Kocienski, Protecting Groups, Georg Thieme Verlag, Stuttgart,

1994. H. Waldmann, Kontakte (Darmstadt) 1991, 2, 33-54. 5 A. Reidel, H. Waldmann, J . Prakt. Chem. 1993,335,109. 4

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18 introduction and Removal of Protecting Groups

L. Panza, S. Brasca, S. Riva, G. Russo, Tetra- . hedron: Asymmetry 1993,4,931-932. 177 L. Panza, M. Luisetti, E. Crociati, S. Riva, j . Carbohydr. Chem. 1993,12,125-130. 178 M. J. Chinn, G. Iacazio, D. G. Spackman, N.J. Turner, S. M. Roberts,]. Chem. Soc., Perkin Trans. 1 1992, 661-662. 179 M. J. Chinn, G. Iacazio, D. G. Spackman, N.J. Tumer, S. M. Roberts, J. Chem. Soc., Perkin Trans. 1 1992, 2045. 180 J. J. Gridley, A. J. Hacking, H. M. I. Osborn, D. G. Spackman, Tetrahedron 1998,54, 14925-14946. 181 J. J. Gridley, A. J. Hacking, H. M. I. Osborn, D. G. Spackman, Synlett 1997, 1397-1399. 182 D. Colombo, F. Ronchetti, A. Scala, L. Toma, J. Carbohydr. Chem. 1992,11,89-94. 183 P. Ciuffreda, F. Ronchetti, L. Toma,]. Carbohydr. Chem. 1990,9,125-129. 184 P. Ciuffreda, D. Colombo, F. Ronchetti, L. Toma,]. Org. Chem. 1990,55,4187-4190. 185 R. Lopez, E. Montero, F. Sanchez, J. Canada, A. Femandez-Mayoralas,/. Org. Chem. 1994,59,7027-7032. 186 C. Chon, A. Heisler, N.Junot, F. Levayer, C. Rabiller, Tetrahedron: Asymmetry 1993, 4, 244-2444. 187 N.Boissiere-Junot, C. Tellier, C. Rabiller,J. Carbohydr. Chem. 1998,17,99-115. 188 M. Woudenberg-van Oosterom, C. Vitry, J. M. A. Baas, F. van Rantwijk, R. A. Sheldon,]. Carbohydr. Chem. 1995, 14, 237-246. 189 N.Junot, J. C. M e s h , C. Rabiller, Tetrahedron: Asymmetry 1995,6,1387-1392. 190 E. W. Holla, J. Carbohydr. Chem. 1990,9, 113-119. 191 F. Nicotra, S. Riva, F. Secundo, L. Zucchelli, Tetrahedron Lett. 1989, 30, 1703-1704. 192 G. Carrea, S. Riva, F. Secundo, B. Danieli,J. Chem. Soc., Perkin Trans. 1 1989, 1057-1061. 193 J.-F. Shaw, A. M. Klibanov, Biotechnol. Bioeng. 1987,29,648-651. 194 A. L. Fink, G. W. Hay, Can. J . Biochem. 1969, 47,353-359. 195 T. Horrobin, C. H. Tran, D. Crout, ]. Chem. Soc., Perkin Trans. 11998, 1069-1080. 196 H. M. Sweers, C. H. Wong, J. Am. Chem. SOC.1986, 108,6421-6422. 197 M. Kloosterman, E. W. J. Mosmuller, H. E. Schoemaker, E. M. Meijer, Tetrahedron Lett. 1987,28,2989-2992. 198 Y. Kodera, K. Sakurai, Y. Satoh, T. Uemura, 176

Y. Kaneda, H. Nishimura, M. Hiroto, A. Matsushima, Y. Inada, Biotechnol Lett. 1998, 20,177-180. 199 K.-F. Hsiao, F.-L. Yang, S.-H. Wu, K.-T. Wang, Biotechnol. Lett. 1995, 17, 963-968. 200 T. Matsui, Y.Kita, Y. Matsushita, M. Nakayama, Chem. Express 1992,7,45-48. 201 A. Bastida, R. Fernindez-Lafuente, G. Fernindez-Lorente, J. M. Guisin, G. Pagani, M. Terreni, Bioorg. Med. Chem. Lett. 1999, 9. 202 0. Kirk, M. W. Christensen, F. Beck, T. Damhus, Biocatal. Biotransform. 1995, 12, 91-97. 203 L. Gardossi, R. Khan, P. A. Konowicz, L. Gropen, B. S. Paulsen, J. Mol. Cat. B: En~ y m1999,6,89-94. . 204 D. Chaplin, D. H. G. Crout, S. Bornemann, D. W. Hutchinson, R. Khan, J. Chem. Soc., Perkin Trans. 11992, 235-237. 205 K. F. Hsiao, S. H. Wu, K. T. Wang, Bioorg. Med. Chem. Lett. 1993, 3, 2125-2128. 206 C. Vogel, S. Kramer, A. J. Ott, Liebigs Ann./ Red. 1997, 1425-1428. 207 R. Lopez, C. Perez, A. Fernandez-Mayoralas, S. Conde, 1.Carbohydr. Chem. 1993, 12, 165-171. 208 R. Csuk, B. I. Glanzer, 2. Naturforsch., B: Chem. Sci. 1988,43,1355-1357. 209 J. Zemek, S. Kucar, D. Anderle, Collect. Czech. Chem. Commun. 1987,52, 2347-2352. 210 M. Kloosterman, M. P. De Nijs, J. G. J. Weijnen, H. E. Schoemaker, E. M. Meijer,J. Carbohydr. Chem. 1989,8, 333-341. 211 E. W. Holla, V. Sinnwell, W. Klaffke, Synlett 1992,413-414. 212 A. Ballesteros, M. Bernabe, C. Cruzado, M. Martin-Lomas, C. Otero, Tetrahedron 1989, 45,7077-7082. 213 H. Waldmann, A. Heuser, Bioorg. Med. Chem. 1994,2,477-482. 214 S. Tomic, J. Tomasic, L. Sesartic, B. Ladesic, Carbohydr. Res. 1987, 161,150-155. 215 S. Tomic, D. Ljevacovic, J. Tomasic, Carbohydr. Res. 1989, 188,222-227. 216 S. Tomic, A. Trescec, D. Ljevakovic, J. Tomasic, Carbohydr. Res. 1991, 210, 191-198. 217 D. Ljevacovic, S. Tomic, J. Tomasic, Carbohydr. Res. 1992,230,107-115. 218 H. Waldmann, A. Heuser, P. Braun, H. Kunz, Ind. I . Chem. 1992, 31B, 799. 219 J. 0. Rich, B. A. Bedell, J. S. Dordick, Biotechn. Bioeng. 1995,45,426-434. 220 M. Woudenberg-van Oosterom, F. van

References I1415 Rantwijk, R. A. Sheldon, Biotechnol. Bioeng. 1996,49, 3'28-333. 221 S. Cai, S. Hakomori, T. Toyokuni,J. Org. Chem. 1992,57, 3431-3437. 222 B. Danieli P. D. Bellis, G. Carrea, S. Riva, Helv. Chim.Acta 1990,73, 1837-1844. 223 M. A. Cruces, C. Otero, M. Bernabe, M. Martin-Lomas, A. Ballesteros, Ann. N. Y. h a d . Sci. 1992,672,436-443. 224 D. B. Sarney, M. J. Bamard, D. A. MacManus, E. N. Vulson, J. Am. Oil Chem. Soc. 1996,73, 1.481-1487. 225 M. Kloostmnan, J. G. J. Weijnen, N. K. De Vries, J. Mentech, I. Caron, G. Descotes, H. E. Schciemaker, E. M. Meijer, J. Carbohydr. Chenz. 1989,8,693-704. 226 K.-Y. Chang, S.-H. Wu, K. T. Wang, Carbohydr. Res. '1391,222,121-129. 227 K.-Y. Chang, S.-H. Wu, K. T. Wang, J . Carbohydr. Chenz. 1991, 10, 251-261. 228 G.-T. Ong. S.-H. Wu, K. T.Wang, Bioorg. Med. Cherw. Lett. 1992,2, 161-164. 229 G. T. Ong: S. H. Wu, K. T. Wang, Bioorg. Med. Chern. Lett. 1992, 2, 631. 230 D. C. Palmer, F. Terradas, Tetrahedron Lett. 1994,35, 1673-1676. 231 S . Bornemann, J. M. Cassells, J. S. Dordick, A. J. Hacking, Biocatalysis 1992,7, 1-12. 232 G.-T. Ong, K.-Y. Chang, S.-H. Wu, K.-T. Wang, Carbohydr. Res. 1994,265,311-318. 233 M.Ferrem, V. Gotor, Monatsh. Chem. 2000, 131,585-616. 234 H. S. Sadidev, N. A. Starkovsky, Tetrahedron Lett. 1969,9,733-736. 235 A. Taunton-Rigby, J . Org. Chem. 1973,38, 977-985. 236 C.-H. Wong, S.-T. Chen, W.-J. Hennen, J.A. Bibbs, Y.-F. Wang, J. L.-C. Liu, M. W. Pantoliano, kl. Whitlow, P. N. Bryan, J . Am. Chem. Sol:. 1990, 112,945-953. 237 F. Moris, V. Gotor, J . Org. Chem. 1992, 57, 2490-2492. 238 L. F. Garcia-Alles, F. Moris, V. Gotor, Tetrahedron Lett. 1993,34,6337-6338. 239 L. F. Garcia-Alles, V. Gotor, Tetrahedron 1995,51,307-316. 240 V. Gotor, F. Moris, Synthesis 1992, 626-628. 241 F. Moris, V. Gotor,J. Org. Chem. 1993,58, 6 53-660. 242 F. Moris, V. Gotor, Tetrahedron 1994,50, 6927-6934. 243 R. V. Nail, M. M. Salunkhe, Synth. Commun. 2000,30, 3115-3120. 244 S . Ozaki, K. Yamashita, T. Konishi, T. Mae-

kawa, M. Eshima, A. Uemura, L. Ling, Nucleosides Nucleotides 1995, 14, 401-404. 245 F. Moris, V. Gotor, Tetrahedron 1992, 48, 9869-9876. 246 L. F. Garcia-Alles, J. Magdalena, V. Gotor, 1.Org. Chem. 1996,61,6980-6986. 247 J. Magdalena, S. Fernandez, M. Ferrero, V. Gotor, Tetrahedron Lett. 1999, 40, 1787-1790. 248 L. F. Garcia-Alles, V. Gotor,]. Mol. Catal. B: Enzym. 1999,6,407-410. 249 F. Moris, V. Gotor, Tetrahedron 1993,49, 10089-10098. 250 V. Gotor, F. Moris, L. F. Garcia-Alles, Biocatalysis 1994, 10, 295-305. 251 M. Mahmoudian, J. Eaddy, M. Dawson, Biotechnol. Appl. Biochern. 1999,29,229-233. 252 A. Uemura, K. Nozaki, J.-I. Yamashita, M. Yasumoto, Tetrahedron Lett. 1989, 30, 3817-3818. 253 A. Uemura, K. Nozaki, J.-I. Yamashita, M. Yasumoto, Tetrahedron Lett. 1989, 30, 3819-3820. 254 H. K. Singh, G. L. Cote, R. S. Sikorski, Tetrahedron Lett. 1993, 34, 5201-5204. 255 D. H. G. Crout, A. M. Dachs, S. E. Glover, D. W. Hutchinson, Biocatalysis 1990, 4, 177-183. 256 D. L. Damkjaer, M. Petersen, J. Wengel, Nucleosides Nucleotides 1994, 13, 1801-1807. 257 L. E. Iglesias, M. A. Zinni, M. Gallo, A.M. Iribarren, Biotechnol. Lett. 2000, 22, 361-365. 258 B. Danieli, P.D. Bellis, G. Carrea, S. Riva, Gazz. Chim. Ital. 1991, 121,123-125. 259 B. Danieli, P. D. Bellis, G. Carrea, S. Riva, Heterocycles 1989, 29, 2061-2064. 260 N. Nakajima, K. Ishihara, T. Itoh, T. Furuya, H. Hamada, J. Biosci. Bioeng. 1999, 87, 105-107. 261 B. Danieli, S. Riva, Pure @Appl. Chem. 1994, G6,2215-2218. 262 B. Danieli, M. Luisetti, S . Riva, A. Bertinotti, E. Ragg, L. Scaglioni, E. Bombardelli, J. Org. Chem. 1995,60,3637-3642. 263 H. Lehmann, 0. Miersch, H. R. Schuttle, Z. Chem. 1975, 15,443. 264 G. Schneider, 0. Miersch, H.-W. Liebisch, Tetrahedron Lett. 1977, 405-406. 265 D. L. Delinck, A. L. Margolin, Tetrahedron Lett. 1990, 31, 3093-3096. 266 A. L. Margolin, D. L. Delinck, M. R. WhaIon, J. Am. Chem. SOC.1990, 112, 2849-2854.

1416

I

78 Introduction and Removal ofprotecting Croups

L. Gardossi, D. Bianchi, A. M. Klibanov,/. Am. Chem. SOG.1991, 113,6328-6329. 268 S. Riva, A. M. Klibanov, J. Am. Chem. SOC. 1988, 110,3291-3295. 269 A. Bertinotti, G. Carrea, G. Ottolina, S. Riva, Tetrahedron 1994, 50, 13 165-13172. 270 E. Santaniello, P. Ferraboschi, S. RezaElahi, Monatsh. Chem. 2000, 131,617-622. 271 S. Riva, R. Bovara, G. Ottolina, F. Secundo, G. Carrea,]. Org. Chem. 1989,54, 3161-3164. 272 V. C. 0. Njar, E. Caspi, Tetrahedron Lett. 1987,28,6549-6552. 273 A. Baldessari, A. C. Bruttornesso, E. C. Gros, Helv. Chim. Acta 1996,79,999-1004. 274 A. Baldessari, M. S. Maier, E. G. Gros, Tetrahedron Lett. 199536,4349-4352. 275 G. Nicolosi, M. Piattelli, C. Sanfilippo, Tetrahedron 1992,48,2477-2482. 276 D. Lambusta, G. Nicolosi, M. Piattelli, C. Sanfilippo, IndianJ. Chem., Sect. B 1993, 32B, 58-60. 277 G. Nicolosi, M. Piattelli, C. Sanfilippo, Tetrahedron 1993,49, 3143-3148. 278 D. Lambusta, G. Nicolosi, A. Patti, M. Piatelli, Synthesis 1993, 1155-1158. 279 K.-F. Hsiao, F.-L. Yang, S.-H. Wu, K.-T. Wang, Biotechnol Lett. 1996, 18, 1277-1282. 280 P. Allevi, P. Ciuffreda, A. Longo, M. Anastasia, Tetrahedron: Asymmetry 1998, 9, 2915-2924. 281 V. S. Parmar, A. K. Prasad, N. K. Sharma, K. S. Bisht, R. Sinha, P. Taneja, Pure Appl. Chem. 1992,64,1135-1139. 282 V. S. Parmar, A. K. Prasad, N. K. Sharma, S. K. Singh, H. N. Pati, S. Gupta, Tetrahedron 1992,31,6495-6498. 283 K. S. Bisht, 0. D. Tyagi, A. K. Prasad, N. K. Sharma, S. Gupta, V. S. Parmar, Bioorg. Med. Chem. 1994,2,1015-1020. 284 M. Natoli, G. Nicolisi, M. Piattelli, Tetrahedron Lett. 1990,31,7371-7374. 285 M. Natoli, G. Nicolisi, M. Piattelli,/. Org. Chem. 1992,57,5776-5778. 286 V. S. Parmar, H. N. Pati, A. Azim, R. Kumar, Himanshu, K. S. Bisht, A. K. Prasad, W. Errington, Bioorg. Med. Chem. 1998, 6, 109-118. 287 A. K. Prasad, H. N. Pati, A. Azim, S. Trikha, Poonam, Bioorg. Med. Chem. 1999,7, 1973-1977. 288 P. Ciuffreda, S. Casati, E. Santaniello, Tetrahedron 2000,56,317-321. 289 F. Balkenhohl, C. von dem Bussche Hunne267

feld, A. Lansky, C. Zechel, Angew. Chem. Int. Ed. En@. 1996,35, 2289-2337. 290 J. S. Fruchtel, G. Jung, Angw. Chem., Int. Ed. En@. 1996, 35, 17-42. 291 L. A. Thompson, J. A. Ellman, Chem. Rev. 1996.96, 555-600. 292 L. Yan, C. M. Taylor, R. Goodnow, D. Kahne, J. Am. Chem. SOC.1994, 116,6953-6954. 293 R. L. Halhomb, H. M. Huang, C. H. Wong, J . Am. Chem. SOC.1994, 116,11315-11322. 294 This nomenclature was introduced by S. L. Flitsch et al. See references 312 and N. J. Turner, Cum. Org. Chem. 1997, 1, 21-36. 295 M. Schuster, P. Wang, J. C. Paulson, C. H. Wong,J. Am. Chem. SOC.1994, 116, 1135-1136. 296 K. Yarnada, S. I. Nishimura, Tetrahedron Lett. 1995, 36, 9493-9496. 297 K. Yamada, E. Fujita, S. I. Nishimura, Carbohydr. Res. 1997,305,443-461. 298 S. Nishimura, K. Yamada,J . Am. Chem. SOC. 1997,119,10555-10556. 299 J. Weiler, H. Gausepohl, N. Hauser, 0. N. Jensen, J. D. Hoheisel, Nucleic Acids Res. 1997,25,2792-2799. 300 D. T. Elmore, D. J. S. Guthrie, A. D. Wallace, S. R. E. Bates,J. Chem. SOC., Chem. Commun. 1992,1033-1034. 301 SPOCC resin is based on the cross-linking of long-chain poly(ethy1eneglycol) (PEG) terminally substituted with oxetane by cationic ring-opening polymerization. 302 J. Rademann, M. Grotli, M. Meldal, K. Bock,J. Am. Chem. SOG.1999, 121, 5459-5466. 303 Bio-beads consists of 1 % cross-linked polystyrene with 1.25 mmol chloromethyl substitution per g of dry resin, respectively benzhydrylamine polymer 1% cross-linked polystyrene with 0.24 mmol NH2 per g of dry resin, Bio-Rad Laboratories (Richmond, CA, USA). 304 H. Gaertner, A. Puigserver, Eur. J. Biochem. 1984,145,257-263. 305 Novasyn KA constits of kieselguhr supported dimethylacrylamide functionalized with sarcosine methylester. 306 B. D. Larsen, A. Holm, /. Pept. Res. 1998.52, 470-476. 307 J. Vagner, G. Barany, K. S. Lam, V. Krchnak, N. F. Sepetov, J. A. Ostrem, P. Strop, M. Lebl, Proc. Nat. Acad. Sci. U.S. A. 1996,93, 81944199.

References

G. Corrales, A. Fernandez-Mayisralas,E. Garcia-Junceda,Y. Rodriguez, Biocataf. Biotransform. 2000, 18, 271-281. 309 MPEG consists of a poly(ethy1eneglycol) 5000 monomethyl ester, see: J. J. Krepinsky, Advances in polymer-supported solution synthesis of oligosaccharides, In Modern Methods i n Carbohydrate Synthtsis (Eds. S. A. Khan and R. A. O'Neill), Harwod Academic Publishers, The Netherlands, 1996, 194-224. 310 C. Schmitz, M. T. Reetz, Org. 1.ett. 1999, I , 1729-173 1. 311 Flitsch, S. L., Lahja, S., andTurner, N. J. Solid phase preparation and enzymic and non-enzymic bond cleavage of sugars and glycopeptides, PCT Int. Appl. 1997, patent EP 9605535, CAN 127:81736. 312 G. Bohm, J. Dowden, D. C. Rice, I. Burgess, J. F. Pilard, B. Guilbert, A. Hauton, R. C. Hunter, N. J. Turner, S. L. Flitsch, Tetrahedron Lett. 1998,39, 3819-3822. 313 B. Sauerbrei, V. Jungmann, H. Waldmann, Angew. Chem., Int. Ed. Engf. 1!)98,37, 1143-1 146. 314 H. Waldmann, B. Sauerbrei, 1J. Grether, 308

Enzyme cleavable linker for solid phase synthesis, (BASF A.-G., Germany), Ger. Offen. 1998, CAN 128:114573. 315 D. Kadereit, H. Waldmann, Chem. Rev., 2001, 101,3367-3396. 316 W. Rapp in: Combinatorid Peptide and NonPeptide Libraries (Ed.: G. rung),VCH, Weinheim, 1996, p. 425. 317 U. Grether, H. Waldmann, Chem. Eur.J. 2001,7,959-971. 318 U . Grether, H. Waldmann, Angew. Chem., Int. Ed. Engl. 2000, 39, 1629-1632. 319 B. F. Cain,J. Org. Chem. 1976,41, 2029-203 1. 320 I. D. Entwistle, Tetrahedron Lett. 1994, 35, 4103-4106. 321 F. Cubain, Rev. Roum. Chim. 1973, 18, 449-461. 322 G. Just, G. Rosebery, Synth.Commun.1973, 3,447-451. 323 D. J. Gravert, K. D. Janda, Chem. Rev. 1997, 97,489-509. 324 E. Bayer, M. Mutter, Nature 1972,237, 512-5 13. 325 J. M. Dust, 2.H. Fang, J. M. Harris, Macromolecules 1990, 23, 3742-3746.

I

1417

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyriqht 0Wiley-VCH Verlaq GmbH, Weinheim 2002

I

19 Replacing Chemical Steps by Biotransformations: Industrial Application and Processes Using Biocatalysis Andreas Liese ‘Bacteria are capable of briiaging about chemical reactions of amazing variety and sublety in an extremely short time ... Many bacteria are ofvery great importance to industry where they peform tasks which would take much time and trouble by ordinary chemical methods.’ Sir Cyril Hinshelwood, 1956L11

19.1

Introduction

Starting with big promises and many expectations in the seventies biocatalytic processes have left the status of a lab curiosity together with many prejudices far behind and are now established on an industrial scale[2].Product examples range from amino acids, sugars, chiral alcohols and amines, and highly functionalized building blocks for pharmaceuticals to bulk chemicals such as acrylamide or propane-1,3-diol. When speaking about biotechnological processes one has to distinguish between fermentation processes and biotransformations. In a fermentation process the desired product is synthesized from nutrients and trace elements by either microorganisms (bacteria, yeasts, fungi) or higher cells such as mammalian or plant cells. The phrase “biotransforrnation”or “biocatalysis”is commonly used to describe a one-step or multi-step transformation of a precursor to the desired product using whole cells and/or (partly)purified enzymes. Whole cell processes are often used for redox reactions using the metabolism of the living cell for cofactor regeneration. In some cases the cell is used as a compartment containing the enzymes in a confinement allowing easier separation of the entire biocatalyst using centrifugation or microfiltration. If one has to deal with membrane-bound enzymes, whole cell biocatalysts are to be preferred. Numerous authors have given overviews over biotransformations used in inA very recent monograph summarizes almost 100 processes including du~try[~-”]. many details on reaction conditions, screening of the biocatalyst or the product application[2].The use of biocatalysis from the viewpoint of a chemist in the laboratory is also summarized in several books. Recent ones are[12-14].

1419

1420

I

7 9 Replacing Chemical Steps by Biotransformations

In this contribution we shall focus on those examples where the biocatalytic step has distinct advantages over the corresponding chemical method, or even has replaced or is about to replace other methods. The reasons may be better regio-, stereo- or chemoselectivity,better product purity or simplified downstream processing. Often the incorporation of biocatalytic steps reduces the amount or toxicity of waste.

19.2

Types and Handling of Biocatalysts

This chapter tries to give a brief introduction to the types of biocatalysts, their requirements and methods of handling them. A more detailed treatment can be found in other chapters of this book or in the literature [15-171. The biocatalyst may be a whole cell or a partly purified enzyme. In the first case the cell may be regarded as a mini-reactor with all necessary cofactors and enzymes to catalyze multiple steps concentrated in one cell. In the second case the main catalytically active species is isolated and purified. For the whole cell systems either prokaryotic cells such as Escherichia coli or eukaryotic cells such as Saccharomyces cereuisiae or Zymomonas mobilis are used. Prokaryotic cells do not posses a nucleus. The nuclear material is contained in the cytoplasm of the cell. Therefore introduction and processing of foreign DNA to obtain a genetically engineered strain is simple. They are relatively small in size (0.2-10pm) and exist mostly as single cells. Eukaryotic cells are higher microorganisms and have a true nucleus separated by a nuclear membrane. They are larger in size (5-30 pm) and sometimes form more complex structures. For both types the bioreactor has to fulfill certain requirements. An adequate supply of nutrients as well as oxygen into the bioreactor has to be assured. Parameters such as pH, oxygen, feed rate and temperature in the bioreactor must be kept within certain limits in order to guarantee optimum growth and/or metabolic activity of the cells. Especially when recombinant microorganisms are employed genetic stability during cultivation has to be observed carefully. Substrates, products and/or solvents required may be toxic for the cells and may therefore have to be added in low amounts to secure a low stationary concentration. The example of a ketone reduction with whole cells of Zygosaccharomyces rouxii shows one possible solution. The toxic substrate is adsorbed on XAD-7 resin (80 g/L resin, resulting in a concentration of 40 g/L reaction volume), and the resin is added to the fermentation broth. The equilibrium concentration in the aqueous phase is approximately 2 g/L. The product is adsorbed on the resin as well, thus providing integrated downstream processingw191.

When purified enzymes are used, basically the same requirements have to be met. The purification may cause additional costs, but contrary to a biochemical characterization it is not necessary to purify the protein to homogeneity. On the contrary, the remaining protein content in a partly purified enzyme may increase its stability. The only requirement is to have a functional pure enzyme, meaning that activities

79.3 Examples I1421

catalyzing undesired side reactions have to be absent. This is the major advantage of purified enzymes over whole cell processes: side reactions may be easily avoided, and substrates that are toxic for the cell or which may not be able to enter the cell can be converted. For enzymes the thermal deactivation or deactivation by interphases (liquid-liquid,liquid-gas) may be limiting. For industrial biotransformations, catalyst recovery and reuse are major issues. This may be desirable either for reasons of downstream processing or for repeated use in order to reduce the specific catalyst costs per kg of product produced. A very simple method is the use o f membrane filtration. Because of the increasing number of membranes from difft2rent materials (polymers, metal or ceramics) this is an attractive alternative. Whereas for whole cells microfiltration or centrifugation can be applied, for the recovery ‘of soluble enzymes ultrafiltration membranes have to be Often immobilization on a support is chosen to increase the catalyst’s stability as well as to facilitate its recovery. The main advantages of immobilization are: easy separation, often increased stability, use of fixed or fluidized bed reactors (for continuous processes). Disadvantages are: loss of absolute activity due to the immobilization process, mass transport limitations, There is no general or best method of immobilization; the protocol has to be developed individually for each catalyst. The most common methods for the immobilization are entrapment in matrices such as alginate beads, cross-linking, and covalent or adsorptive binding to a carrier. A very recent method is the development of cross-linked enzyme crystals (CLECS) (see also Chapter 6 ) [231. A survey of different immobilization methods can be found in in[24. 251

19.3

Examples

The examples presented here are taken from 12]. Only those biotransformations were chosen where a classical chemical step was replaced. The enzymes involved are mainly from the group:; of oxidoreductases (E. C. class 1) and hydrolases (E. C. class 3). There are a few examples of lyases (E.C. class 4) and one example of an isomerase (E. C. class 5). The processes involving oxidoreductases mainly use whole cells because of the problem of cofactor regeneration. The examples are sorted in the order of the main classes of the Enzyme Commission (E. C.). The big letter E denotes the biotransformation in the syntheses schemes.

1422

I

19 Replacing Chemical Steps by Biotransformations

19.3.1

Reduction Reactions Catalyzed by Oxidoreductases (E. C. 1) 19.3.1.1

Ketone Reduction Using Whole Cells of Neurospora crassa (E.C. 1.1.1.1) [26-291

The key step in the synthesis of Trusopt@,which is a topically active treatment for glaucoma, is the enantioselective reduction of S,G-dihydro-G-methyl-4H-thieno[2,3b] thiopyran-4-one-7,7-dioxide (Fig. 19-1). The biological route overcomes the problem of incomplete inversion of the cisalcohol in the chemical synthesis (Fig. 19-2).The reaction is carried out below pH S to prevent epimerization of the (6s)-methylketosulfone in aqueous media. The (R)-3-hydroxy-butyrate(Fig. 19-l),which is responsible for the stereochemistry of the methyl group in the sulfone ring, can be produced by depolymerization of biopolymers, e. g. Biopol from Zeneca. This is a natural polyester produced by some microorganisms as a storage compound. 1) pyridinel

twyl chloride

CH30ZC

COOCH,

3) H202,NaWOT

LiS

1

2

1 = biologically derived homopolymer 2 = (Rp3-hydroxy-rnethylbutyrate

3 = 5,6-dihydro-6-methyl-4H-thieno(2,3b]thiopyran-4-one~7,7-dioxide 4 = 5,6-dihydro-4-hydroxy-6-methyl-4H-thieno[2,3b]thiopyran-7,7-dioxide 5 = trusopt, MK-0507 E = alcohol dehydrogenase,whole cells from Neurosporia crassa

Figure 19-1.

Synthesis ofTrusopt@ (5) via enzymatic ketone reduction (Astra-Zeneca)

79.3 Examples I1423

noMe OTS 0

pyridinel tosyl chlorie

conc. HCI

cH30Zca,)

~

uOMe

NHCOCH

NHCOCH

m s o z c i

oAo Figure 19-2.

~

~

~

s

,.”..,

S

VHCHZCH, o

z

BH~DMS, N TH

z

H

o4 “o

Chemical synthesis of Trusopt@.

19.3.1.2

Ketoester Reduction Using; Cell Extract of Acinetobacter calcoaceticus (E.C. 1.1.1.1)[30-321

The biotransformation (Fig. 19-3)is an alternative to the chemical synthesis via the chlorohydrine and selective hydrolysis of the acyloxy group (Fig. 19-4). After final fractional distillation this synthesis has an overall yield of 41 %. The biotransformation has a yield of 92%. The diketoester can be obtained as shown in Fig. 19-5. G-Benzyloxy-(3R,5S)-dihydroxy-hexanoic acid ethyl ester is a key chiral intermediate for anticho1estei:ol drugs that act by inhibition of hydroxy methyl glutaryl coenzyme A (HMG CoAl reductase.

1

(3R,5s)-2

1 = 6-benzyloxy-3,5-dioxo-hexanoic;mid ethyl ester 2 = 6-benzyloxy-3,5-dihydroxy-hexar1oic acid ethyl ester E = alcohol dehydrogenasefrom Ac,inetobactercalcoaceticus

Figure 19-3.

Synthesis of key intermediate of anticholesterol drugs (BristoCMyers Squibb)

~

1424

I

7 9 Replacing Chemical Steps by Biotransformations OH

0 CH&OO'BU

~

LiHMDS/ - 78 "C

I

cluok

1. Et,EOMe/NaBH, THFiMeOH/78 "C

2. H,OflaOH

/ ("Eu),NOCOMe ___)

CSA

Figure 19-4.

NMP/ 85 "C/ 9 h

Chemical synthesis of key intermediate of anticholesterol drugs.

pyridine

a o J c ,

- HCI

+HN(OMe), 94 %

I

o\

62 %

Synthesis of starting material 6-benzyloxy-3,S-dihydroxyhexanoic acid ethyl ester. Figure 19-5.

19.3.1.3 Enantioselective Reduction with Whole Cells of Candida sorbophila (E.C. 1.l.X.X) 133-351

Here the biotransformation (Fig. 19-6) is preferred over the chemical reduction with commercially available asymmetric catalysts (BH3- or noble-metal-based), since with the chemocatalysts the desired high enantiomeric excess (ee > 98%, 99.8% after purification) is not achievable. Since the ketone has only a very low solubility in the aqueous phase, I kg ketone is added as solution in 4 L 0.9 M H 2 S 0 4to the bioreactor. The bioreduction is essentially carried out in a two-phase system, consisting of the aqueous phase and small droplets made up of substrate and product. The downstream processing consists of multiple extraction steps with methyl ethyl ketone and precipitation induced by pH titration of the pyridine functional group (p& = 4.66) with NaOH. The (R)-aminoalcohol is an important intermediate for the synthesis of P-3-agonists that can be used for obesity therapy and to decrease the level of associated type 11diabetes, coronary artery disease and hypertension.

7 9.3 Examples

1 NHzOH .HCI

1 EtOK. EtOH

2 TsCI, pyndine

2 HCI, MTBE

EDC. THF. H20.pH 5.5, pnilrophenyl acetic acid

-1 BHo-SMen. THF. 1 M H,SO,

ElOH HCI

2 purifyvia phosphate salt, then free base

92 %

75 %

1 = 2-(4-nitro-phenyl)-N(2-oxo-2-pyridin-3-ethyl)-acetamide

2 = (R)-N-(2-hydroxy-2-~ridin-3-yl-ethyl)-2-(4-nitro-phenyl)-acetamide 3 = p-3-agonist E = dehydrogenase, whole cells 01 Candida sorbophila

Figure 19-6.

Synthesis of key intermediate of P-3-agonist (Merck Research Laboratories).

19.3.2

Oxidation Reactions Catalyzed by Oxidoreductases (E.C. 1) 19.3.2.1

Alcohol Oxidation Using Whole Cells of Cluconobacter suboxydans (E. C. 1.1.99.21) [36-381

In 1923 the bacterium Acinetobacter suboxydans was isolated and, starting in 1930, was used for the industrial oxidation of L-sorbitol to L-sorbose in the ReichsteinGriissner synthesis of vitamin C 13'1. Bayer uses the same type of reaction, but instead of Acinetobacter the bacterium Gluconobacter suboxydans is used in the oxidation of Nprotected 6-amino-~-sorbi to1 to the corresponding 6-amino-~-sorbose, which is an intermediate in miglitol production (Fig. 19-7).1-Desoxynojirimycinis produced by chemical intramolecular reductive amination of 6-amino-~-sorbose. In contrast, the

I

1425

19 Replacing Chemical Steps by Biotransformations

T$ :izi

HOfi

~

HO$i

protection

HOfH

OH OH

OH

OH

D-glucose

OH

1

1-amino-D-sorbitol

R' HO.

1) deprotection 2) NaBH4

HO& HO

HO{\

OH 1-desoxynojirimycin

2

90% yield

rniglitol 1 = 1-amino-D-sorbitol (N-protected) 2 = 6-arnino-~-sorbose(N-protected) E = D-sorbitol dehydrogenase, whole cells of Gluconobacter oxidans

Figure 19-7.

Synthesis of key intermediate for miglitol (Bayer).

published chemical synthesis of 1-desoxynojirimycin and its derivatives requires multiple steps and laborious protecting-group chemistry. Miglitol and derivatives thereof are pharmaceuticals for the treatment of carbohydrate metabolism disorders (e.g. diabetes mellitus). 19.3.2.2

Oxidative Deamination Catalyzed by Immobilized D-Amino Acid Oxidase from Trigonopsis uariabilis (E. C. 1.4.3.3) [40-421

This oxidative deamination catalyzed by immobilized enzymes is part of the 7-aminocephalosporanic acid (7-ACA) process. Ketoadipinyl-7-aminocephalosporanic acid decarboxylates in situ in the presence of H202,which is formed by the

7 9.3 Examples

I

1427

3 1 = cephalosporin C 2 = a-ketoadipinyl-7-aminocephalosponnicacid 3 = glularyl-7-aminocephalosporanicacid (7-ACA) E = 0-aminoacidoxidase, immobilizedenzyme from Trigonopsis variablis

Synthesis of glutaryl-7-arninocephalosporanic acid (7-ACA) (Hoechst-Marion-Roussel).

Figure 19-8.

biotransformation step yielding glutaryl-7-ACA. The reaction solution is directly transferred to the 7-ACA production (see Sect. 19.3.4.2 for details and a comparison with the chemical synthesis). 19.3.2.3

Kinetic Resolution by Oxidation of Primary Alcohols Catalyzed by Whole Cells from Rhodococcus erythropolis (E. C. 1.X.X.X) [43-45]

(R)-Isopropylideneglycerol is a useful C3-synthon in the synthesis of (S)-P-blockers, e. g. (S)-metoprolol.Also, (R)-isopropylideneglycericacid may be used as the starting material for the synthesis of biologically active products. The resolution is carried out by selective microbial oxidation of the (S)-enantiomer (Fig. 19-9). The chemical synthesis of (R)-isopropylideneglycerolstarts either from unnatural L-mannitol or from L-ascorbic acid (Fig. 19-10). In comparison to the biotransformation, here stoichiometric quantities of lead tetra acetate are needed.

( WS)-1 1 = isopropylideneglycerol

2 = isopropylideneglycericacid

E = oxidase, whole cells from Rhodococcuserythropolis

Figure 19-9. Synthesis of (R)-isopropylideneglyceroland (R)-isopropylideneglyceric acid (International BioSynthetics).

1428

I

7 9 Replacing Chemical Steps by Biotransformations

From L-mannitol:

OH HO

Pb(0Ack

2

-

0

HO

2

0

CHO

HO

(W-1 From L-ascorbic acid:

Figure 19-10.

C h e m i c a l synthesis of

(R)-isopropylideneglycerol.

19.3.2.4 Hydroxylation o f Nicotinic Acid (Niacin) Catalyzed by Whole Cells ofAchrornobacter

xylosoxidans (E.C. 1.5.1.1 3) 146-481

6-Hydroxynicotinate is a versatile building block used chiefly in the synthesis of modern insecticides. The 6-hydroxynicotinate-producingstrain (Fig. 19-11) was found by accident, when in the mother liquor of a niacin-producing chemical plant precipitated white crystals of 6-hydroxynicotinatewere found. The second enzyme of the nicotinic acid pathway, the decarboxylating 6-hydroxynicotinate hydroxylase becomes strongly inhibited at niacin concentrations higher than 1%, whereas the operation of niacin hydroxylase is unaffected. In contrast to the biotransformation, the chemical synthesis of 6-substituted nicotinic acids is difficult and expensive because of the necessity for the separation of by-products that are produced by nonregioselective hydroxylations. (Jcoo-

** +H20

1

2[H] '12

02

1

HvO

1 = niacin = nicotinic acid = pyridine-3-carboxylate 2 = 6-hydroxynicotinate= 6-hydroxy-pyridine-3-carboxylate E = nicotinic acid hydrolase, whole cells from Achromobacterxylosoxidans

2

> 90% yield

Figure 19-11. Synthesis of 6-hydroxynicotinate

(Lonza).

19.3.2.5

Reduction of Hydrogen Peroxide Concentration by Catalase (E. C. 1.1 1.1.6)

[491

During oxidative coupling to dinitrodibenzyl (DNDB),hydrogen peroxide is formed as a by-product. It is not possible to decompose H202by adding heavy-metal catalysts

19.3 Examples

0 2

+2

pJMe @ (t-BuOK)

NO2

/

NO2

1

2

+

H202

I

+'

1 = nitrotoluene 2 = dinitrodibenzyl (DNDB)

E = catalase, enzyme from microbial source

Figure 19-12.

Degradation o f hydrogen peroxide (Novartis). catalase

Figure 19-13.

I

Flow scheme ofdinitrodibenzyl synthesis.

because only an incomplete conversion is reached. Additionally, subsequent process steps with DNDB are problematic because of contamination with heavy-metal catalyst. The biotransforination is the only relevant method of decomposing the undesired side product Hz02 (Fig. 19-12).The enzyme of choice is catalase derived from a microbial source, which has advantages compared to beef catalase since the activity remains constant over a broad pH range from 6.0 to 9.0, temperatures up to 50 "C are tolerated, and salt concentrations up to 25% do not affect the enzyme stability. The reaction is carried out in a cascade of three continuously operated stirred tank reactors (Fig. 19-13).The H 2 0 2concentration is reduced from 7000 ppm to < 200 ppm in the product solution. The third vessel is aerated with nitrogen to degas the product solu-tion. The dinitrodibenzyl is used as a pharmaceutical intermediate.

1429

1430

I

7 9 Replacing Chemical Steps by Biotransformations 19.3.3

Hydrolytic Cleavage and Formation o f C - 0 Bonds by Hydrolases (E.C. 3) 19.3.3.1 Kinetic Resolution of Clycidic Acid Methyl Ester by Lipase from Serratia rnarcescens (E.C. 3.1.1.3)[50-53]

Trans-(2R,3S)-(4-methoxyphenyl)glycidic acid methyl ester is an intermediate in the synthesis of diltiazem, a coronary vasodilator and a calcium channel blocker with

6

enzymatic process

Meon a::2 aspo

CICH,COOMe

w

NaOMe

CHO

chemical process

PCOOMe

COOMe

I

D,L-threo I

1‘ I

’

0

COOMe

hydrolysis O M /.e--A -

COOH

(2S,3R)-2 99 9% ee

40-45% yield

NaHSO,

-co2

3

S0,Na

COOH

1

(+)-ihreo reduction

4

NMe, .HCI

I

1 = tramp-methoxyphenylmethylglycidale(MPGM) 2 = trans-p-melhoxyphenylglycidic acid 3 = bisuifite adduct after decarboxylalion 4 = diltiazem E = lipase, enzyme from Serraiia marescens

Comparison o f chemical and biocatalytical route t o diltiazem (Tanabe Seiyaku Co., Ltd.).

Figure 19-14.

19.3 Examples

I

1431

antianginal and antihypersensitive activity. It is produced worldwide in excess of > 100 t a-l. In comparison to the chemical route, only 5 steps (instead of 9) are necessary with the biotransformation (Fig. 19-14).The kinetic resolution is carried out in an earlier step with a lower molecular weight compound during the synthesis, resulting in a reduction of waste. By redesigning the synthesis route using a biotransformation, the manufacturing costs of diltiazem were decreased to two thirds of those of the original process including a chemical resolution rs41. The lipase from Serratia rnarcescens has a high enantioselectivity (E = 135) for the (ZR,3S)-(4-rnethoxyphenyl)glycidic acid methyl ester, which acts as a competitive inhibitor. The formed acid (hydrolyzed (+)-methoxyphenylglycidate)is unstable and decarboxylatesto give 4-methoxyphenylacetaldehyde;this aldehyde strongly inhibits and deactivates the enzyme. It is removed by transfer to the aqueous phase by formation of a water-soluble adduct with sodium hydrogen sulfite added to the aqueous phase. The bisulfite acts also as a buffer to maintain constant pH during synthesis. The enantioselective hydrolysis is carried out in an organic-aqueous two-phase reactor (toluene/water), where the phase contact is established by a hydrophilic hollow-fiber membrane (polyacrylonitrile).The lipase is immobilized onto a spongy layer by pressurized adsorption. The productivity is about 40 kg trans-(ZR,3S)(4-methoxypheny1)glycidic: acid methyl ester m-* a-l. This process has been operated since 1993. 19.3.3.2

Kinetic Resolution of Diestier by Protease Subtilisin Carlsberg from Bacillus sp. (E. C. 3.4.21.62)*'si 561

(R)-(2-Methylpropyl)-butanedioic acid 4-ethyl ester is used as a chiral building block for potential collagenase inhibitors (e.g. Ro 31-9790) in the treatment of osteoarthritis. The diester is reacted as a 20% emulsion in 30 mM aqueous NaHC03 using Protease@L 660 or [email protected] L (9% each, with respect to the racemic diester) (Fig. 19-15).The unconverted (S)-diestercan be extracted in a solvent such as toluene and racemized by heating the anhydrous extract with catalytic amounts of sodium ethanolate. The resulting racemic diester can be recycled, thus improving the overall organic phase

aqueousphase

1 = (2methylpropyl)butanedioicacid diethylether 2 = (2methylpropyl)butanedioicacid 4-ethyl ester, Na-form E = hydrolase, subtiiisin Carlsberg from Bacillus sp.

Figure 19-15.

Synthesis o f (F')-(2methylpropyl) butanedioic acid diethylether (Hoffmann La-Roche).

1432

I yield from 45

19 Replacing Chemical Steps by Biotransforrnations

% to 87 %. The reaction was repeatedly carried out on a 200 kg scale with respect to the racemic diester. The enzyme is highly stereoselective even at high substrate concentrations (20%). The chemoenzymatic route starting from the cheap bulk agents maleic anhydride and isobutylene replaced the existing chemical research synthesis for bulk amounts (Fig. 19-16). drug discovery synthesis

=A H~N-COOH

process research synthesis

i;""" COO'BU

t

d

EtOO

COOEt

JE

EtOOO dH

'BuOO

A -?-

r i.

Ro 31-9790

Figure 19-16.

Comparison of drug discovery and process research route of Ro 31-9790.

7 9.3 Examples

I

1433

D,L-1

D-2

t

+

crystallization

D-1

L$ 0

racemization

L- 1 1 = pantolactone 2 = pantoic acid E = lactonase, whole cells from Fusarium oxysporum

Figure 19-17.

Synthesis o f D-pantolactone (Fuji Chemical Industries).

19.3.3.3 Kinetic Resolution of Pantolactonesand Derivatives thereof by a Lactonase from Fusarium oxysporum (E. C. 3.1.1.25) ["I

Pantenoic acid is used as a vitamine BZ complex. D- and L-pantolactoneare used as chiral intermediates in chemical synthesis. The enantioselectivehydrolysis is carried out in the aqueous phase with a substrate concentration of 2.69 M = 350 g L-' (Fig. 19-17).For the synthesis whole cells are immobilized in calcium alginate beads and used in a fured bed reactor. The immobilized cells retain more than 90 % of their initial activity after 180 days of continuous use. At the end of the reaction Lpantolactone is extracted and reracemized to D,L-pantolactone, which is recycled to the reactor. The D-pantenoic acid is chemically lactonized to D-pantolactone and extracted. By applying cells from Brewibacteriurn protophormia the L-lactone is available. The biotransformation eliminates several steps that are necessary in the chemical resolution process (Fig. 19-18). 19.3.3.4 Hydrolysis of Starch to Glucose by Action ofTwo Enzymes: a-Amylase (E.C. 3.2.1.1) and Arnyloglucosidase (E. C. 3.2.1.3) [5s-601

The process is part of the production of high fructose corn syrup. After several improvements, this process (Fig. 19-19)provides an effective way for an important, low-cost sugar substitute derived from grain. At various stages enzymes are applied in this process[", '*I. The corn kernels are softened to separate oil, fiber and proteins by centrifugation. The enzymatic steps are cascaded to yield the source product for the invertase process after liquefaction in continuous cookers, debranching and filtration (Fig. 19-20). Since starches from different natural sources have different compositions, the procedure is not unique. The process ends, if all starch is completely broken down to limit the amount of oligomers of glucose and dextrins. Additionally, recombination of molecules has to be prevented. The thermostable

1434

I

I9 Replacing Chemical Steps by Biotransformations

chemical process

-

enzymatic process

-tgo

enzymatic hydrolysis

1 --tre_: KO resolution

-

0

+

I

0

%

HO

I

extraction

lactonization

-tgo

-tgo

-

-

Figure 19-18. Comparison o f chemical and biocatalytic route for the enantioselective synthesis o f pantolactone. CHzOH

CH,OH

El

&o&o&o&o,

+HzO

*

oligomer units

\O OH

OH

OH

OH

1 CH2OH

1 = starch 2 = glucose E l = a -amylase, enzymefrom Bacillus licheniformis E2 = glucoamylase, enzyme from Aspergillus niger

Figure 19-19.

Synthesis of glucose (Several companies)

OH &OH

OH

2

19.3 Examples 11435

‘ 0

o + *o , OH

so*

OH

OH

OH

+ centrifuge 1. liquefaction 105-115°C pH-6

2. liquefaction 90-95°C

protein, fat

4

\ \

OH

Figure 19-20.

\

J

Flow scheme for the hydrolysis of starch to glucose.

enzyme can be used up to 115 “C. The enzymes need Ca2+ions for stabilization and activation. Since several substances in corn can complex cations, the cation concentration is increased requiring a further product purification, i. e. making it necessary to refine the product. There is no alternative industrial chemical process for starch liquefaction. The worldwide production is about 10’ t a-’, 19.3.4 Formation or Hydrolytic Cleavage of C-N Bonds by Hydrolases (E.C. 3) 19.3.4.1 Enantioselective Acylation of Racemic Amines Catalyzed by Lipase from Burkholderia plantarii (E.C. 3.1.1.3)[63-651

The lipase catalyzes the kinetic resolution of racemic amines, e.g. l-phenylethylamine (Fig. 19-21)[ll]. Products are intermediates for pharmaceuticals and pesticides. They can also be used as chiral synthons in asymmetric synthesis. As acylating agent ethylmethoxyacetate is used, because the reaction rate is more than 100 times faster than that with butyl acetate. Probably an enhanced carbonyl activity induced by the electronegative a-substituents accounts for the activating effect of the methoxy group. The lipase is immobilized on polyacrylate. The lowered activity caused by use of in organic solvent (tert-methylbutylether= MTBE) can be increased

1436

I

I9 Replacing Chemical Steps by Biotransformations

Lo,

NH

NHZ

2

(q-3 > 93% ee

1 = I-phenylethylamine 2 = ethylmethoxyacetate 3 = phenylethylmethoxyamide E = Iipase, enzyme from Burkholdenaplantanr

Figure 19-21.

Kinetic resolution o f phenylethylarnine (BASF).

(about 1000 times and more) by freeze drying a solution of the lipase together with fatty acids (e.g. oleic acid). Because of the use of MTBE a high starting material concentration of 1.G5 M 1-phenylethylaminecan be established. The enantioselectivity is greater than 500. The (R)-phenylethylmethoxyamidecan easily be hydrolyzed to the (R)-phenylethylamine.The unconverted (S)-enantiomer can be racemized using a palladium catalyst. 19.3.4.2

7-AminocephalosporanicAcid Formation by Amide Hydrolysis Catalyzed by Glutaryl Amidase (E.C. 3.1.1.41)[66-691

The second step of the 7-aminocephalosporanicacid (7-ACA)process is the deamidation of glutaryl-7-ACA (Fig. 19-22), the first step is described in Sect. 19.3.2.2. 7-ACA is an intermediate for semi-synthetic cephalosporins. Hoechst Marion Roussel uses the glutaryl amidase immobilized on a spherical carrier. Toyo Jozo and Asahi Chemical immobilize the glutaryl amidase on porous styrene anion exchange resin with subsequent cross-linking with 1% glutardialdehyde. The catalyst is applied in a fixed bed reactor in a repetitive batch mode (70 cycles). Here, an enzymatic process has replaced an existing chemical process for environmental reasons (Fig. 19-23): In the first step, the zinc salt of cephalosporin C is produced, followed by the protection of the functional groups (NH2 and COOH) with trimethylchlorosilane.

1 = glutaryl-7-aminocephalosporanicacid 2 = 7-aminocephalosporanicacid (7-ACA) E = glutaryl amidase, enzyme from Escherichia coli

Figure 19-22. Synthesis of 7-arninocephalosporanic acid (7-ACA) (Asahi Chemical, Hoechst Marion Roussel, Toyo Jozo).

79.3 Examples I1437

chemical process

enzymatic process

0

COOH

J

ZnCPC

++ 0H2

L

- H202 - NH,

solventJTMSCl /

+

HN-SI\

-si/

/

N /o * 0

phoy

CI 0

COOH

\

T c 0 "C

0

COOH

0

hydrolysis ~

H

o

o

c

~

c

o

o

H

H2Np&oy 0

COOH

0

E l = D-aminnacid oxidase

E2 = glutaiyl amidase

Figure 19-23.

Comparison o f chemical and biocatalytical route for the synthesis o f 7-ACA.

The imide chloride is synthesized in the subsequent step at 0 "C with phosphorous pentachloride. Hydrolysis of this imide chloride yields 7-ACA. By replacement of this synthesis with the biotransformation, the use of heavy-metal salts (ZnClz) and chlorinated hydrocarbons as well as precautions for highly flammable compounds can be circumvented. The off-gas quantities were reduced from 7.5 to 1.0 kg. Mother liquors requiring incineration were reduced from 29 to 0.3 t. Residual zinc that was recovered as Zn(NH4)P04is reduced from 1.8 to 0 t. The absolute costs of environmental protection are reduced by 90% per tonne of 7-ACA. Asahi Chemical and Toyo Jozo have produced 7-ACA since 1973 with a capacity of 90 t a-l and Hoechst Marion Roussel since 1996 with a capacity of 200 t a-'.

1438

I

79 Replacing Chemical Steps by Biotransformations

1

2

3

1 = penicillin-G 2 = 6-amino penicillanic acid ( 6-APA) 3 = phenylaceticacid E = penicillin amidase, enzyme from Escherichia coli

Figure 19-24.

Synthesis of 6-amino penicillanic acid (multiple companies).

19.3.4.3

Penicillin C Hydrolysis by Penicillin Amidase from Escherichia coli (E.C. 3.5.1.11) [68-711

6-Amino penicillanic acid (6-APA)is used as the intermediate for manufacturing semi-synthetic penicillins. Companies applying this technology (Fig. 19-24) include Unifar, Turkey; Asahi Chemicals, Japan; Fujisawa Pharmaceutical Co., Japan; GistBrocades/DSM, The Netherlands; Novo-Nordisk, Sweden; Pfizer, USA. The enzyme is isolated and immobilized, often on Eupergit@C(Rohm, Germany). The production is carried out in a repetitive batch mode. The immobilized enzyme is retained by sieves. In case of the Eupergit@Cimmobilized amidase the residual activity is about 50 % of initial activity after 800 batch cycles. Therefore the hydrolysis time after 800 batch cycles increases from initially 60 min to 120 min. The space-time yield is 445 g L-‘ d-*. Phenylacetic acid is removed by extraction and 6-APA can be crystallized. Concentrating the “split” solution and/or the mother liquor of crystallization via vacuum evaporation or reverse osmosis can increase the yield. The production plant operates for 300 days per year with an average production of 12.8 batch cycles per day (production campaigns of 800 cycles per campaign). Asahi Chemical utilizes a penicillin amidase from Bacillus megaterium that is immobilized on aminated porous polyacrylonitrile fibers. The production is carried out in a recirculation reactor consisting of 18 parallel columns with immobilized enzyme. Each column has a volume of 30 L. The circulation of the reaction solution is established with a flow rate of 6 000 L h-’. One cycle time takes 3 h. The lifetime of each column is 360 cycles. Purification of 6-APA is done by isoelectric precipitation at pH 4.2 with subsequent filtration and washing with methanol. 7-Amino deacetoxy cephalosporanic acid (7-ADCA)is also produced by the same technology. Several chemical steps are replaced by a single enzyme reaction (Fig. 19-25). Organic solvents, the use of low temperature (- 40 “C) and the need for absolutely anhydrous conditions, which used to make the process difficult and expensive, are no longer necessary in the enzymatic process.

19.3 €xurnp/es H

1

pc15

-400c

n-butanol

“‘“P% 0

COOH

Figure 19-25.

Chemical process for 6-APA.

19.3.4.4

Kinetic Resolution of a-Amino Acid Amides Catalyzed by Aminopeptidase from Pseudomonas putida (E. C. 3.4.1.1 1) [72-751

Enantiomerically pure a-H-amino acids are intermediates in the synthesis of antibiotics used for parenteral nutrition and for food and feed additives (see also Chapter 12.2). Examples are D-phenylglycine and 4-hydroxyphenylalanine for semisynthetic j3-lactam antibiotics and L-phenylalaninefor the peptidic sweetener aspartame. DSM used this process to produce also L-homophenylalanine, a potential precursor molecule for several ACE-inhibitors. The a-amino amides as substrates for this enantiospecific, biocatalytic amide hydrolysis can be readily obtained from the appropriate aldehydes via the Strecker synthesis (Fig. 19-26). As whole cell catalyst, Pseudornonas putida, which accepts a wide range of substrates, is applied. Subsequent to the biotransformation, benzaldehyde is added, resulting in precipitation of the o-amide Schiff base, which can be easily isolated by filtration. An acidification step leads to the D-amino acid. The L-amino acid can be reused after racemization so that a theoretical yield of 100% D-amino acid is possible. The same process can be used for the synthesis of 100% of L-amino acids by racemizing the Schiff base of the D-amide in a short time using small amounts of base in organic solvents. Using i n vivo protein engineering not only mutant strains of Pseudornonas putida

I

1439

1440

I

7 9 Replacing Chemical Steps by Biotransformations 0

,AH 1

H,N

CN

--zFT + NH,

2

1"-

acetone

1) NH3 2) PhCHO 3) OH- (pH 13) racemization 4) H,O'

D,L-3

i.

2) H,O'

L-4

t = aldehyde 2 = amino nitrile 3 = a-amino acid amide 4 = a-amino acid methyl ester 5 = a-amino acid 6 = a-amino acid amide 7 = schiff base of a-amino acid amide E = aminopeptidase, whole cells from Pseudomonas puhda

Figure 19-26. Production o f L-and D-a-amino acids by kinetic resolution o f a-amino acids amides (DSM).

exhibiting L-amidase and also D-amidase but also amino acid amide racemase activities were obtained. Using these mutants a convenient synthesis of a-H-amino acids with 100% yield would be possible with one cell system. It is noteworthy that only a-H-substrates can be used. By screening, a new biocatalyst of the strain Mycobacterium neoaurum was found, which is capable of converting a-substituted amino acid amides.

1

2

t

racemization

3

4

> 99.5% ee 80% yield

1 = N-acetyl-D.L-methionine 2 = N-acetyl-D-methionine 3 = L-methionine 4 = acetic acid

E = aminoacylase, enzyme from AspergiNus niger

Figure 19-27.

Biocatalytical production of L-rnethionine by kinetic resolution (Degussa)

19.3.4.5 Production of L-Methionine by Kinetic Resolution with Aminoacylase o f A s p e r g i h

oryzae (E.C. 3.5.1.14) [76-791

The N-acetyl-D,L-aminoacid precursors are conveniently accessible through either acetylation of D,L-amino acids with acetyl chloride or acetic anhydride in a SchottenBaumann reaction or via amidocarbonylation[80].For the acylase reaction, Co2+as metal effector is added to yield an increased operational stability of the enzyme. The unconverted acetyl-D-methionineis racemized by acetic anhydride in alkali, and the racemic acetyl-D,L-methionineis reused. The racemization can also be carried out in a molten bath or by an acetyl amino acid racemase. Product recovery of L-methionine is achieved by crystallization, because L-methionine is much less soluble than the acetyl substrate. The production is carried out in a continuously operated stirred tank reactor. A polyamide ultrafiltration membrane with a cutoff of 10 kDa retains the enzyme, thus decoupling the residence times of catalyst and reactants. L-methionine is produced with an ee > 99.5% and a yield of 80% with a capacity of > 300 t a-'. At Degussa, several proteinogenic and non-proteinogenic amino acids are produced in the same way e. g. L-alanine, L-phenylalanine, a-amino butyric acid, L-valine, Lnorvaline and L-homophenylalanine. 19.3.4.6 Production o f D-p-Hydroxyphenyl Clycine by Dynamic Resolution with Hydantoinase from Bacillus breuis (E. C. 3.5.2.2) L8, 81-831

D-p-Hydroxyphenyl glycine is a key raw material for the semisynthetic penicillins such as ampicillin and amoxycillin. It is also used in photographic developers. Racemic hydantoins are synthesized starting from phenol derivatives, glyoxylic acid and urea via the Mannich condensation (Fig. 19-28).The D-specific hydantoinase is applied as immobilized whole cells in a batch reactor. The unreacted L-hydantoins are readily racemized under the alkaline conditions (pH 8) of enzymatic hydrolysis, yielding quantitative conversion. This process enables the stereospecific preparation of various amino acids, such as L-tryptophane, r-phenylalanine, D-valine, D-alanine

1442

I

I 9 Replacing Chemical Steps by Biotransformations

+ A H,N

NH,

J

E

H d

L-1

H d

D-1

1 = 5-(p-hydroxybenzyI)-hydantoin 2 = D-N-carbamoylamino acid 3 = D-4-hydroxyphenylglycine

HO'

HO

D-2

D-3

E = D-hydantoinase,whole cells from Bacillus brews

Figure 19-28.

Synthesis o f o-amino acids (Kanegafuchi).

and D-methionine. Instead of chemical treatment with sodium nitrite, a carbamoylase (EC 3.5.1.77) can also be applied to remove the carbamoyl group. Several other companies have developed patented processes to produce D-hydroxyphenylglycine (Ajinomoto,DSM, SNAM-Progetti,Recordati and others). Here the biotransformation competes with the classical chemical route (Fig. 1929), which employs bromocamphorsulfonic acid (Br-CAS)as the resolving agent. In both routes phenol is used as raw material since p-hydroxybenzaldehyde is too expensive. The hydantoinase process for phenylglycines does not necessarily need an extra racemization step since the hydantoin is racemized in situ at an alkaline pH. Because of the dynamic resolution in the case of this biotransformation, higher yields are reached. 19.3.4.7 Dynamic Resolution of a-Amino-E-caprolactam by the Action of Lactamase (E. C. 3.5.2.11) and Racemase (E.C. 5.1.1.15)[84* "1

Again a dynamic resolution is carried out, but this time the racemization is introduced by an enzyme, a racemase from Achromobacter obae (Fig. 19-30). The lactamase and racemase are applied as whole cells and are fortunately active at the same pH, so that they can be used in one reactor. Reaction conditions enabling

I

79.3 Examp/es 1443

resolution with

Br-CAS 7

J

$” \ /

HO

HNO, c-

E l = D-hydantoinase.whole cells from San//us brevis E2E3 = D-hydantoinase/N-carbamoyl-D-amino acid hydrolase, whole cells, strain Pseudomonassp. contains both enzymes

Figure 19-29. Comparison o f chemical and biocatalytical route for the synthesis o f o-amino acids (Kanegafuchi).

D,L-1

D-1

E2

L-2 > 99.5% ee

1 = a-amino-E-caprolactam(ACLI 2 = lysine E l = L-aminolactarn-hydrolase,whole cells from Cryptococcus laurentii E2= amino-lactam-racemase,whole cells from Achromobacter obae Figure 19-30.

Synthesis of L-lysine (Toray Industries).

chemical racemization would reduce the enzyme stability. L-Lysine was produced

with an ee of 99.5 % at a capacity of 4000 t a-’. This process has been totally replaced by highly effective fermentation methods.

1444

I

7 9 Replacing Chemical Steps by Biotransformations

R’

fJqf

+

H21F>

N /

R3

E -R,H ,

R’Q-Yp& 0

COOH

D(-)-1

2

OOH

3

l a = phenylglycineamide (R’=H, R2=NH,) = PGA 1b = phenylglycinernethyleter(R’=H, RZ=OMe)= PGM 1c = hydroxyphenylglycineamide(R’=OH, R2=NH,) = HPGA I d = hydroxyphenylglycinemethylester(R’=OH, RZ=OMe)= HPGM 2a = 7-amincdeacetoxycephalosporanicacid (&Me) = 7-ADCA 2b = 7-arn1ncdeaceioxymethyl-3-chlorocephalospnic acid (R3=CI)= 7-ACCA 3a = cefaclor (R’=H, R3=CI) 3b = cephalexin (R’=H, &Me) 3c = cefadroxil (R’=OH. &Me) E = penicillin acylase

Figure 19-31.

Synthesis o f p-lactam antibiotics (Chemferm).

19.3.4.8 Synthesis of B-Lactam Antibiotics Catalyzed by Penicillin Acylase (E. C. 3.5.1.1 1) ~86-8gl

The penicillin acylases do not accept charged amino groups. Therefore phenylglycine itself cannot be used at a pH value at which the carboxyl function is uncharged, because the amino group will then be charged. To reach non-equilibrium concentrations of the product, the substrate must be activated as an ester or amide (Fig. 19-31).By this means the amino group can be partly uncharged at the optimal pH value of the enzyme. In biological systems, ATP delivers the activation energy. Using the same synthetic pathway alternatively to 7-ADCA and 7-ACCA, 6-APA derivatives can also be synthesized. The established chemical synthesis started from benzaldehyde and included the fermentation of penicillin (Fig. 19-32).The process consists often steps with a waste stream of 30-40 kg waste per kg product. The waste contains methylene chloride, other solvents, silylating agents and many products from side-chain protection and acylating promoters. In comparison, the chemoenzymatic route needs only six steps including three biocatalytic ones. The biotransformations E l and E2 in Fig. 19-32 can be found in Sect. 19.3.4.3and 19.3.4.4. 19.3.4.9

Synthesis of hetidinone 8-Lactam Derivatives Catalyzed by Penicillin Acylase (E.C. 3.5.1.11)[’Ot ”1

It was thought that the Pen G amidase would exhibit only a limited substrate spectrum, since it does not hydrolyze the phenoxyacetyl side chain of penicillin V. Nevertheless, Eli Lilly shows that the Pen G amidase acylates the amino function of cis-3-amino-azetidinonewith the methyl ester of phenoxyacetic acid (Fig. 19-33).The

19.3 Examples

chemical process

j

benzaldehyde

Penicillium

enzymatic process benzaldehyde

Penicillium

J

~trecker synthesis

J

fermentation

D,L-phenylglycine

penicillin G

1

1

fermentation

Strecker synthesis

penicillin G

1

ring enlargement

classical resolution

D-( -)-phenylglycine

cephalosporin

protection

ring enlargement

D,L-phenylglycineamide ester

cephalosporin

1

El

deacylation

Dane salt

7-ADCA

activation

mixed anhydride

7-ADCA

I

I

protection

Protected7-ADCA

D-(-)-phenylglycineamidd ester

coupling deprotecting

-T coupling

E l = penicillin amidase E2 = amino peptidase E3 = penicillin acylase C o m p a r i s o n of t h e chemical a n d biocatalytical synthesis

1 = cis-3-amino-azetidinone 2 = phenoxy-aceticacid methyl ester 3 = B-lactam intermediate 4 = loracarbef E = Pen G amidase, enzyme from Escherichiacolr

Figure 19-33.

Synthesis of azetidione fi-lactam derivatives

protection

protected 7-ADCA

cefalexin

cefalexin

Figure 19-32.

deacylation

(Eli Lilly).

of cefalexin.

I

1445

1446

I

79 Replacing Chemical Steps by Biotransformations

acylation occurs using methyl phenylacetate (MPA) or methyl phenoxyacetate (MPOA)as the acylating agents. The penicillin amidase is immobilized on Eupergit (Roehm GmbH, Germany). The chemical resolution of the racemic azetidinone is only low yielding. The (2R,3S)-azetidinoneis a key intermediate in the synthesis of the carbacephalosporin antibiotic loracarbef. 19.3.4.10

EnantioselectiveSynthesis of an Aspartame Precursor with Thermolysin from Bacillus proteolicus (E.C. 3.4.24.27) 931

Since the reaction (Fig. 19-34) is limited by the equilibrium the products have to be removed from the reaction mixture to reach high yields. Therefore an excess of racemic phenylalanine methylester (which is inert to the reaction) is added. The carboxylic anion of the protected aspartame forms a poorly soluble adduct with DPhe-OCH3 that precipitates from the reaction mixture. The precipitate can be removed easily by filtration. Final steps of the process are the separation of D-Pheester, removal of protecting groups and racemization of the formed L-amino acid. aAspartame is produced with > 99.9% and a worldwide capacity of - 10,000 t a-', - 2,500 t 6' by enzymatic coupling. The bacterial strain was found in the Rokko Hot Spring in central Japan. Consequently it is very stable up to temperatures of GO "C. The main problem in chemical synthesis coupling of Z-Asp anhydride with COOCH

,

HOOCYCooH HNXZ

L-2

D,L-l

I

I) HCI

1 1 ii) 1 ) H,,

COOCH, \

RH,W

+

Pd/C, MeOH

H O O C q A

aspartame ~~~

1 = phenylalanine methylester 2 = aspartic acid (protected) 3 = a-aspartame (protected) E = thermolysin. enzyme from Bacillus proieolicus

COOCH,

NHZ

Figure 19-34. Biocatalytical synthesis of aspartame (HSC, Holland Sweetener Company).

79.3 Examples 11447

2

1

3 > 95% yield

1 = 2-cyanopyrazine 2 = pyrazine-2-carboxylicacid

3 = 5-hydroxypyrazine-2carboxylicacid EVE2 = nitrilaselhydroxylase,whole cells, strain Agrobacteriumsp. contains both enzymes

Figure 19-35.

Biocatalytical synthesis of 5-hydroxypyrazine-2-carboxylic acid (Lonza).

L-Phe-OCH3 is the by-product formation of P-aspartame. This isomer is of bitter taste and has to be completely removed from the a-isomer. The advantages of the enzymatic route are: (i) No p-isomer is produced, (ii) the enzyme is completely stereoselective, so that racemic mixtures of the substrate or the appropiate enantiomer of the amino acid can be used, (iii)no racemization occurs during synthesis and (iv) the reaction takes place in aqueous media under mild conditions. 19.3.4.11 Hydrolysis of Heterocyclic Nitrile by Nitrilase from Agrobacteriurn sp. (E.C. 3.5.5.1) [94-961

5-Hydroxypyrazine-2-carboxylicacid (Fig. 19-35)is a versatile building block in the synthesis of new antituberculous agents, e. g. 5-chloro-pyrazine-2-carboxylic acid esters. The regioselectivehydroxylation of pyrazine-2-carboxylicacid is catalyzed by a hydroxylase (E2,E. C. 1.5.1.13). This second enzyme is also in the applied suspended whole cells from Agrobacteriurnsp. The biomass is separated by ultrafiltration (cutoff 10 kDa) after the biotransformation. 5-Hydroxypyrazine-2-carboxylic acid is precipitated from the permeate by acidification with sulfuric acid to pH 2.5. In contrast to the biotransformation, the chemical synthesis of 5-substituted pyrazine-2-carboxylicacid leads to a mixture of 5- and 6-substituted pyrazinecarboxylic acids and requires multiple steps. 19.3.5

Formation o f C - 0 Bonds by Lyases 19.3.5.1

Synthesis of Carnitine Catalyzed by Carnitine Dehydratase in Whole Cells (E.C. 4.2.1.89) [', 46s 97-991

L-Carnitine is used in infant health, sport and geriatric nutrition. The biotransformation is catalyzed by carnitine dehydratase in whole cells (Fig. 19-36).(R)-carnitineis produced with > 99.5% conversion of butyrobetaine and > 99.5% ee. The mutant strain has blocked the L-carnitine dehydrogenase and excretes the accumulated product. The purified enzyme could not be used for the biotransformation because of its high instability. Apart from usual batch fermentations, continuous production

1448

I

7 9 Replacing Chemical Steps by Biotransformations Figure 19-36. Comparison of chemical and biocatalytical synthesis o f carnitine (Lonza).

,% CI&COOEt

1

H, Ru-(S)-BINAP

Q”

tO E C, - ,A, - ,~lCOOC , , - , , . + N 3 e M (R) 97% ee

Me3N\ H30t

1

E J

99.9% ee > 99.5% yield

Q”

C O O ~, - , - , , ,N +M ,e

( R)-2 1 = 4-butyro betaine 2 = carnitine E = carnitine dehydratase, whole cells from Escherichia coli

is also feasible since the cells go into a “maintenance state” with high metabolic activity and low growth rate. The cells can be recycled after separation from the fermentation broth by filtration. A chemical resolution process with L-tartaric acid that was developed at Lonza was no longer competitive with the biotechnological route. A more attractive chemical route would be the Ru-BINAP catalyzed asymmetric hydrogenation of 4-chloroacetoacetate (Fig. 19-36). Here an ee of 97% is yielded. 19.3.6

Formation ofC-N Bonds by Lyases (E.C. 4) 19.3.6.1 Synthesis of L-Dopa Catalyzed by Tyrosine Phenol Lyase from Erwinia herbicola (E.C. 4.1.99.2)

The product is applied for the treatment of Parkinsonism that is caused by a lack of Ldopamine and its receptors in the brain. L-Dopamine is synthesized in organisms by decarboxylation of ~-3,4-dihydroxyphenylalanine (L-dopa).Since L-dopamine cannot pass the blood-brain barrier L-dopa is applied in combination with dopadecarboxylase-inhibitors to avoid formation of L-dopamine outside the brain. Ajinomoto produces L-dopa by this lyase-biotransformation with suspended whole cells in a fed batch reactor on a scale of 250 t ax’. Much earlier, Monsanto has successfully scaled up the chemical synthesis of L-dopa (Fig. 19-38).

1

2

L-3

1 = catechol 2 = pyruvic acid 3 =dopa E = tyrosine phenol lyase. whole cells from Erwinia herbicola

Figure 19-37. Biocatalytical synthesis o f L-dopa (Ajinomoto).

HoucHo +

HO

A~HN-COOH

Ac

1

\

2

1 =vanillin 2 = azlactone 3 = Z-enamide 3 = dopa

Figure 19-38. Chemical synthesis of L-dopa (Monsanto)

The enantioselectivehydrogenation of 3,4-dihydroxy-N-acetylamino cinnamic acid is catalyzed by the cationic Rh-biphosphine complex DIPAMP, in which the enantioselectivity is introduced by the chiral phosphine 1104*1051 . The hydrogenation proceeds quantitatively with 94 % ee. The optically pure L-dopa is separated from the catalyst by crystallization. 19.3.6.2

Synthesis o f 5-Cyano Valeramide by Nitrile Hydratase from Pseudomonas chlororaphis 823 (E.C. 4.2.1.84)['06. 1"'

5-Cyanovaleramideis used as intermediate for the synthesis of the DuPont herbicide azafenidine (Fig. 19-39). The whole cells from Pseudornonas chlororuphis are immobilized in calcium alginate beads. The biotransformation itself is catalyzed by a nitrile hydratase that converts a nitrile into the corresponding amide by addition of water. Nitrile hydratases belonging to the enzyme class of lyases (E. C. 4) are not be

1450

I

I9 Replacing Chemical Steps by Biotransformations

1 = adiponitrile 2 = 5-cyano-valerarnide E = nitrile hydratase, whole cells from Pseudomonas chlororaphis

Figure 19-39.

Comparison of chemical and biocatalytical synthesis of 5-cyano-valerarnide

(DuPont) .

confused with the nitrilases belonging to the class of hydrolases (E.C. 3) that hydrolyze nitriles to the corresponding carbon acids. For strain selection it was important that the cells did not show any amidase activity that would further hydrolyze the amide to the carboxylic acid. The biotransformation is carried out in a two-phase system with pure adiponitrile forming the organic phase. A reaction temperature of 5 "C is chosen, since the solubility of the by-product adipodiamide is only 37-42 mM in 1-1.5 M 5-cyanovaleramide.A batch reactor is preferred over a fixed-bed reactor, because of the lower selectivity to 5-cyanovaleramide that was observed and the possibility of precipitation of adipodiamide and plugging of the column. Excess water is removed at the end of the reaction by distillation. The byproduct adipodiamide is precipitated by dissolution of the resulting oil in methanol at z 65 "C. The raw product solution is directly transferred to the herbicide synthesis. By this method 13.6 tonnes have been produced in fifty-eight repetitive batch cycles with 97 % conversion and 96 % selectivity. This biotransformation was chosen over the chemical transformation because of the higher conversion and selectivity, production of more product per catalyst weight (3 150 kg per kg dry cell weight), and less waste. The catalyst consumption is 0.006 kg per kg product. 19.3.6.3

Synthesis of the Commodity Chemical Acrylamide Catalyzed by Nitrile Hydratase from 12] Rhodococcus rodochrous (E. C. 4.2.1.84)

Acrylamide (Fig. 19-40) is an important commodity monomer used in coagulators, soil conditioners and stock additives for paper treatment and paper sizing, and for adhesives, paints and petroleum recovering agents. Since acrylonitrile is the most poisonous of the nitriles, screening for microorganisms was conducted with low molecular weight nitriles instead. Acrylamide is unstable and polymerizes easily; therefore the process is carried out at a low temperature (5 "C). Although the cells, which are immobilized on polyacrylamide gel, and the contained enzyme are very stable towards acrylonitrile,the starting material has to be fed continuously to the reaction mixture because of inhibition effects at higher concentrations. The biotransformation is started with an

19.3 Examples 11451

E @CN

__+

+ H20

1

Figure 19-40.

Biocatalytical

qNH2 synthesis of acrylamide (Nitto 0

Chemical Industry).

2

1 = acrylonitrile

2 = acrylamide

E = nitrile hydratase,whole cells from Rhodococcuserythropolis

acrylonitrile concentration of 0.11 M and is stopped at an acrylamide concentration of 5.6 M. The process is operated at a capacity of 30 000 t a-'. This nitrile hydratase acts also on other nitriles with yields of 100%. The most impressive example is the conversion of 3-cyanopyridine to nicotinamide. The product concentration is about 1465 g L-l.This conversion (1.17 g L-' dry cell mass) can be named "pseudocrystal enzymation", since at the start of the reaction the educt is solid and with ongoing reaction it is solubilized. The chemical synthesis uses copper salt as catalyst for the hydration of acrylonitrile and has several disadvantages: The rate of acrylamide formation is lower than that of acrylic acid formation. The double bond of the starting material and the product causes the formation of by-products such as ethylene, cyanohydrin and nitrilotrispropionamide. Polymerization occurs. Copper needs to be separated from the product (an extra step in the chemical synthesis). The biotransformation has the advantages that no recovering of unreacted nitrile is necessary since the conversion is 100% and no copper catalyst removal is needed. This is also the first case of a biocatalytic conversion of a bulk fiber monomer. 19.3.6.4

Synthesis o f Nicotinamide Catalyzed by Nitrile Hydratase from Rhodococcus rodochrous (E.C. 4.2.1.84)* ' 4 i

Nicotinamide (vitamin B3) is used as a vitamin supplement for food and animal feed. It is the same strain that is also used in the industrial production of acrylamide (see Sect. 19.3.6.3). The biotransformation is carried out on a scale of 3000 t a-l (Fig. 19-41). In contrast to the chemical alkaline hydrolysis of 3-cyanopyridine with 4 % Lyproduct of nicotinic acid (96% yield) the biotransformation works with absolute selectivity and no acid or base is required. The biotransformation (a continuous process) is operated at low temperature and atmospheric pressure. In contrast to the old synthesis route of nicotinamide at Lonza, the new one is environmentally friendly and safe. There is only one organic solvent used throughout the whole process in four highly selective continuous and catalqc reactions. The process water, NH3 and HZare recycled.

1452

I

7 9 Replacing Chemical Steps by Biotransformations

new route

old route

NC

4

+

NH,

u 1

H

4

H

2

1

I

N

-NH,

hydrogenation

L

N

H

2

cyclization over zeolite

(-"y H

1

Pd-catalyzed dehydrogenation

1

ammonoxidation

E = nitrile hydratase, whole cells from Rhodococcus eiythropolis Figure 19-41. Comparison of chemical and biocatalytical synthesis of nicotinarnide (Lonza).

19.3.7 Epimerase 19.3.7.1

Epimerization of Clucosamine Catalyzed by Epimerase from E. coli (E.C. 5.1.3.8) [114-1161

N-Acetyl-D-mannosamineserves as the in situ generated substrate for the synthesis of N-acetylneuraminicacid. Since N-acetyl-D-mannosamineis quite expensive it is synthesized from N-acetyl-D-glucosamineby epimerization at C2. This biotransformation is integrated into the production of N-acetylneuraminicacid (Neu5Ac). By application of N-acylglucosamine 2-epimerase it is possible to start with the inexpensive N-acetyl-D-glucosamineinstead of N-acetyl-D-mannosamine (Fig. 1942). The epimerase is used for the in situ synthesis of N-acetyl-D-mannosamine

7 9.4 Some Misconceptions about lndustrial Biotransformations

,,A L

NHAc

I

1453

H

&

OH

1

OH

2

1 = N-acetyl-D-glucosamine 2 = N-acetyl-D-mannosamine E = GlcNAc 2-epimerase, enzyme from Escherichia coli

Figure 19-42. Biocatalytical epimerization of glucosamine to mannosamine (Marukin Shoyu).

(ManNAc).Since the equilibrium is on the side of the starting material, the reaction is driven by the subsequent biotransformation of ManNAc together with pyruvate to Neu5Ac. The N-acylglucosamine2-epimerase is cloned from porcine kidney, transformed and overexpressed in Escherichia coli. To reach maximal activitiy, ATP and Mg2+need to be added. Since the whole synthesis is reversible, high GlcNAc concentrations are used. The chemical epimerization of GlcNAc is used by Glaxo. The equilibrium of the chemical epimerization is on side of N-acetyl-D-glucosamine(G1cNAc:ManNAc = 4: 1).After neutralization and addition of isopropanol GlcNAc precipitates. In the remaining solution a ratio of G1cNAc:ManNAc = 1: 1is reached. After evaporation to dryness and extraction with methanol the ratio of G1cNAc:ManNAc is shifted to 1:4.

19.4

Some Misconceptions about Industrial Biotransformations

There are a lot of prejudices against biotransformations. The major ones are: Biocatalysts are too expensive. Biocatalysts only work under mild conditions. The first prejudice that biocatalysts are too expensive is only partly true. If the cost per mol or per unit weight is calculated they certainly are expensive. For example, penicillin amidase costs $ 10 OOO/kg on a bulk scale. On the other hand the cost contribution of penicillin amidase in the “splitting”of penicillin G is only $ l/kg of product[117].In the case of L-aspartic acid production the cost contribution of aspartase is even lower, $ O.l/kg. This demonstrates that it is not the absolute catalyst cost but the cost contribution of the catalyst to the final product cost that has to be considered and compared. This is also true for chemical catalysts;e. g., the bulk price of BINAP is $ 4 0 OOO/kg[117].Important parameters influencing the cost contribution are the total turnover number (mol product/mol catalyst) and the turnover frequency (mol product/mol catalyst and unit time). The second prejudice, that biocatalysts only work in an aqueous phase with low concentrations of starting material is also only partly true. The natural environment

1454

I

7 9 Replacing Chemical Steps by Biotransformations Table 19-1.

EC

Highest concentrations applied in industrial biotransforrnations enzyme

1.1.99.21 o-Sorbitol dehydrogenase

substrate

concentration

medium

1.00 M

Aqueous

1.00 M 1.20 M 1.65 M 1.83 M 2.01 M

Aqueous Aqueous MTBE Aqueous Aqueous/ organic Aqueous Aqueous Aqueous 2-Propanol Aqueous/ organic Aqueous

4.2.1.2 3.4.21.62 3.1.1.3 3.5.2.6 4.2.1.84

Fumarase Subtilisin Lipase 0-Lactamase Nitrile hydratase

1-Amino-D-sorbitol (N-protected) Fumaric acid Phenylalanine isopropylester 1-Phenylethylamine y-Lactam Adiponitrile

4.3.1.1 4.1.1.12 3.1.1.25 3.1.1.3 3.1.1.3

L-Aspartase Aspartate P-decarboxylase Lactonase Lipase Lipase

Fumaric acid Aspartic acid Pantolactone Palmitic acid Cyclopentenylester

2.00 M 2.50 M 2.69 M 3.10 M 4.16 M

4.2.1.84

Nitrile hydratase

Acrylonitrile

4.3.1.5

L-Phenylalanineammonia- trans-Cinnamic acid lyase

5.60 M (product) 9.31 M (NH3)

Aqueous

is in general the aqueous phase and ambient temperature. But the examples described above demonstrate that biocatalysts can be also applied in emulsions or even pure organic solvents (Table 19-1). Here, moreover, very high concentrations are reached, e. g. in the case of acrylamide up to 5.6 M.

19.5 Outlook

Despite the progress biocatalysis has made in the last few years its potential is still increasing. By improved screening methods new catalysts will be detected and made available in large amounts by cloning and overexpression. Directed evolution will be . M etabolic engiused to improve properties such as stability or selectivity[”’, neering will be used to analyze and remove bottlenecks in the metabolism or to create novel biocatalysts[l2O1.

References 1 The New Scientist, 1”‘issue, 22 Nov. 1956.

A. Liese, K. Seelbach, C. Wandrey, Zndustrial Biotran$omations; Wiley-VCH,Weinheim, 2000. 3 U. Bornscheuer, Industrial Biotransformations, in H. Rehm, G. Reed, A. Puhler, P. Stadler (eds.),Biotechnology Series Vol. Sb,

2

4

5

Wiley-VCH,Weinheim, 2000,277-294. J. Peters in Biotechnology 2”d edn., Vol8a, H. J. Rehm, G. Reed (eds.), Wiley-VCH, Weinheim, 1998, 391-474. A. Liese, M. Villela Filho, Production of fine chemicals using biocatalysis, C u r . Op. Biotech. 1999, 10 (6), 595-603.

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Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyriqht 0Wiley-VCH Verlaq GmbH, Weinheim 2002

20 Tabular Survey of Commercially Available Enzymes Peter Rasor

Enzymes are catalysts. Nature has designed them to perform specific tasks necessary for the survival of the organism producing the enzyme. The organic chemist tends to name enzymes “biocatalysts”which means nothing more than catalysts of biological origin. These biocatalysts bring some confusion to the well-structured world of organic chemistry: the names are unfamiliar, each enzyme has a variety of names which are all used making it as difficult as distinguishing characters in a Russian novel (e.g. Penicillin G-amidase and Penicillin acylase), when it comes to microogranisms or plants, the origin of enzymes is described in Latin (type face italic), in order to add to the confusion, the names of microorganisms may change over time, for example Pseudomonas cepacia is now Burkholderia cepacia, Candida cylindracea is Candida rugosa, even mammalian sources can be described differently - esterase from hog liver or pig liver, but lipase from porcine pancreas (type face not italic). For identifjmg synonyms or finding out the correct name of an enzyme, the Enzyme Nomenclature Database (EC database) can be searched or downloaded under http:// www.expasy.ch/enzyme/. If the chemist is still not confused and has mastered this hurdle, the manufacturers or suppliers introduce brand names for marketing reasons, and may even change names once in a while. Additionally, not every supplier gives full information on the origin of the biocatalyst and may use old names of microorganisms while other suppliers already use new names. Furthermore, the same biocatalyst by description may behave differently in a specific reaction: for example, lipase from Candida rugosa from Amano (LipaseAY) differs from Lipase MY or OF from Meito Sangyo with respect to activity and stereoselectivitybecause it consists of a number of catalytically active species which differ depending on the production strain used and thus, on the manufacturer.

1462

I

20 Tabular Survey ofCommercially Available Enzymes Table 20-1.

Abbreviation of most commonly used biocatalysts

Abbreviation Lipase from

ANL BCL (PCL) CAL CAL-A CAL-B CLL CRL (CCL) CVL

Abbreviation Lipase from

Aspergillus niger Burkholderia cepacia (formerly Pseudomonas cepacia) Candida antarctica Candida antarctica, type A Candida antarctica, type B Candida lipolytics Candida rugosa (formerly C. cylindracea) Chromobacterium viscosum (identical to Pseudomonas glumae)

PcamL PFL

Penicillium carnembertii Pseudomonasfluorescens

PfragiL PPL ProqL PSL RML (MML)

Geotrichum candidum

TLL (HLL)

Pseudomonasjagi Porcine pancreas Penicillium roquefortii Pseudomonas sp. Rhizomucor miehei (formerly Mucor miehei) Rhizopus oryzae (other names: RNL - Rhizopus niveus, RDL Rhizopus delemar, RJ L Rhizopusjavanicus) Thermomyces lanuginosa (formerly Humicola lanuginosa)

GCL

Abbreviation Esterase from

PLE

ROL

Abbreviation Alcohol dehydrogenase from

Pig liver

YADH TBADH HLADH

Yeast Themoanaerobium brockii Horse liver

Since the full enzyme name according to the EC nomenclature is rather long, the most commonly used enzymes have gotten abbreviations. For esterases and lipases there are certain rules: in most cases, the first (two or three) letters characterize the source, the last the type of enzyme (E for esterase, L for lipases) (see Table 20-1). Alcohol dehydrogenases are treated similarly. All this may explain why many publications give only incomplete information on the exact type of enzyme used in the work described and why many references to enzymes are simply wrong. The author strongly recommends to provide at least the following information: Parameter

Example 1

Example 2

Name of the product

Lipase Type XI11

CHIRAZYME L-2, lyo

Description (if the name of Lipase from Pseudomonas the product is a brand sp. name or non-descriptive)

arctica, type B

Formulation

Powder

Powder

Manufacturer

Sigma

Roche Diagnostics

Lipase from Candida a n t -

20 Tabular Survey ofCornrnercially Available Enzymes Table20-2.

Available screening sets

Enzyme type

Company

Alcohol dehydrogenases Esterases SC lipases Nitrilases Proteases Transaminases (aminotransferases)

ThermoGen, BioCatalytics Altus, Fluka, Roche, ThermoGen BioCatalytics Altus BioCatalytics

In the laboratory protocol, lot. no. and activity (incl. assay no. or assay conditions) must be recorded as well in order to track variation in results because of lot to lot inconsistency. Every development of a biocatalytic reaction starts with a screening for the most appropriate enzyme. Some companies offer screening sets (or kits) containing the most commonly used enzymes (Table 20-2). Some Sets are single use (Altus, ThermoGen) while others contain enough material to perform depending on the scale 5-20 experiments (BioCatalytics, Fluka, Roche). These sets may include enzymes available on industrial scale or on research scale only. The following companies offer screening setlkits for quick enzyme selectiori (Table 20-2). While some companies include only industrial scale enzymes, others contain enzymes only available at lab quantities. Diversa Co. offers an enzyme subscription program for lipases, esterases, nitrilases, cellulases, glycosidases, phosphatases, and transaminases (aminotransferases). Some enzymes of Novozymes A/S (formerly Novo Nordisk A/S) were widely distributed on an experimental stage (SP nnn). Table 20-3 lists the most important Table 20-3.

List of experimental enzymes by Novozymes, current products names and suppliers

~

Old name Characterization SP 361 SP 409

S P 382 SP 435

SP 523

SP 524 SP 525

S P 526

Immobilized enzyme mixture from Rhodococcus sp. containing nitrilase, nitril hydratase,esterase, epoxide hydrolase and amidase activity Immobilized lipase from Candida antarctica, containing type A & B Immobilized lipase from Candida antarctica, type B, rec. in Aspergillus oryzae Lipase powder from Thermomyces lanuginosus (formerly Humicola lanuginosa) Lipase powder from Rhizomucor miehei, rec. in Aspergillus oryzae Lipase powder from Candida antarctica, type B, rec. in Aspergillus oryzae Lipase from Candida antarctica, type A, rec. in Aspergillus oryzae

Current brand name

Availability

discontinued

discontinued Novozym 435 Novo-Nordisk CHIRAZYME L-2, Carrier 2 Roche Diagnostics CHIRAZYME L-8, lyo

Roche Diagnostics

CHIRAZYME L-9, lyo

Roche Diagnostics

CHIRAZYME L-2, lyo

Roche Diagnostics

CHIRAZYME L-5, lyo

Roche Diagnostics

I

1463

20 Tabular Survey ofComrnercially Available Enzymes Table 20-4.

Enzyme producers/suppliers and brief characterization

Company

Adress

Tel./Fax/Email/WWW

Focus/Characterization’

Altus

Altus Biologics lnc. 625 Putnam Avenue Cambridge, MA 02139-4807 USA

Tel.: +1(G17) 299-2900 Fax: +1 (617) 299-2999 Email: [email protected] http://www.altus.com

Manufacturer of stabilized enzymes for use in industrial, biocatalytical,diagnostic, and medicinal applications. No enzyme production itself. Biocatalytical process development.

Amano

Amano Pharmaceutical Co., Ltd. 2-7, 1-chome,Nishiki Naka-ku, Nagoya, 460-8630 Japan

Tel.: +81 (52) 211-3032 Specialtyenzyme producer Fax: +81 (52) 211-3054 for industrial, biocatalytical, http://www.amano-enzyme.co.jp diagnostic, and medicinal applications.

Asahi Diagnostics Division Chemical Hibiya-Mitsui Building Industry Co. 1-2 Yurakucho 1-chome, Chiyoda-ku Tokyo 100-8440 lapan

Tel.: +81 (3) 3259-5776 Fax: +81 (3) 325S-5741 Email: [email protected] http://www.asahi-kasei.co.jp Tel.: +44 (0) 1443843712 Fax: +44 (0) 1443 84 12 14 Email: [email protected] http://www.biocatalysts.com

Specialityenzyme producer for diagnostic and medicinal applications.

Biocatalysts

Biocatalysts Ltd Main Avenue, Treforest lndustrial Estate Pontypridd, Wales, CF37 5UD United Kingdom

BioCatalytics

BioCatalytics Inc. 39 Congress Street, Suite 303 Pasadena, CA 91105-3022 USA

Tel.: +1 (626) 229-0588 Fax: +1 (626) 535-9465 Email [email protected] http://www.biocatalytics.com

Biocatalytical process development. Experimental enzymes for biocatalysis. Limited production capacity. Distributor for Roche in USA and Canada.

Biozyme Laboratories International Ltd.

USA and Canada: 9939 Hibert Street Suite 101 San Diego, CA 92131-1029

Tel.: +1 (858) 549-4484 or (800) 423-8199 Fax: (858) 549-01 38 Email: [email protected]

Specialityenzyme producer for diagnostic and medicinal applications

Producer and distributor of enzymes for use in industrial and diagnostic applications.

USA

All other countries: Biozyme Laboratories Ltd Unit 6, Gilchrist Thomas Estate Blaenavon, South Wales, N P 4 9RL United Kingdom Calbiochem. co., CN Biosciences

Calbiochem-Novabiochem Corporation 10394 Pacific Center Court San Diego, CA 92121 Mailing Address: P.O. Box 12087 La Jolla, CA 92039-2087 USA

Tel.: (+44)1495790678 Fax: (+44) 1495791780 Email: [email protected] http://www.biozyme.com/

Tel.: +1 (858) 4509600 or (800)8543417 Fax: +I (858) 45335 52 Email: [email protected]. [email protected] http: //www.calbiochem.com http://www.cnbi.com

Supplier of enzymes and biochemicals on research scale. Focus on life science, not biocatalysis.

20 Tabular Survey ofComrnercially Available Enzymes I1465 Table 20-4.

(cont.).

Company

Adress

Tel./Fax/Ernail/WWW

Focus/C haracterization’

Diversa

Diversa Corporation 4955 Directors Place San Diego, CA 92121-1609 USA

Tel.: +1 (858) 526-5000 Fax: +1 (858) 5 2 6 5 5 51 Email [email protected] http://www.diversa.com

Discovery and development of industrial enzymes. No general biocatalyst portfolio.

DSM GistBrocades

DSM Food Specialties P. 0. Box 1 2600 MA Delft The Netherlands

Tel.: +31 (15) 2793474 Fax: +31 (15) 2793540 http://www.dsm.nl/dfs/

Enzyme producer for industrial applications (feed &food).

Fluka

see Sigma-AldrichFluka

Genencor

Genencor International, Inc. 200 Meridian Centre Blvd. Rochester, NY 14618-3916 USA

Tel.: +1 (716) 256-5200 Fax: +1 (716) 2566952 Email: [email protected] http://www.genencor.corn

Enzyme producer for industrial applications

Jiilich Enzyme Products

Juelich Enzyme Products GmbH Karl-Heinz-Beckurts-Str. 13 D-52428 Jiilich Germany

Tel.: +49 (2461) 348188 Experimental enzymes for Fax: +49 (2461) 348186 biocatalysis. Limited E-mail [email protected] enzyme production http://www.juelich-enzyme.com capacity.

Lee Scientific

Lee Scientific Inc. 2924 Mary Ave. St. Louis, MO 63144 USA

Tel.: +1 (314) 968-1091 Fax: +1 (314) 968-9851 Email: [email protected]

Specialty enzyme producer. Focus on life science and diagnostics. Some biocatalysts.

Meito Sangyo Co. Ltd.

Fine Chemicals Dept. Meito Sangyo Co. Ltd. Sankeido Bldg., 4 3 - 1 5 , Muromachi, Nihonhashi Chuo-ku, Tokyo 103-0022 Japan

Tel.: +81 (3) 3242-1795 Fax: +81 (3) 3242-1792 Email: [email protected]

Producer and distributor of enzymes for use in industrial and diagnostic applications.

Novozyme A/ s

Europe, Middle East & Africa: Novozymes France S. A. lmmeuble Challenge 92 79, Avenue Frantois Arago 92017 Nanterre Cedex, France Latin America: Novozymes Latin America Limited Rua professor Francisco Ribeiro 683 CEP 83707-660 - Araucaria - Parana Brazil

http://www.leescientific.corn/

http://www.novozymes.com Tel.: +33 146140746 Fax: +33 146140766

Tel.: +55 41641 1000 Fax: +55 416431443

Largest enzyme producer for industrial applications. Distribution agreement with Roche for chiral organic synthesis market.

20 Tabular Survey ofCommercially Available Enzymes Table 20-4.

(cont.).

Company

Adress USA: Novozymes North America Inc. 77 Perry Chapel Church Road Franklinton, N. C. 27525 Postal Address: State Road 1003 P.O. BOX 576 Franklinton, NC 27525 Asia Pacific, Hong Kong: Novozymes Asia Pacific Regional Office 7/F Chinachem Century Tower 178 Gloucester Road, Wanchai

Tel./Fax/Emailm

Focus/Characterization’

Tel.: +19194943000 Fax: +19194943450

Tel.: +852 25193380 Fax: +852 28 77 06 59

Recordati S.p.A.

Via Matteo Civitali, I 20148 Milan Italy

Tel.: +39 (02) 487871 http://www.recordati.it

Manufacturer of industrial enzymes for beta-lactam antibiotics.

Roche Diagnostics

Roche Diagnostics GmbH Roche Molecular Biochemicals Sandhofer Str. 116 68298 Mannheim Germany

Tel.: +49 (621) 7598593 Fax: +49 (621) 7598986 Email: [email protected] http://indbio.roche.com

Specialityenzyme producer for industrial, biocatalytical, diagnostic, and medicinal applications. Broad range of enzymes.

Tel.: +1 (858) 679-4050 or (800) 679-40 50 Fax: (858) 679-1438 Email: [email protected] http://www.seravac.com

Specialityenzyme producer for diagnostic and medicinal applications.

Tel.: (314) 771-5765 Fax: (314) 771-5757 Email: [email protected] http://www.sigma-aldrich.com

Manufacturer and distributor of enzymes and biochemicals on research scale. Very broad range of enzymes (Sigma and Fluka). Limited range of biocatalysts at Aldrich. Within the group, Fluka has the focus on biocatalysts on research scale. Production of selected enzymes up to medium scale.

USA & Canada: Refer to BioCatalytics Inc. Seravac

Seravac USA, Inc. 13220 Evening Creek Drive San Diego, CA 92128 USA

SigmaSigma Co. Aldrich 3050 Spruce Street Fluka (SAF) St. Louis, MO 63103 Mail: P. 0. Box 14508 St. Louis, MO 63178 USA Fluka Chemical LLC. Industriestrasse 25 CH-9471 Buchs Mail P. 0. Box 260 CH-9471 Buchs Switzerland ThermoGen ThermoCen, Inc. 2501 Davey Road Woolridge, IL 60517 USA

Tel.: +41 (81) 7552828 Fax: +41 (81) 756 5449 EMail: [email protected] http://www.sigma-aldrich.com

Tel.: +1 (630) 783-4600 Fax: +1 (630) 783-4909 [email protected] http://www.thermogen.com

Enzyme discovery. Limited enzyme production capacity. Biocatalyhcal process development.

20 Tabular Survey ofCommercially Available Enlymt!s Table 20-4.

(cont.).

Company

Adress

Tel./Fax/Ernail/WWW

Focus/Characterization I

Toyobo Co. Ltd.

Toyobo Co. Ltd. Biochemical Operations Department 17-9 Nihonbashi Koami-cho Chuo-ku Tokyo 103-8530 Japan

Tel.: +81 (3) 3660-4819 Fax: +81 (3) 366&4951 EMail: [email protected] http://www.toyobo.co.jp/e/

Speciality enzyme producer for diagnostic and medicinal applications.

Unitica Ltd.

Medical Products Division Unitika Ltd. 4-1-3, Kyutaro-machi, Chuo-ku, Osaka 541-8566 Japan

Tel.: +81(6) 6281-5021 Fax: +81 (6) 6281-5256 Email : [email protected] http://www.unitika.co.jp/ home-e.htm

Specialty enzyme producer for diagnostic and medicinal applications.

Wako Pure Chemicals Industries, Ltd.

1-2, Doshomachi 3-Chome, Tel.: +81 (6)6203-3741 Chuo-Ku, Osaka 54&8605 Fax: +81(6) 6222-1203 http://search.wako-chem.co.jp Japan

Worthington Biochemica1

Worthington Biochemical Corp.

730 Vassar Ave Lakewood, NI 08701

-

Tel.: +1(732)942-1660 Fax: +1 (732) 942-9270 http://www.worthingtonbiochem.com/

Manufacturer and distributor of enzymes and biochemicals on research scale. Focus on life science, not biocatalysis. Manufacturer and distributor of enzymes and biochemicals. Focus on life science and diagnostics.

1 Industrial applications include detergents, feed and food, pulp & paper, etc.

enzymes and gives the current brand names, wherever possible. Some enzymes have been discontinued at Novozymes but replacements are available from Roche Diagnostics (CHIRAZYME product line). Catalytic antibodies are not yet widely available. Aldrich is offering two aldolase monoclonal antibodies. The major enzyme producers and/or suppliers are listed and briefly characterized in Table 20-4. The author is aware that the list of enzyme producers is not complete. The author has made the attempt to list enzymes that are commercially available (Table 20-5)and thus can be used in biocatalysis. He knows that the list is incomplete and therefore, the reader should not rely solely on this list but rather check the suppliers listed in Table 20-4. Enzyme manufacturers also update their product portfolio continuously, so this list probably needs updating before the book is even i n print. A special word is necessary with respect to the Sigma-Aldrich-Flukaconglomerate: Fluka ha:; taken the lead in biocatalysis, while Sigma serves mostly the life science market. Especially since the Sigma catalog is a book in itself, only enzymes from Fluka are listed. The reader should be aware that the majority of enzymes is available from Sigma as well, and with respect to enzymes not typically used in biocatalysis, the portfolio may be even greater. Explanations to Table 20-5: The table is sorted by the EC number. In most cases the number is given in the

1468

I

20 Tabular Survey ofCommercially Available Enzymes

respective chapter and can be used to find the enzyme in the table. If the EC no. is not known, at least the general reaction of the enzyme class is given according to the EC nomenclature. Underneath the EC name, synonyms are given. The general reaction according EC nomenclature is denoted too. Afterwards, the product (enzyme) names are listed, one entry for each manufacturer per enzyme. If the product is sold under a brand name, this name is listed too. In one enzyme class, the entries are sorted by origin. The availability is characterized in three categories: lab, pilot and industrial scale. It refers to the scale with respect to biocatalytical reactions. The author recognized that this categorization is somewhat arbitrary and in some cases may not be correct because the actual production scale is not generally known. Hopefully, though, it will prove to be useful as a rough guide. Enzyme producers are devoted to certain markets like food & feed, detergents, diagnostics or research. Large enzyme producers such as Novozymes, Genencor or DSM Gist-brocadesare categorized as “industrial”, specialty enzyme producers like Amano, Asahi or Roche Diagnostics serve various markets and thus, scale varies from pilot to industrial. Since the enzyme demand for diagnostics is much lower than for biocatalysis, typical diagnostic enzymes are labeled as “pilot” although the manufacturing process is certainly standardized and therefore, could be call “industrial” as well. Companies serving the life sciences market (e.g. Sigma-Aldrich Fluka, Roche Diagnostics) have manufacturing capacities from small (“lab”)scale to medium scale (here termed as “pilot”).It should also be recognized that the term “pilot scale” in a context other than this table has a different meaning when comparing for example Sigma, Roche Diagnostics, and Novozymes.

20 Tabular Survey ofCommercially Available Enzymes Commercially available enzymes.

Table 20.5.

I

1469

1.1.1.-

Oxidoreductases. Acting on the CH-OH group of donors. With NADI+I or NADPf+) as accedor. Alcohol Dehydrogenase Screening Kit; Origin: microorganism, rec. in E. coli ThemoGen: ThermoCat Alcohol Dehydrogenase Kits

Lab

Ketoreductase, broad-range; Origin: microorganism, rec. in E. coli BioCatalytics: KRED- 1001

Lab

Ketoreductase, broad-range, Ongin microorganism, rec in E coli BioCatalytics

KRED-1002

Lab

Ketoreductase, broad-range, Ongin microorganism, rec in E cob BioCatalytic s

KRED- 1003

Lab

Ketoreductase, broad-range; Origin: microorganism, rec. in E. coli Lab

BioCatalytics: KRED-I004

Ketoreductase, broad-range; Origin: microorganism, rec. in E. coli Lab

BioCatalytics: KRED-1005

Ketoreductase, broad-range; Origin: microorganism, rec. in E. coli Lab

BioCatalytics: KRED-1006

Ketoreductase, broad-range; Ongin microorganism, rec in E coli BioCatalytiLs KRED-1007

Lab

Ketoreductase, broad-range, Ongin. microorganism, rec. in E coli BioCatalytiLs KRED-1008

Lab

Cholesterol Dehydrogenase, Ongin Nocardid sp Amano: Ainano 5 [CHDH-51

Pilot

7-Hydroxysteroid Dehydrogenase, Ongin P\eudomonda sp. Asahi

Pilot

1.1.1.1

Alcohol dehydrogenase.

-

Aldehyde reductase _I

~

_

_

X

~

X

*-

I

/_II

~

*

x

x

"

%

=

Alcohol Dehydrogenase, Ongin. Candida parapsilosis Julich Enzyrne Products

Lab

Fluka

Lab

Biocatalysts Sec ADH 300

Lab

Alcohol Dehydrogenase, Ongin Rhodococcus elythropolis Julicb Enzyrne Products

Lab

Alcohol Dehydrogenase, Ongin yeast Biozyme

Pilot

Alcohol Dehydrogenase, Ongin yeast Fluka

Pilot

Alcohol Dehydrogenase, Ongin yeast Roche Diagnostics Alcohol Dehydrogenase (YADH), lyo

Pilot

Alcohol Dehydrogenase Ohgin yeast Roche Diagriostics Alcohol Dehydrogenase (YADH), susp

Pilot

Alcohol Dehydrogenase, Ongin Zymomonas mobilis Alcohol dehydrogenase (NADP+). Aldehyde reductase (NADPH)

1.1.1.2 An alcohol + NADP(+) = an aldehyde + NADPH

Alcohol Dehydrogenase, Ongin Lactobacillus kefir Fluka

PllOt

1470

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

Alcohol Dehydrogenase,Ongin Lactobacillus kefir Julich Enzyme Products

Lab

Alcohol Dehydrogenase:Ongin Thermoanaerobium brockii nuka

Pilot

1.1.1.5

Acetoin dehydrogenase. Diacetyl reductase

Acetoin + NAD(+) = diacetvl + NADH

Acetoin Dehydrogenase,Ongin Lactobacillus kefir Fluka

Lab

Diketone Reductase, Ongin Lactobacillus kefir Julich Enzyme Products

Lab

1.1.1.6

Glycerol dehydrogenase.

Glycerol + NAD(+) = glycerone + NADH

Glycerol Dehydrogenase,Ongin Bacillus megatenum Asahi

Pilot

Glycerol Dehydrogenase: Ongin Geotnchum candidum Fluka

Lab

Glycerol Dehydrogenase,Ongin Klebsiella pneumoniae (formerly Enterobacter aerogenes) Roche Diagnostics Glycerol Dehydrogenase

Lab

Glycerol Dehydrogenase (GIDH), Ongin nucroorganisms Unitika Glycerol-3-phosphate dehydrogenase (NAD+).

Pilot

1.1.1.a Sn-glycerol3-phosphate+ NAD(+) = glycerone phosphate +

L-iditol2-dehydrogenase. ngin microorganism? Pilot

Sorbitol Dehydrogenase,Ongin sheep liver Fluka

Lab

(S)-lactate + NAD(+) = pyruvate + NADH Biozyme

L-Lactate Dehydrogenase,Ongin. bovine heart Fluka

Pilot

Lab

L(+)-Lactate Dehydrogenase,Ongin hog muscle Roche Diagnostics L(+)-Lact

PllOt

L-Lactate dehydrogenase, Biozyme

Pilot

L(+)-Lactate Dehydrogenase,Origin pig muscle Roche Diagnostics L(+)-Lactate Dehydrogenase (L-LDH)

PllOt

L-Lactate dehydrogenase,0 Biozyme

Pilot

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

I

1471

(cont.).

L-Lactate dehydrogenase,Ongin rabbit muscle Biozyme

Pilot

L-Lactate Dehydrogenase, Ongin rabbit muscle Fluka

Lab I

I

Lactate Dehydrogenase,Ongin Staphylococcus sp Amano

Aniano 3 [LDH-31

Pilot

1.1.I .28

D-lactate dehydrogenase.

(R)-lactate + NAD(+) = pyruvate + NADH

D-lacuc acid dehydrogenase D-lactic dehydrogenase

D-Lactate Dehydrogenase;Origin: Lactobacillus leichmanii Lab

Fluka

D(-)-Lactate Dehydrogenase;Ongin: Lactobacillus leichmannii Pilot

Roche Diagnostics: D(-)-Lactate Dehydrogenase (D-LDH)

D-Lactate Dehydrogenase;Origin: microorganisms Pilot

Toyobo

D-Lactate Dehydrogenase,Ongin nucroorganisms Umtika D-Lactate Dehydrogenase (D-LDH)

Pilot

3-hydroxybutyrate dehydrogenase.

1.I .I .30

B

D-beta-hydroxybutyrate dehydrogenase ~

ycI~-

~~

"_

_rllx, __^__x

I__y-

~

_ x

~

c

"-

(R)-3-hydroxybutanoate + NAD(+) = acetoacetate + NADH c_

y__

1 _ 1

xxx

_x-

.>^_

-*

*-"*

~

I

1

3-Hydroxybutyrate Dehydrogenase

-

Pilot

Asahi

D-3-HydrorybutyrateDehydrogenase;Origin: Pseudomonas sp. Pilot

Toyobo

3-Hydroxybutyrate Dehydrogenase;Origin: Rhodobacter sphaeroides (formerly Rhodopseudomonas sphaeroides) Roche Diagnostics: 3-Hydroxybutyrate Dehydrogenase (3-HBDH), Grade I1

Lab

3-Hydroxybutyrate Dehydrogenase;Origin: Rhodopseudomonas spheroides Lab

Fluka

1.I .I .37

Malate dehydrogenase.

-~

Y l

_I

--

(S)-malate + NAD(+) = oxaloacetate + NADH

Malic dehydrogenase

c_

X r x x x

~

~

1_

_x

Malate Dehydrogenase,Ongin microorganisms Tovoho

L

Pilot

Malate Dehydrogenase,Ongin rmcroorganisms Urukka Malate Dehydrogenase (MDH)

-

Pilot 1 1 1

Malate dehydrogenase,Ongin pig heart Biozyme

Pilot

Malate Dehydrogenase,Ongin porcine heart Fluka Amano. Ammo 3 [MDH-31

Lab Pilot

lsocitrate Dehydrogenase,Ongin porcine heart Fluka

Lab

lsocitrate Dehydrogenase,Ongin. porcine heart Ruka

Lab

1472

I

20 Tabular Survey ofComrnercially Available Enzymes Table 20.5.

(cont.).

Phosphogluconate dehydrogenase (decarboxylating). Phosphogluconic acid dehydrogenase 6-phosphogluconic dehydrogenase 6-phosphogluconic carboxylase 6PGD

1.I .I .44 6-phospho-D-gluconate + NADP(+) = D-nbulose 5-phosphate + CO(2) +NADPH

6-PhosphogluconateDehydrogenase (6PGDH). Ongin Thermoactinomces intermedius Unitika

Pilot

6-PhosphogluconicDehydrogenase,Ongin Torula yeast Fluka

Lab

6-Phosphogluconic Dehydrogenase:Ongin yeaa Fluka

Lab

Glucose 1-dehydrogenase.

1.1.1.47 Beta-D-glucose + NAD(P)(+) = D-glucono-l,5-lactone

+ NAD(P)H

Fluka

Pilot

Glucose Dehydrogenase,Ongin Bacillus sp Amano Amano 2 [GLUCDH-2]

Pilot

Glucose Dehydrogenase,Ongin. Cryptococcus un~guttulatus Ash

Pilot

Glucose Dehydrogenase,Ongin microorganisms

G6PD

D-glucose 6-phosphate + NADP(+) = D-glucono-l,5-lactone 6-phosphate +NADPH

Glucose-6-Phosphate Dehydrogenase Asahi

Pilot

Glucose-6-Phosphate Dehydrogenase (GGPDH), Ongin Bacillus stearothermophilus Unitika

PllOt

Glucose-6-phosphate Dehydrogenase,Ongin baker's yeast Fluka

PllOt

Glucose-6-phosphatedehydrogenase,Ongin Leuconostoc mesenteroides Biozyme

Pilot

Glucose-6-phosphate Dehydrogenase,Ongm Leuconostoc mesenteroides Fluka

Pilot

Glucose-6-PhosphateDehydrogenase, Ongin Leuconostoc mesenteroides Toyobo

Pilot

Glucose-6-phosphate Dehydrogenase, Ongin Leuconostoc mesenteroides Roche Diagnostics Glucose-6-phosphate Dehydrogenase (G6P-DH). susp

PllOt

Glucose-6-phosphate Dehydrogenase,Ongin Leuconostoc mesenteroides, rec in E coli Roche Diagnostics Glucose-6-phosphdte Dehydrogenase (G6P-DH). lyo

PllOt

Glucose-6-phosphate Dehydrogenase,Ongin Torula yeast Fluka

Glucose-6-phosphatedehydrogenase,Origin yeast Biozyme

Lab

Pilot

Glucose-6-phosphate Dehydrogenase,Ongin yeast Fluka

Pilot

Glucose-6-phosphateDehydrogenase,Ongin yeast Fluka

Pilot

Glucose-6-phosphateDehydrogenase,Ongin yeast Roche Diagnostics Glucose-6-phosphate Dehydrogenase (G6P-DH), lyo

Pilot

20 Tabular Survey ofCornrnercially Available Enzymes

I

1473

Table 20.5.

(cont.).

Glucose-6-Phosphate Dehydrogenase(GBPDH),Ongin Zymomonas mobilis Unitika 3-alpha-hydroxysteroid dehydrogenase (B-specific). Hydroxyproc taglandin dehydrogenase 3-alpha-HSD

Pilot

1.1.1.50 Androsterone + NAD(P)(+) = 5-alpha-androstane-3,17-dione+ NAD(P)H.

3-Hydroxysteroid Dehydrogenase Asahi

Pilot

3-alpha-Hydroxysteroid Dehydrogenase(3alphaHSDH), Ongin mcroorganisms Unitika

Pilot

3-alpha-Hydroxysteroid Dehydrogenase, Ongin Pseudomonas testosteroni Fluka

Lab Xanthine + NAD(+) + H(2)O = urat

Xanthine Olehydrogenase Asahi

Pilot

12-alpha-HydroxysteroidDehydrogenasePP ; Origin: Clostridium spec. Jiilich Enzyme Products

Lab

12-alpha-Hydroxysteroid Dehydrogenase, Ongin mcroorganisms Asahi

Glucose oxidase. Glucose oxyhydrase Beta-D-glucose oxygen 1-oxide-reductase Glucose aerodehydrogenase D Glucose- 1-oxidase y_x_

_ _ - c _ r _

>

~

x x x

Glucose Oxidase Seravac Glucose oxidase, Ongin A\pergillus niger Ammo Hyderase I

__

Pilot

1.1.3.4 Beta-D-glucose + O(2) = D-glucono-l,$-lactone+ H(2)0(2)

__ -

Industnal Industnal ~

Glucose oxidase,Ongin Aspergillus niger Amano Hyderase L

I)

__

Industnal I

Glucose oxidase, Ongin A5pergillus niger Biozyme

Pilot

Glucose Oxidase, Ongin Aspergillu$ niger Fluka

Industnal

Glucose oxidase, Ongin A\pergillus niger Novozymes Gluzyme@

Industnal

Glucose Oxidase, Ongin Aspergillus niger overproducer Roche Diagnostics Glucose Oxidase (GOD)

Industnal

Amano

Ammo 2 [GO-2]

Pilot

Glucose Oxidase, Ongin A\pergillus sp Amano Ainano LC [GOLC]

Pilot

Glucose Oxidase, Ongin Aspergillus sp Amano Arnano LD2 [GOLD-21

Pilot

Glucose Oxidase, Ongin Aspergillus sp Toyobo

Pilot

Glucose Oxidase, Ongin microorganism, rec in yeast Roche Diagnostics Glucose Oxidase (GOD)

Pilot

Glucose Oxidase, Ongin Penicillium sp Biocatalysts

Industrial

1474

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

Cholesterol oxidase.

1.1.3.6 Cholesterol + O(2) = cholest-4-en-3-one + H(2)0(2)

Cholesterol-02 oxidoreductase Asahi

Pdot

Cholesterol Oxidase, Ongin Brevibactenum sterolicum, rec in rmcroorganiqm Roche Diagnostics Cholesterol Oxidase

Pilot

Cholesterol Oxidase, Ongin rmcroorganiams Ammo

Amano 6 [CHO-6]

Pilot

Cholesterol Oxidase, Ongin mi~roorganisms Asahi

Pilot

Cholesterol Oxidase, Ongin rmuoorganisms Toyobo

Pilot

Cholesterol Oxidase, Ongin Nocardia erythropolis Fluka

PllOt

Cholesterol Oxidase, Ongin Pseudomonas sp Amano

Ammo 1 [CHO-I]

Pdot

Cholesterol Oxidase, Ongin Pseudomonas sp Ammo

Amano 2 [CHO-2]

Pilot

Cholesterol Oxidase: Ongin Pseudomonas sp Fluka

Pilot

Cholesterol Oxidase, Ongin Streptomyces cinnamomeus Pilot

Asahi

Galactose oxidase.

1.1.3.9 D-galactose + O(2) = D-galacto-hexodialose + H(2)0(2)

Beta-Galactose oxidase

Galactose Dehydrogenase, Ongin Agrobactenum sp Biocatalysts

Pilot

Alcohol oxidase.

1.1.3.13 A pnmary alcohol + O(2) = an aldehyde + H(2)0(2)

Methanol oxidase AOX

Alcohol Oxidase; Ongin Candida sp As&

PllOt

Alcohol oxidase, broad-range: Ongin microorganism, rec in E coli BioCatalytics

BRAO- 100I

Ldb

Alcohol oxidase, Origin Pichia pastons Biozyme

Pilot

Alcohol Oxidase, Ongin Pichia pastons Julich Enzyme Products

Choline oxidase.

Lab

1.1.3.17 Choline + O(2) = betame aldehyde + H(2)0(2)

Choline Oxidase; Origin: Alcaligenes sp Fluka

Pilot

Choline Oxidase, Ongin Arthrobacter globiformis Asahi

Glycerol-3-phosphate oxidase.

Pilot

1.1.3.21

Sn-glycerol 3-phosphate + O(2) = glycerone phosphate + H(2)0(2)

L-GlycerophosphateOxidase Asahi

Pdot

Glycerol 3-phosphate Oxidase, Ongin Aerococcus vindans Fluka

Lab

20 Tabular Survey ofCornrnercially Available Enzyrncs Table 20.5.

(cont.).

I

1475

L-GlycerophosphateOxidase; Origin: Aerococcus viridans Asahi

Pilot

L-Glycerol-3-phosphateOxidase; Origin: microorganism, rec. in E. coli Roche Diagnostics: L-Glycerol-3-phosphate Oxidase (GPO), stabilized

Pilot

L-alpha-GlycerophosphateOxidase; Origin: microorganisms Toyobo

Pilot

L-alpha-Glycerophosphate Oxidase; Origin: Pediococcus sp. Toyobo

Pilot

L-alpha-Glycerophosphate Oxidase; Ongin Streptococcus \p Amano Arnano 2 [GPO-2]

PllOt

wm&

1.I .3.22

Xanthine oxidase.

Xanthine + H(2)O + O(2) = urate + H(2)0(2)

Xanthine oxidoreductase Hypoxanthine oxidase Hypoxanthiiie-xanthine oxidase Schardinger enzyme

Xanthine oxidase, Ongin buttemulk Biozyme

Pilot

Xanthine Oxidase, Ongin buttermilk Fluka

Pilot

Xanthine Oxidase, Ongin cow rmlk Pilot

Fructose 5-dehydrogenase. D-Fmctose dehydrogenase

1.1.99.1 1 D-fructose + acceptor = 5-dehydro-D-fmcto\e

+ reduced acceptor

D-Fructose Dehydrogenase, Ongin Gluconobacter sp Toyobo

Pilot

Formate dehydrogenase.

Formate ~

1.2.1.2

_I”*

nase;

Fluka

Pilot

Formate Dehydrogenase; Origin: Candida boidinii Jiilich Enzyme Products

Lab

Formate Dehydrogenase, rec., Ongin: Candida boidinii, overexpressed in E. coli Roche Diagnostics. Formate Dehydrogenase (FDH), rec

Industrial

Formate Dehydrogenase rec., Ongin E coli nuka

Lab

Formate Dehydrogenase, Ongin microorganisms Unitika Formate Dehydrogenase (FDH)

Pilot

Formate Dehydrogenase, Ongin Pseudomonas sp Fluka

Lab

Formate Dehydrogenase, Ongin Pseudomonas sp Fluka

Lab

Formate Dehydrogenase, Ongin Xilana digitata (formerly Candida biodinii) Roche Diagnostics Formate Dehydrogenase (FDH)

Industnal

Formate Dehydrogenase, Origin yeast Fluka

Aldehyde dehydrogenase (NAD(P)+).

Pilot

1.2.1.5 An aldehyde + NAD(P)(+) + H(2)O = an acid + NAD(P)H

Aldehyde Dehydrogenase, Ongin baker’s yeast Fluka

Lab

1476

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

Aldehyde dehydrogenase, Ongin yeast Biozyme

Pilot

Aldehyde Dehydrogenase; Ongin yeast

Glyceraldehyde 3-phosphate dehydrogenase (phosphorylating).

1.2.1.12

NAD-dependent glyceraldehyde-3-phosphatedehydrogenase Tnosephosphate dehydrogenase GAPDH

D-glyceraldehyde 3-phosphate + phosphate + NAD(+) = 3-phospho-D-glyceroyl phosphate + NADH

Glyceraldehyde-3-Phosphate Dehydrogenase (GapDH); Origin: Bacillus stearothermophilus Unitika

Pilot

Glyceraldehyde-3-phosphate dehydrogenase; Origin: rabbit muscle Biozyme

Pilot

Glyceraldehyde-3-phosphate Dehydrogenase, Ongin rabbit m u d e Fluka

Ldh AD(+) + H(2)O = formate +

Formaldehyde Dehydrogenase, Ongin Pseudomonay putida Fluka Formaldehyde Dehydrogenase, Ongin Pseudomonas sp Toyoho

PyNvlL oxidase

Pyruvate Oxidase , On Asahi

Lab Pilot Pyruvate + phosphate + O(2)+ H(2)O = acetyl phosphate + CO(2) +H(2)0(2) Pilot

Pyruvate Oxidase, Ongin Lactobacillus plantarum, rec E coli Pilot

Bilirubin oxidase.

1.3.3.5 Bilirubin + O(2)= biliverdin + H(2)O

Acyl-CoA oxidase.

1.3.3.6 Acyl-CoA + O(2)= trany-2.3-dehydroacyl-CoA + H(2)0(2)

Acyl-CoA Oxidase, Ongin Arthrobacter sp Asahi

Pilot

Acyl-CoA Oxidase, Ongin mcroorganism~ Amano Ammo 3 [ACO-3]

Pilot

Alanine dehydrogenase. Alanine Dehydrogenase Asahi L-Alanine Dehydrogenase, Ongin Bacillus cereu5 Julich Enzyme Products Alanine Dehydrogenase, Ongin Bacillus ytearothermophilus Unitika Alanine Dehydrogenase (AlaDH)

NH(3) + NADH. Pilot Lab Pilot

L-Alanine Dehydrogenase, Ongin Bacillus subtili\ Fluka

Lab

20 Tabular Survey of Commercially Available Enzymes Table 20.5.

(cont.).

il!

I

-

1477

1.4.1.3

Glutamate dehydrogenase (NAD(P)+).

L-glutamate + H(2)O + NAD(P)(+) = 2-oxoglutarate + NH(3) + NAD(P)H

Glutamic dehydrogenase.

Glutamate dehydrogenase; Origin: beef liver Biozyme

Pilot

Glutamate Dehydrogenase; Origin: bovine liver Fluka

Pilot

L-Glutamate Dehydrogenase; Origin: bovine liver Roche Diagnostics: L-Glutamate Dehydrogenase (GlDH), lyo.

Pilot

Glutamate Dehydrogenase; Origin: microorganisms Toyobo

Pilot

Glutamate Dehydrogenase, Ongin Proteus sp Toyobo

Pilot

1.4.1.9

Leucine dehydrogenase.

L-leucine + H(2)O + NAD(+) = 4-methyl-2-oxopentanoate + NH(3) + NADH

Leucine Dehydrogenase; Origin: Bacillus cereus Pilot

Biocatalysts

Leucine Dehydrogenase; Origin: Bacillus sp Pilot

Toyobo

Leucine Dehydrogenase, Ongin Bacillus stearothermophilus Unitika Leucine Dehydrogenase (LeuDH)

Pilot L-phenylalanine + H(2)O + NAD(+) = phenylpymvate + NH(3) +

L

I

~

^

- -I

_

Phenylala

_x

S

Unitika Phenylalanine Dehydrogenase (PheDH)

Pilot

PhenylalanineDehydrogenase, Ongin Spororarcina sp D-amino acid oxidase.

1.4.3.3 A D-amino acid + H(2)O + O(2) = a 2-0x0 acid + NH(3) + H(2)0(2)

D-Amino Acid Oxidase; Origin: hog kidney Lab

Fluka

D-Amino Acid Oxidase; Origin: hog kidney Lab

Fluka

D-Amino Acid oxidase, Ongin porcine kidney Bioryme

PllOt

D-Amino Acid Oxidase, Ongin Tngonopsis vanabilir Recordati

DAAO Beads

Industnal

D-Amino Acid Oxidase, carrier-fixed, Ongin: Tngonopsis vanabilis Roche Diagnostics D-Armno Acid Oxidase (D-AOD), caner-fixed

lndustnal

D-Amino acid Oxidase, immobilized, Ongin Tngonopsis vanabilis Fluka

Amine oxidase (flavin-containing). Monoamine oxidase Tyramine oxidase Tyraminase Amine oxidase

Industnal

1.4.3.4 RCH(2)NH(2) + H(2)O + O(2) = RCHO + NH(3) + H(2)0(2)

Tyramine Oxidase; Origin: Arthrobacter sp. Asahi

Pilot

1478

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.). ewmm*m*

Dihydrofolate reductase.

-_~ ~ ~ ---_ --~

1.5.1.3 5,6,7,8-tetrahydrofolate+ NADP(+) = 7,8-dihydrofolate + NADPH

- _.

Tetrahydrofolate dehydrogenaqe

"

~

x

Y

I

X

X

Sarcosine Oxidase Asahi

Pilot

Sarcosine Oxidase, Ongin mcroorganisms PllOt

With other acceptors.

1.5.99.

TMADh

Tnmethylamine x_11_____

lyl-"**-

~

-"--%%

~

~

.

+ H(2)O + acceptor = dimethylamine + formaldehyde +reduced acceptor

~

L

~

c

I

_

"

^-"

_"__c

~

Trimethylamlne Dehydrogenase;Ongin Paracoccus spec R

Glutathione reductase (NADPH).

1.6.4.2 NADPH + oxidized glutathione = NADP(+) + 2 glutathione

Glutathione Reductase, Origin baker's yeast Fluka

NADPH dehydrogenase. NADPH diaphorase.

Lab

1.6.99.1 NADPH + acceptor = NADP(+) + reduced acceptor

Diaphorase(NADPH), Ongin Bacillus megatenum Asahi

Pilot

Diaphorase I, Ongin Bacillu5 stearothermophilus Unitika

Pilot

Asahi

PllOt

Uricase;Origin: Bacillus fastidiosus Fluka

Lab

Uricase;Origin: Bacillus sp Toyobo

Pilot

Uricase; Ongin pig liver Biozyme

Dihydrolipoamide dehydrogenase. Lipoanude reductase (NADH) E3 component of alpha-ketoacid dehydrogenase complexes Lipoyl dehydrogenase Dihydrolipoyl dehydrogenaqe

Pilot

1.8.1.4. Dihydrohpoamide + NAD(+) = lipoamide + NADH

Diaphorase(NADH), Ongin Bacillus megatenum Asahi

Pilot

Diaphorase II, Ongin Bacillus stearothermophilus Unitika

Pilot

20 Tabular Survey ofCornrnercially Available Enzymes Table 20.5.

I

1479

(cont.).

Diaphorase,Ongin Clostndium kluyven Fluka

~~~~Laccase A,

Pilot

Unshiol oxidase _3

_I__' -----ww-s*

~

_....*""

-*

4 henzenediol + O(2) = 4 henzosemiquinone + 2 H(2)O ^-xx

*"m-_y_w-/-**__I

u__-

-

_

--

***--_l___i--*

Ongin Agancus hispru5 Julich Enzyme Products

-

I

Laccase C, Ongin: Conolus versicolor Julich Enzyme Products

-

Lab Lab

Laccase,Ongin: rec. nucroorganism Novozymes. DeniLiteTM

---

ascorhate + O(2) = 2 dehydr *xj__

Asahi

--*

__I_

Pilot

Ascorbate Oxidase, Ongin Cucumber Amano. Amano 2 [ASO-21

PllOt

-

Ascorbate oxidase, Ongin Cucurbita sp Biozyme

Pilot

Fiuka

Pilot

Ascorbate Oxidase, Ongin nucroorganisms Amano Aman a

.-.-

Oxidoreductases. Acting on a peroxide as acceptor (peroxidases).

1.11

Brornoperoxidase; Ongin Corallina officinalis Fluka

Lab

Catalase. -

)

I

~

1.11.1.6 ~

~

I

I

x

I

x

x

~

~

~

.

*% - -,

m

y

u

(

I ^ a -

*-m -m M p -----

2 H(2)0(2) = O(2) + 2 H(2)O

Catalase Biocatalysts CATALASE

Industnal

Catalase Seravac

Industrial

Catalase; Origin: Aspergillus niger Amano: Catalase NL "Amano"

Industrial

Catalase, Ongin Aspergillus niger Biozyme

Pilot

Catalase, Ongin Aspergillus niger Fluka

Industrial

Catalase; Origin: Aspergillus niger Novozyrnes: CatazymeB

Industrial

Catalase; Origin: Aspergillus niger Roche Diagnostics: Catalase, technical grade

Industrial

Catalase; Origin: Aspergillus niger, rec. Novozyrnes: TerminoxTM Ultra

Industrial

Catalase, Ongin beef liver Biozyme

Pilot

Catalase, Ongin beef liver Roche Diagnostics

Industnal

-

1480

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

Catalase;Origin: bovine liver Industrial

Fluka

Catalase, immobilized on Euperglt C; Origin: bovine liver Lab

Fluka

Catalase;Origin: Corynebacterium glutamicum Industrial

Roche Diagnostics

Catalase, Ongin Micrococcus lysodeikticus Lab

Fluka

Catalase;Ongin nucroorganisms Pilot

Fluka

Catalase;Ongin microorgmsms Toyobo

Industnal

1.1 1 .I .7

Peroxidase.

Donor + H(2)0(2) = oxidized donor + 2 H(2)O

Myeloperoxidase. X I X X

-_I

x

_ “ I I _ _ _ ~

~

“ I %

1

_

1

Lactoperoxidase,Ongin bovine nulk Biozyme

Pilot

Lactoperoxidase,Ongin bovine milk Fluka

Pilot

Peroxidase, Ongin Copnnus cinereus Novozymes. NovoLym 502

Industnal

Peroxidase, Ongin Copnnus cinereus Novozymes

NS18010

Industnal

Peroxidase, Ongin horse radish Fluka

Industnal

Peroxidase, Ongin horseradish Amano

Amano 2 [PO-21

Pilot

Peroxidase, Ongin horseradish Ammo

Amano 3 [PO-31

Pilot

Peroxidase, Ongin horseradish Biocatalysts

Industnal

Peroxidase, Ongin horseradish Biozyme

Pilot

Peroxidase, Ongin horseradish Roche Diagnostics: Peroxidase (POD), Grade I

lndustnal

Peroxidase, Ongin horseradish Roche Diagnostics Peroxidase (POD), Grade I1

Pilot

Peroxidase , Ongin horseradirh Seravac

Industnal

Peroxidase, Ongin horseradish Toyobo

Pilot

Lab

Chloride peroxidase. Chloroperoxidase

1.11.1.10

2 RH + 2 chlonde + H(2)0(2) = 2 RCI + 2 H(2)O

Chloroperoxidase,Ongin Caldanomyces fumago Fluka

Pilot

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

I

(cont.).

Chloroperoxidase,Ongin Leptoxyphium fumago Julich Enzyme Products

Lab

Lipoxygenase.

1.13.11.12 Linoleate + O(2) = (9Z,IlE)-( 13S)-13-hydroperoxyoctadeca-9,1l-dienoate

Lipoxidase Carotene oxidase Lipoperoxidase

LipoxygenaseII, Ongin. pea, rec in E coli Biocatalysts

Pilot

-_

-

x

Lipoxygenase111, Ongin. pea, rec in E coli Biocatalysts

Pilot

Fluka

Lab

Lipoxidase, Ongin soybeen Biozvme

Pilot

1.13.12.4

Lactate 2-monooxygenase.

(S)-lactate + O(2) = acetate + CO(2) + H(2)O.

Lactate oxidative decarboxylase. Lactate oxidase Lactate oxygenase.

Lactate Oxidase Pilot

Asahi

Lactate Oxidase, Ongin Pediococcus 5p Fluka

Pilot

Oxidoreductases. Acting on paired donors with incorporation of molecular oxygen. With NADH or NADPH as one donor, and incorporation of one atom of oxygen.

1.14.13.-

2-Tridecanone Monooxygenase, Ongin Pseudomonas cepacia Fluka

Lab

Cyclopentanone monooxygenase.

~

~

"

~

1.14.13.16

- ~ -- ~ ~

-

x

~~

Cyclopentanone + NADPH + O(2) = 5-valerolactone + NADP(+) + H(2)0

~

y_"_1___

ne Monooxygenase, Ongin P Cyclohexanone monooxygenase. w.**-**

~ ~- ~ _y

I___

1.14.13.22 Cyclohexanone + NADPH + O(2) = 6-hexanolide + NADP(+) +

Cyclohexanone oxygenase -*--

--

x

" 1

*

x-

~

C"

* "

x

Cyclohexanone Monooxygenase, Ongin Acinetobacter sp Fluka

Lab

Cyclohexanone Monooxygenase; Origin: E. coli overproducer Lab

Fluka

Cyclohexanone Monooxygenase, Ongin Nocardia globemla Lab

Fluka

Cyclohexanone Monooxygenase, Ongin Xanthobacter sp Fluka

Lab

2-hydroxybiphenyl+ NADH + O(2) = 2,3-dihydroxybiphenyI+ NAD+ + H 2 0

Fluka

monooxygenase, Lab

1482

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.). W#w

Camphor 5-exo-methylene hydroxylase Cytochrome p450-cam.

1.14.15.1 (+)-camphor + putidaredoxin + O(2) = (+)-exo-5-hydroxycamphor+

(+)-Camphor Monooxygenase,Ongin Pseudomonas putida Fluka

Lab

Camphor 5-monooxygenase.

oxidizedputidaredoxin + H(2)O

1.14.18.1

Monophenol monooxygenase.

L-tyrosine + L-DOPA + O(2) = L-DOPA + DOPAquinone + H(2)O

Tyrosinase Phenolase Monophenol oxidase Cresolase

Tyrosinase, Ongin mushroom Fluka

Lab

Fluka

Lab

eroxide dismut ~

-

*--

**

__ ---~

-

~

Superoxide Dismuta Umtika

1.15.1.1 2 peroxide radical + 2 H(+) = O(2) + H(2)0(2) i__L_

l

_i

%"

Y

I

Bacillus stearothennophilus Superoxide Dismutase (SOD)

-

*

* *

1_

~

-

Pilot

Superoxide dismutase; Origin: bovine erythrocytes Pilot

Biozyme

Superoxide Dismutase;Origin: bovine erythrocytes Pilot

Fluka

Superoxide Dismutase; Origin: bovine erythrocytes Lab

Roche Diagnostics: Superoxide Dismutase (SOD)

Superoxide Dismutase, Ongin bovine liver Fluka

Pilot

2.1.1.6

Catechol 0-methyltransferase.

S-adenosyl-L-methionine + catechol = S-adenosyl-L-homocysteine +guaiacol

Brenrkatechin-0-methyl-Transferase Fluka

Lab Sedobeptulose 7-phosphate + D-glyceraldehyde 3-phosphate =

Glycoaldehyde transferase -___*_)_-_-I

--_I-

-_I__)__^-

= C _ i - _

Transketolase,Ongin baker's yeast FlUka I

Transketolase ,Ongin E coli

-

~~

I - _ Y 1

x

Y

_x

x

11_^1

_^--

Lab

__

Fluka

Lab

Transketolase,Ongin. E coli K12 (rec ) Julich Enzyme Products

Lab

2.2.1.2

Transaldolase. Dihydroxyacetone transferase Glycerone transferase

Sedoheptulose 7-phosphate + D-glyceraldehyde 3-phosphate = D-erythrose 4-phosphate + D-fructose 6-phosphate "+

Fluka

x

I

~

~-

Lab I

Transaldolase,Ongin E coli K12 (rec ) Julich EnLyme Products

Lab

20 Tabular Survey ofCommercially Available Enzymes 1143

Table 20.5.

(cont.).

Glucosamine-phosphateN-acetyltransferase.

2.3.1.4 Acetyl-CoA + D-glucosmne 6-phosphate = CoA + N-acetyl-D-glucosanune 6-phosphate

Phosphoglucosamine transacetylase Phosphoglucosanune acetylase

Phosphotransacetylase,Ongin Bacillus stearothennophilus Urutlka

Phosphotransacetylase (PTA)

Lab

Camihne acetylase Lab

Gamma-glutamyltranspeptldaseGlutamyl transpeptldase ~

-

-

-(__I-vw

*--s-e-

* e I

(5-L-glutamyl)-peptide + an anuno acid = pepude + 5-L-glutamyl-aminoacid

~~-

-%-

I

I -

gamma-Glutamyltransferase. Ongin beef ludney Biozyme

_

y

I

-

m

I

_

_

~

l

I

I

~

_

I

~ __ ~ l ~ _l m

Pilot

gamma-Glutarnyl Transpeptidase, Ongin hog ludney Fluka

Lab

Muscle phosphorylase A and B Amylophosphorylase Polyphosphorylase

-

in rabbit musc e

((1,4)-alpha-D-glucosyl)(N)+ phosphate = [ ( I ,4)-alpha-D-glucosyl] (N- I)+ alpha-D-glucose I-phosphate -x

-**.'

-

p L -------x l-

-l__l_

Lab UDP-glucose + D-fructose = UDP + sucrose

UDP-glucose-fructose glucosyltransferase Sucrose-UDP glucosyltransferase --**<-

I -

I_)_

Sucrose Synthase, Ongin nce grains Julich Enzyme Products

Lab

1,4-alpha-glucan branching enzyme.

2.4.1.18 Formation of 1,6-glucosidic hnkages of glycogen

Glycogen branchng enzyme Amyb(1.4 to 1,6)transglucosidase Branching enzyme Amylo-( 1,4-1,6)-transglycosylase

Transglucosldase, Ongin Aspergillus niger Amano Transglucosidase L "Amano"

lndustnal

Lactose synthase.

2.4.1.22 UDP-galactose + D-glucose = UDP + lactose.

UDP-galactose-glucose galactosyltransferase N-acetyllactosamine synthase FlUka

Beta-N-acetylglucosamin yl-glycopeptide beta-l,4-galactosyltransferase. Glycoprotein 4-beta-galactosyltransferase Thyroid galactosyltransferase UDP-galactose-glycoprotein galactosyItransferase

-~

------ - -- -- - -

Julich Enzyme Products

UDP-galactose + N-acety-beta-D-glucosanunylglycopeptide = UDP +beta-D-galactosyl-1,4-N-acetyl-beta-D-glucosmnylglycopeptide

"

Beta-l,4-Galactosyltransferase, Ongin S Ijy

2.4.1.38

)

Lab

1484

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

2.4.1 .I51

N-acetyllactosaminide alpha-I ,3-galactosyltransferase.

UDP-galactose + beta-D-galactosyl-(1,4)-N-acetyl-D-glucosaminyl-R = UDP + alpha-D-galactosyl-( 1,3)-beta-D-galactosyI-(1,4)-N-acetyl-D-glucos

Galactosyltransferase

Fluka

Lab

alpha-l,3-Galactosyltransferase,Ongin E coli, rec

Punne nucleoside + phosphate = punne + alpha-D-nbose 1-phosphate

Inosine phosphorylase PNPase _ x x x

I

xxI

1~

Y

l

l

-

I

i

Purine-Nucleosidephosphorylase Ash

Pilot

Purine-NucleosidePhosphorylase,Ongin microorganisms

Transferring nitrogenous groups. Transaminases (aminotransferases).

-

1_7--1__1mm-

~

*a*-i

-.,,-

~

*

I

I

Transaminase, branched-chain, L-specific, Ongin microorganism, rec in E coli BioCatalytics

AT-I02

Lab

Transaminase, broad-range, D-specific, Ongin rmcroorganism, rec In E coli BioCatalytics

AT-I03

Lab

Transaminase, broad-range, L-specific, Ongin rmcroorgannm, rec in E coli

Transaminase A Glutarmc--oxaloacehc transaminase Glutamic-aspartic transarmnase

- Glutamicoxaloacetic transaminase, -

-.,

*

*

<

^*

L-aspartate + 2-oxoglutarate = oxaloacetate + L-glutamate

~

_1

Ongin pig heart

Biozyme

Pilot

Glutamate-OxaloacetateTransaminase,Ongin pig heart (mitochondnal) Roche Diagnostics Glutamate-OxaloaLetateTransaminase (GOT)

Pilot

Glutamic-OxalaceticTransaminase, Ongin porcine heart

Glutarmc--pyruvic transaminase. Glutamc--alanine transarmnase. I_

I~

L-alanine + 2-oxoglutarate = pyruvate + L-glutamate

"~

Glutamate-PyruvateTransaminase, Ongin pig heart Roche Diagnostics Glutamate-Pyruvate Transaminase (GIT)

Pilot

Glutamic-pyruvictransaminase, Ongin pig heart Biozyme

Pilot

Glutamic-PyruvicTransaminase, Ongin porcine heart

exohnase type IV Asahi

Pilot

Hesperidinase,Ongin Penicillium decumbens Amano

Hespendinase "Amano" Conc

Industrial

20 Tabular Survey ofCommercially Available Enzymes 1143

(cont.)

Table 20.5.

Hexokinase, Ongin Saccharomyces sp Toyobo

Pilot

Hexokinase, Ongin yeast Biozyme

Pilot

Hexokinase, Ongin yeast Fluka

Industnal

-

Hexokinase, Ongin. yeast FlUka

Industnal

f

___~

Xlmf

~

~ - -- ~ -

c

~~

stearothermophilus Unitlka Glucokmase (GlcK)

Pilot

Glucokinase, Ongin Zymomonas mobilis

-fructose 6-phosphate =

~---~-~~-~-~

Glycerokinase ATP glycerol 3-phosphotransferase Y

~

x

_ _ I

x

x

Glycerol Kinase Asahi

__

__

ATP + glycerol = ADP + glycerol 3-phosphate I

L

r

X

--

I

~

I - -

_x

I_-

Pilot

Glycerol Kinase, Ongin. Arthrobacter sp Ammo Amano 2 [GK-21

Pilot

_ I

Glycerokinase, Ongin Bacillus stearothemophilus Pilot Roche Diagnostics Glycerokinase, sol I

_

I ~

Pilot

-

Glycerol Kinase, Ongin E coli Fluka

Pilot I

Glycerol Kinase, Ongin rmcroorganisms Toyobo

------

Phosphoenolpyruvate kmase Phosphoenol transphosphorylase ~

-*mix

*--

-

_i-__^~ I *<--I

PllOt

- --

--

--

--

ATP + pyruvate = ADP + phosphoenolpyruvate ~

i

I-r

~

-*-=---

1

__<_(

*-

PllOt Biozyme

Pilot

Pyruvate Kinase, Ongin rabbit muscle Biozyme

Pilot

Pyruvate Kinase, Ongin rabbit muscle Fluka

Lab

Pyruvate Kinase, Ongin Zymomonas mobilis Unitika Pyruvate Kinase (PK)

PllOt

1486

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

"%

2.7.1.72

Streptomycin 6-kinase.

ATP + streptomycin = ADP + streptomycin 6-phosphate

Streptidine kmase Streptomycin 6-phosphotransferase APH(6)

X

Acetate Kinase, Ongin Bacillus stearothermophilus Unitika Acetate Kinase (AK)

"

A

x(

c

-- -

_x

Pilot

Acetate Kinase, Ongin E coli

2.7.2.3

Phosphoglycerate kinase.

ATP + 3-phospho-D-glycerate = ADP + 3-phospho-D-glyceroyl phosphate

Phosphoglycerate Kinase,Ongin Bacillus stearothermophilus Umtika Phosphoglycerate Kinase (PGK)

Pilot

Biozyme

Pilot

Creatine Kinase; Origin: pig heart Pilot

Biozyme

Creatine Kinase; Origin: rabbit muscle Pilot

Biozyme

Creatine Phosphokinase, Ongin rabbit m u d e Fluka

Lab

2.7.4.3

Adenylate kinase.

ATP + AMP = ADP + ADP

Myokmase Adenylic kmase Adenylohnase

2.7.7.8

Polyribonucleotide nucleotidyltransferase. Polynucleotide phosphorylase.

(RNA](N+I) + phosphate = (RNA)(N) + a nucleoside diphosphate

Polynucleotide Phosphorylase, Ongin Bacillus stearothermophilus Unrtika Polynucleotide Phosphorylase (PNPase)

CMP-N-acetylneuraminic acid synthetase CMP-NeuNAc synthetase CMP-bialate pyrophosphorylase CMP-sialate

Prlot CTP + N-acylneurdrmnate = diphosphate + CMP-N-acylneurmnate

Acting on ester bonds. rmcroorganisms Roche Diagnostics CHIRAZYME Lipases & Esterases, Screening Set Industnal Enzymes 2

lndustnal

20 Tabular Survey ofCommercially Available Enzymes 11437

(cont.).

Table 20.5.

3.1.1.1

Carboxylesterase. Ah-esterase B-esterase. Monobutyrase Cocaine esterase

A carboxylic ester + H(2)O = an alcohol + a carboxylic anion

Esterase basic kit FlUka

Lab

-

Esterase, Ongin Bacillus sp Fluka

Lab

Esterase; Ongin Bacillus stearothermophilus Fluka

Lab

Esterase,Ongin Bacillus thermoglucosidasius Fluka

Lab

.

-

I

Esterase, Ongin Candida lipolytica Fluka

Lab

Esterase , Ongin. Candida rugosa Alms

_ _

_

Indusmal

_-

I

-

Esterase, Ongin hog liver Fluka

Indusmal

Esterase, immobilized on Eupergim C, Ongin. hog liver Fluka

-

Pilot

-

Esterase lsoenzyme 1, Ongin hog liver Fluka

_ _ _

I

-

-

_-__

Pilot

-

I

Esterase, Ongin horse liver FIuka -

-_

- - - - - _-

-

Lab

-

-~

Esterase Screening Kit , Ongin microorganism, rec in E coli ThemoGen Quickscreen Esterase IOts

-

-

Lab

I _ _ _

Esterase, Ongin Mucor miehei Fluka -. Esterase; Ongin: pig liver Julich Enzyme Products Esterase PL I

I

I

I

- ~ _ _ _ _ _ _

I -

-

I

___-

___

--

I

Esterase. immobilized, Ongin. pig liver Roche Diagnostics CHIRAZYME E-I, c - f , lyo

--

~I_

I

-

I

--

_-_^

1 1 1

AltuS ~ ~ - ~ - l l _ l _ l l l l l

"

l_-l___l

Pig Liver Esterase, Ongin pig liver Roche Diagnostics PLE, technical grade, susp

I

_-_-

__l__-----l_-I___I1_I

Esterase, Ongin pig liver, fraction 1 Roche Diagnoshcs CHIRAZYME E-I, lyo ~

-

l

_

_

-

l

_

-

_

_

I

I

_

_I

l _ l _ l l

- --

--

___

l l _ l

- -

-

-I_

I

Esterase, Ongin Rluzopus arrhizus Julich Enzyme Products Esterase EL9

~-

I

_

_

_

I

I

I

Esterase, Ongin Rhodotorula pilimanae Julich Enzyme Products Esterase EL5

-

-

--_^_

-

Esterase, Ongin Saccharomyces cerevisiae Fluka ~

---

__

_--

I

I

-

x

-

ll__l

Desacetylesterase, Ongin Therm sp ,rec in E coli Recordati Desa-REC I - I x

--__

I

Esterase, Ongin Thermoanaerobium broclai FlUka

x

Lab

__ _~

I

_--_^

-

I _

-___

__

-

Industrial

__I_

__

I_ I

Industnal - _ _ _ _ I _ _ ^ -

__I^

~

- -1111

I

--

-

I

Industnal l__l

._

I

-

Industnal I

_ _ _ _ - - _ Lab ___

I

Lab

--

__

~

--

~

I

~

----

__

I _

Esterase,Ongin pig liver, fractlon 2 Roche Diagnostics CHIRAZYME E-2, lyo

_I____

___

Lab

Industnal

__ _

I

-

-_

Pig Liver Esterase, Ongin. pig Liver

~

-

-

I

_

I

I

~

Lab

-

Indusmal I

I

Lab

1488

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont )

Triacylglycerol lipase. Lipase Tnglycende lipase Tnbutyrase

-

c x _ _

-~

-

3.1 .I .3

Lioase basic kit Fluka

Lab

Lipase extension kit Fluka

Lab

Monoglyceride Lipase Asahi

Pilot

Lipase, Ongin Achromobacter sp Meito Sangyo. Lipase AL

Industnal

Lipase, immobilized,Ongin Achromobacter sp Meito Sangyo. Lipax ALCIALG

Industnal

Lipase, Ongin Alcaligenes sp Altus

Industnal

Meito Sangyo

Industndl

Lipase PL

Lipase; Ongin Alcaligenes sp Meito Sangyo' Lipase QLL

lndustnal

Lipase, Ongin Alcaligenes sp Meito Sangyo Lipase QLM

Industnal

Lipase, immobilized, Ongin Alcaligenes sp Meito Sangyo Lipase PLCPLG

Industrial

Lipase, immobilized; Origin: Alcaligenes sp. Meito Sangyo: Lipase QLCIQLG

Industrial

Lipase, Ongin Alcaligines sp Roche Diagnostics CHIRAZYME L-10, lyo

Industnal

Lipase, Ongin Aspergillus mger Altus

lndustnal

Lipase, Ongin Aspergillus niger Amano Lipase A "AmdnO" 6

lndustnal

Lipase; Origin: Aspergillus niger Ammo : Lipase AS

Industrial

Lipase; Origin: Aspergillus niger Amano : Lipase DS

Industrial

Lipase; Origin: Aspergillus niger Fluka

Industrial

Lipase;Origin: Aspergillus niger Fluka

Industrial

Lipase, immobilized in Sol-Gel-AK, Ongin Aspergillus niger Fluka

Lab

Lipase, Ongin Aspergillus oryzae Fluka

lndustnal

Meito Sangyo

Industnal

Lipase SL

Lipase , Ongin. Burkholdena ~p Fluka

PllOt

Lipase, Ongin Candida antarctica Fluka

Lab

Lipase A , Ongin Candida antarctica Fluka

Industrial

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

I

1489

(cont.).

Lipase, immobilized; Origin: Candida antarctica Fluka

Industrial

Lipase, immobilized in Sol-Gel-AK; Origin: Candida antarctica Fhka Lipase, immobilized in Sol-Gel-AK on sintered glass; Origin: Candida antarctica Fluka

Pilot Lab

Lipase, type A; Origin: Candida antarctica

AlNS

Industrial

Lipase, type B; Ongin Candida antarctica Alms

Industnal

Lipase B, Ongin Candida antarctica, rec Fluka

Industnal

Ongin. Candida antarctica, type A Roche Diagnostics CHIRAZYME L-5, lyo

Industnal

Lipase, Ongin: Candida antarctica, type A Roche Diagnostics CHIRAZYME L-5, sol

Industnal

Lipase, immobilized, Ongin Candida antarctica, type A Roche Diagnostics CHIRAZYME L-5, c -f , lyo

Industnal

Lipase; Ongin Candida antarctica, type A, rec in Aspergillus oryzae Novozymes NovoCor AD

Industnal

Lipase; Ongin Candida antarctica, type A, rec Novozymes NovozymB 868

Industnal

in

Aspergillus oryzae

Lipase, Ongin Candida antarctica, type B Roche Diagnostics CHIRAZYME L-2, lyo.

Industnal

Lipase, Ongin Candida antarctica, type B Roche Diagnostics. CHIRAZYME L-2, sol

Industnal

Lipase, immobilized, Ongin Candida antarctica, type B ME L-2, c.-f , C2, lyo (Novozym 435)

Industnal

Cdndida antarctica, type B Roche Diagnostics CHIRAZYME L-2, c -f , C3, lyo

lndustnal

Lipase, immobilized, Ongin Candida antarctica, type B Roche Diagnostics CHIRAZYME L-2, c -f , lyo

Industnal

Lipase, Ongin Candida antarctica, type B, rec in Aqpergillus oryzae Novozymes. Nocozym 525 L

Industnal

Lipase, immobilized, Ongin Candida antarctica, type B, rec in Aspergillus oryzae Novozymes NovozymB 435

Industnal

Lipase; Ongin. Candida cylindracae Julich Enzyme Products. Lipase LEI 1

Lab

Lipase, Ongin Candida cylindracea Biocatalyqts

lndustnal

Lipase, Ongin Candida cylindracea Fluka

Industnal

Lipase, Ongin Candida cylindracea Flub

Industnal

Meito Sangyo

Industnal

Lipase MY

Lipase, Ongin Candida cylindracea Meito Sangyo Lipase OF

lndustnal

Lipase, Ongin Candida cylindracea Meito Sangyo Lipase OF%

InduStndl

-

_ _

1490

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

Lipase, immobilized: Ongin Candida cylindracea Meito Sangyo Lipase OFC/OFG

-

-

Lipase, immobilized in Sol-Gel-AK; Ongin Candida cylindracea Fluka

Industrial PllOt Pilot Industnal

Lipase; Ongin Candida rugosa Altus

Industnal

I

Lipase, Ongin Candida rugosa Alms. ChiroCLEC-CR (dry)

lndustnal

Lipase, Ongin. Candida rugosa Alms ChiroCLEC-CR (slurry)

lndustnal

Lipase, Ongin. Candida rugosa Amano. Lipase AY "Amano" 30

lndustnal

Lipase, Ongin. Candida rugosa Amano. Lipase AYS

lndustnal

Lipase, Ongin Candida rugosa (formerly C cylindracea) Roche Diagnostics: CHIRAZYME L-3, lyo

Industnal

-

Lipase, purified, Ongin Candida rugosa (formerly C cylindracea) Roche Diagnostics CHIRAZYME L-3, punfied, lyo

Pilot

Lipase, purified, immobilized, Ongin: Candida rugosa (formerly C cylindracea) Roche Diagnostics. CHIRAZYME L-3, punfied, c -f , C2, lyo

Pilot

Lipase, Ongin Candida utihs Fluka Lipase, Ongin Chromobactenum viscosum Alms Lipase, Ongin: Cromobactenum V ~ S C O S U ~ Asahi Lipase, Ongin: Geotnchum candidum Alms

Lab Industnal Pilot lndustnal

Lipase, Ongin. hog pancreas Fluka

Lab

Lipase, Ongin hog pancreas Fluka

1ndus tnaI

-

Lipase, immobilized in Sol-Gel-AK, Ongin hog pancreas Fluka

Pilot

Pancreatin, Ongin. hog pancreas Industnal Roche Diagnostics CHIRAZYME L-8, sol

Indu stnaI

Lipase 6 , covalently linked to carrier; Origin: microorganisms Fluka

Pilot

Lipoprotein Lipase, Ongin microorganisms Amano Amano 6 [LPL-6]

Pilot

Lipase, Ongin Mucorjavanicus Alms

Industnal

Lipase: Ongin. Mucor javanicus Amano Lipase M "Amano" 10

Industnal

20 Tabular Survey ofCommercially Available Enzymes

I

1491

Table 20.5.

(cont.).

Lipase, Ongin Mucorjavanicus Fluka

Industnal

-

Lipase, Ongin Mucor meihei AltuS I

-

- - Industrial

-

_ I

Lipase, Ongin Mucor nuehei Fluka

-

Lab

-

-

Lipase, Ongin Mucor nuehei Roche Diagnostics CHIRAZYME L-9, lyo

Industnal

Roche Diagnostics CHIRAZYME L-9, sol.

Industnal

Lipase, immobilized: Ongin Mucor rmehei Fluka Lipozymea, immobilized

Industnal

Lipase, immobilized: Ongin: Mucor miehei Roche Diagnostics CHIRAZYME L-9, c - f , C2. lyo

Industrial

Lipase, immobilized, Ongin Mucor miebei Roche Diagnoshcs CHIRAZYME L-9, c -f

Industnal

Lipase, immobilized in Sol-Gel-AK, 0 Fluka

Lab

Lipase, immobilized in Sol-Gel-AK on sintered glass, Ongin Mucor nuehei Fluka

Lab

Lipase, Ongin Mucor nuehei, rec FlUka

Industnal

Lipase, Ongin Penicillium camembertii Ammo. Lipase G "Amano" 50

Industnal

Lipase: Ongin Penicillium roqueforti Fluka

Pilot

Lipase, Ongin Penicillium roquefortii Amano Lipase R

Indubtnal

Lipase; Ongin porcine pancreas Alms

Industnal

Lipase, Ongin porcine pancreas Roche Diagnostics CHIRAZYME L-7, lyo

Industnal

-

Lipase, Ongin porcine pancreas Roche Diagnoshcs. Lipase

-

Pilot

-

Lipase, Ongin Protein engineered in rec. Aspergillus Novozymes Lipolase Ultra -

- -

I

Industnal

- -

-

Lipase, Ongin Protein engineered in rec Aspergillus Novozymes LipoPnme'M

Industnal

__

I

Lipase, Ongin Pseudomonas aeroginosa Altus

Industnal

Lipase, Ongin Pseudomonas cepacia Altus

-

I _

__

Industnal

-

Lipase, Ongin. Pseudomonas cepacia Altus CkoCLEC-PC (dry)

-

lndustnal

-

ChiroCLEC-PC (sluny)

Altus

-

I

Industrial

-

_ I

-

_ I

Lipase, Ongin Pseudomonas cepacia -

I

Amano -

I

Lipase PS-C

- _

I

-

_-

Industnal Industrial _ I

1492

I

20 Tabular Survey ofCornrnercially Available Enzymes Table 20.5.

(cont.).

Lipase, Ongin Pseudomonas cepacia Amano Lipase PS-D

Industnal

Lipase, Ongin. Pseudomonas cepacia Fluka

Industnal

Lipase; Origin: Pseudomonas cepacia Fluka

Industrial

Lipase, immobilized in Sol-Gel-AK; Origin: Pseudomonas cepacia Ruka

Pilot

Lipase, immobilized in Sol-Gel-AK on sintered glass, Ongin Pseudomonas cepacia Fluka

Lab

Lipase, immobilized on Ceramic particles; Ongin Pseudomonas cepacia Fluka

Lab

Lipase, Ongin Pseudomonas fluorescens Ammo LipaseAK

Indurtnal

Lipase: Origin: Pseudomonas fluorescens Fluka

Pilot

Lipase, immobilized in Sol-Gel-AK, Ongin Pseudomonas fluorescens Fluka

Pilot

Lipase, immobilized in Sol-Gel-AK on sintered glass, Ongin Pseudomonas fluorescens Fluka

Lab

Lipase, immobilized on Eupergit C , Ongin Pseudomonas fluorescens Fluka

Lab

Lipase, Ongin. Pseudomonas sp Roche Diagnostics CHIRAZYME L-6, lyo

Industrial

Lipase, immobilized, Ongin Pseudomonas sp Toyobo

Industnal

Lipoprotein Lipase, Ongin Pseudomonas sp Ammo Amano 3 [LPL-31

Pilot

Lipoprotein Lipase, Ongin Pseudomonas sp Toyobo

Pilot

Lipase, Ongin. Pseudomonas stutzen Meito Sangyo Lipase TL

Industrial

Lipase, Ongin Rhizomucor miehei DSM Gist-brocades. Piccantase

Industnal

Lipase, Ongin Rhizomucor nuehei Fluka

Indu\tnal

Lipase, Ongin Rhizomucor nuehei, rec in Aspergillus oryzae Novozymes NovozymB 388

Indwtnal

Lipase, Ongin Rhizomucor nuehei, rec in Aspergillus oryzae Novozymes PalataseB

Industnal

Rhizomucor miehei, rec in Aspergillus oryzae Lipase, immobilized, 0 Novozymes LipozymeB RM IM

Induanal

Lipase, Ongin Rhizopus arrhizus Fluka

Industnal

Lipase, Ongin: Rhizopus delemar Alms

Industrial

Lipase, Origin Rhizopus delemar Fluka

Industnal

Altus

Industnal

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

I

1493

(cont.).

Lipase;Origin: Rhizopus niveus Amano: Newlase F

Industrial

Lipase;Origin: Rhizopus niveus Fluka

Pilot

Lipase, Ongin Rhizopus niveus Julich Enzyme Product? Lipase LE9

Lab

Lipase, Ongin Rhizopus oryLae Altus

Industnal

Lipase, Ongin Rhizopus oryzae Amano Lipase F-APIS

Industnal

Lipase; Origin: Rhizopus oryzae Ammo : Lipase F-DS

Industrial

Lipase, Ongin Rhizopus sp. Meito Sangyo' Lipase UL Lipase, Ongin Thermomyces lanuginosa Altus

Industnal

Lipase, Ongin Thermomyces lanuginosa, rec Aspergillus oryzae Novozymes Novozym 677 BG

Industnal

Lipase;Origin: Thermomyces lanuginosa rec. in Aspergillus oryzae Novozymes: GreasexB

Industrial

Lipase;Origin: Thermomyces lanuginosa rec. in Aspergillus oryzae Novozymes: LipolaseB

Industrial

Lipase, Ongin Thermomyces lanuginosa rec in Aspergillus oryzae Novozymes. NovozymB 27007

lndustnal

Lipase, Ongin Thermomyces lanuginosa rec in Aspergillus oryzae Novozymes NovozymB 398

Industnal

Lipase, Ongin Thermomyces lanuginosa rec in Aspergillus oryzae Novozymes NovozymB 735

lndustnal

Lipase; Origin: Thermomyces lanuginosa rec. in Aspergillus oryzae Novozymes: NovozymB 871

Industnal

Lipase; Origin: Thermomyces sp. (formerly Humicola sp.) Roche Diagnostics: CHIRAZYME L-8, lyo.

Industrial

Lipase; Origin: Thermus aquaticus Fluka

Lab

Lipase; Origin: Thermus flavus Fluka

Lab

Lipase; Origin: Thermus thermophilus Fluka

I.ah

Lipase, Ongin wheat germ Fluka

PIlOt

Phosp Phosphatidylcholine 2-acylhydrolase Lecithinase A Phosphatidase Phosphatidolipase c_

-

yI_xI_y^l_l~

~

-

-i-____

Phospholipase A2, Ongin. Aglustrodon halys Fluka

Phosphatidylcholine + H(2)O = 1-acylglycerophosphoholine + a fattyacid anion -*"

-

_ f _

~

*-*

Lab

Phospholipase A2, Ongin bovine pancrea5 Fluka

Pilot

Phospholipase A2, Ongin hog pancreas Fluka

Pilot

1494

20 Tabular Survey ofCornmercially Available Enzymes

I

Table 20.5.

(cont.).

PhospholipaseA2; Origin: porcine pancreas Biocatalysts

Pilot

PhospholipaseA2, Ongin porcine pancreas Industnal

3.1.1.7

Acetylcholinesterase.

Acetylcholine + H(2)O = choline +acetate

True cholinesterase Choline esterase I Cholinesterase

Acetylcholinesterase;Origin: bovine erythocytes Pilot

Biozyme

Acetylcholine Esterase, Ongin Electrophorus electncus Fluka

Lab 84

3.1.1.8

Cholinesterase.

An acylcholine + H(2)O = choline + a carboxylic acid amon

Pseudocholinesterase Acylcholine acylhydrolase Butyrylcholine esterase Non-specific cholinesterase x-

~

l

y

_

_

_

~

- -~ -

x

X

_

"

~

1

xx

Y

Y

)

i

Butyrylcholineesterase, Ongin horse serum Biozyme

Pilot

ButyrylcholineEsterase, Ongin horse semm Fluka

Lab

3.1.1.11

Pectinesterase.

Pectin + N H(2)O = N methanol + pectate

Pectin methylesterae Pectm demethoxylase Pectin methoxylase ~

~

-

I

_ _c L _

_

I

Pectinesterase

-

-

I _-I ~ __I-jli

~

_ _I ~

xI -_XI_

Novozymes Cellubnx@

I

x

-

~

~

x

x

-

Industnal

Pectinesterase Novozymes

NovoclairTM FCE

lndustnal

Pectinesterase Novozymes

Pectinex BE

-

Industnal

Pectinesterase,Ongin Aspergillus niger Amano

Pectinase P

lndustnal

Ammo

Pectinase PL "Amano"

Industnal

-

Pectin Esterase;Ongin orange peel Fluka

lndustnal

Pectinesterase,Ongin rec microorganism Novozymes

NovoshapeB

Industnal

Pectinesterase,Ongin rec rmcroorganism Novozymes

Pectinex SMASH

Sterol esterase. Cholesterol esterase Cholesterol ester synthase Tnterpenol esterase

lndustnal

3.1.1.13 A steryl ester + H(2)O = a sterol + a fatty acid

Cholesterol Esterase Pilot

Asahi

Cholesterol Esterase, lyo., Ongin Candida rugosa (formerly C cylindracea) Roche Diagnostics Cholesterol Esterase, lyo I_

Pilot

_I_

Cholesterol Esterase, sol., Ongin Candida rugosa (formerly C cylindracea) Roche Diagnostics Cholesterol Esterase, sol.

Pilot

1 1 1

Cholesterol Esterase, Ongin hog pancreas Fluka

Pilot

20 Tabular Survey of Commercially Available Enzymes

I

1495

(cont.).

Table 20.5.

Cholesterolesterase, Ongin pig pancreas Biozyme

Pilot

-

I

-

I

_

Amano. Amano 2 [CHE-Z]

PllOt

Cholesterol Esterase;Origin: Pseudomonas sp. Amano: Amano 3 [CHE-3]

Pilot

Cholesterol Esterase; Origin: Pseudomonas sp. Asahi

Pilot

Cholesterol Esterase, Ongin Pseudomonas sp Toyobo

Pilot

3.1.1.20

Tannase.

Digallate + H(2)O = 2 gallate

Tnacylglycerol + H(2)O = diacylglycerol + a fatty acid anion

Cleanng factor lipase Diglycende lipase Diacylglycerol lipase *

w

-

~

~

-

-

-

-

-

~

~

~

e

a

~

-m -em*

v

~

~~

*-!.

*

p----xyxx

_ ^ L x p

Lipoprotein Lipase,Ongin Chromobactenum viscosum Fluka I

I

Pilot

I

I

I

I

I

I

I

LipoproteinLipase,Ongin Pseudomonas fluorescens Pilot

-

3.1.3.1

Alkaline phosphatase. Alkaline phosphomonoesterase Phosphomonoesterase GIycerophosphatase

An orthophosphonc monoester + H(2)O = an alcohol +phosphate

Phosphatase, alkaline Seravac

Pilot

Phosphatase, alkaline; Origin: Bacillus sp. Biocatalysts

Pilot

Phosphatase alkaline; Origin: bovine intestinal mucosa Fluka

Industrial

Phosphatase alkaline; Origin: calf intestinal mucosa FlUka

Industrial

Phosphatase alkaline, immobilizedon Agarose; Origin: calf intestinal mucosa Fluka

Industrial

Phosphatase alkaline, immobilizedon Agarose, Ongin calf intestinal mucosa Fluka

__

-

I

I

I

I I _

I _ _

-

I

I

-

I _

__

I

Phosphatase, alkaline, Biozyme Phosphatase, alkaline, highly active; Origin: calf intestine, rec. in Pichia pastoris Roche Diagnostics: Phosphatase, alkaline, EIA Grade, highly active Phosphatase, alkaline, Ongin E coli Fluka

---

~

_

_

_

-

Phosphatase, alkaline , Ongin Eschenchia coli Asahi I

I

_

_-_-

-

Phosphatase, alkaline,Ongin calf intestine Roche Diagnostics Phosphatase, alkaline, EIA Grade

_ _

Industnal Industnal I

I

Pilot Industrial Lab Pilot

I

Phosphatase, alkaline; Ongin microorganisms Unitika

Pilot

1496

20 Tabular Survey ofCommercially Available Enzymes

I

(cont )

Table 20.5.

-sw~~u-~&<~~e

Acid phosphatase.

3.1.3.2

Acid phosphomonoesterase. Phosphomonoesterate Glycerophosphatase

An orthophosphonc monoester + H(2)O = an alcohol +phosphate I

~

1

"

c

ase, acid, otato Roche Diagnostics Phosphatase, acid, grade I1

Pilot

Phosphatase, acid. Ongin potatoes Fluka

Pilot

Myo-inositol hexalasphosphate + H(2)O = D-myo-inositol1,2,4,5,6-pent&sphosphate + phosphate

Phytase Phytate 3-phosphatase Myo-inositol-hexaphosphate 3-phosphohydrolase *<_^__*we

__- ___

f^

I

*_"" "

-

x

L

-

I

x

x

x

_

x

Phytase; Ongin Aspergillus niger Amano

Industnal

Phytase; Ongin Peniophora lycii, rec In Asp oryzae Novozymes Bio-feed@Phytase

Industnal

A phosphatidylcholine + H(2)O = 1,2-diacylglycerol+ cholinephosphate

Lipphosphodiesterase I Lecithinase C Clostndium welchn

**

Asahi

Pilot

Phospholipase C, Ongin. Bacillus cereus Fluka

Pilot

Phospholipase C, Ongin Clostndium perfnngens Fluka

Pilot

~

Glycerophosphorylcholine Phosphodiesterase, Ongin microorganisms Asahi

Pilot

Phospholipase D, Ongin Streptomyces chromofuscua Asahi

Pilot

Phospholipase D, Ongin Streptomyces chromofuscu? Fluka

Pilot

Phospholipase D, Ongin Streptomyces sp Asahi

PllOt

Pilot

Deoxyribonuclease 1.

3.1.21 .I

Pancreatic DNase DNase Thymonuclease _-"

~

x_xI

--

-

Endonucleolytic cleavage to 5'-phosphodinucleotide andS'-phosphooligonucleotldeend-products _j

l i l

ribonuclease I,Ongin bovine pancrea5

Guanylonbonuclease Aspergillus oryiae nbonuclease RNase N 1 RNase N2

- -Ribonuclease T1, Ongin Aspergillus oryzae Fluka

Two-stage endonucleolytic cleavage to 3'-pho\phomononucleotides and3'-pho?phoohgonucleotidesending in G-P with 2,3'-cychc pho\phateintermediates. I

-

Lab

I

20 Tabular Survey ofComrnercially Available Enzymes

1497

Table 20.5.

(cont.).

mXX.I*

3.1.27.5

Pancreatic ribonuclease.

Endonucleolytic cleavage to 3'-phosphomononucleotides and 3'-phosphooligonucleotldesendmg in C-P or U-P with 2',3'-cyclicphosphate intermediates

RNase. RNase I Rh'ase A. Pancreatic RNase

_x

-_

"-

_l_l_l

Ribonuclease,Ongin beef pancreas

__

----~-~

x I x I ~ ~ ~ ~ - x I I I x I

Biozyme

Pilot

RibonucleaseA, Ongin bovine pancreas Fluka

__

Pilot

-

-

_ I

Ribonuclease,immobilized on EupergitC, Ongin bovine pancreas Fluka

Lab

Aspergillus nuclease S1.

3.1.30.1

Endonuclease S1 Single-stranded-nucleateendonuclease Deoxynbonuclease S 1

- ~ - ~ -

__x_xa-

~

Amano

Endonucleolytlc cleavage to 5'-phosphomononucleotldeand 5'-phosphooligonucleotideend-products

-_

- -

-m

x

I

~

__m

--_ ~

~

I

-

~

~

-

~

-

~

Enzyme Rp-1

~

-~

x

y

I

~

Industnal

Nuclease P1; Ongin Penicillium citnnum Fluka

Lab

i

Micrococcal endonuclease. -

~

~

-

-

~

-

-

~

=

~

~

-

----~

-

Nuclease micrococcal,Ongin Staphylococcus aureus

~

~

I-_I___

-

~

-

"_I

Endonucleolytic cleavage to 3'-phosphomononucleotide and 3'-phosphooligonucleotideend-products

~

~

~

-

~

" ~ " - ~ - - X _I _ Y

_ v y ~ ~ ~ I I F - ~ - * x x x I ~ l l x I I x l l

Fluka

Lab

3.2.1 .I

Alpha-amylase. 1,4-alpha-D-glucanglucanohydrolase Glycogenase -=--*

-*--

~

Amylase, Ongin Aspergillus niger Ammo' Gluczyme NL4 2 l_li

-

I

Endohydrolysis of 1,4-alpha-glucosidiclinkages in oligosacchandes and polysacchandes

-----I

xIIxuI*-I)*_^-

_ _

Industnal

alpha-Amylase,Ongin Aspergillus oryzae Fluka

Industnal

-

Amylase, O n g n Aspergillus oryzae Ammo

Amylase DS

Industnal

Amylase, Ongin Aspergillus oryzae Ammo Biozyme F10 SD

-

-

Industnal

_x

Amylase, Ongin Aspergillus oryzae Ammo

Biozyme S Conc

Industnal

Taka-Diastase,Ongin Aspergillus oryzae Fluka

Pilot

alpha-Amylase,Origin Bacillus amyloliquefaciens Fluka

Industnal

alpha-Amylase,Ongin Bacillus lichenifomis Fluka

Industnal

alpha-Amylase,Ongin Bacillus suhhlis Fluka

Industnal

Amylase, Ongin Bacillus suhtilis Amano

Amylase A "Amano" Conc

Induanal

alpha-Amylase;Origin: fungus Novozymes: Fungarnylm

Industrial

alpha-Amylase: Origin: hog pancreas Fluka

Lab

1498

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

Amylase; Origin: microbacterium Amano : AMT "Amano"

Industrial

Amylase; Origin: microorganisms Novozymes: AquazymB Amylase; Origin: microorganisms Novozymes: BAN (Bacterial Amylase Nova)

Industrial

alpha-Amylase; Origin: rec. microorganism Novozymes: DuramylTM

Industrial

alpha-Amylase; Origin: rec. microorganism Novozymes: Liquozyme@

Industrial

alpha-Amylase; Origin: rec. microorganism Novozymes: TermamylB

Industrial

alpha-Amylase, Ongin rec nucroorganism Novozymes Termamyl, Type LS

Industnal Industnal

Beta-amylase.

3.2.1.2

1,4-alpha-D-glucan maltohydrolase Saccharogen amylase Glycogenase

beta-Amylase, Ongin barley Fluka

-

-

Pilot

--

-_I_

beta-Amylase, Ongin. sweet potato Fluka

Pilot

Glucan 1,4-alpha-glucosidase.

3.2.1.3

Glucoamylase 1.4-alpha-D-glucan glucohydrolase Amyloglucosidase Gamma-amylase

-

spergillus niger Industnal

Amyloglucosidase; Ongin rec nucroorganism Novozvmes. AMG

Cellulase. ~

"

-

I

_?

Industnal

3.2.1.4

Endoglucanase Endo- 1,4-beta-glucanase Carboxymethyl celluldse _

Hydrolysis of tennmal 1,Clinked alpha-D-glucose residues successively from non-reducing ends of the chams with release

"

"m-m---

A Fluka

x

Hydrolysis of 1,4-alpha-glucosidic linkages in polysacchandes so asto remove successive maltose units from the non-reducing ends of the chains

Endohydrolysis of 1.4-beta-D-glucosidic linkages in cellulose

____ __-

Cellulase NovoLymes Novozym@ 342

Industnal

- -

Cellulase, Ongin Aspergillus niger Ammo Cellulase A "Amano" 3

-

Industnal

Cellulase; Ongin Aspergillus niger Ammo Cellulase DS

Industnal

Cellulase; Origin: Aspergillus niger Fluka

Industrial

Cellulase, Ongin fungus Novozymes Celluzyme@

Industnal

Cellulase, Ongin. Hunucola insolens Fluka

Lab

20 Tabular Survey of Commercially Available Enzymes (cont.).

Table 20.5.

I

1499

Cellulase, Ongin rec mcroorganism Novozymes CarezymeB -

Industnal

-

I

Cellulase, Ongin rec mcroorganism Novozymes: DeniMaxB

-

Industnal

_ _

_-

I

I

I

Cellulase, Ongin Tnchoderma longibrachiatum Fluka _

I

Industnal I

I

Cellulase, Origin Tnchoderma reesei Fluka

Lab

Cellulase, Ongin: Tnchoderma vinde Amano Cellulase T "Amano" 4

Industnal

hoderma vinde

Endo-l A-beta-glucanase Endo-l,3-beta-glucanase Laminannase

- . - - ~~~------~~-y1I_

Endohydrolysis of 1,3- or 1,CIinkages in beta-D-glucans when the glucose residue whose reducing group i s involved in the linkage to be hydrolysed i s itself substituted at C-3

-_x_I

beta-glucanase Novozymes . CerefloB

Industnal

beta-glucanase Novozymes: FinizymB

Industrial

beta-glucanase, heat-stable Novozymes: UltrafloB

Industrial

beta-Glucanase; Origin: Aspergillus niger Fluka

Industrial

beta-Glucanase, Ongin Bacillus subtilis FlUka

Industnal

Inulinase.

3.2.1.7

---

Inulase

e, Ongin Aspergillus niger

Endohydrolysis of 2,l-beta-D-ftuctosidic linkages in inulin _I_(

_-i

-*(

---*

l i

~

Endo-1,&beta-xylenase.

3.2.1.8

1,Cbeta-D-xylan xylanohydrolase

Endohydrolysis of 1,4-beta-D-xylosidic linkages in xylans

Xylanase Novozymes : Pulpzyme'M HC

Industrial

Xylanase; Origin: Aspergillus niger Amano: Hemicellulase "Amano" 90

Industrial

Xylanase, Ongin Aspergillus niger Ammo Hemcellulase "Amano" 90

-

____

I_--

Xylanase, Ongin bactena Fluka I _

-

I I

-

-

I

- -

_ I

Xylanase, Ongin rec rmcroorganism Novozymes PentopanTMMono

-

-

I

I

Xylanase, Ongin rec mcroorganism Novozymes Shearzyme'M

-

Industnal

-

-

I

Industrial

- --

I -

-

-

-

Industnal Industnal

Xylanase; Origin: Tnchoderma viride FlUka

Lab

1500

I

20 Tabular Survey ofCommercially Available Enzymes

(cont ).

Table 20.5.

3.2.1 .I 1

Dextranase. Alpha-l,6-glucan-6-glucanohydrolase Dextranase Novozymes Dextranase

lndustnal

Dextranase,Ongin Chaetonuum erraticum) Amano Dextranase L "Amano"

Industnal

Dextranase,Ongin Paecilomyces lilacinus Ruka Chitinase. Chitodextnnase 1,4-beta-poly-N-acetylglucosamin1dase Poly-beta-glucosanunidase

Pilot

3.2.1 .I 4 Hydrolysis of the 1.4-beta-linkages of N-acetyl-D-glucosanune polymers of clutin

E_

Chitinase, Ongin. bean leaves Julich Enzyme Products Chitinase BB

Lab

Chitinase, Ongin Streptomyces gnseus Ruka

Pilot

Chitinase, Ongin sugar beet Fluka

Lab

Chitinase, Ongin sugar-beet leaves Julich Enzyme Products: Chitinase ZR

Lab Random hydrolysis of 1,4-alpha-D-galactos1duroniclinkages in pectate and other galacturonans

Pectin depolymerase Pectinase

-

Fluka

Industrial

Pectinase, Ongin mould Fluka

Industnal

Pectinase, Ongin Rhizopus sp

Hydrolysis of the 1,4-heta-linkages between N dcetyl-D-ghIcosanune and N-acetylmuranuc acid In peptidoglycan aryotes cell walls

Murmdase

Lysozyme Seravac

Pilot

Lysozyme, Ongin chicken egg white Biozyme

Industnal

Lysozyme, Ongin hen egg white

Sialidase Neuraminldase N-acylneuraminate glycohydrolaie Alpha-neurammidase "_

Hydrolysis of alpha-(2->3)-, alpha-(2->6)-, alpha-(2->8)-glycosidic linkages of terminal sialic residues in oligosacchandes, glycoprotein\, glycolipids, colominic acid and synthetic substrates

c _

Neuraminidase,Ongin Clostndium perfnngens Fluka

Pilot

Neuraminidase,immobilized on Agarose4B, Ongin Clostndium perfnngens Ruka

Pilot

Neuraminidase,Ongin nucroorganisms Unitika

Pilot

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

I

1501

(cont.)

Neuraminidase,Origin Streptococcus sp Toyobo

Pilot

-

Neuraminidase.Ongin Vibno cholerae Fluka

Lab

a*

Alpha-glucosidase. Maltase Glucoinvertase. Glucosidosucrase Maltase-glucoamylase **

XC--I-X_l%^*-Ym-M1

* --**-_

*',-

"

alpha-Glucosidase,Ongin. Aspergillus niger Fluka

-*--

Hydrolysis of terminal, nowreducing 1,4-linked D-glucose residues with release i

r

--c

*

-Ju*x-

Industnal

alpha-Glucosidase,Ongin Bacillus stearothermopbilus Urutlka alpha-Glucosidase

Pilot

alpha-Glucosidase,Ongi Toyobo

Pilot

alpha-Glucosidase;Ongin yeast Biozyme

Pilot

alpha-Glucosidase,Ongin yeast Fluka

Lab

alpha-Glucosidase,Ongin yeast overproducer Industnal

3.2.1.21

Beta-glucosidase. Gentobiase Cellobiase Amygdalase

-Seravac

Hydrolysis of temunal, non-reducing beta-D-glucose residues with release of beta-D-glucose Pilot

-

beta-Glucosidase,Ongin almonds Fluka

Pilot

beta-Glucosidase,Ongin sweet almond Toyobo

Pilot

-

beta-Glucosidase,Ongin sweet almonds Alpha-galactosidase.

3.2.1.22 Melibiose + H(2)O = galactose + glucose

Melibiase

alpha-Galactosidase;Ongin. rec microorganism Novozymes

Alpha-galTM

Industnal

3.2.1.23

Beta-galactosidase. Lactase.

Hydrolysis of terminal. non-reducing beta-D-galactose residues in beta-D-galactosides.

Lactase; Origin: Aspergillus oryzae Amano: Lactase 14-DS

Industrial

Lactase, Ongin Aspergillus oryzae Amano

Lactase F "Amano"

Industnal

beta-Galactosidase,Ongin E coli Fluka

Pilot I

I

beta-Galactosidase,Ongin E coli overproducer Roche Diagnostics beta-Galactosidase

Industnal

beta-Galactosidase,Ongin Eschenchia coli Toyobo

-_

Pilot

I

beta-Galactosidase,Ongin Kluviromyces lactis Recordati beta-Galactosidase, Lattasi beads

Industnal

1502

I

20 Tabular Survey of Commercially Available Enzymes Table 20.5.

(cont.).

beta-Galactosidase,Ongin Kluyveromyces fragilis Fluka

Pilot

Lactase, Ongin Kluyveromyces lactis Novozymes Lactozym@

Industnal

-

beta-Galactosidase, Ongin nucroorganisms Unitika beta-Galactosidase (beta-Gal)

Pilot

Hydrolyw of terminal, non-reducing alpha-D-mannose residues in alpha-D-mannosides _ _ I I _ X X _

_ X " l " X "

~"

- _ I _ ^ L

I

I

_

Y

~

alpha-Mannosldase

Invertase Saccharase

l

/

_

__

--

x

~

*I_

x

Hydrolysis of temunal non-reducing beta-D-fructofuranoside

-D-glucuronoside + H(2)O = an alcohol + D-glucuronate i__

x_

; i

-

;

_

Seravac

Pilot

beta-Glucuronidase: Ongin bovine liver Fluka

Lab

beta-Glucuronidase, Ongin E coli Fluka

Lab

-

beta-Glucuronidase, Ongin E coli K12 Fluka

__

_

Lab

__

beta-Glucuronidase, Ongin: Helix pomatla

Hyaluronidase

x

_rr

x_

"*

"__

~

Hyaluronidase Seravac Hyaluronidase: Ongin bovine testes Fluka Hyaluronidase, Ongin ovine or bovine teste5 Biozyme

^^

-

Random hydrolysis of 1.4-linkages between N-acetyl-beta-D-glucosamine and D-glucuronate residues in hyaluronate 1

-

I

_-

~

*

%

*

*

1

Pilot Lab Pilot

Hyaluronidase, Ongin sheep testes Fluka

Lab

Hyaluronidase, Ongin Streptomyces hyalurolyticus Fluka

Lab

Hyaluronidase, Ongin Streptomyces sp. Amano Amano 1 [HY-I]

Pilot

Hyaluronidase, Ongin Streptomyces sp Ammo Ammo 3 [HY-3]

Pilot

-

20 Tabular Survey ofCommercially Available Enzymes

I

1503

Table 20.5.

(cont.). w&8m

3.2.1.39

J

Glucan endo-l,3-beta-D-glucosidase. (I->3)-beta-glucan endohydrolase Endo-l,3-beta-glucanase hnannase

Hydrolysis of 1,3-beta-D-glucosidic linkages in 1,3-beta-D-glucans

Pullulanase hllulan 6-glucanobydrolase. Limit dextnnase Debranclung enzyme

Starch-debrancbmg enzyme, hydrolyses ( l-6)-alpha-glucosidic linkages in pullulan and starch to form maltotnose

s -s -p*_ -_x

-*

*--

a -e r * ia ---

Pullulanase, Ongin Bacillus sp. Amano Debranchngenzyme "Amano" 8 I

-

_ _ __

_ I

-

I

-

-

-

-

1-1

-

Industnal I

-

Pullulanase, Ongin Bacillus sp Fluka I

-

I

Industnal

_ _

I

"

__

I

Pullulanase.Ongin rec. microorganism

Beta-hexosanunidase Hexosanunidase N-acetyI-beta-glucosamimdase

Hydrolysis of tenrunal non-reducing N-acetyl-D-hexosanune residues in N-acetyl-beta-D-bexosmnides - - - -w m* -b-

Fluka

Lab

3.2.1.81

Agarase.

Hydrolysis of 1,3-beta-D-galactosidic linkages in agarose, giving the teuamer as the predominant product

Agarase, Ongin: Pseudomonas atlantica Fluka

Lab

3.2.3.1

Thioglucosidase.

A thioglucoside + H(2)O = a tbiol + sugar

Myrosinase Sinignnase Sinigrase

Myrosinase, Ongin Senapis alba (white mustard seed) Biocatalysts

Pilot

3.3.2.3

Epoxide hydrolase.

An epoxide + H(2)O = a glycol

Epoxide hydratase Arene-oxide hydratase

Epoxide Hydrolase;Origin: Agrobacterium sp Lab

Fluka

Epoxide Hydrolase;Origin: Aspergillus niger Lab

Fluka

Epoxide Hydrolase;Origin: Rhodococcus rhodochrous Lab

Flub

Epoxide Hydrolase, Ongin Rbcdotorula glutinis Fluka

Lab

Hydrolases. Acting on peptide bonds (peptide hydrolases).

--

m -p --*w --m m

Protease, Ongin Aspergillus melleus Amano. Protease DS

-m*yL

c^i-v_cx-.A~~~Iu--~-~*--

--

3.4%---____l__M(_n--

_--

lndustnal

Protease;Ongin. Aspergillus melleus Amano

Protease P "Amano" 6

Alms

Industnal

_

Industnal

Protease; Origin: Aspergillus niger Amano: Acid Protease A

Industrial

1504

I

20 Tabular Suwey ofCommercially Available Enzymes Table 20.5.

(cont.).

Protease, Origin Aspergillus niger Ammo Acid Protease DS

Industnal

Protease, Ongin Aspergillus niger Julich Enzyme Products

Lab

-

Protease , Ongin Aspergillus oryzae Altus

Industnal

Protease, Ongin Aspergillus oryzae Altus

Industnal

Protease, Ongin Aspergillus oryzae Altur

Industnal

Protease, Ongin Aspergillus oryzae Ammo Protease A "Amano" 2G

IndUStndl

Protease, Ongin Aspergillus oryzae Ammo Protease A-DS

Industnal

-

I

Protease, Ongin Aspergillus oryzae Ammo Protease M "Amano"

Industnal

Protease; Origin: Aspergillus oryzae Jiilich Enzyme Products

Lab

Proteinase 2A, Ongin Aspergillus oryzae Huka

Industnal

Protease, Ongin Aspergillus sp Altus

Industnal

Protease, neutral; Ongin Bacillus amyloliquefaciens Novozymes NeutraseB

Industnal

Protease, Ongin Bacillus lichenifomis Novozymes Bio-FeedB Pro

Indurtnal

Protease, Ongin Bacillus lichenifomis Novozymes NovozymB FM

Industnal

Protease, Endopeptidase ; Origin: Bacillus lichenifomis Novozymes: AlcalaseB

Industrial

Proteinase; Origin: Bacillus lichenifomis Fluka

Industrial

Protease, Ongin Bacillus sp Altus

Industnal

Protease, Ongin Bacillus sp Novozymes NovoCor S

Industnal

Protease, alkaline, Ongin Bacillus sp Novozymes: EsperaseB

Industnal

Proteinase, neutral; Origin: Bacillus sp. Toyobo

Industrial

Endoproteinase; Origin: Bacillus sp.. rec. Fluka

Pilot

Protease; Ongin Bacillus sp , rec Novozymes Novo-ProTMD

Industnal

Protease, Ongin Bacillus sp ,rec Novozymes PyraseB

Industnal

Protease; Origin: Bacillus stearothennophilus Amano: Protease S "Amano"

Industrial

Protease;Origin: Bacillus subtilis Amano : Proleather FG-F

Industrial

20 Tabular Survey ofCommercially Available Enzymes

I

1505

Table 20.5.

(cont.).

Protease;Origin: Bacillus subtilis Ammo: Protease N "Amano"

Industrial

Protease;Origin: Bacillus subtilis Ammo: Protease NL "Ammo"

Industrial

Protease; Origin: Bacillus subtilis Jiilich Enzyme Products

Lab

Proteinase; Origin: Bacillus subtilis Fluka

Industrial

Proteinase; Origin: Bacillus subtilis var. biotecus A Fluka

Industrial

Protease;Origin: Carica papaya L Ammo : Papain W-40

Industrial

Protease, Proline-Specific Endopeptidase; Origin: Flavobacterium sp. Toyobo

Pilot

Protease, neutral to acidic; Origin: Fungus Novozymes: NovoCor P

Industrial

Protease, alkaline; Origin: microorganism, rec. in Bacillus sp Novozymes: Savinase@

Industrial

Protease;Origin: microorganisms DSM Gist-brocades: Fermizyme

Industrial

Protease;Origin: Penicillum sp. Altus

Industrial

Protease; Origin: Protein engineered in rec. Bacillus Novozymes: Kannase'M

Industrial

Protease, alkaline; Origin: Protein engineered, rec. in Bacillus sp. Novozymes: Everlase@

Industrial

Protease; Origin: Rhizomucor miehei, rec. in Aspergillus oryzae DSM Gist-brocades: Optiren

Industrial

Protease, neutral to acidic; Origin: Rhizomucor sp. Novozymes: NovoCo~fBAB

Industrial

Protease; Origin: Rhizopus niveus Ammo : Acid Protease

Industrial

Protease; Origin: Rhizopus oryzae Ammo: Peptidase R

Industrial

Pronase,Ongin Streptomyces gnseus Fluka

Industnal

Fluka

lndustnal

Pronase nonspecific protease,Ongin. Streptomyces gnseus Roche Diagnostics Pronase nonspecific protease

Industnal

Protease, alkalophilic, Ongin Streptomyces sp Toyobo

Cytosol anunopeptidase Leucine anunopeptidase Peptidase S

Pilot

Release of an N-terminal amino acid, Xaa-I-Xbb-, in which Xaa is preferably Leu, but may be other amino acids including Pro although not Arg or Lys, and Xhb may be Pro

--

Leucine Aminopeptidase, cytosol , Ongin hog ladney Fluka Leucine arninopeptidase, Ongin pig ladney Biozyme

~

_

Y

X

1

I

~

---_---

x

"

~

L I

Lab Pilot

1506

I

20 Tabular Survey ofCommercially Available Enzymes

(cont.).

Table 20.5.

a

~~~~~~

Xaa-Pro dipeptidase.

3.4.13.9

X-Pro dipeptldase Proline dipeptidase Imdodipeptidase Prolidase

Hydrolysis of Xaa-I-Pro dipeptides, also acts on aminoacyl-hydroxyproline analogs No action on Pro-I-Pro

Prolidase,Ongin Lactococcus lactis Fluka

Lab

Prolidase, Ongin. pig ludney Biozyme

Pilot

Dipeptidyl-peptidase1.

3.4.14.1

Cathepsin C Cathepsin J Dipeptidyl mnopeptidase I Dipeptidyl transferase.

Release of an N-terminal dipeptide, Xaa-Xbb-I-Xcc, except when Xaa is Arg or Lys, or Xbb or Xcc is Pro

Cathepsin C, sol., Ongin bovine spleen Roche Diagnostics. Cathepsin C, sol

Industnal

Transferred entry: 3.4.1 6.5 and 3.4.16.6.

3.4.16.1

a

CarboxypeptidaseY; Origin: baker’s yeast Fluka

Lab

CarboxypeptidaseY, Ongin yeast

Roche Diagnostics Carboxypeptidase Y,Sequencing Grade

Lab

CarboxypeptidaseA.

3.4.17.1 Peptidyl-L-amino acid + H(2)O = peptide + L-amino acid

Carboxypolypeptidase

CarboxypeptidaseA Pilot

Seravac

CarboxypeptidaseA, Ongin bovine pancreas Fluka

Lab

Carboxypeptidase P Microsomal carboxypeptiddse

--

x

l _ _ _ _ v -

-

Release of a C-temnal residue other than proline, by preferentialcleavage of

x--

X

I

-

~

-

~

~

-

P

Pyroglutamyl-peptidase1.

_

_

.

I -

3.4.19.3 5-oxoprolyl-peptide + H(2)O = 5-oxoproline + peptide

5 oxoprolyl-peptidase Pyrrolidone-carboxylate peptidase Pyrrolidone carboxyl peptidase Pyroglutamyl aminopeptidase

Pyroglutamate Aminopeptidase, Ongin calf liver Fluka

Pilot

Hydrolases. Acting on peptide bonds (peptlde hydrolases). Serlne endopeptidases.

3.4.21.-

Endoproteinase Pro-C, Ongin. microorganism, rec in E coli Fluka

Lab

Chymotrypsin.

alpha-Chymotrypsln

3.4.21.1

-

Chymotrypsin A Chymotrypsin B Alpha-chymotrypsin yI--II--yx--~~-uI-

I ~ x _ x _

-

~

~

~

~”

I

Preferential cleavage Tyr-I-Xaa, Trp-I-Xaa, Phe-I-Xaa, Len-I-Xaa ”

__I

Seravac

--x*-i

~

_- ~

I(

li_

I-^*.,-

-

~-

nX”-we_

Pilot

alphaChymotrypsin. Ongin Bacillus Iicheniformis Alms

Industnal

alpha-Chymotrypsin; Origin: bovine pancreas Fluka

Pilot

20 Tabular Survey ofComrnercially Available Enzymes

I

1507

Table 20.5.

(cont.).

3.4.21.4 Preferentlal cleavage Arg-I-Xaa, Lys-I-Xaa. -x

-m -_ -'___)_-

x

_I*

Trypsin Seravac

Pilot

Trypsin Pilot

Seravac

Trypsin; Origin: bovine pancreas Industrial

Fluka

Trypsin; Origin: pig Pancreas Industrial

Biozyme

Trypsin ;Origin: porcine pancreas Alms

Industrial

Trypsin; Origin: porcine pancreas Biocatalysts: Trypsin

Industrial

Trypsin;Origin: porcine pancreas Biocatalysts: Trypsin 250

Industrial

Trypsin; Origin: porcine pancreas Biocatalysts: Trypsin 250

Industrial

Trypsin ; Origin: porcine pancreas Novozymes: Crystalline Porcine Trypsin

Industrial

Trypsin (Chrymotrypsinas minor constituent), Ongin porcine pancreas Novozymes ITN (Pancreatic Trypsin Novo)

Industnal

Thrombin.

3.4.21.5

Fibnnogenase I _r

~~~-~

Preferential cleavage Arg I-Gly, activates fibnnogen to fibnn andreledses fibnnopeptide A 1 - _I _ x _

1 _ 1

Thrombin, Ongin bovine plasma Fluka

Enteropeptidase. Enterokinase.

_y

~~~

-"

~

-

-

-

Pilot

3.4.21.9 Selectlve cleavage of 6-Lys-I-Ile-7bond in trypsinogen.

Enteropeptldase Seravac

Glutamyl endopeptidase. Staphylococcal senne proteinase V8 proteinase Protease V8 Endoproteinase Glu-C

Pilot

3.4.21.1 9 Preferential cleavage Asp-I-Xaa, Glu-I-Xaa

EndoproteaseGIu-C; Origin: Endophrins Alms

EndoproteinaseGIu-C, Ongin Staphylococcus aureus stram V8

Fluka

EndoproteinaseGIu-C, Ongin Staphylococcus aureus strain V8

Roche Diagnostics. Endoproteinase Glu-C, Sequencing Grade

Pancreatic elastase. Pancreatopeptidase E Pancreatic elastase I

Pilot Lab Lab

3.4.21.36 Hydrolysis of proteins, including elastin Preferential cleavage Ala-I-Xaa

Elastase;Origin: hog pancreas Fluka

Elastase;Origin: pig pancreas Biozvme

Pilot

1508

I

20 Tabular Survey of Commercially Available Enzymes Table 20.5.

(cont.).

Elastase, Ongin porcine pancreas Altus

Industnal

I Lysyl bond specific proteinase

Preferential cleavage L y d X a a , includin v_

~

Endoproteinase Lys-C, Ongin Lysobacter enzymogenes Fluka

~

Lab

Subtilisin.

3.4.21.62 Hydrolysis of proteins with brodd specificity for peptide bonds, and a preference for a large uncharged residue in P1 Hydrolyses peptide mdes Pilot Industnal Bacillus Iichenifonnir Industnal Industnal

Altus

PeptiCLEC-BL (slurry)

- _

Industnal _ I

Subtilisin, Ongin Bacillus lichenifomis Fluka

Industnal

Subtilisin, Ongin Bacillus lichenifomis Roche Diagnostics CHIRAZYME P-1, lyo

Industnal

Subtilisin, Ongin Bacillus lichenifomis Roche Diagnostics CHIRAZYME P-1, sol

In dustn aI

Subtilisin Carlsberg , Ongin Bacillus licheniformis Altus

Industnal

Proteinase K. Endopeptidase K Tntirachium alkaline proteinase Tntirachium album proteinase K

3.4.21.64 Hydrolysis of keratin and of other proteins, with subtilisin-like specificity Hydrolyses peptides amdes

Proteinase N, Ongin Bacillus subtilis Ruka

Industnal

Proteinase K, Ongin Tntirachium album Fluka

Industnal

Proteinase K, immobilized on EupergitC,Ongin Tntirachium album Fluka Proteinase K, lyo., Ongin Tntirachium album Roche Diagnostics Proteinase K, lyo.

Papaya peptidase 1

Papain Biocatalysts

Lab Industnal

Hydrolysis of proteins with broad specificity for peptide bonds, with preference for a residue beanng a large hydrophobic sidechain at the

PROMOD 144L

lndustnal

Papain, Ongin Canca papaya Flukd

Industnal

20 Tabular Survey ofComrnercially Available Enzymes

I

1509

(cont.).

Table 20.5.

Papain, Ongin Canca papaya Roche Diagnostics Papam

Industnal

Papain, immobilizedon Eupergim C, Ongin Canca papaya Fluka

Lab

_--

" C

-

Specificity sinular to that of papam

-

x

_I___

1

>_clwIc--c-

x-

Industnal

Clostndiopeptidase B ~

~

~

Preferentlal cleavage Arg-I-Xaa, including Arg-I-Pro bond, hut not

-xIxxL -

~ ~

a

Clostripain, Ongin Alms

Industnal

Clostripain, Ongin Clostndium histolyticum Fluka

Pilot

_ _

Endoproteinase Arg-C, Ongin Clostndium histolyticum se Arg-C, Sequencing Grade

Fluka

_ _

Lab

-

Pilot

-

-

Endoproteinase Arg-C, Ongin rmce (suhmaxillary glands)

Bromelain m___~

_1

-

~

x

x

I

c

x-_-

~

x

_XX^

~

_r

xxx

_c

x

-

Broad specificity for cleavage of proteins, but strong preference for Z-Arg-Arg-I-NHMec amongst small molecule substrates

-

x

_ " ~ _ _ I _ ~

1

x

-_

x___x

~

~

-_~ - - -~ x1_

x_Ix

Bromelain,Ongin. pinapple stem Alms Bromelain,Ongin pineapple stem

Fluka

Pilot

Pepsin A.

3.4.23.1

Pepsin

^ _ I

yII

-- -

y

f

C

I

__

Preferentlal cleavage hydrophobic, preferably aromatic, residues in PI and PI' positions Cleaves I-Phe-I-Val-2, 4-Gln-I-His-5, 13-Glu-I-Ala-14, 1CAla-I-Leu-15, 15-Leu-I-Tyr-16, I

x

x

_I"

z

x_

c

I _

Pepsin, Ongin hog stomach Fluka

I -

-~~~ ~~_ ~ ) _ _ _

~

Industnal

Pepsin, Ongin pig Ftomach mucosa Biozyme

Pilot

Pepsin ,Ongin porcine pancreas Industnal

Altus

3.4.23.4

Chymosin. Rennin

-

-xl_*%".

1

x

--

Broad specificity similar to that of pepsin A Clots nulk by cleavageof a single bond in casein (kappa cham) ^"

Chyrnosin,Ongin calf stomach Altus

-x

X

I

x - x

"&*

-

_ I - _ ) _ j

Y

_"L

Industnal

1510

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

Microbial collagenase.

3.4.24.3

Clostndium histolytlcum collagenase Clostndiopeptidase A Collagenaye A Collagenase I

Digestion of native collagen in the tnple helical region at Xaa-I-Gly bonds With synthetlc pepudes, a preference is shown for Gly at P3 and PI', Pro and Ala at P2 and P2': and hvdroxvmoline. Ala or Are

Collagenase; Origin: Clostridium histolyticum Fluka

Pilot

Collagenase; Origin: Clostridium histolyticum Fluka

Pilot

Collagenase: Origin: Clostridium sp. Amano: Ammo 1 [CL-I]

Pilot

Collagenase, Ongin Clostndium sp Ammo Ammo s [CL-S]

Pilot

Thermolysin; Ongin Bacillus thermoproteolyticus Altus

PeptCLEC-TR (dry)

Industnal

Thermolysin; Origin: Bacillus thermoproteolyticus Industrial

AItus: PeptiCLEC-TR (slurry)

Thermolysin; Origin: Bacillus thermoproteolyticus Fluka

Pilot

3.4.24.33

Peptidyl-Asp rnetalloendopeptidase. Endoproteinase Asp-N

Cleavage of Xaa-I-Asp, Xaa-I-Glu and Xaa-I-cysteic acid bonds.

Endoproteinase Asp-N: Ongin Pseudomonas fragi (mutant) Fluka

Lab

-

EndoproteinaseAspN, Ongin Pseudomonas fragi (mutant) Roche Diagnostics Endoproteinase Asp-N, Sequencing Grade

Lab

Acting on carbon-nitrogen bonds, other than

Acting on carbon-nitrogen bonds, other than peptide bonds.

Fluka

Industnal

Glutaryl acylase, carrier-fixed,Ongin E coli overproducer Industrial

Roche Diagnostics Glutanl acvlase, camer-fixed (GI-Ac)

Glutaryl-7-ACA Acylase, Ongin microorganlsm, rec in E coh Recordati GAA Beads

Industrial &S

L-asparagine + H(2)O = L-a5partate " _ L 1

1

""

_ I - _~~

~

I -x

L-Asparaginase, Ongin E coli Fluka

Lab

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

3.5.1.2

Glutaminase.

L-glutanune + H(2)O = L-glutamate + NH(3)

--~~-~ rxarw--*anr

_/_Lx”

Acylanudase Acylase

-

---_ Amidase, Ongin Pseudomonas aeruginosa, rec in E coli l ~ l l ~ ~ L _l iw_-i---m

_--_I_

*-m-m-

)I-

- - P l W * - - * - s a _

A monocarboxylic acid amde + H(2)O = a monocarboxylate t NH(3) a

_xn--m--m

_

-

/

_

_

_

_

l

_

l

~

-

~

~

_

_

l

-

i

-

_

I

-

~

_

~

~

~

~

~

~

~

Lab

nukd

~

-

-

~

.

~

~

~

~

~

~

~

-

~

p --a---w **

Urea + H(2)O = CO(2) t 2 NH(3)

”

-

~

-

-

-

~

~

~

~

Urease, Ongin jack bean Industnal

Fluka -*--w--

1

-----*-

I s l a 1 7 -

Urease, Ongin jack bean

-

-

Roche Diagnostics Urease

Industrial

Toyobo __I_x_ ~ ~

Pilot

-

I

- _-

___xl^

1 1 1

I I

I

-

I

I_ I I

Urease, immobilized on Eupergit C, Ongin jack bean

-

Industnal

--

Fluka

Pilot _ _ I _

Lab

3.5.1.1 1

Penicillin amidase. Perucillin acylase

Penicillin + H(2)O = a fatty acid anion + 6-mnopenicillanate

----

-m -m --p --

-*-*

----*-me---*

* -

Penicillin Acylase, Ongin E coli Alms

_-

XI_-

- --

I

_ I

-

I _

-

- _ - --

I

Industnal

__

I

-_-__ -

I_I ^

Penicillin Acylase, Ongin E coh

Altus ChoCLEC-EC (dry)

__

_I___I_

_I

Industnal

___- -

I^___

_ - I -

I

I _

1_

_I-

I _

~~

_-_

Penicillin Acylase, Ongin E coli Altus

ChoCLEC-EC (slurry)

__

I

_ _ _ I _ _ _ _ _

I

_

_

_

I

_

--___

_I_--

I__

I

l

I

_

l

I

I

-_

Industnal

--

Penicillin Amidase, Ongin E coli FlUka

-----

Penicillin Amidase, immobilized on Eupergim C, Fluka _ I _

---

-

I

I

I-

E coli

___

~

-

I

-

Industnal -___ ___ - - -_ Industnal

__

I

-

_

-

I

_

1

1

1 1 1 1

1

Penicillin G Amidase, immobilized, Ongin E coli FlUka l

_

l

_

l

-_

I

I

I

__

-

Penicillin-G Acylase, Ongin E coli Recordati PGA beads, Standard

-

--^-

-

I_

_ I

--

_

I _

Penicillin-G Acylase, Ongin E coli Recordati

PGA beads, Superenzyme

_I__I l_l__l-I

-

_ -I

_lll

Penicillin G Amidase, Ongin E coli overproducer

-

--

_ I

I

1-1-

I

I I

_ ^ -

lndustnal

-

_ I

I

_

Industnal

-

ll_l

_-

-

Industnal _

l

l

l

Industnal

3.5.1.14

Aminoacylase. Histozyme Hippuncase Benzanudase Dehydropeptidase I1

-

An N-acyl-L-anuno acid + H(2)O = a fatty acid amon + an L-anuno acid

-*

* “ , - > _ _ _ _ ( _ J x xm I^__c~~--mn_l * ___I I__-a--v % -

Acylase I, Ongin Aspergillus melleus Fluka

Industnal

1512

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.).

Acylase I, immobilized on Eupergit C, Ongin Aspergillus sp Fluka

Pilot

Ammo. Acylase

Industnal

Acylase I; Ongin hog ladney Fluka

Pilot

Acylase I, Ongin. pig kidney Biozyme

PllOt

Acylase, Ongin Streptomyces chartreusis Fluka

Lab

Acylase, Ongin Streptomyces gnseocarneus Fluka

Lab Streptomyces hachijoensis

Fluka

Lab

Acylase, Ongin Streptomyces toyocaensis Fluka

Lab

Acylase, Ongin Streptomyces zanmyceticus Fluka

Lab

Recordati

Hyda-REC

Industnal

D-Hydantoinase, Ongin Azuk beans Fluka

Lab

D-Hydantoinase 1, carrier-fixed,0 Bacillus thermoglucosidasius, rec in E coh Roche Diagnostics D-Hydantoinase 1, camer fixed

-

lndustnal

D-Hydantoinase, recombinant, immobilized,Ongin E coli Beta-lactamase. Penicillinase Ceohalosoonnase

3.5.2.6 A beta-lactam + H(2\0 = a substituted beta-ammo acid

beta-Lactamase I: Origin: Bacillus cereus Lab

Fluka

beta-Lactamase, Ongin Enternbacter cloacae Fluka

Lab

Creatininase. Creatinine armdohydrolase

3.5.2.10 Creatinine + H(2)O = creatine

Creatininase, Ongin microorganisms Pilot

Asah

Arginase.

-~ -

Arginine amdinase Canavanase ”

3.5.3.1 L-arginine + H(2)O = L-ornithine + urea

Arginase

Biozyme Bovine liver

PllOt

L-Arginase, Ongin bovine liver Huka

Lab

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

I

1513

(cont.).

P

Creatinase.

3.5.3.3 Creatine + H(2)O = sarcosine + urea

Creatine mdinohydrolase

Creatinase, Ongin. Bacillus sp Asahi

Pilot

Creatinase, Ongin Flavobacterium sp Fluka

Pilot

Hydrolases. Acting on carbon-nitrogen bonds, other than peptide bonds. In cyclic amidines.

3.5.4.-

Deaminase;Ongin. Aspergillus melleus Adenosine deaminase.

3.5.4.4 Adenosine + H(2)O = inosine + NH(3)

Adenosine armnohydrolase.

Adenosine Deaminase;Ongin: calf intestinal mucosa Fluka

Pilot

____

I

Adenosine Deaminase, Ongin calf intestine Roche Diagnostics Adenosine Deanunase

Nitrilase.

Industnal

3.5.5.1 A mtnle + H(2)O = a carboxylate + NH(3)

Nitrilase, broad-range;Origin: microorganism, rec. in E. coli BioCatalytics: NIT-I01

Lab

Nitrilase, broad-range;Origin: microorganism, rec. in E. coli BioCatalytics: NIT-I02

Lab

Nitrilase, broad-range, Ongin nucroorgamsm, rec in E coli BiOCatdlytiCS' NIT- 103

Lab

Inorganic pyrophosphatase. Inorganic diphospbahse. Pyropbosphate phosphohydrolase Diphosphate phosphobydrolase ~

x

I

I

I

I

~

x

I

I

a

w

x

~

~

x

I

I

3.6.1.1 Diphosphate + H(2)O = 2 phosphate

x

- i - _ " ~ _ _

Pyrophosphatase,inorganic, Ongin bake Fluka

Lab

Pyrophosphatase,inorganic. Ongin E coli Fluka

Alpha-carboxylase Pyruvic decarboxylase Alpha-ketoacid carboxylase

A 2-0x0 acid = an aldehyde + CO(2)

Pyruvate Decarboxylase,Ongin Zymomonas mobils, E coli (rec ) Julich Enzyme Products

Oxaloacetate decarboxylase. Oxdlate beta-decarboxylase

4.1.1.3 Oxaloacetate = pymvate + CO(2)

Oxalacetate Decarboxylase;Origin: Pseudomonas sp. Fluka

Lab

Oxaloacetate Decarboxylase;Origin: Pseudomonas sp. Asahi

Pilot

1514

I

20 Tabular Survey ofCommercially Available Enzymes Table 20.5.

(cont.)

Acetolactate decarboxylase.

4.1.1.5 *-F*s7--M-

Acetolactate Decarboxylase, Ongin Bac Brevis in rec, Bac subtilis Novozymes

MaturexB

Industnal

Phosphate + oxaloacetate = H(2)O + phosphoen

Phosphoenolpyruvate carboxylase, Ongin maze leaves Biozvme

Pilot

Phosphoenolpyruvate Carboxylase, Ongin maize leaves Fluka

Lab

- -__

Phosphoenolpyruvate Carboxylase, Ongin rmcroorganisms Toyobo

Pilot alanine = phenethylamine +

-_wm-w-I-x

%"-_

--%cw-

~ _ a _ ( _ _ _ - ~ h _ _

I

*

-

-L-

L-Phenylalanine Decarboxylase, Ongin: Streptoco Fluka

Lab

Methionine decarboxylase.

4.1.1.57 L-methionine = 3-methylthiopropanane+ CO(2)

L-Methlonine decarboxylase; Ongin Streptomyces sp Fluka

Lab

Phosphodeoxynboaldolase Deoxynboaldolase

2-deoxy-D-nbose 5-phosphate = D-glyceraldehyde 3-phosphate +acetaldehyde *-----

" L 1 a *

~

__

Fluka I

I

~

_-

--^_II/x_

"

- ---

L-threonine = glycine + ace *-

*-*

I

Lab

-

Threonin Aldolase; Ongin Pseudomonas putida Fluka

Lab

Mandelonitrile lyase. Hydroxynitnle lyase. (R)-oxynitnlase

4.1.2.10

_-

% -

(I?)-Oxynitrilase, Ongin bitter almonds ( h n u s amygdalus) Julich Enzvme Products

Mandelonitnle = cyanide + benzaldehyde

~--

_-I

^

X

-,--

x

I ah

20 Tabular Survey ofCommercially Available Enzymes 11515 Table 20.5.

(cont ).

R-Oxynitrilase rec., Ongin Pichia pastons FlUka

Pilot

Hydroxymandelonitrile lyase.

4.1.2.1 1

(S)-4-hydroxymandelonitnle = cyanide + 4-hydroxyhenzaldehyde

Hydroxynitnle lyase

(S)-Oxynitrilase, Ongin Sorghum bicolor or S vulgare Juhch Enzyme Products

Lab

D-fructose 1,6-bisphosphate = glycerone phosphate + D-glyceraldehyde 3-phosphate

Aldolase. Fructoye-1,6-bisphosphate tnosephosphate-lyase

----

--

--*----+*-

Fluka

Lab

Aldolase; Origin: rabbit muscle Fluka

Lab

Aldolase; Origin: rahhit muscle Roche Diagnostics: Aldolase

Lab

Aldolase; Origin: Staphylococcus aureus Flub

Lab

Aldolase ;Origin: Staphylococcus carnosus Fluka

Lab

Fructose 1,6-bisphosphateAldolase; Origin: Staphylococcus carnosus Jiilich Enzyme Products

Lab

Aldolase, Ongin Thermus aquaucus Fluka

Lab

6-phospho-2-dehydro-3-deoxygalactonate aldolase 6-phospho-2-keto-3-deoxygalactonate aldolase 2-0x0-3-deoxygalactonate6-phosphate aldolase. ------w.-vp*"w<"*-

- x

2-dehydro-3-deoxy-D-galactonate 6-phosphate = pyruvate +D-glyceraldehyde 3-phosphate

__ta-~-

~

6-Phospho-2-dehydro-3-deoxygalactonatealdolase

Dihydroneopterin aldolase.

II-I--Iyx

-- - - - ~ ~ - - - ~ ~ - ~ v

L I - ^ " I I I - ~ ~ x - x - - x L - ~

4.1.2.25 2-amino-4-hydroxy-6-(D-erythro-1,2,3-tnhydroxypropyl)-7,8-dihydr optendine = 2-anuno-4-hydroxy-6-hydroxymethyl-7,8-dihydroptendine +

DihydroneopterinAldolase Fluka

Lab

Industnal

N-Acetyl-neuraminicacid aldolase, Ongin E coli Fluka

Lab

NeuraminicAcid Aldolase, Ongin E coli K12

Julich Enzyme Products

-

Lab

N-AcetylneuraminicAcid Aldolase, Ongin nucroorganisms Toyobo

Pilot

1516

I

20 Tabular Survey ofCammercially Available Enzymes Table 20.5.

(cont.).

N-AcetylneuraminicAcid Aldolase, Ongin microorganisms Unitika N-Acetylneuranunic Acid Aldolase (Nana-Ald)

Pilot

Citntase Citndesmolase

Citrate Lyase, Ongin Enterohacter aerogenes Fluka

2-keto-4-hydroxyglutaratealdolase 2-0x0-4-hydroxyglutaate

Lab

4-hydroxy-2-oxoglutarate= pyruvate + glyoxylate

4-Hydroxy-2-oxoglutarate aldolase ,Ongin E coli Fluka

4-hydroxy-4-methyl-2-oxoglutarate aldolase.

Lab

4.1.3.17 4-hydroxy-4-methyl-2-oxoglutarate = 2 pyruvate

L tryptophan + H(2)O -

Tryptophanase, Ongin microorganisms PI101

Pilot

Carbonate dehydratase. Carbonic dehydratase Carbonic anhydrase I

X

I

-

4.2.1.1 H(2)C0(3) = CO(2) + H(2)O

-

Carbonic anhydrase, Ongin Bhvine erythocytes Biozyme

Pilot

Carbonic Anhydrase,Ongin bovine erythrocyte? Fluka

Lab

Carbonic Anhydrase lsozyme II, Ongin bovine erythrocytes Fluka Chondroitin ABC lyase. Chondroitinase Chondroitin ABC elinunase

4.2.2.4 Elimlndtive degradation of polysacchandes contaming 1,4-beta-D-hexosamnyI and I ,3-heta-D-glucuronosyl or 1,3-alphd-L-iduronosyIlinkages lo disacchandes containing

droitinase ABC; Ongin Proteus vulgans

Pectin lyase.

Phenylalanine ammonia-lyase.

PhenylalanineDeaminase,Ongin Rhodotomla glutinis Fluka

4.2.2.10 hliminative deavage of pectin to give oligosaLchandes with termindl 4-deoxy-6-methyl-alpha-D-galact-4-enuronosyl groups

4.3.1.5 L-phenylalanine = trans-cinnamate + NH(3) Lab

I

20 Tabular Survey ofCommercially Available Enzymes

Table 20.5.

1517

(cont.).

4.4.1.9

L-3-cyanoalanine synthase. "I_ x

_1

~

x

c

L-cysteine + cyanide = H(2)S + L-3-cyanoalanine

-

_*

~

A

Ongin Bacillus stearothennophilus Unitika Alanine Racemase (AlaR)

c

Ammo

Y

" 1

~~

Amano 2 [MUT-21

Pilot

Mutarotase,Ongin pig kidney

Triosephosphateisomerase, Ongin rabbit muscle Biozyme

Pilot

Xylose isomerase.

- -_

D-xylose = D-xylulose x

%

I

~

5.3.1.9

Glucose-6-phosphate isomerase. Phosphoglucose isomerase Phosphohexo5e isomerase Phosphohexomutase Oxoisomerase

_-_

___(

_ - C I -

* * *I

D-glucose 6-phosphate = D-fructose 6-phosphate

i

PhosphoglucoseIsomerase,Ongi Unitlka Phosphoglucose Isomerase (PGI)

Pilot

_ _

PhosphoglucoseIsomerase. Ongin baker's yeast

S-S rearrangase

Rearrangement of both intrachain and interchain diwlfide bonds r

- _

I

~

_%

_"

er

Fluka

Lab I_

Protein disulfide isomerase,Ongin E coh

- ~ - _ _- _ ucomutase. Ongin

sphoglucose mutase

I

- q

_c

rabbit muscle

Acyl-activating enzyme Acyl-CoA synthetase Fatty acid

ATP + a long-chain carboxylic acid + CoA = AMP + diphosphate +

Acyl-CoA Synthetase,Ongin microorganisms Asahi

Pilot

1518

I

20 Tabular Survey ofComrnercially Available Enzymes Table 20.5.

(cont.).

Acyl-CoA Synthetase,Ongin Pseudomonas sp Ammo Ammo 2 [ACS-2]

Pilot

Acyl-CoA Synthetase,Ongin Pseudomonas sp Ammo Ammo3 IACS-31

Pilot

Acyl-coenzyme A Synthetase, Ongin Pseudomonas sp Fluka

Glutamate--ammonia ligase.

Lab

6.3.1.2 ATP + L-glutamate + NH(3) = ADP + phosphate + L-glutamme

Glutamme synthetase

Glutamine Synthetase, Ongin Bacillus stearothermophilus Urutka Glutamme Synthetase (GS)

Lab

NAD(+) synthetase

ATP + deamido-NAD(+) + NH(3) = AMP + diphosphate +

NAD Synthetase Asahi

Pilot

Urease (An-hydrolysing) Urea carboxylase (hydrolysing) ATP--urea amidolyase Urea anudo-lyas 1Ix-

-*-*m_-l

ATP + urea + CO(2) = ADP + phosphate + urea-1-carboxylate

--*--

Urea Amidolyase,Ongin. yeast Toyobo

Pilot

Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyriqht 0Wiley-VCH Verlaq GmbH, Weinheim 2002

Index a

AADH 1048 - carbinolamine 1052 - catalytic mechanism 1051 - protonated imine 1051 AAT - directed evolution 877 - protein engineering 877 - site-directed mutagenesis 877 abscisinic acid 1385 ABTS 1113,1132,1175 ACA acylase 1058 7-ACA 1436 ACE inhibitors 1056 - enalapril 874 acetaldehyde 343,478, 571, 574 acetate kinase 614,615, 902, 904,905,1246, 1319 acetates 397,458 acetic anhydride 473,545 acetoacetates 544 acetoin reductase 1129 acetone 342,352,478,1002 acetone cyanohydrin 544,565 acetone powder 1002 acetonitrile 342, 545 acetophenone 995,1019 acetoxy ketone, 2- 565 acetoxyethers 458 60-acetoxyeudesmanone 1075 acetoxyphosphonates 458 acetoxysulfides 458, 565 acetyl esterase 1378,1383 acetyl kinase 904 acetyl phosphate 614,902,904 N-acetyl-D,r-aminoacids 1441 - racemase 1308 acetylamino glutarate 365 acetylase 1483 acetylcholine acetyl hydrolases 407 acetylcholine esterase 406,407

acetyl-CoA 963,1247 acetylene carboxylatehydratase - malonic semialdehyde 690 - propynoicacid 690 acetylene dicarboxylatehydratase - pyruvate 690 N-acetyl-D-glucosamine1324,1452 - 2-epimerase 1324 acetylhexosamidinase 636 acetyllactosamine 615 N-acetyl-D-mannosamine 1324,1452 9-0-acetylneuraminate 945 N-acetylneuraminate - aldonase 944 - lyase 1324 - synthesis from N-acetyl-D-glucosamineand pyruvate 1324 N-acetylneuraminic acid 194,1452 - aldolase 194 acetylphosphate 615 achiral 352 - diol 344 achromobacter 784 - xylosoxidans 744 acid phosphatase 918,919,921 acid-base conditions 281 acidophilic xylose 1316 acidovorax facilis 707 acids 544 acinetobacter - calcoaceticus 1207, 1220, 1226, 1229, 1231 - calcoaceticus NCIMB 9871 1214,1220,1223 - calcoaceticus NCIMB 9871/TD 63 1225 - sp. 704 - sp. TD 63 1222,1223,1226 AcK 907 aconitate hydratase - cis-aconitate 688 - citrate 688 - isocitricacid 688 AcP 907

1520

I

Index

acrylamide 173,1454 - production 712 acrylonitrile 1454 actinomycetes 594 activation 351 - energy 192 active site 146, 148, 337, 338, 346, 413 - model 352,398,407 - residues 146 activity 342, 473, 571 - of immobilized enzymes 177 acyclic 427 - diacetates 417 - dicarboxylic acid esters 399 - ketones 996 - monoesters 369 acyl - activating enzyme 1517 - donor 473,544 - transfer 820 acylamino acid racemase 1306 - gene cloning 1308 - microbial distribution 1307 - properties 1307 - substrate specificity 1307 acylases 351, 741, 746,747, 752, 753,757, 758,1441 - ACA acylase 1058 - acylase1 1511 - glutaryl acylase 1510 - penicillin acylase 180, 855, 1336, 1337, 1341,1403,1405, 1407, 1444,1511 acylated - alcohols 545 - hemiacetal 558 acylations 336, 342, 346, 348, 416, 473, 474, 478,486,525, 570-572 - enantiosekctive 1435 - reagents 343 acyl-CoA synthetase 1517 acyknzyme 337, 338, 343, 398 - product 337 acyloin condensation 962 acyloxy groups 343 adaptive walks 116 additives l l G , 417, 556, 569, 570 additivity 109, 118 adenosine - aminohydrolase 1513 - deaminase 1513 S-adenosyl-L-homocysteine1290 - hydrolase 1290 S-adenosyl-L-methionine 1290 adenovirus endopeptidase 808

adenylate kinase 615,906, 907 adipodiamide 1450 adiponitrile 1454 adsorption 170 aerobacter 784 affinitychromatography 59 aging 1001 agrarases 1503 agrobacterium 774 - radiobacter 593, 777, 780, 781, 783 - SP. 774,779-782 - tumefaciens 705, 774 AIDS 814 ajinomoto 1442, 1448 alanine 1441 - as a component of cyclosponn A 1283 - production form fumaate 1298 - scanning 106 alanine dehydrogenase 1286 - AlaDH 1049 - characterization 1053 - isolation 1053 - product synthesis 1053 alanine racemase 1283,1517 - from yeast schizosaccharomyces pombe 1283 - genecloning 1283 - isozyme 1283 - reaction mechanism 1284 - stability 1284 - thermolabile enzyme from a psychrophile 1284 - thermostable enzyme from a thermophile 1284 alcalase 1346, 1379 alcaligenes 755 - faecalis 299, 744 - SP. 752,754 - sp. hpase 443,458, 526 - xylosoxidans 788, 789 alcohol dehydrogenases 81,997, 1006, 1009, 1010,1014,1017,1018, 1029,1038,1231 - yeast 996 alcohols 288, 342, 351, 352, 370, 383, 416, 442,458,473,486,525,526, 544, 545,1120, 1122, 1127 alcoholysis 336,342, 343, 348, 416,459,473, 486, 545, 554, 555,558,565 aldehyde dismutase activity 1196 aldehyde reductase 1010 aldehydes 1194 aldofuranose 1317 aldol reactions 931 aldolases 194, 618, 931, 946, 948, 950,953, 954,958,1321,1515

-

N-acetylneuraminic acid aldolase 944,1515

2-dehydro-3-deoxyphosphogalactonate aldo-

lase 1515 deoxyribose-phosphate aldolase 1514 - dihydroneopterin aldolase 1515 - FDP aldolase 931-933,936,938,953,961 - fructose-biphosphate aldolase 1317, 1515 - fucose 1-P aldolase 939 - 4-hydroxy-4-methyl-2-oxoglutarate aldolase 1516 - 4-hydroxy-2-oxoglutarate aldolase 1516 - KDG aldolase 950 - KDO aldolase 946 - KDPG aldolase 949 - KHG aldolase 948 - rhamnose aldolase 939 - TDP aldolase 942 - threonine aldolase 953, 1514 aldose 1-epimerase 1517 algorithms 139 alignment 141 - score significance 144 aliphatic - alcohols 1145 - ketones 1019 alitame 873 alkaline phosphatases 897,908,918-921 alkaline protease 1346 alkenes 1085,1088,1089 alkoxycarbonyl group 458 alkyl-alkyl 525 alkyl-aryl 525 akyl-enzyme 586 alkylphenols 1173, 1189 allantoinases 764766, 793 allenylic alcohols 525 2-allo-hydroxy-~-proline - substrate of proline racemase 1301 D-allose - from D-fructose 1322 allyl alcohols 1004, 1142 allyl bromide 1004 allylic 525 - alcohols 458, 565, 1145 - hydroxylation 1081 - isomerization by 1,3-shift 1282 alzheimer’sdisease 744 amidases 729,735,719,720,730-732,763, 765, 1336, 1436, 1511 - penicillin amidase 1438, 1453,1511 - peptide amidase 197,1510 amidation - ofcarboxylic 716 amidinohydrolases 1512, 1513 -

amidohydolases 742,749,751,752,777,782, 786,1303 - r-asparagin amidohydrolase 1510 - r-glutamine amidohydrolase 1511 amines 569,1035,1037 - kinetic resolution 1435 amino acids - a-amino acid 370,442, 1047, 1048 - - amide 1439 - D-aminoacid 1303 - - N-acetyl-D,L-aminoacid 1441 - - aminotransferase 1287, 1295 - - donor 890 - -ester 405 - - labeled with 13N by means of I3N-NH3 1288 - - oxidase trigonopsis variabilis 1426 - - production of 1286 - - synthesis from 5-substituted D,L-hydantoins through N-carbamoyl-L-aminoacids 1303 - - synthesis with D-amino acid aminotransferase 1296 - - synthesis with formate dehydrogenase 1296 - - synthesis with glutamate dehydrogenase 1296 - - synthesis with glutamate racemase 1295, 1296 - - transferases 889 - L-amino acid 873 - - 6-(L-aminoadipoyl)-L-cysteinyl-D-valine 1308 - -derivative 405 - -ester 412 - - homophenylalanine 874 - - isotopically labeled 874 - - non-naturally occuring - - D-penicillamine 874 - - L-phosphinotrhicin 874 - dehydrogenases 1047 - derivatives 558 - epimerase - - cofactor-independent 1293 - esters 398,412 - - ester derivatives 405 - racemase - - cofactor-independent 1293 - - pyridoxal S’-phosphate-dependent 1283 - - with low substrate specificity 1289 - receptor 1007 - sequence similarity 584 - spimerase - - pyridoxal S’-phosphate-dependent 1283

1522

I

Index amino alcohols 397,442, 1119 a-amino butyric acid 1441 a-amino-c-caprolactam 1442 - racemase - - genecloning 1292 - - isolation and purification form achromobacter obae 1292 - - properties 1292 - - reaction mechanism 1292 2-4-amino-4-carboxybutyl-2-aziridine carboxylic acid - inactivator of diaminopimelate epimerase 1299

7-amino deacetoxy cephalosporanic acid (ZADCA) 1438 6-amino penicillanic acid (6-APA) 1438 amino-2-thiazoline-4-carboxylate - hydrolase - - chemical synthesis of D,r-cysteine 1302 - intermediate in chemical synthesis of D,Lcysteine 1301 - racemase - - bacterial distribution 1301 a-amino-6-valerolactam - substrate of a-amino- c-caprolactam racemase 1292 aminoacylases 194, 744, 747, 754, 755, 756 - aminoacylases 746 - D-aminoacylase 1306 - L-aminoacylase - - of pig ludney 1306 - - substrates 1306 - - thermostable enyzme 1306 - spergillus olyzae 1441 aminoacyl-tRNA 822,842 4-aminobutyrate-2-ketoglutarate transaminase - broad-spectrum herbicide Basta 881 - 7-~-phosphinothricin881 aminocephalospoanic acid 1256,1426,1436 aminocylase 1511 R-1-aminoethylphosphonicacid 1285 3-aminoglutarates 365 aminolysis 599 aminopeptidases 720, 723,802,809, 1506 - aminopeptidase N 1399 - pseudomonas putida 1439 1-amino-D-sorbitol 1454 6-amino-1-sorbitol 1425 6-amino-~-sorbose1425 aminotransferases 874,878, 1484 - alanine aminotransferase 1484 - amino acid transferase 1287, 1295 - c-aminotransferase - - cephalosporin biosynthesis 882

- - glutamate oxidase 882 - aspartate aminotransferase 1484 - glutamic-pyruvictransaminase 875 - leucine aminotransferase 888 - mechanism pyridoxal phosphate 875 - mechanism pyridoxamine phosphate intermediate 875 - mechanism Schiff-baselinkage 875 8-aminovalerate 1301 ammonia lyases 866 ammoniolysis 717 - of carboxylic esters 716 amoxycillin 1441 ampicillin 1441 amygdalase 1501 amylases 1497 - a-amylase 315,654, 655, 1433, 1497 - P-amylase 656, 1498 - glucoamylase 315,656, 1498 amyloglucosidases 1433, 1498 amylopectin 315,654 amyloses 315,654 anaerobic 1130 andida antarctica 413 androstane 1390 androst-2-en-3,17-dione 1160 angiotensin 801,856 anhydrides 554,555 anhydrous organic solvents 412 anilines 1186 animal enzymes 46,48 anisotropic 102 antagonist 1007 anthracene 1100 antibiotics 729, 730, 735 antibody - anti-0-endorphin antibody 1400 anticholesterol drugs 1423 Ap4 hydrolases 923 APG4 1002,1022 apoptosis 813

applied molecular evolution 96 aqueous media 259 arabidopsis thaliana 744 a-L-arabinofuranosidases671 arabinogalactan 674 D-arabinOSeS 1323 archaea 313 arginases 46, 1512 arginine 871 - amidinase 1512 - racemase 1290 aromatic acids 1194 aromatic cyanohydrin acetates 544

arthrobacter 1158 - aurescens 778, 785, 786, 788, 789, 791 - crystallopoietes 774, 778 - SP. 725,791 - sp.lipase 459 artificial - redox coenzymes 1163 - substrate 113 arylallylethers 1087 2-aryloxy propionic acid 571 Asahi Chemicals 1436, 1438 L-ascorbic acid 1427 L-asparaginases 1510 aspartame 831,834,838,852,856,866,859, 873,1337,1439 - precursor 1446 aspartases 866,867, 1325, 1453 - aspartame 866 - for synthesis of D-alanine 1298 - fumaric acid 866, 867 L-aspartase 1454 aspartate - aminotransferase 130,881 - 0-decarboxylase 1454 - - fumaric acid 867 - - L-alanine 867 - - L-aspartic acid 867 - racemase - - bacterial distribution 1297 - - composite active site 1298 - - essential cysteine residues 1298 - - for synthesis of D-alanine 1298 - - ofarchaea 1297 - - properties 1298 - - purification from streptococcus therrnophilus 1297 - - reaction mechanism 1298 - - substrate specificity 1298 - synthesis with maleate cis-trans isomerase and aspartase 1325 aspartic acid 866,1454 - maleicacid 866 - peptidases 808,812 - phosphinotricin 867 aspartoacylases 749 aspergillus 746, 748 - rnelleus 743 - niger 303, 588, 593, 1355 - niger I F 0 4415 1000 - nigerlipase 418,428,443,474, 555, 559 - oryzae 743,746,752,754,758 - oryzae aminoacylase 743 astasia longu 1037 Astra-Zeneca 1422

Index

asymmetric - dihydroxylation 580 - epoxidation 579 - hydrolysis 473 - synthesis 335, 342,412,473 - transformation 458 atenolo1 1087 atomic absorption spectrometry 785 ATP 1245 - regeneration - - by acetate kinase 1319 - - systems 909 ATPases 897 autoregeneration 1129,1246 auxotrophy 119 axial 370 axial-chiral 545 - diols 502 azafenidine 1449 azasugars 934 azetidinone 1444 azide 1035 2-azidoaldehydes 934 azido-2-hydroxypropanal 934 azidolysis 599 azure 1130

b Bacillus 784, 1037 - acylamina acid racemase 1306 - brevis 766, 788,789 - cereus 994 - circulans 774,779 - fastidiosus 765 - lichenijorrnis 1346 - megatenurn 582 - sp. 774,777 - stearothemophillus 76,78, 81, 83,85, 123, 743,747,774,775,777,779,788,789,1012, 1013,1014,1028,1128 - subtilis 123,744, 1353 - subtilis DY 1346 - themoglucosidius 748 - therrnoproteolyticus 78, 1345 Baeyer-Villiger - monooxygenases 1212,1213,1234 - oxidation 1065, 1202, 1204, 1210, 1219, 1222, 1226,1229, 1230 baker’s yeast 308,310, 1016,1020 balstobacter sp. 779,782, 789 BASF 1436 batch reactor 1450 Bayer 1425 beauveria bussiana 308, 588, 593, 1208

1524

I

fndex

beef liver 1037 benzaldehyde 1247 - dehydrogenase 1247 benz[a]anthracenes 1100, 1101 benz[a]pyrene 1100 benzocycloalkenes 1080 benzocyclobutenes 1080 benzoquinones 1130,1132 benzoyl formate 1247 - decarboxylase 1247 benzoyl-phenyl alanine 557 benzyl alcohols 545, 554, 1151, 1152, 1174 5-benzylhydantoin - substrate for hydantoin racemase 1304 benzylic - alcohols 1145, 1171 - alkenes 1171 benzylisoquinoline alkaloids 1259 a-benzyloxyketones 458 6-benzyloxy-(3R,SS)-dihydroxy-hexanoicacid ethyl ester 1423 3-benzyloxy-1,3-propane478 bicyclic - diacetates 369 - lactones 383 - monoesters 360 bicyclo[3.1.O]cyclohexanes 486 bicyclo[3.3.0]octanoIs 545 bifunctional enzyme 1309 BINAP 1453 binaphthols 545 binding motifs 90 biocatalysis 699, 736, 1419 biocatalysts 67, 79, 90, 713 - effects on acid-base conditions 281 - effects on enzyme form 260 - effects on residual water level 264 - effects on solvent choice 276 - effects on temperature 274 - substrate concentrations 274 bioconversion processes 127 biodegradation - alkenes 583 - aromatics 583 biofuel cells 1162 bioinformatics 139 biological - complexity 120 - pathways 153 - phosphorylating agents 899 biosensors 1130 biostoning 667 biotin 19 biotransformations 1419

using whole cells 33 biphasic - reaction systems 1184 - systems 1127,1151 - - medium engineering with organic solvents 832,833 bipyridine complexes 1130 bisabolol 601 BLAST 74, 144 blastobacter sp. 771, 780, 788 bleomycine 1256 !3-blockers 1157 blocks 156 Bower’s compound 602,603 branched chain - aminotransferase (BCAT) - - BCAT 878 - - selection system 878 - - substrate specificity 878 - a-keto acids - - substrate specificity 1053 BrCN-sepharose 359 BRENDA 152,158 brevibacterium - R312 710 - sp. 753 Bristol-Myers Squibb 1423 broad-range - continuous production of L-p-fluorophenylalanine 887 - transaminase - - immobilization 886 - volumetric productivity 887 bromelain 1509 brominated cyclohexenol derivatives 545 4-bromo-1-butene 1088 bromoketones 1025 bromoperoxidase 1142, 1264, 1267, 1479 bromotoluene 1100 7-bromotryptophan 1271 burkholderia - cepacia 413, 1201 - SP. 719 - sp. lipase 503, 526 butanediol 3,487, 1154 butanol 342,352, 370,1008,1009,1152 butanone 1009,1012 butenol 545,1088 butyl hydroperoxide 1143 butyl methyl ether 342, 413,473 butyrate 458 butyrolactones 545 -

C

Ca” 1004 cadaverine 1259 caerulein 856 cakitonin 856 calcium alginate bead 1433 caldariornycesfumago 1262 camphor 1200 - (+)-camphor 1073 - &camphor 1066 candida - albicans 310 - antarctica 295, 296, 546, 716, 1383 - antarctica A lipase 428, 503, 546 - antarctica B lipase 305, 425,435, 443,479, 487, 503, 526,546, 555, 558, 559, 566 - antarctica lipase 294, 335, 347, 418,428, 442,474,487, 559,566 - cylindracea 413,434,1377, 1389 - cylindracea lipase 300,425-428,442,443, 458,459,479,486,487, 503, 526, 545, 546, 554,555,566 - lipolytica lipase 487 - magnoliae 1000,1001, 1010, 1027 - parapsilosis I F 0 0708 1000 - rugosa 413 - rugosa lipase 300, 347,418, 428, 435, 458, 459,474,479, 503, 526, 546,555 - sp.lipase 555 candidate genes 159 canier activation methods 169 capillary electrophoresis 115 caprolactamase 727 capsaicin 1174 N-carbamoylamino acid amidohydrolase 1303 - gene cloning 1304 carbamoyl phosphate 905 carbamoylases 763,767,770,777,780,782, 786,787,790-792,795,1442 S-carbamoyl-L-cysteinehydrolase 1302 carbamyl kinase 905 carbon dioxide 1038 carbonate 335 - dehydratase 1516 carbonyl reductase 993, 1010, 1016 carboxyl esterase 413 carboxylase 73,79 - phosphoenolpyruvate carboxylase 1514 - urea carboxylase 1518 carboxylester hydrolase 335 carboxylic acids 342, 351, 555, 558, 1154 - esters 335,473, 554, 555 carboxymethyl thioester 848

index carboxypeptidases 197,802,809,853,1336, 1345,1352 - carboxypeptidase A 10,11, 1506 - carboxypeptidase P 1506 - carboxypeptidase Y 1506 carebastine 1160 Carlsberg - subtilisin 1508 carnitine 1447 - dehydratase 1447 - - 4-L-butyrobetaine 694 carotenoid biosynthetic pathway 130 carrageenan 571 carrier types 170 - inorganic 171 - organic-biopolymer 171 - organic-synthetic polymer 171 - proteinaceous 171 - synthetic 171 (-)-carve01 1148 (-)-camone 1148 carvotanacetone 1072 caryophyllene 1096 p-casomorphin 1398 caspases 808,812-814 cassette 123 - mutagenesis 69, 105 castanospermine 1386,1387 catalase 1145, 1428, 1479 catalysis by UDP-glucose pyrophosphorylase 615 catalytic - activity 81,87, 89, 473 - antibodies 840, 955, 1467 - residues 88 - triad 585 catechin 1390 catechol 1179,1186 - methyltransferase 1482 catecholase activity 1176 cathepsin 812, 1506 - cathepsinc 1399 cefalexin 1445 cell extract - acinetobacter calcoaceticus 1423 cell-free transcription/translation system 121 cellobiase 1501 cellobiohydrolases 663 cellobiose 1379 cehlases 321, 663, 1498 cellulose 321, 661 - hydrolyzing enzymes 321 celhlosome 665 cephalosporins 1341,1436,1512

1526

I

lndex

ceramide glycanase 1398 cetus process 1132 chain elongation 728 chalcone 1390 chaperonin GroESL - solubilization of glutamate racemase 1293 characterization of immobilized biocatalysts 178 chelation affinity binding - carbohydrate resin 166 - diethylaminoethyl 166 - ionic adsorption 166 chemferm 1444 chemical - modification 86, 335,407 - mutagenesis 99,124 - racemization - - combination with enantioselective hydroysis 1307 - resolution 1431 chemoselectivity 706 chiral - alcohols, primary 486 - auxiliaries 397, 545 - chromium carbonyl complex 502 - glutarates 365 - cis-glycols 1101 - malonates 365 - monoacetates 370 - monoesters 343, 346 - synthesis 131 chitin 325 - binding domain 820 chitinases 326,1500 chlorinated hydrocarbons 342 chlorine 673 chloroacetates 458,473 a-chloroalanine - production 1287 l-chloro-l,2-epoxypropane, 1088 a-chloroketoester 1025 chloroketones 1025 4-chloro-3-oxobutanoate 993, 1000, 1001, 1026 5-chloro-2-pentanone 1020, 1027 chloroperoxidases 581, 1143, 1145, 1262, 1267, 1270,1273,1274,1480 chlorophenyl-acetate 565 chlorotoluene 1100 7-chlorotryptophan 1271 cholecystokinin 856 cholesterol 1158 - esterases 347,413, 426, 434, 458,459, 1343 choline esterase 148 chondroitinase 1516

chroman 1080 chromanone 1390 chromatography 49,757 chromobacterium viscosum lipase 418,426, 474,479,486 chryseomonas luteola 594 chymopapain 1509 chymosin 1509 chymotrypsin 805, 832, 834,839,849,852, 854,855,857,1334,1336,1345,1377,1399 - a-chymotrypsin 335, 336, 342, 347, 398, 399,472,1337,1381,1394 - chymotrypsin-catalyzed 850,858 - equilibrium-controlled synthesis 830 - frozen aqueous systems 837 - medium engineering with organic solvents 833 - prediction by S’subsite mapping 829, 830 chymotrypsinogen 841,842 13-cineole 1073 cinnamic acid 1454 cinnamyl alcohol 1152 citraconase citraconate hydratase - kitramalate 688 - 2-methylmaleate 688 - (R)-2-methylmaleate dehydratase 688 - 2-methylmalic acid 688 citrate dehydratase - cis-aconitate 688 - citrate 688 - isocitric adic 688 citric acid derivatives 369 citronello1 1095 classification 160 - E.C: 809 - of immobilization methods 165 cleavage of P-0-bonds 895 CLEC 342,413,473 clostridium - beijerinckii 1010 - histolyticum 1335 - oroticum 765, 787-789 - uracilicum 780, 781 clostripain 807, 808, 846, 854, 1509 - prediction by S’subsite mapping 829 dostripin 845 CMP-N-acetylneuraminic acid 618 - derivatives 619 - synthetase 618,1486 C-N bonds 1435 C-N ligase 859 C-0 bonds - formation 1430

hydrolytic cleavage 1430 cobalamine 21 cobalt tert-pyridine 1142 cocaine derivatives 397 codon bias 104 codon-levelmutagenesis 101 coenzymes 12,16, 335,991,1108 - coenzyme A (CoA-SH) 18 - regeneration 992 cofactor 12, 16, 1047 - consumption 238 - regeneration 1048 - - FDH/formate 1054 - - glucose DH/glucose 1054 - specificity 82,89 COG database 157 colchicoside 1383 cold surface 269 cold-activeenzyme - alanine racemase 1284 collagenase 873,1335,1510 coUetotrichum gloeosporoides 307 colominic acid 1324 colyophylization 571 comamonas - acidovorans 719,725,780 - SP. 781 - testosteroni 707, 752, 753 - testosteroni N I 1 700 commercial - biocatalysts 65 - enzymes 41 commodity chemical acrylamide 1450 comparative modeling 68,70, 73-75 comparison of immobilization methods 170 conducting polymers 1113 conductive polymers 1186 configured aldofuranose, 2R,3R - conversion by xylose isomerase 1317 confocal fluorescence coincidence analysis (CFCS) 116 consensus - concept 80,81 - sequences 106,1098 conservation patterns 145,146, 154 continuous -culture techniques 119 - flow membrane reactor 342 - operated stirred tank reactor 1429, 1441 - process 1451 control of water activity 269 conversion 196, 344, 346, 348 coordination site 146 core packing 127 -

correlation length 102 corresponding transition 338 cortexolone 1079 corticosteroids 1067 corynebacterium 582,770 - sp. 1210 - strain 1033 - strain STlO 1010 corynesporium cassiicola 592 cosolvents 342,398,412,425 cost contribution 1453 coumarine 1390 counter-ions 281 - loss 283 coupled residue 102 covalent attachment 170 - acylation 168 - alkylation 168 - conjugate addition 168 - deactivation 168 - imine formation 168 creatinase 1513 creatine amidinohydrolase 1513 creatininase 1512 creatinine amidohydrolase 1512 cresolase 1482 - activity 1176 cross-linked - crystals 261 - enzyme crystals 179,342,835 - - cross-linked enzyme aggregate (CLEA) 180 - - lipases 179,180 - - penicillin acylase 179, 180 - - proteases 179,180 - glutaraldehyde 179 cross-links 261,343 cross-linking - amorphous solid 175 - high fructose corn syrup 175 - whole cells 175 crotonase 149 crotonobetainyl-CoAhydratase - butyrobetaine 694 crotonylamino glutarate 365 crown ether derivatives 544 crude preparations 343 crystal structures 398, 413 crystallization 64, 261, 757 cultivation 1003 cunninghamella echinulata 1031, 1229 (+)-curdione 1074 curvularia - inaequalis 1270

1528

I

lndex

lunata 1222, 1226 cyamide hydratase 702 cyanidase 702,703 5-cyano valeramide 1449 cyanoalanine synthase 1517 cyanobacterium 994 cyanogenesis 975 cyanohydrins 289,458,544, 565 - acetates 458 - enantioselective synthesis 974 - follow-up chemistry 985 - ofaldehydes 979 - ofketones 981 - of 3-phenoxybenzaldehyde 976 - preparations of optically active cyanohydrins 974 - safe handling of cyanides 988 cyanopyridine 1451 cyclases 898, 1378 cyclic - dicarboxylic acids 426 - - diesters 360,427 - diesters 365 - dimethanol derivatives 478 - diol diacetates 399 - 1,2-diols 397 - ketones 996 - monoacetates 370 - monoesters 365 - secondary alcohols 545 - secondary dialkanoates 425 cyclitols 936 cycloalkanols 458, 545 cyclodextrins 316,659, 1007 - glycosyltransferases 316, 659 cycloheptane derivatives 370 cyclohexadiene carboxylates 383 cyclohexanes 413,434 - derivatives 349, 397 cyclohexanoid 370 - compounds 397 - dimethanol 486 - methylesters 360 cyclohexanol 545 cyclohexanone monooxygenase 1214,1216, 1263 cyclohexene dicarboxylate 351 cyclohexenoid - monoacetate 370 - monoesters 351, 360, 398 cycloisomerase 1281 cyclopentanoid 370, 398,434 - dimethanol 486 - dimethanol diacetates 425 -

monoesters 360, 434 cyclopentanol 1003, 1021 cyclopentanone monooxygenase 1214 cyclopentenylester 1454 cylindrocarpon destructans 1225 cysteine 1302 - desulfhydrase - - degradation of cysteine by 1302 - peptidases 805,812 - - mechanism of catalysis 807 - synthesis from ~,~-2-amino-A~-thiazoline4-carboxylate 1302 cytochrome - cytochromeC 768 - cytochrome P-450 1066,1262 - cytochrome P-450 monooxygenase 1263 -

d DAHP synthetase 947 dancus carota 1027 Danvinian theory 95 databases 144,160 DCIP 1146 deacetoxycepharosporin C 1308 - hydroxylase 1309 deactivation of the enzyme 343 deacylation 337, 545 deamidation 1436 deaminases 766, 1516 - adenosin deaminase 1513 - cyclodeaminase 871 - histidine deaminase 869 - serine deaminase 871 - threonine deaminase 871 deaminooxytocin 1334 debranching enzyme 1503 decarboxylases - !3-decarboxylase 877 acetolactate decarboxylase 1514 - aspartate decarboxylase 867, 1454 - benzoyl formate decarboxylase 1247 - glutamate decarboxylase 1296,1297 - lysine decarboxylase 1514 - methionine decarboxylase 1514 - oxaloacetate decarboxylase 885, 1513 - phenylalanine decarboxylase 1514 - pyruvate decarboxylase 1513 - tyrosinedecarboxylase 1514 decarboxylation 877 I-decene 1086 definition 163 degradation - enzymatic 725 - ofaromatics 582 ~

Degussa 1441 dehalogenase 149 dehalogenation 999 dehydratases 686,690-692,695 - altronate dehydratase 690 - arabionate dehydratase 690 - captopril 694 - carbapenems 694 - carbonate dehydratase 1516 - L-carnitine 694 - citrate dehydratase 688-690 - diol dehydratase 582 - glycerol dehydratase 692 - (R)-2-hydroxyisobutyricacid 694 - hydroxysteroid dehydratase 695 - isopropylmalate dehydratasr 688 - mannonate dehydratase 690 - methylmalate dehydratase 088 - serine dehydratase 871 - tartaric acid dehydratase 6886 - xylonate dehydratase 690 dehydration 342 dehydrogenases 25,26,288,754,766,896, 937,1108,1146,1454 - acetoin dehydrogenase 14711 - alanine dehydrogenase 1049,1053,1251, 1283,1284,1286,1476,1517 - alcohol dehydrogenases 81,992,994,996, 997,1006,1009,1010,1014 1017,1018, 1029,1038,1109,1120,1115,1120,1194, 1231,1469 - aldehyde dehydrogenase 1196 - amino acid dehydrogenase 1047 - benzaldehyde dehydrogenase 1247 - cellobiose dehydrogenase 1131 - dihydrolipoamide dehydrogenase 82.83, 1478 - dimethylamine dehydrogenase 1478 - enzyme-coupled regeneration 1110 - formaldehyde dehydrogenase 1476 - formate dehydrogenase 23 I , 993,1014, 1048,113G,1184,1287,1295,1475 - fructose dehydrogenase 1126,1475 - glucose dehydrogenase 99:1,994, 1037, 1126,1146,1148,1472 - glucose phosphate dehydrogenase 993, 1126,1472 - glutamate dehydrogenase ?9,80, 1037, 1054,1295 - glyceraldehyde 3-phosphate (phosphorylating) dehydrogenase 1476 - glycerol dehydrogenase 992,1122,1470 - glycerol-%phosphatedehydrogenase 1124, 1470

- hydroxybutyratedehydrogenase 1471 - hydroxyisocaproatedehydrogenase 231, 1015,1058 - hydroxysteroid dehydrogenase 84,85, 996, 1127,1160,1473 - iditol2-dehydrogenase 1470 - isocitrate dehydrogenase 1013, 1039 - isopropylmalate dehydrogenase 1013 - intrasequential regeneration 1110 - lactate dehydrogenase 1012-1014,1036, 1109,1125,1470,1471 - leucine dehydrogenase 1049,1052, 1053, 1477 - maleate dehydrogenase 1471 - rnandelate dehydrogenase 1310 - mannitol dehydrogenase 1126 - methanol dehydrogenase 1038,1147 - NADPH/ NAD(P)+)dehydrogenase 1111, 1286, 1478 - - aldehyde dehydrogenase (NAD(P)+) 1475 - - glutamate dehydrogenase (NAD(P)+) 1477 - - isocitrate dehydrogenase (NADP') 1471 - nicotinic acid dehydrogenase 1252 - other dehydrogenases 1127 - phenylalanine dehydrogenase 1049,1054, 1056,1477 - phosphogluconate (decarboxylating)dehydrogenase 1472 - polyol dehydrogenase 1126 - quinohemoprotein dehydrogenase 1147 - quinoprotein dehydrogenase (QDH) 1146 - sorbitol dehydrogenase 1470 - trimethylamine dehydrogenase 1478 - triosephosphate dehydrogenase 1476 - xanthine dehydrogenase 1473 dehydrogenation 1174 dehydroprogesterone 695 dehydropyrirnidinases 767,774 deleterious mutations 102 deletions 141,143 Delft Cleavage 732 delta sleep inducing peptide 856 dernethylation 1067 demonstration - glucose isomerase 166 4-deoxy-4-fluorofructose1319 deoxy-fluoroglucose 1317, 1319 4'deoxy-4'-fluorosucrose 1319 5-deoxy-~-fluoro-~-xylulose 1317 - production with xylose isomerase 1318 deoxy-glucose 1317 deoxy-~-manno-2-octu~osonate 8-phosphate synthase 946

1530

I

lndex

3-deoxy-~-manno-2-octulosonate aldolase 946 1-deoxynojirimicine 1387 deoxyribonucleases 895,1496 deoxyribose-5-phosphatealdolase (DERA) 950,1321 1-deoxy-5-thio-galactopyranose 935 1-deoxy-5-thio-D-mannopyranose, 935 deoxy-thio sugars 935 depeptidyl peptidase 809 depolymerases 1500 deracemization 288,596,1136,1158,1251, 1255,1256 derivatives 1100 descriptors 154 designer yeast 1241 desoxynojirimycin 1150,1425 detoxification 581 dextranase 1500 dextromethorphan 571 DHAP 932,943 diacetates 346,350-352,369,370,407,417, 478 diacetyl reductases 1028,1129 diacylated diols 346 dialkanoates 425 dialkyl dicarboxylate 370 diaminase 1513 a-diamines 1259 diaminopimelate 1299 - epimerase - - essential cysteine residue 1299 - - evolutionary origin 1299 - - genecloning 1299 - - mechanism-based inactivator 1299 - - properties 1299 - - tertiary structure 1300 - - topology of secondary structural element 1300 diamond lattice model 996 diary1 ketones 1029 dicarboxylic acids 352,1199 - diesters 351 - esters 370,383 dichloroacetate 473 dichloroindophenol 1132 dichloromethane 473,486 dichlorophenacylchlorise 1025 Diels-Alderreaction 1177,1407 diesterases 898,918,923-925 diesters 343,383,398,426 diethyl ether 398,342,473, 545,554 differentiation 343 difluoroacetophenone 1023 digital imaging analysis 115

dihydrolipoamide dehydrogenase 82,83 dihydroorotase 765,768 dihydropyridines 434,1259 dihydropyrimidinases 765,771,774,775, 1512 dihydroxyacetonephosphate 1321 dihydroxyacid dehydratase 691 dihydroxyphenylalanine 1448 diisopropyl 544 - ether 413,417,434,473, 545 - phosphofluoridate 805,816 diketene 544 diketocamphane monooxygenase 1216 diketoester 1423 diltiazem 252,1430 dilution series 119 dimethyl sulfoxide 342,352,360,398,412 dimethylformamide 342,398,412 dimethylhexadiene 1093 dinitrodibenzyl 1428 diols 336,352,579,1120,1122,1129,1137, 1249 - dehydratase 582 dioxygenases 129,1100 dioxygenation 1099 dipeptidases 802,809, 1506 dipeptidyl - aminopeptidase 1506 - peptidases 802 - transferase 1506 dipropionate 473 directed evolution 68,90,95,96,335,407, 1014,1099 - of hydantoin racemase 1305 disaccharides 1134,1142 dismutase 1482 disparlure 600 disulfide - bond 89 - bridge 89,90 dithio - acetal derivative 427 - monoester 427 divergent evolution 1050 DMS 1439 DMSO 597 DNA 124 - ligases 895 - polymerase 72,73,79,329,895 - shuffling 68,107 - shuppling 123 - synthesizer 105 DNases 918,922,923 dodecadiene 1086

domains 149 r-dopa 1448 L-dopamine 1448 dried cells 37 drying agents 273 DSM 1440.1442 dual-action vasopeptidase inhibitor 882 DuPont 1449 dynamic - kinetic resolution 351, 544, 558, 559, 565, 1030,1441,1442 - programing 141, 144 dynorphin 856 e E 348 - value 348, 349 E.C. nomenclature 152 E.C.l.l.l.l 1422, 1423 E.C.1.1.99.21 1425 E.C.I.I.X.X. 1424 E.C.1.11.1.6 1428 E.C.1.4.3.3 1426 E.C.1.5.1.13 1428, 1447 E.C.l.X.X.X 1427 E.C.3 1430, 1435 E.C.3.1.1.3 1430 E.C.3.1.1.25 1433 E.C.3.1.1.41 1436 E.C.3.2.1.1 1433 E.C.3.2.1.3 1433 E.C.3.4.1.11 1439 E.C.3.4.21.62 1431 E.C.3.4.24.27 1446 E.C.3.5.1.11 1438, 1444 E.C.3.5.1.14 1441 E.C.3.5.1.77 1442 E.C.3.5.2.2 1441 E.C.3.5.2.11 1442 E.C.3.5.5.1 1447 E.C.4.1.99.2 1448 E.C.4.2.1.84 1449, 1450, 1451 E.C.4.2.1.89 1447 E.C.5.1.1.15 1442 E.C.5.1.3.8 1452 ebastine 1160 EDP-GlcNAcpyrophosphorylase 616 ee value 344, 352 effect of temperature 204 effectors 26 EF-hand 146 EGF 856 Eigen 96 elastase 813, 1507

Index

equilibrium-controlled synthesis 830 - prediction by S’subsite mapping 830 electrical - power 1185 - wiring 1126 electrochemical - enzyme regeneration 1130 - methods 1098 - oxidation 1251 - reduction 1145, 1199 - regeneration 995, 1112, 1130 electrode 1113, 1126, 1146, 1162 electron - acceptors 1129, 1132 - transfer 1130 eledoisin 856 Eli Lilly 1444 emulsion 342, 975, 1431 - process 1184 enantiocomplementary 416 enantioconvergent synthesis 598 enantiodivergent synthesis 346 enantiogenic 1079 enantiomer - preference 347 - recognition 473 enantiomer-differentitating 344, 345, 348, 352, 399,405,473,545,546, 566 - acylation 486,487, 503, 525, 526, 544 - alcoholysis 545, 554, 555, 557 - hydrolysis 345, 352, 370, 371, 383, 384, 405, 428,434,435,443,458,459,486,557 - process 558 enantiomeric - esters 343 - monoesters 343, 344 - ratio 348 enantiomers 348 enantioselective - acylation 571 - hydrolysis 343 enantioselectivity 103, 129, 131, 348, 370, 1048 - improvement 999 enantiotopic - acyloxy 343,344 - ester groups 336, 343,360, 365 - groups 348, 349, 373,416,473 - - recognition 346,417,425 - hydroxyl groups 336 - methoxycarbonyl groups 343 - selectivity 352 enantiotopos 345 - differentiation 344 -

I””

1532

I

lndex

enantiotopos-differentiating 343, 344, 352, 473,474,546, 566 - acylation 479 - alcoholysis 554, 555 - hydrolysis 351-353, 361, 366, 369, 399, 406,414,418,426 encapsulation 170, 171 - liposome encapsulation 174 - microencapsulation 174 endoglucanase 321,663,1498,1499 endolinkers 1393 endomyces - mugnusii 1020 - resii 1020 endonucleases 129,895, 922,923 endopeptidases 801,808-810 - adenovirus endopeptidase 808 - endopeptidase K 1508 - lysyl endopeptidase 810 - serine endopeptidase 1506 endoproteinases - Arg-C endoproteinase 1509 - Asp-N endoproteinase 1510 - Glu/Asp-C endoproteinase - - equilibrium-controlled synthesis 830 - - prediction by S’subsite mapping 830 - Glu-C endoproteinase 1507 - Glu-C V8endoproteinase - - equilibrium-controlled synthesis 830 - - prediction by S’subsite mapping 830 - Lys-C endoproteinase 1508 - Pro-C endoproteinase 1506 endoxylanase 670, 1499 enhancement 569 enol - acetates 458 - esters 458 enolase G92 enolyl-CoA - hydratase - - enoylase 693 - - (R)-2-hydroxybutanoicacid 693 - hydratase/isomerase 88 - isomerase 149 enterobacter ugglomeruns 1149 entrapment 170,171 - alginate 173 - carrageenan 173, 174 - polyacid 173 - polyacrylamide 173 - polyacrylate 173 - polymer network 173,571 - precipitation 173 entropy calculation 117

enzymatic halogenation 1270 - peptide synthesis 822 - - chemically modified enzymes 835 - - fragment condensation 854 - - genetically engineered enzymes 835 - - immobilized enzymes 834 - - in frozen aqueous systems 836 - - in solvent-free micro-aqueous systems 838 - - planning and process development 851 - - solid-to-solid peptide synthesis 839 - - solvent-modified enzymes 834 - - stepwise chain elongation 853 - - substrate engineering 839 enzymes 158,261,281, 337, 342,343 - activity 189, 190, 722 - availability 1468 - carriers, for 172 - classification 991 - commercially available 1461 - committee 152 - consumption 238 - cross-linked 342 - crystals 343 - database 152 - engineering 95 - experimental by Novozymes 1463 - families 148 - flexibility 127 - form 260 - function 4 - immobilization 193 - isomerization 1282 - kinetics 23, 208 - - catalytic 214 - - competitive inhibition 214 - - initial reaction rates 209 - - international unit 213 - - Michaelis-Menten kinetics 210 - - multiple enzyme systems 230 - - non-competitive inhibition 215 - - ordered bi bi 220 - - ordered bi uni 221 - - ping pong bi bi 221 - - randombibi 220 - - random bi uni 220 - - resolution 558 - - reversibility 217 - - substrate inhibition 227 - - theorell-chance 221 - - two-substrate reactions 218 - - uncompetitive inhibition 216 - membrane reactor 237, 241 -

metal combination 293 - nomenclature 21,1461,1465 - quality requirements 32 - pH-optimum 191 - producers/supplicers 1464, 1468 - productivity 1058 - reactors 174 - - batchreactor 232 - - continuous stirred tank reactor 233 - - mass balances 234 - - pflug flow reactor 233 - - stirred tank reactor 232 - selectivity 190, 280 - specificity 280 - stability 190, 192 - structure 4 - substrate complex 337,850 - suppliers 44,45 - synonym 1468 - temperature stability 191 - tetrahedral intermediated 338 - transition metal combination 294, 295 - typical abbrevitation 1462 - use in organic solvents 31 eosinophil peroxidase 1269 epimerases 149, 1281, 1282,1299,1300 - acetyl glucosamine epimerase 1324 - aldose epimerase 1517 - amino acid epimerase 1293 - E. coli 1452 - isopenicillin N epimerase 1308 - tagatose epimerase 1322 epimerization 565, 1452 - of allo threonine 1290 epoxidation 1067,1084 epoxides 579, 592,1084 - aminolysis 599 - azidolysis 599 - disubstituted 594 - hydrolases 579 - - isolation 582 - - mechanism 584 - - screening 587 - -structure 584,585 - isomerase 582 - monosubstituted 592 - styrene oxide-type 593 - trisubstituted 596 epoxy alcohols 442 epoxyindene 589 equilibration 287 equilibrium 338, 341,748 - conversion 194 - of hydrolysis reactions 264 -

index error - level 113 - prone PCR 99, 105,131 - rates 100 - threshold 103 erythrose - substrate oftylose isomerase 1317 erjthrulose 1317 eschericha coli 68, 70,79, 82, 83, 123, 308, 311, 775,777,782,791,1336 essential 873 ester 335, 336, 351, 545 - formation 416 esterases 158,677, 1487 - acetyl esterase 1378, 1383 - acetylcholin esterase 406, 407, 1494 - butyrylcholine esterase 1494 - carboxyl esterase 413,1487 - catalyzed hydrolyses 352 - cholesterol esterase 347,413,426,434,458, 459, 1343,1494 - cholin esterase 148, 1494 - diesterase 898,918, 923-925 - pectinesterase 1494 - porcine liver 335, 336, 342-344, 346, 351-353,360,361,365,366,369-371,383, 384,398,407,425,472,571,572,840,1383 - rabbitserum 1378 - resolution 983 - sterol esterase 1494 esterification 336, 473, 555, 558 esters 154, 338, 342, 546 - of racemic alcohols 336,435,443,459 - of racemic carboxylic acids 336 ethanol 352, 398,412, 545, 1152 ethanolamine ammonia lyase 871 ethene 1090 ethers 342 ethyl - acetate 342, 473, 478 - chloroacetate 1004 - chlorohydroxybutanoate 1157 - chlorooxobutanoate 1157 - methoxy acetate 473, 1435 - octanoate 473 - thioacetate 458 - thiooctanoate 473 ethylene glycol 1137,1139 eudesmanes 1075 eugenol 1191 Euglenales 1037 eukaryotic - cell 1420 - enzymes 122

Eupergit 342,352, 571, 1438, 1446 evolution 148 evolutional relationship - between diaminopimelate epimerase and aspartate racemase 1299 - between diaminopimelate epimerase and glutamate racemase 1299 evolvable 97 exchange of water with the environment 269 exoglucanase 321,663 exolinkers 1393 exonucleases 922, 923 exopeptidases 802,808,809,853 exosialidase 1500 expanded bed absorption chromatography 1183 experimental enzymes by novozymes 1463 expression 122, 791 - pattern 160 extracellular enzymes 48 extraction of enzymes 47 extreme thermophiles 313 extremophiles 67

f

FAD-containing monooxygenase 1263 FADHZ-dependent halogenase 1268,1275 family shuffling 100, 111 farnesol 1093 FAS 1027 FASTA 74,144 fatty acids 342, 1085, 1325 - synthase 1012 FDH 1058 FDP aldolase 931-933,936,938,953,961 fed batch reactor 1448 (+)-fenchone 1073 fermantation 41 - ofenzymes 46 fermlipase 303 ferredoxin-NADP' 995 ferricyanide 1146 ferrocenes 1113, 1130, 1142, 1146 - alcohols 502 fibrinogenase 1507 fingerprint 83 - motifs 82 first-order rate constants 344 fitness - information 117 - landscape 97, 107 fixed bed reactor 1433 flap 348 flavanone 1390

flavin 991 monooxygenase 1214 - nucleotides 16 - reductase 1268 jlavobacterium 785 - SP. 785,788,789 flavocytochrome enzyme 1188 flavoperoxidase 1143 fluorescence-activated cell sorter (FACS) 116 fluorinated - acetophenones 1007 - alcohols 1024 - ketones 1005,1021, 1024 - malonates 427 - toluenes 1100 FMN 1111 - reductase 1111 focused mutagenesis 99,104,105 folate coenzymes 20 folding process 9 formaldehyde 1038 formate 1038, 1245 - dehydrogenase 231,993,1014, 1048, 1136, 1287,1295 formation - ofamides 716 - of carboxylic esters 472 - of C-N bonds 699 - ofhydantoins 761 - ofP-Obonds 895 formylesters 1199 freeze-concentration effect 838 frontalin 602 frozen aqueous systems 837 P-fructofuranosidase 1502 fructose 1132 - bisphosphate 130,1319 - - aldolase 1317 - - from glucose 1319 - dehydrogenase 1126 - phosphate 1318 - - kinase 1319 fucose aldolase 939 fucose analogue - modified at nonpolar terminus 1323 fucose isomerase 939,941 - three-dimensional structure 1323 fucosidase 129, 636 fucosylated oligosaccharide 1323 fucosyltransferase 624 Fuji Chemical Industries 1433 Fujisawa Pharmaceutical Co. 1438 fumarase - aspartic acid 867 -

malic acid 867 fumarate hydratase - fumaricacid 687 - malic acid 687 fumaric acid 1454 functional - domain 149 - space 106 &sariurn s o h i pisi cultinase 418 fusion 149 -

g galactose 1132, 1141 a-galactosidases 635, 636, 1402, 1501 galactosyltransferases 609, 6l!i, 619, 1484 galacturonic acid 673 ganglioside G M 3 1398 GAOX 1142 gap penalties 142,143 GC 115 GDP-fucose 617 GDP-mannose 617 - pyrophosphorylase 617 gel filtration 56, 57 generalized profiles 155 genetic - algorithms 98,102,112 - code 101,119 - engineering 819 - methods 1010 - modified organisms 45 genome sequences 1098 genotype-phenotype coupling systems 120 gentobiase 1501 geotrichurn candidurn 307, 3101,994, 1002, 1004,1006,1008,1018-1027,1156 - glycerol dehydrogenase 992 - lipase 418,479 geraniol 1069, 1152 geranylacetone 1093 germacrone 1074,1096 - epoxide 1074 gibbereliinic acid 1385 Gist-Brocades/DSM 1438 Glaxo 1453 GlcNAc-phosphate 616 GlcNAc-transferases 609 Glu/Asp-specific endopeptidase (GSE)840 Glu4-oxytocin 1335 glucan glucosidase 1498, 1503 glucanase 1503 glucoamy1ases 315, 656, 1498 glucoinvertase 1501 glucokinase 1319, 1484

gluconobacter oxydans 1149 glucosamine 1452 glucose 1132,1433 - dehydrogenase 993,994,1037, 1126 - derivatives 1316 - isomerase 1313 - oxidase 1145 - phosphate 615,1318 - phosphate dehydrogenase 993,1126 - pyrophosphorylase 909 - sensors 1139 glucosidases 654 - a-glucosidase 315, 321,636,657, 663, 664, 1501 - P-glucosidase 1501 - amyloglucosidase 1433, 1498 - glucan glucosidase 1498, 1503 - thioglucosidase 1503 glucosyltransferases 632, 654 glucuronidases 671, 1502 glutamate 1293 - ammonia ligase 1518 - decarboxylase 1297 - dehydrogenase 79,80, 1037,1054, 1295 - oxidase 1297 - production 1287,1289,1297, - racemase 1295 - - gene cloning 1293 - - properties 1293 - synthesis 1296 - - catalytic 1294 - - for poly-glutamate production 1294 - - from gram-positive bacteria 1294 - - structure and mechanism 1294 - - tertian structure 1294 - - thermostable enzyme from bacillus subtdis 1294 glutaminase 25, 1511 glutamine synthetase 1518 glutamyl - endopeptidase 810 - transferase 1483 - transpeptidase 1483 glutaraldehyde 177 glutarates 398 glutaric acid anhydride 558 glutaryl-7-ACA 1427 glutaryl amidase 1436 glutathione 1337, 1343 - peroxidase 1142 - reductase 82, 83, 1236 glycals 544 glycanases 1398, 1499 glyceraldehyde 1122

1536

I

lndex

glycerol - dehydatase - - PDO 692 - - polyols 692 - derivatives 571 - diacetate 425 - kinase 907,908,909 - monoacetate 417 glycidic acid methyl ester 1430 glycidol 1147 glycine-dependent aldolases 953 glycogenase 1497,1498 glycohydrolases 1500 glycolate oxidase 1137 glycolic acid 1137 glycol monoester enzyme 585 glycopeptide 1349 glycoprotein 1483 - remodeling 641 glycosidases 633,1338 - transferase inhibitors 639 glycosides 637 glycosidic bonds 609 glycosyl transferases 611,612,619, 627-629, 631,909,1394 - cyclodextrin glycosyl transferase 316,659 - inhibitors 639 glycosylation 611 glycosyl-enzyme intermediate 587 glyoxal 1139 glyoxylic acid 1137 goat liver lipase 503 gold 157 Golgi apparatus 609 group preference 344 growing cells 36 growth hormone releasing factor 856 guanidinium chlorid 412 guanidinophenyl ester 843-846,849,850 l-guanyl-32,5-dimethylpyrazole841 guanylate kinase 906

h

haemb 17 HAL 869 Haldane equation 218 halogenases 1268,1275 halogenating enzymes 1267 halogenations 1267,1271,1273,1277 halomethyl ketones 1025 haloperoxidases 1262,1264,1267,1268,1271, 1277 - perhydrolase 1267 Hamming distance 118

hanging drop method 72 hansenula anovnala 1022 Heck reaction 1407 helianthus annus 1385 helicase 1385 helicenediol 502 helix 79 - stabilization 127 heme - containing 1267 - type 1268 hemiacetals 558 - hemithioacetals 291,292,565 hemiaminal 558 hepatitis B S antigen 856 heptane 1084 heptanediol 478 heptyl methyl ether 1067 hesperidin 1383 hesperidinase 1484 heterocyclic 1100 - monoesters 360,370 heterogeneous conditions 343 heterologous expression 1098 heuea brasiliensis 976 hexadecanoate 458 I-hexadecene 1084,1087 hexadiene 1086,1093 hexahydronaphthalenones 1081 hexanal 1151 hexane 349,413,473,545 hexanediol 478 hexanol 545,558,1151,1152 hexoaminidases 1503 hexoketose I-phosphate 1317 hexokinase 948 hexylcyano complexes 1130 hexylchymotrypsin 835 Hidden Markov model 143,144,154,156,160 high - fructose corn symp 1433 - mutation rate 102,103 - product concentrations 129 high-entropy positions 106 high-throughput 119,160 - digital image analysis 130 hinge movement 1050 histidinase 869 histidine 869 - ammonia lyase 869 - a-deaminase 869 - nikkomycin 869 - urocanoicacid 869 HIV 814

protease inhibitors 873, 1159 retropepsin 814 HLADH 994,1033,1196 HMMs 158 HOBT 1175 Hoechst Marion Roussel 1427,1436 Hoffmann La-Roche 1431 Holland Sweetener Company 1446 hollow fiber ultrafiltration membrane 359 homo sapiens 743,744 homoaconitate - hydratase 689 - homocitric acid 689 homoallylic 525 homology 142,145 - domains 150 - modeling 70,73 homonojirimycin 934 homophenylalanine 1439, 144-1 homopropargylic 525 horse liver 1109, 1115 - alcohol dehydrogenases 992,994,996 - esterase 371, 383 - peroxidase 122, 1034, 1262 hot-spots 106 HPLC 115 HSDH 1127 Ht 31(493-515) peptide 856, 857 human - immuno deficiency virus (HIV) 814 - insulin 820,831,834,852, 558 humicola - lanuginosa lipase 435,487, 546 - sp. lipase 443 humulene 1096 hyaluronidase 1502 hyaluronoglucosaminidase 1502 hybrid - enzymes 151 - in vitro-in vivo recombinatbm method 111 hydantoins 761, 762, 767,785, 794, 1441 - racemase - - bacterial distribution 130'3 - - genecloning 1304 - - inactivation by hydantoin derivatives 1305 - - of hydantoin racemase 1305 - - properties 1304 - - purification from Arthrobacter aurescens DSM 3747 1304 - - sulfur compounds 1305 hydantoinases 306,762-765,767-775,777, 780, 784, 786, 787, 791, 792.,795, 1303, 1304,1512 -

- bacillus brevis 1441 hydratases 149,690,692,694 - acetylene carboxylate hydratase 690 - acetylene dicarboxylate hydratase 690 - aconitate hydratase 688, 689 - captopril hydratase 694 - carbapenems 694 - carnitine hydratase 694, 1447 - citraconate hydratase 688 - cyanamide hydratase 702 - dimethylmaleate hydratase 688 - enoyl-CoA hydratase 88,693 - fuconate hydratase 690 - - mannonate dehydratase 690 - - phosphogluconate dehydratase 690 - fumarate hydratase 687 - hydratase/dehydratase 690 - - farnesyl-CoA dehydratase 695 - - 3-hydroxybutyryl-CoAdehydratase 695 - - 3-hydroxydecanoyl-ACP dehydratase 695 - - 3-hydroxypalmitoyl-ACPdehydratase 695 - - isohexenylglutaconyl-CoAhydratase 695 - - itaconyl-CoA dehydratase 695 - - lactoyl-CoA dehydratase 695 - - long-chain enoyl-CoA hydratase 695 - - methylglucatonyl-Cothydratase 695 - hydratase/isomerase - - enoyl-CoA 88 - (R)-2-hydroxyisobutyricacid 694 - maleate hydratase 688 - nitrile hydratase 173,686, 700-704, 707, 710,712,713,719,1449-1451,1454, oleate hydratase 695 - 2-oxopent-4-enoatehydratase 686 hydrated - reverse micelles 263 - salts 343 hydrations 383 hydride transfer 992 hydrindane 1081 hydrocarbon monooxygenase 1263 hydrocarbons 342 hydrocyanation - of carbonyl compounds 974 hydrogen - abstraction 1185 - bonds 76,77,89,90,127 - bond chromatography 59 - peroxide 1129, 1143, 1428 - safe handling of cyanides 988 hydrogenases 994 hydrolases 22.81, 146, 335, 336, 344-346, 351,407,525,581,686,771,919,1277, 1421,1430,1435,1486,1493 -

1538

I

Index

- acetylcholine acetyl hydrolase 407 - adenosine amino hydrolase 1513 - adenosyl homocysteine hydrolase 1290 - amidohydrolase 742,749, 751, 752,777, 782,786,1302,1304,1510,1511 - Ap4 hydrolase 923 - caprolactam hydrolase 1292 - carboxylester hydrolase 335 - catalyzed 545 - cellobiohydrolase 663 - creatine amidinohydrolase 1512, 1513 - epoxide hydolase 579, 582, 584, 585, 1503 - fold 582 - peptide hydrolase 1503, 1506 - pullulan hydrolase 315, 316, 657-659 - serine hydrolase 337 hydrolyases 686 hydrolysis 335, 336, 338, 342, 343, 348-352, 405,414,416,473,486,544, 545,555, 558, 565,699,705,721,723,748,753,1433,1447 - biocatalytic 716 - of amides 716,719,722 - of amino acid amides 720,741 - of carboxylic amides 719 - ofcyanide 702 - of cyclic amides 727 - ofdinitriles 705 - ofhydantoins 761 - of nitriles 699, 700, 703,708, 710, 716 hydronaphthalenone 1081 hydroperoxides 1034 hydrophilic 145 hydrophobic 79,145 - core 145,149 - interaction 89, 90 - interaction chromatography 54 - polymerXAD 1007 hydroquinones 1186 hydroxy acids 1135,1136 - esters 405 hydroxy butyric acid 1256 hydroxy carboxylic acid esters 544,545,554, 1137 hydroxy ketones 544,1122 hydroxy lactones 383 2-hydroxyalkanes 1120 5-p-hydroxybenzylhydantoin - substrate for hydantoin racemase 1304 hydroxybiphenyl 1179 4-hydroxy-2-butanone 1155 (R)-3-hydroxy-butyrate1422 3-hydroxy-2,4-dimethylglutarate 369 6-hydroxydopamine 1259 hydroxyesters 545, 1031

N-hydroxyglutamate 1294 hydroxyhistidine 1255 2-hydroxy-1-indanone 1159 hydroxyisocaproate dehydrogenase 231,1015, 1058 hydroxyl groups 1122 hydroxylases 1309,1447 - decane hydroxylase 1067 hydroxylations 1066,1086,1170,1174,1176, 1179,1186,1189, 1428 hydroxymethyl ketone 1016 hydroxymethyl lactones 442 hydroxymethylisoxazolinebutyrates 442 6-hydroxynicotinicacid 1254 hydroxynitrilases 1515 hydroxynitrile lyases - availability for organic synthesis 976 - catalyzed addition of hydrogen cyanide 976, 978 - crystal structure of 975 - experimental techniques 981 - immobilization 977 - overexpression 976 - recombinant 977 - safe handling of cyanides 988 - substrate acceptance 976 - used for preparative application 975 hydroxyphenyl glycine 1441 - production by hydantoinase 1303 hydroxyphenylalanine 1439 2-hydroxy-~-proline, - substrate of proline racemase 1301 5-hydroxypyrazine-2-carboxylicacid 1447 hydroxysteroid dehydratase - ergosterol 695 - hydroprogesterone 695 hydroxysteroid dehydrogenase 84,85,996 hyperthermophiles 67,79,313 hyperthemophilicarchaeon 73,79 hypohalogenic acid 1268 hysteresis 260, 266 I

ibuprofen 604 igase 822 imidachlorid 1254 imidases 772, 1336 immiscible 342 immobilization 44, 342, 343, 352, 398, 571, 786,1421 immobilized 1431, 1435 - cell 1433 - dye chromatography 61 - enzymes 189,261

index imprinting 263 improvement o f enantioselectivity 999 in silico screening 1098 in situ - analyses of stereospecificityof hydrogen transfer 1286 - cofactor regeneration 626 - racemization 558 in vitro - assembly 105 - evolution 96 - protein biosynthesis 1246 - recombination 98 in vivo - homologous DNA recombination 110 - oxidations 1190 - recombination 124 - selection 113 inclusion bodies 69 - solubilization of glutamate racemase 1293 1,2-indandiols 1159 indanols 545 indigo 1181,1182 indinavir 589 indole 1143,1181 - derivative 1271 5-indolylmethylhydantoin - substrate for hydantoin racemase 1304 industrial - enzymes 41 - reactor 119 influence of pH 27 information theory 106 inhibition 351 inhibitors 26, 398, 998, 1004 inhibitory domains 151 inorganic salts 569 insertions 141, 143 insulin (human) 856 intact microbial cells 1201, L!47 integrated downstream processing 1420 intein 642 inter alcoholysis 545 interfacial - activation 348 - deactivation 342 intermolecular alcoholysis 54-5, 546 International BioSynthetics 3 427 Internet 151 INTERPRO 156 intramolecular alcoholysis 545, 546 inulinase 1499 invariant residues 145 inverse esters 843

invertase 1502 iodine 654 iodoacetophenone 999 iodoperoxidase 1267 ion exchange - chromatography 49,757 - crystallization 757 ion pairs 79 ionic - interaction 89, 90 - liquids 206,412 ionone 1071 ion-pair networks 80 IPTG 69 irreversible enzymatic - nonpeptidases 840 - substrate engineering 840 - synthesis - zymogens 841 irreversible transformation 344, 345 isoamylase 657 isoatisene 1076 isobutylhydantoin - substrate for hydantoin racemase 1304 isobutyrylhydantoin - substrate for hydantoin racemase 1304 isocitrate dehydrogenase 1013, 1039 isoenzymes 351, 352 isolation of enzymes 41 isoleucine 688 isomerases 22,149, 1281, 1282, 1421 - &trans isomerase 1281, 1325 - classification of 1281 - enoyl-CoA isomerase 149 - epoxide isomerase 582 - glucose isomerase 166, 1313, 1517 - glucose-6-phosphate isomerase 1517 - fucose isomerase 939,941, 1323 - maleate isomerase 866, 1324 - phosphoglucose isomerase 1318,1319, 1517 - protein disulfide isomerase 1517 - rhamnose isomerase 939, 941, 1321, 1322 - sugar isomerase 1281 - triosephosphaste isomerase 1320,1517 - tylose isomerase 1317 - xyloseisornerase 1313-1318,1517 isomerization 661, 1281 isooctane 554 isopenicillin - N epimerase 1308, 1309 - Nsynthetase 1308 isoporpylhydantoin

1540

I

Index

inactivator of hydantoin racemase 1305 isoprenoid synthesis 965 isopropanol 352 isopropenyl - acetate 342,473, 478 - alcohol 478 - esters 571 isopropyl ester 360 (R)-isopropylideneglycericacid 1427 isopropylideneglycerol 1157, 1427 isopropylmalate dehydratase 688 isopropylmalate dehydrogenase 1013 isoquercitrin 1383 isoquinoline 1101 isovaleraldehyde 1153 -

k Kanegafuchi 1442,1443 Kaneka process 776 kcat 88 - value 83 k c a t / h 85887 - value 83 KDG aldolase 950 KDO - aldolase 946 - 8-P synthetase 947 - 8-phosphate phosphatase 918 KDPG aldolase 949 KEGG 153 keto acids 1037, 1048, 1135 - conversion to D-amino acid 1287 a,a:keto diesters 398 P-keto esters 383 2-keto sugars 1132 ketopantoyl lactone 1000 ketoreductase 1010, 1469 KHG aldolase 948 kinases 897,898 - acetate kinase 614, 615, 902, 904, 905, 1246,1319,1486 - acetyl kinase 904 - adenylate kinase 615, 906, 907, 1486 - carbamyl kinase 905 - creatine kinase 1486 - fructose phosphat kinase 1319 - glucokinase 1485 - glycerol kinase 907-909, 1485 - guanylate kinase 906 - hexokinase 948,1484 - monophosphat kinase 615 - phosphoenolpyruvate kinase 1485 - 6-phosphofmctokinase 1319,1485 - phosphoglycerate kinase 1486

pyruvate kinase 614,902,907,909,921, 1485 - streptomycin kinase 1486 kinetic 189 - assays 114 - constants 345 - enantiomer separation 486 - resolution 301, 351, 352, 370, 383, 398, 412, 417, 544,565,571, 587,1118,1122,1129, 1136,1142,1427,1430,1431,1433,1439, 1441 - - alcohols with cataylst 1, 296 - - conventional 287,288,293 - - dynamic 287-289,296,298, 302,723 - - enzymatic 287,293 - - of racemic cyanohydrin esters 983 - - of racemic cyanohydrins 983 - - simple 293 kinetically controlled synthesis - kinetics 827 - prediction (of synthesis) by S' subsite mapping 827 - synthesis 634 - - medium engineering with organic solvents 831 klebsiella - pneumoniae 1031 - pneumoniae I F 0 3319 1000 - temgena 719 kloeckera - magna 308,1031 - saturnus 1028 KM 83 - values 88 Knowles' classification 896 kyotorphin 832,856 -

I laccases 1129, 1130, 1162, 1170, 1174, 1479 lactam 1454 - antibiotics 1439, 1444 lactamase 1442,1454,1512 lactase 1501 lactate 1256 - dehydrogenase 1012-1014,1136 - monoxygenase 1481 - oxidase 1256 lactobacillus - brevis 1029 - delbrueckii bulgaricus 1015 - fermenturn 1027 - kefir 994, 1017, 1027 - - alcohol dehydrogenase 992 - SP. 1014

lactococcus lactis 743 lactonase 1454 - fusarium oxyspoium 1433 lactones 335, 336, 338, 342, 371, 383,428, 442,545,546,554,1120 lactonization 383, 545 lactoperoxidase 1262,1269, 1480 lactose 1379 - synthase 1483 laminarinase 1499 landscape ruggedness 102, 1:16,118 large scale 342 - experiments 359 - production 122, 351 - resolution 412 lecithinase A 1493 Leloir pathway 609, 611 lens esculenta 772 leucine dehydrogenase 1049 - characterization 1052 - cloning overexpression 1052 - isolation 1052 - kinetic parameters 1053 - substrate specificity 1052 leu-enkephalin 854456,1398 levomethorphan 571 Lewis acids 624 LH-RH 856 lid 348 ligand database 153 hgases 22,895, 1518 - chain reaction 105 - C-N- 804,859 lignin peroxidase 1269 limited proteolysis 800, 814, 816, 817 limonene 600,1070,1092,1149 R(+)-limonene linum usitatissimurn 976 lipases 167, 179, 180, 181, 2132, 289, 290, 292, 296298,303,335,336,341-344, 370,407, 413,414,417, 418, 425,426,428,434,435, 442,443,459, 473, 474,478, 479,486,487, 503, 525, 526, 544-546, 5584, 555, 558, 559, 565, 566, 569, 571,840, 1354, 1454, 1488 - aspergillus niger lipase 13B9, 1385, 1390 - burkholderia plantarii lipase 1435 - candida antarctica lipase l67, 294, 305, 335, 347, 418, 425,428,442,4/.3,474, 479, 487, 503, 526, 546, 555, 558, 559, 566, 1354, 1381,1384,1388,1390 - candida cylindracea lipase 300,425, 426-428, 442, 443, 458,459,479, 486, 487, 503, 526, 545, 546, 554, 555, 566,

1353-1355,1368,1369,1377,1379,1381, 1383,1385,1388,1392 - candida lipolytica lipase 487 - candida rugosa lipase 300, 347, 418,428, 435,458,459,474,479,503, 526,546,555 - candida sp. lipase 555 - chrornobacterium viscosurn lipase 418, 426, 474,479,486,1355,1379,1385,1388,1390 - fermlipase 303 - geotrichum candidum lipase 418,479 - goat liver lipase 503 - humicola lipase 167,435,443,487,546 - interesterification 167 - lipoprotein lipase 1492, 1495 - mammalian lipase 413 - mucorjavanicus lipase 414,418,1352,1385 - mucor meihei lipase 167, 413, 418,443,479, 487,526,546,559, 566 - mucor sp. lipase 418,425,479 - penicillin lipase 546 - phospholipase 146,147,898, 1493, 1496 - porcine pancreas hpase 335, 346, 413, 414, 417,426,428,435,474,479,486,503, 526, 546,555,559,1354,1355,1369,1377,1381, 1390,1392 - pseudodomonas aeruginosa lipase 347,435, 443 - pseudomonas cepacia lipase 289, 296298,347, 349, 413,414,418,428,435, 442,443,458,459,474,479,487, 526, 544546,555,559, 566,1354,1369,1381, 1388,1390,1391 - pseudodomonase chlororaphis 700,701, 707, 709-712 - pseudomonasfluorescencelipase 294, 295, 347,417,418, 425,428, 443,459, 474, 478, 479,486,487, 503,526,544-546,555,559, 566,1368,1381,1385 - pseudomonas sp. 335,418,426,428,434, 435,443,459, 474, 479, 503, 526, 546, 554, 559,565,566 - RB 001-05 lipase 1403 - resolution 983 - rhizomucorjavanicuslipase 474 - rhizomucor miehei lipase 413 - rhizomucor sp. lipase 418, 503 - rhizopusjavanicus lipase 474, 1377 - rhizopus delemar lipase 418 - rhizopus niveus lipase 1349 - rhizopus oryzae lipase 443 - serratia marcescens lipase 1430 - spergillus niger lipase 1379 - triacylglycerol lipase 1488 - wheat germ lipase 1377, 1379, 1383

1542

I

Index

lipotropin 1335 lipoxygenases 46,84, 1481 liquefaction 661 liquid fermentation 46 lithium chloride 571 liver alcohol dehydrogenase (HLADH) 997 local alignment 150 local fitness landscape 117 log P 279 - value 205 long-jump mutagenesis strategy 102 Lonza 1428,1447,1448,1452 loop flexibility 127 loracarbef 1445 low solubility 1424 low water - biocatalysts 260 - content 342, 343, 398 - media 259 luciferase 1198 lyases 22, 1421, 1447 - acetylneuraminatelyase 1324 - ammonia lyase 866,868,870,1255 - chondroitin ABC lyase 1516 - citratelyase 1516 - histidine ammonia lyase 869 - hydrolyase 686 - hydroxymandelonitrile lyase 1515 - hydroxynitrile lyase 975-978, 981, 988, 1514,1515 - mandelonitrile lyase 1514 - pectin lyase 679, 1516 - phenylalanine ammonia lyase 1516 - tyrosine phenol lyase 1448,1516 lyophilization 343 lyophilized 478 - powder 260,413,473,478 lyoprotection 263 lysine 882, 1443 - aminogroups 343 - production - -total conversion of racemic a-amino-s-caprolactam 1292 - - with a-amino-c-caprolactamhydrolase 1292 lysozyme 1500 lysyl endopeptidase 810 m

maleate 688 dehydratase 688 - hydratase 688 - isomerase 1324 - - fumaric acid 866 -

maleic acid 866 maltase 1501 maltobiose 1379 mammalian lipase 413 mandelate - dehydrogenase 1310 - racemase - - crystal structure 1311 - - divalent metal ions 1310 - - genecloning 1311 - - large-scale production 1310 - - p(bromomethy1)mandelate 1311 - - purification 1310 - - reaction mechanism 1311 - - similarity to muconate lactonizing enzyme 1311 mandelonitrile 975 manganese peroxidase 1143 manihot esculenta 976 mannitol 1427 - dehydrogenase 1126 mannosidase 636,1502 manufacturing cost 1431 Markov chain analysis 102 marmin 1094 Marukin Shoyu 1453 mass transfer effect 176 mathematical 352 - model 344,348 MDB 154 mean-field theory 118 mechanisms 748 mediators 1113, 1139, 1141 melibiase 1501 membrane 1421 - hollow-fiber 1431 - reactor 237, 1286 menthols 1072 Merck Research Laboratories 1425 MEROPS 154 mesaconate hydratase - citramalate 688 - ethylmalic acid 688 - methylfumarate 688 - methylmalate dehydratase 688 - substituted malic acids 688 mesophiles 313 metabolic - engineering 1288 - pathway 150 metabolism 119 metal - chelate affinity chromatography 61 - ions 146,584 --

metallopepidases 808,812,813 metalloproteases 147,148 metalloproteins 154,743 Met-enkephalin 853,856,1335,1345 methane 1089 - monooxygenases 1067 methanol 342,352,360,398,412, 545,1146 - dehydrogenase 1038 methionine 1255,1305,1441,1442 - synthesis from (methylthioethy1)hydantoin 1305 methods of immboilization - covalent attachment 164 - cross-linking 164 - entrapment 164 - non-covalent adsorption 164 methoxy malonic acid dimethyl ester 554 methoxy viologen 1038 methoxycarbonyl phosphate 904 methyl imidoesters 1086 methyl vinyl ketone 1004 methyl viologen 995 methylaspartase 868 methylaspartate 868 - mesaconicacid 868 methylaspartate ammonia lyase 868 - chloroaspartic acid 868 - dialkylethylaspartates 868 - ethylaspartic acid 868 - methylaspartic acid 868 methylbutanol 1152,1153 methylchrymotrypsin 835 methylcitrate dehydratase 689,690 methyl-1,2-dimethoxysuccinate 369 methylesterases 677 methylgeranate 1091 methylglucose 1317 methylhepteneone 1008,1020,1092,1096 methylhydantoinase 765 methyloxobutanoate 1000,1031 methylphenols 1189 methylpropanol, 558 methylpropyl-butanedioicacid ethyl ester 1431 methylthioethylhydantoin - substrate for hydantoin racemase 1304 methyltransferases 72,73,79, 1482 metoprolol 1087,1157 mevalonolactone 602,603 Mg” 1004 Michaelis-Menten - complex 806,827 - constant 24,748 - equation 1005

- kinetics 76 microarrays 160 microbacterium campoquemadoensis 1000, 1001 microbial - enzymes 44,45,47 - lipase 336 micrococcus 784 microemulsion 263 microorganisms 299 microtiter plate-based screening systems 115 miglitol 1425 milbemycin 1082 minimal screening requirement 116 minimum nucleotide sequence identity 110 miscible 342 - organic solvents 412 misconception 1453 Mitsunobuy reaction 1407 model evaluation 75 modeling 73 , modification 999 - of the substrate 998 modular enzymes 149 modularity 151 modules 110 molecular - breeding 96 - oxygen 1111,1129,1145 - sieves 273,554 - traps 825 monensin 1082,1083 monoacetates 350,352,370,398,407,416, 417,425,473,478,486,525,544 monoalkanoates 427 monoalkyl glutarates 558 monodechloroaminopyrrolinitrin-3-halogenase 1268 monoesters 344,346,351,352,398,442, 557 monofluoroacetophenone 1023 monolactones 383 monomethyl 360 monooxygenases 581,582,1170,1176,1198, 1214,1216,1262 - Baeyer-Villingermonooxygenase 1212, 1213,1234 - camphor monooxygenase 1482 - cyclohexanone monooxygenase 1214,1216, 1263,1481 - cyclopentanone monooxygenase 1214,1481 - cytochrome P-450 monooxygenase 130, 1160,1199,1263 - flavin monooxygenase 1181,1214 - hydrocarbon monooxygenase 1263

1544

I

lndex

hydroxybiphenyl monooxygenase 1179, 1481 - lactate monooxygenase 1481 - methan monooxygenase 1067 - monophenol monooxygenase 1482 - progesterone monooxygenase 1482 - tridecanone monooxygenase 1481 monophasic organic solvents 831, 834 - medium engineering with organic solvents 832 monopropyl ester 360 monosaccharides 1133, 1142 monosubstituted hydantoins, 5 - spontaneous racemization 1304 monoterpenes 1069 Monsanto 1448 moraxella sp. 1009 - alcohol dehydrogenase 992 MPEG 343, 571 MPMS 1130 mRNA-protein fusions 121 MSH 856 muconic acid 1248 mucor - circinelloides 307 - griseocyanus 308 - heimalis 1020 - javanicus 1020, 1349 - - dihydroxyacetone reductase 992 - - lipase 414,418 - miehei lipase 413, 418,443,479, 487, 526, 546, 559,566 - racemosus 1025, 1026, 1031 - racemosus kloeckera 307 - sp. lipase 418,425,479 multi - catalytic enzymes 150 - domain proteins 149 - enzymesystem 1288 - substrate enzymes 150 multiple - alignment 143,145 - generations of small libraries 116 muramidase 1500 murine epidermal growth factor 1335 mus musculus 744 mutagenesis 71,123,124, 1067, 1098 mutant - fitness distribution 118 - library 99 - redundancy 99 mutarotase 1517 mutases 897,925, 1281, 1517 mutation rate 118, 119 -

mutator strain 131 rnycobacterium 770 - neoaurum 725,726 - tuberculosis 1241 myeloperoxidase 1262, 1269 myo-inositol derivatives 544 - hexakisphosphate phosphohydrolase 81 myrcene 1073,1092 myrosinase 1503 myxococcusfihs 1268

n NAD hydrolase 639 NAD' 82,83,84,1050 NADH 991,1047, 1290 - dehydrogenases 1111 - oxidases 1111, 1128 - regeneration with formate dehydrogenase 1287 - stereospecifically deuterated 1286 NAD(P)' 82-84, 1108 NAD(P)H 991, 1245 - dependent oxygenase 1203 naphthalene 1100 naproxen 1080 naringine 1384 natural - amino acid esters 412 - diversity 106 - evolution 128 nazlinin 1260 nerol 1069 nerolidol 1092, 1093 nerylacetone 1093 NeuSAc 1452 neuraminidases 636, 1500 neuroligins 148 neutral protease 78 NHAA 1175 nicked proteins 817 nicotiana tabacurn 1037 nicotinamide 1451 - adenine dinucleotide (NAD') 82 - adenine dinucleotide phosphate (NADP') 82 - nucleotides 16 nicotinic acids 1428, 1451 nifknalol 597 nitration 1170, 1187 - ofphenols 1187 nitrilases 700-704, 707, 711, 713, 1513 - agrobacteriurn sp. 1447 - oxynitrilases 200, 975-978, 981, 982, 1514, 1515

Index

nitriles 1447 - hydratases 173,686, 700-702,704,707, 710,712,713,719,1454 - - pseudomonas chlororaphis 1449 - - rhodococcus rodochrous 1450, 1451 nitro 1035 nitrobenzene 1187 nitrogen 486 Nitto Chemical Industry 1451 N-linked 609 NMR spectroscopy 70,75 nocardia 582, 594 nonadienal 1196 non-additive 107 non-convalent adsorption - affinitybinding 167 - chelation 167 - electrostatic binding 166 - ionic adsorption 165, 166 - physical adsorption 165, lt17 non-heme - haloperoxidase 1267 - type 1268 non-homologous recombination methods 110 non-Leloir pathways 609 non-native sugars 1142 non-natural amino acids and ,esters 412, 873 non-ribosomal peptide synthases 150,151 norethisterone acetate 1079 norlaudanosine 1259 norvaline 1441 Novartis 1429 Novo-Nordisk 1438 N-oxides 1036 N+ S acyl transfer - expressed protein ligation 821 nucleases 898,918, 922, 923, 1497 - endonucleases 129,895,922,923 - ribonucleases 825, 835, 856, 857, 859, 918, 923,924,1402,1496 nucleophiles - non-natural 599 nucleoside - monophosphate kinase 61 5 - phosphorylase 638 - phosphate regeneration system 901 - triphosphates 614, 895 nucleotidases 897 nucleotidyl - cyclases 898 - transferases 898 S-nucleotidyl-L-homocys teintt, - production 1291 - total conversion 1291

0

0 2 1111 ochobactrum anthropi 723-725 ocimene 1073 octadiene 1086, 1088 octane 1087 - hydroxylase 1067 octanol 545 octanone 1019 octene 1084-1086 0-demethylation 1067 old yellow enzyme 1037 oleate hydratase - linoleic acid 695 - oleic acid 695 - pamitoleic acid 695 oleic acid 695, 1436 oligonucleotide - cassette mutagenesis 105 - directed codon mutagenesis 125 oligopeptidases 802,810 oligosaccharides 622, 624, 627, 1142 0-linked 609 omapatrilat 882 one base mechanism - for racemization 1285 one pot process 762,777 oninitrilase - safe handling of cyanides 988 operon 159 opposite - enantiopreference 591,593 - regioselectivity 598 optimal mutation rate 103 optimalpH 81 optimization 190 organic - biphasic system 831 - cosolvent 335, 352,360, 370 - media 342 - phasebuffer 284 - polymers 343 organic solvents 128, 203, 204, 259, 342, 346, 348, 351, 398, 413, 473, 474, 487, 503, 526, 545, 570-572, 1005 ornithine 871 ornithine cyclodeaminase 871 orthoester 558 ortholog 157, 159 - search 157 overexpression 1010 ovoperoxidase 1269 oxaloacetate decarboxylase 885 oxamesaconate hydratase

1546

I

Index 4-carboxy-2-oxohexenedioatehydratase 690 oxoprolinases 765 oxazolinones 554,555,558 oxyanion - hole 807 oxidases 26,1108,1170 - acyl-CoA oxidase 1476 - intermediates 338 - alcohol oxidase 1139,1141,1151,1170 oxygen 486 - amine oxidase 1256,1477 oxygenases 73,79,1203 - amino acid oxidase 1254,1477 - lipoxygenase 46,84,1481 - monooxygenase 130,1214,1216,1263 - - galactose oxidase 1474 - - glucose oxidase 1138,1162,1473 - NADH-dependent oxygenase 1203 - toluate dioxygenase 1249 - - glycerol-3-phosphateoxidase 1474 - - glycolate oxidase 1135 oxynitrilases 200,982,1514,1515 - - lactate oxidase 1481 - available for organic synthesis 976 - catalyzed addition of HCN to aldehydes - - trigonopsis variabilis 1426 - ascorbate oxidase 1479 976 - catalyzed addition of HCN to ketones 978 - bilirubin oxidase 1476 - catalyzed biotransformations in aqueous - cholesterol oxidase 1142,1474 medium 981 - choline oxidase 1474 - catalyzed biotransformations in biphasic - cytochrome c oxidase 1129 - diamine oxidase 1259 medium 982 - catalyzed biotransformations in organic me- glucose oxidase 1130,1145 - glutamate oxidase 1297 dium 981 - crystal structure of 975 - glycerophosphate oxidase 1474 - immobilization 977 - glycolate oxidase 1137 - overexpression 976 - lactate oxidase 1256 - recombinant 977 - monoamine oxidase 1256 - substrate acceptance 976 - NADH oxidases 1111,1128,1129 - transhydrocyanation for HCN generation - nucleoside oxidase 1138 - phenol oxidases 1176,1190 982 - used for preparative application 975 - pyranose oxidase 1131,1132 - pyruvate oxidase 1246,1476 oxypressin 1335 oxytocin 856 - sarcosine oxidase 1478 - tyramine oxidase 1477 - ureate oxidase 1478 P P value - xanthine oxidase 1201,1475 oxidations 1133,1134,1262,1264,1425,1427- prediction by S’subsite mapping 828 P-450monooxygenase 130 - alcohol 1425 - ofacids 1245 PAL 870 - 3,Cdihydroxy-trans-caffeic acid 870 - ofalcohols 1108 - dihydroxy-L-phenylalanineammonia lyase - ofaldehydes 1194 palladium complexes 565 - of C-N bonds 1250 palmitic acid 1454 - ofdiols 1121 oxidative palmitoleic acid 1088 pancreas lipase 487 - coupling reactions 1185 pancreatic trypsin inhibitor 841 - deamination 1426 pantenoic acid 1433 oxidoreductases 22,1108,1421,1422,1469, pantolacetone 1433 1479,1481 pantolactone 1454 - cresol oxidoreductase 1188 - ethylphenol oxidoreductase 1189 pantoyl lactone 1000 papain 807,812,834,855, 1353,1508 oxime esters 342,473 - equilibrium-controlled synthesis 830 oxirane acid esters 370 - prediction by S’subsite mapping 829,830 oxobutyrates 1019 papain-like endopeptidases of RNA virus 807 0x0-fatty acids 1199 paptide antibiotics 823 0x0-norleucine acetal 882 oxoperoxidases 1269 paralog 157,158 -

- search 157 parental fitness 118 Parkinson's - disease 761 - syndrome 744 partition 176 - value 827 pathways 159, 160 PCR 68-70,330 - mutagenesis 105 PDB 153 pectic substances 673 pectinase 1500 pectinate 675 pectins 673 - depolymerase 1500 PEG-modified enzymes 834 penicillin 729-732, 735, 775, 1.441 - acylase 855,1337, 1444 - amidase 1438, 1453 - hydroylsis 1438 - penicillin G 1341 - - acylase 1336, 1337, 1341, 1403, 1405, 1407 - - cylase 1378 - penicillin N - - expandase 1308 - - expandase/deacetoxycepharosporin C hydroxlase 1309 - penicillin V 1341 penicillinase 1512 penicilliurn - chrysogenum 307 - roquefortelipase 546 - simplicissimum 1170 pentane 473 pentanediol 478 pentanol 1008 pentanone 1012 PEP 902,904,907 pepsin 1399, 1509 peptidases 800,802,809,817,1505 - amino peptidase 720,723,802,809,1341, 1399 - aspartic peptidase 808, 81;! - ATP-dependent peptidase 815 - carboxypeptidase 10, 11, 197,802,809, 853, 1336,1345,1352,1506 - catalytic mechanism 805 - catalyzed modification 85;' - catalyzed peptide synthesis: 823 - - equilibrium-controlled synthesis 825, 826 - - general manipulations 824 - - kinetically controlled synthesis 826

-

Index

cell-surface peptidase 801 clans 809,811 - classes of peptidases 805 - cysteine peptidase 805, 807, 812 - dipeptidyl-peptidase 809, 1506 - endopeptidase 801,807-810,829,1508 - enteropeptidase 1507 - evolutionary classification 811 - families 809, 811 - glutamyl endopeptidase 1507 - inhibitors - - active site-specific low-molecular-mass inhibitors 816 - - naturally occuring inhibitors 816 - leucyl aminopeptidase 1505 - lysyl endopeptidase 1508 - metallopeptidase 808, 812, 813 - nomenclature 803 - oligopeptidase 802, 810 - peptidyl-Asp methalloendopeptidase 1510 - peptidyl dipeptidase 802,809 - pyroglutamate aminopeptidase 1506 - pyroglutamyl peptidase I 1506 - serine 805,807, 810 - structural probes of conformation of soluble proteins 816 - tripeptidyl peptidase 802,809 peptide - amidase 197,1510 - nucleic acid 1398 - synthesis 197,801 - - chemical peptide synthesis 818,858 - - coupling method 819 - - expressed protein ligation (EPL) 820 - - native chemical ligation 820 peptides 818 - bonds 800,803 - cleavingenzyrnes 801 - glycosylated 1338, 1349, 1352 - hydrolysis 801, 803 - lipidated 1338, 1352 - nucleopeptides 1349, 1352 - phosphorylated 1338,1349,1352 peptidyldipeptidases 802, 809 peptidyltransferase 804, 822 peptidyl-tRNA 822, 842 peracetic acid 1268 perhydralase 1267, 1277 permanent modification 344 permeabilized 37 peroxidases 26,46, 581,1108, 1142,1170, 1185,1262,1264,1267,1269,1352,1479, 1480 - bromo peroxidase 1142, 1264, 1267, 1479

1548

I

Index

chloride peroxidase 1480 chloroperoxidase 581, 1143, 1145, 1262, 1267,1270,1273, 1274, 1480 - eosinophil peroxidase 1269 - flavoperoxidase 1143 - glutathione peroxidase 1142, 1480 - haloperoxidase 1262, 1264, 1268, 1271, 1277 - horseradish peroxidase 122, 1034,1186, 1262 - iodoperoxidase 1267 - lactoperoxidase 1262, 1269, 1480 - lignin peroxidase 1186, 1269 - manganese peroxidase 1143 - ovoperoxidase 1269 - thyroid peroxidase 1269 - soybean peroxidase 1139, 1186 PFAM 156,158,160 Pfizer 1438 PH - dependence 351 - effect 204 - memory 281 - profiles 177 - value 342 phage display 68, 121 pharmacological application 717 phenanthrene 1100 phenanthroline complexes 1130 phenazine 1130,1146 phenolase 1482 phenols 1170,1179,1185,1186 phenothiazine 1130 phenoxy radicals 1170 phenoxypropionic acid 1191 phenyl benzyl oxazolinone 557 phenyl ethanediol 1122 phenyl propanol 1152 phenyl butyl acetate 352 phenylacetaldehyde 1154 - reductase 1033 phenylalanine 866, 870, 1255, 1287, 1441 - ammonialyase 1454 - - cinnamicacid 870 - - coumaric acid 870 - - DOPA 870 - - tyrosine 870 - deaminase 1516 - dehydrogenase 1049 - - cloning 1056 - - heterologous expression 1056 - - sequencing 1056 - - substrate specificity 1054 - - synthesis of allysine ethylene acetal 1056 -

deuterated 1290 isopropylester 1454 - lyases 866 - production 1287 - synthesis with glutamate racemase 1296 phenylcatechol 1180 phenylethanol 565, 1152 phenylethylamine 1435, 1454 phenylethylmethoxyamide 1436 phenylglycidate - inactivator of mandelate racemase 1312 phenylglycine 1439 - deuterated 1290 phenylmethanesulphonylfluoride 816 phenyloxazolinone 473 phenylphenol 1180 phenylpropanoic acid 1154 phenylpyruvic acid 1255 phenylthioethanol 1152 phosphatases 26, 897,918 - acid phosphatase 918,919,921,1496 - alkaline phosphatase 897, 908, 918-921, 1495 - phosphate phosphatase 918 - pyrophosphat phosphatase 909,919,920, 1513 phosphatidylcholine 2-acylhydrolase 1493 phosphodiesterases 898,918,923-925,1496 phosphoenolpyruvate 614,902 - hydratase 692 phosphoesterase 158 phosphofructokinase 1319 phosphoglucomutase 925, 1517 phosphoglucosamine transacetylase 1483 phosphoglucose - isomerase 1319 - - reaction mechanism 1318 - - thermostable enzyme 1318 - X-ray structure 1318 - substituted 1319 phosphohydrolases 898,918, 919 phosphokinases 897 phospholipases 146, 147,898, 1493, 1496 phosphomutases 897,925 phosphopeptide 1349 phosphoresters 896 phosphorous-containing racemic esters 383 phosphoryl transfer 895,896 phosphorylases 612,638,897,901,923,1483 phosphorylating - agents 899,900,910 - enzymes 895 phosphorylations 896,897, 901,907, 918, 920, 1246 -

~

phosphotransacetylases 1483 phosphotransferases 897 photochemical regeneration 1114 photoelectrochemical 1039 photosensitizers 1114 photosynthetic microorganism 994, 1037 phthalyl amidase 1336 phylogenetic 155 - relationships 158 physical 171 phytase 81,1496 pichia pastoris 1151 Pictet-Spengler reaction 1403 pig liver 397 - esterase 46, 335, 336, 342-344, 346, 351, 353, 360,361, 365,366,369-371, 383,384, 398,407,425,472,571,572,840, 1377 pig pancreas 434 - lipase 46, 335, 346,413,414,417,418,426, 428,435,474,479,486,503, 526, 546, 555, 559 pinene 1073 piperidine 1080 - derivatives 425 PK 904,907 pK,value 81 planar chiral - [2.2]paracyclophane 458 - racemic ester 370 planar chiralities 1033 plant - cellcultures 1037 - enzymes 46,48 P U P 352 plasma-atomicemission spectrometry 785 plasmid 70, 71 polyacrylamide gel 1450 polycyclic 1100 polygalacturonases 678, 1501) - endopolygalacturonases 679 - exopolygalacturonases 679 polyghtamate 1294 polyketid 130 - synthases 150, 151 polymerase chain reaction (I’CR) 98 polymerases 71, 72, 79, 329, 895 polymer-modified NAD 1163 polynucleotide 923 - synthetases 898 polyo1 1142 - dehydrogenase 1126 polyphenolase 1176 polysaccharides 622 pooling strategy 117

Index

porcine insulin 1337 position of chemical equilibrium 276 precipitant 72 precipitation 62 - by changing pH 63 - by organic solvents 63 - by salting out 62 - by water-soluble polymer 63 preference 350 prejudice 1453 prenyl diphosphate synthase 85 pre-screening 117 pressure 1002, 1009 - swingreactor 777 PRINTS 156 propanediol derivatives 478 PROCARD 156 process - conditions 192 - design 186 - optimization 186 processing - proteolytic processing 814 prochiral 416,473 - acyclic 426 - - dicarboxylic acid diesters 361 - - diol diacetates 369,414 - - diols 474 - acylated diols 344, 370 - alcohols 571 - anhydrides 473 - cyclic - - dicarboxylic acid esters 353, 399 - - diol diacetates 366, 406 - - diol dialkanoates 418 - - diols in organic solvents 479 - diacetates 407,473 - dialkanoate 416 - diesters 554 - diketones 458 - diols 344, 346, 414, 473 - esters 408 - glutarates 365 - glutaric anhydrides 557 - malonates 365, 398 - substrates 343 product - inhibition 344 - specificity 85 production of enzymes 41 productivity 185 profiles 143, 144, 154, 158, 160 - method 142 progesterone 1066, 1067

1550

I

lndex

prokaryotic cell 1420 proleather 1354 prolidase 1506 proline 871, 1256 - competitive production 1301 - racemase - - free energy profile 1301 - - properties 1301 - - purification 1301 - - reaction mechanism 1301 - - recombinant enzyme of clostridium stick landii 1301 - reductase 1301 - rule 89 prolyl - endopeptidase - - prediction by S’subsite mapping 829 - oligopeptidase 810 PROMISE 154 promoter, T7 69 pronase 1505 propanediol - dehydratase 692 - derivatives 417 propanol 545,994,1003, 1017, 1152, 1154 propargylic 525 - alcohols 1145 propene 1090 propenylphosphonate 1090 properties 171, 172 - benefits 163 - limitations 163, 164 - of immobilized biocatalysts 175 propionases 767, 770,777 propionylphosphate 905 propylene oxide 1134 prosite 155, 156, 158, 160 prostaglandin 1101 prosthetic - centers 154 - groups 584 proteases 26,46, 78, 149, 801, 840, 847, 848, 852,1379, 1503 - alcaIase@ 1431 - akaline protease 1346 - aspartic protease 326 - cysteine protease 326 - endopeptidase 326 - exopeptidase 326 - heat-stable protease 327 - metal protease 147, 148, 326 - serine protease 326, 398,407, 806 - subtilisin Carlsberg 1431 proteasome

- 20s proteasome 815 26Sproteasome 815 PA700 regulatory complex 815 protecting group 365 protection - of amino groups - - acetoxy-benzyloxycarbonyl 1338 - - Arg-OH 1334 - - benzoylphenylalanine 1334 - - phenylacetylbenzyloxycarbonyl 1338 - - phenylacetamide 1403, 1405 - - phenylacetic acid amides 1336, 1341, 1343 - - phthalyl imide 1343 - - pyroglutamyl amides 1341 - - tetra-O-acetyl-galactopyranosyloxycarbonyl 1339 - - tetra-0-acetyl-glucopyranosyloxycarbonyl 1339 - of carboxy groups 1344 - - allylesters 1353 - - benzyl esters 1346 - - butyl esters 1345, 1350 - - bromoethyl esters 1348 - - choline esters 1352 - - cyclopentylesters 1353 - - ethyl esters 1345, 1346 - - heptyl esters 1348 - - methoxyethoxyethylesters 1351 - - methoxyethyl esters 1351 - - methyl esters 1345, 1346, 1350 - - nitrobenzyl esters 1348, 1350 - - phenylhydrazide 1352 - of hydroxy groups 1353 - - acetyl esters 1370,1371, 1375 - - butanoyl esters 1387 - - butyrylesters 1375 - - octanoyl esters 1370 - - pentanoyl esters 1370, 1371 - of thiol groups - - phenylacetamidomethyl 1343 protein 4 - aggregates 342 - comparison 140 - crystallization 68, 70, 75 - engineering 67, 76, 89, 90 - family 143 - peptidebonds 800 - scaffold 88 - structure 73, 74 proteinases 801, 504, 1508 - endoproteinase 830,1506-1510 - inhibitor 816 proteolysis 804, 813, 816 -

-

proteolytic enzymes 154,800,801 protoheme 1142 protonation state 281 protopectin 675 protopectinases 676 protoporphyrin IX 1142 pmnus amygdalus 976 pseudocrystal - enzymation 1451 - process 1254 pseudomonas 582,735,754,756, 784, 1158 - aemghosa 442,728,1205, '1241 - aemginosa lipase 347,435,443 - cepacia 289-298, 413 - cepacia lipase 347, 349,418,428,435, 442, 443,458,459, 474, 479, 487, 503, 526, 544-546,555,559,566 - chlororaphis 707,709, 710, 712 - chlororaphis B23 700, 711, '712 - desmolyticum 774 - fluorescens 291,413,719, 727, 774, 778, 1268 - fluorescens lipase 294,295, 347,414, 417, 418,425,428,443,459,474,478, 479, 486, 487, 503, 526, 546, 559, 566 - lipases 303 - NCIMB9872 1232 - putida 701,705,722,723, 770,775, 780, 782,788,789,1209,1210, 1227,1234 - putida ATCC 17453 1216,1224, 1229, 1232, 1238 - solanacearum 727 - SP. 710, 718, 752,754, 774-, 778, 780, 782, 783,788,789,794,994 - sp. alcohol dehydrogenase 992 - sp. hpase 335,418,426,428,434,435,443, 459,474,479,487, 503, 526, 546, 554, 559, 565,566 - Sp. NCIMB 9872 1207,1228 - sp. strain PED 1017 - striata 774,778 -

testosteroni 996

PSI-BLAST 74 psicose - conversion into allose 1322 psychrophiles 313 pulegone 1072 pullulan 657 - hydrolases - - type1 315,659 - - type I1 316,659 - - type 111 659 pulldanases 1503 - type1 315,657

type I1 315,658 pulp 673 purification 122 - ofenzymes 49 purine-nucleoside phosphorylase 1484 putidaredoxine 1199 putrescine 1259 pyridoxal phosphate 19, 1283 pyroglutamate aminopeptidase 1341 pyrophosphatases 909,919, 1513 pyrophosphate 920 pyrophosphokinases 898 pyrophosphorylases 612,615-618,909 pyrrolecarboxylate - competitive inhibitor of proline racemase 1301 pyrrolidine 1080 pyrrolonitrin 1187 pyrroloquinohe quinone 991, 1038, 1141, 1145,1146 pyruvate 1245 - kinase 614, 902, 907,909, 921 - oxidase 1246 -

9 quasi-species theory 102, 103 quaternary - carbon 442 - structure 785 quinazoline 1101 quinoline 1101 quinones 1176 - methide 1171, 1189 quinoxaline 1101 r racemases 758,763,765,766,771,791,792, 1281,1282 - achromobacter obae 1442 - alanine racemase 1284, 1285 - amino acid racemase 1283,1289, 1293, 1306-1308,1440 - amino caprolactam racemase 1292, - arginine racemase 1290 - aspartate racemase 1297, 1298 - glutamate racemase 1293, 1295, 1296 - hydantoin racemase 1303-1305 - mandelate racemase 1310-1312 - p r o h racemase 1301 racemates 336, 351, 383 - resolution 1251 - separation 412 racemic - acetates 384, 398, 473

1552

I

Index

- acyclic alcohols 487, 503

acylated alcohols 473 - alcohols 336, 416,473, 570-572 - alkanoate 416 - carboxylic acid esters 336, 371, 399, 428 - carboxylic acids 343,473 - esters 408 - ketones 458 racemization 287, 288, 290, 291,293, 294, 297,298, 301, 309,412, 544, 557, 558, 565, 723,727,792,794,1439,1441 - by one-base mechanism 1285 - by two-base mechanism 1285 RAMA 932,934-936 random - mutagenesis 68, 123, 1099 - point mutagenesis 99, 105 - pointmutations 108 randomized oligonucleotides 105 random-priming method 108 Raney copper 711 rare codons 69 rat liver imidase 1336 rate constants 344 rational design 67 reactions - conditions 203 - consecutive 195 - engineering 185 - kinetics 23, 189 - mechanism 991 - parallel 195 reactivity 417, 473, 569 reactors - batch reactor 232 - continuous stirred tank reactor 233 - fixed bed reactor 250 - flow reactor 233 - fluidifizedbed 250 - immobilized enzymes 250 - massbalance 234 - recirculation reactor 250, 1438 - stirred tank reactor 232 recognition 150 recombinant - cells 112, 1011 - DNA technology 819,820,855 - enzymes 795 - gene expression system 69 - genetechnology 68 - pig liver esterase 352 - proteins 69 recombination 107 -

efficiency 108 familyshuffling 100 - singlegene 100 Recordati 1442 - process 776 recovery 342 recursive PCR 105 red yeasts 584 redox - dyes 1130,1142 - mediators 1146 reductases - acetoin reductase 1129 - aldehyde reductase 1010, 1033 - diacetyl reductase 1028, 1129 - dihydrofolte reductase 1478 - dihydroxyacetone reductase 992 - diketone reductase 1470 - flavin reductase 1268 - FMNreductase 1111 - glutathion reductase 82, 83, 1236, 1478 - ketoreductase 1010, 1469 - lipoamide NADH reductase 1478 - oxidoreductase 22, 1108, 1188, 1189, 1421, 1422, 1469, 1479, 1481 - phenylacetaldehyde reductase 1033 - proline reductase 1301 reductions 991 - enantioselective 1424 - equivalents 1185 - ketoester 1423 - ketone reduction 1422 - of C=N bonds 1047 - OfNAD' 1247 reductive amination 1048 - thermodynamic limitation 1050 regeneration 1014, 1108 - coenzyme 992,1245 - electrochemical 995 - enzyme 1126 - hydrogenases 994 - OfATP 902,909, 1246 - OfNADH 1184,1247 - OfNADP' 1122 - of NAD(P)H 993 - of nucleoside triphosphates 901 - system 909 regiospecificity 84, 85, 89 regular expression 155 Reichstein-Grussner synthesis 1425 relationship with water activity 267 rennin 1509 repetitive batch 1450 replacement of chloroalanine 966 -

replacing chemical steps by bicitransformations - industrial application 1419 residence time 232 residual water level 264 resolution 351, 383, 565, 1255 - of racemates - - oxynitrilase as catalyst 982 resting cells 37, 777 retinal 1194 retropepsin 812 reverse - hydrolysis 346 - micelles 343, 833 - phase chromatography 58 - transcriptases 895 rhamnose 673,1384 - aldolase 939 - isomerase 939,941 - - genecloning 1321 - - immobilization 1322 - - metal-mediated hydride-shift mechanism 1322 rhamnulose 1321 rhizomucor - jauanicus lipase 474 - mieheilipase 413 - sp.lipase 503 rhizopus - arrhizus 1031 - delemarlipase 418 - niueus 1348 - oryzaelipase 443 - sp.lipase 418 rhodobacter sphaeroides 1034 rhodococcus 582, 591, 594, 710 - butanica 704 - equi 705,727,728 - erythropolis 728, 993, 1016, 1148 - rhodochrous 704-708 - rhodochrous J l 701,709,712,713 - SP. 700,701,705,710,1207 - sp. N-774 711,712 rhodosporidium toruloides 1029 rhodotorula 588 - glutinis 310, 592, 593, 1031 - minuta I F 0 0920 1000 - mucillaginosa 1025, 1026 - pilimanae 1029 - rubra 752 riboflavin 1383 ribonucleases 825, 918 - ligase 895 - polymerase 329

- ribonuclease A 835,856,857,859,923, 1402 - ribonuclease T1 924, 1496 - ribonuclease T2 923,924 ribonucleotide phosphohydrolase 919 ribosome 822,823 - display 121 ribozyme 804,823 ribulose 1,s-bisphosphatecarboxylase/oxygenase 73,79 Roehm GmbH 1446 rubisco 72 rugged landscapes 119 ruthenium catalysts 565 rutin 1383 rutinose 1379 S

saccharase 1502 saccharification 661 saccharomycescerevisiae 124, 307, 794,1002, 1009, 1022,1027,1241 safety-catchlinker 1405 salicin 1383 salt hydrates 269 saltingout chromatography 62 santonin 1075 saturated salt solution 266 saturation mutagenesis 104, 131 schizosaccharomycespombe 794 sdareol 1076 screen 101 screening 104, 109, 112, 123, 1000 - sets 1463 SDS-PAGE 72 search engine 155 secretory proteins 814 selection 112, 119, 123, 130 - methods 101 selective cleavage 728 selectivity 196, 344,417,473, 569,634 seleno subtilisin 835 selenomethionine - production 1287 sequence - alignment 74, 81-83, 85, 90, 106 - analysis 140 - comparison 140,159 - region 143 - similarity 140 - space 97, 106 sequence-structure-functionrelationships 122 sequential kinetic resolution 397 serine

1554

I

Index

- deaminase 871 - - 2-oxobuturate 871 - - pyruvate 871 - dehydatase 871 - hydrolase 337 - peptidase 807 - - catalytic mechanism 805 - - clan 810 - -family 810 - protease 398,407 - - catalytic mechanism 806 - residue 337 serratia marcensens lipase 487 sertraline 1025 serum albumin 359 1,I-shift - by epimerase 1282 - by racemase 1282 1,2-shift

- by aldose-ketose isomerase 1282 1,3-shift - for allylic isomerization 1282 shikimic 1102 sialic acid 1324 - aldolase 618 sialidase 1500 sialyltransferase (SiaT) 623 silapropanediol derivatives 478 silica 565 - gel 359 single - gene 100 - turnover events 120 site - entropies 106 - tolerances 120 site-directed 71 - mutagenesis 68,69,75, 81, 85, 87, 105, 335,407 sitting drop method 72 sliding window 845 small-scale 342 SMART 156 smart polymers 181 SNAM-Progetti 1442 - process 776 sodium - chloride 349 - cyanide 1086 sol-gel - gel entrapment 181, 343 - lipase 181 - materials 413, 473 - poly(akylsi1oxanes) 181

- poly(hydroxymethylsi1oxanes) 181 solid - biocatalyst 260 - fermentation 47 - statebuffers 284 - support 343 - - bio-beads 1399 - - controlled pore glass 1343 - - kieselguhr-PDMA-resins 1402 - - MPEG-support 1402 - - novasyn KA 1399 - - pepsyn K (polyacrylamide)resin 1398 - - POE 6000 1406 - - SPOCC-resin 1399 - - TentaGel beads 1400 - - TentaGelS beads 1403 solid-phase - peptide synthesis 818,819,852 - synthesis 835 solid-to-solidconversion 854 solketal 1147 solubility 341 solubilized enzymes 263 solubilizing protecting groups 832,852,854 - medium engineering with organic solvents 833 solvents 546 - choice 276 - engineering 417 - hydrophobicity 279 - parameters 279 - polarity 277, 279 somatostatin 856 Sonogashira reaction 1407 sorbiton dehydrogenase 1454 sorbose derivatives - modified at position 5 1317 sorghum bicolor 976 soybean peroxidase 11 39 space-timeyield 196 specificity 103,129,1060 sphingomyelin phosphodiesterase 1496 sphingomyelinase 1496 spiked oligonucleotides 105 spimerases - amino acid spimerase 1283 spirobicyclic amides 1081 spontaneous mutations 124 sporobolomyces - salmonicolor 1027, 1033 - salmonicolor AKU4429 1010 stability 177, 342, 571 stabilization during purification 64 staggered extension process 108

Index

stannylated derivatives 525 staphylococcus epidermidis 1015 starch 653, 1433 - processing enzymes 315 statistical distribution of mutation 101 Stemmer method 108 stemodine 1077 stereochemical control 997 stereochemistry 992, 1127 stereoinversions 288, 1127, 1157, 1256 stereospecific aldol condensation 1317 stereospecificity 84 steroid synthesis 965 steroids 695, 1067, 1127, 1202 stop codon 102 storage of enzymes 64 Strecker reaction 725, 761 streptavidin 1400 streptidine kinase 1486 streptogrisin A 810 streptokinase 1486 streptomyces 755, 756, 1022 - coelicolor 744 - griseus 298 - sp. 754 - strain 1027 streptoverticillium 755 string 159 structural - genomics 140, 160 - tolerant protein 106 structure 6 - comparison 140,141 - units 110 structure-function studies 108 styrene 1200 styrene - anion exchange resin 1436 - oxide 588 substance P 855-857 substituted hydantoin derivatives - precursors for amino acids 1303, 1306 substitution matrix 141, 143 substrate 101 - bindingenergy 76 - concentrations 201, 274, 1005 - mimetics 842, 843, 850, 851 - - anionic substrate mimetics 847 - - cationic substrate mimetics 845 - - hydrophobic substrate mimetics 849 - - kinetic model 844 - - steady-state kinetic 843 - mimetic-mediated synthesis - - clostripain-catalyzed coupling 846, 847

- - fragment condensation 845,846,848, 851,852 - - V8-protease-catalyzed 848 - models 346 - modification 458 - solvation 275 - specificity 68,84,89, 351 subtiligase 835, 856 subtilisin 86, 298, 335, 337, 342, 347,407, 408,472,570,571,817,835,857,1346, 1350,1354,1368, 1378,1379,1381,1383, 1384,1399,1454, 1508 succinases 768,787 succinates 365 succinic acid anhydride 473,486 sucrose 1379 - derivatives 1317 - isomerase 1281 - nucleoside diphosphate 613 - synthase 1317, 1483 - synthetase 625, 909 suicide inhibitors 121 sulcaton N-phenylcarbamate 1095 sulfides 1034,1262-1266 sulfoxidation 1263, 1264 sulfoxides 1034, 1262, 1266 sulfur - compounds 1004 - containing racemic esters 383 - functionalities 486, 1024 sulphenyl ketones 1024 supercritical - carbon dioxide 1006 - fluids 278,412 - solvents 1006 superfamily 1050 superoxide dismutase 1482 surface 343 - saltbridges 127 Suzuki reaction 1407 swinging arm 150 SWISS-PROT 152 synechococcus sp. 994 synergism 666 synthesis 858 - in frozen-aqueous systems 833 synthetases 947 - acyl-CoA synthetase 1517 - CMP-Neu5Acsynthetase 618, 1486 - DAHP synthetase 947 - diphosphate synthetase 85 - fatty acid synthetase 1012 - glutamate synthetase 1293 - glutamin synthetase 1518

1556

I

Index

- isopenicillin synthetase 1308 KDO 8-P synthetase 947 - lactose synthetase 1483 - NAD(+) synthetase 1518 - phosphate synthetase 946 - polynucleotide synthetase 898 - sucrose synthetase 625, 909, 1317, 1483 - tryptophan synthetase 1290 - tyrosyl-tRNA synthetase 76, 77 synthetic - applications 1222 - oligonucleotides 69 -

t o-tagatose 3-epimerase 1322 Tanabe Siyaku 1430 tannase 1495 targeted random mutagenesis 125 tartaric acid 686 - dehydratase 686 tautomerase 1281 TBADH 1033,1196 TDP aldolase 942 temperature 28, 274, 1008 template matching method, 3D 74 temporary 344 terpene 1092 terpinolene 1072 terpyridine complexes 1130 testosterone 1158

tetracyano-iron-l,l0-phenanthroline 1142 tetracyanoquinodimethane (TCNQ) 1130 tetracyclic alcohols 545 tetrahedral intermediates 806,807 tetrahydrofuran 342,473, 545 tetrahydropyran derivatives 425 tetraphenylporphyrin 544 tetrathiofulvalene (TTF) 1130 th. themophilus 124 thermal stability 157 thermitase 1347, 1349, 1353 themoactionomyces vulgaris 1347 therrnoanaerobacter ethanolicus 996,997, 1008, 1010,1012 thermoanaerobium brockii 994,996,1010, 1018,1109,1195,1231 - alcohol dehydrogenase 1120 themococcus - kodakaraensis 79, 80 - kodakaraensis DODl 73 thermodynamic equilibrium 193 thermodynamics 748 thermolysin 78, 472, 813, 834, 1345, 1510 - bacillus proteolicus 1446

therrnophiles 313 therrnophilic organisms 157 thermostabile - DNA polymerase 71 - proteins 67 thermostability 78-81, 89,90, 127,473, 996, 1120 thiacrown 571 ethers 569 thiamine pyrophosphate 19 thioesters 412,442, 565 thioglucosidase 1503 thiol subtilisin 835 thio-NAD 1147, 1163 thionin 1130 three-dimensional - models 75 - protein structures 67 - structure 70, 75, 79, 81, 82, 87,89, 139 threonine - aldolases 953, 1514 - deaminase 871 thrombin 1400, 1507 - inhibitors 873 throughput 113 thymidine 1321 thyroid - galactosyltransferase 1483 - peroxidase 1269 tiered screening 118 tight-bind inhibitor - of alanine racemase 1285 tolerant 116 toluate-l,2-dioxygenase 1249 toluene 473, 545, 571, 1099 toluic acid 1100 topaquinone 1259 Toray Industries 1443 total turnover number 238, 1453 Toyo Jozo 1436 TPL 870 Transacetylases 1483 transaldolases 962, 1482 transaminases 874,886,1047,1484,1260 - a-transaminases - - 2-aminobutane served 880 - - enantiomerically pure arnines 880 - - isopropylamine transaminase 880 - - ketoglutarate transaminase 881 - - phenylethylamine transaminase 880 - - phenylisoproylamine transaminase 880 - (R)-transaminase 877 - (5’-transaminase 877 glutamate-pyruvate transaminase 1484 ~

index ghtamic-oxaloacetic transam inase 1484 triethylamine 558,570 transamination 874 trifluoro esters 342 - decarboxylation spontaneous 884 trifluoro(2-thieny1)ethanone 1025 - equilibrium constant 884 trifluoromethyl ketones 1022 - oxaloacetate 884 triglycerides 417 transcriptases 895 trimethylsilylalanine transcyanation 978 - synthesis from D,r-5-trimethylsilylmethyltransesterification 336,342,343,349,571, hydantoin 1305 574,717 trioctylamine 565 transferases 22,150,625, 1484 triosephosphate isomerase 1320 - acetylglucosaminyl-glycopeptidegalactosyltripeptidyl peptidases 802,809 transferase 1483 triphosphohydrolases 898 - N-acetyllactosaminidea-1,3-galactosyltranTrusopt@ 1422 serase 1484 trypsin 802,831,8429345,849,852, 1334, - acylneuraminate cytidylyltransferase 1486 1336,1345,1352 - amino acid transferase 889 - equilibrium-controlled synthesis 830 - aminotransferase 130,874,875,881,882, - prediction by S’subsite mapping 830 1484 trypsinogen 841 - dipeptidyl transferase 1506 tryptophan 1441 - fucosyltransferase 624 - halogenase 1268 - galactosyl transferase 615,619,1483,1484 - combination production from N-acetyl-D,r- glucosamine-phosphate N-acetyltransferase tryptophan 1307 1483 - production with tryptophan synthase 1290 - glucosidase transferase 639 tryptophanase 1516 - glycosyltransferases 316,611, 612, turnover frequency 1453 627-629,631,639,654,659 909,1394 two-base mechanism - ghtamykransferase 609,1483 - for racemization 1285 - methyltransferases 1482 two-hybrid systems 68 - nucleotidyl transferase 898 two-liquid phase systems 1117,1118 - peptidyl transferase 804,822 two-phase - polyribonuceotide nucleotidyltransferase - reactor 1431 1486 - systems 252,413,425,473 - sialyltransferase 623 tyrosinase 1176,1352,1482 - uridyltransferase 616 tyrosine 1176 transferrin receptor 148 - phenollyase 870 transformation 121 - - enuinia herbicola 1448 - protocols 112 - production 1287 transglucosidase 1483 - synthesis with glutamate racemase 1296 transhydrocyanation 978 P-tyrosinase 1516 - principle of 980 tyrosyl-tRNAsynthetase 76, 77 - procedureof 982 transhydrogenase 993 U transition metal UBA 147 - combination 293 UDP 616 - complexes 365 - N-acetylgalactosamine 616 transition state analogs 120 - N-acetylmuramoyl-r-alanine transketolase 960,1482 - - activator of glutamate racemase 1293 transpeptidases 1483 - N-acetylmuramoyl-r-alanyl-D-glutamate syntriad 338,413 thetase 1293 trichloroethyl - gaIactose 615 - acetate 570 - galactose pyrophosphorylase 615 - esters 342 - galactosylphosphate uridyltransferase tricyclic alcohols 545 616 triethyl citrate 369 - glucose 615 -

1558

I

Index

- glucose dehydrogenase 618 - glucose pyrophosphorylase 616,909 - glucuronic acid 618 ulocladium atrum 582 ultrafiltration 49, 239, 342, 352, 1447 UM-BBD 153 uncoupling reaction 1181 unfolding temperature 81 Unifar 1438 unsaturated fatty acid cis-trans isomerase - purification 1325 unusual sugar derivative - production 1316 urea - amidolyase 1518 - carboxylase 1518 urease 765, 1511 ureidopropionases 767, 770,777 ureidosuccinases 768,787 uricase 766,1478 uridine-4-diphosphoglucose dehydrogenase 1126 UTP 616 Y

V8 protease 840,847,848,852

valencene 1096 E-valerolactones 554 valine 1441 vanadium 1142,1267 - bromoperoxidase 1264 vanillin 1174, 1191 vanillylamine 1174, 1260 vanlev - 6-hydroxy-~-norleucine1054 vasopressin 856 vic-diols 1119 vinyl 478 - acetates 342, 343, 346, 348, 473,478, 486, 502,544, 545,565, 571 - butyrate 473 - carboxylates 558 - esters 342, 343, 571 - laurate 473 - palmitate 473 - propionate 473, 574 vinylogous a-hydroxy carboxylic acids 442 viologene 1113 vitamin B3 1451 vitamin B6-dependent enzymes 875

W

water - activity 205, 265 - budget 271 - concentration 267 - control via vapor phase 272 - mimics 273 - filed porous supports 571 - immiscible solvents 338, 412 - in-oil emulsions 121 - miscible 338 wheat germ lipase 1379 whole cell 1442, 1447 - achromobacter obae 1443 - achromobacter xylosoxidans 1428 - candida sorbophila 1424 - catalyst 708 - cluconobacter suboxydans 1425 - crytococcus laurentii 1443 - ertuinia herbicola 1448 - neurospora crassa 1422 - oxidation 1148 - process 1419 - pseudomonas chlororaphis 1449 - gseudomonasputida 1439 - zygosaccharomyces rouxii 1420 wursters blue 1146 X

XAD-7 resin 1420 ranthobacter agilis 1336 X-ray - crystal structure 347, 348 - crystallography 70 - diffraction 70, 75 xylan 667 - degrading enzymes 324 xylanases 324 xylose 324,1132,1313 xylose isomerase - by site-directed mutagenesis enhancement of thermostability 1316 - divalent metal cation 1314 - from various microorganisms 1316 - gene cloning 1313 - immobilization 1316 - properties 1313 - reaction mechanism 1314 - thermostable enzyme from thermophile 1314 - three dimensional structure 1313 xylosidases 670 xylulose, 1313

Index

Y

Z

YADH 1109 yarrowia Iipolytica 1002, 1009 yeast 125, 591, 1022, 1023, 1027, 1035, 1036,

(-)-zeylena 1102 zinc-fingers 146

1109,1120

alcohol dehydrogenase 992 yield 196 -

zoogloea ramigera 753 zopfiella karachiensis 582 zygosaccharomycesrouxii 1007, 1010

zymogens 801