Biotechnology

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Biotechnology

Second Edition Volume 8a

Biotransfonnations I

@ WILEY-VCH

Biotechnology

Second Edition Fundamentals

Special Topics

Volume 1 Biological Fundamentals

Volume 9 Enzymes, Biomass, Food and Feed

Volume 2 Genetic Fundamentals and Genetic Engineering

Volume 10 Special Processes

Volume 3 Bioprocessing Volume 4 Measuring, Modelling, and Control Products Volume 5 Recombinant Proteins, Monoclonal Antibodies and Therapeutic Genes Volume 6 Products of Primary Metabolism Volume 7 Products of Secondary Metabolism Volumes 8a and b Biotransformations I and I1

Volumes l l a and b Environmental Processes Volume 12 Legal, Economic and Ethical Dimensions

A Multi-Volume Comprehensive Treatise

Biotechnology

Second, Completely Revised Edition Edited by H.-J. Rehm and G. Reed in cooperation with A. Bhler and P.Stadler

Volume 8a

Biotransformations I Edited by D. R. Kelly

CB WILEY-VCH

Weinheim . New York . Chichester . Brisbane Singapore * Toronto

Series Editors: Prof. Dr. H.-J. Rehm Institut fur Mikrobiologie Universitat Munster CorrensstraBe 3 D-48149 Munster FRG

Dr. G . Reed 1029 N . Jackson St. #501-A Milwaukee, WI 53202-3226 USA

Prof. Dr. A . Puhler Biologie VI (Genetik) Universitat Bielefeld P.O. Box 100131 D-33501 Bielefeld FRG

Prof. Dr. P. J. W. Stadler Artemis Pharmaceuticals Geschaftsfuhrung WilhelrnstraBe 10 D-42755 Haan FRG

Volume Editor: Dr. D. R. Kelly Cardiff University of Wales P.O. Box 912 Cardiff, CF1 3TB, Wales UK

This book was carefully produced. Nevertheless, authors, editors and publisher d o 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.

Library of Congress Card No.: applied for British Library Cataloguing-in-Publication Data:

A catalogue record for this book is available from the British Library

Die Deutsche Bibliothek - CIP-Einheitsaufnahme Biotechnology : a multi volume comprehensive treatise I ed. by H.-J. Rehm and G . Reed. In cooperation with A . Piihler and P. Stadler. 2., completely rev. ed. -VCH. ISBN 3-527-28310-2 (Weinheim ...)

NE: Rehm, Hans-J. [Hrsg.] Vol. 8a: Biotransformations I I ed. by D. R. Kelly - 1998 ISBN 3-527-28318-8 OWILEY-VCH Verlag GmbH, D-69469 Weinheim (Federal Republic of Germany), 1998 Printed on acid-free and chlorine-free paper. All rights reserved (including those of translation into other languages). N o part of this book may be reproduced in any form-by photoprinting, microfilm, or any other means-nor transmitted or translated into a machine language without written permission from the publishers. Registered names, trademarks, etc. used in this book, even when not specifically marked as such, are not to be considered unprotected by law. Composition and Printing: Zechnersche Buchdruckerei, D-67330 Speyer. Bookbinding: J. Schaffer, D-67269 Griinstadt. Printed in the Federal Republic of Germany

Preface

In recognition of the enormous advances in biotechnology in recent years, we are pleased to present this Second Edition of “Biotechnology” relatively soon after the introduction of the First Edition of this multi-volume comprehensive treatise. Since this series was extremely well accepted by the scientific community, we have maintained the overall goal of creating a number of volumes, each devoted t o a certain topic, which provide scientists in academia, industry, and public institutions with a well-balanced and comprehensive overview of this growing field. We have fully revised the Second Edition and expanded it from ten to twelve volumes in order t o take all recent developments into account. These twelve volumes are organized into three sections. The first four volumes consider the fundamentals of biotechnology from biological, biochemical, molecular biological, and chemical engineering perspectives. The next four volumes are devoted to products of industrial relevance. Special attention is given here to products derived from genetically engineered microorganisms and mammalian cells. The last four volumes are dedicated to the description of special topics. The new “Biotechnology” is a reference work, a comprehensive description of the state-of-the-art, and a guide to the original literature. It is specifically directed t o microbiologists, biochemists, molecular biologists, bioengineers, chemical engineers, and food and pharmaceutical chemists working in industry, at universities o r at public institutions. A carefully selected and distinguished Scientific Advisory Board stands behind the

series. Its members come from key institutions representing scientific input from about twenty countries. The volume editors and the authors of the individual chapters have been chosen for their recognized expertise and their contributions t o the various fields of biotechnology. Their willingness to impart this knowledge to their colleagues forms the basis of “Biotechnology” and is gratefully acknowledged. Moreover, this work could not have been brought t o fruition without the foresight and the constant and diligent support of the publisher. W e are grateful to VCH for publishing “Biotechnology” with their customary excellence. Special thanks are due to Dr. HansJoachim Kraus and Karin Dembowsky, without whose constant efforts the series could not be published. Finally, the editors wish to thank the members of the Scientific Advisory Board for their encouragement, their helpful suggestions, and their constructive criticism. H.-J. Rehm G. Reed A. Puhler P. Stadler

Scientific Advisory Board

Pro$ Dr. M. J. Beker

Prof Dr. T. K. Ghose

Prof Dr. J. D. Bu’Lock

Prof Dr. I. Goldberg

August Kirchenstein Institute of Microbiology Latvian Academy of Sciences Riga, Latvia

Biochemical Engineering Research Centre Indian Institute of Technology New Delhi, India

Weizmann Microbial Chemistry Laboratory Department of Chemistry University of Manchester Manchester, UK

Department of Applied Microbiology The Hebrew University Jerusalem, Israel

Prof Dr. C. L. Cooney

Prof: Dr. G. Goma

Department of Chemical Engineering Massachusetts Institute of Technology Cambridge, MA, USA

Departement de Genie Biochimique et Alimentaire Institut National des Sciences Appliquees Toulouse, France

Prof Dr. H. W. Doelle

Sir D. A . Hopwood

Department of Microbiology University of Queensland St. Lucia, Australia

Department of Genetics John Innes Institute Norwich, UK

Prof Dr. J. Drews

Prof Dr. E. H. Houwink

Prof Dr. A. Fiechter

Prof Dr. A . E. Humphrey

F. Hoffmann-La Roche AG Basel, Switzerland

Institut fur Biotechnologie Eidgenossische Technische Hochschule Zurich, Switzerland

Organon International bv Scientific Development Group Oss. The Netherlands

Center for Molecular Bioscience and Biotechnology Lehigh University Bethlehem, PA, USA

VIII

Scientific Advisory Board

Prof Dr. I. Karube

Prof Dr. K . Schiigerl

Research Center for Advanced Science and Technology University of Tokyo Tokyo, Japan

Institut fur Technische Chemie Universitat Hannover Hannover, Germany

Prof Dr. M. A. Lachance

Prof Dr. P. Sensi

Department of Plant Sciences University of Western Ontario London, Ontario, Canada

Chair of Fermentation Chemistry and Industrial Microbiology Lepetit Research Center Gerenzano, Italy

Prof Dr. Y. Liu

Prof Dr. Y. H. Tan

China National Center for Biotechnology Development Beijing, China

Institute of Molecular and Cell Biology National University of Singapore Singapore

Prof Dr. J. F. Martin

Prof Dr. D. Thomas

Department of Microbiology University of Leon Leon, Spain

Laboratoire de Technologie Enzymatique Universite de Compiegne Compiegne, France

ProJ Dr. B. Mattiasson

Prof Dr. W. Verstraete

Department of Biotechnology Chemical Center University of Lund Lund. Sweden

Laboratory of Microbial Ecology Rijksuniversiteit Gent Gent, Belgium

Prof Dr. M. Roehr

Prof Dr. E.- L. Winnacker

Institut fur Biochemische Technologie und Mikrobiologie Technische Universitat Wien Wien. Austria

Prof Dr. H. Sahm

Institut fur Biotechnologie Forschungszentrum Julich Jiilich, Germany

Institut fur Biochemie Universitat Miinchen Munchen, Germany

Contents

Introduction D. R. Kelly

1

1 Perspectives in Biotransformation 5 M.Turner 2 Biotransformations - Practical Aspects 25 D. R. Kelly

Hydrolases 3 Biotransformations with Lipases 37 R. Katlauskas, U. Bornscheuer 4 Esterases 193 S.Phvthian 5 C1ea;age and Formation of Amide Bonds 243 D. Hoople 6 Nitriles 277 A . Bunch

Redox Enzymes 7 Alkaloids 327 N Bruce 8 Yeast 363 S. Servi 9 Alcohol Dehydrogenases - Characteristics, Design of Reaction Conditions, and Applications 391 J. Peters 10 Hydroxylation and Dihydroxylation 475 H. Holland 11 Flavin Monooxygenases- Uses as Catalysts for Baeyer-Villiger Ring Expansion and Heteroatom Oxidation 535 D. R. Kelly, f? Wan, J. Tsang

Index 589

Contributors

Dr. Uwe Bornscheuer

Dr. David Hoople

Universitat Stuttgart Allmandring 31 D-70569 Stuttgart Germany Chapter 3

Pfizer Central Research Ramsgate Rd. Sandwich, Kent, CT13 9NJ UK Chapter 5

Dr. Neil Bruce

Prof. Romas J. Kazlauskas

Institute of Biotechnology University of Cambridge Cambridge, CB2 1QT UK Chapter 7

Department of Chemistry McGill University 801 Sherbrooke St. W. MontrCal, Quebec H3A 2K6 Canada Chapter 3

Dr. Alan Bunch

Dr. David R. Kelly

Biological Laboratory University of Kent Canterbury, Kent UK Chapter 6

Cardiff University of Wales PO. Box 912 Cardiff, CFl 3TB UK Chapters 2 and I I

Prof. Herbert L. Holland

Dr. Jorg Peters

Department of Chemistry Brock University St. Catharines, Ontario, L2S 3A1 Canada Chapter I0

Bayer AG Geschaftsbereich Pharma T O Biotechnologie D-42096 Wuppertal Germany Chapter 9

XI1

Contributors

Dr. Sara Phythian

Prof. Michael K. Turner

Natural Resources Institute The University of Greenwich Central Avenue, Chatham Maritime Kent ME4 4TB UK Chapter 4

The Advanced Center for Biochemical Engineering Department of Chemical and Biochemical Engineering University College London Torrington Place London, WClE 7JE UK Chapter 1

Prof. Stefan0 Servi

Dr. Peter W. H. Wan

Dipartimento Chimico Politech Milan Via Marcinelli 7 1-20131Milano Italy Chapter 8

Dr. Jenny Tsang

Department of Chemistry The Robert Robinson Laboratories University of Liverpool PO. Box 147 Liverpool, L69 3BX UK Chapter I1

Department of Chemistry Imperial College of Science, Technology and Medicine Exhibition Road, South Kensington London, SW7 2AY UK Chapter I1

Biotechnology Second, Completely Revised Edition H.-J. Rehm and G. Reed copyrightOWILEY-VCH Verlag GmbH, 1998

Introduction

DAVID R. KELLY Cardiff, UK

1984 was a much heralded year. Fortunately ORWELL’S vision was not fulfilled, but the year was marked by a more modest event; the publication of the first edition of the volume in hand. The second edition of Biotransformations has been completely rewritten and it could not have been otherwise, because the entire subject has changed beyond all recognition in the intervening 14 years. The differences are underscored by the organization of the two editions. In the first edition each chapter is devoted to a class of compounds whereas in the current edition virtually all the chapters are focused on functional groups. The difference arises from the recognition that individual enzymes have both extraordinary selectivity and are able to transform an enormous range of compounds. In many cases these un-natural substrates show little obvious structural kin-

ship with the natural substrate other than the functional group. Pre-1980’s biotransformations developed out of the needs of industry which required enantiomerically pure compounds on a scale which could not be considered using the abiotic asymmetric synthetic tools of the time. Despite considerable advances, large-scale synthesis of enantiomerically pure chiral compounds still largely relies on products from the chiral pool and biotransformations. Even in academia, the magnificent total syntheses of Palytoxin and the Brevetoxins relied on sugars as starting materials. This was perhaps understandable given the similarities between sugars and the final products, yet only recently NKOLAOU and HOLTONused respectively a late stage resolution and camphor as a starting material in the first two total syntheses of Taxol@.

2

Introduction

Continuing demands for enantiomerically pure materials provided the impetus for academic investigation of biotransformations. Moreover, synthetic expertise and the development of analytical chiral chromatography enabled viable investigations of novel substrates, which constitute the majority of the work reported in this volume. This volume is divided into three superchapters. Chapters 1 and 2 provide an introduction to the history of biotransformations and background information for the novice. Chapters 3-6 cover the hydrolytic enzymes: lipases, esterases and hydrolytic enzymes which act on amide and nitrile groups. These are generally the easiest enzymes to use particularly on a large scale. There are probably more papers reporting lipase- and esterase-catalyzed reactions than all other biotransformations (Chapters 3 and 4). Moreover, biotransformations catalyzed by hydrolytic enzymes have been utilized for the manufacture of such large volume products as aspartame (Chapter 5 ) and acrylonitrile (Chapter 6). The third super-chapter covers redox enzymes. The broad range of reactions encompassed is illustrated by the alkaloids, together with aspects of their biosynthesis (Chapter 7). Similarly yeast which are best known for the reduction of ketones are also capable of a wide range of other reactions (Chapter 8). Horse liver alcohol dehydrogenase was extensively investigated by J. B. JONES in a pioneering study, and recent developments such as enzymes from thermophilic bacteria are covered in Chapter 9. Problems with the regeneration of cofactors such as NAD(P)H have largely been overcome, which enables these enzymes to be used on any scale. This chapter also analyzes strategies for the introduction of chirality into targets and some novel reactors (Chapter 9). There are some reactions for which there is no viable alternative to biotransformations. These include remote hy-

droxylation which has played a vital role in the development of the corticosteroids (Chapter 10, cf. Chapter 1) and dihydroxylation which has furnished cis-benzene glycol a unique synthetic intermediate (Chapter 10). Similarly enantioselective Baeyer-Villiger reactions catalyzed by mono-oxygenases are only now being imperfectly imitated by abiotic catalysts (Chapter 11). The topics in Volume 8b will include phosphorylation, carbon+arbon bond formation, glycosidation and the application of biotransformation products in synthesis. Biotransformations cross a constellation of disciplines which include chemistry, biochemistry, microbiology and engineering. Consequently, most of the other volumes in the second edition of Biotechnology cover topics which impact on biotransformations. The primary focus of the current volume is the reactions of un-natural substrates, however, many similar issues are addressed in the volumes covering the products of primary and secondary metabolism (Volumes 6 and 7). An understanding of the physiology and metabolism of microorganisms (Volume 1) and methods for genetic manipulation (Volume 2) are essential for the development of laboratory-scale reactions into viable manufacturing processes. Engineering aspects of biotransformations are covered in the volumes on bioprocessing (Volume 3) and measuring, modeling and control (Volume 4). Will abiotic asymmetric synthesis overtake biotransformations and render them obsolete? Is there a long-term future for natural catalysts constructed from a limited range of amino acids, adapted for metabolic efficiency in vivo when abiotic catalysts can draw on any functional group or structural entity and can be designed for reactions in vitro. There can be no doubt that the efficiency and enantioselectivity of abiotic catalysts will continue to be improved. Moreover, there are signs that the two

Introduction

areas are drawing together as shown by the reactions of poly-leucine and catalytic antibodies (cf. Volume 5 ) . Although the early promise of the latter has not yet been achieved. However, for the foreseeable future enzymes will remain the only catalysts that can be used in tandem in a single pot. Similarly the use of blocked mu-

3

tants enables fragments of multi-enzyme biosynthetic pathways to be exploited. In both cases biotransformations provide unique opportunities to create complex molecular architectures from simple precursors. Cardiff, February 1998

D. R. Kelly


1 Perspectives in Biotransformations

MICHAEL K. TURNER London, UK

1 Introduction 6 2 Perspective 1895-1935 - Organic Chemistry and Industrial Microbiology 6 2.1 PASTEUR, Chiral Resolution and Fermentation 6 2.2 Fermentations and Applied Biocatalysis 8 3 Perspective 1935-1955 - A Fallow Time 10 4 Perspective 1955-1975 - Functionalization of Natural Products 11 4.1 Steroid Manufacture 11 4.2 P-Lactam Manufacture 14 4.3 Enzyme Engineering and the Manufacture of Amino Acids 17 5 A Current Perspective - Collaboration 18 6 References 21

6

I Perspectives in Biotramfonnatiom

1 Introduction Perspectives depend on your point of view. The development of biotransformations as a tool in organic synthesis is only one of many uses of biological catalysis These include applications in the manufacture and preservation of food and drink, in the manufacture and cleaning of textiles, in diagnostic or sensor technology, and in the synthesis of nucleic acids to name only those which dominate the list. These lie outside the scope of this introduction, although research in the brewing industry must be acknowledged as having stimulated many of the classical nineteenth century studies in microbial biochemistry (ROBERTS et al., 1995). This volume should describe the proper use of biochemical methods in the synthesis of defined organic chemicals either for their value as intermediate synthons, or as products an their own right.At the heart of this is the need to synthesize complex molecules which are chiral and selectively functionalized. The principles on which the biochemical methods are based are not new. The reactions described in subsequent chapters stand on precedents which reach back to PASTEUR’S research on the optical activity of organic chemicals. The immediate influence of his research on his contemporaries,and the current state of applied biocatalysis which is reviewed in this volume, even though they are separated by almost 150 years, illustrate the reasons why chemists and manufacturers turn to biocatalysis, and the strands which must be pulled together to create an effective process. The collaboration of chemists and biologists creates the competing process options which engineers must harness and control in largescale processes, but this in itself is not sufficient. There must also be a market for the product of an economic process and at any stage of technical development much more is possible than is economically viable, as this volume will certainly demonstrate. However, markets develop in response to social needs which can change as rapidly as technology, and the choice of an economic process will frequently depend on more than a balance sheet of process costsl’his is certainly true of the application of biocatalysis to organic chemical

synthesis. This introduction will sketch out these influences as they have affected the use of the technology since PASTEUR first demonstrated the microbial resolution of tartaric acid (PASTEUR,1858), the third of the classical methods for resolving racemates, all of which he invented.

2 Perspective 1895-1935 - Organic Chemistry and Industrial Microbiology 2.1 Pasteur, Chiral Resolution and Fermentation In March 1897 PERCYFRANKLAND, the Professor of Chemistry at Mason College in Birmingham (UK), delivered a lecture in memory of LOUISPASTEUR who had died eighteen months 1897). He included in his earlier (FRANKLAND, description of PASTEUR’S research the three classical methods for resolving the enantiomers in a racemate. The first method, in which the two enantiomers separated spontaneously on crystallization, could be applied only in a few instances.The second, in which diastereomeric salts were formed with either the natural alkaloid bases or with one of the resolved enantiomers of tartaric acid, had “proved the master key to some of the most remarkable organic syntheses which have hitherto been realized”. FRANKLAND argued that LADENBURG’S synthesis of the hemlock alkaloid coniine (1) (Fig. l), and FISCHER’S synthesisof sugars were both largely dependent on the successful application of this second method (FRANKLAND, 1897, p. 695).

0 I

%/A

H

1 Coniine

Fig. 1. Coniine.

2 Perspective1853-2935 - Organic Chemistiy and Industrial Microbiology

7

The lecture then describes PASTJWR’Sthird chemistry. The choice which is always implied, method in which a microorganism degrades when there are alternative methods of achievone enantiomer of a racemate leaving the oth- ing the same technical goal, was as valid then er untouched. This too had become “ an inval- as it is now. If a microorganism would efficientuable instrument in experimental science”. He ly catalyze a necessary reaction then that was a continued that sufficient reason to use it. What is striking, is enthusiasm for their use, which “it would be impossible for me here to at- FRANKLAND’S tempt to place on record even the names of should not be attributed simply to the context those numerous optically active compounds of the remarks in a Pasteur Memorial Lecture. The following decade saw the introduction our acquaintance with which is wholly dependent on the subjection of racemoids to the selec- of another aspect of biocatalysis which we tive action of microorganisms, indeed the would recognize as central to its current use. and LOEVENHART (1900), method seems to be well nigh of universal ap- HILL(1897), KASTLE (1%) all showed that the catalplication in the case of all racemoid bodies and POITEVIN which are capable of being attacked by these ysis of hydrolytic enzymes was reversible. POTlow forms of vegetable life, and selective de- TEVIN went further and demonstrated that a compositions of this kind have been effected crude pancreatic lipase would synthesize both by moulds, by bacteria and by the saccha- methyl oleate from methanol and oleic acid in a reaction mixture which was largely organic. 1897). romyces or yeasts” (FRANKLAND, Chirality and living organisms were particu- These reactions should not be given a spurious larly closely linked in the minds of the chem- technological context. The kinetics of enzygeneration but their actu- matic catalysis were a puzzle, and it was imporists of FRANKLAND’S al use of microorganisms in organic syntheses tant to show that they were consistent with the may surprise chemists practicing a century af- law of mass action. However, the surprise death. FRANKLAND’S views, and which was later to greet similar reactions, in ter PASTEUR’S the research on which they were based, were the context of a newly applied biocatalysis, already part of the main stream of organic would have been misplaced at the time when

Fig. 2a,b. A sim lified drawing (SYKES,1895) (a) of PASTEUR’S large-scale fermenter (PASTEUR, 1876, p. 328) (bk The 20 are useful for synthesis. To calculate E, one measures two of the three variables: enantiomeric purity of the starting material (ee,), enantiomeric purity of the product (ee,), and extent of conversion ( c ) and uses one of the three equations below (Eqs. 8 a- 8 ~ )Often . enantiomeric purities are more accurately measured than conversion; in these cases, the third equation is more accurate.

E=

ln[1 - c ( l +ee,)] In[l - c ( l -ee,)]

E=

In [(I - c) (1 - ee,)] In [(1- c ) (1 + ee,)] In[

1+ (ee,/ee,)

In[

1+ (ee,/ee,)

E=

l-ees

1+ee5

] ]

(8b)

(8c)

High E values (-100) are less accurately measured than low or moderate E values because the enantiomeric ratio is a logarithmic function of the enantiomeric purity. When E 100, small changes in the measured enan-

-

63

tiomeric purities give large changes in the enantiomeric ratio. Thus, the survey below avoids reporting E values above 100. In practice, we found that even E values near 50 were sometimes difficult to measure more precisely than +lo. A simple program to calculate enantiomeric ratio using the above equations is freely available at http://www-orgc.tugraz.ac.at. (KROUTIL et al., 1997a). In spite of the fact that these equations include assumptions such as an irreversible reaction, one substrate and product, and no product inhibition, they are reliable in the vast majority of cases, especially for screening studies. Recently, we developed a faster spectrophotometric method for measuring the enantiomeric ratio (JANESand KAZLAUSKAS, 1997a).This method, called Quick E, requires samples of the pure enantiomers. For careful optimization of reactions, three situations require a more careful approach. First, when the biocatalyst is a mixture of enzymes, for example, isozymes, which all act on the substrate, then the calculated E value reflects a weighted average of all the enzymes (CHENet a]., 1982).When these enzymes differ significantly in their affinity for the substrate, then different enzymes will dominate the activity at different substrate concentrations. Thus, the apparent enantioselectivity may vary as the reaction depletes the substrate or when the reaction is carried out with different initial substrate concentrations. When enzymes differ in their stability, apparent enantioselectivities for long vs. short reaction times may differ. To measure the true E value, one must purify the enzymes and measure E separately. Second, when product inhibits the reaction the apparent enantioselectivity can change (RAKELS et al., 1994;VAN TOLet al., 1995a, b). For example, addition of 4 v/v% ethanol to a carboxylesterase NP-catalyzed hydrolysis of ethyl 2-chloropropionate increased the enantioselectivity from 4.7 to 5.4. RAKELSet al. (1994) attributed this change not to changes in the inherent selectivity of the enzyme, but to selective inhibition of one of the enantiomers by ethanol. In another example, VAN TOLet al. (1995a, b) could not recover enantiomerically pure starting material in the PPL-catalyzed hydrolysis of glycidol butyrate even at high conversion. The enantiomeric purity of the re-

64

3 Biotransformations with Lipases

maining glycidol butyrate reached 95% ee at called these reactions sequential kinetic reso70% conversion, but did not increase further lutions, but we favor in situ recycling and reeven at 90% conversion. In other words, the serve the term sequential kinetic resolution apparent enantioselectivity dropped from 20 only for those reactions where both steps ocat 31% conversion to 2.7 at 95% conversion. cur at the same time, such as the acylation of VANTOL et al. (1995a, b) attributed this pla- diols. Like recycling reactions, sequential kinetic teau to product inhibition promoting the reverse reaction for the product enantiomer. To resolutions enhance the enantiomeric purity 1989; Guo et include product inhibition in the quantitative of the products (KAZLAUSKAS, 1991). For analysis, reseachers used more complex equa- al., 1990; CARONand KAZLAUSKAS, tions which take into account the mechanism example, hydrolysis of trans-1,Zdiacetoxycyof lipase-catalyzed reaction (usually ping-pong clohexane proceeds stepwise - first hydrolysis bi-bi). Until now few researchers included to the monoacetate, then to the diol (Fig. 11) 1991). Both reacproduct inhibition in their analysis, but a read- (CARONand KAZLAUSKAS, ily available computer program (ANTHONSEN tions favor the same enantiomer, thus, the two et al., 1996; http://bendik.mnfak.unit.no) sim- resolutions reinforce each other. Maximum reinforcement occurs when both reactions occur plifies this task. Third, when the reaction is reversible, such as at comparable rates with an overall enantiotransesterification, one must include the equi- selectivity of approximately ( E l * E2)/2 (CAlibrium constant for the reaction (CHENet al., RON and KAZLAUSKAS, 1991). In addition, se1987). One can first measure the equilibrium quential kinetic resolutions yield both the constant in a separate experiment and then de- starting material and product in high enantiotermine E from measurements of ee, and eep. meric purity at the same extent of conversion ANTHONSEN et al. (1995) developed a simpler because the “mistakes” remain in the intermeapproach where they determine both K and E diate product (monoacetate in the example in by fitting a series of ee, and eep measurements. Fig. 11). In contrast, single step kinetic resolutions yield high enantiomeric purity for the product at < 50% conversion, but high enantiomeric purity for the starting material re3.1.2 Recycling and Sequential quires > 50% conversion. Kinetic Resolutions C,-symmetric diols are especially well suited to sequential kinetic resolution because both To enhance the enantiomeric purity, the en- steps are likely to have the same enantiopreferiched material can be isolated and resolved rence (Fig. 12). Unsymmetrical diols can also undergo a seagain. This double resolution is called recycling. CHEN et al. (1982) derived an equation to quential kinetic resolution (Fig. 13). predict the optimum degree of conversion in Only one dicarboxylic acid was resolved by recycling reactions and many researchers have lipase-catalyzed sequential kinetic resolution used this strategy (for an example see JOHN- and this was a special case. NODEet al. (1995) SON et al., 1995). BROWNet al. (1993) and VAN’ZTINEN and KANERVA(1997) reported several examples and computer programs for RR I )R R I )R R calculations are available for instance at ss s‘s s‘s‘ http://www-orgc.tu-graz.ac.at. (KROUTIL et al., 1997b). Guo (1993) reported plots to predict OAc the maximum chemical yield in various situa.-.’oAc PCL. hexane-water + * tions. To minimize the work in recycling reactions, several groups used in situ recycling 42%, >99% ee 38%. >99% ee racemic where the two resolutions are carried out stepwise, but without isolation of the intermediate Fig. 11. Sequential kinetic resolution enhances the products (CHENand LIU, 1991; MAJERICand enantiomeric purity of the product through two 1996; SUGAIet al., 1996).Some authors enantioselective steps. SUNJIC,

6

- -

3 Enantioselective Reactions

65

secondary alcohols

aj

OH OH

,LA

OH

PCL, >98% ee for diacetate vinyl acetate Bisht 8 Parmar (1993) Caron 8 Kazlauskas (1993)

PFL (lipase AK), Eoverall>>I00 esterification w/ hexanoic acid Guo et a/. (1990) CAL-B, >99% ee for diol and diester S-ethyl thiooctanoate Mattson, eta/. (1993)

h OH

OH CAL-B, >99% ee for diol and diester S-ethyl thiooctanoate Mattson, et a/. (1993) PCL, Eoverall= 30 vinyl acetate Caron & Kazlauskas (1994)

?H

..JJ3

N3

EoveralF 48, CRL Eover?iI= 7,. PCL hydrolysis of dibutyrate Gruber-Khadjawi eta/. (1996)

primary alcohols

OH Eoverali>loo, CRL, GCL hydrolysis of dibutyrate Gruber-Khadjawi et a/. (1996)

axially chiral diol

PCL, Eoverai1>50 vinyl acetate Sibi & Lu (1994) hydrolysis of diacetate Kawanami et a/. (1994, 1996)

PPL, Eoverall '100 vinyl acetate Guanti 8 Riva (1995)

CEOEoverall >loo hydrolysis of diacetate Kazlauskas (1989, 1991) lnagaki eta/. (1989) Wu et a/. (1985)

Fig. 12. Examples of Cz-symmetricdiols resolved by sequential kinetic resolution include secondary and primary alcohols as well as diols with axial chirality.

hydrolyzed a racemic C,-symmetric tetraester. The non-conjugated ester groups reacted selectively followed by spontaneous decarboxy-

xR

&R

OH PCL, E high PCL, E high vinyl acetate vinyl acetate R = Ph, n-Pr to n-Hx R = OTr, CH20Tr Kim et a/. (1995a) Kim et a/. (1995a) OH

Fig. 13. Sequential kinetic resolution of non-C, symmetric diols.

lation. Interestingly, CRL and RJL favored opposite enantiomers. Although NODEet al. suggested possible racemization of the starting tetraester, which would allow a dynamic kinetic resolution (Sect. 3.7), they did not report yields over 50%. The lack of carboxylic acid examples may be due to more efficient resolution of alcohols by lipases, or to the slow hydrolysis by lipases of monoesters containing a charged carboxylate group (Fig. 14). For substrates with a single functional group, researchers demonstrated a sequential kinetic resolution by in situ hydrolysis of an

66

3 Biotransforrnations with Lipases

COOMe

H

O

W H COOMe COOMe

.olysis arboxylation *

rOH

Me HO oo& H

COOMe

CRL, 32% yield, 100% ee RJL, 20% yield, 90% ee (opposite enantiomers) Node et a/. (1995) Fig. 14. Sequential kinetic resolution of a chiral diacid.

fl

to separate a mixture of meso and racemic diols. The (R,R)-diol reacted to the diacetate, the (R,S)-diol to the monoacetate, and the (S,S)-diol did not react (Fig. 15).

3.2 Asymmetric Syntheses

PCL E o v e r a ~ ~high = acetylation or hydrolysis Wallace et a/. (1992)

Lipase-catalyzed asymmetric syntheses start with meso compounds or prochiral comFig. 15. Enantioselective reactions separated dia- pounds and yield chiral products in up to stereomers as well as enantiomers. 100% yield. In an asymmetric synthesis the enantiomeric purity of the product remains constant as the reaction proceeds and is given ester and reesterification to a new ester (MAC- by ee= (E - l)/(E+ l), where E is the enantioFARLANE et al., 1990).However, reversibility of meric ratio. For example, an enantioselectivity these reactions limited the enhancement of of 50 yields product with 96% ee. Rearrangeet al., 1995). In ment of this equation gives E=(l+ee)/(l-ee), enantioselectivity (STRAATHOF these cases, an in situ recycling reaction (see useful to calculate the enantioselectivity from above) is probably a better way to enhance the the enantiomeric purity of the product. enantiomeric purity. In practice, however, many lipase-catalyzed Enantioselective reactions can also separate asymmetric syntheses undergo a subsequent diastereomers. For example, WALLACEet al. reaction, a kinetic resolution (Fig. 16). For ex(1992) used the (R)-enantioselectivity of PCL ample, hydrolysis of a meso diester first gives

a

> \:

asymmetric synthesis

RS

kinetic resolution

chiral

RS

meso

S'R chiral

b

AcO,

..

5++ i

R'S'

,OAc

--

:

kf

5 ?

minor enantiomer

Fig. 16. Asymmetric syntheses are usually coupled to kinetic resolutions. a Schematic diagram;b PPL-cat(WANGet al., 1984). alyzed hydrolysis of 1,5-diacetoxy-cis-2,4-dimethylpentane


the chiral monoester, but this monoester also reacts giving the meso diol. Although this overhydrolysis lowers the yield of the monoester, it usually favors the minor enantiomer and thus increases the enantiomeric purity of the monoester by kinetic resolution. For the PPL-catalyzed hydrolysis of 1,5-diacetoxy-cis-2,4-dimethylpentane, the enantiomeric ratio for the diacetate to monoacetate hydrolysis was 16

6

67

yielding an enantiomeric purity of 88% ee (WANG et al., 1984). The subsequent kinetic resolution with an enantiomeric ratio of 5 increased the enantiomeric purity to 97% ee, but lowered the yield of monoacetate to -70%. Quantitative analysis of the enantioselectivity in asymmetric syntheses is more difficult than for kinetic resolutions because three variables must be measured: the enantioselectivity of

meso secondary alcohols

HOfi,,,-OBn

OH HO \/O 'H 51% yield, 95% ee OBn PCL, isopropenyl acetate PCL, E >loo, vinyl acetate Laumen & Ghisalba (1994) Harris eta/. (1991)

CAL-B, E >50 vinvl acetate or hydrolisis of diacewte Johnson & B , (1992) ~

prochiral 'alcohol' with remote stereocenter

meso primary alcohols A c O A Ar

OH

u

Q

HO-Ar

PPL. hydrolysis of diacetate PCL, >98% ee, 92-100% yield 88 to >96% ,mwm88 >%% ee, 65,65-80'3/0yield Guanti eta/. (1990b) ' vinyl acetate . Tsuji eta/. (1989), ltoh et a/. (1993~)

02N=Yel

9,

0

R PFL (Amano AK), 97% ee, 31% yield PCL, 88% ee, 21% yield Holdgrijn 8 Sih (1991). Ebiike eta/. (1991) Hirose eta/. (1992, 1995), Salazar & Sih (1995)

HO\ HO,

PCL, vinyl acetate or hvdrolvsis of dibutvrate 4 9 % ee, 8 1 - ~ % yield pcL, 99% ee, 78y0 yield Tanaka etal. (1992) vinyl acetate Mohar etal. (1994) Gais eta'' (1992) also high E with ROL, CVL. RJL 1

meso acid fOOH

PPL, 97% ee. 97% yield hydrolysis of dimethyl ester Nagao eta/. (1989)

prochiral acid

prochiral acid with a remote stereocenter

,COOH

MeOOQSc

-

1 ... . R O *I

PPL, 91% ee. 86% yield hydrolysis of diester Tamai etal. (1994)

>98% ee, 95% yield, hydrolysis of dimethyl ester PCL. Hughes eta/. (1989, 1990, 1993). Smith eta/. (1992) P. aeruginosa lipase. Chartrain etal. (1993)

Fig. 17. Examples of lipase-catalyzed asymmetric syntheses.

68


each step and the relative rate of each step. WANGet a]. (1984) developed the necessary equations, but most researchers only report the enantiomeric purity and yield of the product. For this reason, we will also report only the enantiomeric purity and yield for asymmetric syntheses in this review. Selected examples of lipase-catalyzed asymmetric syntheses are shown in Fig. 17;more are included in the survey of enantioselectivity in the following sections and in an excellent review (SCHOFFERS et al., 1996).The lipase-catalyzed asymmetric syntheses include a wide range of primary and secondary alcohols, as well as carboxylic acids. One example of the advantage of the combined asymmetric synthesis and kinetic resolution is the PCL-catalyzed acetylation of cis-2-cyclohexen-l,4-diol (HARRISet al., 1991), a meso-secondary alcohol. Although the enantioselectivity for first acetylation (asymmetric synthesis) is only 4 and the enantioselectivity for the second acetylation (kinetic resolution) is only 10, the monoacetate was isolated in moderate yield (51%) and high enantiomeric purity (95% ee). Many of the primary alcohol examples are 2substituted 1,3-propanediols, which are versatile synthetic starting materials. The dihydropyridine example (Fig. 17) is a chiral acid, but the acetyloxymethyl group places the stereocenter in the alcohol part of the ester; thus, this prochiral compound can be classified as a chiral alcohol.

3.3 Survey of Alcohols A number of recent reviews also include surveys of hpase enantioselectivity: SIH and WU (1989); CHENand SIH (1989); KLIBANOV (1990); BOLAND et al. (1991); XIE (1991); KAZLAUSKAS et al. (1991); FABERand RIVA(1992); SANTANIELLO et al. (1992); MARGOLIN (1993); MORI (1995); GAIS and ELFERINK (1995); THEIL(1995);SCHOFFERS et al. (1996).The survey below includes only representative examples to give the reader a feel for the type and range of molecules that undergo enantioselective reactions with lipases.

3.3.1 Secondary Alcohols

3.3.1.1 Overview and Models Although lipases show high enantioselectivity toward a wide range of substrates,the most common substrates are secondary alcohols and their derivatives. Researchers have resolved hundreds of secondary alcohols using lipases Selected examples, including asymmetric syntheses of secondary alcohols,are collected below. Based on the observed enantioselectivity of lipases, researchers proposed a rule to predict which enantiomer reacts faster in lipase-catalyzed reactions (Fig. 18; Tab. 7). This rule is based on the size of the substituents and sug-

Tab. 7. Size-Based Rules Similar to Those in Fig. 18 Proposed for Different Lipases Lipase

Comments

Reference

CAL-B CRL PAL PCL PCL PCL PFL PFL PPL PPL RML CE Lipase QL

8 substrates

ORRENIUS et al. (1995a) KAZLAUSKAS et al. (1991) KIMand CHO(1992) LAUMEN (1987) XIEet al. (1990) KAZLAUSKAS et al. (1991) BURGESS and JENNINGS(1991) NAEMURA et al. (1993a, 1995) JANSSENet al. (1991b) LUTZet al. ( 1992) (1989) ROBERTS KAZLAUSKAS et al. (1991) NAEMURA et al. (1996)

86 substrates;reliable for cyclic, but not acyclic,substrates 28 substrates tried to also include primary alcohols and acids 6 substrates 64 substrates 31 substrates 27 substrates 23 substrates 21 substrates 6 substrates 15 substrates 27 substrates


69

so the difference suggests that an electronic effect lowers the enantioselectivity. X-ray structures of transition state analogs containing a secondary alcohol,menthol, bound lipases to CRL identified the alcohol binding pocket Fig. 18. An empirical rule to predict which enan- (CYGLERet al., 1994). This pocket indeed tiomer of a secondary alcohol reacts faster in lipase- resembled the empirical rule: a large hydrocatalyzed reactions. M, medium-sized substituent, phobic pocket and a smaller pocket for the e.g., methyl. L, large substituent, e.g., phenyl. In acy- medium-sized substituent (Fig. 20). A comparlation reactions, the enantiomer shown reacts faster; ison of the structures of the fast- and slowin hydrolysis reactions, the ester of the enantiomer reacting enantiomers of menthol showed that shown reacts faster. the transition state analog for the slow-reacting enantiomer lacks a key hydrogen bond. gests that lipases distinguish between enantio- This observation suggests that enantiomers meric secondary alcohols primarily by com- differ mainly in their rate of reaction, not in paring the sizes of the two substituents. In- their relative affinity to the lipase. Consistent et al. (1997) measdeed, a number of researchers increased the with this idea, NISHIZAWA enantioselectivity of lipase-catalyzed reactions ured the kinetic constants with PCL for two by modifying the substrate to increase the size enantiomers of a secondary alcohol and found of the large substituent for examples see SCILI- similar values for the apparent K,, but very MATI et al., 1988; GOERGENS and SCHNEIDER, different values for k,,,. However, modeling of 1991a, b; KAZLAUSKAS et al., 1991; GUPTAand transition state for ester hydrolysis in CAL-B KAZLAUSKAS, 1993; JOHNSON et al., 1991; KIM suggested that differences in binding may also and CHOI,1992;ROTTICCIet al., 1997). Similar- contribute to the difference in reaction rates of et al., 1995). ly, SHIMIZU et al. (1992) reversed the enantio- the two enantiomers (UPPENBERG Further support for the proposed alcohol selectivity by converting the medium substitubinding site comes from variations in the amient into the large one. To add more detail to this model many no acids within the pocket for the medium subgroups tried to more precisely define the size stituent (Tab. 8). These variations are consislimits of the medium and large substituents tent with differences in the selectivity of lip(PFL: BURGESS and JENNINGS, 1991;NAEMURA ases. For example, smaller amino acids line the et al., 1994, 1995; lipase QL: NAEMURA et al., M region of CRL (Glu, Ser, Gly) than in PCL 1996; PCL: THEILet al., 1995; LEMKEet al., and CVL( = PGL) (His, Leu, Gly). If the back1997) while others have tried to include elec- bone lies in the same place for both lipases, then the smaller side chains in CRL create a tronic effects (PCL: HONIGet al., 1994). Although steric effects are the most impor- larger binding site. Consistent with this suggestant determinant of lipase enantiopreference, tion, CRL catalyzes the hydrolysis of esters of electronic effects also contribute. For example, large alcohols (esters of norborneols; OBERthe CAL-B shows high enantioselectivity to- HAUSER et al., 1987) and esters of tertiary alcoward 3-nonanol ( E > 300), but low enantiose- hols (O'HAGANand ZAIDI,1992), while the lectivity toward 1-bromo-2-octanol (E= 7.6) Pseudomonus lipases do not. Using substrate under the same conditions (Fig. 19). Both an mapping EXLet al. (1992) found t,hatCRL had ethyl and a -CH,Br group are similar in size, a larger alcohol binding site than*-PCL.Further, CRL shows low enantioselectivity toward esters of primary alcohols, while the PseudoOH OH monus lipases show moderate enantioselectivdc6H13 Brdc6H13 ity. All of these characteristics are consistent E = 7.6 E>300 with a larger binding site in CRL. CAL-B, acylation w/ Sethyl thiooctanoate Because the same size rule works for all Orrenius eta/. (1995a), Rotticci eta/. (1997) lipases, CYGLERet al. (1994) suggested that Fig. 19. Electronic effects also change enantiose- structures common to all lipases cause this enantiopreference. Indeed, all lipases follow lectivity. OH

70

3 Biotransformationswith Lipases

b

Fig. 24). Proposed binding site for secondary alcohols in CRL. a X-ray structure of CRL highlighted to show the catalytic machinery (Ser 209, His 449,Glu 341 and the N-H groups of Ala 210 and Gly 124) and the alcohol binding site. b Schematic of the first step of hydrolysis of an ester of a secondary alcohol.The alcohol oxygen orients to form a hydrogen bond with the catalytic His, while the large and medium substituents orient in their respective pockets.

the dphydrolase fold and have similar catalytic machinery. On the basis of X-ray crystal structures of chiral transition state analogs bound to the active site of CRL, OGLER et al. (1994) suggested that the loops that assemble the catalytic machinery also assemble an alcohol binding site that is similar to the rule in Fig.

18. A large hydrophobic pocket open to the solvent can bind the large substituent, while a restricted pocket near the catalytic machinery can bind the medium substituent (Fig. 20). Although all lipases favor the same enantiomer of secondary alcohol, subtilisin favors the opposite enantiomer, but the enantio-

Tab.8. Amino Acid Residues in 11 Lipases Showing the Catalmc Triad (Bold), in the Oxyanion Hole (Bold Italics), and in the Proposed M Binding Site (Italics) Lipase

Consensus Sequence Near Nucleophilic Ser Catalytic H i s GIy-X-Ser-X-Gl y

Catalytic Oxyanion Hole" AsplGlu

CVL, PCL Rh4L HLL PcamL ROL PPLb CAL-B CRL (lipl) GCL (lipII)

-Gly85-His8~er87-Gln88-Gly8e

-Asp263-AspU)> -AspU)l-Asp199-Asp-Asp176 -Asp187-Glu341-Glu354-

-HIs285-Leu28&

-Gly142-Hisl43-Serl44-Leul4S-Glyl46- -His2sI-Leu258-Glyl44-Hisl45-Serl46-Leul47-Glyl48- -Hisz58-Leu259-Gly143-Hisl44Ser14S-Leul46-Gly147- -His25!4-Zle26& -Gly143-H~l44Serl4S-Leu146-Gly147- -His257-Leu258-Glyl5O-Hisl5l-Serl52-Leul53-Glyl54 -His263-Leu264 -~lO3-TrplO4!3erlO5-GlnlO6-GlylO7- -His224-AIa225-Gly207-Glu208-SerZla21&Gly211 -His44%Ser45W -Gly215-Glu2I&Ser217-AIa218-Gly219 -His46>Gly464-

-Glyl&Leul7-Gly81Scr82-Gly82Srr83-Gly834er84-Gly82-Thr83-Gly7&Phe77-Gly39-Thr4& -Gly123-GIy124-Gly131-AIa132-

All lipases use two hydrogen bonds from the amide N-H of the two residues shown in bold italics. In addition, structures of transition state analogs show that RML stabilizesthe oxyanion with a third hydrogen bond from the hydroxyl group of Ser or Thr. Other lipases with a Ser or Thr at this position may also use this third hydrogen bond. The corresponding residues in the human enzyme are identical. 'The minor isozymes of CRL contain either Gly or Ala in place of Ser450. a


3.3.1.2 Candida antarctica Lipase B

selectivity is usually lower (KAZLAUSKASand WEISSFLOCH, 1997). One way to reverse the enantiopreference of CAL-B toward secondary alcohols is to place the alcohol in a different place within the binding site. For example, aminolysis of an allyl carbonate derivative (Eq. 9), replaces the allyl group (Pozo and GOTOR,1993b).

R

. I o%

A

R

0 CAL-B,NHzBn

n-hexyl, Ph, Et; E >50

OANnPh

*R A H

(9)

In this reaction, RCHMeOC(0)- behaves as the acid portion of an ester; thus, the alcohol stereocenter probably binds in the acid binding site. Of course, the secondary alcohol rule no longer applies to this reaction. Pozo and GOTOR(1993b) found that CAL-B favored the alcohol enantiomer opposite to the one predicted in Fig. 18.

E >150. Sethyl thiooctanoate Orrenius et al. (1995a) E = 22, diketene Suginaka eta/. (1996)

71

CAL-B and PCL are usually the most enantioselective lipases toward secondary alcohols, see Figs. 22-24 for CAL-B, Figs. 30-32 for PCL. All examples follow the empirical rule in Fig. 18. CAL-B is more enantioselective toward secondary alcohols where the medium-sized substituent (M in Fig. 18) is relatively small, e.g.,methyl, ethyl, -C=CH, -CH=CH2. Reactions are slower, but still highly enantioselective when M is n-propyl or -CH20CH3, but no reaction occurs when M = i-propyl or tbutyl as in the substrates in Fig. 21 (ORRENIUS et al., 1995a). In contrast, PCL accepts longer n-alkyl chains as the M substituent (Fig. 31). CAL-B also resolves a number of 2-substituted cycloalkanols (Fig. 24). Many truns-

Fig. 21. No reaction w i t h CAL-B.

R= Bn

CAL-B, E >I00 S-methyl thioacetate Trolls& et a/. (1996)

L

E

T

Ph 35 1-naphthyl 57 2-naphthyl 66 CAL-B. diketene Suginaka et a/. (1996)

CAL, vinyl acetate, E >lo0 Hamada eta/.(1996)

LR R = H, n64Hg E = 70, CAL, vinyl acetate

% To Petschen etal. (1996)

X = CI, Br R = H, E = 1.3 E >loo R = SiMe3, E >lo0 acylation w/ Sethyl thiooctanoate Rotticci et a/. (1997)

% A (OH& &OH /OH OH 73 - >99% ee for diacylated products Mattson et a/. (1993, 1996)

Fig. 22. Selected examples Of 2-alkanols resolved by CAL-B.

E 4 0 0 , Sethyl thiooctanoate Frykman eta/.(1993) Orrenius etal. (1994)

OH

72

3 Biotransformations with Lipases OH Jc6H13

dC6H13

// A C

6 H 1 3 CAL-B, E >I50 Sethyl thiooctanoate Orrenius eta/. (1995a)

E=10 CAL-B, E = I 3 vinyl acetate hydrolysis of chloroacetate Kato e l a/. (1996) Kato et a/. (1994)

4,

LR

C M C H Z (CH2)9 ) ~

F3C

CAL-8, E = 78, vinyl acetate tentative absolute configuration Ohtani eta/. (1996)

R = H, n64H9, n-C5H11, n-C~H17 E =50,CAL, vinyl acetate Petschen etal. (1996) OH F3

OH MeO&O-R

OAc

CF3 CAL. hydrolysis of diacetate n = 1, 3, 5; E >>lo0 ltoh eta/. (1996b)

E CH2Ph -8 CH2CH2Ph -20 Partali et a/. (1993)

, . I

F3CL

OH F3CL

I

R

CAL, vinyl acetate, E = 100 R = Me, Et, Pr, Bu, CH20Bn Harnada etal. (1996)

S

4

V

V

R

E Ph 22 - 23 CHzPh 18-22 CH2CH2Ph >I00 CHzCH20Ph >55 Waagen et a/. (1993)

R -

R -

CAL-B, 'E' >lo0 isopropenyl acetate Hoye et a/. ( 1996)

zBr

J C 6 H 1 3 X = CI, Br E = 8-14 acylation w/ S-ethyl thiooctanoate Rotticci et a/. (1997) E=81

R = H, Et. n-C4Hg E= 100, CAL. vinyl acetate Petschen et a/. (1996)

Mattson et a/. (1996)

Fig. 23. Selected examples of acyclic secondary alcohols resolved b y CAL-B.

oriented substituents gave high enantioselectivity, but a cis hydroxyl gave low enantioselectivity. The X-ray crystal structure of CAL-B shows a deeply buried M-pocket (or alcohol stereoselectivity pocket) large enough to accommodate a methyl or ethyl group without changing the conformation of the protein (see Fig. 3).

3.3.1.3 Candida rugosa Lipase In a previous survey of secondary alcohols resolved by CRL, KAZLAUSKASet al. (1991)

found that the secondary alcohol rule in Fig. 18 is not reliable for acyclic alcohols. Out of 31 examples, only 14 followed the rule. Since there are only two choices in predicting the fastreacting enantiomer, even guessing yields 50% correct predictions. Thus, the rule is little better than guessing for acyclic alcohols. More recent examples (Fig. 25) include four examples that follow the rule, three that do not and two with either an uncertain absolute configuration or large and small substituents with similar sizes. Compared to other lipases (especially CAL-B, PCL, and RML) CRL accepts larger substrates and the X-ray crystal structures


e; 4,Q &:;

,.NHBoc

'OH

CAL-B, E >50 S-ethyl thiooctanoate isopropenyl acetate Mattson et a/. (1996) Sundram et a/. (1994)

B

P

,

,

,

73

n = 1-3; E = 2-3

OH

CAL-B, E >50 vinvl acetate vinyl acetate or Nicolos; et a/. (1995a) hydrolysis of diacetate Johnson & Bis (1992)

& &

~& i & X & '

/

/

X = Br, I; E > l o 0 ~ = a - 3 2 E 2100, CAL-B isopropenyl acetate vinyl acetate E = 29, diketene vinyl acetate (1997) Johnson 8 Sakaguchi (1992) Orrenius et a/. (1995b) Suginaka etal. (1996) lgarashi

..OMe S-ethyl thiooctanoate Mattson etal. (1996)

E >200 acylation with Sethyl thiooctanoate

CAL-B, E > 50 CAL-B, E >15 E >200 vinyl acetate isopropenyl acetate vinyl acetate Frykman eta/. (1993) Gustafsson et a/. (1995) Stead et a/. (1996) B a n et a/. (1996)

Fig. 24. Selected examples of cyclic secondary alcohols resolved by CAL-B.

CRL, E high R = Et, C s H l j Ph. CHzCH=CMe, CHOHMe esterification with tributyrin Cambou & Klibanov (1984)

Fig. 25. Selected examples of acyclic secondary alcohols resolved by CRL. Note that the several examples do not follow the empirical rule in Fig. 18. This rule is not reliable for CRL-catalyzed reactions of acyclic secondary alcohols.

CRL, E = 2-4 CRL. E = 10, vinyl acetate vinyl acetate Kaminska et al. (1996) Fiandanese eta/. (1993) follow rule in Figure 18 CRL, E >lo0 CRL, E = 20 to >lo0 vinyl acetate monoacetylation only R = malkyl Pai eta/. (1994)

U 4

C

F

3

CRL, , E >lo0 vinyl acetate Hamada et a/. (1996) hydrolysis of acetate Yonezawa et a/. (1996)

H

O TPh Ph

CRL. E >50 vinyl acetate Nicolosi eta/. (1994b)

PhS+JC5Hl SPh CRL, E >lo0 hydrolysis of acetate Pai et a/. (1994)

7

do not follow rule in Figure 18

74


OAc

PhSJ

MeOACC13

I

R

SPh

CRL, PPL, CE, E >50 hydrolysis of acetate abs. config. not established Chbnevert et a/. (1990)

CRL, E = 30 to *I00 R = P r , aryl, CMe=CH2 hydrolysis of acetate Pai et a/. ( 1994)

substituents w/ similar sizes or unknown abs configuration

Fig. 25. Continued.

R CRL, E = 50 CRL, E *50 hydrolysis of butyrate Klempier et a/. (1990)

CRL. E = 27 hydrolysis Of acetate Cotterillef (I 991

OH

CRL, E >20,slow eta’’ (IgM)

0

CRL. E =61-64 R = CPr, Ph hydrolysis of butyrate Cotterilleta’’ (1991) Banziger eta/. (1993a)

OH

E ~ 1 0 0CRL , EGz:fetEL hvdrolvsis of dibutvrate Crotti’et a/. (1996) Grdber-Khadjawi et a/. (1996)

CRL, E -50 esterification w/ lauric a. Ar = Ph, 4-t-BuPh Comins & Salvador (1993)

. .. Ph

CRL, E = 125 vinyl acetate

U

U

OAc OAc CRL, E high hydrolysis of diacetate Kazlauskas eta/. (1991)

CRL, E =39 CRL, E >50 hydrolysis Of acetate hydrolysis of formate Hoenke eta/. (1993) Akita eta/, (1997)

& @

CRL. E = 10 to >50 esterification or transesterification: Langrand etal. (1985, 1986) et (lga5)* Lokotsch eta/. (1989) Rabiller eta/. (1990) XU eta/. (1995) . hydrolysis: Yamaguchi etal. (1976) Cygler etal. (1994)

TBS

AcO AcO OMe CRL, 298% ee, 61% yield CRL, >98% ee, 48% yield CRL, >98% ee. 61% yield hydrolysis of diacetate hydrolysis of diacetate hydrolysis of diacetate Pearson & Lai (1988) Pearson & Srinivasan (1992) Pearson etal. (1987)

E ~ 1 0 0CLEC-CRL , vinyl acetate Khalaf et a/. (1996)

Fig. 26. Selected examples of cyclic secondary alcohols resolved by CRL. Two examples do not follow the rule in Fig. 18: second row, first structure and third row, second structure.


I ( C H 2 ) " A

E = 100

E = 26 vinyl acetate Wang et a/. (1988) E = 90 trifluoroethyl laurate Morgan et a/. (1991, 1992)

R = Et, E = 2.5 n=O,E=41 trifluoroethyl laurate R = Pr, E = 52 n = 1, E > 100 Stokes & Oehlschlager (1987) R = Bu. E > 100 n = 2, E > 100 R = -&HI1, E = 92 n = 3, E = 80 E=17 R=-C*H18, E > 100 vinyl acetate R = -(CH&,C02Et, E = 70 Wang et a'. ( l 988) R = -(CH&,CH=CHCO*Et, E = 75

-

E = 17 71 E = 47 esterification with octanoic acid trifluoroethyl butyrate Secundo et a/. (1992) Yang eta/. (1995a)

trifluoroethyl laurate Morgan eta/. (1991, 1992) OH

0

OH

L/

-OH E=60-70 trifluoroethyl butyrate Morgan et a/. (1991, 1992)

E = 75-76

E = 60, trifluoroethyl butyrate Ramaswamy & Oehlschlager (1991)

E = 14-15

R = C7H15, Ph. E > 100

E = 28-30

R = C7H15, E = 6 R = Ph, E = 2.5

wc7H15 k E=13

R

E400 methyl propionate, Janssen et a/. (1991) trifluoroethyl laurate, Morgan et a/. (1991, 1992) vinyl butyrate, Ottolina eta/. (1994)

%

trifluoroethyl butyrate R = C7Hj5, E > 100 Morgan et a/. (1991, 1992) R = Ph, E = 51

JJ)

E=15-29 trifluoroethyl laurate Morgan et a/. (1991, 1992)

E = 62 - 71 Morgan et a/. ( l 991 9g2) trifluoroethyl lauratemethyl propionate methyl propionate, Janssen eta/. (1991) Janssen eta/. (1991) I

methyl propionate, Janssen ef a/. (1991) ?H

-&Me

PPL, R = Me, Et, Pr, E >lo0 2,2,2-trifluoroethyl pentanoate Chong & Mar (1991)

75

E=20 methyl propionate Janssen eta/. (1991) E = 400 vinyl acetate Kaminska et a/. (1996)

Fig. 27. Examples of 2-alkanols resolved by porcine pancreatic lipase.

%

E = 20.5 trifluoroethyl laurate Morgan et a/, (1991, 1992)

76


3.3.1.5 Pseudomonas Lipases

show a larger active site. This larger active site may allow acyclic substrates to react in several productive conformations. Some of these conformations may favor the opposite enantiomers. In contrast, cyclic secondary alcohols reliably follow the rule in Fig. 18. Of the 55 substrates in an earlier survey, 51 followed the rule (KAZLAUSKASet al., 1991). All but two of the selected examples in Fig. 26 follow the rule.

The Pseudomonas lipases show high enantioselectivity toward a wide range of secondary alcohols (Figs. 30-32). A previous survey also includes 64 secondary alcohols (KAZLAUSKAS et a]., 1991). Except for increasing the difference in size of the substituents (see Sect. 3.3.1), or lowering the temperature (for an example see SAKAI et al,, 1997), no general method exists to increase the enantioselectivity of PCL-catalyzed reactions. Longer esters or acylating agents (butyrates and above) sometimes give higher enantioselectivity than acetates. Hydrolyses of P-(pheny1thio)- or /3-(methy1thio)acetoxy groups increased enantioselectivity 10-fold (e.g., from 6 to 5 5 ) as compared to hydrolyses of acetates or valerates (ITOHet al., 1991). Thiocrown ethers (e.g., 1,4,8,11-tetrathiacyclotetradecane) increased the enantio-

3.3.1.4 Porcine Pancreatic Lipase All examples of PPL-catalyzed reactions of secondary alcohols follow the rule in Fig. 18. The examples are divided into 2-alkanols in Fig. 27, other acyclic secondary alcohols in Fig. 28, and cyclic secondary alcohols in Fig. 29. Note in Fig. 27 that the cis vs. trans configuration of the double bond in the large substituent strongly influenced the enantioselectivity.

E >lo0 trifluoroethyllaurate Morgan et a/. (1991, 1992)

R = n-Pr, E = 35-50 R = ~ - B uE, = 25 R=n-C5H11.E=35 R = CHzBr, E = 55-60

methyl propionate Janssen eta/. (1991)

methyl propionate Janssen et af. (1991)

-

E = 8. PPL hydrolysis of acetate Treilhou et af. (1992)

D >lo0 Configuration o f ’ * ’ stereocenter set by synthesis;diastereoselective hydrolysis of acetate Mulzer et a/. (1992)

OH C&OP ,h

OH E=4-15

H O W L R

OH “‘vCo-ph

E-1

Partali et a/. (1993)

PPL, Eoverall= 7 - 18 R = H, Me, Et, Hx hydrolysis of diacetate Poppe etaf. (1993)

0

PPL, E = 9-23 R = n-heptyl. OPh, OBn, CHZOBn, CH2CHZOBn hydrolysis of cyclic carbonate Matsumoto eta/. (1995, 1996)

PPL. E = 6 hydrolysis of cyclic carbonate Matsumoto eta/. (1995, 1996)

Fig. 28. Examples of other acyclic secondary alcohols resolved by porcine pancreatic lipase.

Q

PPL. E >50 hydrolysis of butyrate Klernpier et a/. (1990)

OAc

E 4 0 0 , R = Ac hydrolysis gf rneso diacetate Sugai 8 Mori (1988) E>100. R = H vinyl acetate Theil eta/. (1991)

QH

PPL, E >50 hydrolysis of acetate Cotterill et a/. (1988a,b)

6

OH

E >loo. R = But hydrolysis of meso dibutyrate Laurnen & Schneider (1986)

O

PPL, E >300 hydrolysis of acetate Cotterill et a/. (1990, 1991)

6

cI/

R

R = -CH2C=C(CH2)3COzMe R = -CHzCaCH E >loo, vinyl acetate Babiak ef a/. (1990)

PPL, methyl acetate, E -1 1 Hemmerle 8 Gais (1987)

bTBS

high E hydrolysis of acetate Mori 8 Takeuchi (1988)

R = -(CHz)zCH=CH(CH2)2COzMe R = -(CH~)$IC(CH~)~CO~M~

OH

PPL, E = 95; PCL, E = 25 vinyl acetate Takano eta/. (1991)

77


OH

n=3, E = 65 rnethvl DroDionate Janssen eial. (1991)

PPL. E >lo0 vinyl acetate Crotti et a/. (1996)

Fig. 29. Examples of cyclic secondary alcohols resolved b y porcine pancreatic lipase.

selectivity of PCL four-fold (from 9 to 27-37 and from 100 to 400) in the resolution of several secondary alcohols (ITOH et al., 1996a; TAKAGIet al., 1996). ITOH et al. (1993a) observed smaller increases with simple crown ethers.

3.3.1.6 Rhizomucor Lipases Researchers have resolved fewer secondary alcohols using RML (Fig. 33). In triglycerides, RML selectively hydrolyzes esters at the primary alcohol positions (see Sect. 4.2.1), but the examples below show that RML can also hydrolyze esters of secondary alcohols. NORITOMI et al. (1996) increased the enantioselectivity of an RML-catalyzed acylation

of 1-phenylethanol by lowering the temperature. Acylation with vinyl butyrate in dioxane gave E = 19 at 45 "C, but E = 170 at 7 "C. In other solvents (pyridine,THF, triethylamine) temperature did not affect E.

3.3.1.7 Other Lipases Other lipases are also enantioselective toward secondary alcohols and usually follow the rule in Fig. 18.Selected examples are in Fig. 34. NAEMURA et al. (1996) surveyed the enantioselectivity of a lipase from Alcaligenes sp. and found good enantioselectivity toward a range of secondary alcohols (27 examples, mostly MeCH(0H)aryl). The favored enan-

78


X P h PCL. E >lo0 vinyl acetate or hydrolysis of acetate LWfnen et (1988) Laumen & Schneider (1988)

PCL, PFL, E >50,X = H, F hydroysis of acetate Gaia e l a/. (1996)

slow rxn PCL, E >150, vinyl acetate Gaspar & Guerrero (1995) Ferraboschi eta/. (1995b)

PCL, R = Me E >150,Sethyl thiooctanoate Orrenius et a/. (1995a) E = 22, diketene Suginaka et (,996) PCL, PFL, R = Et E = 12-13, vinyl acetate Hamada et a/. (1996)

PCL, E >lo0 hydrolysis of chloroacetate BBnziger et a/. (1993b)

PCL, E = 70 vinyl acetate Kaminska eta/. (1996) OH ,LCN

PCL, hydrolysis of acetate PCL, E >50 PCL, E >50 PFL (lipase K-lo), E >20 E=10-35 vinyl acetate hydrolysis of chloroacetate vinyl acetate ltoh et a/. (1993a, 199W Kim Choi (1992) hang & Paquette (1990) Burgess & Henderson (1990) OH /

L

x N 0 2

, R -1 j

A

P

PFL (Amano AK) E = 48 (THF) vinyl acetate Nakamura etal. (1991) Kitayama (1996)

h

.)'n-CloH21 '!*Ph

R = Ph, Bu, n-Oct, SiMe3

E 720

E = 5-20

PFL (lipase AK). vinyl acetate, Burgess & Jennings (1991) OH L S i M eB u-3t,,)

&

OH E 50, Pseud. lipase or Pseud. cholesterol esterase PCL, >98% ee for diacetate esterification w/ 5-phenylpentanoic acid vinyl acetate Bisht & Parmar (1993) Uejima eta/. (1993) Caron & Kazlauskas (1993)

PFL (Amano AK). E >lo0 n = 0, 1, 5,10 hydrolysis of acetate Scilimati eta/. (1988)

PFL (lipase AK). Gverall >>loo esterification w/ hexanoic acid Guo et a/. (1990)

PFL. 97% ee, 83% yield hydrolysis of diacetate Adje etal. (1993)

Fig. 30. Selected examples of 2-alkanols resolved by Pseudomonas lipases.


79

-t

R PCL, vinyl acetate PCL, R = CICH2, Et, E '50 R = ph, E >loo R = n-Pr. E = 10 PCL. hydrolysis of acetate R=n-C&,E=16 vinyl acetate E = 16- 17.5 kng et (1995) Haase 8 Schneider (1993) Takagi eta/. (19%) Kim 8 Choi (1992) E >100, R 'ndlkyl pcL

CWe

'

&

""w

' k p h

q

R

E >lo0 R =Me, n-C5H11, n-C~H17 PCL, hydrolysis of acetate or 2-(thiornethyl)acetate ltoh et el. (1990), ltoh 8 Ohta (1990)

PFL (lipase K-lo), E >20, vinyl acetate Burgess Henderson (1990)

BU

s E n= 6 M

d

H2&C.ph

* ) b p h

.kPh

Dph E = 5-20

E >20

P h, j

E >20 PFL (lipase AK), vinyl acetate, Burgess 8 Jennings (1991) i e Et n-Pr

OH L OOH- $ * R&OSiMe+Bu R = Me, Et, CH$I, CH=CH2, CH20CH=Cl+ L 0 -

I

R

4

E>100. PCL (SAM-2) hydrolysis of butyrate or chloroacetate Goergens 8 Schneider (1991b)

R&R

4-",iMe3

~

'2

K

N

njgl:

PCL nCBH13 trifiuoroethyl butyrate nC7H15 Aiievi eta/. ( 1 9 ~ ) nCeH1-1 nCsH19 n41oH21 Ph

\

a

E7

27

;X 32 50

34

27 25 58

0

E>100, PCL (SAM-2) Pseud. lipase E ~ 1 0 0CRL, , R = H. Me; R = t-Bu, Ph O O M e esterification wl Sphenylpentanoic acid hydrolysis or acylation Uejima et a/. (1993) PCL, E M O O vinyl acetate or hydrolysis of acetate Goergens 8 Schneider (1991a) Takano ef a/. (1993e)

Fig. 31. Selected examples of enantioselective reactions of Pseudomonus lipases with other racemic acyclic alcohols Note that substituent type and substituent location in the aromatic ring strongly influences the enantioselectivity of some reactions

80


PAL, E >lo0 alcoholysis of chloroacetate w/ hexanol Kato et a/. (1996)

.r

NC

s

PCL, vinyl acetate

G~~~~~ 8 G~~~~~~~ (1995) Kato etal. (1995a)

R = Ph, 4-CI-Ph, 4-Me-Ph,

1-naphthyl, 2-naphthy1, CH2CH2Ph.

R

-

PCL, E typically 10 20 hydrolysis or acylation van Alrnsick et a/. (1989) Hsu et a/.(1990) lnagaki eta/. (1991, 1992)

E high, PCL, vinyl acetate low selectivity toward '*' stereocenter Danieli eta/. (1996)

H02CLPh

HOzC

Me02C

ph

PCL, E >50 PCL, E >50 vinyl acetate or hexanoic anhydride vinyl acetate Sugai 8 Ohta (1991) Chadha 8 Manohar (1995) Chadha 8 Manohar (1995) OH

E >loo, PCL, vinyl acetate X NOz. NH2. NHC(0)R. NHC(0)OR Kanerva 8 Sundholm (1993a.b)

MeOZC MeOzC OR PFL, hydrolysis of acetate tentative abs. config. R = we, Et, E = 30,80 Milton etal. (1995) OH EtOZQLO-I-naphthy PCL, vinyl acetate, E >loo CVL, vinyl acetate, E = 79 Bornscheuer et a/. (1993) Wunsche et a/. (1996) OH@OMe EtOOC

NHBz pcL, = 58 vinyl acetate Barco etal. (1994)

M

OH

e

PCL. E >I00 hydrolysis Desai et al. of (1996) acetate

Fig. 31. Continued.

2

L

R

Et02&ph

l

PCL, vinyl acetate. E >loo R 4 4 e 0 , 2-CH2CH=CHz Bornscheuer et (1993) Wunsche ef a/. (1996)

4

MeOZC OAc

0

PCL. vinyl acetate PCL. vinyl acetate, E >lo0 R = H, E = 16 Bornscheuereta/. (1993) R = *C3H7, E = 7 Bornscheuer eta/. (1993)

R R = ~CaH17.E -100, PCL butanoic R = *C6H13, or succinic E = 60, anhydride PCL Fukusaki eta/. (1991, 1992a)

PCL. E 40. vinyl acetate. Lefker eta/. (1994)

r? % 'A

KAr0'0;.!

EtOZC

0

PCL, E = 7 - 9, vinyl acetate, Lefker eta/. (1994)

Me02C&>% PFL b a s e K-10). E >20. vinvl acetate burgess 8 Henderson (i990)

81


J y

R*0 x 0 dS.Q0 -

PCL. palmitoyl anhydride, E >I00 R mC4, CIO, c16 Chenevert 8 Gagnon (1993)

OH

OH

MeO&O.R

R

Ph CHZPh

RO
lo0 2 5 R = CHZPh. 4-MeOCeH4 Takano etal. (1993~)

CHZCHZOPh 2 Waaaen et a/. , 1993

-

F &OM.

A c O L O

0

OH

A c o d p h PCL,E=50-130 vinyl acetate Lemke et a/. (1996)

4-Me (55j; 4-OMe (>iooj; 4-CI (85jj d - t - ~ u(>loo) 2,3-C4H4 (12) PCL, vinyl acetate, Theil eta/. (1994)

Fig. 31. Continued.

tiomer was the one predicted by the empirical rule in Sect. 3.3.1.

3.3.1.8 Choosing the Best Route The best route to a particular compound is rarely a straightforward choice. In addition to several lipase-catalyzed routes, researchers consider other chemical and biochemical routes. The “best” is often an individual deci-

Hsu etal. (1990) Bevinakatti 8 Banerji (1991) Ader & Schneider (1992)

sion which depends on the intended next steps and available starting materials. The two examples below summarize only the lipase-catalyzed routes to these targets.

3.3.1.8.1 Inositols Researchers have found a number of different routes to enantiomerically pure inositol derivatives. Starting from the achiral myo-ino-

82


racemic cyclic secondary alcohols HO x,Cl

&Cl

0 4 p P h

H

CI PFL (Biocatalysts), E = 53 hydrolysis of acetate PCL, PFL, E >50 PFL, E >50 PFL, E '50 hydrolysis of butyrate, Klempier ef a/. (1990) Cotterill ef a/. (1991)

&

,...Br

OBn \

OBn

PCL, E >50 hydrolysis of acetate Chen eta/. (1992)

h

PCL, E = 60 hydrolysis of acetate or acylation w l AcpO Ghosh & Chen (1995)

M

$ ~

~

~

\

OSiMept-Bu

0

0 PCL, E 4 0 0 hydrolysis of acetate Sugahara et a/. (1991)

S

PCL. E >lo0 vinyl acetate Takano ef a/. (1993d)

e 0 p . t

~

"loo

.*.**

$r

hydrolysis PCL, Eofhigh acetate 0 Siddiqi eta/. (1993) PCL, E >300 pcL, >50 vinyl acetate PCL, isopropenyl acetate vinyl acetate Cotterill eta/. (1991) Ar = Ph, E -90 Ar=4-MeOCsH4, E -30 Merlo eta/. (1993) Biadatti ef a/. (1996) V

&OpEt

pcL,

PCL. E >75 vinyl acetate MacKieth et a/. (1993)

E=13 E>50 E=12 PCL. hydrolysis of chloroacetyl ester Maleuka & Paquette (1991) Borrelly 8 Paquette (1993) Lord ef a/. (1995)

OR

PCL, E = 75 to >lo0 PCL, E = 4 0 -60, R = H, Me PFL, E >50 PFL, E = g vinyl acetate or hydrolysis of acetate R = Trityl. vinyl acetate vinyl acetate slow reaction Thuring ef a/. (1996) MacKeith et a/. (1994) Takano et a/, (1992a, b) R = TBDMS, hydrolysis Of acetate Roberts & Shoberu (1991) Evans et al. (1992)

PCL, E 250, vinyl butyrate R = C(O)NM%, CHpOTBDMS Ema ef a/. (1996)

E = 5-7, PCL. PFL, others vinyl acetate Mitrochkineet a/, (1995a) lgarashi et a/. (1997)

E >loo, PCL, vinyl acetate Takahashi 8 Ogasawara (1996)

Fig. 32. Selected examples of enantioselective reaction of cyclic secondary alcohols catalyzed by Pseudornonas lipases.

~

~


n

83

OH

JOH

v

PCL, E o v e r a ~>2000 hydrolysis of diacetate Caron & Kazlauskas (1991) Laumen et a/. (1989)

PCL, isopropenyl acetate E high, Meng eta/. (1996) OH

/ C, C O O E t

U

PFL, E = 92 vinyl acetate Panunzioet a/. (1997) OH

OH

Q, Q,

w-

"PCL, E >50 vinyl acetate Yang eta/. (1995)

PCL, E = 35 E >loo, PCL, PFL ( lipase AK) PFL (Amano AK), E = 20 vinyl acetate Crotti et a/. (1996) diketene Suginaka et a/. (1996)

OH

E >lo0 I

vinyl acetate Bovara eta/. (1991)

Kj,

C02Me PFL, E 250 hydrolysis of acetate tentative abs. config. Allen &Williams (1996)

,n,:;o oN

0

PCL, E >I00 vinyl acetate Hoenke et a/. (1993)

6

Ph PFL, E = 33 hydrolysis of acetate tentative abs. config. Allen & Williams (1996)

4

,,SPh

-

E = 11 17, R = CH=CH2 E=8, R=Me PCL, vinyl acetate Sugai et a/. (1996)

0

%O

PCL, E >loo, vinyl acetate Yamada & Ogasawara (1995)

C ' bz

E>IOO, PCL, vinyl acetate Sakagami eta/. (1996a,b, 1997)

meso cyclic secondary alcohols

U -

OH HO HO OH OBn >99% ee, 100% yield 6H >99% ee, 89% yield 95% ee, 51% yield >99% ee, 89% yield PCL, acetate lipase from Toyobo, vinyl acetate PCL, vinyl acetate Miyaoka et a/, (1995) Sugahara & Ogasawara (1996) PCL, isopropenyl acetate Harris eta/. (1991) Laumen 8 Ghisalba (1994) Fig. 32. Continued.

6 a$ bR6

84


63

*+yL

OH '"0

OH >99% ee. 79% yield '95% ee, 90% yield, PCL PCL, vinyl acetate isopropenyl acetate Takano eta/. (1993a, b) Johnson eta/. (1991)

U

U

PCL, 88% ee, 100% yield RML, >98% ee, 94% yield vinyl acetate Nicolosi et a/. (1995a)

PCL. ~ 9 8 % ee, 89% yield RML, 95% ee, 93% yield vinyl acetate Patti et a/. (1996)

e;:.

"'OTBDMS

OH

OH

E = l l Ei100 PFL (Amano YS) monoacylation with phenyl esters Naemura ef a/. (1993a)

?H

HO PCL,>98% ee, 91% yield R = NHCbz, NHBoc, OTBDMS R = N3; E not determined isopropenyl acetate Johnson 8 Bis (1995) Johnson ef a/.(1993)

0%

HO OH PCL, 80% ee, 92% yield isopropenyl acetate Bis et a/. (1993)

HO

PCL,>95% ee, 95% yield isopropenyl acetate Johnson et a/. (1993)

PCL. >98% ee, 90% yield isopropenyl acetate Johnson et a/. (1994)

OH

PCL,>98% ee, 89% yield isopropenyl acetate Bis et a/. (1993)

HO

PFL (Amano AK), >99%ee, 96% yield PCL, 98% ee, 23% yield CRL, 94% ee, 10% yield vinyl acetate Toyooka et a/. (1993)

Fig. 32. Continued.

which contain a secondary alcohol stereocenter. The (S)-enantiomer is usually more active than ( R ) ,e.g., 100 times more active in the case of propanolol. For this reason, chemists have developed routes to the (S)-enantiomer, including lipase-catalyzed reactions (Fig. 36) (reviewed by KLOOSTERMAN et al., 1988;SHELDON, 1993). None of these routes have been commercialized. In most examples in Fig. 36 the aryl group is the large substituent, thus the (R)-enantiomer reacts faster. Resolution by acylation of the alcohol is preferred over hydrolysis of the acetate because acylation yields 3.3.1.8.2 P-Blockers the desired (S)-alcohol as the unreacted startMost /3-adrenergic antagonists (p-blockers), ing material. A resolution by hydrolysis reused for the treatment of hypertension and an- quires an extra step to hydrolyze the unreacted gina pectoris, are 3-aryloxy-2-propanolamines ester.

sitol, researchers added protective groups to increase the size of one substituent. Protection yields either a racemate or a mem derivative. Several different lipases showed excellent enantioselectivity. The asymmetric synthesis starting from meso derivatives is probably the best route since it gives both high yield and high enantioselectivity.However, in some cases another route may fit better with subsequent synthetic steps (Fig. 35).

85

3 Enantioselective Reactions 241kanols

% HA

other acyclic secondary alcohols OH

' A 0 * p h

R = Bu, E = 9.5 R

RML, E = 10 hydrolysis of acetate Chan ef a/. (1990)

RML, E = 50- 106 vinyl acetate Karninska ef a/. (1996)

Hx. E > 50

R = ce~,,, E = 7.7 R CloH21, E = 15 R = C=C(CH2hCH3. E >50

= CHzCHMez, E =24

OH O, , ) ,Oe M

P h' RML. E = 19 (45"C), 170 (7°C) vinyl butyrate Noritomi ef a/. (1996)

R = C-Hx.E > 50 R = Ph, E =42

RML, E = 12 hydrolysis of butyrate Partali ef a/. (1993)

e0. Ph

RML, E = 5 hydrolysis of butyrate Waagen et a/. (1993)

RML, esterificationwith hexanoic or octanoic acid

cyclic secondary alcohols

Sonnet (1987)

OH

E >50 E = 4.5 hydrolysis of acetate Cotterill el a/. (1988a,b)

OH

Ph RML.E>50 0 ph RML, E = 20 >50 hydrolysis of acetate hydrolysis Of bub'rate RML' '50 or butyrate Klempier ef vinyl acetate Cotterill ef a/. (1988b) Cotterill e l a/. (1991) Klempier eta/.(1990)

-

z o s .,...,062 RML. E = 11 hydrolysis of propionate Cousins et a/. (1995)

E>50

E=15 E=ll RML, vinyl acetate Carrea ef a/. (1992)

k; + k:

meso secondary diols

also RMLremoved after long by

-

E-40

reaction times

bAC

OAc E -20 RML, E 50 RML, vinyl acetate hydrolysis of acetate ~ i ~efa/,~ (1995a.b) l ~ ~ E i >loo, RML Laurnen 8 Schneider (1986) alcoholysis of tetraacetate w/ n-BuOH Sanfilippo ef a/. (1997

OAc

-95% ee, 30-56% yield RML. CRL, PPL alcoholysis of tetraacetate w/ n-BuOH Patti et a/. (1 996)

Fig. 33. Examples of Rhizomucor miehei lipase-catalyzed enantioselective reactions of the secondary alcohols.

86


racemic alcohols p

F

H

Ph ROL, E -20 vinyl acetate Nicolosi eta/. (1994b)

R = Ph, E > 50 E > 50 R = (€)-MeCH=CH. E = 25 ROL, hydrolysis of acetate Li 8 Harnrnerschrnidt (1993)

E>100, lipase QL hydrolysis of laurate Seki et a/. (1996)

BH CE, E = 27 hydrolysis of acetate Cotterill eta/. (1991)

CAL-A, E = 40 LP237.87, E = 48 (ent) hydrolysis of acetate Mitrochkine et al. (1995b)

6 A . U

CE, E >lo0 hydrolysis of acetate Kazlauskas eta/. (1991)

GCL, E>100 hydrolysis of acetate Hoenke ef a/. (1993)

meso diols

6

BnO., AcO

@< @I< OH

0

GCL, >97% ee hydrolysis of diacetate Hoenke eta/. (1993)

OH

O0

>95% ee, 87 - 94% yield Fusarium cutinase hydrolysis of dibutyrate Durnortier et a/. (1992)

a QH

C02Me ProqL, ROL, E >50 hydrolysis of acetate tentative abs. config. Allen 8 Williams (1996)

...N3

N'j

ANL, >97% ee. 100% hydrolysis of diacetate Hoenke eta/. (1993)

E 4 0 0 , CVL vinyl acetate FernBndez et a/. (1995)

OBn HO PCL, >97% ee, 89% yield vinyl acetate GCL, >97% ee, 60% yield hydrolysis of acetate Hoenke et a/. (1993)

r

Fig. 34. Selected examples of enantioselective reactions catalyzed by other lipases.

87

3 Enantioselective Reactions kinetic resolution of racemic derivatives yielding D-myo-inositol derivative

U

U

OH

U

OH

CRL, E >loo, acetic anhydride PPL or CE,E = 60 - 80 hydrolysis of butyrate Ling et a/. ( I 992) Liu 8 Chen (1989) Gou et a/. (1992)

W

PCL, E >loo, acetic anhydride Ling et a/. (1 992) PPL, E >50,vinyl acetate 10% yield Rudolf 8 Schultz (1996)

CRL, E >loo, acetic anhydride Ling et a/. (1992)

kinetic resolution of racemic derivative yielding L-myo-inositolderivative

asymmetric synthesis starting from meso derivatives

U ,.,$OR

U

?H

OH PCL, E = 43, Pseud. sp. lipase, E >I00 acetic anhydride Ling 8 Ozaki (1993)

& O :H:

OR

R = Bz, Pseudomonas sp. lipase >95% ee, vinyl butyrate Andersch & Schneider (1993) R = Bn, PCL. >99% ee, 89% yield,vinyl acetate Laumen & Ghisalba (1994)

Fig. 35. Lipase-catalyzed enantioselectivereactions of myo-inositol derivatives.

3.3.2 Primary Alcohols of the Type RR'CHCHzOH Lipases usually show lower enantioselectivity toward primary alcohols than toward secondary alcohols. Only PPL and PCL show high enantioselectivity toward a wide range of primary alcohols.

3.3.2.1 Pseudomonas Lipases An empirical rule can predict some of the enantiopreference of PCL toward primary alcohols (WEISSFLOCH and KAZLAUSKAS, 1995). Like the secondary alcohol rule above, the primary alcohol rule is based on the size of the substituents. but, surprisingly the sense of

enantiopreference toward it is opposite. That is, the -OH of secondary alcohols and the -CH,OH of primary alcohols point in opposite directions. One way to rationalize this opposite enantiopreference is to assume the extra CH, in primary alcohols introduces a kink between the stereocenter and the oxygen as suggested in Fig. 37. In this manner the large and medium substituents bind in the same enzyme pockets in both cases. Another possibility is a different binding mode for primary alcohols. Indeed, modeling suggests that the large substituent of primary alcohols does not bind in the same pocket as secondary alcohols (TUOMIand KAZLAUSKAS, unpublished data). Not all primary alcohols fit this rule. In particular, primary alcohols that have an oxygen at the stereocenter (e.g., glycerol derivatives)

88


PCL R = Ac, 87% ee, 40% yield hydrolysis of acetate Matsuo & Ohno (1985) (S)-propranolol

R = H, 96% ee, 44% yield vinyl acetate Wang eta/. (1989)

PCL, >95% ee, 47% yield vinyl acetate Hsu eta/. (1990) Bevinakatti & Banerji (1991) Ader & Schneider (1992)

0

Q O H

O A N K I0 M e Ar

OH

i-Pr

0

o A o M

R

typical p-blocker

PCL, 98% ee, 49% yield vinyl acetate Wijnsche eta/. (1996) 0

R = H, PCL, E = 31 to >50 succinic anhydride Terao etal. (1989) acetic anhydride Bianchi etal. (1988b)

R = hexanoyl

hydrolysis Hamaguchi et a/. (1985)

PPL, 83% ee, 40% yield e hydrolysis of methoxycarbonate Shieh eta/. (1991) R

O

00 d

R = butanoyl, PPL, E >23 hydrolysis Ladner & Whitesides (1984) OB ,n HO&O-PCYl

PCL, 90% ee, 92% yield vinyl acetate Terao eta/. (1988) vinyl stearate Win et a/. (1992) Baba eta/. (1990b) also Breitgoff et a/. (1986)

Fig. 36. Lipase-catalyzed routes to enantiomerically pure precursors of propranolol.

secondary alcohols primary alcohols (no 0 at stereocenter)

Fig. 37. Empirical rules that summarize the enantiopreference of PCL toward chiral alcohols. a Shape of the favored enantiomer of secondary alcohols. b Shape of the favored enantiomer of primary alcohols. This rule is reliable only when the stereocenter lacks an oxygen atom. Note that PCL shows an opposite enantiopreference toward these two classes of alcohols.

do not fit this rule. Of the remaining primary alcohol examples, the rule showed an 89% reliability (correct for 54 of 61 examples). For secondary alcohols, increasing the difference in the size of the substituents often increases the enantioselectivity of PCL and other lipases. Indeed, researchers often introduce a large protective group to increase the enantioselectivity, see Sect. 3.3.1.1. However, for primary alcohols this strategy is not reliable. Upon adding large substituents, the enantioselectivity sometimes increased (LAMPE et al., 1996), sometimes decreased, and sometimes


89

racemic primary alcohols

AcOL I PCL, E >loo, R = i-Pr, 1-Bu. Ph E = 4 - 7, R = Et, n-Bu, rrdecyl, benzyl PCL. hydrolysis of chloroacetyl ester acetylation w/vinyl acetate E = 11, Guevel8 Paquette (1993) Egri et a/. (1996) HO.

U 02yy& OH PCL, E = 27 hydrolysis of palmitate ChCnevert et a/. (1994b)

R = Me, E = 3-5. PCL, PFL R = CH1CH=CI+. R = CH2Ph E = 4-6, PCL vinyl acetate Gais 8 von der Weiden (1996)

CI PCL, selectivity = 3 at C20. Other stereocenters pure. vinyl acetate Ferraboschi eta/. (1996)

0"""

PCL, E -33 vinyl acetate Rosenquist et a/. (1996)

&cOOtB" NHBoc PCL, vinyl acetate, E = 5 Burgess Ho (lgg2)

Ph PCL, E >50 (-40°C) vinyl acetate Sakai eta/. (1997)

PFL (lipase AK), E >lo0 vinyl acetate Sugahara eta/. (1991)

TSHN pcL,

= 31 Edwards et a/. (1996)

E = 20, PFL (Amano AK) PCL, vinyl acetate PCL, E = 40 acylation w/2,2,2-trifluoroethyl butyrate E = 15 hydrolysis of butyrate or rnethanolysis of butyrate ester pallaviciniet a/. (1994) W i n & Walther (1992) VBnttinen & Kanerva (1994,1997) E = 7 - 9. PCL hydrolysis of benzoate Bosetti et a/. (1994). Bianchi eta/. (1997)

E = 90 t0>200. PCL hydrolysis of acetate or acylation w/ vinyl acetate Taniimoto & Oritani (1996)

Fig. 38. Selected examples of PCL-catalyzed enantioselective reactions of primary alcohols (continued).

remained unchanged (WEISSFLOCH and KAZLAUSKAS, 1995). Selected examples of primary alcohols that are resolved or desymmetrized by PCL are shown in Fig. 38. More examples are found in a recent survey (WEISSFLOCH and KAZLAUSKAS, 1995). One group of popular substrates are the meso-1,3-diols which can be asymmetrized either

by hydrolysis of the diester or acylation of the diol. For hydrolysis reactions, LIUet al. (1990) noted that acyl migration in the monoester was fast enough in aqueous solution at pH 7 and above to lower the enantiomeric purity of the product. Acyl migration slows considerably in organic solvents (benzene, chloroform, tetrahydrofuran). For this reason, asymmetrization of diol by acylation can give higher enantio-

90

-


meso and prochiral primary alcohols

U

u

(OH

HO\-

U

HO\-

---

H0 ,

(OH

--.

M

H0+JO 99%ee, 33% yield Xie et a/. (1993)

4

PCL, >98% ee, 92-100% yield, vinyl acetate Tsuji eta/. (1989), ltoh eta/. (1993~)

R = Et, 88% ee, 65% yield Gaucher et a/. (1994)

3

0

"'''\OH

do

PCL. 97% ee, 88% yield PCL, vinyl acetate or hydrolysis of dibutyrate PCL, 99% ee, 78% yield hydrolysis of dibutyrate Pottie et a/. (1991) vinyl acetate >99% ee. 81-94% yield Gais et a/. (1992) Tanaka eta/. (1992) Mohar eta/. (1994) also high E with ROL, CVL. RJL

PCL, >98% ee, 88% yield hydrolysis of diacetate. Lampe et a/. (1996)

Fig. 38. Continued.

HO\-

PCL, 99% ee, 75% yield hydrolysis of diacetate Patel etal. (1992b)

AcO,

,,/

c. 0

PCL, 96%ee, 79% yield hydrolysis of diacetate Xie et a/. (1993)

PCL, 85% ee, 76% yield isopropenyl acetate Kim etal. (1995b)

91


Fig. 39. The presence and configuration Of a bond change and even reverse the enantioselectivit y.

97% ee, 75% yield 95% ee, 63% yield saturated analog: 72% ee, 47% yield saturated analog: 70% ee, 56%yield cis isomer: 21% ee (ent),25% yield cis isomer: 55% ee (ent),44% yield PPL, hydrolysis of diacetate, Guanti et a/. (199Oc)

meric purity. (Also see Sect. 4.2.1.2.1 for discussion of acyl migration in the similar glycerides.)

3.3.2.2 Porcine Pancreatic Lipase No generally applicable rule to predict the fast-reacting enantiomer in PPL-catalyzed reactions of primary alcohols exists. Researchers have proposed several rules, but none are satisfactory (EHRLERand SEEBACH, 1990;HULTIN and JONES,1992;WIMMER, 1992;GUANTI et al., 1992). Two rules even predict opposite enantiomers. An example of the difficulties is shown in Fig. 39. Enantiopreference of PPL reversed upon changing from a trans to a cis configuration of the double bond in the 2-substituted 1,3-propandiol derivatives. PPL favored the (S)-enantiomer with high enantioselectivity for the trans isomer, the (S)-enantiomer with moderate enantioselectivity for the saturated analog, but the (R)-enantiomer with low to moderate enantioselectivity for the cis isomer. This reversal is difficult to explain using only the relative sizes of the substituents. Note

that for secondary alcohols, the enantioselectivity also varied with the configuration of double bonds in the large substituent, but the enantiopreference remained the same (MORGAN et al., 1992). Selected examples of PPLcatalyzed resolutions of primary alcohols are shown in Fig. 40.

3.3.2.3 Other Lipases Only a handful of primary alcohols have been either resolved or desymmetrized by other lipases. Selected examples are shown in Fig. 41.

3.3.2.4 Enantioselectivity of Lipases Toward Triglycerides Triacylglycerides are presumably the natural substrates of lipases, so many researchers have investigated the enantioselectivity of lipases toward triacylglycerides. Triacylglycerides with identical functional groups as the two primary alcohols (sn-1 and sn-3) are prochiral.

kinetic resolution of racernic primary alcohols

PPL R = Me, Et. E = 17 - 21 R = Pr, Bu, E >50 ethyl acetate FernBndez et a/. (1992)

R = n-C&i13 or i-Pr PPL. E=13 E = 13 to >50,PPL purified ppL, E = 17 - 24 hydrolysis Of butyrate hydrolysis or acylation hydrolysis of butyrate Ladner 8 Whitesides Bianchi eta/. (1988~) Quartey eta/. (1996) (1984) Fukusaki eta/.(1992b)

HQ

PPL, E = 21 to >5C vinyl acetate Ar = Ph, 4-MeO-Pt Herrad6n (1994)

Fig. 40. Selected examples of PPL-catalyzed enantioselective reactions of primary alcohols (continued).

3 Biotransformationr with Lipases

92

asymmetric syntheses starting from meso or prochiral primary alcohols

U

HO>

HO,

R=

-1-

AcO-

I

98% ee 76% yield

U

HO, i

R R = mC&l, 96% ee,63% yield R = kPr, 97% ee, 75% yield PPL, hydrolysis of diacetate Guanti e t a . (1989, IQSOa, 1992)

OBn PPL, vinyl acetate 95% ee,87% yield (1996)

HO, A c O G Ar PPL, hydrolysis of diacetate X = H, 97% ee,81% yield 88 to >96% ee,6580% yield X = COMOM, 97% ee,82% yield Ar = Ph, 4-MeC& CMeoC&4 i ,0 X = 30ally1, 89% ee.72% yield CCIC&, 4N@Cg14,4-BnOCsH4, 2-naphthyl,2-thienyl, Mhienyl >98% ee Guanti eta/. (1990b) 52% yield C ' bz purified PPL, 92% ee. 98% yield Ramos-Tomb et a/. (1986) PPL, vinyl acetate Banfi et a/. (1995) Guanti et a/. (1994a,b)

PPL, ~ 9 9 % ee,66% yielc vinyl acetate Marshalko eta/. (1995)

"PN

/HO\-

HO0.-

O ,H P t P o L o R 88% ee.45% yield, PPL Breitgoff et a/. (1986) Kerscher 8 Kreiser (1987)

F PPL, >96%de, 77% yield methyl acetate configuration of *' set by synthesis Lovey et a/. (1994)

O ,H =

:

/

i

:

A

9"Ph Ph PPL, 96% ee, 93% yield acetylation of diol w/ vinyl acetate Oddon 8 Uguen (1997)

3 HO, f

b-

PpL, E -30 86% ee,57% yield 94% ee,78% yield 90% ee,60% yield PPL, 95% ee, 66% yield vinyl acetate hydrolysis of diacetate PPL, hydrolysis of diacetate, Hemmerle 8 Gais (1987) ~~~~~i 8 ~i~~ (1995) (1996)

Fig. 40. Continued.

The stereochemical numbering for triacylglycerides and other glycerol derivatives starts with a Fischer projection of the triacylglyceride with the central hydroxyl group positioned to the left. Numbering the carbons sn-1, sn-2, and sn-3 from top to bottom uniquely identifies each position. Enantioselectivity towards triacylglycerides is based on the ability of a lipase to discriminate between the sn-1 and sn-3 position. Enantiomers thus formed are 1,2- or

2,3-diacylglycerides or 1- or 3- monoacylglycerides (Fig. 42). ROGUKA et al. (1993) surveyed the enantioselectivity of 25 lipases in triacylglyceride monolayers. Some lipases (e.g., from Pseudomonus sp., RML, CRL) showed high selectivity toward the sn-1 position, but the stereoselectivity of other lipases varied with the triacylglyceride: CAL-B showed sn-3 selectivity with trioctanoin, but sn-1 selectivity with tri-

3 Enantioselective Reactions C. antarctica B lipase HO.

.i

93

U

HQ

\/

Re,S50 CRL. E = 1 1 hydrolysis of palmitate Chenevert eta/. (1994b)

AcO ROL (R. delernar) hydrolysis of diacetate R = 0-t-Bu, >99% ee, 95% yield R = OAc, 95% ee, 86% yield R = Ph, 95% ee, 64% yield R = OMOM, 95% ee, 95% yield Tanaka et a/. (1996)

>SiMe3 E >loo,ROL, abs, config not established esterification w/ 5-phenylpentanoic acid Uejima eta/. (1993)

E high, hydrolysis of diacetate R1 = Ph, Me; R2 = H, OMOM Chenevert & Dickrnan (1992,1996)

Fig. 41. Selected examples of enantioselective reactions of primary alcohols catalyzed by other lipases.

94


vinyl, or nitrile. The ability of CRL to catalyze reactions involving tertiary alcohols is consisCH 2 0 A ~ tent with a large alcohol binding site as sugA~O+H gested above in Sect. 3.3.1. Surprisingly, RML CH ~ O A C also catalyzed hydrolysis of an acetate of a terLcsn-3 position tiary alcohol. Fig. 42. Stereochemical numbering for triglycerides BRACKENRIDGE et al. (1993) used an oxalate ester to introduce a less hindered ester group. PPL-catalyzed hydrolysis of the mixed oxalate olein. Selectivity also varied with interfacial ester showed moderate enantioselectivity, altension. STADLERet al. (1995) found large though the actual site of reaction was not dechanges as well as reversals in selectivity using termined. In primary alcohols with quaternary stereoanalogs of triacylglycerides with ether or alkyl groups at the sn-2 position. For ROL, HOLZ- center, the hindered stereocenter lies further from the reactive hydroxyl, so lipase-catalyzed WARTH et al. (1997) rationalized these changes reactions remain fast. Selected examples are in using computer modeling. Fig. 44. HOFand KELLOGG (1996a) proposed an active site model for PFL which proposed a flat pocket for one substituent, but the model is lim3.3.3 Other Alcohols, Amines, ited to diols of the type RR’C(OH)CH20H. and Alcohol Analogs Other examples of quaternary stereocenters are in Sect. 3.4.4 on acids and Sect. 3.3.3.2 on 3.3.3.1 Tertiary Alcohols and alcohols with remote stereocenters. sn-1 position

Other Quaternary Stereocenters

Lipase-catalyzed reactions involving tertiary alcohols are slow, presumably due to steric hindrance. O’HAGANand ZAIDI(1992, 1994a) resolved several acetylenic alcohols with CRL (Fig. 43). O’HAGANand ZAIDIsuggested that the acetylenic moiety in these tertiary alcohols occupies the same site as a hydrogen in secondary alcohols because CRL showed low enantioselectivity toward alcohols containing both a hydrogen and a - C r C H substituent at the stereocenter. No CRL-catalyzed hydrolysis occurred when O’HAGANand ZAIDIreplaced the acetylenic moiety with Me,

E = 32

3.3.3.2 Alcohols with Axial Chirality or Remote Stereocenters Pure enantiomers of axially-disymmetric and spiro compounds are often difficult to make using traditional chemical methods, so lipase-catalyzed reactions are often the best route to these compounds (Fig. 45). Other difficult-to-resolve compounds are those with a stereocenter remote from the reaction site. Nevertheless, lipases often showed high enantioselectivity toward these compounds (reviewed by MIZUGUCHI et al., 1994).

4-CF3-P E = 5.7

CRL, E = 23, R = H RML, E = 13-38, R = H , Me, Et, tentative abs. config. CRL, hydrolysis of acetate O’Hagan a Zaidi (1992, ,994a)

Hx PPL, E = 7, hydrolysis hydrolysis of acetate Brackenridge eta/. (1993) transesterification of chloroacetate Barnier eta/. (1993)

Fig. 43. Lipase-catalyzed enantioselective reactions of tertiary alcohols.


U

6 y

-\'

P L Y

95

l9 .H O , 16 >200

Ho

R I

RxPh

CRL, E =13.5 vinyl acetate abs. config. not established Cheong et a/. (1996)

PFL (Iipase AK) vinyl acetate Hof 8 Kellogg (1994, 1996a)

a ,

HO\. B

Z

HO,

hR

o~

O-SiMe3

CVL, 70% ee, 37% yield hydrolysis of diacetate Watanabe et a/. (1992)

R = i-Pr, GCL, E 9 R = Bn, CHAr; PFL (Amano AK). E = 4-5 vinyl acetate, Berkowitz et a/. (1994)

N, I i:OOn-Pr PCL. E = 32 - 68 isopropenyl acetate Sih (1996). Henegar ef a/. (1997)

HO, Fy. R \Ph PCL, acetic anhydride R = Me, E = 21- 38 R = Et, n-Pr, n-Bu, E >50 Goj et a/. ( 1997)

U

U HO,

tp& PPL, E -13 methyl acetate Lovey et a/. (1994)

HO\ 0 : Bn, E >I00 R = *R R = CgHle E = 82 OH,

0'

L C 0 2 E t

CRL, E = 45, vinyl acetate or hydrolysis of acetate PCL, E >loo, vinyl acetate Khlebnikov eta/. (1995)

F

0H9 .:

.

HO. 0 :

3 '

HO,

oi

-OH E=6-21 E=10 PCL, vinyl acetate Ferraboschi et a/. (1991, 1994a-c)

IxK

PCL, R = Bn or CgH19, E >50 vinyl acetate Ferraboschi eta/. (1991) HO, 0 :

PPL. 92% ee, 77% yield hydrolysis of diacetate Seu & Kho (1992) PPL, 96% ee, 85% yield ltoh eta/. (1993b)

h

PCL, E >50 vinyl acetate Ferraboschi eta/. (1993)

Fig. 44. Lipase-catalyzed enantioselective reactions of primary alcohols w i t h quaternary stereocenters.

Selected examples involving alcohols are summarized in Fig. 46. The prochiral dihydropyridine (Fig. 46, first example in the second last row) is a chiral acid, but is included with the chiral alcohols due to the acetyloxymethyl ester. Lipases do not catalyze hydrolysis of simple esters of these dihy-

dropyridines presumably due to a combination of steric hindrance and lower reactivity (this carbonyl is a vinylogous carbamate). Researchers used the acetyloxymethyl group to introduce a more reactive and more accessible carbonyl group.This strategy places the stereocenter in the alcohol portion of the ester.

96

3 Biotronsformations with Lipases HO\

HO\

"0

RJL, Fluka, >99% ee. 60% yield PPL, 85% ee, 50% yield CRL. 75% ee, 37% yield hydrolysis of butyrate Estermann el a/. (1990)

HO\

R = Ph, 4-BrPh: CAL, E >200

p c $ ~ ~ ~ ; i ~ ,"~ ~~ ~ ~$ ~:p "~p "~o L~o ~~ ~; - ' ~ p ~"t) R = Me: CRL, E = 7 (ent) et al, (1993)

CAL, E ,50 ~

r

vinyl acetate, Hof & Kellogg (1996b)

U

HO,

CRL, 100% ee. 32% yield vinyl acetate lhara eta/. (1990)

CAL-8, E = 66 isopropenyl acetate Johnson et a/,,1995

W

PCL, E > l o 0 vinyl acetate Roccoet a/. (1991)

Fig. 44. Continued.

NHBz

-

CRL, E 5 esterification with lauric acid Gil eta/. (1987)

H'*kC

H

TO,

t-B4-'oH PFL, E = 4 (+)-enantiomerfavored succinic anhydride Fiaud et a/. (1992)

OH

tLO PFL (lipaseAK), vinyl butyrate E = 2040, tentative abs. config Jones ef a/. (1995)

HO

CE, E >I0 C E Ewerall '100 hydrolysisof diacetate hydrolysis of diacetate hydrolysis of dihexanoate Kazlauskas (1989) Kazlauskas(1989.1991) Kazlauskas (1989) lnagaki et al. (1989) Wu eta/. (1985)

GE,Ewerall -7

PCL, 98% ee, 45% yield GAL-B, 92% ee,opposite enantiomer

vinyl acetate Tanaka et a/. (1995)

Fig. 45. Selected examples of lipase-catalyzed enantioselective reactions of axially-disymrnetric and spiro alcohols.

97


-

R = Me or H PFL (Amano AK), E = 60 90 vinyl butyrate PCL or PfragiL, E >50 Csomos et a/. (1996) vinyl acetate Nagai eta/. (1993)

"4

PCL or PFL, E >60 R1=R2=H R1= C@Me, R2 = H R1 = H, R2 = OMe acetylation wl vinyl acetate or hydrolysis of butyrate Hongo et a/. (1997)

PCL or PFL. E = 10-40 acetylation w/ vinyl acetate or hydrolysis of butyrate Hongo et a/. (1997)

Ho\ HO

HO

PCL, E = 41 - 75 R = Me, Et, n-Pr, mBu, Alceligeneslipase, E = 5 - 10 n-C5Hs, CPr H pcL, = Pseud. lipase. E = 14 vinyl acetate vinyl acetate E = 32 - 67, X =CH2, CH2CH2,O Jouglet & Rousseau (1993) hydrolysis of propanoate pcL, = 8o PCL, hydrolysis of butyrate De Amici et a/. (1996) et a'' (lgg4) hydrolysis of propanoate Mizuguchi et al. (1994)

Hou

Me0 PCL, E =-I00 PFL, E = 64 vinyl acetate Me0 Nakanoef a/. (1996)

Me0 E 2100

OMe

CRL, hydrolysis of acetate, Hoshinoet a/. (1994)

L

OH

Ho'*

E >loo. PCL hydrolysis of acetate Taniimoto & Oritani (1996)

0'

PFL (Amano AK), 97% ee, 31% yield PCL, 88% ee, 21% yield Holdgrun & Sih (1991), Ebiikeet a/. (1991) Hirose eta/. (1992, 1995). Salazar & Sih (1995)

Achrornobacter lipase E = 14.5 vinyl acetate Mizuguchi et a/, (1994)

i

OAc

PCL, 298% ee, 98% yield hydrolysis of diacetate Bonini et a/. (1993)

Fig. 46. Lipase-catalyzed enantioselective reactions of alcohols with remote stereocenters.

98


PCL, E = 2.3 acetic anhydride Kuge et a/. (1993)

-I- E = 50

CRL. >98% ee. 62-80% yield Duhamel et a/. (1993) hydrolysis of butryate Renouf et a/. (1997) Zhang & Kazlauskas (unpublished)

Fig. 46. Continued.

3.3.3.3 Alcohols with Non-Carbon Stereocenters The first examples of lipase-catalyzed resolutions involved secondary alcohols where the organometallic was the large substituent (WANGet al., 1988; BOAZ,1989; CHONGand MAR,1991;IZUMIet al., 1992).Acylation in organic solvent was crucial to the success of the resolution of the ferrocenyl derivatives because the corresponding acetate reacts readily with water (Fig. 47). Later examples of lipase-catalyzed resolutions involved organometallics with planar chirality.For the (arene)chromium tricarbonyl complexes (Fig. 48), the Pseudomonas lipases were the most enantioselective. CRL showed opposite, but low E (UEMURA et al., 1994). PCL also resolved several other metal carbonyl complexes (Fig. 49) and ferrocenes (Fig.

50). Surprisingly, the shape of the favored enantiomers in PCL-catalyzed reactions is similar for all the (arene)chromium tricarbonyl complexes, but is opposite for most of the ferrocenes. CRL and CE were the most enantioselective lipases toward phenols containing sulfur or phosphorus stereocenters (Fig. 51). Examples of acids containing phosphorus or sulfur stereocenters are summarized in Sect. 3.4.4.

3.3.3.4 Analogs of Alcohols: Amines, Thiols, and Hydroperoxides 3.3.3.4.1 Amines Lipases also catalyze the enantioselective acylation of amines, although reactions are slower than for alcohols. CAL-B is the most

OH

&inMe3 vinyl acetate PPL, E = 9 - 20, Wang eta/. (1988) PCL (SAM-2), E >loo, Boaz (1989)

CRL. PCL, E >lo0 PPL, RML, E -20 lzumi eta/. (1992)

PPL, E >lo0 trifluoroethylvalerate Chong 8 Mar (1991)

Fig. 47. Secondary alcohols containing an organometallic substituent.


. F R &(cq3

Pseudornonas lipases favor enantiomer shown. CRL favors opposite enantiomer.

.KIH I ..'

&( co)3 PCL, isopropenyl acetate, E >I00 Nakamura eta/. (1990) Uemura et a/. (1994)

PCL, vinyl palmitate, E >I00 RJL, vinyl benzoate, E = 30 CRL, vinyl benzoate, E = 13 (ent) Yamazaki & Hosono (1990) PFL (Amano AK), E = 75 PAL (Toyobo A), E = 35 isopropenyl acetate Uemura etal. (1994)

99

I::

Cr(c013 PFL (Amano AK) E >I00 PCL E = 91 isopropenyl acetate Nakamura etal. (1990) Uemura et a/. (1994)

PAL (Toyobo A) E 33 isopropenyl acetate Nakamura etal. (1990) Uemura et a/. (1994)

dr(co)3 PAL (Toyobo A) E >I00 CRL, E = 7 (ent) isopropenyl acetate Uemura etal. (1994)

RJL, vinyl acetate, E = 6 PCL, vinyl benzoate, E -20 CRL. vinyl benzoate, E -65 (ent) Yamazaki & Hosono (1990) Yamazaki etal. (1991)

Fig. 48. Favored enantiomer in the lipase-catalyzed reaction of ortho-substituted hydroxymethyl benzene chromium (0) tricarbonyl complexes.The Pseudornonas lipases favor the enantiomer with the general structure shown for five examples, while CRL favors the opposite enantiomer in three examples.

h PCL (FERM P-5494)E = 8 vinyl decanoate Yamazaki eta/. (1991)

PCL, E = 8 - 17, vinyl acetate Rigby & Sugathapala (1996)

R = M ~E, = 7.4 R = Ph, E = 14 R = CH~OH,E -20

R = M U , E = >50 R = Ph. E = >lo0

isopropenyl acetate, Uemura etal. (1993)

Fig. 49. Other examples of metal carbonyl complexes resolved by lipases.

100


Fe(q5-C5H5)

Fe(q5-C5H5)

Fe(q5-C5H5)

PCL, vinyl acetate, Nicolosi eta/. (1994a)

pH64" pcL

Fe(q5-C~H5)

vinyl acetate, high E

Nicolosi et a/. (1 992)

Fe(q5-C5H5) PCL, E = 2-3 vinyl butyrate

lzumi & Aratani (1993)

&OH Fe(q5-C+15) PCL, E = 2 CAL-B, RML, CRL, E = 11 - 30 (ent) vinyl acetate Larnbusta et a/. (1996)

Fig. 50. Favored enantiomer in the lipase-catalyzed reaction of 1,2-disubstituted ferrocenes. For the hydroxymethyl-substitutedferrocenes, PCL favors the general structure shown. Note the absolute configuration of the favored enantiomer of the ferrocenes differs from the benzene tricarbonyls in Fig. 48.

.. CE, E = 5 - 2 5 R = CH3, CH'CI, *CqHg, Ph CRL, E = 5 - 7, R = CH3, Ph ANL, E = 9, R = CH3

CE, E = 19 CRL, E = 6

CE, E = 32 CRL, E = 81

hydrolysis of acetate, Serreqi & Kazlauskas (1994, 1995)

Fig. 51. Lipase-catalyzed enantioselective reactions of alcohols containing phosphorus or sulfur stereocenters.

popular lipase, but PCL and lipase from Pseudomonas aeruginosa also showed high enantioselectivity. Most researchers used less reactive acylating agents (e.g., esters or carbonates) to avoid chemical acylation of the more nucleophilic amines. Primary amines of the type NHzCHRR' are isosteric with secondary alcohols. SMIDTet al. (1996) proposed extending the secondary alcohol rule to primary

amines for CAL-B and indeed all of the amines below fit this rule (Fig. 52). BASF AG commercialized the resolution of primary amines by a Pseudomonas lipase-catalyzed acylation (BALKENHOHL et al., 1997). A key discovery was the acylation reagent ethyl methoxyacetate. Activated acyl donors react chemically, while lipase-catalyzed reactions with simple esters or carbonates are usually


slow. Acylation with ethyl methoxyacetate proceeds at least 100 times faster than acylation with ethyl butyrate. The reason for this acceleration is not known, but one possibility is that the oxygen may help deprotonate the amine. Researchers also resolved several amines which do not resemble secondary alcohols.

101

ORSATet al. (1996) resolved a secondary amine with CRL and YANG et al. (1995) resolved a primary amine that is isosteric with a primary alcohol (Fig. 53). Lipases usually do not catalyze hydrolysis of amides. One exception is the CAL-B-catalyzed hydrolysis of N-acetyl 1-arylethylamines, but reaction times were a week or longer

rule to predict favored enantiomer

NH2 ,LCOOEt

E 1-8

R

Ei

CAL-B, E = 74-82 ethyl acetate Sanchez et a/. (1997)

llpase

CAL-B, PAL CAL-B, PAL PAL CAL-B CAL-B, PAL PAL CAL-B, PAL CAL-B CAL-B PAL CAL-B, PAL

y H2 U E high, CAL-B acylation with ethyl octanoate Mattson eta/. (1996)

Gotor etal. (1993), Pozo & Gotor (1993a) Puertas etal. (1993) Reek 8 Dreisbach (1994) Kanerva eta/. (1996), Ohrner eta/. (1996) Jaeger et a/. (1996)

COOEt

COOEt PCL, E >loo, CAL-B, E = 6 trans: PCL, E = 12

CIS:

CAL-B, El = 45, E2 = 68 acylation w/ dimethyl malonate Alfonso et a/. (1996)

cis:

PCL, E = 53, CAL-B, E = 51 trans: PCL, E >I00

COOEt PCL, E = 6, CAL-B, E = 29 trans: PCL, E = 87

cis

2.2.2-trifluoroethyl butyrate, Kanerva eta/. (1996)

Fig. 52. Examples of lipase-catalyzed resolutions of amines.

PAL, E>100 PAL, E >lo0 Jaeger eta/. (personal commun.)

102


CRL, E = 2a PCL, E = a vinyl acetate Acylation w/ diallyl carbonate (-)-product favored Yang eta/. (1995) Orsat et a/. (1996)

CAL-B, hydrolysis E >loo, X = H,2-OMe E = 100, X = 3-Br E = 67-78, x = 4-OMe, 2-F E = 30, X = 3-OMe Chapman et al. (1996)

Fig. 53. Other amines resolved by lipases.

Fig. 54. Resolution of amines as oxalamic esters.

(SMIDT et al., 1996). However hydrolysis of the N-methoxyacetyl-1-arylethylamine was significantly faster with reaction times of only 48 h (WAGEGGet al., in press). CHAPMAN et al. (1996) found an indirect way to resolve amides using oxalamic esters. CAL-B catalyzed hydrolysis of the ester group and showed high enantioselectivity toward the remote stereocenter. The stereocenter now lies in the acid portion of the reacting ester, so the rule above no longer applies. Coincidentally, this reaction

also favors the same amine enantiomer predicted above (Fig. 54).

3.3.3.4.2 Thiols Researchers resolved several thiols, which are the simplest analogs of alcohols (Fig. 55). Resolutions used either hydrolysis or alcoholysis of thiol esters and examples include primary and secondary thiols and even an axially

secondary thiols SH

PCL, E = 14 transesterification of acetate with propanol Bianchi 8 Cesti (1990) PCL, E = 3 hydrolysis of acetate Baba et a/. (199Oc)

PCL, E = 25 hydrolysisof acetate Baba et a/. (199Oc) CAL-B, E = aa transesterification of octanoate with alcohols ohrner et a/. (1996)

primary thiols "S\.

Her-

oE=5

PCL, E = 4 hydrolysis Of Baba et a/. (199Oc)

thiol with axial chirality HS\-

0 E=26

PCL hydrolysis of acetate Patel e f a/.(1992a)

HS,

0 PCL, E = 47 PPL, E >lo0 transesterification of acetate with propanol Bianchi 8 Cesti (1990)

>loo CE, Eovera~~ hvdrolvsis of diacetate -Kiefer eta/. (1994)

Fig. 55. Selected examples of lipase-catalyzed enantioselective reactions of thiols.

3 Enanrioselective Reactions

chiral thiol. Enantioselectivities were similar to those for the corresponding alcohols. BABA et al. (1990~)and OHRNER et al. (1996) noted that acylation of thiols gave no reaction. The acyl enzyme intermediate may not be a strong enough acyl donor to acylate thiols. HO.O

HO-

p

HO.

p

PCL,E = 29 PCL.E = 4 PPL.E = 4 isopropenyl acetate isopropenyl acetate isopropenyl acetate Baba et a/ (1988) Baba et a/ (1988) Hbft et a/ (1995)

103

3.4 Survey of Carboxylic Acids 3.4.1 General Considerations There are fewer examples of lipase-catalyzed enantioselective reactions of carboxylic acids (reviewed by HARALDSSON, 1992). In water, lipases catalyze hydrolyses of various carboxylic acid esters, while in organic solvents, lipases catalyze esterification of acids, transesterification of esters, and aminolysis of esters. Reactions of chiral anhydrides and lactones are discussed below in Sects. 3.5 and 3.6.

OOMe

3.4.2 Carboxylic Acids with a Stereocenter at the a-Position (RR'CHCOOH)

PCI.E = 4 vinyl acetate Baba et a/ ( 1990a)

Fig. 56. Alkyl peroxides resolved by PCL-catalyzed acylations.

3.3.3.4.3 Peroxides Lipases also discriminate between enantiomers of alkyl peroxides, which resemble primary alcohols. Enantioselective acylation of alkyl peroxides yielded unreacted starting material in high enantiomeric excess, but the produced peroxyesters decomposed under the reaction conditions to ketones. The structures in Fig. 56 show the fast-reacting enantiomer.

3.4.2.1 Candida antarctica Lipase B In contrast to its high enantioselectivity toward alcohols, CAL-B usually shows low to moderate enantioselectivity toward carboxylic acids (Fig. 57). Preparation of enantiomerically pure 2-arylpropionic acids, a class of nonsteroidal anti-inflammatory drugs, required two sequential resolutions (TRANIet al., 1995; MORRONE et al., 1995). Starting from 300 g of racemic ibuprofen (Ar =4-i-BuC6H,) TRANI et al. (1995) used two sequential esterifications to make 38 g of (S)-ibuprofen with 97.5% ee.

E = 1 - 20, esterification of acid or transesterification of ester Arroyo & Sinisterra (1994), Morrone et a/. (1995) Trani et a/. (19954, Mertoli et a/. (1996) COOH

Fig. 57. CAL-B-catalyzed enantioselective reaction of carboxylic acids with a stereocenter at the a-position.

/L E = 3-10 arninolysis of ethyl ester Quir6s eta/. (1993)

E = 60 esterification with isobutanol Ozegowski et a/. (1994a)

104


The acyl binding site of CAL-B is a shallow crevice. It is likely that the lower enantioselectivity toward stereocenters in the acyl part of an ester stems from fewer and/or weaker contacts between the acyl part and its binding site. In contrast, the alcohol binding site appears to engulf the alcohol.

3.4.2.2 Candida rugosa Lipase In contrast to CAL-B, CRL shows high enantioselectivity toward many carboxylic acids (Fig. 58) and a rule can predict the e.lantiopreference of CRL-catalyzed reactions of carboxylic acids with a stereocenter at the a-position. Recent reviews also contain a few more

examples of acids resolved by CRL (AHMED et al., 1994;FRANSSEN et al., 1996). One important commercial target are pure (S)-enantiomers of 2-arylpropionic acids, a class of nonsteroidal anti-inflammatory drugs. Although CRL shows high enantioselectivity toward these acids, reaction rates are too slow for commerical use. Chirotech in the UK developed a resolution of naproxen using a Bacillus esterase (QUAXand BROEKHUIZEN, 1994).This process produced 13 tons of (S)-nairoxen in 1996 (STINSON, 1997). Comparing the above empirical rule to the X-ray structure of CRL suggests that the large substituent, L, binds in a tunnel, while the stereocenter lies at the mouth of this tunnel. Indeed, molecular modeling supports this pro-

rule to predict enantiomer favored by CRL Ahmed et a/. (1994) Franssen eta/. (1996) COOH R

EtOOCAOE1

E = 5 (R = H), 40 (R = n-CeH13) CRL. E = 75 esterification with heptanol (E slightly lower for hydrolysis) hydrolysis of diethy1ester Witz &Swrr Holmberg eta/. (1992) . (1995) . .

R = H. Me, Et. CRL, E = 3 to >50 esterification w/ n-alcohols Berglund eta/. (1994)

Me0

CRL

E >loo. hdrolvsis of various esters Gu i t a/. (1986) Battistel et a/. (1991) Hernaiz et a/. (1994)

CRL-CLEC, E = 88 E >loo, esterificationw/ HOCHzSiMe3 Tsai & Wei (1994a,b,c) esterificationw/ BuOH Tsai et a/. (1996) Persichetti et a/. (1996) COOH

CRL, E = 20 hydrolysis of Me ester Kornetani et a/. (unpublished)

CRL-CLEC, E>300 esterificationw/ BuOH Persichetti et a/. (1996)

CRL. E >lo0 esterification Mustranta (1992) . . CRL-CLEC, E = 300 esterification w/ amyl alcohol Persichetti eta/. (1996)

Fig. 58. Selected examples of CRL-catalyzed enantioselective reactions of carboxylic acids with a stereocenter at the a-carbon.


105

$OOH R” LOoH

CRL.E=2-26 transester. of Me ester w/ BuOH Kanerva 8 Sundholm (1993a)

R”’

xoH

-A-

R = i-Pr., CRL. ~. PPL. E >I00 R = tBu, CRL, E = 36 CRL, E = 12 - 83 abs. config. not established hydrolysis of methyl ester transester. of -0CHzCF3 ester w/ BuOH Bhaskar Rao eta/. (1994) Martres et a/. (1994) ~

R = H , E>100 R = Me, E = 13 Ac-NH CRL. E = 13 R = Ph. E not reported hydrolysis of Me ester R = COOMe. E = 11 CRL, E > l o 0 CRL, E >50 hYdrolysis Of Me Or Et ester hydrolysis of Bu ester transester. of Me ester w/ BuOH Schueller ef a/. (1996) Gu 8 Li (1992) Franssen et a/. (1996) Csuk & Dorr (1994)

COOMe

HO COOMe

fl

CRL, 32% yield, 100% ee RJL, 20% yield, 90% ee (opposite enantiomer) hydrolysis of Me ester followed by decarboxylation Node et a/. (1995)

Fig. 58. Continued.

posal (HOLMQUIST et al., 1996; BOTTAet al., 1997). Further modeling rationalized some known exceptions to the empirical rule (HOLMQUIST et al., 1996).When the large substituent is extensively branched, it no longer fits in the tunnel. An alternate binding mode with the substrate outside the tunnel favors the opposite enantiomer. Researchers increased the enantioselectivity of CRL-catalyzed resolution of chiral acids using a number of different methods. Most examples involve either 2-arylpropionic acids, or 2-aryloxypropionic acids, a class of herbicides. For example, G u o and SIHincreased the enantioselectivity of a CRL-catalyzed hydrolysis of the 2-chloroethyl ester of 2-(3-benzoyl)phenylpropanoic acid by adding a chiral amine, dextromethorphan ( G u o and SIH,1989; SIHet al., 1992). Kinetic analysis showed that dextromethorphan increased the enantioselectivity

by inhibiting the hydrolysis of the slow-reacting enantiomer. In another example, COLTON et al. (1995) increased the enantioselectivity of a CRL-catalyzed hydrolysis of the methyl ester of 2-(4-chloro)phenoxypropanoicacid by a purification procedure that involved treating the enzyme with isopropanol (Fig. 59). Other researchers have increased the enantioselectivity of CRL toward 2-aryl- or 2-aryloxypropionic acids by changing the solvent (MIYAZAWA el al., 1992), temperature (YASUFUKU and UEJI, 1995) or pH, by carrying out the reaction in a microemulsion (HEDSTROM et al., 1993), by adding (S)-2-amino-4-methylthio-1-butanol (ITOHet al., 1991) or Triton X100 (a surfactant) (BHASKAR RAOet al., 1994), by linking the oamino group of lysine residues to a solid support (SINISTERRA et al., 1994), by nitration of tyrosyl residues (Gu and SIH, 1992),by purification and cross-linking of crys-

106


crude, E = 2.3-17 isopropanol-treated, E >lo0 hydrolysis of methyl ester Colton et a/.(1995)

crude, E = 4 added dextromethorphan, E = 42 hydrolysis of 2-chloroethyl ester Guo 8 Sih (1989)

Fig. 59. Several methods increase the enantioselectivity of CRL toward acids.

tals of CRL (LALONDE et al., 1995;PERSICHET-commerical CRL. Second, the treatments may (Wu et a]., 1990; change the conformation of CRL. CrystalloALLENMARK and OHLSSON, 1992a, b), and by graphers solved the structures for both an careful addition of water to avoid clumping in “open” and a “closed” form of CRL which diforganic solvents (TSAIand DORDICK, 1996). fered in the orientation of the lipase lid (a surChemists do not know how these changes face helical region). Indeed, LALONDEet a]. increase enantioselectivity on a molecular lev- (1995) found that crystals of the open and el, but two possibili,ties are most reasonable. closed forms differed in their enantioselectivFirst, the treatments may remove or inactivate ity. These two possibilities do not exclude each a contaminating hydrolase with low or oppo- other, so both effects may contribute to the insite enantioselectivity. Indeed, LALONDE et al. creased enantioselectivity. (1995) reported a contaminating protease in TI et al., 1996), by purification

COOH

BzS,),

PCL. E =16 esterification with MeOH Patel eta/. (1991)

COOH A P h PCL. __. -E = 62 ._ hydrolysis of thioester Tan et a/. (1995)

PCL. E = 8 - 26 hydrolysis of ethyl ester Lefker e f a/. (1994) COOH

PCL, CRL, 70-80%ee aminolysis of Me ester Yang eta/. (1995)

COOH MeS +OH PCL, E >lo0 hydrolysis of Et ester Urban eta/.(1990)

PCL, E >lo0 hydrolysis of Et ester Houng et a/. (1996b)

COOH

4 X PCL. E = 31-38,X = F. OH PCL, E = 12,X = Br hydrolysis of Et ester Kalaritis ef a/. (1990)

PCL, E >50 hydrolysis of benzyl ester Yamazaki et a/. (1990)

COOH

PCL, E = 36 hydrolysis of Et ester Kalaritis et a/. (1990)

Fig. 60. Selected examples of PCL-catalyzed enantioselective reactions of carboxylic acids with a stereocenter at the a-carbon.


3.4.2.3 Pseudornonas Lipases PCL also catalyzes enantioselective reactions of acids (Fig. 60). The relative sizes of the substituents cannot account for the enantiopreference, but note that all but one example have a similar orientation of an electron-withdrawing group. O'HAGANand RZEPA(1994) suggested that the high enantioselectivity of PCL toward acids with a fluorine substituent at the a-position may be due to a stereoelectronic effect. In nonenzymic reactions, nucleophilic attack at a carbonyl favors an anti orientation of an electron-withdrawing sub-

b"

MeOOC

CE, E -20 hydrolysis of diester Chenevert et a/. (1994a)

stituent at the a-position. A similar preference in the active site of PCL may also account for the observed enantioselectivity.

3.4.2.4 Other Lipases Fig. 61 summarizes selected examples of enantioselective reactions involving other lipases. The PPL-catalyzed resolution of amino acid esters (HOUNGet al., 1996a, b) used crude enzyme which contains protease contaminants. Researchers observed high enantioselectivity

F I

ArJN)

*"c5H11

0

PPL. E = 31 hydrolysis of Et ester Drioli eta/. (1996)

PPL, E >lo0 hydrolysis of Me or Et ester Ar = Ph, 4-0H-c~H4,indoyl, imidazoyl Houng et al. (1996a,b)

$OOH

COOH HooCdOH

W ANL, E = 26 to >lo0 hydrolysis of octyl ester Ng-Youn-Chen eta/. (1994)

I

.Ao

E = 13, Rhizopuslipase (Saiken) dipropyl ester; not indicated which ester reacts Ushio et a/. (1992)

ANL. E = 19 transesterification of vinyl ester with MeOH Miyazawa eta/. (1992)

COOH

COOH I

COOH

Ph..

107

E >50 RML (Amano MAP-10) transesterification of Me ester with isobutanol Gou eta/. (1993)

RML (Amano MAP-10) hydrolysis of Me ester Fulling & Sih (1987)

RML, E = 2 - 2 0 esterification with MeOH CAL-B, E = 2 13 (ent) esterification with PrOH Mertoli et a/. (1996)

-

COOH

Me0

ROOC"" E = high ROL (Arnano F-AP) Crout et a/. ( i 9 9 3 j ~~~

E -50.

RML. hvdrolvsis of Me ester Botta et a/..(1997)

Fig. 61. Selected examples of lipase-catalyzed enantioselective reactions of carboxylic acids with a stereocenter at the a-carbon.

108


( E > 100) only for amino acids where the alkyl group is -CH,-aryl.These amino acid esters are good substrates for chymotrypsin, thus chyrnotrypsin, a likely contaminant, may contribute to the observed selectivity. A survey of the enantioselectivity of ANL toward carboxylic acids identified a-amino acids as the best resolved class of carboxylic acids (JANES and KAZLAUSKAS, 1997b). Replacement of the positively charged -NH: substituent with -OH or -CH3 lowered the enantioselectivity drastically.

Note that CRL and RML favor opposite enantiomers of 2-arylpropionic acids.

3.4.3 Carboxylic Acids with a Stereocenter at the P-Position Even though the stereocenter is further away, a number of lipases also catalyze enantioselective reactions of carboxylic acids with a stereocenter at the p-position (Fig. 62).

Candida anfarctica lipase B COOH HO

COOH

HOL

A

,COOH

,COOH HOLcI

CAL-6, E = >50 arninolysis of ethyl ester Garcia et a/. (1993)

Cbz-1

0

..

,COOH HOP ’h

CBuOOC

slow

CAL-6, E = 15 CAL-6, E = >I00 hydrolysis of ethyl ester arninolysis of ethyl ester Sanchez et a/. (1997) Garcia et a/. (1993)

Candida rugosa lipase

CAL-6, E = 7-10 esterificationwith isobutanol Ozegowski et a/. (1995a)

CRL. E >50 esterification w/ mBuOH Chattapadhyay & Marndapur (1993)

Pseudomonas lipases HOOC,,pR

f,””H

4 : H

Hh : J

fOH

HO NO2 = PCL, E > 50 4’-CH3’ 5’-CH3 Knezovic et a/. (1993)

PCL, E = 37 Boaz (1992)

PFL (Arnano AK) hydrolysis of2,2,2-trifluoroethyl ester Kato et a/. (1995b)

COOMe Et ph

O E

0

E=42 54 PCL 5.9 Blanco et a/. (1993)

PCL, E -7 PCL, E =5 Yarnarnoto n-butyl amine eta/. (1988) Garcia et a/. (1992)

R = CF3, CH3 E >loo, PFL (Amano AK) = cF3, = ; R = CH3, E >100,cRL hvdrolvsis of diethvl ester Kaio et a/. (1995b)

Fig. 62. Selected examples of lipase-catalyzed enantioselective reactions of carboxylic acids with stereocenter at the P-position.

109


Porcine pancreatic lipase ,COOH +Me

ol-

,COOH M e O b P h

MeOD 4;" \

\

OMe 0 0 E = 6 8 PPL 0 Guibe-Jampel ef a/. (1987) E > 50 Bianchi et a/. (l 988a) Barnier et a/, (1989) E = 10 Wallace et a/. (1990)

R O E

me

4~1

Et >30 C5H11 >50 CH2Ph >45 Ph 3.7 Blanco et a/. (1993)

G C O O M e PPL, 97% ee, 97% yield hydrolysis of diester Nagao et a/. (1989)

Fig. 62. Continued.

3.4.4 Other Carboxylic Acids

3.4.4.3 Remote Stereocenters

3.4.4.1 Quaternary Stereocenters

Lipases occasionally show high enantioselectivity toward carboxylic acids with stereocenters far from the carbonyl (Fig. 65). For example, researchers at Merck Research Laboratories (Rahway, NJ, USA) used Pseudornonas lipases to enantioselectively hydrolyze pro-R ester in a dithioacetal (Fig. 65) yielding the (S)-monoester in enantiomeric purity even

Several examples are shown in Fig. 63.

3.4.4.2 Sulfur Stereocenters Several examples are shown in Fig. 64.

COOH

R

COOH

) , ~e O I

Ph

E = 10 - 25, R = Et, n-Pr, rt-C9H19, CLL. E >loo, R = H. AC CH2CH=CH2 hydrolysis O f Et ester SperO 8, Kapadia (1996) E = 52, R = n-C6Hl, CRL, hydrolysis of Me ester tentative abs. config. COOH COOH Sugai eta/. (1990a,b)

F,A

Ph

PPL, R = f-Bu, CPr, E = 6 - 8 hydrolysis of diester configuration at C not specified Bucciarelli et a/. (1988)

p-TZACN

E=23 CRL hydrolysis of Me esters Kometani et a/. (unpublished) E=89

Fig. 63. Selected examples of lipase-catalyzed enantioselective reactions of carboxylic acids with quaternary stereocenters.

110

3 Biotransformations with Lipases HOOC

I B

..f'**Aror c-CeH

PPL, PFL (Amano AK, K10) E 280 hydrolysis of methyl ester Burgess et a/. (1992)

HOOC

11

Q,/.

HOOC

0

.

HOOC

bS\n-alkyl

CRL. E 66 to >lo0 hydrolysis of methyl ester Allenmark Andersson (1993)

sulfur stereocenter lies two bonds away from carbonyl

PFL (Amano K10) PPL, 91% ee, 86% yield E = 33 to >I00 hydrolysisof diester Ar = Ph. p-X-GH4, 2-Np Tamai e l a/. (1994) hydrolysis of methyl ester Burgess 8 Henderson (1989) Burgess et a/. (1992)

..

PXL (Amano K10) X = H. E >80 X N02. E >40 X = CI. E > I 6 hydrolysis of methyl ester Burgess et a/. (1992)

CRL Ar = 2-Np, E = 27 Ar = p-Tol. E = 6 ~~~~~~~~~~~~~~~

sulfur stereocenter lies three bonds away from carbonyl

Fig. 64. Selected examples of lipase-catalyzed enantioselective reactions of carboxylic acids with sulfur stereocenters.

u

H

CI >98% ee, 95% yield, hydrolysis of dimethyl ester PCL. Hugheset a/. (1989. 1990, 1993), Smitheta/. (1992) P. aeruginosa lipase, Chartrain eta/. (1993)

HOOC-

H O O -

O

O

\ Br

C Y OAC \

Br

OAc \ Br

OAc CRL, hydrolysis of methyl or butyl ester E = 4, ~ 1 0 0>loo. , Bhalerao et a/. (1991)

Fig. 65. Selected examples of lipase-catalyzed enantioselective reactions of carboxylic acids with remote stereocenters.

though the stereocenter lies four bonds from the carbonyl (HUGHES et al., 1989,1990,1993; SMITH et al., 1992;CHARTRAIN et al., 1993).The enantioselectivity dropped for analogs where the stereocenter lies either three or five bonds from the carbonyl. BHALERAO et al. (1991) found that CRL showed surprisingly high enantioselectivity toward several carboxylic acids where the stereocenter lies eight or nine

bonds (but not seven bonds) from the carbonyl. The X-ray crystal structures of CRL suggest that the carboxylic acid binds in a tunnel which bends approximately at C-9 of a fatty acid. The observed enantioselectivity toward stereocenters in this region may be due to this bend. Researchers also resolved several other synthetic intermediates with remote stereocenters.


111

3.5 Anhydrides

However, CAL-B was both regio- and enantioselective (OZEGOWSKI et al., 1994b, 1995a). PCL catalyzed the regioseiective ring open- For 2-methyl glutaric acid, the (R)-enantiomer ing of 2-substituted succinic and glutaric anhy- reacted at the more hindered carbonyl, while drides, but without enantioselectivity (HIRA- the (S)-enantiomer reacted at the less hinTAKE et al., 1989).Reaction occurred at the less dered carbonyl (Eq. 10a). Similarly, the (2R)hindered carbonyl (Fig. 66). enantiomer of 2,3-dimethylglutarates reacted at the more hindered carbonyl while the (2s)enantiomer reacted at the less-hindered carbonyl (Eqs. 10b and c) (OZEGOWSKI et al., 1996). A similar reaction with the five-membered 2methyl succinic anhydride was less enantioselective (not shown). For resolution of syn-2,3-dimethylbutanR = Me, i-Pr, Ph dioic anhydride, PCL was the most enantioregioselectivity 4: 1 to 100:1 no enantioselectivity selective enzyme (Eq. 11) (OZEGOWSKI et al., ring opening w/ ethanol 199%). Hiratake eta/. (1989) Lipase-catalyzed ring opening of prochiral Fig. 66. Regioselective ring opening of anhydrides and meso anhydrides also proceeded with occurred at the less hindered carbonyl. good enantioselectivity (Fig.67).

a.

f\/

i s ~ ~ HOOC ~ ~ O LCOOi-Bu

0

b0

CAL-B isobutanol

0

~-

*

29% yield, 99% ee

HOOC

CAL-B isobutanol

*

i-BuOOC

COOH

(1Oa)

28% yield, 88% ee

COOCBu

i-BuOOC

92% ee, 29% y

(*)

0

+

COOH

74% ee, 30% y

A/

f1r

_..'

f R'i

HOOC COoi-Bu 95% ee, 30% y

+

i-BuOOC

COOH

50% ee, 47% y

isobutanol 96% ee

95% ee

A

O

Fig. 67. Lipase-catabzed ring opening of prochiral and meso anhydrides.

Orr0

x-

O

O n o

B0

O n

-

PCL, E 20 R = Me, OMe. E = 21 CAL-B, E 20 R = Et, n-Pr, iPr, E = 4 - 9 ring opening W/ n-BuOH ring opening w/ i&OH PCL, ring opening w/ n-BuOH Ozegowski eta/.(1995b) Chenevert eta/. (1994a) Yamamoto et a/. (1988, 1990)

112


3.6 Lactones Lactones are important flavor compounds and synthetic intermediates. Researchers have used lipases for the synthesis of enantiomerically pure lactones either directly using reactions involving the lactone link, or indirectly using other reactions that eventually lead to enantiomericallypure lactones. Another application of lipases is the selective formation of macrolides or diolides (cyclic dimers) from hydroxy acids. Without a lipase catalyst, oligomers are the major product. Enantioselective reactions involving the lactone link are summarized in Fig. 68. PPL catalyzed the enantioselective lactonization of a

tiR

wide range of y-hydroxy esters to the fivemembered y-lactones (Eq. 12). Researchers used hydroxy esters, not hydroxy acids, as the starting materials to avoid spontaneous lactonization. The enantioselectivity was moderate to good, but the reaction times were often several days. Hydrolysis of the y-lactones was less enantioselective than lactonization.

PPL also catalyzed the formation of &lactones, but with lower enantioselectivity. Surprisingly, the fast-reacting enantiomer differed

0

Ph E = 15 to >50 Henkel eta/. (1992, 1993)

JL

R = CHzCH2COOEt; E >40 R = CH2CH2COOBn; E >40 lactonization of prochiral diester Gutman & Bravdo (1989)

HO

R = Cd-C8H17; E c 11 Sugai eta/. (1990~)

OH E = 11 Henkel et a/. (1995) 0

HO

Ph

CAL-A & B. E = 8 Henkel eta/. (1993)

C,

'-Ph CAL-A & B, E = 6 Henkel eta/. (1994)

h

R = Et, C e H i j C8Hi7, Ph, Me-C6H4, MeOC6H4, Br-C&; E = 23 - >50 Gutman (1990)

R

R = CH2OH; E = 5 Taylor et a/. ( I 995)

R=H,PCL,E=13-16 alcoholysis of lactone Uemura et a/. (1995)

hydrolysis of lactone R = Et, Pr, C5H11, C7H15; E = 5-9 Blanco el a/. (1988)

R = Me, Et, PCL. E > I 00 hydrolysis Enzelberger et a/. (1997)

hydrolysis of lactone R Et, CH2N3, CH21, CHzCI; E = 4 12 Ha et a/. (1996)

-

hydrolysis of lactone R=C5Hl1,PCL,E=11(ent) Enzelberger eta/.(1997)

C10H21

HO

R = Me; E > 50 Gutman et a/. (1987)

0

ko

' d .

PCL, E high alcoholysis of lactone Furukawa eta/. (1994)

E = 9 - 23, hydrolysis R = mCeH17, -(CHZ),OBn Matsumoto et a/. (1995)

(n = 1-3)

Fig. 68. Lipase-catalyzed enantioselective reactions involving the lactone ring. Unless otherwise noted, all reactions refer to the lactonization of the ester catalyzed b y PPL. In all cases, the faster-reacting enantiomer is shown. The cyclic carbonate resembles a lactone so i t is included in this list.


such lactones without affecting the lactone ring; more examples are included in the surveys above. The advantages of these indirect methods are higher enantioselectivity and faster reaction times. Lipases also catalyze the efficient macrolactonization of hydroxy acids as well as macrolactonization of diacids with diols (Fig. 72). Such macrolactonizations are difficult to perform chemically and require high dilution to minimize the competing oligomerization. Below 45 "C, oligomers are also the main products of the lipase-catalyzed reactions, but at higher temperatures macrolactonization dominates even without high dilution conditions. Many different lipases catalyze such macrolactonizations. G u o and SIH (1988) reported that the free acids give higher yield of macrolactone than the esters for another group of hydroxy acids. LOBELLand SCHNEIDER (1993) reported that only the vinyl esters lactonize efficiently. Although noone knows why lipases favor the formation of macrolactones over oligomers, one possibility is that the hydrophobic binding pocket of lipases favors folded conformations of the hydroxy acid.These folded conformations place the alcohol and acid closer to one another and thus favor intramolecular cyclization. The ring-opening oligomerization of lactones is discussed in Sect. 4.3. The macrolactonization reaction was enantioselective favoring the (R)-enantiomer for lactones (LOBELLand SCHNEIDER, 1993) and dilactones (GUOet al., 1988) (Fig. 73).

for the y- and Glactones. Although the alcohol portion of y- and Glactones is a secondary alcohol, the secondary alcohol rule cannot be used here because the stereocenter lies in a different position as shown in Fig. 69. Attempts to form four- or seven-membered lactones yielded oligomers and polymers as discussed in Sect. 4.3. In some cases, oligomeric side products also formed during the lactonization of six-membered rings. However, ringopening alcoholysis can resolve seven- (FURUKAWA et al., 1994) or four-membered lactones (Fig. 70) (Xu et al., 1996; ADAMet a]., 1997). A more common route to enantiomerically pure lactones is to prepare a lipase precursor (usually an efficiently resolved secondary alcohol) and convert to the desired lactone. Selected examples are shown in Fig. 71 (see also SUGAIet al., 1990b). Another special case is lactones with additional alcohol or acid functional groups. Researchers resolved several Favored conformation along C-0 places carbonyl oxygen and stereocenter syn to one another. /

Ring requires an anti orientation of the carbonyl oxygen and stereocenter.

Fd -

\

I

R

Fig. 69. The secondary alcohol rule cannot be used for lactones because the stereocenter lies in a different position. Acyclic esters adopt a syn conformation along the carbonyl C-alcohol 0 bond. The crystal structure of transition state analogs bound to lipases suggests that this conformation persists in the active site. On the other hand, the lactone ring forces an anti conformation along the carbonyl C-alcohol 0 bond which places the stereocenter in a different part of the enzyme. In particular, the lactone stereocenter appears to lie entirely within the L-pocket of the alcohol binding crevice. Indeed, many of the lactone examples in this section do not follow the secondary alcohol rule.

113

3.7 Dynamic Kinetic Resolutions Kinetic resolution limits the yield of the pure enantiomer to 50%. However, if the substrate racemizes quickly in the reaction mixture, then the yield can be 100%.This resolu-

PQ Fig. 70. Seven- and four-membered ring lactones resolved by lipase-catalyzed alcoholysis.

PCL. E high ethanolysis Furukawa eta/.(1994)

CAL-8, E >I00 PCL or PFL, E = 10-74 alcoholysis wl benzyl alcohol Adam et a/. (1997)

PCL. E = 8 methanolysis Xu eta/. (1996)

114

a


R

E

s

1. Me30+BF42. OH- or H ,

Y

E >loo, R = n-alkyl PCL (SAM 111 Haase 8 Schneide; (1993)

R0

synthetic intermediate

cognac lactone

CRL (lipase AY) Pai et a/. ( I 994)

E >loo. PCL Ferraboschi et a/. (1994a)

(R)-(-)-mevanolactone from slow-reacting enantiomer

?H

fi

,.**#\

9 0

0

flavor lactones

E >loo, R = CHzPh, 4-MeOC6H4 PCL, Takano eta/. (1993~)

b

Po --, R

R

OSiMeZt-Bu

PCL, E >lo0 hydrolysis of acetate Sugahara eta/. (1991)

OJ 0 - 0 PCL. E >75 vinyl acetate eta’. (lgg3)

tion with in siru racemization is called dynamic kinetic resolution or second order asymmetric transformation (for reviews see WARD,1995; STECHER and FABER,1997).The requirements for a dynamic kinetic resolution are: first, the substrate must racemize faster than the subsequent enzymatic reaction, second, the product must not racemize and third, as in any asymmetric synthesis the enzymic reaction must be highly stereoselective. Equations for asymmetric syntheses (Sect. 3.2) also apply to dynamic kinetic resolutions. Normal alcohols and car-

Fig. 71. Indirect resolutions of lactones with lipases. a Lipase-catalyzed resolution of lactone precursors and their conversion to lactones. b Functionalized lactones can be resolved by reaction at the secondary hydroxyl group without affecting the lactone ring.

boxylate esters racemize only with difficulty, so this method is limited to the special structures where racemization is rapid. Most enzymic dynamic kinetic resolutions involved base-catalyzed racemization of esters. Racemization involves deprotonation at the a-carbon, so esters contained various electron-withdrawing substituents. FULLINGand SIH(1987) reported the first enzyme-catalyzed example using a protease. The first lipase-catalyzed examples involved 2-phenyloxazolin-5-ones (Eq. 13;Tab. 9).These

a

115


n

7 various tipases

0

c: cG Gatfield (1984) Makita et a/. (1987) Kodera eta/. (1993) Robinson eta/ (1994)

Py;-dnsop$~; 65 "C

HH>

OH

musk fragrance 15-pentadecanolide also 16-hexadecanolide

HO

Guo 8 Sih (1988)

a_-3

+ dilactone

56% yield

"$

Meito Sangyo CRL(0F-360: 65 "C

+ dilactone

I

HO

Guo 8 Sih (1988)

15% yield

42% yield

8% yield

uo,

+ trimer

Sugai ef a/. (1995) lower yields and slower reaction with PCL

0

44% yield

32% yield

Fig. 72. Lipase-catalyzed formation of macrolactones and rnacrodiolides (cyclic dimers?. Examples not shown include cyclization of 16-hydroxyhexadecanoic acid to a 34-membered diolide (Guo et al., 1988: ZAIDIet al., 1995). 0

Y - e 0 .Ph

R A N

-

0 XQ-Ph R N

lipase

*

hydrolysis or alcoholysis

R

L& 13

PCL. lactonization of vinyl ester, dilactones also formed Lobell 8 Schneider, 1993

Fig. 73. Enantioselective macrolactonization.

(13)

H

derivatives of a-amino acids readily racemize by enolization. For R=Me, Bn, and several others, BEVINAKATTI et al. (1990,1992) used an RML-catalyzed alcoholysis in organic solvents to form esters of N-benzoyl amino acids. Unfortunately, the enantioselectivity was only 3-5. SIH'Sgroup screened a dozen lipases for hydrolysis of the phenylalanine derivative (R = Bn) and found that PPL favored the nat-

116

3 Biorransformations with Lipases

Tab. 9. Lipase-Catalyzed Ring-Opening of 2-Phenyloxazolin-5-ones

E

Reference

alcoholysis hydrolysis hydrolysis hydrolysis

3-5 (S) > 100 (S) > 100 ( R ) 2-12

BEVINAKATTI et al. (1990,1992) Gu et al. (1992) Gu et al. (1992) Gu et al. (1992),CRICH et al. (1993)

alcoholysis I-Bu

5-39 CRICHet al. (1993) > 100 (S) TURNER et al. (1995)

Lipase

Reaction

RML PPL ANL ANL, PPL PXL" RML

R=

Bn, Me, n-Pr, CH2i-Pr Bn Bn Ph, 4-OMePh, CH2CH2Ph,several CH2Ar,CH2i-Pr,CH2CHzSMe alcoholysis 13 different examples

a One of several Pseicdornonas lipases: PCL, Amano AK, or Amano K-10. Most reactions favored the (S)enantiomer, but in some cases the enantiopreference was either ( R ) or (S) depending on the amount of added water.

ural (R)-enantiomer ( E > loo), while A N L favored the unnatural (S)-enantiomer ( E > 100) (Gu et al., 1992; CRICHet al., 1993) (Tab. 9). However, these enzymes were less enantioselective toward other, similar derivatives. Several Pseudomonas lipases (PCL, Amano AK, Amano K-10) at 50°C in t-BuOMe catalyzed methanolysis of a variety of 4-substituted 2phenyloxazolin-5-ones with enantioselectivities of 5-39, usually favoring the (S)-enantiomer. In several cases, the enantioselectivity reversed depending on whether the reaction mixture contained added water o r not.The lipase usually hydrolyzed substrates with larger R groups (e.g., Ph, CH2i-Pr) more selectively than small ones (e.g., Me). For preparative use, CRICHet al. (1993) further resolved the enantiomerically enriched methyl esters of N-benzoyl amino acids by protease-catalyzed cleavage of the ester. TURNER et al. (1995) found

that RML-catalyzed alcoholysis of the t-butyl derivative was highly enantioselective (99.5% ee, 94% yield), but only when the reaction mixture contained a catalytic amount of triethylamine. The authors suggested that the triethylamine inhibits a less enantioselective isozyme. TANet al. (1995) resolved 2-(pheny1thio)propanoic acid by PCL-catalyzed hydrolysis of the thioester in the presence of trioctylamine (Eq. 14). Both the thioester and the trioctylamine promote racemization via an enolate mechanism. VORDEet al. (1996) suggested that even simple esters may racemize in the presence of both CAL-B and 1-phenylethylamine (Eq. 15). They did not detect racemization in the presence of only one of these. For chiral alcohols, INAGAKIet al. (1991, 1992) racemized cyanohydrins by the reversible base-catalyzed addition of HCN to aldehydes. Enantioselective acetylation of the ( S ) -

coo@ 96% ee 99% conversion

COOEt

CAL-B 70°C

.

Ph

CO-NT

n-%H13j\ 45% de 99% conversion

117


cyanohydrin catalyzed by PCL yielded the acetate in good to moderate yields and enantiomeric purity. In general, PCL showed higher enantioselectivity toward cyanohydrins derived from aromatic aldehydes than from aliphatic aldehydes. PCL did not catalyze acylation of the HCN donor, acetone cyanohydrin, a tertiary alcohol, presumably because it is too hindered (Fig. 74). Another similar case is the resolution of hemithioacetals where a thiol adds reversibly to an aldehyde (BRANDet al., 1995) (Fig. 75).

10 mol% anion

70-94%ee 60-100% yield

R = Ph, 4-CI-Ph, p-tolyl n-pentyl, CH2CHPhMe

Fig. 74. Dynamic kinetic resolution of cyanohydrins OH

X = 0, lipase R, E >loo, 90% conversion PCL. E = 34, 100% conversion X = NAc, CAL-B, 69"C,E >loo, 100% conversion absolute configurations tentative van der Deen etal. (1996)

OH

MeOOC-&-R

A c O J s / G R

E = 14 - >40 R = CPr, Bu, n-Oct, CH2CH20SiEt3 PFL, vinyl acetate, tentative abs config., Brand etal. (1995) E = 20 - >40 R = Et, OSiEt3

Fig. 75. Dynamic kinetic resolution of hernithioacetals.

Butenolides racemize readily at room temperature and pyrrolinones racemize at 69 "C (Eq. 16) (VANDER DEENet al., 1996). Lipases catalyzed a highly selective acetylation of one enantiomer in excellent yield. The absolute configurations, assigned tentatively, do not fit the secondary alcohol rule. THURING et al. (1996a) independently reported a similar dynamic kinetic resolution of butenolides (Fig. 76). Palladium-catalyzed in situ racemization of allylic acetates, such as the 1-acetoxy-3-phenyl-Zcyclohexene (Fig. 76), also allowed dynamic kinetic resolution. Slow racemization limited the rate of the reaction, but both the yield and enantioselectivity were good. A potentially more general reaction is the racemization of simple secondary alcohols by temporary oxidation followed by reduction using hydrogen transfer catalysts (Eq. 17) DINH

%+

acl

OAc

0 ~

R

;

PCL, E = 8 - 13, 100% COnVerSiOn R1 = R2 = H; R1 = R2 = Me; R1 = H, R2 = Me; R7=Me,R2=H Thuring eta/. (1996a)

Fig. 76. Dynamic kinetic resolution of butenolides and an allylic acetate.

et al., 1996; LARSSON et al., 1997). Similarly, REETZand SCHIMOSSEK (1996) catalyzed the racemization of amines with palladium during a resolution. A related strategy, although it is not a dynamic kinetic resolution, is to invert the con-

CAL-8, Ru catalyst t-BuOH, acetophenone 70°C, 87 h

rii

PFL, hydrolysis of acetate E = 50, 81% yield tentative abs. config. Allen 8 Williams (1996)

2fJ >99.5% ee 92% isolated yield

118


figuration of one enantiomer (SCHNEIDER and GEORGENS, 1992). VANITINENand KANERVA (1995) resolved 1-phenyl ethanol by PCL-catalyzed acetylation with vinyl acetate yielding a mixture of the (@-acetate and the (S)-alcohol. Treating the mixture as shown in Eq.18 conR = OBut, OAc verted the alcohol to the acetate while inverting the configuration. The net reaction was Fig. 77. PPL-catalyzed hydrolysis of the single converting a racemic alcohol to the (R)-ace- ester group in protected sugars. tate. mixture after resolution Mitunobu inverS,On

OAc 4 P h

>99% ee

OH

'

AcOH, DEAD, PPh3

4 P h

>99% ee

OAc A P h

97% ee 97% yield

4 Chemo-and Regioselective Reactions 4.1 Protection and Deprotection Reactions in Organic Synthesis 4.1.1 Hydroxyl Groups

vantage over chemical methods in these reactions. The most useful reactions are those which selectively protect or deprotect one hydroxyl in the presence of several others. The selectivity of lipases usually parallels the chemical reactivity of the hydroxyls, but with increased selectivity. Thus, in hydrolysis reactions of peracylated sugars, the ester at the anomeric carbon (a secondary hydroxyl) reacts first, followed by the ester at the primary hydroxyl. The remaining esters at the secondary hydroxyls react next. In acylation reactions, the primary hydroxyl group reacts first, followed by the secondary hydroxyls. The relative reactivity among the secondary hydroxyls in either acylation or hydrolysis of the esters remains difficult to predict because it varies with the lipase, reaction conditions, and structure of the sugar. Not all reactions follow generalizations. For example, lipases sometimes acylate a secondary hydroxyl group in the presence of a primary hydroxyl.

The most difficult part of carbohydrate chemistry is the selective protection and deprotection of the various hydroxyl groups. The difficulty stems from their similar chemical reactivity, so researchers have searched for en- 4.1.1.1 Primary Hydroxyl Groups zymic methods to simplify this problem. For in Sugars example, FINKand HAY (1969) investigated the selective deprotection of peracylated sug4.1.1.1.1 Hydrolysis of Esters ars almost 30 years ago, but only more recently have researchers found enzymes and reac- of Primary Hydroxyl Groups tion conditions sufficiently selective for synSWEERSand WONG(1986) found that CRL thetic use (for reviews see WALDMANNand SEBASTIAN, 1994; BASHIRet al.. 1995: THIEM, selectively hydrolyzed the ester of the primary alcohol of methyl 2,3,4,6-tetra-O-pentanoyl-~1995; WONG,1995; RIVA,1996). The simplest examples are sugars with a glycopyranosides of glucose, galactose, and single ester group. For example, PPL catalyzed mannose yielding the corresponding tri-0hydrolysis of the esters in the glucopyranose pentanoates. A later paper included methyl and furanose shown in Fig. 77 (KLOOSTERMAN2-acetamido-2-deoxy-3,4,6-tri-0-pentanoyl-~et al., 1987).However, lipases provide little ad- mannoside (HENNENet al., 1988). The solvent

4 Chemo- and Regioselective Reactions OR, F CRL, 33-50% yield

OR FCRL, 75-90% yield R o O R CRL, ~ 29% yield

'8&$,

OR

OMe

R = pentanoyl, octanoyl

R

119

Re*

R, = pentanoyl OMe R2 = 0-pentanoyl, NHAc

o bOROMe , R = pentanoyl

Fig. 78. Selective hydrolysis of esters at the primary position.

~ ~ 0 CRL, . t8596% yield ~

~

0 CRL. - 63% t yield

AcO

4

L - 0 CRL,5:4 o > M -e) AcO OAc Driboside

PPL

a AcO

AcO p-D2deoxyriboside

a-[r2-deoxyriboside

CRL, 60% yield

Kloostermanet a/. (1987) a-Darabinoside

OAc PDxyloside

a-Dxyloside Hennen eta/.(1988)

Fig. 79. Hydrolysis usually favors the primary position.

was water containing 9% acetone and the isolated yields were good for the glucoside, but moderate for the galactoside and the two mannosides. The corresponding acetyl esters did not react under these conditions and the octanoyl esters formed emulsions which made isolation difficult (Fig. 78). HENNEN et al. (1988) extended this work to the furanosides shown in Fig. 79 using 10% dimethylformamide in buffer. In most cases the ester at the primary hydroxyl reacted selectively. In methyl a-D-deoxyriboside, both the primary ester at C-5 and the secondary ester at C3 reacted at similar rates, while in methyl-P-Dxyloside, the secondary ester at C-3 reacted faster than the primary ester at C-5. KLOOSTERMAN et al. (1987) used PPL to selectively hy-

PH O -H HO

R

OH OHC13CpOKCl?3 PPL, pyridine 45"C, 2 d

drolyze the ester from the primary alcohol in the protected D-riboside (Fig. 79).

4.1.1.1.2 Acylation of Primary Alcohols in Unmodified Sugars For the reverse reaction, acylation, the biggest problem is finding an organic solvent that dissolves the polar sugar, but does not inactivate the lipase. THERISOD and KLIBANOV (1986) were the first to find that warm pyridine dissolved sugars, yet did not denature crude PPL. They used PPL to selectively acylate glucose at C-6 with 2,2,2-trichloroethyl laurate giving 40% conversion after two days with 95% regioselectivity (Eq. 19). Similarly, PPL

0

HO HO %OH 6-0-lauryl glucose 40% conversion 95% regioselectivity

120


selectively acylated the primary alcohol in mannose and galactose, but in fructose, which has primary alcohols at C-1 and C-6, both reacted at similar rates. In 2 :1 benzene/pyridine, WANGet al. (1988) found that CRL also retained activity (Tab. 10).They acylated mannose and N-acetylmannosamine with the more active acyl donor, vinyl acetate. Using oxime esters as acyl doners, GOTORand PULIDO(1991) found that PCL was active in pyridine or 3-methyl-3-pentanol and acylated glucose, L-arabinose, galactose, mannose, and sorbose. All acylations favored the primary hydroxyl groups, but the oxime esters were more selective since GOTORand PULIDOdetected no diacylation. With the thermostable CAL-B PULIDOand GOTOR(1993) raised the temperature to 60°C and used the more convenient solvent dioxane and alkoxycarbonyl oximes as acyl donors. This reagent introduced the carbobenzyloxy (Cbz) protective group among others (Eq. 20). More active acyl donors, such as acid anhydrides and even vinyl esters in pyridine, gave nonselective background reactions.None of the conditions above is suitable for the acylation of disaccharides, presumably because they are too insoluble. Another application of sugar esters is as nonionic surfactants for the food and cosmetic industries (Fig. 80). The advantage of an enzymic route over chemical processes, besides milder reaction conditions and fewer side reactions, would be the ability to label the surfactant "natural" (SARNEYand VULFSON, 1995).

In most countries, products produced from natural starting materials using enzymic catalysts are still considered natural. The acylation reactions above all use expensive acylating agents, toxic solvents, and far too much lipase (sometimes four times the weight of sugar). Although they are convenient on a lab scale, they are not practical for surfactant production. For these applications, researchers directly esterified sugars with fatty acids and optimized the reactor configuration to increase yields and reaction rate (Tab. 10). This section reviews the synthesis of acylated carbohydrates, such as 6-0-acyl glucose and 6-0-acyl alkyl glucosides; Sect. 4.2.1.3 below reviews the synthesis of monoacylglycerols (reviewed by BORNSCHEUER. 1995; FIECHTER, 1992). Although SIENOet al. (1984) reported esterification of sugars and fatty acids in aqueous SOlUtiOn,JANSSEN et al. (1990) found only small amounts of ester, which they extracted using a membrane reactor. Others used polar organic solvents such as 2-pyrrolidone and hindered tertiary alcohols and vacuum or drying agents to remove the water released in the esterification. This removal increased the reaction rate and the yield. CAOet al. (1996, 1997) crystallized the product ester to shift the equilibrium. A suspension of glucose, stearic acid, immobilized CAL-B, and molecular sieves in acetone yielded solid 6-0-stearoyl-D-glucose in 92% conversion after 72 h at 60"C.The acetone created a small catalytic phase, while allowing the

0

68%yield 0

glucose

0

6-0-acyl glucose 6-0-acyl alkyl glucoside monoacylglycerol

Fig. 80. Examples of surfactants prepared by lipase-mediated reactions. R = C7-Cl7 chain.

4 Chemo- and Regioselective Reactions

121

product to precipitate (solubility: glucose in acetone =0.04 mg mL-', glucose stearate = 3.3 mg mL-'). No sugar esters formed in reverse micelles (HAYESand GULARI,1992, 1994). Direct esterification usually works better with longer fatty acids (C14-Cl8) than with medium chain fatty acids (C&C12).

4.1.1.1.3 Acylation of Primary Alcohols in Alkyl Glycosides and Other Modified Sugars Since the poor solubility of sugars in organic solvents is a major limitation of lipase-catalyzed acylations of sugars, many researchers modified the sugars to increase their solubility (Tab. 11). HOLLA(1989) used glycals (sugar precursors) which are more soluble because they lack two hydroxyl groups. Acetalization of sugars with acetone increased the solubility so much that researchers eliminated the solvent and dissolved the sugar acetal in the cosubstrate fatty acid (FREGAPANE et al., 1991). IKEDAand KLIBANOV (1993) complexed glucose with phenylboronic acid (Fig. 81). The complex dissolved in t-butanol and PCL efficiently catalyzed the acylation of the primary hydroxyl group with vinyl or trifluoroethyl butyrate. Solubilization of fructose in hexane with phenylboronic acid allowed selective acylation of the C-1 primary hydroxyl, with no reaction at the C-6 primary hydroxyl (SCHLOTTERBECK et al., 1993; SCHECKERMANN et al., 1995). However, the reaction was 100 times slower than the glucose reaction reported by IKEDAand KLIBANOV. Complexation with boronic acids or acetalization also allow acylation of disaccharides (OGUNTIMEIN et al.. 1993; SARNEY et al., 1994). Alkyl glucosides are more soluble in organic solvents, hence, lipase-catalyzed acylations of these sugar derivatives are simpler than unmodified sugars. In addition, the rate of acylation increases as the size of the acyl group increases. The WONGgroup acylated methyl+D-glucoside using CRL and vinyl acetate in a mixture of benzene and pyridine and several z j methylfuranosides (D-ribose, D-arabinose, or ~-xylose)using PPL and 2,2,2-trifluoroethyl acetate in THF-(WANG et al., 1988; HENNEN et

'*


122

Tab. 11. Lipase-Catalyzed Reactions of Modified Sugars" Sugar Derivative

Acyl Donor

Solvent

Lipase

Rateb Reference

IPG-sugar PBA-cY.-D- and p-Dglucose and others' PBA-fructose

fatty acid vinyl ester

none t-butanol

RML PCL

4.540 4.166

FREGAPANE et al. (1991,1994) IKEDA and KLIBANOV (1993)

fatty acid

hexane

RML CAL-B

0.068

SCHLOT~ERBECK et al. (1993). SCHECKERMANN et al. (1995)

Methyl xylose Methyl glucose Ethyl sugar Alkyl sugar

TFEA vinyl ester vinyl ester fatty acid

THF pyridine/benzene THF/Et,N none

PPL CRL PPL CAL-B

0.007 0.006 1.880 nad

Alkyl sugar

fatty acid

hexane

RML

nad

HENNEN et al. (1988) WANGet al. (1988) THEIL and SCHICK(1991) BJORKLING et al. (1989), ADELHORST et al. (1990) FABRE et al. (1993)b ~~

IPG-sugar: isopropylidene glucose, galactose, or xy1ose;TFEA: 2,2,2-trifluoroethyl acetate; PBA: phenylboronic acid comp1ex;THF tetrahydrofuran; D M F dimethylformamide. mmol sugar ester produced per gram enzyme and hour calculated from literature data. 'D-Galactose, D-fructose, sucrose, lactose, maltose, D-mannitol, D-glucosamine, D-glUCOniC acid. na =data not available. a

OH

PCL

9 %

OH

(r

OH C= PCL

RML

P

h

-

s

e

HO ph-B-0 R = P-OH. 45 min R = U-OH,24 h glucose acetal PBA-glucose complex el a/.(1991) lkeda & Klibanov (1993) ~ ~(1gag) l l Fregapane ~

Fig. 81. Increased solubility of modified sugars in organic solvents simplifies acylation reactions.

al., 1988).THEIL and SCHICK(1991) significantly improved the rate of acylation using ethyl glycosides, crude PPL, and vinyl acetate in a mixture of THF and triethylamine. In all cases, acylation was selective for the primary alcohol group. For surfactant applications, the Novo group catalyzed the direct esterification of alkyl glu-

cosides in molten fatty acid using immobilized CAL-B (BJORKLING et al., 1989; ADELHORST et al., 1990).Typical reactions, for example, Eq. 21, showed excellent yield and good regioselectivity. Upon scale-up, Unilever encountered difficulties with the viscous and heterogenous reaction mixture. Adding 25 vol% t-butanol reduced the viscosity and adding 5 mol% prod-

w 0

n

OH

CAL-B, 70°C

vacuum removal of H20 dodecanoic acid

LJ

* H'o

&

r"l+23

Kcl

0

+OH

OEt

6-0-dodecyl ethyl glucoside,94% yield

0E-t" HO

O+O

1 H23

OEt c11H23

2.6-0-didodecylethyl glucoside. 2.4% yield


uct (e.g., 6-U-laurylglucopyranoside) emulsified the reactants. In a packed bed reactor with a separate pervaporation compartment to remove water, they achieved 90% conversion in 25 h for 40 batch reactions (MACRAE, personal communication, 1996). RML was less regioselective (14% of the diester), but adding hexane improved the regioselectivity (FABREet al., 1993a).PELENCet al. (1993) further combined this process with an a-transglucosidase-catalyzedsynthesis of a-butyl glucoside from maltose and butanol (Eq. 22).

maltose

123

BocNH(CH,),COOCH,CC1, (FABREet al., 1994) and di(2,2,2-trichloroethyl)adipate (FABRE et al., 1993b). CAL-B selectively acylates the primary alcohol in a wide variety of nucleosides. GOTOR and MORIS(1992) found that oxime esters of simple acids or protected amino acids selectively acylated the primary hydroxyl, while oxime carbonates gave the 5 '-U-carbonates. Interestingly, PCL selectively acylated the secondary 3 '-hydroxyl even when the primary alcohol was unprotected (Fig. 82).

OC(0)R

a-transglucosidase n-butanol-

RCOOH vacuum

82% yield

RML also catalyzed the 6-0-selective acylation of a-butyl glucoside with more complex acids, for example, the protected amino acid

0-n-butyl

80% yield

Subtilisin, a protease, also catalyzes the selective acylation of carbohydrates, but this is beyond the scope of this review (Riva et al., C A L - B J HO

OH R

fl

PCL R = H, base = uracil, 5-fluorouractl.5-tnfluoromethyluraciI hexanoic anhydride, Uemura eta/ (1989a). Nozakt ef a/ (1990) acylation with oxime esters R = H, base = adenine, thymidine Nacyl cytosine 0 R = OH, base = adenine. uracil, hypoxanthtne R = Me, mC3H7, mC7HI5, n-CgH19, 1-propenyl. Ph. RAo*Ny Gotor & Moris (1992). Moris & Gotor (1993a) R = CH30, BnO, CH&HO, CH2=CHCH20, Morls & Gotor (1992a,b), Garcia-Alles eta/ (1993) R = Cbz-Gly, Cbz-O-Ala, Boc-p-Ala. Morls & Gotor (1994) CAL-B jHO or PCL CAL-B

&thymidinethymidine CAL-B j

CAL-B 3 H O

Fig. 82. CAL-B selectively acylates the primary position, while PCL favors the secondary position.

OH

OH

base = uracil, N-butyryl cytidine

oxime butyrate or butyric anhydride Moris & Gotor (1993b)

124


EBERLING et al. (1996) used lipase from 1988;CARREA et al., 1989;RIVA,1996). Subtilisin is better suited than lipases for the acyla- wheat germ to hydrolyze all the acetates from tion of disacharides, and often shows comple- the sugar portion of 0-glycosyl amino acid mentary selectivity to lipases (KAZLAUSKAS(methoxyethoxy)ethyl (MEE) esters. The MEE esters were removed later using RJL and WEISSFLOCH, 1997). DANIELIet al. (1995) selectively acylated (see Sect. 4.1.3). For example, a glucosyl serine et al. one of the two primary hydroxyl groups in the derivative is shown in Fig. 84; EBERLING triterpene oligoglycoside ginsenoside R,, us- also deprotected a number of similar coming CAL-B and vinyl acetate or di(2,2,2-tri- pounds, galactose, galactosamine, and xylose for the sugar portion and threonine for the chloroethy1)malonate (Fig. 83). amino acid portion. RIVA et al. (1996) selectively acylated the only primary hydroxyl group in the flavonoid glycosides isoquercitin and naringin using CAL-B (Fig. 85).

+I4

I

4.1.1.2 Secondary Hydroxyl Groups 4.1.1.2.1 Hydrolysis of Acylated Secondary Hydroxyl Groups

Fig. 83. CAL-B selectively acylates one of the two primary hydroxyls.

* OAc j AcO AcO -

J ~ c

COOMEE

73% yield, wheat germ lipase Eberling, etal. (1996)

Fig.84. Wheat germ lipase hydrolyzed the acetyl groups at both primary and secondary positions.

Ho%op

HO-0, L & L O z - - - H OH 0 isoquercitin Riva etal. (1996b)

In peracylated sugars, the most chemically reactive ester is the one at the anomeric position.Although it is more hindered than the primary alcohol ester, the electron-withdrawing effect of the additional oxygen makes the anomeric hydroxyl the best leaving group. The ester at the anomeric position is also the most reactive in lipase-catalyzed deacylations. HENNEN et a]. (1988) found that PPL or ANL in buffer containing 9% dimethylformamide catalyzed selective hydrolysis of the acetate at the anomeric position for pyranoses and furanoses in Fig. 86. In most cases, the yields were above 70%.

P

;;"

CAL-B dibenzylrnalonate

O

H

OH \

0 naringin CAL-B. dibenzylrnalonale Riva eta/. (1996b) or subtilisin, vinyl acetate

Fig. 85. CAL-B selectively acylates the primary position.


125

ysis of both the butyrates at the 2- and 4-positions in the glucose derivative, while several other lipases catalyzed hydrolysis of only the butyrate at the 4-position (KLOOSTERMAN et al., 1989).However, in the mannose derivative, CRL selectively hydrolyzed the acetate at the 4-position (HOLLAet al., 1992),while in the galactose derivative, CRL favored the butyrate at the 2-position (BALLESTEROS et al., 1989) (Fig. 87). PCL selectively hydrolyzed the acetate at the 3-position or 4-position in the structures of Fig. 88 (HOLLA,1989; LOPEZet al., 1994).The

One exception was peracetyl /3-D-glucopyranose, where CRL catalyzed selective hydrolysis of the esters at positions 4 and 6 leaving the triacetyl derivative. If the anomeric position lacks an ester group, then the most reactive ester is the one at the primary hydroxyl. (See Sect. 4.1.1.1 above for details.) When the sugar lacks esters both at the anomeric and at the primary hydroxyls, it is not easy to predict which secondary hydroxyl ester will react most rapidly, even for the same lipase. In the series of anhydropyranoses in Fig. 87, CRL catalyzed hydrol-

CRL 3 O A c AcO OAc

OAc

OAc

PPL

PPL, 75%

n

l?

PPL, 54%

AcO OAc

R

AcO

6

"')y$l

OAc

L L , 63%

OAc OAc

furanoses

OAc

ANL, 50%

Fig. 86. Selective hydrolysis of the acetate at the anomeric position.

Do

mannose derivative

OBut

1

OBut

'Cz, 47% CVL, RML, PCL. 91%

glucose derivative

/jj7

OAc

OAc

II CRL, 85-90%

galactose derivative

By&7 PPL. 65%

4

n

OBut CRL, 77-90%

Fig. 87. Selectivity among secondary hydroxyl groups is hard to predict.

126


yields were 90% in both cases. Note that this selectivity follows the secondary alcohols rule in Fig. 18. OAc

Fig. 88. Regioselectivity sometimes follows the secondary alcohol rule.

Several researchers reported selective reaction at the secondary alcohol position in the presence of a chemically more reactive primary alcohol position. For example, PCL selectively cleaved the hexanoate ester of secondary alcohol in several 2-deoxyribonucleosides (UEMURA et al., 1989b).Subtilisin selectively cleaved the ester at the primary position. In the reverse reaction, PCL also catalyzed the selective acylation of this secondary hydroxyl (see above) (UEMURA et al., 1989a; GARCIAALLESet al., 1993). Protecting the primary alcohol as a hindered ester also allowed selective hydrolysis of the ester at the secondary position in the protected arabinose in Fig. 89 (KLOOSTERMAN et al., 1987).

2-deoxyribonucleotides

To form the monoacetate of 1,4 :3,6-dianhydro-D-glucitol,SEEMAYER et al. (1992) started with the diacetate and selectively hydrolyzed the acetate at the (@-stereocenter. This selectivity fits the secondary alcohol rule discussed in Sect. 3.3.1, but in this case the starting material was derived from a sugar and thus was already enantiomerically pure.

4.1.1.2.2 Acylation of Secondary Hydroxyl Groups THERISOD and KLIBANOV (1987) found that lipases catalyzed the regioselective acylation of the C-2 or C-3 hydroxyl group in C-6 protected glucose. The regioselectivity depended on the lipase. For example, CVL catalyzed the butyrylation of the C-3 hydroxyl of 6-0-butanoyl glucopyranose with trichloroethyl butyrate in THF, while PPL catalyzed butyrylation of the C-2 hydroxyl. Chemical or enzymic methods removed the protecting groups at the 6-position leaving a C-2 or C-3 hydroxyl protected glucose. No lipase acylated at the C-4 hydroxyl. For 6-0-butyryl mannose and galactose the selectivity was low (5: 1 at best, typically 2 :1). Another example of lipase-depen-

protected arabinose CRL or PPL

n

PCL. 58 - 80% yield Uemura et a/. (1989b) OH

/ \

Kloosterman et a/. (1987) U

H&+

$J+ozo&o OH 0 rutin Riva et a/. (1996)

I

O ‘-H ‘0 d - O H HO

0

:

.

OAc PCL (SAM-2) hydrolysis of diacetate Seemayer eta/. (1992)

Fig. 89. Lipases sometimes favor hydrolysis of esters at secondary positions over primary positions. Lipases also show selectivity among secondary positions.


dent regioselectivity is the butyrylation of 1,4anhydro-5-U-hexadecyl-~-arabinitol with trichloroethyl butyrate in benzene (NICOTRA et al., 1989). HLL catalyzed butyrylation of the C-2 hydroxyl, while RJL favored the hydroxyl at C-3 (Fig. 90). For the methyl glycosides, several lipases (PCL, PPL, CRL) selectively acylated the C-2 hydroxyl in 6-U-butyryl methyl a-D-galactopyranoside, but the regioselectivity for the corresponding mannoside was still low, top two structures in Fig. 91. In a series of methyl py-

127

ranosides, CIUFFREDA et al. (1990) found that PCL acylated the C-2 hydroxyl in the D-series of sugars, but the C-4 hydroxyl in the L-series. Acylation was much slower when the reacting hydroxyl group was axial (D-rhamnose and L-fucose derivatives). They suggested that efficient acylation requires an axial-equatorialequatorial arrangement of hydroxyls with acylation occurring at the last equatorial hydroxyl. The regioselectivity of the PCL- and PFLcatalyzed acylation of methyl 4,6-U-benzylidene glycopyranosides depended on the conHLL, 66% yield also PCL. PPL, RML

11

Fig. 90. Selectivity ANL. CVL, R = But among secondary CVL, R = ~ , j PPL, R = But CRL. R = t-BuPhZSi (in CHzCIz) positions when the primary position is Therisod 8 Klibanov (1987) protected.

Q

OH HO

a

II

R. japonicus lipase, 79% yield also CRL Nicotra et a/. (1989)

R = Me, PCL, slow

Jl

R = CHzOBut H CRL

VOMe

PCL, PPL also CRL for R = CHZOBut R = Me, methyl-a-D-fucopyranoside R = CHzOBut, 6-O-butyryl methyI-a-D-galactopyranoside

R = Me. PCL

3

7 OH 0

B

~

L

L

I OMe R = Me, methyl-a-D-rhamnopyranoside R = CHzOBUt, 6-O-butyryl methyl-a-D-mannopyranoside

OH

model of efficiently acylated secondary hydroxyl

HO

R = Me, methyl-a-L-fucopyranoside R = CHzOBut. 6-O-b~tyryl methyl-a-L-galactopyranoside

PCL,PPL R = Me, methyl-a-L-rhamnopyranoside R = CH20But. 6-O-butyryl methyl-a-L-mannopyranoside

Fig. 91. In methyl a - ~and - a-L-glycopyranosides,PCL regioselectively acylated the C-2 hydroxyl group in the D-series (top two structures),but the C-4 hydroxyl group in the L-series (bottom two structures) using trifluoroethyl butyrate in THF (CIUFFREDA et al., 1990). Only the sugars a and b reacted quickly, thus CIUFFREDA et al. suggested that an efficiently acylated sugar contains an axial-equatorial-equatorialarrangement of hydroxyls as shown in the model.

128

3 Biotransformations with Lipases a-anomers

n

p-anomers

glucose ph’:sOMe OMe

PCL or PFL

PCL or PFL

U

I OMe

It PCL

OMe

galactose PCL, R = allyl, no reaction PFL, R = Me, 19% yield, 10 days

PCL. R = ally1,91% Yield, 4 days pFL, = Me, yield* lo days

PFL

OH

OMe

figuration at the anomeric carbons (Fig. 92) (CHINNet al., 1992; IACAZIOand ROBERTS, 1993; PANZAet al., 1993a, b). The a-anomers yielded the C-2 monoester with typical reaction times of 7 h, while the /3-anomers reacted in about 1 h and yielded the C-3 monoester. The ROBERTS group used vinyl acetate as the solvent and acylating agent, while PANZAet al. used a variety of vinyl esters and trifluoroethyl esters in THE The galactose derivatives reacted significantly more slowly, possibly due to steric hindrance. PCL also catalyzed acylation of the C-3 hydroxyl in 6-0-acetyl D-glucal and 6-0-acetyl Dgalactal with vinyl esters (HOLLA, 1989)(Fig. 93). LOPEZet al. (1994) found that the regioselectivity also varies with the nature of the sub-

,,& HO

ll

PCL

Hoo&

II

PCL

Fig. 93. Regioselectivity among secondary hydroxyl groups sometimes follows the secondary alcohol rule.

Fig. 92. The configuration at the anomeric carbon determines the regioselectivity of the acylation of methi1 4,60-benzylidene glycopyranosides. PCL and PFL acylate the C-2 hydroxyl in the a-anomers and the C-3 hydroxyl in the p-anomers, regardless of the orientation of the reacting hydroxyl.

stituent at the anomeric position. PCL catalyzed formation of the 3,4-diacetate of methylP-D-xylopyranoside using vinyl acetate in acetonitrile, whereas the octyl derivative in acetonitrile or hexane gave a mixture of the 2,4and 3,4-diacetates. At short reaction times, the 2-monoacetate predominated. The choice of solvent and reaction conditions is less critical than for sugars because these sugar derivatives are more soluble (Fig. 94). In summary, lipases can react selectively at the different secondary hydroxyls. The selectivity varies with lipase and substrate structure (anomeric orientation, anomeric substituent, orientation of hydroxyl) and one cannot make broad generalizations yet. R=Me OH 3,4rdiacetate R = moctyl, hexane solvent 3 h reaction 2-monoacetate 74 h reaction 3 6 1 mix of 2,4- and 3,4-diacetates PCL, vinyl acetate, L6pez eta/ (1994)

Fig. 94. Regioselectivity varies with the nature of the substituent the anomeric position.

129


4.1.1.3 Hydroxyl Groups in Non-Sugars

tates and related compounds, a generalization in Fig. 96 summarizes some of the observed regioselectivity. In addition, PCL catalyzed the regioselective acetylation of polyphenols with vinyl acetate (Figs. 95 and 97). Both acetylation and deacetylation favor the less hindered positions, thus the two reactions yield complementary products. NICOLOSI et al. (1993) used the deacetylation reaction in the synthesis of a rare O-methyl flavonoid, ombuin. In a number of symmetrical acylated catechols, PPL selectively removed only one acyl group (PARMARet al., 1996, 1997). Lipases CRL and PPL also showed excellent chemoselectivity.They cleaved the phenolic ester, while leaving the benzoate ester intact.

4.1.1.3.1 Phenolic Hydroxyls Several lipases, especially PCL and PPL, catalyze the deacetylation of peracetylated polyphenols by transesterification with n-butanol in organic solvents (Figs. 95-97). Researchers deacetylated by transesterification instead of hydrolysis because the substrates do not dissolve in water. The regioselectivity of lipases toward phenolic hydroxyls usually paralleled their chemical reactivity - less hindered positions reacted more quickly. For flavone ace-

U

u

OBut

OAc 0

U

U OAc

CVL, CRL, ANL transesterification w/ n-butanol, Rubio eta/. (1991)

R = H, 60% R = ()Me, 65% (CRL)

0

0 78%

R = H, 80% R = OAC, 55%

PPL. transestrification w/ rrbutanol, Parmar eta/. (1992)

11

U

II

U

u

CI 3.5: 1

Br

OH H

R = H, Me, Et PCL, vinyl acetate, Nicolosi eta/. (1993)

U RO

U

OR

U

Me0 I OR OR PPL, hydrolysis, R = C(O)CH~CHJ Parrnar eta/. (1996, 1997)

CRL

U

2.4: 1

CRL, PPL

U

I

COOMe

transesterification wl n-butanol Parrnar et a/. (1997)

Fig. 95. Regioselectivity of lipases toward polyhydroxylated benzenes. Lipases favored the less hindered position in both deacetylation of peracetylated phenols by transesterification with n-butanol and in acetylation of phenols with vinyl acetate. Note that the first two examples of PARMAR et al. (1996,1997) show deacylation of the more hindered ester.

130


PCL, Natoli eta/. (1990, 1992)

PPL, Parmar e f a/. (1993a)

Fig.96. Regioselectivity of lipases toward flavone acetates and related compounds. Lipases catalyzed the deacetylation by transesterification with n-butanol. Less hindered acetates react more quickly; a generaliza-

tion for the observed regioselectivity is suggested above.

PCL vinyl acetate acetonitrile 48 h. 45"CW

-OH

(+)-catechin

PCL butanollTHF (+)-catechin 12 h. 45"CW pentaacetati PCL butanol (+)-catechin 36 h. 45°C. pentaacetate

5-monoacetate, 40% 7-monoacetate, 32%

3,3',4'-triacetate, 50%

3-monoacetate, 95%

Fig.97. Regioselective acetylation and deacetylation of catechin. Hydroxyls or acetates at positions 5 and 7 react most quickly, while those at position 3 do not react. Acetylation and deacetylation yield complementary acetates (LAMBUSTA et al., 1993).

4.1.1.3.2 Aliphatic Hydroxyls PPL in acetone selectively acylated the primary hydroxyl group in several diols using trifluoroethyl butyrate (PARMARet al., 1993b).

Deacylation of the corresponding diesters showed apparent selectivity for the secondary hydroxyl, but later work showed that deacylation occurred at the primary position, followed by acyl migration to the secondary position (BISHTet a]., 1996) (Fig. 98).

4 Chemo- and Regioselective Reactions RLOHO

u

H

u

Ho%i-OH

rating the isomers than the original method of flash chromatography (Fig. 100).

-

R = Me, Et. n-Pr. n-BU, f?-C6Hi3,Ph

131

4.1.2 Amino Groups

PPL, high selectivity. trifluoroethyl butyrate Parrnar eta/. (1993b), Bisht eta/. (1996)

Amines react spontaneously with most acylating agents, so few lipase-catalyzed reactions have been reported. GARDOSSI et al. (1991) used dilute solutions and a large amount of lipase to selectively acetylate the camino group in L-Phe-cu-L-Lys-0-t-Bu and L-Ala-Q-L-Lys0-t-Bu with trifluoroethyl acetate. Pozo et al. (1992) used the less reactive vinyl carbonate and CAL-B to form a carbamate, one of the more common amino protective groups (Eq. 23). ADAMCZYK and GROTE (1996) protected amines by PCL-catalyzed acylation using benzyl esters. Lipases are not used to deprotect amines because lipases rarely cleave amides or carbamates, the most common amino protective groups. Proteases such as penicillin G acylase

Fig. 98. Selective acylation of primary alcohols.

In some cases, the configuration of nearby stereocenters changed the selectivity. For example, PCL showed a low selectivity for the less hindered primary hydroxyl in the (R)enantiomer in Fig. 99, but a moderate selectivity for the more hindered primary hydroxyl in the (S)-enantiomer. In another case, CRL acetylated the hydroxyl at the (S)-stereocenter only in the (S,S)-stereoisomer, not in the (S,R)-stereoisomer (Fig. 99). SATTLER and HAUFE(1995) selectively acylated the primary over the secondary alcohol in a mixture of diastereomers. This regioselective reaction is a more convenient way of sepa2: 1

u

HO&

:

OH OH

OH

OBn

P&Ph

OBn

PCL, vinyl acetate Ferraboschi et a/. (1995a)

Fig. 99. Selectivity varies with the configuration of nearby stereocenters.

OH OH

P a P fast no rxn CRL,vinyl acetate monoacetylationonly Levayer eta/. (1995)

no acylation

OH

U

Fig. 100. Selective acylation of one diastereomer simplified separation.

-w

+

'

G

U

OH

acylation O

M

e

H

O

k

regloselectivity>go%, CVL, vinyl acetate Fernandez eta/. (1995)

PCL, acetic anhydride Sattler & Haufe (1995)

-

0 poKoAPh CAL-B

0

Jt-

"'OH

..)lOAPh 58%

h

132


The advantages of lipases over proteases are that they tolerate water-insoluble substrates, do not cleave peptide bonds (a potential side reaction in protease-catalyzed reactions of peptides), and tolerate both L- and D-amino acids. Many groups have used lipases to deprotect carboxyl groups in peptides. BRAWNet al. (1990,1991) cleaved the heptyl ester carboxyl protective group from a wide range of dipeptides using ROL (Amano N). This lipase did not cleave the amino protective groups Cbz, Boc, Aloc, or Fmoc and the heptyl protective group survived conditions used to remove these amino protective groups (hydrogenation, HCl/ether, or Pd(O)/C-nucleophile). Although the crude lipase (Amano N) also hydrolyzed the peptide link, pretreatment with PMSF, a serine protease inhibitor, eliminated this side reaction. Hydrolysis of the heptyl group slowed and sometimes did not proceed when the peptide was hindered and/or hydrophobic. In some v 87% yield of these cases, replacing the heptyl ester with (24) the more reactive 2-bromoethyl ester or the more water-soluble 2-(N-morpholino)ethyI ester or 2-[2-(methoxy)ethoxy]ethyl (MEE) es4.1.3 Carboxyl Groups ter increased reaction rate (WALDMANN et al., Although many chemical methods exist to 1991; BRAUNet al., 1993; KWNZet al., 1994; et al., 1996) (Fig. 101). protect and deprotect carboxyl groups in ami- EBERLING In other cases where hydrolysis catalyzed by no acids for peptide synthesis, many of these are incompatible with sensitive functional ROL (Amano N) was slow, researchers substigroups such as thioesters, phosphate esters, tuted another lipase. During a glycopeptide and polyenes (farnesyl groups). The mild reac- synthesis researchers used RJL to cleave the tion conditions for enzymic reactions makes heptyl protective group (BRAUNet al., 1992, them ideal for reactions involving sensitive 1993). In a similar glycopeptide, RJL did not et al. substrates. Chemists have developed a variety cleave the heptyl ester, so EBERLING of methods, most of which involve proteases, (1996) used the more reactive MEE ester. To esterases, or lipases (WALDMANN and SEBAS- cleave C-terminal proline MEE esters, researchers used HLL (KWNZet al., 1994) (Fig. TIAN, 1994). Only the lipase examples are re102). viewed below. are normally used for deprotection (reviewed by WALDMANN and SEBASTIAN, 1994). However, WALDMANN and NAEGELE (1995) reported an indirect removal of carbamate protective group with an esterase. Upon cleavage of the acetyl group from a p-acetoxybenzyloxycarbonyl-protected peptide with acetylesterase, the carbamate link cleaved spontaneously.Lipases should also catalyze this reaction. PCL catalyzed the hydrazidolysis of a,punsaturated esters such as methyl acrylate (Eq. 24) (GOTORet al., 1990; ASTORGA et al., 1991, 1993). These reactions occur at room temperature with simple esters, while chemical methods require higher temperatures, activated esters, or acid chlorides, and suffer from competing Michael additions.

ROL (Arnano N ) p H 7, 37 'C,9% acetone, 84-97%

Boc-Val-Phe

Fig. 101. Deprotection of carboxyl groups in more hydrophobic peptides requires more reactive or more

soluble esters.


Lipases can also selectively deprotect the two carboxylate groups in glutamic acid. For the uncommon enantiomer glutamic acid diesters (D-), both CRL and CAL-B favored reaction at the less hindered ester. CRL catalyzed the selective hydrolysis of the dicyclopentylester (Wu et al., 1991), while CAL-B catalyzed the selective amidation of the diethyl ester with pentylamine (CHAMORRO et al., 1595). On the other hand, in the more common enantiomer L-glutamate diethylester, CAL-B selectively amidates the more hindered ester (CHAMORRO et al., 1995) (Fig. 103).

To develop immunoassays for drugs, researchers must immunize animals with the drug linked to a protein. In several cases, lipases have simplified this linking process (ADAMCZYK et al., 1994,1995). PCL selectively hydrolyzed one of the ester groups in the diacid linker molecule for both the rapamycin and the digoxigenin derivatives. For the digoxigenin derivatives, ester groups on shorter linkers did not react (Fig. 104). SHARMA et al. (1995) used PPL to selectively monoesterify aliphatic dicarboxylic acids with

?-sugar

RJL (Amano M), 76%

Fig. 102. Deprotection of carboxyl groups using other lipases.

Fig. 103. Regioselective deprotection of glutamic acid esters.

AcO

OAc RJL (Arnano M), 88%

HLL (Arnano CE). 31%

H 2 N 7 O X , 3 DGlu 0 CRL (lipase OF),90%

"

L-GIu CAL-B, 94%

CAL-B. 6:1 PCL

OR

R = Me, Bn Adarnczyk eta/ (1994)

DMe

Adamczyk eta/. (1995)

H raparnycin 42-hemisuccinate esters

Fig. 104. Regioselective deprotection of carboxyl groups.

133

digoxigenin esters CRL

0

0 n = 1. 2. 4 Fukusaki eta/. ( 1 9 9 2 ~ )

134

3 Biotransformations with Lipases faster when R = 4-0Me-C~H4

0

.='

VS.

U

U

COOMe

R

faster when R = alkyl

reacts 2 80 times faster

OAc

R

CRL, R = t-Bu, i-Pr, Et, Me, COOMe Konigsberger et al. (1996)

CRL equal rates when R = Ph Cipiciani eta/. (1996)

Fig. 105. Chemoselective reactions catalyzed by CRL.

butanol. Acid groups containing a carbon-carbon double bond at the qP-position reacted more slowly than acid groups adjacent to saturated chains. Some CRL-catalyzed chemoselective reactions are summarized in Fig. 105.

4.2 Lipid Modifications

YAMANE(1987); BUHLER and WANDREY (1987a, b); NIELSEN (1985).

4.2.1 1,3-Regioselective Reactions of Glycerides

Many lipases, called 1,3-selective lipases, Of the 60 million metric tons of fats and oils catalyze reactions at the primary alcohol poproduced each year worldwide, most are used sitions of glycerol and glycerol derivatives, directly in food, but about 2 million tons while other lipases, called nonselective lipases, undergo chemical processing such as hydroly- react at all three positions (Tab. 12). (See Sect. sis, glycerolysis,and alcoholysis.Current chem- 3.3.2.4 for the glycerol nomenclature.) The ical processes require high temperatures and Rhizomucor and Penicillium lipases are all 1,3pressures which degrade the fats and intro- selective but the Candida lipases include both duce impurities. For example, sodium meth- 1,3-selective and nonselective lipases. A single oxide-catalyzed interesterification of triacyl- microorganism, Candida antarctica produces glycerides also catalyzes Claisen condensa- two lipases: A, which is nonselective, and B, tions and imparts a brown color. Lipases re- which 1,3-selective. For lipase from Pseudoquire milder conditions - lower temperatures, monas fluorexens, some researchers reported near neutral pH - and are also regioselective 1,3-selectivity,while others reported no selecfor the primary vs. secondary positions in tivity. This may be due to the f! cepacia vs. f! glycerol and chemoselective for different fatty fluorescens confusion mentioned in Sect. 1.1.2. acids. Researchers have developed several The 1,3-selectivelipases differ in the degree lipase-catalyzed processes for specialty fats. of selectivity as indicated qualitatively in Tab. These processes exploit either the regioselec- 12. Quantitative measure of selectivity is diffitivity of lipases, the fatty acid selectivity,or the cult in water due to facile acyl migration (Sect. mild reaction conditions. However, lipases are 4.2.1.2.1), but SCHNEIDER'S group measured more expensive, so they will not replace chem- the selectivity for several lipases in nonpolar ical catalysts for processing of low-value fats. organic solvents where acyl migration is slow. For recent reviews on lipase-catalyzed mod- The highly 13-selective ROL (Amano D) reification of lipids see VILLENEUVE and FOGLIA acted at the primary position 76 times faster (1997); BORNSCHEUER (1995); MARANGONIthan at the secondary position, while the rnodand ROUSSEAU (1995); HAAS and JOERGER erately selective CVL and RML reacted 26 (1995); VULFSON (1994); ADLERCREUTZand 11times faster, respectively.The nonselec(1994); PRAZERES and CABRAL (1994); CASEY tive PFL reacted only 1.4 times faster (BERand MACRAE(1992); BJORKLING et al. (1991); GER and SCHNEIDER, 1991a; BERGERet al., MUKHERJEE (1990); BAUMANN et al. (1988); 1992). Note also that lipases with 1,3-selectiv-


135

Tab. 12. Regioselectivityof Some Lipases Toward Hydrolysis or Transesterification of Triacylglycerides ~~

Lipase”

Regioselectivity

ANL CAL-B CAL-A CRL CLL CVL GCL HLL

PPL PcamL ProqL PCL PFL

moderately 1,3-selective 1,3-selective non- or 2-selective nonselective moderately 1,3-selective moderately 1,3-selective nonselective slightly 1,3-selective slightly 1,3-selective 1,3-selective highly 1,3-selective moderately 1,3-selective nonselective non- or 1,3-selective

ROL (Amano D)

highly 1,3-selective

ROL (Amano N) RML

highly 1,3-selective moderately 1,3-selective

RJL

a

Reference MACRAE and HAMMOND (1985);Amano Pharmaceutical

Novo Nordisk

Novo Nordisk; ROGALSKA et al. (1993) Amano Pharmaceutical Amano Pharmaceutical MACRAEand HAMMOND (1985);BERGER et al. (1992) Amano Pharmaceutical MACRAE and HAMMOND (1985);Amano Pharmaceutical MACRAEand HAMMOND (1985);Amano Pharmaceutical MACRAEand HAMMOND (1985) Amano Pharmaceutical Amano Pharmaceutical Amano Pharmaceutical MACRAEand HAMMOND (1985);BERGERand SCHNEIDER (1991a); BERGER et al. (1992);Amano Pharmaceutical (1991a);BERGERet al. (1992); BERGER and SCHNEIDER Amano Pharmaceutical Amano Pharmaceutical BERGER and SCHNEIDER (1991a);BERGERet al. (1992); Novo Nordisk

See Tab. 1 for Lipase abbreviations.

ity toward triacylglycerides still hydrolyze esters of secondary alcohols. For examples see Sect. 3.3.1.6. ROGALSKA et al. (1993) reported that CALA selects for the secondary alcohol position (sn-2) in monolayer films. In other cases, apparent 2-selectivity of a lipase was later attributed t o nonselective hydrolysis combined with acyl migration from the 2-position to the 1,3positions (for an example see BRIANDet al., 1995). ROGALSKA et al. found n o acyl migration under their reaction conditions, but the 2selectivity of this lipase awaits independent verification.

4.2.1.1 Modified Triglycerides Structured triacylglycerides (STs) are triacylglycerides modified in either the type of fatty acid or the position of the fatty acids. The synthesis of STs such as cocoa butter substitutes oc16 0

EOC18 oc16 0

palm oil fraction

and MLM lipids (triacylglycerides with medium chains at sn-1 and sn-3 and a long chain at sn-2) relies on the 1,3-selectivity of lipases. The synthesis of STs enriched in polyunsaturated fatty acids exploits the fatty acid selectivity of lipase and will be discussed in Sect. 4.2.2.2 and 4.2.1.1.3 below.

4.2.1.1.1 Cocoa Butter Substitutes Cocoa butter is predominantly 13-disaturated-2-oleyl-glyceride,where palmitic, stearic, and oleic acids account for more than 95% of the total fatty acids. Cocoa butter is crystalline and melts between 25 and 35 “C imparting the desirable “mouth feel”. Unilever (COLEMAN and MACRAE,1977) and Fuji Oil (MATSUOet al., 1981) filed the first patents for the lipasecatalyzed synthesis of cocoa butter substitute from palm oil and stearic acid (Eq. 25). Both companies currently manufacture cocoa butOCl8 0

oc16 0

+ 3 C8l,

1,3-setectivelipaseF

Eoclal

0% 0

+

Eoclsl

OCl8 0

cocoa butter substitute

+

3c160

(25)

136


ter substitute on the multi-ton scale using a 1,3-selective lipase to replace palmitic acid with stearic acid at the sn-1 and sn-3 positions (for reviews see QUINLAN and MOORE,1993; MACRAE and HAMMOND,1985; MACRAE, 1983). Other suitable starting oils are sunflower, rape seed (ADLERCREUTZ, 1994), or olive oils (CHANGet al., 1990). Recent work on cocoa butter substitutes focused on optimizing this process. MOHAMED et al. (1993) used CAL-B, while others complexed lipases from Pseudomonas species (ISONOet al., 1995) or RJL (BASHEER et al., 1995a,b) with surfactants to either incrLdse reaction rates or selectivity. CHO and RHEE (1993) and CHOet al. (1994) investigated continuous packed bed reactors. Another structured triglyceride is Betapol. a formula additive for premature infants. Saturated fatty acids with chain lengths longer than C18 are poorly absorbed partly because they form insoluble calcium salts.Thus, digestion of a typical vegetable oil such as P O 0 (mixture of 1,2-dioleyl-3-palmityl glyceride and its enantiomer) by pancreatic lipase (a 1,3-selective lipase) yields the poorly absorbed palmitic acid. Human milk, on the other hand, contains OPO (1,3-dioleyl-2-palrnityl glyceride). Digestion yields oleic acid and 2-palmityl monoglyceride, both of which are absorbed more efficiently. Interesterification of tripalmitin with oleic acid using RML at low water activity yields OPO. Evaporation removes excess fatty acids and crystallization removes remaining PPF!

in the sn-1 and sn-3 positions of the glycerol backbone and a long chain in the sn-2 position (reviewed by AKOH,1995). Several chemically synthesized MLMs are commercially available (Tab. 13) but these have a random distribution of fatty acids. MLMs are an efficient food source for persons with pancreatic insufficiency and other forms of malabsorption (BABAYAN, 1987; BABAYAN and ROSENAU, 1991; MEGREMIS, 1991). Pancreatic esterases catalyze hydrolysis of meIium chain triacylglycerides faster than long hain and the resulting 2-monoacylglycerides are absorbed efficiently (JANDACEKet al., 1987). Further, the 2-monoacylglycerides transfer directly from the bloodstream to the cells without forming chylomicrons (KENNEDY, 1991). Triacylglycerides with three medium chain fatty acids (MMMs) are also easily digested, but lack essential fatty acids such as linoleic or linolenic acid, which can be included in MLMs. Another application of MLMs is as lowcalorie fats. Shorter chain fatty acids have fewer calories per unit weight than long chain fatty acids. Stearic acid is poorly absorbed so stearic acid-containing fats impart less energy. Thus, a triacylglyceride containing stearic acid and short chain fatty acids contains fewer calories. Current syntheses of these materials (e.g., Salatrim (SMITHet al., 1994), Caprenin) use chemical catalysts yielding a random distribution of fatty acids, but several researchers made these materials using lipases (Eq. 26) (AKOH,1995; SHIEHet al., 1995; SOUMANOU et al., 1997;MCNEILLand SONNET, 1995).

4.2.1.1.2 Synthesis of MLMs MLMs are triacylglycerides containing medium chain fatty acids (usually C8 :0 or C10: 0) Tab. 13. CommerciallyAvailable Chemically Synthesized MLMs Product

Composition

Company

Captex Neobee Caprenin Salatrim

C8:0, ClO:O,C18 :2 C8:0, ClO:O,LCFA” C6:0, C8:0, C22 :0 C3 :0, C4 :0, C18 :0

ABITEC, Columbus. OH Stepan Co., Maywood, NJ Procter & Gamble, Cincinnati, OH Nabisco Foods, East Hanover, NJ

a

LCFA= long chain fatty acids.


137

For instance, the interesterification of trisun unsaturated fatty acids (20:ln-9, 22: ln-9) 90 (an oil containing 90% triolein) with capric over EPA or DHA. For this reason, incorporaacid using RML yielded 69% COC (also contion of PUFAs into triacylglycerides containtaining CCO and OCC) after 30 h reaction in ing monounsaturated acids was slow. Direct esterification of PUFAs (or their hexane (SHIEHet al., 1995).A two-step process gave significantly higher yields and purer ethyl esters) and glycerol yielded monoacylproducts (SOUMANOU et al., in press). In the glycerides, and some di- and triacylglycerides first step, alcoholysis (see Sect. 4.2.1.3.1) of (KOSUGIand AZUMA,1994; ZUYIand WARD, pure triglycerides or natural fats with 1,3-re- 1993; HARALDSSON et al., 1993). Esterification giospecific lipases (e.g., ROL) yielded sn-2- of isopropylidene glycerol with a mixture of monoglycerides (2-MG) in up to 72% yield EPA and DHA using RML yielded protected after crystallization. In the second step, the monoacylglycerols with a maximum yield of same lipases catalyzed esterification of these 80% (ZUYIand WARD,1994). 2-monoglycerides with caprylic acid. The final product contained more than 90% caprylic acid in sn-1- and sn-3-positions, whereas the 4.2.1.1.4 Other Triglycerides sn-Zposition was composed of 98.5% unsaturated long chain fatty acids. Another potential application of lipases is in the synthesis of “zero-trans” margarines (MARANGONI and ROUSSEAU, 1995). Partial hydrogenation of oils to increase the melting 4.2.1.1.3 Triacylglycerides point also introduces trans-isomers of unsatuContaining Polyunsaturated rated fatty acids. Natural oils contain only cisisomers and the trans-isomers may contribute Fatty Acids (PUFAs) to heart disease. Interesterifications similar to Polyunsaturated fatty acids, PUFAs, for ex- those for the synthesis of cocoa butter also ample, eicosapentaenoic acid (EPA, 20: 511-3) raise the melting points of oils without introand docosahexaenoic acid (DHA, 22: 6n-3), ducing trans-isomers, but the cost is much are essential fatty acids and may be beneficial higher than hydrogenation. An alternative to triacylglyceride modificain cardiovascular and inflammatory diseases (AKOH,1995; KOSUGIand AZUMA,1994; PE- tion is genetic engineering of the metabolic DERSEN and HOLMER, 1995). PUFAS are most pathways in plants that produce the oils, so efficiently absorbed as triacylglycerides. Ele- they produce more of the desired oils. For exvated temperatures and extreme pHs pro- ample, high laurate canola has the melting bemotes side reactions, oxidation, cis-trans isom- haviour of a saturated fat, but retains the un1996). erization, or double-bond migrations in PUFAs saturated oleate chain at sn-2 (KINNEY, (HARALDSSON et al., 1993), so lipases are ideal for syntheses of PUFA-containing triacylglycerides. 4.2.1.2 Diacylglycerides Researchers incorporated EPA or DHA Diacylglycerides (DAGs) occur as 1,3-DAGs into soybean oil to a final content of 10.5 to 34.7% (HUANGand AKOH,1994), into sardine and the racemic mixture of 12-DAG and 2,3oil up to 70% in the presence of ethylene gly- DAG which we abbreviate as 1,2(2,3)-DAGs. col as water mimic (HOSOKAWA et al., 1995), Mixtures of isomeric DAGs are used with into trilinolein up to 70 and 81% using RML MAGSas emulsifiers, but pure regioisomers of and EPA or DHA ethyl esters (AKOHet al., DAGs are most useful as chemical intermedi1993). Re1995),and completely into glyceryl ether lipids ates (BERGERand SCHNEIDER, (HARALDSSON and THORARENSEN, 1994). SHI- searchers used DAGs for the synthesis of phospho- (VANDEENENand DE HAAS,1963) MADA et al. (1995) incorporated up to 51% arachidonic acid into single cell oil from Mor- and glycolipids (WEHRLI and POMERANZ, tierella alpina with CRL. PEDERSON and HOL- 1969),and conjugated DAGs to drugs to create more lipid-soluble prodrugs (GARZON-ABURMER (1995) reported that RML favored mono-

138


BEH et al., 1983,1986; SARAIVA-CONCALVES et tion creates a large surface area and also prevents the glycerol from coating the lipase al., 1989). et al., 1997). RML-catalyzed esterOf the many possible routes to 1,3-DAGs (CASTILLO (Fig. 106) the best one is the esterification of ification with fatty acid methyl esters or ROLglycerol with a free fatty acid (or derivatives catalyzed esterification with fatty acid vinyl like ethyl- or vinyl esters) using a 1,3-selective esters, both yielded 1,3-DAG.Vinyl esters relipase (Tab. 14). To combine the immiscible acted faster than simple esters, but are more glycerol and fatty acid, BERGERet al. (1992) expensive. Both the yields ( > 70%) and the readsorbed glycerol onto silica gel. This adsorp- gioisomeric purity ( > 98%) were high. Al-

?

E

0-C-R

TAG

0-C-R I ,2 (2,3)-DAG

LOH 2-MAG

0-C-R

OH

.*& : acyl migration =-: lipase reaction H: hydrolysis A: alcoholysis E: esterification

OH

OH

0-C-R

1 (3)-MAG

glycerol

et al., Fig. 106. Reactions during the lipase-catalyzed synthesis of DAG (adapted from MILLQVIST-FUREBY 1997). Tab. 14. Lipase-Catalyzed Syntheses of Diacylglycerides(DAGs)”

Acyl Acceptor

Acyl Donor

Glycerol Glycerol

palmitic acid capric acid

Glvrpml --J-----

.

-..r-J----

vinvl ranrvlatp

Glycerol Glycerol Glycerol Glycerol Glycerol

lauric acid oleic acid beef tallow palmitic acid ethyl caproate

Water Ethanol

triolein trilaurin

Reaction System

Yieldb

solid-phase free evaporation MTRF -I----

MTBE 2-butanone solid-phase hexane free evaporation/ precipitate phosphate buffer DIPE

Lipase‘

Reference

80 (1,3) 73 (1,3)

RJL RML

WEISS(1990) KIMand RHEE(1991)

80 (1,3) 28 (1,3) 90 (mix) 63 (1,3) 88 (1,3)

(Amano D) RML CVL I? sp. Ld RML ROL‘

BERGER et al. (1992) JANSSEN et al. (1993b) YAMANE et al. (1994) KWONet al. (1995) MILLQVIST-FUREBY et al. (1996a)

43 (mix) 75 (1,2)

PPL ProqL

PLOUet al. (1996) MILLQVIST-FUREBY et al. (1997)

Pol

R n \/- 3i- / a

--

ROT

RFRGFR et _. -- a1 11997) \----I

a Abbreviations: MTBE: methyl t-butyl ether; DIPE: diisopropyl ether. 1,3: 1,3-DAG; 1,2: 1,2(2,3)-DAG; mix: mixture of 13-DAG and 1,2(2,3)-DAG. See Tab. 1 for lipase abbreviations. Pseudomonas species lipase from Kurita Water Industries, Japan. Authors used lipase from Rhizopus arrhizus which has been reclassified as Rhizopus oryzae.


though ROL is more regioselective, they preferred RML because it had a higher activity.To increase yields, WEISS(1990) and YAMANEet al. (1994) carried out reactions in the solid phase, where crystallization of the diacylglycerol minimized further reactions and shifted the equilibrium toward 1,3-DAGs. The other regioisomer - 1,2(2,3)-DAG - is more difficult to prepare. Hydrolysis or alcoholysis of triacylglycerides initially yields 1,2(2.3)-DAG, but cleavage continues to 2MAG. In addition, the water promotes acyl migration so that 1,2(2,3)-DAG isomerizes to 1,3-DAG. MILLQVIST-FUREBY et al. (1997) formed 1,2(2,3)-DAG in 75% yield by alcoholysis of trilaurin using ProqL. Reaction condition control minimized overhydrolysis, while minimizing water activity avoided acyl migration.

4.2.1.2.1 Acyl Migration in Mono- and Diacylglycerides

139

THONSEN, 1994; MILLQVIST-FUREBY et al., 1996b). Surprisingly, acyl migration is faster in nonpolar solvent like hexane than in polar aprotic solvents like acetone (SJURSNES and ANTHONSEN, 1994; MILLQVIST-FUREBY et al., 1996b).The rate is still hundreds of times slower than in protic solvents. To further reduce acyl migration, one should minimize the water content (MILLQVIST-FUREBY et al., 1996b; HEISLER et al., 1991) and use polypropylene supports for lipases instead of ion exchange resins (MILLQVIST-FUREBY et al., 1996b). Hydrogen phosphate salts, used to control water activity, promoted acyl migration, but sulfate salts did not (SJURSNES et al., 1995). DORSET(1987) even observed an intermolecular acyl migration in crystalline 1,2(2,3)DAGs. Since the equilibrium constant favors the l(3)-MAGS over 2-MAGS by 9: 1, researchers often either ignore or even encourage acyl migration in this case. A 9: l ratio of regioisomers is sufficient for emulsifier applications. For synthetic application a crystallization can give pure l(3)-MAG in many cases.

Acyl migration is the intramolecular transfer of an acyl group to an adjacent hydroxyl group. This reaction isomerizes the regioisomers of mono- and diacylglycerides (Eqs. 27 and 28) thus degrading the regioselectivity of lip- 4.2.1.3 Monoacylglycerides ases. The equilibrium favors acylation of the (MAGs) less hindered primary position in both monoand diacylglycerides,although the equilibrium Monoacylglycerols (or simply monoglycconstant in diacylglycerides is only 1.5 (SER- erides) are the predominant emulsifiers in food, pharmaceuticals, and cosmetics (BAUDAREVICH, 1967). MANN et al.. 1988; SONNTAG, 1982). In addition, MAGs are also building blocks for synthesis of lipids, liquid crystals, and drug carriers (BERGER (27) and SCHNEIDER, 1993). Current MAG manufacture involves continuous glycerolysis of fats 2-MAG 1(3)-MAG and oils at 220-250°C using inorganic alkaline catalysts. Manufacturers avoid unsaturated fats because they burn or polymerize causing a dark color, off-odor, and burnt taste. This process yields technical MAG of -50% purity OC(0)R which is suitable for many applications. Pur1,2(2,3)-DAG 1,3-DAG ification by molecular distillation yields pure MAG: 90% MAG (an equilibrium mixture of Acid or base catalyze acyl migration, but regioisomers), 10% DAG (also mixed reacyl migration remains fast even at neutral pH gioisomers), and 95% ee tentative absolute configuration

25

Fig. 108. PPL-catalyzed polymerization of a diester and a diol.

F3C-o

+

OACF3

PPL 1,3-dirnethoxybenzene vacuum

HO-OH

Fig. 109. PPL-catalyzed polymerization under vacuum gives high molecular weight polyester.

Polymerizations starting with vinyl esters (UYAMAand KOBAYASHI, 1994) or oxime esters (ATHAWALE and GAONKAR, 1994) gave molecular weights up to 7000 (-35 repeat units). CHAUDHARY et al. (1995) lowered the molecular weight of polyesters from 2600 to 800 in supercritical fluoroform by changing the pressure to decrease the solubility of the polymer (see Sect. 2.4). Ring-opening polymerization is a special case of transesterification polymerization which does not release a molecule of alcohol. Lipase-catalyzed polymerization of .s-caprolactone with either PCL or PPL yields a polyester with a molecular weight up to 7700 (67 repeat units) (KNANIet al., 1993; UYAMAand KOBAYASHI, 1993; UYAMAet al., 1993; MACDONALDet al., 1995). Researchers added a small amount of alcohol such as butanol to initiate the polymerization. MACDONALD et al. (1995) suggested that water bound to the enzyme limits the molecular weight of the polymer by reacting with the oligomers (Fig. 110). Similar polymers form upon ring-opening polymerization of the 12-membered ll-undecanolide and the 16-membered 15-pentadecanolide (UYAMAet al., 1995) and also the four-membered P-propiolactones, including substituted p-propiolactones (SVIRKIN et al., 1996; NOBESet al., 1996). Ring-opening polyCAL-8

0 JO,OH

R-

* -

Fig. 110. Ring-opening polymerization of caprolactone.

merization of succinic anhydride with diol gave polymers with degrees of polymerization up to 14 (KOBAYASHI and UYAMA, 1993). Lipases also catalyze the degradation of polyesters (for examples see TOKIWAet al., 1979;NAGATA, 1996; KOYAMA and DOI,1996).

4.4 Other Lipase-Catalyzed Reactions In addition to various hydrolysis and transesterification reactions, CAL-B also catalyzed the “esterification” of carboxylic acids and hydrogen peroxide to peroxycarboxylic acids (BJORKLING et al., 1992; CUPERUS et al., 1994; KIRKet al., 1994). Peroxycarboxylic acids are more reactive than hydrogen peroxide and reacted in situ with olefins to give epoxides (Eq. 34). Similarly, added ketones underwent Baeyer-Villiger oxidation (LEMOULTet al., 1995).

R4 O

73 - 85% yield

(34)

146


5 Commercial Applications and Future Directions

5.1.2 Enantiomerically Pure Chemical Intermediates

The amounts of enantiomerically pure intermediates produced by 1 DSM-Andeno produce (R)-glycidol butyrate using a PPL-cata5.1 Commercial Applications lyzed resolution, but they did not reveal details of the process (LADNER and WHITESIDES, 1984; KLOOSTERMAN et al., 1988). Several groups 5.1.1 Food Ingredients have since studied this reaction and its scaleFood applications of lipases produce the up in more detail (WALTSand Fox, 1990; Wu large amounts of relatively inexpensive prod- et al., 1993;VAN TOLet al., 1995a,b) (Eq. 35). ucts. A Unilever subsidiary in Holland (QuestBASF produces enantiomerically pure Loders Croklaan) produces cocoa butter sub- amines using a Pseudornonas lipase-catalyzed stitute using a RML-catalyzed transesterifica- acylation (BALKENHOHL et al., 1997). A key tion of stearic acid with POP (palm oil mid part of the commercialization of this process fraction or high oleate sunflower oil) (see Sect. was the discovery that methoxyacetate esters 4.2.1.1). Multi-ton production is possible, but reacted much faster than simple esters. Actithe cost-effectiveness of this process depends vated esters are not suitable due to a competstrongly on the cost of cocoa butter and the ing uncatalyzed acylation (Eq. 36). Chiroscience (Cambridge, UK) has scaledneeded oils. Unichema International produces esters up the dynamic kinetic resolution of (S)-rertsuch as decyl oleate, octyl palmitate, isopropyl leucine, an intermediate for the synthesis of myristate, isopropyl palmitate, and PEG400 conformationally restricted peptides and chimonostearate for skin care products using ral auxiliaries (TURNER et al., 1995;MCCAGUE CAL-B-catalyzed esterification (BOSLEY, 1997). and TAYLOR, 1997).(See Sect. 3.7.) Water is removed by vacuum. Unichema calls these “bioesters” because products made by lipase-catalyzed process starting from natural 5 ’1‘3 Enantiomerically Pure materials retain their “natural” designation. Other companies may produce flavor esters Pharmaceutical Intermediates such as isoamyl acetate or geranyl acetate by lipases. Both DSM-Andeno (Netherlands) and TaUnichema also produces biodegradable sur- nabe Pharmaceutical (Osaka, Japan) in collabfactants using lipases. A CAL-B-catalyzed es- oration with Sepracor (Marlborough, MA) terification of ethyl glucoside yields the 6 - 0 have commercialized lipase-catalyzed resolutions of ( )-(2S,3R)-MPGM, a key precursor ester (see Sect. 4.1.1.1.3).

+

(*)-glycidyl butyrate

(R)-glycidyl butyrate

0

II

racemate

R = H, 4Me, 3-OMe

(R)-glycidol

0 L O M e HN

NH2

147


to diltiazem (MATSUMAE et al., 1993, 1994; yields an aldehyde which reacts with the bisulFURUIet al., 1996; HULSHOFand ROKSHAM, fite in the aqueous phase. In the absence of 1989).The DSM-Andeno process uses RML, bisulfite, this aldehyde deactivates the lipase. while the Tanabe process uses a lipase secreted The desired (+)-MPGM remains in the toby Serrutia marcescens Sr418000. In both cases luene phase and circulates back to the crystalthe lipase catalyzed hydrolysis of the unwant- lizer where it crystallizes. Lipase activity drops ed enantiomer with high enantioselectivity significantly after eight runs and the mem( E > 100).The resulting acid spontaneously de- brane must be recharged with additional lipase. Although the researchers detected no composed to an aldehyde (Fig. 111). Details for the Tanabe process are given be- lipase-catalyzed hydrolysis of ( - )-MPGM, low. A membrane reactor and crystallizer com- chemical hydrolysis lowered the apparent bine hydrolysis, separation, and crystallization enantioselectivity to E = 135 under typical of ( +)-(2R,3S)-MPGM. Toluene dissolves the reaction conditions. The yield of crystalline racemic substrate in the crystallizer and carries (+)-(2R,3S)-MPGM is >43% with 100% it to the membrane containing immobilized chemical and enantiomeric purity. Glaxo resolves (lS,2S)-trans-2-methoxycylipase. The lipase catalyzes hydrolysis of the unwanted (-)-MPGM to the acid, which then clohexanol, a secondary alcohol, on a ton scale passes through the membrane into an aqueous for the synthesis of the tricylic p-lactam antiphase. Spontaneous decarboxylation of the acid biotic (STEADet al., 1996). The slow-reacting

HOOC

+ MeOH

lipase from Me0

racemic trans-isomer

toluene-water/NaHSO 3 membrane reactor

Me0 (+)-(ZR.BS)-MPGM ...

OMe spontaneous decarboxylation

3 steps

1

*

"m,

HCI

OMe

+

IMeO'

coz

I

Fig. 111. Commercial synthesis of diltiazem by Tanabe Pharmaceutical uses a kinetic resolution catalyzed by lipase from Serrutiu marcescens.

e;

CAL-B. 37 glL vinyl acetate, 1.7 M triethylamine, 0.16 M ,..OMe cvclohexane. 6-8h

racemic

is

,..OMe MeO.,,

+

>99% ee 36% yield

Fig. 1U. Glaxo resolves a building block for antibiotic synthesis.

w

-*-w antibiotic

148


enantiomer needed for synthesis is recovered in 99% ee from an acetylation of the racemate with vinyl acetate in cyclohexane. Immobilized CAL-B and PFL (Biocatalysts, Ltd.) both showed high enantioselectivity, but CAL-B was more stable over multiple use cycles. Other workers had resolved this alcohol by hydrolysis of its esters with PCL, CRL, or pig liver acetone powder (LAUMEN et al., 1989; HONIGand SEUFER-WASSERTHAL, 1990;BASAVAIAH and KRISHNA, 1994), but Glaxo chose resolution by acylation of the alcohol because it yields the required slow-reacting alcohol directly (Fig. 112).

Researchers reported a number of other kilogram scale routes to pharmaceutical precursors that involve lipases. Selected examples are summarized in Fig. 113.

5.2 Future Directions

5.2.1 Reaction Engineering Large-scale applications, especially in organic solvents, require continued optimization of the reaction rate and enantioselectivity. Im-

OAc Ph-a*()cO N H for side chain of taxol, an anti-cancer drug Patel et al. (1994)

for carbovir. an anti-HIV agent enantiomer used for antihypercholesternicagents MacKeith eta/. (1993, 1994) HO\-

for antifungal agent Saksena et a/. (1995)

n

HOE

elastase inhibitor experimental treatment for cystic fibrosis Cvetovich et a/. 1996

for a thromboxane A2 antagonist Patel eta/. (1992b)

u.

LTD4 antagonist for asthma treatment (did not pass clinical trials) Hughes eta/. (1989, 1990)

I

COOn-Pr PCL, E = 32 - 68 isopropenyl acetate Sih (1996). Henegar et a/. (1997)

Fig. 113. Kilogram-scale routes to pharmaceutical precursors involving lipases.


149

mobilization techniques that prevent denatur- butions to enantioselectivity (most modeling ation and allow lipases to adopt their more programs calculate only enthalpy contribuactive open conformation will continue to be tions), neglect of long-range Coulombic forces during calculations, and an incomplete underimportant. Efficient reactions in organic solvents also standing of the origins of enantioselectivity. Inrequire an optimum water activity. In most deed, results from directed evolution suggest cases, water activity increases activity, but also that residues far from the active site may promotes hydrolysis. To minimize hydrolysis strongly influence enantioselectivity (JAEGER researchers added water substitues such as et al., unpublished data). DMSO or methanol to increase activity (ALMARSSON and KLIBANOV, 1996; HUTCHEON et al., 1997). Dynamic kinetic resolutions, which increase 5.2.3 Directed Evolution of Lipases the maximum yield from 50 to loo%, are curDirected evolution is random mutagenesis rently limited by the inability to racemize the substrate. New reports using transition metals to create a library of mutant enzymes followed to catalyze racemization (DINHet al., 1996; by selection for the desired property. In many LARSSON et al., 1997; REETZand SCHIMOSSEK,cases, it is difficult to select for the desired 1996) look promising and will likely be a focus property (that is, to devise growth conditions which kill mutants without the desired properof future research. ty), so researchers also use screening methods to find desired mutants. Directed evolution is especially useful for cases like solvent tolerance or thermostability where current theories 5.2.2 Modeling and Mutating are inadequate to predict which structural the Selectivity of Lipases changes will give improvement. For example, You and ARNOLD(1996) randomly mutated The X-ray crystal structures of 11 synthet- subtilisin E and screened for increased total ically useful lipases are solved, often in several activity in 60% DMF. The mutant isolated conformations.The current challenge is to use after two rounds of mutation and screening this information to first, predict selectivity showed a 16-fold increase in total activity.The more precisely than the empirical rules and specific activity increased 3-fold and the second, to design mutants with modified selec- amount produced increased 5-fold. A similar tivities. Modeling indeed qualitatively predict- approach on a p-nitrobenzyl esterase ined the observed selectivity of lipases towards creased activity in 30% DMF 50-60-fold after alcohols (SAINZ-DIAZ et al., 1997; ZUEGGet four rounds of mutagenesis and screening 1996). In both cases the al., 1997) and toward carboxylic acids (BOTTA (MOOREand ARNOLD, et al., 1997).HOLMQUIST et al. (1996) explained mutated amino acids were far from the active several exceptions to the carboxylic acid rule site and could not have been predicted using a for CRL using modeling, while HOLZWARTH et rational design approach. Directed evolution al. (1997) explained changes in the selectivity might also be useful to create mutants capable of lipases toward triglyceride analogs. KLEINet to resolve sterically hindered substrates (BORNal. (1997) used modeling to design mutations SCHEUER et al., in press) and to increase the to change the acyl chain length selectivity. enantioselectivity of a lipase as shown recently Blocking the acyl binding site in Rhizopus by REETZet al. (1997). Using 2-methyl decanodelernar lipase with two tryptophan residues ic acid esters as the target substrate, they inincreased RDLs selectivity for short chains creased the enantioselectivity of a lipase from (butanoyl) 80-fold or more. However, several 1to 10. Thermostability is also difficult to increase other mutations did not yield the predicted results. Modeling cannot currently predict enan- rationally and thus is a good target for directed tioselectivity quantitatively. Possible reasons evolution. Random mutagenesis and screening for the failure include omitting entropy contri- yielded a more thermostable subtilisin (SAT-

150


et al., 1996) and lipase (SHINKAI et al., 1996). In contrast, a rational approach to increase the thermostability of Penicillium camembertii lipase by introducing a disulfide link failed (YAMAGUCHI et al., 1996). Although microbiologists have long improved strains using random mutagenesis (UV light or chemical mutagens) combined with screening or selection, the excitement of directed evolution comes from mutagenesis techniques such as error-prone PCR that target a single gene or even region of a gene. This targeting ensures that the improvement occurs only in the biocatalyst of interest and not due to other changes in the genome. The key to any directed evolution project is an efficient screening or selection technique (ZHAOand ARNOLD,1997).The improvement steps during directed evolution can be less than a factor of two, so screening requires an accurate method. Recently, we developed a fast method for measuring enantioselectivity of hydrolases (JANESand KAZLAUSKAS, 1997a) which may be useful for screening mutants with improved enantioselectivity. These screening techniques will also speed the screening of commercial hydrolases. Directed evolution does not replace modeling. Indeed researchers use modeling to rationalize the results of directed evolution and to select target regions for mutagenesis. Even random mutagenesis cannot explore all possible structures. TLER

Acknowledgement We thank the NSERC (Canada) for financial support and Prof. ROLF D. SCHMIDand his group for their warm hospitality during ROMAS J. KAZLAUSKAS stay in Stuttgart (1995-96). Acknowledgement is made to the donors of The Petroleum Research Fund, administered by the ACS, for partial support of this research.

6 References AALTONEN, O., RANTAKYLAE, M. (1991), Biocatalysis in supercritical COz, Chemtech 21,240-248. I?, SAHAMOLLER, C. R. (1997), ADAM,W., GROER, Enzymatic preparation of optically active amethylene p-lactones by lipase-catalyzed kinetic resolution through asymmetric transesterification, Tetrahedron:Asymmetry 8,833-836. M., GROTE, J. (1996), Pseudomonas ceADAMCZYK, pacia lipase mediated amidation of benzyl esters, Tetrahedron Lett. 37,7913-7916. M.. GEBLER,J. C., MAITINGLY, F! G. ADAMCZYK, (1994), Lipase-mediated hydrolysis of Rapamycin 42-hemisuccinate benzyl and methyl esters, Tetrahedron Lett. 35,1019-1022. ADAMCZYK, M., GEBLER,J. C., GROTE,J. (1995), Chemo-enzymic transformations of sensitive systems. Preparation of digoxigenin haptens via regioselective lipase-mediated hydrolysis, Tetrahedron Lett. 36,6987-6990. K., BJORKLING, F., GODTFREDSEN, S. E., ADELHORST, KIRK,0.(1990),Enzyme catalyzed preparation of 6-O-acylglucopyranosides,Synthesis 112-1 15. ADER,U., SCHNEIDER, M. F? (1992), Enzyme assisted preparation of enantiomerically pure P-adrenergic blockers. 111. Optically active chlorohydrin derivatives and their conversion, Tetrahedron: Asymmetry 3,521-524. I?, UGUEN, D. (1993), DesymADJE,N., BREUILLES, metrization of meso-propargylic diols, Terrahedron Lett. 34,46314634. P. (1991), On the importance of the ADLERCREUTZ, support material for enzymatic synthesis in organic media. Support effects at controlled water activity, Eur. J. Biochem. 199,609-614. ADLERCREUTZ, l?(1994),Enzyme-catalyzed lipid modification,Biotechnol. Genet. Eng. Rev. U , 231-254. H., CHIN,L. C., ARAI,K., FUJIADSHIRI, T., AKIYA, MOTO, K. (1992), Lipase-catalyzed interesterification of triglyceride with supercritical carbon dioxide extraction, J. Chem. Eng. Jpn. 25,104-105. AFFLECK, R., Xu, Z . F., SUZAWA, V., FOCHT,K., CLARK, D. S., DORDICK, J. S. (1992), Enzymatic catalysis and dynamics in low-water environments, Proc. Natl. Acad. Sci. USA 89.1100-1104. S. N., KAZLAUSKAS, R. J., MORINVILLE,A. H., AHMED, GROCHULSKI, P., SCHRAG, J. D., CYGLER,M. (1994), Enantioselectivity of Candida rugosa lipase toward carboxylic acids: a predictive rule from substrate mapping and X-ray crystallography, Biocatalysis 9,209-225. K., TAMAURA, AJIMA, Y.,YOSHIMOTO,T..TAKAHASHI, Y., SAITO, Y., INADA,Y. (1985), Polymerization of 10-hydroxydecanoic acid in benzene with polyethvlene elvcol-modified liuase. Biotechnol. Lett. 7.363-30;. .

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Esterases

SARAJ. PHYTHIAN Exeter, UK

1 Introduction 194 2 Pig Liver Esterase 194 2.1 Enantioselective Hydrolysis of Prochiral Diesters 195 2.2 Enantioselective Hydrolysis of meso-Diesters 196 2.3 Kinetic Resolution of Racemic Esters 198 2.4 Regioselective Hydrolysis 206 2.5 ChemoselectiveHydrolysis 207 2.6 Practical Considerations 207 2.7 The Active Site Model for Pig Liver Esterase 207 3 Horse Liver Esterase 209 4 Chicken Liver Esterase 211 5 Acetylcholinesterase 212 6 Cholesterol Esterase 214 7 Microbial Esterases 219 8 Proteases with Esterase Activity 221 8.1 Subtilisin 221 8.2 a-Chymotrypsin 226 8.3 Penicillin Acylase 229 8.4 Papain 233 9 References 235

194

4 Esterases

1 Introduction Esterases are a class of hydrolytic enzymes which, as the name implies, catalyze the formation and hydrolysis of carboxylic acid esters. A variety of esterases from mammalian sources are commercially available, the most widely used in organic synthesis being pig liver esterase (PLE). PLE has the advantage that it exhibits broad substrate specificity while maintaining high stereoselectivity. This, coupled with the fact that PLE is cheap, readily available, and does not require the presence of cofactors, makes it the esterase of choice in any screening program. Examples of the use of other esterases appear to a much lesser extent in the literature and only horse liver esterase (HLE), chicken liver esterase (CLE), acetylcholine esterase (ACE), and cholesterol esterase (CE) will be further considered here. In a number of cases, these esterases have been utilized when PLE has failed to give the required high stereoselectivity. Although less commonly used than PLE, these additional esterases are also easy to handle with only cholesterol esterase requiring the presence of a cofactor in the form of bile salts. In addition to the isolated enzyme esterases, there are a number of examples of whole microbial cells which possess esterase activity. Several proteases also possess esterase activity and are able to catalyze the selective hydrolysis and, in some cases, formation of ester bonds. The most frequently used enzymes of this group are subtilisin, a-chymotrypsin, and, to a lesser extent, penicillin acylase and papain. It is, perhaps, appropriate at this point to consider the differences between esterases and lipases. In nature, esterases catalyze the hydrolysis of carboxylic acid esters while lipases catalyze the hydrolysis of triglycerides, e.g., triolein (Fig. l),to form glycerols and fatty acids. When considering unnatural substrates, a basic “rule of thumb” can be applied. Esters which comprise a complex acid moiety and a simple alcohol moiety, e.g., a methyl ester, are preferentially hydrolyzed by an esterase (Fig. 2a). Esters which comprise a simple acid moiety and a complex alcohol moiety, e.g., an acetate, are preferentially hydrolyzed by a lipase (Fig.

Triolein R = (CHz),CH=CH( CH&CH3 Fig. 1.

Fig. 2.

2b). One other important difference between esterases and lipases is in their physicochemical interaction with substrates. Esterases exhibit normal Michaelis-Menten activity, with an increase in substrate concentration leading to an increase in enzyme activity. Esterases, therefore, operate in true solution, with watersoluble organic co-solvents being employed where necessary. In contrast to this, lipases show little activity until the concentration of the substrate is increased beyond its solubility limit. This is known as interfacial activation and as a consequence lipase catalyzed hydrolyses must be carried out with either high substrate concentrations or in biphasic media employing water-immiscible organic solvents such as hexane.

2 Pig Liver Esterase Pig liver esterase (EC 3.1.1.1) is a serine type of esterase (GREENZAID and JENKS, 1971) which has the biological role of hydrolyzing esters in the pig diet. It consists of a mixture of isozymes but, from the organic chemists’ point of view, can be regarded as a single enzyme

2 Pig Liver Esterase

since all the isozymes exhibit similar stereospecificity (LAMet al., 1988). PLE exhibits a broad substrate tolerance and has been used extensively to hydrolyze a wide range of carboxylic acid esters in the preparation of chiral synthons. The synthetic work employing PLE up to 1990 has been documented in two recent reviews (OHNOand OTSUKA,1989; ZHU and TEDFORD,1990). These reviews comprehensively list a large number of substrates and the reader is directed to these papers for further information concerning specific substrates.

Me02C

1

C02Me

195

1

pLE

MeO2C

CO2H

2 62%,99% ee

2.1 Enantioselective Hydrolysis of Prochiral Diesters

Fig. 3.

The first report of PLE being used in an enzymatic reaction came in 1975,when HUANG et al. (1975) demonstrated the successful application of PLE to the asymmetric hydrolysis of dimethyl P-hydroxy-P-methylglutarate (1) to give the monoester (2) with excellent enantiomeric excess (Fig. 3). The enantioselective hydrolysis of a wide range of substituted malonate esters (3) (Tab. 1) (BJORKLING et al., 1985a, b; HEIDELet al., 1994) and substituted glutarate esters (4) (Tab. 2) (HEROLDet al., 1983; LAMet al., 1986) has been reported. Tab. 1 shows selected examples of the hydrolysis of substituted malonate es-

ters. For a series of 2,2-disubstituted malonates, a reversal of enantioselectivity was observed. For substrates with a short alkyl chain, PLE hydrolysis gave the S-enantiomer whereas substrates with larger alkyl and aryl groups gave rise to the R-enantiomer. The successful enantioselective hydrolysis of the glutarate esters (4) (Tab. 2) demonstrates that chiral recognition by PLE is still possible when the prochiral center is P to the reaction site. A range of 3-(protected amino) glutarates and acylamino glutarates have also been efficiently hydrolyzed using PLE (Tab. 2) (OHNO et al., 1981;ADACHIet al., 1986).

Tab. 1. Asymmetrization of Prochiral Malonates by PLE MeO2CYCO2Me

3 R

PLE

___c

Yc02H

Me02C

HO2C;O ' 2Meor

S Configuration

R

ee

Reference

73 52 58 46 87

BJORKLING et al. (1985a) BJORKLING et al. (1985a) BJORKLING et al. (1985a) BJORKLING et al. (1985a) BJORKLING et al. (1985a) BJORKLING et al. (1985a) BJORKLING et al. (1985b) BJORKLING et al. (1985b) HEIDELet al. (1994)

["/.I

88 45 82 81

196

4 Esterases

-

Tab. 2. Asymmetrizationof Prochiral Glutarates by PLE

M e O 2 X C O 2 M e 4

R

P E

R

R

Configuration

CH3

R

C2H5 n-C3H7 n-CJ413 Ph Bn CeHSCHZCOZNH CH,CONH CH3CH= CHCONH

R R R

R H or Me02C&C02H

H

H02C&C02Me

S

Yield

S S S

R S

ee

[%I

Reference

Pol

86 95 61 18 95 98 95 93 81 60

90 79 50 25 17 42 54 93 93 100

HEROLD et al. (1983) LAMet al. (1986) LAMet al. (1986) LAMet al. (1986) LAMet al. (1986) LAMet al. (1986) LAMet al. (1986) OHNO et al. (1981) ADACHI et al. (1986) ADACHI et al. (1986)

2.2 Enantioselective Hydrolysis of meso-Diesters

substrates as chiral synthons in the synthesis of natural products such as prostaglandins and cyclopentanoid natural products such as Brefeldin A, also carbapenem antibiotics and carboAcyclic rneso-diesters such as 2,3-disubsti- cyclic nucleosides. tuted succinates (5) (MOHRet al., 1983) and For the series of monocyclic meso-diesters 2,4-disubstituted glutarates ( 6 ) (MOHRet al., (7) (Tab. 3) there is a change in chiral recogni1983;CHENet al., 1981) have been hydrolyzed tion by the enzyme as the ring size of the cyclowith PLE to yield monoesters in good chemi- alkane moiety increases (MOHR et al., 1983; cal yield and moderate to excellent optical SCHNEIDER et al., 1984b). When the ring size is yield (Fig. 4). It is interesting to note that in small (n = 1 , 2 ) the S-carboxyl ester is hydroboth cases, incorporation of a hydroxy substi- lyzed whereas when the ring size is increased tuent increased the chiral recognition of the to a cyclohexane ring (n=4), it is the R-carenzyme. boxyl ester group which is hydrolyzed. For the Many examples of the enantioselective hy- cyclopentane ring (n =3) the chiral recognidrolysis of cyclic rneso-diesters have appeared tion is poor representing the “change-over’’ in the literature. This reflects the use of these point, thus resulting in low optical purity. Sim-

R’ = R3 = Me, R2 = H, 91%, 18%ee

Me02C

COzMe

5

R

w

6 Fig. 4.

CO2Me

R20A

Co2R3

PLE

Me02C

R’ = OH, R2 = Me, R3 = H, 92%. 18%ee

CQH

R R

= H , 8596.64% ee = OH, 35%, 98% ee


197

Tab. 3. Asyrnmetrization of Cyclic rneso-1,2-Dicarboxylatesby PLE

n

R'

1 2 3 4

Me Me H H

R2

Yield

ee

Reference

H H

92 98 80 98

100

MOHRet al. (1983) SCHNEIDER et al. (1984b) MOHRet al. (1983) MOHRet al. (1983)

Me Me

["/I

PJ1 94 9 78

Tab. 4. Asymmetrization of Cyclohexene rneso-1,2-Dicarboxylatesby PLE

R

Yield

ee

Reference

CH3 C& n-C3H7 i-C3H7 n-C4H,

99 67 68 5 18

99 27 25 2 13

GAISand LUKAS(1984,1986) ADACHIet al. (1986) ADACHIet al. (1986) ADACHIet al. (1986) ADACHIet al. (1986)

Pol

ilar enantioselectivity was observed for the unsaturated cyclohexene derivative (8)(Tab. 4, R=CH3) as was shown by the saturated cyclohexane derivative (Tab. 3, n=4) with the Rcarboxyl ester group being efficiently hydrolyzed in both cases. However, for the series of cyclohexene derivatives (8), the efficiency of the asymmetrization tailed off rapidly with increasing chain length of the R group (GAISand LUKAS, 1984,1986;ADACHIet al., 1986). For the cyclopentane rneso-diesters (9) (Tab. 5 ) , PLE catalyzed hydrolysis of the carbocyclic analog, dimethyl cis-cyclopentane1.3-dicarboxylate (X=CH,), gave the (1S,3R)monoester albeit with low optical purity (JoNES et al., 1985).The opposite enantioselectivity was observed for the heterocyclic analogs (X=O, S, NBz) giving the (lR3S)-monoesters (JONESet al., 1985; KURIHARA et al., 1985; BJORKLING et al., 1987).

[% 1

A large number of bi- and tricyclic mesodiesters have undergone asymmetrization by PLE, the resulting half acid-esters being used as starting materials in natural products synthesis. 7-0xabicyclo[2.2.1]heptane-2,3-dicarboxylates (10-12)were hydrolyzed with PLE to give the corresponding monoesters (13-15) (Fig. 5) (BLOCHet al., 1985).Hydrolysis of the exo-diesters (10)and (11)gave the monoesters (13)and (14)with good to excellent enantiomeric excess, while the more sterically demanding endo-diester (U) was less selectively hydrolyzed. The series of tricyclic diesters (16) (Tab. 6) were hydrolyzed with diminishing chemical yield as the size of the ester group increased. The most favorable results were obtained with the dimethyl ester (entry 1) (ARITA et al., 1983;ADACHIet al., 1986;ITO et al., 1981).

198

4 Esterases

Tab. 5. Asymmetrization of Cyclopentane rneso-1,3-Dicarboxylatesby PLE

X

R'

RZ

Yield

ee

Reference

CHz

Me

S NBz

H

H Me Me Me

82 98 83 85 39

34 42 46 80 100

JONES et al. (1985) JONES et al. (1985) JONES et al. (1985) KURIHARA(1985) BJ~RKLING et al. (1987)

H H

0

a

[%I

13 86%, 75% ee

10

C02Me

11

12

C0,Me -

[% 1

PLE% C02Me 14 82%, 98%ee

15 87%, 64%ee

Hydrolysis of the diacetates of a series of cyclic meso-diols (18) of varying ring size yielded the corresponding monoesters in low yields and poor enantioselectivity (Tab. 8) (LAUMEN and SCHNEIDER, 1985; SABBIONI and JONES, 1987). However, the nitro-containing cis-1,3diacetate (19) (Fig. 6) proved to be an excellent substrate for PLE giving the monoacetate in high chemical yield and excellent optical purity (SEEBACH and EBERLE, 1986). PLE has also been used to generate planar chirality in (arene)tricarbonylchromium compounds (Fig. 7) (MALEZIEUX et al., 1992). Hydrolysis of the meso-diester (20) proceeded with p r o 4 specificity giving the half ester (21). Following derivatization, analysis by chiral HPLC revealed excellent enantiomeric excesses.

Fig. 5.

PLE can also be used to asymmetrize mesodiols by enantioselective hydrolysis of the corresponding meso-diacetates. For example, enantioselective hydrolysis of the diesters of a series of cyclopentene cis-1,3-meso-diols (17) (Tab. 7) resulted in high chemical and optical yields for the diacetate, although these rapidly decreased with increasing alkyl chain length (LAUMEN and SCHNEIDER, 1984).The resulting cyclopentane monoester is one of the most important chiral synthons used for prostaglandin synthesis.

2.3 Kinetic Resolution of Racemic Esters A variety of substrates have been reported to undergo kinetic resolution by PLE catalyzed hydrolysis. For a number of a-substituted a-hydroxy esters (22), the chiral recognition improves when the R group is switched from a methyl group to a more bulky phenyl group, leading to higher optical purities for both acid and ester at more acceptable conversions (Fig. 8) (MOORLAG et al., 1990,MOORLAG and KELLOGG, 1991).However, for a series of P-substi-


199

Tab. 6. Asymmetrization of Tricyclic meso-Diesters by PLE C02R2

'OzR2

PLE 16

Entry

X

R'

RZ

Yield

ee

Reference

1 2 3 4 5 6 7

-OC(CH3)20-OC(CH3)20-OC(CH,),O-OC (CH3),0 -OC (CH3)20-OC(CH,),O-

CH2 CH, CH2 CHz CH2

CH3 C2H5 n-C3H7 i-C3H7 n-C4H9 CH3 CH3

100 37 1.5 22 4.4 96 100

80 100 45 39 73 77 77

ARITAet al. (1983) ADACHI et al. (1986) ADACHI et al. (1986) ADACHI et el. (1986) ADACHI et al. (1986) ITO et al. (1981) ITO et al. (1981)

0 0

-0-

P I

1%1

Tab. 7. Asymmetrization of Diacylated Cyclopentene meso-Diols by PLE (LAUMEN and SCHNEIDER, 1984)

Rmoe"o PLE

17

R

Yield

ee

["/.I

[Yo

I

86 66 30

86 52 trace Tab. 8. Asymmetrization of Cyclic meso-Diacetates by PLE

R'

RZ

Yield

1 2

Ac Ac

H H

3

H Ac

Ac

54 62 44 40 31

n

4

H

tuted P-hydroxy esters (23),where the chiral center is p t o the reaction site, the chiral recognition was found to be poor regardless of the size of the R group. In this case, high conversion rates were required to obtain high optical

[Yo

I

ee

Reference

44

LAUMEN and SCHNEIDER (1985) LAUMEN and SCHNEIDER (1985) SABBIONI and JONES (1987) LAUMEN and SCHNEIDER (1985) LAUMEN and SCHNEIDER (1985)

["/.I 0

4 8

4

purity and then only for the recovered ester (Fig. 9) (WILSONet al., 1983). For a series of a,a-disubstituted a-amino acid esters (24)(Fig. 10),PLEshowed poor enantioselectivity for a range of alkyl and aryl sub-

200

4 Esterases

_No2 A

c

NO2

V pLE H

V

__c

19

W C O )3

20

89%,98% ee

Fig. 6.

Fig. 7.

W C O )3 21 R = Me, 85%, 91% ee R = Et, 99%ee

R = M?

27%, 42% ee

43%, 17%ee

44%, 83%ee

41%, 86%ee

R = Ph

rac-22

5 1%conversion Fig. 8.

HC)Cco2Me PLE

R

buffer

rap23

Fig. 9.

R P h

r* “x HO K C O z H

R = Et 88%conversion

I

R = MezCHCH2 67%conversion

””

~

R R

13%ee

C02H

45%, 47% ee

PLE, buffer

H2NxCO2Et

rac-24 Fig. 10.

R = CH2CH=CH2 57%, 72% ee R = ”Bu 41%,93% ee

41%, 95% ee 31%,97%ee

Rx sc o z M e 12%,98% ee

X C O 2 M e

R s

26%. 91% ee


stituents with only two exceptions; high optical purities were observed where the R group was either ally1 or n-butyl (KAPTEIN et al., 1993). PLE has been employed in the kinetic resolution of a variety of cyclic mono- and diesters. For example, resolution of the truns-cyclopropanecarboxylates (25) and (26) gave the (1R,3R)-acids (27)and (28),and the unreacted (1S,3S)-esters (29) and (30),albeit with low optical purities (Fig. 11) (SCHNEIDER et al., 1984a). The trans-epoxy dicarboxylate (31) was successfully resolved using PLE. giving the (2R,3R)-ester and the (2S,3S)-acid in good yield and high optical purity (Tab. 9) (MOHR et al., 1987). However, the meso-aziridine (32) proved to be a poor substrate for PLE. No im-

)J

201

provement in enantioselectivity was observed by the addition of the bulky benzyloxycarbonyl group (33)(RENOLD and TAMM,1993). Further substrates which have been successfully resolved using PLE are the cyclopentanone dicarboxylate (M), (TANAKA et al., 1987) the N-acetylamidocyclopentene carboxylate (35) (SICSICet al., 1987), and the tricyclic monoester (36)(NAEMURA et al., 1993) (Fig. 12). In all of these cases, the optically pure substrates were required as starting materials for natural products synthesis. PLE has been applied to the kinetic resolution of a series of racemic E-caprolactones (37) resulting in hydrolysis of the lactone of S-configuration regardless of the alkyl chain length

50% conversion C02Me ( 1R,3R)-27

rac-25

( 1S,3S)-29

85%, 46% ee

80%.40% ee

MeOzC

H02C

COzMe

C02Me

Ma2C

85%, 60% ee

Fig. 11.

C02Me

( 1S,3s)- 3 0 90%, 50% ee

(lR,3R)-28

rap26

x\

4.

50%conversion

Tab. 9. Kinetic Resolution of Racemic Cyclic Diesters by PLE

Acid X

No.

n

~~

(31) (32) (33)

NH NZ

1 0

0

Ester

Yield

ee

1% 1

~~

0

(2R.3R)

(2S.3S)

rac-31-3 3

[% 1

Yield

1

[Yo

ee

Reference

95 27 28

MOHRet al. (1987) RENOLD and TAMM(1993) RENOLD and TAMM(1993)

[% 1

~~

40

95

40

-

202

4 Esterases

PLE

r a c 34

44%. 95%ee

45%, 95%ee

AcHNu C02Me

HO2CUNHAc

PLE

rac-35

A

c

H

NCOzMe ~

+

~

94%.97%ee

86%.87%ee

A

rap36

96%ee

83% ee

Fig. U.

R (Fig. 13) (FELLOUS et al., 1994). This is in contrast to HLE for which there is a reversal in enantioselectivity as the chain length increases. At 60% conversion, PLE gave the R-lactones (37)in 30-35% yield and 33-98% enantiomeric excess. While the majority of kinetic resolutions catalyzed by hydrolase enzymes have been applied to secondary alcohols, there are few reports of the kinetic resolution of quaternary centers, in particular tertiary alcohols and their acylated derivatives. This is because hydrolases

rac-3 7 R =Me, Et, "Pr, *Bu, "Pe, "Hex, "Hep,"Oct

Fig. 13.

30-35%yield 33-98%ee

do not readily accept such sterically demanding substrates. A popular tactic to overcome this problem is to use a tertiary alcohol which bears a second functional group capable of undergoing stereoselective hydrolysis or esterification, thus kinetic resolution is carried out via this second functional group. For example, the hydroxy esters (1)and (2) (Fig. 3) undergo enantioselective hydrolysis via the ester functionality. Examples where direct resolution of a quaternary center has been achieved include the a-substituted P-ketoester (38) (Fig. 14) and the tertiary quinuclidinol ester (40) (Fig. 15). For the P-ketoester (38) incorporating a variety of R groups, PLE catalyzed the enantioselective hydrolysis of the ethyl ester leading to a p-ketoacid which underwent decarboxylation during work up to yield the substituted ketone (39), plus the optically pure p-ketoester (Fig. 14) (WESTERMANN et al., 1993). Resolution of the racemic tertiary alcohol 3hydroxy-3-ethynyl quinuclidine, required in optically pure form for the synthesis of squalene synthase inhibitors, was achieved by PLE catalyzed hydrolysis of the corresponding bu-

p


203

I

OAc

R = Me, "Bu, "Pe, ( C H z ) E N , (CH2)3CN

40-8846 yield, >98%ee

OAc

rac-41

PLE, buffer 50% conversion

Fig. 16.

Fig. 14.

tyrate ester (40) (Fig. 15) (COOPEand MAIN, 1995). The resolution failed when the alkyne group was replaced by either an alkyl or aryl group.This was seen as evidence to support the view that the enzyme recognizes the acetylene substituent as essentially the same as a hydrogen atom, thus allowing hydrolysis to occur. The kinetic resolution of racemic acyclic and cyclic 12-diols can be achieved, in most cases, by the enantioselective hydrolysis of the corresponding diacetates. These compounds are of interest because of their potential use as chiral

35% conversion 35%, 97% ee

rac40

56%conversion 40%, 99% ee

1 PLE, HzO,

2 KOH, H20, MeOH Fig. 15.

1Y

auxiliaries and ligands for hydrogenation catalysts or chiral crown ethers. The racemic truns2,5-disubstituted tetrahydrofuran derivative (41), which was required for use as a chiral building block in the preparation of polyether antibiotics, gave the (2S,5S)-alcohol plus the unreacted (2R,SR)-diacetate in only moderate optical purity (Fig. 16) (NAEMURA et al.. 1993). Similarly, the rigid bicyclic compound bicyclo[2.l.l]heptane-2,5-diacetate (42) gave the ( -)-alcohol plus the ( +)-diacetate in very low optical purity (Fig. 17).Higher enantioselectivity was achieved with the bicyclo[2.2.2]octane2,3-diacetate (43) which gave the ( - )-alcohol plus the (+)-diacetate in good enantiomeric excess (Fig. 18) (NAEMURA et al., 1992). The cyclohexane-1,2-diol derivatives (44) and ( 4 9 required in high optical purity as subunits for the preparation of chiral crown ethers, were prepared by PLE catalyzed hydrolysis of their monoacetates (46)and (47) respectively. Termination of the reaction at around 50% conversion gave the diols (+)(44) and (-)-(45) plus the unreacted acetates (-)-(a) and (+)-(47) in high enantiomeric excess (Fig. 19) (NAEMURA et al., 1991). A series of cycloalkane-l,2-diols were resolved by the PLE catalyzed hydrolysis of the corresponding diacetates (48) (Fig. 20) (CROWet al., 1986). Although in the case of the cyclobutane and cyclohexane 1,Zdiols it was possible to obtain products with high opti-

204

4 Esterases

PLE OAc A A

~

C

OAc

nr42

(+)-42,50%, 15% ee

(-), 40%, 19%ee

Fig. 17.

PLE

Ac* OAc

~

+

&OH I

OAc

OAc

rar43

-$7

(-), 4896, 82% ee

(+)-43, 32%, 85% ee

Fig. 18.

OPh 111

OH

"40Ac

nc46

111

(+)-(

lR,ZR)-44,

48%, 84% ee

(-I-( 1S,2S)-46,

46%, 85% ee

OH

dIAc

rac47

(-)-(lS,ZR)-45,

47%, 78% ee

(+)-(lR,25)-47, SO%, 82% ee

Fig. 19.

cal purity, the pattern of the products from all three substrates was less clear cut with a mixture of products being obtained. CARONand KAZLAUSKAS(1991) suggested that for substrates of this type, i.e., any molecule with two reactive functional groups which can undergo two sequential reactions, the enantiomeric purity of the products could be improved by linking two kinetic resolutions to give a sequential kinetic resolution. In order for the optimum reinforcement of enantioselectivities to occur, the rates of the two steps should be

equal. This was demonstrated by truns-1,2-cyclohexane-diacetate (49) (Fig. 21). When the reaction was carried out in buffer, the first step proceeded 47 times faster than the second step, yielding the diol(50) in only 58% ee at 44 mol%. Addition of a hexane phase slowed the first step by selectively extracting the fastreacting diacetate from the aqueous phase. Using this technique the enantiomeric purity of diol(50) was increased to 94% ee at 34 mol% . Racemic l-phenyl-l,2-ethane diol has been successfully resolved using PLE catalyzed hy-

r

205


50% conversion

"'10 Ac OH 49%. >95%ee 41%,>95%ee lo%,>95% ee

41%, 95% ee

Fig. 20.

a:::a;;c El = 41

Fig. 21.

bo rac-5 1

( T O '//OH H

50,58% ee at 41 mol%

rac49

PLE

28% DMSO, buffer

Ph

33%, 95% ee

$* 85%. 97%ee

s 78% ee

Fig. 22.

drolysis of the corresponding cyclic carbonate (51) (Fig. 22) (BARTON and PAGE,1992). One advantage of using a cyclic carbonate is that no reactive intermediate is produced in the reaction since the by-product is carbon dioxide. Planar chirality can also be recognized by PLE. For example, both the racemic iron carbony1 complex (52) (ALCOCK et al., 1988) and the racemic allenic carboxylic ester (53)(RA-

MASWANY et al., 986) have been resolved efficiently (Fig. 23) It is also possible to resolve racemic esters w .ere the chirality is located on a heteroatom. Chiral phosphine oxides constitute an important class of chiral phosphorus compounds,particularly as precursors of chiral phosphines, which in turn, are widely used as chiral ligands in transition metal catalysts. A series of racemic methyl alkylphenylphosphin-

206

4 Esterases

racs 2

85% ee

85% ee

61% ee

90%ee

rac53 Fig. 23.

oylacetates (54) was hydrolyzed by PLE to 2.4 Regioselective Hydrolysis give the corresponding P-chiral phosphinoylacetic acids and unreacted esters in high enanPLE has been used to catalyze the regiosetiomeric purity (Tab. 10) (KIELBASINSKI et al., lective hydrolysis of the 1-(2,3,5-tri-O-acyl-p1994).There is also a report of the kinetic res- D-arabinofuranosy1)uracil derivative (55) to olution of a-(acety1oxy)phosphonates(Fig. 24) give the 2'-O-acyl monoester (56) in high yield for which PLE gave only poor yields in a bi- (Fig. 25).The monoester was required as a prophasic system (LI and HAMMERSCHMIDT, drug in the treatment of herpes simplex virus 1993). Better results were however obtained type 1 (HSV-1) (BARALDI et al., 1993). with a number of different lipases. Preparation of the P-hydroxy ester (57), required as an intermediate in p-lactam synthesis, was accomplished by regioselective hydrolQCOR~ ysis of the diester (58) using PLE (Fig. 26).This method was found to be slightly superior to OR^ chemical hydrolysis using potassium carbonR' bOR2 ate (BARTON et al., 1993). 0

A

Fig. 24.

Tab. 10. Kinetic Resolution of Racemic Methyl Alkylphenylphosphinoylacetatesby PLE (KIELBASINSKI et al., 1994)

Acid R CH, C A CH,=CH PhCH,

Yield

["/.I

42

41 18 43

Recovered Ester ee

1% I

82

81

-

79

Absolute Config. S S

R S

Yield

1%I 50 45 40 46

ee

1 1

Absolute Config.

OO /

74

> 96

100 80

R R S

R

2 Pig Liver Esterme

207

2.6 Practical Considerations Commercially available PLE has been used to carry out the majority of ester hydrolyses and is generally available as an ammonium sulfate solution (Sigma E3128, suspension in 3.2 M (NH4)*S04solution). The reactions are carried out at room temperature in phosphate buffer at pH 7. To obtain the best results, reacOR OH tions should be maintained at pH 7 using either a pH-stat charged with 0.1 M sodium hy56 55 R = Ac, pentanoyl, droxide solution or by monitoring and adjustmethoxyacetyl , Bz, ing the pH manually. Reactions should be terdecanoyl minated when the calculated amount of base has been consumed. Results can often be imFig. 25. proved by the addition of a co-solvent such as acetone. Using these techniques, reactions can be carried out on a multi-gram scale using a round-bottomed flask equipped with an effiR02C C02Me cient stirrer (HUTCHINSON et al., 1992). Although PLE has been successfully employed to catalyze the enantioselective hydrolysis of a large number of carboxylic acid esters 57 R = H in water, the reverse reaction, acylation of alcohols in organic solvents has proved to be imFig. 26. practical. Unlike lipases which function well in both aqueous and organic media, PLE shows only low and erratic activity in organic solvents (HEISSand GAIS,1995). One method of conferring activity on enzymes in organic sol2.5 Chemoselective Hydrolysis vents is to covalently link the enzyme to polyKELLOGGhas shown that for the ester (59), ethylene glycol monomethyl ether (MPEG). S have reported the prePLE will hydrolyze the ester group in prefer- HEISSand G A ~(1995) ence to the acetylsulfonyl group (Fig. 27) (HOF paration and characterization of MPEG-PLE and KELLOGG, 1995).This is in contrast to re- and give two examples of its application to the ports that lipases show the opposite chemo- enantioselective acylation of meso-diols (60) selectivity (BIANCHIand CESTI,1990; BABAet and (61). In both cases, toluene was used as the al., 1991). PLE has also been shown to hydro- solvent and vinyl acetate as the acylation relyze a phenolic acetate in the presence of an agent, with the corresponding acetates (62) and (63) being obtained in low optical purity aromatic methyl ester (BASAK et al., 1993). (Fig. 28).

Bn7-toH

SAC JCOzEt 59 Fig. 27.

pLE

2.7 The Active Site Model for Pig Liver Esterase f C 0 2 H

The undoubted synthetic potential of PLE to produce chiral acid-ester synthons from a wide variety of prochiral diester substrates, suffered a setback when it became clear that it was impossible to predict the stereochemical

208

4 Esterases

?H

Q

PAC

OH

6 2 52%, 56%ee

60

OH 61

Fig. 28.

g

OH

MPEG-PLE

vinyl acetate, toluene

OH

6 3 2696, 15%ee

drolysis by its attack on the carbonyl group of the ester function present in the substrate. The two hydrophobic pockets differ greatly in size. The larger pocket, designated H,, was initially thought to have dimensions of 4.6 * 3.1 2.3 A with a volume of approximately 33 A3,but this was later revised when H,'s capacity for large groups turned out to be greater than at first thought (TOONEand JONES,1991a,b). Its maximum dimensioqs have now been established as 6.1 .4.6.3.1 A (JONES,1993). The smaller pocket, designated H,, has dimensions of 1.6 2.3; 1.6 A with a volume of approximately 5.5 A3. The hydrophobic pockets accommodate the aliphatic and aromatic hydrocarbon portions of a substrate, and if necessary, can accommodate less polar heteroatom functions such as halogen and ether or ketal oxygen atoms. Polar groups such as hydroxyl, amino, carbonyl, and nitro are not accommodated by these pockets. The two more polar or hydrophilic pockets can accommodate more polar groups. They are located at the front (PF) and back (PB) of the active site, respectively. The dimensions of the binding pockets represent the physical restrictions placed on the substrates binding in the active site by the amino acids of the enzymes. With the exceptions of the area above the model and the rear +

outcome of the hydrolysis reactions. It was often found that within a series of substrates, the stereoselectivity was reversed, for example, the prochiral malonates (3) (Tab. 1) and the homologous series of monocyclic meso-diesters ( 7 )(Tab. 3). Initially it was thought that this stereochemical variability was due to the commercial PLE being composed of a mixture of enzymes, some possessing R and others S stereospecificity preferences. However, separation of PLE into its isozymes still produced the same stereospecificity results (LAM et al., 1988).The conclusion drawn from this was that although PLE is a mixture of isozymes, it behaves as a single enzyme and that the reversal in stereospecificity is, in fact, due to the structure of the enzyme's active site. This led JONES to probe the structure of the active site. Since no X-ray structure was available, JONES built up an active site model using an empirical approach. By using computer graphics, known literature examples were overlaid in order to build up a picture of the volumes and orientations permitted by the active site on steric grounds. From this, a surprisingly simTop perspective ple active site model was obtained (Fig. 29) (TOONEet al., 1990). The model consists of four binding pockets, two of which are hydrophobic (H, and H,) with the other two being more polar in character (PF and PB).Also essential to the active site model is the serine residue of the catalytic triad which initiates hy- Fig. 29.

h

1.G A

3 Horse Liver Esterase

boundary of the P, pocket which are both open, substrates are not able to penetrate these boundaries. In order to see how the active site model works,it is necessary to consider a homologous series of substrates such as (64-66) which, when incubated with PLE gave rise to the series of half acid-esters (6769) (Fig. 30). Two initial considerations must be taken into account. Firstly, hydrolysis of an ester group can only occur when it is located within the spherical locus of the catalytically active serine function. Secondly, binding of hydrophobic groups must occur in the Hs rather than the HL pocket if sterically possible. So, in the case of the cyclobutane diester (a), where hydrolysis of the ester of S-configuration is observed, the substrate adopts the orientation where the S-center ester is located in the serine sphere and the cyclobutane ring is located in the H, pocket.The alternative binding mode required for hydrolysis of the ester of R-configuration would place the cyclobutane ring in the unfavored HL pocket. In the case of the cyclopentane diester (65), which represents the changeover point in terms of stereospecificity, only a slight preference for hydrolysis of the ester of R-configuration was observed.This can be accounted for by the fact that both the ester groups of R- and S-configu-

*C02H

%C02Me

67 >97%ee

64

a C 0 2 M e S

C02Me

65 R

C02Me

a C 0 2 ~ ?

66

Fig. 30.

pLE

--

C

H

s C02Me 68 17% ee R

CO2H

a C 0 2 M e

69,>97% ee

209

ration can be acceptably located in the serine sphere due to the cyclopentane ring being accommodated in both the H, and H, pockets. However, the cyclopentane ring is slightly too large for an optimum fit in the H, pocket thus favoring the HL pocket resulting in the observed 17% ee of the 2R-acid (68). For the cyclohexane diester (66), hydrolysis of the ester group of R-configuration is favored since the cyclohexane group is too large to fit the Hs pocket and is consequently bound in the HL pocket resulting in the corresponding R-acid (69) in >97% ee.

3 Horse Liver Esterase Despite being commercially available, horse liver esterase has received little attention in the literature. It has been most notably employed in the kinetic resolution of racemic small- (BLANCOet al., 1988; FELLOUS et al., 1994) and medium-sized (FOUQUE and RousSEAU,1989) lactones (Tab. l l ) , and racemic bicyclic lactones (GuIBE-JAMPELet al., 1989) (Tab. 12) where, in the majority of cases, HLE gave enantiomeric excesses superior to those obtained with PLE. The kinetic resolution of lactones by enzyme catalyzed hydrolysis involves the hydrolysis of one enantiomer of the lactone to form the corresponding hydroxy acid. After separation from the optically pure lactone, the hydroxy acid can be relactonized thus providing access to both enantiomers of the lactone. In some cases (BLANCO et al., 1988) the production of hydroxy acids was found to inhibit the reaction. However, this could be overcome by the addition of 10% calcium chloride solution causing the partial precipitation of the calcium salts of the hydroxy acids thus driving the reaction to 5560% conversion. From the results in Tab. 11, it is apparent that the Gvalerolactones (entries 1-3) gave rise to lactones of S-configuration with high enantiomeric excesses irrespective of the alkyl group chain length, R. For the medium-ring lactones where n = 3-5 and R=CH3, the lactones of S-configuration were obtained with excellent enantiomeric excesses while the lac-

210

4 Esterases

Tab. 11. Kinetic Resolution of a Series of Racemic Lactones by HLE

Sor R

RorS

Lactone Entry

n

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17

1 1 1 2 2 2 2 2 2 2 2 2 2 2 3 4

R

5

Acid

Yield

ee

Config.

Yield [%]

45

95 78 92 76 84 22 35 4 38 72 53 50 60 63 >95 >95 >99

s

42

64

R

R R R

42

47

s

36 40 40

42 >95 >99

R R R

[%]

40

37 47 44

[%]

S S

ee

[%]

Config. Reference

s

R S S

s

s S S

s s s

FOUQUE and ROUSSEAU (1989) BLANCO et al. (1988) BLANCO et al. (1988) FOUQUE and ROUSSEAU (1989) FELLOUS et al. (1994) FELLOUS et al. (1994) BLANCO et al. (1988) FELLOUS et al. (1994) FELLOUS et al. (1994) BLANCO et al. (1988) FELLOUS et al. (1994) FELLOUS et al. (1994) FELLOUS et al. (1994) FELLOUS et al. (1994) FOUQUE and ROIJSSEAU(1989) FOUQUE and ROUSSEAU (1989) FOUQUE and ROUSSEAU (1989)

Tab. 12. Kinetic Resolution of a Series of Racemic Bicyclic Lactones by HLE (GUISE-JAMPEL et al., 1989)

Lactone n 1 2 3 4

Conversion

Yield

ee

Pol

Po

P o

55 55 55 55

41 42 43 40

1

98 80 3 47

I

Configuration lR, 5s 1R, 5s 1R, 5s 1R, 6 s

4 Chicken Liver Esterase

tones of R-configuration were obtained in moderate to excellent enantiomeric excesses by relactonization of the corresponding hydroxy acids (entries 15-17). In the case of the e-caprolactones, the alkyl chain length, R, was found to influence the enantioselectivity (entries 4-14). In the case where R=CH3, the lactone of R-configuration was obtained in 76% ee (FOUQUE and ROUSSEAU, 1989) and 84% ee (FELLOUS et al., 1994).Theoptical purity of the lactones decreased through the series to propyl (4% ee) and when the alkyl group was larger than propyl, HLE led to the hydrolysis of the opposite enantiomer with a concominant increase in the optical purity with increasing chain length. The reversal of enantioselectivity observed at the “change-over” point where R=C,H,, probably accounts for the discrepancy in the observed configurationsfor entries 7 and 8. In the kinetic resolution of racemic bicyclic et al., 1989) a crude lactones, (GuIBB-JAMPEL preparation of HLE acetone powder gave chiral lactones of greater optical purity than those obtained using either porcine pancreaticlipase (PPL) or PLE (Tab. 12).

Tab. W. 1989)

R

211

A series of racemic methyl 2-alkyl-2-arylesters proved to be excellent substrates for HLE (Tab. 13) (AHMAR et al., 1989). Enantioselective hydrolysis of the R-enantiomers was observed and the reactions were terminated at 4048% or 5&58% conversion depending on whether the acid or ester was required in greater optical purity. HLE acetone powder has also been used to catalyze the asymmetrization of prochiral organosilyl-substituted esters (DE JESO et al., 1990) (Tab, 14). The enantioselectivity was found to improve when DMSO was employed as a co-solvent.

4 Chicken Liver Esterase Chicken liver esterase was employed for the kinetic resolution of trans-1-acetoxy-2-arylcyclohexanes (70) (BASAVAIAH and DHARMA RAO,1994) (Fig. 31) and anti-homoallyl alcoand DHARMARAO, hols (71) (BASAVAIAH 1995)(Fig. 32) after PLE failed to give satisfac-

Kinetic Resolution of a Series of Racemic Methyl 2-Alkyl-2-aryl Esters by HLE (AHMARet al.,

Ar

Conversion Po

1

Yield

ee

Yield

34 45 34 39 34 36 38 36 40 36 35 41

92 53 91 72 88 66 93 66 91 94 91 92

48 38 43 35 35 31 46 36 33 36 42 41

[% 1

VJ1

[% 1

ee

Ph1

~

Me

Ph

Me

p-MeOPh

Me

p-‘BuPh

Et

Ph

‘Pr ‘Pr

Ph p-MeOPh

48

‘Pr

p-c1

46

40 56 42 50 40 58 42 51

46 52

43

>96

47 90 60 > 96 66 >96 92 76 >96 84

212

4 Esterases

Tab. 14. Asymmetrization of Prochiral Organosilyl-SubstitutedEsters (DE JESO et al., 1990)

L /co2R2

Me3Si

FR'C 0 2 R 2

H LE

RZ

Conversion

H Me Me

Et Et Me

100 100 100

[% 1

,C02H bC02R2 R'

0.2 M phosphate* buffer, 31°C

R'

EIC-70

Me3SiL

Yield [Yo

1

[Yo1

10 71 81

68

50

49

2 1-32% 90 - >99% ee

ee

53-67% 30-55% ee

Fig. 31.

71 , R = 'Pr, 'Bu,"Bu, "Pent, "Hex, Ph

20-3 1% 67- >99% ee

60-72% 19-50% ee

Fig. 32.

tory results. Although CLE is commercially available, the authors employed a crude enzyme preparation which was obtained by homogenization of fresh chicken livers in acetone using a food blender.

5 Acetylcholinesterase In nature, the role of acetylcholinesterase (EC 3.1.1.7) is to hydrolyze the neurotransmitter acetyl choline.The common source of commercially available ACE is the electric eel and as a consequence the cost is prohibitive for large-scale preparative synthesis.

The first report of the synthetic utility of ACE was by DEARDORFF et al. (1986) who described its use in the asymmetrization of mesodiacetate (72) (Tab. 15). The resulting alcohol (73) was obtained in excellent yield and enantiomeric excess. Following this, several reports described the use of ACE for the asymmetrization of 5-,6-, and 7-membered ring meso-diacetates (Tab. 15). In the case of the 5-membered ring derivatives, enzymatic hydrolysis resulted in the corresponding alcohols (73), (DANISHEFSKY et al., 1989), (74) (GRIFFITHS and DANISHEFSKY, 1991; LEGRAND and ROBERTS, 1992), and (75) (JOHNSON and PENNING, 1986) in high yield and enantiomeric excess, although there are conflicting reports concerning the absolute configuration of alcohol (74).

5 Acetylcholinesterase

213

Tab. 15. Products from the ACE Catalyzed Asymmetrization of Cyclic rneso-Diesters

No.

Product

Yield [% 1

ee

Reference

[%I

(72) R=Ac (73) R=H

94 89

(74)

95

>95

(75)

80

-

60

50-55 5 ,111 OR2 39

(77) R'=R2=Ac (78) R'=R2=H (79) R'=H, RZ=Ac

OH --

a

Not reported. The opposite enantiomer is reported.

96 nra

DEARDORFF et al. (1986) DANISHEFSKY et al. (1989)

GRIFFITHS and DANISHEFSKY (1991) LEGRAND and ROBERTS (1992)b

JOHNSON and PENNING (1986)

racemic SIHet al. (1992

PEARSON et al. 1987)

>98

-

-

PEARSON et al. (1989)

79

-

JOHNSON and SENANAYAKE (1989)

214

4 Esterases

No stereoselectivity was observed in the case of the cyclohexene derivative, resulting in the racemic alcohol (76) (SIHet al., 1992). In the case of the cycloheptene meso-diacetate (77), enantioselective hydrolysis proved difficult with increased amounts of diol (78) being obtained after prolonged treatment. Optimum results were obtained by terminating the reaction at approximately 50% conversion (8 h) to obtain hydroxy acetate (79) in 39% yield, 100% ee with diacetate (77) being recovered in 5 6 5 5 % yield plus diol (78) in 5% yield. Both the recovered diacetate and diol were subsequently recycled (PEARSON et al., 1987). No reaction was observed for diacetate (80) (PEARSON et al., 1989) but diol (81) was successfully obtained from the corresponding meso-diacetate and was subsequently utilized in a synthesis of compactin analogs (JOHNSON and SENANAYAKE, 1989).

Q1H.,-.- g C02Me

@.

COzMe

'///OH

r a c82

rac83

(+), 37%, 68% ee

(+), 11%, 93% ee

Fig. 33.

Early reports of the use of CE in organic synthesis resulted from initial screenings of a range of hydrolytic enzymes in order to find a suitable catalyst for a specific substrate. For example, enantioselective hydrolysis of esters of Cholesterol Esterase type (82) and (83) with varying alkyl chain The two most common sources of commer- length, identified CE from bovine pancreas as cially available cholesterol esterase (EC giving the most favorable results (Fig. 33) 3.1.1.13) are from bovine and porcine pancre- (PAWLAKand BERCHTOLD, 1987). Similarly, as. Its synthetic utility has been restricted to CE proved to be useful in the preparation of the resolution of bulky substrates similar in optically active D-myo-inositol analogs (84) size to the natural cholesteryl ester substrates. (Fig. 34) (LUI and CHEN,1989), although the The enzyme is only active in the presence of enzyme did not show complete enantio-disbile salts hence its modern name, bile salt-acti- crimination since a small amount (3%) of the vated lipase (BAL). The functions of the bile opposite acetate was produced along with the salts in vivo are fourfold: diol(85) as the major product. Limited success was however observed with (1) to activate CE, CE in the large-scale preparation of chiral bromide (86) (Fig. 35) (CHENAULT et al., 1987).Al(2) function as a biological detergent, though CE gave reasonable results on a 30 g (3) to protect CE from degradation by scale, it proved inferior to other enzymatic pancreatic proteases, and methods on a 50 g scale and consequently the (4) to protect CE from denaturation at the CE method was not optimized. water-lipid interface. Greater success has been achieved in the kiThe natural 3a,7a,l2a-trihydroxy bile salts netic resolution of binaphthols (KAZLAUSKAS, cholate, glycocholate and taurocholate are 1989, 1992) and spirobiindanols (KAZLAUSparticularly effective activators of CE, the KAS, 1992). For example, excellent yields and most commonly employed in synthesis being enantiomeric excesses were obtained for the sodium taurocholate. However, it has been enantioselective hydrolysis of binaphthyl dishown that the choice of bile salt can modulate pentanoate (87) (Fig. 36). The hydrophobic the stereoselectivity of the esterase (MOORE et substrates were hydrolyzed in emulsions of ethyl ether and phosphate buffer containing al., 1995).

6

6 Cholesterol Esterase

CE, D M F / p h o s p h a t e buffer, 0.5% meen

28°C.7 days

ElC

Fig. 34.

8 6 , 7 7 % ,80% ee Fig. 35.

taurocholate. The biocatalyst used for these investigations was pancreas acetone powder prepared from defatted, ground-up pancreas. This was chosen because it is commercially available, inexpensive, and possesses CE activity. More recently, KIEFER et al. (1994) have suc-

rap87 X = O E ~ C - 8 X8 = S

Fig. 36.

*

215

+

84 39%, 86% ee

85 53%, 85% ee

cessfully applied this methodology to the kinetic resolution of a binaphthalene dithiol by enantioselective hydrolysis of the corresponding dipentanoate (88) (Fig. 36). Due to the lack of X-ray crystallographic information concerning the structure of CE, its initial uses tended to be the result of random screening of lipases and esterases. The limited number of examples of its use is testament to the fact that, in most cases, efficient results can be achieved with either PLE or a lipase and these are simpler to use than CE which requires the presence of a bile salt. However, using the limited data available, KAZLAUSKASet al. (1991) have developed a rule for predicting which enantiomer of a secondary alcohol will react faster with CE. The rule predicts which enantiomer of a given acetylated substrate will be hydrolyzed by CE based on the relative sizes of the three substituents of the secondary alcohol. In broad terms, what this rule predicts is that secondary

X = 0,99%,>99% ee X = S, 94%. 98% ee

X = 0,>99%,>99.9% ee X = S, GO%, 98% ee

216

Fig. 37.

4 Esterases

Where M =medium and L = large

alcohols bearing substituents which differ greatly in size should be more efficiently resolved than those where the groups are similar in size. Fig. 37 illustrates the model and the fastest reacting enantiomer of secondary alcohols.

KAZLAUSKASapplied this rule to a number of examples including those which had previously appeared in the literature. Using the measured values of enantiomeric excess and percentage conversion, the enantioselectivity, E, was calculated. This gives an indication of the degree to which the enzyme prefers one enantiomer over the other (CHENet al., 1982). Out of 15 substrates, 14 were predicted correctly, corresponding to an accuracy of >93%. Tab. 16 shows the structures of the fast-reacting enantiomers, the exception being the enantiomer where E = l . The secondary alcohols are arranged so that the larger group is always on the right-hand side, as shown in Fig. 37.

Tab. 16. Fastest Reacting Enantiomers of Racemic Secondary Alcohols with CE as Predicted by KAZLAUSKAS et al. (1991)

w1

ee

43

69

8.8

46

35

2.6

46

45

4.3

Conversion

Substrate

34

1

E

[Yo

6.2

4.6

2.9

Br

6.3

29

65

50

94

> 100

44

56

10

217

6 Cholesterol Esterase Tab. 16. (Continued)

Substrate

GUFTAand KAZLAUSKAS(1993) then went on to demonstrate that for substrates where CE showed poor enantioselectivity (due to little difference in the size of substituents at the stereocenter), the enantioselectivity could be increased by modifying the substrate. Thus, increasing the difference in size between the substituents may help the enzyme to distin-

Conversion

ee

48

59

6.8

53

46

4.4

44

96

> 100

50

-

1

[% 1

Pol

E

guish between the two enantiomers. This strategy was successfully applied to a series of cyclic allylic alcohols. CE has also been used in the kinetic resolution of phosphines and phosphine oxides with phosphorus stereocenters (SERREQI and KAZLAUSKAS, 1994). In this approach, the enzyme did not act directly on the phosphorus stereo-

218

4 Esterases

center but instead catalyzed the enantioselec- oxides may serve as starting points for chiral tive hydrolysis of a pendant acetate group. A Wittig reagents and other reagents requiring a number of lipases plus CE were screened for stabilized methylene anion in a chiral environthe enantioselective hydrolysis of the phos- ment. Recently, CH~NEVERT phine oxide (89). From this, CE was found to and MARTIN(1992) be the enzyme of choice, although the enantio- have demonstrated that CE shows greater chimeric ratio, E, was low (4.8). To find a more ral discrimination in the asymmetrization of enantioselective reaction, a number of related dimethyl cis-cyclopentane-1,3-dicarboxylate substrates, (90-93) were examined (Tab. 17). (94) than that exhibited by a variety of proHydrolyses of both phosphine oxide (89) and teases, lipases, and esterases including PLE, phosphine (90) were modestly enantioselec- which gave the product in only 34% ee. CE tive. Moving the acetoxy group further from gave the half acid-ester (95) in 95% yield and the stereocenter as in (91), decreased the 90% ee (Fig. 38). enantioselectivity (E = 1.3), while replacing the phenyl group of (89) with a naphthyl group as in (92), increased the enantioselectivity by a COzMe COzMe factor of seven (E=32). This same replaceCE ment of phenyl by the naphthyl group in the phosphines (90) and (93) resulted in a small decrease in enantioselectivity. Thus, only (92) C ' OzMe * C'OzH was resolved with high enantioselectivity suggesting that both the P=O moiety and the 94 95 95%, 90% ee naphthyl group are required for high enantioselectivity.It is suggested that these phosphine Fig. 38

Tab. 17. Kinetic Resolution of Racemic Phosphines and Phosphine Oxides with Phosphorus Stereocenters by CE (SERREQI and KAZLAUSKAS, 1994) OAc

I

89 X=O 90 X=lone pair

91

No.

Conversion

(89) (90) (91) (92) (93)

52 40 40 42 51

92 X=O 93 X=lone pair

E

["/.I

53 ( R ) 49 (9 7 (S) 89 ( R ) 43 (S)

49 33 7 61 44

4.8 4.0

1.3 32 3.8

7 Microbial Esterases

7 Microbial Esterases

219

Bacillus subtilis var. niger has been employed in the preparation of chiral cyclohexyl alcohols involving enantioselective hydrolysis In addition to the use of isolated enzymes, a of the corresponding acetates (ORITANIand 1973, 1974, 1980a, b). It has also number of examples of microbial esterases can YAMASHITA, also be found. In the majority of cases, micro- been used by for the enantioselective hydrolybial esterases have been used for the kinetic sis of the acetates of racemic alkynyl alcohols resolution of secondary alcohols by enantiose- (97) (Fig. 41) and a-hydroxy esters, yielding lective hydrolysis of the corresponding ace- optically active acetates and alcohols in 7-90%0 optical purities (MORIand AKAO,1980a, b). tate. One early report by MCGRAHREN et al. The masked a-hydroxy aldehyde (98), re(1977) describes the use of the fungus Rhizo- quired as an intermediate in a prostaglandin synthesis, was conveniently prepared from the pus nigricans to prepare (-)-1-octyn-4-01(96) (Fig. 39), required as an intermediate in a pros- corresponding racemic acetate (99) by hydroltaglandin synthesis, by enantioselective hy- ysis using a Bacillus species (Fig. 42) (TAKAIdrolysis of the benzoate ester. Subsequently, SHI et al., 1982).It was necessary to protect the ZIFFERand coworkers have documented the carboxylic acid function as a t-butyl ester to use of R. nigricans for the enantioselective hy- prevent any undesired hydrolysis at this posidrolysis of over 50 acetates of various cyclic tion. OHTAet al. (1989a) have employed Bacillus and acyclic alcohols (ZIFFERet al., 1983;KASAI et al., 1984,1985;CHARTON and ZIFFER,1987). coagulans for the preparation of optically enThe aim was to prepare chiral alcohols of pre- riched cyanohydrin acetates (100) (Fig. 43). It dictable configurations from racemic esters. was necessary to use a large excess of lyophiFrom the results, a rule was proposed from lized cells in order to reduce the action of the which it was possible to predict the absolute cyanide produced during the reaction which stereochemistry of all the chiral alcohols. The had a detrimental effect on the stereoselectivrule is based on the relative sizes of substitu- ity. Under these conditions the (R)-cyanohyents and states that the enantiomer shown in drin acetate (100) was produced in 30% yield Fig. 40 is the one most rapidly hydrolyzed. Fur- with 100% optical purity. OHTAet al. (1989b) ther work made a quantitative analysis of the have also employed the yeast Pichia miso contributions of steric, electrical, and polariz- IAM 4682 in the preparation of optically enability effect in enantioselective hydrolysis riched cyanohydrin acetates (101) (Fig. 44). In this case, opposite enantioselectivity was ob(CHARTON and ZIFFER,1987).

OAc

2

(-)-96

rac97

Fig. 39.

R' = alkyl, R2 R 2 = H orCH3 Bacillus subtilis

var. niger

Fig. 40.

where R' is larger than R2 Fig. 41.

220

4 Esterases

I

Ac(kCN R CH3 mc-99

Pichio miso IAM 4682

Bacillus sp.

O'Bu (+)-98 32%, 94% ee

L+orBuS (-)-99 46%, 98% ee Fig. 42.

""k"" Ph

CF3

rac-100

rac-101 R = alkyl

( 9 - 1 0 1 28-38%, 9->99% ee Fig. 44.

Pichio miso IAM 4682 has also been employed in the preparation of a-chiral ketones (102) (MATSUMOet al., 1990) (Fig. 45). The methodology involved a novel enantioface differentiation in the hydrolysis of enol esters, resulting in the preparation of chiral six-, eight-, ten-, and twelve-membered-ring ketones with 70-96% enantiomeric excesses. Bakers' yeast (Succhuromyces cerevisiue) is well known for its dehydrogenase activity but less well known is that it possesses esterase activity. Perhaps this is less surprising when one considers that the natural process of gly-

Bacillus coagulans Pichia miso IAM4682,

(R)-100 30%, 100%ee Fig. 43.

served with hydrolysis of the R-enantiomer of a series of 1-cyano-1-methylalkyl and alkenyl acetates being observed. The required (S)-cyanohydrin acetates were obtained in 28-38% yield by recycling of the (R)-cyanohydrin to the racemic starting material via the corresponding parent ketone.

& e.

102 79%,90%ee Fig. 45.

R

rac-103

Fig. 46.

-

COzH H-k-NHAc I R

(R)-103

C02H AcHN-k-H I R

(9

*Y

8 Proteases with Esterase Activity

221

colysis which occurs in fermenting bakers' yeast, involves the formation and cleavage of many phosphate ester bonds. The esterase activity of fermenting yeast was first reported for the preparation of D-N-acetyl amino acid esOAc 106 ters (103) by enantioselective hydrolysis of their racemates (Fig. 46) (GLANZER et al., 1987a). For amino acid esters where the R Penicillium group was either an unbranched alkyl or arylfrequentans alkyl substituent, optical purities of 3-100% IMI 92265 were reported. Subsequently,a series of chiral 1-alkyn-3-01s (104) were prepared from the corresponding racemic acetates (*)-(105) in high optical purity using lyophilized yeast (Fig. et al., 1987b).1-Alkyn-3-01sare 47) (GLANZER useful starting materials in the synthesis of a OH 107 variety of natural products. A recent report has detailed the use of a microbial esterase in the regioselective hydroly- Fig. 48. sis of the triacetate (106) (Fig. 48) (WORONIECKI et al., 1994).The esterase,isolated from Penicillium frequentans IMI 92265 and subsequently immobilized,was used in the regioselective hydrolysis of the triacetate, 1P-diacetoxy-Zacetoxymethyl butane (106). The acetoxyethyl ester was hydrolyzed in preference to the acetoxymethylester groups yielding the alcohol (107),required for a synthesis of the 8.1 Subtilisin antiviral agents penciclovir and famciclovir. Subtilisin (EC 3.4.21.14) is a serine protease that aspecifically hydrolyzes peptides and proteins. It is commercially available as Alcalase" (Novo Nordisk) which is a crude preparation OAc of subtilisin. In addition to its proteolytic activity, subtilisin possesses esterase activity and will specifically hydrolyze the L-enantiomer of amino acid esters. This has been demonstrated by the use of Alcalase" to catalyze the enanR2 = H or Me raClO5 tioselective hydrolysis of the methyl and benzyl esters of racemic amino acids, giving rise to the L-amino acid and the D-amino acid ester in high yield (75-98%) and high optical purity (86100% ee) (CHENet al., 1986).Also, the kinetic resolution of N-acetyl-(R,S)-phenylalanine methyl ester has been achieved by ROPER and BAUER(1983). Other esters which have been successfully resolved using subtilisin are 2-amino-3-(2,2'bipyridiny1)propanoicacid methyl ester (lOS), (9-104 (R) required for a synthesis of peptides with po86 >97% ee 10 >97% ee tential metal-binding sites, (Fig. 49) (IMPERIALI et al., 1993) and a series of psulfonamidoproFig. 47.

1

AcY


-

-

222

4 Esterases

pionic acid esters (lo!)),required as starting materials for a synthesis of P-3 site modified rennin inhibitors (Tab. 18) (MAZDIYASNI et al., 1993). Subtilisin has also been used to catalyze the hydrolysis of esters of a-halo acids (110) in high yields (Fig. 50) (PUGNIERE et al., 1990). The authors also describe the transesterification of the same esters by simple alcohols catalyzed by subtilisin immobilized on alumina. Diastereoselective hydrolysis of racemic dipeptide esters, e.g., (111)has been achieved using Alcalasem (Fig. 51).The reactions were carried out in 40% acetone giving rise to dipeptides of high optical purity with no cleavage of the peptide bonds being observed (CHENet al., 1991). Regioselective hydrolysis of the dibenzyl esters of L-aspartic acid (112) and L-glutamic acid (113) has been achieved with AlcaIaseB

108

(Fig. 52). The reactions were carried out in an acetone/water mixture (1 :3) to improve solubility and to increase the rate of reaction. Selective hydrolysis of the a-benzyl ester group was observed in both cases, giving p-benzyl Laspartate and y-benzyl L-glutamate in 82% and 85% yields, respectively (CHEN and WANG,1987). A study of the regioselective hydrolysis of the 2 '-deoxy-3 ' ,5 '-di-0-hexanoyl pyrimidine nucleoside (114) met with limited success (Tab. 19). While it was found that differences in regioselectivity could be achieved according to the enzyme used (subtilisin favoring hydrolysis of the 5 ' ester while Pseudomonaspuorescens lipase favored hydrolysis of the 3' ester), total regioselectivity was not achieved resulting in mixtures of products. In the case of subtilisin, the desired 5 ' alcohol was observed,

93% ee

98% ee

Fig. 49.

Tab. 18. Kinetic Resolution of a Series of P-SulfonamidopropionicAcid Esters by Subtilisin (MAZDIYASMI et al., 1993)

X NH NCH3 0 0 NCH3

R

CH, CH3 CH3 CH, CHZCH3

Enzyme Subtilisin Carlsberg Alcalasem Alcalasem Subtilisin Carlsberg Alcalasem

Yield

["/I

90 70 80 60 80

ee P o

1

96 75 75 75 >98


a\

223

Alcalase@

CO2R'

110

H2

X = C1 or Br R = H, CH3, CH3CH2 R'= CH3, (CH3)2CH, CH3(CH2),CH2,n = 0,1,2,

L-112 COzBn I

C02Bn I

Alcalase@

Fig. 50.

L-113 Fig. 52.

Cbz-D,L-Ala-L-Phe-OBzl

1

111

however the major product resulted from deacylation of both the 3' and 5' positions with significant amounts of starting material being recovered (UEMURA et al., 1989). Selective manipulation of the C-2 and C-4 functionalities of 1,6-anhydro-2,4-di-O-acetyl3-azido-3-deoxy-~-~-glucopyranose (115) was possible via regioselective hydrolysis (Fig. 53). AlcalaseB catalyzed the hydrolysis of the acetate at the C-2 position resulting in the C-4 monoacetate in 82% yield. Conversely, Candidu cylindruceu lipase catalyzed the hydrolysis of the acetate at the C-4 position (HOLLAet al., 1992). The ability to differentiate between

Alcalase@

Cbz-L-Ala-L-Phe-OH

9296, 88% de

Cbz-D-Ala-L-Phe-OBzl

93%,90% de

Fig. 51.

Tab. 19. The Regioselective Hydrolysis of 2 '-Deoxy-3 ' , 5 '-di-0-hexanoyl Pyrimidine Nucleoside (114) by Subtilisin (UEMURA et al., 1989)

$ x

H

"Pe

Subtilisin 3'

DMF

HO

npeKo 0 114 X

Recovered (114)

["/.I

H

0

3'-alcohol

["/.I

5'-alcohol Pol

3',5'-Diol

1% I

~

H Br F CH3

24

32 7 7

0 0

0

0

31

12 22

28

45

54

71

65

224

4

Esterases

nonanoic acid. This lack of selectivity was overcome by the use of subtilisin Carlsberg, which regioselectively hydrolyzed the terminal ester group, yielding the acids (116)and (117). In addition to its hydrolytic activity in aqueous solution, subtilisin is moderately stable 115 82% and catalytically active in anhydrous organic solvents and consequently can be used for esFig.53. terification reactions. In particular, it has been employed in the regioselective esterification of the C-2 and C-4 hydroxy functions of the D- carbohydrates. This was first reported by RIVA glucopyranose (115)was then exploited in syn- et al. (1988) who selectively acylated glucose thesis by inversion of the c - 2 functionality (121)in anhydrous dimethylformamide using yielding, after chemical elaboration, a D-man- 2,2,2-trichloroethyl butyrate as the acyl donor nopyranose derivative. to give 6-0-butyryl glucose (122)in 60% yield Subtilisin Carlsberg has proved useful in the (Fig. 55). Similarly, KIM et al. (1988) have repreparation of the deprotected rearrangement ported the acylation of N-acetyl-D-mannosaisomers (116)and (117)of the marketed anti- mine (123) in dimethylformamide using isobiotic Pseudomonic acid A (118) (Fig. 54) propenyl acetate as the acyl donor, thus avoid(SIMEet al., 1987).The acid or base catalyzed ing the problem of reaction reversibility (Fig. rearrangement of Pseudomonic acid A (118) 56). Again, acylation of the primary hydroxy yields the two trans-fused bicyclic acids (116) group was observed. and (117).In order to separate the acids (116) Regioselective acylation of disaccharides and (117)and to carry out absolute structure has also been achieved with subtilisin. The didetermination, it was convenient to convert saccharides maltose (124), cellobiose (l25), the acids to the corresponding methyl esters, lactose (126), and sucrose (127) all reacted (119)and (120).However, acid or base hydrol- readily in anhydrous dimethylformamide emysis of the separated esters did not yield the ploying 2,2,2-trichloroethyl butyrate as the acoriginal acids (116)and (117),but instead hy- ylating agent (Fig. 57) (RIVAet al., 1988). In drolyzed both the activated allylic ester and the cases of maltose (124) and cellobiose the terminal ester groups releasing 9-hydroxy (125), acylation occurred exclusively at the C-6' hydroxy group. However, in the case of lactose (m), the enzyme was less discriminating giving acylation predominately at the C-6'

's H

OH

Ho

Subtilisin DMF "

120 R

=

Me

Fig.54.

OH

H

116 R = H 119 R = M e

PrC02CH zCC13

Hw

COz(CH2)&02R OH

121

H

Fig.55.

HO

OH

122 60%

HX 123

OH

H

225


H a - OOH h k

OH

128

1 I

Subtilisin, DMF PrC02CHZCC1,

OH Fig. 56. Fig. 58.

-

H 124

OH

OH

H&+oH OH

OH

125 H

B OH

H

HO

e

126

O

H

OH

’

tose moiety. This is in direct contrast to the chemical acylation where the most reactive position is the C-6 hydroxy group followed by the C-6 ’ hydroxy group. Regioselective acylation of the disaccharide,methyl P-lactopyranoside (128) has been achieved using subtilisin in anhydrous DMF (Fig. 58). In this case, the primary hydroxy group of the non-reducing sugar unit was acylated using 2,2,2-trichloroethyl butyrate as the acyl donor (CAIet al., 1992). Subtilisin has been employed for the regioselective acylation of the aza-sugars castanospermine (129) (MARGOLIN et al., 1990) and 1deoxynojirimycin (130) (DELINCKand MARGOLIN,1990) (Fig. 59). Castanospermine (l29), which possesses four secondary hydroxy groups, was selectively acylated at the C-1 position using vinyl acetate as the acylating agent and pyridine as the solvent. In the case of l-deoxynojirimycin (130),which possesses one primary and three secondary hydroxy groups plus a potentially reactive amino function, selective acylation of the primary hydroxy function oc-

127 Fig. 57.

hydroxy group but reaction was also observed at the C-3’ and C-4’ hydroxy groups. Unexpectedly, sucrose (l27),which possesses three primary hydroxy groups, was acylated exclusively at the C-1 ’ hydroxy group of the fruc-

129 Fig. 59.

H

130

6

226

4 Esterases

curred using 2,2,2-trichloroethyl butyrate as the acylating agent and pyridine as the solvent. No enzymatic acylation of the more reactive amino group was observed. The use of subtilisin in the regioselective acylation of carbohydrates is a consequence of their poor solubility in all but the most polar of organic solvents, such as DMF and pyridine. Most hydrolytic enzymes are inactive in these solvents and although subtilisin has proved useful, it too has limited stability. Thus, several DMF-stable subtilisin BPN’ variants have been developed by WONGand coworkers using site-directed mutagenesis to improve their stability (WONGet al., 1990). Subtilisin 8350 is a mutant derived from subtilisin BPN ‘ via six site-specific mutations. It was found to be 100 times more stable than the wild-type enzyme in aqueous solution at room temperature and 50 times more stable than the wild type in anhydrous DMF. Demonstration of the esterase activity of the mutant enzyme has been shown with the enantioselective hydrolysis of N-protected and unprotected common and uncommon amino acid esters in water showing 85-9870 enantioselectivity for the L-isomer, and in the regioselective acylation of nucleosides in anhydrous DMF with 65-100% regioselectivity at the 5 ‘-position. More recently, FITZand WONG(1994) have reported the use of the subtilisin variant 8399 in the regioselective acylation of N-acetyl-D-mannosamine (123)using vinyl acetate as the acyl donor in 97% DMF and 3% aqueous Tris buffer. Acylation occurred in 65% yield regioselectivity at the primary hydroxy group of N-acetyl-Dmannosamine (123). In addition to enzyme engineering as a means of influencing factors such as enzyme stability and enantioselectivity, solvent engineering has also been employed. For example, in a study of the kinetic resolution of racemic amines by enantioselective acylation, KLIBANOV and coworkers (KITAGUCHI et al., 1989) chose to resolve a-methyl benzylamine (131) using subtilisin in a range of solvents. From this initial screening, anhydrous 3-methyl-3-pentano1 was identified as the solvent showing greatest enantioselectivity yielding the (S)amide (132)in 89% ee (Fig. 60). These conditions were then successfully applied to the kinetic resolution of a number of racemic

Subtilisin PrC02CH2CF3

(s)-132,89%ee Fig. 60.

amines. Solvent engineering has also been employed in the enantioselective hydrolysis of 2chloroethyl esters of N-acetyl-L- and D-amino acids (SUKURAI et al., 1988).

8.2 a-Chymot rypsin a-Chymotrypsin (EC 3.4.21.1) is a serine protease which catalyzes the hydrolysis of amide bonds of proteins of aromatic amino acids such as Phe, Tyr, and Trp. It can also catalyze the hydrolysis of ester bonds and has been used synthetically in the enantioselective and regioselective transformation of a variety of amino acids and structurally related compounds. The enzyme is highly selective for the L-amino acids and this has been exploited in the enantioselective hydrolysis of N-acetyl-DLtryptophan methyl ester (NIEMANNand HUANG, 1951), N-acetyl-DL-tyrosine ethyl ester (NIEMANN et al., 1951), N-acetyl-phenylalanine methyl ester (CLEMENTand POTTER, 1971), ring-substituted phenylalanine ethyl esters (TONGet al., 1971), and protected racemic amino acid esters (BERGERet al., 1973). In each case, hydrolysis of the L-enantiomer occurred giving rise to the L-amino acids and the unreacted D-enantiomers, all with high optical purity.


F ~ rI F *

227

Unlike the common a-amino acid derivatives outlined above which contain chiral tertiary centers, a-substituted a-amino acid derivatives containing quaternary centers, are not ideal substrates for hydrolytic enzymes. Although a-methyl a-amino acid esters (ANANTHARAMAIAH and ROESKE,1982) and a-alka-Chymotrypsin enyl a-amino acid esters (SCHRICKER et al., 1992) have undergone enantioselective hydrolysis catalyzed by a-chymotrypsin giving the corresponding L-amino acids, the reaction 0 0 rates were slow. However, this problem has been overcome by LALONDEet al. (1988) who found that a-nitro a-methyl acid esters (133) were good substrates for a-chymotrypsin with \ \ R the D-enantiomers being preferentially hydrolyzed and the products undergoing spontaneR = H, 36% ous decarboxylation. The unreacted L-enanR = CH3,38% tiomers were obtained in high enantiomeric excess and were subsequently reduced to the Fig. 62. corresponding a-methyl L-amino acids (Fig. 61). These a-methyl a-amino acids have been used to replace natural a-amino acids in peptides, the increased steric bulk at the a-posi0 C02Me tion leading to conformational rigidity as well as resistance to hydrolysis by peptidases. rac-135 a-Chymotrypsin has been used to catalyze cis:trans 81:19 the kinetic resolution of racemic substrates. DIRLAMet al. (1987) applied a-chymotrypsin to the enantioselective hydrolysis of the bena-chymotrypsi n zopyran carboxylic acid derivative (134), required in a synthesis of the aldose reductase

KC’

rac-133 a-chymotrypsin J

L-133 , E = CH2=CH-CHr, Ph, >95%ee Fig. 61.

(+)-(2$3R) 38%, 86% ee

(-)-(3aR,6aR) 35%, 82% ee

Fig. 63.

inhibitor sorbinil (Fig. 62). Although the rate of hydrolysis was slow, the reaction was synthetically useful as it could be carried out on a multi-gram scale. Racemic methyl cis-3-chloromethyl-2-tetrahydrofurancarboxylate(135) has been successfully resolved using a-chymotrypsin (Fig. 63), the resulting lactone and unreacted ester being obtained in high optical purity (UDDING et al., 1993).

228

4 Esterases

One early report of the use of a-chymotrypsin concerned the enantioselective hydrolysis of the prochiral substrates (136) and (137) (Fig. 64) (COHENand KHEDOURI, 1961a, b). These substrates were chosen with a view to defining the structural requirements for the stereospecificity of a-chymotrypsin. Enantioselective hydrolysis of the meso-substrate (138) (Fig. 65) (SCHREGENBERGER and SEEBACH,1986) has also been reported, the resulting half acid-ester being required as a starting material in a total synthesis of the macrodiolide (+)-conglobatin. Enantioselective and regioselective hydrolysis has been used as a method of achieving kinetic resolution by selectively hydrolyzing one ester group in a racemic compound. For example, racemic diethyl-N-acetylaspartate (139) underwent a-chymotrypsin catalyzed hydrolysis of the a-ethyl ester group of only the Lenantiorner, yielding the half acid-ester in 100% ee (COHENet al., 1963).Similarly,the racernic diethyl ester (140)yielded the half acidester (141)in high optical purity (CROUTet al.,

~c~~~

Fig. 64.

I

1993) (Fig. 66). Also, a-chymotrypsin was found to hydrolyze the y-ester of the N-protected a-dehydroglutamate diester (142)(Fig. 67). This was in contrast to papain which hydrolyzed the a-ester (SHINet al., 1990). Optically active sulfoxides have been prepared using a-chymotrypsin (CARDELLICCHIO et al., 1994). In this case, the kinetic resolution of methyl (Z)-3-phenylsulfinylpropenoates (143) with a-chymotrypsin showed higher enantioselectivity than with a variety of lipases (Tab. 20). In an attempt to further improve the enantioselectivity of the reaction, co-solvents

540 times) was used in the final deprotection step of PhCH,CO-Asp-Phe-OMe (FUGANTI et al., 1986) and N-protection of the amino-substituted nucleoside bases adenine, guanine, and cytosine (Waldmann et al., 1994).

4 Esterases and Lipases Which Can Be Used with Amides and Peptides One of the most significant recent developments in the application of hydrolases to synthesis has been the realization that many enzymes are able to function in virtually anhydrous organic solvents and use non-activated esters as acyl donors in amide bond formation. The kinetic amidation of esters catalyzed by proteases capable of forming acyl-enzyme intermediates, for example serine and cysteine proteases, has been discussed earlier (Sect. 2.2). In this mechanism (see Fig. 6) the acyl-

enzyme intermediate formed by initial reaction of ester and enzyme can be intercepted by a nucleophile other than water, as in the normal hydrolysis reaction, to form an alternative product such as an amide, peptide, or a second ester. Even in aqueous media, amines can compete favorably to give high yields of amides. Many lipases and esterases also form acylenzyme intermediates and thus are able to catalyze the same aminolysis reaction of esters with arnines.

4.1 a-Amino Acids and Peptides The application of esterases and lipases to peptide synthesis has several inherent advantages compared to normal protease-catalyzed synthesis. For example, secondary product hydrolysis is minimal because the amide bond formed is not a natural substrate for these enzymes. Likewise, toleration of unnatural substrates is also greater. In addition, esterases and lipases are generally more active and stable than proteases in anhydrous solvents, although reactions in aqueous media are frequently slower. Early work of KLIBANOV'S group (MARGOLINand KLIBANOV,1987) showed that dipeptide formation between hydrophobic amino acid residues could be achieved by porcine pancreatic lipase (PPL)catalyzed reaction of N-acetyl protected acyl L-donor esters (41) with D- or L-amino acid amides (42) (Fig. 21). Other dry solvents such as THF, t-BuOH, MeCN, and i-Pr,O could be used. Similar couplings can be achieved in aqueous systems containing water-miscible solvents. In a study of the PPL-catalyzed synthesis of the dipeptide Z-Phe-Phe-NH, by the reaction of Z-Phe-OEt with Phe-NH,, optimum solvents at 50% aqueous concentration were DMF?DMSO, and MeOH whereas dioxan and acetonitrile were much poorer. Interestingly, MeOH (839'0, 3 h) was far superior to EtOH (48%, 24 h) under these conditions although reactivities of the corresponding starting esters were comparable. PPL was selective for L-acyl donors in the reaction; Z-D-Phe-OEt was not a substrate for either dipeptide formation or hydrolysis by the enzyme. However, the less sterically-demanding D-configuration donor

263

4 Esteruses and Lipases Which Can Be Used with Amides and Peptides

PPL, PhMe

. 41

Fig.21.

45"C, 2d

42 DLeuNHz

43 Ac-Phe-D-ku-NH2, 76% yield

I

See text (MARGOLIN and KLIBANOV, 1987).

Z-D-Ala-OCH,CH,Cl was accepted by PPL in coupling to Ala-NH2, although the reaction was much slower than with Z-L-AlaOCH,CH,Cl (KAWASHIRO et al., 1993). Where acyl donors possess reactive side chains as in Phe-Lys-O'Bu. the Pseudornonas sp. lipase-catalyzed acetylation with trifluoroethyl acetate is completely regiospecific for the E- and not the a-amino substituent in contrast to the normal protease-catalyzed mode of reaction. Other lipases acted in the same way (GARDOSSIet al., 1991). However, Candida antarctica lipase (CAL)-catalyzed amidation of Z-protected glutamic acid diesters showed the opposite regioselectivity, favoring a-amidation of the L-enantiomer but y-amidation of the D-enantiomer (CHAMORRO et al., 1995). These results indicate that lipases can show a high degree of stereoselectivity in amidations of amino acid esters. This is further demonstrated in the latter study by the use of rac-amethylbenzylamine (45) as acyl acceptor, where only the (R)-amine reacted (Fig. 22).

lipases and esterases catalyze the reaction frequently, they are used in supported form and can be recovered with little loss of activity. Enzymes most commonly used include Candida antarctica lipase (CAL), C. cylindracea lipase (CCL), and porcine pancreatic lipase (PPL). In a study of the aminolysis of ethyl octanoate (47) by ammonia, C. antarctica lipase SP435 was found to be the optimum catalyst from a range of twenty hydrolases tried (Fig. 23) (DE ZOETEet al., 1994).The same enzyme was also effective in catalyzing the reaction between methyl 3-(2-furyl)propionate (49) and a range of primary amines (Fig. 24). Suc-

4.2 Other Substrates A wide structural diversity of amines and esters undergo the lipase (or esterase)-catalyzed aminolysis reaction to form amides. Reactions can be carried out in nonpolar organic solvents which, in the case of volatile acyl donor esters such as ethyl or vinyl acetate, may actually be the donor ester. Nonactivated esters such as ethyl acetate give acceptable rates of reaction with most amine substrates, but hindered or poorly nucleophilic amines may require more activated acyl donors such as chloroethyl or trifluoroethyl esters. Many

4A sieves

CAL, ' P r p , 45°C

EtO2C

f

I

46 ( & - h i d e , 91%conversion

Fig. 22. See text (CHAMORRO et al., 1995).

264

S Cleavage and Formation of Amide Bonds

CAL

RNH2 'pr20 30°C

24"c 'BuOH 47

l N H 3 9 c A L 1

50

48 95Wyield

Fig. 23. See text (DEZOETEet al., 1994).

R

Time, h

Yield %

Ally1 "Bu

4

Bn

5

91 83 87

~

cessful reaction of anilines under these conditions, albeit at slower rates, is notable (GOTOR et al., 1993). A further interesting application to simple amines is the PPL-catalyzed formation of the macrocyclic bis-lactam (52)from diester (51) (Fig. 25) (GUTMAN et al., 1992). The stereoselectivity of the reaction has been investigated in both the amine and ester component. In aminolysis of a-substituted esters the degree of enantioselectivity is strongly dependent on the reacting amine, and prediction of structural effects can be difficult. With ammonia itself, high ee's can be obtained with esters of sterically-demanding acids. A good example is the C. anrarctica lipase (CAL SP435)-catalyzedammonolysis of racemic ibu-

Ph

~

77 ~~

profen 2-chloroethyl ester (53) which yielded unchanged (S)-ester (55) of 96% ee (Fig. 26). Similar ammonolysis of less bulky esters such as ethyl 2-chloropropionate and ethyl 2-hydroxypropionate gave only low to moderate ee's (DEZOETEet al., 1994). However, simple a-substituted esters frequently give higher enantioselectivities in the amidation reaction when the amine compo-

PPL, 4A sieves, CH2C12, reflux, 3-4d, 45% yield

Fig. 25. See text (GUTMAN et al., 1992).

38

Fig. 24. See text (GOTORet al., 1993).

H2N-(CH2) lo-NH2

51

4.5

*

52

4 Esterases and Lipases Which Can Be Used with Amides and Peptides

265

H

53

54 ( R ) - h i d e , ee not reported

55 ($-Ester, 96% ee

Fig. 26. See text (DEZOETEet al., 1994).

nent is larger. Here, the lipase from C. cylindruceu (CCL) showed consistent (S)-selectivity towards the ester with moderate to high ee’s for a range of primary amines (QUIROS et al., 1993). Reaction rates were strongly dependent on the electronic and steric effects of the a-substituent, with bromo and ethyl reacting notably slower than chloro (Tab. 5). Little correlation of amine structure with product ee was observed. C. antarcfica lipase (CAL) gave much faster amidation rates with the a-bromoester but was not enantioselective. However, the same enzyme was moderately enantioselective (40-78%) towards the corresponding a-ethyl substrate, yielding the expected ( R ) amides in all cases tried. CCL-catalyzed aminolysis of ethyl 2-chloropropionate by aromatic amines has also been reported (GOTORet al., 1988).Although the weaker nucleophilicity

of anilines compared to primary aliphatic amines required the use of more drastic conditions (60°C, 31-62 h), anilides were still obtained in moderate to good ee’s. Interestingly, aniline gave a much higher yield and ee of product anilide in tetrachloromethane (52% yield, 80% ee) than in hexane (26% yield, 56% ee). The opposite solvent effect was observed with n-butylamine in the same study, yielding amide of 95 % ee in hexane but only 40% ee in tetrachloromethane. Enzyme-catalyzed aminolysis of ethyl 2chloropropionate by racemic a-methyl substituted amines has also been studied. While the lipase (CCL)-catalyzed reaction showed the expected (S)-stereoselectivity towards the ester, use of a protease (subtilisin) favored the (S)-amine. In no case, though, did the enzyme show simultaneous enantioselectivity to both

Tab. 5. Lipase Catalyzed Amidation

Lipase, R’NH, hexane, RT

R R Cl

c1 c1

Br Br Et

R’

Lipase

octyl decyl dodecyl butyl decyl benzyl

CCL CCL CCL CCL CCL CAL

Time

PI

5 5 5.5 89 94 72

~

-;a,..’ R

Conversion

P I

23 35 20 24 38 25

I

H ee P o

70 92 51 90

64

78

I

5 Cleavage and Formation of Amide Bonds

266

56

5 7 ( R)-Ester

58 ( 3 - A m i d e , 85% ee

et al., 1993). Fig. 27. See text (GARCIA

components (BRIEVAet al., 1990). Efficient lipase (CAL)-catalyzed resolution of racemic a-substituted amines by amidation in ethyl acetate has also been reported (REETZand DREISBACH, 1994). In this case the enzyme was (@selective. A further feature of the lipase-catalyzed aminolysis reaction arises because the reactivity of the ester component is considerably increased by formation of an acyl-enzyme intermediate. The effect of this can be to enhance chemoselectivity towards the ester group in presence of other normally more reactive functionality. An example of such “reversed reactivity” has already been seen in the aminolysis of a-haloesters (Tab. 5). Here, aminolysis of the a-bromoester in the absence of enzyme gives nucleophilic displacement of the abromo substituent, and no amide formation occurs (QUIROS et al., 1993). Further examples are: (1) the lipase-catalyzed ammonolysis of cY,p-unsaturated esters where, instead of the

OMe

normal 1,Qaddition reaction, a$-unsaturated amides are frequently formed in high yields (SANCHEZ et al., 1994) and ( 2 )the chemoselective aminolysis of 3,4-epoxyesters (56) (Fig. 27) (GARCIA et al., 1993). The lipase-catalyzed aminolysis of cY,p-unsaturated esters by aliphatic amines, anilines, and hydrazines has similarly been achieved. With unsubstituted a$-acetylenic esters such as ethyl propiolate, reactivity is too high to prevent 1,4-addition of primary aliphatic amines. Anilines, however, form the corresponding acetylenic anilides when the reaction is catalyzed by CCL but, in the absence of enzyme, 1,4-addition is again preferred (PUERTAS et al., 1993). C. antarctica lipase-catalyzed reaction of ethyl propiolate with ammonia also yields the amide (SANCHEZ et al., 1994). Nacylhydrazines react similarly with methyl acrylate or vinyl crotonate to yield hydrazides rather than addition products when catalyzed by Amano PS lipase. Again, 1P-addition oc-

RNH2. CAL. dioxane 30°C. 24-4831

~

4 NH2

6 0 R = H, “Bu,allyl, Bn; 92-98% yield 0 -

5 9

RN ’H2 e

0 61

e 0

0

M

M

4 OMe

CAL, dioxane-

30”C, 24-48h

Fig. 28. See text (PUERTASet al., 1995).

M e O , p , k N & I 0

H

6 2 (R)-amide, R = Et, Pentyl, Ph; 2840% yield, 92-97% ee

5 Others

curs preferentially in the absence of enzyme (ASTORGA et al., 1991). Lipase-catalyzed aminolysis of succinic acid diesters has also been investigated. Reactions are frequently regioselective for one ester group only, leading to high yields of monoamides (60) (Fig. 28). In hexane as solvent, longer reaction times were necessary to achieve the same conversion rates and significant amounts of N-alkylsuccinimides were formed from the three alkylamines used. Presumably, these products arise via acyl-enzyme activation of the product ester group, followed by intramolecular “amidation” with the amide nitrogen. Racemic amines (Fig. 28) show consistent (R)-stereoselectivity and similar solvent effects, although cyclization to succinimides does not occur in this case as the amethylated monoamide products (62) are not accepted as nucleophiles by the CAL active site (PUERTAS et al., 1995). Similar CAL-catalyzed aminolysis of diethyl fumarate to yield trans-monoamides has also been reported (QUIROS et al., 1995). Interestingly, the corresponding cis-isomer, diethyl maleate, gives exclusively the same frunsmonoamides, most likely via prior-Michael/ retro-Michael isomerism of maleate to fumarate.

L-H ydantoinase

267

5 Others 5.1 Hydantoinases and Carbamoylases Hydrolytic cleavage of hydantoins by microorganisms has been known for some time. The enzymes which catalyze this hydrolysis are found in many organisms but only two, carboxymethylhydantoinase (EC 3.5.2.4) and allantoinase (EC 3.5.2.5), hydrolyze natural hydantoin substrates - most are involved in the processing of pyrimidine-derived cyclic ureas. Thus hydantoinase is named systematically as 5,6-dihydropyrimidine amidohydrolase (EC 3.5.2.2) and all these enzymes are placed in the subclass EC 3.5, cyclic amidases (Anonymous, 1984). Both L- and D-specific hydantoinases are known. In the whole cell environment they are frequently found in combination with enantiospecific carbamoylases which hydrolyze the initially formed a-ureidoacids to chiral a-amino acids. Sometimes a third enzyme, a hydantoin racemase, is also present in the system and thus complete processing of a racemic hydantoin (63) to a single amino acid enantiomer (66) is possible (Fig. 29). Even

*

Hydantoin racemase or chemical racemization

I

65 N -Carbamoyl-L-amino acid

I L-Carbamo ylase

1

66 L-Amino acid

Fig. 29. The conversion of racemic hydantoins to L-amino acids.

268


without a racemase, the facile base-catalyzed racemization of many 5-substituted hydantoins (typically at pH 8-10) means that complete turnover to a chiral amino acid may still be achievable by use of mildly basic conditions for the biotransformation. Hydantoins are readily accessible by a variety of synthetic methods, for example, by reaction of an aldehyde or ketone with KCN and (NH,),CO, (HENZEand SPEER,1942), or from a-amino acids by conversion to the urea by potassium cyanate, followed by acid-catalyzed cyclization (SUZUKIet al., 1973). Thus, wholecell hydantoinase systems have found application in industrial processes for the production of chiral amino acids. Since D-specific hydantoinase activity is present in many organisms, the process is particularly useful for the production of D-amino acids. For example, the organism Agrobacterium radiobacter, which contains both a D-hydantoinase and D-carbamoylase, has been used to prepare a wide range of bulk D-amino acids from racemic hydantoins (BOMMARIUS et al., 1992) Amino acids prepared by the method included D-Ala, D-Val, D-Met, D-His, D-naphthylalanine and D(3-pyridy1)alanine (all > 99% ee). As these examples demonstrate, hydantoinases generally possess a wide tolerance of the 5-substituent. Direct 5-phenyl substitution is usually accepted as in a recent study of two commercially available D-specific hydantoinases from thermophilic microorganisms (KEIL et al., 1995). Here, in the absence of a carbamoylase, chemical modification (diazotization) of the intermediate a-D-ureido acids was used to prepare several D- amino acids, including D-phenylglycine,in high ee. However, hydrolysis of 5,5-disubstituted hydantoins, although still highly enantioselective, was much slower (7% yield of D-ureido acid formed from 5-methyl-5-phenyl hydantoin after 72h). Resolution of a phosphonatebased hydantoin intermediate (67) (Fig. 30) in the synthesis of a D-configuration NMDA antagonist (69) by the Agrobacrerium enzyme also illustrates the relaxed specificity of these cyclic amidases (HAMILTON et al., 1993). The carbamoylase enzymes which are sometimes associated with hydantoinases of the same stereospecificity in microorganisms have high regiospecificity for the a-ureido sub-

stituent. An example is the synthesis of the unnatural o-ureidoamino acid, D-citrulline (72). Using the D-hydantoinasekarbamoylase system from a strain of A . radiobacter the hydantoin precursor (70,71) could be directly converted into D-citrulline (72) without cleavage of the side chain ureido group (Fig. 31) DRAUZet al., 1991).

1

0

67

D-H ydantoinase 7 0-8 0% yield

I

68

1. H N 0 2 2. 1 0 N HC1, 95°C

et al., 1993). Fig. 30. See text (HAMILTON

5 Others

269

transition state mimic (hapten) for the reaction, which subsequently can be used to raise antibodies in an animal species (GODING, 1986). However, while the antibody will probably only recognize part of an antigen molecule, small transition state haptens generally do not induce antibody formation and linking to an immunogenic protein such as bovine ser7 0 L-Citrulline hydantoin um albumin is required for the production of antibodies. More recently, in vitro gene library techniques have been developed for the generation of large numbers of different monoclonal antibodies for subsequent hapten affinity binding studies (HUSEet al., 1989). The design of haptens closely parallels the design of transition state enzyme inhibitors, a strategy widely used in medicinal chemistry for drug design. Indeed, catalytic antibodies (abzymes) raised against a particular hapten may also show strong similarities to enzymes capable of catalyzing the same reaction. A re71 D-Citrulline hydantoin cent example is provided by the three-dimensional X-ray structure of an active hydrolytic antibody with a bound phosphonate transition Agrobacterium state hapten (ZHOUet al., 1994). The antibody radiobacter (17E8) catalyzes the hydrolysis of nor-Leu and HZO, pH 8.4,40"C Met phenyl esters and is L-selective. Interestingly, the active site region contains a Ser-His dyad similar to the catalytic triad of serine proteases (Ser-His-Asp). Of further note is that the X-ray structure shows only the L-phosphonate bound to the active site whereas racemic hapten was used for the crystallization experiment. 72 DCitrulline Phosphonate-based haptens have frequently proved effective mimics for the tetrahedral Fig. 31. See text (DRAUZ et al., 1991). transition state intermediate in hydrolasecatalyzed processes. This is illustrated by the use of a phosphonate hapten (77) to generate antibodies for a peptide coupling reaction based on aminolysis of p-nitrophenyl esters 5.2 Catalytic Antibodies (HIRSCHMANN et al., 1994) (Fig. 32). The Antibodies are proteins which are generat- authors' premise that inclusion of a bulky ed by B lymphocytes of the immune system in cyclohexyl substituent in the hapten would response to foreign molecules (antigens). create a hydrophobic binding site in the antiMolecules capable of eliciting an antibody re- body capable of binding other similar-sized sponse vary enormously in size between, for alkyl groups proved correct, and isopropyl, example, a small molecule food toxin and a isobutyl, and benzyl were all accommodated bacterial or viral coat protein. An effective on the acyl donor. However, p-nitrobenzyl or strategy for utilizing antibodies in a catalytic p-chlorophenyl esters were not accepted, imsense to achieve a particular chemical reaction plying a specific antibody interaction with the relies on the synthesis of a small-molecule p-nitrophenyl group. Moreover, the product

270

5 Cleavage and Formation ofAmide Bonds

Antibody A

U

73 D- or L-TrpNH2

O

R

p H 7.0 buffer 5% DMSO

w

75 Proposed tetrahedral 74 D- or L-pnitrop-.enyl ester

intermediate

77 Phosphonamide hapten Fig.32.

See text (HIRSCHMANN et al., 1994).

dipeptides were not substrates for antibodycatalyzed hydrolysis, although both L- and Dreactants could be coupled. This study demonstrates that catalytic antibodies can be generated with very tight specificity for particular structural elements of a substrate but that more relaxed binding sites can also be incorporated in the same antibody. Although catalytic antibody-catalyzed amide bond formation and hydrolysis is still at an early stage compared to the widespread use of enzymes for these biotransformations, there is little doubt that the approach has significant potential for the design and development of

new biocatalysts. Taking peptides as an example, the synthetic potential of an antibody catalyst that is specific for peptide bond formation or cleavage between a particular pair (or perhaps involving a particular sequence) of amino acid residues would be enormous. Already the technique has been shown to be effective for amide bond hydrolysis of diverse substrates such as succinimides (LIOTTA et al., 1993) and primary amides (MARTIN et a]., 1994).This latter study is particularly interesting in that it demonstrates antibody-catalyzed hydrolysis of a nonactivated amide bond without cofactor assistance.

6 References

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SCHELLENBERGER, V., JAKUBKE, H. D., ZAPEVALOVA,TURK,J., PANSE,G. T., MARSHALL, G. R. (1975), StuN. P. (1991), Protease-catalyzed peptide synthesis dies with alpha-methyl amino acids. Resolution using inverse substrates: the influence of reaction and amino protection, J. Org. Chem. 40,953. conditions on trypsin acyl transfer efficiency, Bio- VANDAMME, E. J. (1988). Immobilized biocatalysts technol. Bioeng. 38,104. and antibiotic production: biochemical, genetical SCHMIDT-KASTNER. G., EGERER,P. (1984). Amino and biotechnical aspects. in: Bioreactors, Immobiacids and Peptides. in: Biotechnology, 1st Edn., lized Enzymes and Cells (Moo-YOUNG,M., Ed.), Vol. 8 (REHM,H.-J.. REED,G., Eds.). Weinheim: pp. 261-286, Amsterdam: Elsevier. VCH. VINCE,R., HUA,M. (1990), Synthesis and anti-HIV SCHWARZ. A.. STEINKE, D., KULA,M. A., WANDREY, activity of carbocyclic 2 ',3 '-dideoxy 2,6-disubstiC. (1990). Optimization of enzyme-mediated peptuted purine nucleosides,J. Med. Chem. 33.17-21. tide bond formation, Biotechnol. Appl. Biochem. WALDMANN. H.. SEBASTIAN, D. (1994). Enzymatic 12.188. protecting groups, Chem. Rev. 94.911-937. SEMENOV, A. N., CEROVSKY, V., TITOV, M. I.. MARTI- WALDMANN, H., HEUSER, A., REIDEL,A. (1994). SeNEK. K. (1988), Enzymes in organic synthesis: the lective enzymatic deprotection of amino groups in problem of increasing the yield of product, Colcarbohydrate and nucleoside synthesis, Svnlett,65. lect. Czech. Chem. Commun. 53,2963-2985, G. M. (1994), Use of hyWONG, c. H., WHITESIDES, SHADID, B., VAN DER PLAS,H. c.,B O E S T E N ,H. ~ .J., drolytic enzymes, in: Enzymes in Synthetic CheniKAMPHUIS, J., MEIJER,E. M., SCHOEMAKER, H. E. istry. pp. 41-59. Oxford: Pergamon Press. (1990), The synthesis of L( - ) and D( + ) lupinic WONG,C. H., CHEN,S. T., HENNEN, W. J., BIBBS.J. A., acid, Tetrahedron 46,913-920. WANG,Y. F., LIU,J. L. C., PANTOLIANO, M. W.. SHIN.C. G., SEKI.M., KAKUSHO, T., TAKAHASHI, N. WHITLOW. M., BRYAN,P. N. (1990). Enzymes in (1 993). Dehydrooligopeptides XV. Useful synorganic synthesis: Use of subtilisin and a highly thesis of dehydrodipeptides by the enzymatic stable mutant derived from multiple site-specific coupling of a-dehydroglutamate with various amutations. J. Am. Chem. SOC.112,945-953. amino acid amides using proteases. Bull. Chem. YAMANAKA. T., KAWAMOTO, T.. TANAKA, H., FUKUI, SOC.Jpn. 66,2048-2053. A. (1996), Enzymatic preparation of optically SOLOSHONOK. V. A.. KOKINA. N. A,, RYBAKOVA,A. V., active 3-trimethylsilylalanine, Appl. Microbiol. SHISHKINA. I. P., GALUSHKO. S. V.. SOROCHINSKY, Biotechnol. 4551-55. A. E.. KUKHAR,V. P. (1995). Biocatalytic ap- YOKOIGAWA, K.. SATO,E.,EsAKI,N., SODA,K. (1994). proach to enantiomerically pure p-amino acids, Enantioselective synthesis of N-acetyl-L-methioTetrahedron: Asymmetry 6.160 1- 1610. nine with aminoacylase in organic solvent, Appl. STEINKE. D.. KULA.M. R. (1990). Application of carMicrobiol. Biotechnol. 42,287-289. boxypeptidase C for peptide synthesis, Enzyme YUASA,Y.,WATANABE,T., NAGAKURA. A..TSURLTA, Microb. Technol. 12,836840. H. (1992). An improved synthesis of (S)-aspartylSUZUKI, T., IGARASHI. K., HASE.K., TUZIMURA, K. (7.7-dimethylnorborn-2R-yl)-(S)-alaninemethyl (1973). Optical rotatory dispersion and circular ester, a new high intensity artificial sweetener, dichroism of amino acid hydantoins, Agric. Biol. Tetrahedron 48,3473-3484. Chem. 37.41 1416. ZENONI, M., FUGANTI, C. (1994), Method for acylaSWEENEY, €? J., WALKER, J. M. (1993). Proteolytic ention of the 7-amino group of the cephalosporin zymes for peptide production, in: Methods in Moring, Eur. Parent 0582 102. lecular Biology 16 (BURRELL. M. M., Ed.), pp. ZHOU,Q., HUANG, Z. (1993). Immobilized thermoly277-303. Totowa, NJ: Humana Press Inc. sin and synthesis of precursor of aspartame, InTAYLOR, S. J. C., SUTHERLAND, A. G.. LEE. C., Wrsdian J. Chem. (Sect. B ) 32,35-39. DON, R., THOMAS, s., ROBERTS,S. M.. EVANS,C. ZHOU,G. W., Guo, J., HUANG,W., FLETTERICK, R. J.. (1990). Chemoenzymatic synthesis of ( - )-carboSCANLON,T. S. (1994), Crystal structure of a catavir using a whole cell catalyzed resolution of 2lytic antibody with a serine protease active site. azabicyclo[2.2.1Jhept-5-en-3-one, J. Chem. SOC., Science 265.1059-1064. Chem. Comrnicn. 1120-1121. ZMIJEWSKI. M. J., BRIGGS,B. S., THOMPSON. A. R.. TOGAME, H., INAOKA,T., KOKUBO. T. (1994). I n virro WRIGHT,I. G. (1991). Enantioselective acylation amidation for the preparation of an a-amidated of a beta-lactam intermediate in the synthesis of peptide: enzymatic coupling with prolyl endopeploracarbef using penicillin G amidase, Tetrahetidase. J. Chern. SOC..Chem. Commun., 1107-1108. dron Lett. 32,1621-1622.


6 Nitriles

ALAN WILLIAMBUNCH Canterbury, Kent, UK

1 Introduction 278 2 Chemical Properties of the Nitrile Group 278 2.1 Chemical Synthesis of Nitriles 278 2.2 Chemical Nitrile Transformations 278 3 Naturally Occurring Nitriles 279 3.1 Hydrogen Cyanide and P-Cyanoalanine 279 3.2 Cyanogenic Glycosides 280 4 Biotransformation of the Nitrile Group 282 4.1 Nitrile Hydratases 283 4.2 Nitrilases 295 4.3 Enzymes Capable of Biotransforming Cyanide 299 4.3.1 Fungal Cyanide Hydratases 299 4.3.2 Other Enzymes 300 4.4 Enzymes for the Synthesis and Transformation of Cyanohydrins 301 4.4.1 Oxynitrilases 301 5 Biotechnology of Nitrile Transformations 302 5.1 Use for Chemical Synthesis 302 5.1.1 Transformation of Mononitriles 302 5.1.2 Regiospecific Biotransformation of Dinitriles 308 5.1.3 Stereoselective Biotransformation of Nitriles 308 5.1.4 Commercial Processes 311 5.1.4.1 Biotransformation of Acrylonitrile to Acrylic Acid 311 5.2 Bioremediation of Nitrile Containing Wastes 312 6 Future Developments 313 6.1 Search for Novel Nitrile Biotransforming Activities 313 6.2 Redesign of Existing Enzymes by Protein Engineering 314 6.3 Metabolic Engineering for the Production of Multistep Biotransformations Involving Nitrile Substrates or Intermediates 314 6.4 Final Comments 314 7 References 315

278

6 Nitriles

1 Introduction Nitriles are important molecules in organic syntheses. There are several, often straightforward, routes for their synthesis by chemical methodology, and they can be converted into many other functional groups. Nitriles are also common products of biological systems,in particular plants, where their synthesis is often associated with defense of the producer. Biological catalysts capable of transforming nitriles are also readily found. Although the range of enzymatic activities so far discovered is relatively narrow, many reactions are regio- and stereoselective which make them useful tools for the synthetic organic chemist. Exploitation as biocatalysts is often limited because of structural instability or catalytic specificity. Nevertheless, commercial exploitation of nitrile biotransformations has already been achieved at a bulk-chemical production scale. With our rapidly increasing knowledge of enzyme structure it is possible to predict that a much wider exploitation of such enzymes will be possible in the near future. Protein engineering will be used to overcome the current limitations that thwart the exploitation of nitrile biotransforming enzymes.

2 Chemical Properties of the Nitrile Group In a review of this length it is not possible to cover all aspects of nitriles in detail, but some aspects of the chemical reactivity of nitriles are necessary to place the biological mechanisms used in nitrile biotransformations into context. Excellent summaries of the chemical properties of the nitrile group are given by AHMED and TRIEFF(1983) and by RAPPOPORT (1970). Hydrogen cyanide is the simplest molecule containing this functional group and is well known for its toxic properties, whereas other molecules may contain one or more nitrile groups. The characteristic group present in all nitriles is the carbon nitrogen triple bond; C=N (Fig. 1). This group is very polar and the

Fig. 1. The electronic structure of the nitrile group.

large dipole moment results in a high dielectric constant that explains the high solubility of simple nitriles in water.

2.1 Chemical Synthesis of Nitriles Nitriles can be synthesized chemically in a number of ways. For aliphatic nitriles the most common method is nucleophilic substitution of alkyl halides by sodium cyanide in DMSO at 140-150 "C.Aromatic nitriles can be made using diazonium chemistry, in which an aromatic amine is reacted with sodium nitrite (in aqueous HCI) and copper cyanide (0-5 "C). In plants and microorganisms nitriles are derived from amino acids (see Sect. 3).

2.2 Chemical Nitrile Transformations Nitriles can be chemically transformed in a number of ways. Under acidic or basic conditions the nitrile group can be hydrolyzed to the corresponding carboxylic acid, or partially hydrolyzed with hydrogen peroxide to the corresponding amide. Little selectivity is observed when more than one cyano group is present. Nitriles are reduced to primary amines by hydrogenation in the presence of Raney nickel catalyst. Ester formation is also possible by the reaction of nitriles with primary alcohols, and nitriles can be condensed with hydrogen sulfide, hydroxylamine, acetylene, butadiene, Grignard reagents, and alkenes. Finally, if the nitrile group is adjacent to a carbon bearing a hydrogen, deprotonation with a strong base

3 Naturally Occurring Nitriles

yields an anion which can be alkylated (e.g., with alkyl halides). In contrast, most biologically catalyted processes involve hydrolytic transformations to acids and amides, with only a few reports of reduction of the nitrile group. The chemical mechanism in most cases is addition to the nitrile group. This involves nucleophilic attack on the electron deficient carbon atom, and electrophilic attack on the electron rich nitrogen atom to give imines. Subsequent addition to imines gives the corresponding amines and their derivatives. Biologically catalyzed nitrile transformations seem to use the same mechanism.

279

transformation occurs has not been elucidated and it is still not known whether one or more enzymes are involved (MICHAELS et al., 1965; BRYSKet al., 1969;BUNCHand KNOWLES, 1982; CASTRIC,1981; WISSING,1975, 1983; WISSING and ANDERSEN, 1981). In the bacteria in which it has been studied the synthetic activity is associated with the membrane fraction of cells, although it can be solubilized with detergents. The activity is sensitive to oxygen and can be protected by using reducing agents such as dithiothreitol. An electron acceptor is needed for hydrogen cyanide production by cell-free extracts. Phenazine methosulfate, dichlorophenolindophenol, or ferricyanide can function in this respect (CASTRIC, 1981; BUNCHand KNOWLES, 1982; WISSINGand ANDERSON, 1981). The natural electron acceptor has not been identified, but does not appear to be NAD(P)+, FAD, or FMN. Several routes for the transformation of glycine into cyanide have been proposed (KNOWLES and BUNCH, 1986). There is no evidence, to date, that hy3.1 Hydrogen Cyanide and droxylation of glycine is the first step in the P-Cyanoalanine process. Fungi also use glycine as a precursor for hydrogen cyanide synthesis (BUNCHand Hydrogen cyanide is synthesized directly, or KNOWLES,1980). WARD et al. (1977) have indirectly by many microorganisms, plants, and again shown that hydroxyglycine does not apanimals. Direct synthesis only occurs in micro- pear to be an intermediate, neither is glyoxylic organisms. Bacterial cyanogenesis has been acid oxime. Earlier literature proposed that observed in only a few species. Chromobacteri- the cyanohydrins of glyoxylic acid and pyruvic um viofaceum is a prolific producer of hydro- acid were fungal intermediates for hydrogen gen cyanide and, in common with several spe- cyanide synthesis, but it is now thought that cies of pseudomonads that are cyanogenic, this is unlikely to be the case (WARD,1964; uses the amino acid glycine as the precursor WARDet al., 1971;STEVENS and STROBEL, 1968; (KNOWLES and BUNCH,1986). Using radiolab- TAPPER and MACDONALD, 1974; BUNCHand led glycine (1) it has been shown that the KNOWLES, 1980). methylene carbon atom and the amino nitroPhotosynthetic microorganisms synthesize gen are converted into hydrogen cyanide, with hydrogen cyanide via a different metabolic the carboxyl carbon lost as carbon dioxide route. Cell-free extracts of Chforeffavufgarzs (Fig. 2). The exact mechanism by which this produce cyanide in small amounts when illuminated in the presence of 02, Mn2 ,and peroxidase. Once again amino acids act as the direct precursor, the best being D-histidine. Glycine could not act as a substrate for this system. A similar process appears to be present in the New Zealand spinach plant (GEWITZet al., 1976a. b). PISTORIUSet al. (1977) showed that a D-amino oxidase and a 1 Glycine particulate component (the latter could be reFig. 2. The conversion of glycine into hydrogen placed with horseradish peroxidase or certain cyanide and carbon dioxide by microorganisms. redox metals such as manganese or bound


+

280

6 Nitriles

iron) could perform the complete conversion p-Cyanoalanine synthase has similar propof D-histidine into hydrogen cyanide. These erties in both cyanogenic plants and bacteria, workers also showed that aromatic amino ac- whereas the enzyme used by non-cyanogenic ids could act as substrates for this system. GE- bacteria can have different properties, and in WITZ et al. (1980) showed that imidazole 4some cases is more akin to a cysteine synthase aldehyde (and imidazole 4-carboxylic acid), or serine sulfhydrylase (KNOWLES and BUNCH, carbon dioxide, ammonia, water, and imida- 1986). zole acetic acid are also products when histidine is the substrate. It was proposed that an imino acid is an intermediate in the process (VENNESLAND et al., 1981). A similar mecha- 3.2 Cyanogenic Glycosides nism seems to be used in the blue-green bacterium Anacysfis nidulans, although the enzyme Many plant diseases where fungi are the involved appears to be an L-amino oxidase causative agent involve the liberation of cya(PISTORIUS and Voss, 1982). nide in plant tissues (VENNESLAND et al., C. vulgaris possesses a second system for 1982). Cyanide can be made de novo by the synthesizing hydrogen cyanide, in this case fungus (see above, Sect. 3.1) or by enzymatic from hydroxylamine and glyoxalate (SOLO- attack on cyanogenic glycosides present in the MONSON and VENNESLAND, 1972).The process plant (COTOTELO and WARD,1961).LEGRAS et is stimulated by Mn2+ and ADP (SOLOMON- al. (1990) have comprehensively reviewed the SON and SPEHAR, 1979). It is interesting that range of nitriles produced by living organisms, glyoxylic acid oxime could act as a substrate in and in particular those made by the 2000 plus a reaction stimulated by ADF'. At present the species of cyonogenic plants. metabolic pathway involved has not been The majority of plant nitriles are cyanogenic characterized. glycosides of which over 50 different types In cyanogenic bacteria another nitrile, p- have so far been identified. There are six subcyanoalanine (2) is often synthesized (Fig. 3 ) . groups of these compounds (3-8) which are The bacterium Chromobacterium violaceum based on their biosynthetic origin. Amino acsynthesizes this amino acid from either cys- ids are again used as precursors and include teine, serine, or 0-acetylserine) (BRYSKet al., valine, isoleucine, leucine, phenylalanine, and 1969; RODGERS, 1981,1982). There is evidence tyrosine. The last two groups include cyanothat the production of p-cyanoalanine (2) is genic glycosides with pentene structures (9, used by this organism to remove cyanide from 10) and those whose structures do not fall into its environment, and this nitrile can subse- the preceding five groups. Examples of these quently be converted to aspartic acid for use and nitriles synthesized by animals (11,U)are in the central metabolism (RODGERS,1981; given in Figs. 4 and 5, respectively. Plants also MACADAM and KNOWLES, 1984). The enzyme produce hydroxy nitriles that are esterified that catalyzes the synthesis of p-cyanoalanine with fatty acids. These metabolites have been (2) is found not only in cyanogenic bacteria classified into four groups based on the nature but also in cyanogenic plants (HENDRICKSONof the a-hydroxy nitrile, showing a resemand CONN,1969). In addition, many non-cyan- blance to the aglycones proacacipetalin (5,Fig. ogenic bacteria have this capability (DUNNILL 4) and cardiospermin synthesized from L-leuand FOWDEN, 1965; CASTRICand CONN,1971; cine (MIKOLAJCZAK, 1977). YANESE et al., 1982a, b, 1983). The metabolic pathways used by plants and animals to synthesize cyanogenic glycosides (18) are known in some detail (CONN,1979; WRAYet al., 1983).A generalized pathway for their production from amino acids (13) is shown in Fig. 6. Cell-free extracts can be used for the conversion of L-tyrosine into p-hydroxymandelonitrile and subsequently to the glycoside derivative (MACFARLANE et al.,


From isoleucine

H HO O

G

%

HO

H+ \O-oH CE N

3 Lotaustralin

From leucine

C 3N

HO

4 Sarmentosin

6OH

HO

5 (3-Proacacipetalin 6 ( R)-Epiproacacipetalin

HO

From phenylalanine 7

a R = H; Prunasin b R = P-D-glucopyranosyl; Amygdalin c R = a-L-arabinopyranosyl;Vicianin d R = P-D-xylopyranosyl; Lucumin e R = P-D-apio-D-furanosyl;Oxyanthin

From tyrosine HH

O

S HO

4

8 Triglochinin

CEN

Cyclopentene cyanogenic glycosides

s c q = = ou" J g H

9 Tetraphyllin

HO H HO

o*\'*

Ho&&=c+pH H

HO

HO

Fig. 4. Nitrile glycosides (LEGRAS et al., 1990).

'VH

10 Passitrifaciatin

281

282

6 Nitriles

11 Aerophysin-1 from the sponge Lanthella verongia

Br

B* H&

cyanide are compartmentalized in the black cherry, Prunus serotina (SWAINand POULTON, 1994; SWAINet al., 1992). A complicated series of enzyme catalyzed steps has been proposed that not only allows this plant to use cyanogenic glycosides as a defense mechanism, but also as a potential source of nitrogen during seedling development. Removal of the glucose residues from prunasin (7a) or amygdalin (7b) (see Fig. 4) by specific glucosidases (Fig. 7) (POULTON and LI, 1994) gives the hydroxynitrile (19).Cleavage to benzaldehyde (20) and hydrogen cyanide is catalyzed by the enzyme mandelonitrile lyase (Fig. 7). This last step is reversible in vitro.

OH

C EN

CEN

12 Benzoyl nitrile from myriapods

Fig. 5. Examples of nitriles isolated from animals.

1975). A mono-oxygenase is used in the hydroxylation of the nitrile intermediate (16) and a UDP-glucosyltransferase for the conversion of the hydroxynitrile (17) to the corresponding glycoside (18). It is noteworthy that the enzymes involved with this synthesis appear to form a channeled complex which allows limited access to intermediates added to cell-free preparations (CONN,1979). Such an arrangement of metabolic pathway enzymes is fairly common in plant biochemistry (HRAZDINA and JENSEN, 1992). The cyanogenic glycosides prunasin (7a) and amygdalin (7b)(see Fig. 4) and enzymes capable of their transformation to hydrogen

H

0

18

Although there has been a passing interest in nitrile metabolism for many years, more attention was focused on this area of biochemistry when it was found in the 1950s that nitrile derivatives of natural growth promoting substances could also affect plant development (CHAMBERLAIN and MACKENZIE,1981). At first a-hydroxylation of nitriles proposed, and detected, in plants and animals was thought to

+

0

O

14

13 Amino acid

R yCO G I Nl y c o s i d e

4 Biotransformation of the Nitrile Group

HzO + COZ

E N

C IN

17

16

Fig. 6. Biosynthetic pathway to cyanogenic glycosides in plants (CONN,1979).


283

HCN

~oG1ycoside

Glucosidase

7

0

Lyase ~

f, Y O

C= N

C rN

H

18 eg 7a or 7b

19

20

Fig. 7. General scheme for the enzyme catalyzed release of cyanide from plant cyanogenic glycosides.

be the main route of nitrile metabolism, in In addition, lyases have been isolated which both cases liberating hydrogen cyanide (FAW- catalyze the breakdown of cyanogenic glycosides, into hydrogen cyanide and the correC E T ~et al., 1958; OHKAWA et al., 1972). Subsequently, THIMANN and MAHADEVAN(1964) sponding carbonyl derivative. Before detailing found nitrile hydrolyzing activity in grasses, the biotransformation potential of these encabbage, radish, and members of the banana zymes their biological distribution and biofamily. In the same year HOOKand ROBINSON chemical properties will be described. (1964) isolated a “nitrilase” from a pseudomonas spp. that could hydrolyze the nitrile group of ricinine. Since this time several different types of nitrile biotransformations have 4.1 Nitrile Hydratases been described. To date most of the enzyme catalyzed nitrile These enzymes catalyze the general reaction transformations so far discovered are hy- shown in Fig. 8 and have been the most extendrolytic, there have been few reports of reduc- sively studied of the nitrile hydrolyzing entive processes. Nitrile hydrolyzing enzymes are zymes. Most of the microorganisms so far isodivided into two types. those that convert the lated that have this enzymatic activity belong nitrile group directly to the corresponding car- to the order Actinomycetales (Tab. 1). Howboxylic acid, nitrilase (EC 3.5.5.-) and those ever, many other bacteria have also been isowhich generate an amide, which then is con- lated including pseudomonas, bacilli, Microverted to the corresponding carboxylic acid by coccus spp., Bacteridium spp., an Agrobacterian amidase, nitrile hydratases (EC 4.2.1.-) um, and several fungi, identified as ascomyce(Fig. 8) It should be noted that this nomencla- tes. All these bacteria and fungi are microorture for nitrile hydrolyzing enzymes was only ganisms usually found in soils, where the posintroduced relatively recently (ASANOet al., session of hydrolytic enzymes enables them to 1980b), papers prior to this tended to use the utilize a wide range of naturally occurring term nitrilase for both types of enzyme. polymers and metabolites. The rhodococci in particular have an impressive metabolic diversity and are well suited to life in nutritionally Nitrile harsh environments (WARHURST and FEWSON, hydratase 1994). In plants, this type of enzyme seems to be rare and most reports focus on the conversion of P-cyanoalanine (2, Fig. 2) to its amide Amidase derivative asparagine, a process that has been R- C E N NH3 most extensively studied in the blue lupin (CASTRIC et al., 1972).A similar enzyme could 2H20 Nitrilase be present in cyanogenic millipedes, although there is a strong possibility that the enzyme responsible is actually located in a bacterium Fig. 8. Nitrile hydrolysis by hydratases and nitril- closely associated with these animals (DUFases. FEY, 1981).

284

6 Nitriles

Tab. 1 MicroorganismsPossessing Nitrile Hydratase Activity Microorganism

Reference

~~

Rhodococcus sp. Rhodococcus sp. N774 Rhodococcus butanica ATCC21197 Rhodococcus erythropolis JCM2892 and JCM6823 Rhodococcus rhodochrous J1 Rhodococcus sp. A3270 Rhodococcus sp. C311 Rhodococcus sp. 771

COHENet al. (1990) IKEHATA et al. (1989) KAKEYA et al. (1991) DURANet al. (1993) KOBAYASHI et al. (1989b) BLAKEY et al. (1995) LAYHet al. (1994) NOGUCHI et al. (1995)

Corynebacterium sp. C5 Corynebacterium nitrophilus Corynebacterium pseudodiphteriticum

TANIet al. (1989a) AMARANT et al. (1989) LI et al. (1992)

Brevibacterium sp. R312 Brevibacterium sp.

ARNAUD et al. (1976a) MOREAU et al. (1993)

Nocardidrhodochrous” LL100-21

DIGERONIMO and ANTOINE (1976)

Arthrobacter sp. J1 Arthrobacter sp. IPCB-3

ASANOet al. (1982a) RAMAKRISHNA and DESAI(1982)

Agrobacterium sp. Agrobacterium tumefaciens strain d3

MARTINKOVA et al. (1992) BAUERet al. (1994)

Pseudomonas chlororaphis 102 Pseudomonas chlororaphis B23 Pseudomonas Group I11 NCIB10477

WEIQUANG et al. (1989) NAGASAWA and YAMADA (1989) FIRMIN and GRAY(1976)

Pseudomonas sp.

ROBINSON and HOOK(1964)

Bacillus sp.

ARNAUD et al. (1976a,b, 1977)

Ascomycetes

VAN DER WALT et a!. (1993)

Myrothecium verrucaria

MAIER-GREINER et al. (1991)

a

See comments in text.

Rhodococci Of all the nitrile hydratase activities so far reported the enzymes in this type of bacterium have been the most extensively characterized, both with regard to their biochemical properties and the growth conditions needed for their optimal synthesis.As interest in the biotechnological applications of nitrile transforming enzymes grew it was inevitable that those ca-

CEN

(

21

Acrylonitrile

22 Acrylamide

Fig. 9. The hydrolysis of acrylonitrile to acrylamide.

285

4 Biorransformation of the Nitrile Group

pable of the partial hydrolysis of acrylonitrile (21), to the important commercial product acrylamide (22) (Fig. 9), would be among the first sought for. In a screen for acrylonitrile hydrolyzing activity in microorganisms WATANABE et al. (1987a) isolated many different types capable of this biotransformation. The screen included microbes from the environment and culture collections, assessing their ability to grow on acetonitrile as the sole source of nitrogen, and/or the presence of nitrile hydratase activity in cells grown on a standard partially defined medium. A total of 150 strains belonging to 30 different genera was examined from culture collections of which 8 gave rise to detectable levels of accumulated acrylamide in culture broths. The highest activity found was in Rhodococcus erythropolis IAM 1484. However, the estimated nitrile hydratase activity was too low for commercial usefulness. The contribution of amidase catalyzed removal of acrylamide in the enzyme assay was not assessed. About 1000 isolates from the environment were able to utilize acetonitrile. Six were selected for further investigation based on their growth rate/yield and nitrile hydratase activity. Rhodococcus sp. N-774 was the best acrylamide (22) producing isolate. The enzyme(s) present in the cell-free extracts of this bacterium could also hydrate a wide range of other nitriles. Given a reported activity of 46.9 pmol min-' mg protein-' with acrylonitrile (21) as the substrate for the enzyme, it is interesting to note that chloroacetonitrile, succinonitrile, and n-butyronitrile all gave higher activity at the concentration tested .The activity reported for benzonitrile and 3-cyanopyridine, the only aromatic compounds used in the study, were very low in comparison. Cyanoacetic acid did not appear to be able to function as a substrate. In the partially defined medium the nitrile hydratase specific activity increased in parallel with the increase in biomass concentration in batch cultures. On the cessation of growth the nitrile hydratase activity remained stable for many hours. It is possible to speculate that the rhodococci strains had the highest activity in these studies because of their characteristic, constitutive production of many catabolic enzymes. In many other bacteria such activities may only be expressed under certain nutrition-

al conditions. Even so, WATANABEet al. (1987b) have shown it is still possible to obtain better activities by optimizing growth conditions. Growth in a fully defined medium with ammonium sulfate as the sole source of nitrogen was very poor. The most marked improvement on growth was seen when complex nitrogen sources were added to the fully defined medium. For maximal nitrile hydratase activity in such cells the vitamin thiamine, and either an optimal level of Fez' or Fe3' need to be present. Other metal ions such as Co2 ,Niz , and Mn2+had little effect. In addition, an optimum temperature of 30 "C and adequate aeration is essential for maximum activity. In cell-free extracts the nitrile hydratase activity was maximal at 35 "C and at pH 7.7, although the pH profile was broad. At temperatures greater than 30 "C the preparations of the enzyme quickly lost activity. Cell-free extracts could be stored at 3°C at pH 7.5 for several days without loss of activity. In contrast, maximal accumulation of acrylamide occurred when the reaction temperature was decreased from 30°C to O"C, although, not surprisingly, the initial reaction velocity also declined. Improved productivity could be obtained by increasing the concentration of cells in the reaction mixtures with maximal yields of 30% (w/v) acrylamide being recorded. Immobilization of cells in acrylamide gels resulted in poor productivities in contrast to the non-immobilized systems. The authors implied that acrylonitrile (21) and acrylamide (22) (see Fig. 9) have deleterious effects on the nitrile hydratase, which could be reduced by operating the biotransformation at lower incubation temperatures. In the search for a nitrile hydratase that had good activity with aromatic nitriles Rhodococcus rhodochrous J1, was isolated and selected as being the most active (NAGASAWA et al., 1988b).It catalyzes the synthesis of the vitamin nicotinamide (24) from 3-cyanopyridine (23) (Fig. lO).This bacterium was isolated from soil samples using enrichment culture on a variety of aliphatic and aromatic nitriles. The biotransformation is one of the most dramatic examples of the capability in this area of biotechnology, where during the time course of the reaction (6 h) insoluble crystals of 3-cyanopyridine (23) disappear to be replaced by crystals of +

+

286

6 Nitriles

23 3-Cyanopyridine

24

Nicotinamide

Fig. 10. The hydrolysis of 3-cyanopyridine to nicotinarnide.

nicotinamide (24). The optimum pH for the whole cell biotransformation was between 7 and 9, a sharp decrease of activity was seen if the pH was much below this range. In contrast to the Rhodococcus sp. N-774 transformation of acrylonitrile (21,Fig. 9), the optimum temperature for this reaction by whole cells was 30 "C,although the enzyme was once again less stable at higher temperatures. This indicates that 3-cyanopyridine (23) and nicotinamide (24) are not causing the sort of inhibition proposed in the biotransformation of acrylonitrile. Up to 9 molar 3-cyanopyridine could be totally converted over 22 h to nicotinamide, and a maximum concentration of 12 molar nicotinamide could be achieved. No nicotinic acid could be detected in the product, even though amidase activity was present in the cells. The primary sequence of the Rhodococcus sp. N-774 nitrile hydratase has been determined by IKEHATA et al. (1989). Unpublished observations by WATANABE, referred to in this paper, reported the enzyme to be constructed of two subunits, M , 27000 and M , 27500. One unit of each is present in the intact enzyme complex. It was also stated that the enzyme contained pyrrolquinoline quinone (PQQ) and non-haem iron. Earlier and subsequent work indicated that the enzyme, in either the intact cell or cell-free extracts of Rhodococcus strains N-771 and N-774, had to be activated by near-UV light (NAKAJIMA et al., 1987; NAGAMUNE et al., 1990a, b). There was some evidence that the inactivation of the enzyme in the dark required oxygen and was temperature dependent. The enzyme in cell-free extracts could not be inactivated in the dark after photoactivation (NAGAMUNE et al., 1990a).

IKEHATA et al. (1989) purified the two subunits from the N-774 strain and the amino acid sequences were determined. Probes for the aand P-subunits were prepared and used to identify the region of chromosomal DNA coding for the structural genes. A 2070 bp piece of DNA was cloned into an Escherichia coli and shown to contain the two genes. No nitrile hydratase activity towards acrylonitrile (21, Fig. 9) could be observed unless the proteins produced were activated by treatment with urea, followed by dialysis. Even then the activity present was low, probably due to problems with folding of the protein during it's synthesis in the E. cofi,rather than a lack of cofactor such as PQQ. These workers also showed that a sequence of DNA, in addition to that which codes for the two structural genes, was needed for transcription or translation of the nitrile hydratase gene cluster. NAGASAWA and YAMADA (1989) proposed a mechanism for nitrile hydrolysis to the corresponding amide. The nitrile hydratase from Rhodococcus sp. N-774 was purified by ENDOand WATANABE (1989) and the one from Rhodococcus sp. N771 by NAGAMUNE et al. (1990a).The Rhodococcus sp. N-774 enzyme was reported as being very unstable, despite the use of a putative stabilizer n-butyrate, and, therefore, all procedures were performed at 0 4 "C.This probably accounts for most of the activity lost during the six steps, giving an 11.5-foldpurification to produce two bands, corresponding to the subunits of the enzyme on SDS-PAGE. Crystals of the nitrilase were generated from this preparation and the.subunits separated by HPLC on a C4 reverse phase column. The size of the two subunits a and P were 28500 Da and 29000 Da, respectively. N-terminal amino acid sequencing of the two subunits showed no homology with each other. The purified enzyme in solution was not inactivated by storage in the dark. In a subsequent paper NAGAMUNE et al. (1990a) purified the dark inactivated nitrile hydratase from Rhodococcus sp. N-771 using a similar procedure to ENDOand WATANABE (1989). They achieved a 17-fold purification, and better retention of activity. Once again two subunits were identified (a27500 Da and P 28000 Da). In studies on the stability of the active and inactive nitrile hydratase (the inactive enzyme was activated for analysis) it was


shown that the inactive enzyme was more stable during 800 h storage at 5 "C.It has now been shown that the photoactivation site is located on the a-subunit (TSUJIMURA et al., 1996). Both forms of the enzyme contained two atoms of iron. The optimum temperature of the active enzyme was 30°C and there was a clear pH optimum of 7.8, the enzyme having no activity at a pH less than 5 or greater than 10.5. The active enzyme preparations became unstable when stored above 35 "C,below pH 6, or greater than pH 7.8. The purified enzyme displayed the same substrate reactivities as reported earlier by WATANABE et al. (1987a).This range of activity is different from that exhibited by nitrile hydratases from other microorganisms, as was the observation that a narrower range of metals were capable of inactivating this enzyme. In common with nitrile hydratases inactivated by light was the observation that the UV-Visible absorption spectrum of the inactivated enzyme was very different from the active enzyme. There was little evidence of heme or flavin molecules being present in the purified preparations. In the paper by NAGAMUNE et al. (1991) there is a report referring to unpublished observations that indicates that the gene(s) coding for the nitrile hydratase of Rhodococcus sp. N-771 has been cloned and shown to be identical, in terms of the nucleotide sequence, to that found for the nitrile hydratase of Rhodococcus sp. N-774. These workers describe the production of crystals of the Rhodococcus sp. N-771 enzyme that should allow good resolution of structure by X-ray crystallography. R. rhodochrous J1 was originally investigated for its nitrilase activity (NAGASAWA et al., 1988b;MAITHEWet al., 1988;KOBAYASHI et al., 1989a). NAGASAWA et al. (1988c, d) demonstrated that a nitrile hydratase activity was present in this organism and that it was induced by cobalt. 2-Buteneamide (crotonamide) needed to be present in order for cobalt to have its effect. None of the other metals tested, including iron, had such an effect on nitrile hydratase induction or activity. The protein was purified and shown to account for more than 20% of the cellular protein. One or two bands were seen on polyacrylamide gels, relating to the nitrile hydratase activity. The purified enzyme catalyzed the hydration of benzonitrile (25) to

287

benzamide (26) (Fig. 11). It was shown that the molecular mass of the purified enzyme was about 530000 Da, made up of 10 a- and 10 Psubunits of 29000 Da and 26000 Da, respectively. The enzyme complex contained 5.7 atoms of cobalt per mole, which were tightly bound. Using a fed batch approach MAUGER et a). (1988, 1989) showed that whole cells of this Rhodococcus were able to biotransform a range of aromatic nitriles including benzonitrile (25),2,6-difluorobenzonitrile, thiophenecarbonitrile (27), and indoleacetonitrile (28) (Fig. 12) to the, corresponding amides. Up to 1 kg of product could be obtained with a 100% conversion of substrate in the reactors.

CEN

25 Benzonitrile

26

Benzarnide

Fig. 11. The hydrolysis of benzonitrile to benz-

amide.

H

27

Thiophenecarbonitrile

28 Indoleacetonitrile

Fig. U. Heterocyclic nitriles hydrolyzed by Rhodococcus rhodochrous J1.

During further studies on the factors needed for optimal nitrile hydratase activity in this bacterium NAGASAWA et al. (1991) showed that vitamin B12could not act as cobalt source for the synthesis of the enzyme and that a variety of aliphatic nitriles and amides (crotonamide was still the best) could induce the enzyme. Several carbon sources were evaluated for enhancing enzyme production, from which

288

6 Nitriles

glucose was selected. In contrast, changing the nitrogen source for growth had some dramatic effects. Some supplements substantially improved the biomass yield in the reactors, and enzyme activity. Of the media evaluated one gave a surprising result when supplemented with urea. Using yeast extract as the sole source of carbon and nitrogen for growth, but at a reduced level, urea greatly stimulated enzyme induction without affecting growth. It was suggested that crotonamide was a better inducer than other amides and nitriles tested because it was metabolized at a significantly slower rate. If this amide was fed at intervals during the incubation, a much higher activity of nitrile hydratase resulted. There was some evidence that nitrile hydratase activity could be related to the overall nitrogen metabolism of the bacterium. When ammonia was present lower activity resulted. Urea being only slowly converted to ammonia for growth was thus a much better source for cellular nitrogen. The improvement of activity obtained by careful growth medium design was truly amazing increasing 32 000-fold over the level obtained in the initial growth medium. Using the optimized growth medium NAGASAWA et al. (1991) proceeded to purify the cobalt containing nitrile hydratase from this bacterium. Although only 22% of the original activity was recovered after the purification procedure, the fact that the nitrile hydratase accounted for up to 30% of the total cell protein meant that crystallization of the enzyme was easily possible. These workers confirmed the mass properties of the intact enzyme complex and the two subunits, showing that they were not linked by disulfide bonds. A variety of organic acids in addition to n-butyric acid could stabilize the activity of preparations. n-Butyric acid also improved the temperature stability of the enzyme. The broad substrate specificity of the enzyme was also extensively investigated showing that although aliphatic nitriles were better substrates good transformation rates and yields could be obtained with aromatic nitriles. It was shown, using these enzyme preparations, that one cobalt atom is likely to be shared by each pair of subunits and the purified enzyme had a noticeably pink tinge. Since Na,S,O, was a potent inhibitor of the enzyme, it was proposed that the nitrile hydratase con-

tained Co3+.The enzyme was also strongly inhibited by HgClz and AgN03, but other reagents that potentially react with thiol groups did cause inhibition. Relatively low concentrations of hydrogen cyanide also inhibited the enzyme. N-terminal analysis of the subunit peptides showed some similarities with nitrile hydratases synthesized by other bacteria. In particular the P-subunit seemed to have closely related sequences. Nevertheless, there was quite a lot of variability in the a-subunit sequences, which may reflect a good deal of diversity when the full sequences are determined, a prospect supported by the lack of cross-reactivity between antisera prepared from each type nitrile hydratase. With reference to unpublished work, KoBAYASHIet al. (1991a) refer to the fact that R. rhodochrous J1 produced a lower molecular weight nitrile hydratase to the one just described. It was stated that these two enzymes, referred to as H-NHase and L-NHase, had different physicochemical properties and substrate specificities, although both contained cobalt. In addition H-NHase could be selectively induced with urea, while cyclohexanecarboxamide induced only L-NHase (KOBAYASHI et al., 1992a). Both forms were induced by crotonamide. Probes derived for isolating the Rhodococcus sp. N-774 nitrile hydratase were used to identify putative nitrile hydratase genes in R. rhodochrous J1. Two DNA fragments were identified using Southern hybridization. Analysis of the peptides for the HNHase, and predicted amino acid composition from the nucleotide sequence of the isolated chromosomal DNA fragments, were by and large in good agreement. A further experiment on the a-subunit of the L-NHase gave similar results. Although there was a good deal of variability in the N-terminal portion of the (a-subunit, the internal amino acid sequences were up to 79% identical. Active enzymes could be obtained in an E. coli host, but only if it was grown in the presence of added cobalt. An excellent account of the comparison between nitrile hydrolytic enzymes from a variety of bacteria is given by KOBAYASHI et al. (1992b).They highlight the fact that the R. rhodochrous J1 nitrile hydratase has the extra property of being much more resistant to inhibition by acrylamide than the Rhodococcus sp. N-774 en-


zyme. It is fascinating how such closely related structures, with regard to their amino acid sequence, can have such a variety of different properties, a clear target for closer examination to reveal potential protein engineering strategies for this and other enzymes. More recently, details have become available concerning the nature of the metal components of iron containing nitrile hydratases in inactive and photoactivated preparations. NAGAMUNE et al. (1992) and HONDA et al. (1992), using Mossbauer electron paramagnetic resonance and magnetic susceptibility techniques, have shown that one of the iron atoms in the inactive Rhodococcus sp. N-771 complex is oxidized during the photoactivation process. If the iron atom is oxidized it is likely that there will be a specific acceptor for the electron. HONDAet al. (1994) have shown that stabilizing agents, such as n-butyric acid, can affect the absorption spectra of nitrile hydratases. They suggest that in their system such a stabilizer is not necessary to preserve the stability of inactivated preparations. It was shown that the presence of n-butyric acid increases the efficiency of photoactivation. The data also indicated that tryptophan residues in the enzyme play an important role in photoactivation, possibly in the energy transfer facet of the process. It is currently uncertain where, if at all, PQQ is associated with this process. Intriguingly, NOGUCHI et al. (1995) using Fourier transform infra-red spectroscopy to investigate the difference between active and inactive nitrile hydratase, detected the possible presence of nitrous oxide in the enzyme complex, coordinated with the iron atoms. They showed that this was not due to contamination during the purification procedure. It was proposed that N O acts as the electron acceptor in the photoactivation process. It can thus be seen that many questions remain to be answered with regard to the mechanism by which nitrile hydratase catalyzes the hydrolysis of nitriles to amides, particularly the role that metals play. In addition, the biological organization of the genes involved with nitrile biotransformation in rhodococci, especially their relationship to areas of metabolism such as nitrogen assimilation, will be interesting to elucidate. HASHIMOTO et al. (1991) cloned a piece of DNA closely associated with the ni-

289

trile hydratase genes in Rhodococcus sp. N774, expressed it in an E. coli and showed it had amidase activity. In a later paper (HASHIMOTO et al., 1992) showed that a transfer of the nitrile hydratase and associated amidase genes to R. rhodochrous ATCC 12674 resulted in much better production of both enzymes, than when E. coli was used as a host. It has subsequently been found that up-stream and downstream elements of DNA sequences containing the amidase and nitrilase genes, are needed for efficient production of the two enzymes. It was proposed that the down-stream sequence could be involved with the incorporation of iron (or perhaps PQQ) into the nitrile hydratase. This down-stream sequence had components that are present in proteins requiring ATP for their function. Whether such a requirement is needed by the protein(s) coded for by this piece of DNA remains to be clarified. The new constructs had a 6-fold higher level of both amidase and nitrile hydratase, when compared to Rhodococcus sp. N-774 (HASHIMOTO et al., 1994). In comparison with the rhodococci mentioned above relatively little information is available on the nitrile hydratase activity present in other species. However, the few reports that have been made indicate similarities and additional biotransformation potential of this group of bacteria. In a screen for microorganisms capable of growth on 2-cyanoethanol and benzonitrile, as sole sources of carbon, KAKEYAet al. (1991) selected Rhodococcus burunica ATCC 21197 as the best strain. ECaprolactam (29) (Fig. 13) was essential for the induction of the enzymatic activities observed. The bacterium was able to biotransform a wide range of substituted benzonitriles, usually with good yields, to the corresponding acids. Of particular note was the observation that orrho-substituted derivatives, previously apparently resistant to enzymatic hydrolysis, were transformed predominantly to amides. n

Fig. 13. cCaprolactam, an important inducer of nitrilase and nitrile hydratase activity.

29 E-caprolactam

290

6 Nitriles

Some regioselective nitrile hydrolysis was observed when isophtalonitrile was the substrate. The authors proposed that it was steric hindrance, rather than the electronic effects of substituents, that determined the efficiency of the biotransformation. Using a-arylpropionitriles they showed that this bacterium was capable of stereoselective transformation. There was some evidence that both nitrile hydratase and nitrilase activities were present, giving rise to the synthesis of (R)-amides and @)-acids. Given the many similarities between the DNA sequences coding for nitrile hydratases in not only rhodococci but also other microorganisms, plus the number of inducers that can be needed for good enzyme induction, a screen using DNA probes should be more efficient than traditional methods for detecting other nitrile hydratase producers. DURAN et al. (1993) used this approach to screen 31 microorganisms, representing many different types from a variety of environments, using the nitrile hydratase genes from Rhodococcus sp. N774 as a probe. Chromosomal sequences showing positive hybridization were obtained with the two R. erythropolis strains. It was shown that once again E-caprolactam (29, Fig. 13) was required for the induction of the nitrile hydratase. It was interesting to note that the substrate specificity of the new enzyme was much broader than the Rhodococcus sp. N-774 nitrile hydratase, being able to use aromatic as well as aliphatic nitriles The amidase activities were also a good deal higher than present in the bacterium from which the probe was made, showing that this method has the potential to detect a variety of nitrile hydratases. Nevertheless, it is important to note that traditional screening methods are also capable of

nc-30

31

yielding interesting finds. BLAKEY et al. (1995) have recently isolated Rhodococcus sp. AJ270 from a river bank associated with former industrial activity. This bacterium is capable of biotransforming, sometimes with regio- and stereospecificity, a wide range of aliphatic mono- and dinitriles, aromatic mono- and dinitriles as well as heterocyclicaromatic nitriles. No inducer was mentioned for activity to be observed. Reference to unpublished data indicates that no nitrilase is present. It will be interesting to see if the nature of the nitrile hydratase of this isolate is similar to the ones already described. If it is, it will be intriguing to know what structural differences in the closely related enzymes determine their substrate specificity.This would again be an ideal model system for the application of protein engineering. Corynebacteria The first report of a nitrile biotransformation by a Corynebacterium was made by MrM U M et al. (1969).The Corynebacteriumnitrilophilus species could metabolize mono- and dinitriles MARTINKOVA et al. (1992) more recently found a nitrile hydratase activity in a similar bacterium. Again using acetonitrile in et al. (1971) isoenrichment culture, FUKUDA lated a Corynebacterium capable of growing on a variety of nitriles, including racemic aminopropionitrile (Ma) (Fig. 14) which could be totally converted to approximately equal amounts of L- and D-alanine (32a and 33a). Whole cells of the isolate were also able to convert L-alanine (32a) to D-alanine (33a), presumably via a racemase enzyme. When racemic amino isovaleronitrile (3Ob) was used as a substrate L-valineamide (31b) was the main

32

a R = Me; b R = CHMe2; c R = H

Fig. 14. Corynebacterium hydrolysis of amino nitriles

33

4

Biotransformation of the Nitrile Group

291

product plus some racemic valine. No race- perature stable than many other nitrile hydramase activity was shown to be present, that tases at 40 "C, though it still lost much of its accould interconvert L- and D- valine (32b and tivity. The best substrate tested for the nitrile 33b). Therefore this bacterium apparently had hydratase was n-valeronitrile. It had poor activity with the aromatic nitriles tested. Nickel, both nitrile hydratase and nitrilase activity. TANIet al. (1989a, b) used Corynebacterium mercury, and copper ions inhibited the ensp. C5 to produce trans-4-cyanocyclohexane-l- zyme. The addition of iron had little effect carboxylic acid (35) from the corresponding even though some was lost during the purificadinitrile, frans-1,4- dicyanocyclohexane (34) tion procedure. Using reported protocols for (tDCC) (Fig. 15). They showed by separating determining the presence of PQQ in enzymes the two activities that the acid (35) was pro- it was shown that this nitrile hydratase also duced by a nitrile hydratase and an amidase. contained this cofactor. The final nitrile hydraThe activity could be enhanced, by about 50%, tase preparations were 130-fold purer, comby supplementing the growth medium with pared to 220-fold for the amidase. Surprisingly, iron (mainly effecting the nitrile hydratase) the best substrate for the amidase was not nbut not a variety of other metals. Cobalt addi- valeramide but the amide of the mononitrile tion actually lowered the activity present in intermediate (35) in the conversion of tDCC cells by 50%. The nitrile hydratase activity was (34)to the dicarboxylic acid product (36).The quickly lost from stationary phase cells grown amidase had a temperature optimum of in batch culture, although changing the nitro- around 50°C and had a broad pH optimum. gen source from ammonia to peptone could in- This enzyme was much less sensitive to thiol crease the stability of both the nitrile hydra- group inhibitors than amidases from other nitase and the amidase in stationary phase cells. trile hydratase containing bacteria. LI et al. (1992) have shown in CorynebacferiThe substrate for this biotransformation, tDCC (M), could greatly increase the activity um pseudodiphteriticum that methacrylamide of both enzymes if present as the sole source of was a good inducer of nitrile hydratase activnitrogen in batch culture. If this dinitrile (34) ity. The enzyme appeared to loose activity in or a variety of nitriles and amides were added culture at temperatures above 27 "C. Using to media containing peptone as the main nitro- purified preparations of the enzyme they suggen source for growth, few were able to in- gested that it had a molecular weight of 80000 crease the activity of the nitrile hydratase and Da with three subunits having two different amidase. The addition of dinitriles, including molecular weights of 25 000 Da and 28 000 Da. tDCC, lowered the activity of these enzymes. The temperature optimum of the nitrile hydraThere was evidence that in resting cells the nitase was 25"C, with a p H optimum of 7.5. It trile hydratase activity was unstable. This could was strongly inhibited by silver and mercury be overcome by the addition of organic acids ions, plus phenyl mercuric acetate. Iodoaceto preparations during purification. Isovaler- tate, copper ions, and EDTA also caused inhiate and caprylate were more effective than n- bition. There is therefore evidence of some diverbutyrate. The nitrile hydratase was shown to have a molecular weight of 61 400 Da made up sity in Corynebacterium enzymes, as well as of two subunits, each estimated at 26900 Da. It similarities in structure compared to those enhad a pH optimum of 8-8.5 and was more tem- zymes present in other bacteria.

6 C IN

C EN

--0 $"

Corynebacterium sp. c5 sp.c5

C IN

-6"

Fig. 15. Corynebacterium sp. C5 hydrolysis of cyclohexane dinitriles.

--

?02H 32H

292

6 Nitriles

ent order of K , and V,,, values was obtained. Brevibacteria The hydrolysis of nitriles via amides to acids This paper confirmed that the amidase is an inwas observed in Brevibacterium sp. R312, ducible enzyme, in contrast to the nitrile hywhich remains the best studied of this type of dratase, whose activity can be modulated by bacterium (ARNAUD et al., 1976a, b; JALLA- the presence of amides. It was uncertain whether the organic acid products of the GEAS et al., 1978a). Mutants of this microbe have been made which have either the nitrile amidase reaction or certain amides could hydratase or the amidase genes disrupted. The repress the biosynthesis of the amidase et al., 1984a). Using the mutant acetonitrile hydrolyzing activity was complete- (MAESTRACCI, et al. ly inhibited by n-bromosuccinamide and diiso- strain Brevibacterium sp. 19, MAESTRACCI propylfluorophosphate. It lost activity rapidly (1984b) showed that acrylonitrile (21,Fig. 9) above 40°C (ARNAUD et al., 1977). The ami- and three other nitriles tested, were competidase was in contrast a very heat stable enzyme tive inhibitors of the amidase enzyme. Acryhaving a temperature optimum between 60 lonitrile (21) was shown to inhibit the activity and 70°C (JALLAGEAS et al., 1978b).A mutant of the nitrile hydratase of Brevibacterium sp. lacking much of the activity of the wild type 312 at high substrate concentrations (0.2 M or strain (loss of aliphatic amide hydrolyzing cap- greater). Using an amidase negative mutant it ability) was still able to hydrolyze L-a-amino was shown that amides could reduce the bioamides (31,Fig. 14) (JALLAGEAS et al., 1979a,b; synthesis of the nitrile hydratase. Cyanide was ARNAUD et al., 1976c, 1980; KIENY-L'HOMMEa more potent inhibitor of this enzyme (Buret et al., 1981). A combination of gas chromato- al., 1984b). graphy and proton NMR was used to follow MILLERand KNOWLES (1984) showed that enzymatic conversion of a wide variety of ni- both the nitrile hydratase and amidase of triles and amides (JALLGEAS et al., 1979b; BUI Brevibacterium sp. 312 are intracellular enet al., 1984~). The biotechnological potential of zymes not closely associated with the memthese enzymes was apparent (BUIet al., 1982). brane, and thus both the nitriles and amides BUI et al. (1984a) isolated a nitrile hydratase must be freely transported into the cells during defective mutant of Brevibacterium sp. 312. growth. They used an interesting selection procedure FRADETet al. (1985) partially purified the where loss of capability to biotransform chlo- nitrile hydratase from a amidase negative muroacetonitrile to the potentially toxic product tant of Brevibacterium sp. 312, with the princichloroacetic acid was used to indicate enzyme pal objective of removing intracellular prodeficient mutants. Brevibacterium strain 19 teases. The preparation was immobilized onto was isolated which had lost the expected range a DEAE-cellulose support and resulted in an of nitrile hydratase activities when compared active system with a slightly lower pH optito the wild type.The mutation was stable for at mum and slightly higher temperature optileast 50 generations. Analysis of the enzymes mum than observed with the free enzyme. revealed that the nitrile hydratase of the deriv- About 40% of the original activity was lost on ative strain had a wide range of K , and V,,, immobilization. Continuous production and values for different substrates. Cyanide was a excellent conversion of propionitrile to the et al. good substrate for the enzyme. As the chain amide was observed at 5°C. TOURNEIX length of the aliphatic nitriles tested increased, (1986) showed that N-methylacetamide and the K , decreased. The enzyme worked well N,N '-dimethylacetamide repressed nitrile hywith a-hydroxy or amino nitriles (30,Fig. 14) in dratase production. N-methylacetamide is an which cases the best V,,, values were also ob- inducer but not a substrate for the amidase in served. Benzonitrile (25,Fig. 11) was also a this bacterium. When the nitrile hydratase was good substrate. On the basis of this and previ- purified to homogeneity it was shown to have ous work the authors proposed a catalytic two kinds of subunits of 27000 and 27500 Da mechanism involving amino and sulfhydryl which formed an aggregate of 90000 Da. PQQ groups plus potentially a hydroxyl group at the was detected and the enzyme contained Fe3'. active site. The amidase activity of Brevibacte- Others have shown that the iron is coordinatrium sp. 312 was also fairly broad, but a differed to sulfur and nitrogen or oxygen (NELSON


293

et al., 1991),and that an iron coordinated water molecule has a role in nitrile hydration (BRENNAN et al., 1994a; DOANet al., 1994). In addition, the presence of PQQ in the enzyme has been questioned, the spectral properties of preparations being due to changes in the coordinated iron (BRENNAN et al., 1994b). The Nterminal sequence of 20-25 amino acids of the subunits was identical to that reported by ENDOand WATANABE (1989) for Rhodococcus sp. N-774 (NAGASAWA et al., 1986; DURANet al., 1992a). During studies on the genetic arrangement of the two enzymes MAYAUX et al. (1990) have gained evidence that the enantioselective amidase present in Brevibacterium sp. 312 is genetically coupled to the nitrile hydratase. Promoter sequences that could be recognized by Brevibacterium sp. A4 and E. coli RNA polymerase were isolated from Brevibacterium sp. R312 (DURAN et al., 1992b; AZZAet al., 1994). An adiponitrile transforming strain, Brevibacterium sp.ACV2, is a mutant of Brevibacterium sp. R312. Although it apparently has the same macro structure of the wild type it has different properties with regards to both substrate specificity and its response to pH (MOREAU et al., 1993,1994b).

trile (25), benzamide (26, Fig. 11) or benzoic acid. When grown on benzonitrile (25) such cells showed little oxygen uptake when acetonitrile, acetamide, or acetate were added to them. It was also shown that the acetamidase had a broad pH optimum, while the “acetonitrilase” had a sharp optimum at pH 7. Putative metal chelators inhibited the activity of the “acetonitrilase”. The benzonitrilase had a pH optimum of 7.0-7.4 and was present in benzamide grown cells, as was benzamidase activity. Difficulties were encountered getting active preparations of these two enzymes (COLLINSand KNOWLES, 1983). LINTONand KNOWLES (1986) showed that several amides and nitriles could only act as nitrogen sources for growth. These included acrylonitrile (21) and acrylamide (22)(see Fig. 9) plus the dinitriles succinonitrile and glutaronitrilet. It was shown that the “acetonitrilase” and acetamidase induced by growth on acetonitrile and acetamide could use a wide range of substrates. Amidase activities present in cells grown on a variety of other amides showed a similar range of activities. The ratio of “acetonitrilase” to acetamidase activities was similar following growth on a range of nitriles and amides.

Nocardia Several species of Nocardia have been reclassified as rhodococci, but as there is some evidence that the enzyme profiles of the best studied strains are different from other species that have been investigated, and because the literature until quite recently uses some of the old terminology, they will be considered as a separate group (GOODFELLOW et al., 1990). These bacteria have been shown to have both nitrilases and nitrile hydratase activities. Nocardia rhodochrous LL 100-21 is able to grow on a wide range of amides and nitriles, sometimes as both sole carbon and nitrogen sources. Both aliphatic and aromatic nitriles and amides can be utilized (DIGERONIMO and ANTOINE, 1976; COLLINSand KNOWLES, 1983; LINTONand KNOWLES, 1986; VAUGHAN et al., 1988). Acetonitrile was converted via the amide to acetic acid, while benzonitrile (25, Fig. 11) was apparently converted directly to benzoic acid. The acetonitrile grown cells had little capacity for the metabolism of benzoni-

Arthrobacter This is the last of the bacterial groups belonging to the order Actinomycetales to be described in detail. ASANOet al. (1980a) resolved the nitrile hydrolyzing activity of Arthrobacter sp. J1 (YAMADA et al., 1979) into a nitrile hydratase and an amidase. The activity of the nitrile hydratase decreased rapidly in the early growth phase in batch cultures growing on acetonitrile (ASANOet al., 1982b). A 290-fold purification of the enzyme gave a native enzyme with an estimated molecular weight of 420000 Da constructed from two types of subunits with molecular weights of 24000 Da and 27000 Da. The enzyme had a pH optimum of 7.1 and was inactivated by incubation at 55 “C. Silver nitrate, mercuric chloride, and cyanide were able to completely inhibit the activity of the enzyme, whereas p-chloromercuribenzoate and iodoacetate caused partial inhibition at the concentrations tested. The effect of the last two, and of mercuric salts, could be abolished in the presence of 2-mercaptoetha-

294

6 Nitriles

nol. The addition of metal ions, including iron (cobalt was not tested), had little effect. Activity was observed with several aliphatic nitriles, including acrylonitrile (21,Fig. 9). In another isolate RAMAKRISHNA and DESAI(1982) suggest that some Arthrobacter strains can make two types of nitrile hydratase, however the evidence for this was only preliminary. Pseudomonads Glutaronitrile, a dinitrile, was shown to be degraded by a Pseudornonas (YAMADAet al., 1980). Growth on glutaric acid was inhibited by the presence of a wide range of nitriles, but not glutaronitrile. Analysis of the culture filtrates showed that the metabolic route involved in the conversion of the dinitrile proceeded via amide and acid intermediates. In a subsequent paper, using a different Pseudomonas isolate, YANASEet al. (1985) purified a nitrile hydratase active against n-butylyonitrile but not with the “natural” nitrile, P-cyanoalanine (2,Fig. 3). The nitrile hydratase of Pseudomonas chlororaphis B23, a bacterium with very high acrylonitrile (21)to acrylamide (22)(see Fig. 9) activity (see also WEIQIANG et al.. 1989), was purified to homogeneity after induction of the enzyme with methacrylamide (NAGASAWA et al., 1987). n-Butyric acid was used to stabilize the enzyme which rapidly lost activity above 35°C. The enzyme showed a broad substrate specificity towards aliphatic nitriles. Of especial interest was the observation that isobutyronitrile was a strong inhibitor of the enzyme but it was not detectably biotransformed.The native enzyme had a molecular weight of 100000 Da, and was constructed from four identical subunits of 25 000 Da. Four iron atoms were associated with the enzyme and could not be removed by dialysis. Apparently no acid labile sulfur was present in the enzyme. The authors showed that there were significant differences in the response of the enzyme to a variety of inhibitors when compared to the Arthrobacter sp. J1 enzyme, and that antigenically it was unrelated. Using inhibitors that react with carbonyl groups, and which strongly inhibit the nitrile hydratase, SuGIURA et al. (1988) claimed that these results show an interaction between the low-spin et al. (1991) have Fe3+ and PQQ. NISHIYAMA cloned the DNA involved with the production

of the nitrile hydratase subunits, and the amidase, into E. coli. Active nitrile hydratase and amidase were produced by the recombinant host, unlike with Rhodococcus sp. N-774, but only if the entire operon was cloned. A 38000 Da protein appeared to be involved in the production of active enzymes. Although there is significant homology between the nucleotide sequences coding for the nitrile hydratase and amidases in Rhodococcus sp. N-774 and I? chlororaphis B23, the sequences have many differences.

Agro bacteria During a screen for microorganisms capable of detoxifying low-level nitrile wastes O’GRADY and PEMBROKE (1994) recently isolated an Agrobacteriurn which was capable of biotransforming a range of mono- and dinitriles. Particularly noteworthy is the ability to hydrolyze indole-3-acetonitrile (28, Fig. 12) to the plant auxin indole-acetic acid. Using acetonitrile as the sole source of carbon and nitrogen for growth, a two-step hydrolysis of the nitrile via acetamide was implied. Both nitrile hydratase and amidase activities were detected, the activity of the latter enzyme being 170 times greater. Neither enzyme appeared to be produced during growth in the absence of nitriles. The involvement of microorganisms in the production of plant hormones has been documented (MORRIS,1986), and seems to be essential for the virulence of bacteria in their host plants. In a return to earlier observations KOBAYASHI et al. (1995) have studied in detail the enzymes involved with the synthesis of indoleacetic acid from the natural plant nitrile indole-3-acetonitrile (28,Fig. 12) in the closely related plant pathogen Agrobacteriurn and plant symbiont Rhizobium. Interestingly, they found that crotonamide, a good inducer of nitrile hydratases in some bacteria, inhibited production in many agrobacteria. Similar experiments with rhizobia isolates showed that crotonamide addition to growth media had little effect on the activity of either the nitrile hydratase or the amidase. Selecting the Agrobacterium with highest nitrile hydratase activity, and e-caprolactam (29,Fig. 13) as the inducer, the enzyme was purified to homogeneity. It was found to have a molecular weight of 102000 Da, made up of 4 identical subunits,


and thus resembled the enzyme from Pseudomonas chlororaphis B23. Strangely, the enzyme appears to contain both iron and cobalt. It had a very low K , of 7.9 pM. Fungi Given that nitrile hydratase activity has been implicated in fungi, and the widespread biotechnological use of these microbes, it is surprising that few detailed studies have been made. KUWAHARA et al. (1980) showed that a Fusarium solani isolate was able to use monoand dinitriles as a sole source of nitrogen for growth, some could be used as both the sole carbon and nitrogen source. When succinonitrile was used as a substrate, cyanopropionamide and succinamide accumulated in the culture medium. Amides did not accumulate with the other nitriles tested. Therefore, there is a possibility that a nitrile hydratase and a nitrilase were present in the cells. In the early 1970s there was a report that cyanamide (37),an agricultural fertilizer, could be converted into urea (38) by Myrothecium verrucaria (Fig. 16). A nitrile hydratase has subsequently been isolated from this fungus (MAIER-GREINER et al., 1991). Growth on cyanamide (37),or chemically related compounds, is necessary for the synthesis of nitrile hydratase. The purified enzyme had a molecular weight of 170000 Da and is made up of six subunits of about 27000 Da. A temperature of 50 "C and a pH of 7.7 gave the best activity.The enzyme is very specific for cyanamide (37). Studies with inhibitors indicate that a cysteine residue is important for activity and that metal chelators also reduce the enzyme activity. The nitrile hydratase could be cloned into E. coli where active enzyme was synthesized. There was little sequence similarity with other microbial nitrile hydratases. Finally, there is some indication that nitrile hydratases may be present in a variety of ascomycetous yeasts, but as yet no attempt has

Fig. 16. The conversion of cyanamide into urea by Myrothecium verrucaria.

295

been made to isolate the enzyme from these microorganisms (VANDER WALTet al., 1993).

4.2 Nitrilases Although the distribution of nitrilases appears to be wider than that of nitrile hydratases, they have been studied in less detail. This is probably due to two major reasons. Firstly, early work on the exploitation of nitrile hydrolyzing enzymes concentrated on the production of acrylamide, nitrilases convert nitriles directly to the carboxylic acid without the release of an amide as intermediate (see Fig. 8). Secondly, problems have been encountered with the purification of these enzymes. Nevertheless, nitrilases can have useful biotechnological potential, and they may be more important in nitrile metabolism in nature than the nitrile hydratases (KOBAYASHI and SHIMIZU, 1994). Rhodococci The first report of nitrilase activity in these bacteria was made by HARPER(1977a) who observed benzonitrile metabolism in Rhodococcus rhodochrous NCIMB 11216 (formerly Nocardia rhodochrous). The purified enzyme showed time dependent activation in the presence of benzonitrile that was a function of enzyme concentration, temperature, and pH. The process involved the aggregation of 47,000 Da subunits into a 12-subunit aggregate, which had optimal activity at pH 8.0. Preparations of the enzyme showed low stability at 0 ° C with total loss of activity on freezing. A variety of chemicals that react with thiol groups abolished the activity of the enzyme (HARPER, 1977b). The nitrilase was specific for nitrile groups directly attached to a benzene ring. Orrho-substituted benzonitriles, except when fluorine was used, could not act as substrates. Meta- and para-substituted benzonitriles could be hydrolyzed. Benzamide (26, Fig. 11) could not

37 Cyanamide

38 Urea

296

6 Nitriles

be used for growth by this bacterium, neither that aliphatic nitriles, in particular isovaleronidid the purified nitrilase produce benzamide as trile and isobutyronitrile, were better inducers an intermediate (HARPER,1977b).It is interest- of the enzyme than benzonitrile (25,Fig. 11). ing that the same author subsequently showed None of the organic amides or acids tested actthat another Rhodococcus isolate, unlike the ed as inducers. But it is interesting to note the previous bacterium, could use p-hydroxyben- effect of adding extra cobalt, which lead to zonitrile and possessed a nitrilase that was good growth but poor nitrilase activity.Thisormore stable and did not undergo an activation ganism also contains a cobalt requiring nitrile process (HARPER,1985).The enzyme from this hydratase (see Sect. 4.1). Subsequently it was organism also had a much broader pH range shown that c-caprolactam (29,Fig. 13) was a over which it was active, had a molecular better inducer than isovaleronitrile and that weight of 560Ooo Da (12 subunits of 46OOO Da), *hiscompound also increased nitrilase activity 1 a variety of other rhodococci. It is noteworand was sensitive to reagents that react with thiol groups. In addition, this enzyme uds able thy that R. rhodochrous NCIMB 11215 beto use some ortho-substituted benzonitriles as haved differently in these experiments, showsubstrates, plus the substituted hydroxyben- ing a different ratio of activity to the substrates tested in comparison to the other bacteria zonitrile herbicides bromoxynil and isoxynil. More recently it has been shown that R. rho- (NAGASAWA et al., 1990~). An isolate designatdochrous NCIMB 11216 is also capable of ed as an Arthrobucter (BANDYOPADHYAY et al., asymmetric hydrolysis of ruc-2-methylbuty- 1986) showed that two nitrilases were present. ronitrile and other chiral nitriles (GRADLEY Therefore, this may also be the case for some and KNOWLES, 1994). It was possible to induce rhodococci.The nitrilase purified from R. rhothe nitrilase activity with either benzonitrile dochrous J1 had a molecular weight of 76000 (25,Fig. l l ) , propionitrile, or propionamide as Da and is constructed from two identical subthe sole source of carbon and nitrogen for units. It contained no iron or cobalt and no growth. Amidase activity was only observed other metal content was detected (KOBAYASHI, when cells were grown on propionitrile or pro- et al., 1989a). Provided that the enzyme was pionamide. Cell suspensions grown on propi- stored with glycerol and dithiothreitol, it was onitrile could not respire benzonitrile (25)and stable at low temperatures. It had a temperavice versa. No activation of the nitrilase en- ture optimum for activity of 45°C and a pH zyme appeared to occur in cell-free extracts of optimum of 7.5.The enzyme was very sensitive the bacterium prepared from propionitrile to chemicals that react with thiol groups. In a grown cells. With ruc-2-methylbutyronitrile as more extensive investigation of the substrate a substrate for cell-free extracts a kinetically specificity of this enzyme it was shown to have resolved transformation of the two isomers a similar profile to R. rhodochrous NCIMB was observed, with the (S)-isomer being the 11216. In a comparison of antigenic properties first to be hydrolyzed. Whole cells showed the of nitrilases from other rhodococci, and the nisame specificity, but nonselectively metabo- trile hydratase from this same bacterium, no lized the acid products. Some improvement cross reactivity was observed between anticould be made in the kinetic resolution when bodies raised against the R. rhodochrous J1 cell-free extracts were incubated at 4"C, rather nitrilase and the other enzymes.The biotransthan at the growth temperature (30°C). Using formation potential of R. rhodochrous J1 is imwhole cells it was shown that a range of C-2 pressive. Using whole cells KOBAYASHI et al. substituted substrates could be stereoselec- (1989a) used a fed batch process (because of tively transformed, the best kinetic resolution substrate inhibition) to convert p-aminobenbeing observed with ruc-2-methylhexanitrile zonitrile (39) into p-aminobenzoic acid (40) where the ( )-enantiomer is initially favored (Fig. 17), with yields of up to 110 g L-' and (GRADLEY et al., 1994). 100% substrate conversion. Using a similar NAGASAWA et al. (1988a) isolated another R. system, yields of 390 g L-' acrylic acid and rhodochrous, strain J1, from soil and deter- 260 g L-' methacrylic acid were obtained mined the optimal culture conditions for max- from the corresponding nitriles as substrates imal nitrilase activity. These workers showed (NAGASAWA et al., 1990b).

.

+


39

40

Fig. 17. The conversion of p-aminobenzonitrile into p-aminobenzoic acid by Rhodococcus rhodochrous J1.

297

lently bound intermediate is formed with the nitrilase. It was proposed that this was a thimidate or acyl-enzyme consistent with the reaction sequence outlined in Fig. 8. Aliphatic o-amino nitriles can be biotransformed by nitrilases in rhodococci to the corresponding amino acids (BHALLA et al., 1992). Unlike the other rhodococcal enzymes,the nitrilase from this did not apparently form a multi-subunit structure, but the same assessment of enzyme activation as performed by HARPER (1977b) was not carried out.

Rhodococci that have been shown to contain both nitrilase and nitrile hydratase activities tend to biotransform aromatic nitriles via Gram-Negative Bacteria the nitrilase (LINTONand KNOWLES,1986; VAUGHAN et al., 1988; VAUGHAN et al., 1989) Pseudomonas whilst metabolizing aliphatic nitriles via the niIn an unidentified Pseudomonas isolate trile hydratase present. Some rhodococci con- ROBINSON and HOOK(1964) showed that the tain nitrilases that are only active against aro- naturally occurring nitrile ricinine (41) could matic nitriles. R. rhodochrous K22 can use cro- be hydrolyzed to the corresponding carboxylic tonitrile as a sole source of nitrogen for acid (42) (Fig. 18). The authors mention that growth. It contains an aliphatic nitrilase with a stability problems occurred during attempts to molecular weight of 650000 Da made up of 15 purify the enzyme. Nevertheless, they were or 16 identical subunits. No metals appear to able to show that thiol group reagents inhibitbe necessary for its activity. Again, if stored in ed activity and proposed a mechanism for the 1964). glycerol and dithiothreitol, the enzyme was catalysis (HOOKand ROBINSON, stable with a temperature optimum of 50°C. The enzyme lost activity below pH 5. A wide Alcaligenes faecalis variety of aliphatic and aromatic nitriles could This gram-negative bacterium was isolated act as substrates for the enzyme. Acrylonitrile, from media where isovaleronitrile was used as succinonitrile, and glutaronitrile gave particu- a growth substrate. It was subsequently shown larily high activities in comparison with cro- to be able to hydrolyze indole-3-acetonitrile et al., 1990).The nitritonitrile. It is also noteworthy that mono hy- (28, Fig. 12) (MAUGER drolysis of glutaronitrile, to yield 4-cyanobu- lase activity could be induced by a variety of tyric acid, was possible (KOBAYASHI et al., nitriles, but isovaleronitrile gave the highest 1990a). No antigenic cross-reactivity between activities. None of the amides tested induced this nitrilase and those from other similar bac- the enzyme. Metal supplements to the basic teria was observed (KOBAYASHI et al., 1990b). growth medium did not influence the final nitIsovaleronitrile, a poor substrate for growth, was a good inducer of the enzyme (KOBAYASHI et al., 1991a).Following cloning of the nitrilase gene it was predicted that only one cysteine residue was in the amino acid sequence. The gene sequence was not identical but had many similarities with that of Klebsiella ozaenae (see Me Me below). Minimizing conformational changes in the enzyme by replacing the cysteine with ala41 42 nine or serine, it was shown that such mutaRicinine tions abolish the activity of the enzymes (KoBAYASHI et al., 1992a). STEVENSON et al. (1990) Fig. 18. The conversion of ricinine into the correhave gained evidence, using ion-spray mass sponding carboxylic acid by an unidentified Pseudospectrometry and acid quenching, that a cova- monas isolate.

298

6 Nitriles

rilase activity observed. The enzyme could not hydrolyze aliphatic nitriles, even though they could act as inducers, and preferentially used arylnitriles as substrates. The specific activity of the purified enzyme was much higher than that observed with other purified nitrilases (NAGASAWA et al., 1990a) and the active enzyme had a molecular weight of 260000 Da, consisting of six identical subunits. No metals appeared to be associated with the pure enzyme. The temperature and pH optima for the enzyme were 45 "C and 7.5, respectively, and the enzyme was moderately unstable. Once again a thiol group was implicated in the activity. There was no cross-reactivitybetween antibodies raised against this enzyme and the nitrilase from R. rhodochrous J1. YAMAMOTO et al. (1991) have shown that another isolate of this bacterium is able to stereoselectively convert a racemic mixture of mandelonitrile to produce R-( - )-mandelic acid. Of the inducers tested n-butyronitrile was the most effective. Over the time course studied the authors concluded that a stereoselective nitrilase was involved in the transformation, a racemase being implied to explain the less than 100% yield of product obtained. Klebsiella ozaenae MCBRIDE et al. (1986) showed that the manmade nitrile herbicide bromoxynil (43) could be metabolized by numerous soil bacteria (Fig. 19).These workers concentrated their efforts by studying the metabolism of this compound by one isolate, Klebsiella ozaenae. They showed that in cell-free extracts the nitrilase from this bacterium was highly specific for the herbicide. The gene for the protein is plasmid encoded and could be transformed into an E. coli host. Bromoxynil had to be present in growth media to select for cells retaining the

43 Bromoxynil

44

Fig. 19. The metabolism of the herbicide bromoxynil.

plasmid borne gene. It was shown that the gene coded for a peptide of around 40000 Da (STALKERand MCBRIDE,1987). On purification the enzyme was shown to be able to form dimers of 72000 Da. It had a pH optimum of 9.2 and an optimum temperature for activity of 35 "C. Once again thiol group reagents inhibit the activity of the enzyme (STALKERet al., 1988b). The gene coding for the nitrilase has been successfully expressed in tobacco plants, giving them resistance to the herbicide bromoxynil (STALKER et al., 1988a). Acineto bacter Immobilized cells of an Acinetobacter were able to convert a-aminopropionitrile (30a) to predominantly L-alanine (32a). a-Aminoacetonitrile (30c) could also act as a substrate for these cells being converted into the amino acid glycine (32c) (see Fig. 14).The apparent K, for the nitrilase activity was rather high (21 mM for immobilized cells, 34 mM for non-immobilized cells, MACADAM and KNOWLES, 1985). YAMAMOTO et al. (1990b) showed that this type of bacterium is also capable of other stereospecific biotransformations. Rac-2-(4'-isobutylpheny1)propionitrile was metabolized by whole cells of a Acinetobacter isolate to produce (S)-( )-2-(4'-isobutylpheny1)propionic acid, a non-steroidal anti-inflammatory drug. Little of the other isomer was synthesized during the time course of the experiment, Apparently the purified enzyme (molecular weight 560000 Da) could be constructed from two types of subunit, although this remains to be confirmed. The enzyme could use a wide range of aliphatic and aromatic nitriles 2-(4 '-Isobutylpheny1)propionitrile was a relatively poor substrate. Some amino acid sequence homology was found between this enzyme and the Kfebsieffa ozaenae nitrilase. The nitrilase of Acinetobacter was fairly stable in comparison with other nitrilases and had a temperature optimum of 50°C and maximal activity at around pH 8.0. Reagents that react with thiol groups inhibited the enzyme (YAMAMOTO and KOMATSU, 1991).

+

Fungi There have been very few reports on the nitrilases produced by fungi. In an early screen for this enzymatic activity by THIMANN and


(1964) six species of Fusariurn, MAHADEVAN two strains of Aspergillus niger, and one Penicillium chrysogenum isolate showed an ability to convert indoleacetonitrile into the corresponding acid. The A. niger enzyme was sensitive to reagents that react with thiol groups and no free amide could be detected during the reaction . FUKUDA et al. (1973) have shown that the yeast Torulopsis candida was able to use a variety of nitriles as a source of nitrogen for growth. The L-a-hydroxy acids could be synthesized from rac-hydroxy isocaproic acid and hydroxyisovaleric acid by whole cells of this yeast.There was no mention of amide intermediates being formed. Claims have been made that a variety of fungi (including a psychrophilic cyanide producer) can also produce chiral compounds from chiral nitriles, in this case amino acids (ALLENand STROBEL,1966, STROBEL, 1964; STROBEL, 1967). However, in some cases there is some doubt as to whether the activity is physiologically and biochemically significant (BUNCH and KNOWLES, 1980). Hsu and CAMPER (1976) showed that Fusarium solani was capable of degrading the aromatic nitrile ioxynil. HARPER(1977c), using another isolate of this fungus, showed that a nitrilase was present with a molecular weight of 620000 Da, consisting of six subunits. The enzyme was fairly unstable, had a broad pH range over which it was active, and could use a wide range of aromatic nitriles as substrates. Plants THIMANN and MAHADEVAN ( 1964) detected indoleacetic acid nitrilase activity in a limited, but significant,range of plant types. Properties of the enzyme partially purified from barley leaves indicated that the enzyme involved was similar in many respects to those found in microorganisms. It was active with 2-, 3-, and 4cyanopyridine (23, Fig. 10) plus a range of substituted benzonitriles and could also hydrolyze, albeit more slowly, several aliphatic nitriles (MAHADEVAN and THIMANN, 1964). Surprisingly,it is only relatively recently that plant nitrilases have received much more extensive investigation. The new research was stimulated by studies into the synthesis of the plant hormone indoleacetic acid, where the relative importance of various synthetic routes

299

had not been assessed. Plants are able to synthesize nitriles (see Sect. 3) and it has been shown that indole-3-acetonitrile(28, Fig. 12) is et al. (1992) a natural plant product. BARTLING cloned the gene coding for the indoleacetonitrilase from Arabidopsis fhalianainto an E. coli. Sequencing the gene showed that it had many similarities to the nitrilase found in Klebsiella ozaenae. The estimated size of the basic enzyme component was about the same as the bacterial enzyme, although whether multimers were formed was not determined. BARTELand FINK(1994) have cloned four nitrilases from A . rhaliana, three of which are very similar. It was shown that the “NIT” genes are each expressed preferentially in different plant tissues. There was some evidence that one of the nitrilase genes is expressed following bacterial pathogen infiltration into vegetative tissue. Using the same plant BARTLING et al. (1994) observed two nitrilase activities, one soluble, the other membrane associated and expressed only at certain stages of plant development.

4.3 Enzymes Capable of Biotransforming Cyanide 4.3.1 Fungal Cyanide Hydratases DUBEYand HOLMES(1995) have recently reviewed the enzymes obtainable from microorganisms that can catalyze the transformation of hydrogen cyanide. Three basic mechanisms exist that involve attack by an oxidase, a hydrolase, or assimilation into another molecule (Fig. 20). Once again a relatively diverse range of microorganisms are capable of hydrogen cyanide metabolism and the subject has been reviewed several times (for example, see KNOWLES and BUNCH,1986; HARRISet al., 1987). Fungal hydrogen cyanide metabolism seems to most commonly involve the enzyme cyanide hydratase. This enzyme adds water to the nitrile group to generate formamide, which is then converted to formate, and usually to carbon dioxide, by other enzymes (Fig. 20). FRY and MILLAR(1972) demonstrated this activity in the plant pathogenic fungus Sremphylium

300

6 Nitriles

Cyanide hydratase

+ J O H

Cyanide

Cyanide oxidase 02,

-----)

co2

I

H- C E N

H20

NAD(P)H

I

(CNO)

dihydratase

2H2O

CO2

-t

NH3

C02

+

NH3

Cyanase

Fig. 20. Enzymes involved in the metabolism of hydrogen cyanide by bacteria.

loti, which attacks the plant bird’s-foot trefoil (Zeneca plc) Biological Products. The enzyme liberating hydrogen cyanide from its cyano- was estimated as forming up to 22% of the solgenic glycosides. RISSLERand MILLAR(1977) uble protein present in fungal cells induced showed that the activity of this enzyme is relat- with hydrogen cyanide,and had a pH optimum ed to other metabolic activities in the fungus, of 8.5. Reduced activity at higher pH values notably the cyanide-insensitive alternate res- may be related to the concentration of the cyapiratory system. Recent studies have focused nide ion. Using denaturing and non-denaturon the role that cyanide hydratase plays in pro- ing gel electrophoresis it was shown that the tecting this and other plant pathogens that lib- native molecular mass was between 300erate hydrogen cyanide during the pathogenic 400000 Da, with subunits of 43000 Da. The nitrilase from Fusarium oxysporum and E soprocess. WANGet al. (1992) studied the inducibility lani (see Sect.4.2) forms similar aggregates. No of cyanide hydratase by hydrogen cyanide in nitrilase activity was found in the crude ex19 fungal isolates. Gloecercospora sorghi, a tracts of E lateritium. Most of the gene coding pathogen of the plant Sorghum bicolor was se- for the cyanide hydratase was expressed in an lected for further study. These workers puri- E. coli host and the gene showed good sefied this enzyme to homogeneity yielding a quence homology with the bromoxynil nitrisingle 45000 Da protein, which could be re- lase from Klebsiella pneumoniae (STALKER et solved into three isozymes by gel electropho- al., 1988a), and some homology with the nitriresis each with a different pI.The K , of the un- lase in Alcaligenes faecalis JM3 (KOBAYASHI et resolved “purified” preparations for hydrogen al., 1993). It is interesting that the cysteine rescyanide was 12 mM, thus showing a relatively idue, implicated in the catalytic mechanism of low affinity for the substrate. The enzyme had the nitrilases, is in one of the regions showing an optimum pH of 7-8 for activity, and Ag+, homology between the two types of enzymes. Zn2+, and CU” were all inhibitory. Com- No sequence homology was found between pounds that react with thiol groups also inhib- the cyanide hydratase and the nitrile hydraited the enzyme, with the exception of iodoac- tase from Rhodococcus sp. 774 or Myrotheetate. Metal chelating agents did not effect the cium verrucaria.WANGand VANETTEN (1992) activity of preparations, but sodium dodecyl- showed that the gene coding for the cyanide sulfate, at the concentrations tested, caused ir- hydratase from Gloecercospora sorghi showed reversible inactivation. A fairly specific poly- some, but not as much, homology with the clonal antibody was raised against the purified Klebsiella pneumoniae nitrilase.These workers protein that inactivated the cyanide hydratase. expressed the enzyme in an Aspergillus niduNo investigation of alternative substrates for fans host. the enzyme was reported. Similar observations were made by FRYand MUNCH(1975). CLUNESS et al. (1993) purified cyanide hy- 4.3.2 Other Enzymes dratase from the cyanide tolerant fungus Fusarium lateritium used in the cyanide-degradPseudomonas jluorescens NCIMB 11764 ing product CYCLEAR manufactured by ICI when growing on cyanide has the capacity to


produce several enzymes capable of using cyanide as a substrate (KUNZet al., 1994;see Fig. 20). HARRISand KNOWLES (1983) showed that one of these enzymes is an oxygenase, using NAD(P)H as a reducing agent in the conversion of cyanide to carbon dioxide and ammonia. A similar enzyme could be involved with the conversion of thiocyanate to carbonyl sulfide by Thiobacillus thioparus (KATAYAMA et al., 1992). DORR and KNOWLES (1989) have shown that this bacterium also has a cyanase activity which may be involved in this conversion. The enzyme is induced when this bacterium grows on metal cyanide complexes (ROLLINSON et al., 1987). By comparing reaction product stoichiometries, cofactor requirements, and using kinetic analysis (under different assay conditions) KUNZ et al. (1994) showed that there may be two other types of enzyme present, one a cyanide nitrilase (dihydratase), the other a cyanide hydratase. MEYERS et al. (1993) have purified a cyanide dihydratase present in Bacillus pumilus C1 and have shown that the enzyme is not only structurally different to nitrilases so far described, but also cannot use acetonitrile as a substrate.

301

re face

fl

si face

45

46

Fig. 21. Enzyme catalyzed cyanohydrin formation.

the isozymes of this enzyme and performed initial X-ray diffraction studies. Operating the enzyme with organic solvents not miscible with water can reduce non enzymic catalyzed synthesis of cyanohydrins, yielding a product with high enantiomeric excess. In other plant sources, such as Sorghum bicolor, (S)-oxynitrilases can be found. It is interesting that the two types of oxynitrilase differ structurally as well as in the products they make (VANSCHARRENBURG et al., 1993;WOKER et al., 1992).The ( R ) oxynitrilase has an FAD prosthetic group which is missing in the other enzyme. It is also noteworthy that the (S)-oxynitrilase can only at present be isolated in small amounts, a clear target for recombinant gene technology. In addition, this enzyme only adds hydrogen cyanide to aromatic and heteroaromatic aldehydes. The use of catalytic systems operating in 4.4 Enzymes for the Synthesis and organic solvents is particularly useful when dealing with substrates that are poorly water Transformation of Cyanohydrins soluble. Ketones can also act as substrates for This has been reviewed recently in an excel- the (R)-oxynitrilases, although the range of lent article by EFFENBERGER (1994), that fo- chemicals acceptable to the enzyme is much et al., 1993; cused specifically on the use of such enzymes less than for aldehydes (ALBRECHT and HEID,1995). The (S)-oxyfor the synthesis and transformation of opti- EFFENBERGER cally active cyanohydrins. Therefore only a nitrilase apparently cannot use ketones as substrates. Alternative enzymes to the (R)-oxynitbrief resume of the subject will be given here. rilase synthesized by Prunus amygdalus are available. A new type has recently been reported by WAJANTet al. (1995) which was iso4.4.1 Oxynitrilases lated from the cyanogenic fern Phlebodium These enzymes were used for one of the first aureum.This enzyme is present as a multisubasymmetric syntheses with biological catalysts. unit complex, and there are at least three isoThey catalyze the conversion of aldehydes and zymes of the enzyme. Unlike other oxynitrilashydrogen cyanide to cyanohydrins (Fig. 21). es the enzyme was not inhibited by sulfhydryl The plant Prunus amygdalus contains an (R)- or hydroxyl modifying chemicals. It also does oxynitrilase,which catalyzes addition to the re- not contain FAD. In plants that naturally synthesize cyanohyface of the aldehyde. It has a typically low substrate specificity, and high enantioselectivity drins, the free hydroxyl group can be conjugat(EFFENBERGER et al., 1995; WARMERDAM et ed with glucose (CONN,1979).Lipases are also al., 1996).LAUBLE et al. (1994) have crystalized able to conjugate this hydroxyl group in this

302

6 Nitriles

case with various aliphatic carboxylic acids, leading to the possibility of “one-pot’’ syntheses of optically active products using chemical or enzyme catalyzed synthesis of cyanohydrins (EFFENBERGER, 1994; INAGAKIet al., 1992).

5 Biotechnology of Nitrile Transformations It is relatively straightforward to synthesize nitriles using chemical technology, and they have important roles as “synthons” for the construction of commercial products. However, the opportunities for generating useful compounds from nitriles can be greatly enhanced using biocatalysts which are capable of regioselective hydrolysis of molecules containing more than one nitrile group (or an additional functional group that is susceptible to hydrolysis by nonbiological technologies), and stereoselective synthesis from chiral or prochiral nitriles. In many cases partial hydrolysis to amides is also possible. The technology for enzyme based nitrile transformations has been available for some time. Many notable successes using this type of technology have been reported, including the generation of products at concentrations commonly encountered using chemical technology. Biological systems are often considered incapable of such yields. Below is a review of what applications have been reported in the literature. It is extremely likely that this represents only a small percentage of what is known to be possible using nitrile hydratases and nitrilases, the remaining information being commercially sensitive. Not included in this section is the cloning of nitrile hydrolyzing enzymes into plants to confer resistance to herbicides used for the elimination of unwanted plant competitors. The early stages of the development of this type of technology is dealt with in Sects. 4.1 and 4.2 that specifically deal with the individual types of enzymes.

5.1 Use for Chemical Synthesis Tab. 2 summarizes the non-stereoselective nitrile hydrolyses performed by microbial enzymes, that have been described in the literature, which evaluated the substrate which specific microorganisms and cell-free preparations could biotransform. In addition to this list are one-off studies on the transformation of specific nitriles. These are detailed in Sects. 4.1 and 4.2. In some respects the list in Tab. 2 is not very inspiring. However, the vast majority of the data published was probably generated by researchers making use of commercially available nitriles, rather than testing the full potential of the biocatalysts with a wider range of non-commercially produced molecules. FABER (1994) summarizes the product yields that can be obtained using biotechnology. This book and that by ROBERTS et al. (1995) are an excellent guide to the factors that should be taken into account when using biotechnology, and the potential benefits that can be obtained.

5.1.1 Transformation of Mononitriles This subject has been reviewed in Sects. 4.1 and 4.2 and by others (THOMPSON et al., 1988; NAGASAWA and YAMADA,1992; KOBAYASHI and SHIMIZU, 1994).The main features to highlight with regard to biotechnological applications are firstly the potential for high product yields and secondly the capability of stopping the hydrolysis of the nitrile group at the amide stage. The latter can be achieved by either reducing/eliminating amidase activity from biocatalyst preparations, or by the use of amidase inhibitors (MAESTRACCI et al., 1984a, b). Acrylamide (22,Fig. 9) and acrylic acid synthesis are the best known commercial processes using this technology, where the process yields can be in excess of 60% (w/v). Currently the process accounts for nearly 30% of the world’s production of acrylamide (22).The system also has the attractive feature that it is “cleaner” than the alternative chemical route, avoiding the use of metal catalysts and generating fewer impurities. As penalties for environmental

5 Biotechnology of Nitrile Transformations Tab. 2. Examples of Nitrile Biotransformationsby Microorganisms Microorganisms Capable of Nitrile Transformation" Nitdeb

Nitrile Hydratase Activity

Nitrilase Activity

CH3-C= N acetonitrile

Y; FS; PC23; RRJ1; B312; NRLL; AG

FS

CH,-CH,-CGN

Y; FS; PC23; RRJ1; NRLL; PS13; AG

proprionitrile

CH3-(CHz),-C=N

butyronitrile

Y; FS; PC23; RRJ1; B312; NRLL; PS13; AG

CH3-(CH,),-C=N

valeronitrile

Y; PC23; RRJ1; B312

CH3-(CH,),-C=N

capronitrile

PC23; RRJl

CH, -(CH,), -C=N pelargonitrile Y Me,CH-C=N

Y; PC23; RRJ1; B312

tg, -CE N

Y; PC312; RRJ1; B312

&EN

2 1

Acry loni tri le

EtO-

Y; PC23; RRJ1; B312; AG; RR774; BCH2; AIP

RRK22

RRK22

E N

3-Ethoxyacrylonitrile RRK22

c1

A

N

2-Chloroacrylonitrile RRK22 RRK22 3-Aminocrotonitrile

A

PC23: RRJ1; B312; NRLL CN

RRK22; RRJl

Methacrylonitrile

-EN

PC23: RRJl

Crotonitrile RRJl

RRK22

303

6 Nitriles

304 Tab. 2.

Continued Microorganisms Capable of Nitrile Transformation”

Nitrileb

NGC-CH,-C-N

malononitrile


Nitrilase Activity

RRJl

RRK22

FS; B312; NRLL; AG

N=C-(CH&-C=N

succinonitrile Y; FS; B312; NRLL

N=C-(CH,),-C=N

glutaronitrile FS; B312; AG

N=C-(CH,),-C=N

adiponitrile

FS; B312 RRK22

Fumaroni trile RRK22

B312

HCN

B312

HzNCN (37)

MV PC23; RRJ1; B312

HO-EN

OH

AEN

B312

Lactonitrile

MeO-

CN

E t o r G N

RRJ1; B312 PC23; RRJl

0

PC23; RRJl

PC23; RRJI; B312;AG PC23; RRJ1; B312; AG

5 Biotechnology of Nitrile Transformations

305

Tab. 2. Continued Microorganisms Capable of Nitrile Transformation" Nitrileb


'""x"" E

Nitrilase Activitv

RR21197

N

R361 R O i E N E N 59 R =

H,Bn, Bz, MEM PC23; RRJl

(27) X=S or 0

P

E

RRJl

N

0""

RRJl

RRJ1: B312; NRLL; AG

NRLL; RR215; AJ1

RRJ1: AG

RRJl

0-FS rn-FS; RR215; RR216 p-FS; RR215; RR216 rn- and p-RRJ1

0-FS rn-FS; RR215; RR216 p-FS; RR215; RR216; AJ1

306 Tab. 2.

6 Nirriies Continued Microorganisms Capable of Nitrile Transformation”

Nitrileb


Nitrilase Activity

MeoDEN m- and p-RRJl

NH,

(39)

m- and p-RRJI

0-, m-

(43)Brornoxynil

Br I

HO

and p-RRJI

p-RRJ1

0-RR215; FS m-RR215; FS; RR216 p-RR215; FS; RR216 RR215; FS; KO

Br m-and p-RR.215; FS; RR216

0-, m-, andp-RRJ1

0-, m-

0-RR215 m-RR215; FS; RR216 p-RR215; FS; AJ1; RR216

and p-RR215; FS; RR216

m- and p-RRJ1

RRJl

m-and p-RRJI

0-FS; RR215 rn-RR215; FS; RR216 p-RR215


307

Tab. 2. Continued

c-

Microorganisms Capable of Nitrile Transformation"

Ni trile

N

CZN

23

"7""


Nitrilase Activity

0-RRJ1 rn-RRJ1 p-RRJ1

0-RR215;FS; RR216 m-RR215; FS; NRLL; RR216; RRJl p-RRJ1; RR215; FS; AJ1; RR216

RRJl

RRJl

N

RRJ1; AG; RH

FS

\

H (28) Indole-3-acetonitrile ~~

~

~

~

Microbial abreviati0ns:AC.i Agrobacterium sp.;AIP Arthrobacter sp. IPCB3;AJl Arthrobacter sp. J1: B312 Brevibacterium 312; BCH2 Brevibacterium sp. CH2; FS Fusarium solani; KO Klebsiella ozaenae; MV Myrothecium verrucaria; NRLL Nocardia rhodochrous LL100-21; PS13 Pseudornonas S13; PC23 Pseudornonas chloraphis B23: RH Rhizobium sp.; R361 Rhodococcus sp. SP361; RR21197 Rhodococcus rhodochrous ATCC 21197; RR215/216 R. rhodochrous 215/216; RRJl R. rhodochrous J1; RRK22 R. rhodochrous K22; Y a

yeast sp.

Non-systematic names for structures shown as recorded in cited literature.

damage resulting from manufacturing steadily increase in many parts of the world, this adds an extra important benefit to the use of biotechnology (RAMAKKRISHNAand DESAI, 1993). Equally impressive yields have been obtained from other mononitrile hydrolyses. For example, using a nitrilase from Rhodococcus rhodochrous J1, over 10% (w/v) of the vitamin p-aminobenzoic acid (40) could be obtained from p-amino benzonitrile, (39) (see Fig. 17) (KOBAYASHIet al., 1989a). Other important biochemicals are obtainable in even higher yields (FABER,1994). A constant search is underway for enzymes with different properties that allow further biotechnologies t o be developed. COHN and WANG(1995) have recently evaluated a Rhodococcus for the transformation of heteroaro-

matic nitriles to amides and acids. LANGDAHL et al. (1996) have isolated a strain of Rhodococcus erythropolis from marine sediments that has the highest reported tolerance to acetonitrile (900 mM) and some uncharacteristic nitrile transforming capabilities, thus giving this nitrile hydratase containing bacterium new possible uses for chemical synthesis and bioremediation of nitrile pollutants. Others are finding new uses for previously isolated biocatalysts. For example, LIUet al. (1995) have used a Rhodococcus nitrilase as the basis of a microelectrode system for the selective detection of organonitriles. It was possible to detect between 0.1 and 3.3 mM benzonitrile with a linear response, and the system was tolerant to the presence of other aromatic compounds. However, it is the use of nitrile biotransform-

308

6 Nitriles

ing enzymes for regio- and stereoselective synthesis that has probably the greatest potential for the future.

5.1.2 Regiospecific Biotransformation of Dinitriles It was shown in Tab. 2 that dinitriles can act as substrates for hydrolytic enzymes. Investigations have shown that microbes are capable of transforming one or both of the nitrile groups (KOBAYASHI et al., 1988). COHENet al. (1990) have reported that a fairly wide range of aliphatic and aromcatic molecules of this type can be selectively transformed. In Sect. 5.2 it will be shown that this capability can be exploited for the detoxification of wastes containing fumaronitrile, a dinitrile that is a potent biocide. Using an immobilized Rhodococcus it was shown that isophtalonitrile could be converted to the cyanoacid. The intermediate amide could also be synthesized, at lower yields, by changing the incubation conditions for the reaction. Using terephthonitrile, a symmetrical dinitrile, one nitrile group was again selectively transformed. However, when the corresponding diamide was used as a substrate for the amidase in this bacterium, both amide groups were converted to the corresponding acid. When testing the conversion to acids of a range of other aromatic dinitriles variable selectivity was observed, but in all cases it was poorer than in the previous example. Using a strategy based on earlier observations by others, these workers neatly showed that with aromatic fluorodinitriles selective production of the corresponding monoamide could be achieved. When employing fluorinated aromatic dinitriles where one is orfho-substituted with respect to an amine group (47), single amide isomers of the orfho-nitrile group could be obtained (48) (Fig. 22). Finally, dinitriles can be metabolized by the fungus Trichoderma sp. MB 519 with the liberation of hydrogen cyanide, as well as hydrolysis of the nitrile group. This could represent a “nitrile lyase” activity in this organism. No details were given with regard to the possible selectivity of the reaction (KUWAHARAand YANASE,1985).

47

48

Fig. 22. Selective hydrolysis of aromatic dinitriles.

5.1.3 Stereoselective Biotransformation of Nitriles This is potentially the most important biotechnological application of nitrile hydrolyzing enzymes. It is worth noting that stereoselectivity is possible in the nitrile hydratase biotransformation route both with regard to the initial hydrolysis by the nitrile hydratase and the subsequent hydrolysis of the amide by the amidase. FUKUDA et al. (1971,1973) have shown that microorganisms can transform racemic a-aminonitriles (30)and racemic a-hydroxynitriles. In the former type of molecule a mixture of Land D-amino acids (32,33) were obtained (see Fig. 14). MACADAM and KNOWLES (1985) using a putative Acinetobacter showed that it was possible to produce up to 87% of L-alanine (32a) from a-aminopropionitrile (30a), via nitrilase hydrolysis. Interconversion of the two enantiomers of a-aminopropionitrile aided the high yields of L-amino acids (32). Other workers showed that in the absence of such an activity only a 50% yield of the L-enantiomer was possible (ARNAUDet al., 1980). A similar result was obtained with a suspected nitrilase on racemic a-hydroxyisovaleronitrile and racemic a-hydroxycapronitrile in the yeast Torufopsis candida GN405. This organism was unable to convert both enantiomers of a-aminonitriles to amino acids. KAKEYAet al. (1991) performed a kinetic resolution of arylpropionitriles using the enzymes present in Rhodococcus bufanica ATCC 21197. Using a-arylpropionitriles containing sterically bulky, electron-withdrawing and electron-donating groups they showed


that in all cases (R)-amides of high enantiomeric excess were synthesited, along with (S)carboxylic acids. It was proposed that this bacterium contained both nitrile hydratase and nitrilase enzymes. However, they also showed that an amidase was present that preferentially converted (S )-amides to the corresponding carboxylic acids.These observations show that stereoselective conversions are possible using both microbial nitrile hydratases and nitrilases. GRADLEY and KNOWLES (1994) and GRADLEY et al. (1994) examined the biotransformation potential of a nitrilase activity present in R . rhodochrous NCIMB 11216. Using whole cells of this bacterium induced by growth on either benzonitrile (25, Fig. 1l), propionitrile, or propionamide it was shown that different biotransformation capabilities could be expressed. Using propionitrile-, but not benzonitrile-induced cells, it was shown that both enantiomers of 2-methylbutyronitrile and 2methylbutyric acid could be transformed. These cells were unable to metabolize racemic 2-methylbutyramide. Cell-free extracts of this bacterium revealed that (S)-2-methylbutyronitrile was biotransformed faster (about 3-fold) than the (R)-enantiomer. A kinetic resolution of the potential products was, therefore, possible. Using a variety of C-2 substituted nitriles it was shown that the best kinetic resolution was possible with racemic 2-methylhexanitrile, although racemic 2-phenylpropionitriie was also reasonably well resolved in comparison with racemic 2-methylbutyronitrile. Mandelonitrile is a natural product and can be biotransformed to benzaldehyde and hy-

rac-49

309

drogen cyanide by oxynitrilases (see Sect. 4.4.1). LAYHet al. (1992) using soil bacterial isolates obtained from cultures grown on a variety of aromatic nitriles, showed that it was possible to obtain enantioselective hydrolysis of 0-acetylmandelonitrile (stable under the incubation conditions needed) by nitrilases. All the bacteria were identified as pseudomonads. From racemic mixtures of this compound all isolates preferentially formed (R)(-)-0-acetyl mandelic acid, varying ratios of the two isomers were produced by the different bacteria. It was important that the microorganisms used had low esterase activity to avoid deacetylation and breakdown of this compound via other metabolic routes. Enantioselective hydrolysis of nitriles by a Pseudomonas has also been observed by MASUTOMO et al. (1995). In this case the transformation was the result of a nitrile hydratase having poor enantioselectivity for the nitrile (49), and an amidase that only hydrolyzed the (S )-enantiomer of the racemic amide (50) to give the (S)-acid (51) and recovered (R)-amide (52) (Fig. 23). The reverse type of selectivity has been observed with 3-substituted glutaronitriles where an (S)-selective nitrile hydratase, followed by a nonselective amidase, can be used to transform one of the nitrile groups to synthesize (S)-acids (CROSBYet al., 1994; GODTFREDSEN and ANDRESEN, 1986). This technology has been used for the synthesis of pharmaceuticals (YAMAMOTO et al., 1990). Increasingly, enantioselective amidases from non-nitrile hydrolyzing as well as nitrile hydrolyzing bacteria are being used for selective transformation of chemically synthesized

rac-50

Fig. 23. Exploitation of a non-enantioselective nitrile hydratase and an enantioselectiveamidase.

310

6 Nitriles

ysis of the malononitrile (56)(Fig. 25) gave the amidocarboxylic acid (58) as the sole product in 92% yield. The intermediate diamide (57) was detected in the reaction mixture, but not the corresponding amidonitrile, which indicates that nitrile hydrolysis is faster than amide hydrolysis (YOKOYAMA et al., 1993). Conversely, in the Rhodococcus sp. SP361 mediated hydrolysis of the dinitrile (59) (Fig. 26) the nitrile hydrolysis was slow and selective whereas the amide hydrolysis was fast and nonselective. This yielded the carboxylic acid nitrile (61) (83% ee) as the sole product (BEARDet al., 1993). Given the frequently observed broad substrate specificity of such enzymes it is clear that they are likely to find more extensive uses for the production of enantiomerically pure chemicals in the future. Stereospecific synthesis of nitriles is also possible using enzymes in-

amides. For example, CISKANIK et al. (1995) have shown that the amidase from the nitrile transformer Pseudomonas chlororaphis B23 exhibited enantioselectivity for several aromatic amides. In this case the (S)-enantiomers were the preferred substrates. YAMAMOTO et al. (1996) used an amidase from Comamonas acidovorans KPO-2771-4, which has the unusual ability of favoring the (R)-enantiomer of racemic amides. In this case the activity was exploited for the production of (R)-( -)-ketoprofen (54), an anti-inflammatory drug (Fig. 24). The resolution of racemic nitriles by hydrolytic enzymes can only give a 50% yield of product, unless in situ racemization occurs. However, enantioselective hydrolysis of prochiral nitriles has the potential to yield a single enantiomerically pure product in 100% yield. R. rhodochrous ATCC 21197 mediated hydrol-

&Hdbg;H2 dH2 /

/

/

/

/

__t

5 4 ( R)-(-)-Ketoprofen

53

/

55

Fig. 24. Enantioselective amide hydrolysis in the resolution of ketoprofen.

"Bu

L=IY

N

C='

nitrile hydratase Fast

b

I

56

57

58

Fig. 25. Selective amidase hydrolysis of prochiral dinitriles.

pro-(S)-Selective nitrile hydratase

Slow

CfN

Non-selective amidase

*

Fast 0

59

C EN

*

B n o V O H 0

60 not detected

Fig. 26. Selective nitrile hydratase hydrolysis of prochiral dinitriles.

61


volved with natural nitrile metabolism. VAN DER WERF et al. (1994) have reviewed the use of oxynitrilases and other lyases in such applications.

5.1.4 Commercial Processes

311

needs. FINNEGAN et al. (1992) observed that at the date of their review the starting nitriles for the potential biological synthesis of commercially important amides, such as benzamide, formamide, chloroacetamide, propionamide, salicylamide, nicotinamide, and phenacetamide, were more expensive than the products. This is unlikely always to be the case, and as additional requirements of industry develop there is every chance that biotechnological methods will become extensively used. The most important commercial use of this technology to date is the conversion of acrylonitrile (21)to acrylamide (22) (see Fig. 9) in the Nitto process. It illustrates the exciting potential for the use of biocatalysts and serves as a typical example of biological process design. Most noteworthy is the fact that it could now account for up to 30% of the world’s total production of acrylamide.

Clearly the use of biocatalysts for the commercial synthesis of chemicals is possible. As with any process, the viability of using biotechnology has to be assessed with regard to current technology and present and future legislation. The specificity of enzymes can be an important factor when product purity is preeminent, as in the pharamaceutical industry. In addition, the potential for cleaner synthesis, reducing waste treatment, and energy costs can also make the use of biocatalysts in a process attractive. It is not appropriate to discuss optimization strategies for maximizing the efficient use of biocatalysts in this review. Where the technology does not have to compete 5.1.4.1 Biotransformation of strongly with existing methodologies, produc- Acrylonitrile to Acrylic Acid tivity targets can be much lower than is usually One of the earliest patents relating to this obtained with chemical methods. However, if et al. the process is competing with such systems the process was lodged by COMMEYRAS task of matching product concentrations in (1975). They compare the biological process bioreactors can be rather daunting. All the with chemical technology claiming better relimost significant biotransformations of nitriles ability and percentage conversion for the biouse water as a cosubstrate. Hydrolytic reac- catalysts used. Both the range of bacteria cations are frequently the easiest type of biocat- pable of the transformation and the range of alytic activity to use for high yielding process- amides that could be made using these organes, as it is often possible to use the enzymes in- isms were highlighted. The Nitto Chemical volved “uncoupled” from the other metabolic Company in Japan subsequently published a processes of the cell. For example, if reduced number of patents relating to the basic use of cofactors are also needed for a reaction, the the technology and subsequent improvements use of isolated enzymes can become too ex- to overcome operational problems. WATApensive, and intact cell systems more difficult NABE et al. (1979) outline the process using to operate. Many of the microorganisms that bacteria from the genus Nocardia or Corynecontain nitrile hydrolyzing enzymes are natu- bacteria. Yields of 10-20% (w/w) were reportrally designed to survive under the conditions ed, but under conditions where maximum often found in high intensity bioreactors. Con- yields were obtained the biocatalysts lost acsequently, even for bulk-chemical production, tivity making their reuse difficult. They deit is not surprising that the benefits of using scribe a method for continual production by biocatalysts can outweigh the disadvantages in polyacrylamide immobilized bacterial cells some processes. In an increasing number of placed in columns with multiple inlet feeds. cases the factors preventing their use are the These workers also noted that the life of the cost differences between starting material and biocatalyst could be extended by operating the product, often due simply to the fact that the process at 15°C or lower, which reduced the production of starting materials has been tra- inhibitory effects of both the substrates and ditionally geared towards chemical technology products. A separate patent filed at around the

312

6 Nitriles

same date claimed that the addition of dialde- lysts and/or bioreactors that can exploit their hydes to cells before use of the reactors pre- use. Fortunately, progress is being made in the vented the polymerization of amide products acquisition of both of these requirements. by components of the bacterial cells (WATANABE and SATOH,1980). A number of subsequent patents covered the wider range of microorganisms that could be used and refine- 5.2 Bioremediation of Nitrile Conments on the original process (YAMAGUCHI et taining Wastes al., 1980, WATANABE et al., 1981a, b; YAMAGUCHI et al., 1981; KANEHIKO et al., 1987) have Hydrogen cyanide and nitrile contaminashown that various amides and organic acids tion of environments is not unusual. Old gas can also act as stabilizers of the nitrile hydra- work sites illustrate the way that such contamtase activity in intact cells and isolated en- ination can severely effect the recycling of land zymes. Throughout the development of the for new uses. Although this review is primarily process the improvement of the biocatalyst concerned with the use of nitrile transforming has been achieved by optimizing the growth enzymes for chemical synthesis, it is approprimedium used to produce the active catalyst ate in a section on biotechnological applicaand by finding ever better bacterial strains. In tions to briefly mention the use of these biothe third generation nitrile hydratase used for catalysts in the remediation of polluted enviacryl-amide production employing cells of R. ronments. MUDDER and WHITLOCK(1983, 1984) rhodochrous J1 an astonishing 65.6% (wlv) acryl-amide in the reaction mixture can be showed how microorganisms can be used to produced by cells which have over 50% of treat wastes from lead mining operations contheir soluble protein made up of the nitrile hy- taining cyanide, cyanide complexed to metals, drolyzing enzyme, operating at temperatures and thiocyanate in the presence of other inorup to 50°C (KOBAYASHI et al., 1992b). NAGA- ganic molecules. So good was the resultant SAWA and YAMADA (1995) place this system in product that the treated water could be used the context of new processes using microor- for trout farming.A key organism in the mixed ganisms as sources of catalytic activities for the cultures was Pseudomonas paucimobilis which production of commodity chemicals, revealing presumably attacked the cyanides using enthe potential for industries to apply biotech- zymes mentioned in Sect. 4. This technology is nology. Current research continues to show another example of how fairly unpleasant that the techniques for developing the enzy- chemical procedures for treating cyanide matic synthesis of acrylamide can also be used wastes can be replaced with more user and ento help exploit other similar bacteria (LEE et vironmentally friendly procedures (GREEN al., 1993; BAUERet al., 1996; KOBAYASHI et al., and SMITH,1972;HARDEN et al., 1983).The dis1996). MOREAUet al. (1994a) have mutated covery of fungi that form cyanide hydratase cells of Brevibacterium sp. R312 to improve has enabled the development of an alternative the enzymic conversion of adiponitrile to adip- technology for dealing with cyanide wastes. ic acid, one of the main raw materials used in ICI (Zeneca p/c) Biological Products has used the synthesis of nylon 6,6. Given the advances such enzymes as the basis of their “CYCnow being made in both molecular biology LEAR” system for treating hydrogen cyanide and bioreactor development it would not be containing wastes. It can be manufactured in a too optimistic to predict that a much wider variety of forms (e.g., pellets, powders, solurange of commercially viable nitrile biotrans- tions), to suit various requirements. 1 kg is able formations will be available in the near future to treat 50 kg of hydrogen cyanide, reducing (BUNCH, 1994). Immobilized cell technology is the concentration to about 10 ppm in a few well suited to the transformation of water sol- hours. A drawback to the use of these enzymes uble nitriles and amides. However, it is often is that they can be sensitive to high concentrathe case that transformation of poorly water tions of metal cyanide complexes, particularly soluble substrates is required. This will require those of iron and copper. But with the right apthe development of solvent resistant biocata- plication this product offers an excellent safe

6 Future Developments

method for treating wastes. DUBEY and HOLMES (1995) have recently reviewed the full range of enzymes that can be used to biotransform hydrogen cyanide and illustrate how the technology can be used for large scale treatment of wastes generated by the mining and gas industries. These are interesting examples as, in the case of the gasworks sites, the cyanides in the contaminated samples were present with phenols and polyaromatic hydrocarbons. It was shown that a mixed community of microorganisms could work together to treat all these compounds. It is possible to find some microorganisms that are able to remediate wastes containing hydrogen cyanide and nitriles (BABUet al., 1994). An isolate of Pseudomonas putida was able to utilize acetonitrile and sodium cyanide as a sole source of carbon and nitrogen. Using immobilized cells the authors showed the conversion of these compounds into carbon dioxide and ammonia. WYA'ITand KNOWLES(1995) have recently described an elegant strategy for treating waste streams that contain nitrile mixes that no single microorganism is capable of metabolizing. Their focus was on the waste streams produced from the manufacture of acrylonitrile which contains 9 major organic components, including the potent biocide fumaronitrile. One of the most important features of the work described was that although traditional enrichment techniques could not find any organisms that were capable of the complete conversion of fumaronitrile into metabolizable products, mixtures of microbes could perform this transformation. Their work is encouraging because it implies that sites Contaminated with simpler nitrile mixtures should be easy to treat. Observations suggest that as in this case nitriles added to environments like the soil, either accidentally or as applications of nitrile herbicides and pesticides, can be removed (Hsu and CAMPER,1976; DONBERG et al., 1992). However, sometimes the metabolism of nitriles may result in the production of cyanide as an intermediate in the process (TOPPet al., 1992).

313

6 Future Developments 6.1 Search for Novel Nitrile Biotransforming Activities During the last few years there has been a great deal of interest in the actual numbers of living organisms on our planet. Most estimates of the earth's biodiversity indicate that we have so far isolated and studied relatively few of the species that exist. This is particularly true of the microbial world, where in some cases it has been proposed that up to 95% remain to be discovered. Since microorganisms have provided so many different nitrile biotransforming enzymes already, it is possible to speculate that many more useful biocatalysts exist. Before the development of molecular probes for detecting putative genes coding for such proteins, the search for novel activities could be slow. Development of rapid throughput screens using both molecular biology and chemical analyses will greatly aid the search for new enzymes. In the published literature the indications of likely success can already be inferred (see Sect. 4.1 on the use of probes for nitrile hydratase enzymes). Given the fact that many genes can be efficiently expressed in foreign hosts, there will increasingly be no need to develop new processes employing the original strains for generating useful amounts of a biocatalyst for studies on their biotransformation potential. One area that must be a priority for more attention is that of fungal nitrile hydratases, nitrilases, and cyanide hydratases. Given the clear indications that the activities exist, it is surprising how little research has been apparently focused on these organisms. Perhaps the perceived view that fungi are difficult to grow in laboratory culture explains this situation. If this is the case, it is largely misconceived, given the notable range of biotechnological processes that use fungi as a source of biocatalysts. For example, it has recently been shown that fungi possess the capability of performing one of the most valuable types of chemical syntheses known (OIKAWAet al., 1995). A biologically based Diels-Alder reaction has been sought for many years, and yet again the fungi have

314

6 Nitriles

been the first microbes in which such an activity has been found. By acquiring a wider range of nitrile biotransforming enzymes it should be possible to both extend immediately the range of biotechnological applications that are possible, and aid the design of novel enzymes and whole cell systems from the gene banks obtained.

6.2 Redesign of Existing Enzymes by Protein Engineering This area of biochemistry is one of the most exciting both with regard to the understanding of protein structure and function, and the generation of new biotechnological products. As has been reported in Sects.4.1 and 4.2, it is becoming possible to apply some of the most sophisticated techniques to nitrile transforming enzymes as more are purified and crystallized for structural studies. BORK and KOONIN (1994) have shown that there are significant protein sequence similarities between nitrilases, cyanide hydratases, and aliphatic amidases, especially nitrilases that have distinct substrate specificities.They claim this as a new family of carbon-nitrogen hydrolases and have identified both an invariant cysteine residue and a highly conserved glutamate residue as being important in the catalytic mechanism of the enzymes. As more enzymes of this putative group are studied these similarities in protein structure are being confirmed (BROWN et al., et al., 1992;LEVYSCHIL et al., 1995;BHALLA 1995; Novo et al., 1995).These data will eventually reveal which structural properties of these proteins are responsible for the kinetic properties of nitrile transforming enzymes. Coupled with studies that are identmng, for example,the factors that enable biocatalysts to operate at higher temperatures and have better stability in organic solvents, a new generation of enzymes drawing on the natural gene pool for their design, should be obtainable. One example that illustrates this potential has recently been reported. DUFOURet al. (1995) have engineered a peptide nitrile hydratase activity into the cysteine protease papain. It shows what can be achieved when a protein has been extensively studied and its structure

characterized. It is noteworthy that these workers propose that in addition to the cysteine residue, a glutamate and a histidine side chain may also be involved in the catalytic mechanism.

6.3 Metabolic Engineering for the Production of Multistep Biotransformations Involving Nitrile Substrates or Intermediates Many of the genes coding for nitrile hydrolyzing enzymes have been identified and characterized. In many cases nature has already arranged the genes coding for enzymes that convert nitriles into metabolizable intermediates, or useful commercial products. For bioremediation applications of nitrile biotransforming technology the use of mixed cultures, rather than a “super microbe” containing multiple heterologous genes, is likely to remain the norm for some time (e.g., see LISTERet al., 1996). However, when using combinations of biocatalysts to synthesize chemicalsthere is often an advantage in having control over the expression of each enzyme involved to simplify the process design and maximize productivity. An elegant example of this is the synthesis of hydromorphone from morphine. BRUCEet al. (1995) have shown how careful engineering of a pathway can be exploited for biosynthesis. Their work reveals many of the important features that have to be taken into account when designing a recombinant system.

6.4 Final Comments Overall there has been a great deal of progress in determining the biocatalysts that are naturally available for the transformation of the nitrile group. Many uses and potential uses have been found and there is every reason to expect that this range will increase extensively over the next few years.As additional information is gained on the structure-function relationships in such biocatalysts, especially the catalytic mechanisms involved, there is the distinct possibility that nitrile biotechnology

7 References

will lay many of the guidelines needed to develop biotransformation systems.A lot is owed to the sustained effort of those who have worked in this exciting area of biotechnology.

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Chem. 239,42574260. serotina, Plant Physiol. 106,437-445. RODGERS,P. B. (1981), Cyanide degradation by SWAIN,E., LI, C. P., POULTON, J. E. (1992), DevelopChromobacterium violaceum, in: Cyanide in Bioment of the potential for cyanogenesis in maturlogy (VENNESLAND, B., CONN,E. E., KNOWLES, C. ing black cherry (Prunus serotina Ehrh.) fruits, J., WESTLEY, J., WISSING,F., Eds.), pp. 301-310. Plant Physiol. 98,1423-1428. London, New York: Academic Press. TANI,Y., KURIHARA, M., NISHISE,H., YAMAMOTO, K. RODGERS,P. B. (1982), Cyanide metabolism and p(1989a). Bioconversion of dinitrile to mononicyanoalanine formation by washed, non-prolifetrile, a transexamic acid intermediate, by Corynerating cultures of Chromobacterium violaceum: bacterium sp., Agric. Biol. Chem. 12,3143-3149. studies with radiolabeled cyanide, J. Gen. Micro- TANI,Y., KURIHARA, M., NISHIISE,H. (1989b). Charbiol. 128,2983-2989. acterization of nitrile hydratase and amidase, ROLLINSON, G., JONES,R., MEADOWS, M. P., HARRIS, which are responsible for the conversion of diniR. E., KNOWLES, C. J. (1987),The growth of a cyatriles to mononitriles, from Corynebacterium sp., nide-utilizing strain of Pseudomonas fluorescens Agric. Biol. Chem. 12,3151-3158. in liquid culture on nickel cyanide as a source of TAPPER,B. A.. MACDONALD, M. A. (1974), Cyanonitrogen, FEMS Microbiol. Lett. 40,199-205. genic compounds in cultures of a psychrophilic SOLOMONSON, L. P, SPEHAR, A. M. (1979), Stimulabasidiomycete, Can. J. Microbiol. 20,563-566. tion of cyanide formation by ADP and its pos- THIMANN, K . V., MAHADEVAN, S. (1964). Nitrilase I. sible role in the regulation of nitrate reductase. J. Occurrence, preparation and general properties of Biol. Chem. 254,2176-2179. the enzyme,Arch. Biochem. Biophys. 105,133-141. SOLOMONSON, L. P., VENNESLAND, B. (1972), Proper- THOMPSON, L. A., KNOWLES, C. J., LINTON,E. A., ties of a nitrate reductase of Chlorella, Biochim. WYATT,J. M. (1988), Microbial biotransformations of nitriles, Chem. Br. (Sept.), 900-902. Biophys. Acta 267,544-557.

7 References

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TOPP,E., XUN,L., ORSER. C. S. (1992), Biodegrada- WARD,E. W. B. (1964). On the source of hydrogen tion of the herbicide bromoxynil (3,5-dibromo-4cyanide in cultures of the snow mold fungus, Can. J. Bot. 42,319-327. hydroxybenzonitrile) by purified pentachloropheA. N. no1 hydroxylase and whole cells of Flavobacteri- WARD,E. W. B., THORN,G. D., STARRATT, um sp. strain ATCC 39723 is accompanied by cya(1971),The amino acid source of HCN in cultures nogenesis, Appl. Environ. Microbiol. 58,502-506. of a psychrophilic basidiomycete, Can. J. MicrobiTOURNEIX, D., THIERY,A., MAESTRACCI, M., AR01. 17,1061-1066. J. R. NAUD,A.,GALZY,P. (1986), Regulation of nitrile- WARD,E. W. B., STARRATT,A. N., ROBINSON, hydratase synthesis in a Brevibacterium sp., An(1977),Studies of the pathway of HCN formation from glycine in a psychrophilic basidiomycete, tonie van Leeuwenhoek 52,173-182. Can. J. Bot. 55,2065-2069. TSUJIMURA, M., ODAKA,M., NAGASHIMA, S.,YOHDA, C. A. (1994). BiotransA. M., FEWSON. M., ENDO,I. (1996), Photoreactive nitrile hydrat- WARHURST, formations catalyzed by the genus Rhodococcus, ase: The photoreaction site is located on the aCrit. Rev. Biotech. 14,29-73. subunit,J. Biochem. 119,407-413. E. G. J. c., VAN DENNIEUWENDIJK, A. VANDER WALT,J., BREWIS, E.A.,PRIoR,B. A. (1993), WARMERDAM, M. C. H., KRUSE,C. G., BRUSSEE, J. (1996). SyntheA note on the utilization of aliphatic nitriles by sis of (R)-2-hydroxy-3-enoic and (S)-2-hydroxyyeasts, Syst. Appl. Microbiol. 16,330-332. 3-enoic acid esters, J. R. Neth. Chem. SOC. 115,20. VANDER WERF, M. J., VAN DER TWEEL,W. J. J., KAMPPHUIS,J., HARTMANS, S.. DE BONT,J. A. M. (1994), WATANABE,I., SATOH, Y. (1980), Brit. Patent 2 045 747 A. The potential of lyases for the industrial producI., SATOH,Y., TAKANO, T. (1979), Brit. tion of optically active compounds, TZBTECH U, WATANABE, Patent 2 018 240 A. 95-103. I., Satoh,Y.,Takano,T. (1981a). Brit. PaVAN SCHARRENBERG, G. J. M., SLOOTHAAK, J. B., WATANABE, tent 2 076 820 A. KRUSE,C. G., SMITSKAMPWILMS, E., BRUSEE,J. I., SATOH,Y., TAKANO, T. (1981b), Brit. (1993), The potential of (R)-oxynitrilases for the WATENABE, Patent 2 076 821 A. enzymatic synthesis of optically active cyanohyI., YOSHIAKI, S., ENOMOTO, K. (1987a), WATANABE, drins, Med. Chem. 32,1619. Screening, isolation and taxonomical properties C. J. VAUGHAN, P. A,, CHEETHAM. P. S. J., KNOWLES, of microorganisms having acrylonitrile-hydrating (1988), The utilization of pyridine carbonitriles activity, Agric. Biol. Chem. 51,3193-3199. and carboxamides by Nocardia rhodochrous I., SATOH,Y., ENOMOTO,K., SEKI,S., WATANABE, LL100-21, J. Gen. Microbiol. 134,1099-1 107. SAKASHITA, K. (1987b), Optimal conditions for VAUGHAN, P. A,, KNOWLES, C. J., CHEETHAM, P.S. J. cultivation of Rhodococcus sp. N-774 and for con(1989), Conversion of 3-cyanopyridine to nicotinversion of acrylonitrile to acrylamide by resting ic acid by Nocardia rhodochrous LL100-21, cells, Agric. Biol. Chem. 51,3201-3206. Enzyme Microb. Technol. 11,815-823. S., DENGXI,W., GUANGLIN, Y. (1989), VENNESLAND, B., PISTORIUS, E. K., GEWITZ,H. S. WEIQIANG, Studies on Pseudomonas chlororaphis 102 having (1981), HCN production by micro-algae, in: Cyaacrylonitrile-hydrating activity, Znd. Microbiol. B., CONN,E. E., nide in Biology (VENNESLAND, 19,15-18. KNOWLES, C. J., WESTLEY, J., WISSING, F., Eds.), pp. WISSING, F. (1975), Cyanide production from glycine 349-363. London, New York: Academic Press. by a homogenate from a Pseudomonas species, J. I? A,, CONN,E. E., SOLOVENNESLAND, B., CASTRIC, Bacteriol. 121,695-699. J. (1982), Cyanide meMONSON,M. V., WESTLEY, WISSING, F. (1983), Anaerobic column chromatogratabolism, Fed. Proc. 41,2639-2648. WAJANT,H.,FORSTER, S., SELMAR, D., EFFENBERGER, phy in the presence of detergents and its application to a bacterial HCN-producing enzyme, J. MiF., PFIZENMAIER, K. (1999, Purification and charcrobiol. Methods l , 31-39. acterization of a novel (R)-mandelonitrilase lyase K. S. (1981),The enzymology from the fern Phlebodium aureum, Plant Physiol. WISSING,F.,ANDERSEN, of cyanide production from glycine by a Pseudo109,1231-1238. monas species. Solubilization of the enzyme, in: WANG,P.,VANETTEN,H. D. (1992), Cloning and proCyanide in Biology (VENNESLAND, B., CONN,E. perties of a cyanide hydratase gene form the E., KNOWLES, C. J., WESTLEY, J., WISSING. F., Eds.), phytopathogenic fungus Gloeocercospora sorghi, pp. 275-288. London, New York: Academic Press. Biochem. Biophys. Res. Comm. 187,1048-1054. B., KULA,M. R. (1992), WANG,P., MATTHEWS, D. E.,VANETTEN,H. D. (1992), WOKER,R., CHAMPLUVIER, Purification of (S)-oxynitrilase from Sorghum biPurification and characterization of cyanide hycolor by immobilized metal ion affinity chromadratase from the phytopathogenic fungus Gloeotography on different carrier materials, J. Chrocercospora sorghi, Arch. Biochem. Biophys. 298, matogr. 584,85-92. 569-575.

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W R A Y , ~DAVIS, ., R. H., NAHRSTEDT,A.(1983),Bio- YAMAMOTO, K., OISHI, K., FUJIMATSU, I., KOMATSU, synthesis of cyanogenic glycosides in butterflies K.-I. (1991), Production of R-( - )-mandelonitrile from mandelonitrile by Alcaligenes faecalis ATCC and moths: incorporation of valine and isoleucine 8750, Appl. Environ. Microbiol. 57,3028-3032. into linamarin and lotaustralin by Zygaena and YAMAMOTO, K., OTSUBO, K., MATSUO,A., HAYASHI, Heliconius species, Z. Naturforsch. 38C, 583-588. T., FUJIMATSU, I., KOMATSU, K.-I. (1996),ProducWYAIT, J. M., KNOWLES, C. J. (1995),The develoption of R-(-)-ketoprofen from an amide comment of a novel strategy for the microbial treatpound by Comamonas acidovorans KPO-2771-4, ment of acrylonitrile wastes, Biodegradation 6, Appl. Environ. Microbiol. 62,152-155. 93-107. YAMADA, H., ASANO,Y., HINO,T. (1979), Microbial YANASE,H., SAKAI,T.,TONOMURA, K. (1982a),Purification and crystallization of a p-cyanoalanine utilization of acrylonitrile, J. Ferment. Technol. 57, forming enzyme from Enterobacter sp. 10-1, Ag8-14. ric. Biol. Chem. 46,355-361. YAMADA,H., ASANO,Y., TANI,Y. (1980), Microbial K. (1982b).Some utilization of glutaronitrile, J. Ferment. Technol. YANASE,H., SAKAI,T.,TONOMURA, 58,495-500. properties of a P-cyanoalanine forming enzyme from Enterobacter sp. 10-1,Agric. Biol. Chem. 46, YAMAGUCHI, Y., WATANABE. I., SATOH,Y. (1980), 363-369. Brit. Patent 2 048 877 A. K. (1983),PurifiYAMAGUCHI, Y., WATANABE, I., SATOH, Y. (1981). YANASE,H., SAKAI,T.,TONOMURA, cation, crystallization and some properties of pBrit. Patent 2 054 563 A. YAMAMOTO, K., KOMATSU, K.-I. (1991),Purification cyano-L-alanine-degradingenzyme in Pseudomoof nitrilase responsible for the enantioselective nus sp. 13, Agric. Biol. Chem. 47,473-482. hydrolysis from Acinetobacter sp. AK 226, Agric. YANASE,H., SAKAI,T.,TONOMURA, K. (1985).MicroBiol. Chem. 55,1459-1466. bial metabolism of nitriles in Pseudomonas sp.,J. YAMAMOTO, K.. UENO,Y., OTSUBO, K., KAWAKAMI, Ferment. Technol. 63,193-198. M., SUGAI. T., OHTA,H. (1993),AsymK., KOMATSU, K.4. (1990),Production of S-( +)- YOKOYAMA, metric hydrolysis of a disubstituted malononitrile ibuprofen from a nitrile compound by Acinetoby aid of a microorganism, Tetrahedr0n:Asymmebacter sp. strain AK226. Appl. Environ. Mitry 4,1081-1084. crobiol. 56,3125-3129.

Redox Enzymes


7 Alkaloids

NEIL C. BRUCE Cambridge, UK

1 Introduction 328 2 Tropane Alkaloids 332 2.1 Tropane Alkaloid Biosynthesis 332 2.2 Microbial Metabolism of Tropane Alkaloids 335 3 Benzylisoquinoline Alkaloids 338 3.1 Benzophenanthridine Alkaloids 338 3.2 Morphine Alkaloids 339 3.2.1 Morphine Alkaloid Biosynthesis 342 3.2.2 Microbial Metabolism of Morphine Alkaloids 344 3.2.3 Transformations of Morphine Alkaloids by Pseudomonas putida M10 346 3.2.4 Biological Production of Hydromorphone and Hydrocodone 350 3.2.5 Microbial Transformation of Heroin 351 4 Monoterpenoid Indole Alkaloids 352 5 Summary 356 6 References 357

328

7 Alkaloids

1 Introduction

ever, results from a recent examination of materials from the tomb of the Royal Architect KHAseem to refute earlier observations (BISPlants have the ability to produce tens of SET et al., 1994).It was not until the 19th centuthousands of highly complex secondary me- ry that the active compounds, alkaloids, were tabolites to assist their survival in the environ- isolated from the opium. Morphine was also ment, many of which protect the plant from the first alkaloid to be identified and crystalin 1805. This predators. Man has exploited these com- lized by the chemist SERTURNER pounds of self-defence as sources of medicinal was a significant achievement as not only was agents, poisons, and potions since time imme- it the first time that a nitrogenous base had morial. Throughout the world different com- been isolated from a biological source, but it munities have discovered plants with phar- xas also the first time that such a substance ad been shown to be intrinsically basic. This macological properties, and many useful drugs have their origins in indigenous ethnop,iarma- finding formed the basis of one of the earliest cologies. Some notable examples include: the definitions of an alkaloid which was attributed (HESSE,1981; roots of the mandrake plant, known for their to the pharmacist W. MEISSNER sedative properties from the time of HIPPO- PELLETIER,1983). The two authors, HESSE (1981) and PELLETIER (1983), differ on the exCRATES (ca. 400 BC) and also used as a deadly poison during Elizabethan times (MANN, act date of the coining of the term (1818 and 1989);the leaves of the coca plant, which were 1819 being cited, respectively) and the derivachewed as an aid to stamina and as part of ce- tion, but the general meaning was taken to be remonies in South America over 5000 years an “alkali like” compound of plant origin, or as ago (VANDYKEand BYCK,1982); and plants BENTLEY(1954) interpreted, a “vegetable alwhose hallucinogenic properties were used in kali”. This was extended by WINTERSTEIN and the preparation of “magic potions” by the Az- TIET(1910) to include a four part definition tec Indians. The compounds responsible for stating a “true alkaloid” can be characterized these physiological effects in man were isolat- by: (1) the possession of a nitrogen atom as ed during the 19th and early 20th centuries and part of a heterocyclic system; (2) a complex were identified as scopolamine, cocaine, and molecular structure; (3) significant pharmacoamides of lysergic acid, respectively. SOCRA- logical properties; (4) its origin from the plant kingdom (cited in PELLETIER, TES’ death in 399 BC was the result of con1983). sumption of hemlock (Conium maculatum) The majority of alkaloids fit this four part which contains the alkaloid coniine (1) (Fig. 1; definition; however, a number of exceptions HENDRICKSON et al., 1970), while CLEOPATRA exist.The compounds samandarine (2) (Fig. l ) , used extracts from Egyptian henbane samandarone, and cycloneosamandarine, iso(Hyoscyamus muticus) during the last century lated from the skin glands of the European fire BC to dilate her pupils and increase her beau- salamander (Salamandra maculosa Laurenti) ty. Likewise, medieval European women used all exhibit the usual properties of an alkaloid extracts of deadly nightshade (Atropa bella- substance, but do not fit the definition of a donna) in their beauty preparations, hence the “true alkaloid” owing to their animal origin. name bella donna, “fair lady”. There are numerous examples of alkaloids furOther historical uses include extracts from nished by animal, including batrachotoxinin A the bark of Cinchona officinalis which have (3), a steroidal alkaloid from the Colombian been employed as antimalarials. Extracts de- arrow-poison frog (Phyllobates aurotaenia), rived from the opium poppy Papaver somni- bufotenine (4), a tryptamine-type alkaloid ferum comprise another group of important from the common European toad, ( - )-deoxypharmacologically active compounds which nuphradine and (-)-castoramine (5) from the possess powerful analgesic properties. It has Canadian beaver (Castorfiber L.), and muscobeen reported that extracts of the milky latex pyridine (6) from the scent gland of the musk material that exudes from the cut unripe seed deer (Moschus moschiferus). Alkaloids have capsule of the opium poppy were used by the also been identified from arthropod, bacterial, early Egyptians for medicinal purposes; how- and fungal origins. For example, the quinazole

.

I Introduction

1 Coniine

4 Bufotenine

Me 8 Pyocyanine

2 Samandarine

5 Castoramine

H 9 Agroclavine

329

3 Batrachotoxinin A

6 Muscopyridine

OMe 10 Colchicine

7 a R = Me; Glomerine b R = Et; Homoglomerine

Me0 11 Mescaline

Fig. 1. See text.

alkaloids, glomerine (7a) and homoglomerine (7b) discharged from the dorsal glands of the European millipede (Glomeris marginata),the deep-blue colored alkaloid pyocyanine (8), isolated from the bacterium Pseudomonas aeruginosa,and agroclavine (9),produced by the fungi Claviceps purpurea and Aspergillus fumigatus. Other examples of alkaloids exist which also do not adhere to the criteria stipulated in the four part definition of an alkaloid. For example, the alkaloids colchicine (10) (autumn crocus, Colchicurn autumnale L.) and mescaline (11) (Lophophora williamsii) do not possess nitrogen as part of a heterocyclic system.Also colchicine (10) is essentially neu-

tral and, therefore, does not conform to the original definition of an alkaloid. PELLETIER (1983), however, provides a reasonable summary of an alkaloid’s properties as being an “alicyclic compound containing nitrogen in a negative oxidation state which is of limited distribution among living organisms”. Over loo00 compounds fall within this definition (SOUTHON and BUCKINGHAM, 1989) and new alkaloids are continually being reported from various sources. These represent approximately 20% of all known natural products: however, only about 30 of these with biological activity are commercialized (FARNSWORTH, 1990).

330

7 Alkaloids

Unlike any other group of compounds the alkaloids exhibit a vast array of skeletal types and are classified accordingly. A typical example is the scheme used by HESSE(1981), who describes 11 classes of heterocyclic alkaloids, differentiating by the nature of the carbon skeleton, e.g., the pyrrolidine and isoquinoline alkaloids. The majority of alkaloids are amino acid-derived, although terpenes, steroids, purines, and nicotinic acid can also act as building blocks of, e.g., aconitine, solanidine, caffeine, and nicotine, respectively. If the anabolic route of an alkaloid is known, this can be used to classify the compound (DALTON, 1979). The tropane, and pyrrolidine alkaloids, for instance, are all derived from ornithine, a derivative of arginine, and thus grouped together under this scheme. Alkaloids have provided a wealth of pharmacologically active compounds; approximately 25% of the drugs used today are of plant origin. These are administered either as pure compounds or as extracts and have often served as model structures for synthetic drugs, e.g., atropine (13)for tropicamide, quinine for chloroquinine, and cocaine (12) (Fig. 2) for procaine (KUTCHAN,1995). Screening of plant extracts for pharmacologically active compounds still continues and results in new drug discoveries; recent examples include the anticancer drugs taxol from the western yew, Taxus brevifofiu,and camptothecin from Cumptotheca ucuminatu.Alkaloids are generally regarded as speciality chemicals; approximately 300-500 metric tons of quinine and quinidine are produced each year; ajmalicine (98) production amounts to about 3600 kg, while compounds like vincristine (94) and vinblastine (95) (Fig. 21) are produced in the kilogram range. The annual market value of the major alkaloids has been estimated to be in the range of several hundred million dollars (VERPOORTE et al., 1993). The important pharmacological activity of many alkaloids has spurred chemists to make many derivatives of these natural compounds. The chemical preparation of such semisynthetic alkaloids has resulted in the production of drugs with improved properties, such as the addition of a 1Chydroxy group to the morphine alkaloid structure which has been found to dramatically increase potency (JOHNSON

12 Cocaine

0

13 Hyoscyamine (Atropine is racemic hyoscyamine)

14 Scopolamine

Fig. 2. Examples of tropane alkaloids.

and MILNE,1981). However, the synthesis of such compounds is often difficult to achieve on a commercial scale due to the chemical complexity of the starting material, cost, and environmental issues, in addition to the precursors being in limited supply. Biotransformations can offer a number of advantages over conventional chemical processes. The specificity of enzyme-catalyzed reactions, e.g., allows the stereospecific transformation of defined functional groups. However, biotransformations of alkaloids, unlike their steroid counterparts, have yet to meet their potential on an industri-

I Introduction

a1 scale. This is in part due to the lack of suitable enzymes and partly because no alkaloidbased drug commands a significant share of the therapeutic market, unlike the steroids. The rate at which new drugs derived from natural products are entering the therapeutic market has declined significantly over recent periods in contrast to synthetic molecules, possibly due to the difficulty of modifying these often complex chemicals for the development of new drugs and the difficulty of producing these natural products in a pure form. The difficulties associated with the development of new drugs are being addressed by the ever increasing interplay of chemistry and biology. Undoubtedly combinatorial approaches (AMATO,1992) and genetic engineering will play an important role in the development of new drugs. The use of recombinant DNA technology is beginning to have substantial impact on biotransformation processes, resulting in the development of new approaches using biological systems. Recent advances in the understanding of the genetic and biochemical basis of alkaloid biosynthetic pathways are now beginning to make biotransformations of complex alkaloid molecules more plausible. The expression of plant enzymes, which are often present at very low levels in the plant, in heterologous hosts such as bacteria allows detailed examination of mechanisms of reaction which are often unknown in synthetic organic chemistry. It is possible to add to the genetic repertoire of a plant by incorporating genes from other species allowing the possibility of producing unique compounds with potential biotechnological applications. Microorganisms have been used for the large-scale production of high-value chemicals for many years, and the use of microbial processes to make analogs of naturally occurring alkaloids is achievable. It is now possible to assemble hybrid transformation pathways in microbes using structural genes cloned from different organisms which mediate enzymic processes which are not indigenous to the host organism. These “patchwork” pathways can have the advantage of removing unwanted side reactions, they allow the possibility of increasing the activity of a cell by altering regulatory processes, and avoid the need to supply expensive exogeneous co-

331

factors. Furthermore, it is now theoretically possible to manipulate alkaloid biosynthetic pathways to improve yields and to extend pathways to synthesize new bioactive molecules. Metabolic engineering of plants offers the capability of altering the pattern of alkaloid accumulation in the plant; in addition, the ability to house and express recombinant genes in plants from other organisms offers the potential of both extending pathways and allowing the biological synthesis of semisynthetic derivatives. It is now possible to design strategies to alter the metabolic flux in a variety of organisms, such as the introduction of extra copies of genes encoding enzymes which form bottlenecks in pathways affords a way to attain increases in yields of plant secondary products, or more globally through the expression of one or more regulatory genes (for reviews see BAILEY,1991; NESSLER, 1994; HUTCHINSON, 1994; KUTCHAN, 1995). Due to their complex structures, alkaloids are still most efficiently produced by the plant and the future success of metabolic engineering of plant secondary products is dependent on having a good understanding of the biochemistry and regulation of the pathways under consideration. Plant cell culture has been invaluable as a means of providing suitable biomass for the elucidation of pathways for secondary metabolites, particularly for alkaloid synthesis. Cell culture has also been examined for biotransformation purposes (reviewed by VERPOORTE et al., 1993) and extensively investigated as a means of producing plant secondary products on a large scale. Unfortunately, the level and manner of production of alkaloids in plants does not necessarily correlate with production in cell cultures. The use of plant cell cultures for biotransformations of alkaloids will not be considered in detail this chapter. The purpose of this review is to introduce some of the newer technologies which are beginning to make an impact on the area of alkaloid transformations. It also aims to introduce some of the more recent developments in the biochemistry and genetic understanding of biosynthetic and catabolic routes of some of the more pharmacologically important alkaloids in different organisms, without which the rational design of any recombinant alkaloid biotrans-

332

7 Alkaloids

formation processes would not be feasible. The alkaloids discussed include the tropane alkaloids, the benzylisoquinoline alkaloids, the benzophenanthridine alkaloids, the morphinan alkaloids, and the monoterpenoid indole alkaloids.

recent study suggested that the alkaloid has insecticidal properties at naturally occurring concentrations due to potentiation of insect octopaminergic neurotransmission (NATHANSON et al., 1993).

2.1 Tropane Alkaloid Biosynthesis The biosynthetic pathways for the tropane alkaloids have been studied in considerable detail and are associated with nicotine biosynThe tropane alkaloids occur in the Solana- thesis, since the N-methyl-A'-pyrrolinium catceae family, but they are also found in the ion (15) is a precursor to both classes of alkaplant families Erythroxylaceae, Convolvula- loids. The formation of the tropane nucleus ceae, Proteaceae, and Rhizophoraceae. Their from ornithine and acetoacetate was first incommon structural element is the azabicyc- vestigated in plants using radioactive tracers as lo[3.2.l]octane system, and over 150 tropane long ago as 1954, but it was not until the early alkaloids have been isolated. The 3-hydroxy 1980s that a biosynthetic scheme was finally aromatie ester derivatives form the parent al- elucidated in Erythroxylon coca (Fig. 3; LEETE, kaloids, examples of which include cocaine 1983). The pathway for biosynthesis of hyos(12) (Erythroxylon coca, coca plant), hyoscy- cyamine (13) and scopolaminc (14) is quite amine (13), (Hyoscyamus niger, henbane), complex, since not only is an acetone unit reatropine (13) (Atropa belladonna, deadly quired for the formation of tropinol(19), but a nightshade), and scopolamine (14) (Scopola second converging pathway is necessary for carniolica) (Fig. 2). It appears that in most the conversion of phenylalanine (20) to tropic cases atropine (13) is formed by racemization acid (23) (Fig. 4). Recent work with root culof hyoscyamine (13) during extraction. tures of Datura stramonium, suggests that hyLong before the elucidation of their struc- grine (17) is not an intermediate, but an off tures, the pharmacological properties of sever- shoot from the main pathway (ROBINSet al., al tropane alkaloids were exploited. Atropine 1997).The biosynthesis of cocaine (12) is simi(13), which typifies the action of tropane alka- lar to that of hyoscyamine (13).The N-methylloids, causes antagonism to muscarine recep- ation and cyclization of ornithine-derived putors (parasympathetic inhibition) (CORDELL, trescine gives the N-methyl-A'-pyrrolinium 1981). cation (W), which condenses with acetoacetylThese receptors are responsible for slowing CoA (16). Methylation of the free carboxylate of the heart rate, vasodilation, dilation of the group followed by ring closure, reduction of pupil, and stimulation of secretions. The heart the ketone group, and benzoylation results in rate altering properties of atropine (13) have the formation of cocaine (12).The benzoic acled to its use in the initial treatment of myocar- id is derived from phenylalanine (20). dial infraction. Tropane alkaloids have also The molecular biology of tropane alkaloid been used to treat peptic ulcers, prevent mo- synthesis is being studied extensively and tion sickness, and as components of pre-an- holds considerable potential for alkaloid bioesthetic drugs. Cocaine (12) is perhaps the transformations. An elegant example is the best known of all the tropane alkaloids mainly construction of a transgenic species of Atropa because of its use as an illicit drug; it is a pow- belladonna that was able to accumulate the erful central nervous system stimulant and ad- important pharmaceutical scopolamine (14) renergic blocking agent, and its hydrochloride instead of hyoscyamine (13) (YUNet al., 1992). salt has been used as a local and surface anes- The final two steps in the pathway for the biothetic in face, eye, nose, and throat surgery synthesis of scopolamine (14) (Fig. 5) are cata(GERALD, 1981).The function of cocaine (12) lyzed by 2-oxoglutarate-dependent hyoscyin leaves of the coca plant was unknown until a amine 6p-hydroxylase (hyoscyamine[6p]-di-

2 Tropane Alkaloids

2 Tropane Alkaloids

@

15 N-Methylpyrrolinium

-.

I

Me

@ NH3

333

20 Phenylalanine

nSCoA 1 0

16 Acetoacetyl-CoA

0

J

,

~

OH

1

Ph

2 1 Phenylpyruvic acid

GPh

17 Hygrine

?

HO

0

18 Tropinone

19 Tropinol

--

HO

2 2 Phenyllactic acid

23 Tropic acid

Fig. 4. Biosynthesis of tropic acid (ROBINSet al., 1994).

t

showed that catalysis of these two steps carried out was by the same enzyme (HASHIMOTO et al., 1987).Analysis of key enzymes of metabolic pathways at the molecular genetic level assists clarification of complex biochemical mechanisms, and hydrolysis and epoxidation of the tropane ring was later confirmed unequivocally by molecular cloning and expression of the structural gene of the 6P-hydroxylFig. 3. Biosynthesis of tropane alkaloids (ROBINS ase in a heterologous host (MATSUDAet al., 1991; HASHIMOTO et al., 1993b);A. belladonna et al., 1994). accumulates hyoscyamine (13) instead of scopolamine (14) because it lacks the 6P-hydroxylase. The cDNA encoding the 6P-hydroxylase oxygenase; EC 1.14.11.11).This enzyme first from H. niger was transferred into Agrobactehydroxylates hyoscyamine in the 6P-position rium tumefaciens and introduced into A. bellaof the tropane ring (24). which is followed by donna.The regenerated transgenic plants were epoxidation. The use of purified 6P-hydroxy- found to contain elevaled levels of scopollase from root cultures of Hyoscyamus niger amine (14) (YUNet al., 1992).The change in 13 Hyoscyamine

334

7 Alkaloids

H yoscyamiiie 6R-hydroxy lase 2-Oxoglutarate Succinate

JJ 13 Hyoscyamine

t

I 0 2

co2

Ascorba te

K

R 24 6R-Hydrosyhyoscyamine

14 Scopolamine

Fig. 5. See text.

alkaloid composition in the transgenic A . bef- eospecificities were found in cultured roots of fadonna was considerable, with scopolamine H. niger (HASHIMOTO et al., 1992). These two distinct enzyme activities reduc(14) being almost the only alkaloid present in the aerial parts of the plant. It was thus posed tropinone (18) to 3a-hydroxytropane (19) sible to isolate pure scopolamine (14) by re- (tropinol, tropine) and 3P-hydroxytropane crystallization of the total alkaloid fraction, in- (pseudotropinol, $-tropine, pseudotropine), stead of conventional differential extraction respectively. Marked differences were oband chromatography. Analysis of expression of served between the two reductases in their afthe 6P-hydroxylase gene by measurements of finities for tropinone (18),substrate specificity, levels of mRNA and Western blot analysis of and in the effects of amino acid modification protein extracts from various tissues showed reagents. The cDNA clones for the two tropithat enzyme expression in scopolamine pro- none reductases have been expressed in Esducing species of Hyoscyamus was lacking in cherichia coli and sequenced (NAKAJIMA et al., the stem or leaves, being localized in the roots 1993). Preparation of various chimeric forms of these plants, and explains why it has not of these two enzymes led to the identification been possible to produce these alkaloids in sig- of the domain conferring the stereospecificity nificant quantities by cell culture (HASHIMOTO of the reaction (NAKAJIMAet al., 1994). et al., 1991;MATSUDA et al., 1991).HASHIMOTO These elegant experiments demonstrate a et al. (1993b) have now engineered transgenic key to future alkaloid biotransformation proA. belladonna hairy root cultures that express cesses by the manipulation of biosynthetic the H . niger gene encoding hyoscyamine 6P- routes in plants with the use of recombinant hydroxylase which exhibited up to 5 times DNA technology not just for alkaloids but also higher activity. These transgenic roots may other secondary products. It is becoming posprove to be useful for enhancing scopolamine sible to design strategies to advantageously productivity in vitro. Recombinant strains of manipulate the metabolic flux in organisms, or Escherichia coli expressing the gene encoding to decrease or increase the production of phyhyoscyamine hydroxylase were also capable of tochemicals; however, with regard to biotranstransforming hyoscyamine (13) to scopol- formations, it is the possibility of manipulating amine (14) (HASHIMOTO et al., 1993a; LAYet pathways by altering enzyme function by dial., 1994). rected mutagenesis and extendinglaltering exTropinone reductase acts at a branch point isting pathways by heterologous gene expresof biosynthetic pathways leading to a variety sion to alter the spectrum of plant alkaloids of tropane alkaloids. It is an NADPH-de- which is particularly challenging and exciting. pendent enzyme which reduces the 3-keto Although the genetic tools for manipulating group of tropinone (18) (ROBINSet al., 1994). biosynthetic pathways in plants lag behind Two tropinone reductases with different ster- those for prokaryotic organisms, the success of

2 Tropane Alkaloids

scopolamine production in transgenic plants will, hopefully, encourage interest and further development.

2.2 Microbial Metabolism of Tropane Alkaloids Microorganisms possess an incredible variety of metabolic pathways which enables them to degrade a plethora of natural and manmade organic compounds. The elucidation of alkaloid dissimulating pathways has considerable potential for the identification of biotransformation routes for new and existing therapeutic compounds (BRUCEet al., 1995). The most extensively studied tropane alkaloid, in terms of microbial metabolism is atropine (13). Several bacterial species have been shown to possess an esterase that catalyzes the esterolytic hydrolysis of the atropine molecule, to form tropinol (19) and tropic acid (23) (Fig. 6). The interest in atropine esterase lies in its similarity to mammalian serine proteases and its use as a possible model of mammalian cholinergic receptors. RORSCHet al. (1971) reported the isolation of a number of Pseudomonas strains capable of utilizing atropine as a sole source of carbon and nitrogen. The atropine esterase from one of these strains, Pseudomonas putida PMBL-1, has been purified and extensively characterized (VAN DER DRIFT, 1983; VAN DER DRIFTet al., 1985a, b, 1987).This esterase showed activity with both enantiomers of hyoscyamine (13), but not with cocaine (12) (RORSCHet al., 1971),despite the close similarity of structure of those compounds. NIEMERet al. (1959) and NIEMERand BUCHERER(1961) reported a breakdown route of atropine (13) by Corynebacterium belladonnae, which involves the formation of tropinol (19) and tropic acid (23) by esterase action, followed by dehydrogenation, ring opening, and deamination of the tropane ring (Fig. 6). The first step in their proposed route of tropine catabolism involves a tropine dehydrogenase. Activity, however, was only demonstrated in the reverse direction. The step postulated by NIEMERand BUCHERER(1961) which follows tropinone formation involves

335

ring cleavage and the formation of tropinic acid (25), though no enzymes or cofactors were identified. Isolation of a picrate derivative of methylamine from whole cell incubations with tropinone, indicated that nitrogen debridging was taking place. More recent investigations into the microbial metabolism of atropine showed that a strain of Pseudomonas sp. (termed AT3) isolated from the rhizosphere of atropine plants was able to utilize tropinol(l9) as a sole carbon and nitrogen source (LONG et al., 1993). Growth studies revealed a diauxic growth pattern. When this organism was supplied with atropine (13) and an exogenous nitrogen source, tropic acid (23) was utilized during the first phase of growth and the heterocyclic moiety, tropinol(19), was utilized in the second. The enzymes responsible for tropinol (19) degradation appeared to be strongly repressed during the first phase of growth. Under nitrogen limitation, however, the nitrogen must be stripped from the tropane ring before growth can occur and under these conditions tropinol (19) was utilized in the first growth phase. Pseudomonas sp. AT3 initiated the degradation of tropinol (19) by attacking the nitrogen atom, yielding a dinitrophenyl hydrazine positive intermediate, identified as 6hydroxycyclohepta-l,4-dione(28), which was oxidized by an NAD +-dependent dehydrogenase activity to cyclohepta-1,3,5-trione (29). The subsequent cleavage of this compound resulted in the formation of 4,6-dioxoheptanoic acid (succinylacetone) (30) which was, in turn, the substrate for a second hydrolase yielding succinate (31) and acetone(32) (BARTHOLOMEW et al., 1993,1996). BARTHOLOMEW et al. (1995) identified an NADP+-dependent tropinol dehydrogenase in cell-free extracts of Pseudomonas sp. AT3 that was induced by growth on atropine (13), tropinol(l9) or tropinone (18).The product of the reaction was tropinone (18) and the reaction was shown to be freely reversible.The dehydrogenase showed activity only with tropino1 (19) and nortropinol (27); no activity was detected with a number of closely related compounds including atropine (13), scopine, pseudotropinol, ecgonine (34)(Fig. 7) and 6-hydroxycyclohepta-1,4-dione (28), which suggests thal this inducible enzyme is involved in the metabolism of tropinol(l9) in Pseudomo-

7 Alkaloids

336

13 Atropine

23 Tropic acid

19 Tropinol

I

18 Tropinone

25 Tropinic acid

[?!

COZH

31 Succinate

b&-5? "" OH 6-Hydroxycyclo-

dehydrogenase

28 6-Hydroxycyclohepta1,4-dione

acid

(t4H

27 Nortropinol

0

26 2,6-Dioxopimelic

0

32 Acetone

4,6-Dioxoheptanoate hydrolase H2O

Cyclohepta-l,3,5trione hydrolase

29 C yclohepta-l,3,5-trione

0

30 4,6-Dioxoheptanoic acid

Fig. 6. Proposed pathway for the degradation of atropine (13)by Pseudornonas sp.AT3 (BARTHOLOMEW et al., 1996).---* Pathway proposed by NIEMERand BUCHERER (1959) for C. belludonna.

2 TropaneAlkaloids

337

MeN nus sp. AT3. However, the occurrence of 6-hydroxycyclohepta-1,4-dione(28) during the metabolism of tropinol seemed to dispute this (BARTHOLOMEW et al., 1993).An elegant set of experiments with tropinol (19) and pseudotropinol labeled with deuterium in the C-3 position provided an answer (BARTHOLOMEW et esterase 12 Cocaine al., 1995).The labeled alcohol group at C-3 was shown to remain intact past the point of rePhCO2H moval which indicated that tropinone (18) is not an intermediate in the pathway of tropinol C02Me (19) metabolism.What then is the role of tropinol dehydrogenase in the metabolism of trop3 3 Ecgonine inol? Tropinone (18) serves as a growth subOH methyl ester strate for Pseudomonas sp. AT3 and it is likely to be encountered in nature, along with atroI Ecgonine H20 pine (13) and tropinol (19), as it is an intermemethyl diate in the biosynthesis of the tropane alkaesterase loids in plants (LANDGREBE and LEETE,1990). MeOH A mutant strain of Pseudomonas sp. AT3 blocked in 6-hydroxycyclohepta-1,4-dionedehydrogenase activity was grown on tropinone; this resulted in the accumulation of 6-hydroxycyclohepta-1,4-dione (28), an indication that tropinone (18) is metabolized via the same route as tropinol(l9) and that its keto group is reduced in the process. Thus, the tropinol deEcgonine hydrogenase may function primarily as a reepimerase ductase in order to channel tropinone (18) and nortropinone into the pathway of tropinol(l9) metabolism in Pseudomonas sp. AT3. The elucidation of the pathway for microbial metabolism of the related alkaloid cocaine (U) has proven to be slightly more elusive. A strain of Pseudomonas maltophilia (termed MBllL) was isolated from samples taken in Pseudoand around a pharmaceutical company that ecgonyl processes cocaine. Z? maltophilia M B l l L was capable of utilizing cocaine (12) as its sole synthetase source of nitrogen and carbon for growth. The bacterium possessed an inducible cocaine esterase which converted cocaine (12)to ecgonine methyl ester (33) (Fig. 7), and benzoic acid. Both degradation products supported COSCoA growth of Z? maltophilia MBllL, although only cocaine induced high activities of the coOH caine esterase (BRITTet al., 1992).Benzoic acid was further metabolized via catechol and 36 Pseudoecgonyl CoA the 3-oxoadipate pathway; however, the pathway for the metabolism of ecgonine methyl es- Fig. 7. Proposed pathway for the metabolism of coter (33) was not further elucidated. caine (12)by k? fluorescens MBER and C. acidovor-

k 1

It

ans MBLF (LISTERet al.. 1995).

338

7 Alkaloids

A mixed microbial culture was also isolated there are over 2500 compounds known to be from the same site as Z? maltophilia MBllL, derived from the alkaloid (S)-reticuline (43) comprising a strain of Pseudomonas jluores- (Fig. 8) (KUTCHAN et al., 1991) and a number cens (termed MBER) and a strain of Coma- of these, such as morphine (52)(Fig. 10) and its monas acidovorans (termed MBLF) which derivatives, have significant commercial value. was able to utilize cocaine as a sole source of (S)-Reticuline (43) is the pivotal intermediate carbon and nitrogen for growth. Fig. 7 shows for the synthesis of an amazing variety of difthe proposed initial degradative route of the ferent alkaloid families including: morphinans, tropane moiety of cocaine by this mixed cul- protoberberines, phthalide isoquinolines, proture. Analysis of the growth substrate spect- topines, and benzophenanthridines. The enrum of the individual components of the zymes mediating the pathway from tyrosine to mixed culture indicated a mutualistic relation- (S)-reticuline (43) have been described in ship between the two microorganisms. C. aci- some detail (Fig. 8; FRENZEL and ZENK,1990). dovorans MBLF was shown to possess an in- The pathway is initiated by the modification of ducible cocaine esterase which calalyzed the two molecules of tyrosine to form dopamine hydrolysis of cocaine (12)to ecgonine methyl (37) and 4-hydroxyphenylacetaldehyde (38) ester (33) and benzoic acid. F! fluorescens (LOEFFLERet al., 1987; STADLERet al., 1988). MBER was then able to utilize these two hy- Tyrosine decarboxylase is thought to be indrolysis products, reducing their concentra- volved in providing precursors for alkaloid tion, and thus enabling C. acidovorans to grow synthesis in the opium poppy (Papaver somnion lower and less inhibitory levels of benzoate. ferum L.). cDNAs for the enzyme have been P fluorescens MBER was observed to initiate isolated and expressed in E. coli (FACCHININ the metabolism of ecgonine methyl ester (33) and DE LUCA,1995);a genomic clone of the tyby hydrolysis yielding ecgonine (34) which was rosine decarboxylase gene has been obtained further metabolized via pseudoecgonine (35) which could also be expressed in E. coli, since and pseudoecgonyl-CoA (36). Subcellular the gene lacked any introns which facilitated studies resulted in the identification of an ec- its expression (MALDONADO-MENDOZA et al., gonine methyl esterase, an ecgonine epimer- 1996). Bacterial cell cultures expressing rease, and pseudoecgonyl CoA synthetase which combinant tyrosine decarboxylase were able were induced by growth on ecgonine methyl to accumulate tyramine and dopamine (37) in ester (33) or ecgonine (34). Further metabo- the medium after the addition of exogenous lism of the ecgonine moiety was postulated to tyrosine and dopa, respectively. involve debridging with the production of a carbonyl containing intermediate (LISTERet al., 1995).The pathway for the degradation of ecgonine (34) is different from that of the related tropine alkaloid, atropine (13), in 3.1 Benzophenanthridine Pseudomonas AT3 (BARTHOLOMEW et al., Alkaloids 1993,1996) where the melabolism of tropinol (19) is initiated by attack at the nitrogen atom. The benzo[c]phenanthridine alkaloids are a specific group of isoquinoline alkaloids which are found mainly in the plants belonging to the Papaveraceae family. These alkaloids have provided active ingredients for treatments of gastric ulcers, warts, papillomas, and condylomas (KUTCHAN, 1994). The initial steps of the biosynthetic pathway for the benzo[c]phenanthridine alkaloid, sanguinarine (51),have been The benzylisoquinoline alkaloids are an ex- studied in some detail by ZENKand his tremely large and wide ranging group of alka- coworkers and are summarized in Fig. 9 (for loids which possess a diverse range of phar- reviews see KUTCHANet al., 1991; KUTCHAN, macological properties. Within this family 1994; ZENK, 1994).

3 Benzylisoquinoline A1kaloids

m y

3 Benzylisoquinoline Alkaloids

339

HopNH* -

HO

HO

37 Dopamine

HO

1

2

H '

~

/

HO

/

HO

@

39 (S) - N orcoc la urine

HO 38

4'-Hydroxyphenylacetaldehyde

Me0 H HO

Me0

-

40 ( S ) - C o c l a u r i n e

:-loMe-H:Tce

Me0

0

3

MH "'

A/ 4

43 (S)-Reticuline

e

5

Me0

\

4

HO

/

HO

41

42 (S)-3'-Hydroxy-Nmethylcoclaurine

4'

/

4 1 (S) -N - M eth y 1coc la uri ne

1 (S)-Norcoclaurine synthase 2 (S)-Adenosyl-L-methionine: (S)-norcoclaurine-6-0 -methyltransrerase 3 (S)-Adenosyl-L-methionine: (S) -coclaur ine-N -met h y 1trans f erase, 4 Phenolase, 5 (S)-Adenosyl-L-methionine: Fig. 8. Biosynthesis of the tetrahydrobenzylisoquinolinealkaloid (S)-reticuline (43) (KUTCHAN,1994).

The first step in the pathway to sanguinarine (51), is catalyzed by the berberine bridge enzyme (Fig. 9). This enzyme is of particular interest to chemists as it catalyzes the oxidative cyclization of the N-methyl group of (S)reticuline (43) to form the berberine bridge carbon, C8, of (S)-scoulerine (44).The stereochemistry of this transformation has recently been elucidated by BJORKLUND el al. (1995). cDNA encoding the berberine bridge enzyme was isolated from an Eschscholtzia californica cDNA library ( D I ~ R I Cand H KUTCHAN, 1991, 1995).The enzyme was not expressed in an ac-

tive form in E. coli; however, expression in yeast resulted in low levels of activity.The berberine bridge enzyme was found to be 0,dependent but did not have any requirement for soluble cofactors. There is a commercial need for the sanguinarine (51), which has antimicrobial activities.

3.2 Morphine Alkaloids The morphine alkaloids are perhaps the most studied group of alkaloids owing to their

340

7 Alkaloids

-

HO

’ ‘

43 (S)-Reticuline

44 (S) - S c o u 1e r i n e

48 P r o t o p i n e

47 (S)-cis-N-methylstylopine

t

7%?( 0

’I

‘

0

0

’I

45 ( S ) - C h e i l lan t h i f ol in e

46 (S)-Stylopine

O)

49 6 - H y d r ox ypro to pi n e

50 Di h y d r o s a n g u i n a r i n e

51 S a n g u i n a r in e

Berberine bridge enzyme, 02 (S)-Cheilanthifoline synthase (S)-Stylopine synthase (S)-Adenosyl-L-methionine: (S)-tetrahydroprotoberberine cis-N-methyltransferase (S)-cis-N-Methyltetra-hydroprotoberberine 14-hydroxylase Pro topine hydroxylase Spontaneous Dihydrobenzophenanthridine oxidase Fig. 9.

Biosynthetic pathway for sanguinarine (51)from (S)-reticuline (43).

important medicinal qualities. Morphine (52), codeine (53) (Fig. lo), and heroin (ma) (Fig. 19) for example, are potent analgesics. Subtle changes in the structures of morphine can alter the pharmacological activity quite considerably; methoxylation of the C3 carbon reduces potency and acetylation of the C3 and C6 hy-

droxyls increases the psychotic effect (BRYANT,1988). Chemists have striven to produce morphine analogs with diverse properties. The morphine molecule is, however, highly complex and possesses an abundance of functional groups. This has made transformation of morphine very difficult using conven-


5 2 R = H;Morphine 5 3 R = Me: Codeine 5 4 R = Et; Ethylmorphine

341

55 Thebaine

56 R = H ; Hydromorphone 57 R = Me: Hydrocodone

60 Diamorphine

6 1 R = H; Dihydromorphine 62 R = Me: Dihydrocodeine

64 Nalorphine

65 a R = allyl; Naloxone b R = CH2-cyclopropyl: Naltrexone

NMe

5 8 R = H; Oxymorphone 5 9 R = Me: Oxycodone

6 3 Neopinone

66 Pholcodeine Fig. 10. Opiate drugs.

tional chemical techniques, requiring the protection and subsequent deprotection of functional groups. Examples of important semisynthetic opiate analgesics include hydromor-

phone (56), nalbuphine, dihydrocodeine (62), and oxycodone (59) (Fig. 10). Morphine alkaloids have other pharmacological uses, also being antitussives (hydrocodone (57) and phol-

342

7 Alkaloids

codeine (66)) and narcotic antagonists used in the treatment of respiratory depression resulting from opiate overdose (naloxone (65a) and naltrexone (65b)). Naltrexone (65b) has recently gained recognition in the USA as also being useful in the treatment of severe alcohol addiction (MASONet al., 1994; VOLPICELLI et al., 1995). Since a third of emergency hospital admissions in the UK are due to alcohol abuse, and an average of US$ 100 billion per year is spent in the USA on medical bills or lost working days due to alcohol abuse, such morphine alkaloids have an obvious value. At present, these compounds are synthesized chemically from morphine (52), codeine (53), or thebaine (55) oblained from opium, which is the air-dried milky latex exudate of the incised seed capsule of Papaver somniferurn, the opium poppy. These industrial-scale multi-step syntheses are complex, using expensive metal catalysts and producing toxic waste streams, which require further treatment prior to release into the environment. Product yields are often moderate to low; e.g., the yield of hydromorphone (56) from morphine (52)is approximately 49%, while the yield of hydrocodone (57) from morphine (52) or thebaine (55) is 54% (HAILES,1993). Thebaine (55) amounts to a mere 6.5% of alkaloids found in opium; its limited supply makes hydrocodone (57) a high cost product, in turn limiting its availability. Such processes, with all their associated problems, are an obvious target for the design of alternative biological production routes, yet, to date, there are no viable biological alternatives with which they may be replaced. A considerable amount of effort has, therefore, been expended on developing a commercially feasible total synthesis of important morphinan alkaloids with varying degrees of success. The chemistry of the morphine alkaloids has been the subject of a number of excellent reviews (SZANTAY et al., 1994). Opium alkaloid production by plant cell culture has been studied extensively, as this would avoid the problems of variations in climate which affect opium yields; while alkaloid production has been observed (KUTCHAN et al., 1983) the production of morphine (52) or codeine (53) has not yet been achieved (VERPOORTE et al., 1993).

3.2.1 Morphine Alkaloid Biosynthesis The route proposed for the biosynthesis of morphine alkaloids from tyrosine via the benzylisoquinoline alkaloids (ROBINSON and SuGASAWA, 1931; BARTON et al., 1963) has largely been confirmed (reviewed by SZANTAYet al., 1994; Fig. 11). This has been elucidated by a combination of radiolabeled precursors and enzyme studies. Unlike the biosynthetic pathway for the tropane alkaloids very little is currently known about the molecular genetics of morphine biosynthesis . (S)-Norcoclaurine (39) originates from the stereospecific condensation of dopamine (37) and 4-hydroxyphenylacetaldehyde (38) (Fig. 8). The transformation of (S)-norcoclaurine (39) to (S)-reticuline (43) entails one hydroxylation and two 0-methylation sleps. ZENKand coworkers have elucidated the enzymatic steps and stereoselectivity of these reactions using 13C-labelingexperiments. GERADY and ZENK (1993) have demonstrated that it is (R)-reticuline (69) and not (S)-reticuline (43) that is the substrate for salutaridine synthase (Fig. 11). This enzyme which catalyzes the carbon-carbon coupling of C-12 and C-13 from (R)-reticd in e (69) and its congeners, has posed a particular problem for organic chemists. The enzyme was identified in the flowering stage of I? somniferum and was shown to be microsoma1 bound and to require NADPH and 0, for activity (ZENKet al., 1989).The absolute configuration of (S)-reticuline, therefore, has to be changed during the biosynthetic process and it is thought that this transformation proceeds via an iminium salt (67) (BATTERSBY et al., 1965).Although HE and BROW(1993) suggest that the NADPH-dependent enzyme catalyzing this conversion might be acting on the enamine (68)(Fig. 11). The enzyme that catalyzes this conversion has been purified to homogeneity from f? somniferurn seedlings (DEEKNAMKUL and ZENK,1990).

343

3 Benzylisoquinoline Alkaloids Me0

Me0

69 (R)-Reticuline

43 (S)-Reticuline

JT

MeoQ HO c---

c -

N Me Me0

NMe Me0

M eO

71 S a l u t a r i d i n e

Me

OH

0

0

\

70

Me0 HO N Me

Me0

-

OH

72

Salutaridinol

5 2 R = H; Morphine 53 R = Me; Codeine

NMe

Me0

5 5 Thebaine

73

63

Codeinone

Fig. 11. Biosynthesis of morphine (52) in fl somniferum (SZANTAYet al., 1994).

Neopinone

344

7 Alkaloids

3.2.2 Microbial Metabolism of Morphine Alkaloids There is little previously reported work on the microbial metabolism or biotransformation of morphine alkaloids. Early work was performed mostly with fungi and was concerned with the biotransformation of such compounds with the aim of producing more powerful and effective analgesics (reviewed by VINING,1980). IIZUKAet al. (1962) demonstrated the ability of the basidiomycete Trametes sanguinea to convert thebaine (55) to 14P-hydroxycodeinone (74) by monooxygenation and demethylation, followed by reduction (Fig. 12). i? cinnabarina was shown to produce the same compounds; however, only trace amounts of 14P-hydroxycodeine (76) were de-

tected, instead the majority of the 14P-hydroxycodeine was present as the N-oxide derivative ( G R ~ G Eand R SCHAUDER, 1969). YAMADAet al. (1962) later reported the conversion of codeinone (73) to codeine (53), 14P-hydroxycodeine (76), and 14P-hydroxycodeinone (74) by i? sanguinea. Substrate specificity studies revealed that the dehydrogenase present in i? sanguinea was relatively specific for its substrate, being able to reduce the ketone group of codeinone (73) and 14P-hydroxycodeinone (74), but not that of neopinone (63), pseudocodinone, or the structurally related steroid-4-ene-3-ones (YAMADAet al., 1963). In addition, i? sanguinea was later shown to be able to transform morphine (52) in addition to codeinone (73) and thebaine (55) (YAMADAet al., 1963), although no products could be identified at that stage.

Me0

55 Thebaine

Me0

/

\

J

7 4 14-Hydroxycodeinone

75

0

63 Neopinone

\

N Me

o 7 6 14 P-Hydroxycodeine

53 Codeine

0

73 Codeinone

Fig. l2. Transformations of thebaine (99,and neopinone (63),by I: sanguinea.


More recently, a number of fungal strains were found to transform morphine (52) to 2,2 '-bimorphine (77)(pseudomorphine) (Fig. 13; STABLERand BRUCE,unpublished data).

52 Morphine

77 2,2'-Bimorphine, y-morphine, pseudomorphine

1

78 ( 1OS)-u-hydroxy-2,2 '-bimorphine

1 79 ( lOS,lO'S)-u,a'-dihydroxybimorphine

Fig. 13. Dimerization of morphine (52) by C. didy-

mum.

345

This observation prompted closer examination of this activity in cell-free extracts of Cylindrocarpon didymum. The enzyme activity was purified by ion exchange, hydrophobic, and gel filtration chromatographies and detailed characterization work suggested that the enzyme was a laccase (p-diphenol oxidase; EC 1.10.3.2). Commercially available fungal laccases were also found to convert morphine (52) to 2,2'-bimorphine (77).Under slightly acidic conditions C. didymum was found to transform 2,2 '-bimorphine (77) further to the mono- (78) and dihydroxy derivates (79).The latter appeared to be degraded further by C. didymum in whole-cell incubations though the reaction product remained elusive prompting the thought that degradation may not be biological in this instance. Both the laccase and hydroxylase activities were shown to be induced by the presence of morphine (52); however, C. didymum was incapable of utilizing morphine (52) as a carbon source for growth and showed no activity with codeine (53) which is methoxylated at C-3, suggesting that a free phenolic group at C-3 is required for activity. Bacteria also have been shown to possess morphine alkaloid transforming abilities. LIRAS et al. (1975) used enzyme preparations from Pseudomonas testosteroni containing aand @-hydroxysteroiddehydrogenases to produce morphinone (83) (Fig. 15) and codeinone (73)from morphine (52) and codeine (53), respectively, in the presence of NAD +. Low levels of 14-hydroxymorphine (82) (Fig. 15) were also observed and it was speculated that this was formed by a spontaneous reaction involving morphinone (83), a fairly unstable compound.The conversion of morphine (52) to 14hydroxymorphine (82) was also observed in resting cell suspensions of an Arthrobacter sp. (LIRASand UMBREIT,1975). N-demethylation of the morphine alkaloids is an important reaction step as it provides pivotal intermediates for the synthesis of narcotic analgesics. such as naloxone (65a) and naltrexone (65b) (Fig. 10). Synthetic methods for these therapeutic morphine alkaloids are not satisfactory as they are difficult or hazardous to achieve, and they suffer from low and variable yields. Microbial N demethylation of morphine alkaloids has also been reported; studies in the 1980s saw the

346

7 Alkaloids

53 Codeine

80 N -Hydroxymethyl codeine

8 1 Norcodeine

Fig. 14. N-Demethylation of codeine (53) by Cunninghamella sp.

production of norcodeine (81)from codeine (53) by cell-free extracts of the fungus Cunninghamella (Fig. 14). The enzyme responsible for the reaction was identified as a membranebound cytochrome P-450 monooxygenase (GIBSON et al., 1984).These findings seemed to suggest that N-demethylation proceeds via attack of molecular oxygen on the N-methyl group (a-C oxidation) to produce a carbinolamine, N-hydroxymethyl codeine (80) (Fig. 14). The N-hydroxymethyl codeine intermediate (80) is unstable, and hence breaks down to norcodeine (81). More recent studies have demonstrated the ability of Mucus piriformis to N-dealkylate thebaine (55) to its nor-derivative (MADYASTHA and VIJAYBHASKER REDDY, 1994). The N-demethyl-ation of morphine alkaloids is not a phenomenon reserved solely for fungi, however. Streptomyces griseus has also.been shown to convert codeine (53)to norcodeine (81) (KUNZet al., 1985) (Fig. 14). The observations made by these studies were unable to advance knowledge of the catabolic pathways involved in the microbial metabolism of morphine alkaloids nor provide an adequate source of suitable enzymes for use in biotransformation systems. Indeed, many of the reaction products observed were probably due to the broad substrate specificity of the enzymes involved (VINING, 1980).

3.2.3 Transformations of Morphine Alkaloids by Pseudornonas putida M10 Morphine alkaloids possess a complex structure which makes them more resistant to

microbial degradation than other more simple molecules. However, morphine alkaloids are synthesized naturally and it is likely that an environment containing such compounds will also possess microbes capable to degrade them. If an organism is able to utilize a compound as a carbon or energy source, it must possess the enzymes necessary for its metabolism. It was with this reasoning that selective enrichments were carried out on samples from waste streams of an opiate processing plant. A mixed microbial culture was isolated, from which three different organisms were purified and identified as Methylobacterium sp., Alcaligenes xylosoxidans, and Pseudomonas putida designated strain M10 (BRUCEet al., 1990). I? putida M10 appears to be able to use morphine (52) or codeine (53) to support growth, albeit extremely slowly; growth commenced after a lag phase of 200 h with a doubling time of 180 h (BRUCEet al. 1990 HAILESand BRUCE, 1993). Two enzymes which are thought to initiate the degradation of morphine and codeine, morphine dehydrogenase and morphinone reductase, were identified and purified from this organism. Expression of morphine dehydrogenase and morphinone reductase was found to be constitutive in I? putida M10 (BRUCEet al., 1990,1991). Th'is was an unexpected finding, as the majority of catabolic enzymes in microorganisms are inducible and the constitutive expression of such enzymes would, presumably, place an unnecessary metabolic burden on the cell. It was assumed that the selection procedure employed may have favored those organisms with a faster growth rate and may have produced a constitutive mutant of f? putida M10. Although both morphine dehydrogenase and morphinone re-


ductase clearly participate in the transformation of morphine and codeine in vivo in I? putidu M10, and both enzymes appear to be specific for alkaloid substrates, it has, as yet, not been possible to demonstrate introduction of carbon from morphine alkaloids into central metabolism via this route, and it is, therefore, impossible to conclude that conversion of morphine (52) to hydromorphone (56) (Fig. 15) by morphine dehydrogenase and morphinone reductase is the true physiological function of these two enzymes. The oxidation of morphine (52)by washedcell incubations of I? putida MI0 was shown to give rise to a large number of transformation products including morphinone (83),hydromorphone (56)(dihydromorphinone), 14P-hydroxymorphinone (U),and dihydromorphine (61)(Fig. 15) (LONGet al., 1995). Similarly, in incubations with codeine (53) as substrate, codeinone (73), hydrocodone (57), 14P-hy-

61 Dihydromorphine

347

droxycodeinone (74),and dihydrocodeine (62) (Figs. 10,12) were identified as transformation products (LONGand BRUCE,unpublished information), while incubations with oxymorphone (58) (14P-hydroxydihydromorphinone) as substrate gave a major transformation product identified as oxymorphol (85) (Fig. 16; LONGet al., 1995). Some of these transformation products have significant commercial value. Semisynthetic opiates such as hydromorphone (56) and hydrocodone (57) are widely used analgesics;dihydromorphine (61)and dihydrocodeine (62)are also narcotic analgesics, while 14P-hydroxymorphine (82)and 14P-hydroxycodeine (76) could be used as pivotal intermediates in the synthesis of other analgesic agents. The chemical manufacture of these derivatives requires protection of functional groups during the synthetic process which can result in poor yields.While these semisynthetic opiates are often potent analgesics, their avail-

52 Morphine

82 14-Hydroxymorphine

Morphine dehydrogenase NADPH

56 Hydromorphone

83 Morphinone

8 4 14-Hydroxymorphinone

Fig. 15. Transformation of morphine (52) by E! putidu M10 (LONGet al., 1995).

348

7 Alkaloids

L

5 8 Oxymorphone

86 Codeine N-oxide

NADPH

Morphine dehydrogenase

M

NADP 3

%

HO

N /

M

e

H

87 Isocodeine

OH

85 Oxymorphol Fig. 16. Transformation of oxymorphone (58) by P putida M10 (LONGet al., 1995).

ability is often limited by the high cost of the chemical process. Furthermore, precursors such as thebaine (55) are often in limiting supply which also raises the cost of these compounds. Thus, utilizing microorganisms to transform naturally occurring morphine alkaloids to more potent derivatives could be advantageous. As hydromorphone (56) and hydrocodone (57) are important semisynthetic opiates the use of morphine dehydrogenase and morphinone reductase as biocatalysts for the production of these analgesic compounds was investigated. Morphine dehydrogenase was purified 1200-fold to electrophoretic homogeneity from €? putida M10 by two selective affinity chromatography steps (BRUCEet al., 1991). Analysis of the native enzyme by gel filtration chromatography gave a MW of 32000 Da,

Fig. 17. Codeine N-oxide (86)and isocodeine (87).

while SDS-PAGE showed the enzyme to be a monomer with an approximate MW of 31 000 Da. Morphine dehydrogenase was shown to be specific for the oxidation of morphine (52) and only a few other morphine alkaloids possessing a 7,8-unsaturated bond. It was shown to have a greater affinity for codeine (53) than morphine (52), while dihydrocodeine (62) was seen to be a poor substrate, as it lacked a 7,8unsaturated bond. This affinity was reflected by the K,,, values for morphine (52), codeine (53), and dihydromorphine (61) which were found to be 0.46,0.044, and 2.91 mM, respectively (BRUCEet al., 1991). Oxidative activity was also exhibited towards ethylmorphine (54), nalorphine (64) and codeine-N-oxide (86) (Fig. 17). The stereospecificity of morphine dehydrogenase was demonstrated by its inability to oxidize the codeine epimer, isocodeine (87) (Fig. 17).The enzyme exhibited a lack of oxidase activity towards primary, secondary, aromatic, or cyclic alcohols, or the structurally related steroids, testosterone and androsterone, making it quite distinct from the hydroxysteroid dehydrogenases of I? testoste-


roni described previously (LIRASet al., 1975). In the reverse direction, morphine dehydrogenase reduced hydrocodone (57), oxymorphone (58), oxycodone (59), neopinone (63), naloxone (65a), naltrexone (65b),and codeinone (73) (Figs. 10,12,17;BRUCE et al., 1994). lipurida M10 was found to carry a large natural plasmid, estimated by restriction digest analysis as being approximately 165 kb in size. The presence of this plasmid was shown to be essential for the metabolism of morphine (52), as plasmid curing experiments removed the ability of I?putida M10 to transform morphine (WILLEYet al., 1993).The structural gene for morphine dehydrogenase, morA, was cloned from this mega-plasmid using a degenerate oligonucleotide probe based on the N-terminal amino acid sequence of purified protein. Recombinant morphine dehydrogenase was expressed constitutively at approximately 5% soluble cell protein in E. cofi from the pseudomonad promoter and ribosome binding site located upstream of morA (WIUEYet al., 1993). Sequence analysis showed that morphine dehydrogenase from I? purida M10 was closely homologous to 18 proteins, further defining a distinct superfamily of oxidoreductases which have diverse functional acitvities (BRUCEet al., 1994). Morphine dehydrogenase particularly resembles two bacterial, 2,5-dioxo-~-gluconic acid reductases, and two eukaryotic proteins of unknown function. The structural archetype of the superfamily is a p/cu arrangement (WILSONet al., 1992). The relationships within this superfamily of oxidoreductases are extensive and complex; however, on the basis of residue conservations/exchanges the nicotinamide coenzyme binding and substrate reduction occur in all the enzymes by broadly analogous mechanisms. Constitutively expressed morphinone reductase activity was demonstrated in cell-free extracts of li purida M10. Low levels of hydromorphone (56) and hydrocodone (57) accumulated when extracts were incubated with morphine (52) or codeine (53), respectively. These compounds were produced by the reduction of the 7,8-unsaturated carbon-carbon bond of morphinone (83) or codeinone (73) by an NADH-dependent enzyme which catalyzed an essentially irreversible reaction (HAILES and BRUCE, 1993).Purification to apparent ho-

349

mogeneity was achieved by a single affinity chromatography step using an immobilized dye, Mimetic Yellow 2 (FRENCHand BRUCE, 1995).Morphinone reductase was found to be a homodimeric flavoprotein with subunits of M, 41 OOO, binding non-covalently one molecule of FMN per subunit. It was also highly specific for its alkaloid substrate and has been shown to catalyze the reduction of only morphinone (83),codeinone (73), neopinone (63), and 2-cyclohexene-l-one; it is not active against morphine (52), codeine (53), or isocodeine (87).The steroids progesterone and cortisone were potent competitive inhibitors and steady-state kinetic studies suggested a ping pong (substituted enzyme) kinetic mechanism (FRENCH and BRUCE, 1995). Interestingly, morB, the structural gene for morphinone reductase,was cloned from genomic DNA, not from the mega-plasmid, as was morA, by the use of degenerate oligonucleotide probes based on elements of the amino acid sequence of the purified enzyme (FRENCH and BRUCE,1995).Sequence analysis of morA and surrounding DNA revealed an insertion sequence motif of the kind found in Rhizobium sp., which promotes interesting speculation as to the origin of these genes (BRUCE, unpublished data). Analysis of the upstream DNA sequence of morB indicated the presence of an E. coli-type ribosome binding site motif and an almost perfectly conserved rrp gene promoter motif, TTGACA-N,,-TTAAG, almost identical to that of morA. The presence of these regulatory motifs enabled the constitutive expression of morphinone reductase when the morB clone was transformed into E. cofi (FRENCH and BRUCE,1995). Sequence analysis and structural characteristics indicated that morphinone reductase was related to the flavoprotein d p barrel oxidoreductases, and was particularly similar to Old Yellow Enzyme of Saccharomycesspp. and the related estrogen binding protein of Candida afbicans (FRENCHand BRUCE,1995). Expressed sequence tags from several plant species show high homology to these enzymes, suggesting the presence of a family of enzymes conserved in plants and fungi. Although relaled bacterial proteins are known, morphinone reductase appears to be more similar to the eukaryotic proteins.

350

7 Alkaloids

3.2.4 Biological Production of Hydromorphone and Hydrocodone The commercial synthesis of morphine is still not feasible, nor is the production of opiates by plant cell culture. Thus a combination of biocatalysis and chemical synthesis may provide an alternative route for the production of some of the most effective analgesics. An industrial-scale enzymic process for the production of hydromorphone (56) and hydrocodone (57) could offer a significant improvement over existing synthetic methods for the production of these compounds. At present such a biotransformation is envisaged as being a whole-cell system, rather than a cellfree or immobilized enzyme system, as the nicotinamide cofactors required by morphine dehydrogenase and morphinone reductase

may be prohibitively expensive. Thus it was decided to investigate the use of recombinant strains of bacteria that express morphine dehydrogenase and morphinone reductase as a means of producing hydromorphone (56). Several expression constructs were prepared wich either contained single copies of morA and morB together on a single plasmid or on separate compatible plasmids (FRENCH et al., 1995).Two whole-cell biotransformation systems were studied to examine the potential effect of enzyme levels and ratios on the amounts and types of products obtained. HPLC analysis of the transformation mixtures demonstrated that the recombinant strains of E. coli were able to transform morphine (52) principally to two compounds: dihydromorphine (61) and hydromorphone (56) (FRENCH et al., 1995; BRUCEet al., 1995). Similarly, hydrocodone (57) and dihydrocodeine (62)were

Morphine dehydrogenase 83 R = H; Morphinone 7 3 R = Me; Codeinone

52 R = H; Morphine 53 R = Me; Codeine

NADP

NADPH

Morohine dehydrogenase 61 R = H; Dihydromorphine 62 R = Me; Dihydrocodeine

O

W

56 R = H; Hydromorphone 57 R = Me; Hydrocodone

Fig. 18. Redox reactions of morphine (52) and codeine (53)(FRENCH et al., 1995).

3 BenzylisoquinolineAlkaloids

the main products of transformations of codeine (53), with reactions proceeding at a slightly lower rate than with morphine (52). Dihydromorphine (61) and dihydrocodeine (62) accumulated because hydromorphone (56) and hydrocodone (57)are poor substrates for morphine dehydrogenase (Fig. 18). Although dihydromorphine (61) has analgesic properties, it can be synthesized relatively easily and so its synthesis is not desirable in such biotransformation systems.The primary objective of these biotransformations was to produce hydromorphone (%), hence any side reactions which resulted in the “loss” of valuable opiate material in the form of unwanted byproducts were undesirable. The high concentration of substrate and reduction in the level of morphine dehydrogenase with respect to morphinone reductase were thought to keep the equilibrium of the morphine dehydrogenase reaction towards morphinone (83) (rather than towards morphine) such that it was largely unable to carry out the reduction of hydromorphone (56) to dihydromorphine (61) which now enables yields of hydromorphone (56) approaching 80% to be achieved. A second point of note in such biotransformations is the disappearance of opiate material in the form of a blackbrown precipitate, thought to be the result of the spontaneous degradation of morphinone (83), which has been reported to be particularly unstable in 1954).This may be aqueous media (BENTLEY, overcome in future biotransformations if the level of morphinone reductase can be increased to that morphinone (83) is never present in solution for very long, but rather converted quickly to hydromorphone (56).

3.2.5 Microbial Transformation of Heroin The requirement of enzymes active against heroin for use in a biosensor for the detection of illicit drugs led to the isolation of soil bacteria capable of growth with heroin (88s) as a source of carbon and energy. One of these isolates, Rhodococcus sp. H1, was found to be able to grow on heroin (ma), and possess an et inducible heroin esterase (Fig. 19;CAMERON

351

88

a R1 = R2 = Ac; Heroin b R1 = H, R2 = Ac; 6-Acetylmorphine 5 2 R1 = R2 = H; Morphine

Fig. 19. Heroin metabolites from the action of heroin esterase.

al., 1994; RATHBONE et al., 1996b). Growth studies demonstratedthat Rhodococcus sp. H1 hydrolyzed the 3-acetyl ester group from the heroin molecule to yield 6-acetyl morphine (88b); this product was then excreted into the supernatant.When the heroin (8th) was nearly exhausted,the 6-acetyl morphine (=) was reinternalized into the cell and the 6-acetyl ester group was hydrolyzed to give morphine (52).This transformation finds value by the fact that, when coupled with morphine dehydrogenase, it has potential use for the detection of heroin, since the NADPH liberated by the action of morphine dehydrogenase with morphine (52)can be linked to a number of transducer systems. This subject has been examined in detail elsewhere (HOLTet al., 1995, 1996;RATHBONEet al., 1996a, b). The structural gene, her, encoding heroin esterase from Rhodococcus H1 has been cloned and overexpressed in E. coli, and this recombinant strain is capable of transforming heroin (Ma) to morphine (52). The her gene has been sequenced and comprised a 969 bp open reading frame, encoding a 322 amino acid polypeptide. The native heroin esterase is a tetramer with a M, of 137000 ( ~ T H B O N Eet al., 1996b). Sequence analysis and structural characteristics indicated that heroin esterase is related to the serine hydrolase family of proteins.

352

7 Alkaloids

doid secologanin (90).This enzyme has been purified from Catharanthus roseus (TREIMER and ZENK,1979; PFITZNER and ZENK,1989) and Rauwolfia serpentina (HAMPP and ZENK, 1988) and characterized in some detail. When the purified enzyme is immobilized it has been The synthesis of strictosidine (91)from trypt- found to be highly stable and has facilitated amine (89)and the terpene secologanin (90) the synthesis of gram quantities of strictosiand (Fig. 20) is of importance as strictosidine (91) dine (91)on a laboratory scale (PFITZNER acts as a precursor for a very large and diverse ZENK,1987). The precursors tryptamine (89) group of terpenoid indole alkaloids (for re- and secologanin (90)are commercially availviews see CORDELL, 1974;HERBERT, 1994).The able, therefore the potential to produce inuse of extracts from the plants Rauwolfia and dole alkaloids by biomimetic processes exists. Vinca for medicinal applications led to the Since strictosidine synthase can only be proidentification of vincamine (99) (Fig. 21) in the duced in limited quantities an obvious ap1950s from Vinca minor L., which is used as a proach to generating more enzyme was to vasodilator. This was followed by the isolation overexpress the structural gene for this enand characterization of vindoline, vincaleuko- zyme in an appropriate host organism. To this blastine, and catharanthine (111) (Fig. 24) end the cDNA from R. serpentina for strictosifrom Catharanthus roseus (KUTCHAN, 1994). dine synthase was isolated and cloned and ex3a-(S)-Strictosidine (91)is a pivotal interme- pressed in E. coli (KUTCHANet al., 1988; diate to over 1200 monoterpenoid indole alka- KUTCHAN,1989). The recombinant organism loids, many of which have highly complex was found to be also capable of producing structures. Furthermore, this alkaloid family strictosidine (91) when tryptamine (89) and gives rise to some important therapeutic com- secologanin (90)were added to cultures. The pounds such as ajmaline (92),quinine, strych- cDNA sequence for strictosidine synthase has nine (93),vincristine (94),vinblastine (99,and been expressed in an active form in transgenic yohimbine (97)(Fig. 21). Because 3a-(S)-stric- tobacco which opens the way for manipulating tosidine (91)is a critical intermediate in all the monoterpenoid indole alkaloid producing et al., 1991). monoterpenoid alkaloid pathways, the enzyme plants (MCKNIGHT Ajmaline (92)is found in the roots of R. serwhich catalyzes its formation, strictosidine synthase, has become an important target for pentina and is clinically important in its use as genetic manipulation. Strictosidine synthase an anti-arrhythmic agent. It appears to reduce catalyzes the stereospecific condensation of the maximum rate of rise of cardiac muscle actryptamine (89)with the monoterpenoid iri- tion potential without affecting the resting po-

4 Monoterpenoid Indole Alkaloids

W

N HN

89 Tryptamine

H

Strictosidine synthase

-

OH

91 3cr-(S)-Strictosidine

Fig. 20. Biosynthesis of 3a-(S)-strictosidine(91).

4 Monoterpenoid Indole Alkaloids

Ajmaline group

Strychnine group

353

Catharanthine group HO

92 Ajmaline

94 R = CHO; Vincristine 95 R = Me; Vinblastine

Vincamine group c--OH

Et

91 Strictosidine

\

Sarpagine group

99 Vincamine

Heteroyohimbine group

98 Ajmalicine

Yohimbine group

97 Yohimbine

96 Sarpagine

OH

Fig. 21. Important alkaloids derived from strictosidine (91).

tential and has proved useful in the treatment of both ventricular and supraventricular arrhythmias (CREASEY, 1994). Semisynthetic derivatives such as N-propylajmaline have also received clinical application. The mechanism of action by which these alkaloids are thought to exert their effect is by blocking calcium and sodium channels.The biosynthesis of ajmaline (92)is, however, not well understood and early reports on 'the pathway were reviewed by CORDELL(1974) and were proposed by consideration of structural relationships within this group of alkaloids; however, progress is now being made regarding the enzymology of the pathway that leads from strictosidine (91)

to ajmaline (92) (Fig. 22; HERBERT1994; KUTCHAN, 1994;reviewed by STOCKIGT,1995). A highly selective esterase catalyzes the conversion of polyneuridine aldehyde (101)to 16epi-velosimine (102)(PFITZNER and STOCKIGT, 1983) which is further converted to vinorine (103b)by a synthetase in the presence of acetyl-CoA. Cell culture work with R. serpentina suggests that vinorine (103b)is a precursor for vomilenine (104),and subcellular enzymology has provided evidence that NADPH-dependent reductases convert vomilenine (104) to acetylnorajmaline (106a).An esterase finally removes the original protecting 0-acetyl group yielding ajmaline (92).

354

7 AIkaloids

100 4,5-Dehydrogeissoschizine 101 Polyneuridine aldehyde

91 Strictosidine

1

PNA - Esterase eM- -?-?

H\V

*OH

\

104 Vomilenine

c

1

Reductase,

NADPH

;;g

103 102 16-epi-Velosimine a R = H; Deacetylvinorine; b R = Ac; Vinorine

Q t r a z g

Methyl-

%

OH

105 1,2-Dihydrovomilenine

0

k OH H

Y H Me

106 92 Ajmaline Acetyl- a R = Ac; Acetylnorajmaline esterasec b R = H; Norajmaline

.*

5

H

OH

Fig. 22. Biosynthetic pathway leading to ajmaline in Rnuwolfia (KUTCHAN,1994).

A considerable amount of research has been directed towards the biosynthetic formation of vincristine (94) and vinblastine (95) because of their clinical importance in the treatment of leukemia. They are dimeric alkaloids that are formed from the precursors vindoline (110b) (Fig. 23) and catharanthine (111) (Fig. 24) and are found at very low concentrations in the leaves of the Madagascan periwinkle plant, Carharanthus roseus G. Don (Apocynaceae), also known as Vinca rosea L. Semisynthetic derivatives of these alkaloids have been made and are also in standard clinical use as components of cocktails of drugs employed for cancer chemotherapy (CREASEY, 1994). These al-

kaloids arrest cell division in metaphase and the mitotic spindle undergoes attrition and finally disappears. It is thought that this antimitotic action is the cause of the cytotoxic action of these drugs against cancerous cells.Two biosynthetic pathways have been proposed for vindoline (110b),one of which is shown in Fig. 23. In the alternative proposal tabersonine (107) undergoes three sequential hydroxylations, followed by N- and O-methylation to give 17-0-deacetylvindoline (110a) (KUTCHAN, 1994). Vinblastine (95) is formed by the coupling of vindoline (110b) and catharanthine (111); while the dimeric alkaloid is present only at very low concentrations in the

355

4 Monoterpenoid lndole Alkaloids

H ydroxylase ----_---__________________________ + H

602Me

1 07 Tabersonine

SAM

108 a R = H; 11-Hydroxytabersonine R = Me: 1I-Methoxytabersonine

SAHX~ OMethylation

.

C02Me

Hydroxylase

t

110 Acetyl-SCoA a R = H; 17-ODeacetylvindoline CoASH b R = Ac; Vindoline OAcety 1transferase

X

109

SAMXa SAH b R = Me =

WMethylation

Fig. 23. Two proposed biosyntheticpathways in the late stages of vindoline formation in Cafharanthus (KUTCHAN,1994).

plant, the precursors are found in relatively high abundance and can be easily extracted. Formation of vinblastine (95) from vindoline (110b)and catharanthine (111) can be achieved chemically with a modified Polonovski reaction in which 3 ',4'-anhydrovinblastine (113)is formed (Fig. 24; LANGLOIS, et al., 1976; KUTNEY et al., 1976). 3'4'-Anhydrovinblastine (113) can be converted chemically to vinblastine (95) (MANGENEY et al., 1979), while enzymic conversion by plant cell-free extracts has also been reported (KUTNEY et al., 1982). A biotransformation route for the coupling of vindoline (110b)and catharanthine (111) has been achieved with horseradish peroxidase with either the addition of FMN and MnCl, or Hz0 2 (GOODBODYet al., 1987). The same coupling mechanism is believed to occur in both cases as it is thought that light activation of FMN may generate superoxide radicals and is believed to be similar to that of the modified Polonovski method. The peroxidases, in the presence of hydrogen peroxide, presumably oxidize the catharanthine (111) to a reactive

peroxide which is then subject to nucleophilic attack by vindoline (110b),and 3 '4'-anhydrovinblastine (113)is formed by sodium borohydride reduction of the iminium intermediate. Immobilized cell-free extract from C. roseus cell cultures has been used to catalyze the coupling of vindoline (110b) and catharanthine (111) to 3 '4'-anhydrovinblastine (113) and leurosine (KUTNEY et al., 1988). C. roseus tissue culture has been employed for biotransformations of 3 '4'-anhydrovinblastine (113) to other bisindole alkaloids. Most of the plant cell work on the production of alkaloids has been performed with C. roseus cell cultures. Hairy root cultures of C. roseus have also been explored as a potential source of vindoline (110b) and catharanthine (111) (BHADRAet al., 1993).

356

7 Alkaloids

5 Summary 11 1 Catharanthine

I

110b Vindoline

112 lminium intermediate

113 3',4'-Anhydrovinblastine

Fig. 24. Synthesis of

(1W.

3 ',4'-anhydrovinblastine

This chapter has sought to make the point that a good biochemical and genetic understanding of alkaloid metabolism is essential if biotransformationsof alkaloids are to be anything other than fortuitous. There is little doubt about the need to continue to elucidate new alkaloid biosynthetic and catabolic pathways, particularly if the diversity of available biocatalysts is to increase. Metabolic engineering will have a considerable influence on the development of alkaloid biotransformation systems in the future; while this field is still a very young one, much has been accomplished, and this is beginning to have an impact on alkaloid synthesis.The combination of increasing commercial interest and advances in understanding the genetic and biochemical basis of metabolic pathways is now producing a more rational approach to engineering biotransformation systems. New genetic tools are continually being developed which makes the manipulation of particular genes or chromosomes increasinglyeasier, and we can look forward to genetically manipulating plants enabling alkaloid transformations to occur in vivo. Understanding cells as integrated systems will be increasingly important if the whole process of genetically engineering organisms for biotransformation purposes is to become less empirical. If potential bottlenecks in pathways are to be identified there is the need to develop effective systems for the measurement of metabolic flux. It seems quite reasonable to postulate that engineering of hybrid pathways will enable the use of any enzyme on an industrial scale plausible regardless of its cofactor requirement. Furthermore, it is not inconceivable to envisage a time when it will be possible to design a biological route to a desired compound and then build in a stepwise manner a biotransformation pathway of choice.

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8 Yeast

STEFANO SERVI Milano, Italy

1 Introduction 364 2 Yeast Biotransformations 364 2.1 Reproducibility 365 2.2 Operational Conditions 366 2.2.1 Fermentation Conditions Using Carbohydrates as Energy Source 366 2.2.2 Fermentation Conditions Using Ethanol as Energy Source 367 2.2.3 Resting Cells 369 2.2.4 Cell Immobilization 369 2.2.5 Organic Solvents 369 3 Yeast Enzymatic Activities 370 3.1 Reducing Enzymes 370 3.1.1 Reducing Enzymes from the Fatty Acid Synthetase Complex 373 3.2 Enzymes Promoting the Formation of C-C Bonds 378 3.3 Hydrolytic Enzymes 379 4 Selectivity 379 4.1 Selectivity by Substrate Modification 380 4.2 Selectivity by Selective Enzyme Inhibition 382 5 Genetically Modified Yeasts 383 6 Product Recovery 383 7 References 384

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8 Yeast

1 Introduction The use of enzymatic systems in organic synthesis has become a fundamental tool for effecting selective, efficient, and economical transformations with the goal of using fewer aggressive reagents, less organic solvents, lower temperature, and shorter reaction pathways (JONES,1986).Applications are found especially in the production of chiral materials in enanet al., tiomerically pure form (SANTANIELLO 1992) and in the regio- and chemoselective transformation of complex multifunctional substrates. This is exactly what enzymatic systems are able to do: discriminate between enantiomers, diastereoisomers, chemically identical functional groups in the same molecules, different functionalities. Recently, the possibility of having access to purified enzymes with known catalytic capacity in larger amounts through genetically modified microorganisms has increased applications of these biocatalysts in synthesis. It is well known that hydrolytic enzymes can effectively be used as reagents due to the fact that they are secreted extracellularly and thus can be easily collected, their stability can be increased by various methods of immobilization/confinement and, most important, they do not require cofactors. Many different types of these enzymes offer a wide range of catalysts allowing the resolution of almost any kind of racemic secondary alcohol. The possibilities are increased in efficiency if dissymmetrization of prochiral compounds can be effected. These methods find increasing applications in transformations of industrial interest, although the strict economical requirements of industrial processes seldom allow companies to profit from these new methodologies. In fact, most biocatalytic applications concern the resolution of a racemic mixture: the undesired enantiomer still needs to be racemized or disposed of. Other isolated enzymes like oxidoreductases are seldom used in practical transformations. Cofactor regeneration and enzyme costs and stability are still partially unsolved problems. In reductions, whole-cell biocatalysts are still mostly preferred (SERVI,1992). Chances to find the required catalytic capacity by simple screening are very high. However, only bakers’

yeast is accepted by synthetic organic chemists as a selective reagent.The vast synthetic capacities and their application in synthesis have been described in exhaustive recent reviews (KIESLICH, 1976;REHMand REED,1984;SERVI, 1990; CSUKet al., 1992; DRAUZand WALDMA”, 1995). In this chapter the principle of the catalytic activities of bakers’ yeast and their possible applications in practical biotransformations are considered.

2 Yeast Biotransformations Bakers’ yeast (Saccharomyces cerevisiue) and brewers’ yeast (Saccharomyces carlsbergensis) are industrial organisms produced for the food industry in huge amounts (TRIVEDIet al., 1986). Their main metabolic pathways (KOCKOVA-KRATOCHVILOVA, 1990) have attracted more studies than those of any other microorganism. These studies are mainly devoted to the understanding of the production of enzymes responsible for the metabolism of carbohydrates during growth and fermentation and to fundamental biochemical studies. Together with the use of yeast in baking and brewing, the possibility of metabolic pathway modification has recently focused the attention on the possibility of using genetically modified organisms for the production of particular compounds by fermentation (ROMANOS et al., 1992). For a long time, the application of biomass in biocatalysis was dedicated to investigate the activity displayed by yeast on non-natural substrates. Recently,this type of research has been reinforced, and bakers’ yeast has proven to be an extremely valuable tool as a source of enzymes for biocatalytic applications.The reason for its success is quality, which is appreciated by chemists, but probably does not constitute interesting arguments for microbiologists or biochemists. Bakers’ yeast is a very economical source of enzymes. It can be easily grown in an open jar without sterile conditions. It has a high reducing capacity and does not require addition of cofactors. It is a multienzymatic system which will eventually give rise to multistep sequential

2 Yeast Biotransfonnations

365

reactions. Selectivity can be obtained by a ture elucidation and stereochemical assignnumber of empirical variations of numerous ment of a natural product caused RIDLEYto parameters and, most importantly, by careful try a yeast-mediated biotransformation to secure the required product (BARTONet al., selection of the substrate. The presence of a non-conventional sub- 1975; DEOLet al., 1976). The success of that strate in the complex enzymatic system can transformation and subsequent work done by cause a number of different responses at the the same authors favored the revision of most same time, which will be responsible for often of the known work on yeast biotranformation as well as new synthetic applications (RIDLEY unexpected substrate transformations. 1975; CRUMBIE et al., 1978). Yeast has played a special role in the devel- and STRALOW, opment of biocatalysisduring the last 20 years. Several hundreds of articles have been pubMany organic chemists have tried to solve lished in organic chemistry journals in the last chemical problems by using yeast, mainly con- 25 years which were mostly devoted to syncerning the production of chirality.Thus, it can thetic applications. Many of these articles debe said that yeast has had an important tutori- scribe work of general interest but of limited al impact on modem biocatalysis. Among the synthetic utility due to low selectivity and limlarge number of academic applications of ited yield of products.Today,problems of accuyeast some biotransformations are of absolute racy, mechanism,and selectivity are being consynthetic significance. Industrial applications sidered as central points in bakers' yeast catalare also important. Recently,a surge in interest ysis, which starts a third period of interest. In particular, the following general quesmainly for selectivity and mechanistic aspects has caused new attention to this kind of bio- tions have been addressed: transformation.It is evident that there is a well - reproducibility of yeast biotransformaestablished knowledge of the basic metabolic tions (capacities of different commercial pathways in yeast fermentation. But for the yeasts, standard definition); biotransformation of non-natural substrates - operational conditions (growing yeast vs. (often referred to as non conventional subresting cells, different sources of reducing strates) there is only very little and non-syspower); tematic information available. Attempts to ra- identification of the enzymatic activities tionalize yeast activity toward those substrates present isolation of enzymes,determinahave failed so far due to the complexity of the tion of their stereochemicalpreferences, interaction between the substrate and the livstudies on the synthesis of the enzymes in ing cells. yeast); Publications dating from the beginning of - selectivity (by substrate modification,sethis century to the 1970s describe the basic calective enzyme inhibition,effect of organpacities of yeast, with no special attention to ic solvents,effect of immobilization of the resolution of synthetic problems.The work of this period was in part outlined in wellcells); - genetically modified yeasts; known review articles (FRIEDMA", 1931; - product recovery. NEUBERG, 1949).The transformation of simple ketones (MCLEODet al., 1964) and of cyclic substrates with various microorganisms (KIESLICH, 1976) have been reported. Due to the more recent developmentsin analytical chem- 2.1 Reproducibility istry and the interest in enantiopure substancThe concept of reproducibility in biotranses of well defined purity and absolute configuration, most of that work has to be revised.The formation with whole-cell biocatalysts cannot focus on the synthetic aspect dealing mostly meet the standards as those used in organic with the preparation of chiral building blocks synthesis. Reproducibilityrequires at least the has opened a surge in interest in yeast-mediat- use of the same strain grown exactly in the ed biotransformations. The need of a com- same way and at the same cellular density pound in enantiomerically pure form for struc- using identical substrate concentrations. The

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8 Yeast

yeasts which were used in most of the procedures published were either fresh pastes from the local grocery or dried yeast from the supermarket shelf. It is commonly known that there is a great deal of difference in the fermentation properties of yeasts from different brands in bread making and brewing. No wonder that they also display different catalytic activity in the biotransformation of non-conventional substrates. When reductive transformations are involved, reaction rates and stereochemistry of reduction are sometimes slightly different from one yeast brand to another (PEREIRA, 1995). These differences presumably mainly depend on the ability of the yeast to produce enough coenzyme in order to keep up with the transformation. Changes in the expression of enzymatic activity have not been reported. The observed differences may be due not only to different strain compositions, but also to ageing or to the carbohydrate content of the paste. It is the capacity to produce coenzymes and not the limited availability of a particular enzyme that is considered to be rate-determining. In the presence of several reducing enzymes acting on the same substrate with different rate and selectivity,the limitation of the coenzyme can cause evident differences in enantiomeric composition. In order to give reproducible results it would suffice to always describe a control experiment by using standard dried yeast which is commercially available (Sigma type 11). Recently, experimental procedures for the preparation of various 3hydroxy esters have been collected and the authors’ procedures have been repeated by the editors using this kind of dried yeast. For all the transformations reported good agreement between the controlled procedure and the one reported with local yeast has been observed (ROBERTS, 1992). High interest in yeast catalysis is given to biotransformations which have been rarely or never found before with isolated enzymes or whole-cell transformations. The outcome of the biotransformation is often the result of several enzymatic steps, and small differences in catalytic activity may cumulate to justify the surprisingly different capacities attributed to yeast on the same substrate in similar conditions (TAKESHITA and AKUTSU, 1992; METH-COHN et al., 1994). Published reports often describe reactions occur-

ring with limited yields under poorly defined experimental conditions. Reproducibility of these reactions has sometimes been denied. However, conflicting results have been published only in a few cases (EASTONet al., 1995; RAMARAOet al., 1990);in other cases inability to reproduce published results has not been reported. Finally, in some reports, the experimental procedures describe experiments which have hardly anything to do with biocatalysis since the yeast is treated under conditions in which enzymes are denatured (BAIKet al., 1994). The nature of the chemoenzymatic mechanism through which product formation occurs has sometimes been revealed (FUGANTI et al., 1990).

2.2 Operational Conditions Operational conditions in yeast biotransformations are very important in that they definitely determine the quality and the quantity of products. The efficiency of transformation ultimately depends on operational parameters. Performance evaluation can be done through the calculation of the productivity number PN which is the amount of product in mmol per dry weight of catalyst in kg per hour (SIMONet al., 1985). High productivity numbers indicate better volume-time yields and usually easier isolation of the product. With purified enzymes a PN of 30000 can be attained; using biomass containing only a very limited amount of enzyme per unit weight of cells, a PN between 20 and 100 can be expected. For the yeast reduction of 3-0x0 butanoate (SEEBACH et al., 1984) PN=50. Many yeast transformations have a lower PN.

2.2.1 Fermentation Conditions Using Carbohydrates as Energy Source In usual biotransformation conditions the substrate is added to fermenting yeast using carbohydrates as energy source at temperatures between 25 and 36°C. Usually the mixture is stirred for 30-60 min before the sub-

2 Yeast Biotransformations

strate is added as such or as an ethanol solution. During the biotransformation which can last from several hours to several days, additional yeast andlor carbohydrates are added at fixed intervals. During the initial incubation period, carbohydrates are used as hydrogen donors for the production of the NAD(P)H, which is necessary for the reduction. However, substrate reduction is only an accidental event in the cell economy, i.e., that several reactions catalyzed by different enzymes occur before the reducing power of the cell is transferred to the substrate. The energy supply for the reduction comes from carbohydrates through the metabolic pathway usual for the microorganism in those conditions, and the reducing enzymes are only expressed at the end of the pathway. At that point the substrate is in competition with the natural substrates (acetaldehyde) for catalytic reduction. The desired reduction is usually not very efficient compared to the endogenous consumption of reducing power. Using yeast as microbial reagent in reductions, hundreds of moles of ethanol are produced per mole of reduced substrate, i.e., only a fraction of the produced NAD(P)H is used for the biocatalytic step (BUCCIARELLI et al., 1983; MCLEOD et al., 1964). The reducing power generated in this type of biotransformation is usually very high. During the fermentation secondary metabolites are produced such as surface-activeagents and carbon dioxide which will make product recovery more difficult and decrease purity. The pH value usually decreases from an initial value of 7 down to 3 according to the reaction conditions. Although the equilibrium between NAD(P) and NAD(P)H (and hence reduced and oxidized forms of substrates) is considerably influenced by pH, apparently the internal pH of cells appears not to be affected by the external pH. Nevertheless,pH has a significant effect on the outcome of the biotransformation (SAKAIet al., 1991;FUGANTI et al., 1984). The reduction of 3-0x0-esters to (S)- and (R)-3-hydroxyesters is known to proceed through the intervention of NADPH-dependent enzymes (HEIDLASet al., 1988). The aspect of cofactor regeneration from the oxidized form in Saccharomyces cerevisiae has been studied. It has been proposed that one of

367

Glucose

1

NADP+

NADPH

Cluco~e-6-P

6-P-Gluconolactone C-6-PDH

Ribulose-5-P T K d NADPH

i

I

6-P-Gluconate NADP'

Fig. 1. Part of the NADPH regeneration system in yeast using carbohydratesas energy source.

the pathways by which carbohydrates (glucose) are utilized for the production of NADPH can be described in Fig. 1 (KATAOKA et al., 1992). During fermentation the amount of glucose used is about 5 times higher compared to the cofactor actually used in the reduction of the ketoester. Reduction continues even after complete consumption of glucose. This has been attributed to the utilization of ethanol formed during the cofactor regeneration (KOCKOVA-KRATOCHVILOVA, 1990;KOMETANI et al., 1996).

2.2.2 Fermentation Conditions Using Ethanol as Energy Source It is well known that, upon exhaustion of saccharides, yeast is able to utilize the ethanol produced during the preceding phase: ethanol is oxidized to acetate and then to carbon dioxide. Utilization of ethanol as the sole carbon source is commonly used in yeast identification. Most yeast species are able to grow on ethanol. In biocatalysis ethanol has been used as substitute for carbohydrates in the yeast reduction of ethyl acetoacetate and of 2-0x0-propanol (KOMETANI et al., 1996).This possibility has recently been investigated and considered as a favorable alternative to the classical reduction in fermenting yeast using carbohydrates. The specific consumption rate of ethanol was about twice that of the specific reduction rate of the keto ester. The reaction proceeds under aerobic conditions but is completely blocked

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8 Yeast

under aerobic conditions where the carbohydrate-mediated reduction occurs.The yields of the reaction and the stereochemical outcome in the two systems were comparable.The reaction in presence of ethanol is considered to be clean and does not form by-products Moreover, the enzymatic activities displayed are apparently the same in the two systems - at least for the two examples reported. Tho different mechanisms of cofactor regeneration have been proposed for the reduction of acetol which is catalyzed by an NADH-dependent alcohol dehydrogenase and for the pketoester reductases which are NADPH dependent. In the first case, ethanol is oxidized to acetate which enters the tricarboxylic acid cycle

Ethanol

y,

respiratory chain

Acetate

N

A

H

producing two moles of NADH (Fig. 2) (KoMETANI et al., 1993).?his reaction is apparently not influenced by strong aeration. Conversely the reduction of ethyl acetoacetate which needs NADPH as cofactor is proposed to occur through a modified mechanism in which the NADP' is regenerated at the expense of the conversion of malate to pyruvate (Fig. 3). In this case, experimental conditions proved that the efficiency of the reaction is strongly influenced by aeration. These observations have been applied on a large scale by construction of a bubble-columnreactor which favors strong aerating conditions (KOMETANI et al., 1989).The product yields are improved with respect to traditional fermentation conditions.

N

A

M

co2

H

fig. 2. Proposed mechanism of the regeneration cycle for NAD+ in the yeast reduction of acetol to propylene glycol using ethanol as energy source.

7-x

NAD+

Acetate NADH

TCA cycle

respiratory chain

-i Malate

NADP

* co2 Pyruvate

NADPH /

Fig. 3. Proposed mechanism of the regeneration cycle for NADP+ in the yeast reduction of 3-0x0-butyrate to 3-hydroxy-butyrate using ethanol as energy source.

2 Yeast Biotransforrnations

2.2.3 Resting Cells The use of resting cells is an alternative to fermentation conditions: the main advantage is that the reaction usually is much cleaner. In this procedure the pressed (or lyophilized) yeast is used as it is, or it is activated for 30min. The suspension is filtered or centrifuged, washed with water, and resuspended in water or buffer in a stirred flask before the substrate is added. A small amount of glucose can be added in order to prolong viability and increase the reducing capacity. Under these reaction conditions the reduction rate might be lower or the amount of required biomass might be higher. However, under these operational conditions the phase of microbial growth and the biotransformation are separated with the advantage that each step can be optimized individually. In the highly enantioand diastereoselective reduction of ethyl-20x0-cyclopentane carboxylate, 25 g of biomass are required per g of substrate (PN=5) (SEEBACH et al., 1992). In the fermenting yeast reduction of ethyl acetoacetate 2 g of yeast per g et of substrate were used (PN=50) (SEEBACH al., 1984).In general, advantages in product recovery are evident; cofactor regeneration is made at the expense of carbohydrate storage of cells, and the amount of internal glycogen is limited. Its availability has been measured (SLAUGHTER and NOMURA,1992). A simple procedure making use of pressed yeast suspended in water has been compared with the classical fermenting approach. It gave favorable space-time yields with simple ketones as substrates (BUCCIARELLI et a]., 1983). These authors used about 5 times higher amounts of yeast in the resting cell procedure, but the substrate concentration was about the same. The procedure of using resting cells is particularly

1

2

Fig.4. Reduction of a C=C double bond with resting cells on a multi kg scale.

369

suitable for laboratory-scale preparations. However, the method has been applied to the reduction of ketoisophorone (1)to (6R)-2,2,6trimethyl-1,Ccyclohexanedione (Z), a useful precursor of the synthesis of optically active carotenoids on a rather large scale (Fig. 4) (LEUENBERGER et al., 1976). 13 kg of ketoisophorone are transformed in 17 d, with isolated yields higher than 80%.

2.2.4 Cell Immobilization Cell immobilization techniques have been developed and advantages are claimed compared to traditional fermentation processes. Comparisons of different techniques have been reported (HILGE-ROTMANN and REHM, 1990). Examples of industrial processes, especially in brewing, using immobilized yeast are numerous (MENSOUR et al., 1996). In yeast catalysis cell immobilization coupled with the use of organic solvents has been reported as advantageous in increasing stereochemical discrimination (NAKAMURA et al., 1988; CHRISTEN and CROUT,1987).Using this technology, a continuous process with a flow of substrates passing through a fixed bed of immobilized biocatalyst is convenient on a large scale (ODAand OHTA, 1992).Product recovery is also simplified.Yeast cells immobilized on calcium alginate are stable for several weeks (KIERSTANand BUCKE,1977).The efficiency of recycling of the immobilized cells has been shown to reach a maximum at the 4th cycle and to decrease with successive use (NAOSHIMAand HASEGAWA, 1987).Although cell immobilization is reported to improve the enantiomeric excess of the reduced products (NAKAMURA et al., 1985), application of this technique in biocatalysis seems to offer little advantage over other procedures, since yields are usually lower and recovery of the biomass does not significantly affect costs in the transformation of substrates of high value compared with products of normal fermentation processes (e.g., alcohol).

2.2.5 Organic Solvents The use of organic solvents in yeast biotransformation can be more advantageous

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8 Yeast

than conventional water suspensions. Organic solvents can increase the conversion in the biotransformation of poorly water soluble substratedproducts. Since product inhibition usually is a serious problem in both, biocatalysis using whole cells and applications of isolated enzymes, the presence of the organic solvent can decrease the substrate/product concentration in the pure water phase thus minimizing the inhibitory effect. Undesired side reactions in which water is involved, like hydrolysis of substrates or products, can also be prevented or minimized. A reduction of mass transfer limitation can also be expected following cell adhesion to the organic phase, increasing substrate availability to cells or simply by increasing cell wall permeability.The increase in product enantiomeric excess (ee) which is sometimes observed can be the consequence of the influence of varied substrate concentrations within the cell and thus of a kinetic effect causing increased differences in reaction rate between two possible competing enzymes, or a simple selective inhibitory effect on enzymes with opposite stereochemical preferences. The effect of organic solvents on the ee of the products obtained has been studied with selected substrates (NAKAMURA et al., 1988, 1991b; NAOSHIMA et al., 1992). Product recovery might become easier if surfactants can be avoided in .the biotransformation. The permeability of the plasma membrane, the part of the membrane through which solute uptake is controlled by cells, can be damaged without the total destruction of cell integrity by the solvent, resulting in cell permeabilization. The communication between enzymes and substrates in this type of cell is altered giving access to enzymes which are still in a situation similar to the one they had in living cells. Yeast cells in this condition are, however, no more viable and cell content will not be renewed. In certain particular conditions using solvents and surfactants a single phase containing solubilized cells can be obtained (HOCHKOPPLER et al., 1989). These preparations can still be active for use as biocatalysts. It has been proved that in systems in which cells are practically soluble, they are still viable. In this system, however, recovery of low molecular-weight organic compounds is practically impossible. If organic solvents are to be

used, it is important to use a solvent which can assure cell viability since proteins are rapidly degraded within dead cells (SALTER and KELL, 1995; SIKKEMA et al., 1994).

3 Yeast Enzymatic Activities It is well known that living cells are able to produce a very high number of different enzymes synthesized according to the organism’s requirements. Some hundreds of them are acting at the same time. From the fact that yeast is such a generous, easily grown, non-toxic organism, it appears natural that enzymatic activities have been particularly investigated in yeast. For many of these enzymes substrate specificity, kinetic constant, and conditions regulating their synthesis are known. From the point of view of application of yeast in biocatalysis it is important to identify the individual enzymes which are acting on substrates when these are subjected to yeast action. This would permit the selection of experimental conditions favoring the action of the specific enzyme and increasing selectivity. The knowledge of the enzyme at work also constitutes the basis for the development of new genetically modified reagents with altered or new catalytic properties.

3.1 Reducing Enzymes The most often applied yeast capacities are carbonyl and prochiral C=C double bond reductions. These activities are particularly abundant in yeast (WARDand YOUNG,1991). Some known oxidoreductases recognized in yeast are listed in Tab. 1. Most of these reducing enzymes act on soluble substrates, they may have very narrow substrate specificity and often require that the substrates are temporarily derivatized as in the case of S-CoA esters of carboxylic acids.Typical substrates for whole-cell biotransformations are low molecular-weight hydrophobic substances on which the majority of the en-

3 Yeast Enzymatic Activities

371

zymes listed inTab. 1will not act.The presence edge available,increased efficiency is often obof these enzymes not acting on lipophilic sub- served. A simple variation in oxygenation strates is very important in that they are in- could sometimes change the outcome of the volved in the glycolytic pathway and, there- reduction or help in identifying the presence fore, their concentration in cells is proportion- of a certain enzyme.Aerobic conditions are real to the ability of increasing NAD(P)H pro- ported to increase G-6-PDH and D-LDH duction. (3-6-PDH, 6-P-gluconate DH, and while depressing activity of MDH (malate) probably ketopantoyl lactone reductase are and ADH (HEISEand PIENDL,1973). Since et G-6-PDH is involved in the regeneration of involved in NADPH production (KATAOKA NAD(P) in vivo strongly aerobic conditions al., 1992). The possibility of regulating at least the should in some cases increase the cell reducing known enzymatic activity is probably not an efficiency. A large effect in bakers’ yeast reeasy task because of the scarce information duction of C=C double bonds with increased available and the complexity of the system. oxygen concentration has been reported (Fuet al., 1992). Moreover, complex experimentalconditions or GANTI et al., 1988b;HOGBERG At least three different ADH have been the selection of special strains will somewhat decrease the advantages of using this kind of identified in yeast. ADH I is present during exorganism. However, with appropriate knowl- ponential growth on glucose.ADH I1 is an oxidative enzyme and is active during late exponential growth on glucose and ethanol. ADH I11 is a mitochondria1 enzyme. Yeast alcohol Tab. 1. Some Oxidoreductases in Yeast dehydrogenases are enzymes of limited synthetic utility in biocatalysis. Although they are YADH(ADH 1,ADH I1,ADH 111) able to reduce aldehydes of a rather wide Xylitol DH structural range, only a few small methyl keMethylglyoxale reductase (MR) tones are reduced. The reduction of aldehydes Acetoin DH to primary alcohols is of little interest. The kiDiacetyl reductase netic resolution of racemic aldehydes by this Isocitrate DH Homoserine DH means has, however, been proposed (TSUBOI Shikimate DH et al., 1988; KAWAHARAet al., 1988,TOPet al., 17/3-Hydroxysteroid DH 1988).YADH has been invoked in the reducKetopantoic acid reductase tion of pyruvate to @)-lactate (VANEYSand Enoate reductases (enone reductases) WLAN, 1956). A-1-sterol DH Immobilized YADH has successfully been Glucose DH employed in the selective reduction of the carSaccharopine reductase bony1 group in the cqpunsaturated vinyl bro6-P-glumonate DH mide (3)to the correspondingalcohol (4) (Fig. Glucose-6-PDH Lactate DH (LDH) 5 ) (BHALERAO et al., 1993). Whole-cell bio-

Mannitol DH Glyoxylate reductases (I and 11) Acetoin reductase Malate DH Histidinol DH Threonine DH PIsopropyl malate DH Ketopantoyl lactone reductase FAS complex Fumarate reductase Succinate DH a-AminoadipateDH a-Glycero phosphate DH a-Ketoglutarate DH Glycerol DH

BrA

3

BrA

4

Fig. 5. Selective reduction of an a,&unsaturated ketone with YADH.

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8 Yeast

YADH

YADH

fast

'low

1

Enoate reductase

transformation of this compound would invariably give the saturated alcohol as the only product in this case. The role of YADH in bakers' yeast transformation can be interesting in multi-step transformations which will not take place without the initial oxidative or reductive intervention of this enzyme. It is the C=C double bond reduction in allylic alcohols to give the saturated alcohols.The putative enzyme involved in this reaction is enoate reductase which is only able to reduce a,@-unsaturated carbonyl compounds (FUGANTI et al., 1988b).Allylic alcohols are not reduced by isolated enoate reductases (SIMON et al., 1985).In bakers' yeast, double bond reduction occurs after the initial oxidation of the substrate to the corresponding aldehyde: evidence for the mechanism in Fig. 6 has been reported (FuGANTI et al., 1988b).A further example of the hidden participation of YADH in the reduction catalyzed by yeast is the observed transformation of a-diketones to the corresponding diols in Fig. 7. YADH catalyzes the initial reduction of the methylketone to the secondary alcohol. The hydroxy ketone formed is not further reduced by the same enzyme; instead it is reduced by glycerol DH. The net reduction observed in yeast might, therefore, be the consequence of the sequential intervention of two distinct enzymes (BESSEet al., 1993). Lactate dehydrogenase occurs in microorganisms with both stereochemical preferences. L-LDH is present in yeast and is responsible for the reduction of pyruvate to L-lactic acid which is accompanied by D-lactic acid. L-LDH is inhibited by oxamate, while the D-enzyme is not. Selective inhibition of one of the enzymes in the presence of the other should be possible. L-LDH from different sources has been stud-

CHO

Fig. 6. Mechanism of yeast reduction of activated C=C double bond.

ied and is considered a synthetically useful enzyme for biocatalytic applications, although the rate of reduction of pyruvate analogs is lower by several orders of magnitude (KIM and WHITESIDES, 1988). Other enzymes which can act on a-0x0 acid derivatives are the group of amino acid reductases which are not conveniently used in a whole-cell system. Glycerol dehydrogenases are also present in yeast with D- and L-selectivity (MERKELet a]., 1982; ABERTYN et al., 1994a,b).They can be responsible for the reduction of keto acids and esters (NAKAMURA et al., 1988; NAKAMURA, 1992), hydroxy ketones (BESSEet al., 1993),but probably not diketones. Ketopantoyl lactone reductase (conjugated polycarbonyl reductases) reduces ketoesters and ketones besides its natural substrate (HATAet al., 1989; NAKAMURA et al., 1993). Hydroxysteroid dehydrogenases are responsible for the observed ability of yeast to reduce carbonyl groups in steroids and probably keto groups in large cyclic compounds.

0

Fig. 7. Enzymes involved in the yeast conversion of a-diketones to diols.


Some of the oxidoreductases present in yeast can act in the oxidation directi0n.A double kinetic resolution of acetol by successive reduction-oxidation has been reported (KoMETANI et al., 1996). In other cases the oxidation of secondary alcohols was used for the kinetic resolution of racemic secondary alcohols (FANTIN et al., 1993).The oxidation usually requires a rather long reaction time. The bakers’ yeast-mediated oxidation of sulphides to homochiral sulfoxides (BUST et al., 1990) and the preparation of R-p-tolylsulfoxide of high enantiomeric excess has been reported. The biotransformation requires careful manipulation of the microorganism (BEECHERi t al., 1995).

373

3.1.1 Reducing Enzymes from the Fatty Acid Synthetase Complex Fig. 8 shows a schematic picture of the fatty acid synthetase cycle responsible for the in vivo synthesis of fatty acids (WALSH,1979a). Among others, the capacity to reduc 3-0x0thioester groups and activated C=C double bonds are exploited synthetically on structurally analogous substrates. Both the 3-0x0-esters and the C=C double bond reduction are considered to be of special interest for the production of chiral hydroxy esters, valuable chiral synthons, and of building blocks chiral due to the presence of a methyl bearing carbon.

R I

ACP acyltransferase

d

Fig. 8. Schematic representationof the fatty acid synthetase cycle highlighting the two reducing enzymes utilized in biocatalysis with whole-cell organisms.

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8 Yeast

At least two different oxidoreductases are found in yeast which are able to reduce 3-0x0esters with opposite stereochemical preferences (HEIDLASet al., 1988). The formation of enantiopure 3-hydroxy esters is an important practical problem due to the value of this and related compounds as chiral synthons. A number of structurally related 3-0x0-esters of (S)absolute configuration (L-series) can be obtained with yeast and other microorganisms with different conditions and efficiencies with high to very high enantiomeric excesses. Simple structural variation permits changing of the stereoselectivity of the reduction. Since the opposite enantiomer is available either by depolymerization of natural polyhydroxybutyrate (SEEBACH and ZUEGER,1982) or by hydroxylation of butanoic acid with Cundidu rugosa (HASEGAWA et al., 1981),the application to synthesis is particularly suited. In practice, both enantiomers of 3-hydroxybutyrate and valerate are available in enantiomerically pure form through biotransformation or fermentation. Fig. 9 shows some examples in which 3-hydroxy esters of high enantiomeric purity have been obtained in synthetically useful yields. The preparation of a-alkyl-@-hydroxy esters in the form of enantiomerically pure single diastereomers is not easily accomplished by reduction of the corresponding racemic ketones. In this case the carbonyl group is usually also reduced with high enantiospecificity, irrespective of the configuration of the adjacent chiral center. In fact, many examples reported always list hydroxy esters with the

Seebach e f al. 1985 Dillon e f a1 1991 Wipf et a1 1983

-OH Sih eta1 1983

OH

0

OK Hirama et al. 1983

Fig. 9. Structural features and stereochemistry of the reduction of 3-0x0-acid derivatives.

same relative configuration at the newly formed secondary alcohol (L-series). Enzymes with different stereochemical preferences for 3-0x0-esters and 2-alkyl-30x0-esters have been isolated (HEIDLAS et al., 1988; SHIEHet al., 1985; SHIEHand SIH, 1993; SIHet al., 1983;NAKAMURA et al., 1991a).They are NADPH-dependent enzymes and able to catalyze the reduction of 0x0-esters of different types. Fig. 10 shows that the two substrates (5) and ( 6 ) are in rapid equilibrium through the enol form (7). The L-enzyme 1 preferentially binds to the S-ketoester (5),whereas the L-enzyme 2 preferentially binds to the R-ke-

syn-(ZR,3S) Fig. 10. Stereopreferences of two yeast enzymes which reduce a-alkyl-P-ketoesters.


toester (6). In both cases the hydride equivalent is delivered to the Re face of the carbonyl group to give the respective products (SHIEH and SIH,1993; WARD,1995). Enantiomerically pure anti and syn hydroxy esters are obtained with the purified enzymes. In yeast the two enzymes have been estimated to be approximately present in the relative amount of 1:1.The ratio of the k,,,lK, for the two enzymes should account for the stereochemical outcome of the transformation. When R1 = ally1 and RZ= ethyl the result is a 77:23 anti:syn ratio. The ability to reduce C=C double bonds has been attributed to the enoyl reductase enzyme of the fatty acid synthetase (CORNFORTH, 1966; DUGAN et al., 1970). The asymmetric reduction of a prochiral C=C double bond is a general method for the preparation of chiral compounds of type X. This reaction requires that the double bond is activated with strongly polarizing groups. The enzyme responsible for this catalytic activity is not commercially available. This reducing capacity of bakers’ yeast has been also found in a number of other microorganisms, especially anaerobic bacteria of the Clostridiurn type (SIMONet al., 1985). They have extraordinary reducing capacities with very high productivity numbers (10 to 100 times higher than the corresponding reactions with yeasts) which permits the reduction of large amounts of substrate with very little biomass. Reduction of the alkene bond occurs with trans-hydrogen delivery (SEDGWICK and MORRIS, 1980), and the chirality of the product obtained has been studied. The stereochemical preference is opposite for E and 2 double bonds. Low enantiomeric excesses are obtained if the 2-Econversion is favored under the reaction conditions. Enoate reductase from yeast is not able to reduce a,p-unsaturated acids or esters, unless X is a halogen. Activating groups of the substrates are -CHO or NOz.Allylic alcohols are substrates since they usually are partially transformed with yeast cells into the corresponding aldehydes by the alcohol dehydrogenases present (FUGANTI et al., 1988b). Fig. 11 shows the two different types of triply substituted olefines which are substrates for yeast enoate reductase. They give rise to methyl alkanols (or corresponding products) of two different chirality senses. Examples of

375

A

X = H, D, CH3, C1, Br, CF3 A = CHO, CHzOH, CH(OCH312, C02Rt N O 2 Fig. 11. Substrates for yeast enoate reductase.

the two types of products obtained are outlined in Tab. 2. The sense of chirality observed has been studied by several authors (SIMON et al., 1985; KOUL et al., 1995). A simple working model which needs to accomodate the small hydrogen group into an appropriate orientation can be drawn. It is in agreement with the observed stereochemistry of the reduction of the two types of reduced structures and the double bond stereochemistry. No mechanistic implications are involved in this model. The model only assumes that hydrogen delivery occurs in a trans fashion as it has been proved with deuteriated a$-unsaturated aldehydes (Fig. 12).

T2n

Fig. 12. Working model for the enoate reductase reduction of triply substituted double bonds.

376

8 Yeast

Tab. 2. Preparation of Chiral Compounds by Yeast Reduction of Activated C=C Double Bonds Substrate

Product

Reference

(Fuganti ef al. 1975)

(Gramatica ef al. 1988)

(Sato ef al. 1988)

W

C

c1

H

(Fuganti ef al. 1988)

O

c1

c1

c1

C02Me

CI

c1

c1 p

C02Me

l$ c1

(Utaka ef al. 1987)

C02Me

C1 - - C \ C 0 2 M e CI

c1

(Utaka ef al. 1987)

@

(Leuenberger ef al. 1979)

AcO

0

0

EtOzC 4

EtOzC&M .e

M

e

(Leuenberger ef al. 1979)

OMe

OMe

A

O

(Gramatica ef al. 1987)

OH OH

CHO

(Gramatica et al. 1987)

(Hogbcrg ef al. 1992) (Ohta ef al. 1989)


The activity in yeast is also present in the yeast enzyme concentrate type I1 from Sigma (FRONZA et al., 1996).Experiments carried out on different substrates showed that the enzyme is a NADPH-dependent protein. Application of this preparation is, however, much less convenient than use of the whole-cell catalyst. The yeast-mediated reductive step of conjugated endocyclic double bonds has been reported to occur with concomitant resolution at the chiral center removed from the actual bond reduced. The enantioselective reduction of racemic (8) in Fig. 13 gives access to R-go-

I t

8

Ph& O h

Fig. 13. Reduction of an endocyclic C=C double bond with chiral discrimination at a remote center.

377

niothalamin (9) (FUGANTIet al., 1994). Exocyclic double bonds in a$-unsaturated ketones are also reduced. The selectivity of the reduction of the carbonyl group in the presence of the C=C double bond has been studied. Usually the carbonyl group is preferentially reduced, but if strongly activating groups are present formation of the saturated alcohol is observed. Interestingly, the selective reduction of the C=C double bond in the presence of a conjugated carbonyl group has been reported. Compounds of type (10) (R’=Me) are preferentially reduced to the saturated ketones (KAWAIet al., 1995; SAKAIet al., 1991). The unsaturated ketone (loa) is almost completely transformed into the saturated ketone (lla) before any saturated alcohol (l2a) appears in the mixture (Fig. 14). This result is of interest since it allows the selective preparation of the saturated ketone (llb), of the impact flavor of raspberries. Starting with (lob) of natural origin the biotransformation permits formation of natural (FUGANTI et al., 1993) raspberry ketone. The reduction was also performed using as catalyst a yeast enzyme concentrate (Sigma type 11) in the presence of NADPH or NADH. When (4R)- and (4S)-[4-*H]-NADPH or NADH were used the experiments showed that both cofactors were utilized, but with opposite stereochemistry, thus suggesting the presence of two different enone-reductases. The same type of selective C=C double bond reduction in

12

a R = H; R = Me b R = OH:R = H

13 Fig. 14. Selectivity in the reduction of a$-unsaturated ketones.

378

8 Yeas1

the presence of a carbonyl group has been reOther C-C bond forming activities can inported (see the example in Fig. 4; LEUENBER-stead be directly exploited in bakers’ yeast and GER et al., 1976). constitute unique examples in biocatalysis for the use of whole-cell biocatalysts for this purpose. They are sterol cyclase and pyruvate de3.2 Enzymes Promoting the carboxylase. Sterol cyclase is responsible (HOSHINO et al., Formation of C-C Bonds 1991) for the cyclization of the epoxide (17)in Although several aldolases, transketolases, Fig. 16 to lanosterol (18). The same reaction acyl-coA synthases and other activities re- can be done on several analogs and requires sponsible for the formationhreaking of C-C pretreatment of yeast with ultrasound. Native bonds are present in yeast (WALSH,1979b), bakers’ yeast can affect the transformation they cannot in general be exploited in bioca- only marginally (KYLERand NOVAK, 1992).A talysis for the biotransformation of unnatural similar enzymatic capacity has been observed substrates.This is due to their narrow substrate with other sources (ABEet al., 1993; XIAOand specificity and to the prevalent utilization of PRESTWICH, 1991). the enzymes and of the natural substrates in the yeast metabolic cycles, if wild-type strains are used. However, fructose diphosphate (FDP) aldolase was overexpressed in Saccharomyces cerevisiae. The whole-cell organism was used in biocatalysis with phenylacetaldehyde (14) (Fig. 15) as unnatural substrate (COMPAGNO et al., 1993).Interestingly, the cells provide the cosubstrate (dihydroxyacetone phosphate, DHAP) and the phosphatase required to hydrolyze the initially formed phosphate ester (15).Although the product (16)is obtained in only low yields due to the concomitant utilization of the substrates by yeast, the system appears to be of great potential value.

Fig. 16. Lanosterol cyclization mediated by yeast sterol cyclase.

OH

15 R = Po 3 2 16R=H OR

Fig. 15. A recombinant yeast overproducing FDP aldolase transforms phenyl acetaldehyde (14). The system does not require added DHAP and phosphatase.

Pyruvate decarboxylase is a thiamine diphosphate-dependent enzyme catalyzing the decarboxylation of pyruvate to acetaldehyde or the transfer of a C,-unit of pyruvate onto acetaldehyde with formation of the acyloin. This enzymatic system has been exploited in biocatalysis using benzaldehyde (19) as substrate (Fig. 17) (NEUBERG and OHLE,1922). The hydroxy ketone (20) initially obtained in the condensation reaction between benzaldehyde and acetaldehyde formed in situ from pyruvate decarboxylation is used in the indus-

4 Selectivity

R-

CJ%

20

Fig. 17. Acyloin condensation.

trial preparation of L-ephedrine on a multi-ton scale. For longer reaction times it is reduced to the diol(21). Other aromatic aldehydes (LONG et al., 1989; OHTAet al., 1986) are good substrates as well as cinnamaldehyde (FUGANTI and GRASSELLI, 1977) and furylacrolein (FuGANTI et a]., 1988a).Incorporation of higher a0x0 acids has been explored allowing the formation of higher homologs of the phenylacetyl carbinols, albeit with lower efficiency if compared to pyruvate (FUGANTI et al., 1988b).Purified yeast pyruvate decarboxylase has been used with several aldehydes and pyruvate derivatives (C2-units) as donors.The corresponding acyloins are usually obtained with lower yields compared with the whole-cell system (CARDILLO et al., 1991; CROUTet al., 1991). Other decarboxylases with potential synthetic applications are found in other organisms (WARDet al., 1992).

3.3 Hydrolytic Enzymes

379

pholipases (WITTet al., 1984), amino acylases, (GLANZERet al., 1986, 1987b), phosphatases (TREVELYAN, 1966) have been recognized. Their synthetic utility is usually limited if the applications are compared with the use of lowcost easily accessible hydrolytic enzymes of wide substrate specificity and stereochemical preferences (see Chapter 3, this volume). In some specific applications the resolution step of racemic alcohols-esters is rather efficient (GLANZER et al., 1987a). In other cases the unexpected hydrolytic activity cooperates with a reductive step in the production of homochiral intermediates (PEDROCCHI-FANTONI and SERVI,1991). More often the hydrolytic activity is undesired causing by-product formation (MAMOLL 1938). Peptide bond formation using immobilized viable bakers' yeast in reversed micelles has been reported (FADNAVIS et al., 1990). Recently, yeast phosphatases have been exploited for the hydrolysis of natural phytic acid (22) in Fig. 18 to ~-myo-inositol-l,2,6-trisphosphate (23) (BLUMet al., 1995) or for the hydrolysis of phosphatidic acid from phospholipid mixtures (D'ARRIGOet al., 1994).

4 Selectivity The primary goal of all transformations in biocatalysis is to exploit the high selectivity displayed by enzymes. The particular situation of yeast biotransformation is such that often selectivity is lower than desirable. It is well known that this results from the concomitant action of several enzymes of opposite stereochemical preferences acting on the same sub-

OR

2 2 R = F'02H21 Yeast Hydrolytic enzymes, abundant in many or23 R = H phosphatases ganisms are present in yeast. Proteases (ACHSTETTER and WOLF, 1985), lipases (SCHOUSFig. 18. Hydrolysis of phytic acid catalyzed by BOE, 1976),esterases (PARKKINEN, 1980),phos- yeast phosphatases.

380

8 Yeasf

strate with different kinetics. Non-conventiona1 substrates are actually accepted by several enzymes present in yeast, and the result of the biotransformation depends on a series of factors which affect enzyme production or activation and cofactor regeneration. Yeast biotransformations acquire synthetic significance when the product can be recovered in high enantiomeric purity and without the need of purification from the starting material and byproducts or isomeric compounds.The possibility to increase selectivity has been extensively studied and is the object of current research in this field. Tools for the improvement of selectivity, which is mainly devoted to enantioselectivity and diastereoselectivity are in practice all the operational conditions which usually can affect growth and metabolism of a microorganism. The most effective results have been obtained by substrate modification. In the reduction of prochiral carbonyl compounds, a well-known simple rule has been devised to forecast the stereochemical outcome of the secondary hydroxy alcohol obtained. It is known as Prelog’s rule (PRELOG, 1964).It simply states that hydride equivalent addition to the planar prochiral carbonyl group will occur from the re face giving the alcohol of S absolute configuration (Fig. 19).The rule which has been deduced after the experimental results obtained with Curvufuriufufcutu applies rather

I

well to the results obtained with bakers’ yeast, with a consistent number of exceptions. It has been observed with several oxidoreductases and with yeast that prochiral ketones flanked with structurally similar groups give products of low enantiomeric purity. In contrast, the more different the two groups are the higher the discrimination. This result is explained by assuming two possible orientations at the enzyme active site, depending on the size of the two groups flanking the carbonyl group. Alternatively,similar substrates with marked different S and L groups will be more selectively recognized by one of two enzymes acting with opposite stereochemical preferences.

4.1 Selectivity by Substrate Modification Substrate modification is a powerful tool in improving the selectivity of enzymatic reactions. A well known application is in the reduction of keto esters: modification of the alcohol esterifying the acid can invert the stereochemical outcome of the reaction. Comparison of the yeast reduction of 3-0x0-valerates in Fig. 20 shows that the ee of the obtained alcohol clearly depends on the structural features of the ester. This simple modification can be advantageous in synthesis (NAKAMURA et al., 1979). 1986; DESHONGet al., 1986; FRATER,

re-face addition

R = C H 3 5%ee R = C z H S 40%ee R = CsHI7 95% ee ( 9-alcohol

Fig. 19. Prelog’s rule for carbonyl reduction.

Fig. 20. Dependence of enantiomeric excess from the chain length of alcohols esterifying 3-0x0-valerates.

4 Selectivity

381

This concept appears to be of general appli- sulphurization of the resulting compound. This cability: better selectivity is usually obtained concept has initially been applied by JONES when the two parts flanking the carbonyl (DAVIES and JONES, 1979) for the preparation group are well differentiated. of the 3-hydroxyoctane of (S)-configuration The same structural modification has little mediated by HLADH. It has been extended to influence on the reduction of a-substituted p- cyclic compounds in which the preparation of keto esters: invariable mixtures of diastereo- the single enantiomer of a compound with two isomers are obtained. chiral centers in yields higher than 50% is posAnother structural modification which can sible due to the dynamic kinetic resolution inhave a beneficial effect on the enantiomeric ex- volving the enol form of the keto ester through cess of the alcohols obtained is the reduction of which racemization of the slower reacting carbonyl groups embedded in a sulphur ether- enantiomer occurs (HOFFMANN et al., 1981; ocyclic structure. By this, reduction of the car- FUJISAWA et al., 1992) or to the high-yielding bony1 is highly enantiospecific and the open- preparation of homochiral methylalkanols chain compound can be easily obtained by de- (HOGBERG et al., 1992) (Fig. 21). 0

II

A-p-

R

CHO

Yeast

&-

R

CHO

Fig. 21. Using sulphur etherocycles to improve reduction efficiency.

R

d

--

O

H

382

8 Yeas1

4.2 Selectivity by Selective Enzyme Inhibition

lieved to occur through the intermediate formation of the two isomeric hydroxy ketones. In the attempt to control the overall reaction in the reduction of (24) it has been proved that As already mentioned in this chapter, the the preparation of compound (26) in high unsatisfactory outcome of a biotransformation yield, enantiomeric excess, and selectivity is in which a chiral discrimination is involved, is possible by using the combined action of often the consequence of the action of more methyl-vinyl ketone as additive (50 mM) and a than one enzymatic activity. Control of these pretreatment at 50°C of the yeast (Fig. 22). activities can be in principle effected either by Thus it appears that in this biotransformation selectively enhancing the production of one of there are at least three different enzymes inthe enzymes at work - not an easy task in such volved. The reduction of (26) to (27) is inhibita complex system - or by selective inhibition ed by the addition of methyl-vinyl ketone. of another enzymatic activity. This possibility Since the putative enzyme in this reaction is only requires introduction into the reaction glycerol DH, it would be interesting to corremixture of an appropriate inhibitor in ade- late the purified enzymatic activity with the quate amounts (LANZILLOTTA et al., 1975). presence of the inhibitor. Reduction of (24) to This operation can be done by using a series (25) is prevented by preheating treatment: in of low molecular-weight reactive molecules this operation the selective inactivation of a whose inhibitory activity has been generically specific enzyme should occur (NAKAMURA et demonstrated even though the mechanism of al., 1996). In the reduction of P-diketones, aninhibition is not known. The list of compounds other class of compounds often considered as is rather broad and comprises allylic alcohol, bakers’ yeast substrates, the effect of several allyl bromide, vinyl-methyl ketone, cyclohexa- additives on the product configuration has none, ethyl chloroacetate, carboxylic acids, sec- been compared. Although complete stereoondary alcohols like 2-propanol, cyclopenta- chemical control is not achieved in the case no1 etc. (NAKAMURA, 1992; NAKAMURA et al., shown in Fig. 23, it has clearly been shown that 1995;PEDROCCHI-FANTONI et al., 1992;TICOZZIin the reduction of (28) the addition of allyl aland ZANAROTTI, 1988; FORNI et al., 1994).The cohol favors Re face hydride equivalent delivuse of these compounds has proved to be ef- ery with the formation of the S-alcohol(30) in fective in the improvement of the enantiomer- good enantiomeric excess, while allyl bromide ic excess for yeast reduction of 1,3-diketones, favors the formation of a product of opposite a- and p-keto acid derivatives, ketones and absolute configuration (29), albeit in lower others. The additive is usually employed in enantiomeric excess (FORNIet al., 1994). more than stoichiometric amounts with respect to the substrate and to the protein involved in catalysis. The choice of the inhibitor is completely empirical. The data obtained in various laboratories can hardly be used for extending the knowledge to other substrates since there usually is an overlapping of more than one effect (organic solvent, yeast pretreatment, temperature). Allylic alcohol is a substrate for YADH which oxidizes it to the corresponding aldehyde. This compound is considered as a severe inhibitor of this enzyme. This additive might be responsible for lowering the activity of this specific enzyme. 2-propanol and other secondary alcohols might be involved in cofactor regeneration (ITOH, 1982). The reduction of 12-diketonesto vic-diols is Fig. 22. Selective enzyme inhibition and selectivity very efficient (BESSEet al., 1995) and is be- in the yeast reduction of a-diketones.

F3

3

F3C

F3C

1

J

29

-Br

28

-OH

6 Product Recovery

ically modified yeast will still retain properties which made the wild type organism more attractive than others. Recently, an interesting report along this line - although on the oxidative direction - has appeared. The catalytic repertoire of bakers’ yeast has been expanded to include enantioselective Baeyer-Villiger oxidations (STEWART et al., 1996). The cyclohexanone monooxygenase gene from Acinetobacter sp. was inserted into a yeast expression vector and this was used to create a new microorganism 15C(pKR001) able to perform oxidative transformations (Fig. 24; see Chapter 11, this volume).

30

Fig.23. Effect of different inhibitors on the stereoselectivity in the reduction of P-diketones.

15C(pKROO 1 )

R

31

5 Genetically Modified Yeasts Although there is an increasing number of reports in the literature concerning the introduction and faithful expression of foreign genes in yeast (WILLIAMSON, 1985; EVANSand ATTFIELD,1989; ROMANOS et al., 1992), very little is presently known about how these laboratory-developed organisms will behave in a practical production situation. Such reports usually refer to the production of specific proteins (GOFF et al., 1984; MACHEROUX et al., 1991) or the modification of the basic yeast metabolism for the accumulation of a specific compound (PORROet al., 1995; COMPAGNO et al., 1993).The goal of having different yeast strains able to express specific reductase activities accompanied by the required cofactor might be met, and such a possibility would greatly improve the practical utility of yeast biotransformation reductions. It must be remembered that even though the same enzymatic capacities can be convenientely overexpressed in a number of microorganisms, genet-

383

32

Fig. 24. Enzymatic Baeyer-Villiger oxidation of cyclohexanone derivatives with a genetically modified yeast.

Whole cell-mediated Baeyer-Villiger reactions were carried out on a 1 mmol scale, and several cyclic ketones were converted in 20-30 h into the corresponding lactones with good yields. Oxidation of prochiral4-substituted cyclohexanones (31) produced lactones (32)with very high enantioselectivity. With low cell density, ketone reduction constituted only a minor side-reaction. In contrast to the biotransformation with Acinetobacter sp. cells (a class I1 organism), the use of this yeast does not require an induction period with cyclohexanone to produce the monoxygenase.

6 Product Recovery Isolation of product after yeast biotransformations can be effected by usual means: direct extraction of the mixture is possible if emul-

384

8 Yeast

I., ROBERTS,S. M., sion can be separated by sedimentation or cen- BEECHER,J., BRACKENRIDGE, TANG,J., W x m r n , A. J. (1995), Oxidation of trifugation. A practical way to avoid centrifumethyl p-tolyl sulfide with bakers' yeast: preparagation is stirring the mixture with an equal tion of a synthon of the mevinic acid-type hypoamount of organic solvent and let the mixture cholesteremic agents, 1. Chem. SOC.Perkin Trans. stand for a while. Subsequently, the lower I , 1641-1643. phase is separated by suction, and the organic BESSE,P., BOLTE,J., FAUVE,A., VES-BRE, H. phase is filtered through a celite pad. The bio(1993), Bakers' yeast reduction of a-diketones: mass-containing phase is submitted to the investigation and control of the enzymic pathway, Bioorg. Chem. 21,342-345. same procedure again. If difficulties are experH. (1995), Bakers' ienced after a preliminary solvent extraction, BESSE,P., BOLTE,J.,VESCHAMBRE, yeast reduction of a-diketones: a four-hour exthe mixture can be steam-distilled if the prodperiment for undergraduate students, J. Chem. uct has appropriate physical properties. When EdU. 72,277-278. dealing with hydrophilic or partly water-soluBHALERAO, U. T., DASARADHI, L., MURALIKRISHNA, ble compounds, the mixture can be treated N. W. (1993), A novel chemo-enzyC., FADNAVIS, with polymeric resins of adequate polarity matic enantiospecific synthesis of (S)-coriolic (Amberlite, XAD7 or similar). This method acid mediated via immobilized alcohol dehydrogcan also be used for non-hydrophilic comenase of bakers' yeast, Tetrahedron Lett. 34, 1982). The separation of the pounds (VOSER, 2359-2360. G.,SPIESS,B., resin beads from the suspension containingthe BLUM,C., KARLSSON,S., SCHLEWER, N. (1995), Synthesis of optically acREHNBERG, often abundant biomass can be easily effected tive ( + )-~-3,4,5-tn-O-phenylcarbamoyl-myo-inby filtration through a sintered glass filter of ositol from phytic acid, Tetrahedron Lett. 36, appropriate porosity.

7 References ABE,I., ROHMER, M., PRESTwIm,G. D. (1993), Enzymatic cyclization of squalene and oxidosqualene to sterols and triterpenes, Chem.Rev. 93,2189-2206. ACHSTETITR. T., WOLF,D. H. (1985), Hormone processing and membrane-bound proteinases in yeast, Yeast 1,139-142. ALBERTYN, J.. HOHMA", S . , ~ V E L E I J.NM., , PRIOR, B. A. (1994a), Gene GDPl encoding glycerol-3phosphate dehydrogenase is essential for growth under osmotic stress in Saccharomyces cerevisiae and its expression is regulated by the hog pathway, Arch. Int. Physiol. Biochim. Biophys 102, B31. J., HOHMA", S., THEVELIN, J. M., PRIOR, ALBERTYN, B. A. (1994b), GDP1, which encodes glycerol-3phosphate, is essential for growth under osmotic stress in Saccharomycescerevisiae,and its expression is regulated by the high-osmolarity glycerol response pathway, Mol. Cell.BioL 14,41354144. BAIK,W., HAN,J. L., LEE,N. H., KIM,B. H., HAHN,J. T. (1994), Selective reduction of aromatic nitrocompounds to aromatic amines by bakers' yeast in basic solution, Tetrahedron Len. 35,3965-3968. D. H. R.,BROWN, B. D., RIDLEY, D. D., WIDBARTON, DOWSON, D. A., KEYS, A. J., LEAVER, C. J., (1975), The structure of daucic acid, 1.Chem. SOC.Perkin Trans.I , 2069-2076.

7239-7242. BUCCIARELLI, M., FoRNI,A.,MORE'ITI,I.,ToRRE, G. (1983), Asymmetric reduction of trifluoromethyl and methyl ketones by yeast; an improved method, Synthesis,897-899. D. M.,PARTINGTON, E. T., BUST, P. H., MARECAK, S ~ AP. ,(1990), Enantioselective sulfoxidation of a fatty acid analog by bakers' yeast, J. Org. Chem. 55 (22), 5667-5669. CARDIUO,R., SERVI,S., TINTI, C. (1991), Biotransformation of unsaturated aldehydes by microorganisms with pyruvate decarboxylase activity, Appl. Microb. Biotechnol. 36,3&303. CHRISTEN,M., CROW,D. H. G. (1987), Enzyme reductions of pketoesters using immobilized yeasts, in: Bioreactor and Biotransformations (MOODY,G. W., BAKER,€? B.,Eds), pp. 213-218. Amsterdam: Elsevier. COMPAGNO, C., T ~ MA., , W ZB.IM., , MARTEGANI, E. (1993a), Bioconversion of lactose/whey to fructose diphosphate with recombinant Saccharomyces cerevisiae cells, Biotechnol.Bioeng. 42,398400. C., SPERANZA, G., PANOSETTI,E., MACOMPAGNO, NIITO, P., RANZI,B. M. (1993b), The use of engineered yeast cells to perform aldol condensation, Biotechnol.Len. 15,1205-1210. J. W. (1959), Biosynthesis of fatty acids CORNFORTH, and cholesterol considered as a chemical process, J. Lipid Res 1,3-28. H., HUTCHINSON, D. W., CROW, D. H. G., DALTON, MYAGOSHI, M. (1991), Studies on pyruvate decarboxylase - acyloin formation from aliphatic, aromatic and heterocyclic aldehydes, J. Chem. SOC. Perkin Trans.I, 1329-1334.

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WARD,R. S. (1995), Dynamic kinetic resolution, Tetrahedron: Asymmetry 6,1475-1490. WARD,0. P., YOUNG,C. S. (1991), Reductive biotransformations of organic compounds by cells or enzymes of yeast, Enzyme Microb. Technol. U , 482493. WARD,0.I?, WILCOCKS, R., PROSENT, E., COLLINS, S., DEWDNEY, N. J., HONG,Y. (1992), Acyloin formation mediated by benzoylformate decarboxylases, in: Microbial Reagents in Organic Synthesis (SERVI, S., Ed.). Dordrecht: Kluwer Academic Publishers. WILLIAMSON, D. H. (1985), Cloning in brewers’ yeast Saccharomyces cerevisiae, in: Molecular Biology and Biotechnology (WALKER, J. M., GINGOLD, E. B., Eds.), pp. 102-118. The Royal Society of Chemistry, England. WIPF,B., KUPFER,E., BERTAZZI, R., LEUENBERGER, H. G. W. (1983), Production of (+)-(S)-ethyl 3hydroxybutyrate by microbial reduction of ethyl acetoacetate, Helv. Chim. Acta 66,485488. WIT, W., SCHWEINGRUBER, M. E., MERTSHING, A. (1984), Phospholipase B from the plasma membrane of Saccharomyces cerevisiae. Separation of two forms with different carbohydrate content, Biochim. Biophys. Acta 795,10&116. XIAO,X . Y.,PRESTWICH, G. D. (1991), Enzymatic cyclizations of 26-hydroxy-2,3-oxidosqualenesand 29-hydroxy-2,3-oxidosqualenesgives 19-hydroxylanosterols and 21-hydroxylanosterols, Tetrahedron Lett. 32,6843-6846.


9 Dehydrogenases Characteristics, Design of Reaction Conditions, and Applications

JORGPETERS Wuppertal, Germany

1 Introduction 393 2 Advantages and Disadvantages of Whole-Cell and Enzymatic Transformations 396 3 Regeneration of Nicotinamide Coenzymes 397 3.1 Introduction 397 3.1.1 Cost of Coenzymes 398 3.1.2 Origin of Coenzymes 399 3.1.3 Requirements of an Economical Coenzyme Regeneration System 399 3.1.4 Stability of Nicotinamide Coenzymes 400 3.2 Principles for the Regeneration of Nicotinamide Coenzymes 400 3.2.1 Nonenzymatic Regeneration 400 3.2.2 Substrate-Coupled Regeneration 401 3.2.3 Enzyme-Coupled Regeneration 402 3.3 Survey on Coenzyme Regeneration Methods and their Assessment 402 3.3.1 Regeneration of NAD(P)H 403 3.3.2 Regeneration of NAD(P) 405 3.4 Immobilized Coenzymes and Reactor Concepts 407 3.4.1 Coupling Methods 407 3.4.1.1 Coupling of the Coenzyme to Cross-Linked Enzymes 407 3.4.1.2 Coupling of the Coenzyme on a Carrier 408 3.4.2 Entrapment Methods 409 3.5 Conclusions 410 4 Assessment of Strategies for the Synthesis of Chiral Hydroxy Compounds 411 4.1 Introduction 41 1 4.2 Survey of Different Strategies for the Synthesis of Chiral Hydroxy Compounds and their Assessment 41 1 4.2.1 Enantioselective Reduction 412 4.2.2 Enzymatic Hydrolysis of the Racemic Hydroxyacid Ester Compound 413 4.2.3 Enzymatic Hydrolysis of the Racemic Ether Derivatives of the Hydroxy Compound 414

392

5

6

7

8 9

9 Dehydrogenases - Characteristics,Design of Reaction Conditions, and Applications

4.2.4 Enzymatic Hydrolysis of the Racemic Acyl Derivatives of the Hydroxy Compound 415 4.2.5 Transesterification Reactions 416 4.2.6 Acylation Reactions 418 4.3 Conclusions 419 Application of Alcohol Dehydrogenases in Organic Solvents or Cyclodextrins 420 5.1 Introduction 420 5.2 Solubilities and Stabilities of Substrate(s) and Product(s) 421 5.3 Modern Concepts for the Application of Alcohol Dehydrogenases Using Organic Solvents or Cyclodextrins 422 5.3.1 Oxidoreductions Using Organic Solvents 423 5.3.2 Oxidoreductions in the Presence of C dodextrins 425 5.4 Conclusions 427 Kinetics and Stereochemistry of Alconol Dehydrogenases 427 6.1 Introduction 427 6.2 Stereochemistry and Stereoselectivity of Alcohol Dehydrogenases 427 6.3 Reaction Mechanisms of Alcohol Dehydrogenases 428 6.3.1 Reaction Mechanisms 428 6.3.2 Substrate, Product, and Cross Inhibition 429 6.3.3 Impact of Reaction Mechanism and Type of Inhibition on the Selection and Modeling of Enzyme Reactors 431 6.4 Resolution of Racemic Hydroxy Compounds with Alcohol Dehydrogenases 434 6.4.1 Access to Both Enantiomers of a Hydroxy Compound 434 6.4.2 Introduction of Two Chiral Centers in One Reaction Step 435 6.5 Conclusions 436 Applications of Alcohol Dehydrogenases 436 7.1 Commercial Dehydrogenases, their Characteristics and Applications 436 7.1.1 Horse Liver Alcohol Dehydrogenase 437 7.1.2 Yeast Alcohol Dehydrogenase 439 7.1.3 Thermoanaerobium brockii Alcohol Dehydrogenase 441 7.1.4 L-Lactate Dehydrogenases 442 7.1.5 D-Lactate Dehydrogenases 442 7.1.6 Glycerol Dehydrogenases 443 7.1.7 Glutamate Dehydrogenases 444 7.1.8 Leucine Dehydrogenases 445 7.1.9 Phenylalanine Dehydrogenases 446 7.1.10 Alanine Dehydrogenases 446 7.1.11 Hydroxysteroid Dehydrogenases 447 7.2 Noncommercial Dehydrogenases, their Characteristics and Applications 448 7.2.1 Hydroxyisocaproate Dehydrogenases 448 7.2.2 Pig Liver Alcohol Dehydrogenase 452 7.2.3 Mucor javanicus Alcohol Dehydrogenase 452 7.2.4 Pseudomonas sp. Alcohol Dehydrogenase 453 7.2.5 Curvularia falcata Alcohol Dehydrogenase 453 7.2.6 Lactobacillus kefir Alcohol Dehydrogenase 454 7.2.7 Candida parapsilosis Carbonyl Reductases 454 7.2.8 Rhodococcus erythropolis Carbonyl Reductase 456 7.2.9 Candida boidinii Alcohol Dehydrogenase 456 7.2.10 Other Alcohol Dehydrogenases and Carbonyl Reductases 457 Conclusions and Outlook 458 References 460

I Introduction

1 Introduction There is an increasing trend in the pharmaceutical and agrochemical industries, to develop products containing enantiomerically pure materials.This trend was accelerated by a decision of the American Food and Drug Administration (FDA) in May 1992. Safety information is now demanded for individual stereoisomers of products submitted for approval and although racemates will still be continued to be approved on a case by case basis, information on each of the enantiomers is required. This significantly increases the cost of generating the necessary data for approval of racemates. This regulatory environment has led to an increase in the number of single isomer drugs being approved. In the current world market for pharmaceuticals, it is estimated that around 20% ($50 billion) is accounted for by optically pure drugs.This percentage is expected to increase to over 30% by the year 2000. As a result of the growth in demand for chiral actives, the market for chiral intermediates is currently estimated in the region of $ 1billion per annum worldwide, and growing fast. This has provided an enormous impetus for the development of enantioselective chemical transformations, such as the reduction of carbonyl compounds. A range of reagents for the asymmetric reduction of carbonyl compounds is available. They can be divided into two groups: stoichiometric and catalytic reagents (for reviews see: SINGH,1992; HARADAand MUNEGUMI, 1991; NISHIZAWAand NOYORI,1991). Although in many cases high enantioselectivity can be achieved, these reagents still have limitations; Nonactivated carbonyl compounds may not be reduced or only slowly with low enantioselectivity (e.g., alpine borane (l),they can show a limited substrate tolerance (e.g., chlorodiisopinocampheylborane (2), the synthesis of some of these agents is laborious (e.g., R,R-2,5dimethylborolane (3); MASAMUNE et al., 1986), they have to be used at low temperatures ( -78°C to - 1OO”C, BINAL-H) or at high pressure (e.g., 100 bar hydrogen, BINAP-Ru (4); NOYORI, 1989,1990).The oxazaborolidine (5) reduces aryl alkyl ketones with high enan-

393

tioselectivity,but gives poorer results with other ketones (DOUGLAS et al., 1996). Although a number of chemical agents for the reduction of carbonyl compounds is available (Fig. l), from both the academic and industrial viewpoints, it is likely that biotransformations will have an important role in the preparation of chiral intermediates over the medium term. There is considerable interest in the application of enzymes or whole cells for the reduction of carbonyl compounds, since they act under mild conditions and show high regioand stereoselectivity (HOSONOet al., 1990). The first report of the use of yeast cells as a catalyst for asymmetric reductions was the reduction of a carbonyl compound by fermenting bakers’ yeast in 1918 by NEUBERG and LEWITE (see Chapter 8). A broad range of bacteria, yeasts, and fungi has been used for the asymmetric reduction of carbonyls (CHRISTEN et al., 1992;AZERADand BUISSON, 1992; BESSE et al., 1994; GUNTHERand SIMON, 1985; WARHURST and FEWSON, 1994; YAMADA and SHIMIZU, 1988). Today, about 3500 enzymes are known (Enzyme Nomenclature 1992,Academic Press), of which about 70% are cofactor-dependent and about 15% are commercially available. The above mentioned microbial reductions are performed by redox enzymes (EC l).The substrate that is oxidized is regarded as hydrogen donor. The recommended name is dehydrogenase or reductase. The term oxidase is only used in cases where 0, is the acceptor, and peroxidases use H,O, as acceptor. More than 650 oxidoreductases are known (DAVIES et al., 1989) of which about 90 are commercially available.About 80% of all redox enzymes use nicotinamide adenine dinucleotide (NAD 6a, NADH 7a) as coenzyme redox “partners” and a further 10% the corresponding phosphates (NADP 6b, NADPH 7b) (Fig. 2). In most organisms, NAD is employed in oxidative catabolism linked to ATP production whereas NADPH is utilized for reductions involved in the biosynthesis (anabolism) of cell structures or energy storage (e.g., steroids or fatty acids). The abbreviations NAD(H) and NADP(H) are used to simultaneously refer to the oxidized and reduced forms of the same coenzyme (i.e., 6a/7a; 6b/7b), whereas NAD(P) and NAD(P)H refer to the oxidized and reduced

394

9 Dehydrogenases

- Characteristics,Design of Reaction Conditions, and Applications

1 (R)-3-Pinanyl-9- bora bicyclo[3.3.l]nonane

2 (-)-Chlorodiisopinocampheylborane

\\\I"

I H 3 (R.R)-2,5-Dimethylborolane

4 2.2'-Bis(dipheny1phosphino) - 1 ,l'-binaphthyl-ruthenium

Ph

forms respectively of both coenzymes (i.e., 6a/6b;7a/7b).NAD(P)(H) refers to all four coenzymes. NAD (6a) is a pyridinium salt and hence is frequently denoted as NAD +,whereas NADH is neutral according to this specification. However, except under the most acidic conditions the pyrophosphate bridge bears two negative charges and the phosphate group of NADP(H) will also be deprotonated, hence overall the molecule has negative charge. Recent literature tends not to show the charges with the abbreviations and this convention will be adhered to in this chapter unless the presence of the charge is essential to the argument. This has the advantage that it avoids ambiguities inherent in the consensus abbreviations, e.g., NAD(H), NAD(P)(H). The classification of NAD(P)-dependent oxidoreductases is shown in Tab. 1.This group is also the most important for preparative applications (WANGand KING, 1979). Flavins (FMN, FAD) and pyrroloquinoline quinone (PQQ) are involved more rarely. Oxidoreductions cover about 20-25% of the publications and patents (SEEBACH, 1990;FABER,1995).The Chapman & Hall Biotransformation Database (1996) contains about 1040 reactions catalyzed by dehydrogenases. The number of reactions published on dehydrogenases over the past 20 years is summarized in Fig. 3. Typical dehydrogenase reactions are depicted in Fig. 4. It is not intended to give an exhaustive review of all redox reactions catalyzed by alcohol dehydrogenases. These are much easier accessible in reaction databases on biotransformation (e.g., Chapman & Hall, SynopSys). Rather, this chapter is written from the viewpoint of a synthetic chemist, who wants to reduce a given carbonyl compound by means of a biotransformation. Several questions will be addressed consecutively in the following sections:

- What are the advantages of isolated enzymes versus whole cells?

- What are best systems for the regeneration of expensive coenzymes?

- What are the alternative routes to the di-

5 (R)-Oxazaborolidine

Fig. 1. Some reagents used in enantioselective chemical reductions of carbonyl compounds.

rect reduction of a carbonyl compound and how are they assessed? - Can oxidoreductases be used in organic solvents? What alternatives are there?

1 Introduction

-

II

6

RO OH

H o OH

aR=H;NAD b R = PO,”; NADP

395

NH2

7

a R = H; NADH b R = PO3*-; NADPH

Fig. 2. Nicotinamide adenine dinucleotide coenzymes.

Tab.1. Classification of NAD(P)-Dependent Oxb doreductases According to Enzyme Nomenclature EC Number

Donor

Number of Enzyme Subsubclasses

1.1.1 1.2.1 1.3.1 1.4.1 1.5.1 1.6.1 1.8.1 1.10.1 1.12.1

-CH-OH CHO,C=O -CH-CH-CH-NHZ -CH-NHNAD sulfur diphenols hydrogen

208 48 35 17 21 1 4 1 1

1.11.1 1.13.11 1.13.12

peroxidases dioxygenases monooxygenases

+

This chapter will focus on the isolated alcohol dehydrogenases whereas microbial transformations and oxygenases will be addressed in the preceding and the followingchapters,respectively. Parts of the topics addressed here have been reviewed in Vol.6a of the First Edition of Biotechnology (REHMand REED,1984) and by JONES(1986), YAMADAand SHIMIZU

11 36 8

Which types of reactors can be used for continuous enzymatic synthesis? Is the stereochemical outcome of an enzymatic reduction predictable? What are the kinetic limitations and how can they be circumvented? Which commercial and noncommercial dehydrogenases are available? What are their characteristics?Which of them are most recommended for a primary screening? Which dehydrogenases have already been applied on a large scale?

c 0

3

Fig. 3. Number of publications on reactions catalyzed by dehydrogenases.

396


Fig. 4. Reduction reactions catalyzed by dehydrogenases.

(1988), HUMMELand KULA (1989), WONG (1989), SEEBACH(1990), WARD and YOUNG (1990), WONGand WHITESIDES (1994), DRAUZ and WALDMANN(1995), and FABER(1995).

and coworkers (1964) found that (S)-alcohols were formed with moderate selectivities. Subsequently, many papers were published addressing asymmetric reductions catalyzed by bakers’ yeast (cf. Chapter 8, this volume). Bakers’ yeast shows a very broad substrate spectrum and in many cases good or high enantio- and diastereoselectivity, however, problems are often encountered (Fig. 5). Biological systems are genetically variable. The enzymes involved in redox reactions are not necessarily the same in different strains of the same organism. In order to reproduce published results it is essential to employ exactly the same strain (CHRISTEN and CROUT,1988; CHENet al., 1984). For example, in the reducIn the course of the first systematic investi- tion of 4-chloro-3-oxobutanoic acid methyl esgations on the stereochemistry of bakers’ yeast ter catalyzed by bakers’ yeast from Oriental reductions of carbonyl compounds, MACLEOD Yeast Co. (NAKAMURA et al., 1985) an (R)-se-

2 Advantages and Disadvantages of Whole-Cell and Enzymatic Transformations

Genetic Variabllity cell wall

chromosomes

isoenzyme/

Metabolism

Transport

Fig. 5. Problems encountered with microbial reductions of carbonyl compounds.

2 Advantages and Disadvantages of Whole-Celland Enzymatic Transformations

397

lectivity was obtained, whereas the same re- products is to work with resting cells. Howevduction catalyzed by bakers’ yeast from Red er, the large amount of biomass present in the Star Co. yielded the (S)-enantiomer (ZHOUet reaction mixture reduces the overall yield and often makes product recovery problematic. al., 1983). Moreover, the cultivation conditions (USHIO Due to a high excess of glucose or sucrose et al., 1986),the age, and the metabolic status added as auxiliary substrates for coenzyme reof the organisms have to be the same to repro- generation, by-products are formed which ofduce published results because of physiologi- ten impede product purification. Only a small cal variabilities (EHRLERet al., 1986). The ex- amount (typically 0 5 2 % ) of the carbon pression of enzymes, which are responsible for source is involved in coenzyme regeneration. a desired biotransformation, depends on the The solvent selection for whole-cell biocultivation conditions of the organisms. For transformations in organic media is problemeconomical reasons, the cell produces some atic, since for each pair of solvent and organenzymes only, if an essential nutrient is not ism a specific cytotoxicity is found (SALTER present in the medium. Some enzymes are and KELL,1995). The well-established log P controlled by “catabolite repression”, which concept (the octanol: water partition coeffimeans that in the presence of glucose their for- cient; LAANEet al., 1987) for enzymatic conmation is repressed, whereas during growth on versions does not correlate with the cytotoxicanother carbon source, these enzymes are ex- ity of organic solvents. Thus, mixtures of solpressed. Last but not least, the final product of vents optimized for a given organism have to a biochemical pathway can also affect the ex- be used in microbial conversions. pression of enzymes involved in this pathway The above described aspects of whole-cell (end product repression). biotransformations are summarized in Tab. 2 Transport of carbonyl or hydroxy com- and are compared with enzymatic systems.The pounds into and out of cells is often a problem. main advantages of microbial over enzymatic Product recovery can be troublesome, if the conversions are the low price of the catalyst product is not excreted from the cells. Chiral and the avoidance of cofactor recycling. On transport can lead to problems if racemic sub- the other hand, enzymatic systems do have strates are used. Cells often contain several de- considerable advantages since they are well hydrogenases with opposite stereoselectivity defined, controllable, scalable, and can be op(“iso”-enzymes) which compete for the same erated continuously. substrate. As a result, the stereoselectivity is reduced. SHIEH(1987) and NAKAMURA et al. (1991) demonstrated for a P-ketoacid ester that two (S)- and two (@-specific dehydrogenases are present in bakers’ yeast. Depending on the origin and cultivation conditions of the cells, variations in yield (10-85%) and enantioselectivity (40-98%) were observed in the case of reductions of P-ketoacid esters with bakers’ yeast (POPPEand NOVAK,1992; CHRISTEN and 3.1 Introduction CROUT,1987; WARDand YOUNG,1990; BUISSON et al., 1992). Moreover, the substrate Nicotinamide adenine dinucleotide (NAD, and/or product may be metabolized resulting 6a) and the analogous 2‘-phosphate (NADP, in reduced overall yields. 6b) (Fig. 2) are involved in redox reactions catThe productivity of microbial conversions is alyzed by alcohol dehydrogenases. Two elecusually low since the majority of nonnatural trons and a proton (hydride) are transferred substrates are toxic to the living cell and are from the reduced coenzyme NAD(P)(H) to the therefore tolerated only at low concentrations carbonyl compound in a stoichiometric reac(0.1-0.3%). If the electron transport chain is tion. Using coenzyme recycling, the expensive not involved in a reduction reaction, one pos- cofactor is needed only in catalytic amounts sibility to avoid the toxicity of substrates or leading to a drastic reduction in cost of dehy-

3 Regeneration of Nicotinamide Coenzymes

398

9 Dehydrogenases - Characteristics, Design of Reaction Conditions, and Applications

Tab. 2. Advantages and Disadvantages of Whole-Cell and Enzymatic Transformations Criteria

Whole Cells

Enzymatic

Availability Reaction conditions Optical purity Reproducibility Competing enzymes By-products Permeability/mass transfer Coenzyme regeneration Effective coenzyme concentration Price of catalyst Process Productivity Overall yields Substrate concentration Product recovery Compatibility with organic solvents

good mild medium -high variable Yes often limiting not necessary unknown low batch low low low troublesome problematic cytotoxic

often limited mild high constant no less or no non-limiting necessary well defined medium - high batch or continuous medium - high medium - high medium - high easy established

drogenase-catalyzed reactions. Additionally, coenzyme regeneration can accomplish three major objectives. First, it can influence the position of equilibrium of a redox reaction. A thermodynamically unfavorable reaction can be driven by coupling with a favorable cofactor regeneration reaction. Second, regeneration requires only catalytic amounts of coenzyme preventing the accumulation of by-products which may inhibit the reduction reaction. Third, coenzyme regeneration can simplify the downstream processing of the reaction mixture. This section deals with different aspects of methods for coenzyme regeneration.

3.1.1 Cost of Coenzymes Nicotinamide adenine dinucleotides (NAD (H), NADP(H)) are expensive compounds (Tab. 3). If these coenzymes were used in stoichiometric amounts, enzymatic reductions with dehydrogenases would be prohibitively expensive.Therefore, the application of dehydrogenases on the industrial scale requires effective methods for coenzyme regeneration. The economic barrier to large-scale reactions posed by cofactor costs has been recognized for many years (WILLNERand MANDLER, 1989; CHENAULT and WHITESIDES, 1987; CHENAULT et al., 1988). Turnover number (TN) is defined as the number of moles of product formed per mole of cofactor or enzyme per unit time. This number is indicative of the productivity of the pro-

Tab. 3. Prices of Nicotinamide Coenzymes Coenzymes

Price per Mole [U.S. $1 (depending on purity, JONES and BECK,1976)

Price per Mole [U.S. $1 (mean catalog prices, 1997)

NAD+ NADH NADP' NADPH

1.7 to 50 6 to 53 21 to 169 177 to535

710 3 25 215

(6a) (7a) (6b) (7b)


cess. Total turnover number (TTN) is defined as the total number of moles product formed per mole of coenzyme or enzyme during the course of the entire reaction. Hence, the TI" reflects the loss of coenzyme and enzyme throughout the process, and thus it is a useful estimate of the operational cost of the coenzyme or the enzyme. In order to overcome economical limitations,TTN > 100 are desirable. For a batch synthesis,the cost of the coenzyme or enzyme is calculated on the basis of its initial cost. In contrast, for a continuous process, the cost may be calculated on the basis of the activity loss over the entire process time. When estimating the cost of a given reaction per unit time, the rate of the reaction must also be considered. The regeneration cost is defined as the cost of the components like enzymes, reagents, and coenzymes required to regenerate one mole of coenzyme per unit time (WONGand WHITESIDES, 1994). The highest 'TTN can be achieved, if

(EGUCHIet al., 1983).Additionally, a combined chemical and enzymatic route for the synthesis of NAD(P) has been developed which may be useful for the preparation of NAD(P) analogs (TRAUBet al., 1969; KORNBERG, 1950;WALTet a1.,1984).

3.1.3 Requirements of an Economical Coenzyme Regeneration System CHENAULT and WHITESIDES (1987) compiled the following criteria for an ideal coenzyme regeneration system:

- Compatibility with the synthetic reaction.

- Commercial availability of inexpensive -

- the substrate concentration is as high as

possible determined by the substrate solubility, product inhibition, and enzyme stability, - the concentration of coenzyme and enzyme to achieve acceptable reaction rates is as low as possible, - the coenzymes and enzymes are highly stable. Thus, both the initial costs of the cofactor and the enzyme and the efficiency of their regeneration and utilization determine their contribution to the cost of the product.

399

-

and stable enzymes with high specific activity. Simple and cheap auxiliary substrates. The corresponding products should not interfere with the isolation of the product of interest, and should not influence the stability of enzymes and coenzymes. High turnover numbers (TN) and total turnover numbers (lTN). The equilibrium constants of the coupled coenzyme regeneration system should be as high as possible. The analysis of the yield should not be effected by the substrates or products of the coenzyme regeneration.

As a rule of thumb, 103-104 cycles are sufficient for laboratory-scale redox reactions, whereas at least 10'-lo6 cycles are desirable for industrial-scale synthesis. The exact turnover numbers required depend on the initial 3.1.2 Origin of Coenzymes cost of the coenzyme and the value of the Nicotinamide adenine dinucleotide (NAD, product of interest. A high turnover number 6a) is presently isolated from yeast (SAKAIet requires a high selectivity for the formation of al., 1973),and its 2'-phosphate NADP (6b) is enzymatically active coenzyme. 1,CDihydronormally synthesized by enzymatic phosphor- NAD(P) (7a, 7b) is the only enzymatically acylation of NAD using NAD-kinase (EC tive form, but 1,2- and 1,6-dihydroisomers are 2.7.1.23) and ATP (MURATAet al., 1979;HAY- frequently formed by direct chemical reduction. If this occurs at a significant rate the rate ASHI et al., 1979). NAD(P)H can be prepared from NAD(P) by three different methods: of enzyme reductions is decreased. For inchemically (LEHNINGER, 1957), enzymatically stance, if 50% of the original coenzyme activity (RAFTER and COLOWICK,1957; SUYE and is to remain after 1000 cycles, the regeneraYOKOYAMA, 1985), or by microbial reduction tion reaction must be 99.3% regioselective.

400

9 Dehydrogenases

- Characteristics, Design of Reaction Conditions, and Applications

In the case of industrial applications of dehydrogenases, lo6 turnovers are desired, resulting in a requirement for a regioselectivity of 99.99993%. Chemical, electrochemical, photochemical, or enzymatic methods (see below) show different selectivities in the formation of 1P-dihydro NAD(P) .Especially electro- and photochemical methods suffer from poor regioselectivities.Thus, the requirement for very high regioselectivity necessitates enzymatic catalysis, particularly in the recycling of NADH (7a) and NADPH (7b)CHENAULT and WHITESIDES, 1987; CHENAULTet al., 1988; HUMMEL and KULA,1989). +

3.1.4 Stability of Nicotinamide Coenzymes Nicotinamide coenzymes contain three labile bonds which may be the target of general or specific acid- or base-catalyzed hydrolysis: - the N-glycosidic bond between nicotin-

amide and ribose,

- the phosphodiester bond between the two riboses,

- the bond between ribose and phosphate

(NADP).

OPPENHEIMER (1982) and CHENAULT and WHITESIDES (1987) compiled data on the influence of different buffers on the stability and the possible products from the hydrolysis of NAD(P)(H). Under basic conditions, the reduced nicotinamide coenzymes are stable, whereas under acidic conditions they are unstable. In contrast, the oxidized coenymes are stable under acidic and labile under basic conditions (WONGand WHITESIDES, 1981; OPPENHEIMER, 1982). As a compromise, NAD (6a) and NADH (7a) are used at pH 7-73, whereas NADP (6b) and NADPH (7b)are used at pH 8-8.5. Moreover, it is very important that the enzymes used for synthesis and for coenzyme regeneration are also active and stable at the above mentioned pH ranges (see Sect. 7.1). The mechanisms of the decomposition of NAD(P)H were studied in great detail (CoLOWICK et al., 1951; OPPENHEIMER and KAPLAN, 1974;WONG and WHITESIDES, 1981).The

N-glycosidic bond between nicotinamide and ribose is the most labile bond of the molecule. Base-catalyzed hydrolysis of this bond results in the removal of the nicotinamide ring, and thus the coenzymatic activity is destroyed. On the other hand, the general acid-catalyzed protonation of the nicotinamide ring at position C , is followed by a rapid rearrangement to a cyclic ether product. Additionally, the 2 '-phosphate group of NADP(H) catalyzes this decomposition intramolecularly. The half-lives for NADH and NADPH in 0.1 M phosphate (pH 7, 25°C) are 27 h and 13 h, respectively (WONG and WHITESIDES, 1981). Organic buffers like imidazole, TRIS, HEPES, and triethanolamine destroy the reduced coenzymes at a reduced rate resulting in a three times longer half-life. The major pathway for the decomposition of NADP (6b), is nucleophilic addition at C , of the nicotinamide ring yielding a 1,4-dihydropyridine structure (JOHNSON and SMITH,1976; BIELLMAN et al., 1979; EVERSE et al., 1971). In TRIS-buffered solution at pH 7-8, NAD(P)+ is stable for at least 14 d. There is a strong dependence of the stability of the oxidized coenzymes on the concentration of the buffer. At low molarity (50 mM), the half-life in TRIS buffer is lower compared to buffer of high molarity (500 mM). Poly(ethy1ene) glycol coupled NADP+ is very stable at pH 8 in TRIS buffer of any molarity. Half-lives of more than 240 d have been reported (PETERS,1990).

3.2 Principles for the Regeneration of Nicotinamide Coenzymes 3.2.1 Nonenzymatic Regeneration Nonenzymatic methods for coenzyme regeneration can be divided into chemical, electrochemical, and photochemical methods. Chemical reduction of NAD(P)+ using a reducing agent like sodium dithionite (Na2S,0,) suffers from a very low total turnover number of less than 100 (JONESet al., 1972). Moreover, this agent can deactivate enzymes presumably by modification of thiol groups in the protein (RAMIOand LILIUS,1971). The major disadvantages of most electrochemical and photo-


chemical regeneration methods are low regioselectivity (MANDLERand WILLNER,1986; JONES and TAYLOR, 1976; LEGOYet al., 1980; JULLIARD et al., 1986),leading to coenzyme inactivation (see Sect. 3.1.3), occurrence of side reactions, and low total turnover numbers of less than lo00 (CHENAULT and WHITESIDES, 1987; CHENAULT et al., 1988). The main difficulty is the simultaneous transfer of two electrons since otherwise the formation of dimers occurs. Therefore, mediators like viologen or rhodium complexes have to be used. However, SIMONet al. (1985) stated that electro- and photochemical methods should not be underestimated. SCHUMMER et al. (1991) reported the synthesis of polyfunctional (R)-2-hydroxycarbonic acids on the preparative scale with resting cells of Proreus vulgaris by using electrochemical regeneration. RUPPERTet al. (1987), STECKHANet al. (1990), WESTERHAUSEN et al. (1992), and WIENKAMP and STECKHAN (1982) reported the electrochemical regeneration of NAD(P)(H) by using a bipyridine rhodium-(I) complex as electron transfer agent.The bipyridine rhodium complex can be coupled to poly(ethy1ene) glycol and thereby retained in an enzyme membrane reactor. D r c OSIMO et al. (1981) described the electrochemical regeneration of NAD(P)H using methyl viologen and flavoenzymes. However, enzyme denaturation on the surface of the electrode or enzyme inactivation by redox agents may occur. In summary, enzymatic methods meet most of the requirements for an ideal coenzyme regeneration system as described in Sect. 3.1.3. The difficulties encountered with the enzymatic coenzyme regeneration are mainly competitive or mixed inhibition by the substrate or product (KULA et al., 1980 LEE and WHITESIDES, 1986). In the following sections, the two general concepts for coenzyme regeneration using enzymes are presented (HUMMEL and KULA, 1989). Sect. 3.3 will then focus on the most attractive methods for the recycling of reduced and oxidized nicotinamide coenzymes.

401

3.2.2 Substrate-Coupled Regeneration In the substrate-coupled approach both the main reaction and the regeneration of the coenzyme are catalyzed by a single enzyme (Fig. 6, DH1 =DH2). The auxiliary substrate (reduced substrate 2) of the coenzyme regeneration reaction serves as the hydride donor. In order to shift the equilibrium of the reaction to the desired position, the hydride donor, usually a low cost alcohol, is applied in large excess. However, the system will run into the equilibrium of the coupled system. For the reduction of acetophenone using 2-propanol as auxiliary substrate, the equilibrium was reached at 60% yield (PETERSet al., 1993d). In the case of the reduction of 2-acetylnaphthalene, using 2-propanol as hydride donor, the equilibrium could be shifted by the use of cyclodextrins (see Sect. 5.3.2, Fig. 20) and, thereby, the total yield was increased to 93% of 1-(2-naphthyl)-ethanol (ZELINSKI, 1995). In the continuous synthesis, yields of more than 80% were obtained and total turnover numbers for the coenzyme were in the range of lo4. However, in this application the native coenzyme was not recovered from the product solution but used in substoichiometric amounts. The total turnover number for the enzyme was greater than lo6 and the productivity was 2.5 times as high as the comparable system with enzyme-coupled cofactor regeneration (formate dehydrogenase system). This demonstrates the potential of substrate-coupled systems for coenzyme regeneration. Due to the lack of a well-established method for NADPH regeneration, substrate

Oxidized substrate - 1

DH

NAD(P)H

Reduced product 1

NAD(P)

1 Reduced

Oxidized *\ product 2 DH 2

substrate 2

Fig. 6. Coenzyme regeneration methods (DH: dehydrogenase): substrate-coupled approach (DH, =DH2) and enzyme-coupled approach (DH, +DHZ).

402


coupling is mainly used for this purpose. However, there are some drawbacks of this method: - The auxiliary substrate has to be present

in a large excess, which may lead to problems in product recovery or which may cause enzyme inactivation or inhibition. - Enzyme inactivation may also occur as the result of the accumulation of a highly reactive carbonyl species like acetaldehyde or cyclohexenone. A special enzyme reactor employing a membrane, which is only permeable to gases (comparable to Gore-Tex), may avoid most of these drawbacks.This type of reactor was successfully used to improve the cofactor recycling by the yeast alcohol dehydrogenase/ethanol system in the stereoselective reduction of pyruvate (8) to D-lactate ( 9 ) catalyzed by lactate dehydrogenase (VANEICKEREN et al., 1990) (Fig. 7).

3.2.3 Enzyme-Coupled Regeneration In the case of the enzyme-coupled approach (see Fig. 6), the reduction of the main substrate and the coenzyme regeneration reaction are catalyzed by two different enzymes (LEVYet al., 1957).The enzymes employed should have sufficiently different substrate specificities to avoid interferences. The enzyme-coupled cofactor recycling has the following major advantages:

9

- The position of the equilibrium of a re-

dox reaction can be influenced. Thermodynamically unfavorable reactions can be driven by coupling with a favorable cofactor regeneration reaction. - High turnover numbers can be achieved (>600000) (SCHMIDT et al., 1987a). - The activity ratio has to be optimized for each pair of enzymes to ensure neither lack nor excess of reduction equivalents (PETERS,1993). However, there are also some disadvantages with this type of cofactor recycling method:

- Cross inhibition between the main sub-

strate and the coenzyme regeneration system may occur (PETERS,1993;WANDREY and WICHMANN, 1985;see also Sect. 6.3.2). - A second enzyme has to be used generating additional costs. - The characteristics of the second enzyme, like pH and temperature optima, have to be compatible with the main reaction.

3.3 Survey on Coenzyme Regeneration Methods and their Assessment Methods for the regeneration of NAD(P) (H) have been discussed in great detail by several authors (WANGand KING,1979; WONG and WHITESIDES, 1981, 1994; BADERet al., 1984;WONGet al., 1985;SIMON et al., 1985;LEE and WHITESIDES,1985;CHENAULT and WHITE-

Gas membrane

Fig. 7. Coenzyme recycling using a gas membrane reactor. LDH, lactate dehydrogenase; YADH, yeast alcohol dehydrogenase.


403

SIDES, 1987; CHENAULT et al., 1988; HUMMEL and KULA, 1989; POPPE and NOVAK,1992; FABER,1995). In this section, only the most useful methods are discussed and assessed with respect to their applicability in batch and continuous synthesis. Furthermore, recently developed new methods for nicotinamide coenzyme recycling are highlighted.

3.3.1 Regeneration of NAD(P)H In Tab. 4, the major disadvantages of a selec1 tion of available cofactor recycling systems are 0 0.2 0.4 0.b 0.8 listed, and each method is assessed with respect to its applicability in batch or continuous CPCR / (CPCR + FDH) processes. For the regeneration of native and poly- Fig.8. Conversion as a function of the activity ratio (ethylene) glycol-bound NADH (PEG- of Candida parapsilosis carbonyl reductase (CPCR) NADH), the formate dehydrogenase (FDH) to formate dehydrogenase (FDH). The coenzyme from Candida boidinii (SCHUTTEet al., 1976) regeneration has to be efficient to achieve a high At the optimal activity ratio of 0.2, the rahas been widely and successfully applied turnover. tio of activities of CPCR and FDH is 1 :4(PETERS, (SHAKED and WHITESIDES, 1980;WICHMANN et 1990,1993). al.. 1981;SCHMIDT et al., 1987a. b; VASIC-RACKI et al.. 1989). The enzyme catalyzes the oxidation of inexpensive formate (HCOO-) to carbon dioxide (CO,). The reverse reaction is formate dehydrogenase does not accept hardly detectable (SCHUTTEet al., 1976). NADP(6b). Therefore, the FDH reaction can shift the Although several NADP-dependent forequilibrium of unfavorable reactions to the mate dehydrogenases have been discovered product side. Both, the auxiliary substrate and (YAMAMOTO et al., 1983; ANDREESEN, 1974; the coproduct are innocuous to enzymes and LJUNDAHL and ANDREESEN, 1975; KRUMHOLZ are easily removed from the reaction. There- et al., 1987; TANNENBAUM, 1956), only the refore, product isolation is easy. FDH has a cently described mutant formate dehydrogebroad pH optimum of activity so that it can nase from Pseudomonas sp. 101 (TISHKOV et easily be implemented in coupled enzymatic al., 1993a, b) accepting NADP as coenzyme synthesis.The major disadvantages are the low was applied in batch and continuous synthesis. specific activity of 4 U mg-' and the product Using this enzyme for the regeneration of inhibition by NADH (7a). However, these NADPH, acetophenone was reduced by the drawbacks can be circumvented or minimized: NADPH-dependent alcohol dehydrogenase the former by using immobilized or mem- from Lactobacillus sp. (EC 1.1.1.99) at a spacebrane-retained systems (SHAKEDand WHITE- time yield of 8 g-' d-' and a conversion rate SIDES, 1980;HUMMEL et al., 1987),the latter by of 90-95% (SEELBACH, 1994).The new FDH is adjusting the optimal ratio of enzymatic activ- quite stable showing no loss in activity over ities for the main reaction and the coenzyme one year at 4°C (SEELBACH et al., 1996). Unrecycling reaction (PETERS,1993) (Fig. 8). fortunately, this enzyme is not yet commercialOverall, the formate/FDH system is the ly available. A further disadvantage is the relamost economical method for regeneration of tively high K , value of the mutant FDH for NADH for batch and continuous processes. NADP (320 pM).Additional work has to be The total turnover numbers (TTN) are ranging done to engineer FDH mutants with improved typically from lo3 to lo5 (SCHMIDTet al., kinetic properties. In conclusion, the new for1987a,b; HUMMEL et al., 1987). Unfortunately, mate: NADP dehydrogenase (EC 1.2.1.2) has 1.0

404 Tab. 4.

9 Dehydrogenases - Characteristics, Design of Reaction Conditions, and Applications Methods for in situ Regeneration of NAD(P)H

Methods

Major Disadvantages

Assessment

Glucose/glucose dehydrogenase

K , value for PEG-NADP+ loo0 times that of native NADP+ - complication of product isolation caused by gluconate - high amounts of K + or Na+ necessary to stabilize enzyme

useful regeneration system for native NAD(P)H in batch or continuous mode of operation

Glucose-6-P/ glucose-6-P dehydrogenase

- high costs for glucose-6-phosphate ( 5500 $ mole-') - coenzyme inactivation by 6-phospho-gluconate

useful only for lab-scale reactions. Regeneration of NADH and NADPH.

-

-

- no efficient usage of the coenzyme - complication of product recovery caused by 6-phosphogluconate

- complication of product recovery caused by 6-sulfo-glu-

circumvents some limitations of the former method. Useful for preparative applications in continuous reactors. Regeneration of NADP and NADPH.

Ethanol/alcohol and aldehyde dehydrogenase

- relatively complicated and unstable multienzyme system

useful only for lab-scale reactions. Regeneration of NADH and NADPH.

Formate/formate: NAD+ dehydrogenase

- low specific activity (3 U mg-') - inactivated by autooxidation - product inhibition caused by (PEG-)NADH

simple and stable system for regeneration of (PEG-)NADH in batch or continuous mode of operation up to the process scale. Very good compatibility with other enzymes.

Formate/formate: NADP' dehydrogenase

- enzyme not commercially available yet

simple and stable system for regeneration of NADPH in batch or continuous mode of operation. Very good compatibility with other enzymes.

sec. Alcohol/ Thermoanaerobium brockii alcohol dehydrogenase

- deactivation by autooxidation possible - relatively expensive enzyme (58 centdunit)

simple and stable system for regeneration of (PEG-)NADPH in batch or continuous mode of operation. Very good compatibility with other enzymes.

Glucose-6-sulfate/glucose-6-P dehydrogenase

conate - substrate is only accepted by glucose-6-phosphate dehydrogenase from yeast

with low TTN - oxygen-sensitivity of alcohol- and aldehyde dehydrogenases - low reactivity of the aldehyde dehydrogenase with NADP+ - only activated carbonyl substrates are reduced at good yields, which may cause enzyme inactivation

PEG: poly(ethy1ene) glycol; TTN: total turnover number; K,: Michaelis-Menten constant; Kip: product inhibition constant.


405

a large potential for use as a regeneration Overall, the TBADH is recommended for system for NADPH. the recycling of NADP(H) (6b, 7b) for the folThe secondary alcohol dehydrogenase from lowing reasons. First, the enzyme is commerThermoanaerobium brockii (TBADH) ac- cially available and is stable in a polypropycepts a broad range of secondary alcohols and lene enzyme membrane reactor. Second, the NADP+ as coenzyme (LAMEDand ZEIKUS, specific activity is almost 10 times as high as 1981). The enzyme is commercially available the formate dehydrogenase. Favorably, the K , and contains a very low NADP-2'-phospha- values for native and PEG-bound NADP(H) tase activity (PETERS,1990). NADP(H) is hy- are low (10 pM),and the product inhibitions drolyzed at the 2'-ribose phosphate bond at a by 2-butanone and (PEG-)NADPH are also rate of 0.1% d-' in the continuous synthesis of low. Third, the TBADH is compatible with NADPH (PETERSand KULA,1991). This en- high amounts of organic solvent (e.g., 30% zyme has been used in the reduction of numer- isopropanol) and stable at elevated temperaous carbonyl compounds using the substrate- tures. coupled approach for coenzyme recycling (KEINANet al., 1986a, b, 1987, 1992; LAMEDet al., 1981; KELLYand LEWIS,1991). Since this enzyme tolerates up to 30% isopropanol, the substrate-coupled regeneration of NADPH is 3.3.2 Regeneration of NAD(P) highly favorable. The oxidized nicotinamide coenzymes are However, the TBADH is also an excellent system for the regeneration of native and po- used for the synthesis of carbonyl compounds ly(ethy1ene) glycol-bound NADPH (PETERS, from the corresponding racemic hydroxy com1990; PETERSand KULA,1991). In a model pounds. Both, enzymatic and nonenzymatic system, glutamate dehydrogenase (GluDH) methods have been reported for their regenerwas employed for the synthesis of glutamate ation (for a review see WONGand WHITESIDES, from a-ketoglutarate. A complete set of kinet- 1994). However, enzymatic methods seem to ic data was recorded, and the effectivity of the be preferred because of their compatibility regeneration of PEG-NADPH by the with biological systems and their simplicity. TBADH was studied in a batch reactor. The The regeneration of oxidized nicotinamide cooptimal activity ratio of GluDH/(GluDH+ enzymes is somewhat problematic due to the TBADH) was 0.6. At this ratio, 95% of the to- thermodynamics which are often unfavorable. tal coenzyme is reduced. The conversion of a- Additionally, product inhibition by the reketoglutarate is affected negatively, only at a duced coenzyme may cause problems (see beTBADH portion of less than 20%. For com- low). In Tab. 5 the major disadvantages of a selecparison, in the leucine dehydrogenaseiFDH system studied by WICHMANN et al. (1981) on- tion of available cofactor recycling systems are ly 10% PEGiNADH is present at the optimal listed and each method is assessed with respect activity ratio of 1 :1.The reason for this differ- to its applicability in batch or continuous proent behavior is the product inhibition of the cesses. The alcohol dehydrogenase from PseudoFDH by PEG-NADH. In contrast, the TBADH shows only slight product inhibitions rnonas sp. (see Sect. 7.2.4), which is not comby 2-butanone and (PEG-)NADPH. The loss mercially available, has potential for the recyof coenzyme in the enzyme membrane reactor cling of NAD+ (SHENet al., 1990; BRADSHAW is a function of the absolute coenzyme concen- et al., 1992a,b). A range of simple ketones can tration (KULA and WANDREY,1987) and, be used as hydrogen acceptors which are retherefore, should be as low as possible. The K, duced at high velocity (up to 270 U mg-'). value of the TBADH for native and PEG- However, the kinetic limitations of this enbound NADPH is 10 pM. Hence, only 0.2 to zyme have to be explored in more detail. For recycling of NADP', the secondary al0.3 mM NADP(H) are usually sufficient to ensure a maximum usage of the catalytic activ- cohol dehydrogenase from Thermoanaerobium brockii (TBADH) is problematic due to ities in the reactor.

406


Tab. 5. Methods for in situ Regeneration of NAD(P)+

Methods

Major Disadvantages

Assessment

AlcohollPseudomonas sp. alcohol dehydrogenase

- enzyme is not commercially available

sec. Alcohol/ Thermoanaerobium brockii dehydrogenase

- strong product inhibition by (PEG-)NADP+

not useful due to the severe product inhibition by (PEG-)NADP+.

Acetaldehyde/ yeast alcohol deh ydrogenase

- self-condensation of acetaldehyde

not useful for the regeneration of NAD + due to side reactions and inactivation by acetaldehyde.

a-ketoglutarate/ glutamate dehydrogenase

- glutamate may complicate product isolation

simple and stable system for regeneration of (PEG-)NAD(P)H. Good compatibility with other enzymes.

Pyruvate/lactate dehydrogenase

- pyruvate tends to polymerize in solution and reacts with

simple and stable system for regeneration of NAD +.Side reactions may happen.

(KipNADP):50 yM; K, (NADPH): 10 pM)

- enzyme inactivation by highly reactive acetaldehyde

NAD+ in a process catalyzed by lactate dehydrogenase

PEG: poly(ethy1ene) glycol; K,: Michaelis-Menten constant; K,p: product inhibition constant.

the low product inhibition constant Kip for NADP+ of 50 pM (PETERS,1990; PETERSand KULA,1991). The K, value of NADPH is 10 pM. Therefore, the oxidation of alcohols producing the reduced coenzyme is the favorable reaction. Acetaldehyde and the commercially available alcohol dehydrogenase from bakers' yeast (YADH) have also been used to regenerate NAD+ from NADH (LEMIEREet al., 1985). The total turnover numbers were 103-104. However, the deactivation of the enzyme by the highly reactive acetaldehyde and the self-condensation of acetaldehyde outweigh the advantages of the inexpensive enzyme and the volatility of the reagents involved. The most widely applied method for regeneration of oxidized nicotinamide coenzymes involves the glutamate dehydrogenase (GluDH) which catalyzes the thermodynami-

cally favorable reductive amination of inexpensive a-ketoglutarate to give L-glutamate (LEE and WHITESIDES, 1986; CARREAet al., 1984). Both the substrate and the product are innocuous to enzymes. The commercially available and inexpensive GluDH accepts NADH, NADPH, and the respective poly(ethylene) glycol-bound coenzymes (PETERS, 1990) and has a reasonably high specific activity of 40 U mg-' protein. The products of the reductive amination of a-ketoglutarate, (PEG)NAD(P)+ and L-glutamate do not inhibit the reaction strongly (PETERS,1990). The ratio of the product inhibition constant to the Michaelis-Menten constant (Kip/Km)will determine the efficacy of the reaction (LEE and WHITESIDES,1986).At a Kip/Kmratio of greater than 1,the reaction may proceed to acceptable conversions. If this ratio is less than 1, the reaction can never proceed efficiently.The Ki,IK, ratio for the substrate and product of the reductive


amination of a-ketoglutarate to L-glutamate is 10.5. For the (PEG-)NADPH and (PEG-)NADP’ the Ki,/K,,, ratio is about 30 (PETERS, 1990). Hence, product inhibition by L-glutamate and (PEG-)NADP+ does not play a significant role for the efficacy of the regeneration reaction. The disadvantage of this regeneration method is that L-glutamate may complicate the product isolation from the reaction mixture. This has to be addressed case by case. Pyruvate and the commercially available and inexpensive lactate dehydrogenase (LDH) have been used for recycling of NAD (BEDNARSKIet al., 1987; CHENAULTand WHITESIDES, 1987).The LDH is stable and has a high specific activity of about 1000 U mg-’ protein. However, the redox potential is less favorable and, in contrast to the GluDH, LDH does not accept NADPH. A further disadvantage is that pyruvate tends to polymerize in solution and reacts with NAD’ in a process catalyzed by LDH. Despite these drawbacks, the pyruvate/LDH system has been applied in enzymatic oxidation reactions of 10-100 mmol of material. +

3.4 Immobilized Coenzymes and React or Concepts Enzyme-catalyzed reactions are often accompanied by the following disadvantages:

- Enzymes may not be sufficiently stable

under the reaction conditions employed. Some lose their catalytic activity due to autooxidation, due to proteolytic degradation andlor denaturation by the solvent, and others due to mechanical shear forces. - Isolated enzymes are water-soluble and have to be used repeatedly because of their cost (SUCKLING and SUCKLING, 1974).They have to be separated from the reaction mixture without destroying their catalytic activity. - The productivity, expressed as the spacetime yield, is often low due to the limited tolerance of enzymes to high concentrations of substrate(s) and product(s).

407

Some of these problems may be overcome by “immobilization”of the enzyme. However, a special problem is encountered with NAD(P)(H)-dependent dehydrogenases since the coenzyme has to associate and dissociate freely to and from the active site(s) of the enzyme(s). Therefore, co-immobilization of the dehydrogenase and the cofactor is necessary to make the system practical (GESTRELIUS et al., 1975). However, either dehydrogenase or coenzyme or both may decompose leading to the replacement of the immobilized system. If the K , value for the coenzyme is in the micromolar range and the substrate-coupled approach for coenzyme regeneration is employed (see Sect. 3.2.2), a column reactor can be used with a small amount of free coenzyme dissolved in the mobile phase (KEINAN et al., 1986a).This situation is particularly efficient, if the reaction mechanism is an ordered bi-bi- or Theorell-Chance mechanism where the coenzyme is bound first to the active site, followed by the substrate. These mechanisms were found for a broad range of alcohol dehydrogenases (see Sect. 6.3). The immobilization techniques most widely used for coenzyme-dependent dehydrogenases are shown in Fig. 9. Immobilization techniques for coenzymes may be divided into coupling and entrapment methods.

3.4.1 Coupling Methods For dehydrogenases, coupling methods can be further divided into methods for coupling the coenyme on cross-linked enzymes or methods for coupling the enzyme and coenzyme on a carrier.

3.4.1.1 Coupling of the Coenzyme to Cross-Linked Enzymes Enzymes can be attached to each other by covalent bonds which is termed “cross-linking”. Therefore, bifunctional reagents like glutardialdehyde, dimethyl adipimidate, dimethyl suberimidate, and others can be used. The coenzyme can be bound to cross-linked enzymes using a spacer long enough so that the coenzyme has access to the active sites of both en-

408

9 Dehydrogenases - Characteristics,Design of Reaction Conditions,and Applications

VJ

to Cross-Llnkod -E

0 0

0 0

outside

w

0-

PEG-^^ coenzyme

substrate oproduct o

magnetic stirrer bar

dialysis bag

Ct4tOIyriS

(MEEC)

zymes thereby transferring the hydride. These requirements are very difficult to meet in practice. LORTIE et al. (1989) reported on the coimmobilization of horse liver alcohol dehydrogenase (HLADH), albumin, and NAD using glutaraldehyde as the cross-linking agent. Long-chain aldehydes were prepared using this immobilized H L A D W A D system in a packed bed reactor (glass column). The conversions were performed in organic solvent (hexane) saturated with buffer. Since proteins and coenzymes are insoluble in the water-immiscible hexane they remained in the aqueous phase. The long-chain alcohols (substrate) diffused into the aqueous phase, underwent enzymatic conversion, and the product +

Methods for immc-ilizatiin of coenzymes. a Coupling methods. b Entrapment methods

(aldehyde) finally diffused back into the organic phase. This system was limited by the mass transfer rates between the organic phase and the water containing porous particles. LORTIEand coworkers generalized this result to all types of fixed bed enzyme reactors in biphasic systems. However, the regeneration of the coenzyme was not a rate limiting step.

3.4.1.2 Coupling of the Coenzyme on a Carrier Another option for immobilizing coenzymes is to bind both the dehydrogenase(s) and the coenzyme to an insoluble carrier like cellulose,


409

A simple form of a membrane reactor which dextrans, starch, chitin, agarose, or synthetic copolymers (CARR and BOWERS,1980; LEE does not require any special equipment is and CHEN,1982).The activation of the hydrox- termed “Membrane-Enclosed-Enzymatic-Cayl groups of polysaccharides is usually done by talysis” (MEEC). The enzyme(s) and the polyreaction with cyanogen bromide leading to the mer-bound coenzyme are entrapped in a dialyformation of reactive imidocarbonates. Syn- sis bag, mounted on a gently rotating magnetic et al., 1987). thetic polymers can be activated using partial stirring bar (BEDNARSKI Modifications of the enzyme membrane rehydrolysis of the acetate groups followed by activation with epichlorohydrin. A range of actor concept make use of charged ultrafiltrapre-activated polysaccharides and synthetic tion membranes in order to retain the native and ISHIMATSU, polymers (Biosynth, Eupergit, etc.) may be ob- coenzyme in the reactor (IKEMI tained from commercial sources. Also in this 1990;IKEMIet al., 1990;ROTHIGet al., 1990a,b; et al., 1996a, b). The application of case, a spacer long enough to allow the coen- NIDETZKY zyme to have access to the active sites of the charged ultrafiltration membranes in memenzyme(s) has to be used. Horse liver alcohol brane reactors is limited to the production of dehydrogenase (HLADH) and an NADH an- noncharged products. In contrast, nanofiltraalog N6-[(6-aminohexyl)carbamoyl-methyl]- tion membranes can be used to retain the coNADH, have been successfully co-immobi- enzyme in the reactor whereas the reaction lized to Sepharose 4B (GESTRELIUS et al., product(s) can freely pass the membrane and KRAGL,1997).The separation 1975).The coenzyme could be regenerated at a (SEELBACH cycling rate of 3400 cycles h-’ using the subof coenzyme and reaction product(s) is due to strate-coupled approach. Both thermal and the differences in the molecular weight withstorage stability of the HLADH were in- out use of a negative net charge of the memcreased substantially. No soluble coenzyme brane. For further details on the applications of dehad to be added to the substrate solution. hydrogenases using the enzyme membrane reactor technology, the reader is referred to Sects. 5 and 7. Many reports on the preparation of poly3.4.2 Entrapment Methods mer-linked coenzyme derivatives are available and MOSBACH, 1974; ZAPELLI et al., Polymer-linked coenzyme derivatives are of- (LARSSON and BRIGHT,1977;BUCKMANN et ten introduced into enzyme reactors with co- 1975;FULLER et al., 1985; BUCKMA” enzyme-dependent processes, since they can al., 1981a, b; OKUDA and CARREA, 1989). be retained upon ultrafiltration or dialysis and 1987,1988;BUCKMANN The dextran-bound coenzymes have been are recycled (see Fig. 7; WANGand KING,1979; KULA et al., 1980; KATAYAMA et al., 1983; demonstrated to have a serious problem of WANDREYet al., 1984; LEUCHTENBERGER et leakage, because of the instability of the isoa]., 1984b; WANDREYand WICHMANN, 1985; urea linkage between coenzyme derivatives KULA and WANDREY, 1987; SCHMIDTet al., and dextran (AXENet al., 1967). However, 1987a, b; HOSONO,1988; VASIC-RACKI et al., more stable dextran derivatives can be synthe1989;KRAGLet al., 1989;STECKHAN et al., 1990; sized by using bromohydroxypropyl derivaKISE and HAYASHIDA, 1990). Small substrate tives of dextrans (LEE et al., 1978).These deand product molecules can freely pass the rivatives are used mostly in enzyme electrodes membrane (e.g.,polyamide, polyethersulfone), or as stationary phases in affinity chromato1974). but the large enzyme(s) and the polymer- graphy (LOWEand MOSBACH, The polyethyleneimine and polylysine delinked coenzyme cannot. Synthetic membranes of defined pore size covering the range rivatives of coenzymes (ZAPELLIet al., 1975) between 500 and 300000 Da are commercially have the disadvantage of the electrostatic available at reasonable cost. Additionally, a va- interactions between the multicharged polyriety of synthetic membrane shapes are on the mers and enzymes, thus interfering with their market such as membrane discs or hollow fi- effectiveness as active coenzymes.The relative reaction rates compared to native NAD were bers.

410


2% to 25% (polyethyleneimine) and 2% to organic solvents have been worked out by (1990), KRUSE et al. 60% (polylysine) depending on the enzyme KISE and HAYASHIDA (1996), LIESEet al. (1996), and LIESEet al. (in tested. The synthesis of polyacryl copolymers has press). These concepts circumvent the immothe advantage that parameters like solubility, bilization of the coenzyme by using hydrophocharge and the functional group can be varied bic microfiltration membranes and two-phase just by changing the monomer (FULLERand systems, an emulsion membrane reactor or a BRIGHT,1977). Therefore, synthesis is much differential circulation reactor with continueasier. However, polyacryl-bound NAD ous extraction, respectively. For more details showed only 30% of theVmaxvalue of the na- please refer to Sect. 5. tive coenzyme (FULLER et al., 1980). Another approach was established by BUCKMANN and coworkers using poly(ethy1ene) glycol derivatives with a molecular 3.5 Conclusions weight of 3000 to 20000 and modified coEnzymatic reductions catalyzed by NAD(P)enzymes.The yield of the coupling of N(1)-(2aminoethy1)-NAD with the carboxy-PEG dependent dehydrogenases require stoichioderivative was 90% (BUCKMANN et al., 1981a). metric amounts of nicotinamide coenzymes, The alkylation of NAD(P)(H) with 1,4-bis- which are very expensive compounds. The ap(2,3-epoxypropoxy)-butane was investigated plication of dehydrogenases on a preparative by STEIN(1990).Thismethod allows the direct scale, therefore, requires effective methods for alkylation of NAD(P)H in a one-step reaction, the regeneration of these coenzymes. Today, coenzyme regeneration of both whereas all other described alkylation methods are multi-step reactions starting from the NADH and NADPH are very much straightoxidized coenzymes. Moreover, the coenzy- forward. Therefore, the cofactor is no longer matic activities of the NADP(H) derivatives the dominant cost in preparative reductions. were almost identical to the native coenzymes. Usually, the costs of the enzyme(s) and other The NAD(P)H derivatives carry a hydrophilic reagents is much greater than the cost of the spacer and a terminal epoxy group, which can coenzyme. Nicotinamide coenzymes are labile combe coupled easily to PEG-NH, resulting in a stable, covalent bond between PEG and the pounds, but, if they are used under optimal coenzyme derivative.For most of the enzymes conditions, the half-life can be extended to tested, the coenzymatic activity of PEG- more than 14 d. Coupling of NAD(P)(H) to NAD(H) is comparable to the native co- poly-(ethylene) glycols even stabilizes the coenzyme (BUCKMANN et al., 1981a; KATAYAMAenzymes, by which means the half-life can be et al., 1983,1984). However, there are also ex- extended to more than 240 d. Enzymatic methods for coenzyme regeneraceptions (BUCKMANN et al., 1987). The PEGcoenzyme derivatives have been successfully tion are preferable over nonenzymatic methapplied in the above mentioned enzyme mem- ods since they meet most of the requirements brane reactor (KULAet al., 1980;KATAYAMA et for an ideal recycling system. Two different al., 1983;WANDREY et al., 1984;LEUCHTENBER-principles can be distinguished, the substrateGER et al., 1984; WANDREY and WICHMANN,and the enzyme-coupled approach. The en1985; KULAand WANDREY, 1987; SCHMIDT et zyme-coupled method is advantageous since al., 1987a,b; HOSONO, 1988;VASIC-RACKI et al., even unfavorable reactions can be driven to 1989; KRAGLet al., 1989; STECKHAN et al., the product side by coupling with a favorable 1990).PEG-NAD(H) as well as other PEG de- cofactor regeneration reaction. The most economical method for (PEG-)rivatives are commercially available from spinoffs of the German research centers in Braun- NADH-recycling is the formate/formate dehyschweig and Julich (see Tab. 15;ASA Spezial- drogenase system, which was used up to the process scale. enzyme GmbH, Julich Enzyme Products). For the regeneration of (PEG-)NADPH, Modern reactor concepts for the application of dehydrogenases and native coenzymes in the alcohol/alcohol dehydrogenase system +

4 Assessment of Straregiesfor the Synthesis of Chiral Hydroxy Compounds

from Thermoanaerobium brockii or the formate/formate :NADP dehydrogenase system are recommended. The a-ketoglutaratelglutamate dehydrogenase system catalyzes the thermodynamically favorable reductive amination of a-ketoglutarate. Both, NAD' and NADP' as well as PEG-NAD(P) can be regenerated effectively. The use of enzyme membrane reactor technology in conjunction with poly(ethy1ene) glycol-linked coenzymes has made it possible to scale up enzymatic processes to a preparative scale (LEUCHTENBERGER et al., 1984b; BOMMARIUS et al., 1995; KRAGL et al., 1996a). Amongst others this technology was successfully used for the synthesis of different L-amino acids and a range of chiral hydroxy compounds of high value (for examples see Sects. 5 and 7). +

4 Assessment of Strategies for the Synthesis of Chiral Hydroxy Compounds 4.1 Introduction

411

droxyacids or their corresponding esters (HASEGAWA et al., 1981a, b; AKITAet al. 1984a, b, 1982; BELANet al., 1987; FAUVEand VESCHAMBRE, 1987, 1988; LARCHEVEQUE et a]., 1988; MORI,1981,1989; KEINANet al., 1990). In this section, the enzymatic synthesis of a chiral hydroxycarboxylic acid ester is taken as an example to discuss and assess different strategies for the synthesis of a chiral hydroxy compound in general. The criteria for the assessment are the following: The position of a resolution of a racemic compound in the synthetic route should be as early as possible. Additional steps should be as few as possible. The products of an enzymatic resolution of a racemate should be separable (e.g., extraction, distillation). The synthetic route, especially to the desired enantiomer, should be as short as possible. The enantiomeric purity of the product should be more than 80% ee. The space-time yield of the enzymecatalyzed reaction should be high enough to allow a productivity of several tons per year. The catalyst should be stable and accessible in reasonable quantities.

The employment of chiral building blocks is the most economic strategy for the synthesis of enantiomeric pure compounds (EPC)(GRAF, 4.2 Survey of Different Strategies 1985),which play an increasing role in the synthesis of pharmaceutically active compounds, for the Synthesis of Chiral Hydroxy agro and veterinary chemicals. The market for Compounds and their Assessment chiral building blocks is currently estimated at about 1.8 billion dollar sales (POLASTRO, In the following sections, the most attractive 1991), containing chiral hydroxy compounds alternatives for the formation of a chiral hyas an important subgroup. Chiral hydroxyacids droxyacid ester are discussed with respect to or esters have been proved to be especially the above listed criteria. However, there are versatile synthons, due to their useful dual further routes which are less attractive in functionality (HASEGAWA et al., 1981a,b; HAS- terms of the number of additional synthetic EGAWA and OHASHI, 1996). A broad range of steps. In this example, the compound of interpharmaceutical compounds (e.g., carbapenem est is the tetrahydropyranyl derivative of (3.9)antibiotics, statin analogs, L-carnitine, leuco- methyl-3-hydroxy-butanoate (S)-(13) (Fig. 10). trienes/prostaglandins and benzothiazepine compounds), pheromones, agrochemicals,aromatic compounds, and inhibitors (e.g., compactin) can be synthesized starting from P-hy-

412

9 Dehydrogenases- Characteristics,Design of Reaction Conditions,and Applications

cially used for the synthesis of (S)-and (R)-hydroxyacids or ester (HASEGAWA and OHASHI, The direct enantioselective reduction of a p- 1996; Kaneka Corporation, Japan). In the literature, a number of isolated enketoacid ester using isolated enzymes or microorganisms does not lead to an increased zymes have been described catalyzing the number of steps, since there are no separation asymmetric reduction of p-ketoacids or esters. problems of different isomers if the enantio- Bakers' yeast, Saccharomyces cerevisiae, contains four NADPH-dependent p-ketoacid remeric excess is high enough (Fig. 10). A wide range of microorganisms have been ductases (HEIDLASet al., 1988; SHIEHet al., discovered to catalyze the asymmetric reduc- 1985; SHIEH,1987; NAKAMURA et al., 1991). tion of p-ketoacids or esters (POPPEand No- Another NADPH-dependent dehydrogenase VAK,1992; CHRISTEN and CROUT,1987; WARD active on p-ketoacid esters was isolated from and YOUNG,1990; BUISSON et al., 1992). De- another yeast, Hansenula silvicola, by NASSENpending on the reaction conditions, additives, STEIN et al. (1992).These enzymes are not comand the source of the organism, the yield and mercially available. The extreme thermophilic enantioselectivity of bakers' yeast reductions bacterium Thermoanaerobium brockii conof p-ketoacids or esters varies between 10% to tains a NADPH-dependent secondary alcohol 85% and 40% to 98'30, respectively. Besides dehydrogenase with a very broad substrate bakers' yeast, several other yeasts and fungi spectrum reducing, amongst other substrates (Hsu et al., 1 9 8 3 ; A ~ et l ~al., ~ 1982, 1984a; p-ketoacids or esters. This enzyme is very USHIO et al., 1986; AZERADand BUISSON, stable and commercially available. All of the above mentioned oxidoreductas1992; BUISSONet al., 1992; HASEGAWA et al., 1981a;HASEGAWA and OHASHI, 1996) and bac- es are NADPH-dependent enzymes.However, teria (MACONIand ARAGOZZINI, 1989; CHRIS- two novel reductases from the yeast Candida TEN et al., 1992) can also reduce p-ketoacids or parapsilosis and the bacterium Rhodococcus their corresponding esters stereoselectively. erythropolis have been described recently, reSome of these microorganisms are commer- ducing a broad range of p-, y-,and Gketoacid esters using NADH as coenzyme (PETERSet al., 1992, 1993a). Both enzymes, however, are not commercially available yet, but can be isolated quite easily (PETERSet al., 1993b;ZELINSKI et al., 1993). The carbonyl reductase from Candida parapsilosis (CPCR) is especially stable under reaction conditions over extended periods of time (ZELINSKI, 1995). It was DH shown in the CPCR-catalyzed reduction of carbonyl compounds that, in a continuous process, high space-time yields (118 g L-' d-' can R (S)-11 be achieved (ZELINSKI, 1995). The above listed isolated enzymes catalyzing the stereoselective reduction of p-keto12 DHP acids or esters are highly enantioselective, yielding more than 95% ee. In total, the direct reduction of a p-ketoacid or ester by either microbial or enzymatic means is straightforward. Most of the above mentioned criteria are met. Moreover, the en(S)-13 zymatic processes do not form undesired side products complicating the work-up of the reFig. 10. Direct reduction of a 0-ketoacid using a NAD(P)H-dependent dehydrogenase and protec- action mixture. The desired stereoselectivity tion with dihydropyran (DHP) to give the tetrahy- can be chosen by selecting the appropriate microorganism or isolated enzyme. dropyran (THP) derivative (S)-(13)

4.2.1 Enantioselective Reduction

1

VOMe

413

4 Assessment of Strategies for the Synthesis of Chiral Hydroxy Compounds

4.2.2 Enzymatic Hydrolysis of the Racemic Hydroxyacid Ester Compound In this approach, the p-ketoacid ester (10)is reduced chemically yielding a racemic hydroxy-acid ester ruc-(11).Two different routes can be used to synthesize the compound of interest using lipases as catalysts (Fig. l l a , b). First, the undesired enantiomer can be hydrolyzed resulting in the (S)-hydroxyacid ester (S)-(11) and the (R)-hydroxy acid ( R ) (14) (Fig. lla). The latter is either rejected or has to be esterified and reoxidized to the orig-

inal P-ketoacid ester (10). The lactonization of the p-hydroxyacid (R)-(14) may be avoided by using buffered solutions which stabilize the hydroxycarboxylic acid by forming a salt bridge. In total, one enzymatic resolution step is necessary to synthesize the desired (S)-hydroxyacid ester (S)-(11) and two additional steps have to be performed to esterify and reoxidize the undesired hydroxyacid (1 2 additional steps). Second, the desired enantiomer can be hydrolyzed giving the (S)-hydroxyacid and the (R)-hydroxyacid ester (Fig. llb). The latter is either rejected or has to be reoxidized to the original p-ketoacid ester. Since, in this exam-

+

1 Esterification; 2 Oxidation

R uOH

(R)-14

R 10

ruc-1 1

THPG Rd

R

12 DHP, pTsOH O

M

e

OMe ( S ) - l l

I

Oxidation

uoMe uoMe uOMe

NaBHr

R

Esterase

R

rac-1 1

10 THPG Rd

(R)-11

(S)- 14

1 Esterification; 2 12 DHP, pTsOH

O

M

e

Fig. 11. Enzymatic hydrolysis of the racemic hydroxyacid ester compound. a The undesired enantiomer is hydrolyzed, b the desired enantiomer is hydrolyzed.


414

ple, the hydroxyacid ester is used for the following synthetic steps, the desired (S)-hydroxyacid (S)-(14) has to be esterified with methanol and protected. In total, two steps (resolution step and esterification) are needed to synthesize the desired (S)-hydroxyacid ester (S)-(13),and one additional step has to be performed to reoxidize the undesired hydroxyacid ester (R)-(11) (2+ 1 additional steps). In both cases, the separation of the hydroxyacid and ester products may be achieved by extraction.

4.2.3 Enzymatic Hydrolysis of the Racemic Ether Derivatives of the Hydroxy Compound In this case, the resolution of the racemate occurs after derivatization of the hydroxy

group of the racemic hydroxyacid ester with dihydropyran (DHP). Therefore, this reaction variant does not meet the criterium (1) (resolution of racemates as early as possible). Four different strategies are possible. First, the acetal of the undesired enantiomer is hydrolyzed (Fig. 12a). Second, the acetal of the desired enantiomer is hydrolyzed (Fig. 12b). The latter is highly unfavorable, since four additional steps are generated in total. In the first case, only two additional reaction steps have to be performed, since the product of the enzymatic resolution step can be processed further without modification, and the undesired ( R ) hydroxyacid ester (R)-(11) is reoxidized to the corresponding P-ketoester (lo), Third, the hydrolysis takes place at the ester function of the desired (S)-enantiomer (Fig. 12c).The coexistence of a carboxylic acid and an acid-labile dihydropyran acetal may be a problem. This can be solved possibly by using a buffered solu-

Oxidation i

1 NaBH,

RU

O

M

e

RW

r

THPO

2 12 DHP, pTsOH

O

M

e

( R ) - 11

R

10

rac-13

(S)-1 3 1 Hydrolysis; 2 Oxidation

1 NaBH,

R=OMe 10

THPQ R

b)

(S)-1 3

2 12 DHP, pTsOH

THF'O

(R)-13

rac-13

12 DHP, pTsOH

LR

XOMe (S)-1 1


1 Hydrolysis; 2 Oxidation

415

THPO uOMe

THPo

1 NaBH4

RU

O

M pTsOH e

K

10

rac-13

THPQ RA

(R)-13

Esterase

(S)-1 4

Esterification O

M

e

TWO pTsOH 10

4

(R)-14

THPQ rac-13 (S)- 13

Fig. 12. Enzymatic hydrolysis of the racemic ether derivatives of the hydroxy compound. a Acetal cleavage of the undesired enantiomer, b acetal cleavage of the desired enantiomer, c ester hydrolysis of the desired enantiomer, d ester hydrolysis of the undesired enantiomer.

tion. In this case, an additional esterification reaction with methanol has to be performed subsequently (2 + 1 additional steps). Fourth, hydrolysis takes place at the ester group of the undesired (R)-enantiomer (Fig. 12d). This sequence is favorable to the third variant, since the valuable product stream is shorter. However, the number of additional steps is the same (1+2).

4.2.4 Enzymatic Hydrolysis of the Racemic Acyl Derivatives of the Hydroxy Compound In this approach, the racemic hydroxy group of the hydroxyacid ester is esterified followed

by an enzymatic resolution step. The hydrolysis of the acyl group (e.g., butyric or acetic acid) can take place next to the stereogenic center, whereas in case 4.2.2 the hydrolysis takes place at the more distant carboxyl group. Again, either the undesired (3 + 1 additional steps) (Fig. 13a) or the desired enantiomer (2+2 additional steps) (Fig. 13b) can be hydrolyzed. Although the total number of additional steps is equal, the latter alternative is more favorable, since one recycling step is moved to the low-valued product stream. In the second case, two additional steps are necessary to obtain the desired intermediate, the (S)-hydroxyacid ester. Two more steps, a deprotection and a reoxidation, have to be done to recycle the undesired product.

416


Oxidation

ruc-15

10

4

(R)-1 1

Es terase

1 NaBH4

R

OMe (S)-15

1 Deprotection; 2 12 DHP, pTsOH

OMe

1 Deesterification; 2 Oxidation

1

RuOMe (R)-15

1 NaBH4

Ru

0

M

e 2 Esterificatior R

rac-15

10

12 DHP. pTsOH

T H e

Rd

R

O

M

( S ) - 11

e

Fig. 13. Enzymatic hydrolysis of the racemic acyl derivatives of the hydroxy compound. a Hydrolysis of the undesired enantiomer, b hydrolysis of the desired enantiomer.

4.2.5 Transesterification Reactions The lipase-catalyzed transesterification of the racemic hydroxyacid ester offers advantages compared to the ester hydrolysis (4.2.2), since the differences in the polarity of esters of higher alcohols (e.g., 3,5,5-trimethylhexanol) can be used for the extractive separation of the enantiomers. Four different strategies are possible leading to the desired product, the dihydropyran derivative of a P-hydroxyacid ester.

In the first route, the desired enantiomer of the racemic hydroxyacid ester ruc-(11)is transesterified leaving the undesired enantiomer as the methyl ester (R)-(11)(Fig. 14a). Only two additional steps are involved, the enzymatic resolution and the reoxidation of the (R)-hydroxyacid methyl ester (R)-(11). The second alternative is less attractive (Fig. 14b) since one additional step is necessary compared to the first route (1 additional step to the correct enantiomer, 2 additional steps to recycle the undesired enantiomer). In this


417

Oxidation

(R)-1 1

rac-1 1

10

THPO-

1 Transesterification; 2 12 DHP, pTsOH

(S)-1 6

R

1 Oxidation; 2 transesterification

THPQ

12 DHP, pTsOH

(S)-1 1

R

b)

( S ) - 13

1 Hydrolysis; 2 Oxidation;

1 NaBH4

THPO

pTsOH 10

rac-13

THPQ Rd

1 Transesterification; 2 12 DHP, pTsOH

o

M

c

(5')- 17

418


1 Transesterification; 2 Hydrolysis; 3 Oxidation;

THPO JJOl?

1 NaBH,

RM

O

M

2 12 DHP, pTsOH

,

10

(R)-17

Esterase

rac- 13

a

(S)-13

Fig. 14. Transesterification reactions. a Transesterification of the desired enantiomer, b transesterification of the undesired enantiomer, c transesterification of the desired DHP-enantiomer, d transesterification of the undesired DHP-enantiomer.

case, the undesired enantiomer is transesterified. The resulting ester alcohol has to be exchanged against methanol prior to the reoxidation step in order to recycle the undesired enantiomer. Two other alternative routes resolve the two enantiomers at a later step, where the dihydropyran protection group is already coupled. Again, either the desired or the undesired enantiomer can be transesterified. The corresponding synthetic routes are shown in Figs. 14c and 14d. As in the previous two cases, the number of additional steps is 2 and 3, respectively.

4.2.6 Acylation Reactions In this variation, the undesired enantiomer of the P-hydroxyacid ester is selectively acylated (Fig. 15).The recycling of the acylated enantiomer may be difficult since not only the acyl group, but also the methyl ester group, will be hydrolyzed. In case of rejection of the undesired enantiomer this route may be attractive. The stereoselectivity of the reaction may be high since the acylation is taking place next to the stereogenic center.

1 Selective hydrolysis; 2 Oxidation;

AcO uOMe

12 DHP, pTsOH

THPQ R3

0

M

(S)-1 1

e

(S)-13 Fig. 15. Selective acylation of the undesired enantiomer of the racemic P-hydroxyacid ester.

419


4.3 Conclusions Tab. 6 shows a summary of the above discussed alternative synthetic routes leading to a chiral p-hydroxyacid ester. The major criteria for the assessment of the different routes are the total number of additional steps and the position of the resolution of enantiomers (except route 4.2.1). From the chemical process development viewpoint, the direct enantioselective reduction is clearly the route of the highest priority, since no additional steps are involved. Most of the criteria mentioned in Sect. 4.1 are met.The desired stereoselectivity can be chosen by selecting the appropriate microorganism or isolated enzyme. A variety of microorganisms with different stereoselectivities catalyzing the reduction of p-ketoacid esters have been described in the literature. Alternatively, a number of different enzymes can be used for the enantioselective reduction of P-ketoacid esters. According to BISELLI et al. (1995), the further development of a process basically consists of three steps:

(1) Investigation of the reaction system 0 Selection of a suitable enzyme 0 Aspects of the properties of the enzyme: - Dependency of enzyme activity on substrate concentration, temperature, and pH, - dependency of enzyme stability on pH, temperature, and oxidizing agents, - dependency of enzyme selectivity on the reaction conditions, - reactor concepts and considerations on the use of organic solvents or cosolvents (see Sects. 3 and 5 ) . 0 Other properties of the reaction system: - Equilibrium constant of reversible reactions, - factors influencing side reactions (catalyzed and non -catalyzed), - effects of pH and temperature on the reaction and solubility of substrates and products. (2) Investigation of the reaction kinetics 0 Kinetics of the enzyme-catalyzed reaction(s)

Tab. 6. Assessment of Strategies for the Synthesis of a Chiral Hydroxy Acid Ester

+

~

Total Number of Number of Steps to Additional Steps the Racemic Educt

Priority

0+0

0

1

1

4.2.2a 4.2.2b

1+ 2 2+1

3 3

2 2

2a 2b

4.2.3a 4.2.3b

1+1 2+2

2 4

3 3

2a 3

4.2.4a 4.2.4b

3+ 1 2+2

4 4

2 2

3 3

4.2.5a 4.2.5b 4.2.5~ 4.2.5d

1+1 1+ 2 1+1 1+ 2

4.2.6

1+ 2

Variants Additional Steps to Desired Enantiomer Additional Steps for Recycling of Undesired Enantiomer ~

~~

4.2.1

~

~

2a 2b 2a 2b 3

2

2a


420

Kinetics of the non-catalyzed reaction@) 0 Characterization of mass transfer phenomena which are often encountered with liquid-liquid two-phase systems or immobilized enzyme systems. (3) Reactor design and reaction modeling 0 Reactor kinetics (combination of kinetic models of enzymatic and noncatalyzed reactions and their mass balances) 0 Reactor performance as a function of substrate concentration,enzyme and coenzyme concentration,and residence time, 0 Optimization of the reactor performance based on the following criteria: conversion rate, stereoselectivity, space-time yield, enzyme and coenzyme consumption. 0

5 Application of Alcohol Dehydrogenases in Organic Solvents or Cyclodextrins 5.1 Introduction Nearly all reactions in preparative organic chemistry are performed in organic solvents since most of the organic compounds are poorly soluble or even insoluble in water. Furthermore, the high boiling point and high heat of vaporization make the removal of water expensive and tedious. Additionally, side reactions such as hydrolysis, racemization,decomposition, or polymerization are often encountered when using water as solvent. Therefore, synthesis of most organic compounds is performed in organic solvents. The synthetic chemists demand, first of all, enzymes tolerating organic solvents and, secondly novel reactor concepts circumventing solubility problems of substrates and products. Enzymes are most active in their natural environment. However, this does not mean necessarily that enzymes only function in water.

Indeed, it has been found for a range of enzymes that only a limited amount of water has to be present to maintain full catalytic activity et al., (ZAKSand KLIBANOV,1988;GRUNWALD 1986; ITOHet al., 1992; ANTONINI et al., 1981). LAANEand coworkers (1987) investigated the effect of the partition coefficient (P)of a solvent containing a poorly water-soluble substrate on the activity and stability of a variety of lipases.The log P concept was confirmed by KISEand HAYASHIDA (1990) and ITOHet al. (1992) for horse liver alcohol dehydrogenase and other NAD(P)(H)-dependent oxidoreductases In Tab. 7 the log P values of a range of commonly used organic solvents are listed. Even though log P is a rather crude parameter lacking a detailed mechanistic basis, the log P value gives a helpful guideline for the use of organic solvents in enzymatic catalysis. Solvents with a log P below 2 were not suitable for enzymatic systems, because they strongly distort the essential water-enyme interaction, thereby inactivating or denaturing the enzyme. The toxicity of solvents with log P values between 2 and 4 is unpredictable. Prolonged presence of the enzyme could be expected in solvents with a log P above 4, such as heptane, octane, and higher alkanes as well as different phthalates subsituted by long alkyl chains. As outlined in Sect. 4, alcohol dehydrogenases show several advantages compared to lipases for the production of enantiomerically pure alcohols: (1) Direct enantioselective reduction with no additional steps; (2) 100% conversion as opposed to 50% conversion for the racemic resolution catalyzed by lipases. However, these advantages are only applicable without restriction if the compound which is to be reduced is water soluble. If the carbonyl compound is only poorly soluble in =mowater, low total turnover numbers (l" le productfmole cofactor) are obtained. In order to overcome economical limitations, a TTN > 100 is desirable.Additionally,large volumes of aqueous phase are necessary to solub i k e the substrate, complicating the work-up of the product solution. Redox enzymes have been used rarely in organic solvents compared to lipases. CARREA and coworkers (1979) reported that immobilized Phydroxysteroid dehydrogenaseshowed an inactivation rate of only 0.16% d-' during

421

5 Application of Alcohol Dehydrogenases in Organic Solvents or Cyclodextrins Tab. 7. Log PValues of Commonly Used Organic Solvents

Solvent

Log P Value

Solvent

Log P Value

Dimethylsulfoxide N,N-Dimethylformamide Ethanol Acetone Methyl acetate Tetrahydrofuran Ethyl acetate Pyridine Propyl acetate Butyl acetate Chloroform Pentyl acetate

- 1.3

Toluene Octanol Dibutylether Carbon tetrachloride Cyclohexane Diphenylether Octane Nonane Dibutyl phthalate Dodecane Didecyl phthalate Dilauryl phthalate

2.50 2.90 2.90 3.00 3.50 4.30 4.50 5.10 5.40 6.60 11.7 13.7

continuous reuse for two months in a water-ethylacetate solvent. LARSSON et al. (1987) described that horse liver alcohol dehydrogenase (HLADH) is stable in a detergentlisooctane reverse micelle system for at least two weeks after an initial loss of activity. GRUNWALD et al. (1986) found that deposition of HLADH and coenzyme onto the surface of glass beads is a generally applicable concept for using coenzyme depending oxidoreductases in water-immiscible organic solvents. They reported also very high cofactor turnover numbers for these substrate-coupled systems of lo5 to lo6. JONESand SCHWARTZ (1982a, b) summarized effects of organic solvents on HLADH. KISE and HAYASHIDA (1990) found that the log P concept is also applicable to nicotinamide coenzyme-dependent oxidoreductases. Additionally, a number of recent publications deal with different reactor technologies for the continuous application of dehydrogenases in organic solvents, two-phase systems, or cyclodextrin-based systems (KISE and HAYASHIDA, 1990; ZELINSKI, 1995; KRUSE et al., 1996; LIESEet al., 1996; KRAGLet al., 1996a;LIESEet al., in press). This section deals with the application of coenzyme-dependent dehydrogenases in organic solvents covering novel reactor concepts for the continuous synthesis of poorly soluble chiral molecules.

- 1.0 -0.24

- 0.23

0.16 0.49 0.68 0.71 1.20 1.70 2.00 2.20

5.2 Solubilities and Stabilities of Substrate(s) and Product(s) Coenzymes are highly polar compounds and, therefore, completely insoluble in a lipophilic medium. Using the enzyme-coupled approach for coenzyme recycling, the nicotinamide cofactor has to dissociate from one enzyme, diffuse to the other enzyme, and associate to its active site, and vice versa.The diffusion of the coenzyme from one enzyme to the other may be restricted by using apolar organic solvents. In contrast, using the substrate-coupled regeneration, the coenzyme stays in the vicinity of the active site of only one enzyme, which facilites the use of organic solvents. However,in both cases the enzyme has to be surrounded by a small layer of water molecules allowing the association and dissociation of the coenzyme. Hence, the poorly water-soluble substrate and product have to pass this layer of water molecules. Therefore, considering the use of organic solvents, the solubility and the stability of the substrate(s) and product(s) in water should be known. As already said in the introduction, side reactions such as hydrolysis, racemization, decomposition, or polymerization are often encountered if water is used as solvent.

422


For example, 4-0x0-hex-5-in carboxylic acid ester (19) reacts in water or in the presence of sodium formate forming a yellow colored aromatic compound (20) (Fig. 16). In the presence of ammonium formate, which may also be used as hydride source in the formate dehydrogenase reaction for coenzyme recycling, other side reactions occur, of which a pyridine compound (21)is predominantly formed. Similar reactions leading to (hetero-)aromatic compounds have been already described in the literature (BALASUBRAMANIAN et al., 1980; BOWDENand JONES,1946). However, the described reactions of 4-0x0-hex-5-in carboxylic acid ester (19) are not observed in organic solvents. This demonstrates that the investigation of the stability of the substrate and product in water is necessary for the successful application of alcohol dehydrogenases in preparative organic synthesis.

5.3 Modern Concepts for the Application of Alcohol Dehydrogenases Using Organic Solvents or Cyclodextrins The repeated use of expensive biocatalysts and their separation from products and reactants are major goals in continuous enzymatic synthesis. For isolated enzymes, either immobilization on an insoluble support or retardation using ultrafiltration membranes are used to accomplish these goals and applied up to large-scale industrial synthesis. Recently, BOMMARIUS (1993) gave a comprehensive overview regarding reactor systems used for enzymatic transformations. Using enzymes immobilized on a support, the following disadvantages have to be considered: 0

0 0

Additional costs for the carrier and additional chemicals have to be balanced against the increase of stability. Co-immobilization of enzyme(s) and coenzyme may be troublesome (see Sect. 3). Loss of enzyme activity often occurs during immobilization.

Hy + - f o c H 3

k water

H3c COOCH,

COOCH,

Fig. 16. Decomposition of 4-0x0-hex-5-in carboxylic acid ester in water and in the presence of ammonium formate.

5 Application of Alcohol Dehydrogenases in Organic Solvents or Cyclodextrins 0

0

0

Mass transfer especially between the organic phase and the water containing porous particles of the support is limited (LORTIEet al., 1989). Low volumetric enzyme activities often limit the application of immobilized enzymes for the conversion of poor substrates. Process stability is often limited due to sterility problems.

For these reasons, the following sections focus on membrane-based reactor concepts.

423

Additionally, dehydrogenases are often deactivated by lipophilic compounds forming a water-organic interface. Thus, for dehydrogenases, aqueous/organic two-phase systems are not recommended. Recently, a range of novel enzyme membrane reactor concepts have been proposed which are specially designed for the application of coenzyme-dependent oxidoreductases using organic solvents (KRUSE,1995; KRUSEet al., 1996; LIESEet al., 1996 and LIESEet al., in press):

In continuous enzymatic processes, two major strategies are employed to achieve high total turnover numbers ('ITN). First, the TTN can be increased by using immobilized coenzymes (e.g., PEG-NAD(H), see Sect. 3.4.2). Second, the substrate can be used at very high concentrations, whereas the coenzyme concentration is kept at the lowest possible level in order to saturate the enzyme. However, this approach fails in the case of poorly water-soluble substrates and products.

Enzyme membrane reactor with continuous product extraction This concept is based on the combination of three main cycles (Fig. 17).The enzyme membrane reactor (EMR) represents a continuously operated loop reactor supplied with a hydrophilic hollow fiber ultrafiltration membrane.The enzyme(s) are retained in the reactor, while substrate, coenzyme, and products can freely pass the membrane. The product stream enters the continuous, countercurrent extraction module equipped with a hydrophobic, microporous membrane. In this module, the product is extracted into hexane and, subsequently, distilled yielding the product fraction and the hexane fraction, which is again recycled into the extraction module.

Fig. 17. Schematic presentation of the enzyme membrane reactor with continuous product extraction.

HCOOH

5.3.1 Oxidoreductions Using Organic Solvents

S

(NAD)

424

9 Dehydrogenases - Characteristics,Design of Reaction Conditionr,and Applications

7 3 s type of reactor has been successfully employed for the continuous reduction of several aromatic substituted ketones to the corresponding chiral (S)-alcohols (KRUSE et al., 1996) using the NAD-dependent alcohol dehydrogenase from Rhodococcos erythropolis (HUMMELand GOTIWALD,1991). For coenzyme regeneration the formate dehydrogenase from Cundidu boidinii was used (see Sect. 3). Conversion rates greater than 65%, spacetime yields between 60 and 105g L-' d-',purities between %% and 99%, total turnover numbers between 158 and 1350, and enantiomeric excesses greater than 99% have been achieved. Product quantities of 20 to 90 g were obtained. However, the substrates employed must have a residual minimum solubility in water of about 10 mM or more in order to use this type of reactor. Additionally, the affinity of the alcohol dehydrogenase for the substrate should be very high (low, KM values) in order to achieve high conversion rates and to optimally use the enzyme activity.

sists of a loop containing a circulation pump and an ultrafiltrationmodule.The product outflow is recirculated to the stirred emulsion vessel, where, due to the partition coefficients,the product is extracted into the organic phase. The aqueous phase is recharged with substrate. Overall, the aqueous phase is repeatedly charged due to the partition coefficients with substrate in range of the maximal solubility, whereby the product is continuously extracted i ito the organic phase. Three main advantages of this reactor design are evident: (1) The product is concentrated in the organic phase ( =substrate). No additional extraction is necessary for product recovery. (2) The oxidoreductase is not in direct contact with the organic phase, thereby preventing the enzyme from inactivation at the interface. (3) The total turnover number will be increased significantly.

Emulsion enzyme membrane reactor The concept of the emulsion enzyme reactor As a model reaction, LIESEand coworkers (Fig. 18) is based on the combination of a (1997) performed the enantioselective reducstirred emulsion vessel and a second enzyme tion of 2-octanone to (S)-2-octanol catalyzed membrane reactor (LIESEet al., in press). The by the carbonyl reductase from Cundida parastirred vessel is used to charge the feed solu- psilosis (CPCR) (PETERS et al., 1992, 1993b; tion with substrate and contains a hydrophilic ZELINSKI et al., 1993) in the emulsion enzyme membrane separating the aqueous phase from reactor. The haif-life of the enzyme under rethe organic phase. The substrate-charged solu- action conditions was determined to be 31 d, tion then enters the enzyme membrane reac- corresponding to an inactivation rate of 1.7% tor containing an enzyme system performing d-l. A total turnover number of 124 and a the desired reduction. This second reactor con- space-time yield of 11 g L-' d-' at a conver-

Fig. 18. Schematic presentation of the emulsion enzyme membrane reactor. 1 Membrane reactor (stirred emulsion vessel equipped with hydrophilic membrane), 2 enzyme membrane reactor (equipped with an ultrafiltration membrane), 3 PUP, 4 air trap.

425

5 Application of Alcohol Dehydrogenases in Organic Solvents or Cyclodextrins

sion of 91% were reached. Using a conventional enzyme membrane reactor, a total turnover number of only 13.6 was reached during continuous production of (S)-Zoctanol over a period of 4 months. The applied mole of substrate is no longer determined by the maximal solubility of the substrate in water, but instead, by the volume of the organic phase. The appropriate coenzyme concentration is determined by the K , value of the CPCR. Both factors result in an increased total turnover number which may be increased further by one order of magnitude by changing the organic phase from 1% (v/v) to 10% (v/v).

5.3.2 Oxidoreductions in the Presence of Cyclodextrins Cyclodextrins are cyclic polysaccharides containing ~ ( 1 - 4 glucose ) units. The most important cyclodextrins consist of 6,7, or 8 glucopyranoside units and are referred to as a-, p-, y-, and Gcyclodextrins (22-25). The structure of all cyclodextrins consists of a torus with a hydrophilic outside and a hydrophobic cavern (Fig. 19). The diameter of this cavern depends on the number of glycopyranoside units and ranges from 490,620, and 790 pm for a-,p-,

and y-cyclodextrins (22-24),

respectively.

Thus, hydrophobic guest molecules like, e.g.,

benzol (a-cyclodextrin (22)), naphthalene (pcyclodextrins (23)) or steroids (y-cyclodextrin (24)) can enter the cavern forming a water-soluble host-guest complex (SAENGER,1980; SZEJTLI, 1982; WENZ,1994). The concept of using cyclodextrins as co-solvents for the application of oxidoreductases makes use of this properly of cyclodextrins. The host-guest complexes are homogeneously dissolved in water. Therefore, the above mentioned instability of most oxidoreductases at the interface of two immiscible solvents and, additionally, mass transfer problems are circumvented by the use of cyclodextrins. Cyclodextrins as co-solvents for poorly water-soluble substances have been utilized mainly for microbial transformations of steroids (HESSELINK et al., 1989;JADOUN and BAR, 1993a, b; SCHLOSSER et al., 1993; SINGER et al., 1991).Only a few publications deal with enzymatic transformations in the presence of cyclodextrins (OTEROet al., 1991; PEKICand LEPOJEVIC, 1991; JADOUN and BAR,1993a; WOERDENBAG et al., 1990). As first proposed by PETERS (1993), the application of oxidoreductases using cyclodextrins as co-solvents for poorly water-soluble compounds is possible and offers considerable advantages (ZELINSKI, 1995).

-0

“fi;l ~11111

OH no]

HO

2 2 n=6; a 23a n=7:

24 n=8; y 25 n=9; S

Fig. 19. of cyclodextrins.

Rd

23 a R = H; P-Cyclodextrin b R = Me; heptakis(2,6-di0-methyl)-P-cyclodex trin

b

426


As a model reaction, the chiral reduction of the poorly water-soluble 2-acetylnaphthalene (27) to the corresponding (S)-l-(2-naphthyl)ethanol (28) catalyzed by the novel carbonyl reductases from Rhodococcus erythropolis (RECR) and Cundida purapsilosis (CPCR; PETERS et al., 1992, 1993b; ZELINSKI et al., 1993) was used for the application of coenzyme-dependent dehydrogenases in cyclodextrin solution. The results of this investigation (ZELINSKI,1995) are discussed in the following section in more detail, since the cyclodextrin concept may be broadly applicable. Both, the substrate and the product form inclusion complexes with the cyclodextrin molecules (23)(Fig. 20). Therefore, the equilibrium of the ketone reduction depends on the Kdirs values of the cyclodextrin equilibria with substrate and product. In order to ensure a constant substrate concentration, the velocity of the host-guest equilibrium has to be faster than the enzymatic reaction.The affinity of the enzyme for the substrate should be higher than the complex stability constant Kdiss.The Kdissvalue of the complex of heptakis-(2,6,di0-methyl)-P-cyclodextrin (DIMEB) and 2-ac-

26

NmH

etylnaphthalene (26)is about 1 mM, whereas the K,,, value of the CPCR is only 3 pM. Hence, the enzyme has access to the substrate and the active centers are completely saturated at 1 mM concentration (Kdisr). It was shown for 2-acetylnaphthalene (27) that the addition of the P-cyclodextrin derivative DIMEB (23b)enhances the solubility of this poorly water-soluble compound by a factor of 150 (0.35 mM solubility in water versus 52 mM solubility in DIMEB solution). Both enzymes, CPCR and RECR, reduced 2-acetylnaphthalene (27)to the corresponding (S)-1(2-naphthy1)-ethanol (28) at a high enantiomeric excess of >99%, which can be recovered by extraction with n-hexane without losses. The carbonyl reductases, as well as the formate dehydrogenase used for coenzyme regeneration, were stable during continuous synthesis in an enzyme membrane reactor.The deactivation rate of 3% d-' is of the same order of magnitude as in non-cyclodextrin media. The conversion rates during continuous synthesis reached 79% and 82% for the enzyme-coupled and substrate-coupled coenzyme regeneration systems, respectively. The

27

\I

/ 2 9

23b

/

28

23b

Fig. 20. The enzymatic synthesis of (S)-l-(2-naphthyl)-ethanolusing P-cyclodextrin as co-solvent.Both the substrate and the product form host-guest complexes with cyclodextrin.

6 Kinetics and Stereochemistryof Alcohol Dehydrogenases

space-time yield was determined to be 118 g L-’d-’.The reactor concept established by KRUSE(1995) and LIESE(1994) may be suitable for the continuous recycling of the cyclodextrin co-solvent.

5.4 Conclusions The stirred tank reactor has been used for a long time in industry and is still the reactor for most chemical processes. The special characteristics of enzymes, however, sometimes demand novel types of bioreactors.As will be addressed in the next section, the choice of a suitable enzyme reactor largely depends upon the type of inhibition. For instance, a plug flow reactor such as a packed bed reactor may be advantageous for enzymes inhibited by their reaction product(s).The liquid impelled loop reet al., 1987) is specifically deactor (TRAMPER signed for biocatalysis in nonconventional media. Up to now, only a limited number of reactor et al., 1996a, b) concepts are known (KRAGL which specifically address the problems of using alcohol dehydrogenases and poorly watersoluble substrates in organic solvents mentioned in Sect. 5.1. The enormous potential of dehydrogenase-catalyzedredox reactions will become more accessible, if the above described reactor concepts are further developed and applied on the pilot scale.

6 Kinetics and Stereochemistry of Alcohol Dehydrogenases 6.1 Introduction The reaction mechanism, the stereochemistry, and the kinetics of reactions catalyzed by alcohol dehydrogenase are of special importance with respect to the following aspects: 0

The stereochemistry can be used for the classification of alcohol dehydrogenases

427

and for the prediction of the enantioselectivity of a given reduction. This gives a helpful guideline to the user of commercial and well-characterized noncommercial oxidoreductases. Kinetic constants are very important for understanding the kinetic limitations of a given enzymatic reaction and, therefore, for circumventing them. Furthermore, the kinetic characteristics of an enzyme are the basis for modeling of enzyme reactors and important for the scaleup to the technical scale. Special parameters, like the substrate and product inhibition constants, have additional relevance in the selection of the appropriate type of reactor. A given alcohol dehydrogenase is characterized by its enantioselectivity.The kinetic resolution of racemates catalyzed by alcohol dehydrogenasescan be used to get access to the opposite chirality. Normally, dehydrogenases generate only one stereogenic center at a time. In the case of a-acidic carbonyl compounds,two stereogenic centers can be introduced into a molecule at a time by making use of the keto-enol tautomeric equilibrium and the differences in the kinetics of the two a-substituted enantiomers.

6.2 Stereochemistry and Stereoselectivity of Alcohol D ehy drogenases During the course of the reduction of a carbony1 compound, the dehydrogenase delivers the hydride preferentially either from the si or the re side of the ketone to give, for simple systems, the (R)- or (S)-hydroxy compounds, respectively. For most cases, the stereochemical outcome of the reaction can be predicted from a simple model which is referred to as “Prelog’s rule” (PRELOG,1964) (Fig. 21). This model is based on the stereochemistry of microbial reductions using Curvuluriu fulcutu cells. It states that the dehydrogenase delivers the hydride ion from the re face of a prochiral ketone predominantly resulting in the formation of (S)-alcohols.

428


b/

Dehydrogenase NAD(P)

Fig. 21. Prelog’s rule for the asymmetric reduction of carbonyl compounds.In most cases this is equivalent to re-face attack. S, small residue; L, large residue.

The majority of the commercially available dehydrogenases and microorganisms used for ketone reductions follow Prelog’s rule, but “anti-Prelog” dehydrogenases are also described in the literature (Tab. 8). Additionally, both NADH and NADPH contain two diastereotopic hydrogens (pro-R and p r o 4 hydrogen) that can be transferred as a hydride to an oxidized substrate. In principle, any NAD(P)H-dependent oxidoreduction should fall into one of these four types of stereospecificity. For the classification of other dehydrogenases the reader is referred to the literature (RETEY and ROBINSON, 1982). In order to determine the specificity of an enzyme with respect to the pro-R or p r o 4 diastereotopic hydrogens of NAD(P)H, ‘H-NMR is considered to be most convenient. The pro-R hydrogen has a chemical shift of 2.77 ppm and the p r o 4 hydrogen has a shift of 2.67 ppm. Therefore, one can examine the chemical shift of the [4-*H]-NAD(P)Hproduced by enzyme-catalyzed oxidation of a deuterated substrate in the presence of NAD(P)+. The reverse reaction starting from [4R-’H]- or [4S’HI-NAD(P)H and a carbonyl compound can also be used to determine the stereospecificity concerning the coenzyme.

In general, the secondary alcohol dehydrogenase from Thermoanaerobium brockii (TBADH) obeys Prelog’s rule with “normalsized” ketones leading to (S)-alcohols, but the stereospecificity was found to be reversed with small substrates (KEINANet al., 1987; KELLY and LEWIS,1991). This behavior was also observed with other alcohol dehydrogenases (ZELINSKI, 1995).The selective recognition of small molecules having no ionic groups is generally a difficult task for an enzyme, since only small energetic differences exist between the two possible diastereomeric transition states (WONG,1989). However, the recently described carbonyl reductase from Candida parapsilosis (CPCR) is able to differentiate even between the two enantiomers of 2-butanol (PETERS,1993).

6.3 Reaction Mechanisms of Alcohol Dehydrogenases 6.3.1 Reaction Mechanisms The determination of the reaction mechanism of an enzymatic reaction consists of two parts: first, the kinetic mechanism and second, the catalytic mechanism. The kinetic mechanism describes the order of binding of the substrate(s) and the order of the dissociation of the product(s). On the other hand, the catalytic mechanism describes in detail the chemical reactions at the active site and identifies the rate limiting step of the enzyme-catalyzed reaction (DIXONand WEBB,1973). Alcohol dehydrogenase-catalyzed reactions are two-substrate reactions. These reactions can be classified according to LASCH(1987) into two categories: (1) Sequential mechanism - all substrates bind to the enzyme before the first product dissociates. This type of mechanism can be further divided into the random mechanism (binding of substrates occurs randomly) and the ordered mechanism (binding of substrates occurs in ordered fashion).


429

Tab. 8. Stereochemistry of the Hydride Transfer: Classificationof Alcohol Dehydrogenases

Type

Hydrogen Transfer

Dehydrogenase from

Enantioselectivity

Coenzyme

Commercial Availability

El

pro-(R)/si-face

Pseudomonas sp. Lactobacillus kejir

(R)

NAD(H) NADP(H)

-

EZ

pro-(S)/si-face

Mucor javanicus

(4

NADP(H)

-

E3

pro-(R)/re-face

Yeast Horse liver Thermoanaerobium brockii Candida parapsilosis Rhodococcus erythropolis Curvularia falcata

(S1

NAD(H) NAD(H) NADP(H) NAD(H) NAD(H) NADP(H)

unknown

(S1

~~

E'l

pro-(S)/re-face

( 2 ) Ping-pong mechanism - one or more products dissociate from the enzyme before all substrates are bound. Reaction mechanisms can be distinguished from each other by studying the substrate and product inhibition patterns (CLELAND, 1986). The enzyme kinetics of two-substrate reactions, activation and inhibition effects, and reaction mechanisms have been reviewed by CARRand BOWERS (1980). All of the alcohol dehydrogenases characterized so far fall into the first category showing a sequential mechanism with ordered binding of substrates and dissociation of products. In this case, NAD(P) associates first with the active site before the hydroxy compound is bound forming a ternary enzyme-coenzymesubstrate complex. After the hydride transfer occurred, first the carbonyl compound and then the reduced coenzyme (NAD(P)H) dissociates from the active site of the enzyme. This reaction is reversible for most dehydrogenases, although in some cases (formate dehydrogenase from Cundida boidinii) the equilibrium is far from one side (SCHUTTE et al., 1976).

-

+ + + -

-

A special variation of the ordered sequential mechanism is the so called TheorellChance mechanism. In this case, the coenzyme (NAD+) also binds first to the active site of the enzyme, but association of the hydroxy compound does not lead to the formation of a ternary complex. The dissociation of the carbony1 compound occurs immediately after the hydroxy compound is bound. Finally, the reduced coenzyme leaves the active center. This type of ordered sequential mechanism was also found for a number of alcohol dehydrogenases (e.g., carbonyl reductase from Rhodococcus erythropolis; ZELINSKI, 1995).

+

6.3.2 Substrate, Product, and Cross Inhibition The rate of enzyme-catalyzed reactions is often influenced by either substrate, product, or substances which are not involved in the catalyzed reaction (KULAet al., 1980). When the rate is decreased, there is said to be inhibition and the substance causing the effect is

430

9 Dehydrogenases- Characteristics, Design of Reaction Conditions,and Applications

called an inhibitor. Conversely, an effector (1) Ki/Kmratio > 1:The reaction may prowhich causes an increase in the rate is known ceed to high conversions.The product as an activator. concentration should be maintained at a low level to overcome the inhibition. In dealing with inhibition, there are a number of important distinctions which must be (2) KJK, ratio < 1:The reaction can not recognized. Reversible inhibition can be rapidproceed effectively. ly overcome by removal of the inhibitor. On the other hand, irreversible inhibition causes Taking the carbonyl reductase from Cundian irreversible change in the enzyme which du purupsilosis (CPCR) as an example (PEeliminates the catalytic activity of the enzyme. TERS et al., 1993d), the ratio of the product inFrequently this involves covalent bonding of hibition constant of the reduced coenzyme the inhibitor to the enzyme. For example, the K,(NADH) and the K , value of the oxidized formate dehydrogenase from Cundidu boidinii coenzyme (K,(NAD+)) is only 0.2. On the oth(FDH) was irreversibly inhibited by 4-chloro- er hand, the enzyme is only slightly inhibited 3-oxobutanoate ethyl ester (PETERS,1993). by 2-butanone (Ki/Kmratio =26). Therefore, in This inhibition was most probably due to an al- the case of the CPCR the oxidation of alcohols kylation of the sulfhydryl group in the active is not inhibited by the carbonyl product, but center of the FDH. severely inhibited by the reduced coenzyme. In reversible inhibition, three situations can Hence, for kinetic reasons this enzyme does be distinguished: not oxidize alcohols but reduces carbonyl compounds. At this point it is worthwhile to - Non-competitive inhibition: The degree of stress that the strong product inhibition caused inhibition is not a function of the concen- by NADH can be easily circumvented by tration of the substrate. coupling to an appropriate coenzyme regener- Competitive inhibition:The degree of ination method (either substrate- or enzymehibition is a function of the concentration coupled approaches are possible; PETERS, of the substrate, where the inhibition is 1993;ZELINSKI, 1995).About 80% conversion of substrate is achieved by using either regenreduced as the concentration of the suberation method in a continuous stirred tank restrate increases. This type of inhibition is actor (compare Fig. 8; ZELINSKI, 1995). Meththe result of inhibitor and substrate ods to overcome product inhibition caused by molecules competing for the same site of the carbonyl or hydroxy compounds are disthe enzyme. - Uncompetitive inhibition:The degree of cussed in Sect. 6.3.3. Coupling of enzymes, e.g., a reductase and inhibition is a function of the concentration of the substrate, where the inhibition the formate dehydrogenase (FDH) for coenzyme regeneration, may cause additional inhiis increased at increased concentrations bition phenomena termed cross inhibition. of the substrate. Substrate(s) or product(s) of the first reaction Many enzyme-catalyzed NAD(P)(H)-de- may inhibit the second enzyme and vice versu. pendent redox reactions suffer from inhibition For example, the formate dehydrogenase from caused by the product. Especially in the oxida- Cundidu boidinii (FDH) was employed for cotion of alcohols, the carbonyl product tends to enzyme regeneration in a coupled enzyme bind to the enzyme more strongly than do the system with the carbonyl reductase from Cunalcohols. But the product inhibition caused by didu parupsilosis (CPCR). In this system, the the coenzyme is also of considerable impor- CPCR reduced lactaldehyde dimethyl acetal tance. A detailed analysis of product inhibition to the corresponding (S)-hydroxy compound related to the reaction rate revealed that the (PETERS,1993;PETERSet al., 1993a). Lactalderatio of product inhibition constant (KJ to the hyde dimethyl acetal reversibly inhibited the Michaelis-Menten constant of the substrate FDH. The inhibition could be circumvented by (K,) will determine the efficacy of the oxida- decreasing the concentration of lactaldehyde tion process (LEEand WHITESIDES, 1986).The dimethyl acetal below a critical value of 0.2 M thereby restoring the activity of the FDH. For following cases can be distinguished:

6 Kinetics and Stereochemistry of Alcohol Dehydrogenases

other coupled enzyme systems (leucine dehydrogenaseFDH and mandelate dehydrogenaselFDH) a range of cross inhibitions were observed (VASIC-RACKI et al., 1989; KRAGLet al., 1996a,b).

6.3.3 Impact of Reaction Mechanism and Type of Inhibition on the Selection and Modeling of Enzyme Reactors

431

well as the kinetic parameters and limitations plays an essential role in the design of an enzymatic process. In order to circumvent these limitations, the appropriate designs with respect to the reactor and the process itself have to be chosen. Additionally, several other considerations have to be made as already outlined in Sect. 4.3.

Choice of the appropriate reactor In dealing with different reactor concepts, there are a number of important distinctions which must be recognized. Either a batch or a continuous reactor system may be employed. A batch enzyme reactor (stirred tank reacThe development of an enzymatic process involves four aspects: (1) reaction conditions tor, STR),which is frequently used in chemical (e.g., pH, temperature, concentrations), (2) processes, is often limited in the space-time process design (e.g., reactors, product recov- yield which can be achieved and, in general, is ery), (3) thermodynamics (e.g., equilibrium not preferred for alcohol dehydrogenase-cataconstant, and (4) kinetics (e.g., K,, Ki, and lyzed reactions. The substrate concentration at V,,, values). The thermodynamic parameters the beginning of the reaction should be as high describe the equilibrium of a given reaction, as possible in order to achieve the highest whereas the kinetic constants give the velocity space-time yields, probably leading to subfor reaching conversion at equilibrium. The strate inhibition. The reaction products accuHuldune equation links the thermodynamic mulate more and more as the reaction proceeds leading to limitations in the conversion and kinetic parameters with each other: due to product inhibitions. Dealing with poorly soluble substrates (see Sect. 5 ) these can only be used at very dilute concentrations leading to unacceptable low space-time yields and large volumes causing additional problems in with: V,,,' Maximum velocity of the reaction downstream processing of the reaction mixKmP Miachelis constant of the product V,,,' Maximum velocity of the reverse tures. A range of continuously operating enzyme reaction reactor concepts have been published and reK,S Substrate inhibition constant KmS Michaelis constant of the substrate viewed recently (BISELLIet al., 1995;KRAGLet al., 1996b; BOMMARIUS, 1993). Two different Today, modeling of enzyme reactors espe- types can be distinguished: The plug flow reaccially in the case of alcohol dehydrogenases or tor (PFR) and the continuous stirred tank rereductases is well known and straightforward actor (CSTR). In Tab. 9 several characteristics (IKEMI and ISHIMATSU, 1990; IKEMl et al., 1990; of these reactors are compared to the batch KISE and HAYASHIDA, 1990; KRAGL, 1996; stirred tank reactor, which is frequently used KRAGLet al., 1996a; KULAand WANDREY, in chemical processes. The PFR exhibits de1987; LEUCHTENBERGER et al., 1984b; PETERS creasing substrate and increasing product conand KULA, 1991; ROTHIGet al., 1990a, b; centrations with reactor length. In contrast, the SCHMIDT et al., 1987b; VASIC-RACKI et al., CSTR exhibits at all positions the same low 1989; WANDREYand WICHMANN, 1985; WICH- substrate and high product concentrations. ReMANN et al., 1981; ZELINSKI, 1995). Modeling garding the time course of the conversion of enzymatic reactions allows the precise pre- under steady state conditions, the substrate diction of the behavior of an enzyme reactor concentration is low and the product concenand is, therefore, necessary for scaling up a tration is high for both types of continuous reprocess. Knowledge of the thermodynamic as actors.

432


Tab. 9. Comparison of the Characteristics of Reactors for Continuous Biocatalysis versus the Batch Stirred

Tank Reactor

Characteristics 1. Reaction medium Single liquid phase

Two or more phases

2. Kinetic constraints Substrate inhibition Product inhibition 3. Catalyst constraints Maximum particle concentration

Mechanical damage Low stability

Batch STR

PFR

CSTR

possible possible

possible possible

possible possible

poor satisfactory

poor satisfactory

good

low

high low unlikely

low

possible

possible

Considering the kinetic limitations of a given reaction, this difference between PFR and CSTR is important for the choice of the appropriate type of reactor. In those cases, where the product inhibition considerably limits the reaction rate, the PFR should be employed, since this reactor shows an increasing product concentration with reactor length. For example, the reductive amination of terf-leucine catalyzed by leucine dehydrogenase is inhibited by tert-leucine.Therefore, a PFR was installed for the continuous production process (KRAGLet al., 1996b).In cases, where the substrate inhibition plays a major role, the CSTR is advantageous due to the low substrate concentration at all positions in the reactor. Immobilization of enzymes on an insoluble support or a membrane allows continuous operation as a PFR. Immobilization of enzymes can be regarded as an established technology (ZABORSKY, 1973; HASSELBERGER, 1978;WISEMAN,1985; CARR and BOWERS,1980; MooYOUNG,1988). Disadvantages are limitations caused by mass transfer (fluid to carrier surface) and diffusion (carrier surface to catalyst) which are due to the heterogeneous reaction, additional costs for the immobilization, and development of individual immobilization procedures for each enzyme, loss of enzyme activity during immobilization, and the problem to supply fresh enzyme without interruption of the process. On the other hand, the enzyme membrane reactor (KULAand WANDREY, 1987; KULAet al., 1980; WANDREYand WICHMANN, 1985; WANDREY et al., 1984) offers several advanta-

poor

possible unlikely

ges. The enzyme is retained behind an ultrafiltration membrane and, thereby, “immobilized”. The system works as a continuously stirred tank reactor. A plug flow behavior is only possible if several reactors form a cascade. There are no additional costs for immobilization and the enzyme can be used at its full catalytic activity. The inactivated enzyme(s) can be substituted easily. Since the reaction occurs homogeneously, no mass transfer limitations reduce the reaction yield. The product stream leaving the reactor is proteinand pyrogen-free. Sterility is often difficult to achieve in an immobilized enzyme reactor. In contrast, the enzyme membrane reactor can be sterilized by chemical methods (e.g., ethanol, peracetic acid, formaldehyde) or steam. However, the enzyme membrane reactor suffers also from some disadvantages. Compared to an immobilized enzyme reactor, the investment costs are higher and the stability of the enzymes is normally lower. Enzyme membrane reactors are commercially available ranging from 10 mL bench top versions (Bioengineering, Switzerland), 50 mL reactors for process development to large scale hollow fiber or stacked flat membrane moduls for industrial productions (e.g., Amicon, Filtron, Millipore, Pall, Romicon, Sartorius, Sepracor, and others). Modeling of enzyme membrane reactors The performance of an enzyme membrane reactor may be calculated by means of measured kinetics and simultaneous calculation of mass balances of each reactant (BISELLIet al.,

6 Kinetics and Stereochemistry of Alcohol Dehydrogenases

433

1995). By selecting suitable conditions for the tion based on the reaction mechanism (seproduction process, a large number of experi- quential bi-bi- or Theorell-Chance mechaments may be avoided thus saving both time nisms, see Sect. 6.3.1) and includes the amount and costs. This approach is especially useful in of enzyme taking part in the reaction. the case of reactions with two or more enThe Michaelis-Menten term only describes zymes and parallel or consecutive reactions one-substrate reactions. Since dehydrogenases coupled via common reactants, where the catalyze two-substrate reactions (1. alcohol/ interdependence of the reactants otherwise ketone, 2. coenzyme) - in a simplified model might no longer be grasped. This is often the two one-substrate terms are multiplied giving case in dehydrogenase reactions with in situ a complete rate equation of the alcohol oxidacoenzyme regeneration by the enzyme- or sub- tion reaction termed “Michaelis-Menten doustrate-coupled approaches. The simulation of ble substrate kinetics” (WICHMANN et al., 1981; coupled enzyme reactions based on the meas- BISELLIet al., 1995). Additional terms are inured kinetics and mass balances can be esti- cluded considering the product inhibition and mated by numerical integration of the diffe- the substrate inhibition (BISELLIet al., 1995). rential mass balance equations by means of For the reverse reaction the corresponding the Runge-Kutta method (HOFFMANNand rate equation has to be written. The velocity HOFMANN, 1971).For this purpose, the simula- term v is then expressed as the difference of tion program SCIENTISTTM (MS”-Win- the reverse reaction (R2) and the forward redowsTM, MicroMath, USA) or equivalent action (R,)multiplied by the enzyme concensoftware can be used. Modeling of enzyme re- tration (E). This simple approach regards the actors is based on mass balance equations of a oxidation and reduction reactions as two indecontinuously stirred tank reactor (CSTR). The pendent processes, although they are catalyzed accumulation of product can be described by at the same active site of the enzyme. The mathe decrease of substrate concentration as a jor disadvantage of this simplified model is function of time: that the equilibrium constant, Keg cannot be calculated from the kinetic constants (BISELLI, Accumulation =Convection + Reaction 1991).Therefore, the applicability of this model is restricted to reactions with an equilibrium on the product side (SEELBACH, 1994; KRUSE 1995). Another approach describing the reaction V velocity uses the rate equation based on the rewith: T = action mechanism (SEGEL,1975;BISELLIet al., F 1995). In this case, oxidation and reduction reIt reactor volume [L] actions are not regarded as independent proF: substrate feed rate [L h-’1 cesses, but instead, the model accounts for the So inlet substrate concentration [MI competition of both reactions for the same acS outlet substrate concentration [MI tive site. A complete kinetic characterization T mean residence time [h] of the enzyme(s) is necessary to define the rev reaction velocity term action mechanism(s). The exact rate equation then accomplishes the calculation of the equiThe mass balance term consists of the accu- librium constant, which is not possible in the mulation term, a convection term, and a reac- simplified model (see above). Hence, the modtion velocity term. The convection terms de- el based on the reaction mechanism allows scribe the changes of the concentration of each simulation of behavior of the enzyme systems reactant within the residence time in the reac- much more precisely. tor. The reaction velocity term takes into acFor more detailed information regarding count the position of the equilibrium and the the determination of enzyme kinetics, reactor velocity. modeling, and the formulation of mass balancThe reaction velocity term consists of either es for coupled enzyme systems the reader is rethe Michaelis-Menten term or the rate equa- ferred to the literature (BISELLIet al., 1995;

434


BISSWANGER, 1979; HOWALDTet al., 1986; formate to carbon dioxide), another approach HUMMELet al., 1986; IKEMIand ISHIMATSU, giving access to the opposite enantiomer of a 1990; KATAYAMA et al., 1983; KRAGLet al., given hydroxy compound is possible. First, the 1996a; KULAet al., 1980; LIESEet al., 1996; carbonyl compound (30) is reduced chemically LIESEet al., in press; OKADAand URABE1987; to the corresponding racemic hydroxy comPETERSand KULA,1991;SCHMIDT et al., 1987b; pound ruc-(31).Then, in a second step, one enantiomer is oxidized selectively by the dehyVAN HOOIDONK and BREEBAART-HANSEN, et al., 1989; WICHMANN et drogenase giving the desired enantiomer ( R ) 1971; VASIC-RACKI (31) (Fig. 22) and again the carbonyl comal., 1981; ZELINSKI, 1995). pound (M), which can be recycled. When the enzyme is completely stereospecific, the oxidation of the racemic hydroxy compound terminates automatically at the “50%-of-reaction’’ 6.4 Resolution of Racemic point when all the reactive enantiomer has Hydroxy Compounds with Alcohol been transformed. Therefore, recycling of the carbonyl compound theoretically allows Dehydrogenases achievement of up to 100% yield of transformation. A number of examples making use of the 6.4.1 Access to Both Enantiomers above described methodology have been pubof a Hydroxy Compound lished over the years (JONESand TAYLOR, 1976; JONES, 1985; LEMIERE et al., 1985; HUM1991; PETERSand KULA, In practice, the synthesis of both enantiom- MEL and GOTTWALD, ers of a chiral hydroxy compound may be de- 1992; LIESE,1994; LIESEet al., 1996; FABER, 1994) (Fig. 23). sirable for a number of different reasons. On 1995; WONGand WHITESIDES, One prerequisite for the economic kinetic the other hand, only a limited set of dehydrogenases with opposite enantioselectivity and resolution of alcohols via selective oxidation is overlapping substrate specificities are avail- the effective regeneration of NAD .The highable for the direct reduction of a given carbo- ly recommended cofactor recycling systems nyl compound which is the most favorable for NAD are the glutamate/glutamate dehyroute to a chiral hydroxy compound (cf. Sect. drogenase and the pyruvate/lactate dehydrogenase systems (see Sect. 3). The other prere4). Since most dehydrogenase-catalyzed reac- quisite is to overcome product inhibition tions are reversible (exception: e.g., the for- caused by the carbonyl compound by using mate dehydrogenase-catalyzed oxidation of suitable reactor concepts (LIESEet al., 1996). +

+

(S) -Selective dehydrogenase

R‘

(S) Selective

dehydrogenase

-OH

Fig. 22. Use of a single dehydrogenase to prepare both enantiomers of an alcohol.


OH HLADH,pH9

*

33

NAD recycling, 5 0 8 oxidation

rac-3 2

OH (1R)-32

Fig. 23. Enantioselective alcohol oxidation.

6.4.2 Introduction of Two Chiral Centers in One Reaction Step Normally, oxidoreductases can only introduce one stereochemical center at a time by the stereoselective reduction of a carbonyl compound. On the contrary, oxidoreductases accepting a-acidic carbonyl compounds, like

OEt

435

a-branched P-ketoacid esters, can be used to introduce two stereogenic centers in only one reaction step. a-Branched P-ketoacid esters easily racemize via the keto-enol tautomeric equilibrium, whereas the reduced compounds do not racemize in water. Therefore, it is possible to perform a diastereoselective reduction with this type of substrates. Fig. 24 shows the four possible diastereomers of the reduction of 2-alkyl-3-oxocarboxylicacid ester. 2-alkyl-3-hydroxyacidesters (35) are useful chiral building blocks for the synthesis of a range of compounds such as pheromones, macrolides, P-lactam antibiotics, and other natural products (SEEBACH et al., 1987;NAKAMURA, 1992).The reduction of the corresponding 2-alkyl-3-ketoacid esters (34) was performed by using two NADPH-dependent oxidoreductases from bakers’ yeast (SHIEH,1987;SHIEH et al., 1985; NAKAMURA et al., 1991) and two NADH-dependent oxidoreductases from Candida parapsilosis and Rhodococcus erythropolis (PETERS et al., 1993a; ZELINSKI, 1995). One of the bakers’ yeast enzymes called Llenzyme produces syn-(2R, 3S)-(35) alcohol

DH

( 2 S ) - 34

sy I t

anti

36

( 2 R ) - 34

Fig. 24. Stereochemistry of the diastereo- and enantioselective reduction of racemic 2-methyl-3-oxobutanoic acid ethyl ester. anri-(2S,3S)-35:produced by the NADPH-dependent oxidoreductase from Georrichum candidurn (98% de). syn-(2R,3S)-35: produced by the NADPH-dependent bakers’ yeast oxidoreductases:L1-enzyme (99% de) and L2-enzyme (82% de). the NADH-dependent carbonyl reductases from Candida parapsilosis (98% de) and Rhodococcus eryrhropoiis (95% de).

436

9 Dehydrogenases- Characteristics, Design of Reaction Conditions, and Applications

with a diastereomeric excess (de) of 99% (KAWAIet al., 1994a),whereas the other called L2-enzyme yields a mixture of syn-(2R, 3s)(35)and anti-(2S, 3S)-(3S) products (82% de; NAKAMURA, 1992). The carbonyl reductases from Candida parapsilosis (CPCR) and Rhodococcus erythropolis (RECR) reduce 2-methyl-3-oxobutanoic acid ethyl ester (34) with diastereomeric excesses of 98% and 95%, respectively, to the corresponding syn-(2R, 33)-(3S) (PETERSet al., 1993a;ZELINSKI, 1995). Anri-(2S, 33)-(35) was synthesized using the isolated NADPH-dependent oxidoreductase from Geotrichurn candidum (KAWAIet al., 1994b), whereas the other two diastereomers, syn-(23, 3R) and anti-(2R, 3R), have not been synthesized enzymatically until now.

7 Applications of Alcohol Dehydrogenases

6.5 Conclusions

In most cases when a (bio-)chemist is trying to convert a given substrate, screening for a suitable enzyme is of major importance. Normally, before an extensive microbial screening program is initiated, the known enzymes are first tested. Therefore, this section may be of special relevance for synthetic chemists, since most of the criteria necessary to preselect and use a dehydrogenase are given in tables. In the case of the noncommercial dehydrogenases, suitably equipped microbiology and biochemistry laboratories should be available to handle and supply considerable amounts of biomass and isolated enzymes. Therefore, the microorganism, tissue, or plant containing the enzyme is given, and suppliers of commercial dehydrogenases are listed. Moreover, the section deals with selected applications of dehydrogenases, where either large-scale continuous synthesis has been performed, or the product is of commercial interest, or a broader synthetic applicability of a chiral hydroxy compound is given.

The development of an enzymatic process involves the following aspects: (1) reaction conditions, (2) process design, (3) thermodynamics, and (4)kinetics. Therefore, kinetic aspects play an important role during the development of an enzymatic process. As outlined in this section, the stereochemistry can be used to classify oxidoreductases and to predict the stereochemical outcome of a reaction. This is of special relevance for a synthetic chemist looking for a catalyst of a given enantioselectivity. Furthermore, the kinetics of oxidoreductases are of considerable importance for the use of these enzymes in organic synthesis. The kinetic mechanisms of oxidoreductases (ordered bi-bi- sequential mechanism and the Theorell-Chance derivative) are necessary for the understanding of the kinetic limitations caused by noncompetitive, competitive, and uncompetitive types of inhibition. Knowledge of the kinetic parameters is the basis for overcoming kinetic limitations, for evolving a model, and predicting the behavior of a suitable enzyme reactor. Enormous efforts have been made in the past to establish suitable reactors specially designed for oxidoreductases and to model their behavior. Finally, exemplary applications are discussed making use of the kinetic resolution of racemic hydroxy compounds.

In this section, commercial and noncommercial dehydrogenases and reductases which are useful in organic synthesis are discussed with respect to the following criteria: - Summary of substrate spectra and coen-

zyme dependencies,

- biochemical characterization including

temperature/pH optima and stability, stereospecificity, compatibility with organic solvents and detergents (reversed micelles), inhibitors, and the specific activity, - compatibility with enzyme- or substratecoupled coenzyme regeneration methods.

7.1 Commercial Dehydrogenases, their Characteristics and Applications Commercial dehydrogenases useful in enzymatic synthesis are available from several manufacturers (Tabs. 10 and 15). Except for


horse liver alcohol dehydrogenase (HLADH), some hydroxysteroid dehydrogenases (HSDH), and the L-lactate dehydrogenase from bovine muscle, all other dehydrogenases mentioned in the subsequent sections are isolated from microorganisms.

7.1.1 Horse Liver Alcohol Dehydrogenase HLADH (EC 1.1.1.1) is probably one of the best known enzymes. It is a very useful enzyme in organic synthesis because of a number of characteristics which are discussed in the following section. The enzyme exhibits a broad substrate specificity and a narrow stereospecificity (Tab. 10).About one third of the publications on alcohol dehydrogenases cited in the Biotransformation Databases (Chapman &

Hall, Synopsys) deal with HLADH-catalyzed oxidoreactions. Thus, it is one of the most widely used dehydrogenases in biotransformations. The enzyme consists of two nearly identical subunits each containing two zinc ions. Thus, the enzyme is susceptible to inactivation by metal chelating reagents. Inhibition may also be observed at high concentrations of ions. HLADH is stable at ambient temperature. Temperatures above 30 "C increasingly inactivate the enzyme (GORISCHand SCHNEIDER, 1984). HLADH can be immobilized on solid supports like CNBr-activated sepharose (GORISCH et al., 1984) to increase its stability. It is impossible to define a single overall pH optimum because the effects of pH on the interaction of the enzyme with alcohol and carbonyl substrates, and with NAD(H), are all different (SUNDand THEORELL, 1963).The enzyme shows a broad pH stability (5-lo), al-

Tab. 10. Substrate Spectra of Selected Dehydrogenases and Reductases Useful in Organic Synthesis

E c

w

YADH HLADH CFADH MJADH CPCR RECR PADH PLADH TBADH L-LDH D-LDH D/L-HicDH GlyDH HSDH LKADH

S. cerevisiae Horse liver C.falcata M . javanicus C. parapsilosis R. erythropolis f! spec. Pig liver T brockii Bovine muscle L. leichmanii L. caseilconfusus G. candidum different sources L. kefir

NADH NADH NADPH NADPH NADH NADH NADH NADPH NADPH NADH NADH NADH NADH NAD (P)H NADPH

437

to 50 8.5-9.5 to 30 8.5-9.5 20 7.3

50 25

25

75

60 50

HSDH

AlaDH

LeuDH PheDH GluDH

1.5-8

6.4-7

25

25

UP

UP

UP

7.2 7.2 to65 7.5-8

25 25

re-face si-face re-face si-face

7

11.5 10-1 1 8.3

10.5

8-9

si-face si-face si-face

si-face

5.5-9.5 5.69.8 7.5-8.5 7

re-face

7.5-9.5

7-9 7.5

6

re-face

6.69.5

8

8.7-9 8.7-9 7.8-9 9

re-face

5-10

8-9

StereoPH PH Optimum Stability specifi(oxidacity tion)

J J

J

J

J

J

SpecificActivity [U mg-' protein] Reduction Reaction

chelators, SH blockers, 2.7 heavy metal ions chelators, heavy metal > 300 ions, SH blockers 450 Oxidation Oxidation 300 SH blockers 8(25"C) Zn*+,Na +,Li +, 25 SH blockers, DTE heavy metal ions, 5-350* SH blockers Hg-, Cu-ions,p-CI-mer13 curibenzoate 60-120 65 50

Compatibility Compatibility Inhibitors with Organic with DeterSolvents gents

HL: horse liver,TB: Thermoanaerobium brockii, Phe: Phenylalanine,Y yeast, Gly: glycerol,Ala: Alanine, Leu: leucine, L lactate, DTE: dithioerythritol, HS: hydroxysteroid,Glu: glutamate, *: Specific activity varies between different HSDHs

40

uoto75 8.2

30 30

L-LDH D-LDH TBADH GlyDH

7.15

10-25

30

YADH

7

25

HLADH 37

Enzyme T T PH Optimum Stability Optimum ["C] (reduc["CI tion)

Tab. 11. Biochemical Characteristicsof Selected Commercial Dehydrogenases

s-

!i

F

>linear ketones > > cyclic ketones > > primary alcohols. However, a,P-unsaturated ketones and ketones, where both substituents are larger than ethyl, are not accepted as substrates. Based on kinetic data, KEINAN and coworkers (1986a) proposed a model of the substrate binding site of TBADH. Two hydrophobic pockets of different geometry bind the small (methyl or ethyl) and the large alkyl chain (up to C,H,,), directing the carbonyl function such that the hydride transfer can occur to the re face. This was developed into a four domain binding model by KELLYand LEWIS(1991).

7.1.4 L - L a c t a t e Dehydrogenases L-Lactate dehydrogenases (EC 1.1.1.27) have been isolated from several sources: rabbit and pig muscle, pig or beef heart, beef liver, mouse testes, heart and kidney, chicken liver and heart, spiny dogfish, lobster tail, Limulus polyphemus, and the bacterium Bacillus stearorhermophilus. Some of these LDHs have been crystallized, and the three-dimensional structure was solved (see EVERSE et al., 1982 for an overview). Recently, lactate dehydrogenase was crystallized in the Russian Space Station Mir in a microgravity environment (KOSZELAK et al., 1996). This inexpensive and very active enzyme (450-550 U mg-') is stable at its temperature optimum of 25 "C provided a suitable protection agent against autooxidation has been added (e.g., dithiothreitol). The thermal stability can be enhanced drastically upon immobilization on a solid support (e.g., polyacrylamide; HICKSand UPDIKE,1966; WIELAND et al., 1966).The equilibrium of the reaction favors L-lactate production (Keg= 2.76 * lo-'* M; BERGMEYER, 1983), which is

one reason for the frequent application of LDHs for NAD+ regeneration (see Sect. 3). The pH optima are pH 7.2 and pH 8.9 for the reduction and oxidation reactions, respectively.The enzyme is stable at pH 7 and 25 "C. Inhibition of the L-LDH can be observed at high pyruvate concentrations. Addition of sodium and chloride ions increases the apparent K,, value for pyruvate at concentrations of 0.2 M (BERGMEYER, 1983). The cloned L-LDH from B. stearothermophilus as well as the rabbit muscle enzyme have been successfully coupled with enzymatic (FDH) and electrochemical systems for coenzyme regeneration (BURet al., 1989; CASYet al., 1992;KIMand WHITESIDES, 1988;BOURDILLON et al., 1988). The NADH-dependent L-LDH is specific for 2-oxocarboxylic acids (esters are not accepted) converting them to (S)-hydroxyacids, which are highly valuable chiral building blocks. Only aliphatic residues are substrates of the enzyme (KIMand WHITESIDES, 1988). The L-LDH from B. stearothermophilus is more specific for short chain a-ketoacids, whereas the enzyme from testes also accepts larger substrates. The stereospecificity is generally very high ( > 99% ee).

7.1.5

D-Lactate

Dehydrogenases

D-lactate dehydrogenases (EC 1.1.1.28) have been isolated from bacteria such as Leuconostoc mesenteroides, Staphylococcus epidermis, Lactobacillusfuseus (DSM 20196), and Lactobacillus leichmanii. As with L-LDH, the D-LDH from L. mesenteroides is inexpensive and exhibits a high specific activity (300 U mg-').The enzyme is stable at its temperature optimum of 25°C provided a suitable protective agent against autooxidation has been added (e.g.,dithiothreitol or P-mercaptoethanol). The pH optima are pH 7 and pH 9 for the reduction and oxidation reactions, respectively. The enzyme is stable at pH 6 and 25 "C (BERGMEYER, 1983). The D-LDH has been successfully coupled with the formate dehydrogenase (FDH) from Candida boidinii which was employed for coenzyme regeneration (KIMand KIM,1991).Either co-immobilization (SIMONet al., 1989) or


443

retainment behind a membrane (enzyme The pH optimum of the reduction of dihymembrane reactor; WANDREY et al., 1982a, b; droxyacetone is between pH 7.5 and 8 and for HUMMEL et al., 1984) may be used to achieve a the oxidation reaction pH 9. GlyDH is activatcontinuous process. ed by ammonium and potassium ions, but inThe D-LDH reduces specifically 2-oxocar- hibited by zinc, sodium, and lithium ions. Furboxylic acids at the expense of NADH to the ther inhibitors are N-ethylmaleiimide, iodocorresponding (R)-2-hydroxycarboxylic acids acetamide, dithioerythritol, and P-mercaptowith high enantioselectivity (GHISALBA et al., ethanol. 1989; SIMONet al., 1989; BRADSHAW Both the glycerol dehydrogenases from E. et al., 1991; KIMand KIM,1991). It is often desirable aerogenes and from G. candidum accept 2to have access to both enantiomers of a given propanol as co-substrates allowing the subcompound. L- and D-LDH are more or less strate-coupled regeneration of NADH (NAKcomplementary to each other in that the oppo- AMURA et al., 1988,1992). Moreover, the biosite enantiomer of a number of substrates can chemical characteristics of GlyDH are combe obtained. patible with formate dehydrogenase, which is D-LDHfrom L. mesenleroides shows a sub- often employed for coenzyme regeneration et al., 1987). strate tolerance which is substantially narrow- (CHENAULT er than that of the L-enzyme, as 2-oxoacids Many a-hydroxy ketone substrates have with side chains longer than three carbons are been found for this enzyme that can be either converted at a reduced rate. However, the en- cyclic or acyclic (LEE and WHITESIDES, 1986; et al., 1987). In the oxidative direczyme accepts 2-0x0-3-phenylpyruvic acid and CHENAULT 2-0x0-4-phenyl-pyruvic acid as substrates (SI- tion a broad range of substrates are accepted MON et al., 1989; BRADSHAW et al., 1991; KIM (Tab. 13). Hydroxy aldehydes are, however, and KIM,1991).The D-LDHfrom L. leichmanii not accepted as substrates. The GlyDH from reduces also 2-0x0-4-phenylpyruvic acid (GHI- E. aerogenes was used for the kinetic resoluSALBA et al., 1989), whereas the D-LDH from tion of racemic l-phenyl-1,2-ethanediol in a S. epidermis is able to accept a-ketobutyric ac- continuously operated enzyme reactor id, a-ketopentanoic acid, phenylpyruvate, and equipped with a hydrophobic hollow fiber a-ketocyclopropyl acetic acid (KIM and KIM, membrane for continuous product extraction 1991) as well as 4-benzyloxocarbonylamino-2- (LIESEet al., 1996). Oxidation of alcohols by oxobutanoic acid (BENTLEY et al., 1995) with alcohol dehydrogenases often suffers from severy high enantioselectivities ( > 99% ee). vere inhibition by the ketone product, which can be overcome by continuous product extraction. The K , value of GlyDH for l-phenyl-1,2-ethanediol is 10.8 mM, whereas the in7.1.6 Glycerol Dehydrogenases hibition constant K ifor the corresponding ketone is only 0.25 mM, indicating a strong prodGlycerol dehydrogenase (EC 1.1.1.6) naturally interconverts glycerol to dihydroxyacetone ((R)-1,2-diol).The enzyme has been isolat- Tab. 13. Substrates Accepted by Glycerol Dehyed from bacteria (Enterobacter aerogenes, Cel- drogenases (Oxidative Direction) lulomonas sp., Lactobacillus sp.; SLININGER et Substrate Relative rate [%] al.. 1982) and fungi (Geotrichum candidum; NAKAMURA et al., 1988). GlyDH is stable at its 1.2-Propanediol 100 temperature optimum of 25 "C and at a pH of 2.3-Butanediol 100 37 7.5 and can be used in organic-aqueous bi- 1,3-Butanediol 20 phasic media after immobilization (NAKAMU- Ethylene glycol 18 RA et al., 1992; Japanese Patent JP 05328-984, Inositol 17 1993;LEEand WHITESIDES, 1985;CHENAULT et 1,4-Butanediol 17 al., 1987).The equilibrium strongly favors the 2-Propanol Sorbitol 3 reduction reaction (Keq=5.1 . M: BERG- Ethanol 1 MEYER, 1983).

444

9 Dehydrogenases - Characteristics,Design of Reaction Conditions,and AppIications

uct inhibition which leads to a very inefficient conversion.However, applying the novel differential circulation reactor with continuous product extraction allowed conversion of 50% with respect to the racemate at an enantiomeric excess of greater than 99% of the (S)-enantiomer. This underlines the necessity of novel reactor concepts when dealing with enzymes (see Sect. 5). For synthetic purposes, the GlyDH from Cellulomonas sp. is superior due to its higher specific activity and lower cost (LEE and WHITESIDES, 1986).2- and 3-ketocarboxylicacid esters are reduced to the corresponding (2R)- and (3R)-hydroxycarboxylicacid esters by the glycerol dehydrogenasefrom G. cundidum (NAKAMURA et al., 1988,1992; PATELet al., 1992). For example, ethyl 4-chloro-3-oxobutanoate (40)was reduced stereoselectivelyto the corresponding ethyl 4-chloro-(3S)-hydroxybutanoate (41) using the GlyDH from G. cundidum (55% yield, 99% ee; NAKAMURA et al., 1988; Fig. 26). The product is a key intermediate in the synthesis of L-carnitine. Recently, another secondary alcohol dehydrogenase was isolated from G. cundidum W F 9101 accepting (S)-diols,3-ketocarboxylicacid esters, and a-branched derivatives as substrates (MORIet al., 1996). However, this enzyme reduces the carbonyl function to an ( 8 ) hydroxy function and is thus not identical with the above described GlyDH. The enzyme has been used by MORIand coworkers for the preparation of (R)-diols from racemic diols.

7.1.7 Glutamate Dehydrogenases Glutamate dehydrogenase (GluDH) (EC 1.4.1.3) belongs to the amino acid dehydrogenases catalyzing the reductive amination of aketocarboxylic acids to the corresponding L(R)-amino acids. In the reductive amination,

GlyDH

P Cl

O

E

t

OEt

c1

40

Fig. 26. See text.

the enzyme converts 2-oxoglutarate to L-glutamate which is one of the key reactions in the biosynthesis of amino acids. L-Glutamate is produced on a several hundred ton scale by a fermentation process and mainly used as a food additive (IZUMet al., 1978; SODAet al., 1983). In the oxidative direction the L-enantiomer is converted to the a-ketocarboxylic acid leaving the D-enantiomer,which can be prepared in this manner (see also Sect. 6.4.1).The equilibrium of the GluDH reaction is greatly in favor of glutamate formation (Keg= 1.5 . to 2.5 L mole-' depending on pH, ionic strength, and temperature). Unusually, the enzyme accepts both NADP(H) and NAD(H). Glutamate dehydrogenasehas been isolated from the livers of numerous species (human, mouse, rat, horse, dog, etc.) and from other tissues like kidney, heart, and brain. The enzyme from beef liver is commercially available. The enzyme consists of minimally six polypeptide chains with molecular weights between 50 and 60 kDa. Hence, the molecular weight of the active enzyme ranges between 310 and 350 kDa. GluDH is characterized by a temperature optimum of 40°C and pH optima of 7.3 and 8.0 for reduction and oxidation, respectively. The enzyme is stable at temperatures up to 25°C and at a pH of 7. These features allow the coupling of the enzyme to most of the enzymes used for coenzyme regeneration (see Tab. 14). Moreover, GluDH can be immobilized on solid supports to increase its stability (COPPER and GELBARD, 1981). Above pH 7 GluDH is activated by adenosine diphosphate (ADP), which weakens coenzyme binding to the active site, whereas the products of the reductive amination, glutamate and NAD(P)+, slightly inhibit the enzyme. Calcium ions also inhibit the enzyme. The product inhibition caused by 2-oxoglutarate, ammonium ions, and NAD(P)H for the oxidative deamination is more severe. Therefore, besides the equilibrium favoring the glutamate formation, the reductive amination is also the preferred direction for kinetic reasons. Dextran-bound NAD (Gu and CHANG, 1988) as well as polyethylene glycol-bound NAD(P)(H) is accepted by GluDH (PETERS, 1990) with about 4-fold lower affinity compared to the native coenzymes.

-

+

4 1


445

Tab. 14. Biochemical Characteristics of Selected Enzymes for the Regeneration of NADH

Enzyme Formate dehydrogenase Glucose-6-phosphatedehydrogenase Glucose dehydrogenase Horse liver ADH Thermoanaerobium brockii ADH

T Optimum [“CI

55

30

25 25

70

Glutamate and ammonium ions stabilize GluDH in a concentration-dependent manner, whereas 2-oxoglutarate leads to increased destabilization with increased concentration (PETERS, 1990) ADP also exhibits stabilizing effects (LEHMANN, 1969). Other amino acid dehydrogenases, like phenylalanine dehydrogenase, also show comparable stabilizing and destabilizing effects of substrates and products (SCHMIDT, 1987). Besides 2-oxoglutarate, which is the best substrate, GluDH accepts other mono- and dicarboxylic acids (STRUCKand SIZER,1960). Hence, the enzyme is useful to synthesize unnatural amino acids like L-a-aminoadipic acid, (2R, 3R)- and (2R, 3S)-2-amino-3-fluoro-glutaric acid (VIDAL-CROS et al., 1989) or D-amino acids (NAKAJIMA et al., 1988). Furthermore, GluDH was used for the preparation of 13N-labeled L-amino acids (COPPERand GELBARD, 1981).

T Stability [“CI

pH Optimum

25

7.5-8.5 7.8 7.6-8.2

30

25

37

10-85

8-9 7-9

Specific Activity [Ums-’l 4

650 250

1-2 17 (25°C)

As with GluDH, the equilibrium of the reaction is by far on the side of aminated products (Keq=9 . 10l2 at pH 11). The enzyme shows pH optima at 8.5 to 9.5 and at 11.5 for reductive amination and oxidative deamination, respectively (SCHUTTEet al., 1985).The temperature optimum is at 60°C and the enzyme is stable at temperatures up to 50°C and pH values between 5.6 and 9.8. The enzyme has been crystallized successfully (SCHUTTEet al., 1985). LeuDH normally catalyzes the reductive amination of branched-chain a-keto acids to the corresponding a-amino acids like leucine, but it is also useful for the synthesis of unnatural amino acids such as rerr-(S)-leucine, (S)-/3hydroxy-valine, and very hydrophobic branched chain (S)-amino acids (SCHUTTE et al., 1985; WANDREY, 1986;WANDREY and Bossow, 1986; G u and CHANG,1990: HASUMIet al., 1996; KRIXet al., 1997). fen-Leucine finds increasing use as templates or catalysts in asymmetric synthesis as well as in peptidic medicinal compounds, such as anti-AIDS, anti-arthritic, and anti-cancer drugs (BOMMARIUS et al., 1995). 7.1.8 Leucine Dehydrogenases LeuDH was intensively studied with respect to Leucine dehydrogenase (LeuDH) (EC its kinetic characteristics (KRAGL et al., 1.4.1.9) has been isolated from Bacillus species 1996b). Recently, the LeuDH from B. cereus such as B. sphaericus DSM 396 (HUMMEL et (DSM 626) was cloned and overexpressed acal., 1981;WANDREY et al., 1982c;H~NsoNet al., tively in E. coli to yield approximately 30% of 1990a), B. cereus (SCHUTTEet al., 1985).B. sub- the total soluble protein (STOYAN et al., 1997). tilis (HANSONet al., 1990b), and the thermo- The unnatural substrate trimethylpyruvate phile B. sfearothermophilus (OHSHIMA et al., slightly inhibits LeuDH, which also acts as a 1985). In contrast to GluDH, LeuDH is only noncompetitive inhibitor of the formate dehyactive with NAD(H).The enzyme accepts both drogenase employed for coenzyme recycling polyethylene glycol-bound NAD(H) (WICH- (refer to Sect. 6.3.2 for cross inhibition). The MANN et al., 1981; WANDREY et al., 1982c; Ku- product, tert-leucine, shows a very strong prodLA and WANDREY, 1987; HANSON et al., 1990b; uct inhibition on LeuDH, hence a plug flow reKRAGLet al., 1996b) and dextran-NAD’ (Gu actor (low product concentration in the reactor) would be advantageous (see Sect. 6.3.3). and CHANG,1990).

446

9 Dehydrogenases - Characteristics, Design of Reaction Conditions,and Applications

Based on the kinetics of the LeuDHiFDH system a continuous industrial process is operating at Degussa AG (Germany) producing high quality tert-(S)-leucine on a multi-100-kg scale (BOMMARIUS, 1993).Space-time yields of up to 366 g L-' d-' at a mean conversion of 93% were reported (KRAGL et al., l993,1996a, b). L-Leucine is produced at a 0.5 kg d-' scale (LEUCHTENBERGER et al., 1984b). It is worthwhile to mention that the process also yields equally high total turnover numbers (see Sect. 3) with native cofactor instead of the PEG-bound NAD(H) (WANDREY and Bossow, 1986). Therefore, the cofactor costs are no longer a limiting factor for enzymatic redox reactions (KRAGLet al., 1996b). Moreover, LeuDH from the thermophile B. stearothermophilus is more resistant to both thermal and chemical denaturation than the mesophilic enzyme. It has been shown that the thermostable LeuDH is better suited in the continuous production of L-leucine in an enzyme membrane reactor compared to its counterpart with mesophiles (OHSHIMA et al., 1985).

7.1.9 Phenylalanine Dehydrogenases NAD(H)-dependent phenylalanine dehydrogenases (PheDH) (EC 1.4.1.20) have been et isolated from Brevibacterium sp. (HUMMEL al., 1984,1986;KULAet al., 1986),Rhodococcus sp. M4 (HUMMEL et al., 1987; SCHMIDT et al., 1987a; BRADSHAWet al., 1991), Bacillus sphaericus (ASANOet al., 1990),and a thermostable enzyme was cloned from Thermoactinomyces intermedius into Escherichia coli (TAKADA et al., 1991).The latter enzyme consists of 6 subunits of 366 amino acids and exhibits homology to other PheDHs (56% to B. sphaericus, 42% to Sporosarcina ureae, 47% to LeuDH from B. stearothermophilus). The enzyme was cloned into E. coli JM109 with an expression of 8.3% of the soluble protein (TAKADA et al., 1991). The PheDH isolated from Sporosarcina sp. is commercially available (Sigma). The enzymes from Brevibacterium and Rhodococcus accept polyethylene glycol-bound NAD(H) and can be coupled to enzymes for cofactor re-

cycling (HUMMELet al., 1986;KULAet al., 1986; SCHMIDT et al., 1987a,b). Phenylalanine dehydrogenase from Rhodococcus sp. is characterized by a temperature optimum of 50 "C and pH optima of 8.5 to 9.5 and 10 to 11 for the reduction and oxidation direction, respectively.The enzyme is reasonably stable at temperatures up to 30°C (5% inactivation d-') and can be used continuously in an enzyme membrane reactor at a spacetime yield of up to 456 g L-' d-' a turnover number of up to 600000, and 95% conversion (HUMMEL et al., 1986;SCHMIDT et al., 1987a,b; HUMMELand KULA,1989). Phenylalanine is used in the synthesis of the low-calorie sweetener aspartame (8000 t a-', 1990)and is currently produced by a fermentation process. It is also a useful chiral synthon (DRAUZet al., 1982). Besides its natural substrate, 2-oxo-3-phenylpropanoic acid, PheDH from Rhodococcus sp. accepts a range of other aromatic 2-0x0acids for the preparation of unnatural amino acids, e.g., p-halogenated phenylalanine,(S)-2amino-4-phenylbutanoic acid, and (R)-Zhydroxy4phenylbutanoic acid (HUMMEL et al., 1987;BRADSHAW et al., 1991). PheDH from B. sphaericus (strain collection number SCRC R79a) was also explored for synthesis and shown to be compatible with the FDH for coenzyme regeneration (ASANO et al., 1990).The enzyme was used for the preparation of different natural and unnatural L-amino acids, e.g., phenylalanine, tyrosine, 4-(fluoro-phenyl)alanine, 2-amino-4-phenylbutyricacid, 2-aminononanoic acid, and 4-vinyl-~-phenylalanine. Furthermore, the enzyme has been cloned and overexpressed in E. coli.

7.1.10 Alanine Dehydrogenases Alanine dehydrogenase (AlaDH) (EC 1.4.1.1) catalyzes the reversible NADH-dependent reductive amination of pyruvate, and has an important role in the carbon and nitrogen metabolism of various microorganisms. The enzymes from different mesophilic and thermophilic Bacillus strains, halophilic bacteria, and other bacteria (Pseudomonas,Streptomyces, Thermus) have been isolated and characterized in detail (for references see OHSHI-

7 Applications ofAlcohol Dehydrogenases MA et al., 1990 HONORAT et al., 1990).The enzymes from Bacillus subtilis and B. sphaericus have been demonstrated to accept polyethylene glycol-bound NAD(H) and can be coupled to enzymes for cofactor recycling (WANDREY et al., l981,1982a,b; OHSHIMA et al., 1989). Alanine dehydrogenases from the mesophilic B. sphaericus and B. subtilis and the thermophilic B. sphaericus strains (IF0 3525, and DSM 462) are quite similar to each other: The high pH optima for both oxidative deamination and reductive amination at pH 10.5 and 8.2, respectively, a strong inhibition of the activity by HgCl, and CuC12,and high specificity for L-alanine and NAD+ in the oxidative deamination reaction are common characteristics of these enzymes. However, the thermophilic AlaDH is more stable against thermal and chemical denaturation and thus may be more suitable for continuous applications. The enzyme is stable against heating at 75°C and pH-resistant between pH 5.5 and 9.5. However, the specific activity at 37°C is lower (13 U mg-l) compared to the mesophilic enzyme (30-40 U mg-’ for B. subtilis AlaDH). AlaDH from B. megaterium ATCC 39118 (HONORAT et al., 1990) is also a stable enzyme with a half-life of 22 d when incubated at 30°C. The enzyme is characterized by an optimum temperature of 52°C and pH optima at pH 8 and 10.5 for reduction and oxidation, respectively. AlaDH accepts a range of substrates for both reductive amination and oxidative deamination reactions. Pyruvate and L-alanine are the preferred substrates. Besides other substrates, a-ketoglutarate, hydroxypyruvate, 3bromopyruvate, and a-ketoisolvalerate as well as D-alanine, L-serine, and aminobutyrate are converted by AlaDH (HONORAT et al., 1990). AlaDH has been used for the continuous production of L-alanine in an enzyme membrane reactor (WANDREY et al., 1981, 1982a; LEUCHTENBERGER et al., 1984b), for the synthesis of the essential amino acid L-valine (93-100% yield) starting from 3-methyl-20x0-butyric acid using glucose dehydrogenase for NADH regeneration (MONOTet al., 1989; HONORAT et al., 1990), and for the continuous preparation of 3-fluoro-~-alanine(OHSHIMA et a]., 1989), which is a potentially useful compound for the synthesis of antibacterial and antiviral agents and insecticides.

447

7.1.11 Hydroxysteroid Dehydrogenases Commercially available hydroxysteroid dehydrogenases (HSDH) are presently manufactured from Pseudomonas testosteroni (ATCC 11996), E. coli, Pseudomonas sp., Bacillus sphaericus, and Streptomyces hydrogenans. Other 3a-HSDHs have been described from Cellulomonas turbata FERM P-9059 (KISE and HAYASHIDA, 1989) and rat liver (TALALAY, 1962). The enzymes catalyze the regiospecific oxidation of a hydroxy group or the regio- and stereospecific reduction of a ketone functionality of a steroid. The regiospecificity regarding the natural substrate is designated by the name of the particular enzyme (e.g., 3aHSDH, EC 1.1.1.50; 3P-HSDH, EC 1.1.1.51; ~cY-HSDH,EC 1.1.1.159; 12a-HSDH, EC 1.1.1.176; 3a,20P-HSDH, EC 1.1.1.53). The properties of the hydroxysteroid dehydrogenases from I? testosteroni, S. hydrogenans, and rat liver have been compiled by TALAY (1962). ?? testosteroni induces 3a- and 3PHSDHs when grown on testosterone as carbon source. The enzyme is strongly inhibited by heavy metal ions and p-chloromercuribenzoate, but preincubation with NAD can protect the enzyme from inactivation. 3P-HSDH is powerfully stabilized by low concentrations of estradiol-17P. Both enzymes are inactive with NADP(H). The Michaelis-Menten constants for the steroid substrate are between lo-’ M and lo-’ M. The 3a-HSDH oxidizes only 3a-hydroxysteroids of the C,,,C2,, and C,, (bile acid) series.Typica1substrates are androsterone (42) (Fig. 27) (&: 1.5 FM), 3a-hydroxy-5P-androstan-l7-one,3a-hydroxy-SPpregnan-20-one, and deoxycholic acid.

42 Androsterone Fig. 27. See text.

448

9 Dehydrogenases - Characreristics, Design of Reaction Conditions, and Applications

et al., 1987; KISEand HAYThe 3P-HSDH oxidizes 3P-hydroxysteroids DRUECKHAMMER 1989; RIVAet al., 1989). For instance, of the C,, and C,, series. Moreover, the en- ASHIDA, zyme also accepts 17P-hydroxysteroidsof the bicyclo[3.2.0]heptan-6-one systems were reCIS,CI9, and C,, series and certain 16P-hy- duced with HSDH with very low selectivity ( < 10% ee), when substituents in the adjacent droxysteroids. The NAD(H)-dependent 20P-HSDH from 7-position were small. The situation changed S.hydrogenans shows a pH optimum between completely when the steric requirements of 6.4 and 7 for the reduction direction and ac- the substrate were increased by additional cepts among others cortisone, cortisol, and methyl or chloro substituents. Then, HSDH progesterone as substrates.The 3a-HSDH iso- became a very specific (90-98% ee) catalyst lated from rat liver differs from the above and other enzymes (TBADH, HLADH) were mentioned enzymes in that it accepts both unable to reduce these substrates ( B u m et al., NAD(H) and NADP(H). The activity of the 1985; DAVIESet al., 1986).Another example is enzyme is equal with both coenzymes, al- 30-HSDH which was used for the manufacture of optically active alcohols from ketones, though the K , values are different. Generally, the reactions catalyzed by especially ethyl 4-~hloro-(3S)-hydroxybutaHSDHs are reversible at neutral pH. The oxi- noate (41) (Fig. 26) (KISE and HAYASHIDA, dation is favored at a basic pH, whereas the re- 1989). The above described dehydrogenases are duction is favorable at an acidic pH. The HSDHs can be immobilized on Sepha- commercially available from manufacturers rose C L 4 B (CARREA et al., 1979; RIVAet al., listed in Tab. 15. 1986) or Eupergit C (RIVAet al., 1988;DAVIES et al., 1986), are compatible with reversed micelles (ANTONINI et al., 1981;HILHORST et al., 7.2 Noncommercial 1983), and can be used in water-organic solvent two-phase systems (CREMONESI et al., Dehydrogenases, their 1975; CARREAet al., 1979, 1988; CARREA, Characteristics and Applications 1987).The most suitable two-phase system for Noncommercial dehydrogenases useful in 20P-HSDH was determined to be water-butyl acetate (CREMONESI et al., 1975). Sepharose- enzymatic synthesis are available from differimmobilized 3P-HSDH has been shown to re- ent microorganisms. The enzymes listed in tain 60% of its initial activity after 6 months of Tabs. 10 and 16 can be regarded as well-characterized with respect to their biochemical charcontinuous synthesis in water-ethylacetate. The biochemical characteristics of hydroxy- acteristics and their substrate spectra. Howevsteroid dehydrogenases allow the coupling to er, in order to make use of these catalysts, facilenzymes for cofactor regeneration (CREMONE- ities have to be in place to cultivate the microSI et al., 1975; CARREA et al., 1979,1984,1988; organisms on a reasonable scale and to isolate the respective enzymes. Therefore, the usefullHILHORST et al., 1983;RIVAet al., 1988). Due to the high regio- and stereoselectivity ness of the catalysts listed in Tab. 16 is limited and tolerance regarding different side chains to well-equipped laboratories. (BERGMEYER, 1983) on the steroid skeleton, the HSDHs have great potential in the synthesis of steroids (CREMONESI et al., 1975;RIVAet al., 1986,1988; CARREA et al., 1988), bile acids 7.2.1 Hydroxyisocaproate (TALALAY, 1962;CARREA et al., 1984),and oth- Dehydrogenases er steroid derivatives (CARREAet al., 1979; The hydroxyisocaproate dehydrogenases CARREA, 1987;HILHORST et al., 1983).However, particularly 3a-HSDH and 3 s 20P-HSDH (HicDH) (EC 1.1.1.-)catalyze the reduction of accept also nonsteroid substrates and have 2-oxocarboxylic acids to the corresponding 2been shown to be useful catalysts for the re- hydroxycarboxylic acids at the expense of duction of bulky mono- and bicyclic ketones NADH. NADPH is not accepted as coenzyme. (DEAMICIet al., 1991; BUTTet al., 1985, 1987; In contrast to the lactate dehydrogenases


449

Tab. 15. Some Manufacturers of Commercial Dehydrogenases or Reductases Company

Germany Boehringer Mannheim GmbH E. Merck KGaA Juelich Enzyme Products ASA Spezialenzyme GmbH

United Kingdom British Drug Houses Hughes and Hughes (Enzymes) Ltd. Sigma Chemical Co., Ltd. Whatman Biochemical Ltd.

USA Boehringer Mannheim Inc. PL. Biochemicals Inc. Worthington Biochemical Co. Enzymol International Inc. Diversa (formerly Recombinant Biocatalysis Inc.) Japan Amano Pharmaceutical Co., Ltd.

Address

Notes

Sandhofer Str. 116 D-68298 Mannheim, Germany Frankfurter StraBe 250, Postfach 41 19 D-64271 Darmstadt, Germany Waldhofer Stralje 102 D-69123 Heidelberg, Germany Mascheroder Weg l b D-38124 Braunschweig, Germany

research and analytical enzymes pharmaceutical and industrial enzymes enzymes, PEG-NADH, and fine biochemicals enzymes, PEG-NADH, and fine biochemicals

Poole Sorset BH12 4NN, UK Elms Industrial Estate Church Road, Harold Woods Romford, RM3 OHR,UK Norbiton Station Yard Kingston-upon-Thames Surrey KT2 7BH, UK Springfield Mill Maidstone, Kent, UK

research and analytical enzymes analytical and industrial enzymes research and analytical enzymes isolation of a range of research enzymes

9115 Hague Road; P.O. Box 50414 Indianapolis, IN 46250-0414, USA 1037 West McKinley Ave. Milwaukee, Wisconsin 53205, USA Freehold New Jersey 07728, USA 6927 Americana Parkway Columbus, Ohio 43068, USA 10665 Sorrento Valley Road San Diego, CA 92121, USA

research and analytical enzymes research and analytical enzymes research and analytical enzymes research and analytical enzymes research and industrial (thermostable) enzymes

1-21-chome, Nishiki, Naka-Ku Nagoya, Japan

research and industrial enzymes

which also catalyze similar oxidoreductions, IF0 12964 (YAMAZAKI and MAEDA,1986b), the HicDHs accept substrates with longer side whereas L-HicDH is specificfor (S)-Zhydroxychains. While LDHs optimally reduce pyru- acids and was prepared from Lactobacillus vate, the HicDHs show the highest affinitiy to- confusus (RAO and ROTH,1988; SCHUTTEet wards 2-ketoisocaproate (Km:60 pM; HUMMEL al., 1984; HUMMEL et al., 1990). The D-HicDH et al., 1985). from L. casei and the L-HicDH from L. con&As with lactate dehydrogenases, hydroxy- sus have been successfully cloned (JOHNCOLisocaproate dehydrogenases are available with LINS, GBF Braunschweig, Germany). opposite enantioselectivities but quite similar The pH optimum of the D-HicDH isolated substrate specificities. D-HicDHs are specific from the lactic acid bacterium S. faecalis was for (R)-2-hydroxyacids and were isolated from 5-7 and 8-9 for reduction and oxidation reacLactobacillus casei ssp.pseudoplanatrum DSM tions, respectively (HOSONO et al., 1990; HUM20008, L. casei ssp. rhamnosus (LEUCHTENBER-MEL et al., 1985). The enzyme is stabilized by GER et al., 1984a; WICHMANN et al., 1984;HUM- SH protecting agents like P-mercaptoethanol. MEL et al., 1985), and Streptococcus faecalis The enzyme from L. cnsei is unusually stable

35

36-40

CPCR

up to 45 5.5-7.0 up to 40 7

up to 25 7-8.5

up to 37 5.54.5

8-9 8-8.5

9-10.5

9-10

3.3-9.7 5.5-9

6 8

6-8

8.5-9.5

8-10

-

si-face re-face

re-face

re-face

si-face

re-face si-face re-face si-face si-face

J

limited

limited limited

limited limited

Compatibility with Organic Solvents

J

~

chelators, metal ions, SH blockers chelators, metal ions, SH blockers chelators, metal ions, SH blockers Hg2 no or weak inhibitions

palmitoyl-CoA EDTA

Compatibility Inhibitors with Detergents

~~

570 810

1800

220

1.6

270 360 s - ' 99

Specific Activity [U mg-' protein] Reduction Reaction

C F Curvularia falcata, MJ: Mucor javanicus, CP: Candida parapsilosis, P Pseudomonas, CB: Candida boidinii, Hic: hydroxyisocaproate, LK: Lactobacillus kejir, RE: Rhodococcus erythropolis, P L pig liver, GC: Geotrichum candidum, n.r.: not reported.

D-HicDH 50 L-HicDH 50

3 8 4

RECR

CBADH 45

25

n.r.

5-1 0 7 >7 7 4.5-8.5

-

10 n.r. 7-10 7.5-8.5 10

- 6.5

n.r.

PH PH StereoOptimum Stability specifi(oxidacity tion)

PH Optimum (reduction)

6.7 6.5-7.5 unstable - 6 at 1 6 2 5 7 99.5% ee). The propanol as co-solvent. In most cases the ex- space-time yield was 21 g L-' d - ' at a convercess 2-propanol or product acetone does not sion of 97%. interfere with product isolation. CPCR is stable under reaction conditions (2% 2-propanol) with a inactivation rate of only 0.2% d-' (ZELINSKI, 1995).

456


7.2.8 Rhodococcus erythropolis Carbonyl Reductase A novel strictly NAD(H)-dependent carbonyl reductase from Rhodococcus erythropolis DSM 743 (RECR) (EC 1.1.1.1) exhibiting a broad substrate specificity and high enantioselectivity was recently described by PETERSand coworkers (ZELINSKI et al., 1993;PETERS et al., 1992).A technical-grade enzyme (19 U mg-') can be isolated quite easily at 54% yield. The enzyme is characterized by a temperature optimum between 38 "C and 44 "C (reddction) and is stable under reaction conditions up to 37 "C between pH 6 and 8. The pH optima are 5.5-6.5 and 9-10 for reduction and oxidation reactions, respectively. RECR is inhibited by chelating agents, heavy metal ions, and sulfhydry1 blocking agents. Sulfhydryl protecting agents, e.g., 1,4-dithiothreitol do not affect the activity. In contrast to CPCR, RECR is unstable in the presence of detergents such asTriton X-100 and isopropanol. The acidic pH optimum of RECR for the reduction reaction is unfavorable for coupling to most enzymes used for coenzyme regeneration. For instance, formate dehydrogenase is active between pH 7.5 and 8.5, but is rapidly inactivated below pH 6 (SCHUTTE et al., 1976). At pH 8, where most enzymes used for coenzyme regeneration are active and stable, RECR exhibits only 40% of its maximum activity. Therefore, the substrate-coupled approach for coenzyme regeneration is to be preferred for applications of RECR. However, it has been shown that RECR and FDH can be used in multienzyme systems (PETERS et al., 1993d;ZELINSKI, 1995). RECR delivers the pro-(R) hydride of the nicotinamide ring to the re face of the carbonyl yielding (S)-hydroxy compounds (ZELINSKI et al., 1993).The enzyme follows a sequential ordered bi-bi Theorell-chance mechanism, and a hypothetic model of the substrate binding site has been deduced from kinetic measurements. The model can be used for the prediction whether or not a given carbonyl compound is accepted by the enzyme. The substrate spectrum of RECR includes 3-, 4-, and 5-oxocarboxylic acid esters, cu-substituted 3-oxoacid esters, a variety of aliphatic 2-,

3-, and 4-ketones, aromatic ketones, ketoacetales, and diketones. Ketoacids and cyclic ketones are not substrates for RECR. Aldehydes are only poorly reduced. The enzyme oxidizes primary and secondary alcohols. RECR was successfully applied for the preparation of a range of synthetically useful chiral synthons (PETERS et al., 1993d;ZELINSKI et al., 1993) like (2S, 5s)-hexanediol (50% yield, > 99% de), ethyl 4-chloro-(3S)-hydroxy-butanoate (41) (Fig. 26) (24% yield, >99% ee), ethyl (2R, 3S)-Zrnethyl 3-hy,'roxy-butanoate (35) (Fig. 24) (49% yield, 95% de), and (S)-l-(2-naphthyl)ethanol (28) (Fig. 20) (70% yield, > 99% ee).

7.2.9 Candida boidinii Alcohol Dehydrogenase A secondary alcohol dehydrogenase from Cundidu boidinii (CBADH) (EC 1.1.1.1) has been described by SCHO-ITE et al. (1982), but the enzyme was scarcely used for the reduction of carbonyl compounds. The tetrameric enzyme is NAD(H)-dependent and characterized by a temperature optimum of 45°C. It is stable at temperatures up to 40"C.The pH optima are pH 8-10 and 7 for oxidation and reduction, respectively. CBADH is stable between pH 8.5 and 9.5. The enzyme is inhibited by heavy metal ions such as HgZ+,Znz+, Cd2+,Cuz+,Co2+,and Mn2+.Reagents reacting with SH groups (iodoacetate, CN- p-chloromercuribenzoate) have a strong inhibitory effect, indicating an essential SH group(s). Chelating agents such as 2,2'-bipyridyl and 1,lO-phenanthroline inhibit CBADH strongly indicating a possible role of a metal ion in enzyme activity or stability. It is important to notice that SH protecting agents like P-mercaptoethanol strongly inhibH CBADH noncompetitively. The biochemical characteristics of CBADH allow the coupling to enzymes used for coenzyme regeneration such as formate dehydrogenase (ZELINSKI, 1995). The enzyme generally delivers the hydride to the re face of the carbonyl yielding (S)-hydroxy compounds. As with other dehydrogenases (TBADH, KEINANet al., 1987; TEADH,


457

PHAMand PHILLIPS, 1989) an interesting reverThe enzyme was used for the preparation of sal of stereochemistry is seen with smaller sub- several chiral hydroxy compounds at the gram strates where the hydride is partially delivered scale (50%-83% yield) (ZELINSKI, 1995), e.g.. and (2R, 3R)-butanediol (99% ee) and methyl also to the si face to give mixtures of (R)(S)-alcohols (SCHUTTEet al., 1982; ZELINSKI,(3s)-hydroxybutanoate (95% ee). 1995). (S)-2-butanol is converted at a velocity of 16 U mg-' (K,,,:0.42 mM), whereas (R)-2butanol is oxidized at a reaction rate of 103 U 7.2.10 Other Alcohol mg-' (K,,,:0.099 mM). Methyl 3-oxobutanoate Dehydrogenases and Carbonyl is reduced at very high enantioselectivity of >99% ee to the corresponding methyl (3s)- Reductases hydroxybutanoate. A number of other oxidoreductases have CBADH accepts a broad range of subbeen described in the literature and sometimes strates. A range of 3- and 4-oxocarboxylic acids and esters, aliphatic mono- and diketones, and they have been used for the synthesis of chiral some cyclic ketones are reduced by CBADH. hydroxy compounds. These enzymes and the Aliphatic 2-ketones are good substrates corresponding references are listed in Tab. 17. whereas 3- and 4-ketones are converted at re- However, these enzymes remain to be fully duced rates. Aromatic ketones and aldehydes characterized or developed for large-scale synare either not accepted or poor substrates of thesis. As with other reductases these enzymes the enzyme. Secondary alcohols with chain have potential for further development in orlength between C3 to C6 as well as cyclic alco- ganic synthesis. hols such as cyclobutanol, cyclopentanol, and cyclohexanol are oxidized to the corresponding ketones. Tab. 17. Other Noncommercial Oxidoreductases with Great Future Potential in Organic Synthesis Enzyme

Organism

Coenzyme

Substrate(s) Enan- Coupling Reference (reported in tioto Regenereferences) selec- rating tivity Enzymes

Aldehyde reductase (S)-Diacetyl reductase

Sporobolomyces salmonicolor Saccharomyces cerevisiae

NADP(H)

3-oxoacid esters aliphatic 2ketones, diketones

(R)-Diacetyl reductase RSADH

Saccharomyces cerevisiae Rauwolfia serpentina

Aromatic acid reductase Polyol dehydrogenase a-Acetoxy ketone reductase Alcohol dehydrogenase

Nocardia asteroides Candida utilis Saccharomyces cerevisiae Geotrichum candidum

n.r.

n.r. NADP(H)

NADP(H) NAD(H)

NADP(H)

re

J

re

si 2-oxoacid esters 10-0x0-geran- n.r. iol, acyclic monoterpenes, prim. alcohols 3-hydroxyn.r. benzoic acid sugar n.r. alcohols a-acetoxy re ketones ketoacid re esters

J

SHIMIZU et al. ( 1990) HEIDLAS et al. (1991) HEIDLAS and TRESSL (1990) HEIDLAS and TRESSL (1990) IKEDAet al. (1991)

KATOet al. (1991) BORYSENKO et al. (1989) ISHIHARA et al. ( 1994) KAWAIet al. (1994a,b) BINGFENGet al. (1995)

458


Tab. 17 Continued

Enzyme

Organism

Coenzyme

Substrate(s) Enan- Coupling Reference (reported in tioto Regenereferences) selec- rating tivity Enzymes

TEADH

Thermoanaerobium ethanolicus

NADP(H)

NTADH

Nicotiana tabacum Candida utilis

n.r.

aliphatic and re cyclic ketones, 3-, 4-ketoacid esters, aldehydes, prim./sec. alcohols n.r. cycloalkanones polyols n.r.

CUADH

NAD(H)

SpSADH

Sporobolomyces NADP( H) salmonicolor

NAADH

Nocardia asteroi- NADP(H) des (JCM 3016) Cellulomonas NADP(H) turbata KE-31 A cetobacter n.r. pasteurian us Rhizopus arrhizus NADP(H) Fischer

CTADH APADH RAADH

SSADH

Sulfolobus soyataricus

NAD(H)

3-oxocarn.r. boxylic acid esters aromatic acid n.r. menthone

n.r.

2,3-epoxyn.r. propanol cyclic ketones, n.r. aromatic double bounds, prim. alcohols, vanillyl alcohol re aliphatic ketones, sec. alcohols

PHAMand PHILLIPS (1989,1990) BRYANT and LJUNGDAHL (1981)

SUGAet al. (1986) BORYSENKO et al. (1989) SHIMIZU et al. (1990) KATOet al. (1991) LISE and HAYASHIDA (1990) GEERLOF and DUIRE (1991) FAVERO et al. (1981)

RELLAet al. (1987)

nx.: not reported.

8 Conclusions and 0utlook In the past decades, more than 1000 articles dealing with alcohol dehydrogenases have been published (see Fig. 3). The usefulness of these enzymes in organic synthesis has been reported in numerous publications, and the most attractive route for the synthesis of chiral hydroxy compounds is the direct reduction (see Sect. 4). However, despite these facts, only very few applications of alcohol dehydrogenases on the industrial scale are known until now (BOMMARIUS, 1993).There are a number of different reasons for this.

As a consequence of using isolated alcohol dehydrogenases, the coenzyme has to be regenerated effectively. For a long time, this has been regarded as prohibitive for the largescale application of alcohol dehydrogenases. Nowadays, a number of effective coenzyme recycling systems for NAD(P)+ and NAD(P)H are available that have been discussed in detail. Enzymatic cofactor regeneration systems are superior to nonenzymatic methods so far, although especially electro- and photochemical methods should not be underestimated. The formate/formate dehydrogenase system is the most convenient and useful regeneration system, which has been applied up to the large scale. Formate dehydrogenase using NAD as +

8 Conclusionsand Outlook

459

coenzyme is a commercial enzyme.This system bination with organic solvents or solubility enis also now available for the regeneration of hancers like cyclodextrins are now available. A NADPH as well, although the mutant FDH very important aspect when applying alcohol from Pseudomonas sp. remains to be commer- dehydrogenases is the consideration of kinetcialized. ics, inhibition patterns, and the reaction mechA prerequisite for an economically feasible anism.To a great extent, the kinetic limitations process using oxidoreductases is to achieve a due to substrate or product inhibition(s) govhigh total turnover number. For this purpose, a ern the choice of the appropriate reactor denumber of different immobilization strategies sign (see Sect. 6.3.3). When dealing with alcoand reactor concepts have been developed. hol dehydrogenases, the batch stirred tank reFor enzymes which bind the coenzyme very actor, which is widely used in the chemical intightly no additional methods for the retain- dustry, is not appropriate in most cases (TRAMment of the cofactor are necessary (ZACHARI- PER,1996). Knowledge of the kinetic parameou and SCOPES,1986). It is also possible to ters and the reaction mechanism allows the achieve high total turnover numbers for the modeling of reactors, which can be used to prenative coenzyme by using substrates especially dict the behavior and to scale up to the indusof high or low water solubility (KRAGLet al., trial scale. Last but not least, a variety of oxidoreduct1996a, b). In order to circumvent the need for coen- ases which are either commercially available zyme regeneration, transformations with or, if not, are well-characterized enzymes have whole cells can be used (see Chapter 8). How- been discussed with respect to biochemical ever, cells, especially bakers’ yeast, contain a characteristics, compatibility with organic solvariety of oxidoreductases with opposite enanvents and coenzyme regeneration systems, tioselectivities (CHENet al., 1984;HEIDLAS and their substrate spectra and stereoselectivities. TRESSL,1990;HEIDLAS et al., 1988,1991; NAK- Besides, applications of these oxidoreductases AMURA et a]., 1985, 1991; SHIEH et al., 1985; have been discussed where, either large-scale USHIOet al., 1986).Moreover, the accessibility continuous applications have been performed, of large amounts of biomass often restricts the or the product is of commercial interest, or a use of whole cells in chemical laboratories. For broader synthetic applicability of a chiral hythese and other reasons, the use of defined, iso- droxy compound is given. lated oxidoreductase systems for the reduction In summary it can be stated that today a vaof carbonyl compounds seems to be superior riety of alcohol dehydrogenases and carbonyl reductases are described in detail in the literato biotransformations with whole cells. Carbonyl substrates are often poorly water- ture which allow the reduction of virtually any soluble and, therefore. chemical syntheses are class of carbonyl substrate and the preparation performed predominantly in organic solvents. of both enantiomers of a hydroxy compound. On the other hand, enzymes and coenzymes Most of the alcohol dehydrogenases menare only soluble in water containing solvents. tioned in Sect. 7 have already been used in Therefore, alcohol dehydrogenases have been continuously operating reactors. The problem regarded for a long time as incompatible with of reducing poorly water-soluble substrates the use of organic solvents.The log P concept, with coenzyme depending alcohol dehydrowhich was established by LAANEand cowork- genases has been started to be solved by develers (1987) for the prediction of the compatibil- oping novel reactor concepts. ity of enzymes and organic solvents, was conBiotransformation has been successfully apfirmed by KISE and HAYASHIDA (1990) and plied to the development of cleaner producITOHet al. (1992) for NAD(P)(H)-dependent tion processes. The specificity and mild condialcohol dehydrogenases. Modern reactor con- tions of biotransformations are such that, cepts have to be implemented allowing the where biotransformations replace existing combination of poorly water-soluble sub- chemical processes, the new processes often strates and water-soluble enzymes and coen- have significantly lower environmental impact 1993). zymes. A number of interesting new reactors (JOHANN, The breakthrough for a broader application specially designed for oxidoreductases in com-

460


of alcohol dehydrogenases in industry seems to depend first on the commercial availability of a larger number of enzymes exhibiting both broad substrate spectra and high enantioselectivities. Secondly,the introduction of the above mentioned novel reactor designs into the chemical industry seems to be a prerequisite to achieve economically feasible processes with alcohol dehydrogenases.

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10 Hydroxylation and Dihydroxylation

HERBERT L. HOLLAND St. Catharines, Ontario, Canada

1 Introduction 477 1.1 Hydroxylation 477 1.1.1 History and Applications 477 1.1.2 Hydroxylase Enzymes 477 1.1.3 Regioselectivity of Hydroxylation 478 1.2 Dihydroxylation 480 1.2.1 Dioxygenase Enzymes 481 1.2.2 Epoxidation and Epoxide Hydrolases 481 1.3 Biocatalysts for Hydroxylation and Dihydroxylation 1.3.1 Whole Organisms 481 1.3.2 Isolated Enzymes 482 1.3.3 Genetically Engineered Organisms 482 2 Hydroxylation Reactions 483 2.1 Whole-Cell Biotransformations 483 2.1.1 General Aliphatic Compounds 483 2.1.2 Steroids 488 2.1.3 Terpenes 491 2.1.4 Alkaloids 495 2.1.5 Antibiotics 496 2.1.6 Dealkylation Reactions 496 2.1.7 Aromatic Ring Hydroxylation 499 2.2 Biotransformations with Isolated Enzymes 502 2.2.1 Lipoxygenases 502 2.2.2 Peroxidases 504 2.2.3 Steroid Hydroxylases 505 2.2.4 Other Enzymes 505 3 Dihydroxylation Reactions 506 3.1 Dioxygenase-Catalyzed Reactions 506 3.1.1 Arenes and Heteroarenes 506

481

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3.1.2 Dihydroxylation of Other C=C Bonds 511 3.2 Monooxygenase-Catalyzed Reactions 51 1 3.2.1 Arenes and Heteroarenes 511 3.2.2 Dihydroxylation of Other C=C Bonds 512 4 Summary 516 5 References 516

477

1 Introduction

1 Introduction

oids, heralded a decade of intensive investigation into the ability of other microorganisms to perform similar reactions on a wide range of Biotransformations involving hy droxylation steroid substrates. Concomitant with this efreactions have gained steadily in scope and fort, and continuing unabated to this day, was significance since the first successful microbial the study of the use of microbial biocatalysts steroid hydroxylations of the 1950s.The more for the hydroxylation of other groups of naturecent development of biocatalytic dihydroxy- ral products and synthetic substrates. In the lations has provided a unique tool for the former area, hydroxylation of terpenes has preparation of chiral diols from aromatic sub- provided potentially new materials for the perstrates and a valuable method for the conver- fume industry, while the hydroxylation of alkasion of alkenes to chiral vicinal diols. Subse- loids and antibiotics provides new materials quent to the last review in this series (KIES- for biological screening.The hydroxylation of LICH,1984), these reactions have been dis- synthetic organic substrates has applications in cussed in detail by HOLLAND(1992a, 1994a) the regiospecific syntheses of chiral and nonand form a significant portion of many more chiral alcohols. In addition to these applicageneral reviews of biocatalysis (JONES,1986; tions, the use of biocatalytic hydroxylation for HAWKINS, 1988;DAVIES et al., 1989;HOLLAND, study of the metabolism of pharmaceutical 1992b; FABER,1992; SERVI,1992; FABER and and other xenobiotic materials has been extenFRANSSEN, 1993;TURNER, 1994;AZERAD,1995; sively developed, and dealkylation reactions DRAUZand WALDMANN, 1995;HANSON, 1995). involving hydroxylation a to heteroatoms The present chapter will review the present have been similarly exploited. All these topics status of biocatalytic hydroxylation and dihy- are considered in detail in Sect. 2. droxylation reactions as tools for organic chemistry and to this end covers the literature to the end of 1995. 1.1.2 Hydroxylase Enzymes

1.1 Hydroxylation

1.1.1 History and Applications The now classical discovery (PETERSONet al., 1952) that Rhizopus arrhizus converts progesterone and related steroids to the corresponding 1la-hydroxy derivatives (Fig. l), intermediates in the production of corticoster-

In the absence of evidence to the contrary it is widely assumed that biological hydroxylations are performed by cytochrome P-450dependent monooxygenases (cyt. P-450), in which the one atom of oxygen from dioxygen is incorporated in the substrate and the other is incorporated into water.The number of cases where this has been unambiguously established for microbial hydroxylations is very small (HOLLAND, 1992a),and in the light of an 0

Progesterone

1 1-a-hydroxy-progesterone

Fig. 1. Conversion of progesterone to 11 a-hydroxy-progesterone by Rhizopus arrhizits (PETERSON et al., 1952).

478


increasing number of reports of hydroxylations involving other enzyme systems (see Sect. 2.2 and below) this assumption no longer commands the confidence that it once did. It is still reasonable to assume, however, that the majority of biocatalytic hydroxylations, particularly those carried out by eukaryotic microbial systems, are the result of cyt. P-450 action. The complexity of membrane-bound cyt. P-450 enzyme complexes and their general intractability with respect to isolation has ensured that most preparative biocatalytic hydroxylations are performed by whole-cell biotransformation techniques. The use of isolated cyt. P-450 enzymes for hydroxylation has been confined largely to mechanistic work, centered initially on the soluble camphor hydroxylase, cyt. P-450caM, of Pseudomonas putida (e.g., LOIDAet al., 1995 and references therein) and more recently extended to other cyt. P-450 systems (KRAINEV et al., 1993;HALPERT and HE, 1993;BLACK et al., 1994;IWASAKI et al., 1995a; ALTERMAN et al., 1995; MODIet al., 1995;IWASAKI et al., 1995b),or to screening of crude extracts for their biotransformation capabilities (TROWER et al., 1988). Other enzymes that have been reported to carry out hydroxylations include methane and other alkane monooxygenases (KIENERand ZIMMERMANN, 1992;LIUet al., 1993), naphthalene dioxygenase (GIBSON et al., 1995),monophenol oxygenase (HSIUNG et al., 1992),horseradish peroxidase (OKADA et al., 1988;AKASAKA et al., 1995), soybean lipoxygenase (LUND et al., 1992), chloroperoxidase (MILLERet al., 1995;ZAKS and DODDS,1995),and cyclooxygenase (ZHANG et al., 1992a).These will be considered in detail in Sect. 2.2. Enzyme commission nomenclature distinguishes two major subclasses of oxidoreductases in which dioxygen is incorporated into the substrate (EC 1.13 and 1.14). In 1984, all subsubclasses from 1-10 for both of these subclasses were abandoned, and hence the first extant subsubclass in each case is 11. In subclass 1.13, no external hydrogen donor is required for the oxygenation reaction. There are three subsubclasses:in 1.13.11 and 1.13.12 two atoms or one atom of oxygen are incorporated into the substrate, respectively. Subsubclass 1.13.99 is for enzymes which have not been fully characterized. Many enzymes in subclass

1.13 effect cleavage of aromatic rings and comparable reactions and hence fall outside the ambit of this review, however one relevant example is lipoxygenase (EC 1.13.11.12). Enzymes in subclass 1.14 act on two hydrogen donors with incorporation of one or two oxygen atoms from dioxygen into one or both donors. The subsubclasses are distinguished by the type of hydrogen donor and/or the number of oxygen atoms incorporated into the substrate. In subsubclass 1.14.11,2-oxoglutarate is oxidized to succinate and carbon dioxide and the other oxygen atom is incorporated into the substrate (e.g., thymine 7-hydroxylase EC 1.14.11.6). E C 1.14.12 covers enzymes which use NADH or NADPH as one donor and incorporate two atoms of oxygen into the other donor (e.g., benzene dioxygenase EC 1.14.12.3).At present all higher numbered subsubclasses (except the miscellaneous, 1.14.99) cover reactions in which only one atom of oxygen is incorporated into one hydrogen donor. Thus they may be distinguished solely on the basis of the second donor: for example, EC 1.14.13, NADH or NADPH (e.g., phenol hydroxylase EC 1.14.13.7); EC 1.14.14, flavoproteins; E C 1.14.15, iron-sulfur proteins, including heme-thiolate proteins such as the cyt. P-450s (e.g., alkane 1-monooxygenase, EC 1.14.15.3); EC 1.14.16, reduced pteridines; and EC 1.14.17, ascorbate. Many important hydroxylases and dihydroxylases (including some cyt. P-450s) fall into the miscellaneous subsubclass EC 1.14.99,which reflects the lack of knowledge about these enzymes (Anonymous, 1984).

1.1.3 Regioselectivity of Hydroxylation A problem inherent in whole-cell biotransformation, particularly when seeking a specific site or stereochemistry of reaction in a defined substrate, is the inability to reliably predict the regio- and stereochemistries of oxidation of a given substrate by a new organism, or even of a new substrate by a microorganism whose oxidative biotransformations of different molecules have previously been examined. The first problem is the more difficult, and has been ad-

1 Introduction

479

dressed by screening of microorganisms likely tries of the latter are determined for several to perform the desired transformation and fungal varieties by the position of existing oxysubsequent development of empirical rela- gen substituents in the substrate through tionships between the mode of hydroxylation which binding to the enzyme can occur, hyand the phylogenetic position of individual droxylation occurring at a fixed distance, usumicrobial strains (AZERAD,1993; ABRAHAM, ally 6-88,, away from substrate carbonyl 1994). The second problem is more amenable groups (JONES,1973). Interpretation of the reto analysis and several models, specific to lationship between substrate binding centers single microorganisms and defined groups of and the site of hydroxylation of steroids is substrates, have been developed to predict the complicated by the fact that binding can occur outcome of microbial hydroxylation reactions. in several different orientations, a phenomeThese have been recently reviewed by HOL- non first suggested for pragmatic reasons by BRANNON et al. (1967) and later confirmed by LAND (1995a). Hydroxylations of amides by Beauveria sul- site-directed mutagenesis studies (IWASAKI et furescens ATCC 7159 (also known as Sporo- al., 1995b). trichum sulfurescens and Beauveria bassiana) The regio- and stereochemistries of the hyoccur in nonpolar residues located predomi- droxylation of terpenes by Streptomyces purnantly 4-7 8,away from a polar binding site as purescens (LIUet al., 1988) and of alkaloids by described in the model of Fig. 2 (ARCHELAS et Streptomyces griseus (SARIASLANI and ROsAzal., 1984;FOURNERONet al., 1987,1989a).based ZA,1984) have also been rationalized by modon an earlier version proposed by JOHNSON et els involving a defined spatial relationship between substrate binding and hydroxylation al. (1968). In view of the importance of the substrate sites,but little similarity exists between the two class, steroid hydroxylations have been exten- models. One of the more frequently reported sites of sively studied and may be classified into two groups: those hydroxylationsoccurring at posi- hydroxylations is that involving attack at bentions a or vinylogous to carbonyl groups in zylic or allylic carbons. Such positions are actiwhich the regio- and stereochemistry of reac- vated towards radical abstraction, thought to tion is controlled by mechanistic factors dictat- be the primary but not exclusive mode of aced by the stereoelectronics of electrophilic ad- tion of cyt. P-450 oxygenases (BACHet al., et al., 1995), and in the case of dition to enols (HOLLAND, 1984; HOLLAND et 1995;NEWCOMB al., 1989); and those occurring at unactivated benzylic centers can also be activated by eleccarbon atoms. The regio- and stereochemis- tron abstraction from the aromatic ring. This latter phenomenon is involved in the mode of action of lignolytic fungi (FIELDet al., 1993), and is thought to be responsible for the predominance of benzylic hydroxylation in biotransformations of aromatic hydrocarbons by Mortierella isabellina (HOLLAND et al., 1988). A model for the latter process has been developed (Fig. 3), which is based on analysis of the benzylic hydroxylation of over 40 substrates (HOLLAND et al., 1993). Although the various models for hydroxylation reactions by single microorganisms discussed above do not show any close similar[C] = cyclic structure ities, some common features do emerge. All in[B] = binding site for amide oxygen volve a three-point interaction of hydroxyla[O]= site of hydroxylation tion and binding sites, with optimal distances Fig. 2. Stereochemical model for hydroxylationby (where specified) of 4-7 A between binding Beauveria sulfurescens ATCC 7159 (ARCHELAS et and hydroxylation positions. There is a recurring suggestion that efficient substrate binding et al., 1987,1989a). al., 1984;FOURNERON

480


A: aromatic binding pocket B: aliphatic binding region [ O ] : oxidizing site dimensions in Angstrom

8 8

A

8 8 8

4

.----4 : -0

6.5

)

) 0 9 9

7

.1.5 I

3

Fig. 3. Model for the benzylic hydroxylase of Mortierella isabellina (HOLLAND et al., 1993).

requires both polar and nonpolar interactions 1.2 Dihydroxylation with the enzyme, and the existence of multiple binding modes for a single substrate is also Microbial dihydroxylation may be defined common. The larger question of whether these as the conversion of a carbon-carbon double factors are common to a wider range of micro- bond into a vicinal diol. This process, although bial systems and isolated hydroxylases re- known since the early days of steroid bioconmains to be established, although similar data versions in the 1950s, has only been developed for a range of cyt. P-450enzymes give cause for as a useful biotransformation in the last optimism in this regard (STAYTON and SLIGAR, decade. It is broadly divided into two groups 1990; HARRISand LOEW,1995; LOIDAet al., (Fig. 4): the direct conversion of arenes or hetet al., 1995b). eroarenes into cis-dihydrodiols, a reaction per1995;MODIet al., 1995;IWASAKI

Dioxygenase

Dehydrogenase *

Rearrangement

Hydrolysis

m

Monooxygenase -t

w 0

Dioxygenase Fig. 4. Routes for dihydroxylation.

Hydrolysis

a:: a:: t

OH

I Introduction formed by dioxygenase enzymes; and the biotransformation of isolated alkenes by a process involving epoxidation followed by enzymic or nonenzymic hydration to generate the vicinal diol. However, this classification is qualified by the fact that monooxygenase activity can. through an intermediary arene oxide, lead to the formation of trans-dihydroarenediols, and dioxygenases can also transform non-aromatic alkene substrates.

481

alkene epoxidation can result in the isolation either of the epoxide or of the corresponding vicinal diol. Among the factors that influence the stability of an epoxide in a biological system are the pH of the medium and the presence or absence of epoxide hydrolase enzymes. Under acidic conditions, non-enzymic epoxide hydrolysis can become the dominant mode of reaction, whereas under neutral or basic conditions the activity of an epoxide hydrolase can be instrumental in determining the nature of the final product. Biotransformation 1.2.1 Dioxygenase Enzymes of epoxides by isolated epoxide hydrolases is beyond the scope of this chapter, but the role The use of arene dioxygenases for the pro- of in situ microbial epoxide hydrolases in the duction of crs-dihydroarenediols has devel- conversion of alkenes to vicinal diols will be et considered in Sect. 3.2.2. oped from the pioneering work of GIBSON al. (1968) to the stage that over 150 such comet al., 1995). pounds are now known (STABILE The arene dioxygenase enzyme complex, like the cyt. P-450 monooxygenase system. is not 1.3 Biocatalysts for Hydroxylation amenable to simple isolation and use in a pur- and Dihydroxylation ified form. Conventional mechanistic work Relatively few useful hydroxylations or diwith arene dioxygenases has been limited, but the requirement for molecular oxygen has hydroxylations are performed by isolated enbeen established and in one example, trypto- zyme preparations, but in these cases the naphan 2,3-dioxygenase (EC 1.13.11.11), forma- ture of the catalytic agent is generally well tion of a dioxetane intermediate has been pro- understood (see Sect. 1.3.2). Arene dihydroxylation is performed by et al., 1993). posed (LEEDS Accumulation of czs-dihydrodiol metab- enzymes whose natural function is the activaolites normally requires an organism in which tion of a hydrocarbon substrate for subsequent the subsequent enzyme in the arene degrada- biodegradation reactions. However, in spite of tion pathway of the wild-type strain, a cis-dihy- our knowledge of the properties and biological droarenediol dehydrogenase, is suppressed or functions of many oxygenase enzymes, microabsent, and several mutant organisms have bial hydroxylations and epoxidations are genbeen obtained which fulfill this criterion. An erally carried out by enzymes whose natural alternative approach is the expression of an function is unknown or, at best, uncertain. It is arene dioxygenase in a host organism that thought that, in parallel to the situation found does not contain genes for the remainder of in mammalian biochemistry, many such oxidathe arene degradation pathway, and these al- tions are an expression of a general detoxification process, but there is little direct evidence ternatives are examined in Sect. 1.3. to support this assumption. In other cases, the hydroxylation reaction is known to be the first 1.2.2 Epoxidation and Epoxide step in an oxidative degradation process that involves using the substrate as an energy Hydrolases source for primary metabolism. Epoxidation of carbon-carbon double bonds in aromatic or non-aromatic systems can be performed, inter alia, by cyt. P-450monooxy- 1.3.1 Whole Organisms genases (HOLLAND, 1992a). Epoxidation of arenes can result in the formation of phenolic In the majority of hydroxylation and dihyor trans-arenedihydrodiol products, whereas droxylation biotransformations, the nature of

482


Of the other enzymes capable of hydroxylathe oxygenase enzymes concerned, coupled with their extensive cofactor requirements, tion reactions, perhaps the best understood is means that employment of whole-cell catalysts the non-haem iron protein methane monois necessary. The fact that a single enzyme or oxygenase (EC 1.14.13.25) from Mefhylococrange of isozymes from among the thousands cus capsulatus. This enzyme, which requires present in a bacterial or fungal cell can oxidize NADH and hydroxylates a range of small alet al., 1991; LIUet al., 1993) an organic compound in a selective manner is kanes (DEIGHTON a remarkable but fortunate event in that it pro- has been extensively characterized (ROSEN1994). Other non-haem vides the chemist with a wide range of tools for ZWEIG and LIPPARD, iron oxygenases are less well characterized, carrying out hydroxylation reactions. The range of organisms known to carry out but those from Pseudomonas oleovorans also hydroxylation reactions is vast: cyt. P-450 perform hydroxylation reactions (CASPIet al., monooxygenases are among the most wide- 1981;WUBBOLTS et al., 1995). Recently, a gene spread enzymes in nature, being found in all or- from this species for alkane R-hydroxylase ganisms from bacteria to humans. For practical (EC 1.14.15.3)has been expressed in E. coli to purposes, however, hydroxylation reactions are give catalytically active, soluble protein vesimost often carried out by bacterial or fungal cles (SHANKLIN et al., 1997). cultures. The techniques involved have been Fungal monophenol oxygenase, used in the described elsewhere in detail (HOLLAND, form of a crude preparation, required Fe3' for 1992b; LEAK,1994), and sources of useful mi- maximum efficiency (HSIUNGet al., 1992), crobial cultures have been tabulated recently while hydroxylation by mammalian cyclooxy(HOLLAND,1992a; LEAK, 1994; DRAUZand genase appears to be specific for products of WALDMANN, 1995). Dihydroxylations that in- the arachidonic acid cascade (ZHANGet al., volve an intermediate epoxidation are carried 1992a). The abilities of naphthalene dioxygeout in a manner analogous to microbial hy- nase (GIBSON et al., 1995) and other dioxygedroxylations,but arene cis-dihydroxylations of- nases (TAKIKAWA et al., 1983) to perform ten require more exacting biotransformation monohydroxylations is well known, and apconditions, and relatively few organisms useful pears to be a function of the nature of the for the latter biotransformation are available dioxygenase active-site iron-oxo complex. The from culture collections at the present time. identity of the latter species is, however, unknown at the present time and mechanistic details for the arene dioxygenases are obscure. Dihydroxylation of alkenes via epoxidation 1.3.2 Isolated Enzymes by isolated monooxygenases has been studied Of the isolated enzymes reported to carry but is severely limited in scale (HOLLAND, out hydroxylations only horseradish peroxi- 1992a),while dihydroxylation by isolated arene dase, chloroperoxidase, and soybean lipoxyge- dioxygenases has been carried out mainly for nase are commercially available at the present mechanistic purposes (RESNICK et al., 1994). time. Horseradish peroxidase is restricted to the low-yield formation of phenol in nonaqueous solvent (AKASAKA et al., 1995) and to the 1.3.3 Genetically Engineered benzylic hydroxylation of highly activated aro- Organisms matic substrates (OKADAet al., 1988); chloroperoxidase from Caldariomyces fumago also Expression of mammalian hydroxylase enperforms benzylic hydroxylations (MILLER et zymes in microbial systems has been develal., 1995; ZAKSand DODDS,1995). Soybean oped as a means to facilitate study of these enlipoxygenase is implicated only indirectly in zymes. Steroid hydroxylations, for example, the hydroxylation of cholesterol at C-7 carried have been performed following the expression out in the presence of linoleic acid (LUNDet of mammalian steroid hydroxylases in microal., 1992). Other processes involving lipoxyge- organisms and these systems have been used nases and peroxidases are considered in Sects. to examine the binding requirements for hydroxylations at C-2 and C-7 (IWASAKI 2.2.1 and 2.2.2, respectively. et al.,

2 Hydroxylation Reactions

483


1995b).Human cyt. P-450 3A4, which catalyzes hydroxylations of progesterone at C-6p and C-16a, has been expressed in yeast (MEH2.1 Whole-Cell Biotransformations MOOD et al., 1995), while C-17a and C-21 hydroxylases have been expressed in Kluyveromyces lactis (SLIJKHUIS et al., 1990). Vitamin D 2.1.1 General Aliphatic Compounds derivatives have been hydroxylated by rat kidSome of the simplest substrates amenable to ney cyt. P-450 expressed in E.coli (AKYOSHI et al., 1994), metabolism of the hypoglycemic tol- biocatalytic hydroxylation are the fatty acids, butamide has been studied using human liver and formation of chiral lactones from saturatcyt. P-450s expressed in yeast (BRIANet al., ed fatty acids has been reported using a Mucor 1989), and the expression of site-directed species capable of converting dodecanoic acid mutants of mouse coumarin 7-hydroxylase in into 11-hydroxy-y-dodecalactone (MEYERand yeast has been used to examine substrate bind- LADNER,1989). Unsaturated fatty acids can ing requirements of cyt. P-450 systems (IWASA- also be hydroxylated: a Pseudomonas species converts oleic acid into (R,R)-7,10-dihydroxyKI et al., 1995a). A second application for genetic manipula- 8-(E)-octadecenoic acid (Hou and BAGBY, tion involving hydroxylase enzymes is the ex- 1990; KNOTHEet al., 1992), and the fungus pression of a microbial hydroxylase in a micro- Gaeumannomyces graminis has been reported bial host lacking an enzyme capable of react- to carry out hydroxylation of several unsatuand OLIW, ing with the desired product of hydroxylation. rated fatty acids at C-8 (BRODOWSKY E.coli lacking alcohol dehydrogenase activity 1992). The conversion of oleic acid into 10has been used as host for monoxygenases from hydroxystearic acid by hydration of the olefinPseudomonas putida, and in this way efficient ic bond is not a hydroxylation per se, but is benzylic hydroxylations of methylpyridines efficiently carried out by Sphingobacterium et al., 1994), (ZIMMERMANN et al., 1992) and terminal hy- NRRL B-14797 (KANESHIRO et al., 1994), Flavodroxylations of ethyl groups in heterocyclic Pseudomonas (FARBOOD 1995), and other unaromatic substrates (KIENERand ZIMMER-bacterium (KAWASHIMA, identified bacteria (GOCHOet al., 1995). MANN, 1992) have been achieved. The production of chiral p-hydroxyacids The genetics revolution has had its greatest impact on oxidative biotransformations in the such as p-hydroxyisobutyric acid by hydroxyarea of arene dioxygenase-catalyzed reactions. lation of isobutyric acid is well known. (S)The applications are twofold, namely the use ( + )-p-hydroxy-isobutyric acid is produced of mutant organisms that allow for accumula- commercially (48% yield) in Japan using f! 1992). Many tion of the desired cis-dihydrodiol products, putida (PANSEand DESHPANDE, and the expression of arene dioxygenases in other organisms are also capable of this oxidamicrobial hosts that do not possess enzymes tion, e.g., immobilized Bullera alba I F 0 1030 capable of product degradation. The former which gives higher yields than using free cells method was used in the initial application of (KANDAet al., 1993).Alternatively, (R)-(-)-pdioxygenases for arene cis-dihydroxylation, hydroxy-isobutyric acid is accessible by hydeveloped mainly through the use of mutant droxylation of isobutyric acid using Candida 1992a). strains such as GIBSON’S f! putida 39/D (GIB- rugosa (LEE et al., 1996;HOLLAND, Hydroxylations of amino acids are not SON et al., 1970) and DALTON’S F! putida UV4 (BOYDet al., 1995a), and still finds extensive common, but the conversions of L-proline to application (e.g., STABILE et al., 1995). The ex- cis-4-hydroxy-~-prolineby Helicoceras oryzae and SERIZAWA, 1993) and of Lpression of dioxygenase activity in a non- (MATSUOKA arene-degrading organism such as E. coli is a aspartic acid to L-threo-hydroxyaspartic acid more recent development, but such systems by Papularia arundinis (NISHIDEet al., 1995) have now been used for production of arene have been reported. cis-dihydrodiols by WUBBOLTSand TIMMIS Modification of bioactive compounds pro(1990),Bou~et al. (1994), GIBSON et al. (1995), vides a continued rationale for microbial hydroxylations. Recent examples include the and WHITEDet al. (1994).

484


hydroxylation of the immunosuppressant an- hibitors of squalene synthase (MIDDLETON tagonist FR-900520 at C-21 by Streptomyces et al., 1995). hygroscopicus ATCC 55 166 (TREIBERet al., 1992) and hydroxylations of the avermectin derivatives (1) at C-27 by Nocardia autotrophica (CHARTRAIN et al., 1990) and (2) at C-28 by Saccharopolyspora erythrea (Fig. 5) (ARISONet al., 1992). Studies involving cholesterol synthesis inhibitors have shown that the HMG-CoA reductase inhibitors (4) are produced from lovastatin analogs (3) by hydroxylation using Nocardia MA6455 (WILLIAMSON et al., 1989a) or Actinomycetes species (MARCIN et al., 1991), while hydroxylation of 14C-5 by Mucor hiemalis has been used to prepare 7 Squalestatin S 1, Zaragozic acid A labeled /3-hydroxycompactin ( 6 ) (WALLACE et al., 1993). The squalene synthase inhibitor, squalestatin (7) (Fig. 6), was hydroxylated in low yield by Actinomycetes at the positions indicated to give products that were also in23

8R 9R OH

= H; deoxypodophyllotoxin = OH; epipodophyllotoxin

J

1 R = glycone, C22C23 unsaturated 2 R = H, C2z-C23 saturated R

3 R = CH3 4 R = CHzOH 5R=H 6R=OH Fig. 5. Averrnectin and cornpactinanalogs.

Fig. 6. Microbial hydroxylation of bioactive compounds (MIDDLETON et al., 1995;KONDOet al., 1989; CHENand Doss, 1991,1992). t hydroxylation positions


485

Other recent examples of the microbial hy- press rat kidney cyt. P-450,24(AKYOSHI et al., droxylation of bioactive compounds include 1994). Stereospecific hydroxylation of a series the quantitative conversion of deoxypodo- of 15-deoxyprostanoids (17) at C-18 or C-19 phyllotoxin (8) to epipodophyllotoxin (9) (Fig. by Rhizopus arrhizus ATCC 11145, Calonec6) by a Penicillium species (KONDO et al., tria decora ATCC 14767, or Aspergillus ochra1989), the modification of angiotensin I1 re- ceus 4TCC 18500 serves also as a model for ceptor antagonists (10) by hydroxylations as the dosolute stereochemistry of mammalian et al., indicated using Streptomyces species (CHEN metabolism of prostanoids (HOLLAND and Doss, 1991,1992), and the preparation of 1990a).The well-established use of fungal biothe antidepressant pyrimidine derivatives (12) transformation to produce mammalian drug and (13) by hydroxylations of compound (11) metabolites is also exemplified by the benzylic using Cunninghamella echinulata I F 0 4443 hydroxylation of triprolidine (18) at the aryl (Fig. 7) (NAKAGAWA et al., 1990a) or Strepro- methyl group in 55% yield by Cunninghamella myces lavendulae SANK 64687 (NAKAGAWAelegans ATCC 9245 (HANSEN et al., 1988). et al., 1990b), respectively. The estrogenic fungal metabolite zearalenone (14) is converted to the (8’s)-hydroxy derivative (15) in 32% 24 yield by Streptomyces rimosus (EL-SHARKAWY and ABUL-HAJJ, 1988). Hydroxylation of vitamin D3 (16, Fig. 8) at C-25 by Amycolata autotrophica (TAKEDA et al., 1994) or A . saturnea (OMURA et al., 1990) mimics the mammalian metabolism of this substrate, while hydroxylation at C-24 can be carried out using E. coli transformed to ex~1 n

R2

HOP’’

16 Vitamin D3

NQ

12 R’ = OH, R2 = H 13 R 1 = H , R2 =OH

rac-17 R = H; ll,lS-dideoxy-PGEl R = OH; 15-deoxy-PGEl

0

14 R = H;Zearalenone

15 R = OH;(8’3-hydroxyzearalenone Fig. 7. Microbial hydroxylation of bioactive comet al., 1990a. b; EL-SHARKAWYFig. 8. Hydroxylation that mimics mammalian mepounds (NAKAGAWA and ABUL-HAJJ, 1988). tabolism.

486


Microbial hydroxylation as a means for the preparation of chiral alcohols continues to be investigated. Rhizopus arrhizus ATCC 11145 hydroxylates a series of bicyclic compounds (19 and 20) (Fig. 9) at the designated allylic positions with high yield and stereoselectivity (OUAZZANI et al., 1991; ARSENIYADIS et al., 1991,1994), and several fungi are reported to carry out efficient diastereoselective allylic hydroxylation of the enantiomers of bicyclic ketone (21) (HAMMOUMI et al., 1993). One of these (Mucorplumbeus CBS 110-16) was later used for the preparation of compound (R)-22, a chiral building block for terpene synthesis, by allylic hydroxylation of the corresponding mono-alcohol (ARANDAet al., 1995). Chiral terpene synthons have also been produced by hydroxylation of 2,2-dimethylcyclohexanone by Pseudomonas purida to give the (S)-alcohol (23) (YAMAMOTO et al., 1990) and by hydroxylation of a series of saturated and unsaturated bicyclo[2.2.l]heptanes (24) to give either ex0 or endo alcohols (YAMAZAKI and MAEDA, 1985a,b, 1986).4-(R)-Hydroxychroman (25) is produced from chroman in low yield but high enantiomeric excess (ee) ( 298%) by hydroxylation using Mortierella isabellina ATCC 42 613 (HOLLAND et al., 1991a).

rac- 19

22

One of the classic hydroxylating microorganisms, Beauveria sulfurescensATCC 7159, continues to be investigated for its ability to hydroxylate new substrates. In addition to the studies leading to the active site model of Fig. 2, this fungus has recently been used for the hydroxylation of amides of type (26, Fig. 10) (ARCHELAS et al., 1986),p-lactams (ARCHELAS et a]., 1988a), carbarnates (VIGNEet al., 1991), the activation of benzamide-substituted adamantanes (27) (JOHNSON et al., 1992a) and other cyclic hydrocarbons (JOHNSON et al., 1992b), and the hydroxylation of a series of N-arylpiperidines such as (28) (FLOYDet al., 1993; PARSHIKOV et al., 1993).In the latter series, the designated transformation of structure (28) is notable for its inclusion in a handbook of preparative biotransformations (FLOYDet al., 1995). B. sulfurescens was also investigated for its ability to hydroxylate cyclohexylcyclohexane (29), but the most efficient organism for production of the diequatorial 4,4’-diol (30) proved to be a Cunninghamella species (DAVIES et al., 1986). Hydroxylations of unsubstituted saturated hydrocarbons are not frequently reported. More often, as in the case of hydroxylation of the 1- and 4-diamantanols

21

20

23

24

Fig. 9. The preparation of chiral alcohols as synthetic intermediates. t hydroxylation positions

25

t


26n=1 R -3 / G R 29R=H 3 OR=OH

487

in the absence of any substituent in the substrate. Substrates such as toluene, ethylbenzene, indan, and tetralin undergo benzylic hydroxylation by the fungus Mortierella isabellina ATCC 42613, and this process has been the subject of intensive study by HOLLAND et al. (l985,1987,1988,1990b, 1990c, 1991b),culminating in the proposal of the model shown in Fig. 3 (HOLLAND et al., 1993).The selectivity of the benzylic hydroxylation carried out by M.isabellina has also been studied using compound (35,Fig. 11) (BACIOCCHI et al., 1995) and the results are consistent with the model of Fig. 3. Benzylic hydroxylation of tetralin and related compounds (SIKKEMA and DE BONT, 1991) and of methylnaphthalenes (OSUMI and

$jR 9

3 1 R = H,OH

28

B 36

35

2

3 2 R = NHCOCH3 or CH20H

f 33

-c3, z

,8^a

R \

37 R = "Pr, 'Pr,"Bu, 'Bu

34

t hydroxylation positions

(31)by Rhizopus species (predominantly at the C-7 and C-9 positions, respectively), the substrate contains an oxygen substituent (LIPAVSKA et al., 1982).This is exemplified by the hydroxylations of substituted adamantanes (32and 33) and of cyclododecanone (34) by Cephalosporium aphidicola (HANSON and PARVEZ, 1995;FAROOQ and HANSON. 1995). In contrast, hydroxylation of aromatic hydrocarbons often occurs at a benzylic position

38

QJotco

0

Fig.10. Alicyclic hydroxylations.

koH

RO

39

40

n = 1,2 41 R = H

43

42 R = O H

Fig. 11. Benzylic hydroxylations.

488


TAKESHITA, 1990a) can also be catalyzed by to give (R)-benzylic alcohols in high ee (BOYD bacteria. M. isabellina also hydroxylates a va- et al., 1991a), and converts dihydrobenzofuran riety of substituted aromatic (HOLLAND et al., to the (S)-alcohol(43) (BOYDet al., 1993a). 1987,1990a) and heteroaromatic (AGARWAL et al., 1994) substrates at the benzylic position; Aspergillus niger and Cunninghamella elegans 2.1.2 Steroids perform benzylic hydroxylations of several The microbial hydroxylation of steroids substituted 1,2,3,4-tetrahydroquinolinessuch as (36) (CRABBet al., 1994), but other organ- continues to receive much attention, and has isms, such as Helminthosporium sp. NRRL been reviewed periodically by MAHATOand 4671, perform only terminal and subterminal MAJUMDAR(1993). A recent review on the mihydroxylation of the alkyl group of substituted x-obial transformation of 5a-steroids is also aromatic compounds such as (37) (HOLLAND ,ancerned, inter alia, with hydroxylations (VOISHVILLO et al., 1994a). The present comet al., 1994a). Benzylic hydroxylation of dialkylpyridines mentary will focus on new techniques for mihad been reported as a method for the prepar- crobial steroid hydroxylation, the discovery of ation of the alcohols (38) and (39), using Beau- new microorganisms for this process, its appliveria bassiana and Curvularia species, respec- cation to new substrate groups, and mechanistively (EGOROV et al., 1985; ZEFIROV et al., tic studies. Immobilization of biocatalyst has been em1993).Other microorganisms,particularly Pseudomonas and Rhodococcus species, convert ployed in attempts to improve productivity of aryl methyl groups directly to carboxylic acids, steroid hydroxylations. Spores of Aspergillus presumably via hydroxylation.This latter reac- ochraceus (BIHARIet al., 1984) or Rhizopus tion has been developed as an efficient meth- nigricans (VIDYARTHI and NAGAR,1994) enod for the production of a variety of hetero- trapped in polyacrylamide gel can be used rearomatic and aromatic carboxylic acids from peatedly for the 1la-hydroxylation of progesthe corresponding methyl-substituted com- terone (see Fig. 1) while immobilized Curvupounds (KIENER, 1992a, b: HOEKS,1992; KIE- laria lunata (MAZUMDER et al., 1985) or CochNER and GLOECKER, 1993; WATANABEet al., liobolus lunatus (UNDISZet al., 1991) can be 1994; UEHARA et al., 1993; TAKEUCHI et al., similarly employed for llp-hydroxylation. The 1993; ISHIKAWAet al., 1995: TAKAHIRO et al., 14a-hydroxylation of progesterone is per1995). Oxidation of methyl to carboxylic acid formed efficiently by M. isabellina immobilhas also been employed in the production ized in calcium alginate beads (HOLLAND et of chiral acids such as ((R)-40) by oxidation al., 1992). of the corresponding isopropyl ether using The use of inducers to improve the yield of Rhodococcus rhodochrous (CLIFFORD et al., llp-hydroxylation by Cochliobolus lunatus 1989). has been studied (GROHet al., 1991;HORHOLD A series of benzylic hydroxylations has been et al., 1992; UNDISZ et al., 1992) and an analoreported in which the dioxygenase activity of a gous biotransformation has been improved by Pseudomonas species is implicated. This was SEDLACZEK et al. (1993) by the application of first reported by WACKEITet al. (1988) for the pressure to the fermentation vessel. This latter conversion of indan to 1-(R)-indanol by to- technique has also been applied to the 15aluene dioxygenase and to 1-(S)-indanol by hydroxylation reaction performed by Penicilnaphthalene dioxygenase; these enzymes also lium stoloniferum (PETZOLDT, 1985).The llpcarry out benzylic hydroxylations of indene hydroxylation of cortexolone by Curvularia and of 1- and 2-indanone (RESNICK et al., lunata has been subjected to mathematical 1994). Indene and its homologs (41) are con- analysis to optimize substrate loading (CHEN verted to the corresponding (R)-benzylic alco- and WEY,1990),and the same reaction carried hols by P putida UV4 in high enantiomeric out by Cochliobolus lunatus has been optipurity (BOYDet al., 1989, 1990; AGARWAL et mized by continuous adjustment of the bioal., 1990). The same species also hydroxylates transformation medium composition (SCHULTthe corresponding saturated benzocarbocycles ZE et al., 1992).


489

Hydroxylations of steroids by plant cell cul- 12) (TURUTAet al., 1991). Similarly, llp-hytures have been recently studied using Digita- droxylation of the steroid (45) by Cunninghalis lanata (PARKet al., 1995),Marchantia poly- mella elegans can be optimized by selection of morpha (HAMADA et al., 1989), and a variety the appropriate ester group at C-21, maximum of unicellular algae (FIORENTINO et al., 1991), yields (75%) being obtained using the monobut have not reached the efficiency of analo- succinate (USZYCKA-HORAWA et al., 1990). gous microbial processes. Useful new microbiHydroxylations at A-nor, A-homo (CRABB al hydroxylations include an efficient 1la-hy- and RATCLIFFE,1986), and D-homosteroids droxylation by Absidia repens (TRUCKEN- (CAMPBELL et al., 1986) by Cunninghamella BRODT et al., 1991), the 9a-hydroxylation of elegans have been examined and found to A5-3p-hydroxysteroidsusing Circinella species occur principally at C-7 and C-9a in a pattern (VOISHVILLO et al., 1994b, c) and the 14a- not dissimilar from that observed for convenhydroxylation of various steroids of both the tional steroids. Hydroxylations of 4- and 17androstane and pregnane series (MADYASTHAazasteroids also proceed in a conventional and SRIVATSAN, 1987; MAHATOet al., 1989; manner, finasteride (46) and related comNAKAKOSHI et al., 1993; MADYASHTA and pounds being hydroxylated at C-11a by SeleJOSEPH,1993; ASADA,1994; YOSHIOKA et al., nastrum capricornittum (ARISONet al., 1993a, 1994a,b; Hu et al., 1995). b) and the oxazoline (47) being hydroxylated Testosterone and progesterone (see Fig. 1) at C - l l p by Curvularia lunata NRRL 2380 remain favorite substrates for the investigation (Ass], 1989). Hydroxylations of brassinosterof new steroid hydroxylations. The former is oids (48) (ADAMet al., 1 9 9 1 ;Vo lc ~ eal., t 1993) hydroxylated predominantly at C-15a by and of a withaferin A derivative (FUSKA et al., Botryosphaerica obtusa (SMITHet al., 1990) 1990) by Cunninghamella echinulata occur at and at 6/3 and 7 a by Phycomyces blakesleeanus c-12p. (SMITHet al., 1989a).Hydroxylation of progesHydroxylations of estrogens have not been terone occurs at C-8 using Corynespora melo- frequently reported, but the ether (49) is hynis CBS 16260 (KRISCHENOWSKI and KIES- droxylated at C-6p by Aspergillus alliaceus LICH,1993), C-6p and C - l l a using Cephalo- (Fig. 13) (WILLIAMSON et al., 1989b) and at sporium aphidicola (FAROOQ et al., 1994), C-16a by Streptomyces roseochromogenes (FERC-16a using Sepedonium ampullosporium RER et al., 1990).A similar paucity of bile acid (SMITHet al., 1989b), C-7a using Botryos- hydroxylations exists, the only recent reports phaerica obtusa (SMITHet al., 1989c), C-6p, being hydroxylation of lithocholic acid (50) at -9a, -14a, and -15 using Apiocrea chrysosper- C-lSp by Cunninghamella echinulata (YAKULT ma (SMITHet al., 1988) and C-7p using Acre- HONSHA, 1985) and at C-7p by Fusarium equiet al., 1989). monium strictum (YOSHIHAMA seti (XIONGand FA, 1995).The bisnorcholenol The microbial hydroxylation of new or (51) is hydroxylated at C-7a by various Bounusual steroidal substrates illustrates the tryodiplodia and Botryosphaerica strains in up flexibility of biotransformation for the prepar- to 40% yield (DESPREAUX et al., 1986). ation of hydroxysteroids. Cholesterol is hySeveral aspects of the mechanism of microdroxylated at C-25 by a Streptomyces species bial steroid hydroxylation have been investi(SUZUKIet al., 1995) and vitamin D, at the gated by HOLLAND and co-workers. The rearsame position by Amycolata autotrophica rangement of radical intermediates was exam(SASAKI et al., 1992).The modification of sub- ined by using the pregnadiene (52) as a substrate structure in order to control either the strate for C-11 and C-7 hydroxylation, and site or yield of hydroxylation has been system- rearranged products were obtained following atically investigated in several cases. The pres- biotransformations by Rhizopus stolonifer and ence of a dimethylketal at C-20 in substrates Mucor griseocyanus (HOLLANDand RIEMsuch as (44) hampers 14a-hydroxylation such LAND,1985). The involvement of cyclopropyl that biotransformation of steroid (44) by Cur- groups in such rearrangements was examined vularia lunata, which produces both llp- and using compounds (53) and (54) as substrates 14a-hydroxy products from the corresponding for hydroxylation at C-6 by Rhizopus arrhizus, dione, involves only 11p-hydroxylation (Fig. but no ring opening was observed (HOLLAND

490


OH

U

44

46 Finasteride

45

H

47

R

= C(0)

R = C(0)O; Brassinolide

Fig. U. Steroid hydroxylation.

do & (yp 49 3-DMethylestradiol

0

/

7

50 Lithocholic acid

-

0

6

52 F'regna-4,8-diene

51

53

Fig. 13. Estrogens, bile acids, and mechanism probes.

6

54

491


et al., 1990b). The hydroxylation of testosterone at C-2p by Gnomonia fructicola ATCC 11430 was shown to proceed under stereoelectronic control via oxidation of an intermediate A2-4-dienol(HOLLAND et al., 1989), a situation analogous to the C-6p hydroxylation of this substrate by R. arrhizus (HOLLAND, 1984).

2.1.3 Terpenes During the preceding decade, terpenes have been the most intensively investigated group of substrates for microbial hydroxylation, over 200 such reactions having been reported for mono-, sequi-, and diterpenoids. Hydroxylation of triterpenes remains a relatively unexplored area. The following discussion will focus on new developments in the area of terpene hydroxylations during this period. Biotransformation of monoterpenes by Borrvtis cinerea has been studied as part of an investigation of the biotransformation of flavor compounds during grape infestation by this fungus. Hydroxylations of citronellol (55)

(BRUNERIE et al., 1987a), citral (56) (BRUNEet al., 1987b), and geraniol (57) (Fig. 14) (BOCKet al., 1988) occur predominantly at the allylic positions as shown, but are accompanied by other bioconversions. Hydroxylations of the cineoles (58) and (59) have been studied by several groups with the aim of producing chiral alcohols from these common symmetrical precursors. 1P-Cineole (58) is hydroxylated predominantly at C-8 by Streptomyces griseus (ROSAZZA et al., 1987), but gives mainly C-2 endo product on biotransformation with Bacillus cereus (LIUet al., 1988) or Aspergillus niger (MIYAZAWA et al., 1991a). Hydroxylations of 1,8-cineole (59)similarly occur endo to the carbocyclic ring at C-6 using Bacillus cereus (LIU and ROSAZZA, 1990), but take place at the enantiotopic C-7 position using Glomerella cingulata (MIYAZAWA et al., 1991b) or Pseudomonas fZava (CARMANet al., 1986). Substrate enantioselectivity has also been shown during the allylic hydroxylation of (+)piperitone by Rhizoctonia solani to give the hydroxypiperitone (60) in high ee (MIYAZAWA et al., 1993a) and by Fusarium culniorum, RIE

,A6 t t

55 Citronellol

60

8

56 R = C H O citral 57 R = C H 2 0 H ; geraniol

mc-6 1

58 1.4-Cineole

62

Fig. 14. Hydroxylation of monoterpenes and related compounds. t hydroxylation positions

t

endo

59 1.8-Cineole

6 3 Rose oxides

492

I0 Hydroxylation and Dihydroxylation

which hydroxylates the racemic cis-lactone 1994). Other hydroxylations of cyclopropyl (61) at the indicated site with low enantio- substrates, such as that of the synthetic terpenselectivity (NOBILEC et al., 1994). A high de- oid (71)by Rhizopus oryzue (KUTNEYet al., gree of enantioselectivity ( > 8 5 % ) is also 1994), occur without rearrangement. shown during hydroxylation by Beauveria Biotransformation of monoterpenes by sulfirescens at the methyl groups of the syn- plant cell cultures can provide a route to oxythetic racemic substrate (62) (ARCHELAS et al., genated derivatives. The hydroxypiperitone 1988b), but cis- and trans-rose oxides are both (60, Fig. 14) is formed in 40% yield from ( -)hydroxylated by Aspergillus niger to give com- piperitone by Catharunthus roseus (HAMADA pound (63) with no substrate stereodifferen- et al., 1994), while fenchone (72, Fig. 15) is hytiation (MIYAZAWA et al., 1995a). droxylated at the designated sites by cells of Hydroxylations occurring with rearrange- Eucalyptus perriniana (ORIHARA and FURUYA, ment are of interest for mechanistic reasons: 1994). In the latter example the products are recent examples include the conversion of ( R ) - isolated as glycosidic derivatives. pulegone (64) to alcohol (65) by Botrytis allii In the past decade sesquiterpenes have re(Fig. 15) (MIYAZAWA et al., 1991~);of (1S)-3- ceived more attention than any other single carene (66) to alcohol (67) using Mycobacre- substrate group for hydroxylations, reflecting riunz smegrnatis DSM 43061 (STUMPFet al., the importance of these compounds and their 1990);and of isopinocampheol(68) to the diols derivatives as bioactive compounds and as fla(69 and 70) by several fungi, notably Rhizopus vor and fragrance agents. The antimalarial sesarrhizus and Mortierella isabellina (ABRAHAM, quiterpenes artemesinin (73) and arteether

Qo-Qo

OH

64 (&-Pulegone

65

$-$ HO

66 (lq-3-Carene

A

68 Isopinocampheol

71

Fig. 15. Hydroxylation of cyclic terpenes. t hydroxylationpositions

69

72 Fenchone

67

70


(74), derived from Chinese medicinal herbs, can be hydroxylated at C-la (Fig. 16) (HUFFORD et al., 1995) or C-90 (RAMUand BAKER, 1995) by Cunninghamella elegans ATCC 9245; at C-3a by Penicilliunz chrysogenum ATCC 9480 or Aspergillus niger ATCC 10549 (LEEet al., 1989,1990); or C-14 by Beauveria sulfures77 R = H; a-Ionone cens (Hu et al., 1991, 1992). Cunninghamella 78 R = O H echinufata NRRL 3655 has been used to convert the antitumor lactone quadrone (75) to the 8a-hydroxy derivative (76) in up to 25% yield (GOSWAMI et al., 1987a). Hydroxylations of simple fragrance sesquiterpenoids are exemplified by the conversion of a-ionone (77) to 3-hydroxy derivatives (78) by Aspergillus niger (Fig. 17) (YAMAZAKI et al., 79 R = H; a-Damascone 1988) or Mucor plumbeus (AZERADand 80 R=OH HAMMOUMI, 1991), and of a-damascone (79) to derivative (80) by Botrytis cinerea (SCHOCH Fig. 17. Fragrance sesquiterpenes. et al., 1991). Flavor components derived from farnesene (81) can be produced by hydroxylation of the sulfone adduct (82) (Fig. 18) (ABRAHAM et al., 1992), while hydroxylation of trans-nerolidol (83) and related sesquiter-

73 R = 0; Artemesinin

6

493

494


substrate groups, the cedranes and the eudesmanes. In the former area, several hydroxylations of cedrene (84) and cedrol (85) have been reported (Fig. 19). Cedrene (84) is hydroxylated at C-3, C-12, and C-15 by Beauveria sulfurescens 62474sites tiple (ABRAHAM by (LAMARE Corynespora et al., et 1987), al., 1987) casiicola but and hydroxylaatDSM multion by Rhodococcus sp. KSM-7358 occurs in high yield and specificity at C-10 (KOBUTAet al., 1992;TAKIGAWA et al., 1992). Cedrol(85) is hydroxylated predominantly at C-3a by Glomerella cingulata (MIYAZAWAet al., 1995b), Beauveria sulfurescens, and Aspergillus niger (LAMARE et al., 1987) at C-3P by Cephalosporium aphidicola (HANSON and NASIR,1993), but again at multiple sites by Corynespora casiicola DSM 62474 (ABRAHAM et al., 1987). Cephalosporium aphidicola also transforms 8epicedrol (85, stereochemistry at C-8 inverted), to the 3P-hydroxy derivative (GANDet al., 1995). Hydroxylation of a series of eudesmanes (86) has been examined by GARCIAGRANADOS et al. (1991a, b, 1993): biotransformation by Rhizopus nigricans occurs mainly to give the derivatives (87), whereas Curvularia lunara also produces products arising from hydroxylation at an isopropyl methyl group.

3

WI5 3@

H S

H

S

dIH

Hydroxylations of medicinally active diterpenes have been used to prepare new com-

HQh,.

..&

..\ti

8 . ---

88 Stemodin

HOP"

-

O 'H 89 Aphidicolin

OH

&o

\

90

84 Cedrene

85 Cedrol

R'

RHO & 9 1 Gibberellins 86 R ' = H , O H o r O , R = H 87 R ' = H, OH or 0, R = OH

Fig. 19. Alicyclic sesquiterpenes.

R = CH3, CHzOH

R' = CH2 or H, CH3

Fig. 20. Diterpenes and derivatives.


pounds for biological evaluation. The structurally related compounds stemodin (88) and aphidicolin (89) have received particular attention with eight hydroxylated derivatives of the former having being isolated from incubations with Rhizopus arrhizus, Cunninghamella echinulata, Polyangium cellulosum, and Streptomyces species (Fig. 20) (BADRIAand HUFFORD, 1991; HUFFORDet al., 1991), or Cephalosporium aphidicola (HANSONet al., 1994). Biotransformation of aphidicolin and derivatives by its source organism, C. aphidicola, has been extensively studied by HANSONet al. (1995). Biotransformations of synthetic diterpenoids such as (90) analogous to those isolated from the traditional Chinese medicinal herb Tripterygium wilfordii have been examined by MILANOVA and co-workers (MILANOVA and MOORE,1993; MILANOVA et al., 1994, 1995) and over 10 different hydroxylated products isolated from incubations with Cunninghamella and Syncephalastrum species. Microbial hydroxylation of structurally modified gibberellin precursors has been studied in Gibberella fujikuroi (ALI et al. 1992; FRAGAet a1 1992, 1994) and Cephalosporium aphidicola (BOA-

92

R = a-OH, P-CH3; Sclareol

93 R = CH2; Man001

94 Manoyl oxides

Fig. 21. Labdane diterpenes.

495

VENTURA et al., 1994): the hydroxylation of gibberellins such as (91) at C-15a can be performed by Rhizopus stolonifer (FRAGAet al., 1993). Hydroxylation of the labdane diterpenes sclareol (92), manool (93), and a series of isomeric cyclized derivatives (94) have been examined. Ring A hydroxy derivatives are formed from sclareol (92) by hydroxylation at C-2a or C-3p using Septomyxa affinis ATCC 6737 (Fig. 21) (KOUZIand MCCHESNEY, 1990), while Mucor plumbeus hydroxylates sclareol (92) at C-3p in high yield, and also converts manool (93) to the 2a-hydroxy derivative (ARANDA et al., 1991). Biotransformations of the manoyl oxides (94) by Rhizopus nigricans (GARCIA-GRANADOS et al., 1990a), Fusarium moniliforme, or Cunninghamellaelegans (GARCIA-GRANADOS et al., 1995) give more complex product profiles, but hydroxylations at C-3 and C-7 are observed.

2.1.4 Alkaloids In spite of the range of biological activities shown by this substrate group, there have been few recent reports of the microbial hydroxylations of alkaloids. The indole alkaloid eburnamonine (95) is hydroxylated in low yield at C-6a and C-18a by Mucor circinelloides (Fig. 22) (ADACHIet al., 1992),and hydroxylation of Arisrorelia alkaloids (96 and 97) has been systematically examined using a range of fungi (DOBLERet al., 1995). Hydroxylation of alkaloid (96) by Cunninghamella species occurs at C-19, while Mucorplumbeus hydroxylates (97) at the adjacent site, C-14, to give product 98 in 23% yield. The latter microorganism has also been used for conversion of dihydroquinidine (99) to the derivative (100) in high yield (AZERAD,1993). Hydroxylations at C-14 of morphine (101) and codeine (102) can be carried out using Pseudomonas putida (LONGet al., 1995) and Streptomyces griseus (HARDER and KUNZ,1989), respectively, and the semisynthetic alkaloid N-heptyl physostigmine (103) is hydroxylated at various positions indicated on the heptyl chain by Actinoplanes, Acremonium, and Verticillium species (SO et al., 1995).

496


.

H

9 5 Eburnamonine

9 7R

96

=H

9 8 R = OH

0

I

9 9 R = H; Dihydroquinidine 1 0 0 R = OH

1 0 1 R = H; Morphine 1 0 2 R = CH,; Codeine

1 0 3 N-Heptyl physostigmine

Fig. 22. Alkaloids. t hydroxylation positions

2.1.5 Antibiotics A major focus in the study of the biotransformation of antibiotics has involved milbemycin and related substrates of general structure (104, Fig. 23; cf. 2, Fig. 5). Various Streptomyces species have been identified that convert R of

structure (104) from H to OH (MIYAKOSHI et al., 1989; RAMOSTOMBOet al., 1989; NAGAKAWA et al., 1990c, 1994), while many other microorganisms perform hydroxylation of one or both substituent alkyl groups R ' (LAWRENCE et al., 1989; NAKAGAWA et al., 1989a, b, 1990d, e, 1993, 1995; SCHULMAN et al., 1993; ARISON et al., 1995). Other hydroxylations of antibiotics include conversion of monensin (105) to compounds (106) and (107) by Sebekia benihana NRRL ll l l 1 (Fig. 24) (VAUFREY et al., 1990) and of the C-19a methyl group of immunomycin (108) by Streptomyces sp. ATCC 55281 to give the hemiketal(lO9) (Fig. 25) (CHENet a]., 1994).

2.1.6 Dealkylation Reactions . .

OH

104 R = H or OH,

Fig. 23. Milbernycins.

R' = alkyl

Hydroxylation of carbon situated a to a heteroatom such as oxygen or nitrogen generally results in dealkylation of that heteroatom via an unstable intermediate hemiacetal or hemiaminal. That this reaction is frequently observed in mammalian drug metabolism has


497

OH

H Sebekia benihana

-\H

H 106

NRRL 11111

105 Monensin H

Fig. 24. Biotransformation of monensin.

been a major incentive for the investigation of microbial dealkylation reactions, and in an ongoing study FOSTER et al. (1991a, 1992a) have examined the biotransformation of a series of drugs by Cunninghamella echinulata and reported 0-demethylations of methoxyamphetamines (110). Colchicine (111) is specifically demethylated at the C-3 methoxy group by two Bacillus species (Fig. 26) (POULEVet al., 1995) and the demethylation of several immunosuppressants of the immunomycin type

L- $ y n

52--

Me0

-

OMe

Streptoxnyces sp. .1111\

H

107

(108, Fig. 25) at C-31 by Actinoplanacete ATCC 53771 has been reported (ARISONet al., 1990a, b, c; INAMINE et al., 1990). O-Demethylation of an analogous substrate by Streptomyces sp. ATCC 55387 results in the formation of a new hemiacetal (112) (SHAFIEE et al., 1994). The phenol (113) is produced in 75% yield from biotransformation of the corresponding methyl ether schizandrin by Mortierella isabellina (KANETANI et al., 1991) and similar regio-

Meo%oH

0

HO

ATCC 55281

108

Fig. 25. Oxidation of an allylic methyl group in an immunomycin.

498


\

Me0

11 1 Colchicine

110

112

Me0 I

OH

113 Gomisin T,5 ' - 0 demethyl schizandrin

114 R = H; Eudesmin 115 R = OCH,; Magnolin

.

Fig. 26. 0-Dernethylation.

selectivity is observed in the 4'-O-demethylation of eudesmin (114) and magnolin (115) by Aspergillus niger (MIYAZAWA et al., 1993b) and Spodoptera litura (MIYAZAWA et al., 1995~). Microbial removal of alkyl groups from nitrogen has also been studied as a model for mammalian metabolism (SEWELL et al., 1984), and Cunninghamella strains identified that perform N-dealkylation of propranolol (116, Fig. 27) (FOSTERet al., 1992b), pyrilamine (117) (HANSENet al., 1987), and furosemide (118) (HEZARIand DAVIS,1992). Beauveria bassiana can be used for N-demethylation of diazepam (119) (GRIFFITHSet al., 1993) and Mucor piriformis performs the N-dealkylation of a series of thebaine analogs (120) in high yield (MADYASTHA and REDDY, 1994).Various ergolin derivatives (121) can be efficiently demethylated by Streptomyces species (HUMMEL-MARQUADT et al., 1992).

In all the above examples dealkylation is presumed to involve an intermediate hydroxylated species, but in only one example, conversion of leurosine (122) to the aminal (123) by Aspergillus terricola or A . ochraceus in 35-50% yield, has an intermediate of this type been isolated (Fig. 28) (GOSWAMIet al.. 1987b). The existence of an analogous intermediate in the conversion of (*)-carnithe to the L-(-)enantiomer (124) by selective oxidation of the optical antipode by Acinetobacter calcoacericus ATCC 39647 is suggested by isotopic labeling studies (DITULLIO et al., 1994). Dealkylations of other heteroatoms have been reported recently, including the conversion of dimethylsulfonium compounds to thiomethyl ethers by various Desulfobacterium species (HANSEN and VANDER MAAREL, 1994) and the biotransformation of tributyltin by a range of microorganisms, yielding di- and


499

I OMe

fI

4

118 Furosemide

1 1 7 Pyrilamine

116 Propranolol

I

COlH

Me

.J$?

&\ H\

R"

I

1 1 9 Diazepam

1 2 0 Thebaine analogs

N

R

1 2 1 Ergolin derivatives

Fig. 27. N-Dealkylation.

124 (L)-(-)-Carnitine

122 R = H;Leurosine 123 R = O H

monobutyltin products (ERRECALDE et al., 1995). but the enzymic processes involved in these reactions are unknown.

2.1.7 Aromatic Ring Hydroxylation Formation of phenols by the microbial oxidation of aromatic substrates is a common mode of biotransformation for this substrate group

Fig. 2.8. Aminal formation.

(HOLLAND, 1992a). The discussion below will focus on recent developments in the microbial formation of mammalian phenolic metabolites, the formation of phenolic derivatives of natural products, the oxidation of simple aromatic substrates, polycyclic aromatic hydrocarbon (PAH) metabolism, and the formation of phenols from heterocyclic aromatic compounds. In the former area the formation of the catechol(l25, Fig. 29) from 3-0-methylestrad-

500


iol (49, Fig. 13) by Aspergillus alliaceus involves both 0-demethylation and hydroxylation at C-4 (WILLIAMSON et al. 1989b); and Ndemethylation accompanies hydroxylation in the conversion of N-methylcarbazole (126) to the phenolic products (127) and (128) by Cunninghamella echinulata (YANG and DAVIS, 1992). The latter organism has been used extensively as a model for mammalian drug metabolism, and converts propranolol (116, Fig. 27) to 4-hydroxypropranolol (FOSTERet al., 1992b),prenalterol(l29) to the corresponding 4-hydroxyderivative (130) in 72% yield (PAsumo et al., 1987), tranylcypromine (131) to phenol (132) (FOSTERet al., 1991b) and phenazopyridine (133) to the 2- and 4-hydroxy derivatives (134) (Fig. 29) (FOSTERet al., 1991~). In the natural product area, biotransformation of the isomeric Aristotelia alkaloids (96, Fig. 22) by Cunninghamella species gives the corresponding 7-hydroxy derivatives in up to 64% yield (DOBLERet al., 1995). Hydroxylations of flavone (135) (Fig. 30) by a range of fungi (IBRAHIM and ABUL-HAJJ, 1990) and of 5-hydroxyflavone (136) by Strepromyces fill-

vissimus (IBRAHIM and ABUL-HAJJ, 1989) occur predominantly at the 4 ' position, as does hydroxylation of a range of flavone and isoflavone analogs by Absidia blackesleeana (ABUL-HAJJ et al., 1991). The latter reactions have been used to deduce a model for the interaction of flavonoids with the hydroxylase enzyme of A . blakesleeana. Hydroxylation of simple aromatic substrates has been studied as a method for the regiospecific synthesis of phenolic products. Biphenyl (137) and terphenyl (138) are hydroxylated at the 4,4' and 4,4" positions, respectively,in high yield by Aspergillus parasiticus (ABRAMOWICZ et al., 1990 MOBLEY and DIETRICH, 1991),reactions that have been performed on up to 600 g of substrate. Aniline can be converted to 2- and 4-aminophenol by various Aspergillus strains (BURKHEAD et al., 1994), and 4-hydroxy-o-toluidine produced from o-toluidine in good yield by Nocardia asteroides I F 0 3384 (YAMADAet al., 1990) or Fusarium verticillioides (YOSHIOKA et al., 1990; YAMADA et al., 1991).A series of phenols (140) can be produced from the corresponding alkyl

OH

R

125

126 R = CH3, R' = H 127 R = CH3, R' = O H 128 R = H , R ' = O H

131 R = H; Tranylcypromine 132 R = O H Fig. 29. Aromatic hydroxylation (125-134).

129 R = H Prenalterol 130 R = O H

133 R = H 134 R = H,OH


135 R = H 136 R = O H

141 R = halogen, OH

139 R = H 140 R = O H

137 R = H

138 R = Ph

501

8

OH

142

3

143 Fluoranthene

6

144 Pyrene

Fig. 30. Aromatic hydroxylation (135-144).

N-phenylcarbamates by Beauveria sulfurescens ATCC 7159 (VIGNEet al., 1987,1991). Phenylacetic and -propionic acids are converted to phenols by various microorganisms. 2-Hydroxyphenylacetic acid is formed by Humicola or Chaetomium species (STAUDENMAIER et al., 1994) while the 3-hydroxy isomer is produced by Rhizoctonia solani (STAUDENMAIER et al., 1993a).Beauveria bassiana can be used to prepare the 5-hydroxyphenylacetic acids (141) from the corresponding 2-substituted substrates (STAUDENMAIER et al., 1993b, c). 2-Phenylpropionic acid is hydroxylated in the para position by Streptoymyces rimosus ATCC 10907 (KUGE et al., 1991), while the corresponding phenoxyacid is similarly converted by Streptomyces hygroscopicus (COOPERet al., 1990). Biotransformations of polycyclic aromatic hydrocarbons continue to be examined in connection with toxicity studies and the production of analytical standards for metabolite identification. Recent examples include the conversion by Cunninghamella elegans of phenanthrene to the phenol (142) (isolated as the P-glucoside) (CERNIGLIA et al., 1989) and of fluoranthene (143) to the 3- and 8-phenols

(POTHULURI et al., 1990), and the conversion of pyrene (144) to 1-, 6-, and 8-phenolic products by Aspergillus niger (WUNDERet al., 1994) and Penicillium species (LAUNENet al., 1995). A focus of recent attention in the area of phenol formation by biotransformation has been the use of heterocyclic aromatic substrates. Efficient hydroxylations of nicotinic acid (145) at C-6 (Fig. 31) (HOECKSand VENETZ, 1991; YAMADAet al., 1992a; NAGASAWA, 1993; KIENERet al., 1993a) and at C-2 (KIENERet al., 1993b) have been developed; analogous products can also be obtained from 3-cyanopyridine (146) (YASUDAet al., 1993,1995;SASAKIet al., 1994a, b; UEDAet al., 1994a, b). Picolinic acid (147) is hydroxylated at C-6 by Alcaligenesfaecalis (KIENERet al., 1992a, 1993c; YAMADAet al., 1992b) and the corresponding nitrile (148) can also be converted to the 6-0x0 derivative in high yield (KIENER,1992c;KIENERet al., 1993~). Efficient hydroxylations of other substituted pyridines can also be accomplished: a series of 3-alkylaminomethylpyridinesare hydroxylated at C-6 by Arthrobacter oxydans (ISHIKAWAet al., 1993) and hydroxylation of 6-methylnicotinic acid at C-2 is carried out by Alcaligenes eutrophus (KIENERet al., 1994).

502


6

OR

145 R = C O z H 146 R=CN

sion of fatty acids and related substrates containing a (Z, Z)-lP-diene unit into a chiral 5hydroperoxy-1,3-diene (Fig. 32). This material can be reduced to the corresponding alcohol with retention of configuration, providing a route for the asymmetric hydroxylation of suitably constituted substrates. The most commonly used enzyme of this type is soybean lipoxygenase,which has been used for the oxidation of linoleic and linolenic acids to chiral hydroperoxyacids such as (153),an intermediate in the synthesis of phospholipid hydroperoxides (Fig. 33) (BABAet al., 1990;NAKAHARA et al., 1990 YONEDAet al., 1992). Optimization of the soybean lipoxygenase reaction on a scale suitable for the production of gram quantities of the 13-(S)-hydroperoxide (153)from linoleic acid has been achieved (MARTINIet al., 1994). This enzyme is also able to oxidize unnatural fatty acid-related substrates to chiral hydroperoxides; the conversion of compound

147 R = C02H 148 R=CN

H 149

150

15 1

152

Fig. 31. Hydroxylation of pyridines and 1,4-pyrazines.

Other heterocyclic aromatic substrates can also be converted to phenols in high yield. Hydroxylations of quinoline at C-2, -3, -6, or -8 can be achieved using Pseudomonas species (BOYDet al., 1993b;KAWASHIMA and SUEYOSHI, 1992,1993) and pyrazine carboxylic acid is converted to compound (149) or (150)by different strains of Alcaligenes (KIENER,1993;KIENERet al., 1994) and to (151)by Pseudomonas acidovorans (KIENER,1992d; KIENERet al., 1994).2,5Dimethylpyrazine (152) is hydroxylated at C-3 in 90% yield by Rhodococcus erythropolis (KIENERet al., 1992b). Isoquinoline, quinoxaline, and quinazoline all form phenolic products on biotransformation with Pseudomonasputida UV4 (BOYDet al., 1993b).

2.2 Biotransformation with Isolated Enzymes 2.2.1 Lipoxygenases The lipoxygenases are non-haem, iron-containing dioxygenases that catalyze the conver-

io2

FR

HiR' GOH

FR

H

5

R'

OH

Fig.32. Lipoxygenase-catalyzed hydroxylation.


503

157

Fig. 33. Soybean lipoxygenase-mediated hydroperoxidations.

(154)to (155)and of (156)to (157)proceeds in 1992). Porcine 12-lipoxygenase has, however, high yield and enantioselectivity (DUSSAULT been employed for the conversion of the conand LEE, 1995). The use of a prosthetic acidcontaining binding group facilitates the preparation of chiral diols of type (158) by the soybean lipoxygenase-catalyzed oxidation of intermediate esters using the route shown in Fig. 34 (DATCHEVA et al., 1991). This concept has been extensively used by SCHELLER et al. (1995) in a systematic investigation of the substrate selectivity of soybean lipoxygenase with respect to the alkyl substituent R of Fig. 34. n = 3-7 Biotransformations of unsaturated fatty C H acids by other plant lipoxygenases have been R investigated by GRECHKIN et al. (1991a, b). Linolenic acid (159) is converted initially to the hydroperoxide (160)by potato tuber lipox1 SBLO ygenase, but to the isomer (161)and derived 2 Reduction products by lipoxygenase preparations from 3 Hydrolysis flax, wheat, and corn (Fig. 35). A lipoxygenase from the mushroom Pleurotus pulmonarius converts linoleic acid to the hydroperoxide (153,Fig. 33) (ASSAFet al., 1995). The preparative use of lipoxygenases from mammalian sources has not been extensively developed, although a review of the function of these enzymes suggests a wider role for Fig. 34. Synthesis of chiral alcohols using soybean them in biotransformations (YAMAMOTO, lipoxygenase (SBLO).

=. I

I

504


159 Linolenic acid

160

161 Fig. 35. Potato tuber, flax, wheat, and corn lipoxygenase-mediated hydroperoxidations.

jugated arachidonic acid isomer (162) to the convert the epoxyeicosatrienoic acid (164)to 14-(S)-hydroperoxide (163) (Fig. 36) (LABEL- the 11-(R)-hydroxy derivative (165) (Fig. 37) LE et al., 1990), and cyclooxygenase used to (ZHANGet al., 1992a).

2.2.2 Peroxidases

l4

I

162

Porcine 12-Lipoxygenase

In spite of the ability of peroxidases to oxidize a wide range of functionalized organic compounds (HOLLAND,1992a), there have been few reports of the application of these enzymes for simple hydroxylation reactions. Horseradish peroxidase is reported to catalyze the oxidation of benzene to phenol by hydrogen peroxide when the substrate is used as the medium for the conversion (AKASAKA et al., 1995), and to catalyze conversion of the diaminobiphenyl (166) to the benzylic alcohol (167) (Fig. 38) (OKADAet al., 1988). Chloroperoxidase catalyzes the benzylic hydroxylation of simple substituted toluenes (MILLERet al., 1995) and other phenyl alkanes (ZAKSand DODDS,1995), the latter reaction showing a substrate-dependent stereoselectivity.The protein-bound haem unit microperoxidase has also been shown to catalyze hydrogen peroxide-dependent N-demethylation of dimethylaniline (MASHINO et al., 1990).

yco2* Fig. 36. Porcine 12-lipoxygenase-mediated hydroperoxidation.

164 Fig. 37. Hydroxylation by cyclooxygenase.

165


505

R

'

166 R - H 167 R = O H

I

Fig. 38. Hydroxylation by horseradish peroxidase.

.

2.2.3 Steroid Hydroxylases Although the bulk of preparative steroid hydroxylations has been performed using whole-cell biocatalysts (see Sect. 2.1.2), useful steroid-hydroxylating enzyme preparations have recently been obtained from Cochfiobofus funatus (JAENIG et al., 1992; VITASet al., 1995) and Aspergiffusfumigatus (SMITHet a]., 1994). Both are cyt. P-450-dependent systems and have been used for biotransformations of progesterone, the former having 1lp-hydroxylating activity and the latter producing 7p-, lla-, and 15p-hydroxylated products.

4

,OH

169

2.2.4 Other Enzymes Cytochrome P-450-dependent enzymes pos170 Dihydrosanguinarine sess the potential to carry out a wide range of hydroxylation reactions. This is illustrated by Fig. 39. The biosynthesis of dihydrosanguinarine. the reactions performed by a cyt. P-450-enriched enzyme extract from Streptomyces OH griseus that performs a wide variety of such processes, including aromatic and aliphatic hydroxylations, and 0- and N-dealkylations (TROWER et al., 1988). Other cyt. P-450-mediated reactions carried out by isolated enzymes include the biosynthetic conversion of protopine (168) to dihydrosanguinarine (170) via rearrangement of the aminal (169),a process carried out by a microsomal preparation of Eschschoftziacafifornica (Fig. 39) (TANAHASHI and ZENK,1988),and 0-demethylation at C-31 of the immunosuppressant FK-506 (cf. 108, Fig. 25) by an enzyme preparation from Streptomyces rimosus (SHAFIEE et al., 1995). Other isolated enzyme systems can also per17 1 Aphelandrine form hydroxylations. Fungal monophenol oxidase (HSIUNGet al., 1992) and -mushroom Fig. 40. Aphelandrine (17171)(TODOROVA et al., 1994).

506


A wide range of mono-substituted benzenes can be converted to the corresponding cis-2,3diols, the more recent examples being summarized in Tab. 1; other substituted examples are available from the bromo- or iododiols via palladium-catalyzed substitution reactions (BOYDet al., 1991b). Diols derived from toluene and the halobenzenes have been used as chiral starting materials for synthetic purposes, notably in the synthesis of various conduritols (CARLESS and OAK,1991a, b; CARLESS, 1992a, b) and in a wide-ranging synthetic program by HUDLICKY and co-workers, encompassing the syntheses of (-)-zeylena (HUDLICKY et al., 1989a), D- and L-erythrose (HUDLICKY et al., 1989b), pyrollizidine alkaloids (HUDLICKY et al., 1990a),L-ribonic y-lactone (HUDLICKY and PRICE,1990), various cyclitols (HUDLICKY et al., 1990b, 1991a, b, 1994a), (-)-specionin (HUDLICKY and NATCHUS,1992) and (+)3.1 Dioxygenase-Catalyzed lycoricidine (HUDLICKY et al., 1994b). Reactions Di-substituted benzenes are also suitable substrates for the dioxygenation reaction of Pseudomonas. I? testosteroni converts phthalic 3.1.1 Arenes and Heteroarenes acid to the 4,5-dihydro-4,5-diol (MATSUBARA et al., 1990, 1991). I? putida 39/D converts 2The oxidation of aromatic substrates to give bromostyrene to the diol (172)with an ee of and HUDLICKY, cis-1,2-dihydroxycyclohexa-3,5-dienes (see Fig. 92% (Fig. 42) (K~NIGSBERGER 4) is a characteristic reaction of the dioxy- 1993),while the three isomeric chlorostyrenes genase enzyme system of prokaryotic micro- give the corresponding diols (173)with ee’s of organisms. The reaction has been known for >98% (Cchloro), 54% (5-chloro), and 15% et al., some time, but new substrate types and organ- (6-chloro), respectively (HUDLICKY isms capable of carrying out this conversion 1993). These latter data are consistent with continue to be discovered. Although the pro- the model for dihydroxylation of substituted cess has not recently been reviewed in its monocyclic arenes by I? putida UV4 proposed entirety, the conversion of fluoroaromatic by BOYDet al. (1995) and shown in Fig. 41, in compounds (RIBBONS et al., 1987) and reac- which the enantioselectivity of oxidation is tions carried out by Pseudomonas putida UV4 controlled by the steric bulk of the substituents. have been summarized (BOYDet al., 1995).A The value of this model has been demonstrated summary of the stereoselectivity is shown in in the preparation of a series of substituted arene 3,4-dihydrodiols and 2,3-dihydrodiols of Fig. 41. The product derived from benzene, czs-1,2- unusual configuration by using the bulky iodo dihydroxycyclohexa-3,5-diene,has been used group as a “director” for the regio-and enantioas a starting material for the synthesis of a selectivity of dioxygenation, followed by its number of natural products, including pinitols reductive removal (BOYDet al., 1994). The formation of 1,2-dihydrodiolsfrom sub(LEYand STERNFELD, 1989), inositols (LEYet al., 1990), conduritols (JOHNSON et al., 1991), stituted arenes appears to be limited to benand methyl shikimate (JOHNSON et al., 1993). zoic acid and related substrates (HOLLAND, Conditions have been developed that permit 1992a); this phenomenon is seen in the conthe continuous production of this diol by Pseu- version of 2-trifluoromethylbenzoic acid to domonas in organic solvents (VAN DEN TWEEL the diol (174) by Pseudomonas aeruginosa et al., 1995). et al., 1987;NISHIMURA and KAWAKAMI, 1990). (SELIFONOV polyphenol oxidase (BURTON et al., 1993) convert a variety of phenols to catechols, while an analogous enzyme is implicated in the o-hydroxylation of aphelandrine (171)by macerated Aphelandra tetragonu root (Fig. 40) (TODOROVA et al., 1994).The non-heme iron enzyme xylene monooxygenase from Pseudomonas oleovorans converts a variety of substituted toluenes to the corresponding benzyl alcohols (WUBBOLTS et al., 1995).

3 Dihydroxylation Reactions

507


I > Br>CH3> F > H

%OH

HO""

'

9

Q

OH

HO

Major

Minor

Major

Minor

Fig. 41. Predictive rules for dihydroxylation of substituted monocyclic arenes by Pseudomonas putida UV4 (BOYDet al., 1995); L: large, S: small.

Conversion of tri-substituted monocyclic arenes to cis-dihydrodiols has been reported only in the context of the biotransformation of substituted acids, including the conversion of

172

173

3-flUOrO- (MARTINet al., 1987) and 3-chlorophthalic acids (EVANSet al., 1990) to the corresponding diols (175) by Z? testosteroni and I? pufida, respectively, and biotransformation

174

Fig. 42. Diols from dihydroxylation of substituted benzenoids.

175

176

508


Tab. 1. Dihydroxylation of Monosubstituted Benzenes

Substituent

Pseudomonas putida Strain

F

uv4 39lD uv4 39D uv4 39lD uv4 39lD uv4 39lD uv4 39lD

c1 Br

I CH3 CzHs CH=CHZ C= CH C(CH3)=CH2 C ~ S CF3 CH,CH,Br OCH3 OCzHs CH,OAc

Tji 0

xy

HO

C02CH3 COCH, CHZSCH3 CN CSC-Ph

39lD unspecified uv4 uv4 NCIB 12 190 9lD uv4 uv4 uv4 unspecified

ee

["/.I 60

> 98 > 98 > 98 > 98 > 98 > 98 > 98 > 98 >98

>98 >98 >98

Reference BOYDet al. (1991~) HUDLICKY et al. (1994b) BOYDet al. (1991~) HUDLICKY et al. (1988,1989b, 1992,1994b) BOYDet al. (1991b) HUDLICKY et al. (1992,1994b) BOYDet al. (1991b, 1994) HUDLICKY et al. (1994b) BOYDet al. (1991~) HUDLICKY et al. (1988) BOYDet al. (1991~) HUDLICKY et al. (1988,1989a) SCHOFIELD (1989) HUDLICKY et al. (1988) BESE'ITI et al. (1989) BOYDet al. (1995) BOYDet al. (1991~) SCHOFIELD (1988) STABILE et al. (1995) BOYDet al. (1995) ASTLEY et al. (1993) BOYDet al. (1991~) TAYLOR (1990)

39lD

MADERand TAUTVYDAS (1990)

FM 803

JOHNSON and MONDELLO (1991)

NCIB 11680 NCIB 11767 uv4 uv4 uv4 Pseudomonas sp. BM2

BLACKER et al. (1993) BLACKER et al. (1993) BOYDet al. (1995) BOYDet al. (1995) BOYDet al. (1995) GEARYet al. (1990)

Pseudomonas sp. BM2

GEARY et al. (1990)

509


of 3,4- and 3,5-difluorobenzoic acids by I? putida JT 103 to give the 12-diols (176) (ROSSITER et al., 1987). Bacterial biotransformation of bicyclic arenes also involves the action of dioxygenase enzymes. Biodegradation of tetralin by a Corynebacterium species proceeds via the dihydrodiol(l77) (Fig. 43) (SIKKEMA and DE BONT, 1993), but no analogous products are obtained from biotransformation of tetralin by R putida.The latter gives cis-dihydrodiol metabolites (178 and 179) of benzocyclobutene (BOYDet al., 1991a), in contrast to the formation of the isomer (180) by Rhodococcus (GRUND,1993a) or Pseudomonas ATCC 55 196 (GRUND,1993b). Substituted naphthalenes can give mixtures of dihydrodiols on dioxygenation. 2- Methylnaphthalene is reported to give only the diol (181) on biotransformation using Pseudomonas FERM P-9632 (Fig. 44)(OSUMIand TAKESHITA,1990b) or R putida NCIB 9816 (DELUCA and HUDLICKY, 1990),but is transformed to a mixture of diols (181) and (182) by F! putida 39/D (DELUCAand HUDLICKY, 1990);The biotransformation of 2-methoxynaphthalene is more complex, as isomers (183), (184), and (185) are produced in ratios that depend on

the relative activities of toluene, naphthalene, and biphenyl dioxygenases in the organisms concerned (WHITEDet al., 1994). Dioxygenation of heterobicyclic arenes has been studied by BOYDet al., who examined the biotransformations of benzofuran (1993a), dihydrobenzofuran (1993a), benzothiophene (1993b), and a series of nitrogen-containing substrates (1993~)by I! putida UV4. While dihydrobenzofuran was subjected mainly to benzylic hydroxylation, benzofuran gave the (S,S)-diol(lM) in 34% yield (Fig. 45) together with compound (187), the latter arising via dioxygenation of the heteroarene ring. In contrast, benzothiophene gave a stable dihydrodiol derivative (188). Dihydroxylations of quinoline, isoquinoline, quinoxaline, and quinazoline by I! putida UV4 occurred exclusively in the carbocyclic rings to give chiral products (189-193), but diol(l94) was formed as a racemic mixture (Fig. 46). A single report of the dioxygenation of a tricyclic arene involves conversion of dibenzocyclobutene to the diol (195) by R putida UV4 (BOYDet al., 1991a). In addition to dihydrodiol products, the bacterial metabolism of arenes can also produce catechols. The latter products may be formed

HyyJ @ cp HD OH

W

\

HO

OH

OH

OH

177

HO

178

179

180

Fig.43. Diols from dihydroxylation of bicyclic benzenoids. OMe

OH 181 R = Me 183 R=OMe

182 R=Me 184 R=OMe

Fig. 44. Diols from dihydroxylation of naphthalenes.

OH 185

510


OH OH

H

OH 186

d

187

O

H

188

Fig. 45. Products from dihydroxylationof benzofuran and benzothiophene.

189

190

N

. .‘/OH -OH

191

-

“OH OH OH

OH

193

192

194

‘

195

Fig. 46. Products from dihydroxylation of nitrogen heterocycles and dibenzocyclobutene.

196

197

200 X = o-,m-, pC1, eBr

198 n = 1,2

201

199

x=o,s

202

Fig. 47. Products from dihydroxylation of indenes, benzopyrans, benzothiopyrans, styrenes, and norbornenes.


by a combination of dioxygenase and dehydrogenase activities, such cases being exemplified by the conversion of benzene to catechol by Rhodococcus species (TSUJIKIet al., 1993) and that of p-toluic acid to 2,3-dihydroxy-ptoluic acid by Arfhrobacfer FERM P-11173 (SUSUKIet al., 1991). Alternatively, catechols can arise solely as a result of dioxygenase action on substrates that possess a leaving group at the site of oxidation. Examples of the latter conversions include the biotransformation of 2.5-dichlorophenol to dichlorocatechol by the toluene dioxygenase in P pufida FL (SPAINet al., 1989) and the conversion of 2-aminobenzenesulfonic acid to catechol-3-sulfonic acid by Alcafigenes (JUNKER et al., 1994).

3.1.2 Dihydroxylation of Other C=C Bonds The first indication that dioxygenase enzymes could attack non-aromatic carbon-carbon double bonds was provided by the observation by WACKETT et al. (1988) that toluene dioxygenase of P pufida 39/D converted indene to the cis-(lS,2R)-dio1(196) in 35% yield but low (30%) ee. The enantiomeric product (197) was obtained in higher ee using microbial strains containing the naphthalene dioxygenase system (Fig. 47) (GIBSON et al., 1995) or P pufida NCIMB 8859 (ALLENet al., 1995), but the cis-(lS,2R)-diol (196) was again produced from indene (in > 98% ee) by P putida UV4 (BOYDet al., 1989). The latter organism also produces optically pure diols (198) from the appropriate benzocycloalkene (BOYDet al., 1989,1990) and from the analogous heterocyclic substrates (199) (BOYDet al., 1993b). Oxidation of styrenes by P putida 391D gave, in addition to cis-dihydroarene diols, the diols (200) in a configuration dependent on the location of substitution (HUDLICKY et al., 1993; KONIGSBERGER and HUDLICKY, 1993). The sole example of dioxygenase-catalyzed oxidation of a non-aromatic substrate involves conversion of norbornadiene to the cis-exodiol (201) by Pseudomonas sp. BM2 (GEARY et al., 1990). Benzonorbornadiene gave no analogous products, and 7-phenylnorbornadiene (202) gave only products arising from oxidation of the aromatic ring.

511

3.2 Monooxygenase- Cat a1yzed Reactions Dihydroxylations catalyzed by monooxygenases proceed via an epoxidation reaction, by the route outlined in Fig. 4. The intermediate epoxide is rarely isolated in whole-cell biotransformations, being rapidly converted by enzymic or non-enzymic hydration to a vicinal diol, formed with trans stereochemistry in the case of cyclic substrates. The enzymic reaction involves an epoxide hydrolase that can impose enantio- or regioselectivity on the process, but the more frequently observed nonenzymic hydration generally follows the rules of mechanistic chemistry.

3.2.1 Arenes and Heteroarenes The formation of arene trans-dihydrodiols has been extensively studied in the context of the metabolism and toxicity of aromatic hydrocarbons (HOLLAND, 1992a). Much of the microbial work in this area has been carried out by using various Cunninghamella species, exemplified by a recent report by POTHULURI et al. (1990) of the biotransformation of fluoranthene (143, Fig. 30) by C. elegans ATCC 36 112 to give the trans-diols (203-205) (Fig. 48).

203 R = H, R' = O H 204 R = O H , R ' = H 205 R = R ' = H Fig. 48. Diols from the dihydroxylation of fluoranthene.

512


pound (208) by Aspergillus (Fig. 50) (ISMAILIALAOUIet al., 1992). Similar reactions can also be carried out by plant cell cultures, exemplified by the conversion of 3-carene (66, Fig. 15) Conversion of alkenes to vicinal diols fre- to the diol (209) by Nicotiana tabacum and quently occurs during the biotransformation Catharanthus roseus (HIRATAet al., 1994). of terpenes. Several dihydroxylations of un- These reactions frequently, however, give low functionalized monoterpenes have been re- yields of product, a problem that can often be ported, including the conversion of myrcene circumvented by the addition of a polar deriv(206) to diol (207) by Diplodia gossypina ative group to the substrate. The addition of a ATCC 10936 (Fig. 49) (ABRAHAM and STUMPF, phenylcarbamate group to geraniol or nerol 1987) and of (R)-pulegone (64,Fig. 15) to com- results in substrates (210) and (211)(Fig. 51),

3.2.2 Dihydroxylation of Other C=C Bonds

206 Myrcene

207

Fig. 50. Diols from the dihydroxylation of pulegone and 3-carene (64 and 66, Fig. 15).

Fig. 49. Alkene dihydroxylation.

210 2,3-trans 2 1 1 2.3-cis

214

2 09

208

212

213

215

Fig. 51. Dihydroxylation of alkenic monoterpenes and derivatives.

216


respectively, which can be converted to the corresponding diols (212) or (213) by Aspergillus niger LCP 521 in a pH-dependent biotransformation: at pH 2 the 6-(S)-diol (212) is obtained following nonenzymic epoxide hydrolysis, whereas at pH 6 the 6-(R) isomer (213) is formed as a result of epoxide hydrolase action (FOURNERON et al., 1 9 8 9 b ; Z ~et~ al., ~ c1991a, b). Analogous results have been obtained from 7-geranyloxycoumarin (214) (ZHANGet al., 1991b;MULLERet al., 1994) and from the citronellol derivative (215) (ZHANGet al., 1992b). In the former example, the intermediate 6(R),7-epoxide could be isolated from biotransformations by Nocardia alba and Bacillus cereus (MULLERet al., 1994). Substrate activation can also be achieved by formation of a sulfolene derivative such as (216) obtained from reaction of myrcene (206, Fig. 49) with sulfur dioxide. Hydroxylation and dihydroxylation of compound (216) by various microorganisms, including the formation of the corresponding 6,7-diol by Rhizopus arrhizus and Aspergillus niger, is facilitated in this way (ABRAHAM and ARFMANN, 1992). Analogous dihydroxylations of sesquiterpenes also occur. Trans-nerolidol (83, Fig. 18) is converted to diols (217) and (218) (Fig. 52) by A. niger ATCC 9142, and cis-nerolidol(219) gives diol (220) (Fig. 53) (ARFMANN et a]., 1988). Diols (217) and (220) can also be obtained from biotransformation of trans- and cis-nerolidol (83 and 219), respectively, using Streptomyces cinnamonensis, where they are accompanied by the corresponding epoxide products (HOLMESet al., 1990). In contrast,

513

H I

HO

HO 217

Fig. 52. Diols from the dihydroxylation of tmnsnerolidol(83, Fig. 18).

Fusarium solani DSM 62416 converts transnerolidol (83) to alcohols primarily by hydration, although geranylacetone (221) is converted to the diol (222) in low yield (Fig. 53) (ABRAHAM and ARFMANN, 1989). Biotransformations of the sesquiterpene abisabolol (223) by Aspergillus niger (Fig. 54) (MIYAZAWA et al., l990,1992a, b) and Glomerella cingulata (MIYAZAWA et al., 1995d) give a series of products (224-227) arising from dihydroxylations in time-dependent yields. The diterpenoid (228) is converted to diol (229) and epoxide (230) (among other products) by long-term biotransformation using Rhizopus nigricans (Fig. 55) (GARC~A-GRANADOS et al., 1990b). Epoxides are formed as major products (without subsequent hydrolysis to diols) during the biotransformation of drimenol (231) by Mucor plumbeus (ARANDA et al., 1992) and of caryophyllene (232) by Diplodia

PH 2 19 cis-nerolidol 0

2 2 1 Geranylacetone

-

220 OH 0

222

Fig. 53. Diols from the dihydroxylationof cis-nerolidol(219) and geranylacetone (221).

514

p p


HO

P

-

OH

OH 224

2 25

226

227

223 a-Bisabolol

HO Fig. 54. Polyols and tetrahydrofurans from the hydroxylation of a-bisabolol(223).

2 2 8 R = CH2 2 2 9 R = OH,CHzOH 230R=

2 3 1 Drimenol

232 Caryophyllene

0 A

Fig. 55. Epoxidation substrates.

gossypina (ABRAHAM et al., 1990a) or Chaetomium cochliodes (ABRAHAM et al., 1990b). Dihydroxylations of non-terpenoid substrates have also been reported. Beauveria sulfurescens ATCC 7159 converts the olefin (233) to diol(234) in a reaction thought to parallel the hydroxylation of compound (235) to (236) (Fig. 56) (JOHNSON et al., 1992a). Likewise Mortierella isabellina ATCC 42 613, which performs the benzylic hydroxylation of aromatic hydrocarbons, produces a series of diols (237-242) from the analogous unsaturated

233 234 235 236

R = CH;, R = OH, CH;,OH R=Hz R = H,OH

Fig. 56. Parallel pathways for hydroxylation and dihydroxylation.

Mm 3 Dihydroxylation Reactions

515

substrates (HOLLAND et al., 1994b). The isomeric forms of diol 237 are also produced by biotransformation of cis- and trans-l-phenylMe I-propene by Pyricularia oryzae (Fig. 57) (NUKINA et al., 1994). The absolute stereochemis244 PrecoceneII tries of the diols (237-242) are consistent with their formation by a route involving enantioselective epoxidation by a benzylic hydroxylase followed by nonenzymic hydration of the resulting epoxide (HOLLAND et al., 1994b); the epoxides (243) can be isolated following biotransformation of the corresponding olefinic substrates by Mortierella isabellina (AGARWAL et al., 1994). 245 R' = H, R2 = O H The conversion of precocene I1 (244) to the 246 R' = OH, Rz = H cis- and trans-diols (245) and (246) by a crude extract of Streptomyces griseus has been re- Fig. 58. Dihydroxylation of precocene I1 (244). ported (Fig. 58) (TROWERet al., 1988). The analogous substrate (247) is efficiently transformed in high optical purity to the diol(248), a chiral synthon for the production of antihypertensive agents, by Mortierella rammanianu (Fig. 59) (PATELet al., 1994a), and other microorganisms give analogous products (PATEL 247 et al., 1994b). The use of isolated enzymes in dihydroxylations has not been extensively developed, but chloroperoxidase can convert para-chlorostyrene to the corresponding diol in low yield (COLONNA et al., 1993), and gives the diols

Ncm 248

Fig. 59. Dihydroxylation of a benzopyran (247).

& R\

\

237 R = H 238 R = C H 3

~

' I

241 n 242 n

(CH2)" I O = 1

2 3 9 R = CH3 240 R = C2H5

H

[:ao

2 49

Fig. 60. Diol from the chloroperoxidase-mediated dihydroxylation of 1-methylcyclohexene.

243 X , Y

= CH,

N

=2

Fig. 57. Parallel pathways for benzylic hydroxylation and dihydroxylation.

(239, Fig. 57) and (243, Fig. 60) (of undetermined stereochemistry) from the corresponding unsaturated substrates (Fig. 60) (ZAKSand DODDS,1995).

516


4 Summary The use of biotransformation for the production of chiral alcohols and diols has successfully built on and expanded its classical foundation of steroid biotransformation to encompass a wide variety of substrate groups and product types. Studies of the microbial metabolism of aromatic compounds have similarly developed into and consolidated the role of biotransformation in the production of phenols and dihydroarenediols. The continued investigation of new substrates, enzymes and microorganisms can only enhance the utility of oxidative biotransformations in such diverse areas as the environmental biodegradation of xenobiotics, the production of drug metabolites, and the generation of chiral materials for the chemical industry.

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11 Flavin Monooxygenases Uses as Catalysts for Baeyer-Villiger Ring Expansion and Heteroatom Oxidation

DAVID R. KELLY Cardiff, UK

PETERW. H. WAN London, UK

JENNY TANG Liverpool, UK 1 Introduction 536 2 Abiotic Baeyer-Villiger Reactions 537 3 “Biological” Baeyer-Villiger Reactions 541 3.1 Flavin Monooxygenases,Identification and Isolation 541 3.2 Cyclohexanone Monooxygenase (CHMO) from Acinefobacfersp. NCIMB 9871 548 3.3 Large Scale Baeyer-Villiger Reactions 568 4 Asymmetric Sulfoxidations by Flavin Dependent Monooxygenase Enzymes 569 4.1 Introduction 569 4.2 Asymmetric Sulfoxidations by Monooxygenase Enzymes Occurring in Bacteria 570 4.2.1 Acinetobacter calcoaceticus NCIMB 9871 Cyclohexanone Monooxygenase (CHMO) 570 4.2.2 Asymmetric Sulfoxidations by Flavin Dependent Monooxygenase Enzymes Present in Other Microorganisms 574 4.3 Asymmetric Sulfoxidations by Flavin Dependent Monooxygenase (FMO) Enzymes Occurring in Mammalian Tissue 575 5 Conclusions 577 6 References 577

536

11 Flavin Monooxygenases

1 Introduction The three principal flavins found in nature are riboflavin (Vitamin B2)(1) (Fig. l),flavin mononucleotide (FMN) (2), and flavin adenine dinucleotide (FAD) (3).The latter two act as prosthetic groups for flavoproteins and there are well established pathways for their interconversion. The flavins are bright yellow pigments (as are flavoenzymes) which are uniquely important because they undergo one electron reduction to give stable semiquinone radicals. Hence, they are able to mediate between the common two electron oxidations (e.g., NAD(P)/NAD(P)H) and one electron oxidations carried out by heme or iron-sulfur cluster proteins. WALSH(1980) has described flavins as the biological redox cross-roads although coenzyme Q is also able to act in this role, albeit in a more limited way. Another criterion that distinguishes flavoenzymes from

1 R = H; Riboflavin, Vitamin B2 2 R = PO,2-; FMN 3 R = Adenosyl pyrophosphate; FAD

Fig. 1. Natural flavins.

many other redox enzymes is that the flavin remains tightly bound to the active site throughout the catalytic cycle. Indeed FMN (2) is sometimes covalently bound and FAD (3) frequently so (e.g., succinate dehydrogenase or 6hydroxy-D-nicotine oxidase; MAUCH et al., 1990).The consequence of this, is that both the substrate and redox cofactors must be delivered to the active site. The most important uses of flavoenzymes in biotransformations are as oxygenases. These are classified into two groups: monooxygenases and diooxygenases. Monooxygenases, also referred to as “mixed function oxygenases”, catalyze the insertion of one oxygen atom from dioxygen into a substrate, the other oxygen atom undergoing reduction to water, whereas dioxygenases catalyze the insertion of both oxygen atoms from dioxygen into a substrate. The broader aspects of the Enzyme Commission nomenclature for oxygenases are discussed thoroughly in Chapter 10, Sect. 1.1.2. The flavin linked monooxygenases which are the focus of this chapter are assigned to EC 1.14.13. For example, cyclopentanone monooxygenase which catalyzes the oxidation of cyclopentanone to Gvalerolactoneis EC 1.14.13.16 which is systematically named as cyclopentanone, NADPH: oxygen oxidoreductase (5-hydroxylating, 1,2-lactonizing). Dioxygenases usually contain iron and are involved in processes such as prostaglandin or leukotriene biosynthesis and the conversion of aromatic compounds to 12-diols (cf. Chapter 10, this volume). The monooxygenases employ a wider range of prosthetic groups/cofactors including iron (e.g., cytochrome P,,& copper (e.g., dopamine P-hydroxylase), pterins, and flavins, together with a two electron reductant such as NAD(P)H or vitamin C. Many of these systems utilize oxygen in the form of a flavin hydroperoxide (5) (Fig. 2) (GHISHLA et al., 1978). In the normal catalytic cycle, FAD (3) undergoes reduction to the dihydroflavin (4) which is oxidized by dioxygen to the 4a-hydroperoxyflavin (5). This probably occurs by electron donation from the dihydroflavin (4) to oxygen to give the semiquinone radical and superoxide radical anion which undergo rapid radical recombination. In the presence of a substrate, oxygen transfer occurs to give the hydroxyflavin (6) which eliminates water to

537

2 Abiotic Baeyer-Villiger Reactions

nNqYo RqPP

N

NAD(P)H

NAD( P)

N- H

I/

RqPP

I n ; & N .

NYo O

4H

0 2

RqPP

“ H ”

RqPP

6

L1

projuct

.

u

O’ H

5

Fig. 2. The catalytic cycle for the reaction of flavins with oxygen. RAPP: Ribityl adenosine pyrophosphate (FAD)

complete the catalytic cycle by regenerating from the standard repertoire of peracids is the FAD (3). In the absence of substrate the 4a- epoxidation of alkenes. In the next section the hydroperoxyflavin ( 5 ) eliminates hydrogen mechanism of the abiotic Baeyer-Villiger reperoxide to regenerate FAD (3) (MERENYI action will be discussed as a prelude to that of and LIND,1991). 5-Alkylated flavins catalyze the “biological” Baeyer-Villiger reaction. Enabiotic Baeyer-Villiger reactions by reaction zyme catalyzed Baeyer-Villiger reactions have with hydrogen peroxide to give an analog of been reviewed (DAVIESet al., 1989; DRAUZ 1995; AZERAD,1995; WONG the 4a-hydroperoxyflavin ( 5 ) (MAZZINIet al., and WALDMANN, 1994) and recent develop1996, 1997). The 4a-hydroperoxyflavin (5) and WHITESIDES, and TURNER, 1992; reacts with aldehydes to give carboxylic acids. ments discussed (ROBERTS This is simply a variant of the Baeyer-Villiger ROBERTSand WILLETTS,1993; FANGet al., reaction in which hydrogen rather than an al- 1995; CARREAet al., 1996; WILLETTS,1997). kyl group migrates. However, this reaction is The oxidation of sulfides to sulfoxides by flaalso responsible for the production of light by voenzymes has also been reviewed (OTTOLINA et al., 1996; HOLLAND, luciferase (EC 1.14.14.3) in many microorgan- et al., 1995; COLONNA 1997). isms (VILLAand WILLETTS, 1997; FRANCISCO et al., 1996). Further aspects of the mechanism are discussed in the section on cyclohexanone monooxygenase (WALSH and CHEN, 1988; GHISLAand MASSEY, 1989). The practically minded synthetic organic chemist should simply view flavoenzymes as a source of “chiral hydroperoxide” with a reactivity comparable to that of perbenzoic acid. Indeed they oxidize ketones to esters and lacThe reaction of cyclic ketones with peroxytones (Baeyer-Villiger reaction), sulfides to monosulfuric acid to give lactones was first sulfoxides and sulfones, phosphines to phos- discovered at the turn of the century (BAEYER phine oxides, etc. The only reaction missing and VILLIGER, 1899,19OOa,b). The subject has


538

11 mavin Monooxygenases

been intensively reviewed (HASSAU, 1957; 1978).A verifiable consequence of this mechaKROW1981,1993) and hence only a broad out- nism is that the intra-annular oxygen of the line stressing mechanistic aspects will be given lactone is derived from the peracid or peroxhere. The general mechanism is illustrated by ide and the carbonyl oxygen is derived from the oxidation of cyclohexanone(7) to ocapro- the ketone.This has been confirmed for the reet al., 1956), lactone (10) (Fig. 3). In the classical mecha- actions of peracetic acid (BUNTON nism, nucleophilic addition of a peroxide (8) to the conversion of testosterone to testololactthe ketone (7) gives the “Criegee” intermedi- one by a monooxygenase from Penicillium 1963), and and LIEW,1983) which lilacinum (PRAIRIEand TALALAY, ate (9a) (cf. SCHREIBER then rearranges by migration of an alkyl group whole organism conversions of 2-tridecanone to the proximal oxygen of the peroxide linkage to undecyl acetate (Fig. 12) (BRITTONet al., with concomitant reformation of the carbonyl 1974). For most peracids and ketones the reaction group. In both the abiotic (WEBERet al., 1966) and cyclohexanone monooxygenase (SCHWAB, is first order in both these components and 1981; SCHWAB et al., 1983) catalyzed Baeyer- generally rearrangement of the tetrahedral Villiger reactions the migrant group retains its intermediate is rate determining, but for aryl configuration. In the abiotic reaction it is ketones substituted with electron donating generally the group best able to stabilize a pos- groups addition of the peroxide is rate deterand itive charge that migrates (i.e., tertiary > sec- mining, particularly at low pH (PALMER et al., 1974).The reaction is ondary > primary > methyl) (WINNIKand FRY,1970; WINNIK STOUTE,1973), although there are numerous general acid catalyzed and is promoted by exceptions due to conformational factors, electron withdrawing groups in the hydroperwhich are discussed below (ARVAIet al., 1992; oxide (TOKENet al., 1991). Acyclic ketones do not readily undergo HAMLEY et al., 1991; GRUDZINKSKI et al., Baeyer-Villiger reactions with carboxylic peracids. Bis(trimethylsily1)peroxomonosulfate (CAMPOREALE et al., 1990) and Caro’s acid (readily available from commercial potassium caroate; HOOS0,K OxoneTMor Curoxm) react with simple acyclic ketones such as acetone to give esters, but the reaction proceeds via dioxiranes (11) (ADAMand HADJIARAPO1993; ADAMet al., 1989;MURRAY, 1989). GLOU, BAEYERand VILLIGERused Caro’s acid and with extraordinary prescience proposed dioxirane intermediates almost 90years before they were isolated for the first time (BAEYERand 1899). However, dioxiranes (11) are VILLIGER, only formed with peroxides derived from very strong acids and/or with ketones that are extremely resistant to rearrangement. Thus far they appear to play no part in enzyme catalyzed Baeyer-Villiger reactions. It was noted above,that in the Criegee intermediate (9a) there is a preference for the migration of carbon centers best able to bear a positive charge. However, stereoelectronic 1997; analysis and theoretical studies (OKUNO, CARDENAS et al., 1997;SINGHet al., 1996) indicate that there should be a preference for miFig. 3. The gross mechanism of the Baeyer-Villiger gration of the group best able to overlap with reaction. the antibonding orbital of the peroxide bond.

8;


This was neatly demonstrated by intramolecular Baeyer-Villiger reaction of the keto peracid (U) which gave exclusively the lactone (14) formed by migration of the methylene group in preference to the methine group (Fig. 4). The bond which lies anti-periplanar to the peroxide bond is able to overlap with the antibonding orbital of the peroxide bond and hence this results in preferential methylene migration (CHANDRASEKHAR and ROY,1987, 1994). Enantioselective peracid formation using Candida antarctica lipase and hydrogen peroxide enabled the lactone (14) to be prepared with modest enantiomeric excess (21% ee, 54% yield) (LEMOULT et al., 1995). The value of the Baeyer-Villiger reaction in synthesis can scarcely be understated as attested by the table of examples in KROW'Sreview (1993) which fills 420 pages! One less obvious, but nevertheless important application is the use of Friedels-Craft acylation followed by Baeyer-Villiger oxidation and hydrolysis to install a phenolic hydroxyl group (Fig. 5 ) (TERANISHI et al., 1994; NAKATSUKA et al., 1987; YEAGERand SCHISSEL, 1991). This three step

I

t 0 .

14

H

l2 A

13

15

Fig. 4. IntramolecularBaeyer-Villiger reaction.

16

19

539

RCO3H

18

Fig. 5. Tandem Friedels-Craft acylation - BaeyerVilliger reaction.

sequence is frequently more effective than direct oxidation which often results in over oxidation to quinones and other by-products. Similarly benzaldehydes can be converted to phenyl formates, whereas aliphatic aldehydes are oxidized to carboxylic acids. In the laboratory, peracids are generally the reagents of choice for the Baeyer-Villiger reaction, albeit that this is often accompanied by epoxidation of alkenes (MENDELOVICI and GLOTTER,1992). rn-Chloroperoxybenzoic acid is used most frequently (GARLASCHELLI et al., 1992; KOCHand CHAMBERLIN, 1989), but concerns about its stability have restricted commerical availability until recently. Magnesium monoperphthalate has been advocated as a safer alternative and it is generally easier to remove by extraction from a reaction mixture (MINOet al., 1997; HIRANO et al., 1995,1996). These reagents are convenient on a laboratory scale, but less so on a pilot plant or a manufacturing process (COLEMAN et al., 1997; FLISAK et al., 1993). This has spawned efforts to replace peracids with other peroxides and to generate peracids or other peroxides in situ from hydrogen peroxide or oxygen. For large scale manufacturing processes, heterogenous catalytic processes are preferred. These generally consist of a solid catalyst plus hydrogen peroxide (CLERICI, 1993) or dioxygen and an aldehyde (KANEDAet al., 1994a; YANet al., 1997). Typical catalysts include hydrotalcite (UENOet al., 1997), titan-

540


ium silicate molecular-sieves (BHAUMIK et al., 1996), and magnesium oxide supported metal complexes (YANet al., 1997). Hydrogen peroxide is a poor reagent for the Baeyer-Villiger reaction unless a strong acid is present (MLOCHOWSKI and SAID, 1997). This acts either as a general acid catalyst which facilitates addition to the carbonyl group and rearrangement of the Criegee intermediate or it is converted to a peroxy derivative which is the active reagent. For example, phenylseleninic acid upon treatment with hydrogen peroxide gives phenylperoxyseleninic acid which is a potent Baeyer-Villiger reagent (SYPER,1989; TAYLOR and FLOOD,1983; cf. GUZMAN et al., 1995).In some cases Criegee intermediates (9) can be used as reagents for Baeyer-Villiger reactions of other ketones. One of the more stable is the adduct (21) (Fig. 6) formed from chloral (20) and hydrogen peroxide which has a reactivity comparable to that of peracids (GANESHPURE and ADAM,1996). The reaction of aldehydes with dioxygen promoted by light generates a species which is capable of the epoxidation of alkenes (KANEDA et al., 1992, 1994b; S C H O ~ L Eand R BoLAND,1997) and the Baeyer-Villiger reaction (LI et al., 1996).The reactive species is almost certainly the peracid (23)(Fig. 7) formed by autoxidation of the aldehyde (22), although this is not accepted by some workers (MASTRORILLI et al., 1995).The reaction is also promoted by metal catalysts (LI et al., 1996; Mu-

21

20

Fig. 6. Stable “Criegee intermediates”.

lH KOA* 0 2

0

Lightor

R

22

and YAMADA, 1995; YAMADA et al., 1991;MURAHASHI et al., 1992) and in this case the peracid may be an intermediate but not the active reagent. Many other reagents have been advocated for the Baeyer-Villiger reaction including sodium percarbonate-trifluoroacetic acid (OLAHet al., 1991), peroxomonophosphoric acid (PANIGRAHI and NAYAK,1982), borax-HzOz (PANDEand GUPTA,1995), and hydrogen fluoride in water and acetonitrile (ROZENet al., 1995), but these offer few advantages over peracids on a laboratory scale. Until recently the only enantioselective Baeyer-Villiger reactions were those catalyzed by enzymes.The first report of an abiotic enantioselective Baeyer-Villiger reaction was the use of chiral platinum(I1) complexes (e.g., 24; Fig. 8) which act as Lewis acids (FRISONE et al., 1993), however, the enantiomeric excesses were generally poor (Gusso et al., 1994;STRUKUL et al., 1997).The metal complex catalyzed reaction of aldehydes and oxygen (BOLMet al., 1993) and subsequent Baeyer-Villiger reaction has been adapted into an enantioselective procedure by using copper(I1) oxazoline complexes (e.g., 27 Fig. 9). Cyclohexanones, cyclopentanones (BOLMet al., 1994), and cyclobutanones (BOLMand SCHLINGLOFF, 1995;BOLM et al., 1997a, b) are all converted into lactones with respectable to excellent enantiomeric excesses using only 1mole % of the catalyst (27). The standard conditions for the Sharpless asymmetric epoxidation of allylic alcohols, i.e, KAIYAMA

catalyst

Fig. 7. Formation of peracids in situ.

23

24

bnprb,q,,p 0

HzOz HClO?

rac-25

S?

2 6 58% ee

Fig. 8. Pt( 11) complex catalyzed Baeyer-Villiger reactions.

$;q b;?

3 “Biological”Baeyer-Villiger Reactions

541

Ti(O’Pr),, di-isopropyl tartrate and t-butyl hydroperoxide are also capable of effecting enantioselective Baeyer-Villiger oxidation of cyclobutanones. However, the appeal of using this well known procedure, is diminished by the need to use a large excess of reagent over a prolonged reaction time (LOPPet al., 1996).

-

-

3 “Biological” Baeyer-

27

0-N L’

Villiger Reactions

f

28

3.1 Flavin Monooxygenases, Identification and Isolation

R

29 65% ee,

41% yield

Fig. 9. Cu(I1) oxazoline complex catalyzed Baeyer-Villiger reactions.

NADPH NADP

The earliest report of a “biological” Baeyer-Villiger reaction is probably the conversion of progesterone (30) (Fig. 10) to testololactone (34) by several mbXmganiSmS. In one case the isolated yield was 70% (FRIEDet al., 1953;

H

H

3 1 17-OAcetyltestosterone

NADP NADPH

H

35 17-a-Hydroxyprogesterone

32 Testosterone

34 Testololactone

Fig.10. Baeyer-Villiger reactions of steroids

H

33 4-Androstene -3,17-dione

542


PETERSON et al., 1953) and the reaction has luciferase from Photobacterium phosphoreum been run with 4-hydroxy-androst-4-ene-3,17-shows the opposite regioselectivity, i.e., it oxidione (cf. 33) on a multigram scale (SARIASLA- dizes 2-tridecanone to dodecanoic acid, preNI and ROSAZZA, 1984).The conversion of pro- sumably via methyl dodecanoate (VILLAand 1997). gesterone (30)to testosterone (32)by partially WILLETTS, Other early examples which demonstrate purified extracts from Cylindrocurpon radicicola requires NADPH and dioxygen (RAHIM the breadth of compounds which can be acand SIH, 1966) and the same requirements commodated by these enzymes are the ring exwere found for the conversion of 4-andros- pansions of 2-pentyl cyclopentanone (40) (Fig. tene-3,17-dione (33)to testololactone (34)by 13) (SHAW,1966) and questin (42) (Fig. 14) partially purified extracts from Penicillium lilucinum (PRAIRIEand TALALAY, 1963). Purified “steroid” monooxygenase from Cylindrocarpon rudicicola consists of two subunits (M, 115,OO) each binding a molecule of FAD and has an absolute requirement for NADPH and dioxygen.The predominant activity is as a progesterone monooxygenase (EC 1.14.99.4) as indicated by a very low K , (0.41 FM) for progesterone and other 20-ketosteroids (ITAGAKI, 1986a).However, it also acts as a 4-androstene3,17-dione monooxygenase (EC 1.14.99.12), Glomerella fusmioides but with much lower activity ( K , 40 FM) (ITAGAKI,1986b).In contrast the steroid monooxygenase from Rhodococcus rhodochrous acts exclusively as a progesterone monooxygenase, although it contains FAD and has absolute requirements for NADPH and 0, as do the other enzymes (MIYAMOTO et al., 1995). In recent work it has been shown that cytochrome P450CYP17 which normally hydroxylates C-17 of progesterone (354, also Fig.11. Ring A cleavage. forms 17-0-acetyltestosterone (31)in a minor side reaction (MAKand SWINNEY, 1992; SWINNEY and MAK,1994). Another early observation was Baeyer-Villiger oxidation of ring A of “lanosterone type ketones” (e.g., 36,Fig. 11; LASKIN et al., 1964) which has proved to be a common degradation pathway (SHIRANE et al., 1996). 2-Tridecanone (38)(Fig. 12) is very common Fig. U . Degradation of tridecanone (38) in the in nature where it acts as an insect repellant biosphere. for many plants (e.g., wheat and tomatoes). Many microorganisms are able to degrade it to undecyl acetate (39)(FORNEY et al., 1967;FORNEY and MARKOVETZ, 1968,1969). Hydrolysis and oxidation yields undecanoic acid which is further degraded by the P-oxidation pathway. Pseudomonas The enzyme responsible (isolated from PseuSPP. domonas cepacia; BRITTONand MARKOVETZ, 40 41 1977) contains FAD and requires NADPH and 0, for activity (BRITTON et al., 1974).The Fig.13. SHAW (1966).

3 “Biological” Baeyer-Villiger Reactions

543

example, the enantiomer of 25diketocamphane (46)according to the commonly used system is 3,6-diketocamphane (53)! The rationale is that carbons at the same positions in the two enantiomers have the same number. Both enantiomers of camphor induce the 42 Questin formation of a single cytochrome P450,,, monooxygenase (EC 1.14.15.1) which surprisingly hydroxylates both enantiomers at corresponding positions. There are separate dehydrogenases for the enantiomeric 5-exo-hydroxycamphors (45), ent-(45),but there is little selectivity between the enzymes for each enantiomer (GUNSALUSand MARSHALL, 1971).The first enantiospecific reaction in the series is the Baeyer-Villiger reaction of 2,5-diketocamphane (46) catalyzed by 2,5-diketocamphane 1,2-monooxygenase (2,5-DKCMO). COzH OH Early studies indicated that this consisted of two (CONRAD et al., 1961,1965) or three com43 Desmethylsuloclirin ponents (Yu and GUNSALUS, 1969) which required iron for activity. Careful purification Fig. 14. FUJIIet al. (1988). enabled the isolation of an NADH oxidase ( M , 36000; EC 1.8.6.1) (TRUDGILL et al., 1966) and a monooxygenase ( M , 78000), consisting (FUJIIet al., 1988).The cases described above of two nonidentical subunits which bind one share one distressing factor. None of these molecule of FMN. As expected for a flavin systems has been exploited in the biotransfor- monooxygenase, iron was not required for ac1986). The lacmation of unnatural substrates, yet they un- tivity (TAYLORand TRUDGILL, doubtedly have untapped potential. This is tone (47) has never been isolated from this clearly demonstrated by the following exam- Baeyer-Villiger reaction because it undergoes ples which have achieved the transition from spontaneous p-elimination to give the ketocarmysterious biological phenomena to reagents boxylic acid (48). However, treatment of (+)camphor (44)with 2,5-DKCMO does give the of real value in organic synthesis. Treatment of Pseudomonas putida with analogous lactone (54) (Fig. 16). 2J-Diketo(+)-camphor (44)yields virtually a complete camphane (46) and 3,6-diketocamphane (53) sequence of isolable metabolites (45, 46, 48, are not easily accessible hence the correspond49) (Fig. 15) (BRADSHAW et al., 1959).The en- ing enantiomers of camphor are often used to zymes responsible are induced by growth on assay the monooxygenases. The Baeyer-Villiger reaction of the ketoboth (+)-camphor (44) and (-)-camphor (52) and the genes are located on the 165 MDa carboxylic acid (48) (2-0x0-A’-4,5,5-trimethyltransmissible CAM plasmid (RHEINWALD et cyclopentenylacetic acid) requires both a al., 1973).The pathway for the degradation of coenzyme A ester synthetase and an oxygen( -)-camphor is essentially identical to that of ase which has been purified to homogeneity ( +)-camphor, (albeit obviously with enantio- (Mr 106000). It consists of two identical submeric intermediates) and different enzymes. units which bind a single molecule of FAD. The pathways merge at the achiral ketodicar- Baeyer-Villiger activity is absolutely specific boxylic acid (51) (GUNSALUS and MARSHALL, to the coenzyme A ester of the ketone (48)and 1971). Readers are cautioned that the conven- requires both NADPH and dioxygen (OUGtion for numbering the carbon atoms of HAM et al., 1983). The degradation of (-)-camphor (52) has intermediates in the degradation pathway for ( -)-camphor (52) follows perverse logic. For been studied in much less detail than that of

;rs yyOcaL "$$

544

I 1 Flavin Monooxygenases

\

0 2

NADPH NADP

NAD

NADH

46 2,s-Diketocamphane

4 5 5-em-Hydroxycamplior

44 (+)-Camphor

0

Jfo2

2,5-DKCMO NADH NAD

0

CO2H

.Ji

p-el imi nation

I I

HOzC

NADP

47

48

49

H20

0

NADPH

CO2H Oxidation

Oxidation

3 Acetate 1 Isobutyrate

50

5 2 (-)-Camphor

5 3 3,6-Diketocampliane = enr-46

46 25-Diketocamphane = ent-53

Fig. 15. The catabolism of (+)-camphor (44) by Pseudomonusputidu ATCC 17453 (OUGHAM et al., 1983).

+

( )-camphor (GUNSALUSand MARSHALL, tivity. 3,6-Diketocamphane 1,6-dioxygenase 1971) and many aspects probably require rein- (3,6-DKCMO) which catalyzes in vivo the

vestigation. As noted above the initial hydroxylation and oxidation have low enantioselec-

Baeyer-Villiger reaction of 3,6-diketocamphane (53) to the ketocarboxylic acid em-(&)


545

If a mixture of microorganisms (e.g., a soil sample) are grown on a single carbon source, evolutionary selection and/or adaption will frequently yield a species able to survive by 44 (+)-Camphor

54

Fig. 16. 2,SDKCMO catalyzed Baeyer-Villiger reaction.

I

55 Fenchone is very similar to 2,5-diketocamphane 1,2monooxygenase (2,5-DKCMO) but is enanCoomebacterium sp. tiospecific for ( - )-camphor (52) and hence or peracetic acid presumably also for 3,6-diketocamphane (53). It also consists of two components: an NADH oxidase and an FMN linked monooxygenase consisting of two electrophoretically identical subunits (M,76000) binding one molecule of FMN.This enzyme is able to utilize the NADH oxidase isolated with 2,5-DKCMO (JONES et al., 1993). The camphor monooxygenases of this strain (Pseudomonusputidu NCIMB 10007) were exploited by ROBERTS, WILLETTS, and coworkers in a series of investigations with unnatural substrates (see Tab. 2). The crude enzyme extract consisting of 2S-DKCMO with 3,6-DKCMO and 2-0x0-A3-4,5,5-trimethylcyclopentenylacet- Fig. 17. CHAPMAN et al. (1965). ic acid monooxygenase could be used as a selective catalyst by using NADH or NADPH as a cofactor, respectively.After separation of the NADH and NADPH mediated Baeyer-Villiger activity the two fractions were designated M01 (monooxygenase 1) which consisted of 2,5-DKCMO plus 3,6-DKCMO and M 0 2 which contained 2-0x0-A3-4,5,5-trimethylcy- 58 1,8-cineole clopentenylacetic acid monooxygenase. Rhodococcus sp. C 1 There are no doubt further monooxygenor 2,s-DKCMO 59 ases involved in the degradation of terpenes which are ripe for exploitation. For example, the Corynebacterium sp. mediated BaeyerVilliger reaction of fenchone (55) (Fig. 17) gives a product ratio which is quite different from that of peracids (CHAPMAN et al., 1965). An organism isolated from soil under a eucalyptus tree and 2,5-DKCMO both effect ring lactonization of l,&cineole (58) to give an elactone (59) which spontaneously rearranges 61 60 to the useful y-lactone (61) (Fig. 18) (WILLIAMS and TRUDGILL, 1989). Fig. 18. WILLIAMS and TRUDGILL (1989).

I

546


metabolizing the carbon source. This is known as elective culture. Elective culture on cyclopentanol yielded a Pseudomonas sp. (NCIMB 9872) which was able to oxidize cyclopentanol (62)to glutarate (66) (Fig. 19) and presumably on to acetate by an inducible set of enzymes. Cell extracts showed NAD linked alcohol dehydrogenase activity, Baeyer-Villiger ring expansion of cyclopentanone (63) linked to dioxygen and NADPH consumption, lactonase activity towards 6-valerolactone (a), and dioxygen and NADH or NADPH linked oxidation of 5-hydroxyvaleric acid (65) to glutaric acid (66) (GRIFFINand TRUDGILL, 1972). Purified cyclopentanone monooxygenase (CPMO; EC 1.14.13.16)consists of four subunits ( M , ca. 200.00) binding 2-4 molecules of FAD. It is absolutely specific for NADH and requires dioxygen and has a very low K , for cyclopentanone (1 km; GRIFFINand TRUDGILL,1976; TRUDGILL, 1990b). CPMO has been exploited OH

64

65

0 2

NAD

NADH 66

Fig. 19. Metabolism of cyclopentanol (62) to glutaric acid (66).

for the Baeyer-Villiger reaction of unnatural substrates, although the stereoselectivity is similar to that of cyclohexanone monooxygenase(CHM0). Elective culture on cyclohexanol was also used to select for species able to oxidize cyclohexanol to adipic acid in a sequence homologous to that shown in Fig. 19.Two species: Nocardia globerula CL1 (NORRISand TRUDGILL, 1971) and Acinetobacter sp. NCIMB 9871 (DONOGHUE and TRUDGILL, 1975) were isolated which could achieve this transformation, and cyclohexanone monooxygenases (CHMO; EC 1.14.13.22) with comparable properties were purified from both (DONOGHUE et al., 1976). Both consisted of a single polypeptide chain binding one molecule of FAD and as with CPMO the Baeyer-Villiger activity had an absolute requirement for NADPH and dioxygen. The CHMO from Nocardia globerula CL1 consisted of two electrophoretically distinguishable forms ( M , 56,00-58,00) with virtually identical properties (NORRISand TRUDGILL, 1976). Unfortunately, stock cultures of A! globerula CL1 were lost and so no further development has been possible.The procedure for the extraction of the Acinefobacter sp. NCIMB 9871 CHMO has been optimized (TRUDGILL, 1990a) and cultures of the organism are maintained in about a dozen laboratories.The enzyme and the organism are currently the most widely used reagents for biological Baeyer-Villiger reactions. In early publications the NCIMB 9871 strain is described as Acinetobacter calcoacericus, but nowadays the species name is not used. Unfortunately, Acinetobacter sp. NCIMB 9871 is a class 2 pathogen which means that it cannot be grown without specialist microbiological containment facilities. However, the gene for CHMO has been expressed in bakers’ yeast (STEWART, 1997). The Baeyer-Villiger activity with 4-substituted cyclohexanones (STEWART et al., 1996b) was similar to that reported previously with purified CHMO (TASCHNER and BLACK,1988;TASCHNER et al., 1993) and with 2-substituted cyclohexanones (STEWART et al., 1996a) (which have not been transformed with Acinetobacter sp. NCIMB 9871 CHMO); the results were similar to those obtained with Acinetobacter TD63 (ALPHAND et al., 1996).


547

Acinetobacter TD63 was isolated from elec- no1 to acetophenone (74) (Fig. 21). Both this tive culture on 1,2-trans-cyclohexan-1,2-diol species and Nocardia T5 then metabolize (67). Dehydrogenase and monooxygenase ac- acetophenone (74) by a Baeyer-Villiger oxitivity were detected in cell extracts (Fig. 20), dation to give phenyl acetate (75), which which indicated pathways similar to those for the degradation of cyclopentanol (62) and cyclohexanol. The majority of the metabolic activity proceeds via Baeyer-Villiger oxidation of the ketoalcohol (69) and a minor pathway consists of a parallel set of reactions commencing with the diketone (68).This organism is not 73 1-Phenylethanol able to grow on the metabolites of cyclohexano1 degradation, which is attributed to the absence of a lactonase (cf. 64 to 65). This feature Arrlirobacter sp. only is an advantage for the biotransformation of NADH some unnatural substrates because it prevents further degradation (DAVEYand TRUDGILL,

NAD=l

1977).

The microbial decomposition of l-phenylethanol (73) involves two distinct pathways. In one pathway (utilized by Nocardia T5) catechol formation is followed by ring cleavage and further degradation by the P-oxoadipate pathway. An Arthrobacter species is able to enantioselectively oxidize (S)-1-phenyletha-

NADPH

Arthrobacter sp.

75 Pheiiyl a c e u t e

U o H 7 6 Phenol

H2

72

0

71

Fig. 20. Metabolism of cyclohexan-1,2-diol (67) to adipic acid (71).

Fig.21.

CRIpPs

(1975), C R l P P s et al. (1978).

548

11 FIavin Monooxygenases

undergoes rapid ester hydrolysis (CRIPPS, 1975) and oxidation to catechol (77) which enters the P-oxoadipate pathway (CRIPPSet al., 1978). A similar Baeyer-Villiger reaction forms part of the degradation pathway for 4-ethylphenol (JONESet al., 1994). Baeyer-Villiger oxidation of questin (78) to desmethylsulochrin (79) (Fig. 22) by Aspergillus terreus requires two proteins, NADPH, and

78 Questin

COzH OH

79 Desmethylsulochrin Fig. 22. FUJIIet al. (1988).

1

Beauveria bassiam

Fig. 23. Baeyer-Villiger degradation and de-esterification of raspberry ketone (80).

dioxygen. The instability of the enzyme was partially overcome by the addition of polyols and and the detergent Tween 80 (FUJIIet al., 1988). In a search for a new biocatalytic route to raspberry ketone (80), it was discovered that it was degraded by Beauveria bassiana via a Baeyer-Villiger reaction to tyrosol (81) (Fig. 23) (FUGANTI et al., 1996). The reaction proceeds with retention of configuration at the migrating methylene group (FRONZAet al., 1996) and only very closely related ketones are transformed (DONZELLI et al., 1996).

3.2 Cyclohexanone Monooxygenase (CHMO) from Acinetobacter sp. NCIMB 9871 The mechanism of the Baeyer-Villiger reactions of CHMO appears to be ter-ter (RYERSON et al., 1982).NADPH binds to the holoenzyme, reduces FAD (3) to FADH (4) (Fig. 2), and is then released. Oxygen is reduced to give the 4a-hydroperoxyflavin (5)and cyclohexanone is bound. Baeyer-Villiger ring expansion gives the lactone and a hydroxyflavin (6) which eliminates water to regenerate FAD (3). If no substrate is present, the 4a-hydroperoxyflavin (5) is still formed, but it spontaneously eliminates hydrogen peroxide to regenerate FAD (3) (WALSHand CHEN,1988; GHISLAand MASSEY, 1989). Acinefobacter sp. NCIMB 9871 CHMO has a non-covalently bound, fairly loosely bound FAD prosthetic group which is easily lost ( K , 40 nM) during purification. Sequence data (Tab. 1)and inhibitor studies indicate that CHMO has a single active site (WRIGHTet al., 1994a;LATHAM and WALSH,1987).The yellow protein has been purified to homogeneity (TRUDGILL, 1990a) and consists of a single peptide of 542 amino acids (CHENet al., 1988; KELLYet al., 1996a) with two disulfide bridges and no metal ions. There are no X-ray crystal structure determinations for enzymes that catalyze the Baeyer-Villiger reaction, although crystals have been obtained for 2,5- and 3,6DKCMOs, the enzymes involved in the degradation of camphor (MCGHIE and LIITLECHILD,1996). Comparison of the amino acid

549

3 “Riological”Raeyer- Klliger Reactions

Tab. 1. N-Terminal Sequences of Flavoenzymes which Catalyze t h e Baeyer-Villiger Reaction (KELLY e t al., 1996a)

Pseudomonas putida

6-DKCMO

Pseudomonas putida, 23-DKCMO Rhodococcus rhodo irous, SMO

i

Rhodococcus coprophilus WT- 1,

C.radicola.SMO Residue number - 1 to -13

1 2 3 4 5

Ale Glu Trp Ala Glu Glu Phe Asp

6

7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28

Val

Leu Val

Thr Ala Gin( Thr) Thr Ile His Gly(Thr) Val ASP

Ala Val Val

[Met] Ser Gln LeU[LYSI Met ASP We ASP Ah Ile Val

Thr, Thr, Met, Thr, Thr,Met, Thr, Thr, Glu, Gin, Leu, Gly, Met Asn Asn Ser Val Asn ASP LYS

Leu

Met Asn GlY Gln His pro

Vd Leu Leu

Val Val Val

Asp

Arg

Met Gln

Ala GlY Phe Phe GlY

Thr

Pn,

TYr ASP

Leu

Pn, Thr Arg Thr Ala Arg Gln Met

Ala Met Glu

Thr

GlY Leu Ile Phe His Pro TYr Met TYr Pro GIY

LYS Ser Ala Ala

Gln

Asp FADNADPH 1 FMN/NADH Identical residues in the FADNADPH or FMNNADH linked enzymes are marked in bold, very closely related residues are marked with italics. Residues in square brackets are those determined by analysis of the gene which differ from those determined in the current study. Residues in curved brackets are alternative assignments, due to ambiguity in the determination.

I

1

550


W,PP sequence of CHMO with sequences from other flavoproteins indicates a potential nicotinamide binding site between residues 176 and 208 (WIERENGA et al., 1986) and an FAD binding site at the amino terminus between residues 6 to 18. This latter location is typical of flavoproteins and the characteristic sequence of three glycines, closely followed by two glycines is present. This is a subset of the GXGXXG consensus (WIERENGA and HOL, 1983) found in many nucleotide binding proI 82 R teins (MCKIEand DOUGLAS, 1991). Alignment of the 13amino acids in the putaRqPP tive FAD binding region of CHMO, with human and bacterial glutathione reductases, places the five glycines and a further two amino acids (Asp-7, Ile-11) in identical positions for all three enzymes.Two other positions were 3 si-face 0 either leucine or isoleucine, one either phenylalanine or tyrosine and another valine or alanine. In total seven perfect fits and four partial W,PP fits were observed in thirteen amino acids in three enzymes (CHENet al., 1988). The X-ray crystal structures of human glutathione reductase (KARPLUS and SCHULTZ, 1987; ERMLERet al., 1991) and bacterial p-hydroxybenzoate hydroxylase (another FAD linked monooxygen83 ase; SCHREUDER et al., 1988;KEUMet al., 1990) have been determined and this region is folded Fig. 24. Diastereotopic faces of FAD (3) and a carin a right-handed pap unit with FAD bound to ba-analog (83). the protein surface on the si-face. On this basis, the hydroperoxide group should be attached to the re-face of the flavin ring. In p-hydroxybenzoate hydroxylase the dimethyl benzene NADPH (82), hydride donation from NADPH moiety of the isoalloxazine ring lies at the en- (82) to FAD (3) should be selective for one trance to the active site cavity. In glutathione face. If the analog (83) is bound to an enzyme reductase the re-face is covered by a tyrosine the stereochemistry of donation can be deteruntil NADH is bound. In both enzymes a long mined from the 'H-NMR spectrum of the libhelix points towards the N-1-02a region of the erated reduced prosthetic group analog (MANisoalloxazine which bears the majority of the STEIN et al., 1986). Using this technique donacharge in the corresponding radical and anion. tion was shown to occur to the the re-face of et al., 1988) as is common The dipole of the helix stabilizes charge build- FAD (3) (MANSTEIN for NAD(P) linked flavoenzymes (GHISLA and up in this region and helps to remove NADP' 1989; SUMNER and MAITHEWS,1992). from the active site by charge repulsion. On MASSEY, the basis of this evidence it can be reasonably The stereochemistry of donation of hydride assumed that the substrate accesses the hydro- from C-4 of NADPH to FAD bound to peroxide from the side of the dimethylbenzene CHMO has not been determined, but all external flavoenzymes analyzed thus far donate the moiety (WIERENGA et al., 1983). The two faces of the isoalloxazine ring of pro-R hydrogen as shown in Fig. 24 (You et al., FAD (3) are diastereotopic (Fig. 24) and given 1977,1978). Each of the groups involved in this area the rigidity of the binding to the protein and the presence of a putative binding site for have proposed models for the active site of


551

CHMO and many of these have been real. (1996) have defined a sophisticated cubic viewed recently (WILLETTS, 1997). TASCHNERactive site model based on the reactions of was the first to show a correlation between the over 40 ketones (Fig. 25). configuration of the Criegee intermediate The author has proposed an active site modformed with bicyclic ketones and cyclohexa- el based solely on the configuration of the nones. On the basis of the X-ray crystal struc- Criegee intermediate (Fig. 26). All Baeyertures of glutathione and p-hydroxybenzoate Villiger reactions (chiral or achiral) can be dihydroxylase it was proposed that the hydro- vided into two types. Those which proceed via peroxide group was attached to the re-face of a Criegee intermediate with an (S)-configuraFAD (cf. 5, si-face hydroperoxide) ( TASCHNERtion and those that proceed via a Criegee et al., 1992). FIJRSTOSS initially proposed a cu- intermediate with an (R)-configuration. In this bic active site model with an excluded zone for analysis the configuration is not assigned from bicyclic ketones (ALPHAND and FIJRSTOSS,the priorities based on the usual Cahn-Ingold1992b) which was subsequently applied to Prelog notation, but takes the order of precedcyclohexanones (ALPHANDand FURSTOSS,ence peroxide > hydroxyl (or ether) > mi1992a) and refined for 2-substituted cyclohex- grating group > non-migrating group. If this anones (ALPHAND et al., 1996). OTTOLINA et corruption of the usual system is unsatisfacto-

L

’“YP

FI

RF

n

Front

v

Fig. 25. CHMO active site model (OITOLINA et al., 1996).

Side

\

n

552


Sor si

Aclnetobacter sp

. NCIMB 9871 PE

XanthobaCter autobophlcus

NCIMB 1081 1 WC Pseudomnas Sp. NCIMB 9872 PE Nooardla sp. NCIMB 11399 WC NV-2 WC (a black yeast) Pseudomonas puffda NCIMB 10007 M02 PE "Sharpless"Baeyer-Villiger. Lopp & d . , 1996

R or re Non-migrating group

I

85 2.5-diketocamphane 1.2-monaxygenaseand 3.6-dlketocamphane 1.2-monaxygenasefrom Pseudomnas puffda NCIMB 10007 (Moll Bolm ( s.9-coppercomplex. Bolm et al.. 1997

.

Fig. 26. R': flavin or acyl residue, RZ:flavin or proton. Schematic representations of enantiomeric intermediates for Baeyer-Villiger reactions.

ry then the same analysis can be made for the face across which the migrating group moves. Using the usual priorities: peroxide > hydroxyl (or ether) > non-migrating group the faces can be defined as si or re. These are equivalent to (S)-(84) or (R)-(85),respectively, as designated above. This analysis can be made whatever the mechanism of the subsequent reaction of the Criegee intermediate. The analysis gains relevance if anti-periplanar migration is a prerequisite for migration and if the configuration is fixed for a given system. If these crite-

ria are satisfied all that remains is to establish the diastereoselectivity for addition to the ketone. The ketone (86) (Fig. 27) has one highly accessible face and one highly hindered face and hence the diastereoselectivity of addition is guaranteed (KELLY et al., 1995a, b). When this ketone (86) was subjected to all the enzymes and reagents in Fig. 26, the lactones (87, 88) were formed as expected by the model. Examples of the Criegee intermediates for (S)configuration transformations are shown in Fig. 28. These have been selected to illustrate cases where the diastereoselectivity of addition of the peroxide can be justified on reasonable chemical grounds. Thus the cyclobutanones (89,92) undergo addition from the exoface.The strained ring ensures that both methylene and methine migration can occur. The cyclohexanones (95) undergo attack from the

6

I

87

Sor si

&

I

86

R or re

&

88

Fig. 27. Enantioselective Baeyer-Villiger reaction of the prochiral tricyclic ketone (86).


553

“K *Ro R0 - (yo m89

......

111

OH

t

JIMMT

.MNw-

93

92

R

0

9s

a

94

OOR e s OH

y

z

98

qR -*

R

98

Kelly etal.. 1996a (see also table 2)

91

90

t.“s”

&$

.P

qR OA

*

97

Taschner & Black, 1988 Taschner etal.. 1993

7-1

Taschner & Peddada, 1992

R

R

100

z?>oH 99

0

0 101

OOR

102

KOnigsberger e t d . 1991

BnO

103

Fig. 28. Examples of (S)-configurationCriegee intermediates of Acinetobacter sp. NCIMB 9871 CHMO. The dashed line indicates the migrating bond.

equatorial direction which is typical for larger and weaker nucleophiles. The bicyclo[2.2.1]hept-2-ene-7-one (98) undergoes attack from the side of the alkene bond as expected on stereoelectronic grounds.The enantioselectivity of the enzyme catalyzed Baeyer-Villiger reaction of bicyclo[2.2.l]heptan-2-ones is generally

poor. These substrates also give poor selectivity with peracids (HAMLEY et al., 1991;GRUDZINSKI et al., 1978; MEINWALD and FRAUENGLASS, 1960). The unsaturated analog (101) is an exception. Attack from the endo-face is facile and only one enantiomer is transformed with good enantiomeric excess, although some reserva-

554

I I Flavin Monooxygenases

tions need to be maintained because this is a whole cell biotransformation. The exact mechanism by which the 4a-hydroperoxyflavin (5) (Fig. 29), transfers oxygen

is still a matter of intense debate. It is appreciably more reactive than would be expected on the basis of the pK, of the departing alcohol (6) (BACHet al., 1994;BACHand Su, 1994).The

RAPP

H20

It

H

3

O

4

RAPP

PPP

I

104

RAPP I

106 Fig. 29. Accepted and proposed mechanisms for the CHMO catalyzed Baeyer-Viiliger reaction illustrated with the tricyclic ketone (86).


generally accepted sequence is shown in the top cycle of Fig. 29. A mechanism has been postulated (Fig. 29, lower cycle) in which the pK, of the peroxide is reduced by hydrogen bonding (105) and the ring system acts as an electrophile to trap the alkoxide (106). This intermediate has all the requisite orbitals aligned to promote migration in a stereodefined manner (KELLY,1996). Intermediates comparable to the trioxane (107)are formed during the ozonolysis of alkenes in alcohol solvents (SCHREIBER and LIEW,1983).An important consequence of this mechanism is that the (S)-Criegee intermediates (84) (Fig. 26) can only be formed on the si-face of the flavin ring and the (R)-Criegee intermediates (85) can only be formed on the re-face of the flavin ring. Perusal of the lists in Fig. 26 shows that the enzymes fall into two classes. The enzymes which utilize the (S)-Criegee intermediates (84) have FAD prosthetic groups and utilize

NADPH whereas those that utilize the ( R ) Criegee intermediates (85) utilize FMN as a prosthetic group and NADH as a reductive cofactor. This is supported by N-terminal analysis (Tab. 1). The significance of these speculations awaits clarification by determination of the X-ray crystal structures of these enzymes (MCGHIEand LITTLECHILD, 1996). Tab. 2 shows virtually all known biological Baeyer-Villiger reactions. The key names for on-line searchs of current work are S. M. ROBERTS,A. J. WILLETTS,and R. FURSTOSS, plus D. R. KELLY,C. J. KNOWLES, H. L. HOLLAND,and J. D. STEWART. The major compilations (DAVIESet al., 1989; DRAUZand WALDMA”, 1995) and recent reviews should be consulted for further details (ROBERTS and TURNER, 1992;ROBERTS and WILLETTS,1993;AZERAD,1995; FANGet al., 1995; CARREAet al., 1996;WILLEITS,1997;HOLLAND, 1997).

Tab. 2. Baeyer-Villiger Biotransforrnations

Monocyclics Gagnon etal., 1995b

R = “C& Acinetobacter s p . NCIMB 9871 WC M 0 1 P. puttda NCIMB 10007 M02 P.puttda NCIMB 10007 R = ‘C& Acinetobacter s p . NCIMB 9871 WC M 0 1 P. putida NCIMB 10007 M02 P. putida NCIMB 10007 R = Bn Aclnetobacter s p . NCIMB 9871 WC M 0 1 P.puCtda NCIMB 10007 M02 P. putldn NCIMB 10007 R = CHzOBn Acinetobacter s p . NCIMB 9871 WC M 0 1 P. putfda NCIMB 10007 M02 P. putida NCIMB 10007 R=X Aclnetobacter s p . NCIMB 9871 WC MO1 P. p~&& NCIMB 10007 M02 P. puttda NCIMB 10007

555

Config.,% ee, % yield (9. 17,68 ( 8 . 6 9 , 100 GC ( 8 ,54.93 GC

(9.82.57 (9. 15.40 (9.20.26

(9,55.89 ( 9 . 7 4 . 7 4GC ( 8 , 9 0 , 9 5GC (9.95,83 (8.7.38 (9, 14.69

‘R

556 Tab. 2.


Continued ~~

~~

Adger et al.. 1995, 1997 P. puttcia NCIMB 10007 M02 noh 2 2 2 1 1 1 1 2

R "C6H13

"C8H17 CHzCqEt CyCqEt

cyoAc

C&OSf%uMez CyOMEM

cycyoAc

*

b

ee %. yield %, config. 72. 36. S

77, 34, s 93. 30. R >98,27, R >98. 16, R 45. 18, R 90. 45. R 83. 34. R

65,30, R 61,49. R 89, 33, S 75.37, s 65, 19. S 22. 25. S 88. 47, S 75. 13, S

ee %.yield % 77, 15 88.33 84.39

ee %. yield Yo

MO 1 P.putlda NCIMB 10007a

58, 16 74.34 90. 11

9, 14 48.48 22.35

M02 P. putfda NCIMB 10007a

95.40 92,35 95.29

84.26 75.51 59.44

Acfnetobacter sp. NCIMB 9871 WC

ee N R 4% GC 92.30 GC 82,22 GC 60,40 GC

ee NR, 55 GC ee NR. 52 GC ee NR, 52 GC 71 ee. 40 GC

Acfnetobacter sp. TD63 WC

95,lOGC 90.30 GC 70,26 GC 44.45 GC

ee NR, 68 GC ee NR. 50 GC ee NR. 55 GC

"C8HI7

"C6H13 "C8H17

"C4% "C6H13 "C8H17

"C5H1 1 "C7H15 "C9H19 nCllH23

"C5H1 1 "C7H15

"C9H19 "CllH23

'

R

Organism or enzyme fraction P. putlda NCIMB 10007WC a

"C6H13

"C4%

A. .e\\

R

n

ee %. yield %. conflg.

aGagnon etal.. 1994 bAlphand etal.. 199Oab

R "C4%

&'a'\R

R

18.38 34.53 62.49

47. 50 GC

3 “Biological”Baeyer-Villiger Reactions

557

Tab. 2. Continued

Besetal.. 1996

I

(cf Grogan et al.. 1994)

% ee, % yield by GC 78.34 : 40,61 75, 52 : 3 8 . 4 3 ~

Pseudomonas sp. NCIMB 9872 WC Pseudomonas sp. NCIMB 9872 CPMO PE

oh

R

Mger e t d . , 1995, 1997 Beset d..1996

I

Pseudomonas sp. NCIMB 9872 CPMO PE

% ee. % yield by CC 42.61 : 68.39

aAlphand etal.. 1996

b Stewart et al.. 1996a

R Me Et Bu C6H13

C9Hl9 Ph Bn

Acinetobacter TD63 WC a Lactone : Ketone % ee,% yield. config. 61. 35. S: 35. 52. R 38. 60. S: >98. 10, R >98.6. ND : 10.37. ND 98. 23. S: 35. 25. R 85, 26, S: 42, 32. R >98, 40. R : 86.48. S >96. 22. R : 78.28. S

P.putfda NCIMB 10007 M02 * Lactone : Ketone % ee. % yield. config 38, 50. S: 51. 44. R 25, 52, S : 22, 26. R 74, 58. S: >98, 36. R 81, 28, S: 37. 54. R 64,18. S: 15.75. R >98, 13. R : 17, 80. S 28. 66, R : 83. 29. S

I

I I

R

I

I

Me Et h ‘h Bu my1

Acinetobacter NCIMB 9871 CHMO expressed in bakers’yeast Lactone : Ketone % ee, % yield, conflg 49.50. S:ND 95.79. S: >98. 69. R 97.54. S: 92.66, R >98.41. R : 96.46, S >98,59. S: 98.64. R 98. 59. R : >98. 58, S

I1 Flavin Monooxygenases

558

Tab.2. Continued a Taschner & Black, 1988 b Taschner et al., 1993 Aclnetobacter sp. NCIMB 9871 CHMO (cfstewart etal.. 1996b)

0

0

R, config, % ee. % yield Me, ( 8, >98 a Et,(97,>98. 84 h,(87, >98. 80b "Bu, (R)?. 52. 70 'Pr. (91. >98.60 'Bu, (9. >98. 17 OH, (R). 9.6. 73 OMe. ( 8 7 . 7 5 . 7 6 a CH20H. ( 9 ? . 9 8 , 8 0

9

%..

0

Taschner & Black, 1988; Taschner & Chen. 1991

.,\ti

0

d

Aclnetobacter sp. NCIMB 987 1 CHMO

a

* 0

> 98%ee. 73%yield

Taschner & Black, 1988 Acfnetobacter sp. NCIMB 987 1 CHMO

*

,,b..Q

""*e- ,,.I..

0

> 98%ee. 27%yield

111

0

--OH

-OH

-a*

Taschner & Black, 1988 Acinetobacter SD. NCIMB 9871 &HMO

I ( 0

0

'0

>98%ee. 88%yield

Taschner & Black, 1988

~

Acinetobacter sp. NCIMB 9871 CHMO

0

> 98%ee, 25%yield

0

3 "Biological"Baeyer-Villiger Reactions Tab. 2.

Continued

-F

--

0""

Alphand & Furstoss. 1992a

Acinetobacter sp Acinetobacter s p

. NCIMB 9871 WC

80%yield 95%yield

.TD 63 WC

rac

Alphand & Furstoss. 1992a

Acfnetobacters p Acinetobacter s p

A

. NCIMB 9871 WC . TD 63 WC

P

rac

bo

*

73%yield 66%yield

Acinetobacter sp. NCIMB 9871 CHMO Abril etal.. 1989 75%yield

Y --

v -

v -Alphand & Furstoss. 1992a

*

Mc

Acinetobacter s p Acinetobacter sp

. NCIMB 9871 WC .TD 63 WC

curuularialunata NRRL 2380 WC

*

Ouazzant-Chahdi et al.. 1987

OH rac

All > 98%ee; % yield 45 : 20 43 : 20

#!fo&A A OH

S

OH

97%ee. 83%yield : ee NR, 63%yield

559

II Flavin Monooxygenases

560

Tab. 2. Continued

Alphand & Furstoss. 1992b rat Acinetobacter sp Acfnetobacter sp

. NCIMB 987 1WC

*

Fractions from P. p W a NCIMB 10007

% ee, % field 50 : 95 (86%) 89.53 GC : 99,47 GC 82.57 GC : 100.43 GC 10. 17GC: 72, 13GC

wc

MO 1 2,5-DKCMO 3.6-DKCMO

a Shipton et al , 1992 b Crogan et al.. 1993a c Kelly et al.. 1996a d Alphand et d ,1989 e Alphand & Furstoss, 1992b f Camell & Willetts. 1990. 1992

rac

g L ~ M& Knowles, 1994 cf Wrlght et al.. 1994b Aclnetobacter sp . NCIMB 9871 WC Aclnetobacter sp . NCIMB 9871 CHMO Acinetobacter sp . NCIMB 9871 CHMO Acfnetobacter sp . NCIMB 9871 CHMO PE Aclnetobacter sp . NCIMB 9871 CHMO WC Acfnetobacter sp .TD 63 Pseudomoms putlda NCIMB 10007 M02 Cwvularfahtermedta WC Pseudomonasputida AS 1 WC AcfnetobacterJunlt WC Rhodococcus coprophilus WT 1 WC Rhodococcus fasciens WT 13WC

0

19 : 20 21 : 25

.TD 63 WC

Gagnon et al., 1994; Beecher et aL , 1996

rac

Au >95% ee; % yield

0 ee, O h yield 93.7 : 99.2 (67) 94.3 : 98.9 (98) a*c 92 : 95b >98.50 GC : >98. 50 GC >95,30 : >95.33 >95.28 : >95,35 35,72 GC : 95.28 GC 86 : 98 (95%) 89 : 99.6 (86%) a*g 87.4 : 99.3 (79) a 89.1 : 99.6 (67) a 89.1 : 99.0 (63) a

Oh

asc

561


Tab. 2. Continued

A&tobacter

Peracids

sp. NCIMB 987 1 WC

P. puffria NCIMB 10007 WC rat

..

Grogan et al 1993b

*9

>96O/6ee, 28%: >96%ee,28%yield rac 1 : 10

* : /

80% ee: 95%ee (63%yield)

Grogan et al., 1993a. 1993b

*

do (yJ: YOee. Oh yield

Fractions from P. putlda NCIMB 10007

wc

80 : 95. (731 89 : >75, (100 GC) 60 : 95. (100 GC)

M01 M02

rat

ND : 29%ee,63%yield 1 : 20 41%yield. ee NR ND : rac. 56%yield

Achtobacter sp. NCIMB 9871 WC

Pseudomoms sp. NCIMB 9872 WC Peracids

ao rac

+npr

2 . k

z

-

CQ

,“Pr

Wright et aL , 1994b

Xanthobacter autotmphfcus

*

m o s “Pr

NCIMB 1081 1 WC

0

Oh ee. % yield 100, 43.4 : 100, 56.6


562

Tab. 2. Continued

rac Aclnetobacter SD. NCIMB 987 1

Peracids

o aracf o

>98%yield 31%yield : 96%ee 30% yield rac 61Oh yield : ND

wc

Petit & Furstoss, 1993 Acfnetobacter s p

. NCIMB 9871 WC 97%ee. 35%yield : > 98%ee. 35%yield

(Yo

Petit & Furstoss, 1993

Aclnetobacter sp

rac

en* rac

.NCIMB 9871 WC

0

*

>98%ee. 33%yield : >98%ee, 41% yield

0

Petit & Furstoss. 1993. 1995 Acfnetobacter sp

. NCIMB 9871 WC 90%ee. 35%yield : >98%ee. 32%yield

0

Alphand & Furstoss. 1992b

rac Acinetobacter sp Acfnetobacter s p


% ee;% yield 61.30 : >95. 18 53.24 : >95, 12

0

Alphand & Furstoss. 199213

rac Aclnetobacter s p Acfnetobacter sp

. NCIMB 9871 WC

. TD 63 WC

All >95Oh ee; % yield 27 : 35 21 : 37

0

563


Tab. 2. Continued Kbnigsberger et QL, 1990

*

C u l M m m n destructans ATCC 11011 wc

-

rac

27%ee : 28%ee (32%yleld)

Alphand & Furstoss, 1992b

rac Acfnetobacter s p Acinetobacter s p


Petit & Furstoss. 1993 Acfnetobacter s p

0

% ee:% yield 86,27 : >95.22 92. 20 : >95.26

. NCIMB 9871 WC

* v r\

33%ee. 60%yield : >98%ee. 18%yield

Petit & Furstoss. 1993 Acinetobacter s p

rac

. NCIMB 9871 WC 0

70?!ee. 33%yield : >98%ee, 33%yield

Bicyclo[2.2. llheptanones

La-

0

aAbril e t a f . . 1989 b Sandey & Willetts 1989

*

Acinetobacter sp. NCIMB 987 1 CHMO Pseudomom sp. NCIMB 9872 WC

Peracids

0

rac 81% : o%= 38 : 1 (100%yield CC) rac 9 : 1 (100%yield GC)

Grogan et al.. 1993a P. putfda NCIMB 10007 M02

14%ee : ee NR

0

564


Tab. 2. Continued Grogan et al., 1993ab Gagnon et al., 1994

Acinetobacter Sp . NCIMB 9871 CHMO Fractions from P.putlda NCIMB 10007

wc

MO 1 2.5-DKCMO 3.6-DKCMO

% ee,% yield

racemic : racemic 53,60 GC : 70.40 GC 53% ee : ee NR 60.20 GC : 20.80 GC >90.48 GC : >90,52 GC

Khigsberger et aL , 1990 Cyllndrocarpon destructans ATCC 11011 WC

-0

rac, 34% yield

Gagnon et al.,1995a Fractions from P. putlda NCIMB 10007

0

MO 1 2.5-DKCMO 3.6-DKCMO

0

>95% ee. 11%yield 0% yield >95% ee. 33%yield GC

Gagnon et al., 1995a

Ac%

0

P. putida NCIMB 10007 and extracts % yield, all >95%ee

wc MO 1 M02 2.5-DKCMO 3.6-DKCMO

0

27 35 39 cc 35 0

Gagnon et al.. 1995a

Bn?+

0

P. putida NCIMB 10007 and extracts % ee. % yield wc 95,ll MO 1 95.39 M02 87.50 GC

0


Tab. 2.

Continued

Gagnon etal., 1995a

rac

OH

*

Horse liver alcohol dehydrogenase P. putfda NCIMB 10007 M 0 1

Ac%

0

>95% ee. 15% yield GC

Acinetobacter sp. NCIMB 9871 FC

ee N R 50% yield GC : ee NR 15% yield GC

KOnigsberger et aL. 1991

0 Acinetobacter sp. NCIMB 9871 CHMO WC

BnO

OH 99% ee 35% yield

85% ee 26% yield

100 : 1 (11% yield) 1:4

Acfnetobacter sp. NCIMB 9871 WC Peracids

Acinetobacter sp. NCIMB 9871 WC Br

Lwitt etal., 1990b

*

89% ee 8% yield

J5 f b - 0

0

Br

Br

36%. >95% ee : 30%. >95O/6ee Acinetobacter sp. NCIMB 9871 CHMO Abril e t a l . . 1989

> 89% yield

0

565

566


Tab. 2. Continued

0

Acfnetobacter sp. NCIMB 9871 CHMO

Abril et al.. 1989

40%: 40%yields Acfnetobacter sp. NCIMB 9871 CHMO Abril etal.. 1989

0

* 67% : 9%

0 Taschner & Peddada. 1992

I R

R

Acfnetobacter sp. NCIMB 9871 CHMO

*

H, 80,62

Me, >98,70 Et. 93,83 CH20CH2* >98*74

8 I R

R

R % ee. % yield

R O h ee,% yield

0

(CH,),, 97.80 (CH,),, >98.78 (CH,),, >98.57 (CHo)s, 87,55

Polycyclics

Acfnetobacter s p

. NCIMB 9871 PE

Xanthobacter autotrophlcus NCIMB 1081 1 WC Pseudomonas sp . NCIMB 9872 PE Nocardfa sp. NCIMB 11399WC NV-2 WC (a black yeast) M02 P.putfda NCIMB 10007 PE

ee. all 100% yield GC >98 87.5 >98 >98 34 >98

Oh

567


Tab. 2. Continued

% ee. %yield

%02,AcOH. water

rac, 91

wc

21.36 60.75

P.putldn NCIMB 10007grown on (+/-)-camphor

M02. CE P. putlda NCIMB 10007grown on (-)-camphor WC (Kent) WC (Exeter) M02. CE P. putIda NCIMB 10007 grown on (+)-camphor WC (Kent) WC (Exeter) MO 1 2.5-DKCMO, PE 3.6-DKCMO. PE M02. CE et d.1997a ( S.$copper bis(oxazolinecomplex), Bolm

56.5. ND 30.39 89.78 5.5. ND 13.39 >98,72 >98,80 >98.86 77.82 91.62

Co-factor recycling and multi-enzymesystems Acinetobacter sp.

0

11.4g. lOOmmoles

rac 81% yield NADPH

NAD a

HO H&

Glucose-6-phosphatedehydrogenase

HO 0

Abril e t d . . 1989

HO*LOH H HO Glucose-6-phosphate

568


Tab. 2.

Continued

mc

86% ee, 27% yield : ee NR. 14% yield

Acfnetohter sp. NCIMB 9871 PE

0

0

P. punda NCIMB 10007 M01 a Grogan et al.. 1992, 1993b b Gagnon et aL , 1994

rac

*

NADH recycling enzyme Pseudomonas sp . NCIMB 9872 alcohol dehydrogenase Horse liver alcohol dehydrogenase

a

0

Q

*

Gagnon et aL. 1994

rac

% ee. % yield

Pseudomonas cepacia lipase Horse liver alcohol dehydrogenase Pseudomonas putida NCIMB 10007 M01

Acfnetobacter sp. NCIMB 9871 PE

OH

93.9 GC : 100.33 GC

56% ee, 16% yield

Gagnon et al.. 1995a rac

% ee, % yield 79, 59 GC : >98*41 GC 79. 58 CC : 97.42 GC

*

Horse alcohol10007 dehydrogenase P. p u wliver a NCIMB M01

Aco&

0

>95% ee. 15%yield GC

3.3 Large Scale Baeyer-Villiger Reactions Baeyer-Villiger reactions on a large scale present problems even with abiotic reagents (COLEMAN et al., 1997;FLISAK et aI., 1993).The

principal problem with enzyme catalyzed Baeyer-Villiger reactions is cofactor recycling, particularly of NADPH. Generic solutions to this problem are described in Chapter 9, this volume, and examples from the laboratory are shown at the end of Tab. 2. Several sulfides and the bicyclic ketone (107) (Fig. 30) were oxid-

569

4 Asymmetric Sulfoxidations by Flavin Dependent Monooxygenase Enzymes

Ps. Formate pufjda deli) NCIMB drogenase 10007 M 0 1 EX-107

0

w-0

108

d: 109

7 2% ee : 98% ee, > 98% conversion Fig. 30. Large scale Baeyer-Villiger reaction of bicycIo[3.2.0]hept-2-en-6-one(107) (PASTA et al., 1996).

ized by the M 0 1 fraction from Pseudomonas putida NCIMB 10007 and polyethylene glycol

The Baeyer-Villiger oxidation of 4-methylcyclohexanone (110) (Fig. 31) was carried out bound NADH retained within a membrane using Acinetobacter sp. NCIMB 9871 CHMO with a molecular weight cut-off of 3000. NAD and a formate dehydrogenase engineered to was reduced by formic acid catalyzed by for- accept NADPH. The dioxygen concentration mate dehydrogenase. Virtually, all the activity of the stirred reaction was too low to facilitate of the system was retained over four runs efficient conversion and the reactor was aeratwhereupon a substantial decrease in the deed without bubbles through a thin wall silicon gree of conversion occurred (PASTAet al., tube. The enantiomeric excess of the lactone 1996).The ketone (107)and the lactones (108, (111)was very high (RISSOM et al., 1997) and 109) have been used in the synthesis of (+)- comparable with that found in the laboratory multifidene, ( + )-viridiene (marine brown al- scale reaction (TASCHNER and BLACK,1988; et al., 1993). A similar process has gae pheromones) (LEBRETON et al., 1996, TASCHNER 1997),cyclosarkomycin (ANDRAU et al., 1997), been run with cyclohexanone or phenyl methyl sulfide, Acinerobacter sp. NCIMB 9871 and innumerable prostaglandin syntheses. CHMO, and polyethylene glycol bound NADPH retained within a membrane as described above. Thermoanaerobium brockii dehydrogenase was used to catalyze the regeneration of NADPH with isopropanol as the re110 ductant (SECUNDO et al., 1993a).

()

0

Acinetobacter sp. NCIMB 9871 CHMO NADPH, Formate dehydrogenase, IlCOzH

4 Asymmetric Sulfoxidations by Flavin Dependent Monooxygenase Enzymes 4.1 Introduction

0

Fig. 31. RISSOM et al. (1997).

Flavin monooxygenase enzymes catalyze the transformation of sulfides to sulfoxides. This use of enzymes to perform sulfoxidations has been of considerable interest due to the importance of sulfoxides as chiral auxiliaries in

570

I I Flavin Monooxygenases

asymmetric synthesis. Classical methods for 1988, 1992, 1997). Large scale oxidations of the preparation of chiral sulfoxides include the sulfides are described together with large scale Sharpless/Kagan asymmetric sulfoxidation, in Baeyer-Villiger oxidations. Many non-flavoenzymes are capable of oxiwhich tertiary butyl hydroperoxide is used in dizing sulfides to sulfoxides, most notably the the presence of a titanium complex (Ti(Oil'r)J, et al., diethyl tartrate, and water to afford sulfoxides iron-heme chloroperoxidases (COLONNA with a high degree of enantiomeric purity 1988,1992).These are discussed in HOLLAND'S (SATO et al., 1997; CASHMANet al., 1993; reviews (1988,1992,1997). BRUNELand KAGAN,1996). Chiral oxaziridines have also been shown to perform asymmetric sulfoxidations with a high degree of 4.2 Asymmetric Sulfoxidations stereoselection (DAVISet al., 1993;DAVISand by Monooxygenase Enzymes CHEN,1992).Treatment of bovine serum albumin (BSA) with sodium metaperiodate or hy- Occurring in Bacteria drogen peroxides furnishes a reagent (a pseudo-enzyme?!)which converts sulfides to sulf- 4.2.1 Acinetobacter calcoaceticus oxides enantioselectively. Moreover, kinetic resolution of the sulfoxides by further oxida- NCIMB 9871 Cyclohexanone tion to the sulfone resulted in excellent enan- Monooxygenase (CHMO) tiomeric excesses (>90% ee) (SUGIMOTO et The chemistry of CHMO is described in deal., 1981). BSA and dimethyldioxirane can be exploited in a similar way to give sulfoxides tail in the prior section on Baeyer-Villiger reactions. The active reagent is almost certainly with up to 84% ee (COLONNA et al., 1991). While such methods provide efficient means the 4a-hydroperoxyflavin (5) (Figs. 2 and 32) of performing asymmetric sulfoxidations,they which reacts in essentially the same way as a have proved effective only within a limited peracid (e.g., ALI et al., 1997). As with the range of substratesThus, the use of enzymatic Baeyer-Villiger reaction of this species the systems to yield chiral sulfoxides that are not precise nature of the oxygen transfer step is accessible via chemical means is an attractive poorly understood (BACHet al., 1994; BACH et al., 1995), although alternative. The flavin dependent monooxy- and Su, 1994; BONCHIO genase enzymes that have been reported to the mechanism and kinetics have been perform asymmetric sulfoxidations can be di- thoroughly studied (RYERSONet al., 1982; vided into two broad categories, those that are WALSHand CHEN,1988; GHISLAand MASSEY, present in bacteria and those that are found in 1989). Early studies indicated that CHMO catamammalian tissues. The bacterial enzymes have been extensively reviewed (O-ITOLINAet lyzes the oxidation of sulfides to sulfoxides and WALSH, al., 1995; COLONNA et al., 1996; HOLLAND, (LIGHTet al., 1982; BRAUCHARD 1985; LATHAMet al., 1986; LATHAMand WALSH,1987).CHMO generally shows a preference for Baeyer-Villiger reaction over sulRqPP fide oxidation as demonstrated by the BaeyerVilliger oxidation of 4-thiacyclohexanone (112)(Fig. 33) (LATHAM and WALSH,1987). The enantioselectivity of oxidation of prochiral sulfides by CHMO under standard conditions and relative rate studies (Tab. 3) (CARREAet al., 1992; SECUNDO et al., 1993b) enabled an active site model to be proposed (OTTOLINAet al., 1995; COLONNA et al., 1996; HOLLAND, 1997).The active site model is based Fig. 32. Proposed mechanism for sulfoxidation by on the cubic space approach, whereby the active site of the monooxygenases is modeled by Acinetobacter sp. NCIB 9871 CHMO.

4 Asymmetric Sulfoxidations by FIavin Dependent Monooxygenase Enzymes

112

113

Fig. 33. Preferential Baeyer-Villiger oxidation by Acinefobacfersp. NCIB 9871 CHMO (LATHAM and WALSH,1987).

a series of binding pockets. The dimensions of these binding pockets are determined by separating the products (sulfoxides) into two groups according to their absolute configuration ( R or S ) and superimposing the minimized energy conformations of the two groups

571

of sulfoxides to give rise to the three dimensional model of the active site (Fig. 34). The three dimensional model is composed of three binding pockets, a main pocket, a small hydrophobic pocket, and a large hydrophobic pocket. In addition, the model allows the catalytic site to accommodate the hydroperoxide such that the approach of the oxygen atom to the substrate is from the top face of the model. The following guidelines must be applied when using the model to predict absolute configurations: (1) The sulfur atom must be aligned along the S=O axis in the catalytic site. (2) The substrate molecule is divided into two fragments, a large fragment and a small fragment. If the aromatic rings (large fragment) occupy the main

Fig. 34. Cubic active-site model for sulfoxidation by Acinetobacter sp. NCIB 9871 CHMO. The catalytic site is represented by a circle, the main (M), hydrophobic large (HL).and hydrophobic small (H,) pockets are also shown.

572

I1 FIavin Monooxygenases

Tab. 3. Cyclohexanone Monooxygenase Catalyzed Oxidation of Sulfides to Sulfoxides (OTTOLINA et al., 1995)

Sulfide

Conee Configversion [YO]uration

Ph-S-Me Ph-S-Et Ph-S-nPr Ph-S-Vinyl Ph-S-'Pr

88 86 54 73 90

99 47 68 99 12

Bn-S-Me Bn-S-Et Bn-S-"Pr Bn-S-"Bu Bn-S-'Bu Bn-S-'Pentyl Bn-S-Hexyl

97 80 90 98 90 95 75

54 67 96 15 90 80 56

S S S

p-(Me)-Bn-S-Me p-(Et)-Bn-S-Me p-('Pr)-Bn-S-Me p-('Bu)-Bn-S-Me

97 91 88 87

5 8 4 23

R R R R

o-(Me)-Ph-S-Me rn-(Me)-Ph-S-Me p-(Me)-Ph-S-Me p-(Me)-Ph-S-Et p-(Me)-Ph-S-iPr P-Naphthyl-S-Me

90 90 94 89 99 50

87 40 37 89 86 53

R R

Ph-S-CH2-CN 90 Ph-S-CHZ-CH2-CN 61 p-(Me)-Ph-S-CH>-CN 95 p-(Me)-Ph-S-CH2-CHz-CN 71 Ph-S-CHZ-CH2Cl 75 p-(Me)-Ph-S-CH2-CH2C1 65 o-C1-Ph-S-Me 35 p-C1-Ph-S-Me 78

92 14 98 61 93 93 32 51

s

t-Bu-S-Me

99

R

R R S

R R R

S S

s

s

(4) In cases where the aromatic rings do not fit the main pocket, they will occupy the large hydrophobic site. The model was used to predict and to draw comparisons with the results obtained for sulfoxidations performed by the cyclohexanone monooxygenase. Predictions using the model were found to be in agreement with the experimental results obtained. The absolute configuration of phenyl alkyl sulfoxides where the alkyl chain was methyl, ethyl, and vinyl, respectively, were found to be R , with enantiomeric excesses ranging from 47%-99% (Fig. 34). In the case where the alkyl side chain was npropyl, there were found to be two possible

a

S S

s R S

S

S

s s R

U

98

pocket, the resulting sulfoxide will possess the (R)-configuration. (3) If the small fragment of a substrate molecule occupies the small hydrophobic pocket the resulting sulfoxide will possess the (R)-configuration. However, if the small fragment of the molecules does not fit into the small binding site, the fragment will then occupy the main pocket to afford sulfoxides that possess the (S)-configuration.

C

Fig. 35. Active site model depicting the preferred interactions of the phenyl propyl sulfides (OTTOLINA et al., 1996). et al., 1995;COLONNA


conformations for the substrate, the first being where the phenyl ring occupies the main pocket while the n-propyl chain occupies the small hydrophobic pocket (Fig. 35a) and the second being where the phenyl ring occupies the large hydrophobic pocket and the n-propyl chain occupies the main pocket (Fig. 35b).The model suggests that the latter of the two possibilities would be the favored conformation, resulting in the sulfoxide possessing the (S)-configuration. Experimental results obtained for the sulfide gave an enantiomeric excess of 68% in favor of the (S)-configuration. When the n-propyl side chain is replaced by an isopropyl group the enantiomeric excess dramatically decreased to 12%, in favor of the (R)configuration. The low enantiomeric excess observed is a result of the unfavorable fit of the isopropyl group into the small hydrophobic pocket (Fig. 3%). The model has been used to correctly predict the stereochemical outcome of other substrates, ranging from benzyl alkyl sulfides, p-alkylbenzyl methyl sulfides, alkylphenyl alkyl sulfides, cyano and chloro substituted phenyl alkyl sulfides to dialkyl sulfides (OITOLINA et al., 1995; COLONNA et al., 1996; HOLLAND, 1997). The oxidation of 1,3-dithioacetals via chemical means (SharplesslKagan method) has not been successful, giving rise to products with low enantiomeric excesses. In contrast, the oxidation of 1,3-dithiane (114) with the cyclohexanone monooxygenase afforded the corre-

573

sponding (R)-sulfoxide in good yield and with excellent enantioselectivity (Tab. 4) (COLONNA et al., 1995). Further studies based on the oxidation of racemic 1,3-dithiane monosulfoxide revealed that the (S)-sulfoxide was oxidized to the monosulfone at a faster rate than the (R)enantiomer, thereby resulting in kinetic resolution of the enantiomers. Similar yields and enantioselectivities were observed on the oxidation of 1,3-dithiolane (115) and bis(methy1thio)methane (116) to the corresponding ( R ) monosulfoxides. CHMO is inactivated by thiolactones (120-123)(Fig. 36). Typically, 10-100 turnovers of the thiolactones (120, 121) occur before the

120

121

122

123

Fig. 36. Thiolactone inhibitors of CHMO.

Tab. 4. Oxidation of 1,3-Dithioacetals to the Corresponding Monosulfoxides and Monosulfones by Acineet al., 1995) tobacter sp. NCIMB 9871 CHMO (COLONNA Yield

(R)-Monosulfoxide

[%I

ee [%]

Monosulfone Yield [Yo 1

1.3-Dithiane (114) 1.3-Dithiolane(115)

bis(Methy1thio)methane(116)

n

svs

n

94

>98

6

0 T

92

>98

8

sv%

574


enzyme is irreversibly inactivated (LATHAM and WALSH,1987).The thiolactones (122,123) were designed to model the lactones (108,109) (Fig. 30) as enantioselective inhibitors, to determine if the enantiomeric ketones (107) were processed at different active sites. Both thiolactones inhibited formation of the lactones (108,109)equally indicating that there is only one kinetically significant active site in CHMO (WRIGHTet al., 1994a). CHMO also oxidizes selenides to selenoxides (LATHAMet al., 1986), amines to amine N-oxides, alkyl boronic esters into borates with retention of configuration (LATHAMand WALSH,1986; cf. AHRENSet al., 1991), and phosphines into phosphine oxides (WALSHand CHEN,1988) with retention of configuration (KELLY, WILLETTS,and HOLLAND, unpublished data).

4.2.2 Asymmetric Sulfoxidations by Flavin Dependent Monooxygenase Enzymes Present in Other Microorganisms The oxidation of phenyl methyl sulfide (124) with Acinetobacter sp. NCIMB 9871 CHMO and with Pseudomonas sp. NCIMB 9872 whole cells (containing CPMO) gives the (R)- and (S)-sulfoxides (125a, l26a) (Fig. 37), respectively, with virtually 100% enantiomeric excesses whereas in Baeyer-Villiger reactions these enzymes have congruent enantioselectivity. This argues that the constraints for the two reactions are quite different (KELLYet al., 1996b).

6 ..-

00.2

/

R

R

124

125

R

126

Fig. 37. Oxidation of methyl phenyl (p-tolyl) sulfide (W).

Camphor grown Pseudomonas putida NCIMB 10007possesses the ability to perform asymmetric sulfoxidations with moderate enantioselectivity (Tab. 5 ) (BEECHERet al., 1994). It has been suggested that the ability of Pseudomonas putida to catalyze sulfoxidations can be attributed to either or both of the NADH dependent or the NADPH dependent monooxygenases occurring in the microorganism. Preliminary cytochrome P450 monooxygenase inhibition studies have indicated that sulfoxidation is unlikely to be due to the NADH dependent cytochrome P450 monooxygenase enzyme (camphor 5-exo-hydroxylase) present in camphor grown P putida. New activities can be found in surprisingly mundane places. Bakers’ yeast oxidized methyl p-tolyl sulfide (124b)to the corresponding sulfoxide (125b) in 60% yield and 92% ee (BEECHER et al., 1995; TANGet al., 1995). This simple procedure is certain to find many applications. In the 1970s MCCAPRAand HART(1976) reported the use of bacterial luciferase (Phofobacterium phosphoreum) for the oxidation of methyl decyl sulfide to the corresponding sulfoxide (absolute configuration unknown). Interest in bacterial luciferase results from its ability to emit light in the presence of NADH, FMN reductase, and decanal. Luciferase follows the normal sequence for the Baeyer-Villiger oxidation of aldehydes to carboxylic acids (Figs. 2 and 29). Indeed the 4a-hydroperoxyflavin ( 5 ) was detected by I3C-NMR in this enzyme (GHIsLa et al., 1978) and the enzyme catalyzes Baeyer-Villiger reactions of tridecanone (VILLAand WILLE’I-TS, 1997). However for

Tab. 5. Oxidation of Alkyl Aryl Sulfides Catalyzed by Pseudomonas puridu Washed Whole Cells (BEECHER et a]., 1994).All Sulfoxides are (s)-(-) Sulfide Substrate

ee

1% 1

Yield [Yo 1

Methyl phenyl sulfide Ethyl phenyl sulfide Isopropyl phenyl sulfide Methyl p-tolyl sulfide Isopropyl p-tolyl sulfide

27 6 61 64 91

87 83 29 98 87


reasons which are unclear, the breakdown of the peroxide-aldehyde adduct (cf. 104, 106; Fig. 29) to give hydroxyflavin (6) and the carboxylic acid is accompanied by the emission of light (490 nm) (FRANCISCO et al., 1996; AHRENS et al., 1991) which is not observed in the Baeyer-Villiger reactions or sulfide oxidation.

575

studied in some detail (ZIEGLER,1988; POULand ZIEGLER,1979; JONESand BALLOU, 1986; BEATYand BALLOU, 1981a, b) and follows the normal sequence for flavoenzymes (Figs. 2 and 29). Microsomal FMO has a FAD prosthetic group, it is NADPH dependent and a functional active site model has been proposed (CASHMAN, 1995). Simple dialkyl and alkyl aryl sulfides give predominantly the (S)sulfoxide, however, in the presence of n-octyl4.3 Asymmetric Sulfoxidations by amine, the rate of oxidation is often increased and sometimes the enantioselectivity is invertFlavin Dependent Monooxygenase ed. This apparently results from binding of n(FMO) Enzymes Occurring in octylamine to a space in the enzyme, which otherwise accommodates the sulfide. Other Mammalian Tissue members of the FMO family display different Pig liver flavin dependent monooxygenase stereoselectivities. Comparative studies performed on the oxi(hepatic FMO) was first isolated and purified to homogeneity by ZIEGLER in the late 1970s dation of various sulfides by rabbit liver cy(ZIEGLER and POULSEN,1978; CASHMAN, tochrome P450 and pig liver FAD dependent 1995). One of the first applications of the mi- monooxygenases showed that the latter of the crosomal FMO was in the oxidation of amines two enzymes exhibited a greater degree of to the corresponding N-oxides. However, later stereoselectivity (Tab. 6) (TAKAHASHI et al., studies on the microsomal FMO proved that it 1978; WATANABE et al., 1980,1982;FUKUSHIMA was a far more versatile enzyme than original- et al., 1978). ly envisaged. In addition to performing oxidaSulfoxidations using rat liver cytochrome tions on secondary and tertiary amines, (ZIEG- P450 and hog liver FAD dependent monooxyLER, 1980, 1988; DAMANI,1988; CASHMAN, genases have also been extensively studied 1988) the microsomal FMO was found to be (LIGHTet al., 1982).When ethyl p-tolyl sulfide capable of catalyzing the oxidation of a broad was the substrate, the rat liver cytochrome spectrum of substrates encompassing hydra- P450 monooxygenase was found to give prezines (PROUGH et al., 1981), phosphines (SMY- dominantly the (S)-sulfoxide while the use of SER and HODGSON, 1985), boron containing hog liver FMO resulted in the formation of the compounds (JONESand BALLOU,1986), sul- (R)-sulfoxide as the major product. Hog liver fides (ZIEGLER,1980, 1990), and selenides FMO was also shown to oxidize the nonster(ZIEGLERet al., 1992).Further studies showed oidal anti-inflammatory agent sulindac (127) that in addition to the hepatic form of FMO, (Fig. 38) to the corresponding (R)-sulfoxide there also existed a pulmonary form of FMO (128)with a high degree of enantioselectivity found in lung tissue that possessed different (LIGHTet al., 1982). properties to the hepatic FMO (WILLIAMS et The stereoselectivity of the cyclohexanone al., 1984;TYNES et al., 1985;HLAVICA and GOL- monooxygenase (CHMO) from Acinetobacter LY,1991; LOMRIet al., 1993).These two forms sp. NCIMB 9871 has been compared with that were subsequently referred to as FMO 1 of hog liver FMO (LIGHTet al., 1982).These (hepatic form) and FMO 2 (pulmonary form); studies were based on the use of ethyl p-tolyl later additions to the family of mammalian sulfide as the substrate and showed that the FMOs included FMO 3, FMO 4, and FMO 5. microsomal FMO was able to carry out sulfoxThe gene for pig liver FMO 1 was cloned and idations on the substrate with opposing stereoexpressed in E. coli (LOMRIet al., 1993), how- specificity to CHMO. Thus, while CHMO gave ever unexpectedly, the stereoselectivity of sul- predominantly the (S)-sulfoxidein 73% ee, the fide oxidation by cloned pig liver FMO was microsomal FMO from hog liver gave the ( R ) less than with natural FMO. The catalytic sulfoxide in 85% ee. Further oxidation of the mechanism of the pig liver FMO has been sulfoxides to the corresponding sulfones was SEN

576


Tab. 6. Biotransformation of Sulfides by Isolated Enzymes (HOLLAND, 1992)

Rabbit Liver Cytochrome P-450 ee

Substrate

["/.I

Config.

54

-

["/.I

66

Config.

R

-

-

-

32

S

20

12 50 96

S

47

14

-

R

R

-

-

-

10

64

R

127 Sulindac

sulfide

It

ee

-

22

Reduction in vivo

Pig Liver FAD Monooxygenase

Oxidation Pig liver FMO

128 Suliiidac

sulfoxide

Fig. 38. Redox reaction of sulindac (LIGHTet al., 1982).

also investigated. It was found that considerably less sulfone was formed with the microsoma1 FMO than with the cyclohexanone FMO.

Overnight incubations of racemic sulfoxide with the microsomal FMO showed preferential oxidation of the (S)-sulfoxide to the corresponding sulfone and enrichment of the residual sulfoxide (substrate) with the @)-enantiomer. The low level of sulfone formed when using microsomal FMO can be accounted for by the fact that sulfoxidation using the microsoma1 FMO results in the formation of predominately (R)-sulfoxide and thus there is a low abundance of the (S)-sulfoxide, the required substrate for further oxidation. The potential of flavin monooxygenases to stereodifferentiate between geminal lone pairs on a prochiral sulfur atom and geminal sulfur atoms on prochiral carbon atoms has been investigated with 2-methyl-1,3-benzodithiole (Fig. 39) (CASHMAN et al., 1992). Chemical oxidation of 2-methyl-1,3-benzodithiole (129) to the monosulfoxide resulted in the formation of the trans-enantiomers (1S,2S)-(132) and (1R,2R)-(133).In contrast, oxidation using the fungus Aspergillus foetidus gave predominantly the cis-sulfoxides (87% yield) with an enantiomeric excess of 93% in favor of the (1S,2R) enantiomer (131). Similarly intact cultures of Pseudomonas putida UV4 gave cis-(1S,2R)sulfoxide (131) as the major product (100% ee). The enzyme responsible for sulfoxidation is thought to be either a flavin containing monooxygenase or a dioxygenase. Oxidations using hog liver and rabbit lung flavin-containing monooxygenases exhibited a similar preference for the pro-R sulfur atom and the for-

577

6 References

pro-S

lt

pro-R

129

(+)-cis-(1R,,S)

(-)-cis-(1S,2R) (-)-tr3f?S-(1St2S) (+)-tmns-( 1R,2R)

130

13 1

132

133

(CASHMAN et al., 1992). Fig. 39. Structures of cis- and trans-2-alkyl-l,3-benzodithiole-l-oxides

mation of the cis-(lS,2R)-sulfoxide (131),with enantiomeric excesses of 86% and loo%, respectively (CASHMAN et al., 1989,1992).

5 Conclusions Flavin monooxygenases display a reactivity similar to that of peracids, except that they are not capable of epoxidation. The reactions are highly enantioselective and can be performed on a large scale. Steroid mono-oxygenases and luciferase have considerable untapped potential. Yeast flavoenzymes and CHMO expressed in yeast may reduce the practical problems which attend the use of these enzymes. Flavin monooxygenases are the nearest thing to chiral hydrogen peroxide!

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Index

A AA see Aspergillus acylase I abzymes 269f acetone, production volumes 1910-1929 9 N-acetyl cysteine, resolution of 18 acetylcholine, hydrolysis by acetylcholinesterase 212 acetylcholinesterase 212ff - asymmetrization of cyclic meso-diesters 212f - - products 213 acid anhydrides, in lipase-catalyzed acylations 58 Acinefobacfer, as reagent for biological BaeyerVilliger reactions 546 - Baeyer-Villiger biotransformations 555ff - cyclohexanone monooxygenase. Criegee intermediates 553 - - mechanism of the Baeyer-Villiger reactions 548 - immobilization of 298 - isolation from elective culture 547 - nitrilase activity 298 Acinetobacfer jnnii, Baeyer-Villiger biotransformations 560 Acinefobacter sp. NCIMB 9871, sulfoxidations by 574 acrylamide production. enzyme process 302 acrylic acid production, enzyme process 302 - from acrylonitrile 311f acrylonitrile, hydrolysis to acrylamide 284f active site model, for pig liver esterase 207ff - for sulfoxidation, by Acinefobacter 570ff - of cyclohexanone monooxygenase 551 acylases 244ff - microbial -. lactam hydrolysis 257 acylation, of racemic hydroxyacid esters 418f acylation reactions, use of acyl donors 56ff acyl donors, in lipase-catalyzed acylations, enol esters 56ff acyl migration. in mono- and diacylglycerides 139 adamantanes, hydroxylation of 487 ADH see alcohol dehydrogenases adsorption, of lipases 51f

agrobacteria, amidase activity 294

- nitrile hydratase activity 294f - synthesis of indoleacetic acid 294

Agrobacterium radiobacter. D-hydantoinaselcarbamoylase system, D-citruiline formation 268f ajmaline 352f - biosynthetic pathway, in Rairwolfia 353f - structure 353 - treatment of arrhythmias 353 alanine dehydrogenases, applications of 447 - characteristics of 447 - reductive amination of pyruvate 446f Alcalase see subtilisin Alcaligenes faecalis, nitrilase activity 297f alcohol dehydrogenases (ADH), applications of 436ff - CBADH 456f - CFADH 453f - classification of 429 - cross inhibition 430f - emulsion enzyme membrane reactor 424f - enantioselectivity of 427ff - enzyme membrane reactor. with continuous product extraction 423f - HLADH 437ff - introduction of two chiral centers 435f - kinetics of 427ff - LKADH 454 - MJADH 452f - PADH 453 - PLADH 452 - product inhibition 430 - reaction mechanisms, ping-pong mechanism 429 - - sequential mechanism 428f - resolution of racemic hydroxy compounds 4348 - - introduction of two chiral centers 435f - stereochemistry of 427 - substrate inhibition 429f - TBADH 441f - use in cyclodextrins 420ff

590

Index

- use in organic solvents 420ff

alcohol dehydrogenase (secondary), from Thermoanaerobium brockii 403 alcohol dehydrogenases of yeast see yeast alcohol dehydrogenases (YADH) alcohol oxidation, enantioselective 435 alcohol oxidation reaction, rate equation 433 alcohols, acyclic, reactions with Pseudomonas lipases 79ff - analogs of 98f - axially-disymmetric 94ff - chiral, formation by microbial hydroxylation 486 - - formation by soybean lipoxygenase 503 - enantioselective oxidation of 435 - primary -, lipase enantioselectivity 87ff - secondary -, as lipase substrates 68ff - spiro compounds 94ff - tertiary -, enantioselectivity of lipases 94 - with non-carbon stereocenters 98 - with remote stereocenters 97 alcoholysis, of triacylglycerides 141 alkaloids see also benzylisoquinoline alkaloids, monoterpenoid indole alkaloids, morphine alkaloids, tropane alkaloids - as drugs, development of new - 330f - biosynthetic pathways 330 - biotransformations 327ff - classification 328 - from animals 328f - from bacteria 328f - from fungi 328f - from plants 330f - - use of cell culture 331 - - use of metabolic engineering 331 - historical uses 328 - hydroxylation of 495f - semisynthetic derivatives of, cytotoxic action 354 - skeletal types 330 - structures of 329 alkanols, resolution by PPL 75ff alkenes, dihydroxylation of 512 amidase activity, in brevibacteria 292 - of agrobacteria 294 - of penicillin acylase 229 - of pseudomonads 294 - stereoselectivity 308 amide bond cleavage, antibody-catalyzed - 270 amide bond formation 243f - antibody-catalyzed - 270 - hydrolysis of side chain mimics 254f - kinetic control 249 - larger peptides, non a-amino acid substrates 256f - optimum water levels 252 - regioselectivity 253f

-

thermodynamic control 249

- use of immobilized enzymes

255

- with non-activated esters as acyI donors - with unnatural substrates 253ff

262

amides, C-terminal -, hydrolysis of 245 amide 256f 244f - hydroxylations of 479 - N-terminal -, hydrolysis of 246 - synthesis, strategies for 250 amines 98ff - lipase-catalyzed deacylation of 131f - lipase-catalyzed resolutions of 101 amino acid amides 244f amino acids, biocatalytic resolution of N-acetyl cysteine 18 - cleavage of C-terminal residues, by carboxypeptidases 247 - manufacture of 17ff - resolution by acylase I 246ff - unnatural -, enantioselective hydrolysis 248 a-amino acids, coupling of single amino acids, solvent effects 250 - - structural effects 250 - enantioselective hydrolysis of the N-phenylacetyl derivatives 261 D-amino acids, formation from racemic hydantoins 268 L-amino acids, manufacture by N-acetyl hydrolysis 19 - production by immobilized aminoacylase, flow diagram 19 7-amino desacetoxycephalosporanic acid, synthesis of cephalosporin antibiotics 258 amino groups, protection of, by phenylacetyl 261 aminoacylase, production of L-amino acids 19 p-aminobenzonitrile, conversion to p-aminobenzoic acid 296f aminolysis, lipase-catalyzed 263ff 6-aminopenicillanic acid (6-APA), manufacture of 15f - synthesis of penicillin antibiotics 258 aminopeptidase, of Pseudornonas putida, asymmetric hydrolysis of lupinic acid amide 245f - - resolution of racemic a-amino acids 245 ampicillin, formation by penicillin G acylase 259 anhydrides, regioselective ring opening, by lipases 111 animals, alkaloids from 328 antibiotics, hydroxylation of 4% antibodies, catalytic - 269f - monoclonal -, hapten affinity binding studies 269 antibody-catalyzed arnide bond formation 270 antibody-catalyzed amide bond hydrolysis 270 6-APA see 6-aminopenicillanic acid arene dioxygenase-catalyzed reactions 483

- delicious octapeptide - formation by papain

Index arenes, dihydroxylation of 481 formation of dihydrodiols 506 heterobicyclic -, dioxygenation of 509 aromatic hydrocarbons, benzylic hydroxylation 479f aromatic ring hydroxylation 499ff Arthrobacter, nitrilase activity 293 - nitrile hydratase activity 293f ascorbate manufacture, by sorbitol oxidation 19 L-ascorbic acid see vitamin C aspartame 250f - large-scale process for 251 - synthesis, phenylacetyl protection 262 aspartyl proteases, characteristics 248 Aspergillus acylase I ( A A ) 246ff - hydrolysis of N-terminal amides 246ff - resolution of a-amino acids 246 asymmetric sulfoxidations, by flavin dependent monooxygenase enzymes, bakers’ yeast 574 - - mammalian tissue 575f - - Photobacterium phosphoreurn 574 - - Pseudomonas putida 574 - - Pseudomonas sp. 574 - by monooxygenase enzymes, occurring in bacteria 570ff asymmetric syntheses, lipase-catalyzed - 66ff atropine 332 - degradation of, pathway 335f - microbial metabolism of 335ff atropine esterase 335 avermectin derivatives, hydroxylation of 484

-

B Bacillus coagulans, esterase 219 Bacillus subtilis, esterase 219 bacteria, alkaloids from 328 Baeyer-Villiger biotransformations, cofactor recycling and multi-enzyme systems 567f - by Acinetobacter junii 560 - mediated by Corynebacteriurn amidase 291 - Criegge intermediates in - 540 - by Curvularia intermedia 560 - by Cylindrocarpon destructans 563 - by HLADH 565 - multi-enzyme systems for 567 - by Nocardia 566 - by PCL 568 - by Pseudomonas putida 555ff - by Rhodococcus 560 - by TBADH 568 - by Xanthomonas autotrophiciis 561, 566 - of bicyclo[2.2.l]heptanones 563ff - of bicyclo[3.2.0]heptanones 560ff - of bicyclo[4.2.0]octanones 562f - of monocycles 555ff - of polycyclics 566f

591

Baeyer-Villiger oxidation, of cyclohexanone derivatives with a genetically modified yeast 383 Baeyer-Villiger reactions, abiotic - 537ff - biological -, fenchone 545 - - in camphor metabolism 543f - - listing of 555ff - - questin 548 - - raspberry ketone 548 - - ring cleavage of questin 543 - - steroids 54lff - - tridecanone 542 - - unnatural substrates 546 - - use of Acinetobacter 546 - Criegee intermediates 538, 540, 551 - enantiomeric intermediates 552 - flavoenzymes catalyzing - 549 - gross mechanism 538 - intramolecular 539 - large scale - 568f - mechanisms for cyclohexanone rnonooxygenase catalyzed - 554 - N-terminal sequences of flavoenzymes 549 - of steroids 541 - peracids in 538f - reagents for 539f Baeyer-Villiger ring expansion 535ff bakers’ yeast 364f - esterase 220 - expression of cyclohexane monooxygenase in 546 - multiple activities 31 - sulfoxidations by 574 Beauveria sulfurescens, as the classic hydroxylating microorganism 486 - stereochemical model for hydroxylation 479 benzamide, by hydrolysis of benzonitrile 287 benzenes, disubstituted -, dioxygenation reaction of 506 - monosubstituted -, dihydroxylation of 508 - polyhydroxylated -, lipase-catalyzed deacylation 129 benzenoids, bicyclic -, dihydroxylation of 509 - substituted -, dihydroxylation of 507 benzophenanthridine alkaloids 338f - biosynthesis 339 benzopyran, dihydroxylation of 515 benzylic hydroxylation, by Mortierella isabellina 487f benzylisoquinoline alkaloids 337ff Betapol, interesterification of 136 bioactive compounds, hydroxylation of 484 biocatalysis, use in chemical synthesis 11 biocatalytic processes, current large-scale processes 20 - production volumes 20 bioremediation. of nitrile pollutants 307 - of wastes 312f

592

Index

biotransformations, advantages over abiotic asymmetric catalysts 26 - by enzymes, multi-enzyme systems 33 - - practical aspects 26, 32f - by whole cells 13, 26ff, 31f, 396ff, 483ff - experimental procedures for 27ff - - hydrolase-catalyzed reactions 27ff - - oxidoreductase-catalyzed reactions 29 - literature 27ff - on-line services 27 - perspectives Iff - - 1895-1935 6ff - - 1935-1955 lOff - - 1955-1975 l l f f - practical aspects 25ff - process options 6 - screening of microorganisms for 31 - with isolated enzymes 502ff - with yeast 363ff biphenyl, hydroxylation of 500 p-blockers, lipase-catalyzed reactions 84 brevibacteria, amidase activity 292 - nitrile hydratase activity 292 Brevibacteriurn, nitrile hydratase defective mutants 292 brewers’ yeast 364 bromoxynil, metabolism of 298 butanol, production volumes 1910-1929 9 C CAL see Candida antarctica lipase CAL-B see Candida antartica B lipase camphor, catabolism by Pseudornonas putida 544 - pathway for the degradation of 543f Carnprotheca acurninata 330 camptothecin, from Carnptotheca acurninata 330 Candida antarcrica B lipase (CAL-B) 42f - enantioselective reactions of carboxylic acids 103f - enantioselectivity toward secondary alcohols 71f - general features 44 - resolution of acrylic secondary alcohols 72 Candida antarctica lipase (CAL) 263f - amidation by 263 - aminolysis by 263ff - ammonolysis by 264ff Candida boidinii alcohol dehydrogenase ( C B A D H ) ,applications of 457 - characteristics of 456f - formate dehydrogenase 403 Candida cylindracea lipase (CCL) 263f - aminolysis by 265 Candida parapsilosis carbonyl reductases, applications 455 - characteristics of 454f

Candida rrrgosa lipase (CRL) 41ff

- catalytic machinery 70 - enantioselective reactions of carboxylic acids

104

- enantioselectivity toward secondary alcohols

72ff

- general features 44 - resolution of acyclic secondary alcohols 73

resolution of cyclic secondary alcohols 74 E-caprolactam, induction of nitrile hydratase 289f scaprolactone, lipase-catalyzed polymerization 145 carbamoylase 267ff carbohydrate chemistry, selective protection of hydroxyl groups 118 carbohydrates, as energy source in yeast fermentation 366f - regioselective esterification, by substitution 224 carbonyl compounds, asymmetric reduction, Prelog’s rule 427f - enantioselective chemical reductions of. reagents 394 - microbial reductions of 396f carbonyl reductases, enantioselectivity of 455 - novel - 454f - of Candida parapsilosis 454f - of Rhodococcus erythropolis 456 - substrate specifity of 455 - substrate spectrum of 456 carbonyl reduction, Prelog’s rule 380 carboxyl groups, lipase-catalyzed protection in peptide synthesis 132 carboxylic acids, aromatic -. by methyl hydroxylation 488 - lipase-catalyzed enantioselective reactions of 103ff - with a stereocenter at the a-carbon 103ff - - lipase resolution 107 - with a stereocenter at the P-position, lipase resolution 108 - with quaternary stereocenters, lipase resolution 109 - with remote stereocenters 109f - - lipase resolution 110 - with sulfur stereocenters 109f - - lipase resolution 110 carboxypeptidase A (CP-A) 247f - preferred cleavage sites 249 carboxypeptidases, cleavage of C-terminal L-amino acid residues 247 L-carnitine, synthesis by PADH 453 catalytic activity, of lipases, increase in organic solvents 51ff - of yeast 366 catalytic antibodies 269f catechols, formation of 509f -

Index Catharanrhus roseus, biosynthetic pathways for vindoline formation 354f - strictosidine synthase 352 CBADH see Candidn boidinii alcohol dehydrogenase C-C bond formation, by yeast sterol cyclase 378 C=C double bond reduction, by yeast. mechanism 372 - prochiral by yeasts 370f - selective -, by yeasts 377 C = C double bonds, asymmetric reduction by yeast enzymes 375ff - non-aromatic -, dihydroxylation of 511 - preparation of chiral compounds by yeast reduction 376 CCL see Candida cylindracea lipase CE see cholesterol esterase cephalosporin C, two-step enzymatic side chain removal 16 cephalosporin derivatives, conversion from penicillins 16f cephalosporin synthesis 258ff CFADH see Citrvitlnria falcarn alcohol dehydrogenase cheese-making, use of lipases 40f chemical intermediates. enantiomerically pure use of lipases 146 chicken liver esterase 211f chiral alcohols, formation by microbial hydroxylations 486 chiral compounds, preparation by yeast reduction of activated C=C bonds 37Sf chiral technology, biotransformations in 26 chloroperoxidase. commercial availability 482 - hydroxylation by 504 CHMO see cyclohexanone monooxygenase cholesterol esterase (CE) 214ff - fastest reacting enantiomers of racemic secondary alcohols with 216f - from bovine pancreas 214 - kinetic resolution of phosphines and phosphine oxides 217 required presence of bile salts 214 Chromobacteriitni viscosiirn lipase (CVL) 41ff chymotrypsin 226ff - hydrolysis of ester bonds 226ff - increased stability of 229 - kinetic resolution of racemic substrates 227f - preferred cleavage sites 249 - stereospecificity 228 chymotrypsin catalyzed hydrolysis 228 citral. hydroxylation of 491 citric acid, growth of production volumes 10f - production volumes 1910-1929 9 citronellol, hydroxylation of 491 D-citruhe formation. by Agrobncreriurn rridiobacrer hydantoinase 268f

-.

-.

593

CLEC lipases 52 CLECs see cross-linked enzyme crystals clostripain, preferred cleavage sites 249 cocaine 332 - microbial metabolism of 337f cocaine esterase 337 cocoa butter substitutes 146 - use of lipase in manufacture of 135f codeine, hydroxylation of 496 - N-demethylation by Citnninghamella 345f - potent analgesic 340 - redox reactions of 350 codeine N-oxide, structure 348 coenzyme regeneration, characteristics of selected enzymes 445 - dehydrogenases 399f - enzyme-coupled - 402 - methods 402ff - nonenzymatic - 400f - substrate-coupled - 401f - use of LDHs 442 - use of YADH 440 cofactor recycling, Baeyer-Villiger biotransformations 567f cofactor regeneration 368 colchicine, demethylation of 497 coniine, hemlock alkaloid 328f - synthesis of 6 cortisol 13 Corynebacteria, nitrile hydratase activity 290ff Corynebacterium, arnidase 291 - Baeyer-Villiger reaction, mediated by 545 - hydrolysis of amino nitriles 290f - hydrolysis of cyclohexane dinitriles 291 coupling methods, for dehydrogenases 407 CP-A see carboxypeptidase A Criegee intermediates 538, 551ff - as reagents for Baeyer-Villiger reactions 540 - of cyclohexanone monooxygenase, of Acinetobncter 553 CRL see Candida rrrgosn lipase cross inhibition, of alcohol dehydrogenases 429f cross-linked enzyme crystals (CLECs) 52 - in peptide synthesis 256 cross-linked enzymes 407f crude preparations, of lipases 43f culture collections, addresses 32 - listing 32 Ciirvitlnria falcafn alcohol de hydrogenase (CFADH). characteristics of 453 Crrrvrrlnria infernredin, Baeyer-Villiger biotransformations 560 Cirrvirlarin hrnatn. Baeyer-Villiger biotransformations 559 CVL see Chromobocreriiini viscosirm lipase cyanamide, conversion to urea 295 cyanide, biotransformation of 299ff

594

Index

cyanide hydratases 299ff - fungal -, basic mechanisms 299 - - use in bioremediation of wastes 312f cyanide wastes, fungal cyanide hydratase, use of 312f @-cyanoalanine synthesis, by cyanogenic bacteria 280 - - by cyanogenic plants 280 0-cyanoalanine synthase, properties 280 cyanogenic glycosides 280ff - biosynthetic pathway in plants 282 - general scheme for the release of cyanide 283 - metabolic pathways in plants 280 cyanohydrins, biosynthesis 301 - biotransformation of 301f - enzyme-catalyzed formation 301 cyclic diesters, asymmetrization of, by acetylcholinesterase 213 cyclic diesters (racemic -), kinetic resolution by pig liver esterase 201 cyclodextrins, alcohol dehydrogenases in 420ff - as co-solvents 426 - oxidoreductions in 425ff - product complexes with 426 - structure of 425 - substrate complexes with 426 cyclohexane dinitriles, hydrolysis of, by Corynebacterium 291 cyclohexanone monooxygenase (CHMO) 570ff - active site model 551 - Baeyer-Villiger reaction, mechanisms for 554 - Criegee intermediates of Acinefobacfer 553 - from Acinetobacter sp., mechanism of the Baeyer-Villiger reactions 548 - of Acinetobacter calcoaceticris 570ff - oxidation of sulfides to sulfoxides 572 - stereoselectivity of 575 - thiolactone inhibitors of 573 cyclohexanone monooxygenases, expression in yeast 546 cyclohexan-1,2-diol, metabolism to 547 cyclohexene derivatives, asymmetrization by pig liver esterase 197 cyclooxygenase, hydroxylation by 504 cyclopentane dicarboxylates, asymmetrization by pig liver esterase 198 Cylindrocarpon destrucmns, Baeyer-Villiger biotransformations 563 cvtochrome P-450-de~endentenzvmes 505 citochrome P-450-dependent monooxygenase (cyt. P-450) 477f - expression in Escherichin coli 483 - expression in yeast 483 D DAGs see diacylglycerides a-damascone. hydroxylation of 493

dealkylation see also demethylation dealkylation reactions 496ff dehydro-amino acids 253f dehydrogenases 391ff - - see also oxidoreductases - coenzyme recycling 402 - coenzyme regeneration system, requirements of 399 - commercial - 436ff - - biochemical characteristics of 438 - - manufacturers of 449 - enantioselective reduction. of P-ketoacid esters 412 - - of @-ketoacids 412 - enzymatic transformations 396ff - enzyme-coupled regeneration 402 - noncommercial 448ff - - biochemical characteristics of 450 - substrate spectra 437 - total turnover number 399 - turnover number 398f - typical reactions 394ff - whole-cell transformations 396ff delicious octapeptide amide, synthesis 256f demethylation, of colchicine 497f - of immunomycin 497 - of magnolin 498 DHA see docosahexaenoic acid diacylglycerides (DAGs), acyl migration in 139 - formation from glycerol 142 - general features 137ff - lipase-catalyzed syntheses of 138 L,L-diastereomers, protease-catalyzed synthesis 253 diastereotopic faces, of flavin adenine dinucleotide 550 diesters (rneso).enantioselective hydrolysis by pig liver esterase 196 diesters (prochiral -), enantioselective hydrolysis of 195 digoxigenin derivatives 133 dihydroxylation 475ff - definition of 480f - of alkenes 512 - of alkenic monoterpenes 512f - of arenes 481 - of benezenes 508 - of benzenoids 507ff - of benzopyran 515 - of fluoranthene 511 - of geranylacetone 513f - of heterobicyclic arenes 509f - of naphtalenes 509 - of nerolidol 513f - of non-aromatic C=C bonds 511 - of precocene 515 - of sesquiterpenes 513f

Index

- routes 480f - use of isolated enzymes 482 - use of whole-cell catalysts 481f

dihydroxylation reactions, dioxyenase-catalyzed -, non-aromatic C=C bonds 511 - dioxygenase-catalyzed -, arenes 506ff - - heteroarenes 509f - monooxygenase-catalyzed - 511ff a-diketones. conversion to diols by yeast 372 - yeast reduction of, use of selective enzyme inhibitors 382 diltiazem, commercial synthesis, use of lipases 147 dinitriles. aromatic -, selective hydrolysis of 308 - prochiral -, selective amidase hydrolysis of 310 - - selective nitrile hydratase hydrolysis of 310 - regiospecific biotransformation of 308 dioxygenase, dihydroxylation of arenes 506ff - dihydroxylation of heteroarenes 509f dioxygenase enzymes 481 directed evolution, of lipases 149f disaccharides, regioselective acylation by subtilisin 224f diterpenes, hydroxylation of 494f - labdane diterpene hydroxylation 495 docosahexaenoic acid (DHA) 137 drugs, of plant origin, camptothecin 330 - - taxol 330

E

eicosapentaenoic acid (EPA) 137 elective culture, definition of 546 - isolation of Acinetobacter 547 emulsion enzyme membrane reactor, scheme 424f endopeptidases. nomenclature 244 enoate reductase, of yeast 372 - - reduction of triple substituted double bonds 375 - - substrates for 375 enol esters, in lipase-catalyzed acylations S6ff entrapment, of lipases 51f entrapment methods 409f enzymatic activities, of yeast 370ff enzyme activity, optimization of. by heterologous expression 26 - - by site selective mutagenesis 26 - solvation effects on 252 enzyme engineering 17f enzyme inhibition, selective -, improvement of yeast biotransformations 382f enzyme kinetics, turnover number 31 enzyme membrane reactors, modeling of 432f - technology 409 with continuous product extraction 423f enzyme nomenclature 29f

595

- classification system 30

enzyme reactors, batch stirred tank reactor 431f 432 - continuous stirred tank reactor 431f - modeling of 431 - plug flow reactor 431f enzymes, biotransformations of unnatural substrates 32 - cofactor-dependent -, commercially available 393 - immobilization of, in peptide synthesis 255 - multi-enzyme systems 33 - tolerance to organic solvents 32f EPA see eicosapentaenoic acid ephedrine, formation of 9f D-ephedrine manufacture. by aldol condensation 19 epoxidation, by cyt. P-450 monooxygenase 481 epoxide hydrolases 481 Escherichia coli, expression of cyt. P-450 483 - overexpression of heroin esterase 351 - penicillin acylase 15 - penicillin G acylase 258 esterase activity, of penicillin acylase 229 esterases 28, 193ff, 250 - differences from lipases 194 - formation of amides 263ff - by aminolysis 263ff - in peptide synthesis 262 - microbial -, Bacillus coagirlans 219 - - Bacillus subtilis 219 - - bakers' yeast 220 - - Penicillirrm freqirentans 221 - - Pichia miso 219f - - Rhizopirs nigricans 219 - stability in anhydrous solvents 262 estrogens, hydroxylation of 489 estrone, combined synthesis 14 ethanol, as energy source in yeast fermentation 367f - production volumes 1910-1929 9 exopeptidases, nomenclature 244

- characteristics of

F

FAD see flavin adenine dinucleotide farnesene, hydroxylation of 493 fats, lipase-catalyzed processes for specialty fats 134 fatty acids, hydroxylation of 483 - in vivo synthesis of 373 - selectivities of lipases, saturated fatty acids 142ff - - unsaturated fatty acids 143f fatty acid synthetase complex, of yeast, reducing enzymes 373f fatty acid synthetase cycle, schematic representation 373

596

Index

FDH see formate dehydrogenase fenchone, Baeyer-Villiger reaction, Corynebacterium mediated - 545 ferrocenes, lipase-catalyzed reaction of 100 flavin adenine dinucleotide (FAD) 549ff - diastereotopic faces of 550 - structure 536 flavin dependent monooxygenases (FMO), in mammalian tissue, pig liver 575 - - pig lung 575 flavin hydroxyperoxide, structure 537 flavin mononucleotide (FMN), structure 536 flavin monooxygenases 535ff - identification of 541ff - isolation of 541ff flavoenzymes, catalyzing Baeyer-Villiger reactions, N-terminal sequences of 549 - use in biotransformations 536 fluoranthene, dihydroxylation of 511 - hydroxylation of 501 FMN see flavin mononucleotide FMO see flavin dependent monooxygenases food ingredients, production using lipases, cocoa butter substitutes 146 - - flavor esters 146 formate dehydrogenase (FDH), from Candida boidinii 403 - from Pseudomonas sp. 403 Friedels-Craft acylation 539 fumaronitrile, conversion into metabolizable products 313 fungi, alkaloids from 328 - cyanide hydratases 2991 - immobilization of spores 488 - nitrilase activity 298f - nitrile hydratase activity 295

G

genetic engineering. of yeast 383 - - hydroxylase enzymes 482f Geotrichum condidurn, glycerol dehydrogenase of 443 geraniol, hydroxylation of 491 geranylacetone, dihydroxylation of 513 gibberellins, hydroxylation of 494 glucose, selective acylation by PPL 119 glutamate dehydrogenases. applications of 445 - characteristics of 444f - reductive amination of 2-oxoglutarate to L-glutamate 444f - regeneration of NAD(P) 406f glutamic acid, lipase-catalyzed deprotection of carboxylate groups 133 glutarates, prochiral -. asymmetrization by pig liver esterase 195f glycerides, 1,3 regioselective reactions 134

glycerol, esterification with fatty acids 142

- formation of DAGs 142 - formation of MAGS 142 - production volumes 1910-1929 9

glycerol dehydrogenases, applications of 443f

- characteristics of 443f - Geotrichum candidum 443 - substrates accepted by, oxidative direction 443

glycerolysis, of triacylglycerides 141 glycine, conversion into hydrogen cyanide 279 glycosides. cyanogenic - 280ff 1

Haldane equation 431 hapten, phosphonamide 269f - transition state mimic 269 hapten affinity binding sites, of monoclonal antibodies 269 hemlock alkaloid 328 heroin, microbial transformations of 351 - potent analgesic 340 heroin esterase, heroin metabolites formed by 35 1 - overexpression in Escherichia coli 351 heterologous gene expression, for optimization of enzyme activity 26 HLADH see horse liver alcohol dehydrogenase HLE see horse liver esterase horse liver alcohol dehydrogenase, applications of 439 - Baeyer-Villiger biotransformations 565 horse liver alcohol dehydrogenase (HLADH) 437ff - characteristics of 437 horse liver esterase (HLE), asymmetrization of prochiral organosilyl-substituted esters 21 If - kinetic resolution of, racemic lactones 210 - - racemic methyl 2-alkyl-2-aryl esters 211 horse liver esterase (HLE) 209ff horseradish peroxidase, commercial availability 482 - hydroxylation by 504f hycoscyamine 332 hydantoinases 267 hydantoins, racemic -, conversion to L-amino acids 267 hydantoin substrates, hydrolysis by, allantoinase 267 - - carboxymethylhydantoinase 167 hydrocarbons, aromatic -, benzylic hydroxylation 479f hydrocodone 341, 347 - biological production of 350f hydrogen cyanide 279f. 299f - bacterial metabolism 300f - synthesis 279f

Index -

-

- by fungi 279 - by photosynthetic microbes

279f

- of citral 491 - of citronellol 491 - of codeine 496

597

alp-hydrolase fold, of lipases 45 hydrolase-catalyzed reactions, examples 27ff - of cyclic terpenes 492 hydrolases, in yeast 379 - of a-damascone 493 hydrolysis. of triacylglycerides 141 - of diterpenes 494f hydromorphone 341, 347 - of farnesene 493 - biological production of 350f - of fatty acids 483 hydroperoxide formation. by lipoxygenases 502ff - of fluoranthene 501 - of geraniol 491 hydroxy compounds see also hydroxyacid esters - of giberellins 494 - chiral -, preparation by PADH 453 - of a-ionone 493 - - resolution with alcohol dehydrogenases 434ff - of isobutyric acid 483 - - synthesis of 411ff - of labdane diterpenes 495 - enzymatic hydrolysis of racemic acyl deriva- of milbemycins 496 tives 415f - of monensin 496f - - racemic ether derivatives, 414f - of monoterpenes 491f hydroxy esters, chiral - 373 - of morphine 496 hydroxyacid esters, chiral -, synthesis of 411ff - of nicotinic acid 501f - enantioselective reduction 412 - of phenanthrene 501 - racemic -, enzymatic hydrolysis of 413f - of polycyclic aromatic hydrocarbons 501 - - selective acylation 418f - of progesterone, by Rhizopus arrhizus 477 - - transesterification reactions 416ff - of pyrazines 502 hydroxyacids, enantioselective reduction 412 - of pyrene SO1 hydroxyisocaproate dehydrogenases, applications - of pyridines 501f of 451 - of quinoline 502 - characteristics of 448ff - of sesquiterpenes 492ff hydroxyl groups, in non-sugars 129ff - of steroids 479 - primary lipase-catalyzed reactions 118ff - - by plant cell cultures 489 - secondary -, lipase-catalyzed deacylations 124ff - - estrogens 489f - - lipase-catalyzed regioselective acylation 126ff - - progesterone 488ff. 505 - - model of acylation 127 - - testosterone 489 - selective protection of 118 - of substituted adamantanes 487 hydroxylase enzymes, mammalian -, expression in of terpenes 479,491ff microbial systems 482 - of terphenyl 500 hydroxylation 475ff - of vitamin D3 485 - - see also aromatic hydroxylation, benzylic hy- regioselectivity of 478ff droxylation - use of isolated enzymes 482 - by Beauveria sulfurescens, alicyclic - 486 use of whole-cell catalysts 481f - - stereochemical model 479 hydroxylation and dihydroxylation, parallel path- by chloroperoxidase 504 ways for 514f - by cyclooxygenase 504 hydroxylation reactions, biotransformation with - by genetically engineered organisms 482f isolated enzymes 502ff - by horseradish peroxidase 504f - by microbial cultures 482 - by lipoxygenases 502f - whole-cell biotransformations 483ff - by peroxidases 504f hydroxyls, lipase-catalyzed deacylation, of alipha- enzymes 477f tic - 130f - formation of phenols 499ff - - of phenolic - 129 - history 477 hydroxysteroid dehydrogenases, applications of - microbial -, preparation of chiral alcohols 486 448 - occurring with rearrangement 492 - characteristics of 447f - of alkaloids 495f hyoscyamine, biosynthesis of 332 - of amides 479 hyoscyamine 6-p-hydroxylase 332ff - of antibiotics 496 - of aromatic rings 499ff - of avermectin derivatives 484 I - of bioactive compounds 484f immobilization, of Acinerobacrer 298 - of biphenyl 500 - coupling methods for 407ff

-.

598

Index

- in yeast biotransformation 369 - of coenzymes, coupling to cross-linked

enzymes 407f

- of enzymes 18 - - in peptide synthesis 255 - of fungal spores 488 - of lipases 52 - of nicotinamide coenzymes, by entrapment

methods 409

- - coupling on a carrier 408f - of penicillin G acylase, formation of loracarbef

259f of yeast 369 immunomycin, demethylation of 497 indoleacetic acid, synthesis by agrobacteria 294 inositol, enantiomerically pure derivatives 81 interesterification of margarine 137 interfacial activation, of lipases 40, 45f a-ionone, hydroxylation of 493 isobutyric acid, hydroxylation of 483 isocodeine, structure 348 isoquercitin, selective acylation by lipase 124 -

K ketoprofen, enantioselective amide hydrolysis 310 Klebsiella ozaenae, nitrilase activity 298 kyotorphin, large-scale production 252

L

labdane diterpenes see diterpenes p-lactam antibiotics, commercial synthesis, use of lipases 147f - manufacture 15ff lactam hydrolysis, by microbial acylases 257 plactams, amide hydrolysis of 19 - ester hydrolysis of 19 lactate, production volumes 1910-1929 9 lactate dehydrogenase (LDH), of yeast 372 D-lactate dehydrogenases 442f L-lactate dehydrogenases 442 Lactobacillus kefir alcohol dehydrogenase ( L K A D H ) ,applications of 454 - characteristics of 454 - synthesis of chiral hydroxy compounds 454 Lnctococcus tactis, prolyl dipeptidyl-aminopeptidase 256 lactone precursors, lipase-catalyzed resolution 114 lactones, lipase-catalyzed enantioselective reactions 112f lactones (racemic-), kinetic resolution by horse liver esterase 209f lanosterol cyclization, mediated by yeast sterol cyclase 378

LDH see lactate dehydrogenase L-leucine aminopeptidase 246f leucine dehydrogenases, application of 445f - characteristics of 445 - reductive amination by 445f leucine production, with leucine dehydrogenase 446 lipase-catalyzed acylations, of hydroxyl groups 126ff - of modified sugars 121ff - of unmodified sugars 119ff - use of acid anhydrides 58 lipase-catalyzed amidation 265 lipase-catalyzed aminolysis 263ff - formation of an acyl-enzyme intermediate 266 - formation of amides 263ff - of diethyl fumarate 267 - of succinic acid diesters 267 lipase-catalyzed asymmetric syntheses, by PCL 67f - by PPL 67f - enantiomeric purity 66ff lipase-catalyzed deacylations, of aliphatic hydroxyls 130f - of amines 131f - of hydroxyl groups 124ff - of phenolic hydroxyls 129 - of polyhydroxylated benzenes 129 lipase-catalyzed enantioselective reactions, involving the lactone ring 112f - mvo-inositol derivatives 87 - of acyclic alcohols 79ff - of alcohols, with remote stereocenters 97 - of axially-disymmetric alcohols - of carboxylic acids 103ff - - with a stereocenter at the P-position 108 - - with quaternary stereocenters 109 - - with remote stereocenters 110 - - with sulfur stereocenters 110 - of cyclic secondary alcohols 82ff - of primary alcohols 89ff - - with quaternary stereocenters 95 - of secondary alcohols, RML 85 - of spiro alcohols 96 - of tertiary alcohols 94 - of thiols 102 lipase-catalyzed reactions, asymmetric synthesis 66ff - P-blockers 84 - catalytic mechanism 45, 47 - enantiomeric purity 63ff - formation of macrolactones 115 - in situ recycling 64 - of ferrocenes 100 - of triacylglycerides 135 - polymerizations, condensation reactions 144 - - ring-opening polymerizations 145

Index -

- transesterification reactions 144f

reaction media, aqueous solutions 47ff - organic solvents 50ff - reverse micelles 48f - - supercritical fluids 60ff - ring opening of 2-phenyloxazolin-5-ones 116 - sequential kinetic resolution 64 - synthesis of, diacylglycerols 138 - - MLM lipids 136f - - monoacylglycerides 140f - use of acyl donors 56ff - water content of reaction mixture, thermodynamic water activity 59 - - total water content 58f lipase CLECs 52 lipase enantiopreference, empirical rules 69, 88 - size-based rules 68 lipase hydrolysis, in aqueous media 47f - in organic solvents 48ff - in reverse micelles 48f lipase selectivity, modeling of 149 - mutating of 149 lipases 28f, 37ff, 250 - acyl chain binding site 47f - alcohol binding site 47f - availability 40ff - bacterial - 42f - catalytic amino acid triad 70 - chemoselective reactions 118ff - classification, by protein sequence 43 - commercial -, protein sequence alignments 43 - - source and names 42 - - world market 41 - commercial applications, enantiomerically pure chemical intermediates 146 - - enantiomerically pure pharmaceutical intermediates 146f - - food ingredients 146 - crude preparations 43f - directed evolution, random mutagenesis 149f - - screening 149f - enantiomeric ratio 63 - enantioselective reactions involving the lactone ring 112f - enantioselectivity toward triglycerides 91ff - entrapment of 51f - fatty acid selectivity, saturated fatty acids 142ff - - unsaturated fatty acids 143f - fungal - 42f - in cheese making 40f - in commercial synthesis of diltiazem 147 - indirect resolutions of lactones 114 - in laundry detergents 40f - in organic solvents, catalytic activity 51ff - - CLECS 52 - - covalent immobilization 52 - - covalently modified - 53

599

enantioselectivity 54ff lipid-coated - 53f recovery from 51 surfactant-coated - 53f in peptide synthesis 262 in supercritical fluids 60ff - in tricyclic p-lactam antibiotic synthesis 147f - increase of enantioselectivity in organic solvents 54ff - interfacial activation of 40, 45f - large-scale application, optimization of 148f - mammalian - 42f - nomenclature 41ff - nonselective - 134ff - primary alcohols as substrates for 87ff - regioselective reactions 118ff - regioselective ring opening of anhydrides 111 - resolution of amines 101 - resolution of metal carbonyl complexes 99 - secondary alcohols as substrates for 68ff - 1,3-selective - 134ff - stability in anhydrous solvents 262 - structure, dP-hydrolase fold 45 - supercritical COz reactor 62 - X-ray crystal structures 46 lipid-coated lipases 53f lipid modifications, lipase-catalyzed - 134ff - 1,3-regioselective reactions of glycerides 134f - - acyl migration in mono- and diacylglycerides 139 - - cocoa butter substitutes 135f - - diacylglycerides 137ff - - esterification of glycerol with fatty acids or fatty acid esters yielding I(3)-MAGs 142 - - glycerolysis of triglycerides to l(3)-MAGS 141 - - hydrolysis or alcoholysis of triglycerides to 2MAGs 141 - - modified triglycerides 135 - - monoacylglycerides (MAGs) 139 - - synthesis of MLMs 136f - - triacylglycerides containing polyunsaturated fatty acids (PUFAs) 137 lipoxygenases, catalysis of hydroxylations 502f - definition of 502ff - mushroom - 503f - porcine - 503f - potato tuber - 503 - soybean - 502f L KADH see Lactobacillus kefir alcohol dehydrogenase loracarbef, formation by immobilized penicillin G acylase 259f Lowry assay 43 luciferase, bacterial -, oxidation of sulfide to sulfoxide 574 -

-

600

Index

L-lupinic acid 245f lupinic acid amide, asymmetric hydrolysis 246

- naltrexone 345 - biosynthesis in Papaver somniferurn 342f -

M

macrolactones, lipase-catalyzed formation 115 magnolin, demethylation of 498 MAGs see monoacylglycerides malonates, prochiral -, asymmetrization by pig liver esterase 195 mammalian drug metabolism, aromatic hydroxylation 500 mammalian tissues, flavin dependent monooxydases 575f margarine, interesterification of 137 menthol, enantiomers 69 metabolic engineering, nitrile transformations 314 metal carbonyl complexes, resolved by lipases 99 metalloproteases, characteristics 248 methyl alkylphenylphosphinoylacetates (racemic -), kinetic resolution by pig liver esterase 206 Methylococcus capsulatus, non-heme iron protein methane monooxygenase 482 microorganisms, sources of 31f milbemycins, hydroxylation of 496 MJADH see Mucor javanicus dehydrogenase MLM lipids 135ff - synthesis of 136f monensin, hydroxylation of 496 monoacylglycerides (MAGs), formation from glycerol 142 - formation from triglycerides 141 - general features 139 - lipase-catalyzed syntheses 140 monoclonal antibodies, hapten affinity binding sites 269 mononitriles, biotransformations of 302, 307f monooxygenases, asymmetric sulfoxidations in bacteria 570ff - cytochrome P-450-dependent - 477f - dihydroxylation reactions 511ff monoterpenes, alkenic -, dihydroxylation of 512f - biotransformation of, by plant cell cultures 492 - hydroxylation of 491f monoterpenoid indole alkaloids 352ff morphine, biosynthesis in Papaver sornniferum 342f - dimerization of 345 - hydroxylation of 496 - potent analgesic 340 - redox reactions of 350 - transformation by Pseudornonas putida 347 morphine alkaloids 339ff - as potent analgesics, codeine 340f - - heroin 340 - - morphine 340f - - naloxone 345

microbial metabolism of 344ff

- microbial transformations of heroin 351 - production of, hydrocodone 350 - - hydromorphone 350 -

transformation by bacteria 345

- - Pseudornonas putida 346ff

morphine dehydrogenase 346ff of Pseudomonas putida, characteristics of 348f - structural gene for 349 morphine reductase 346f - constitutive expression of 349 morphinone reductase, structural gene for 349 Mortierella isabellina 486f - benzylic hydroxylase model 480 - benzylic hydroxylation 487f Mucor javanicus alcohol dehydrogenase (MJADH), applications of 452f - characteristics of 452f multi-enzyme systems, Baeyer-Villiger biotransformations 567 - in biotransformations 33 mushroom lipoxygenase, hydroperoxide formation 503 rnyo-inositol derivatives, lipase-catalyzed enantioselective reactions of 87 Myrothecium verrucaria, nitrile hydratase activity 295 -

N

NAD see nicotinamide adenine dinucleotide NADH, regeneration of, characteristics of selected enzymes 445 NAD(P), regeneration in situ 405ff - - use of glutamate dehydrogenase 406f NAD(P)-dependent oxidoreductases, classification of 394f NAD(P)H, regeneration in situ 403ff NADPH regeneration system, in yeast using carbohydrates as energy source 367 N A D + regeneration cycle, in yeast reduction 368 naloxone, synthesis of narcotic analgesics 345 naltrexone, synthesis of narcotic analgesics 345 - treatment of severe alcohol addiction 342 naphthalenes, dihydroxylation of 509 naringin, selective acylation by lipase 124 neopinone, transformations by Trarnetes sanguinea 344 nerolidol, dihydroxylation of 513 nicotinamide 285 - hydrolysis of 286 nicotinamide adenine dinucleotide coenzymes, structure 395 nicotinamide adenine dinucleotide (NAD), origin of coenzymes 399

Index nicotinamide coenzymes see also NAD(P), NAD(P)H - half-lives 400, 410 - immobilization methods 408 - immobilization of, by coupling methods 407ff - - by entrapment methods 409f - prices of 398 - regeneration 397ff - - by coupling on a carrier 408f - - by coupling to cross-linked enzymes J07f - - by entrapment methods 409f - - methods 401ff - - of enzyme-coupled - 402 - - of immobilized coenzymes 407ff - - of substrate-coupled - 401ff - - principles for nonenzymatic - 400ff - stability 400 nicotinic acid, hydroxylation of 501 nitrilase activity, in plants 299 - of Acinetobacter 298 - of Alcaligenes faecalis 297f - of Arthrobacter 293f - of fungi 298f - - Torulopsis candida 299 - of Klebsiella ozaenne 298 - of Nocardia 293 - of Pseudomonas 297 - of rhodococci 295 nitrilases 283, 295ff - pH optimum 296 - purification of 296 - temperature optimum 296 nitrile containing wastes, bioremediation of, use of mixed cultures 312 nitrile group, biotransformation of 282ff - chemical properties 278 - electronic structure 278 nitrile hydratase activity, bacterial, growth medium design 288 - in Rhodococcus erythropolis 285 - in Rhodococcus rhodochrous 285ff - of microbes 283ff - - agrobacteria 294f - - Arthrobacter 293f - - brevibacteria 292 - - corynebacteria 290f - - Myrothecium verrucaria 295 - - Nocardia 293 - - pseudomonads 294 - - rhodococci 284ff nitrile hydratases 283ff - cobalt content 288 - induction by E-caprolactam 289f - induction by urea 288 - iron containing - 289 - of Rhodococciu, characteristics 286ff - - primary sequence 286

- - structure 286ff - photoactivated preparations

601

289

- rhodococci 284ff - - Rhodococcus rhodochrous 285ff

- stereoselective activity 308 nitrile hydrolysis, by hydratases 283 - by nitrilases 283 nitrile pollutants, bioremediation of 307 nitriles 277ff - chemical synthesis of 278 - chemical transformations of 278f - enantioselective hydrolysis of 309 - heterocyclic -, hydrolysis of 287 - isolated from animals 282 - naturally occurring- 279ff - stereoselective biotransformation of 308ff nitrile transformations, biotechnology of 302ff - by microorganisms, listing of 303ff - commercial processes, acrylic acid production 311f - redesign of enzymes for, by protein engineering 314 - search for novel activities 313 - use of metabolic engineering 314 Nocardia, Baeyer-Villiger biotransformations 566 - nitrilase activity 293 0 Ochrobactrum anthropi amidase 246f opiate analgesics 340ff - semisynthetic - 341 opiate drugs, chemical synthesis of 342 - structures 341 opiates, semisynthetic -, hydrocodone 347 - - hydromorphone 347 opium, of Papaver somniferum 342 organic chemicals, synthesis by biological catalysis 19 organic solvents, alcohol dehydrogenases in 420ff - in yeast biotransformations 369f - lipases in 5Off - - increase of catalytic activity 51ff - - increase of enantioselectivity 54ff - - water content of reaction mixture 58ff - log P values of 420f - oxidoreductions in 423ff oxamniquine, manufacture of 14 - - by benzylic hydroxylation 19 oxidoreductase-catalyzed reactions, examples 29 oxidoreductases see also dehydrogenases, reducing enzymes - NAD(P) dependent -, classification 394f - NADPH-dependent - 412 - nomenclature 478 - noncommercial -, future potential in organic synthesis 457f

602

Index

- of yeast, listing of 371

oxidoreductions, in the presence of cyclodextrins 425ff - using organic solvents 423ff oxymorphone, transformation by Pseudomonas putida 347f oxynitrilases, enannoselectivity of 301 - Prirnics amygdalus 301

P

PADH see Pseudomonas sp. alcohol dehydrogenase papain 233ff - ester hydrolysis 233f - esterification of N-protected amino acids 235 - formation of amides 244f - hydrolytic resolution of unnatural amino acid derivatives 234 - preferred cleavage sites 249 Papaver somniferum, opium 342 PASTEUR,large-scale fermenter 7 - methods for resolving the enantiomers in a racemate 6ff PASTEURMemorial Lecture 6f PCL see Pseudomonas cepacia lipase penicillin acylase 229ff - amidase activity 229 - esterase activity 229 - kinetic resolution of, phenyl acetyl esters of secondary alcohols 231 - - primary and secondary alcohols 230 - - pyridyl acetate esters of secondary alcohols 231f - of Escherichia coli 15 - substrate tuning 231 - use in selective protecting group strategy 232 penicillin G acylase (PGA), from Escherichia coli 258f - formation of ampicillin 259 - immobilized, formation of loracarbef 259f penicillins, conversion to cephalosporin derivatives 16f - obtained from fermentation 15 - semisynthetic - 15 - synthesis 258ff Penicillium frequentans, esterase of 221 pepsin, preferred cleavage sites 249 peptidases see proteases peptide bond formation, kinetically controlled-, by Alcalase 251 - - coupling involving dehydroamino acids 253f - - coupling involving proline 253 peptide coupling, with minimal protection 255 peptide formation, kinetically controlled -, in anhydrous alcohols 251 peptide synthesis, use of CLECs 256

- use of immobilized enzymes 255

peptides, protease-catalyzed formation 247

- protease-catalyzed hydrolysis 247ff

peracids, in Baeyer-Villiger reactions 538f peroxidases, hydroxylation by 504f PGA see penicillin G acylase pharmaceutical intermediates, enantiomerically pure, use of lipases 146 pharmaceuticals, regulatory environment 393 - world market 393 phenanthrene, hydroxylation of 501 phenols. formation by microbial oxidation of aromatic substrates 499ff phenylacetyl, as protecting group, in peptide chemistry 261 phenylacetylcarbinol, formation of 9f phenyl acetyl esters of secondary alcohols, kinetic resolution by penicillin acylase 231 phenylalanine dehydrogenases, applications of 446 - characteristics of 446 2-phenyloxazolin-5-ones, lipase-catalyzed ring opening 116 phosphines and phosphine oxides, kinetic resolution with cholesterol esterase 217f phytic acid. hydrolysis by yeast phosphatases 379 Pichia miso, esterase of 219 pig liver, flavin dependent monooxygenases 575 pig liver alcohol dehydrogenase (PLADH). applications of 452 - characteristics of 452 pig liver esterase (PLE) 194ff - active site model 207ff - asymmetrization of, cyclic meso-1,2-dicarboxylates 196f - - cyclohexene meso-1,2-dicarboxylates 197 - - cyclopentane meso-1,3-dicarboxylates 197f - - diacylated cyclopentene meso-diols 198f - - prochiral glutarates 195f - - prochiral malonates 195 - - tricyclic meso-diesters 197ff - chemoselective hydrolysis by 207 - enantioselective hydrolysis of, meso-diesters 196ff - - prochiral diesters 195f - kinetic resolution of, quaternary centers 202 - - racemic esters 198ff - - racemic methyl alkylphenylphosphinoylacetates 206 - - racemic acyclic and cyclic 1,2-diols 203 - - racemic cyclic diesters 201 - - racemic E-caprolactones 201f - - secondary alcohols 202 - - tertiary alcohols 202 - polyethylene glycol monomethyl ether (MPEG) linked -, enantioselective acylation of mesodiols 207

Index

practical considerations 207 recognition of planar chirality 205 regioselective hydrolysis by 206 use in the generation of planar chirality. in tricarboxylchromium compounds 198 pig lung, flavin dependent monooxygenases 575 PKA see porcine kidney aminoacylase PLADH see pig liver alcohol dehydrogenase planar chirality, recognition by pig liver esterase 205 plant cell cultures, hydroxylation of steroids 489 plants, alkaloids from 330f - - use of cell culture 331 - - use of metabolic engineering 331 - nitrilase activity 299 PLE see pig liver esterase polycyclic aromatic hydrocarbons, hydroxylation of 501 polyesters, lipase-catalyzed degradation 145 - lipase-catalyzed polymerization 144 polymerizations, lipase-catalyzed - 144f polyunsaturated fatty acids (PUFAs) 137 - lipase selectivities 143f porcine kidney aminoacylase (PKA) 2458' - hydrolysis of N-terminal amides 246ff - resolution of a-amino acids 246f porcine lipoxygenase, hydroperoxide formation 503f porcine pancreatic lipase (PPL) 263f - catalysis of asymmetric syntheses 67f - dipeptide formation 262 - enantioselective reactions catalyzed by primary alcohols 91f - general features 44 - resolution of, acyclic secondary alcohols 76 - - 2-alkanols 75 - - cyclic secondary alcohols 77 - selective acylation of glucose 119 potato tuber lipoxygenase, hydroperoxide formation 503 PPL (porcine pancreatic lipase) 42f precocene 11, dihydroxylation of 515 Prelog's rule 427f - asymmetric reduction of carbonyl compounds 428 - for carbonyl reduction 380 primary alcohols, acylation in, modified sugars 121ff, 121ff - - unmodified sugars 119ff - enantioselective reactions, catalyzed by PCL 89 - - catalyzed by PPL 91f - kinetic resolution by penicillin acylase 230ff - lipase-catalyzed enantioselective reactions, alcohols with quaternary stereocenters 95 - lipase enantioselectivity 87ff product inhibition, of alcohol dehydrogenases 429f -

603

productivity number, definition of 366 product recovery, from reverse micelles 49 - yeast biotransformations 383f progesterone, conversion to testosterone 542 - hydroxylation of 488, 505 - - by Rhizopus arrhizus 477 - 11-a-hydroxylation 12f prolyl dipeptidyl-aminopeptidase, from Lnctococcus lncris 256 propranolol, enantiomeric purity of 84 - lipase-catalyzed routes to 88 prostaglandin synthesis, use of cyclopentane monoester 198 protease-catalyzed reactions 247ff proteases 244ff - aspartyl proteases 248 - classification 247 - metalloproteases 248f - nomenclature 244 - - for the cleavage of 249 - preferred cleavage sites 249 - serine proteases 248 - synthesis of L,L-diastereomers 253 - thioproteases 248 - with esterase activity 221ff protein engineering, of nitrile transforming enzymes 314 Prunrcs amygdalus, oxynitrilase of 301 pseudomonads, amidase activity 294 - nitrile hydratase activity 294 Pseudomonas, atropine metabolism 335f - cocaine metabolism 337 - formate dehydrogenase 403 - nitrilase activity 297 Pseudomonas cepacia lipase (PCL) 41ff - Baeyer-Villiger biotransformations 568 - catalysis of asymmetric syntheses 67f - enantiopreference of 87 - - empirical rules 88 - enantioselective reactions catalyzed by primary alcohols 89f - enantioselective reactions of carboxylic acids, with a stereocenter at the a-carbon 106 - general features 45 Pseudomonas lipases 76f - enantiopreference of 87 - enantioselective reactions of cyclic secondary alcohols 82ff - enantioselective reactions with racemic acyclic alcohols 79ff - resolution of 2-alkanols 78 Pseudornonas putida, aminopeptidase, asymmetric hydrolysis of lupinic acid amide 245f - - resolution of racemic a-amino acids 245 - Baeyer-Villiger biotransformations 555ff - camphor metabolism 543f

604

Index

- dihydroxylation of substituted monocyclic

arenes 506f - dioxygenase reactions 506 - morphine dehydrogenase 348f - transformations of morphine alkaloids 346ff Pseudomonas sp., sulfoxidations by 574 Pserrdomonas sp. alcohol dehydrogenase (PADH). applications of 453 - preparation of chiral hydroxy compounds 453 - synthesis of L-carnitine 453 PUFAs see polyunsaturated fatty acids pyrazines, hydroxylation of 502 pyrene, hydroxylation of 501 pyridines, hydroxylation of 501f pyridyl acetyl esters of secondary alcohols, kinetic resolution of 232 pyrimidine nucleosides, regioselective hydrolysis, by subtilisin 223

Rhizopus arrhizus, hydroxylation of progesterone 477 Rhizopus nigricans, esterase of 219 Rhizopus oryzae lipase (ROL) 41ff rhodococci, nitrilase activity 295 - nitrile hydratase activity 284ff Rhodococcus coprophilus, Baeyer-Villiger biotransformations 560 Rhodococcus eryrhropolis carbonyl reductase, applications 456 - characteristics of 456 Rhodococcus fasciens, Baeyer-Villiger biotransformations 560 riboflavin, structure 536 ricinine, bioconversion to carboxylic acid compounds 297 RML see Rhizomucor miehei lipase ROL see Rhitoprcs oryzae lipase

Q

S

quaternary stereocenters, of alcohols 94f - of carboxylic acids 109 questin, Baeyer-Villiger oxidation 548 - ring cleavage of 543 quinoline, hydroxylation of 502

R

random mutagenesis, of lipases 149f rapamycin derivatives 133 raspberry ketone, Baeyer-Villiger degradation by Beauveria bassiana 548 - formation of 377 Rauwolfia serpenrina, biosynthetic pathway of ajmaline 353f - strictosidine synthase 352 reactor systems, choice of, for enzymatic processes 431ff reductases see also dehydrogenases, oxidoreductases - substrate spectra 437 remote stereocenters, of alcohols 97 - of carboxylic acids 109 resting cells, in yeast biotransformations, C=C double bond reduction 369 reticuline 338f - biosynthesis 339f reverse micelles, lipases in 48f - product recovery from 49 - stabilization by anionic surfactants 48f Rhizomucor lipases, resolution of secondary alcohols 77 Rhizomucor miehei lipase (RML) 42f - enantioselective reactions catalyzed by secondary alcohols 85 - general features 44

Saccharomyces carlsbergensis see brewers' yeast Saccharomyces cerevisiae see bakers' yeast sanguinarine, biosynthetic pathway 338ff scopolamine, biosynthesis of 332ff screening 31 - of lipases 149f secondary alcohols, acyclic -, resolution by PPL 76 - as lipase substrates 68ff - - binding site 70 - cyclic -, enantioselective reactions catalyzed by Pseudomonas lipases 82ff - - resolution by CRL 74 - - resolution by PPL 77 - enantioselective reactions catalyzed by RML 85 - enantioselectivity of, CAL-B 71f - - CRL 72ff - kinetic resolution of, by penicillin acylase 230ff - - by pig liver esterase 202 - phenyl acetyl esters of 231 - pyridyl acetyl esters of 232 - resolution of acyclic -, by CAL-B 72 - - by CRL 73 secondary alcohols (racemic-), fastest reacting enantiomers with cholesterol esterase 216 selenides, oxidation to selenoxides by cyclohexanone monooxygenase 574 sequential kinetic resolution, lipase-catalyzed 64 serine proteases, characteristics 248 sesquiterpenes, dihydroxylation of 513f - hydroxylation of 492f site-selective mutagenesis, for optimization of enzyme activity 26 solvation effects, on enzyme activity 252

Index

solvent engineering 226 soybean lipoxygenase, commercial availability 482 - hydroperoxide formation 502f - synthesis of chiral alcohols by 503 stereocenters, of alcohols, non-carbon - Y8 - - quaternary - 94f - - remote - 97 - of carboxylic acids 103ff - - quaternary - 109 - - remote - 109f - - sulfur - 109f stereoselectivity pocket of lipases 48 steroid hydroxylases 505 steroid manufacture 1Iff steroid nucleus. biotransformation by whole-cell fermentations 13 - microbial oxidation of 12 steroids, Baeyer-Villiger reaction 541 - C-H bond of, microbial oxidation of 12 - combined chemical and microbiological synthesis, of estrone 14 - hydroxylation of 19, 479. 488ff, 505 - - by plant cell cultures 489 - natural -, used in the manufacture of pharmaceuticals 12 - prochiral reduction of 19 - side chain oxidation of 19 - synthesis of, potential use of hydroxysteroid dehydrogenases 448 stigmasterol, biotransformation of 13 strictosidine. biosynthesis of 352 strictosidine synthase 352 structured triacalglycerides (STs) 135 strychnine structure 353 substrate inhibition. of alcohol dehydrogenases 429f substrate modification, improvement of enzyme selectivity 380f substrate tuning 231 subtiligase, cyclization of linear peptides 256 subtilisin 221ff - acylation of, castanospermine 225 - disaccharides 124 - - sucrose 225 - esterase activity of 221ff - peptide bond formation. kinetically controlled- 251 - preferred cleavage sites 249 - regioselective hydrolysis of a pyrimidine nucleoside 223 - selective acylation of carbohydrates 123f - solvent engineering 226 - stability in anhydrous organic solvents 224 - variants 226 subtilisin Carlsberg 251 - acylation of primary amines 257

605

- kinetic resolution of P-sulfonamidopropionic

acid esters 222 sucrose, acylation by subtilisin 225 sugar esters, food use as nonionic surfactants 120 sugars, lipase-catalyzed acylations of, modified 121ff - - unmodified - 119 sulfides, biotransformation by isolated enzymes 576 - oxidation to sulfoxides by cyclohexanone monooxygenase 572 P-sulfonamidopropionic acid esters, kinetic resolution by subtilisin 222 sulfoxidations see also asymmetric sulfoxidations - asymmetric -, flavin dependent monooxygenase enzymes 569ff - active-site model 570ff - proposed mechanism for 570 sulfur etherocycles, improvement of reduction efficiency in yeast 381 sulfur stereocenters of carboxylic acids 109 sulindac, redox reaction 576 supercritical C 0 2 reactor, diagram of 62 supercritical fluids, lipase activity in 60ff surfactant-coated lipases 53 surfactants, prepared by lipase-mediated reactions 120 - stabilization of reverse micelles 48f

T Tanabe process, of diltiazem synthesis 147 taxol, from Tax~tsbrevifolia 330 Taxits brevifolin 330 TBADH see Thermonnnerobium brockii alcohol dehydrogenase terpenes see also diterpenes, monoterpenes. sesquiterpenes - cyclic -, hydroxylation of 492 - hydroxylation of 479, 491ff terphenyl, hydroxylation of 500 tertiary alcohols, kinetic resolution by pig liver esterase 202 - lipase-catalyzed enantioselective reactions 94 testosterone, hydroxylation of 489 thebaine, transformations by Trnmeres snngrtinea 344 Thermonnaerobium brockii alcohol dehydrogenase (TBADH).applications of 441f - Baeyer-Villiger biotransformations 568 - secondary - 403 - thermostability of 441 - tolerance to organic solvents 441 thermodynamic water activity, control methods 59f thermolysin, preferred cleavage sites 249

606

Index

thermostability, of lipases, directed evolution of 149f thiolactone inhibitors, of cyclohexanone monooxygenase 573 thiols, lipase-catalyzed enantioselective reactions of 102f thioproteases, characteristics 248 Torulopsis candida, nitrilase activity 299 total turnover number, definition 399 Trametes sangrtinea, transformations of, neopinone 344 - - thebaine 344 transesterification, of racemic hydroxyacid esters 416ff triacylglycerides, containing medium chain fatty acids (MLMs) 136 - containing PUFAs 137 - glycerolysis to l(3)-MAGS 141 - hydrolysis or alcoholysis to MAGS 141 - lipase-catalyzed reactions 135 - structured - (STs) 135 tridecanone, degradation of 542 triglycerides, enantioselectivity of lipases 91ff triolein, structure 194 tropane alkaloids 331ff - biosynthesis of, hyoscyamine 3328 - - scopolamine 332ff - - tropic acid 333 - metabolic pathways, atropine 335f - - cocaine 337 - microbial metabolism of 335 - parasympathetic inhibition 331f - structures 332 tropic acid, biosynthesis of 333 tropinol dehydrogenase 335ff tropinone reductases 334 trypsin, preferred cleavage sites 249 turnover number, definition 31 - definition of 398f V vinblastine, structure 353 - treatment of leukemia 354 vincamine, structure 353 - use as vasodilator 352 vincristine, structure 353 - treatment of leukemia 354 vindoline, biosynthetic pathways, in Catharanthus 354f vitamin C (L-ascorbic acid), chemical synthesis of 10 - manufacture of 8 vitamin D3 hydroxylation 485

W wastes, nitrile containing -, bioremediation of 312f whole-cell biotransformations, dehydrogenases 396ff - hydroxylation reactions 483ff - of steroid nucleus 13 - practical aspects 26ff, 31f whole-cell catalysts, for dihydroxylation 481f - for hydroxylation 481f

x

Xanrhobacter autotrophicus, Baeyer-Villiger biotransformations 561, 566

Y YADH see yeast alcohol dehydrogenases

yeast see also bakers’ yeast, brewers’ yeast

- as economical source of enzymes 364f - C-C bond formation 378 - C=C double bond reduction 369ff, 375ff - - mechanism 372

- enoate reductase, reduction of triple substituted

double bonds 375

- - substrates for 375 - expression of cyt. P-450

483

- fatty acid synthetase cycle 373

- genetically modified - 383 - hydrolytic enzymes 379 - improvement of reduction efficiency, use of sulfur etherocycles 381

- NADPH regeneration system 367 - reducing enzymes, from the fatty acid synthet-

ase complex 373f

- reducing enzymes of 370ff - reduction of a-diketones 382 - reproducibility of biotransformation 365f - resting cells, biotransformations by 369 - sources of 31f - - influence on catalytic activity 366 - sterol cyclase, lanosterol cyclization 378 - viability 29

yeast alcohol dehydrogenases (YADH) 371,439ff

- see also alcohol dehydrogenases - applications of 440f - characteristics of 439f - enantioselective reduction of enones 440 - selective reduction of an a$-unsaturated

ketone 371f

- sources of 439ff - use for coenzyme regeneration 440

yeast biotransformations 363ff

- enzymatic activities 370ff - operational conditions 366ff - preparation of chiral building blocks 365

Index

- product recovery 383f - reproducibility of 365f

selectivity of 379ff - - improvement by selective enzyme inhibition 382f - - improvement by substrate modification 380f - use of cell immobilization techniques 369 - use of genetically modified yeast 383 - use of organic solvents 369f - use of resting cells, C=C double bond reduction 369 - using ethanol as energy source, NADP + regeneration in the reduction of 3-0x0-butyrate to 3hydroxy-butyrate 368 -

607

- - NAD' regeneration in the reduction of ace-

to1 to propylene glycol 368 yeast cells, asymmetric reductions by 393 yeast fermentation, endogeneous consumption of reducing power 367 - energy source, carbohydrates 366f - - ethanol 367f yeast oxidoreductases, listing of 371 yeast phosphatases, hydrolysis of phytic acid 379 yeast reductions, NAD + regeneration cycle 368 yohimbine, structure 353

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