Enzyme Catalysis in Organic Synthesis
Edited by K. Drauz and H. Waldmann
Second Edition
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Enzyme Catalysis in Organic Synthesis
Edited by K. Drauz and H. Waldmann
Second Edition
Related Titles from Wiley-VCH B. Cornils, W. A. Herrmann (Eds.)
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
D. E. DeVos, I. F. J. Vankelecom, P.A. Jacobs
Chiral Catalysts Immobilization and Recycling 2001, I S B N 3-527-19295-2
U. Th. Bornscheuer, R. J . Kazlauskas
Hydrolases in Organic Synthesis Regio- and Stereoselective Biotransformations 1999, I S B N 3-527-30104-6
R. A. Sheldon, H. van Bekkum
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
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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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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
I
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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12.1 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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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: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
I
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.
I
65
66 2 Production and Isolation of Enzymes
I
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
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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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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.
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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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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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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
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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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3 Rational Design ofFunGtiona1 Proteins
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
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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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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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3 Rational Design of Functional Proteins
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
90
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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.
References
K. M. Ulmer, Protein Engineering, Science 1983,219(4585),666-671. z K. 0. Stetter, Extremophiles and their adaptation to hot environments, FEBS Lett. 1999,452(1-2), 22-25. 3 S. Fujiwara, M. Takagi, T. Imanaka, Archaeon Pyrococcus kodakaraensis KOD1: ap1
plication and evolution, Biotechnol. Annu. Rev. 1998,4, 259-284. 4 W. P. Stemmer, Rapid evolution of a protein in vitro by DNA shuffling, Nature 1994, 370(6488), 389-391. 5 G. P. Smith, Filamentous fusion phage: novel expression vectors that display cloned
References I91 antigens on the virion surface. Science 1985, 228(4705),1315-1317. 6 T. Murai, M. Ueda, M. Yamamura, H. Atomi, Y. Shibasaki, N. Kamasawa, M. Osumi, T. Amachi, A. Tanaka, Construction of a starch-utilizing yeast by cell surface engineering, Appl. Environ. Microbiol. 1997,63(4),1362-1366. 7 A. D. Napper, S. J. Benkovic, A. Tramontano, R. A. Lerner, A stereospecific cyclization catalyzed by an antibody, !:cience 1987, 237(4818),1041-1043. 8 S. Fields, 0. Song, A novel genetic system to detect protein-protein interactions, Nature 1989, 340(6230), 245-246. 9 R. W. Roberts, J. W. Szostak, RNA-peptide fusions for the in vitro selection of peptides and proteins, Proc. Natl. Acad. Sci. USA 1997.94(23),12297-12302. 10 J. A. Kolkman, W. P. Stemmer, Directed evolution of proteins by exon shuffling, Nat. Biotechnol. 2001, 19(5),423-428. 11 U.T. Bornscheuer, M. Pohl, Improved biocatalysts by directed evolution and rational Biol. protein design, Cum. Opin. Ch200-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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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
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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
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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
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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
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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.
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References 1 J. R. Knowle:;, Nature, 1991, 3513, 121-124. 2 H. Cid, Bid. Res. 1996,29, 101--126. 3 R. Bott, R. Boelens, Curr. Opin Biotechnol.,
1999, 10, 391-397. S.K. Burley, Nat. Struct. Bid. Suppl., 2000, ' 932-934. 5 W. G Hol, Nat. Struct. Bid. Suppl., 2000, 7, 964-966. 6 A. R. Williamson, Nat. Struct. 13iol. Suppl., 2000,7,953. 7 S. Dry, S. McCarthy, T. Harris, Nat. Struct. Bid. Suppl., 2000,7, 946949. 8 K. Hofmann, D. Bucher, A. V. Kajava, ]. Mol. Biol., 1998, 282, 195-208. 9 J.F. Gibrat, 'TMadej, S. H. Bryant, Curr. Opin. Struct. Biol., 1996,6, 377-385. 10 P. Bork, T.J. Gibson, Methods Enzymol., 1996,266,162-184. 11 K. Hofmann, Brief: Bioinformat., 2000,1 , 167-178. 12 S. R. Eddy, Bioinformatics, 19911, 14, 755-763. 13 K. Karplus, C. Barrett, R. Hughey, Bioinformatics, 1998, 14, 846-856. 14 T. F. Smith, M. S. Waterman, J. M d . Biol., 1981, 147,195-197. 15 S. Henikoff, CUT. Opin. Strud. Biol., 1996, 6,353-360. 16 M. l'ingron, M. S. Waterman, J. Mol. Biol., 1994,235,l-12. 17 M. 0. DayhlDff, R. M. Schwart:c, B. C. Orcutt in: M. Atlas ofprotein Sequena: and Structure, (Ed.: hl. v. DayhoJ), Vol. 5, National Biornedical Research Foundation, Washington DC 1978, pp. 345-352. 18 S. Henikoff; J.G. Henikoff, Pim. Natl. Acad. Sci. U S A, 1992,89,10915-10919. 19 S. H.enikoff; J.G. Henikoff, Proteins, 1993, 17,49-61. 20 G. Vogt, T. Etzold, P. Argos, ] Mol. Biol., 1995, 249, 816-831. 21 M. Gribskov, A. D. McLachlari, D. Eisenberg, Proc. Natl. Acad. Sci. U :jA, 1987,84, 4355-4358. 22 R. Luthy, I. Xenarios, P. Bucher, Protein Sci., 1994, 3, 13!)-146. 23 E. Birney, J . D. Thompson, T. J. Gibson, Nucleic Acids Res., 1996, 24, 2?30-2739. 24 J. D. Thompson, D. G. Higgins, T. J. Gibson, Comput.Appl. Biosci., 19!)4, 10, 19-29. 25 P. Bucher, K. Karplus, N. Moeri, K. Hofmann, Comput. Chem., 1996, 20, 3-23. 4
26 W. R. Pearson, Methods Mol. B i d , 2000,
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27 S. F. Altschul, W. Gish, W. Miller, E. W.
Myers, D. J. Lipman, 1.Mol. Biol., 1990,215, 403-410. 28 S. F. Altschul, T. L. Madden, A. A. Schaffer, J. Zhang, Z. Zhang, W. Miller, D. J. Lipman, Nucleic Acids Res., 1997, 25, 3389-3402. 29 M. Waite, Biochim Biophys Ada, 1999, 1439, 187-197. 30 K. Degtyarenko, Bioinformatics, 2000, 1G, 851-864. 31 C. M. Lawrence, S. Ray, M. Babyonyshev, R. Galluser, D. W. Borhani, S. C. Harrison, Science, 1999,286,779-782. 32 K. Ichtchenko, Y. Hata, T. Nguyen, B. U11rich, M. Missler, C. Moomaw, T. C. Sudhof, Cell, 1995, 81, 435-443. 33 G. Muller-Newen, W. Stoffel, Biochemistry, 1993,32,11405-11412. 34 G. Muller-Newen, U.Janssen, W. Stoffel, Eur.]. Biochem. 1995,228, 68-73. 35 J. A. Gerlt, P. C. Babbitt, CUT. Opin. Chem. B i d , 1998, 2,607-612. 36 A. G. Murzin, C u r . Opin. Struct. B i d , 1998, 8,380-387. 37 P. C.Babbitt, J. A. Gerlt, Adv. Protein Chem., 2000,55,1-28. 38 I. D. Campbell, M. Baron, Philos. Trans. R. Soc. London Ser. B Bid. Sci., 1991,332, 165-170. 39 C. P.Ponting, J. Schultz, R. R. Copley, M. A. Andrade, P. Bork, Adv. Protein Chem., 2000, 54, 185-244. 40 S. Henikoff, E. A. Greene, S. Pietrokovski, P. Bork, T. K. Attwood, L. Hood, Science, 1997,278,609-614. 41 C. Khosla, P.B. Harbury, Nature, 2001,409, 247- 25 2. 42 R. Bentley, J.W. Bennett, Ann. Rev. Microb i d , 1999, 53, 411-446. 43 H. D. Mootz, M. A. Marahiel, CUT. Opin. Biotechnol., 1999, 10, 341-348. 44 D. E. Cane, C. T. Walsh, Chem. B i d , 1399, 6, R319-325. 45 R. N. Perham, Ann. Rev. Biochem., 2000,69, 961-1004. 46 H. D. Mootz, D. Schwarzer, M. A. Marahiel, Proc. Natl. Acad. Sci. U S A, 2000, 97, 5848-5853. 47 L. Katz, R. McDaniel, Med. Res. Rev., 1999, 19, 543-558.
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48 A. E. Nixon, M. Ostermeier, S. J. Benkovic,
Trends Biotechnol., 1998, 16, 258-264. 49 NC-IUBMB, Recommendations ofthe Nomenclature Commitee of the International Union ofbiochemistry and Molecular Biology on the Nomenclature and Classijkation of Enzymes, Academic Press, New York, NY 1992. 50 A. Bairoch, Nucleic Acids Res., 2000, 28, 304305. 51 A. Bairoch, R. Apweiler, Nucleic Acids Res., 2000,28,45-48. 52 M. Kanehisa, S. Goto, Nucleic Acids Res., 2000,28,27-30. 53 S . Goto, T. Nishioka, M. Kanehisa, Nucleic Acids Res., 2000, 28, 380-382. 54 L. B. Ellis, C. D. Hershberger, E. M. Bryan, L. P. Wackett, Nucleic Acids Res., 2001, 29, 340-343. 55 H. M. Berman, J. Westbrook, 2. Feng, G. Gilliland, T. N. Bhat, H. Weissig, I. N. Shindyalov, P. E. Bourne, Nucleic Acids Res., 2000,28,235-242. 56 K. N. Degtyarenko, A. C. North, J. B. Findlay, Nucleic Acids Res 27 (1999) 233-236. 57 T. M. S. a. D. Group, 1999. 58 N. D. Rawlings, A. J. Barrett, Nucleic Acids Res., 2000, 28, 323-325. 59 X. Cousin, T. Hotelier, K. Giles, P. Lievin, J. P. Toutant, A. Chatonnet, Nucleic Acids Res., 1997, 25, 143-146. 60 K. Hofmann, Trends Guide Bioinfom., 1998, 18-21. 61 T. K. Attwood, Int. J. Biochem. Cell Biol., 2000,32,139-155. 62 A. Bairoch, Nucleic Acids Res. Suppl., 1991, 19,2241-2245. 63 K. Hofmann, P. Bucher, L. Falquet, A. Bairoch, Nucleic Acids Res., 1999, 27, 215-219. 64 P. Bucher, A. Bairoch, Droc. Int. Con$ Intell. Syst. Mol. Biol., 1994, 2, 53-61. 65 A. Bateman, E. Birney, R. Durbin, S. R. Eddy, K. L. Howe, E. L. Sonnhammer, Nucleic Acids Res., 2000, 28, 263-266.
J. Schultz, R. R. Copley, T. Doerks, C. P. Ponting, P. Bork, Nucleic Acids Res., 2000, 28,231-234. 67 T. K. Attwood, M. D. Croning, D. R. Flower, A. P. Lewis, J. E. Mabey, P. Scordis, J. N. Selley, W. Wright, Nucleic Acids Res., 2000, 28,225-227. 68 J. G. Henikoff, E. A. Greene, S. Pietrokovski, S. Henikoff, Nucleic Acids Res., 2000, 28, 228-230. 69 R. Apweiler, T. K. Attwood, A. Bairoch, A. Bateman, E. Birney, M. Biswas, P. Bucher, L. Cerutti, F. Corpet, M. D. Croning, R. Durbin, L. Falquet, W. Fleischmann, J. Gouzy, H. Hermjakob, N. Hulo, I. Jonassen, D. Kahn, A. Kanapin, Y. Karavidopoulou, R. Lopez, B. Man, N. J. Mulder, T. M. Oinn, M. Pagni, F. Servant, Nucleic Acids Rex, 2001, 29, 37-40. 70 A. C. Wallace, N. Borkakoti, J. M. Thornton, Protein Sci., 1997, 6, 2308-2323. 71 A. C. Wallace, R. A. Laskowski, J. M. Thornton, Protein Sci., 1996, 5, 1001-1013. 72 A. Bernal, U. Ear, N. Kyrpides, Nucleic Acids Res., 2001,29, 126-127. 73 M. J. Danson, D. W. Hough, Trends Microbiol., 1998, 6, 307-314. 74 M. Y. Galperin, E. V. Koonin, Genetica, 1999, 106,159-170. 75 R. L. Tatusov, D. A. Natale, I. V. Garkavtsev, T. A. Tatusova, U.T. Shankavaram, B. S . Rao, B. Kiryutin, M. Y. Galperin, N. D. Fedorova, E. V. Koonin, Nucleic Acids Res., 2001,29,22-28. 76 E. V. Koonin, A. R. Mushegian, P. Bork, Trends Genet., 1996, 12, 334-336. 77 B. Snel, G. Lehmann, P. Bork, M. A. Huynen, Nucleic Acids Res., 2000, 28, 3442-3444. 78 K. Hofrnann, P. Bucher, Trends Biochem. Sci., 1996, 21, 172-173. 66
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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166
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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6 Immobilization ofhzymes
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
I
169
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 (
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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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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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
280
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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-
I
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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'
288
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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.
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.
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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.
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9 Enzymatic Kinetic Resolution
40 S-T. Chen, W-H. Huang, K-T. Wang,]. Org.
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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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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
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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
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I
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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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
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%)
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
o425
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, 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
o99 % 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]
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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] Me98 % 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
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.
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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 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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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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11 Hydrolysis and Formation of C-0 Bonds
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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,
/. Am. Chem. SOC.1997, 119,11137. (b) K. Haneda, T. Inazu, M. Mizuno, R. Iguchi, K. Yamamoto, H. Kurnagai, S . Aimoto, H. Suzuki, T. Noda, Bioorg. Med. Chem. Lett. 1998, 8, 1303. (c) M. Mizuno, K. Haneda, R. Iguchi, I. Muramoto, T. Kawakami, S . Aimoto, K. Yarnamoto, T. Inazu, /. Am. Chem. SOC. 1999,121,284. 329 T. J. Tolbert, C.-H. Wong,]. Am. Chem. SOC. 2000,122,5421. 330 (a) C. T. Walsh, /. Bid. Chem. 1989,264, 2393. (b) D. H. Williams, B . Bardsley, Angew. Chem. Int. Ed. Engl. 1999, 38, 1172. (c) D. E. Cane, C. T. Walsh, C. Khosla, Science 1998, 282, 63.
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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657
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
I
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
I
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
I
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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123 W. T. H. Chang, D. W. Thayer, Can. J. Micro-
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125 L. Huang, C. W. Forsberg. D. Y. Thomas,
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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 [ 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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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
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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
I
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 55 720 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
References 1 A. von Baeyer, Liebigs Ann. Chem., 1861,
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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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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
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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> [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
I
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.
References Bohley, Natunvissenschaften 1995, 82, 544-550. 2 A. J. Kenny, N. M. Hooper in: Degradation of Bioactive Substances, Physiology and Pathology (Ed.: J. H. Henriksen), CRC Press Inc, 1991, pp. 47-79. 3 A. J. Kenny, C. M. Boustead (Eds.) Cell-surface Peptidases i n Health and Disease, Bios Scientific Publishers, Oxford, 1997, p. 384. 4 A. J. Barret, N. D. Rawlings, J. F. Woessner (Eds.).Handbook ofProteolytic Enzymes, Academic Press, San Diego, 1998. 5 B. Zhang, T. Cech, Chern. Biol. 1998,5, 539-553. 1 P.
H. Kleinkauf, H. v. Dohren, Eur. J. Biochem. 1990, 192,l-15. 7 M. Schelhaas, H. Waldmann, Angew. Chem., Int. Ed. Engl. 1996, 35, 2056-2083. 8 H. Waldmann, D. Sebastian, Chem. Rev. 1994,94,911-937. 9 P. Hermann, Biomed. Biochem. AGta 1991, 50, ( l O / l l ) , 19-31. 10 C. Walsh, Enzymatic Reaction Mechanism, Freeman, San Francisco, 1979. 11 B. M. Dunn in Proteolytic Enzymes: A Practical Approach, (Eds.: R. J. Beynon, J. S. Bond), IRL, Oxford, 1989, pp. 5781. 6
860
I
1.2 Hydrolysis and Formation ofC-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
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I
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
I
875
876
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
I
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
879
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 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
J. Damell, H. Loddish, D. Baltimore, Molecular Cell Biology, 2"d ed., Scientific American Books Inc., New York, 1990. 2 Nomenclature Committee of the International Union of Biochemistry, Enzyme Nomenclature, Academic Press, Orlando, 1984. 3 P. D. Boyer, E. G. Krebs, D. S. Sigman, The Enzymes 3rded., Academic Press, New York, 1970-1992, Vol. I-XX. 4 D. Schomburg, M. Salzmann, (GFB-Gesellschaft fur Biotechnologische Forschung), Enzyme Handbook, Springer-Verlag,Berlin, D, 1990. 5 A. Fersht, Enzyme Structure and Mechanism, 2"d ed., W. H. Freeman and Co., New York, 1985, p. 235. 6 P. D. Boyer, E. G. Krebs, Control by Phosphorylation (The Enzymes) 3rded., 1986, Vols. XVII and XVIII. 7 E. C. Ball, Energy Metabolism, Addison-Wesley, Reading, MA, USA, 1973. 8 J. R. Knowles, Ann. Rev. Biochem. 1980,49, 877-919. 9 A. Pradines, A. Klaebe, J. Perie, F. Paul, P. Monsan, Enzyme Microb. Technol. 1991, 13, 19-23. 10 H. Flodgaard, H. Klenow, Biochem.]. 1982, 208,737-742. 11 H. Nakajima, 1. Tomioka, S. Kitabatake, D. Dombou, K. Tomita, Agric. Biol. Chem. 1989,53,615-623. 12 S. Kitabatake, M. Dombou, I. Tomioka, H. Nakajima, Biochem. Biophys. Res. Commun. 1987,146,173-178. 13 H. Nakajima, H. Kondo, R. Tsumtani, M. Dombou, I. Tomioka, K. Tomita, ACS Symp. Ser. 1991,466, 111-120. 14 G. M. Whitesides, C. H. Wong, Angew. Chem. Int. Ed. Engl. 1985,24, 617-638. 15 J.A. Gerlt in The Enzymes (Ed.: D. S . Sigman), Vol. XX, Academic Press Inc., San Diego, 1992, p. 95-139. 1
C. Walsh, Enzymatic Reaction Mechanism, W. H. Freeman and Co., New York, 1979, p. 213. 17 W. P. Jencks in Handbook ofBiochemistry (Ed.: H.A. Sober), The Chemical Rubber Co., Cleveland, 1970, p. 1-185. 18 R. J. Kazlauskas, G. M. Whitesides, J. Org. Chem. 1985,50,1069-1076. 19 R.A. Alberty,J . Biol. Chem. 1968,243, 133771343, 20 K. Shikama, K.-I. Nakamura, Arch. Biochem. Biophys. 1973, 157,457-463. 21 G. Rosing, E. C. Slater, Biochim. Biophys. Acta 1972,267,275-290. 22 A. L. Lehninger, Biochemie, 2"d ed., VCH, Weinheim, 1985. 23 D. Kesse1,J. Bid. Chem. 1968,243, 4739-4744. 24 R. A. Hiles, L.V. Henderson, J . Biol. Chem. 1972,247,646-651. 25 D. C. Watts in The Enzymes (Ed.: P. D. Boyer), Vol. VIII, Academic Press, New York, 1973, p. 383-455. 26 J. F. Morrison in The Enzymes (Ed.: P. D. Boyer), Vol. VIII, Academic Press, New York, 1973, p. 457-486. 27 M. Szymona, W. Ostrowski, Biochim. Biophys. Acta 1964,85, 283-295. 28 R. A. Smith, M. C. Theisen in Carbohydrate Metabolism (Ed.: W. A. Wood), Vol. IX, Academic Press, New York, 1966, p. 403-407. 29 L. M. Cagen, H. C. Friedmann, J . Biol. Chem. 1972,247,3382-3392. 30 L. Raijaman, M. E. Jones in The Enzymes (Ed.: P.D. Boyer),Vol. IX, Academic Press, New York, 1973, p. 97-119. 31 E. R. Stadtman, in Methods Enzymol. (Ed.: S. P. Colowick and N. 0. Kaplan), Vol. I, Academic Press Inc., New York, 1955, p. 596-599. 32 R. L. Anderson, M. Y. Kame1 in Carbohydrate Metabolism (Ed.: W. A. Wood), Vol. IX, Academic Press, New York, 1966, p. 392-396. 16
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54 L. Butler, Biotechnol. Bioeng. 1977, 19,
591-593.
55 Y.3. Shih, G.M. Whitesides,]. Org. Chem.
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928
I
13 Formation and Cleavage of P - 0 Bonds
76 C. H. Wong, D. G. Drueckhammer, J . Org.
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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
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
97 : 3
a o OH OH
H
OOH GOH o
H
>97 ; 97 : 97 : 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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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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117 U. Hanefeld, A. J . J. Straathof, J . J. Heijnen,
/. Mol. Catal. B: Enzym. 2001, 11, 213-218.
118 G. Rotcenkovs, 119
L. T. Kanerva,]. Mol. Catal.
B: Enzym. 2000,11, 37-43.
M.Inagaki, J. Hiratake, T. Nishioka, I. Oda.
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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
+
*{
- 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/
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
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
in the region of less than 10% of that with the native cofactor. However, it was shown that these analogs could have at least some potential. References
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Enzyme Catalysis in Organic Synthesis Karlheinz D r a w and Herbert Waldmann Copyright 0Wiley-VCH Verlag GmbH, Weinheim 2002
1170
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P. Pasta, G. Carrea, N. Gaggero, G. Grogan,
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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
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.
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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
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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.
9
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,
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
31, 909. H. Katagiri, K. Kitahara,]. &. Chem. Soc. Jpn. 1936, 12, 1217. 4 E. L. Tatum, W. H. Peterson, E. B. Fred, Biochem.J. 1936,30,1892. 5 G. Rosso, K. Takashima, E. Adams, Biochem. Biophys. Res. Commun. 1969, 34, 134. 6 K. Yonaha, T. Yorifuji, T. Yamamoto, K. Soda,J. Ferment. Technol. 1975,53, 579. 7 N. Esaki, C. T. Walsh, Biochemistry 1986,25, 3261. 8 S. A. Wasserman, E. Daub, P. Grisafi, D. Botstein, C. T. Walsh, Biochemistry 1984, 25, 5182. 9 B. Badet, C. T. Walsh, Biochemistry 1985,24, 1333. 10 K. Inagaki, K. Tanizawa, B. Badet, C. T. Walsh, H. Tanaka, K. Soda, Biochemistry 1986,25,3268. 11 K. Yokoigawa, H. Kawai, K. Endo, Y. Lim, N. Esaki, K. Soda, Biosci. Biotechnol. Biochem. 1993,57,93. 12 K. Hoffmann, E. Schneider-Scherzer, H. Kleinkauf, R. Zocher, J. B i d . Chem. 1994,269,12710. 3
T. Seow, K. Inagaki, T. Tamura, K. Soda, H. Tanaka, Biosci. Biotechnol. Biochem. 1998, 62, 242. 14 Y. Okubo, K. Yokoigawa, N. Esaki, K. Soda, H. Kawai, Biochem. Biophys. Res. Commun. 1999,256,333. 15 S. Kim, Y. Gyu, J. Biochem. Mol. Biol. 2000, 33, 82. 16 T. Uo, T. Yoshimura, N, Tanaka, K. Takegawa, N. Esaki,]. Bacteriol. 2001,183, 2226. 17 A. Galkin, L. Kulakova, H. Yamamoto, K. Tanizawa, H. Tanaka, N. Esaki, K. Soda, /. Ferment. Bioeng. 1997, 83, 299. 18 A. Galkin, L. Kulakova, T. Yoshimura, K. Soda, N. Esaki, Appl. Environ. Microbiol. 1997,63,4651. 19 C. T. Walsh, J. B i d . Chem. 1989,264, 2393. 20 E. Ferrari, D. J. Henner, M. Y. Yang, Biotechnology 1985, 3, 1003. 21 J. Wild, M. Lobocka, W. Walczak, T. Klopotowski, Mol. Gen. Genet. 1985, 198, 315. 22 N. G. Galakatos, C. T. Walsh, Biochemistry 1989,28,8167. 23 H. Toyama, K. Tanizawa, M. Wakayama, Q. Lee, T. Yoshimura, N. Esaki, K. Soda, Agnc. B i d . Chem. 1991, 55, 2881. 24 H. Toyama, K. Tanizawa, T. Yoshimura, S. 13
References I1327 Asano, H. -H. Lim, N. Esaki, K. Soda,J. Bid. Chem. 1991,266,13634. 25 H. C. Dunathan, Proc. Natl. Acad. Sci. U S A 1966, 55, 713. 26 S.-J. Shen, H. G. Floss, H. Kumagai, H. Yamada, N. Esaki, K. Soda, S. A. Wasserman, C. T. Walsh,J. Chem. SOC.Chem. Commun. 1983, 82. 27 G. J. Cardinale, R. H. Abeles, Biochemistry 1968,7, 3970. 28 S. A. Ahmed, N. Esaki, H. Tanaka, K. Soda, Biochemistry 1986,25, 385. 29 W. S. Faraci, C. T. Walsh, Biochemistry 1988, 27, 3267. 30 S. Sawada, Y.Tanaka, S. Hayashi, M. Ryu, T. Hasegawa, Y. Yamamoto, N. Esaki, K. Soda, S. Takahashi, Biosci. Biotechnol. Biochem. 1994,58,807. 31 J. P. Shaw, G. P. Petsko, D. Ringe, Biochemistry 1997, 36, 1329. 32 C. G. Stamper, A. A. Morollo, D. Ringe, Biochemistry 1998, 37, 10438. 33 A. Watanabe, Y. Kurokawa, T. Yoshimura, T. Kurihara, K. Soda, N. Esaki, J. Bid. Chem. 1999,274,4189. 34 B. Badet, K. Inagaki, K. Soda, C. T. Walsh, Biochemistry 1986, 25, 3275. 35 N. Esaki, H. Shimoi, N.Nakajima, T. Ohshima, H. Tanaka, KSoda, J. Bid. Chem. 1989,264,9750. 36 T. Ohshima, K. Soda, Eur. J . Biochem. 1979, 100, 29. 37 K. Yonaha, K. Soda, Biochem. Engin. Biotechnol. 1986, 33,95. 38 R. Wichmann, C. Wandrey, A. F. Buckmann, M. -R. Kula, Biotechnol. Bioeng. 1981, 23,2789;A. S. Bommarius, M. Schwarm, K. Stingl, M. Kottenhahn, K. Huthmacher, K. Drauz. Tetrahedron: Asymmetry, 1995,6, 2851. 39 W. Hummel. M.-R. Kula, Eur. J. Biochem. 1989, 184, 1. 40 Y. Asano, A. Nakazawa, Agnc. Bid. Chem. 1987,51,2035. 41 Y. Asano, A. Yamada, K. Kato, Y. Yamaguchi, K. Hibino, K. Kondo,J . Org. Chem. 1990,55,5567. 42 Y. Kato, K. Fukumoto, Y. Asano, Appl. Microbid. Biotechnol. 1993, 39, 301. 43 T.Ohshima, C. Wandrey, M.-R. Kula, K. Soda, Biotechnol. Bioeng. 1985, 27, 1616. 44 K. Soda, K. Yonaha in: Biotechnology 7a (Eds.: H.-J.Rehm, G. Reed), VCH Verlagsgesellschaft, Weinheim, 1987, 616.
A. Hashimoto, S. Kumashiro, T. Nishikawa, T. Oka, K. Takahashi, T. Mito, S. Takashima, N. Doi, Y. Mizutani, T. Yamazaki,]. Neurochem. 1993,61, 348. 46 A. Hashimoto, T. Nishikawa, T. Oka, K. Takahashi, J. Neurochem. 1993.60, 783. 47 T. Matsui, M. Sekiguchi, A. Hashimoto, U. Tomita, T. Nishikawa, K. Wada, j . Neurochem. 1995,65,454. 48 S. Filc-DeRicco,A. S. Gelbard, A. j. Cooper, K. C. Rosenspire, E. Nieves, Cancer Res. 1990, SO, 4839. 49 K. Yonaha, H. Misono, T. Yamamoto, and K. Soda,J. Biol. Chem. 1975,250,6983. 50 K. Tanizawa, Y. Masu, S. Asano, H. Tanaka, K. Soda,J. Bid. Chem. 1989,264,2445. 51 A. Galkin, L. Kulakova, V. Tishkov, N. Esaki, K. Soda, Appl. Microbiol. Biotechnol. 1995, 44,479. 52 M. L. Scott in: Organic Selenium Compounds: Their Chemistry and Biology (Eds.: D. L. Klayman, W. H. H. Gunther), john Wiley & Sons, New York, 1973,629. 53 A. Shrift in: Organic Selenium Compounds: 7heir Chemistry and Biology (Eds.: D. L. Klayman, W. H. H. Gunther), john Wiley & Sons, New York, 1973,763. 54 N. Esaki, H. Shimoi, H. Tanaka, K. Soda, Biotechnol. Bioeng. 1989, 34, 1231. 55 Y. Sakamoto, S. Nagata, N. Esaki, H. Tanaka, K. Soda,J. Ferment. Bioeng. 1990, 69, 154. 56 S. M. Roberts, N. J. Turner, A. J. Willetts, M. K. Turner in: Introduction to Biocatalysis Using Enzymes and Micro-organisms, Cambridge University Press, New York, 1995, 34. 57 J. E. Bailey, Science 1991, 252, 1668. 58 L. 0. Ingram, F. Alterthum, K. Ohta, D. S. Beall in: Developments i n Industrial Microbiology Indust. Microbiol., Suppl. No. 5), 1990,31, pp. 21-30. 59 K. Soda, T. Osumi, Methods Enzymol. 1971, 17B, 629. 60 K. Inagah, K. Tanizawa, H. Tanaka, K. Soda, Agnc. Biol. Chem. 1987,51, 173. 61 T. Yorifuji, K. Ogata, K. Soda,J. Bid. Chem. 1971,246, 5085. 62 Y. -H. Lim, K. Yokoigawa, N. Esaki, K. Soda, J. Bacteriol. 1993, 175, 4213. 63 K. Reynolds, J. Martin, S.-J.Shen, N. Esaki, K. Soda, H. G. Floss, J . Basic Microbiol. 1991, 31, 177.
45
u.
1328
I
77 lsomerizations
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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.
1334
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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.
1336
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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
I
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