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.
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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
Biotechnology Second, Completely Revised Edition H.-J. Rehm and G. Reed copyrightOWILEY-VCH Verlag GmbH, 1998
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
f
8
1 Perspectives in Biotransformations
MICHAELIS and MENTENset out their steady state analysis of enzymatic reactions (MICHAELIS and MENTEN, 1913). The first benefit for the chemical industry from the nineteenth century microbiological developments which followed PASTEUR'S creation of a scientific microbiology (e.g.,PASTEUR, 1876) is usually associated, quite properly, with the medical advances. Equally important for industry was the effect on large-scale fermentations.This did not lie in the engineering studies which he attempted for the design of fermenters (Fig. 2) and which had little immediate influence on the industry (SYKES,1845). Of much greater importance was the realization that the course of a fermentation depended on the organism which it contained. This had a dramatic impact on the reliability of the brewing industry in particular. It also laid the foundation for the development of other fermenta-
H,, Raney Ni HO
~
tion products, which by 1920 were supplying the chemical industry with hydroxyacids and solvents such as acetone and glycerol (CHAPMAN,1921), as well as the textile and brewing industries with enzymes such as amylases and proteases (NIEDLEMAN, 1990).
2.2 Fermentations and Applied Biocatalysis Although it is now usual to separate fermentation from applied biocatalysis a broad introduction ought not to do so. A viable microorganism is still the preferred catalyst in many bioconversions. The chemical distinction between the fermentation of glucose (2)into ethanol, and the oxidation of D-sorbitol (D-glucitol) (3) to L-sorbose (4) (Fig. 3) is one of
!IoH Acetobacter suboxydans
OH
CHO
3 D-Sorbitol (D-Glucitol)
HO
4 L-Sorbose
)=.
0
COOH
H+
Kh4n04
1
- OH
- 0
OH
1
OH
OH
COOH
OH
0
OH
2 D-Glucose
HO
OH
7
5
6
*Ho*
8 L-ascorbic acid, Vitamin C
HO
OH
Fig. 3. The manufacture of vitamin C.
2 Perspective 1895-1935
- Organic Chemistry and Industrial Microbiology
9
degree rather than one of kind. In the fermenOf these processes ethanol, even in its tative conversions with Succharomyces cerevi- manufacture as an organic solvent, predates siue to produce ethanol, the organisms are al- PASTEUR. It owes its early development as an lowed to grow on the substrates before switch- industrial chemical to innovations in the cheming to the conversions proper. Similarly in the ical engineering of the distillation process 1935),but the continued unrelilatter instance Acefobacfersuboxyduns is al- (UNDERWOOD, lowed to grow on the D-sorbitol(3) before the ability of the fermentation stage is usually research in process switches to its oxidation. The essential credited with starting PASTEUR’S difference is that there are several catalytic microbiology. Other products benefited directsteps between substrate and product in the ly from the results of that microbiology which former example while there is only one in the made sterile fermentation sufficiently reliable latter. The usual distinction made is that the to form the basis of a manufacturing process. chemical structures of the substrate and the Innovations in engineering design were also product resemble one another in a bioconver- essential before it was possible to operate othsion, but do not in a fermentation.This is a fine er large-scale processes for hydroxyacids and distinction, which if it were pressed too far for solvents which, in some instances, were would miss some important lessons about the highly flammable. All of the problems assoapplication of biocatalysis to organic synthe- ciated with sterile fermentations had to be 1971):how to manufacture ses. Both conversions are characteristic of the solved (HASTINGS, manufacturing processes which arose from the research which FRANKLAND described. A range of industrial fermentation processes for organic chemicals was reviewed by CHAPMAN in 1921 (Tab. 1). In the decade after CHAPMAN’Sreview, the industrial foundations were laid for the essentially fermentative processes of the oxidation of D-sorbitol (3) to L-sorbose (4) (BERTRAND, 1904) and the synthesis of 9 10 phenylacetylcarbinol ((R)-1-phenyl-1-hydroxyPhenylacetylcarbinol D-Ephedrine propan-2-one) (9) (Fig. 4) from benzaldehyde Fig. 4. Early fermentation products. and pyruvate (NEUBERG and OHLE,1922).
Tab. 1. Scale of Manufacture of Fermentation Products (1910-1929) Fermentation Product Ethanol Glycerol Acetone Butanol D- and L-Lactate Citric acid
Basis for comparison of manufacture Country
Year
US manufacture
1917
Annual tonnage
0.17 x lo6 (BEDFORD, 1921) German manufacture 1917 25 000 (CHAPMAN, 1921;HABER,1971) UK and Canadian manufacture 1918 approx.3000 (CHAPMAN, 1921; HABER,1971) Butanol production should be double that of acetone 1909 1500 UK consumption UK imports 1916 720 (CHAPMAN, 1921) 1929 3200 US manufacture (PRESCOTT and DUNN,1949)
I Perspectives in Biotransformations
10
a vessel of 125 m3 capacity with suitable seals and gaskets to allow its sterilization with steam at 121°C; how to lay out pipework without dead legs where contaminating organisms accumulate; how to design valves and pumps which provide a sterile seal when transferring inocula; how to manage foaming and, if this were not enough, there is the fire hazard from flammable exhaust gases. The solution to these problems came from a collaboration between microbiologists and engineers which went far beyond what was required for the production of beer and wine. The innovations were driven by the demands of industries as diverse as textiles, foods and beverages, and munitions. The markets for the two remaining processes, for L-sorbose (4) (seeFig. 3) and for phenylacetylcarbinol (9) (Fig. 4), were as the pharmaceuticals ascorbic acid (8) (see Fig. 3) and D-ephedrine (10) (Fig.4).The first (REICHSTEIN and GROSSNER, 1934) transforms a synthesis from an inconvenient starting material, L-xylosone (11) (Fig. 5), into one (see Fig. 3) whose input, D-glucose (2), is cheap and convenient. This, and the innovative chemistry of the acetal protection of the carbohydrates (5, 6) (see Fig. 3), make it a particularly good example of how microbiology can be integrated into a chemical synthesis. In the second the crude product obtained by fermenting benzaldehyde with a sugar solution is condensed with methylamine and reduced to form Dephedrine (10) (Fig. 4) (HILDEBRANDT and KLAVEHN, 1930). Both take the use of microbiology beyond the kinetic resolutions of racemates which FRANKLAND described into the
I!,-
0 HO
OH
OH
1 1
zgr CaCl,
more profitable field of asymmetric synthesis. Neither would be practical without the earlier improvements in fermentation technology.
3 Perspective 1935-1955 - A Fallow Time After this promising start the technology failed to grow. Apart from the two exceptional examples mentioned above it languished as a method for large-scale manufacture. What CHAPMAN foresaw in his review was superceded by alternative processes based on petrochemistry.This is clearly illustrated in the manufacture of ethanol where fermentation was replaced by the hydration of ethylene (TURNER,1995), and in the manufacture of acetone and butanol (SANTANGELO and DURRE,1996). Only where petrochemistry could not devise an alternative approach did the fermentation processes continue to grow. The fermentation of citric acid is a good example whose output has grown by an average of 8% a year since 1920 (Fig. 6), and the processes for ascorbic acid (8) (see Fig. 3) and D-ephedrine (10) (see Fig. 4) manufacture have remained essentially as they were originally developed. What is striking is that no new processes were introduced with the signal exceptions of the fermentation for the secondary metabolites such as penicillins G and V (35a, 35b) (see Fig. 13) and streptomycin A which had antibiotic activity.
CN
+H OH
OH
H2°
~
un
HO
OH 12
L-Xylosone
Fig. 5. Chemical synthesis of vitamin C.
-un 13
HCl 4O-5O0C
HO
OH
8 L-ascorbic acid,
Vitamin C
4 Perspective 1955-1975 - Functionalization of Natural Products
lo6 I Q,
m
0)
c C
10’
-
0
c,
m3
I
E
z
lo4 1o3
1920 1940 1960 1980 200
Year
Fig. 6. The growth in the manufacture of citric acid.
The prevailing view in the 1950s is encapsulated in the listing, in one of the standard textbooks of organic chemistry (FINAR,1956), of the available methods for resolving racemates. The conversion of the enantiomers into diastereomers, PASTEUR’S second method, is the one of choice. It was widely applicable with a number of well-tested methods. In contrast the biochemical separations with microorganisms are noted to suffer from several disadvantages: they require dilute solutions of the racemate, one enantiomer is always destroyed, and the other may be partially so, and it is necessary to find a suitable organism. These are still the common and often justifiable complaints of the chemist, nevertheless the jaundiced view of the process is very different from FRANKLAND’S enthusiastic description. One other enzymatic method for resolving amino acids is listed, in which papain catalyzes the formation of an insoluble N-acyl-anilide derivative (BERGMANN and FRAENKEL-CONRAT, 1937; ALBERTSON, 1951), a method reminiscent of the current manufacturing process for aspartame (OYAMA, 1992). The potential for innovation was frustrated and two explanations for the changed attitudes since PASTEUR’S death are worth advancing. The successful application of biocatalysis requires inputs from both chemistry and biology. The first half of the twentieth century saw the fragmentation of previously unified disci-
11
plines. An organic chemist of FRANKLAND’S generation would have been more familiar with the techniques of microbiology than those who followed him. The divisions between organic chemistry, microbiology, and biochemistry, which were essential for the advances of each of these disciplines, were precisely the opposite of what is required for the successful application of biocatalysis in chemical synthesis. In the nineteenth century one individual could cover all; even by the time the processes for ascorbic acid (8) (see Fig. 3) and D-ephedrine (10) (see Fig. 4)manufacture were introduced this was less likely. Organic chemists turned to the problems of petrochemistry, and, where necessary, chemical methods of asymmetric synthesis,leaving to the biochemist and the microbiologist much of the natural product chemistry which was once their stock-in-trade. As a result chemists would turn to biochemical methods almost as a last resort. A second reason may have been a shortage of suitable markets for the chiral products for whose synthesis applied biocatalysis is so effective.This was to change with the further development of a pharmaceutical industry based on analogs of complex natural products.
4 Perspective 1955-1975 - Functionalization of Natural Products There followed a period of about 20 years in which the problems set by steroid and by plactam chemistry stimulated a further period of process development. The method remained the specialized ground of a few groups, but their achievements were of great commercial importance.
4.1 Steroid Manufacture The pharmaceutical development of the steroids was possible only because of the invention of methods for transforming plant products such as diosgenin (14) (Fig. 7; MAR-
12
1 Perspectives in Biotransformations
n
1 5 Hecogenin
HO
Fig. 7. Natural steroids used in the manufacture of pharmaceuticals. KER, 1940).In the first phase of this work entry into a full range of products was blocked by the failure to oxidize the C-11 position which would allow the synthesis of analogs of corticosteroids. The search for a solution took two parallel strands.
In one a search was made for a natural plant product where the C-ring of the steroid nucleus was already functionalized. This was successful in the discovery of hecogenin (15) (Fig. 7) (MARKERand APPLEZWEIG, 1949; SPENSLEY, 1952). Although the molecule is oxidized at C-12 the functionality is easily moved to the adjacent C-11 position. The alternative process developed from studies of the microbial oxidation of the steroid nucleus. Although microorganisms were known to oxidize the hydroxyl groups on the nucleus, direct oxidation of the carbon skeleton was first observed in 1947. Cholesterol (16) was oxidized to cholestenone (17),7-ketoand 6-methylheptanone (19) cholesterol (U), derived from cleavage of the side chain (HORVATH and KRAMLI,1947). Some chemists were dubious of the microbial catalysis but proper controls confirmed the transformation (KRAMLI and HORVATH, 1949) (Fig. 8).There followed the steady development of a range of important steroid transformations of which the 11-ahydroxylation of progesterone (20a) (Fig. 9) was the first to be reported (PETERSEN and MURRAY, 1952).Although the hydroxyl group has the wrong orientation compared to cortisol (21)(Fig. 9) (11-a rather than 11-p)this is readily inverted. Subsequently other micro-
1 6 Cholesterol
Fig. 8. The first microbial oxidation of a steroid C-H bond.
1 9
4 Perspective 19s-1975 - Functionalization of Natural Products
20
aR=H bR=OH
2 1 Cortisol
Fig. 9. Progesterone (20a) and cortisol (21).
organisms were found which inserted the hydroxyl group with the correct orientation but their regioselectivity for the oxidation only at C-11 was less strict and other products were formed, notably the 9-(Yand 14-a products. The screens for catalysis in these processes were based on what was then the new method of paper chromatography (ZAFFARONI et al., 1950). As an analytical method it allowed the separation of closely related molecules, and this is crucial for the analysis of the microbial transformation. Although the throughput in the screen could not have been large this did facilitate the problem of locating the correct organisms, and so overcame one of the major hurdles in the use of the technology. In the early 1970s political pressures in Mexico on the supply of diosgenin (14) (see Fig. 7) collected from the wild led to a large increase in its price (LENZ,1983) and to an increase in the use of steroids derived from soya oil. From these stigmasterol (22) (Fig. 10) provided a useful alternative to the chemical routes to progesterone (20a) (see Fig. 9) and thence to the cortiosteroids through microbial oxidation. A microbial oxidation of cholesterol
13
which degraded the 17-side chain to a 17-keto nucleus (SIHet al., 1965) provided a route for the synthesis of the estrone (33) (see Fig. 11) nucleus. This was established as an industrial process in Japan in 1976 shortly after the same oxidation had been applied to p-sitosterol(23) (DJERASSI, 1976), which otherwise lay unused as a waste product from soya oil after the extraction of stigmasterol (22) (Fig. 10). By the mid- 1970s the wide range of microbial transformations which were known to affect the steroid nucleus included hydroxylation, reversible oxidation of the alcohols, desaturation and aromatization, Baeyer-Villiger oxidation in both the rings and in the side chain, and aldol fission of the side chain itself (SIH et al., 1965; JONES,1973; JONESet al., 1975). All are essentially whole-cell fermentations not dissimilar in concept from the earlier processes for L-sorbose (4) (see Fig. 3) and phenylacetylcarbinol (9) (see Fig. 4) synthesis. The value of these transformations lies in their regioselectivity. Their development was prompted by the inability of chemistry to match this performance. Although the microbial processes are slow, and operate at low concentrations of substrate the value of the products as synthons for further chemical modification is sufficient to overcome these shortcomings. Remote functionalization with chemical derivatives was considered as a possible alternative (BRESLOW, 1980) but it could not compete with the microbial fermentations. The pharmaceutical value of the steroids also led to methods for their manufacture by
2 2 Stigmasterol 23 22.23-Dihydrostigmasterol = 6-Sitosterol
Fig. 10. Stigmasterol (22) and p-sitosterol(23).
1 Perspectives in Biotransformations
14
1 CH3OH
2 SOCl*
0
o3
1 Hz C=CHz, AlCl
. )
CH3C02
4
C H 3 4 CO
2 Et3N
25
24
1 (BzO),
5 26
NaOCH,
CZHSOZC
28
27
3 steps
Fig. 11. Combined chemical and microbiological synthesis of estrone (33).
total synthesis. These processes are particularly useful in giving access to the 18-methyl steroids (i.e., 13-ethyl). The synthesis of estrone (33) (Fig. 11) is an interesting example of a composite chemical and microbiological synthesis. Two convergent chemical sequences (24 to 26 and 27 to 29) yielded the prochiral diketone (30),which was reduced enantioselectively by Rhizopus arrhizus at the pro-(S)-ketone group to give the ketoalcohol (31) in 70% yield. Robinson annulation and elaboration of the aromatic ring gave estrone (33) (NOMINE, 1980). The technology developed for the steroid hydroxylations was applied to the manufacture of the antiparasitic agent oxamniquine
(34b). In this synthesis the organism Aspergillus sclerotiorum hydroxylates a benzylic carbon attached to a quinoline nucleus (34a) (RICHARDS, 1974) (Fig. 12).
H 3 4
aR= bR=OH
2 A.
I
sclerotiorum
Fig. U. Manufacture of the antiparasitic agent oxamniquine.
4 Perspective 1955-1975 - Functionalitation of Natural Products
15
practical value. An effective enzyme with specificity directed at penicillin V (35b) was discovered much later in the fungus Pleurotus and ROEHR,1976). Fermentation produces only two penicillins, osrreurus (SCHNEIDER Following the discovery of a chemical methG and V (35a, 35b) in good yield. 6-Aminopenicillanic acid (35c, 6-APA) which is the key od for the removal of the 7-~-aminoadipyl synthon in the manufacture of the semisyn- side chain from cephalosporin C (36a) (see thetic penicillins, is only ever a minor fermen- Fig. 14) with NOCl (MORINet al., 1962) a protation component (Fig. 13). Nor is it easily ac- cess with PCl, was developed for the peniciland cessible by simple chemical hydrolysis because lins G and V (35a, 35b) (WEISSENBURGER 1970).The choice between the p-lactam ring is unstable at extremes of VAN DER HOEVEN, pH. The search for a convenient route to 6- the chemical and the enzymatic processes for APA (3%) became a spur for the further appli- 6-APA (35c) manufacture was then finely balanced among the interested companies for cation of biocatalysis. The first indication that a microbial method several reasons. Competition between the varmight be possible came from Japan (SAKAGU- ious patents acted as a block on one or the other of the processes; there was no established CHI and MURAO,1950), but a successful process had to await the discovery of strains of enzymology for hydrolyzing penicillin V (35b) Escherichiu coli containing an amidohydro- which was the alternative fermentation prodlase, now usually known as penicillin acylase, uct; and there was a lower level of expertise in which would selectively cleave the 6-amido handling large-scale enzymology compared to linkage without affecting the cyclic amide in chemistry. Despite the fact that the pharmaceutical inet al., 1960; ROLINthe p-lactam ring (HUANG SON et al., 1960;KAUFMANN and BAUER,1964). dustry was familiar with fermentation processThe enzyme is an effective catalyst only for the es their subsequent handling of the fermented hydrolysis of penicillin G (35a); penicillin V products was almost exclusively chemical. (35b) is hydrolyzed, but at less than 20% of the Where reliable long-term operation is essenrate of penicillin G (35a).Alternative enzymes tial the conservative attitudes in process dewere discovered, for example in Streptornyces sign usually reject process innovation. On ecoluvendufue (BATCHELOR et al., 1961),but their nomic terms, with relaxed procedures for yield of catalytic activity was too low to be of waste disposal, there is little to choose between the two processes. The principle cost input is the penicillin itself, so that the economics of the process hang largely on the losses of plactam nucleus. Under these circumstances a processing industry will turn to technology with which it is most familiar. Some publications from this period also describe the use of the amidohydrolase to synthesize a new amide linkage on the free amino group of the penicillin and cephalosporin nuet al. 1960; TAKAHASHI et cleus (KAUFMANN al., 1972) but these generally failed because the yield from the unfavorable equilibria were never sufficiently large to compete with the much more effective chemical derivatization. A number of other p-lactam conversions illustrate the interest in the use of enzymes to manipulate structures whose chemical stability is limited. The hydrolysis of the 3-acetoxymec R = H, 6-Aminopenicillanic acid ( 6 - M A ) thy1 group of cephalosporin C (Ma) (Fig. 14) Fig. 13. Penicillins obtained from fermentation. with an esterase from Rhodosporidium toru-
4.2 P-Lactam Manufacture
16
I Perspectives in Biotransforrnations
1974). However attempts (e.g., WALTON, 1963) to find a single enzyme which would remove the 7-~-aminoadipylside chain to produce 7aminocephalosporanic acid (39,7-ACA) were unsuccessful, and the two-enzyme approach with the combined action of an amino acid ox36 idase and an amidohydrolase (Fig. 14) was only available some years later as a possible alR . toruloides ternative (SHIBUYAet al., 1981; ICHIKAWA et al., 1981). An attempt to devise a route from the peniTriginopsis variabilis cillin nucleus into a full range of cephalosporins Fusarium solanum and cephamycins (Fig. 15) also failed. Efficient chemical processes will convert penicillins (40) into 3-methyl cephalosporins (41) (CHAUVEITE et al., 1971), but neither the chemical nor the enzymatic oxidation of the methyl group is effective. The chemical processes (e.g., WEBBERet al., 1969) are difficult to operate at a large scale, and the biological hydroxylation with a dioxygenase is highly specific for the natural D-aminoadipyl side chain (TURNER et al., 1978).Subsequent enzymatic conversion to the carbamate (43) was also impractical because of the limited titer of the enzyme in its et source, Streptomyces clavuligerus (BREWER al., 1980) (Fig. 15). These few examples from the manipulation of the p-lactams illustrate not only the interest COZ H generated in biochemical methods by a new 38 set of chemical problems, but also the difficulPseudomonas spp. ties which faced the development of an enzymatic process. They were readily accepted 0 t where there were no chemical alternatives, but it was difficult to replace existing processes particularly where at their first discovery there 0 were obvious shortcomings in the enzymatic approach, whether through low catalyst titer, inconvenient substrate specificity,or even genCOZ eral unfamiliarity with the technology. The need to provide a multidisciplinary team to re39 7-aminocephalosolve these problems of process technology is sporanic acid, 7-ACA often difficult to justify when their resource Fig. 14. Two-step removal of the ?'-~-arninoadipoyl might be more profitably targeted at new side chain from cephalosporin C (36a). product discovery. Two of the processes were successfully commercialized: those for the manufacture of 6APA ( 3 9 ) (see Fig. 13),and the ring-expanded loides (yellow yeast) allows a range of deriva- cephalosporin (41) (Fig. 15) derived from it, tives to be prepared from the 3-hydroxylme- and for the hydrolysis of the 3-acetoxymethyl thy1 derivative (36b) (SMITHand LARNER, group of cephalosporin C (36a) (see Fig. 14).
c;
1
I
H3NF&0Ac
4 Perspective 1955-1975 - Functionalization of Natural Products
H I
Chemical ring expansion
f-'
B
t
1
60, H
4 1
Biocatalytic or chemical oxidation
R R0N v & o H CO, H
42
Biocatalytic 0-carbamoylation
H I
t
CO2 H
0
43
Fig. 15. The conversion of penicillins to cephalosporin derivatives.
4.3 Enzyme Engineering and the Manufacture of Amino Acids At their inception the steroid and p-lactam transformations were all processes which PASTEURmight have recognized. The conver-
17
sion of the substrate was either catalyzed by the terminal phase of a microbial fermentation, or with washed cells harvested from the fermentation. In this form the enzymatic catalyst was present as a solid phase which could readily be separated from the reaction mixture, even if many contaminants were also eluted from the cells. It is also a measure of the added value dependent on these catalytic stages that the catalyst could be used once only and then be discarded. A more economic process reuses the catalyst, but greater savings arise more from process simplification than from the catalyst itself. Separation processes are expensive, and the cost of recovering a product from a catalyst which is at best the total solids output from a fermentation is likely to be larger than the cost of the catalyst itself. Moreover the purity of the recovered product as an intermediate synthon for pharmaceutical manufacture must be very high, and certainly free of detectable protein from the biocatalyst. The ideal reaction is one which is heterogeneous, with the catalyst and the reagents in different phases so that they are easy to separate. This drives process design towards the use of enzymatic catalysts immobilized in a second phase, and preferably one which is solid. Although there were some antecedents for this concept their application to large-scale biocatalysis was new, and their study was central to this phase of the research. Their development expanded rapidly in the late 1960s (Fig. 16), and became the topic of what was almost a separate discipline of research: enzyme engineering (for reviews see ZABORSKI, 1972; CHIBATA, 1978).The significance for manufacture was in separating the application of biocatalysis from fermentation. New forms of immobilized catalysts encouraged new engineering concepts for process design (LILLYand DUNNILL, 1971) which lowered the cost of the biocatalytic step. While reactant concentrations remained low by the standards of many other chemical processes, the process intensity, in terms of units of product per m3 reactor volume per hour, was much improved. The process savings were useful for the pharmaceutical industry, but they were of greater importance for the application of biocatalysis in the manufacture of less valuable
1 Perspectives in Biotransformations
18 240-
220 c
a
2
200
-
140-
(0
Year
Fig. 16. The increase in studies of immobilized enzymes after 1960 (from CHIBATA, 1978).
I
N -acetylation
4 5
N -deacety lation Aspergillus oryzae racemization
0
4 7
46
Fig. 17. The resolution of N-acetyl methionine.
products. It became sensible to consider the use of biocatalysis in the resolution of compounds such as the amino acids (Figs. 17 and 18). This became the first large-scale use of immobilized enzyme technology (CHIBATA, 1978), but the development of an immobilized enzyme process for 6-APA (35c) (see Fig. 13) manufacture was soon to follow, replacing the cruder whole cell process (SAVIDGE, 1984). Immobilized enzyme technology is presently limited to those biocatalytic processes whose chemistry is simple, and it is not an essential prerequisite of an effective and economic process. This is clear from the continuing use of even the whole-cell biotransformations already listed in this introduction, not to write of others introduced more recently and discussed elsewhere in this volume. The value of such processes lies in the chemistry which can be catalyzed, e.g., where the processes are oxidations or reductions dependent on a complex set of intracellular reactions which mediate the action of oxygen or of organic reductants and cofactors. By 1975 biocatalysis had established itself in a small handful of reactions for organic synthesis (Tab. 2). Of these two were catalyzed by immobilized enzymes. It is notable that all of these processes were developed in the pharmaceutical industry where the three necessary disciplines of chemistry, biology, and engineering had a long history of collaboration.
5 A Current Perspective Collaboration A number of factors are now driving the increased use of biocatalysis. One is quite simply that the range of biocatalytic processes now commercialized is sufficiently wide that a chemist can feel confident in concluding that there are few novel technical barriers to overcome. Although a few processes associated with the food industry also operate on a very large scale, the annual production levels in many other processes now exceed lo3 tonnes (Tab. 3). The problems which remain are more likely to be practical or economic; indeed some
5 A Current Perspective - Collaboration
19
control panel
tank for racemiration
Fig. 18. Flow diagram for the continuous production of L-amino acids by immobilized aminoacylase (from CHIBATA, 1978).
Tab. 2.
Biological Catalysis in the Synthesis of Organic Chemicals (1975)
Reaction type
Process
Catalyst
Steroids Hydroxylation Side chain oxidation Prochiral reduction
Corticosteroid manufacture 17-Ketosteroid manufacture Total synthesis of (13-ethyl)-steroid nucleus
Metabolizing cells Metabolizing cells Metabolizing cells
P-Lactams Amide hydrolysis Ester hydrolysis
6-APA manufacture 3-Hydroxymethyl cephalosporin derivatives
Immobilized enzyme Non-viable cells
Sorbitol oxidation Aldol condensation Benzylic hydroxylation N-Acetyl hydrolysis
Ascorbate manufacture D-Ephedrine manufacture Oxamniquine manufacture L-Amino acid manufacture
Metabolizing cells Metabolizing cells Metabolizing cells Immobilized enzyme
of the problems discussed in earlier sections, particularly over the synthesis of 6-APA (35c) (see Fig. 13) and even for 7-ACA (39) (see Fig. 14), are now resolved in favor of biocatalytic methods. The revolution in molecular genetics which has allowed the genetic base of an organism to be reconstructed has made possible what even quite recently would have been judged impossible. Enzymes and organisms
which were once intractable are now targets for development. The dominant influence for innovation remains the pharmaceutical and agrochemical industries where chirality and molecular shape have a critical effect on the value of the final product, and where the methods of modern biology are well established.
20
I Perspectives in Biotransformations
Tab. 3. Current Examples of Large-Scale Biocatalytic Processes
Process
Catalyst
Product
Hydrolases
Amyloglucosidase Nitrile hydratase Penicillin amidohydrolase
Glucose Acrylamide 6-APA
Hydantoinase Pseudomonas sp. Dehalogenase Sorbitol dehydrogenase Hydrolyase Cyclopentadione reductase Ketosulphone reductase
4-Hydroxyphenylglycine Cysteine (S)-2-Chloropropionate L-Sorbose Carnitine (13-Ethyl)-estradiol Hydroxysulphone Synthon for Trusopt Isoglucose Phenylacetylcarbinol L-Dopa Malate Aspartate Aspartame Insulin P-Cyclodextrin
Resolution (with racemization) (w/o racemization) Oxidation Reduction (sec-alcohol) Isomerization C-C synthesis Achiral precursors Peptide synthesis Glycosyl transfer
Glucose (xyl) isornerase Pyruvate decarboxylase Tyrosine phenol lyase Fumarase Aspartate ammonia lyase Thermolysin Trypsin Cyclodextrin glucanotransferase
Other sectors of the chemical industry have adopted the technology more warily, although there are some interesting exceptions; for example, a few companies use enzymes in the synthesis of polymers (POKORAand CYRUS, 1987;BINNSet al., 1993).One factor which may help to increase the extent of its adoption is the increasing pressure to deliver a cleaner chemistry. Although this is not the prerogative of biocatalysis it is nevertheless an important feature of it. Water which is replacing petrochemical solvents should also be viewed as a limited and expensive resource to be recycled, and certainly not used as a convenient diluent for the discharge of toxic by-products.The aim is to achieve a manufacturing- process which can account for all of its waste-products, and minimize their output. The manufacture of cysteine from 2-chloroacrylic acid rather than from the hydrolysis of hair (SANOand MITSUGI, 1978), of acrylamide without the use of a copper catalyst and high temperature (NAGASAWA et al., 1993), or of isopropylmyristate (MILLERet al., 1988), are recent examples which all point to the immediate benefits of
Annual Tonnage 10 x lo6 8 x lo3 10x 103 1000 500 2000 50 x 103 150 1 (?> 1-3 8 x lo6 300-500 50 50 400 2000 41 800-1500
this cleaner chemistry. In some instances the marketing of these compounds also benefits from the “natural” cachet which is associated with biocatalytic methods of synthesis. One barrier in the transfer of this technology to the wider chemical industry is the lack of the necessary mix of disciplines within the industry. Its biological expertise is weak. Without a good biological input the specificity and selectivity of enzymes becomes a weakness of the technology. Its catalysts appear less generally applicable than do those drawn from a more familiar background. The process of screening for and isolating a catalyst with the correct performance, which is standard practice for biologists, is a problem for chemists. Moreover, unless they are well designed, the intensity of many processes which enzymes catalyze is low so that at first sight the spacetime yield in a reactor compares poorly with the chemical alternatives. To a chemist without an understanding of the biology these do still appear forbidding hurdles. Another barrier is that biologists often pursue inappropriate targets because they pay in-
6 References
sufficient attention to the advances in chemical processing which will displace biocatalytic procedures. New methods of asymmetric synthesis, and chromatography on chiral supports will be powerful challenges to the application of biocatalysis. They may displace current methods just as petrochemistry displaced fermentation in the manufacture of ethanol (TURNER, 1995). Recent developments in genetic and biochemical engineering have made many biological catalysts more easily available and more productive in use, and the increased process intensity of biocatalytic transformations does now allow them to be incorporated into traditional chemical processes. It is now important to show the organic chemists and the chemical engineers in the chemical industry that biocatalysis, which converts material inputs into products, will fit well into the overall processes. The question is how this might be done given the lack of a component discipline in the industry. For most companies the application of biocatalysis is likely to remain an occasional occupation. It is just one of a number of catalytic options which are open so that to build the necessary biology into the company itself would run the risk of creating a department whose skills could only be intermittently employed. The alternative would be to collaborate with some outside agency which could temporarily bring the skills required. The aim must be to use the external skills to build a sufficient knowledge of the process in the chemical company so that the further operation of the catalytic step requires only a minimum of biological expertise which could be readily transferred to the company’s own employees. This is a combined process of education and demonstration which should also allow the viability of the process to be assessed both at a technical and at an economic level. One can add that this is a much more general problem in the development of new technologies as the knowledge base from which they could grow expands at its current rate. Putting disparate technologies back together to create new business opportunities is not a trivial task. Without some idea of where those opportunities lie, and of how they might be fulfilled, there will be new delays in the continued
21
development of the technologies and of their associated experimental sciences. With these thoughts in mind one should wish that this volume will be read not only by many who are familiar with the application of biocatalysis in some appropriate sector of industry, but also by a few in those sectors where it would presently be considered inappropriate. If the technology is to thrive it must break new ground and open up new possibilities for its commercialization. It will then achieve a proper status alongside other methods of catalysis.
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22
I Perspectives in Biotransformations
DJERASSI,C. (1976), Problems of manufacture and distribution. The importance of steroidal contraceptives: technical versus political aspects, Proc. R. SOC.London, B 195,175-186. FINAR,I. L. (1956), Organic Chemistry,Vol.2,Stereochemistry and the chemistry of natural products. London: Longmans, Green. FRANKLAND, P. (1897), Pasteur Memorial Lecture, J. Chem. SOC. 71,683-743. HABER,L. F. (1971), The Chemical Industry, 19001930: International growth and technological change. Oxford, UK: CIarendon Press. HASTINGS, J. J. H. (1971), Development of fermentation industries in Great Britain, Adv. Appl. Microbiol. 14,145. HILDEBRANDT, G., KLAVEHN, W. (1930), Ephedrine. German Patent 584,459. HILL,A. C. (1897), Reversible zymohydrolysis, J. Chem. SOC.73,634-658. HORVATH, J., KRAMLI,A.(1947),Microbiological oxidation of cholesterol with Azotobacter, Nature 160,639. HUANG, H.T., ENGLISH, A. R., SETO,T.A., SHULL, G. M., SOBIN,B. A. (1960), Enzymatic hydrolysis of the side chain of penicillins, J. Am. Chem. SOC.82, 3790-3791. ICHIKAWA, S., SHIBUYA.Y., MATSUMOTO, K., FUJILT., KOMATSU, K., KODAIRA, R. (1981), Purification and properties of 7/3-(4-carboxy-butanamido)cephalosporinic acid acylase produced by mutant derived from Pseudomonas, Agric. Biol. Chem. 45,2231-2236. JONES, E. R. H. (1973),The microbiological hydroxylation .of steroids and related compounds, Pure Appl. Chem. 33,39-52. JONES,E. R.H., MEAKINS, G. D., MINES,J. O., PIAGNELL, J. H., WILKINS, A. L. (1975). Microbiological hydroxylation. Part XIX. The action of an ant fungus (‘Acromyrmex fungus’) on oxygenated androstanes, pregnanes and cholestanes, J. Chem. Soc. Perkin I , 1552-1554. KASTLE,J. H., LOEVENHART, A. S. (1900). Concerning lipase, the fat-splitting enzyme and the reversibility of its action, Am. Chem. J. (Baltimore) 24, 49 1-525. KAUFMANN, W., BAUER,K. (1964). Variety of substrates for a bacterial benzyl penicillin-splitting enzyme, Nature 203,520. KAUFMANN, W., BAUER,K., OFFE,H. A. (1960), Enzymatic cleavage and resynthesis of penicillins, in: Antimicrobial Agents & Chemotherapy (GRAY, S. G., Eds.), pp. 1-5. P., TABENKIN, B., BRADLEY, New York: Plenum Press. KRAMLI,A., HORVATH,J. (1949), Microbial oxidation of sterols, Nature 163,219. LENZ,G. R. (1983), Steroids, in: Kirk-Othmer Encyclopedia of Chemical Technology, 3rd. Edn., 21, 645-729. New York: John Wiley & Sons.
LILLY,M. D., DUNNILL, P. (1971), Biochemical reactors, Process Biochem. 6(8), 29-32. MARKER,R. E. (1940), Sterols. CVIII The preparation of dihdroandrosterone and related compounds from diosgenin and tigogenin, J. Am. Chem. SOC.62,2621-2625. MARKER,R. E., APPLEZWEIG, N. (1949), Steroidal sapogenins as a source for cortical steroids, Chem. Eng. News 27,3348-3349. MICHAELIS,L., MENTEN,M. L. (1913),The kinetics of invertin action, Biochem. Z. 49,333-369. MILLER,C., AUSTIN,H., POSORSKE, L., GONZLEZ,J. (1988), Characteristics of an immobilized lipase for the commercial synthesis of esters, J. Ass. Am. Oil. Chem. 65,927-931. MORIN,R. B., JACKSON, B. G., FLYNN, E. H., ROESKE, R. W. (1962), Chemistry of cephalosporin antibiotics. I. 7-aminocephalosporanic acid from cephalosporin C, J. Am. Chem. SOC.84,3400-3401. NAGASAWA,T., SHIMIZU, H., YAMADA, H. (1993),The superiority of the third-generation catalyst, Rhodococcus rhodochrous J1 nitrile hydratase, for industrial production of acrylamide. Appl. Microbiol. Biotechnot. 40,189-195. NEUBERG, C., OHLE,H. (1922), Biosynthetic carbon chain union in fermentation processes, Biochem. Z. l28,610-618. NIEDLEMAN, S. L. (1990), The archeology of enzymology, in: Biocatalysis (ABRAMOWICZ, D. A., Ed.), pp. 1-24. New York Van Nostrand Reinhold. NOMINE,G. (1980), La place de bioconversions dans I’acces industriel aux steroides, Bull. SOC. Chim. FK (1-2, Pt. 2) 18-23. OYAMA,K. (1992), The industrial production of aspartame, in: The Commercial Manufacture and Application of Optically Active Compounds (COLLINS, A. N., SHELDRAKE, G. N., CROSBY, J., Eds.), pp. 237-248. New York: John Wiley and Sons. PASTEUR, L. (1858), Memoire sur la fermentation de I’acide tartrique, C. R. Acad. Sci. (Pans) 46. 615-618. PASTEUR, L. (1876). Etudes sur la biere, ses maladies, causes qui les provoqicent, procedts pour la rendre inalttrable, aver une thkorie nouvelle de la fermentation. Paris: Gautiers-Villars. D. H., MURRAY, H. C. (1952). Microbial PETERSON, oxygenation of steroids at carbon 11, J. Am. Chem. Soc. 74,1871-1872. POKORA, A. R., CYRUS,W. L. (1987), Phenolic developer resins, U.S. Patent 4,641,952. POTTEVIN,H. (1906). Actions diastasiques reversibles. Formation et dedoublement des ethers-sels sous I’influence des diastases du pancreas, Ann. Inst. Pasteur 20,901-923. PRESCOTT, S. C., DUNN,C. G. (1949), Industrial Microbiology. New York: McGraw Hill.
6 References
23
REICHSTEIN,T., GROSSNER, A. (1934), Eine ergiebige SMITH,A., LARNER,R. W. (1974), Enzymatic hydrolysis of 3-acyloxymethylceph-3-em-4-carboxSynthese der L-Ascorbinsaure (Vitamin C), Helv. ylic acids or salts thereof, British Patenr 1,531,212. Chim.Acta 17,311-328. P. C. (1952), Cortisone from CommonRICHARDS, H. C. (1974), 2-Aminoalkyl tetrahydro- SPENSLEY, wealth sisal, Chem. Drug. 158,84-86. quinolines. U. S.Patenr 3,821,228. ROBERTS, S. M.,TURNER, N. J., WILLETTS, A. J.,TuR- SYKES,W. J. (1895),The indebtedness of brewers to M. PASTEUR, J. Fed. Inst. Brewing 1,498-525. NER. M. K. (1995). Inrroduction to biocatalysis T.. YAMAZAKI, Y., KATO,K.. ISONO,M. using enzymes and micro-organisms, 195 pp. TAKAHASHI, (1972), Enzymatic synthesis of cephalosporins, Cambridge: Cambridge University Press. ROLINSON, G. N., BATCHELOR, F. R., BU~TERWORTH, J. Am. Chem. SOC.94,4035-4037. M. K. (1995), Biocatalysis in organic chemD., CAMERON-WOOD, J., COLE,M., EUSTACE,G. TURNER, istry (part I): past and present, Trends Biorechnol. C., HART,M. V., RICHARDS, M., CHAIN,E. B. 13,173-177. (1960), Formation of 6-aminopenicillanic acid by TURNER,M. K., FARTHING, J. R.,BREWER.S. J. (1978), enzymatic hydrolysis, Nature 187,236-237. The oxygenation of [3-methyl-3H]-desacetoxySAKAGUCHI, K., MURAO,S. (1950), A preliminary cephalosporin C [7P-(S-~-arninoadipamido)-3report on a new enzyme, “penicillin amidase”, methyl-ceph-3-em-4-carboxylicacid] to [3-hyJ. Agric. Chem. SOC.Jpn. 23,411. droxy-methyl-3H]-desacetyl-cephalosporin C by SANO,K., MITSUGI, K. (1978), Enzymatic production of L-cysteine from ~~-2-A’-thiazoline-4-carboxyl- 2-oxoglutarate-linked dioxygenases from Acremonium chrysogenum and Streptomyces clavuic acid by Pseudomonas thiazolinophihcrn: optiligerus, Biochem. J. 173,839450. mum conditions for enzyme formation and enzyUNDERWOOD, A. J. V. (193.5), The historical developmatic reaction, Agric. Biol. Chem. 42,231.5-2321. ment of distilling plant, Trans. Znsr. Chem. Eng. 13, SANTANGELO. J. D., DURRE,P. (1996), Microbial pro34-62. duction of acetone and butanol: can history be reWALTON,R. B. (1963), Search for microorganisms peated?, Chimica Oggi 14(5), 29-35. producing cephalosporin C amidase, Dev. Ind. SAVIDGE. T. A. (1984), Enzymatic conversions used Microbiol. 5.349-353. in the production of penicillins and cephalospoWEBBER,J. A., VAN HEYNINGEN, E. M.,VASILEFF, R. rins, in: Biotechnology of Industrial Antibiotics T. (1969), Chemistry of cephalosporin antibiotics. (VANDAMME, E. J., Ed.), pp. 171-224. New York: XVII. Functionalization of desacetoxycephaloMarcel Dekker. sporin. The conversion of penicillin into cephaloSCHNEIDER. W. J., ROEHR,M. (1976), Purification sporin, J. Am. Chem. SOC.91,5674-5675. and properties of penicillin acylase of Bovista WEISSENBURGER, H. w. o.,VAN DER HOEVEN, M. G . plumbae, Biochim. Biophys. Acta 452, 177-185 (1970), An efficient nonenzymatic conversion of (note that the organism is incorrectly identified in benzyl-penicillin to 6-aminopenicillanic acid, this paper). Recl. Trav. Chim. Pays-Bas 89,1081-1084. SHIBUYA, Y.. MATSUMOTO, K., FUJII,T. (1981), Isola0.R. (1972). Immobilized Enzymes. Cletion and properties of a 7P-(4-carboxy-butanami- ZABORSKI, veland, OH: Chemical Rubber Company Press. do)-cephalosporinic acid acylase producing bacZAFFARONI, A., BURTON,R. B., KEUTMANN, E. H. teria,Agric. Biol. Chem. 45,1561-1567. (1950), Adrenal corticol hormones: analysis by SIH, C. J., LEE, S. S.. TSONG,Y. Y., WANG,K. c., paper partition chromatography and occurrence CHANG,F. N. (1965), An efficient synthesis of esin the urine of normal persons, Science 111.6-8. trone and 19-norsteroids from cholesterol, J. Am. Chem. SOC.87.2765-2766.
Biotechnology Second, Completely Revised Edition H.-J. Rehm and G. Reed copyrightOWILEY-VCH Verlag GmbH, 1998
2 Biotransformations Practical Aspects
DAVID R. KELLY Cardiff, UK
1 Introduction 26 2 The Literature of Biotransformations and Reliable Experimental Procedures 27 3 Enzyme Nomenclature 29 4 Enzyme Kinetics 31 5 Getting Started 31 5.1 Whole Organisms 31 5.2 Enzymes 32 6 References 33
26
2 Biotramformatiom - PracticalAspects
1 Introduction
lutions or in organic solvents, and the work-up is easier. Some enzymes (e.g., oxidoreductases, Chapter 9, this volume) require recycling of Nowadays, the synthetic organic chemist is expensive co-factors and it is not uncommon faced with a surfeit of preparative methods for the cost of the cofactor recycling system to and synthetic targets, which at one time be vastly more than that of the enzyme cataseemed unimaginably difficult, are now re- lyzing the desired transformation. Conversely, ported routinely, e.g., palytoxin and Taxol@. most hydrolases only require pH control, Despite these salients the conversion of achi- which is easily achieved with a commercial ral compounds into enantiomerically pure chi- pH-stat. The novice to the biotransformations area ral compounds and the separation of racemic mixtures on a preparative scale, still routinely may be daunted by the apparent difficulties present a challenge. There have been many ad- posed by unfamiliar techniques. However, any vances in abiotic asymmetric transformations, organic synthesis laboratory is easily capable but few methods are generally applicable and of running both lipase-catalyzed ester hydrolyamenable to use on a large scale.The Sharpless sis reactions and yeast reductions. Yeast reducasymmetric epoxidation is a notable exception tions can easily be run in schools with minimum equipment. My eight year old daughter (KATSUKI and MARTIN,1996). Biotransformations are one tool in chiral has helped me prepare sucrose solutions which technology which must be placed alongside she added to yeast reductions! The major advantage of biotransformations asymmetric synthesis, traditional resolution techniques, and utilization of the “chiral pool”. over abiotic asymmetric catalysts is that a sucEach of these approaches has its much quoted cessful process can tap the vast resources virtues and hidden vices. A practising organic available in biotechnology. These include opchemist is well advised to adopt pragmatic timization of enzyme activity by site selective promiscuity, towards the various alternatives, mutagenesis, transfer of the gene into high tempered only by a craving for the highest productivity organisms (heterologous expression), and overexpression by incorporation of enantiomeric purity (MORI,1989). There are two major classes of biotransfor- a promoter into the gene. A good example of mation: whole cells and enzymes. Whole cells heterologous expression is the transfer of the are generally cheap, but some experience with gene for cyclohexanone mono-oxygenase microbiological techniques is required (except (which catalyzes Baeyer-Villiger reactions) for bakers’ yeast).The isolation of the product from the class 2 pathogen Acinetobacter sp. 1997). is complicated by the presence of the biomass NCIMB 9871 into a yeast (STEWART, and side reactions, including metabolism of the This occurs without change in the enantioseet al., 1996a,b) and has ensubstrate can be a problem. Moreover, large lectivity (STEWART volume reaction mixtures are frequently re- abled the enzyme to become commercially quired to transform small amounts of material. available. Most enzymes have a “scientific life cycle” On the other hand, many of the more extraordinary reactions, achieved by biotransforma- which starts with the demonstration of a partions are due to membrane-bound enzymes, ticular activity, proceeds through purification which cannot be easily isolated and hence the and isolation of the enzyme, identification of use of whole cells is mandatory, e.g., the ll-a- the gene, sequence determination, crystallizahydroxylation of progesterone (Fig. 9; Chap- tion, X-ray structure determination and heterter 1, this volume; PETERSENand MURRAY, ologous expression. At some stage in this process an enzyme manufacturer will realize that 1952). Enzymes are more selective, require simpler there is a market for the enzyme and it will beapparatus,can be run in more concentrated so- come commercially available.
2 The Literature of Biotransformations and Reliable Experimental Procedures
2 The Literature of Biotransformations and Reliable Experimental Procedures
27
There are many texts suitable for the novice and teaching purposes (FABER,1995; HANSON, 1995) and notably ROBERTS et al. (1995) provides useful practical details. Many of the prima facie cases of extraordinary selectivity achieved with enzymes can be rationalized from an appreciation of their structures (FERSCHT, 1985; PALMER, 1984) and catalytic 1987,1997; The development of on-line services has mechanisms (PAGEand WILLIAMS, revolutionized the rate at which information BUGG,1997). Recent applications of biotranscan be retrieved (BENTON, 1996). Searches that formation in industrial processes are covered used to take hours can now be done in seconds. by COLLINSet al. (1992) and early work by Many of these services have restricted access ROSE(1980) and in Chapter 1 of this volume. There are of course many reviews of bioor require a fee, but equally many are free and the web enables them to be accessed over na- transformations in the secondary literature, tional boundaries. For example, the National but as the field has developed all encompass1995) have become less Centre for Biotechnology Information (http:I/ ing reviews (AZERAD, www.ncbi.nlm.nih.gov) provides links to ge- common. Early reviews by JONESgive good nomic, peptide sequences, bibliographic and overviews of the synthetic application (JONES, many other databases. Similar information and 1986) and stereoselectivity of enzymes (JONES, links are provided by the European Bioinfor- 1993). Reviews of individual areas are of matics Institute (http://www.ebi.ac.u k / ) . More- course described in the relevant chapters of over, searches in these databases can be linked Volumes 8a and b, however, some such as oxi1992, 1997) and reduction so that query results in one database can be dation (HOLLAND, et al., 1996) are sufused to search another. The University of Min- (FANGet al., 1995;CARREA nesota has an embryonic site devoted entirely ficiently broad to be worth mentioning here. Many of the problems and fears occasioned to biotransformations (http://www.1abmed.umn. eddumbbdl). At present (January ’98), this by working with a new technique can be overcontains very limited information, but has use- come by using a precise experimental description. The biotransformations equivalent of Orful links to elsewhere. The “Biotransformations” CD-ROM (KIES- ganic Syntheses is the series Preparative Biotransformations (ROBERTS et al., 1992) which LICH et al., 1998) contains over 40000 biotransformations, compiled by the Warwick Bio- reports checked procedures in a standard fortransformations Club and KIESLICH from 1971 mat. DAVIESet al. also gives practical proceonwards, but it is very expensive except for in- dures obtained from the authors of original research papers. Organic Syntheses has pubstitutional use. Databases are fine for primary literature, lished a total of 8 biotransformation preparabut most do not abstract books. Moreover, tions (Figs. 1 and 2).These are highly represenbooks are generally able to give a more critical tative of the broad range of biotransformaappreciation of a given area. Currently, the tions which are in common use. Figs. l a and b show typically esterase-catamost comprehensive and up to date compilation of biotransformations is Enzyme Catalysis lyzed hydrolyses of acetate esters. Two of them in Organic Synthesis (DRAUZ and WALDMANN,are conversions of prochiral diesters to chiral 1995). Prior to this DAVIES et al. (1989), REHM monoesters, whereas the cleavage of the and REED(1984), KIESLICH (1976), and Ro- monoester (3) mandates the mildest conditions because of the potential for p-eliminaSAZZA (1982) are fairly comprehensive, and highlights from a range of groups are reported tion. Fig. l c shows a resolution by preferential hyin the proceedings of a CIBA Fundation Symposium (PORTERand CLARK, 1985). Enzymat- drolysis of one enantiomer of the racemic subic (but not whole organism) biotransforma- strate (7). The hydrolysis of esters of secontions are described with numerous tables of dary alcohols are appreciably more difficult examples by WONGand WHITESIDES (1994). than those of primary alcohols. Consequently,
-
2 Biotransformations - Practical Aspects Electric eel acetylcholine esterase 9-12h Deardorff etal.. 1995
A
AcO
1
8 1
Wheat germ lipase
0
pH 5 . 7 days Paquette e t d . 1995
AcO
4 s
HO
3
2 96Oh ee. 86-87% yield
Acov
"YY
298% ee. 84% yield
4 5 T , 48-7231 Schwartz et aL. 1990
rac 7
y/ SiY0Me2
HO
P O A c
" B u g F
11
Lipase Ammo
AK
Beresis et d.1997
P
9w0ee 48Y0yield
DH 7.0.60% conversion Kalaritis et al.. 1990
-
A AcO 12 >95%ee. 48% yield
14
~
--F
6
9 93% ee 49% yield
%SiPhMe2
Lipase Amano P-30 OEt
0
4702
Pig liver esterase
5
9
4 60% yield
go2
Ebele et aL,1990
*
P HO
SiPhMe2 13 >95%ee. 46% yield
" B U G OH F IS 53-68%ee 16 98Yoee 53% yield 34% yield
. B u d
OEt
Fig. 1. Hydrolase-catalyzed reactions in Organic Synthesis.
it is often worthwhile to use a less stable ester such as the chloroacetate (7). Some enzymes such as lipases (Chapter 3, this volume) function in water-immiscible solvents, e.g., pentane (Fig. Id), although the reactivity is only a few % of that in aqueous media. This enables reactions to be achieved that would not be possible in water, because of hydrolysis, e.g., transesterification. These equilibrium processes can be made practically irreversible by using a donor ester such as vinyl aceta-
te (11) in which the alcohol rearranges (self-immolation) after donation of the acyl moiety. In general, ester cleavages in which the acyl moiety is complex are less common and reliable than those in which the acyl group is small (e.g., acetate). The reason for this is that the acyl group is transferred to the enzyme, whereas the alcohol moiety is never covalently bound to the enzyme. Virtually all lipases and most esterases operate on natural substrates in which the acyl moiety is a simple straight
29
2 The Literature of Biotransformations and Reliable Experimental Procedures Bakers’yeast, HzO,sucrose, 25-30°C Seebach e t d . 1985
OEt
S
18 85% ee. 59-76940yield
17
Bakers‘ yeast, H ,O.sucrose, 30°C M o d & M o d 1989
*
20 97% ee. 47-52% yield
19
Horse liver alcohol dehydrogenase NAD. FMN, pH 9.20”C Jones & Jakovac. 1985
(Cl
*
* R
22 >97%ee. 72-77%yield
21
Fig. 2. Oxidoreductase-catalyzed reactions in Organic Synthesis.
chain. Fig. l e shows an extraordinary example in which a fluorosubstituent is efficiently distinguished by the lipase. The enantiomeric excess of the ester (15) and the acid (16) can be controlled by the degree of conversion. In the example shown in Fig. le, conversion beyond 50% improves the enantiomeric excess of the unreacted ester (15),whereas as if the reaction is run to 40% conversion the enantiomeric excess of the acid (16) is improved although the yield is decreased. Fig. 2 shows oxidoreductase reaction reported in Organic Syntheses.The easiest whole cell biotransformations are those mediated by yeast (CSUKand GLANDER,1991; SERVI,1990; and Chapter 8. this volume), and the reduction of ethyl acetoacetate (17) is the easiest of these. If the yeast reduction of a given substrate proves difficult, we use a mixture of the substrate and ethyl acetoacetate (17) to check the viability of the yeast and/or specific metabolic poisoning. Enzyme-catalyzed oxidations are more difficult to achieve than the corresponding reduction, because reduction is normally thermodynamically favored. In the example in Fig. 2c oxidation of the alcohol is
linked to reduction of NAD which in turn is oxidized by the cheap reagent flavin mononucleotide (FMN). Coupling to FMN forces the reaction in the direction of alcohol oxidation, and selectivity is further assured by hemiacetal and lactone formation which prevents the “second” hydroxyl group from being oxidized. All the of the examples in Fig. 2 follow the most common stereochemical course for carbony1 reductions which is delivery of hydride to the re-face of the carbonyl to give (S)-alcohols (Prelog’s rule, Chapters 8 and 9, this volume). In Fig. l c it is the pro-S hydroxymethylene group which is oxidized.
3 Enzyme Nomenclature In the early days of enzymology the assignment of enzyme names was left in the hands of the investigator. In most cases names were assigned on the basis of activity. Usually the ending “ase” was added to the name of the substrate, e.g., urease which catalyzes the hydroly-
30
2 Biotransformations - Practical Aspects
sis of urea to ammonia and carbon dioxide. However, as in any human endeavor; oddities, inexactitudes and obscurities arose and proliferated. Probably the two worst examples (which are still in use today) are catalase and old yellow enzyme. The International Union of Biochemistry Nomenclature Committee was set up to organize and classify the bewildering array of names for enzymes (Anonymous 1984,1992; Tab. 1). Much like systematic names for organic compounds they are generally regarded as a good idea in principle, but are much abused in practice. The classification is based on catalytic activity, hence proteins from different sources (e.g., mammalian and bacterial enzymes) with similar activity are classified under the same heading. All reactions are reversible and it is a principle of the classification that all enzymes within a given class or subclass are assigned as catalyzing a reaction in a given direction, even if this “direction” has not been demonstrated in practice for a given enzyme. For example, enzymes which catalyze the oxidation of alcohols to ketones and the reduction of ketones to alcohols are classified as alcohol dehydrogenases, i.e., there are no aldehyde reductases in the systematic naming system. This often leads to confusion, because for most alcohol-ketone redox couples linked to NAD(P)H the thermodynamic preferred direction is reduction of the ketone. However, to mitigate this the recommended names for enzymes usually relate
to the physiologically preferred direction of the reaction rather than the systematic direction. In the systematic system each enzyme is denoted by a four number code, with each successive number in the code representing a further sub-division of the number preceding it in the code. For example, EC 1.1.1.1is alcohol dehydrogenase. The first number is the class number, which here denotes an oxidoreductase, the second that it acts on alcohols, the third that it is NAD- or NADP-linked, and the final number is a unique designator for each enzyme in the subsubclass. In this particular case it indicates that the enzyme is NADlinked. Other enzymes within EC 1.1.1 are EC 1.1.1.2, which is similar to the enzyme described above, except that it is NADP-linked, EC 1.1.1.3 which is homoserine dehydrogenase and EC 1.1.1.4 which is (R,R)-butanediol dehydrogenase. Sometimes not all aspects of an the enzyme’s chemistry is known and in some of the smaller groups these are assigned arbitrarily as 99. For example, five enzymes which are involved in the biosynthesis of prostaglandins are assigned to EC 5.3.99.1-5, where the initial “5” indicates an isomerase, the “3” an intramolecular oxidoreductase, and the “99” is for this miscellaneous subsubclass. The nomenclature committee has had no compunction about deleting subclasses which are no longer appropriate, but thankfully these are never reused.
Tab. 1. The 5 Main Classes of Enzyme as Classified by the Nomenclature Committee (adapted from DAVIES et al., 1989)
Class
Number Identified, Available
1) Oxidoreductases 2) Transferases
650.90 720,90
3 ) Hydrolases 4) Lyases
636,125 255,35
5 ) Isomerases 6) Ligases
120,6 80,5
Totals
2461,351
Reaction
redox processes transfer of a group from one molecule to another, e.g., acyl or phosphate hydrolysis reactions,e.g., esterases, lipases, and amidases cleavage of C-C, C - 0 and C-N bonds other than by hydrolysis or redox reactions;typically these reactions involve elimination to, or additions to double bonds isomerizations such as epimerization and racemization intermolecular bond formation coupled with the cleavage of ATP or other nucleotide triphosphates
5 Getting Started
4 Enzyme Kinetics Enzyme-catalyzed reactions follow Michaelis-Menten kinetics or one of the related rate models. Briefly (and with some gross simplifications), the rate of reaction is a function of substrate concentration, until it reaches a rate determined by the maximum rate of processing of the substrate by the active site. To avoid problems with product inhibition rates are determined from the initial rate of the reaction, i.e., at 1-5% conversion. The key parameters are V, which is the theoretical maximum rate of the reaction and K , which is the concentration of substrate at half maximum rate (i.e., Vmax/2). The exact significance of K , depends on the nature of the kinetics of the reaction, however, it can be broadly interpreted as the affinity of the enzyme for the substrate. For good substrates K , will be a low value (typically micromolar) and of course V,,, will be large. If the concentration of the enzyme [El is known, it is possible to calculate k,,, = V,,,/(E] which is the number of substrate molecules processed by the enzyme per unit time k,,, is also known as the turnover number.Typically these are in the range 1-500/s. Great care should be taken in determining and interpreting kinetic parameters, particularly with unnatural substrates. In order to calculate accurate values of K , and V, from initial rate data it is important to be able to use concentrations which are a significant proportion of K,. This is frequently impossible with unnatural substrates which have high values of K,, and in consequence the calculations are unstable, i.e., small changes in the measured values cause large changes in the calculated values of the kinetic parameters. The reader is referred to other texts for more detail (FERSCHT, 1985;PALMER, 1984) and for a rigorous exposition (CORNISH-BOWDEN, 1975).
5 Getting Started 5.1 Whole Organisms Yeast may be obtained from any grocery store. We have tried many suppliers over the
31
years and generally find that canned “active dried yeast” is best in terms of activity and convenience. This takes the form of small spheres which are easily weighed out. Fresh moist yeast has to be purchased on the day of use, contains a lot of water so that it is difficult to estimate how much yeast is present and at least in our hands is less active. As a word of caution may drug stores sell yeast extracts or dried yeast which is intended as a dietary supplement. This material is dead! Most microbiologists are happy to provide samples of microorganisms (as slants on agar) and many are freely available from culture collections for a small fee (Tab. 2). However, care must be taken not use organisms which are pathogenic unless the requisite facilities and personnel are available. The techniques for growing cultures are fairly straightforward, but the average chemist is strongly advised to recruit the help of a microbiologist in the early stages. A good outline of the techniques involved is available (ROBERTS et al., 1995). Research papers concentrate on a given activity of an microorganism, but it is less widely recognized all “wild” microbes have a complete set of metabolic enzymes. So, e.g., yeast are capable not only of ketone reduction, but also ester hydrolysis, cyclization of squalene epoxide analogs to lanosterol analogs, acyloin condensation and alkylation of enolates (Chapter 8, this volume). New activities continue to be discovered. Bakers’ yeast oxidizes methyl p-tolyl sulfide to the corresponding sulfoxide in 60% yield and 92% ee (BEECHERet al., 1995; TANGet al., 1995). If a given reaction is chemically reasonable and thermodynamically feasible there is good chance that an organism can be found to catalyze it and the products will have an enantiomeric excess. In the natural world racemic mixtures are exceptional and enantioselectivity is unexceptional. If the resources are available screening of microorganism collections by growth on media containing the substrate frequently yields candidate species. A combinatorial approach is to use water or soil from a biologically rich or stressed source such as contaminated land as an innoculum for a media containing the substrate of interest (elective culture). The source may be even closer to hand. A black fungus, NV-2, which enantiose-
32 Tab. 2.
2 Biotransformations - Practical Aspects Addresses of Culture Collections
Abbreviation
Address
American Type Culture Collection, 12301 Parklawn Drive, Rockville, Maryland 20852, USA Centraalbureau voor Schimmelcultures, Julianalaan 67,2628 BC, Delft, The Netherlands CBS International (Commonwealth) Mycological Institute, Ferry Lane, Kew, Surrey,TW9 3 M , IMI (formerly CMI) England, UK Deutsche Sammlung von Mikroorganismen und Zellkulturen, Mascheroder Weg lb, DSM D-38124 Braunschweig, Germany Institute of Fermentation, Osaka 17-85Jusohomachi 2-chome, Yodogawaku, Osaka 532, IF0 Japan National Collections of Industrial and Marine Bacteria, 23 St Machar Drive, Aberdeen NCIMB (formerly NCIB) AB2 lRY, Scotland, UK National Collection of Yeast Cultures, Food Research Institute, Colney Lane, Norwich, NCYC Norfolk NR4 7UA, England, UK Northern Regional Research Laboratory, Agricultural Research Service, USDA, NRRL 1815 N. University Street, Peoria. Illinois 61604, USA All-Union Collection of Microorganisms, Department of Culture Collection, VKM Institute of Biochemistry and Physiology of Microorganisms. Academy of Sciences, Pushchino, Moscow, Region 142292, Russia ATCC
lectively oxidizes sulfides to sulfoxides was found on a damp wall behind a wardrobe in student accommodation (KELLYet al., 1996).
5.2 Enzymes Enzyme preparations range from crude extracts through to highly purified crystalline materials. All enzymes degrade with time, particularly if they contain extraneous material. However, when stored sealed in a refrigerator the typical half life is no less than that of butyl lithium. As with other reagents the reactivity is profoundly influenced by reaction conditions. pH control is very important for achieving maximal activity. If the pH is not expected to change greatly during the course of the reaction then buffers are adequate, otherwise for reactions such as ester hydrolysis where there are large changes in pH then a pH-stat must be used. All buffers of the same pH are not equal! Some enzymes, particularly those that bind nucleotide phosphates are inhibited by phosphate buffers which compete at the nucleotide binding site. Most enzyme-catalyzed reactions benefit by being run above room temperature. The often quoted 37°C is really only appropriate for
mammalian enzymes, but 30-35 “C greatly increases the rate of many reactions. This is probably due to the increased solubility of the substrate as much as the thermodynamically controlled increase in enzyme conversion. Enzymes from thermophilic bacteria nearly always operate better at elevated temperatures (40-50°C)and, moreover, the enantioselectivity for the reduction of 2-pentanone by Thermoanaerobium brockii alcohol dehydrogenase changes from formation of (S)-2-pentanol at 15°C to (R)-2-pentanol at 37°C (YANGet al., 1997). A major problem with the biotransformation of unnatural substrates is that the substrate is frequently only slightly soluble in water. Most enzymes will tolerate up to 15’A ethanol or isopropanol as a co-solvent without an appreciable effect on reactivity. For oxidoreductases these will also act as the ultimate source of reductive capacity for NAD(P)H recycling systems. On the other hand enzymes (or even yeast) will function in water immiscible solvents such as hexane, dichloromethane and chloroform which is valuable for “reverse hydrolysis reactions”. It is believed that under these circumstances the enzymes retain a layer of water molecules on the surface and interstices which are essential main-
6 References
taining tertiary. Nevertheless the activity of enzymes is generally much less in organic solvents than in water and the specificity may be changed (WESCOTT and KLIBANOV, 1994). Much of this is due to the problems of dispersing the enzyme homogeneously in a media in which it is insoluble (KLIBANOV, 1997). Other techniques for modifying enzyme reactivity have been reviewed (DAVIES et al., 1989; RoBERTS et al., 1995;WONG and WHITESIDES, 1994). One of the many virtues of using enzymecatalyzed biotransformations is that the reactions proceed under more or less identical conditions. This means that multi-enzyme systems can be constructed in which a series of intermediates are processed by a constellation of enzymes. Simple examples of this technique are used routinely in cofactor recycling systems for NAD(P)H (Chapter 9, this volume) and for nucleotides (Volume 8b). More recently, the availability of large quantities of enzyme by heterologous expression in high productivity organisms has enabled complex natural products such as Vitamin B,, to be created in one pot from simple precursors (ROESSNER and SCOTT,1996). This offers possibilities which are currently unimaginable by any other approach.
6 References Anonymous (1984, 1992), Enzyme Nomenclature: Recommendations of the Nomenclature Committee of the International Union of Biochemistry on the Nomenclature and Classification of EnzymeCatalyzed Reactions. Orlando, FL: Academic Press. AZERAD, R. (1995),Application of biocatalysts in organic synthesis, Bull. Chem. Soc. France 132, 17-51. BEECHER, J., BRACKENRIDGE, I., ROBERTS.S. M., TANG,J., WILLEITS,A. J. (1995), Oxidation of methyl p-tolyl sulfide with bakers’ yeast: preparation of a synthon of the mevinic acid type hypocholestemic agents,J. Chem. SOC.,Perkin Trans. I , 1641-1643. BENTON, D. (1996), Bioinformatics - principles and potential of a new multidisciplinary tool, TIBTECH 14,261-272. BERESIS, R. T., SOLOMON, J. S., YANG,M. G., JAIN,K. F., PANEK,J. S. (1997), Synthesis of chiral ( E ) -
33
crotylsilanes: [3R- and 2S-]-(4E)-methyl3-(dimethylphenylsilyl)-4-hexenoate, Org. Synth. 75, 78-88. BUGG,T. A. (1997), An Introduction to Enzyme and Coenzyme Chemistry, p. 247. Oxford: Blackwell Science. CARREA, G., OTTOLINA, G., PASTA, €?, RIVA,S. (1996), Synthetic applications of NAD(P)(H)-dependent enzymes,Ann. N. Y Acad. Sci. 799,642-649. G. N., CROSBY, J. (Eds.) COLLINS, A. N., SHELDRAKE, (1992). Chirality in Industry. The Commercial Manufacture and Applications of Optically Active Compounds. Chichester: John Wiley & Sons. A. (1975), Principles of Enzyme CORNISH-BOWDEN, Kinetics, London: Butterworth. CSUK, C., GLANDER, B. J. (1991).Bakers’ yeast mediated biotransformations in organic chemistry, Chem. Rev. 91,49-97. DAVIES, H. G., GREEN, R. H., KELLY, D. R., ROBERTS, S. M. (1989), Biotransformations in Preparative Organic Chemistry. London: Academic Press. DEARDOFF, D. R., WINDHAM, C. Q., CRANEY, C. L. (1995), Enantioselective hydrolysis of cis-3,5-diacetoxycyclopentene: (1R,4S)-(+ )-4-hydroxy-2cyclopentenyl acetate, Org. Synth. 73,29-35. DRAUZ,K., WALDMANN, H. (1995), Enzyme Catalysis in Organic Synthesis,A Comprehensive Handbook, Weinheim: VCH. EBERLE,M., MISSBACH, M., SEEBACH,D. (1990) Enantioselective saponification with pig liver esterase PLE: (lS,2S,3R)-3-hydroxy-2-nitrocyclohexyl acetate, Org. Synth. 69, 10-18 (also: Org. Synth. Collective 8,332-338). FABER,K. (19954, Biotransformations in Organic Chemistry, 2nd Edn., Berlin, Heidelberg, New York: Springer-Verlag. FANG,J. M., LIN,C. H., BRADSHAW, C. W., WONG,C. H. (1995), Enzymes in organic-synthesis - oxidoreductions, J. Chem. SOC., Perkin Trans. I , 967-978. FERSCHT, A. (1985), Enzyme Structure and Mechanism, New York: Freeman. HANSON, J. R. (1995), An Introduction to Biotransformations in Organic Chemistry. Oxford: Freeman. HOLLAND, H. L. (1992),Synthesis with Oxidative Enzymes, pp. 255-31 1 . New York: VCH. H. L. (1997), Investigation of the carbonHOLLAND, and sulfur-oxidizing capabilities of microorganisms by active-site modeling, Adv. Appl. Microbiol. 44,125-165. JONES,J. B. (1986), Enzymes in organic synthesis, Tetrahedron 42,3351-3403. JONES, J. B. (1993), Probing the specificity of synthetically useful enzymes, Aldrichimica Acta 26, 105-1 12. I. J. (1985) Preparation of chiJONES, J. B., JAKOVAC, ral, nonracemic y-lactones by enzyme catalyzed
34
2 Biotransformations
- Practical Aspects
+
oxidation of meso-diols: ( )-(lR,6S)d-oxabicyclo[4.3.0]nonan-7-one, Org. Synth. 63, 10-17 (also: Org. Synth. Collective 7,406-410). KALARITIS, P., REGENYE, R. W. (1990) Enantiomerically pure ethyl ( R ) - and (S)-2-fluorohexanoate by enzyme-catalyzed kinetic resolution, Org. Synth. 69, 10-18 (also: Org. Synth. Collective 8, 258-262). KATSUKI, T., MARTIN,V. S. (1996), Catalytic asymmetric epoxidation of allylic alcohols, Org. React. 48,l-130. KELLY,D. R., KNOWLES, C. J., MAHDI, J. G.,TAYLOR, I. N., WRIGHT,M. A. (1996), The enantioselective oxidation of sulfides to sulfoxides with Acinetobacter sp., NCIMB-9871, Pseudomonas sp., NCIMB-9872, Xanthobacter autotrophicus DSM431 (NCIMB-10811) and the black yeast NV-2, Tetrahedr0n:Asymmetry 7,365-368. KIESLICH,K. (1976), Microbial Transformations of Non-Steroid Cyclic Compounds. Stuttgart: Thieme. KIESLICH, K., CROUT,D. H. G., DALTON. H., SCHNEIDER,M. (1998), Biotransformations, CD-ROM. London: Chapman & Hall (1996 and updated annually). KLIBANOV, A. M. (1997), Why are enzymes less active in organic solvents? Trends Biotechnol. 15, 97-101. MORI,K. (1989), Synthesis of optically active pheromones, Tetrahedron 45,3233-3298. MORI,K., MORI,H. (1989), Yeast reduction of 2,2dimethylcyclohexane-1,3-dione: (S)-( )-3-hydroxy-2,2-dimethylcyclohexanone,Org. Synth. 68, 56-63 (also: Org. Synth. Collective 8,312-315). PAGE, M.. WILLIAMS,A. (Eds.) (1987), Enzyme Mechanisms. London: RSC. PAGE,M., WILLIAMS,A. (1997), Organic and Bio-Organic Mechanisms, p. 298. Harlow: Longman. PALMER, T. (1984), Understanding Enzymes. Chichester: Ellis Horwood. PAQUETTE, L. A., EARLE,M. J., SMITH,G. F. (1995). (4R)-( + )-tert-butyldimethylsilyloxy-2-cyclopenten-1-one, Org. Synth. 73,36-43. PETERSON, D. H., MURRAY, H. C. (1952), Microbial oxygenation of steroids at carbon 11, J. Am. Chem. SOC.74,1871-1872. PORTER. R., CLARK,S. (1985). Enzymes in organic synthesis, in: CIBA Foundation Symposium I l l . London: Pitman. REHM.H.-J.. REED,G. (Eds.) (1984), Biotechnology, 1st Edn., Vol. 6a (KIESLICH,K., Ed.). Weinheim: Verlag Chemie. ROBERTS, S. M., WIGGINS, K., CASY,G.. PHYTHIAN, S. J., TODD, C. (Eds.) (1992), Preparative Biotransformations: Whole Cells and Isolated Enzymes in Organic Synthesis. Chichester: John Wiley & Sons.
+
ROBERTS, S. M.,TURNER, N. J., WILLETTS,A. J.,TuRNER,M. K. (1995). Introduction to biocatalysis: using enzymes and micro-organisms. Cambridge: Cambridge University Press. ROESSNER, C. A., Scorr,A. I. (1996), Genetically engineered synthesis of natural products: alkaloids to corrins, Ann. Rev. Microbiol. 50,467-490. ROSAZZA, J. P. (1982), Microbial Transformations of Bioactive Compounds, Vol. 1. Boca Raton, FL: CRC Press. ROSE,A. H. (1980), Microbial enzymes and bioconversions, Vol. 5 in the series Economic Microbiology. London: Academic Press. SEEBACH, D., SUTTER,R. H., WEBER,R. H., ZUGER, M. F. (1985), Yeast reduction of ethyl acetoacetate. Ethyl (S)-3-hydroxybutanoate, Org. Synth. 63,l-7 (also: Org. Synth. Collective 7,215-220). SCHWARTZ, A., MADAN,P., WHITESELL, LAWRENCE, R. M. (1990), Lipase-catalyzed kinetic resolution of alcohols via chloroacetate esters: ( -)-(lR,2S)trans-2-phenylcyclohexanol and ( )-(1S,2R)trans-2-phenylcyclohexanol,Org. Synth. 90, 1-9 (also: Org. Synth. Collective 8,516-521). SERVI,S. (1990), Bakers’ yeast as a reagent in organic synthesis, Synthesis, 1-25. STEWART, J. D. (1997). Chemist’s perspective on the use of genetically engineered microbes as reagents for organic synthesis, Biotechnol. Gen. Eng. Rev. 14,67-143. STEWART, J. D., REED,K. W., ZHU,J., CHEN,G., KAYSER, M. M. (1996a), A designer yeast that catalyzes the kinetic resolutions of 2-alkyl-substituted cyclohexanones by enantioselective Baeyer-Villiger oxidations, J. Org. Chem. 61,7652-7653. STEWART, J. D., REED,K. W., KAYSER, M. M. (1996b), Designer yeast - a new reagent for enantioselective Baeyer-Villiger oxidations, J. Chem. SOC., Perkin Trans. I , 755-757. TANG, J., BRACKENRIDGE, I., ROBERTS,S. M., BEECHER,J., WILLETTS, A. J. (1995), Bakers’ yeast oxidation of methyl para-tolylsulfide: synthesis of a chiral intermediate in the preparation of the mevinic acid-type hypocholestemic agents, Tetrahedron 51,13217-13238. WESCOTT, C. R.. KLIBANOV,A. M. (1994),The solvent dependance of enzyme specificity, Biochim. Biophys. Acts l206,l-9. WONG, C.-H., WHITESIDES, G. M. (1994), Enzymes in Synthetic Organic Chemistry, Tetrahedron Organic Chemistry Series, Vol. 12, pp. 169-170. Oxford: Pergamon. YANG, H., JONSSON, A., WEHTJE, E., ADLERCREUTZ, P., MATTIASSON, B. (1997),The enantiomeric purity of alcohols formed by enzymatic reduction of ketones can be improved by optimization of the temperature and by using a high co-substrate concentration, Biochem. Biophys. Acta 1336,51-58.
+
Hydrolases
Biotechnology Second, Completely Revised Edition H.-J. Rehm and G. Reed copyrightOWILEY-VCH Verlag GmbH, 1998
3 Biotransforrnations with Lipases
ROMASJ. KAZLAUSKAS Montreal, Quebec, Canada
UWE T. BORNSCHEUER Stuttgart, Germany
1 Availability and Structure of Lipases 40 1.1 Introduction 40 1.1.1 Lipases in Cheese-Making and Laundry Detergents 40 1.1.2 Commercial Lipases 41 1.1.3 Classification of Lipases by Protein Sequence 43 1.1.4 General Features of PPL, CRL, RML, CAL-B, and PCL 43 1.1.4.1 PPL 44 1.1.4.2 CRL 44 1.1.4.3 RML 44 1.1.4.4 CAL-B 44 1.1.4.5 PCL 4 4 1.2 Structure of Lipases 45 1.2.1 Lipases are cr/p Hydrolases 45 1.2.2 Lid or Flap in Interfacial Activation 45 1.2.3 Substrate Binding Site in Lipases 47 2 Water, Organic Solvents, and Other Reaction Media 47 2.1 Hydrolysis in Aqueous Solutions and Two Phase Mixtures of Water and Organic Solvent 47 2.2 Lipases in Reverse Micelles 48 2.3 “Dry” Organic Solvents 50 2.3.1 Increasing the Catalytic Activity in Organic Solvents 51 2.3.1.1 Adsorption and Entrapment 51 2.3.1.2 Covalent Immobilization 52 2.3.1.3 Cross-Linked Enzyme Crystals - CLECs 52 2.3.1.4 Covalently Modified Lipases Soluble in Organic Solvents 53 2.3.1.5 Lipid- or Surfactant-Coated Lipases 53 2.3.1.6 Choosing the Best Organic Solvent for High Activity 54
38
3 Biotransforrnations with Lipases
2.3.2 Increasing the Enantioselectivity in Organic Solvents 54 2.3.3 Acyl Donor for Acylation Reactions 56 2.3.4 Water Content and Water Activity 58 2.4 Supercritical Fluids 60 3 Enantioselective Reactions 63 3.1 Kinetic Resolutions 63 3.1.1 Quantitative Analysis 63 3.1.2 Recycling and Sequential Kinetic Resolutions 64 3.2 Asymmetric Syntheses 66 3.3 Survey of Alcohols 68 3.3.1 Secondary Alcohols 68 3.3.1.1 Overview and Models 68 3.3.1.2 Candida antarctica Lipase B 71 3.3.1.3 Candida rugosa Lipase 72 3.3.1.4 Porcine Pancreatic Lipase 76 3.3.1.5 Pseudomonas Lipases 76 3.3.1.6 Rhizomucor Lipases 77 3.3.1.7 Other Lipases 77 3.3.1.8 Choosing the Best Route 81 3.3.1.8.1 Inositols 81 3.3.1.8.2 p-Blockers 84 3.3.2 Primary Alcohols of the Type RR’CHCH,OH 87 3.3.2.1 Pseudomonas Lipases 87 3.3.2.2 Porcine Pancreatic Lipase 91 3.3.2.3 Other Lipases 91 3.3.2.4 Enantioselectivity of Lipases Toward Triglycerides 91 3.3.3 Other Alcohols,Amines, and Alcohol Analogs 94 3.3.3.1 Tertiary Alcohols and Other Quaternary Stereocenters 94 3.3.3.2 Alcohols with Axial Chirality or Remote Stereocenters 94 3.3.3.3 Alcohols with Non-Carbon Stereocenters 98 3.3.3.4 Analogs of Alcohols: Amines, Thiols, and Hydroperoxides 98 3.3.3.4.1 Amines 98 3.3.3.4.2 Thiols 102 3.3.3.4.3 Peroxides 103 3.4 Survey of Carboxylic Acids 103 3.4.1 General Considerations 103 3.4.2 Carboxylic Acids with a Stereocenter at the a-Position (RR’CHCOOH) 103 3.4.2.1 Candida antarctica Lipase B 103 3.4.2.2 Candida rugosa Lipase 104 3.4.2.3 Pseudomonas Lipases 107 3.4.2.4 Other Lipases 107 3.4.3 Carboxylic Acids with a Stereocenter at the p-Position 108 3.4.4 Other Carboxylic Acids 109 3.4.4.1 Quaternary Stereocenters 109 3.4.4.2 Sulfur Stereocenters 109 3.4.4.3 Remote Stereocenters 109 3.5 Anhydrides 111 3.6 Lactones 112 3.7 Dynamic Kinetic Resolutions 113 4 Chemo- and Regioselective Reactions 118 4.1 Protection and Deprotection Reactions in Organic Synthesis 118 4.1.1 Hydroxyl Groups 118
I Availability and Structure of Lipases
4.1.1.1 Primary Hydroxyl Groups in Sugars 118 4.1.1.1.1 Hydrolysis of Esters of Primary Hydroxyl Groups 118 4.1.1.1.2 Acylation of Primary Alcohols in Unmodified Sugars 119 4.1.1.1.3 Acylation of Primary Alcohols in Alkyl Glycosides and Other Modified Sugars 121 4.1.1.2 Secondary Hydroxyl Groups 124 4.1.1.2.1 Hydrolysis of Acylated Secondary Hydroxyl Groups 124 4.1.1.2.2 Acylation of Secondary Hydroxyl Groups 126 4.1.1.3 Hydroxyl Groups in Non-Sugars 129 4.1.1.3.1 Phenolic Hydroxyls 129 4.1.1.3.2 Aliphatic Hydroxyls 130 4.1.2 Amino Groups 131 4.1.3 Carboxyl Groups 132 4.2 Lipid Modifications 134 4.2.1 1,3-Regioselective Reactions of Glycerides 134 4.2.1.1 Modified Triglycerides 135 4.2.1.1.1 Cocoa Butter Substitutes 135 4.2.1.1.2 Synthesis of MLMs 136 4.2.1.1.3 Triacylglycerides Containing Polyunsaturated Fatty Acids (PUFAs) 137 4.2.1.1.4 Other Triglycerides 137 4.2.1.2 Diacylglycerides 137 4.2.1.2.1 Acyl Migration in Mono- and Diacylglycerides 139 4.2.1.3 Monoacylglycerides (MAGS) 139 4.2.1.3.1 Hydrolysis or Alcoholysis of Triglycerides to 2-MAGS 141 4.2.1.3.2 Glycerolysis of Triglycerides to l(3)-MAGS 141 4.2.1.3.3 Esterification of Glycerol with Fatty Acids or Fatty Acid Esters Yielding 1(3)-MAGs 142 4.2.2 Fatty Acid Selectivity 142 4.2.2.1 Saturated Fatty Acids 142 4.2.2.2 Unsaturated Fatty Acids 143 4.3 Oligomerization and Polymerizations 144 4.4 Other Lipase-Catalyzed Reactions 145 5 Commercial Applications and Future Directions 146 5.1 Commercial Applications 146 5.1.1 Food Ingredients 146 5.1.2 Enantiomerically Pure Chemical Intermediates 146 5.1.3 Enantiomerically Pure Pharmaceutical Intermediates 146 5.2 Future Directions 148 5.2.1 Reaction Engineering 148 5.2.2 Modeling and Mutating the Selectivity of Lipases 149 5.2.3 Directed Evolution of Lipases 149 6 References 150
39
40
3 Biotransformations with Lipases
1 Availability and Structure of Lipases 1.1 Introduction Both lipases and esterases catalyze the hydrolysis of esters, but only lipases catalyze hydrolysis of water-insoluble esters such as triglycerides. For example, lipases catalyze the hydrolysis of triolein to diolein (Eq. 1). In addition, lipases also catalyze the hydrolysis of a broad range of natural and unnatural esters, while retaining high enantio- or regioselectivity. This combination of broad substrate range and high selectivity makes lipases an ideal catalyst for organic synthesis. Chemists use lipase-catalyzed biotransformations to prepare enantiomerically pure pharmaceuticals and synthetic intermediates (Sect. 3), to protect and deprotect synthetic intermediates (Sect. 4.1), to modify natural lipids (Sect. 4.2) as well as for more specialized uses. A survey of these reactions is the main focus of this review. Besides high selectivity and broad substrate range, another major advantage of lipases for synthetic reactions is that they act efficiently on water-insoluble substrates. Lipases need this ability because the natural substrates of lipases - triglycerides - are insoluble in water. Lipases bind to the water-organic interface and catalyze hydrolysis at this interface. This binding not only places the lipase close to the substrate, but also increases the catalytic power of the lipase, a phenomenon called interfacial activation. Most lipases are poor catalysts in the absence of an interface such as an organic droplet or a micelle. A conformational change in the lipase probably causes the interfacial activation (see Sect. 1.2.2). In contrast,
“4 0
(
0
triolein
+H20
a triglyceride
PH 7
1,2- or 2.3-diolein a diglyceride
R = (CH 2)7CH=CH(CH2)7CH 3
oleic acid a fatty acid
efficient reactions with proteases often require chemical modification of the substrate to increase water solubility. A number of books on lipases, or with large sections on lipases and extensive reviews, are available (ALBERGHINA et al., 1991;BoLAND et al., 1991; BORGSTROM and BROCKMAN, 1984; COLLINS et al., 1992; DRAUZand WALDMANN, 1995;FABER,1997; ROBERTS, 1992-1996; POPPE and NOVAK,1992; SCHMIDand VERGER,in press; GANDHI,1997; SHELDON, 1993; JAEGER :t al., 1994; KAZLAUSKAS, 1994; WOOLLEY and ‘ETERSEN, 1994; WONGand WHITESIDES, 1994; THEIL,1997). More specialized reviews on lipases will be cited in the appropriate sections.
.
1.1.1 Lipases in Cheese-Making and Laundry Detergents Cheese manufacturers use lipase-catalyzed hydrolysis of milk fat to enhance flavors, accelerate cheese ripening, and to manufacture cheese-like products (for reviews containing sections on cheese-making and detergents see VULFSON,1994; HAAS and JOERGER,1995; CHEETHAM, 1993;BERRYand PATERSON, 1990). Traditional cheese-making adds extracts containing lipases to the raw cheeses to impart characteristic flavors. For example, extracts of the pregastric gland of a calf imparts a buttery and slightly peppery flavor, while a kid extract imparts a sharp flavor and a lamb extract imparts a strong “dirty sock” flavor. In addition, microbes responsible for cheese ripening secrete lipases. For example, lipase from Penicillium roquefortii liberates short and medium chain fatty acids which add flavor both directly and by serving as precursors for Glactones and methylketones. Modern cheese-makers can substitute commercial lipases (e.g., lipases
I Availability and Structure of Lipases
from Aspergillus niger or Rhizomucor miehei) for the pregastric gland extracts and for the microbes. Also, addition of lipases to cow’s milk mimics the flavor of goat’s or sheep’s milk. Addition of lipases to cheese followed by incubation at high temperature yields a concentrated cheese flavor that can be used to flavor sauces and other prepared foods. Some detergents include microbial lipases (e.g., lipase from Humicola lanuginosa) to aid removal of fat stains, but the advantage accumulates only after multiple washing.The wash cycle is too short for significant hydrolysis, but the lipase remains on the fat in the subsequent drying where it hydrolyzes the fats. The next wash cycle removes these fats. Lipases may also prevent redeposition of fats on textiles. Using lipases for biotransformations is a smaller market (
41
searchers continue to refer to this lipase as r! fluorescens lipase and Fluka sells SAM-I1 under the name of lipase from Pseudomonas fluorescens. Unfortunately, ATCC 21 808 has been again renamed to Burkholderia cepacia. For this review, we will continue to use Pseudomonas cepacia. A recent reclassification of the Rhizopus fungi renamed R. niveus, R. delemar, and R. javanicus all as Rhizopus oryzae (reviewed by HAAS and JOERGER,1995). Consistent with this reclassification, the lipases isolated from R. delemar, R. javanicus, and R. niveus have identical amino acid sequences and lipase from R. oryzae differs only by two conservative substitutions (His134 is Asn and Ile234 is Leu in Rhizopus oryzae lipase, ROL). In spite of these similarities, Amano sells three different lipases from this group, and they show slightly different selectivities, perhaps due to cleaving the prolipase at different positions (UYTENBROCK et al., 1993).The prolipase contains extra amino acid residues to guide folding and secretion of the lipase. After folding, proteases cleave the extra amino acid residues to give the mature lipase. This cleavage may not 1.1.2 Commercial Lipases always occur at the same amino acid residue. Note that even when the microorganism Lipases occur in plants, mammals, and micro- classification is settled, the same species may organisms where the biological role of lipases produce different lipases. Amano sells lipase is probably digestive. Most biotransformations A H from Pseudomonas cepacia which differs use commercial lipases, about 70 of which are from lipase P in the amino acid sequence in 16 available.Tab.1 lists the most popular of these. of 320 residues.These two lipases had opposite Pancreatic cholesterol esterase is included selectivity for a dihydropyridine substrate with the lipases because its sequence and bio- (HIROSEet al., 1995). chemical properties are identical to bile salt For the purposes of this review, we will simstimulated lipase (Hur and KISSEL,1990;NILS- plify the lipase names. The properties of all commercial preparations of lipase from CandiSON et al., 1990). Lipases are usually named according to the da rugosa seem similar, for this reason we will microorganism that produces the lipase. The refer to all of them as CRL. Amano P, purified classification, and thus name, of a microorgan- forms of this lipase, and SAM-I1 (Fluka) all ism can change as researchers learn more come from microorganism ATCC 21808. We about it. Likewise, the name of the lipase will refer to all of these as PCL, even if the sometimes changes which can be frustrating to authors did not. The amino acid sequence and organic chemists accustomed to molecules biochemical properties of lipase from Pseudowhose name rarely changes. For example, monas glumae and lipase from ChromobacteriAmano researchers first classified the micro- um viscosum are identical (TAIPAet al., 1995; organism that produces “Amano P” (ATCC LANGet al., 1996),and we will refer to both of 21 808) as Pseudomonas fluorescens,but have these as CVL. We will refer to all the Rhizopus since reclassified it as f! cepacia. For this rea- lipases as ROL. For the other lipases, we will son pre-1990 papers on this lipase refer to it as use the abbreviation shown in Tab. 1 or the full f! fluorescens lipase. Confusingly, some re- name.
Rhizopics oryzae
Candida antarctica A Candida antarctica B
Aspergillus niger Candida lipolytica Penicilliurn roquefortii Bacterial lipases‘ Pseudomonas cepacia
Pseudomonas cepacia Pseudomonas Jluorescens Pseudomonas fragi Chromobacterium viscosum Pseudomonas glumae Pseudomonas sp. Bacillus thermocatenulatcis Alcaligenes sp.
ROL
CAL-A CAL-B
ANL CLL ProqL
PCL-AH PFL PfragiL CVL‘
Burkholderia cepaciad
a
’
The amino acid sequences of lipases R. delemar, R. javanicccs,and R. niveus are identical.The sequence of lipase from R. oryzae differs by only two conservative substitutions. SP 525 is a powder containing 40 wt% protein, while Novozym 435 is the same enzyme immobilized on macroporous polypropylene (1 w/w% protein). For a while Novo Nordisk supplied a lipase SP 382 from Candida sp.This lipase was a mixture of lipases A and B from Candida antarctica. ‘Boehringer Mannheim sells two lipases from Pseudomonas species: Chirazymea L-4 and Chirazymem L-6. It is not clear from the product literature which of the Pseudomonas lipases they correspond to. Lipase from microorganism ATCC 21 808. Early reports classified this microorganism as Pseudomonasfluorescens, later as P cepacia, most recently as Burkholderia cepacia. Neither the microorganism nor the lipase has changed by the change in name. ‘The amino acid sequence and biochemical properties of lipase from Pseudomonas glumae and lipase from Chromobacterium viscosum are identical (TAIPAet al., 1995; LANGet al., 1996).
BTL2
PCL
Altus Biologics (ChiroCLEC-PC), Amano (P, P-30, PS, LPL-80, LPL-2OOS), Boehringer Mannheim (Chirazymea L-l), Fluka (SAM-11), Sigma Amano (lipase AH) Amano (lipase AK), Amano (lipase YS), Biocatalysts Ltd. lipase B, Wako Pure Chemical (Osaka) Sigma, Genzyme, Asahi Chemical, Biocatalysts Ltd. Amano (K-10) Boehringer Mannheim Meito Sangyo (lipase QL)
Boehringer Mannheim (Chirazymea L-8), Novo Nordisk (SP 524, Lipolase@) Amano (lipase G) Amano (lipase M) Boehringer Mannheim (Chirazymea L-9), Amano (MAP), Novo Nordisk (SP 523, Lipozyme@),Fluka Amano (lipase F), Amano (lipase D), Amano (lipase N), Fluka, Sigma, Seikagaku Kogyo Co. (Japan) Boehringer Mannheim (Chirazymem L-5). Novo Nordisk (SP 526)’ Boehringer Mannheim (Chirazymea L-2), Novo Nordisk (SP 525 or Novozym 435)’ Sigma Amano (lipase A, AP), Rohm, Novo Nordisk (Palatasem) Amano (lipase L) Amano (lipase R)
Thermomyces lanuginosa Penicillium cyclopium Mucor javanicus Mucor miehei
Geotrichum candidum Humicola lanuginosa Penicillium camembertii Rhizomucor javanicus Rhizomucor miehei
GCL HLL PcamL RJL RML R. javanicus, R. delemar, R. niveus”
Altus Biologics (ChiroCLEC-CR), Amano (lipase AY), Meito Sangyo (lipase MY, lipase OF-360), Boehringer Mannheim (Chirazyme@L-3)
CRL
PPL CE (BSSL)
Amano, Boehringer Mannheim (Chirazyme@L-7), Fluka, Sigma Genzyme, Sigma
Commercial Source and Name
Candida cylindracea
Other Names
Mammalian lipases porcine pancreas pancreatic cholesterol esterase Fungal lipases Candida rugosa
Abbreviation Origin of Lipase
Tab. 1. Selected Examples of Commercially Available Lipases
I Availability and Structure of Lipases
1.1.3 Classification of Lipases by Protein Sequence Naming lipases according to their microbial source sometimes obscures structural similarities. A better classification uses protein sequence alignments (Tab. 2 ) , which is also consistent with the 3-D structures of lipases (see Sect. 1.2).The mammalian (pancreatic) lipases form one group, the fungal lipases form two the Cundidu rugosa and the Rhizomucor families - and the bacterial lipases also form two the Pseudomonus and the Staphylococcus families. The Cundidu rugosa family includes CRL, GCL and, even though it is a mammalian lipase, pancreatic cholesterol esterase. These lipases are large (60-65 kDa). Note that Cundidu unturctica lipase B does not belong to this family, even though it comes from a Cundidu yeast. The Rhizomucor family includes lipases from a wide range of fungi: the Rhizopus lipases, the Rhizomucor lipases, Penicillium carnembertii lipase, HLL, CAL-B. These lipases are all small (30-35 kDa). The Pseudomonus lipases are also small and include all the Pseudomonus lipases and CVL. The Stuphylococcus lipases are medium-sized (40-45 kDa), but none are commercially available. One lipase in this group, a thermostable lipase from Bacillus thermocutenulutus, is available from Boehringer Mannheim since 1997. A number of lipases remain unclassified. For some, e.g., ANL, the amino acid sequence is not known, for others, e.g., CAL-A, the sequence is known (HOEGHet al., 1995),but it shows little similarity to the other lipases.
The most useful lipases for organic synthesis are: porcine pancreatic lipase (PPL), lipase from Pseudomonus cepucia (Amano lipase PS, PCL), lipase from Cundidu rugosa (CRL), and lipase B from Cundidu unturcticu (CAL-B). For lipid modification, lipase from Rhizomucor miehei (RML) is the most important. For this reason, we emphasize these five lipases in this review. Note that the synthetically useful lipases include examples from all the classifications in Tab. 2 except the Staphylococcus family. Two examples - RML and CAL-B come from the Rhizomucor family.
1.1.4 General Features of PPL, CRL, RML, CAL-B, and PCL Researchers use crude, rather than purified lipases, in most biocatalytic applications for two reasons. First, crude enzymes are less expensive. Microbes secrete lipases into the growth medium. To isolate the lipase, manufacturers simply remove the cells and concentrate. Crude preparations often contain other proteins, but they usually contain only one hydrolase. Two values for wt% protein in crude lipases are listed below. The higher value is the Lowry assay on the crude sample. This assay may overestimate protein content due to interferences in the Lowry assay by sugars and other additives.The lower value refers to the Lowry assay after precipitation of the proteins with trichloroacetic acid (WEBERet al., 1995a).This assay will underestimate the protein content if the proteins do not precipitate completely.The second reason researchers often use crude
Tab. 2. Classification of Commercial Lipases According to Similarities in Protein Sequence” Characteristics
Examples
Mammalian (pancreatic) lipases
50 kDa
PPL
Fungal lipases Candida rugosa family Rhizomucor family Unclassified
59-65 kDa 2 9 4 2 kDa
CRL, GCL, CE CAL-B, RML. ROL, PcamL, HLL ANL. CAL-A, CLL
Bacterial lipases Pseudomonas family Staphylococcus family
30-35 kDa 68-73 kDa
PCL, PFL, CVL BTL2
Classification ~
~~~~
~
a
43
Classification according to CYGLER et al. (1993) and SVENDSEN (1994) with some additions.
44
3 Biotransformations with Lipases
preparations is that they often work better than purified enzymes.The crude preparations contain sugars and other inert carriers which increase the surface area and stabilize the lipases, especially for reactions in organic solvents. A bound calcium ion stabilizes the 3-D structure of PCL, for this reason, crude preparations of PCL often contain added calcium salts. Because of the ill-defined nature of the product, most commercial lipases remain proprietary products. Suppliers sometimes create different preparations of the same lipase intended for different application. In addition, lipases from different suppliers may be identical due to cross-licensing agreements or may be different due to separate patents on different, strains of the same species. Crystallographers have solved the X-ray crystal structures of all five lipases (see below).
1.1.4.1 PPL Porcine pancreatic lipase has a molecular weight of 50 kDa. PPL from Sigma contains 8-20 wt% protein (WEBERet al., 1995a).Of all the commonly used lipases for synthesis, PPL is the least pure. Microbial lipases, even when they are not recombinant lipases, are purer because microbes secrete the lipase into the medium. Removal of the cells and precipitation of the lipase yields almost pure lipase. In contrast, PPL must be isolated from pancreas or bile which contains numerous hydrolases. SDS gel electrophoresis of crude PPL from Sigma shows four or five major proteins. Cholesterol esterase, trypsin, and chymotrypsin are likely contaminating hydrolases. Several groups reported increased enantioselectivity upon purification of PPL (e.g.. RAMOS-TOMBO et al., 1986; COTTERILL et al., 1991; QUARTEY et al., 1996;see also BORNEMAN et al., 1992).
1.1.4.2 CRL Commercial samples contain 2-11 wt% protein (WEBERet al., 1995a), the rest is sugars and inert carriers. Gel electrophoresis shows a single protein with molecular weight of 63 kDa when stained with Coomassie blue, but more sensitive staining reveals small amounts of other proteins. Molecular biologists have
cloned five different isozymes of CRL from the Candida rugosa yeast (LOTTIet al., 1993), which all have similar molecular weights. Researchers have not yet expressed the clones of CRL because C. rugosa uses an unusual codon for serine which leads to incorrect translation of the gene in other microorganisms. This means that commercial samples are not a recombinant protein and may contain more than one isozyme. Protein chemists have isolated several different lipases from commercial samples (RUA et al., 1993;CHANCet al., 1994).Differences in glycosylation of these lipases may contribute to these differences. In addition, some purification procedures appear to change the conformation of the lipase (WU et al., 1990; COLTON et al., 1995). One group also reported a small amount of contaminating protease (LALONDE et al., 1995). In spite of this complexity, commercial CRL is a useful and reproducible biocatalyst. Altus Biologics, Inc. sells cross-linked crystals of purified CRL as CLEC-CR (LALONDE et al., 1995).
1.1.4.3 RML RML has a molecular weight of 33 kDa (HuGE-JENSEN et al., 1987) and commercial material contains 25-57 wt% protein (WEBERet al., 1995a). RML is a recombinant lipase produced in Aspergillus fungus (HUGE-JENSEN et al., 1989).
1.1.4.4 CAL-B CAL-B has a molecular weight of 33 kDa and commercial material contains 16-51 wt% protein (WEBERet al., 1995a). CAL-B is a recombinant protein produced in Aspergillus fungus (HOEGHet al., 1995). CAL-B shows little or no interfacial activation and hydrolyzes long chain triglycerides only slowly. For this reason, it may be better classified as an esterase. It shows very high activity and high enantioselectivity toward a wide range of alcohols. Its enantioselectivity is low toward carboxylic acids.
1.1.4.5 PCL PCL is 320 amino acids long with a molecular weight of 33 kDa. Amano lipase P or PS is
-
1 Availability and Structure of Lipases
45
the industrial grade which contains 1-25 wt% protein as well as diatomaceous earth, dextran, and CaCl,. LPL-80 and LPL-200s are diagnostic grades that contain glycine. LPL-200s is pure protein containing some glycine. SAM-I1 from Fluka differs from lipase P or PS only in the purification method. Four groups have cloned and expressed PCL starting from dif3 4 5 6 7 8 ferent Pseudomonas strains, but the amino acid sequences of all four are very similar (for Fig. 1. Schematic diagram of the dp-hydrolase fold. original references see HOMet al., 1991: IIZUMI Oxyanion: residues that stabilize the oxyanion, Nu: et al., 1991;JORGENSEN et al., 1991;NAKANISHInucleophilic residue; for lipases and esterases this is a et al., 1991; for reviews see GILBERT,1993; serine. SVENDSEN et al., 1995). Expression of active lipase required stoichiometric amounts of an additional protein which probably guides the proper folding of the prolipase (HOBSON et al., der in all lipase amino acid sequences and ori1993). Commercial PCL is probably not a re- ent in the same three-dimensional way in all combinant protein. PCL shows interfacial acti- the structures as shown schematically in Fig. 1. vation with an increase in activity of -25 in The 3-D orientation of the catalytic machinery the presence of an interface (CURTISand KAZ- is approximately the mirror image of that in LAUSKAS, unpublished data). A single step pur- the subtilisin and chymotrypsin families of ification yields crystalline PCL, but this pure proteases. material is no longer active in organic solvents The catalytic mechanism for lipase-cata(BORNSCHEUER et al., 1994a). Cross-linking of lyzed hydrolysis is similar to that for serine the crystals gives CLEC-PC ( A h Biologics, proteases. First, the ester binds to the lipase Inc.), which are active in organic solvents (see and the catalytic serine attacks the carbonyl Sect. 2.3.1.3). XIE(1991) reviewed the applica- forming a tetrahedral intermediate (Fig. 2). tion of this lipase to organic synthesis. Collapse of this tetrahedal intermediate releases the alcohol and leaves an acyl enzyme intermediate. In a hydrolysis reaction, water attacks this acyl enzyme to form a second tet1.2 Structure of Lipases rahedral intermediate. Collapse of this intermediate releases the acid. Alternatively, an1.2.1 Lipases are alp Hydrolases other nucleophile such as an alcohol can attack the acyl enzyme thereby yielding a new Although lipases differ significantly in their ester (a transesterification reaction). In most amino acid sequences, all 11 lipases whose cases, it appears that formation of the acyl structures have been solved show similar 3-D enzyme is fast; thus, deacylation is the ratestructures (Tab. 3) (for reviews see CAMBILLAUdetermining step. and VANTILBEURGH,1993; CYGLER et al., 1992; DEREWENDA, 1994; DEREWENDA et al., 1994c; DODSONet al., 1992; RANSAC et al., 1996).This fold, called the alp-hydrolase fold (OLLISet 1.2.2 Lid or Flap in Interfacial al., 1992). consists of a core of eight mostly- Activation parallel @-sheets, which are surrounded on both sides by a-helices.The connectivity of the The X-ray structures of lipases usually show sheets and helices is the same in all a/p-hydrol- the “closed” conformation where a lid or flap ases (Fig. 1). (a helical segment) blocks the active site. HowLipases are serine esterases. The catalytic ever, X-ray structures of lipases containing machinery consists of a triad - Ser, His, and bound transition state analogs or bound lipids Asp( Glu) - and several oxyanion-stabilizing show the “open” conformation where the lid is residues. These residues occur in the same or- opened to permit access to the active site. For
46
3 Biotransformations with Lipases
Tab. 3. X-Ray Crystal Structures of Lipases Lipase
Comments
pdb Code"
Reference
Mammalian pancreatic lipases humanPL with colipase and phospholipid humanPL with colipase and phosphonate humanPL closed form horsePL closed form pigPL with colipase and surfactant open form with bile salts CE (BSSL)b
llpa llpb none 1hpl leth none
VAN TILBEURGH et al. (1993) EGLOFFet al. (1995a, b) WINKLER et al. (1990) BOURNE et al. (1994) HERMOSO et al. (1996) WANGet al. (1997)
Cundidu mgosu family CRL closed form CRL open form CRL' linoleate complex CRL sulfonate complexes CRL phosphonate complexes
ltrh lcrl none llpn, llpo, llpp llpm, lips
GCL
lthg
GROCHULSKI et al. (1993) GROCHULSKI et al. (1994b) GOSHet al. (1995) GROCHULSKI et al. (1994a) GROCHULSKI et al. (1994a); CYGLER et al. (1994) SCHRAG et al. (1991)
closed form
Rhizomucor family CAL-B open form CAL-B phosphonate complex CAL-B Tween 80 complex RML closed form RML phosphonate complex PcamL HLL ROL (R. delemar, R. niveus) ROL
disordered lid closed and partially open forms homology model
ltca, ltcb, ltcc llbs llbt 3tgl 4tgl,5tgl ltia ltib ltic, llgy
UPPENBERG et al. (1994) UPPENBERG et al. (1995) UPPENBERG et al. (1995) BRADYet al. (1990) BRZOZOWSKI et al. (1991); DEREWENDA et al. (1992) DEREWENDA et al. (1994a) DEREWENDA et al. (1994a, b) DEREWENDA et al. (1994a, b); KOHNOet al. (1996)
none
BEERet al. (1996)
Pseudomonus family CVL (PGL) closed form
ltah. lcvl
PCL PAL
loil, 21ip, 31ip none
NOBLEet al. (1993,1994); LANGet al. (1996) KIMet al. (1997); SCHRAG et al. (1997) MISSETet al. (1994)
open form homology model
a Accession code for the Brookhaven protein data bank. Human pancreatic cholesterol esterase is identical to human bile salt stimulated lipase in milk. The structure is for the bovine pancreatic cholesterol esterase. A homology model for pancreatic cholesterol esterase from salmon has also been reported (GJELLESVIK et al., 1994). Isozyme of CRL.
this reason, researchers believe a lipid-induced change in the lid orientation causes interfacial activation. Lipases show poor activity toward soluble substrates in aqueous solution because the lid is closed. Upon binding to a hydrophobic interface such as a lipid droplet, the lid opens and the catalytic activity of the lipase increases. In addition, the opening of the lid places one of the oxyanion-stabilizing residues into the catalytic orientation. Cutinase and acetylcholine esterase, which show no interfacial ac-
tivation, lack a lid and contain a preformed oxyanion hole (MARTINEZ et al., 1992,1994). However, the interfacial activation mechanism may be more complex. A number of lipases (e.g., lipase from Pseudomonas aeruginosa, CVL, and CAL-B) do not show interfacial activation even though they contain a lid. Lipase from Staphylococcus hyicus shows interfacial activation with some substrates, but not with others (for reviews see RANSAC et al., 1996;VERGER, 1997).
2 Water, Organic Solvents, and Other Reaction Media Flu 341
47
I
free enzyme
' H - p : H i s 449
HP
H.
Ser209
HN,
NI
Ala210 ,Gly124 Gly 123
N,
\
H i s 449
HN,
I'l\
,Gly124 Gly 123
Glu 341
(H i s 449
'
s e i 209
"';-H.y
HN,
Ala 210 /Gly 124 Gly 123
N,
Fig. 2. Hydrolysis of a butyrate ester catalyzed by lipase involves an acyl enzyme intermediate and two different tetrahedral intermediates. Formation of the acyl enzyme involves the first tetrahedral intermediate, Tdl.Alcohol is released in this step, thus, this step determines the selectivity of lipases toward alcohols. Release of the acyl enzyme involves the second tetrahedral intermedlate,Td2.When deacylation limits the rate, this step determines the selectivity of the lipase toward acids.The amino acid numbering corresponds to the active site of lipase from Candida rugosa, CRL.
1.2.3 Substrate Binding Site in Lipases X-ray crystal structures of transition state analogs bound to the active site of lipases have identified distinct binding sites for the alcohol and acid portion of esters. The alcohol binding site is similar in all lipases. It is a crevice containing two regions - a large hydrophobic pocket which is open to the solvent and a small pocket that faces the floor of the crevice. As discussed in Sect. 3.3.1, the shape of this pocket sets the stereoselectivity of lipases toward secondary alcohols. The alcohol binding site corresponds to the S1' site in proteases. The binding site for the acid portion of the ester varies considerably among the lipases. In CRL the acyl chain binds in a tunnel long enough to accommodate at least an 18 carbon chain. In RML and CAL-B this region is only a short trough o n the surface. In all three structures the a-carbon of an acyl chain binds just
below the large hydrophobic region of the alcohol binding site. Substituents at the a-carbon would extend into the hydrophobic pocket. This acyl binding region, formed by the tunnel or trough and the hydrophobic pocket, corresponds to the S1 site in proteases (Fig. 3).
2 Water, Organic Solvents, and Other Reaction Media 2.1 Hydrolysis in Aqueous Solutions and Two Phase Mixtures of Water and Organic Solvent The simplest lipase-catalyzed reaction is the hydrolysis of esters in water or biphasic mixtures of water and an organic solvent. Although proteases require a soluble substrate, lipases do not. A second phase is even desir-
48
3 Biorransfonnarions with Lipases
Fig. 3. Proposed substrate binding site in three synthetically-useful 1ipases.The catalytic Ser lies at the bottom of a crevice with the catalytic His on the left. Although the details of this crevice differ for each lipase, each crevice contains a large hydrophobic pocket (light gray) and smaller pocket (medium gray), labeled “stereoselectivity pocket”.This crevice is the alcohol binding site and the two pockets resemble the empirical rule discussed in Sect. 3.3.1.1.The regions that bind the acyl chain of the ester differ significantly among the three structures. For CRL, the acyl chain binds in a tunnel, the mouth of which is shown in dark gray. In CAL-B and PCL, the acyl chain binds in the large hydrophobic pocket of the crevice. Pictures were drawn with RasMac v2.6 using the Brookhaven protein data bank files cited in Tab. 3.
able because it activates most lipase by 10 to 100-fold, probably due to lid opening as discussed above in Sect. 1.2.2. Liquid substrate can also act as the organic phase. A general experimental procedure for a lipase-catalyzed reaction is as follows. Add 100 mg of liquid ester such as acetate or butyrate (or 1 mL of a solution of ester in a waterimmiscible solvent such as toluene or ethyl ether) to 5 mL of 50 mM phosphate buffer at pH 7. Monitor reaction by pH stat until reaction reaches 30% conversion. If a pH stat is not available, monitor by TLC or GC and use 100 mM buffer. Work up reaction, measure enantiomeric purity of both starting material and product, and calculate E using (Eq. 8) in Sect. 3.1.1 below. Preparative Biotransforma1992-1996) contains many detions (ROBERTS, tailed and tested procedures.
-
2.2 Lipases in Reverse Micelles Reverse micelles are the simplest way to run lipase-catalyzed reactions in almost pure or-
ganic solvent, but researchers rarely use reverse micelles for preparative reactions due to difficulties in workup due to the surfactant. Reverse micelles consist of a bulk organic phase containing aqueous droplets stabilized by surfactant. The lipase remains soluble and active in the water, while substrates and products dissolve in the organic phase (Fig. 4). The amount of aqueous phase is small, so that lipases can catalyze transesterification and ester synthesis reactions under these conditions. A second advantage is a large interfacial area between the micelles and the organic phase, which eliminates mass transfer limitations. These advantages simplify the kinetic analysis of lipases (HANet al., 1987;WALDEet al., 1993; STAMATISet al., 1995) and other enzymes (BOMMARIUS et al., 1995).Reverse micelles are also transparent and, therefore, suitable for spectrophotometric studies. HOLMBERG (1994) and BALLESTEROS et al. (1995) recently reviewed enzymic reactions in microemulsions. Anionic surfactants, in particular AOT, are best for lipase-catalyzed reactions because nonionic surfactants can inhibit lipases and
2 Water, Organic Solvents, and Other Reaction Media
d\ryJo- <
a
49
0
AOT (bis(2-ethylhexyl)sodium sulfosuccinate), an anionic surfactant
Br-+ surfactant
-
’‘ CTAB (cetyltrimethyl ammonium bromide), a cationic surfactant
‘
= !
N
T
I
nonpolar
polar
tail
head
3-35 nm-
C ~ H I ~ ~ ( O C H ~ C H ~ ) O H Triton X-100, a nonionic surfactant
Fig. 4. Reverse micelles or water in oil microemulsions.a Reverse micelles contain an inner aqueous phase stabilized by surfactant in a bulk organic phase. b Surfactants used to stabilize reverse micelles include anionic (e.g., AOT), cationic (e.g.,CTAB) and nonionic (e.g.,TritonX-100) surfactants.
can also react in transesterification reactions when the surfactant contains a free hydroxyl group (SKAGERLIND et al., 1992). A cationic surfactant (CTAB) decreases the maximal rate of a ROL-catalyzed hydrolysis of triolein by a factor of 50 compared to AOT (VALISet al., 1992). For preparative work, reverse micelles have several disadvantages. First, recovery of products from surfactant-containing organic solvent can be difficult. To simplify recovery, researchers added gelatin to the aqueous phase. Simple filtration recovers the lipase-containing aqueous phase. Several groups used CVL to make simple esters (REESet al., 1991,1993, 1995;BACKLUND et al., 1995;UEMASU and HINZE, 1994) and CVL or Pseudornonas sp. lipase (Genzyme) to resolve secondary alcohols (DE JESUSet al., 1995). CRL was inactive under these conditions. Interestingly, CVL within the gel remained active at -20°C (Rees et al., 1991). Another method to recover products from reverse micelles is to disrupt the emulsion with a temperature change into an oil-rich and a water-rich phase (LARSSON et al., 1990). Another disadvantage is that esterifications and transesterifications in reverse micelles often have lower yields than in other systems. For example, BORZEIX et al. (1992) compared the RML-catalyzed synthesis of butyl butyrate in hexane, in a two-phase mixture of water-hexane, and in AOT-stabilized reverse micelles in hexane.The rate of ester synthesis was
similar in all three systems, but the yield was significantly lower in the reverse micelles. Lipid modification in reverse micelles, especially synthesis of monoacylglycerides, also yielded less product than other reaction systems (CHANGet al., 1991;HAYESand GULARI, 1991; BORNSCHEUER et al., 1994b; HOLMBERG et al., 1989a; SINGHet al., 1994a, b). In addition, the surfactants used to stabilize reverse micelles can also denature lipases, but optimizing the water-to-surfactant ratio can minimize denaturation (FLETCHER et al., 1985;HANand RHEE, 1986;KIMand CHUNG, 1989;VALISet al., 1992). Although some enzymes show “superactivity” and changes in selectivity in reverse micelles (MARTINEK et al., 1982), lipases show only small changes in selectivity. BELLOet al. (1987) noted that CRL, which normally shows little fatty acid chain length selectivity, favored longer chain lengths in the transesterification of triglycerides in reverse micelles. HEDSTROM et al. (1993) reported significantly increased enantioselectivity of the CRL-catalyzed esterification of ibuprofen in reverse micelles as compared to hexane (E>100 vs. 3) (Eq. 2). Many other treatments and reaction conditions also increase the enantioselectivity of this reaction (see Sect. 1.3.4.2). Lipase from Penicillium sirnplicissimuni showed low selectivity (relative initial rates of 6-7) toward menthol enantiomers in reverse micelles (STAMATIS et al., 1995).
50
3 Biotransformations with Lipases
COOH
COOi-Pr CRL, i-PrOH i-BU
AOT/hexane, E >I00 hexane. E = 3 i-Bu
i
2.3 “Dry” Organic Solvents Researchers reported lipase-catalyzed esterifications in organic solvents containing approximately 10% water more than 50 years ago (SYM,1936;SPERRY and BRAND,1941), but most of this work was forgotten. In 1976, Unilever researchers patented a process for cocoa butter substitute using a lipase-catalyzed transesterification of lipids in hydrocarbon solvents (COLEMANand MACRAE,1977; see Sect. 4.2). KLIBANOV’S group discovered many other examples of enzyme-catalyzed reactions in organic solvents. KLIBANOV’S group further demonstrated that enzymes require only small amounts of water for activity (ZAKS and KLIBANOV,1984, 1985; CAMBOUand KLIBANOV,1984; for reviews see KLIBANOV, 1989, 1990 KOSKINENand KLIBANOV,1996). KLIBANOV’s work convinced others that enzyme-catalyzed reactions are not only possible, but also sometimes more convenient, in organic solvents. Today, researchers report slightly more reactions in organic solvents than in water. One advantage of reactions in organic solvents is the ability to do an esterification reaction instead of hydrolysis. Although lipases favor the same prochiral group in both cases, the two reactions yield opposite enantiomers. For example, acetylation of 2-benzyl glycerol yields the (S)-monoacetate, while hydrolysis of the diacetate yields the (R)-monoacetate. The lipases react at the pro-R position in both cases (Fig. 5). PCL, vinyl acetate
S
OBn A~o,,&,oA~
PPL phosphate buffer, pH 7
B
W
Another advantage of organic solvents is the potential to change the selectivity of the lipase in different solvents, sometimes called solvent (or medium) engineering. For example, the regioselectivity of the transesterification of a 2-octyl-1,4-dihydroxybenzeneester reversed from favoring the 4-position in cyclohexane to the 2-position in acetonitrile (RUBIO et al., 1991) (Fig. 6 ) . RUBIOet al. rationalized the reversal in selectivity according to differences in substrate solvation. Cyclohexane solvates the octyl group well, thus, the ester at the less hindered 4-position reacts. Acetonitrile solvates the octyl group poorly, thus the substrate binds in a manner that places the octyl group within the hydrophobic lipase active site. For this reason, the ester at the 1-position now reacts more rapidly. Consistent with this explanation, HALLING (1990) found that although the observed reaction rate can vary in different solvents, the true specificity constants of the enzyme vary only slightly after correcting for the activity of the substrate in different solvents. Note that substrate solvation changes do not explain changes in enantioselectivity, since both enantiomers are solvated equally in achiral solvents. However, solvation of enzyme-enantiomer complexes may differ and these differences may account for changes in enantioselectivity (see Sect. 2.3.2). Another reason to avoid water is to prevent decomposition of water-sensitive compounds such as organometallics or to simplify the
0 6 ~
AcO&OH
-
92% yield, 94% ee Terao et a/. (1988) Wang 8 Wong (1988)
OBn 75% yield, 91% ee HOAOAC Breitgoff eta/. (1986) R
Fig. 5. Although lipases favor the same prochiral group in both cases, acylation of the meso alcohol and hydrolysis of the meso ester yield opposite enantiomers.
2 Water, Organic Solvents, and Other Reaction Media favored 5-fold in cyclohexane
u?But
PCL transesterification w/ n-butanol Rubio eta/. (1991)
OBut
CBH17
R
favored 2-fold in acetonitrile
51
accelerates the reaction. Several groups reported that heating the reaction mixture with microwaves is more effective than simple heating (GELO-PUJIC et al., 1996;CARRILLO-MUNOZ et al., 1996;PARKER et al., 1996).
2.3.1.1 Adsorption and Entrapment
Lipase powders are insoluble in organic solvents and can be recovered by simple filtration at the end of the reaction. Unfortunately, even after optimizing the solvent and the water conworkup of hydrophilic compounds which are tent, catalysis is often thousands of times slowdifficult to recover from water. Some organic er than in water or water-organic solvent mixchemists favor reactions in organic solvents tures. One reason for the drop in activity is difsimply because they are less familiar with reac- fusional limitations, that is, the substrate cannot reach the lipase molecules in the center of tions in water. the particle.Another reason for the drop in activity is denaturation of the lipase during lyophilization.The simplest solution to both these 2.3.1 Increasing the Catalytic problems is adsorbing the lipase on an insoluble support such as Celite. Adsorption both inActivity in Organic Solvents creases the surface area and also avoids lyoEnzymes suspended in organic solvents are philization of the lipase. In a typical procedure, PCL (0.4 g) was disless active than enzymes dissolved in water. For crystalline subtilisin, SCHMITKE et al. solved in buffer (15 mL), mixed with insoluble (1996) attributed this drop in activity to ap- support (4.0 g), and dried at room temperature et al., 1992; proximately equal contributions from (1) (BIANCHIet al., 1988b; INAGAKI and SUNDHOLM, 1993b). Reactions changes in the pH optimum, (2) changes in KANERVA substrate solvation, and (3) low thermodynam- catalyzed by PPL adsorbed on Celite were ic activity of water. Noncrystalline subtilisin is 7-20 times faster than for crude PPL (BANFIet even less active, probably due to denaturation al., 1995). Sugars added to the buffer further during lyophilization. NOONEhas done such a increased the activity of immobilized PCL by a and KLIBANOV (1993) careful study for lipases, but additional pos- factor of 2-3. DABULIS sibilities for lipases are (1) diffusional limita- also found similar rate increases for PCL, et al. (1991) found tions (ability of the substrate to reach the ac- while SANCHEZ-MONTERO tive site) and (2) lid orientation. To minimize that the rate of heptyl oleate formation catadiffusional limitations, researchers disperse lyzed by CRL depended on the type of carbothe lipase on supports with high surface area hydrate. Rates increased upon adding lactose, or modify the lipase so that it dissolves in or- but decreased upon adding fructose, glucose, et al. ganic solvents. To ensure an open orientation sucrose, or sorbitol. SANCHEZ-MONTERO of the lid, researchers add lipids or surfactants suggested the differences may be due to the to the lipase. Salt hydrates or other techniques ability of sugars to change the activity of water. can optimize the water activity (Sect. 2.3.4) Indeed, many commercial samples of lipases and organic phase buffers (BLACKWOOD et al., contain large amounts of inert materials such 1994) or solid KHCO, (BERGERet al., 1990) as sugars or Celite (sometimes >95 wt%)),so can control the pH. Addition of water in a the available surface area is already large. manner that slows agglomeration of the en- CRL adsorbed on Celite showed increased zyme particles increased the rate of reaction stability toward acetaldehyde, a product of esby approximately a factor of 10 for CRL (TSAI terifications with vinyl esters (KAGA et al., and DORDICK, 1996).Heating the reaction also 1994). Adsorption of PCL on an acrylic resin Fig. 6. Regioselectivity changes in different solvents.
52
3 Biotransformations with Lipases
(Amberlite XAD-8) also increased the cata- 1995) or via a poly(ethyleneglyco1) linker lytic activity >200-fold (Hsu et al., 1990).Sev- (AMPONet al., 1994), or entrapping the lipase eral adsorption-immobilized lipases are com- in urethane prepolymers (KOSHIROet al., mercially available from Boehringer Mann- 1985). Covalent immobilization often increheim: CAL-B immobilized by adsorption on ases the thermal or operational stability of the macroporous acrylic resin and RML immobi- lipase, but does not activate the lipase. lized by adsorption on microporous phenolic anion exchange resin. Covalently immobilized lipases are also available. Adsorption-immobi2.3.1.3 Cross-Linked Enzyme lized lipases may desorb from the support in water; thus, covalently immobilized lipases Crystals - CLECs should be used in water. Cross-linked enzyme crystals are crystals Because lipases often show higher catalytic activity in the presence of insoluble organic (typically 1-100 pm diameter) of pure enzyme substrates, several groups have adsorbed or cross-linked with glutaraldehyde (ST. CLAIR 1996).Two lipase entrapped lipases in hydrophobic matrices. and NAVIA.1992;MARGOLIN, For example, hydrolysis of mixtures of tetra- CLECs are commercially available from Altus methoxysilane and alkyltrimethoxysilanes, Biologics Inc. (Cambridge, MA, USA): lipase RSi(OCH,),, in the presence of lipases forms from Candida rugosa (CLEC-CR) and Pseusol-gel entrapped lipases. Lipases entrapped dornonas cepacia (CLEC-PC) (LALONDEet 1995). Lipase CLECs rein hydrophobic sol-gels also show up to 100- al., 1995; LALONDE, fold increased activity in organic solvents main active and insoluble in organic solvents (REETZet al., 1995, 1996a, b; REETZ,1997). and, unlike adsorbed lipases, in water-organic Enantioselectivities remained unchanged in solvent mixtures. In addition, CLEC lipases most cases. The sol-gel entrapped lipases are are more stable to high temperatures and realso easily recovered and reused with no loss tain their activity over many cycles of use. For et al. (1995) reused CLECin activity. Fluka awarded the Reagent-of-the- example, LALONDE year-1997 prize to MANFRED REETZfor his dis- CR 18 times (recovering 20% of the activity). covery of the sol-gel immobilized lipases. Oth- whereas crude CRL lost virtually all activity er workers found smaller improvements in after one cycle. Mechanical losses and lipase similar systems (SATOet al., 1994; KAWAKAMIinactivation contributed approximately equaland YOSHIDA, 1995). Sol-gel-immobilized lip- ly to the loss in CLEC-CR activity. CLEC-CR ases also work in aqueous solutions (REETZet also showed improved enantioselectivity toal., 1995). The mechanism for activation is ward 2-arylpropionic acids; for ketoprofen the probably lid opening as suggested below for enantioselectivity increased from 5.2 to 64 et al., 1996; LALONDEet al., the lipid- or surfactant-coated lipases (Sect. (PERSICHE~TI 1995).LALONDE et al. (1995) attributed the in2.3.1.5). crease to the removal of a nonselective esterase during purification, but a conformational change may also contribute. CLECs prepared 2.3.1.2 Covalent Immobilization from different conformations of CRL (open Covalent immobilization creates a more vs. closed) also differed in their enantioselecstable link between the lipase and the support, tivity. Coating the crystals with surfactants inbut requires more effort than adsorption (for creased the activity of the lipase in organic solet al., 1996).After taking into reviews see AKITA,1996; BALCAOet al., 1996). vents (KHALAF The most common method is the cross-linking account the protein content, the activity was of adsorbed lipases with glutaraldehyde. Boeh- 2-90 times greater than crude lipase. The surringer Mannheim sells several lipases immobi- factant may maintain the water balance or it lized by this method. Other examples include may facilitate transfer of hydrophobic sublinking to an epoxy-containing resin (BERGER strates through the tightly bound layer of waand FABER,1991), to polystyrene via a cys- ter. teinyl-S-ethyl spacer (STRANIXand DARLING,
53
2 Water, Organic Solvents, and Other Reaction Media
2.3.1.4 Covalently Modified Lipases Soluble in Organic Solvents
n r\l
m
zw;;
n
n g E2g m 2-=m' vi 2 n 2 G =-.; nnz mz z nGm
Another way to increase the activity in organic solvents is to modify the lipase so that it dissolves in the organic solvent. KIKKAWA (1989) coupled polyethylene glycol (PEG) to the free amino groups of PCL and lipase from Pseudomonus frugi. The modified lipases were soluble in benzene, toluene, and chlorinated hydrocarbons and catalyzed the formation of lactones from ethyl 16-hydroxyhexadecanoate and the resolution of 2-phenylethanol. Addition of hexane precipitated the PEG-lipase thereby simplifying recovery (KODERAet al., 1994). Using the same principle, KODERAet al. (1994) coupled PFL to a comb-shaped polymer yielding a more stable and more active lipase. PEG-modified CRL was significantly more stable in organic solvents than unmodified CRL (BASRIet al., 1995;HERNAIZ et al., 1996).
-
:%m"
w
2E
$2
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\o
m
m-mm,C3.vr
- w Z % zSgG*YgTl ;+--=.+cj, m-m - 2 w + w w
4 w m m O j i z 4 a p ~
Q z552zz$Q ;z$ z > 4 4 w 0 0 z a ~ " t r c k % = 8 g
Y Y < O O < Y ? O -
J
~ ~ < < m o o u o m 5~ z
c
U
. d
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Z
V
"V
a
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2 2 2
j
n
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2 p: .' L-1 u s ? P d2aaa&-&LlAi 3 2Zg8gggg82gg g t z
2.3.1.5 Lipid- or Surfactant-Coated Lipases
00 dp:
. 5
C
One of the simplest ways to increase the activity of a lipase in organic solvent is to coat the lipase with a lipid or surfactant before lyophilization (Fig. 7; Tab. 4). For example, a water-insoluble complex containing approximately 10 wt% protein formed upon mixing aqueous solutions of ROL and a nonionic amphiphile (didodecyl N-D-glucono-L-glutamate (OKAHATA and IJIRO,1988,1992). Researchers estimated that 150k 30 amphiphile molecules surrounded each lipase molecule. The modified lipase was soluble in most organic solvents and was >lo0 times more active than suspended enzyme in the synthesis of di- and triglycerides from lauric acid and monolaurin. Other researchers reported similar increases in reaction rate with other lipases and other lipids or surfactants. Adding a hydrophobic interface before lyophilization may prevent the denaturation which often accompanies lyophilization. Further, the hydrophobic interface may open the lid of the lipase (MINGARRO et al., 1995,1996). The open form of lipases is more active due to a more accessible active site and the more suit-
.
.-* 0 m
u
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mgg2 g5;;::3" 2& 6~-o-,-o". - -5.5.52 - s u u= rn
m z 0 2Ti.G; g s z z z % +z 2z . 2 +!-J= = 2 2 :. w p.5'5 . - 5 % & w w w 6 a w w , L
-
0)
a
b
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:,
.-,as *.= g g:s:s:3
h w w w n n - - . - + . d 0 0 0 o&?<.?
5N
2
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s
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.-
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z
2
.,rnoo .-.-._ 0n .=g jg's'<$ZE's'sE~E 5 " zzSSm-,;gmmm 5
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sm
2 zrx n=a c arnL wLrnrxcnhmc. *aCL mwL .rn.p."." ad L* .Cd L ; * 8 rn
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9
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54
3 Biotransformations with Lipases OH OH OH ase 0
lipid
didodecyl KD-glUCOno-L-glUtamate 2ClzGG
glutamic acid dioleyl ester ribitol amide 2CtsA%E
Fig.7. Schematic of lipid coated lipase and structure of lipids used for the coating.
able orientation of the oxyanion-stabilizing residues. The open form is then “frozen” removing the water by lyophilization or precipitation. Consistent with this explanation, surfactant treatment increased the catalytic activity of lipases RML, GCL, and CRL, all of which contain lids, but not cutinase, which lacks a lid. Surfactants also activated crosslinked crystals of lipases (KHALAF et al., 1996), but the mechanism is unclear since the crosslinking presumably prevents significant conformational changes. Other workers used imprinting to increase enantioselectivity as well as reaction rate. Lyophilization of CRL with (R)-1-phenylethanol followed by lipid coating increased the enantioselectivity from 5.5 to 77. Heating or storage for a few days abolished the increased enantioselectivity (OKAHATA et al., 1995a). Surfactants also activated cross-linked crystals of lipases (KHALAF et al., 1996),but the mechanism is unclear since the cross-linking presumably prevents significant conformational changes.
2.3.1.6 Choosing the Best Organic Solvent for High Activity Finding the best organic solvent for a lipasecatalyzed resolution is still a trial and error process, but nonpolar solvents are usually better than polar solvents. LAANE(LAANE,1987; LAANEet al., 1987) divided solvents into three groups according to their log P value - the logarithm of their partition coefficient between octanol and water. Lipases, as well as other biocatalysts, showed low activity in solvents with logP values less than 2, which includes polar solvents like methanol, acetone, pyridine, and diethyl ether. Biocatalytic activity
was difficult to predict for solvents of moderate polarity (24) such as decane and diphenyl ether. Good solvents for lipase-catalyzed reactions include hexane, vinyl acetate, and toluene. Other researchers correlated activity with solvent parameters such as the dielectric constant or the dipole moment (FITZPATRICK and KLIBANOV, 1991; BORNSCHEUER et al., 1993), but logP gave the best correlation. Researchers believe that lipases must retain an essential shell of water to remain active in organic solvents. Nonpolar solvents do not affect this shell of water, while polar solvents strip this essential shell thereby inactivating the lipase. Adding water to the polar solvent helps to retain activity (see Sect. 2.3.4 on optimizing the amount of water in an organic solvent). Clumping or aggregation of the lipase upon addition of water can lower the activity.
2.3.2 Increasing the Enantioselectivity in Organic Solvents Lowering the temperature sometimes increases enantioselectivity. For example, SAKAI et al. (1997) increased the enantioselectivity of a PCL-catalyzed acylation in diethyl ether from E = 1 7 at 30°C to E=84-99 at -40°C. On the other hand, YASUFUKU and UEJI (1995, 1996, 1997) increased the temperature to increase enantioselectivity. The enantioselectivity of a CRL-catalyzed reaction of 2-phenoxypropanoic acids increased from E = 5 at 10°C to E=33 at 57°C.
2 Water, Organic Solvents, and Other Reaction Media
Changing from water to organic solvent often changes lipase enantioselectivity, as does changing from one organic solvent to another. Many researchers used this “medium engineering” to optimize reactions in organic solvents. For example, MORIet al. (1987) reported that PPL showed no enantioselectivity in the hydrolysis of seudenol acetate in aqueous solution, but JOHNSTON et al. (1991) reported moderate enantioselectivity (E= 17) in the acetylation of seudenol with trifluoroethyl acetate in ethyl ether.The enantioselectivity of the CAL-B-catalyzed acetylation of seudenol with vinyl acetate varied from 8-32 depending on the solvent and the water content.The highest enantioselectivity was in dry benzene (ORRENIUSet al., 1995b). Occasionally, the enantioselectivity even reverses upon changing the solvent. For example, CRL esterified the (R)-enantiomer of a chiral acid, 2-phenoxypropionic acid, with butanol in carbon tetrachloride (E = 16), but the (S)-enantiomer in acetone (E= 1.6) (UEJIet al., 1992).In another example, PCL acetylated the (R)-enantiomer of a secondary alcohol, methyl 3-hydroxyoctanoate, with vinyl acetate in methylene chloride ( E = 5 ) , but the (S)-enantiomer in hexane (E=16) (BORNSCHEUER et al., 1993). In the most dramatic example, Amano lipase AH hydrolyzed the pro-R ester of a dihydropyridine derivative in cyclohexane (E-20), but the p r o 3 ester in diisopropyl ether (E> 100) (HIROSE et al., 1992). Finding the molecular basis of the enantioselectivity changes is difficult because the energies involved are small. Rarely does the enantioselectivity change by more than a factor of 10, almost never by a factor of 100.A factor of 10 corresponds to a AAG’ of 1.4 kcal mol-’, a relatively weak interaction. Nevertheless, these enantioselectivity changes are often enough to tip the balance from a useless reaction to a useful one. For this reason, many researchers have tried to understand why enantioselectivity changes in different solvents. They proposed at least four different explanations, but none can predict changes in enantioselectivity reliably (reviewed by CARREA et al., 1995). First, KLIBANOV proposed that differences in the solvation of the enzyme-substrate transition state complex determine the selectivity
55
(RUBIOet al., 1991);KE et al., 1996).For example, they explained the variation in enantioselectivity for the lipase-catalyzed hydrolysis of a prochiral diester (Fig. 8) (TERRADAS et al., 1993). They suggested that the substrate, especially the hydrophobic naphthyl group, binds tightly to the active site in polar solvents giving high enantioselectivity, but binds loosely, or not at all, in nonpolar solvents giving low enantioselectivity.To estimate the differences in solvation, the authors used the solvent logP and found a good correlation with enantioselectivity. Other researchers reported similar correlations (recent examples are given by EMAet al., 1996; HOF and KELLOGG, 1996a), but PARIDA and DORDICK (1991) found only partial correlation in a CRL-catalyzed esterification of 2hydroxy acids and, in this case, the enantioselectivity increased in less polar solvents. CARREAet al. (1995) found no correlation of enantioselectivity and logP in PCL-catalyzed acylation of several secondary alcohols. The lack of correlation suggests that log P may be a poor measure of enzyme-substrate solvation in these cases. Recently, KE et al. (1996) used a more sophisticated approach to estimate solvation of the enzyme-substrate complex. They estimated the portion of the substrate bound to the enzyme and calculated the activity coefficient of this fragment. This approach predicts the enantioselectivity of crystalline proteases in organic solvents,but not for amorphous proteases in organic solvents. Second, researchers suggested that solvent changes the active site by binding in it or near it (NAKAMURA et al., 1991; SECUNDO et al., 1992; HIROSE et al., 1992). X-ray structures of Pseud. lipase
E = 2.6 in CC14
E >30 in acetonitrile
O
a
0
Fig. 8. Enantioselectivity changes in different sol-
vents.
56
3 Biotransformationswith Lipases
protease crystals soaked in solvents indeed 2.3.3 Acyl Donor for Acylation showed solvent molecules (hexane or acetonitrile) bound to the active site (FITZPATRICK et Reactions al., 1993;YENNAWAR et al., 1995). OTTOLINA et The ideal acyl donor would be inexpensive, al. (1994) found that the activities of lipases (PCL, CVL, PPL, CRL, RML) increased by as acylate quickly and irreversibly in the presmuch as a factor of eight in (R)-carvone as ence of lipase, and be completely unreactive in compared to (S)-carvone. The enantiomeric the absence of lipase. No acyl donor fulfills all solvents presumably form different complexes three criteria. For transformations of inexpenwith the lipase. However, the enantioselectiv- sive chemicals (e.g., modified lipids, Sect. 4.2), ity toward secondary alcohols did not differ in cost is most important, so researchers use acids the two solvents. ARROYOand SINISTERRA and simple esters (e.g., methyl, glyceryl). Acy(1995) reported small changes in enantioselec- lations with these donors are often slow and tivity of CAL-B toward a carboxylic acid (ke- reversible with an equilibrium constant near one. To drive reactions to completion, retoprofen) in (R)-vs. (S)-carvone. Third, VAN TOL et al. (1995a, b) suggested searchers removed the water or alcohol by et al., 1989), azeothat the changes in enantioselectivity are due evaporation (BJ~RKLING et al., 1992), mito a combination of errors in measuring enan- tropic distillation (BLOOMER et al., tioselectivity and changes in solvation of the crowave heating (CARRILLO-MUNOZ substrate. The endpoint method for measuring 1996), or chemical drying agents such as moenantioselectivity can give erroneous results lecular sieves or inorganic salts (KVITTINGEN when product inhibits the reaction (Sect. et al., 1992). In other cases, crystallization of 3.1.1). After correcting for inhibition and the the product drives the reaction (MCNEILLet thermodynamic activity of the substrates, VAN al., 1991). For resolution reactions of fine chemicals, TOL et al. (1995a, b) found that the enantioselectivity of a PPL-catalyzed acylation of gly- researchers use activated acyl donors (Fig. 9). cidol with vinyl butyrate did not change in hex- Lipase-catalyzed acylations with these donors ane, diisopropyl ether, tetrachloromethane, are one to two orders of magnitude faster than and 2-butanone (E=5.5). Without these cor- with acids or simple esters. In addition, activatrections, the apparent enantioselectivity varied ed acyl donors shift the equilibrium constant in favor of acylation. In the case of enol esters between 20 and 2.7 in a single reaction. Fourth, researchers correlated the increased and acid anhydrides, acylation is practically irenantioselectivity of subtilisin in different sol- reversible. An irreversible reaction is imporvents with the increased flexibility of the ac- tant for kinetic resolution of alcohols because tive site on the nanosecond time scale (BROOS the reverse reaction degrades the enantiomeret al., 1995).Noone has yet made such a corre- ic purity of the remaining starting material thereby lowering the efficiency of the resolulation for lipases. tion (see Sect. 3.1.1). activated esters 0
trifluoroethyl butyrate
S-ethyl thiooctanoate
enol esters
Fig. 9. Examples of activated acyl donors for irreversible acylation of alcohols.
0
0
biacetyl monooxime acetate anhydrides
R = H, vinyl acetate diketene acetic acid R = CH3, isopropenyl acetate anhydride R = OEt, 1-ethoxyvinylacetate
succinic acid anhydride
2 Water, Organic Solvents, and Other Reaction Media
R'o/'CF,
'
m\
PPL, EtzO c E=8O
R = I I - C H23 ~~
3
97% ee (S)-(+)-sulcatol insect pheromone
90% ee
(3)
enantioselective acylations appeared in 1988 (WANGand WONG,1988; WANGet al., 1988; LAUMENet al., 1988; TERAOet al., 1988). Hoechst AG patented the resolution of alcohols using vinyl esters in 1988. Since that time researchers have resolved hundreds of alcohols using this method. For example, BERKOWITZ et al. (1992) efficiently resolved glycals for the synthesis of artificial oligosaccharides (Eq. 4)Most lipases except CRL and GCL tolerate the liberated acetaldehyde. Acetaldehyde slowly inactivates these two probably by formation of a Schiff base with a Lys residue (WEBERet al., 1995b).Acetone from isopropenyl acetate is less reactive and may not inactivate CRL or GCL, but this has not been investigated. Another alternative is 1-ethoxyvinyl acetate which liberates ethyl acetate (KITA et al., 1996;SCHUDOK and KRETZSCHMAR, 1997). Although the acylation of alcohols by enol esters, such as vinyl acetate, is indeed irreversible, another equilibrium can cause reversibility and lower the enantiomeric purity of the remaining alcohol (LUNDHet al., 1995).Since the reaction mixture contains small amounts of water, the lipase can catalyze hydrolysis of the product acetate ester to the alcohol plus acetic acid. Hydrolysis of the faster-reacting acetate lowers the enantiomeric purity of the remaining alcohol. To minimize this hydrolysis LUNDHet al. recommend dry conditions and an excess of vinyl acetate. In addition, stopping the reaction before 50% conversion, separating the ester and alcohol, and subjecting the alcohol to a second esterification will also minimize hydrolysis. OH
PCL dimethoxyethane
(*I
+
'\
Researchers first used activated esters where the alcohol is a better leaving group. For example, STOKESand OEHLSCHLAGER (1987) acylated sulcatol with trifluoroethyl laurate and recovered the unreacted alcohol in 97% ee (Eq. 3). However, for another secondary alcohol DE AMICI et al. (1989) could not recover the unreacted starting material in high enantiomeric purity even though the transesterification was enantioselective. They attributed this difficulty to the reversibility even for this activated ester. The less expensive trichloroethyl esters are less convenient because the product trichloroethanol is difficult to remove (bp. 151"C).The thioester S-ethyl thiooctanoate drives the reaction both because the thiol is a good leaving group and because the ethanethiol is easily removed by evaporation (OHRNER et al., 1992; FRYKMAN et al., 1993), but working with volatile thiols requires extra care. The oxime esters react faster than simple esters and even enol esters (GHOGAREand KUMAR,1989,1990),but the nonvolatile oxime may complicate separations. Several other leaving groups (not shown) are less useful: cyanomethyl esters release the toxic formaldehyde cyanohydrin while 2-chloroethyl esters do not activate the ester enough. The most useful activated acyl donors are enol esters, such as vinyl acetate or isopropenyl acetate. The product alcohol tautomerizes to a carbonyl compound, thereby driving the reaction and eliminating potential product inhibition.The first reports of lipase-catalyzed acylations with enol esters appeared in 1986-1987 (SWEERSand WONG,1986; DEet al., 1987) and the first GUEIL-CASTAING
57
Ph =-97% ee
(4)
I
>97% ee
V
0-
58
3 Biotransformations with Lipases
Acylation with diketene, a cyclic enol ester, is fast and has the advantage that it produces no by-products (BALKENHOHL et al., 1993b; JEROMIN and WELSCH,1995; SUGINAKA et al.,
OH Aph+ o
0
e
1996). However, the reported enantioselectivity was slightly lower than that for vinyl acetate, possibly due to nonenzymic acylation. For example, the acylation of 1-phenylethanol with diketene showed an enantioselectivity of 12-80 (Eq. 5), while vinyl acetate showed an enantioselectivity > 100 with the same enzyme (LAUMEN et al., 1988;NISHIOet al, 1989). Acid anhydrides also irreversibly acylate alcohols (BIANCHI et al., 1988b), but the release of carboxylic acid may decrease the enantioselectivity of the reaction. For example, CRL-
3vl
MeOdoMe
c c
OH
enantioselectivity of a CAL-B-catalyzed acylation of several secondary alcohols was highest with 2-chloroethyl butyrate (HOFFet al., 1996). Vinyl butyrate, butyric anhydride, and 2,2,2-
acetic ;Ei;g-ydri: CRL base
none
E
23
MeOJ
trichloroethyl butyrate showed 5-35 times lower enantioselectivity. In most cases, the key to high enantioselectivity is to avoid chemical acylation. Sometimes acyl donors with longer chains (butyrates and above) show higher enantioselectivity than those with shorter chains like acetate (for examples see SONNET, 1987; STOKESand OEHLSCHLAGER, 1987; HOLMBERG et al., 1989b; G u o et al., 1990; YAMAZAKI and HOSONO,1990; EMAet al., 1996).
OMe
MeOLOMe
OH
OAc
KHC03 240
catalyzed acetylation of a bicyclic secondary alcohol with acetic anhydride was moderately enantioselective (E = 19), but in the presence of solid potassium bicarbonate the enantioselectivity increased dramatically to E =240 (Eq. 6) (BERGERet al., 1990).In polar solvents, uncatalyzed acylation by acid anhydrides can lower the overall selectivity. TERAOet al. (1989) used succinic acid anhydride to both acylate an alcohol and to simplify the separation of unreacted alcohol and ester. Simple extraction separated the neutral alcohol from the charged succinate half ester. KANERVAand SUNDHOLM (1993a) compared the enantioselectivity of PCL-catalyzed acylation with butyric anhydride, vinyl butyrate, and trifluoroethyl butyrate. All showed similar enantioselectivity,but for one substrate the rate of reaction with trifluoroethyl butyrate was very slow. On the other hand, the
2.3.4 Water Content and Water Activity The amount of water in the reaction mixture strongly influences the reaction rate, and to a lesser extent the enantioselectivity, of lipasecatalyzed reactions in organic solvent. Polar solvents require typically 1-3% added water for optimal activity, while nonpolar solvents require only 0.05-1 % added water. Enzymes require this minimum amount of water to maintain their structure and flexibility (RUPLEY et al., 1983; AFFLECK et al., 1992;BROOSet al., 1995). Initially, researchers optimized the amount of water for a lipase-catalyzed reaction in organic solvent by measuring the total water content with Karl-Fischer titration. More recently, researchers found that the thermody-
2 Water, Organic Solvents, and Other Reaction Media
59
namic water activity (a,) is a better measure of pairs of salt hydrates can be added directly to the amount of water, especially when compar- the reaction to buffer the water activity. Altering different reaction conditions (HALLING, natively, preequilibrated silica particles of 1990,1994,1996; BELLet al., 1995). For exam- known a, may be used (HALLING, 1994). Howple, the optimum reaction rate for RML in sol- ever, these approaches are not practical on a vents ranging from 3-pentanone to hexane large scale due to cost, low water activity-bufoccurred at the same thermodynamic water fering capacity, and difficulties in recovering activity, a, =0.55, but at widely differing total the catalyst. One improvement is to add a siliwater content (VALIVETY et al., 1992b). Simi- cone tube containing a saturated salt solution larly, optimal activity of RML immobilized on (e.g., a, =0.75 for water saturated with NaCl). different supports occurred at a, = 0.55, but at The circulating salt solution can both take up different total water content (OLADEPO et al., and release water through the silicone (WEHT1994,1995).Polar solvents require more added JE et al., 1993,1997).On an industrial scale, the water than nonpolar solvents because polar best route may be an a, sensor (several are solvents competed more effectively with the li- commercially available; HALLING,1994) compase for the available water. BELLet al. (1995) bined with either drying by recirculation of the suggested that water activity is like tempera- headspace gases through a drying column or ture, while water content is like heat content. water addition (KHANet al., 1990). These huTwo systems may have the same water activity midity sensors can also monitor the water acor temperature, but at the same time differ in tivity continuously during reactions (GOLDthe water content or heat content. BERG et al., 1988, 1990 KHANet al., 1990; Of the available methods for controlling the BORNSCHEUER et al., 1993; LAMAREand LEthermodynamic water activity (Tab. 5) the sim- GOY, 1995). Lipases differ in the amount of water needplest are equilibration of the reaction components with salt solutions of known a, or pairs ed to maximize the rate of esterification beof salt hydrate. Equilibration occurs, albeit tween decanoic acid and dodecanol in hexane slower, even without direct contact between (VALIVETY et al., 1992a). RML and ROL were the reaction components and the salt solutions most active at low a, (optimum: 0.32< or salt hydrates.The pairs of salt hydrates (e.g., a, < 0.55), HLL, CRL, and PCL required a, CuSO, .5 H,OICuSO,. 3 H,O gives a, =0.32 close to one. Sequence comparison of HLL at 25 "C; Na,P,O,. 10H,O/Na,P,O, gives a, = and ROL suggested that changes in charged 0.56 at 35 "C) act as water buffers taking up or residues in the "hinge and lid" region may be releasing water to the reaction components. significant in low a, tolerance. Different reacHALLING(1992) lists water activity values for tions may also have different optima. The overall rate of esterification between glycerol 48 salt hydrate pairs. It is more difficult to maintain constant wa- and oleic acid using RML did not change sigter activity in reactions that consume or pro- nificantly with changes in water activity, but duce water (e.g., esterification between an acid the synthesis of diolein from monoolein was and alcohol) because water must be removed fastest at a,=0.5 and triolein synthesis was or added during the reaction. Salt solutions or fastest at low values of a, (DUDALand LORTIE, Tab. 5. Methods to Control Water Activity in Organic Media
Method
Reference
GODERIS et al. (1987); ADLERCREUTZ (1991); et al. (1992b); BLOOMER et al. (1991); VALIVETY et al. (1996) ROSELL Equilibrate with saturated salt solutions via silicone tubing WEHTJE et al. (1993,1997) Equilibrate with salt hydrates HALLING (1992); KVITTINGEN et al. (1992); KIMand CHOI(1995) Equilibrate with wet silica gel HALLING (1994) Measure a, with sensor and control drying of headspace KHANet al. (1990) Equilibrate with saturated salt solutions
60
3 Biotransformations with Lipases
1995).It may also be useful to change a, as the reaction proceeds. For example, the initial phase of a PCL-catalyzed esterification of decanoic acid and dodecanol proceeds faster at high a,, but a lower a, at later stages gives higher yields (SVENSSON et al., 1994). Water activity also influenced the enantioselectivity of lipase-catalyzed reaction, but not in a consistent manner. For the resolution of 2methyl alkanoic acids with CRL (Eq. 7) the enantioselectivity was higher at higher water activity (HOGBERGet al., 1993; BERGLUND et al. 1994). However, for a resolution of seudeno1 increasing water activity had different effects in different solvents (ORRENIUS et al., 1995b). In most cases, the enantioselectivity was higher at low water activity. For example, the enantioselectivity in hexane was 20 at a,<0.11, but dropped to less than 10 at a, > 0.75. However, for dichloromethane and t-amyl alcohol the enantioselectivity was independent of water activity and for vinyl acetate and 3-pentanone the enantioselectivity was
n-CsH3 5
+ n-C12H2,0H
CRL cyclohexane salt hydrate pairs
-
E no control 7
&
0.15
0.76
ment costs and more complex reaction engineering. Of the several possible supercritical fluids, most researchers use carbon dioxide because it is nonflammable, nontoxic, cheap, and reaches the supercritical state at low temperature (31.1 "C). Moreover, its solvating properties are comparable to acetone. To dissolve polar substrates in supercritical carbon dioxide (SCCO,), researchers either added a small amount of polar solvent such as dichloromethane, acetone, or t-butanol (CAPEWELL et al., 1996) or they used techniques developed previously for organic solvents, e.g., complexation of fructose with phenyl boronic acid or immoet bilization of glycerol on silica gel (CASTILLO al., 1994). KAMATet al. (1995) suggested that fluoroform may be a better supercritical fluid for an enzyme-catalyzed reaction because carbon dioxide reacts with the lysine residues on an enzyme to make carbamates. In addition, supercritical fluoroform is a better solvent than SCCO,.
C00n-C12H25 1
n-CsHi5 A
+
COOH
(7)
n-CsH(i
23 95
higher at high water activity. This variation reflects the fact that the effects of different solvents on enantioselectivity are still not well understood (see Sect. 2.3.2).
2.4 Supercritical Fluids As the temperature and pressure of a liquid are raised above the critical point, separate phases of liquid and gas disappear into a single phase called a supercritical fluid. Supercritical fluids have densities and dissolving powers near those of a liquid, but the viscosities near that of a gas. The advantages of supercritical fluids are rapid mass transfer due to the low viscosity, simple downstream processing by evaporation, the elimination of organic solvents, and the ability to change solvation properties by changing the pressure. The disadvantages of supercritical fluids are higher equip-
NAKAMURA et al. (1986) first showed that lipases remain active in supercritical fluids. ROL catalyzed the interesterification of triolein and stearic acid to 8% conversion in SCCO,. Since then researchers examined a wide range of reactions, especially lipid modifications (Tab. 6). The observed changes in conversion, enantioselectivity, or lipase stability are similar to those in organic solvents. Several reviews of enzyme-catalyzed reactions in supercritical fluids have appeared (BALLESTEROS et al., 1995; AALTONEN and RANTAKYLAE,1991;NAKAMURA, 1990 HAMMOND et al., 1985). Most reactions in Tab. 6 used immobilized RML in SCCO, at -40°C and 150 bar in batch systems and the research focused on optimizing the activity and stability of the lipase. For example, MARTYet al. (1992) varied the water content to maximize the rate of esterifi-
-
NAKAMURA et al. (1986); CHIet al. (1988) DODDEMA et al. (1990);JANSSENS et al. (1992) ERICKSON et al. (1990) MARTY et al. (1990,1992) DUMONT et al. (1992) MILLER et al. (1990)
KNEZand HABULIN (1992) BERNARD et al. (1992) MARTINS et al. (1992) VERMUE et al. (1992) KAMATet al. (1992,1993) CHULALAKSANANUKUL et al. (1993) CASTILLO et al. (1994) CHAUDHARY et al. (1995) IKUSHIMA et al. (1993)
B, 35-50"C. 150 bar C, 35-80°C, 80-140 bar, IPI B, 40°C. 90-290 bar C, 33-50°C, 110-170 bar, B, 5 0 T , 150 bar S-C, 35"C, 79-107 bar B, SO'C, 125 bar
C, 40°C, 100 bar, IPI
B, 40"C, 84-167 bar B, 5 0 T , 125 bar B, 35"C, 140 bar C, 60°C. 125-200 bar B, 40-50°C, 107 barb B, 40°C, 140 bar B, 40°C, 150 bar B, 50"C,60-340 bar' B, 31-40°C, 76-193 bar B, 36-7OoC, 100-170 bar B, 40°C, 200 bar
trioleinlstearic acid
ethyl acetateli-amyl alcohol
trilaurin/palmitic acid oleic acidlethanol
myristic acidlethanol
trilaurinlmyristic acid
trilaurinloleic acid methylester
myristic acidlethanol
oleic acidloleyl alcohol
myristic acid/ethanol
butyric acidlglycidol
ethyl acetatelnonanol methacrylatel2-ethyl hexanol
propyl acetatelgeraniol
e.g., fructose-PBAloleic acid
adipate/l,4-butanediol
oleic acidlcitronellol
3-hydroxy esterlvinyl acetate
2"-alcoholslacylation
ROL
RML ROL RML
RML
ROL
RML
RML
RML, ANL
RML
PPL
RML
Several
RML
RML
PPL
CRL
PCL PCL
RANTAKYLAE and AALTONEN (1994) JACKSON et al. (1947)
B, 50°C. 100-150 bar B, 65"C, 275 bar
B, 40-7OoC, 200-345 bar
randomization of, e.g., palm olein
soybean oil/e.g., glycerol or methanol
RML
CAL-B
CAL-B
a B: batch reactor; C: continuous reactor; S-C: semi-continuous reactor; IPI: integrated product isolation; PBA: phenyl boronic acid; IA: isopropenyl acetate;TA: triacetylglycerol. Ethane, ethylene, fluoroform, propane, sulfur hexafluoride were also studied. Fluoroform also.
JACKSON and KING(1997)
MlCHOR et al. (1996)
B, 35-7OoC, 150 bar
menthol or citronellol/IA,TA
ibuprofen
CRL, Esterase EPlO
BORNSCHEUER et al. (1996); CAPEWELL et al. (1996) CERNIA et al. (1994a, b)
BERNARD and BARTH(1995)
ADSHIRI et al. (1992)
References
Process Conditions"
Reaction
Examples of Lipase-Catalyzed Reactions in Supercritical Carbon Dioxide
Lipase
Tab. 6.
s
62
3 Biotransfonnations with Lipases
cation of oleic acid with ethanol. The optimum water amount increased upon addition of a small amount of ethanol to the reaction.As expected, MARTY et al. (1992) observed no diffusion limitations,nor did MILLERet al. (1990) in a similar reaction, but BERNARD and BARTH (1995) observed a partial diffusion limitation. Conversion and residual activity of PCL were improved by adding molecular sieves to the reaction (CAPEWELL et al., 1996),maximum conversion was influenced by pressure and temperature (NAKAMURA et al., 1986; CHI et al., 1988).initial rates were twice higher in SCC02 compared to n-hexane, which was attributed in part due to different solubility of the substrates in the two solvents (MARTY et al., 1990, 1992). Most comparisons suggest that the enantioselectivity of lipases in SCC02 is similar to or slightly lower than in organic solvents (CAPEWELL et al., 1996;MARTINS et al., 1992;MICHOR et al., 1996). Pressure changes the solvating power of a supercritical fluid and several groups found that pressure changes the selectivity of a lipet al. (1995) found that the ase. IKUSHIMA enantioselectivity of a CRL-catalyzed acetyla-
tion of (*)-citronello1 in SCC02 varied with pressure and suggested that pressure changes may change the conformation of the lipase. On the other hand, RANTAKYLAE and AALTONEN (1994) found no changes in enantioselectivity for the RML-catalyzed esterification of ibuprofen with n-propanol. CHAUDHARY et al. (1995) controlled the molecular weight of polyester formed in a PPL-catalyzed transesterification of 1,Cbutanediol and bis(2,2,2-trichloroethy1)adipate by changing the pressure of supercritical fluoroform.As the pressure increased, supercritical fluoroform dissolved longer polymer chains and the molecular weight of the product increased. Batch supercritical reactors allow analysis only at the end of the reaction. To monitor while the reaction is in progress, MARTYet al. (1990, 1992) used a reactor with a sampling loop and a saphire window for visual monitoret ing. To avoid taking samples,BORNSCHEUER al. (1996) monitored formation of acetaldehyde in the acylation of a 3-hydroxy ester with vinyl acetate through a high-pressure flowthrough cell at 320 nm (Fig. 10). The on-line data agreed with off-line values up to 60% conversion.
Fig. 10, Schematic diagram of a supercritical COzreactor with a high pressure flow through cell for online et al., 19%). measurement of the formation of acetaldehyde (BORNSCHEUER
3 Enantioselective Reactions
3 Enantioselective Reactions 3.1 Kinetic Resolutions
3.1.1 Quantitative Analysis In a kinetic resolution, the enantiomeric purity of the product and starting material varies as the reaction proceeds (reviewed by KAGANand FIAUD,1988). Thus, comparing enantiomeric purities for two kinetic resolutions is meaningful only at the same extent of conversion. To more conveniently compare kinetic resolutions, CHARLES SIH’Sgroup developed equations to calculate their inherent enantioselectivity (CHENet al., 1982, 1987; review by SIHand Wu, 1989).This enantioselectivity, called the enantiomeric ratio, E, measures the ability of the enzyme to distinguish between enantiomers. A nonselective reaction has an E of 1, while resolutions with E > 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
3 Enantioselective Reactions
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
3 Biotransformations with Lipases
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
3 Enantioselective Reactions
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 Enantioselective Reactions
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
3 Enantioselective Reactions
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
3 Biotransformations with Lipases
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.
3 Enantioselective Reactions
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 Biotransformations with Lipases
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
3 Enantioselective Reactions
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
3 Biotransformations with Lipases
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.
3 Enantioselective Reactions
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
3 Biotransformations with Lipases
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
3 Enantioselective Reactions
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<
E
SiMe3
56-61 PCL, E >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
3 Biotransformations with Lipases
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.
~
~
3 Enantioselective Reactions
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
3 Biotransformations with Lipases
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
3 Biotransformations with Lipases
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
3 Biotransformations with Lipases
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
3 Enantioselective Reactions
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
-
3 Biotransformations with Lipases
meso and prochiral primary alcohols
U
u
(OH
HO\-
U
HO\-
---
H0 ,
(OH
--.
M
H0+JO <)
PCL, 98% ee, 82% yield PCL, 95% ee, 54% yield isopropenyl acetate vinyl acetate Tsuji et al. (1989) ChCnevert 81Desjardins (1996)
U
Ho\
OEt
' "'0 PCL. 86% de, 87% yield vinyl acetate Hatakeyama eta/. (1994)
HO-Ar
HO&\oEt
phfio&OR 91% ee, 75% yield, PCL R = acyl; hydrolysis of diester Breitgoff et a/. (1986)
PCL, 98% ee, 91% yield vinyl acetate Ohsawa et a/. (1993)
R = H; acylation of diol Wang et a/. (1988), Terao et a/. (1988), Baba et a/. (1990b), Witzeta/. (1992)
HO\R L O A c PCL, hydrolysis of diacetate R = Me, >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
3 Enantioselective Reactions
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,S<\ 0 0 F R = Me. E = 7-18 R = n-Pr, CH2CH=CH2, E = 4-6 CAL-B, 98% ee, 71% yield (E -20) R = CH2Ph, E = 949 vinyl acetate CAL-B, vinyl acetate HLL favors opposite enantiomer Gais 8 von der Weiden (1996) Saksena et a/. (1995)
CRL, 95% ee, high yield CRL, 95% ee, 70% yield vinyl acetate vinyl acetate Sat0 eta/. (1992) Burgess 8 Henderson (1991) HO\
---
U
CRL. E = 10 vinyl acetate Nair 8 Anilkurnar (1996)
O ,H
--.
w
OTBDMS CRL. 97% ee, 94% yield vinyl acetate Chenevert & Courchesne (1995) Rhizornucor lipases
94% ee, 93% yield RML (Amano MAP) hydrolysis of dibutyrate Witz et a/. (1992)
HQ
E = 12-53,RML E high, RJL acylation w/ 2-phenyloxazolin-5-one hydrolysis of dibutyrate Bevinakatti 8 Newadkar (1993) Esterrnann et a/, (1990)
Rhizopus lipases
Aspergi//usniger lipase
?
Rhizopus sp. lipase, E >50 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
3 Biotransformations with Lipases
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.
3 Enantioselective Reactions
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
3 Enantioselective Reactions
-
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
3 Biotransformations with Lipases
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.
3 Enantioselective Reactions
. 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
3 Biotransformations with Lipases
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
3 Enantioselective Reactions
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
3 Biotransformations with Lipases
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
3 Biotransformations with Lipases
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.
3 Enantioselective Reactions
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
3 Biotransforrnations with Lipases
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 Enantioselective Reactions
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
3 Biotransformations with Lipases
( 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
3 Enantioselective Reactions
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.
3 Enantioselective Reactions
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 Biotransformations with Lipases
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.
3 Enantioselective Reactions
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
3 Biotransformations with Lipases
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
3 Enantioselective Reactions
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
3 Enantioselective Reactions
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
3 Biotransformations with Lipases
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
3 Biotransforrnations with Lipases
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
'*
3 Biotransformations with Lipases
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
4 Chemo- and Regioselective Reactions
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
3 Biotransformations with Lipases
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.
4 Chemo- and Regioselective Reactions
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
3 Biotransformations with Lipases
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.
4 Chemo- and Regioselective Reactions
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 Chemo- and Regioselective Reactions
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
3 Biotransformations with Lipases
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
3 Biotransformations with Lipases
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.
4 Chemo- and Regioselective Reactions
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-
4 Chemo- and Regioselective Reactions
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
3 Biotransformations with Lipases
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.
4 Chemo- and Regioselective Reactions
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 con- tion 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
3 Biotransformations with Lipases
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.
4 Chemo- and Regioselective Reactions
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 <1% glycerol. Pure MAGs in polar solvents. To suppress acyl migration, have better emulsifying properties than mixed researchers use aprotic solvents such as ke- acylglycerols (NAGAOand KITO,1990). Yearly tones, ether, or toluene (SJURSNES and AN- European production is 28,000 metric tons of
palm oil triolein tricaprin cod liver oil tripalmitin trilaurin triolein palm oil beef tallow triolein triolein palmitic acid oleic acid lauric acid lauric acid oleic acid oleic acid vinyl laurate 17-OH stearic acid oleic acid lauric acid ethyl oleate palmitic acid vinyl palmitate
water ethanol butanol isopropanol ethanol ethanol water glycerol glycerol glycerol glycerol glycerol glycerol glycerol glycerol glycerol IPG glycerol glycerollPBA glycerol glycerol glycerol glycerol IPG
1 1 1 1 1 1 1 2 2 2 2 3 3 3 3 3 3 3 3 3 3 3 3 3
reverse micelles ethanol butanol/two phase isopropanol MTBE ethanol phosphate buffer reverse micelles precipitatelno solvent reverse micelles precipitatelno solvent precipitatelno solvent molecular sievelno solvent reverse micelles reverse micelles molecular sieve/hexane no solvent MTBE hexane reverse rnicelles reverse micelles acetone hexane pentane
Reaction System (26) 78d (28) 84' (29) 87d (25) 75' (32) 97d (25) 7Y' (22) 67' (30) 30 (72) 72 (50) 50 (96) 96 (31) 95 (25) 74 (18) 55 (20) 62 (24) 72 (26) 80 (30) 90 (28) 84 (14) 42 (4) 11 (22) 68 (20) 61 (31) 95 ROL PFL HLL Ps. sp. L ROL' PCL PPL ROL' PFL CVL CVL PcamL PcarnL CRL ROL PcamL RML RML RML RML PCL CAL-B ROL PCL
(1988) HOLMBERG and OSTERBERG ZAKS and GROSS(1 990a) MAZURet al. (1991) Zuvi and WARD(1993) MILLQVISTet al. (1994) MILLQVIST-FUREBY et al. (1997) PLOUet al. (1996) HOLMBERG et al. (1989a) MCNEILL et al. (1990) CHANG et al. (1991) BORNSCHEUER and YAMANE (1994) WEISS(1990) YAMAGUCHI and MASE(1991) HAYESand GULARI (1991) (1994) HAYESand GULARI AKOHet al. (1992) PECNIK and KNEZ(1992) BERGERand SCHNEIDER (1992) STEFFEN et al. (1992) SINGH et al. (1994a) BORNSCHEUER et al. (1994b) PASTOR et al. (1995a) KWONet al. (1995) BORNSCHEUER and YAMANE (1995)
a Abbreviations: MTBE: methyl t-butyl ether; DIPE: diisopropyl ether; IPG: isopropylidene glycerol; PBA: phenyl boronic acid; for lipase abbreviations see Tab. 1. Method 1: hydrolysis or alcoholysis of triacylglycerides to 2-MAGS; method 2: glycerolysis of triacylglycerides yielding mixtures of l(3)MAGS and 2-MAGS; method 3: esterification or transesterification of glycerol with fatty acids or esters yielding l(3)-MAGS. Overall yields of MAG relative to glycerolysis (per mol TAG), hydrolysis (per mol TAG), or esterification (per mol glycerol) are given in brackets. 2-MAG. No clear data, probably mixture of l(3) and 2-MAG.'Authors used lipase from Rhizopus arrhizus which has also been reclassified as Rhizopus oryzae. SeeTab. 1 for other Rhizopus species also included in R. oryzae.
Acyl Donor
Acyl Acceptor
Method"
Tab. 15. Lipase-Catalyzed Syntheses of Monoacylglycerides (MAG)"
4 Chemo- and Regioselective Reactions
141
technical MAG and 42000 metric tons of pure 4.2.1.3.2 Glycerolysis of MAG. Researchers have developed three lipase- Triglycerides to l(3)-MAGS catalyzed routes to MAGS (1) hydrolysis or alcoholysis of triacylglycerides, (2) glycerolysis A disadvantage of the method above is the of triacylglycerides, (3) esterification or trans- waste of the two fatty acids from the triacylesterification of glycerol with fatty acids or glyceride. A more efficient approach is to use esters (Tab. 15) (reviewed by BORNSCHEUER,glycerol as the alcohol thereby using all three 1995). The first method yields 2-MAGs, while fatty acids (Eq. 30) (YAMANE et al., 1986). One methods two and three usually yield an equi- can use nonselective lipases for this reaction librium mixture of MAGs, from which the pre- because even a 1,3-selective lipase yields one dominant 1(3)-MAGs can be isolated in good 2-MAG and two 1(3)-MAGs. In practice, reacyield. tion conditions usually promote acyl migration so that an equilibrium mixture of 9 : l l(3)MAG and 2-MAG is formed. Glycerolysis of triacylglycerides in the liquid 4.2.1.3.1 Hydrolysis or Alcoholysis phase typically yields only 30-50% MAG, due to an insufficiently favorable equilibrium. To of Triglycerides to 2-MAGS shift the equilibrium, MCNEILLet al. (1990) crystallized the MAG. First, the glycerolysis Hydrolysis or alcoholysis of triacylglycer- was carried out in a liquid-liquid emulsion of ides catalyzed by a 1,3-selective lipase yields glycerol and triacylglyceride, then cooled to 2-MAG (Eq. 29; Tab. 15). Hydrolysis gave crystallize l(3)-MAG. Yields with this method moderate yields of 2-MAGS (78%) (HOLM- increased to 70-99% using different reactions (MCNEILL et al., 1990b;MCNEILLand YAMANE, BERG and OSTERBERG, 1988). A C ~migration I and YAMANE, 1994; FERprobably limited the yield by forming l(3)- 1991; BORNSCHEUER and FONSECA, 1993,1995; MYRNES MAGs which underwent further hydrolysis. REIRA-DIAS On the other hand, alcoholysis can be carried et al., 1995). This crystallization also increases out in nonpolar solvents where acyl migration the relative amount of l(3)-MAG over 2is slower. For this reason, alcoholysis of triacyl- MAG. Reaction temperature is critical and glycerides gave higher yields (75-97%) than must be kept just below the melting temperahydrolysis. In addition alcoholysis reactions ture of the monoacylglycerides, e.g., for beef are often faster because there is no change in tallow, optimum temperature was 42 "C yieldpH during the reaction and less inhibition of ing 72% l(3)-MAG using PFL or CVL (Mcthe lipase by free fatty acids. Addition of ex- NEILLet al., 1990). In continuous processes, recess alcohol shifts the equilibrium toward searchers used membranes or off-line extracMAG formation (MILLQVIST et al., 1994;MILL- tion to remove the MAG in place of crystalet QVIST-FUREBY et al., 1997; ZAKSand GROSS, lization (KOIZUMIet al., 1987; STEVENSON al., 1993; CHANG et al., 1991; GANCET, 1990). 1990a;ZUYIand WARD,1993).
OC(0)R LOC(0)R + 2 ROH L ~ ~ triglyceride
OC(0)R Eoc(o)R OC(0)R triglyceride
( water ~ or) alcohol
1,3-se1ectivelipase ~
lipase +
2 E z l OH glycerol
OH F O C ( 0 ) R + 2 RC(0)OR LOH fatty acid 2-MAG or ester
OC(0)R 3 [OH OH 1(3)-MAG
+ 2-MAG
(29)
142
3 Biotransformations with Lipases
4.2.1.3.3 Esterification of Glycerol with Fatty Acids or Fatty Acid Esters Yielding l(3)-MAGS
added a separate cold compartment where the l(3)-MAG crystallized. This reaction system gave excellent yields ( >75%, often 90% ) and high regioisomeric purity ( 95%). WEISS (1990) increased the surface area by suspendEsterification of glycerol with a fatty acid or ing solid fatty acid or ester in glycerol, but this a fatty acid ester also yields MAGs without required large excesses of glycerol. Another way to increase the solubility of wasting fatty acids (Eq. 31). Reaction mixtures contain polar protic reactants that promote glycerol and avoid subsequent acylation of the acyl migration, so esterification yields an equi- MAGs is to protect the hydroxyls in glycerol as librium mixture of MAGs.To shift the reaction the acetonide (Eq. 32) (WANGet al., 1988; toward MAG formation, researchers removed OMARet al., 1989; PECNIKand KNEZ,1992; water or alcohol using vacuum or molecular AKOH,1993; HESSet al., 1995; BORNSCHEUER Sieves (MILLER,1988;GANCET, 1990;YAMAGU- and YAMANE, 1995). Both yields and isomeric CHI and MASE,1991; ERGAN et al., 1990; KIM purities were close to loo%, but this method and RHEE, 1991; MILLQVIST-FUREBY et al., required extra chemical steps. In principle this 1996a). route can also yield enantiomerically pure
glycerol
fatty acid or ester
'o,f(i)-1.2-Oisopropylidene glycerol
fatly acid or ester
1(3)-MAG
co+
-
-
water or alcohol
wateror alcohol
The low solubility of the glycerol in non- MAGs, but the enantioselectivity of lipases is polar organic solvents slows the reaction and too low (see Sect. 3.3.2.1). promotes a side reaction - further acylation of In a similar approach, researchers used phethe more soluble MAGs to DAGs.To minimize nylboronic acid (PBA) to dissolve and protect this problem researchers used different sol- glycerol in an esterification with uncommon fatvents (JANSSEN et al., 1993a;KWONet al., 1995; ty acids such as (S)-17-hydroxystearic acid (STEFPASTOR et al., 1995a, b; AKOHet al., 1992), re- FEN et al., 1992,1995;MULTZ~CH et al., 1994). verse micelles (BORNSCHEUER et al., 1994b; SINGHet al., 1994a, b; HAYESand GULARI, 1991, 1992, 1994; FLETCHER et al., 1987), solvent-free reactions (YAMAGUCHI and MASE, 4.2.2 Fatty Acid Selectivity 1991; WEISS,1990), or hollow-fiber membrane reactors (VAN DER PADTet al., 1992; YAMANE 4.2.2.1 Saturated Fatty Acids et al., 1983; HOQet al., 1985). As expected, polar solvents dissolve more Most lipases show little preference for difglycerol and favor formation of MAGs, while ferent saturated fatty acids. They catalyze hynonpolar solvents dissolve less glycerol and drolysis or esterification of fatty acids with favor formation of DAGs. BERGER and chain lengths between C2 and C18 with similar SCHNEIDER (1992,1993)dispersed the glycerol rates (Tab. 16). Although each lipase shows on silica gel to increase the surface area and some fatty acids selectivity, this selectivity is
4 Chemo- and RegioselectiveReactions
143
Tab. 16. Fatty Acid Selectivitiesof Lipases
Lipase
Selectivity
References
CAL-B, CRL, HLL, PCL. PPL, RML
little (usually ~ 4 ) "
ProqL ROL GCL
S, MBL fatty acids M to L fatty acids discriminates against erucic acid (cisAl3 C22) discriminates againsr polyunsaturated fatty acids discriminates against fatty acids containg a cisA4 or cisA6 bond selective for cisA9 double bonds
Amano Enzyme Company RANGHEARD et al. (1989) BERGER and SCHNElDER (1991b) Amano Enzyme Company Amano Enzyme Company SONNET et al. (1993)
RML Lipase from Brassica napus (rapeseed) GCL
PEDERSEN and HOLMER (1995) HILLSet al. (1990a) JENSEN (1974);BAILLARGEON and MCCARTHY (1991); CHARTON and MACRAE (1993)
a Within a factor of four for C4, C6, C8. C10, C12. C14, C16, C18. CRL is selective for C4 by a factor of 4.7 over C6. RML did not react with C2.
small (usually less than a factor of 4). Of the lipases that do show significant selectivity, ProqL favors small to medium chain length fatty acids, while ROL favors medium to long chain fatty acids. Like other selectivities,fatty acid selectivity is best measured not by separate rate measurements of pure substrates, but by competition experiments where the lipase chooses among different substrates (RANGHEARD et al., 1989; BERGER and SCHNEIDER, 1991b).Fatty acid selectivities change slightly with reaction conditions (ADLERCREUTZ, 1994),most likely due to changes in the solvation energies of the fatty acids (JANSSEN and HALLING, 1994).Using sitedirected mutagenesis KLEINet al. (1997) created an ROL mutant with high selectivity and activity toward tributyrin over tricaprylin (80fold) and triolein ( > 80-fold). Several other mutants, also predicted to have increased selectivity toward short chain fatty acids, showed only modest changes in selectivity and sometimes significant decreases in activity (JOERGER and HAAS,1994;ATOMIet al., 1996).
4.2.2.2 Unsaturated Fatty Acids Lipases show much higher selectivity toward unsaturated fatty acids. For example, GCL favors fatty acids having a cisA9 bond by
a factor of 20 or more, 100 to 1 for oleic vs. stearic acids (JENSEN, 1974; BAILLARGEON and MCCARTHY,1991; CHARTONand MACRAE, 1993). Nippon Oils & Fats produces ultrapure unsaturated fatty acids by a GCL-catalyzed process. In another example, lipase from the seeds of Brassica nupus (rapeseed or canola plant) discriminates against fatty acids containg a cisA4 or cisA6 bond by a factor of 14 to 50 (HILLSet al., 1990a). Researchers used this discrimination to enrich seed oils in y-linoleic acid (HILLS et al., 1989,1990b). CRL discriminatesagainst erucic acid (cisAl3 C22) and y-linoleic acid (cisA9, cisAl2 C18) (ERGANet al., 1991; TRANIet al., 1992). In esterification with lauryl alcohol, RDL discriminates against DHA, but in esterification with small alcohols (e.g., ethanol), RDL showed little discrimination (SHIMADAet a]., 1997). Several lipases moderately favored polyunsaturated fatty acids (PUFAs) over saturated fatty acids, but no quantitative data are available. The usefulness, biotransformations, and biotechnological applications of PUFA have been reviewed recently (GILLand VALIVETY, 1997a, b). Fish oils contain up to 30% polyunsaturated fatty acids mainly in form of triacylglycerols. Researchers have used lipases to enrich oils in PUFAs. For example, YADWAD et al.
144
3 Biotransformations with Lipases
(1991) used a ROL-catalyzed glycerolysis of cod liver oil (9.6% PUFA) to yield 1(3)-monoacylglycerideswith 29% PUFA (see also ZAKS and GROSS,1990b). Hydrolysis of fish oils with CRL or GCL enriched the fatty acid product in PUFAs to 3 0 4 5 % . Unilever has tested this process at the pilot scale (MCNEILL et al., 1996; MOOREand MCNEILL,1996). PEDERSON and HOLMER(1995) reported that RML favored monounsaturated fatty acids (20: ln-9,22: ln9) over EPA or DHA. The incorporation of PUFAs into triacylglycerides is mentioned above in Sect. 4.2.1.1.3. Lipases CAL-B, RML, and PPL catalyze hydrolysis and transesterification reactions of the C18 furanoid fatty acid in Fig. 107 (LIEKENJIE and SYED-RAHMATULLAH, 1995).
droxydecanoic acid to the corresponding polyester. At 55 "C in the presence of 3 A molecular sieves, O'HAGANand ZAIDI(1994b) obtained a polyester with a molecular weight of 9000 (50 repeat units) (Eq. 33).
-
In the simple condensation of a diacid with a diol, OKUMURA et al. (1984) reported ANLcatalyzed formation of pentamers and heptamers. BINNSet al. (1993) used a two-stage reaction to condense adipic acid with butanel,.l-diol. The first condensation formed oligomers, after isolation these oligomers were coupled in the second condensation to form a polyester with an average of 20 repeat units. Most researchers used transesterification reactions to form the ester link. Early reports Fig. 107. Unusual fatty acid accepted by several showed only oligomers such as pentamers or lipases. hexamers (MARGOLIN et al., 1987; GUTMAN et al., 1987;GERESHand GILBOA, 1990,1991;KNANI and KOHN,1993; PARK et al., 1994; CHAUD4.3 Oligomerization and HARY et al., 1995) even when using activated esters such as 2,2,2-trichloroethyl. However, Polymerizations WALLACE and MORROW(1989a, b) obtained Lipases can catalyze polymerizations of polyesters with degrees of polymerization up to polyesters by catalyzing formation of ester 25 by using highly purified monomers. In addilinks (for reviews see GUTMAN, 1990; KOBA- tion, they used exactly two moles of diester for YASHI et al., 1994; LINKO and SEPPALLA,1996). each mole of diol because only one enantiomer The three main approaches are (1) simple con- of the diester reacted (Fig. 108). In later work MORROWsuggested that the densation of diacids with diols or hydroxy acids with themselves, (2) transesterification of release of the alcohol, even a poorly nucleoeither hydroxy esters or diesters with diols, and philic alcohol like 2,2,2-trichloroethanol, limits (3) ring-opening polymerization of lactones. the molecular weight of the polymer. Release The second two approaches are more com- of alcohol may also promote desorption of wamon. The most useful lipases are CRL, PCL, ter from the enzyme which also limits the moPPL, and RML. The potential advantages of lecular weight by permitting hydrolysis of eilipase-catalyzed polymerizations are their ther the starting diester or the product polyeset al., 1995). To minimize this stereoselectivity and narrow range of molecu- ter (BRAZWELL lar weights. Until now most polymerizations problem, researchers carried out polymerizayielded polymers of too low molecular weight tions under vacuum in high boiling solvent to et al., (usually 1000-7000), but the most recent re- remove the released alcohol (BRAZWELL ports include examples of high molecular 1995; LINKOet al., 1995a, b). For example, a PPL-catalyzed transesterification of bis(2,2,2weight (40 000). Only a few groups reported simple conden- trifluoroethy1)glutarate with 1,4-butanediol sation reactions. Either PCL (AJIMAet al., reached a molecular weight of 39000 (>200 1985) or CRL (O'HAGANand ZAIDI,1993, repeat units) under vacuum, while only 2900 1994b) catalyzed the condensation of 10-hy- without vacuum (Fig. 109).
-
145
4 Cherno- and Regioselective Reactions
(*)-trans
I
HO-OH
+
>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
3 Biotransformations with Lipases
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
5 Commercial Applications and Future Directions
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
3 Biotransformations with Lipases
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.
5 Commercial Applications and Future Directions
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
3 Biotransformations with Lipases
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.
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ical methods. 3. Optical purity enhancement in ethers of 3-methoxy-l,2-propanediol,Tetrahedron:Asymmetry 4,2265-2274. enzymic asymmetric catalysis, J. Am. Chem. SOC. WAGEGG, T., ENZELBERGER, M. M., BORNSCHEUER, 106,3695-3696. J. J., MOMONGAN, M., BERGU.T., SCHMID, R. D. (in press),The use of methoxy WANG,Y.F., LALONDE, BREITER, D. E., WONG,C.-H. (1988). Lipase-cataacetoxy esters significantly enhances reaction lyzed irreversible transesterifications using enol rates in the lipase-catalyzed preparation of optiesters as acylating reagents: preparative enantiocal pure 1-(4-chloro-phenyl) ethyl amines, J. Bioand regioselective syntheses of alcohols, glycerol technol. derivatives, sugars, and organometallics, J. Am. WALDE,P., HAN,D., LUISI,P. L. (1993). SpectroscopChem. SOC.110,7200-7205. ic and kinetic studies of lipases solubilized in reWANG,Y.-F., CHEN,S.-T., LIU,K. K.-C.. WONG,C.-H. verse micelles, Biochemistry 32,40294034. (1989), Lipase-catalyzed irreversible transesterWALDINGER, C., SCHNEIDER, M., BO-ITA, M., Coification using enol esters: Resolution of cyanoRELLI, F., SUMMA,V. (1996),Aryl propargylic alcohydrins and syntheses of ethyl R-2-hydroxy-4hols of high enantiomeric purity via lipase-cataphenylbutyrate and (S)-propranolol, Tetrahedron lyzed resolutions, Tetrahedron: Asymmetry 7, Lett. 30,1917-1920. 1485-1488. F., ZHANG, H., NAEGELE,E. (1995), Synthesis of WANG,X. Q., WANG,C. S.,TANG,J., DYDA, WALDMANN, X. J. C. (1997), The crystal structure of bovine the palmitoylated and farnesylated C-terminal bile salt activated lipase - insights into the bile lipohexapeptide of the human N-ras protein by employing an enzymically removable urethane salt activation mechanism, Structure 5,1209-1218. protecting group, Angew. Chem. (Int. Edn. Engl.) WARD,R. S. (1995), Dynamic kinetic resolution, Tetrahedron:Asymmetry 6,1475-1490. 34.2259-2262. N., SUGAI,T.,OHTA,H. (1992), PreparaWALDMANN, H., SEBASTIAN, D. (1994), Enzymic pro- WATANABE, tion of enantiomerically enriched compounds ustecting group techniques, Chem. Rev.94,911-937. ing enzymes. Part 17. Enzymatic preparation of WALDMANN. H.. BRAUN,P., KUNZ,H. (1991), New glycerol-related chiral pool possessing rerf-alkoxy enzymic protecting group techniques for the congroup, Chem. Lett., 657460. struction of peptides and glycopeptides, Biomed. Biochim. Acta 50, S243-S248. WEBER,H. K., STECHER,H., FABER, K. (1995a). WALLACE, Some properties of commercially available crude J. S., MORROW, C. J. (1989a), Biocatalytic lipase preparations, in: Preparative Biotransforsynthesis of polymers. Synthesis of an optically mations (RoBERTs,S.M.,Ed.),pp.5:2.1-5:2.10. active, epoxy-substituted polyester by lipase-cataH., FABER,K. (1995b), Senlyzed polymerization, J. Polym. Sci: Part A: Polym. WEBER,H. K., STECHER, sitivity of microbial lipases to acetaldehyde Chem. 27,2553-2567. WALLACE, formed by acyl-transfer reactions from vinyl esJ. S., MORROW, C. J. (1989b), Biocatalytic ters, Biotechnol. Left.17,803-808. synthesis of polymers. 11. Preparation of (AABB]x polyesters by porcine pancreatic lipase-cat- WEHRLI, H. P., POMERANZ,~. (1969), Synthesis of galactosyl glycerides and related lipids, Chem. Phys. alyzed polymerization, J. Polym. Sci: Part A: Lipids 3,357-370. Polym. Chem. 27,3271-3284. I., ADLERCREUTZ, P., MATWALLACE, J. S., REDA,K. B., WILLIAMS, M. E., MOR- WEHTJE,E., SVENSSON, TIASSON, B. (1993). Continuous control of water ROW,C. J. (1990), Resolution of a chiral ester by lipase-catalyzed transesterification with polyactivity during biocatalysis in organic media, Biotechnol. Tech. 7,873-878. (ethylene glycol) in organic media, J. Org. Chem. WEHTJE,E., KAUR,J., ADLERCREUTZ, P., CHAND,S., 55,3544-3546. MAITIASSON, B. (1997). Water activity control in WALLACE,J. S., BALDWIN.B. W., MORROW,C. J. enzymatic esterification processes, Enzyme Mi(1992), Separation of remote diol and trio1 stereoisomers by enzyme-catalyzed esterification in orcrob. Technol. 21,502-510. ganic media or hydrolysis in aqueous media, J. WEISS,A. (1990), Enzymic preparation of solid fatty acid monoglycerides, Fat Sci. Technol. 92, Org. Chem. 57.5231-5239. WALTS,A.E., Fox, E. M. (1990), A lipase fraction for 392-396. A. N. E., KAZLAUSKAS, R. J. (1995), resolution of glycidyl esters to high enantiomeric WEISSFLOCH, Enantiopreference of lipase from Pseudomonas excess, US Patent 4923810 to Genzyme (Chem. cepacia toward primary alcohols, J. Org. Chem. Abstr. 113: 113879). 60,6959-6969. WANG,Y. F., WONG,C.-H. (1988). Lipase-catalyzed Z. (1992), A suggestion to the PPL active irreversible transesterification for preparative WIMMER, site model dilemma, Tetrahedron 48,8431-8436. synthesis of chiral glycerol derivatives, J. Org. Chem. 53,3127-3129. WINKLER, F. K., D’ARcY,A., HUNZIKER, W. (1990), WANG,Y.-F., CHEN,C.-S., GIRDAUKAS, G., SIH,C. J. Structure of human pancreatic lipase, Nature 343, (1984). Bifunctional chiral synthons via biochem771-774.
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WIRZ,B., SPURR, F! (1995). Enantio- and regioselec- Xu, J. H., KAWAMOTO, T., TANAKA, A. (1995), Hightive monohydrolysis of diethyl2-ethoxysuccinate, performance continuous operation for enantioselective esterification of menthol by use of acid anTetrahedron:Asymmetry 6,669-670. WIRZ, B., WALTHER, W. (1992), Enzymic preparation hydride and free lipase in organic solvent, Appl. of chiral 3-(hydroxymethy1)piperidine derivaMicrobiol. Biotechnol. 43,639-643. tives, Tetrahedron:Asymmetry 3,1049-1054. Xu, J., GROSS,R. A,, KAPLAN,D. L., SWIFT,G. WIRZ,B.,SCHMID, R.,FORICHER, J. (1992),Asymmet(1996), Chemoenzymatic synthesis and study of ric enzymatic hydrolysis of prochiral2-0-allylglypoly(a-methyl-P-propiolactone) stereocopolycerol ester derivatives, Tetrahedr0n:Asymmetry 3, mers, Macromolecules 29,45824590. 137-142. YAMADA,O., OGASAWARA, K. (1995), Lipase-mediated preparation of optically pure four-carbon diJ. (1993), Facile WIRZ,B., BARNER, R., HUEBSCHER, chemoenzymatic preparation of enantiomerically and triols from a meso-precursor, Synthesis pure 2-methylglycerol derivatives as versatile (Stuttgart), 1291-1294. trifunctional C4-synthons, J. Org. Chem. 58, YADWAD,~. B., WARD,0.P.,NORONHA, L. C. (1991), 3980-3984. Application of lipase to concentrate the docosaWONG,C.-H. (1995), Enzymatic and chemo-enzyhexaenoic acid (DHA) fraction of fish oil, Biomatic synthesis of carbohydrates, Pure Appl. technol. Bioeng. 38,956-959. Chem. 67,1609-1616. YAMAGUCHI, S., MASE,T. (1991), High-yield syntheWONG,C.-H., WHITESIDES, sis of monoglyceride by mono- and diacylglycerol G. M. (1994), Enzymes in Synthetic Organic Chemistry. New York: Pergalipase from Penicillium camembertii U-150, J. Fermon Press. ment. Bioeng. 72,162-167. WOOLLEY, P., PETERSEN, KOMATSU,A., MOROE,T.(1976), OpS. B. (Eds.) (1994), Lipases: YAMAGUCHI,~., tical resolution of menthols and related comTheir Structure, Biochemistry, and Application. pounds. Part 111. Preliminary fractionation of miCambridge: Cambridge University Press. Wu, S.-H., ZHANG,L.-Q., CHEN,C.-S., GIRDAUKAS, crobial menthyl ester hydrolases and esterolysis by commercial lipases. J. Agric. Chem. SOC.Jpn. G., SIH,C. J. (1985), Bifunctional chiral synthons via biochemical methods. VII. Optically active 50,619-620. 2,2 '-dihydroxy-1,1 '-binaphthyl, Tetrahedron Letf. YAMAGUCHI, S.,TAKEUCHI, K., MASE,T.,OIKAWA, K., MCMULLEN, 26,4323-4326. T. et al. (1996), The consequences of engineering an extra disulfide bond in the PenicilWu,S. H.,Guo, Z.W., SIH,C.J. (1990),Enhancing the lium camembertii mono- and diglyceride specific enantioselectivity of Candida lipase-catalyzed ester hydrolysis via noncovalent enzyme modificalipase, Protein Eng. 9,789-795. tion, J. Am. Chem. SOC.112,1990-1995. YAMAMOTO, K., NISHIOKA,T., ODA,J., YAMAMOTO,~. Wu, S.-H., CHU,E-Y., CHANG,C.-H., WANG,K.-T. (1988), Asymmetric ring opening of cyclic acid anhydrides with lipase in organic solvents, Tetra(1991), The synthesis of D-iSOghtamine by a chemoenzymatic method, Tefrahedron Lett. 32, hedron Lett. 29,1717-1720. 3529. YAMAMOTO, Y., IWASA,M., SAWADA,S., ODA, J. Wu, D. R.,CRAMER, S. M.,BELFORT, G. (1993), Kinet(1990), Asymmetric synthesis of optically active ic resolution of racemic glycidyl butyrate using a 3-substituted Gvalerolactones using lipase in organic solvents, Agric. Biol. Chem. 54, 3269multiphase membrane enzyme reactor: experiments and model verification, Biotechnol. Bioeng. 3274. YAMANE,T. (1987), Enzyme technology for the lipids 41,979-990. industry: An engineering overview, J. Am. Oil WUNSCHE, K., SCHWANEBERG, U., BORNSCHEUER, U. Chem. SOC.64,1657-1662. T., MEYER,H. H. (1996), Chemoenzymatic route S. (1983), Continto P-blockers via 3-hydroxy esters, Tetrahedron: YAMANE,T., HOQ,M. M., SHIMIZU, Asymmetry 7,2017-2022. uous synthesis of glycerides by lipase in a microXIE,Z.-F. (1991). Pseudomonas fluorescens lipase in porous membrane bioreactor, Ann. N. I! Acad. asymmetric synthesis, Tetrahedron:Asymmetry 2, Sci. 434,558-568. 733-750. S. (1986), YAMANE,T., HOQ,M. M., ITOH,S., SHIMIZU, Glycerolysis of fat by lipase, J. Jpn. Oil Chem. SOC. XIE, Z.-F., SUEMUNE, H., SAKAI,K. (1990), Stereochemical observation on the enantioselective hy35,625-631. drolysis using Pseudomonas fluorescens lipase, YAMANE,T.,TAE KANG,S., KAWAHARA, K., KOIZUMI, Tetrahedron:Asymmetry 1,395402. Y. (1994), High-yield diacylglycerol formation by XIE,Z.-F., SUEMUNE, H., SAKAI, K. (1993), Synthesis solid-phase enzymatic glycerolysis of hydrogenated beef tallow, J. Am. Oil Chem. SOC. 71, of chiral building blocks using Pseudomonas fluorescens lipase-catalyzed asymmetric hydroly339-342. sis of meso-diacetates, Tetrahedr0n:Asymmetry 4, YAMAZAKI, Y., HOSONO,K. (1990), Facile resolution 973-980. of planar chiral organometallic alcohols with
6 References lipase in organic solvents, Tetrahedron Lett. 31, 3895-3896. YAMAZAKI, T., OHNOGI, T., KITAZUME, T. (1990), Asymmetric synthesis of both enantiomers of 2trifluoromethyl-4-aminobutyric acid, Tetrahedron:Asymmetry 1,215-218. YAMAZAKI, Y., MOROHASHI, N., HOSONO, K. (1991), Lipase-mediated homotopic and heterotopic double resolutions of a planar chiral organometallic alcohol, Biotechnol. Lett. 13,81-86. YANG,H., CAO,S. G., HAN,S. I?, FENG,Y., DING,Z.T. et al. (1995a). Optical resolution of (R,S)-2-octano1 with lipases in organic solvent, Ann. N. Y Acad. Sci. 750,250-254. YANG,F., HOENKE, C., PRINZBACH, H. (1995b), Biocatalytic resolutions in total syntheses of purpurosamine and sannamine/sporamine type building blocks of aminoglycoside antibiotics, Tetruhedron Lett. 36,5151-5154. YASUFUKU, Y., UEJI,S. (1996), Improvement (s-fold) of enantioselectivity for lipase-catalyzed esterification of a bulky substrate at 57°C in organic solvent, Biotechnol. Tech. 10,625-628. YASUFUKU, Y., UEJI,S. (1995). Effect of temperature on lipase-catalyzed esterification in organic solvent, Biotechnol. Lett. 17,1311-1316. YASUFUKU, Y., UEJI,S. (1997), High temperature-induced high enantioselectivity of lipase for esterifications of 2-phenoxypropionic acids in organic solvent, Bioorg. Chem. 25,88-99. YENNAWAR, H. I?, YENNAWAR, N. H., FARBEII, G. K. (1995), A structural explanation for enzyme memory in nonaqueous solvents, J. Am. Chem. SOC. 117,577-585. YONEZAWA. T., SAKAMOTO, Y., NOGAWA, K., YAMAZAKI, T., KITAZUME, T. (1996), Highly efficient synthetic method of optically active l,l,l-trifluoro-2-alkanols by enzymatic hydrolysis of the corresponding 2-chloroacetates, Chem. Lett., 855-856.
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Biotechnology Second, Completely Revised Edition H.-J. Rehm and G. Reed copyrightOWILEY-VCH Verlag GmbH, 1998
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
2 Pig Liver Esterase
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-
2 Pig Liver Esterase
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
2 Pig Liver Esterase
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
2 Pig Liver Esterase
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
2 Pig Liver Esterase
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
8 Proteases with Esterase Activity
-
-
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
8 Proteases with Esterase Activity
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
8 Proteases with Esterase Activity
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.
8 Proteases with Esterase Activity
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
5<.
COzEt
AcHN
MeOz
COzEt
a-Chymotrypsin
C02Et AcHN
Et02&cozH CqH
100% ee
OTs
141 30%. 98% ee
Fig. 66. HOzC-CHz-CH- - r C O z M e
I
t
COZMe 1 3 8
NHCbz
a-chymotrypsin
MeOzC-CH2-CH=
F-
COzMe 142
NHCbz
Papain COzMe
49%, 77% ee
Fig. 65.
I
rac-140
a-chymotrypsin
HO2C J J
OTs
139
136 R = AcNH 137 R = OH a-chymotrypsin
I
Et02&Co2Et
MeO2C-CHz-CH=
COzH NHCbz
Fig. 67.
229
8 Proteases with Esterase Activity
Tab. 20. Kinetic Resolution of (Z)-3-Phenylsulfinylpropenoate(143) with a-Chymotrypsin (CARDELLICCHIO et al., 1994)
I?
0
F’h0SCH=CH-C02Me
a-chymotry psin
Ph’
*.!
CW CH-CO2Me
Ph’
+
E
C1.P CI-I-COZH
143 Entry
1 2 3 4 5 6 7
Configuration Z Z Z
Z
Z Z
Z
Ester ee
Co-solvent None 10% DMSO 20% DMSO 10% t-BuOH
43 31 63 15
10% t-BuOMe 20% t-BuOMe
27 91
20% t-BuOH
such as dimethyl sulfoxide, r-butanol, and t-butyl methyl ether, were added to the reaction medium. This showed that the addition of a suitable co-solvent, e.g. r-butyl methyl ether can enhance the enantiomeric excess values for the reaction. Indeed, one of the drawbacks of using serine proteases in organic synthesis can be their limited activity in organic solvents. While it is known that serine proteases can tolerate high concentrations of hydrophilic organic solvents, e.g., aliphatic alcohols and acetonitrile, the same is not true for methanol or for dimethylformamide, where the highest tolerated concentration was found to be 50%. In view of this, CEROVSKY and JAKUBKE (1994) have prepared a more stable form of a-chymotrypsin by cross-linking the enzyme by treatment with glutaraldehyde. In subsequent peptide syntheses, the specific activity of the water-insoluble polymer was found to be comparable to the native enzyme in a medium containing more than 50% (v/v) of dimethylformamide. This has obvious implications for the use of a-chymotrypsin in esterification reactions. Some mechanistic and active site studies have been undertaken for both natural and modified a-chymotrypsin (COHEN,1969;WEST et a]., 1990).
Acid ee
1% 1
P o
1
52
45
7 72 60
73
80
65
8.3 Penicillin Acylase Penicillin acylase (EC 3.5.1.11) is a serine type of esterase which possesses both esterase and amidase activity, selectively hydrolyzing the phenyl acetyl moiety from both esters and amides. It is commercially important for its amidase activity in the hydrolysis of benzyl penicillin to 6-aminopenicillanicacid (6-APA) (Fig. 68). Although the enzyme will tolerate
Y
Benzyl penicillin Penicillin acylase
o
Fig. 68.
t
6-APA
230
4 Esterases
only minor changes in the structure of the acyl group, it exhibits a wide tolerance in the structure of the alcohol or amine portion of the molecule. This wide tolerance has meant that penicillin acylase has found some application in the kinetic resolution of primary and secondary alcohols. For example, a series of phenyl acetyl esters of substituted 2,2-dimethyl-1,3dioxolane-4-methanols (144) were incubated with penicillin acylase from E. coli, immobilized on Eupergit beads, to give the corre-
sponding alcohols in moderate to high optical purity. The compounds shown in Tab. 21, possessing a structural similarity to penicillin G, were hydrolyzed with high enantiomeric excess (FUGANTIet al., 1987,1988). Chiral2-furylcarbinols (145) were prepared with high optical purities by enantioselective hydrolysis of the corresponding phenyl acetyl esters (146) catalyzed by penicillin acylase (Tab. 22) (WALDMANN, 1989).Previously, enzymatic resolutions employing lipases or ester-
Tab. 21. Kinetic Resolution of Phenyl Acetyl Esters of Substituted 2,2-Dimethyl-1,3-dioxolane-4-methanols (144) by Penicillin Acylase (FUGANTI et al., 1987,1988)
R'
RZ
R3
Absolute Configuration
ee
H Me H H Me H H H Me
H H H H Me Me H H H
H H Me CHzOH H Me Me Et Et
4s 4s 4s, 5 s 4s, 5 s 4s. 5R 4R 4R, 5 s 4R, 5 s 4R, 5 s
60 90 50 52 65 33 46 80 90
(4,5-anti) (4,5-anti) (4S-syn) (4,5-syn) (4S-syn) (4,5-syn)
[%I
Tab. 22. Kinetic Resolution of Phenyl Acetyl Esters of 2-Furylcarbinolsby Penicillin Acylase (WALDMANN, 1989)
146
145
R
ee
Configuration of Alcohol
COzMe CN CH3
82 72 80
S
1% I
R R
8 Proteases with Esterase Activity
ases only gave the desired furan derivatives with useful enantiomeric excesses if a double resolution strategy or hydrolysis with low conversion was applied. Both of these methods resulted in low yields of products. Using penicillin acylase, gram quantities of the alcohols (145) could be prepared in 3540% yield at 60% conversion. BALDARO et al., (1993) have shown that penicillin acylase can successfully catalyze the enantioselective hydrolysis of phenyl acetyl esters of a variety of secondary alcohols which are key intermediates of pharmaceutical interest (Tab. 23). It was found that only the a-hydroxy esters gave high enantiomeric excesses at 50% conversion (entries 6 and 8) and, in a number of cases, termination of the reaction at 50% conversion resulted in alcohols of low enantiomeric excess (entries 1 4 ) . Other substrates gave alcohols of high enantiomeric excess but only when the reactions were terminated at <25% conversion (entries 5, 7, and 9). The conclusion drawn by the authors was that although resolution can occur efficiently with some substrates, the general requirements in terms of steric and polar character of the substrate, are more subtle than for other hydrolytic enzymes. In order to improve the enantioselectivity of penicillin acylase catalyzed hydrolysis of ra-
231
cemic secondary alcohols, POHLand WALD(1995) have introduced the concept of “substrate tuning”. This involves introducing a nitrogen atom into the group which is recognized by the enzyme which, in practice, involves replacing the phenyl acetyl group with a pyridyl acetyl group. The incorporation of a nitrogen atom into the group which is responsible for chiral recognition, might lead to alternative polar interactions within the active site which in turn might lead to a different binding mode with the possibility of enhanced enantioselectivity. Also, further fine tuning might be possible by utilizing 2-, 3-, or 4-pyridyl acetates. The reaction velocity for a series of pyridyl esters (146) showed that 4-pyridyl acetates were better substrates than 3-pyridyl acetates which in turn were better than 2-pyridyl acetates (Tab. 24). Comparison of the enantioselectivity between a pyridyl acetate and a phenyl acetate is interesting (Fig. 69). From the pyridyl acetate (147), (R)-1-phenylethanol was obtained with 73% ee at 40% conversion, whereas the corresponding phenyl acetate (148) gave the same alcohol with only 28% ee under the same conditions. This technique of “substrate tuning” represents one method by which the enantioselectivity of reactions catalyzed by penicillin G acylase might be enhanced. MANN
Tab. 23. Kinetic Resolution of Phenyl Acetyl Esters of Secondary Alcohols by Penicillin Acylase (BALDARO et al., 1993)
Entry 1
2 3 4 5 6 7 8 9 10
R’
R*
Et vinyl Me benzyl Et CO,Et C02Et C02Me CHZCHZCI CHZCO2Et
Me Me phenyl Me phenyl benzyl phenyl ethyl phenyl phenyl phenyl
Conversion
[%I
Configuration of Alcohol
ee [Yo 1
50 50 50
S S
26 10 40 56 94 92 94 90 > 98 36
50
12 50 25 50 18 32
R
S
R
S S S R
S
232
4 Esterases
Tab. 24. Kinetic Resolution of Pyridyl Acetyl Esters of Secondary Alcohols by Hydrolysis with Penicillin G Acylase (POHLand WALDMANN, 1995)
Penicillin G acylase OR Entry 1
Substrate
146
ROH
OR
Conversion
["/.I
mo$
ee
["/I
40
N / 2
40
3
40
4
25
5
30
6
30
A number of the synthetic applications of penicillin acylase have exploited its selectivity for the phenyl acetyl group by developing its use in selective protecting group strategy. BALDARO et al. (1988) used the phenyl acetyl group to protect both the amine and carboxyl
groups of benzylpenicillin sulfoxide (149) which subsequently underwent ring expansion to the cephalosporin nucleus (150) (Fig. 70). Penicillin acylase was used to remove both of the protecting groups in a single step, albeit in low yield. WALDMANN (1988) has shown that
8 Proteases with Esterase Activity
233
R = N 73% ee R = CH 28% ee
147 R = N 148 R = CI-I
Fig. 69.
150 15%
penicillin acylase can be successfully employed in the selective removal of the phenyl acetyl group from a number of monosaccharides (Fig. 71).
Penicillin acylase
R = Ph-7
0
4OAc
y70R ''%o
Ac& Ac
RoOAc
96%
41%
pk 42%
Fig. 71.
= R=H
..\
8.4 Papain Papain (EC 3.4.22.2) is a thio-protease which is known to exhibit a broad substrate specificity in hydrolytic reactions. In general, bulky aliphatic and aromatic substrates are preferred as substrates and this requirement is easily met in the case of amino acids by the use of common amino acid protecting groups. For example, WANGet al. (1978) have described the use of papain for the kinetic resolution of racemic N-t-butyloxycarbonyl-O-benzyl-L-serine by the enantioselective hydrolysis of the corresponding methyl ester. This afforded the L-amino acid in 72% yield plus the recovered D-amino acid in 81% yield after mild hydrolysis of the ester. SHINet al. (1991,1995) have utilized papain as a mild method for the ester hydrolysis of (151) (Fig. 72) and dehydrodipeptide esters (152) (Fig. 73). It has also been used for the regioselective hydrolysis of the N-protected adehydroglutamate diester (142) (Fig. 67) (SHIN et al., 1988). In the case of the dimethyl ester (142), regioselective hydrolysis of the a-ester was observed. This is in contrast to a-chymotrypsin which hydrolyzed the y-ester (SHINet al., 1990).
234
4 Esterases
The papain catalyzed hydrolysis of the Nprotected glycosylated amino acid ester (155) has been described where selective hydrolysis of the a-methyl ester was observed (Fig. 75) (ISHIIet al., 1990). Similarly, selective removal of the (methoxyethoxy)ethyl ester (MEE) protecting group from a-and P-glycosidic conjugates (156)and (157)has been accomplished in high yield using papain (Tabs. 25 and 26) (EBERLING et al., 1996).
Mx Papain
R= =H
Fig.72. 152
CbAAA-AAOR
Mx
R == H
Papain
Fig.73.
Papain has also been utilized in the hydrolytic resolution of unnatural amino acid derivatives (DRUECKHAMMER et al., 1988). The Nprotected furylglycine methyl esters (153)and (154),required for the synthesis of optically active synthons for alkaloids, were studied, although only compound (154)gave the ester in high enantiomeric excess (Fig. 74) Papain
%C02Me
O H H O
155
Fig.75.
*
+C02M,
NHR
NHR
153,R=BzO 1 5 4 , R =EtOCO
Mx
R= =H
Papain
-
NH R
(R)-153 , 40%ee (R)-154,>97% ee
Fig. 74. Tab. 25. Selective Hydrolysis of the (Methoxyeth0xy)ethyl Ester (MEE) Group from a-Glycosidic Conjugates (156) (EBERLING et al., 1996)
Xaa
PG
Yield
Ser Ser Thr
Aloc Fmoc 2
75 90 72
["/.I
9 References
235
Tab. 26. Selective Hydrolysis of the (Methoxyeth0xy)ethyl Ester (MEE) Group from P-Glycosidic Conjugates (157) (EBERLING et al., 1996)
Xaa
PG
R'
R2
R3
Yield
Ser Thr Ser Thr Ser Thr
Aloc Aloc Aloc Aloc Aloc Aloc
OAc OAc H H OAc OAc
H H OAc OAc H H
CH,OAc CHZOAC CH,OAc CHZOAC H H
96 30 82 87 90 90
["/.I
N. W., CROUT, D. H. G., HENDERSON, C. M., The reverse reaction, esterification, has also ALCOCK, THOMAS, S. E. (1988), Enzymatic resolution of a been reported for papain. Here, immobilized chiral organometallic ester - enantioselective hypapain was used to catalyze the esterification drolysis of 2-ethoxycarbonylbuta-1,3-dienetricarof N-protected amino acids (CHENand WANG, bony1 iron by pig-liver esterase, J. Chem. SOC., 1988). The bulky groups Cbz, Boc, and Moz Chem. Commun., 146-747. were used to protect the amino acids while ANANTHARAMAIAH, G. M., ROESKE,R. W. (1982), simple alcohols, methyl, ethyl, n-propyl, i-proResolution of a-methyl amino esters by chymopyl, n-butyl, and benzyl alcohols were used as trypsin, Tetrahedron Lett. 23,3335-3336. the nucleophiles, the reactions being carried ARITA,M., ADACHI, K., ITO, Y., SAWAI, H., OHNO, M. (1983),Enantioselective synthesis of the carbocyout in ethyl acetate. The resulting esters were clic nucleosides ( - )-aristeromycin and ( - )obtained in 17-83% yield. This papain catalyneplanocin-A by a chemicoenzymatic approach, zed method offers a mild alternative to conJ. Am. Chem. SOC.105,40494055. ventional chemical methods.
Acknowledgement I would like to thank Drs. N. B. BASHIRand J.T. SIMEof Zylepsis Ltd. for proof reading this manuscript and for their helpful comments and suggestions.
9 References ADACHI, K., KOBAYASHI, S., OHNO, M. (1986),Chiral synthons by enantioselective hydrolysis of mesodiesters with pig-liver esterase - substrate-stereoselectivity relationships, Chimia 40,311-314. M., GIRARD, C., BLOCH,R. (1989). EnzyAHMAR, matic resolution of methyl 2-alkyl-2-arylacetates, Tetrahedron Lett. 30,7053-1056.
BABA,N., MIMURA. M., ODA,J., IWASA, J. (1991), Lipase-catalyzed stereoselective hydrolysis of thiol ester, Chem. Abstr. 114,184892~. BALDARO, E., FAIARDI, D., FUGANTI, C., GRASSELLI, P., LAZZARINI,A. (1988), Phenylacetyl oxymethylene, a carboxyl protecting group removable with immobilized penicillin acylase, useful in benzyl penicillin chemistry, Tetrahedron Lett. 29, 46234624. BALDARO, E., D'ARRIGO.P., PEDROCCHI-FANTONI, G., ROSELL, C. M., SERVI,S. et al. (1993). Pen-G acylase catalyzed resolution of phenylacetate esters of secondary alcohols, Tetrahedron:Asymmetry 4,1031-1034. BARALDI, P. G., BAZZANINI, R., MANFREDINI, S., SIMONI,D.,ROBINS, M. J. (1993),Facile access to 2'0-acyl prodrugs of 1-(P-D-arabinofuranosy1)5(E)-(2-bromovinyl)uracil (BVARAU) via regioselective esterase-catalyzed hydrolysis of 2',3',5 '-triester, Tetruhedron Lett. 34,3177-3180. BARTON, P., PAGE,M. I. (1992),The resolution of racemic 1,2-diolsby the esterase-catalyzed hydroly-
236
4 Esterases
sis of the corresponding cyclic carbonate, Tetraheand 6-deoxy-6-fluorolactoside,J. Org. Chem. 57, dron 48,7731-7734. 3431-3437. BARTON, D. H. R., CL~OPHAX, J., GATEAU-OLESKER,CARDELLICCHIO, C., NASO,F.,SCILIMATI, A. (1994), A., GBRo, S. D., TACHDJIAN, C. (1993). Synthesis An efficient biocatalyzed kinetic resolution of of 3-a-alkoxy-4-/3-substituted-2-azetidinones from methyl (Z)-3-arylsulphinylpropenoates, TetraheL-(+)-tartaric acid, Tetrahedron 49,8381- 8396. dron Lett. 35,46354638. BASAK, A., BHAITACHARYA, G., NAG,A. (1993), Se- CARON, G., KAZLAUSKAS,R. J. (1991), An optimized lectivity in enzyme-catalyzed reactions - prefsequential kinetic resolution of trans-1,2-cycloerential hydrolysis of a phenolic acetate in the hexanediol, J. Org. Chem.56,7251-7256. presence of an aromatic methyl ester by pig-liver CEROVSKY, V., JAKUBKE,H.-D. (1994), Peptide-synesterase, Biotechnof. Lett. 15,469470. thesis catalyzed by cross-linked a-chymotrypsin BASAVAIAH, D., DHARMA RAO,P. ( 1994),Enzymatic in water dimethylformamide solvent system, Bioresolution of trans-2-arylcyclohexan-1-01susing catalysis 11,233-240. crude chicken liver esterase (CCLE) as biocata- CHARTON, M., ZIFFER,H. (1987), Contributions of lyst, Tetrahedron:Asymmetry 5,223-234. steric, electrical, and polarizability effects in BASAVAIAH, D., DHARMA RAO,P. (1995),Synthesis of enantioselective hydrolyses with Rhizopus nigrienantiomerically enriched anti-homoallyl alcocans - a quantitative-analysis, J. Org. Chem. 52, hals mediated by crude chicken liver esterase 2400-2403. (CCLE), Tetrahedr0n:Asymmetry 6,789-800. CHEN,S.-T.,WANG,K.-T. (1987),The synthesis of pBERGER, A., SMOLARSKY, M., KURN,N., BOSSHARD, benzyl L-aspartate and y-benzyl L-glutamate by H. R. (1973),A new method for the synthesis of enzyme-catalyzed hydrolysis, Synthesis,581-582. c@tkallyactive a-amino acids and their N-a-de- CHEN,S.-T.,WANG,K. T. (1988),Papain-catalyzed esrivatives via acylamino malonates, J. Org. Chem. terification of N-protected amino-acids,J. Chern. 38,457. SOC.,Chem. Commun.,327-328. BIANCHI, D., CESTI,P. (1990), Lipase-catalyzed ster- CHEN,C. S., FUJIMOTO, Y., SIH,C. J. (1981), Bifunceoselective thiotransesterification of mercapto tional chiral synthons via microbiological methesters,J. Org. Chem. 55,5657-5659. ods. 1. Optically-active 2,4-dirnethylglutaric acid BJORKLING, F.,BOUTEWE, J., GATENBECK, S., HULT, monomethyl esters, J. Am. ehem. SOC.103,3580K., NoRIN,T.,SZMULIK, P. (1985a), Enzyme cata3582. lyzed-hydrolysisof dialkylated propanedioic acid CHEN,C. S., FUJIMOTO, Y., GIRDAUKAS, G., SIH,C. J. diesters, chain-length dependent reversal of (1982), Quantitative analyses of biochemical kienantioselectivity,Tetrahedron 41,1347-1352. netic resolutions of enantiomers, J. Am. Chem. BJ~RKLING, F., BOUTEWE, J., GATENBECK, S., HULT, SOC.104,7294-7305. K., NORIN,T.(1985b),Enzyme-catalyzed hydroly- CHEN,S.-T.,WANG,K.-T.,WONG,C.-H. (1986). Chisis of dialkylated propanedioic acid diesters, synrally selective hydrolysis of D,L-amino acid-esters thesis of optically pure (S)-a-methylphenylalaby alkaline protease, J. Chem. SOC.,Chem. Comnine, (S)-a-methyltyrosine and (S)-a-methyl-3,4rnun., 1514-1516. dihydroxyphenylalanine, Tetrahedron Lett. 26, CHEN,S.-T., CHEN,S. Y., HSIAO, S. C., WANG,K.-T. 4957-4958. (1991). Diastereoselective hydrolysis of peptide BJORKLING, F., BOUTEWE,J., HJALMARSSON, H., esters by an alkaline protease -preparation of raHULT,K., NORIN, T. (1987). Highly enantioseleccemization free peptides, Int. J. Pept. Protein Res. tive route to (R)-proline derivatives via enzyme 37,347. catalyzed-hydrolysisof cis-N-benzyl-2,5-bismeth- CHENAULT, H. K., KIM,M.-J., AKIYAMA, A., MIYAoxy carbonylpyrrolidine in an aqueous dimethylZAWA,T.,SIMON, E. S., WHITESIDES, G. M. (1987), sulfoxide medium, J. Chem. SOC., Chern. ComEnzymatic routes to enantiomericallyenriched 1rnun., 1041-1042. butene oxide,J. Org. Chem.52,2608-2611. BLANCO,L.. GUIBE-JAMPEL, E., ROUSSEAU,G. CHENEVERT, R., MARTIN, R. (1992),Enantioselective (1988), Enzymatic resolution of racemic lactones, synthesis of (+) and (-)-cis-3-aminocyclopenTetrahedron Lett. 29,1915-1918. tane carboxylic acids by enzymatic asymmetrizaBLOCH,R., GUIBB-JAMPEL, E., GIRARD,G. (1985), tion, Tetrahedron:Asymmetry 3,199-200. Stereoselective pig-liver esterase-catalyzed hy- CLEMENT, G. E., POITER,R. (1971),Enzymatic resodrolysis of rigid bicyclic meso-diester - preparalution. An organic-biochemical laboratory extion of optically pure 4,7-epoxytetrahydrophthaperiment,J. Chem. Educ. 48,695-696. lides and hexa-hydrophthalides, Tetrahedron Lett. COHEN, S. G. (1969),Active site and stereospecificity 26,40874090. of a-chyrnotrypsin, Trans. N. Y Acad. Sci. 31, CAI,S., HAKOMORI, S.,TOYOKUNI,T. (1992),Applica705-7 19. tion of protease-catalyzed regioselective esterifi- COHEN, S. G., KHEDOURI,E. (1961a), Requirements cation in synthesis of 6 ‘-deoxy-6’-fluorolactoside for stereospecificity in hydrolysis by a-chymo-
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SHIN,C., KAKUSHO,T.,ARAI, K., SEKI,M. (1995), De- UEMURA,A., NOZAKI,K.,YAMASHITA, J.,YASUMOTO, hydrooligopeptides. 18. Enzymatic hydrolysis and M. (1989), Regioselective deprotection of 3 ' , 5 'coupling of dehydrodipeptide esters containing 0-acylated pyrimidine nucleosides by lipase and a-dehydroamino acid residue by using papain, esterase, Tetrahedron Lett. 30,3819-3820. Bull. Chem. SOC.Jpn. 68,3549-3555. WALDMANN, H. (1988). The phenylacetyl (PHAC) SICSIC,S.,IKBAL,M., LE GOFFIC,F.(1987), Chemoengroup as enzymatically removable protecting zymatic approach to carbocyclic analogs of ribofunction for peptides and carbohydrates - selecnucleosides and nicotinamide ribose, Tetrahedron tive deprotections with penicillin acylase, Liebigs Lett. 28,1887-1888. Ann. Chem., 1175-1180. SIH, C. J., GLJ,Q. M., HOLDGRUN, X., HARRIS,K. WALDMANN,H. (1989),A new access to chiral2-furyl(1992). Optically active compounds via biocatacarbinols by enantioselective hydrolysis with penlytic methods, Chirality 4.91-97. icillin acylase, Tetrahedron Lett. 30,3057-3058. SIME,J. T., POOL,C. R.,TYLER,J. W. (1987), Regiose- WEST,J. B., HENNEN, W. J., LALONDE, J. L., BIBBS,J. lective enzymic-hydrolysis in the isolation of isoA., ZHONG, Z. et al. (1990), Enzymes as synthetic mers of Mupirocin, Tetrahedron Lett. 28, 5169catalysts - mechanistic and active-site considera5172. tions of natural and modified chymotrypsin, J. SUKURALT., MARGOLIN, A. L., RUSSELL, A. J., KLIBAm. Chem. SOC.112,5313-5320. ANOV, A. M. (1988), Control of enzyme enantio- WESTERMANN, B., SCHARMANN, H. G., KARTMANN, I. selectivity by the reaction medium, J. Am. Chem. (1993), PLE-catalyzed resolution of a-substituted SOC.110,72367237. P-ketoesters, application to the synthesis of ( + )TAKAISHI, Y., YANG,Y.-L., DITULLIO, D., SIH, C. J. nitramine and ( - )-isonitratnine, Tetrahedron: (1982), Bifunctional chiral synthons via microbioAsymmetry 4,2119-2122. logical methods. 2. Optically-active a-hydroxy WILSON,W. K., BACA,S. B., BARBER, Y. J., SCALLEN,T. aldehydes. Tetrahedron Lett. 23,5489-5492. J., MORROW, C. J. (1983), Enantioselective hydrolyTANAKA, S., OHAISHI,H., SUEMUNE, H., SAKAI,K. sis of 3-hydroxy-3-methylalkanoicacid esters with (1987), at The 107th Annual Meeting, Pharmapig-liver esterase, J. Org. Chem.48,3960-3966. ceutical Society of Japan. WONG,C.-H., Ho, M.-F., WANG,K.-T. (1978), PreparTONG, J. H., PETITCLERC, C., D'LoRIo,A., BENOITON, ation of optically pure N-tert-butyloxycarbonylN. L. (1971), Can. J. Biochem. 49,877. 0-benzyl-L-serine and its antipode, J. Org. Chem. TOONE, E. J., JONES,J. B. (1991a), Enzymes in organ43,3604-3610. ic synthesis. 49. Resolutions of racemic monocy- WONG,C.-H., CHEN,S-T., HENNEN, W. J., BIBBS,J. A., clic esters with pig-liver esterase, Tetrahedron: WANG,Y.-F.et al. (1990), Enzymes in organic synAsymmetry 2,207-222. thesis - use of subtilisin and a highly stable muTOONE, E. J., JONES,J. B. (1991b), Enzymes in organic tant derived from multiple site-specific mutation, synthesis. 50. Probing the dimensions of the large J. Am. Chem. SOC.1l2,945-953. hydrophobic binding region of the active-site of WORONIECKI, S. R., ARMITAGE, I? A., ELSON,S. W.. pig-liver esterase using substituted aryl malonate FORD,B. D., SIME,J.T. (1994),A highly regioselecsubstrates, Tetrahedron:Asymmetry 2,1041-1052. tive esterase from Penicillium frequentans IMITOONE, E.J., WERTH,M. J., JONES,J. B. (1990), En92265, Biocatalysis 8,309-320. zymes in organic synthesis. 47. Active-site model ZHU,L.-M., TEDFORD, M. C. (1990), Applications of for interpreting and predicting the specificity of pig-liver esterases (ple) in asymmetric synthesis, pig-liver esterase, J. Am. Chem. SOC. l U , 4946Tetrahedron 46.658741611. 4952. ZIWER,H., KASAI,K., IMUTA,M., FROUSSIOS, C. UDDING, J. H., FRAANJE, J., GOUBITZ, K., HIEMSTRA, (1983), Microbially mediated enantioselective esH., SPECKAMP, W. N. (1993), Resolution of methyl ter hydrolyses utilizing Rhitopus nigricans - a cis-3-chloromethyl-2-tetrahydrofuran carboxynew method of assigning the absolute stereolate via enzymatic hydrolysis, Tetrahedron: chemistry of acyclic 1-arylalkanols, J. Org. Chem. Asymmetry 4,425432. 48,3017-3021.
Biotechnology Second, Completely Revised Edition H.-J. Rehm and G. Reed copyrightOWILEY-VCH Verlag GmbH, 1998
5 Cleavage and Formation of Amide Bonds
DAVID W. T. HOOPLE Sandwich, Kent, UK
1 Introduction 244 2 Acylases, Peptidases 244 2.1 Transformation of Simple a-Amino Acid Derivatives 244 2.1.1 Formation of Amides 244 2.1.2 Hydrolysis of C-Terminal Amides 245 2.1.3 Hydrolysis of N-Terminal Amides 246 2.2 Peptides 247 2.2.1 Coupling of Single a-Amino Acid Units 250 2.2.1.1 Solvent and Structural Effects 250 2.2.1.2 Regioselectivity and Unnatural Substrates 253 2.2.1.3 Modified Enzymes 255 2.2.2 Larger Peptides 256 2.3 Non a-Amino Acid Substrates 256 3 Penicillin Acylase 258 3.1 Reactions of Penicillin and Cephalosporin Substrates 258 3.2 Other Substrates 260 3.3 Protection of Amino Groups 261 4 Esterases and Lipases Which Can Be Used with Amides and Peptides 262 4.1 a-Amino Acids and Peptides 262 4.2 Other Substrates 263 5 Others 267 5.1 Hydantoinases and Carbamoylases 267 5.2 Catalytic Antibodies 269 6 References 271
244
5 Cleavage and Formation of Amide Bonds
1 Introduction
The properties and synthetic applications of amidases have recently been reviewed (DRAUZ and WALDMANN, 1995; BOESTENet al., 1995; WONGand WHITESIDES, 1994) and were also treated in Vol. 6a of the 1st Edition of the Biotechnology series (SCHMIDT-KASTNER and EGERER,1984).
This chapter covers the amidase-catalyzed formation and hydrolysis of the amide bond, with particular emphasis on recent developments. Here, the term amidase, or amide hydrolase, is used in its broadest context to encompass all hydrolases which catalyze these reactions and thus N-acylases and proteases, as well as lipases and catalytic antibodies, are also included. The efficient use of many lipases and esterases to catalyze direct amide bond formation between esters and amides (kinetic amiThe regio- and stereochemical manipulation nolysis) is an interesting synthetic development and is covered separately (Sect. 4). Of of a-amino acids by hydrolases is of prime imnote also is the now widespread use of en- portance in peptide chemistry. The whole speczymes in organic solvents which has proved trum of these hydrolytic enzymes, including particularly useful in amidase-catalyzed amide lipases and esterases (Sect. 4), has found wide or peptide bond formation, where unwanted application in the area, ranging from the resosecondary product hydrolysis is often greatly lution of single amino acids to the coupling and hydrolysis of much larger peptide fragreduced compared to aqueous systems. Enzyme Commission nomenclature distin- ments (Sect. 2.2.2). guishes between hydrolases acting on peptidic bonds (EC 3.4) and other amide bonds (EC 3.5). In 1984 all of the sub-subclasses EC 2.1 Transformation of Simple 3.4.1-10 were abandoned. Enzymes cleaving a-Amino Acid Derivatives peptide bonds (peptidases,proteases) were divided into two sets of sub-subclasses. E C This section covers reactions of single amino 3.4.11-19 covers peptidases (exopeptidases, acids not connected with the formation or hycarboxy- and aminopeptidases) which cleave drolysis of peptides. single amino acids or dipeptides from the ends of peptide chains, whereas EC 3.4.21-24 covers proteinases (endopeptidases, proteolytic 2.1.1 Formation of Amides enzymes, peptidyl-peptide hydrolases). which Most work concerned with the formation of have no preference for terminal residue cleavage. Preferred systematic names are shown in amides from single amino acids has focused bold type (BARRETTand MCDONALD, 1986). naturally enough on the coupling reactions Enzymes which cannot be allocated to a spe- with further amino acid units to form peptide cific sub-subclass are assigned as an interim bonds, and there is now a large volume of litermeasure to 3.4.99 (Anonymous, 1984). ature covering the area. In contrast, amidaseMost natural amino acids (including all those catalyzed amide formation with non-amino directly coded by mammalian DNA) have the acid-derived substrates has been much less L-configuration as specified by FISCHER.investigated. Early work showed that racemic N-acetylL-amino acids in which a side chain is the group with third priority in the Cahn-Ingold-Prelog phenylalanine could be resolved by direct assignment (NH2 > CO,H > R > H) have enantioselective enzyme-catalyzed coupling the (S)-configuration. Cysteine is exceptional with p-toluidine to yield the L-toluidide (1) because the sulfur atom elevates the priority from which pure L-phenylalanine was obof the side chain above the carboxylic group tained in 82% yield after acid hydrolysis and hence L-cysteine has the (R)-configura- (HUANGand NIEMANN, 1951). The enzyme tion. used in this case was papain, a cysteine pro-
2 Acylases, Peptidases
2 Acylases, Peptidases
tease of broad specificity.Similarly,papain was found to catalyze direct amide bond formation between Boc-protected alanine and 2-aminoethanol to give the ethanolamide (2) (CANTACUZENE and GUERREIRO, 1989). Both of these examples (Fig. 1) serve to illustrate how hydrolytic enzymes can be made to work in reverse, under thermodynamic control by choosing reaction conditions which drive the equilibrium to product formation, either by precipitation of product as in the first example, or by the use of an organic solvent with minimal water content to disfavor product hydrolysis in the second. The aminoacylase (acylase I) from porcine kidney (PKA) has been similarly used in organic solvents to resolve a-amino acids by enantioselective acylation rather than by its more usual role in amide hydrolysis (Sect. 2.1.3). Several racemic a-amino acids (Met, Ala, Leu, Ser. Val) were enantioselectively
245
acetylated in high yield and 100% optical purity in ethyl acetate solution using sodium acetate as acetate donor. Further work (with Met) showed that the water content was crucial - at 3.2% an optimum yield (90%) of N-acetyl-Lmethionine resulted which dropped sharply as the water content was increased to 5.4%. Other acyl groups (N-formyl, N-propionyl) could be introduced in the same way (YOKOIGAWA et al., 1994).
2.1.2 Hydrolysis of C-Terminal Amides
Aminopeptidases are exopeptidases whose natural function is to release single amino acids from the amino end of a peptide chain. A number of these hydrolases, most notably the aminopeptidase from Pseudomonas purida, have been widely applied to the resolution of racemic a-amino acids by enantioselective hydrolysis of C-terminal primary amides. The enzyme from Z? purida is highly specific for Lamino acid amides (3) and requires the presence of the a-hydrogen. However, it accepts a very wide structural variation of the a-substituent, ranging from simple alkyl groups (Fig. 2) (FEENSTRA et al., 1990;Roos et al., 1992) to the much larger natural product L-lupinic acid (7) (Fig. 3), a principal metabolite of the phytohormone trans-zeatine (SHADIDet al., 1990). 1 Ac-Phe-p-toluidide All of these products were formed in high ee. The enzyme can also tolerate N-substitution of Papain, pH 4.6, 4OoC, the amide, as demonstrated by the effective 7 days, 86-89%yield use of N-methoxyamides as substrates (Roos et al., 1992).In this latter example, the unreacted N-methoxy-D-amide was not isolated but instead was converted directly into the D-amino acid by in situ Schiff's base formation to aid separation from the L-amino acid, followed by acid hydrolysis. 0 Other amidases have been found to tolerate a-substitution of the chiral carbon atom. An amidase from Mycobacrerium neoaurum ATCC 2 Boc-Ala-ethanolamide 25795 gave a range of ap-disubstituted L-amino acids from the corresponding racemic priPapain (immobilized), mary amides in 78-99% ee. The highest enanCHZClz, 37OC. 18h, 78% yield tioselectivities resulted when a-methyl was Fig. 1. See text (HUANG and NIEMANN, 1951: CAN- combined with a larger a-substituent such as isopropyl or benzyl. When both a-substituents TACUZENE and GUERREIRO, 1989).
246
5 Cleavage and Formation ofAmide Bonds
H2N 0
R1
P. putida
aminopeptidase
@h0@ -^ R f I
-
H3N s
-r
HZN%~,
0
3 racamide
4 L-amino acid
R'
0
5 D-amide
R = Tr,~Bu,Ph, Bn, BnCH2, R1 = H (Feenstra et al., 1990) R = 'Pr,allyl, cyclopentyl, R' = OMe; L-amino acids 98-100%ees (Roos etal., 1992 Fig. 2. See text.
6
7
L-Lupinic acid,
45% yield, >95% ee
8
Fig. 3. Asymmetric hydrolysis of lupinic acid amide (SHADID et al., 1990).
were small, as in the a-methyl-a-ethyl analog, the (S)-acid product ee dropped to 80%. Interestingly, direct attachment of phenyl to the aposition was compatible with methyl as the second a-group (95% ee of (S)-acid) but not with allyl in place of methyl - here no reaction occurred after 10 days.To account for these results the authors propose an active site model for the amidase in which a small hydrophobic region is able to accommodate a maximum of a C-3 unit, whereas the second a-substituent can be much larger. Substitution of the amino terminus is not tolerated (KAPTEINet al., 1993). Examples of other L-selective amidases which have found application in synthesis are given in Fig. 4 -these include the amidase from
Ochrobactrum anthropi, used to resolve the primary amide of a-methyl-3,4-dichlorophenylalanine (9) (KAFTEIN et al., 1994) and Lleucine aminopeptidase, used to resolve the amide derivative of piperazine-2-carboxylic acid (lo), an important component of several drug prototypes such as HIV-protease inhibitors and NMDA antagonists (BRUCEet al., 1995).
2.1.3 Hydrolysis of N-Terminal Amides Use of acylase I from porcine kidney (PKA) and the fungus Aspergillus sp. (AA) is now a well established method for the resolution of a
2 Acylases, Peptidases
6” c1
9 ( S)-a-Methyl-3,4-dichloro-phenyl alanine, 0. anrhropi amidase, pH 5.3,40”C,15h,93%ee
Boc I
10 (S)-4-Boc-piperazine-2-carbo~lic acid, L-leucine aminopeptidase (Sigma). pH 8.2, rt, 7d, ee not reported but product converted to (S)-amide, 64%ee Fig. 4. See text (KAFTEINet al., 1994; BRUCEet al., 1995).
247
Carboxypeptidases are exopeptidases that formally catalyze the cleavage of C-terminal L-amino acid residues from peptides (see Sect. 2.2) and have also found application to these resolution reactions (Tab. 1, entries 15-17). Typically, the N-acetyl, chloroacetyl, methoxyacetyl, and trifluoroacetyl amides are used, the advantage of the substituted-acetyl derivatives being that these more electrophilic amide carbonyls are more reactive to hydrolysis and frequently give faster reactions. A further advantage is that the unreacted D-amides can be chemically deprotected more easily than the parent acetamides, thus giving ready access to enantiomerically pure D-amino acids. Alternatively, the D-acetamide can be racemized by heating with acetic anhydride/acetic acid to provide starting material for a second resolution cycle (PIRRUNGand KRISHNAMURTHY, 1993). Tab. 1 gives selected examples which illustrate the structural diversity of substrates accepted by acylase I and carboxypeptidase A. The strong stereochemical preference for amides having the bulkier CZsubstituent in the (S)-configuration is emphasized by the resolution of 2-trifluoromethylalanine (entry 12); however, the same authors report that the corresponding mono-fluoromethylalanine amide shows no enantiodiscrimination between 2substituents (KELLERand HAMILTON, 1986). Also noteworthy is the diamide substrate (entry 6) where only the a- and not the P-amide is cleaved. This a-regioselectivity is a general feature of these enzymes, as well as in many aamino acid reactions catalyzed by proteases (Sect. 2.2).
wide range of natural and unnatural a-amino acids by enantioselective hydrolysis of their N-acyl derivatives. Both enzymes are highly L-selective, yet possess a wide substrate tolerance of the amino acid side chain both in terms of steric size and in the range of functional groups accepted. This is well illustrated by the early work of GREENSTEIN and co-workers (GREENSTEIN and WINITZ,1961) and more re- 2.2 Peptides cently in an extensive study of over 50, mostly unnatural, a-amino acids by the WHITESIDES’ The protease-catalyzed formation and hygroup (CHENAULT et al., 1989). In this later drolysis of peptides have been extensively rework, the substrate specificity of both acylases viewed (e.g., BOESTENet al., 1995; WONGand 1994;GILLet al., 1996).Proteases was shown to be complementary -while both WHITESIDES, enzymes accept straight-chain alkyl substrates are divided into four sub-subclasses: serine (EC 3.4.21.X), thioproteaswith or without unsaturation,AA was more re- proteases active towards aromatic or branched alkyl es (EC 3.4.22.X), aspartyl proteases (EC amino acids than PKA. Conversely. PKA ac- 3.4.23.X), and metalloproteases (EC 3.4.24.X). cepted a-methyl-a-amino acids whereas A A The key mechanistic features of each are as follows: did not.
248
5 Cleavage and Formarion ofAmide Bonds
Tab. 1. Examples of Enzyme-Catalyzed Enantioselective Hydrolysis of Unnatural Amino Acids
Entry" R'
1 2 3 4 5 6 7 8 9 10 11 12 13 14
C&L H,C=CH H2C=CHCHz CH,(CHJio C,H,CH( Me) AcNHCH~ OH CF4CHd2-3 (Me),SiCHz eryfhro-p-MeOC6H,CH(oH) ~,~-(HO)ZC~H~ CF,
15 16 17
Me,CH PhCH2 (7-Fluoroindol-3-yl)CHz
RZ R3 H H H H H
Enzymeb Reference
Ac Ac CICH,CO ClCH2C0 ClCH2C0 H Ac H PhCH =CHCO H Ac H Ac H Ac H CICHzCO Me CF,CO (7,7-dimethylnorborn-2R-yl)CHzH Ac Cyclooctatetraenyl-CH, H Ac Me CF,CO Me CF,CO H CF3C0
PKA
-c
PKA AA AA PKA PKA PKA PKA
-c
PKA
-c
AA CP-A CP-A CP-A
CHENAULT et al. (1992a) ITAYA et al. (1994) CHENAULT et al. (1992b) MORIand OTSUKA (1985) MORIand IWASAWA (1980) BIRNBAUM et al. (1952) PINGet a1 (1992) OJIMA et al. (1989) YAMANAKA et al. (1996) INOUE et al. (1993) BAKERet al. (1995) KELLERand HAMILTON (1986) YUASAet al. (1992) PIRRUNG and KRISHNAMURTHY (1993) TURK et al. (1975) TURK el al. (1975) LEEand PHILLIPS (1991)
a All L-amino acid ee values >95%. AA, acylase I from Aspergillus sp.; CP-A, carboxypeptidase A: PKA, porcine kidney acylase I. Acylase I source not given in reference.
(1) Serine proteases -contain the catalytic triad Asp, His, Ser. Amide hydrolysis proceeds via nucleophilic attack of a serine hydroxyl group on the amide carbonyl to form a covalent acyl-enzyme intermediate with loss of the amine component. The nucleophilicity of the serine hydroxyl is enhanced by the adjacent histidine residue, which acts as a general base. Subsequent reaction of this intermediate with a water molecule yields the product acid. The serine proteases are divided by sequence homology into the chymotrypsin family (e.g., trypsin), the subtilisin family, and an undefined group which shows no sequence homology. (2) Thioproteases - sometimes called cysteine proteases. These proteases follow a similar pathway to the serine proteas-
es except that the nucleophile is a thiolate anion from the cysteine residue of the active site. Thus the acyl-enzyme is now a thioester. Common thioproteases are papain (from papaya latex), ficin (from figs), bromelain (from pineapple), cathepsin (from mammals), and bacterial peptidases such as clostripain. (3) Aspartyl proteases - so-called because a pair of aspartic acid residues are involved in the cleavage step. These act as a general base/general acid to activate a bound water molecule which attacks the amide carbonyl. Pepsin is an example used in synthesis. (4) Metalloproteases - these require a divalent metal cation, frequently zinc, which is bound to specific amino acid residues and the amide carbonyl oxygen.The attacking water molecule is
-
249
2 Acylases, Peptidases
again activated by a carboxylate anion. No acyl-enzyme intermediate is formed in this case. Several metalloproteases have been used in synthesis - these include acylases, leucine aminopeptidase, carboxypeptidase A, and thermolysin.
Amino terminus
Carboxylate terminus
L
5.4 -+, Y t R
H I N
O
R
H I
O
5
Proteases, or peptidases as they are sometimes called, can be highly specific in forming HI O R H O R or cleaving a bond to a particular amino acid or type of residue (e.g., hydrophobic, basic, or terminal) or they may have broader specificity. Acyl donor Acyl acceptor For example, trypsin normally cleaves only (nucleophile) bonds adjacent to basic residues such as argiCleavage site nine and lysine, whereas papain has much wider specificity. Exopeptidases such as carFig.5. Nomenclature for the cleavage of peptidboxypeptidase Y and aminopeptidase M cleave ases (SCHECHTER 1967). and BERGER, single amino acids from the carboxyl and amino termini, respectively, and endopeptidases act on internal bonds. A nomenclature for the residue selectivity have been reviewed (SWEENEYand WALKER, of proteases is shown in Fig. 5.The amino acid 1993). There are several advantages of using proresidue bearing the carbonyl moiety of the amide bond undergoing cleavage is designated teases in peptide synthesis: mild conditions, P, and that bearing the amine moiety Pi. Dis- freedom from racemization, minimal protectal amino acid residues are designated sequen- tion of reacting fragments, and a very high detially P2,P3, P4,etc. and P;, P;, Pi, respectively. gree of regio- and enantioselectivity. Synthesis Each of the residues that contributes to the se- can be carried out either under thermodynamlectivity is assumed to be bound to sites in the ic or kinetic control, as depicted in Fig. 6. In the enzyme designated S1,S;,etc. (SCHECHTER and thermodynamically controlled process, which BERGER,1967). Preferred cleavage sites of is the reverse of hydrolysis, the equilibrium has proteases commonly used in synthesis are list- to be moved to the right by modifying the reed in Tab. 2. The individual characteristics and action conditions to favor product formation. detailed cleavage specificities of several pro- For example, use of organic solvents with low teases commonly used in peptide synthesis water content, biphasic systems, and product
Tab. 2. Preferred Cleavage Sites of Proteases Used in Synthesis Enzyme
Catalytic Site
Sequence Cleaved (the Amino Terminus is on the Left)
a-Chymotrypsin Trypsin Subtilisin Proline Iminopeptidase Carboxypeptidase Y Papain Clostripain Pepsin Thermolysin Carboxypeptidase A
Ser Ser Ser Ser
-Trp(Phe,Tyr,Leu,Met)-Xaa
Ser CYS CYS ASP metallo metallo
-Xaa-Xaa-OH -Phe(Leu,Val)-Xaa-Xaa -ArgXaa -Phe(Glu,Leu)-Leu(Phe) -Phe(Gly,Asp,Leu)-Leu(Phe, Val) -Xaa-(Asp,Glu,Phe,Leu)-OH
-Lys(Arg)-Xaa
-Trp(Phe,Tyr,Leu,Met)-Xaa -Pro-Xaa
250
5 Cleavage and Formation ofAmide Bonds
Thermodynamic control
Kinetic control
Enz
0
RK
Enz
0
ANX Enz
R Fig. 6. Strategies for the synthesis of amides.
precipitation by careful selection of protecting groups have all been used in this way. In contrast, the kinetic aminolysis reaction proceeds via a covalent acyl-enzyme intermediate which can either be hydrolyzed to the acid by water or amidated by an added nucleophile such as an amine or second amino acid fragment.Thus, while the approach is limited to those proteases which form acyl-enzyme intermediates, this can also include many lipases and esterases which form such intermediates from esters. An obvious advantage here is the lack of amidase activity of lipases and esterases, which can avoid the potential secondary product hydrolysis inherent in aqueous protease-catalyzed couplings (Sect. 4). Manipulation of reaction conditions is still required in the protease-catalyzed kinetic approach to minimize secondary product hydrolysis. Several reviews are available (SEMENOV et al., 1988; GILLet al., 1996).
2.2.1 Coupling of Single a-Amino Acid Units Much work has been done in recent years to examine the various factors which influence enzyme-catalyzed peptide couplings. In particular, following the commercial success of the sweetener aspartame (Asp-Phe-OMe), interest has focused on the coupling of single amino
I H
acids to form di- and tripeptides. The use of a wide range of organic solvents, coupled with the comparatively recent amidation of esters (kinetic approach) has opened up many new possibilities for peptide synthesis such as the use of lipases and esterases and low water systems to avoid undesired product hydrolysis.
2.2.1.1 Solvent and Structural Effects In a low water (4%) solvent study of the thermolysin-catalyzed thermodynamically controlled synthesis of aspartame the product yield varied widely depending on the solvent used. Hindered aliphatic alcohols such as tamyl alcohol (tert-amyl alcohol) gave the best yield (71.6%0),while MeOH was much poorer (0.75%). Polar aprotic solvents like DMF and DMSO failed to yield any product whereas less polar solvents such as ethyl acetate gave moderate yields (42%) (NAGAYASU et al., 1994b). Similar trends have been noted in other systems (e.g., KISEand FUJIMOTO, 1988).The same reaction could be carried out in water alone to give a quantitative yield of product when a 4-fold excess of nucleophile (PheOMe) was used with immobilized thermolysin (ZHOUand HUANG,1993). Product precipitation occurred in this case, thus driving the equilibrium towards completion.
251
2 Acylases, Peptidases
A further important aspect of enzyme-catalyzed peptide coupling is illustrated by the large-scale process for aspartame where, because of the high stereospecificity of the enzyme for L-amino acid residues, racemic starting materials can be used to generate the desired L,L-diastereomer. After product isolation, unreacted D-starting materials are racemized and recirculated (CHEETHAM,1994). However, in the absence of the corresponding L-enantiomer, D-amino acid fragments may still be accepted as acyl acceptors by some enzymes, although such couplings are usually significantly slower. Most proteases do not tolerate D-configuration acyl donors. A detailed study of kinetically controlled peptide formation in anhydrous alcohols by the industrial protease alcalase (major component: subtilisin Carlsberg) showed that, while only L-acyl donors were accepted by the enzyme, specificity towards the acyl acceptor was much wider (CHENet al., 1992). Several D-amino acid esters or amides could be coupled in high yield (Tab. 3). The authors also examined the role of the alcohol solvent. In ethanol, transesterification of the acyl-enzyme intermediate by the solvent competes with aminolysis but not in r-butanol, leading to higher yields and shorter reaction times in the latter. Enzyme stability was also very dependent on the polarity of the alcohol, being relatively poor in Tab. 3.
Entry 1 2 3 4
5 6 7 8 9 10 11 12 13 14 a
methanol (nearly half of the activity lost after 35 min), moderate in ethanol and t-butanol (half activity remaining after approximately 5 days) and high in t-amyl alcohol (stable for several weeks).The effect of the water content on yield was dramatic - in the synthesis of the dipeptide Moz-Phe-Leu-NH,, (entry 4) the yield in anhydrous t-BuOH (<0.1% H,O) was reduced to 48% when the solvent contained 4.86% H,O. Other points of note from this study are the successful couplings of non-amino acid-derived nucleophiles such as phenyl hydrazine (entry 11) and even the poorly nucleophilic p-nitroaniline (entry s), and the high regioselectivity of the enzyme for the a-carboxyl in the unprotected diacid derivative Boc-DL-Asp-OBn (entry 11).Such regioselectivity is commonly shown by proteases in peptide coupling, and frequently side chain residues do not need to be protected. The kinetic activation of thermolysin by alcohols has recently been reported (ALAMet al., 1996). On ecological grounds, water should be the preferred solvent for kinetic peptide bond formation (as with many thermodynamically based couplings) but in practice the solubility of most protected amino acid esters is too low, requiring at least some water-miscible co-solvent to be present. One way to get round the problem is to use water-solubilizing protecting groups based on ionizable substituents and
Kinetically Controlled Peptide Bond Formation by Alcalase Acyl Donor”
Nucleophile”
Moz-Phe-OMe Ala-NH, Moz-Phe-OMe Ala-NH, Moz-Phe-OMe D-AI~-NH, Moz-Phe-OBn Leu-NH, Moz-Phe-OBn D-Leu-NH, Moz-Phe-OMe Phe-NH, Moz-Phe-OMe D-Phe-OMe Ba-Leu-OBn p-NH?-C,H, -NO, Moz-Asp(Bn)-OBn D-AI~-NH, Ba-Met-Leu-OBn Phe-NHBn roc-Ba-Asp-OBn H,NNHPh roc-Z-Ala-OBn Thr-NHNH, Z-Tyr-OMe D-Arg-OMe Z-D-Phe-OMe Ala-NH,
Solvent
Product”
Time
Yield
EtOH t-BuOH t-BuOH t-BuOH t-BuOH t-BuOH t-BuOH EtOH t-BuOH EtOH EtOH t-BuOH t-BuOH EtOH
Moz-Phe-Ala-NH, Moz-Phe-Ala-NH, Moz-Phe-D-Ala-NH, Moz-Phe-Leu-NH, Moz-Phe-D-Leu-NH, Moz-Phe-Phe-NH, Moz-Phe-D-Phe-OMe Boc-Leu-p-NH,-C,H,-N02 Moz-Asp(Bn)-D-Ala-NH, Boc-Met-Leu-Phe-NHBn Boc-Asp-NHNHPh Z-Ala-Thr-NHNH, Z-Tyr-D-Arg-OMe Z-D-Phe-Ala-NH,
4 2 2 3 2.5 3.5 3.5 12 2 6 5 2 2 12
79 86 87 94 89 82 91 25 86 85 95 65 81 0
PI
Moz,p-methoxybenzoyloxycarbonyl; Bn, benzyl; Z , benzyloxycarbonyl;Boc, t-butoxycarbonyl
[”/.I
252
5 Cleavage and Formation of Amide Bonds
these have proved very effective in some cases (FISCHERet al., 1994). For example, using maley1 protected tyrosine ethyl ester (12)purely aqueous media could be used for kinetic-based couplings at up to 1.5 mol L-' concentration in the large-scale production of kyotorphin (13) (Fig. 7). N-formyl derivatives of amino acid esters have also been found to have improved solubility relative to carbamate-type
0
12 Maleyl-Wr-OEt
1 Arg-OEt, pH 9.5 a-chymotrypsin 2 aq HzS04,5O"C, 72h
et al., 1994). Fig. 7. See text (FISCHER
protecting groups such as Boc, Z and Fmoc. The N-formyl group is easily installed by acetic-formic anhydride generated in situ (AGO, HCOpH) and removed by mild acid hydrolysis (FLOERSHEIMER and KULA,1988). Many studies have been conducted to determine optimum water levels for particular protease-catalyzed reactions (CALVETet al., 1993; HWANCet al., 1995). While enzymes cannot work under totally anhydrous conditions, the amount of essential water can be extremely small ( c0.1%). Frequently, optimum water levels between 1-5% are found to give good yields for peptide couplings for many proteases and in a variety of solvents, with minimal secondary product hydrolysis. The water content can be reduced in thermodynamically controlled couplings by using small hydrogen-bonding solvents such as formamide and ethylene glycol which act as "water mimics" and thereby replace most of the water essential for enzyme function (KITAGUCHI and KLIBANOV, 1989). In many direct acid-amine couplings catalyzed by thermolysin in anhydrous t-amyl alcohol (e.g., Z-Gly-Gly-Phe-OH + H-Leu-NH, to form Z-Gly-Gly-Phe-Leu-NH,), the very fast reaction rate at 4% water could be reproduced with a mixture of 1% water with 9% formamide (the rate with 1%water alone was nearly 200 times slower than at 4%). Secondary product hydrolysis was greatly reduced in these systems compared to the 4% water, t-amyl alcohol system. Solvent inhibition of enzyme activity can be advantageous. Particular concentrations of water-miscible organic solvents ( 5 0 4 0 % ) cause inhibition of the amidase activity of serine and cysteine proteases, but the residual esterase activities are still sufficiently high to catalyze peptide bond formation by kineticcontrolled ester aminolysis.Secondary product hydrolysis was not observed in these solvent mixtures but occurred in water alone (BARBAS et al., 1988). The issue of enzyme-solvent interaction is a complex one. Certainly, marked conformationa1 changes can occur on changing from an aqueous to an organic environment, and this is frequently reflected by changes in enzyme specificity (e.g., CROUTet al., 1993). Selectivity can even be reversed by solvent effects, as
2 Acylases, Peptidases
illustrated in a recent study of the lipoprotein lipase (Pseudornonas sp.)-catalyzed kinetic acylation of L-Ser-P-naphthylamide by the active ester trifluoroethyl butanoate (EBERTet al., 1996). In t-amyl alcohol, acylation occurred preferentially on the amino group but in pyridine the regioselectivity was reversed. The authors conclude that changes in intermolecular hydrogen bonding between substrate and solvent may affect chemoselectivity as the solvent is changed from hydrogen bond donor to acceptor. Solvation effects on intramolecular hydrogen bonding in the substrate may also be important. Another possible solvent effect on enzyme reactivity is the inclusion of solvent molecules in the enzyme’s active site and this has been observed in an X-ray study of subtilisin grown in acetonitrile (FITZPATRICK et al., 1993). Relaxed subtilisin specificity for D- and Lamino acid esters in organic solvent compared to water has also been noted (MARGOLIN et al., 1987). However, while the size of side chain residues which can be accommodated may vary between solvent systems, encouraging the experimental manipulation of reaction conditions to optimize individual processes, most proteases retain their overall L-selectivity regardless of the reaction medium.
2.2.1.2 Regioselectivity and Unnatural Substrates
253
coupling of either D- or L-proline with a range of donor esters (e.g., 14, Fig. 8). The proline carboxylate protecting group could be ester or amide. whereas Ala, Phe and Lys(COCF,) were tolerated at P,. In the latter case, coupling of Z-Lys(COCF,)-OMe with ProOBn gave a dramatic increase in yield from 30% to 83% when dialysis-purified enzyme was used. Moreover, the yield of the coupling was strongly dependent on water content; addition of only 1% water reduced the yield significantly and 10% gave no product at all (CHENet al., 1994). Dehydro-amino acids can also be coupled by protease catalysis under kinetic conditions. In a study of the amidation of a-dehydroglutamate diesters (16) (Fig. 9) by various amino acid amides the regioselectivity of amidation was found to vary between the a- and y-ester groups depending on the protease used. Papain favored formation of the a-amide (17) and a-chymotrypsin the y-amide (18). The yield was highly dependent on the structure of the leucine C-terminal group with primary
14 Moz-Phe-OMe
The protease-catalyzed synthesis of pure L,L-diastereomers from racemic amino acid reL- or DPro-NH2, actants has already been referred to (CHENet alcalase, ‘BuOH, al., 1991). In general, proteases are much more specific for L-residues in the acyl donor (PI) 25°C. 2d than at the acyl acceptor (Pi) although some enzymes such as subtilisin (Bacillus subrilis) are able to incorporate D-residues at PI (MARCOLIN et al., 1987). However, there are many examples of the reaction of D-amino acid esters or amides as nucleophiles in kinetically controlled peptide bond formation (RICCA and CROUT,1989). Couplings involving proline are often more difficult than those of other 15 Moz-Phe-Pro-NH2,46%yield amino acids because the amine group is secon( Moz-Phe-D-Pro-NH2,45% yield) dary, however the relaxed specificity of alcalase in anhydrous r-BuOH allowed successful Fig. 8. See text (CHENet al., 1994).
254
5 Cleavage and Formation of Amide Bonds 0
z, N
enzyme
OMe
p H 8, 35T, 24h
16 (2)-a-dehydroGlu(0Me)-OMe
LY.%
-
Enzyme
17 a-ku-anilide
18 yku-anilide
Papain a-Chymotrypsin
7 7%
not given
7%
64%
Fig. 9. See text (SHINet al., 1993).
amide, phenylhydrazide, and benzylamide all giving much poorer yields than the anilide, although regioselectivity followed the same trend. This raises the interesting possibility of some additional P; or P; hydrophobic binding occurring with anilides which could be exploited in other couplings of these enzymes (SHIN et al., 1993). With natural substrates, selectivity for coupling at the a-amino or carboxylic acid substituents is frequently absolute, and potentially competing side chain groups do not usually interfere (Tab. 4). Lysine is an interesting exception which gave complete camidation when used as acyl acceptor in the subtilisin Carlsberg-catalyzed reaction of Ac-Phe-OEt with H-Lys-O'Bu. In contrast, a-chymotrypsin was partially selective for a-amidation ( a :E = 7 : 3), confirmed by the reverse hydrolysis reac-
tion with this enzyme, for which only the aamide was a substrate (KITAGUCHI et al., 1988). Hydrophobic enzyme-substrate interactions are usually important in protease-catalyzed peptide couplings, and frequently the most efficient reactions with many proteases are those where both PI and P; are hydrophobic. This is not true of all proteases as, for example, proteinase K is widely tolerant of the PI substituent (except Pro) but prefers hydrophilic Pi amino acid amides for the nucleophilic component (CEROVSKYand MARTINEK, 1988).The differing structural requirements of some proteases is further illustrated by peptide coupling reactions with the racemic unnatural amino acid allylglycine (FERNANDEZ et al., 1995).In this study, proteases from Aspergillus oryzue and A . sojue, as well as pronase E and
Trypsin
H-Ala-pNA
Boc
NH
19
pH8,aqDMSO,
I
2 5°C
BOC
NH2
20 Proline stereochemistry L D
Fig. 10. Hydrolysis of side chain mimics (ITOHet al., 1996).
0
-
H
Reaction time, h
Yield
2 12
80
%
76
2 Acylases, Peptidases
P
255
nagarse, were able to use allylglycine (Ag) as the acyl donor (ruc-Z-Ag-OMe) but not as acceptor (rac-Ag-OEt). Conversely, a-chymotrypsin could use ruc-Ag-OEt as acceptor but not rac-Z-Ag-OMe as donor. In contrast to the relaxed substrate specificity of many proteases, trypsin is generally highly specific for arginine or lysine at the PI position. This selectivity has been used in an interesting way through the use of "inverse substrates", in which a basic side chain mimic of these residues is incorporated as the ester leaving group on the acyl donor, rather than at the amino acid chiral center (ITOH et al., 1996). Several dipeptide analogs were prepared in good yield by the method; both D- and L-prolinyl esters (19) reacted well as acyl donor (Fig. 10) whereas D-alanyl and D-leucinyl p-nitroanilides gave only slight reaction as acceptors. Although p-amidinophenyl esters are intrinsically reactive (the authors noted a low level of reaction in the absence of enzyme), several of the reactions were very rapid, implying good recognition by the enzyme. p-Guanidinophenyl esters have been used in a similar way (SCHELLENBERGER et al., 1991).This type of approach, whereby enzyme recognition is directed to an alternative site in the substrate, has much potential for the design of new enzymatic coupling methods utilizing new specific enzymes and catalytic antibodies.
2.2.1.3 Modified Enzymes The immobilization of enzymes on solid supports has been used in peptide synthesis for some time. The technique provides greater ease of handling and recovery plus improved enzyme stability which frequently permits the use of higher temperatures. The enzymes may be chemically linked to the support, trapped within a semipermeable membrane or adsorbed onto a support from an aqueous solution by pH control, or with other additives or by lyophilization for use in organic solvents. Supports frequently used include celite (LoZANO et al., 1995), Amberlite XAD-7 (NAGAYAW et al., 1994b), polymethacrylates (ZHOU and HUANG, 1993),and Sepharose (GAERTNER et al., 1991). Proteases can also be derivatized
256
5 Cleavage and Formation ofAmide Bonds
stepwise addition of amino acid units. A good example is the recently reported 10-step synthesis of “delicious octapeptide” amide in 39% overall yield (GILLet al., 1995). Here two individual tetrapeptide units were built up by stepwise amidations using amino acid esters, utilizing the individual selectivities of different proteases (Fig. 11). Not surprisingly, enzymes of restricted specificity have found application to peptide synthesis. Prolyl endopeptidase, requires L- or Dproline as PI, but will not accept L- or D-proline as Pi. Prolyl endopeptidase was used in reverse under thermodynamic conditions to couple Gly-NH, quantitatively to the C-terminal proline of a nonapeptide precursor in the synthesis of luteinizing hormone releasing hormone Glu-His-Trp-Ser-Tyr-Gly-Leu-ArgPro-Gly-NH2 (LH-RH) (TOGAME et al., 1994). Similarly, a prolyl dipeptidyl-aminopeptidase from Lactococcus lactis (PepX) gave specific peptide bond formation under kinetic conditions between C-terminal Pro dipeptides of the type Xaa-Pro-X and a wide range of Pi residues. X could be ester, amide, or even Phe, and C-terminal protection of the nucleophile was not necessary due to the high specificity of the 2.2.2 Larger Peptides enzyme. Chemically modified enzymes such as In protease-catalyzed dipeptide synthesis, thiolsubtilisin, in which amidase activity has enzyme selection can frequently be made on been considerably reduced relative to the estethe basis of the known P,-P; cleavage specific- rase activity by converting the active site Ser’’’ ity. In couplings of larger peptide units, resi- to Cys, have also been used effectively in the dues more remote from the cleavage site may synthesis of large peptides (MIHARAet al., also affect reactivity. Effects may be adverse, 1995). The subtilisin mutant subtiligase has as in the thermodynamic coupling of Z-Ala- been used for the cyclization of linear pepAla-Phe-OH with H-Phe-Ala- Ala-OMe where tides. From a systematic study of chain length replacement of the P; and P; Ala residues by the authors identified a minimum requirement Gly gave a much lower yield (ABDELMALAK, of 12 residues for cyclization - smaller pep1992), or they may be beneficial in, for exam- tides gave hydrolysis or dimerization. Five cyple, the accommodation of unnatural amino clic peptides (range of 12-31 residues) were et acids at more remote sites (WONGet al., 1990). prepared in yields of 36%-85% (JACKSON Most solvent-based enzyme-catalyzed peptide al., 1995). synthesis to date has focused on relatively small peptides of up to four amino acids and much of the methodology developed has yet to 2.3 Non a-Amino Acid Substrates find application to larger peptides. Of course, Acylase or peptidase-catalyzed formation of this is not always straightforward. Larger peptides often have poor solubility and, even in amides from non a-amino acid substrates has low water media, there is always the potential only been used to a limited extent due to the for secondary hydrolysis at unwanted sites. success of the corresponding lipase or esteHowever, by careful selection of enzyme spec- rase-catalyzed kinetic aminolysis reaction, ificities, larger peptides can be constructed by which has the inherent advantage of minimal on polyethylene glycol by cyanuric chloride activation to give modified enzymes which have high solubility in nonpolar organic solvents (GAERTNER et al., 1991). A recent report (PARADKAR and DORDICK, 1994) describes the extraction of a-chymotrypsin from aqueous into organic solution by ion-pairing with a surfactant. The catalytic activity was significantly higher than when the free enzyme was suspended in the same solvent and only four times less than in aqueous buffer. Cross-linked enzyme crystals (CLEC’s) have also proved very useful catalysts and are commercially available. Here, the microcrystalline protease catalysts are cross-linked with a bifunctional reagent such as glutaraldehyde which confers good thermal and long term stability and facilitates recovery of biocatalyst. For example, the thermolysin CLEC has been successfully used in synthesis of peptides of widely differing size from the aspartame precursor Z-Asp-Phe-OMe to coupling of PheNH2 to the oxidized P-chain of insulin (PERSICHETTI et a]., 1995).
2 Acylases, Peptidases
I
Z-Lys(Z)-OEt
Z-Glu(0Et)-OEt
1
Gly-OEt. chymopapain
Z-Lys(Z)-Gly-OEt
I
Ser-OEt, chymopapain
Z-Glu(0Et)-Ser-OEt
Asp( OAll) -0All Papaya protease IV or chymopapain
Z-Lys(Z)-Gly -Asp(OAll)-OAll
1 I
257
I
Leu-OEt. Pronase E
Z-Glu(0Et)-Ser-Leu-OEt
Glu(OAll)-OAll S ubtilisin
Z-L~~(Z)-Gl~-Asp(OAll)-Gl~(OAll)-OEt Ally1 ester deprotection
1 1
Ala-NH2 Proteinase K
Z-Glu(0Et)-Ser-Leu-Ala-NHz Z-deprotection
1
Z-LyS(Z)-Gly-ASp-Glu-OEt
V8 protease
Glu(OEt)-Ser-Leu-Ala-NH2
Z-Lys(Z)-Gly-Asp-Glu-Glu(OEt)-Ser-Leu-Ala-NH2
1
Deprotection steps
Lys-Gly-Asp-Glu-Glu-Ser-Leu-Ala-NH2 2 1 "Delicious octapeptide" amide Fig. 11. See text (GILLet al., 1995)
secondary hydrolysis of product amide by these esterolytic enzymes. The protease subtilisin Carlsberg was found to catalyze stereoselective (S)-acylation of a range of primary amines, using the reactive acyl donor trifluoroethyl butyrate (KITAGUCHI et al., 1989). Highest ee's were for more sterically-demanding substituents on the amine a-carbon. For example, a-methyltryptamine gave the (S)amide with 99% ee whereas a-methylbenzylamine gave 85% ee. Anhydrous 3-methyl-3pentanol appeared a unique solvent for the reaction - initial rates and product ee's were significantly lower in other solvents tried.
Examples of lactam hydrolysis by microbial acylases have been reported, the best known of which is the enantioselective hydrolysis of the bicyclic lactam derivative (22) (Fig. 12). Two distinct organisms were found to hydrolyze the separate enantiomers of (22) to give either enantiomer of the cyclopentene (23)in 45% yield and >98% ee (TAYLOR et al., 1990). Of note are the high substrate concentrations (50 g L-') tolerated by these biocatalysts (Enza-1 and Enza-20) and the short reaction times (3 h). The homochiral products were used to prepare the antiviral carbocyclic nucleoside carbovir (VINCEand HUA,1990) and a
258
5 Cleavage and Formation of Amide Bonds
bicyclic p-lactam derivative (24) (Fig. 13), whereas the saturated analog was a poor substrate (EVANS et al., 1991b).
1
(lS,4R)-22
(1R,4S)-23
Enza-1.
Rhodococcus equi
NCIB 40213
1 (1R.a)-22
The applications of penicillin G acylase (penicillin amidohydrolase, EC 3.5.1.11) in synthesis (BALDARO et al., 1992) and the enzymatic synthesis of penicillin analogs (LUENGO, 1995) have been reviewed.
3.1 Reactions of Penicillin and Cephalosporin Substrates Enza-20, Pseudomonas solanacearum
NCIB 40249
(1S94R1-23
et al., 1990). Fig. 12. See text (TAYLOR
E~C-24
3 Penicillin Acylase
Fig. 13. See text (EVANS et al., 1991b).
GABA-agonist, cis-3-aminocyclopentanecarboxylic acid. Interestingly, in this latter work, the saturated bicyclic lactam analog of (22) was not a substrate for Enza-1 or Enza-20 (EVANSet al., 1991a). Similarly Enza-1 catalyzed the enantioselective hydrolysis of the
Penicillin G acylase (PGA) from E. coli is widely used in industry for the manufacture of 6-aminopenicillanic acid (6-APA) (27) by deacylation of penicillin G (25).Typically the reaction is performed batchwise at 3540 "C, pH 7.5-8.0 using immobilized PGA (VANDAMME, 1988)packed into a column, which can be reused up to 600 times (MATSUMOTO, 1992). Chemical rearrangement of penicillin G (25) gives the cephalosporin analog (26) (CHAUVETTE et al., 1971), which can also be hydrolyzed by PGA to yield 7-amino desacetoxycephalosporanic acid (7-ADCA) (28). Both 6-APA (27) and 7-ADCA (28) are key intermediates for the synthesis of penicillin and cephalosporin antibiotics by chemical acylation of the amino group (Fig. 14). Thus far, there is no commercial process for the direct removal of the side chain from cephalosporin C, however, a two-step enzymatic process is currently under development (MATSUMOTO, 1992; CONLON et al., 1995; see also Chapter 1, this volume). Penicillin G acylase (PGA) from E. coli is the most commonly used penicillin acylase in synthesis. The active site residue is serine, but there is no adjacent histidine residue as is commonly found in serine proteases (DUGGLEBY et al., 1995). Acylases showing specificity for other penicillin side chains are also known (penicillinV acylase: phenoxyacetyl and ampicillin acylase: D-phenylglycyl side chains). Specificity is frequently tight, as illustrated by
mNpx B
0
25 Penicillin G
3 Penicillin Acylase
259
Chemical rearrangement *
&
C02H
26 Penicillin acylase
Penicillin acylase
Fig. 14. See text.
the D-phenylglycyl-specific enzyme from Xunthornonus sp. which does not accept benzyl penicillin as a substrate (BLINKOVSKY and MARKARYAN, 1993). E. coli PGA catalyzes the hydrolysis of phenylacetyl amides or esters, but can act in reverse to form these derivatives by kinetic acylation of amines with suitable phenylacetyl donors in a similar way to the serine and cysteine proteases discussed earlier (Sect. 2.2). Amides (29) act as acyl donors in the acylation of 6-APA (27) to give ampicillin (30)(Fig. 15) (KAASGAARD and VEITLAND, 1992). PGA has high specificity for the phenylacetyl group, as illustrated by the selective hydrolysis of a protected aminothiazolyl-cephem (31) (Fig. 16) (ZENONI and FUGANTI,1994) and accepts only slightly modified analogs or structural mimics such as p-hydroxyphenylacetyl, phenylpropionyl, 2-furylacetyl, and isobutoxyacetyl. Substitution of the phenylacetyl a-carbon by -OH, -0CH0, and -NH2 is also tolerated. In addition, the enzyme has a high L-selectivity which has found use in resolution processes. An example is the enantioselective acylation of a p-lactam intermediate (33) used in the synthesis of the carbacephalosporin antibiotic loracarbef (35) (Fig. 17) (ZMIJEWSKI et al., 1991).
Hbx 5. C
0
I
O20
27 6-APA
30 Ampicillin Fig. 15. See text (KAASGAARDand VEITLAND, 1992).
260
5 Cleavage and Formation ofAmide Bonds
I
31
J
COzH
P I
Fig. 16. 1994).
COzH
32
H'
Immobilized PGA, pH 6, 28°C. 24h
rac-3 3
0 COzMe 3 4
0
Ar
Yield Q
ee 9%
Ph PhO
41 44
96 97
O+p+,, NH3 H
+--
(ZENONI and FUGANTI,
Ar
0
COzMe
See text
C0,O 35 Loracarbef Fig. 17.
See text
(ZMIJEWSKI et al.. 1991).
3.2 Other Substrates The specificity of PGA for the PhenYlacetYl group is not confined to p-lactams. A wide range of other N- and O-PhenYlacetYl derivatives are accepted by the enzyme and are hYdrolyzed with L-stereoselectivity.Unlike PKA, substrates are not restricted to a-amino acids.
N-phenylacetyl derivatives of both p- and y-amino acids (SOLOSHONOK et a]., 1995; MARGOLIN, 1993) are resolved by PGA-catalyzed hydrolysis to give (R)-amino acids (38.39) and recovered (S)-amide (37)(Figs. 18 and 19). Under forcing conditions (45"C, 2 d) the (S)-amide (37)is also hydrolyzed (MARGOLIN, 1993).
3 Penicillin Acylase
37 ( 9 - h i d e , >95% ee
36
261
38 (R)-Amino acid
~r = Ph, 4-F-, 2-F-, 4 - 0 , 4-MeG,C6H4; 3,4,5-(Me0)3-C&~ Fig. 18. Enantioselective hydrolysis of N-phenylacetyl3-aminoacids (SOLOSHONOK et al., 1995).
39 (R)-amino acids R = ethynyl; >96%ee R = allenyl; 75% ee R = vinyl; 78% ee Fig. 19. y-Amino acids produced by PGA-catalyzed enantioselective hydrolysis of the N-phenylacetyl derivatives (MARGOLIN, 1993).
acetamides was attempted but only the 4methyl derivative was a substrate. Likewise, achymotrypsin only accepted the methyl esters of 4-methyl and 4-ethyltryptophans and not the larger 4-dimethylallyl substituent. PGAcatalyzed hydrolysis of ruc-N-phenylacetyl derivatives of the tryptophan analogs gave (S)acids (40) and recovered (R)-amides. Again, the (R)-amides were also hydrolyzed upon prolonged reaction with PGA.
3.3 Protection of Amino Groups The same high L-enantioselectivity is achieved with N-phenylacetamides of a-amino acids, and this has been used in the resolution of a range of 4-alkyl tryptophans (Fig. 20) (NE-ITEKOVEN et al., 1995). Initially, acylase I-catalyzed hydrolysis of the corresponding
H
40 (9-Tryptophan derivatives R = H, Me, Et, CH2CH=CH2-; >98%ee PGA, MeOH:H20, 92:8, pH 7.6
Fig. 20. a-Amino acids produced by PGA-catalyzed enantioselective hydrolysis of the N-phenylacetyl derivatives (NEITEKOVENet al., 1995).
Several examples of the PGA-catalyzed introduction and cleavage of the N-phenylacetyl protecting group are included in comprehensive reviews covering the whole area of enzymatic protecting groups (WALDMANN and SEBASTIAN, 1994; GREENEand WUTS, 1991). The high specificity of PGA for the N- or 0-phenylacetyl substituent, compared to other amide or ester groups makes phenylacetyl a very useful protecting group, particularly in peptide chemistry where undesired cleavage of peptide bonds does not occur. The enzyme tolerates high levels of organic solvents and functions at near neutral pH in both the introduction and removal of phenylacetyl. Moderate to good yields of N-phenylacetyl di- and tripeptides can be obtained by PGAcatalyzed N-terminal coupling with phenylacetic acid, although the reaction is sensitive to the steric bulk of the N-terminal residue. For example, N-terminal Gly, Ala, and Ser react, but more sterically-demanding residues do not. However, the size of the second amino acid unit of the peptide acyl acceptor is also
262
5 Cleavage and Formation of Amide Bonds
important as shown by the lack of reactivity towards amide formation with Met-Tyr or TyrGly, whereas Met-OEt and Tyr-OEt are good substrates (PESSINAet al., 1988). Although active ester-type peptide coupling of Nphenylacetyl amino acids is prone to slight racemization compared to carbamate-protected analogs such as Boc or Z, the problem can be avoided by use of enzyme-catalyzed couplings, and N-phenylacetyl has found much application to peptide synthesis when used in this way. Recent examples include a synthesis of leucine enkaphalin 1-butyl ester Tyr-GlyGly-Phe-Leu-O'Bu, where N-phenylacetyl was both introduced and cleaved by PGA catalysis in aqueous solution (DIDZIAPETRIS et al., 1991). Papain and a-chymotrypsin were used for individual coupling steps. Reaction times for PGA-catalyzed deprotection of the Nphenylacetyl intermediates in this synthesis increased with the size of the peptide, an observation in keeping with the steric constraints noted by PESSINA et al. (1988). Other applications include the use of Nphenylacetyl protection for a synthesis of aspartame, where immobilized PGA (recyclable >40 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
5 Cleavage and Formation of Amide Bonds
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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BRIEVA, R., REBOLLEDO, F., GOT OR,^. (1990), Enzymatic synthesis of amides with two chiral centers, J. Chem. SOC.,Chem. Commun., 1386-1387. BRUCE,M. A., ST. LAURENT, D. R., POINDEXTER, G. S., MONKOVIC, I., HUANG, S., BALASUBRAMANIAN, N. (1999, Kinetic resolution of piperazine-2-carboxamide by leucine aminopeptidase, Synrh. Commun. 25,2673-2684. CALVET,S., CLAPES,P.,TORRES,J. L., VALENCIA, G., P. (1993), Enzymatic FEIXAS,J., ADLERCREUTZ, synthesis of X-Phe-Leu-NH, in low water content systems: Influence of the N-a protecting group and the reaction medium composition, Biochim. Biophys.Acta 1164,189-196. CANTACUZENE, D., GUERREIRO, C. (1989), Optimization of the papain-catalyzed esterification of amino acids by alcohols and diols, Tetrahedron 45, 741-748. CEROVSKY, V. (1990), Free trypsin-catalyzed peptide synthesis in acetonitrile with low water content, Biotechnol.Lett. U , 899-904. K. (1988),Peptide syntheCEROVSKY, V., MARTINEK, sis catalyzed by native proteinase K in water-miscible organic solvents with low water content, Collect. Czech. Chem. Commun. 54,2027-2041. CHAMORRO, C., GONZALEZ-MUNIZ, R.,CONDE,S. (1995), Regio- and enantioselectivity of the Candida antarctica lipase catalyzed amidations of Cbz-L- and Cbz-D-glutamic acid diesters, Tetrahedron:Asymmetry 6,2343-2352. P. A., RYAN, C. W., CHAUVETE, R. R., PENNINGTON, I. G., VAN COOPER,R. D. G., JOSE,F. L., WRIGHT, HEYNINGEN, E. M., HUFFMAN,G. W. (1971), Chemistry of cephalosporin antibiotics XXI. Conversion of penicillins to cephalexin, J. Org. Chem. 36,1259-1267. CHEETHAM,F! S. J. (1994). Case studies in applied biocatalysis - from ideas to products, in: Applied Biocatalysis (CABRAL, J. M. S., BEST,D., BOROS, L.,TRAMPER, J., Eds), pp. 47-109. Chur: Harwood Academic Publishers. CHEN,S.T., HSIAO,S. C., WANG,K.T. (1991). Enantioselective peptide-bond formation using alcalase, Bioorg. Med. Chem.Lett. 1,445. CHEN,S.T., CHEN,S.Y., WANG,K.T. (1992), Kinetically controlled peptide bond formation in anhydrous alcohol catalyzed by the industrial protease alcalase,J. Org. Chem. 57,69604965. CHEN,S. T., CHEN,S. Y., U o , C. L., WANG,K. T. (1994), Proline as a nucleophile in kineticallycontrolled peptide synthesis catalyzed by alcalase in 2-methyl-2-propanol. Bioorg. Med. Chem. Lett. 4, 443-448. CHENAULT, H. K., DAHMER,J., WHITESIDES, G. M. (1989), Kinetic resolution of unnatural and rarely occurring amino acids: enantioselective hydrolysis of N-acyl amino acids catalyzed by acylase 1, J. Am. Chem. SOC.111,6354-6364.
272
5 Cleavage and Formation ofAmide Bonds
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Biotechnology Second, Completely Revised Edition H.-J. Rehm and G. Reed copyrightOWILEY-VCH Verlag GmbH, 1998
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
3 Naturally Occurring Nitriles
+
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.,
3 Naturally Occurring Nitriles
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).
4 Biotransformation of the Nitrile Group
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
4 Biotransformation of the Nitrile Group
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-
4 Biotransformation of the Nitrile Group
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 ni- tase 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 differ- ed to sulfur and nitrogen or oxygen (NELSON
4 Biotransformation of the Nitrile Group
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,
4 Biotransformation of the Nitrile Group
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).
.
+
4 Biotransformation of the Nitrile Group
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
4 Biotransformation of the Nitrile Group
(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
4 Biotransformation of the Nitrile Group
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
Nitrile Hydratase Activity
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
Nitrile Hydratase Activity
'""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
Nitrile Hydratase Activity
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
5 Biotechnology of Nitrile Transformations
307
Tab. 2. Continued
c-
Microorganisms Capable of Nitrile Transformation"
Ni trile
N
CZN
23
"7""
Nitrile Hydratase Activity
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
5 Biotechnology of Nitrile Transformations
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
5 Biotechnology of Nitrile Transformations
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.
315
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