lipases and lipase-catalyzed esterification reactions in nonaqueous ...

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CATALYSIS REVIEWS Vol. 44, No. 4, pp. 499–591, 2002

LIPASES AND LIPASE-CATALYZED ESTERIFICATION REACTIONS IN NONAQUEOUS MEDIA S. Hari Krishna and N. G. Karanth* Fermentation Technology & Bioengineering Department, Central Food Technological Research Institute, Mysore 570013, India

ABSTRACT Enzymatic reactions in nonaqueous solvents offer new possibilities for the biotechnological production of many useful chemicals using reactions that are not feasible in aqueous media. In the recent years, the use of enzymes in nonaqueous media has found applications in organic synthesis, chiral synthesis or resolution, modification of fats and oils, synthesis of sugar-based polymers, etc. The use of lipases in esterification reactions to produce industrially important products such as emulsifiers, surfactants, wax esters, chiral molecules, biopolymers, modified fats and oils, structured lipids, and flavor esters is well documented. The interest in using lipases as biotechnological vectors for performing various reactions in both macroand microaqueous systems has picked up tremendously during the last decade. This review covers important features of lipases and lipase-catalyzed esterification reactions including the kinetics and stability of lipases, and modeling aspects.

*Corresponding author. Fax: þ 91-821-517-233; E-mail: [email protected] 499 DOI: 10.1081/CR-120015481 Copyright q 2002 by Marcel Dekker, Inc.

0161-4940 (Print); 1520-5703 (Online) www.dekker.com


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HARI KRISHNA AND KARANTH

I.

INTRODUCTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

501

II.

LIPASES: A GENERAL ACCOUNT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Occurrence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Nature of Lipases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Interfacial Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Lipases vs. Esterases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Physico-Chemical Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . G. Inhibitors and Activators . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

503 503 503 506 507 508 509 510

III.

SOLVENT SYSTEMS AND WATER ACTIVITY . . . . . . . . . . . . . . . . . . . . A. Solvent Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Aqueous Reaction Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Water: Water-Miscible Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Water: Water-Immiscible Systems . . . . . . . . . . . . . . . . . . . . . . . . . 4. Nonaqueous Reaction Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Anhydrous Reaction Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Supercritical Fluids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7. Reverse Micellar Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8. Solvent-Free Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9. Gas-Phase Catalysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10. Ionic Liquids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Water Activity: A Key Parameter in Lipase Catalysis . . . . . . . . . . . . . 1. Role of Water and Water Activity . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Determination of Water Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Effect of Water Activity on Lipase Catalysis . . . . . . . . . . . . . . . . 4. Control of Water Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Regulation of Water Activity Through Water Removal . . . . . . .

513 516 516 518 519 519 520 520 522 522 523 523 524 524 525 525 526 527

IV.

LIPASE-CATALYZED ESTERIFICATION REACTIONS . . . . . . . . . . . . . A. Modification of Fats and Oils . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Cocoa Butter Equivalents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Fats with Improved Spreadability . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Structured Lipids (Highly Digestive Triglycerides) . . . . . . . . . . . B. Wax Esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Surfactant Esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Organic Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Kinetic Resolution of Chiral Compounds . . . . . . . . . . . . . . . . . . . 2. Selective Acylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Resolution of Sterically Hindered Compounds . . . . . . . . . . . . . . . E. Biopolymer Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Polycondensation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Ring-Opening Polymerization of Lactones . . . . . . . . . . . . . . . . . . 3. Ring-Opening Copolymerization of Lactones . . . . . . . . . . . . . . . . 4. Control of Polmer Terminal Structure End-Functionalization . . . F. Flavor Esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

528 529 530 530 530 532 533 534 535 538 541 541 543 544 545 545 545

V.

KINETICS AND PROCESS OPTIMIZATION . . . . . . . . . . . . . . . . . . . . . . . A. Kinetics of Lipase-Catalyzed Esterification . . . . . . . . . . . . . . . . . . . . . . .

547 547


ESTERIFICATION REACTIONS IN NONAQUEOUS MEDIA

B. C.

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Mass Transfer Influences on Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . Process Optimization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

554 555

VI. STABILITY OF LIPASES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Enzyme Stabilization in Nonaqueous Media . . . . . . . . . . . . . . . . . . . . . . B. Medium Engineering . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . C. Biocatalyst Engineering . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Microwave Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

556 557 557 558 558

VII.

MODIFIED LIPASES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Immobilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . B. Chemical Modification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Cross-Linking . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Modification with Polyethylene Glycol . . . . . . . . . . . . . . . . . . . . . 3. Modification with Polymers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Modification of Amino Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Lipid- and Surfactant-Coated Enzymes . . . . . . . . . . . . . . . . . . . . . C. Modifications by Lyophilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Ion-Pairing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . E. Molecular Imprinting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . F. Trapping in Presence of Interfaces . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . G. Protein Engineering . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Rational Design . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Directed Evolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Modeling of Lipase Selectivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Combinatorial Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

559 559 560 560 561 561 562 562 563 564 564 564 565 565 566 568 573

VIII.

CONCLUDING REMARKS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

573

ACKNOWLEDGMENTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

575

REFERENCES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

575

Key Words: Lipase; Esterification reactions; Nonaqueous media; Organic solvents; Biopolymers; Emulsifiers; Biosurfactants

I. INTRODUCTION Historically, enzymatic catalysis has been carried out primarily in aqueous systems. However, water is a poor solvent for nearly all reactions in preparative organic chemistry. Attempts to place enzymes in systems other than aqueous media date back to the beginning of last century.[1 – 3] In the earlier attempts, water-miscible organic solvents (e.g., ethanol or acetone) were added to aqueous solutions of enzymes. As long as a high water content was available, enzymes retained the catalytic activity, although much less so than in water. The next step was the use of biphasic mixtures in which an aqueous solution of enzyme is emulsified in a water-immiscible solvent (e.g., isooctane or heptane). Substrates are added to the system where they diffuse to the aqueous phase, undergo


502


enzymatic conversion, and the products diffuse back to the organic phase. To facilitate mass transfer, the size of water droplets may be reduced to result in microemulsions. To stabilize these microemulsions, surfactants may be added to form reverse micelles.[4] A more recent development was to employ nearly nonaqueous solvents as media for enzymatic reactions.[5 – 12] Such solvents containing traces of water have been successful in keeping the enzyme activity at high levels. This approach emphasized by Klibanov has stimulated tremendous research efforts in the direction of achieving various kinds of synthetic reactions with high efficiency enzymatically. A further important step[13] was to use enzymes (both soluble and immobilized) in anhydrous organic solvents. Enzymatic reactions in nonaqueous solvents offer new possibilities for the biotechnological production of many useful chemicals using reactions that are not feasible in aqueous media. The use of enzymes in nonaqueous media has found applications in organic synthesis,[5] chiral synthesis or resolution,[14] modification of fats and oils,[15] synthesis of sugar-based polymers,[16] and gas phase biocatalysis.[17] Hydrolytic enzymes can be employed to usefully carry out synthetic reactions if the equilibrium position of the reaction is shifted sufficiently to give a high product yield.[18] At present, there is considerable interest in the use of enzymes as catalysts in organic synthesis.[19 – 26] Among the reactions that are studied are hydrolysis, esterification, and transesterification catalyzed by hydrolytic enzymes such as lipases, esterases, and proteinases. Lipases (EC 3.1.1.3, triacylglycerol hydrolases) have diverse functions in the degradation of food and fat, and are shown to synthesize aliphatic,[27 – 30] aromatic,[31] and other[32 – 34] esters in nonaqueous and biphasic systems. They have also qualified as valuable drugs against digestive disorders and diseases of the pancreas, and find applications in biotechnology (mainly as detergent additives) and as catalysts for the manufacture of specialty oleochemicals and for organic synthesis. Lipases, when employed to catalyze esterification and transesterification reactions in organic solvents, have shown pH memory,[35,36] increased enzyme activity and stability at elevated temperatures,[37,38] regiospecificity and stereoselectivity, and may be affected by water activity.[39] Most importantly, lipases do not require cofactors for activity. The advantages mentioned earlier enable their use in the synthesis of certain specialty chemicals and pharmaceutical intermediates. The use of lipases in esterification reactions to produce industrially important products such as emulsifiers, surfactants, wax esters, chiral molecules, biopolymers, modified fats and oils, structured lipids, and flavor esters is very attractive. Although these reactions can also be carried out using inorganic metal-derived catalysts, the interest in using enzymes as biotechnological vectors for performing various reactions in both macro- and microaqueous systems has picked-up tremendously during the last decade. In fact, about one-third of all biotransformations reported till-date have been performed with lipases.[24,40] On an average, at least one paper dealing with biocatalysis in organic solvents is published every day.



503

A detailed literature survey indicates that although several research and few review articles have been published on lipases in the past, no exhaustive review encompassing all aspects of lipase-catalyzed esterification reactions is available. In addition, characteristics of lipases, solvent systems, and lipase-catalyzed reactions have been dealt with separately and are scattered in the literature. Modeling of lipase catalysis and lipase modifications for effective application has received much less attention when compared to physico-chemical and biological aspects. The present review covers important features of lipases and solvent systems, lipase-catalyzed esterification reactions including the kinetics and stability of lipases, and modeling aspects.

II. LIPASES: A GENERAL ACCOUNT Lipases are the hydrolase group of enzymes, which catalyze the hydrolysis of glyceride ester bonds. The fatty acid released is transformed to water or molecules having a free hydroxyl group or related moiety (nucleophile). They are also termed as acylglycerolases, acyl hydrolases, or triacylglycerol hydrolases.

A. Occurrence Lipases occur widely in animals, plants, and microorganisms,[41] where their biological role is probably digestive (e.g., initiation of lipid/glyceride metabolism), enabling the exploitation of the rich energy store that lipids represent. Lipases from a variety of sources are commercially available (Table 1). While microbial lipases are available, the advent of genetic engineering techniques has made possible the manufacture of an increasing number of lipases from recombinant bacteria and yeasts. For example, a “detergent lipase” is produced commercially at large scale (.100 ton/annum) through fermentation of Aspergillus oryzae into which the Humicola lanuginosa lipase gene was expressed.[42,43]

B. Nature of Lipases Majority of lipases are extracellular, acidic glycoproteins of molecular sizes between 20 and 60 kDa,[44,45] although some form aggregates or are present as multimers in solution (accounting for the high molecular weights).[46] Most purified lipases contain 2 – 15% carbohydrate. The primary structures of lipases from mammalian,[47] bacterial,[48] and fungal[49] sources have been determined,


504

Table 1.

Lipase Sources and Suppliers Sources of Lipases

Supplier Amano Pharmaceutical Co., Nagoya, Japan (www.amano-enzyme.co.jp) Biocatalysts, Pontypridd, Wales, UK (www.biocatalysts.com)

Biocon India, Bangalore, India (www.biocon.com)

Detergent lipases, and other lipases

ThermoCat, Quick Screen, Esterase kits Various lipases, custom made enzymes, salt-immobilized enzymes Various lipases and esterase, Rhodotorula pilimanae, R. arrhizus, Pig liver esterase, R. niveus, C. rugosa


Boehringer Mannheim, Penzberg, Germany (Now Roche Diagnostics, merged with Roche group) (www.roche.com/diagnostics) Novozymes A/S, Novo Nordisk, Bagsvaerd, Denmark (www.novozymes.com) Diversa Corp., San Diego, CA (www.diversa.com) (Innovase LLC, a joint venture of Diversa with Dow Chemical Co, (www.dow.com) Thermogen Inc., Woodridge, IL (www.thermogen.com) Biocatalytics Inc., Pasadena, CA (www.biocatalytics.com) Juelich Enzyme Products GmbH, Wiesbaden, Germany (www.juelichenzyme.com)

Aspergillus niger, C. rugosa, C. lipolytica, G. candidum, H. lanuginosa, M. javanicus, Penicillium sp., P. roquefortii, Pig liver esterase, P. fluorescens, R. delemar, R. javanicus, R. niveus Lipomod AC, Lipomod RD, Animal, C. lipolytica, Ch. viscosum, G. candidum, H. lanuginosa, M. javanicus, M. miehei, P. cyclopium, P. roquefortii, P. fluorescens, R. delemar, R. javanicus, R. japonicus Lipases for leather and detergent industries, Biolipase conc., Lipase FAP, Pregastric esterase, Lipoprotein lipase, Cholesterol esterase, Biolipase A C. rugosa, C. antarctica, Ch. viscosum, G. candidum, H. lanuginosa, M. javanicus, M. miehei, P. cyclopium, P. roquefortii, Porcine pancreas, Pseudomonas, P. fluorescens, R. delemar, R. javanicus, R. japonicus, Chirazyme screening set Lipolase (detergent lipase), and lipases from several sources


Genzyme Biochemicals, Spingfield Mill, Kent, UK (www.genzyme.com) Gist-Brocades, Delft, Holland (www.gist-brocades.nl) (Now part of DSM group: www.dsm.com) Enzyme business of GB is acquired by Genencor (www.genencor.com) Hoechst, Frankfurt, Germany (Now Aventis, after merger with Rhone-Poulenc SA) (www.aventis.com) Nagase & Co Ltd., Ohama, Japan (www.nagase.co.jp) Merck, Darmstadt, Germany (www.merck.com) Roehm, Darmstadt, Germany (www.roehm.de) Seppim, 61500, Sees, France (www.sfrl.fr) Serva, Heidelberg, Germany (Invitrogen group) (www.serva.de)

Porcine pancreas, C. lipolytica, M. javanicus, P. roquefortii, R. arrhizus, R. niveus, C. rugosa, Wheat germ Ch. viscosum, Lipoprotein lipase, C. rugosa, Pseudomonas sp. M. miehei, R. arrhizus, Piccantase (for dairy) Lipomax, Lumafast (Detergent lipases)

Pancreatin Rhizopus Porcine pancreas Aspergillus oryzae, A. niger, B. subtilis, Pancreas Lipase Porcine pancreas, Rhizopus sp., Wheat germ


Sigma – Aldrich – Fluka (www.sigma-aldrich.com)

505


506


which show that the number of amino acids in them range from 270 to 641. Based on the active site, lipases have been classified as serine hydrolases.[50] Structural characteristics include an a/b-hydrolase fold and a catalytic triad consisting of a nucleophilic serine and an aspartate or glutamate that is hydrogen bonded to a histidine. Four substrate binding pockets were identified for triglycerides: an oxyanion hole and three pockets accommodating the fatty acids bound at positions sn-1, sn-2, and sn-3. Oxyanion hole is formed by two backbone amides of a residue in the N-terminal region of the lipase and the Cterminal neighbor of the catalytic serine. The differences in size and the hydrophilicity/hydrophobicity of these pockets determine the specificity of a given lipase. According to the geometry of the binding site, three subgroups may be defined: (i) a crevicelike binding site (e.g., Rhizomucor miehei ), (ii) a funnellike binding site (e.g., Pseudomonas cepacia, Candida antarctica B), and (iii) a tunnellike binding site (e.g., Candida rugosa ). A tunnellike binding site is more likely to accept substrates with long-chain fatty acids compared to bulky substrates. The opposite should apply to lipases with a crevice- or funnellike binding site. All the lipases are members of “a/b-hydrolase fold” family, which show a common architecture composed of a specific sequence of a-helices and b-strands[51] with some variations. Greater variation may be expected for lipases with molecular weights in the lower and higher ends.[52] X-ray crystallographic studies of pancreatic lipase[47] and R. miehei lipase[53] indicate that serine is the nucleophilic residue essential for catalysis, which is chemically analogous but structurally different from that in the serine proteases. The presence of lid covering the active site along with interfacial activation has been generally used to distinguish between lipases and esterases, and to classify an enzyme as a true lipase. However, a number of enzymes, having a lid but not exhibiting interfacial activation, are shown to be exceptions.[54] Examples are lipases from Pseudomonas glumae,[55] Pseudomonas aeruginosa,[56] C. antarctica B,[57] and pancreatic lipases from coypu[58] and the guinea-pig.[59] Lipase from Staphylococcus hyicus showed interfacial activation only with some substrates.[60] Interfacial activation and/or the presence of a lid domain are therefore inadequate to determine whether a specific esterase belongs to the lipase family. As emphasized by Verger, lipases might be pragmatically redefined as carboxyl-esterases that catalyze the hydrolysis/synthesis of long-chain acylglycerols.[61] The term “long-chain” in this context is not well defined, but glycerol esters with an acyl chain length of $10 carbon atoms can be regarded as lipase substrates, with trioleoylglycerol (triolein) being the standard substrate. Hydrolysis of glycerol esters with an acyl chain length of , 10 carbon atoms with tributyrylglycerol (tributyrin) as the standard substrate usually indicates the presence of an esterase.[62] It should be noted that most lipases are highly capable of hydrolyzing these esterase substrates.



507

C. Interfacial Activation Substrate activation at the interface and the lipase undergoing a change to an activated form upon contact with the interface are the basic theories available. Explanations for the former include higher concentrations of substrates in the vicinity of the interface than in the bulk solvent,[63] more suitable conformations or orientations of lipid molecules for chemical reaction,[64] and poor hydration of lipid molecules.[65] Explanations for the latter include the existence of separate adsorption and catalytic sites so that the lipase becomes catalytically active only after binding to the interface,[66] or a conformational change of the lipase upon approaching the interface,[67] or a reorientation of the short one-turn a-helix that covers the active site.[47] Initial x-ray structures[68] revealed that the presence of lid is a general phenomenon with lipases, hence a common activation mechanism was suggested. However, as outlined earlier, this cannot be generalized. Some lipases such as gastric lipases rapidly become denatured at the interface with a pure tributyrin emulsion.[61] In fact, interfacial activation may be viewed as a depressed action on monomeric esters rather than an increased interfacial activity on aggregated substrates. The phenomenon of interfacial activation exhibited by classical lipases (e.g., human pancreatic lipase) is probably linked to the conformational change from closed lid to open form in the presence of water – lipid interface. It is also important to recognize the fact that the hydrolysis of a monomeric substrate might also require the lid to open without the role of interfacial activation. The status of lid and interfacial activation of lipases in organic solvents was investigated by Louwrier et al.[69] using R. miehei lipase as a model. The enzyme did not exhibit interfacial activation in organic solvents which it did in aqueous solutions, suggesting that the a-helical lid must be essentially in the closed position. To test whether the insolubility of the lipase in organic media is a hindrance, the enzyme was covalently modified with polyethylene glycol (PEG). Though this modified lipase was soluble in organic solvent, it did not undergo any interfacial activation irrespective of the hydrophobicity of the interface. Hence, the lipase activity actually observed in organic solvents must have been due to a small fraction of the enzyme with the open lid, which exists in a dynamic equilibrium with the closed lid enzyme in aqueous solution. This study points to potential opportunities for enhancing lipase activity in organic solvents by preopening the lid.

D. Lipases vs. Esterases Although lipases can hydrolyze and form carboxylic ester bonds like proteases and esterases, their molecular mechanisms differ considerably. Enzymes that hydrolyze the shorter-chain carboxylic ester bonds of water-soluble molecules


508


are termed esterases, whereas lipases display a much broader substrate range. Lipases hydrolyze p-nitrophenyl palmitate, but esterases do not, while both can hydrolyze p-nitrophenyl acetate. In contrast to esterases, whose activity is dependent on substrate concentration, lipase activity is directly correlated with the total substrate area but not with the substrate concentration.[61] Esterases, and a few lipases, do not display a lid structure. Prominent candidates are cutinase from Fusarium solani pisi[70] and the lipase B from C. antarctica.[57] The growing knowledge of protein structures stimulated attempts to classify enzymes according to the nature of their fold. Hydrolases are found in the a/b-hydrolase fold group. All lipases (fungal, bacterial, and pancreatic) and majority of the esterases subscribe to this fold. Esterases are split into three groups: (i) the cutinase group (belonging to the flavodoxin like-fold group), (ii) the esterase, and (iii) acetylcholine esterase groups (the latter two having an a/b-hydrolase fold). Both esterases and lipases generally display a broad substrate specificity and thus cannot be classified based on their function. Fojan et al.[71] provided a novel differentiation tool, for closely related esterases and lipases, based on detailed comparison between amino acid composition and protein surface electrostatic distribution. The global sequence alignment of esterases and lipases showed no links between them. Cutinase, which exhibit optimal activity on partially soluble as well as emulsified substrates, belongs to a separate group of hydrolases in between esterases and lipases on sequence level as well. The lipoprotein lipases cluster outside these groups as another group. The occurrence of tyrosine residues ˚ subset around the active site Ser is 2-fold higher in esterases in the region of 10 A compared to lipases. Lipases display a statistically significant enhanced occurrence of nonpolar residues close to the surface, clustering around the active site at high solvent accessibility values, which may be attributed to an increase in valine, leucine, and isoleucine residues. Both the enzyme groups display optimum activity when the active site is slightly negatively charged. Esterases exhibit their optimum charge around pH 5.5 –6.5 while for lipases it is around pH 8.0 –9.0.[72]

E. Specificity Lipases can be classified into three major groups based on their ability to hydrolyze glycerides. The first group (e.g., lipases of Rhizopus and Rhizomucor ) are unable to hydrolyze at secondary positions, but cleave only the terminal positions of triglycerides and are therefore termed as 1,3-specific. These can be regarded as lipases capable of hydrolyzing primary and to a smaller extent secondary esters, since their substrate ranges are not limited to triglycerides. Some lipases hydrolyze both primary and secondary esters (nonspecific) and belong to the second group. The third group consists of those few lipases that are positionally nonspecific but exhibit fatty acid selectivity, cleaving only ester bonds wherein the fatty acid is of a particular type (e.g., lipases from Geotrichum



509

candidum and ungerminated oat seeds preferentially hydrolyze the esters of 9,10-unsaturated fatty acids). Lipases may also exhibit chain length specificity (e.g., cheese flavor enhancing lipases hydrolyze esters of short-chain, but not medium- and long-chain fatty acids). Jensen et al.[73] have provided an informative account on determination of lipase specificity. Most of the lipases convert esters of medium- (C4) to long-chain (C16) saturated fatty acids. Some of them efficiently hydrolyze fatty acid esters as long as C22 (e.g., R. miehei lipase) or are specific for unsaturated fatty acids like the cis (D-9) specific (G. candidum lipase B). All the microbial lipases hydrolyze triglycerides at sn-1 or sn-3 position, but only a few also at sn-2 position (C. antarctica lipase A, G. candidum lipase B, Penicillium simplicissimum, and C. rugosa lipases). The sn-1(3) stereospecificity of lipases toward triglycerides vary. For example, with trioctanoin as a substrate, lipases from Pseudomonas sp. and P. aeruginosa exhibit sn-1 preference, while C. anatarctica lipase B shows sn-3 preference, with high stereospecificity. All other microbial lipases show medium or low sn-1(3) stereospecificity toward trioctanoin (lipases from Rhizopus oryzae, R. miehei, and C. rugosa hydrolyze trioctanoin preferably at sn-1 with low stereoselectivity). Specificities of various lipases are given in Table 2. The broad synthetic potential of lipases is largely due to the fact that they (in contrast to most other enzymes) accept a wide range of substrates and tolerate organic solvents. They can be employed for either hydrolysis or synthetic reactions depending on the solvent system (Sch. 1). A wide range of substrates other than triglycerides (such as aliphatic, alicyclic, bicyclic and aromatic esters, and even esters based on organometallic sandwich compounds) are accepted by lipases. When the racemic esters or substrates with several hydroxyl groups are fed, lipases react with a high degree of enantio- and regioselectivity. A wide range of thioesters and activated amines can also be substrates for lipases, which expands their potential tremendously.[24]

F. Physico-Chemical Properties Enzymes, in general, are water-soluble (with the probable exception of some membrane-bound enzymes). Most animal lipases exhibit pH optima on the alkaline side (pH 8.0– 9.0).[74] However, depending upon the substrate used, the presence of salts and the kind of emulsifiers present, the optimum may also be shifted to the acidic range.[75] Acid lipases have been found in the lysosomes of a variety of mammalian tissues.[76] Most microbial lipases display maximum activity at pH values ranging from 5.6 to 8.5[74] and maximum stability in the neutral pH range. With regard to temperature, most lipases are optimally active between 30 and 408C, while two lipases exhibited much higher temperature optima: R. miehei (708C) and C. antarctica B (608C). The thermostability of lipases varies considerably according to their origin: animal and plant lipases are usually less thermostable than


510

HARI KRISHNA AND KARANTH Table 2.

Present Name

MW* (kDa)

Burkholderia glumae Burkholderia glumae Burkholderia cepacia Pseudomonas mendocina — — — — — — — — —

33 33 33 33 33 29 48 57 43 20 45 43 22

Candida rugosa — — —

60 35 60

Rhizomucor miehei Thermomyces lanuginosa Penicillium camembertii —

30 30 30 41

Lipase Source Bacterial Chromobacterium viscosum Pseudomonas glumae Pseudomonas cepacia Pseudomonas putida Pseudomonas pseudoalcaligenes Pseudomonas aeruginosa Pseudomonas fluorescens Pseudomonas fragi Bacillus thermocatenulatus Bacillus subtilis Staphylococcus epidermidis Staphylococcus aureus Fusarium solani (Cutinase) Fungal Candida cylindracea Candida antarctica B Candida antarctica A Geotrichum candidum

Specificity of Lipases

Mucor miehei Humicola lanuginosa Penicillium cyclopeum Rhizopus arrhizus (R. oryzae, R. delemar ) Aspergillus oryzae Aspergillus niger Animal Porcine pancreas lipase Guinea-pig pancreatic lipase

— —

30 35

— —

50 48

Human pancreatic lipase Human gastric lipase Human lipoprotein lipase

— — —

50 50 55

Specificity sn-1,3 Nonspecific Nonspecific sn-1,3 sn-1,3 sn-1 (for trioctanoin)

sn-1,3 (thermoalkalophilic) Nonspecific Nonspecific — Nonspecific sn-1,3 trans-specific cis-D9 (unsaturated fatty acids) sn-1,3 Nonspecific sn-1,3 sn-1,3 (phospholipase A1 activity) sn-1,3 sn-1,3 sn-1,3 sn-1,3 (phospholipase A1 activity) sn-1,3 sn-3 (acid stable)

the microbial extracellular lipases.[77] It has been reported that a highly thermophilic strain of Pseudomonas releases a lipase which is heat stable at 1008C.[78] The heat stability of lipases depends on the presence of the substrate, probably because substrate removes excess water from the immediate vicinity of the enzyme and thus restricts its overall conformational mobility. G. Inhibitors and Activators It is recognized that free fatty acids and alcohols tend to inhibit lipasecatalyzed hydrolysis reactions.[79] The fatty acid molecules are thought to accumulate at the interface, thereby blocking access of the enzyme to unreacted



Scheme 1.

Lipase-catalyzed reactions.

511


512


triglyceride molecules,[80] while the low molecular weight alcohols are believed to disrupt the three-dimensional architecture of the lipase. Bile salts enhance the activity of some lipases (e.g., Phycomyces nitens ).[81] The lipase from Rhizopus arrhizus[82] is inhibited by bile salts, but lipases from pancreas,[83] Pseudomonas,[78] Chromobacterium,[84] Streptococcus,[85] and Rhizomucor[86] are not. Pancreatic lipase is inactive with respect to emulsified triglycerides in the presence of micellar concentrations of bile salts. Human pancreatic lipase complex with a specific lipase-anchoring protein (colipase), present in the exocrine pancreatic juice, displays a high specificity towards insoluble triglycerides.[87] Until recently, gastric lipolysis was assumed to be negligible. But, it is now clear that the gastric and pancreatic lipases act in synergy.[88,89] The use of an inhibitor of digestive lipases helps to reduce dietary fat adsorption and holds great promise as an antiobesity agent. Tetrahydrolipstatin, derived from lipstatin produced by Streptomyces toxytricini is a potent inhibitor of pancreatic and gastric as well as cholesterol ester hydrolase. Presently, Roche (Basel, Switzerland) markets this compound as an antiobesity agent (Xenicalw-Orlistat). Many light metal cations are known to influence the lipase action on their substrates. The presence of calcium ions usually increases the reaction rates.[90] While sodium ions have been reported to increase the activities of soluble pancreatic and Aspergillus wentii lipases, they partially inhibit the activity of two lipases from Aspergillus niger.[90] Reversible or irreversible inhibition has been observed upon exposure to heavy metal cations, depending on ion concentration and exposure time.[91,92] Ferrous ions were observed to increase the hydrolytic activity of a lipase from A. niger[90] and Streptococcus faecalis[85] whereas ferric ions inactivated lipases from A. niger,[85] Chromobacterium,[93] Pseudomonas,[94] and S. faecalis.[85] The positive effects of metal ions could be due to the formation of complexes with ionized fatty acids, which change their solubilities and behavior at interfaces, whereas negative effects can be attributed to competitive binding at the active site.[95] Often the lost activity can be restored again by the addition of metal-chelating agents.[92] Nonaqueous solvents can have a variety of effects on enzymes: they may bind specifically, compete with substrate binding, dissociate multimers, shift the equilibrium between two enzyme conformations, alter the helical structure, react with the enzyme, or affect the rate of the catalytic reaction in a number of other ways.[96] Lipase-catalyzed hydrolysis of olive oil was inhibited by the presence of organic solvents because of the competition of the solvent molecules with the triglycerides for adsorption at the interface.[97] Lipase from porcine pancreas was reported to be stable for several hours at high temperatures (,1008C) when the reacting medium was a mixture of tributyrin and heptanol with a water content below 0.4%.[37] This high thermostability can be because the conformational mobility of the enzyme, which is necessary for partial unfolding of the lipase, becomes more restricted when water content is low. This unfolding is the first step in thermal



513

inactivation.[98] When dehydrated, lipases lose their ability to catalyze interesterification reactions involving bulky alcohols, probably because they begin to lose the conformational mobility needed to bind these species in the active center. A layer of bound water (or water hydration shell) plays a key role in maintaining the structural integrity and catalytically active conformation of lipases.[99,100] Therefore, the existence of trace amounts of water in the immediate vicinity of the lipase is a prerequisite for successful functioning of a lipase in organic solvents.[101] This requirement explains why lipases are less active in solvents that are miscible with water, which impart conformational changes leading to inactivation because they extract bound water from the proteinaceous backbones of the enzymes.[102] In microaqueous systems, the bound water in the lipase preparation is of higher significance than the total water content.[103] For lipases from R. miehei and Rhizopus sp., water-binding capacity depends not only on temperature but also on hydrophilic characteristics of the immobilization support[103] and of the additives employed in the immobilization procedure.[104] More information on lipases from different sources, their selectivities, and other details are available in the literature.[41,105 – 107]

III.

SOLVENT SYSTEMS AND WATER ACTIVITY

Enzymes occupy a unique position in synthetic chemistry due to their selectivities and faster catalytic rates under ambient reaction conditions. Nevertheless, synthetic chemists have been reluctant to employ enzymes as reagents in organic synthesis, although there is a need for high selectivity in synthetic chemistry. The reason for this reluctance has been the conventional notion that the enzymes function only in aqueous solutions and most organic compounds are insoluble in water. Furthermore, the water removal is tedious and expensive due to its high boiling point and high heat of vaporization. Side reactions such as hydrolysis, racemization, polymerization, and decomposition are often facilitated in aqueous medium. Now, it has become clear that enzymes can function in organic systems also as they do in aqueous media.[5,14,108 – 110] With respect to lipases, it is beneficial to employ them in organic solvents. Since many of the reactions, which are responsible for the denaturation of enzymes, are hydrolytic reactions and therefore require water, it is obvious that enzymes should be more stable in an environment of low water content.[111] For example, porcine pancreatic lipase is active for many hours at 1008C in a 99% organic medium but it is rapidly denatured at this temperature when placed in water.[37] Completely anhydrous solvents are not capable of supporting enzymatic activity. Some amount of water is always necessary for the enzyme to retain its


514


native structure responsible for catalysis. The answer to the crucial question concerning how much water is required to retain catalytic activity is enzyme dependent. For example, a-chymotrypsin needs only 50 molecules of water per enzyme molecule to remain catalytically active,[112] which is much less than is required to form a monolayer of water around the enzyme. Other enzymes like subtilisin and various lipases are similar in their requirement for trace quantities of water.[36] In other cases, however, much more water is required: polyphenol oxidase, for example, requires the presence of about 3:5 £ 107 molecules of water.[35] The water present in a biological system can be separated into two physically distinct categories. The majority of the water (.98%) serves as a true solvent (bulk water), whereas a small fraction of it is tightly bound to the enzyme’s surface (bound water). It is the bound water that is crucial for the enzyme structure rather than bulk water. The realization that enzymes can retain, and in some cases improve, their high degree of reaction specificity in nearly anhydrous media has dramatically improved the prospect of employing enzymes in synthetic organic chemistry. From a biotechnological point, there are several advantages in employing enzymes in organic media:[5,109,113] 1. Increased solubility of nonpolar substrates and products. 2. Shifting thermodynamic equilibria to favor ester synthesis over hydrolysis. 3. Yields in organic media are generally better due to omission of tedious extractive procedures during product recovery. 4. Ease of product recovery from low boiling, high vapor pressure solvents. 5. Enzymes can be recovered by simple filtration. 6. Immobilization is often unnecessary because free enzymes are insoluble in organic solvents. If desired, simple adsorption is enough for immobilization. Leaking is not possible in lipophilic environment. 7. Suppression of many water-dependent side reactions such as hydrolysis, racemization, and polymerization. 8. Elimination of microbial contamination. 9. Inhibition of the enzyme caused by lipophilic substrates/products is minimized because their local concentration at enzyme’s surface is low. 10. Possibility of altering substrate- and enantio-selectivities. 11. Enhanced thermostability. Many strategies have been utilized to shift the thermodynamic equilibrium of lipase catalysis: organic solvents of different kinds, water activity repressing agents such as polyols and salts, water-free or almost water-free media such as anhydrous solvents, pure liquid substrates, and the gas phase. Some important applications of lipases in industry and research are summarized in Table 3.


Industry Dairy Natural Oils Bakery, brewery, food Leather Detergent Flavor and fragrance Surfactants Drugs and Pharmaceuticals Polymer

Applications of Lipases in Biotechnology Industry

Process

Product and Application

References

Hydrolysis Transesterification of oils for fats Hydrolysis Hydrolysis Hydrolysis Esterification and transesterification Intramolecular esterification Glycerolysis of fats and oils Acylation of sugar alcohols Resolution of racemic alcohols/esters Polyesterification of diesters with diols Transesterification of monosaccharides

Hydrolysis of milk fat Cocoa butter Improvement of flavor and quality in beverages, meat, fish products Removal of fats from skins Removal of oil stains and lipid spots Synthesis of natural flavor esters Macrocyclic lactones for perfumery Monoacyl glycerols for surfactants Sugar monoacyl esters for surfactants Building blocks for chiral drugs and insecticides Oligomers of alkyols (polyester intermediates) Acrylate esters

[114] [115] [116] [117] [42] [118,29,30] [119] [120] [121] [122 – 124] [125] [126]


Table 3.

515


516


A. Solvent Systems Solvent systems as the reaction media for enzymatic synthesis may be categorized as: (1) aqueous; (2) water: water-miscible (monophasic aqueous–organic system); (3) water: water-immiscible (biphasic aqueous–organic system); (4) nonaqueous (monophasic organic system); (5) anhydrous; (6) supercritical fluids; (7) reversed micelles; (8) solvent-free systems; (9) gas-phase, and (10) ionic liquids.

1. Aqueous Reaction Media A typical lipase-catalyzed reaction in aqueous media is ester hydrolysis. In 1993, about 120,000 ton of monoglycerides and mixtures of mono- and diglycerides were manufactured through glycerolysis of triglycerides.[43] After short-path distillation, monoglycerides (which serve as biocompatible emulsifiers in food, cosmetic, and pharmaceutical preparations) having .90% purity are obtained. Lipase-catalyzed production of di- and monoglycerides by the partial hydrolysis of triglycerides has been researched extensively, but the key problem is formation of mixtures of mono- and diglycerides. Merits and demerits of various enzymatic processes are given in Table 4. In the commercial scenario, none of these procedures has the ability to compete with the chemical route. Although Penicillium roquefortii lipase provided . 90% yield in single step glycerolysis, the economic data were not satisfactory. However, lipases are advantageous for the synthesis of partial glycerides containing labile substituents such as 80 -apo-bcarotinoic acid, which would not withstand chemical procedures.[127] Lipases have also been used to produce soaps and key chemical intermediates like fatty acids and alcohols. Miyoshi Yushi (Nagoya, Japan) is probably the only company in the world, which produces large amounts of soap through lipase-catalyzed hydrolysis of fats and oils with C. rugosa lipase. Nippon Oils & Fats (Tokyo, Japan) is probably the only firm that uses C. rugosa lipase for the preparation of highly pure unsaturated fatty acids (oleic, linoleic, and linolenic, etc.).[128] Due to the tedious raffination steps required to remove thermal degradation products and difficulty in processing heat-sensitive oils such as fish oils via chemical route (which requires 200–3008C and 200–300 bar pressure), oleochemical majors like Unilever, Henkel, Proctor & Gamble and others have explored lipase-catalyzed fat splitting as an alternative to generate the raw material for production of fatty alcohols. However, viable technologies have not yet materialized due to several drawbacks.[43] The addition of lipases to detergent formulations has also been investigated due to their ability to remove fat stains[129] and also due to in situ generation of peracid bleach by perhydrolysis.[130] Novo Nordisk has been the first industry to market Lipolase (a recombinant lipase from H. lanuginosa expressed in A. oryzae ).[42] This was probably the first genetically engineered protein to obtain permission by European regulatory bodies for use in consumer products and be discharged into


Reaction Hydrolysis or alcoholysis Esterification

Merits and Demerits of Monoglyceride Synthesis

Lipase 1,3-Specific lipase Any type of lipase

Reactants Triglyceride Glycerol and fatty acid

Isopropylidene-protected glycerol Glycerolysis and improved process engineering

1,3-Specific lipase

Triglyceride

One step glycerolysis

Highly selective lipase from P. roquefortii

Triglyceride

Remarks Acyl migration may lead to total hydrolysis Results in mixtures of mono- and diglycerides as monoglycerides are better nucleophiles than glycerol Leads to pure monoglycerides, but requires tedious protection and deprotection Use of membrane reactor with counter-current extraction allows separating monoglycerides from reaction mixture. They can also be (cryo)precipitated. Provides good results. But process engineering is complex compared to a chemical process Allows formation of 90% monoglycerides. But the economic data are not satisfactory


Table 4.

517


518


the environment after use. At present, lipases are used in several detergent formulations. The major commercial products are: Lipolase (Novo Nordisk); Lipomax, from Pseudomonas pseudoalcaligenes recloned and expressed in the same organism (Gist-Brocades), Lumafast, from Pseudomonas mendocina cloned and expressed in Bacillus sp. (Genencor). All these products show optimal secondary washing behavior. Cutinase (a long-chain cleaving esterase) exhibit an optimal performance in the removal of wax esters found in lipstick stains. Peracids of a chain length C8 – C12 are good oxidants under slightly alkaline conditions of washing. Owing to their limited stability, however, they cannot be mixed into a detergent formulation. The in situ lipase-mediated generation of peracids has been studied as an alternative. But native lipases exhibit a marginal activity to perhydrolyze esters in presence of water/hydrogen peroxide, probably due to the reason that hydrolysis of the initial peracid-acyl enzyme is faster in alkaline pH than its formation.[130] Nihon Seishi Co., a Japanese paper manufacturer, has developed a process by which the triglycerides of raw lumber are hydrolyzed by lipase, which results in better pitch control rendering easier processing of the lumber to low-grade paper. This process is carried out at a scale of several hundred tons of lumber per day.[131] Ester synthesis has also been performed in aqueous media.[132,133] It has been shown that the 1,3-regiospecific lipase from Candida deformans catalyzed the esterification of free fatty acids in aqueous media while the lipase from Candida parapsilosis could only catalyze alcoholysis (but not esterification) in aqueous media. In addition, the ability of seven lipase preparations to catalyze methyl ester synthesis in aqueous media was investigated and three enzymes (R. miehei, Rhizopus delemar, and C. deformans ) were found to catalyze the esterification.[132] Janssen et al.[133] reported the enzymatic synthesis of carbohydrate esters in aqueous media that, however, gave generally very low ester yields.

2. Water: Water-Miscible Systems The enzyme and the substrate/product are dissolved in a monophasic solution consisting of water and a water-miscible organic co-solvent, such as dimethylsulfoxide (DMSO), dimethyl formamide (DMF), tetrahydrofuran (THF), dioxan, acetone or one of the lower alcohols. Systems of this type are used mainly for the transformation of lipophilic substrates, which are sparingly soluble in an aqueous system alone and which would therefore react at low reaction rates. Buffer solutions are also used in place of water. Butler[96] reviewed the research on water-miscible reaction media. As a rule, most water-miscible solvents can be used up to 10% of the total volume, but in some enzyme/solvent combinations, 50 –70% of co-solvent may be used. If the proportion of the solvent system exceeds this value, the essential bound water is stripped from the enzyme’s surface leading to deactivation. Only rarely do enzymes remain catalytically active in water-miscible organic solvents



519

with extremely low water content. These cases are limited to unusually stable enzymes such as subtilisin and some specific lipases.[134]

3. Water: Water-Immiscible Systems These systems contain two macroscopic phases namely the aqueous phase containing the dissolved enzyme, and a second phase of a nonpolar organic solvent such as hydrocarbons, ethers, or chlorinated hydrocarbons.[135] These systems may be advantageous to achieve a spatial separation of the biocatalyst from organic phase. The biocatalyst is in a favorable aqueous environment and not in direct contact with the organic solvent, where most of the substrate is located. The limited concentrations of organic material in aqueous phase cannot cause enzyme inhibition. Furthermore, the removal of product from the enzyme surface drives the reaction towards completion. In such biphasic systems, the enzymatic reaction occurs only in the aqueous phase. Hence, a mass transfer of the reactants to the enzyme and products from the enzyme and between the two phases is necessary.[136] It is obvious that shaking or stirring represents a crucial parameter in such systems. Enhanced agitation would improve the mass transfer but on the other hand, it may lead to deactivation of the enzyme. Nevertheless, water–organic solvent two-phase systems have been employed successfully to transform highly lipophilic substrates such as steroids,[113] fats,[137] and alkenes.[138] The use of water-immiscible reaction media for enzymatic syntheses has been described and reviewed by Klibanov.[5] Detailed description of these biphasic systems and their implications in reactor design has been presented by Lilly and coworkers.[136,139]

4. Nonaqueous Reaction Media The reaction system is considered nonaqueous if no water more than the absorptive capacity of the reaction medium is added. Owing to the obvious advantages offered by these systems, an increasing interest in the problems of biocatalysis in nonaqueous media has been evident in recent years. Replacing all of the bulk water (which occupies .98% of enzyme environment, in general) by a water-immiscible organic solvent leads to a suspension of the solid enzyme in a monophasic organic solution.[140] Although the biocatalyst seems to be dry, it must have the necessary residual water to remain catalytically active. Such systems proved that they are extremely reliable, versatile, simple, and easy to use. Positive effects from the use of water-immiscible organic solvents may further be enhanced by adding stabilizing compounds to enzyme solution, enzyme modification or immobilization.[27,101,141] Correct solvent selection will influence the efficacy of most enzyme-mediated reactions in nonaqueous systems. Several attempts have been made to facilitate solvent selection by defining a physicochemical parameter that would describe the technical potential of an organic


520


solvent for use as a reaction medium. In order to provide a measure for the hydrophobicity and the compatibility of an organic solvent with high enzyme activity, many parameters such as the Hildebrandt solubility parameter (d ), the dielectric constant (1 ), partition coefficient (P ), and the dipole moment (m ) have been proposed. Of these, the most reliable results were obtained by using the logarithm of the partition coefficient (log P ) of a given solvent between octanol and water.[142] Some of the solvent compatibilities based on log P have been given in Table 5. The log P values not available in the literature can be calculated from the hydrophobic fragmental constants.[143] As can be deduced from Table 5, water-miscible hydrophilic solvents such as DMF, DMSO, acetone, and lower alcohols are usually incompatible, whereas water-immiscible lipophilic solvents such as alkanes, ethers, aromatics, and haloalkanes retain enzyme activity due to the fact that they do not strip-off the essential bound water from enzyme. Further details on different aspects of the use of organic solvents can be found in the literature.[36,109,111,140,144,145]

5. Anhydrous Reaction Media While both nonaqueous and anhydrous reaction media consist of bulk organic phase, the anhydrous reaction systems comprise bulk organic phase and enzymes that have been deliberately dehydrated to remove free water. It is astonishing to know that enzymes display remarkable novel properties when placed in anhydrous solvents. The term anhydrous is taken to mean that the water content is below 0.01%, which is near the sensitivity limit of the classical Karl –Fischer method.[13] It has been firmly established over the last decade that not only do enzymes work actively in anhydrous media but also acquire remarkable properties such as enhanced thermostability,[37,146] radically altered substrate[112] and enantiomeric[134] specificities, molecular memory,[146] and the ability to catalyze unusual reactions.[35] Much of the work reported in these media has been carried out using a-chymotrypsin and subtilisin, while the use of lipase in anhydrous media has been limited.

6. Supercritical Fluids Instead of a lipophilic organic solvent, supercritical gases such as carbon dioxide ðT crit ¼ 318C and Pcrit ¼ 73 barÞ; which exhibit solubility properties similar to that of hexane, can be used as solvent or co-solvent for the enzymatic transformation of lipophilic organic compounds.[147,148] Carbon dioxide and fluoroform can be used as supercritical solvents by compressing them at their critical temperatures. Use of hydrolases in this type of system has been prominent. Different types of reactions were performed using these systems; for example, esterification,[147] transesterification,[149] alcoholysis,[149] and hydrolysis.[150]


Log P

Water-miscibility

Compatibility of Solvents with Enzymatic Activity Examples

2 2.5 to 0

Completely miscible

DMSO

0–2

Partially miscible

Dioxan

2–4

Low miscibility

Chloroform, hexane

.4

Immiscible

Heptane, isooctane

Effect on Enzyme Activity May be used to solubilize lipophilic substrates in concentrations up to 50% v/v without affecting the enzyme Limited use due to rapid enzyme deactivation Causes weak enzyme distortion. May be used with caution, since activity is often unpredictable Causes no enzyme distortion and ensures high retention of activity


Table 5.

521


522


The advantages of these systems are absence of toxicity in comparison with toxic organic solvents, easy product separation, and low viscosity. Some disadvantages are the considerable initial investment for high-pressure equipment and the problem of enzyme deactivation due to depressurization step. Kamat et al.[151] reviewed the use of supercritical fluids for enzyme-catalyzed reactions. 7. Reverse Micellar Systems Micellar enzymology concerns the reactions catalyzed by enzymes entrapped in hydrated reverse micelles of surfactants (detergents, phospholipids) in organic solvents. Reverse micelles form spontaneously when surfactants are dissolved in nonpolar organic solvents. Reverse micelles are closed, almost ˚ in diameter), the outer spherical aggregates of surfactant molecules (15– 20 A shell of which is formed by hydrophobic “tails” of surfactant molecules whereas the inner core is composed of polar “heads” of these molecules. An important property of reverse micelles is their ability to solubilize considerable amounts of water (up to 70 water molecules per surfactant molecule) and other polar compounds (electrolytes in particular). Solubilized water is located in the inner core of micelle and forms a microdroplet separated from the bulk organic solvent by a layer of surfactant molecules. Depending on the amount of water added, the diameter of the inner aqueous cavity of hydrated reverse micelles is of the order ˚ . The properties of this micellar water markedly differ from those of 10 – 100 A bulk water in respect of such parameters as polarity, viscosity, acidity, nucleophilicity, etc. The retention of the enzymatic function in such microheterogeneous media is not surprising in the case of lipolytic enzymes, since the presence of an interface is an obligatory condition for their functioning. The enzyme action in traditional aqueous media is quite different from that in micelles. More detailed information about protein solubilization in reverse micelles has been presented in comprehensive reviews.[152,153] Martinek et al.[4] reviewed micellar enzymology and enzymatic biotransformations. More recently, Stamatis et al.[154] provided an informative review on reverse micellar catalysis with particular reference to lipases. 8. Solvent-Free Systems It is sometimes desirable to eliminate the use of organic solvent, particularly to the synthesis of food-grade flavors and additives. Several advantages are associated with the solvent-free systems. The absence of solvent facilitates downstream processing since fewer components would be present at the end of the reaction, thus minimizing the production cost. Solvent-free system would also permit the use of high substrate concentrations. The solvent-free system could be a reaction mixture solely composed of the substrates (alcohol and acid) in equimolar



523

ratios or alcohol used in large excess. In the latter case, the excess alcohol was used to solubilize the acid substrate and insoluble substances and to enable efficient stirring of the reaction medium. However, problems arise when both substrates are solids and also when mass transfer limitations are associated due to the high viscosity of the substrates. We have carried out solvent-free synthesis of isoamyl esters of short-chain organic acids (acetic, propionic, butyric, and isobutyric) and found that mass transfer was not limiting for short-chain acid substrates. These substrates are highly water-soluble and were found to dissolve in the microaqueous layer surrounding the immobilized lipase and were available for the reactions (unpublished results). However, for long-chain fatty acids like oleic, palmitic, and stearic acids, the problems of mass transfer may be a bottleneck during solvent-free synthesis.

9. Gas-Phase Catalysis Davison et al.[155] discussed several types of vapor-phase enzyme reaction schemes that are theoretically possible and formulated nomenclature and methodologies for nonaqueous biocatalysis. Catalysis of vapor-phase reactions employing whole cell systems (e.g., biofiltration of organic vapors) has been well studied.[156] The majority of these systems, however, are not intended for the synthesis of fine chemicals, but for removal and/or destruction of organic gases and odors. Comparatively, progress for synthetic reactions is lacking. Lamare and Legoy[17] and Parvaresh et al.[157] have investigated lipase-catalyzed transesterification reactions in the gas phase and discussed various characteristics associated with solid – gas biocatalysis. Barton et al.[158] reported vapor-phase enzymatic synthesis of ethyl esters in bench-scale reactors and were successful in achieving considerable yields. However, limitations due to the types of organic vapors used were noticed. In practice, enzyme activity in such systems increased substantially as water activity increased with concomitant decrease in thermostability.

10. Ionic Liquids Ionic liquids (or molten salts) are liquids composed entirely of ions (Sch. 2). They are liquid over a broad range of temperature, possess essentially no vapor pressure, a high ionic conductivity, and a large electrochemical window, and are extremely good solvents for a wide range of organic, inorganic, and polymeric compounds. They are also simple and inexpensive to manufacture and easy to recycle, and their properties can be fine-tuned by changing anion or the R group of the cation ([159] and references cited therein). This unique solvent system combines the advantages of nonaqueous medium while being moderately polar. Cull et al.[160] carried out a two-phase biotransformation in which the ionic


524


Scheme 2. Ionic Liquid: e.g., 1-alkyl-3-methyl imidazolium salt. (If R ¼ Butyl and X ¼ BF4 ; the ionic liquid is water-miscible; if R ¼ Butyl and X ¼ PF6 ; it is water-immiscible).

liquid acts as substrate reservoir and the whole cell biocatalyst (Rhodococcus R312) is present in the aqueous phase. Erbeldinger et al.[161] reported the thermolysin-catalyzed synthesis of Z-aspartame in ionic liquids containing 5% water. Madeira Lau et al.[159] reported the first example of the use of a free enzyme in an ionic liquid in the absence of water. Lipase of C. antarctica has been shown to catalyze alcoholysis, ammoniolysis, and perhydrolysis reactions in this media. Reaction rates were observed to be comparable with or better than those obtained in organic solvents. It is envisaged that ionic liquids could have benefits for performing biotransformations with highly polar substrates like carbohydrates, which are only sparingly soluble in common organic solvents. Recently, Kim et al.[162] reported markedly (25 times) enhanced enantioselectivity of lipase in ionic liquids.

B. Water Activity: A Key Parameter in Lipase Catalysis Water plays several roles in the enzyme structure and function:[163] (a) action on enzyme structure by contributing to all noncovalent bonding, (b) alteration of protein structure by disruption of hydrogen bonds, (c) facilitation of reagent diffusion, and (d) influencing in the equilibrium conversion (as substrate or product).

1. Role of Water and Water Activity In enzymatic esterifications, water is formed during the reaction, which often leads to a lag phase where the amount of water is too low to support the reaction.[164] This will be followed by a period where most of the reaction takes place, after which the reaction rate decreases again, due to accumulation of water, favoring hydrolysis. During a transesterification reaction, this is not a problem since water is not formed during acyl transfer. However, the equilibrium will be influenced in this case also due to the reverse reaction, although not a hydrolysis. If the water level is too high, rates can also be reduced because of the tendency of the suspended enzyme particles to aggregate together, leading to diffusional limitations. The amount of water giving near optimal condition is often in quite



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a narrow range. It has been reported that less than a monolayer of water is necessary to retain the activity of a-chymotrypsin and subtilisin.[36] To quantify the amount of water present in the reaction mixture, the thermodynamic water activity (aw) is used. The water activity governs the degree of hydration of the enzyme and gives a direct indication of the mass action of water. Water activity (aw) may be defined as the ratio of the water vapor pressure over the substance ( p ) to that over pure water ( p0): aw ¼ p=p0 :[165]

2. Determination of Water Activity From the relationship between aw and relative humidity, one can estimate aw in a solution or a material by measuring the relative humidity in a confined space, which is in equilibrium with the solution or material. This is done with hygrometers, which can be built[166] or purchased commercially.[167] It is important to calibrate a hygrometer carefully using saturated solutions of standard salts.[168] There are, however, other methods to estimate aw in a sample using the partial pressure of water vapor.[169] However, these may not be feasible for estimation of aw in organic solvents. Halling[170] described a new method to measure aw in organic solvents. Mathematical methods to determine aw have also been developed and used.[171]

3. Effect of Water Activity on Lipase Catalysis Water activity influences not only reaction rate, but also enantioselectivity[172] and equilibrium conversion.[173] Valivety et al.[174] studied the kinetics of esterification catalyzed by lipases from R. miehei and C. rugosa and established the fact that water also acts as a competitive inhibitor for both the alcohol and acid. Lee and Kim[175] investigated the effect of aw on enzyme hydration and reaction rate in organic solvents and developed a theoretical kinetic model,[176] which was found to be in good agreement with the experimental observations. The relationship between water and protein stability has been the subject of numerous scientific reports. There are many ways to depict or picture the hydration of a protein molecule. One that has proved to be most useful both for interpreting the denaturation of proteins and the activity of enzymes, is the water adsorption isotherm, which describes the relationship between water content and aw.[166,177] For most globular proteins, it is a sigmoidal curve, with one break point at aw ¼ 0:2 and another at aw ¼ 0:8: The region of low aw has been described as a tightly bound monolayer of water molecules on the protein surface, the intermediate region as a bi- and multi-molecular layer, and the region of aw . 0:8 as water condensed in capillaries.[178] A aw of 0.8 corresponds to a water content of ,25% w/w, at which point the sample still appears rather dry.


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4. Control of Water Activity Taking into account the role of aw in esterification reactions, several techniques have been employed to arrive at near optimal conditions for regulating aw. Enzymatic reactions carried out with initially adjusted aw (by preequilibrating the reactant solution and enzyme preparation with the water vapor of saturated salt solutions) were first described by Goderis et al.[39] However, if water is produced or consumed in the reaction, aw will change, which can cause adverse effects. One way to control aw during a reaction is by continuously adjusting aw in the headspace above the reaction mixture.[179] Another possibility is to have a vessel with a saturated salt solution in contact with the reaction mixture, via the gas phase, so that the salt solution continuously absorbs or releases water vapor to keep the aw constant. Kuhl and Halling[180] have demonstrated the use of pairs of salt hydrates to control aw during the chymotrypsin-catalyzed peptide synthesis. The salt hydrates present in the reaction mixture are able to control the aw by absorbing or releasing water molecules as the need arises and thereby act as buffers of water activity. Kvittingen et al.[164] reported the successful use of salt hydrates to maintain optimal aw in a lipase-catalyzed esterification of butyric acid with butanol. Halling[181] compiled the data on various salt hydrates of possible application for aw control in organic media. The exchange of water is rather slow with many salt hydrates and the system is limited to salts that are nontoxic. Enzyme reuse also becomes difficult because the enzyme and the solid salt are to be separated. Wehtje et al.[182] and Svensson et al.[183] passed the salt solution through a silicone rubber tube immersed in the reaction medium containing the immobilized enzyme to continuously adjust the aw of the reaction medium (Fig. 1).

Figure 1. Experimental set-up of the water activity-controlling device: (1) silicone tubing, (2) reaction medium, (3) enzyme preparation, and (4) magnetic bar (from Ref. [183]).



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5. Regulation of Water Activity Through Water Removal Use of Molecular Sieve Water generated during the reaction can be removed by adding molecular sieves.[184,185] Ibrahim et al.[186] investigated continuous synthesis of geranyl esters in a packed column reactor attached to a molecular sieve column in series. Under the optimum water-controlled conditions, they were able to achieve nearly 100% conversion of geranyl laurate. The disadvantage in using molecular sieve is that it needs reactivation (at 2508C) before reuse.

Pervaporation Pervaporation is a promising technique for water removal involving the transport of liquid through a homogeneous, nonporous membrane with simultaneous evaporation of permeates.[187] Transport phenomena in pervaporation are more complex compared to other membrane processes, involving a threestep process: sorption into the membrane at the upstream side, diffusion through the membrane, and desorption into the vapor phase at the downstream side. Van der Padt et al.[188] applied pervaporation technique, for the first time, for enzymatic synthesis of triglycerides, using a porous cellulose acetate hollow-fiber membrane but it was difficult to scale up and the enzyme immobilization step was time consuming. Okamoto et al.[189] reported pervaporation-aided esterification of oleic acid with ethanol using p-toluene sulfonic acid as the catalyst. Both the studies showed that membrane separation of water produced from the reaction mixtures favorably shifts the equilibrium towards synthesis. Kwon et al.[190] successfully applied the pervaporation technique by using a nonporous polymeric membrane (cellulose acetate) for the lipase-catalyzed esterification of oleic acid with n-butanol in isooctane and found that pervaporation selectively separated water from the reaction mixture.

Removal of Water by Evacuation There are a significant number of systems, which involve the use of liquid substrates that are nonvolatile compared with water. In such systems, the possibility of shifting the equilibrium towards synthesis by partially evacuating the headspace above the liquid reaction mixture has been demonstrated by several workers.[15,102,191,192] Water removal in organic phase enzymatic reactions has also been done by continuously sparging nitrogen through the reaction medium[193] and the gas was reused after being stripped of its water content in an adsorber.


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Reactor Designs for Water Activity Regulation While the techniques mentioned earlier can be used to favor reverse hydrolytic reactions, they have some disadvantages. The pervaporation technique can be used for continuous aw control but would be too expensive on large scale since it requires a partial vacuum and the use of a condenser for water removal. Salt hydrates have a limited buffering capacity. Consequently, they require frequent regeneration, which restricts their use on an industrial scale. Unlike sparging (of air or nitrogen), which can only be operated as a batch process, the use of saturated salt solutions can be operated in continuous mode since the salt solution can be recirculated after concentration or resaturation to make up for the dilution. However, such a reactor configuration described by Wehtje et al.[182] may not be suitable for use on a large-scale continuous operation.[194] The utility of their reactor design is also limited by the poor permeability of the silicone rubber tubing used for recirculation of the saturated salt solution. Recently, a twin-core packed-bed reactor[195] and a packed-bed hollow fiber reactor[194] incorporating salt hydrate pairs and salt solution, respectively, have been reported. The former reactor consisted of an easily removable inner core of salt hydrate that was separated from an outer core of immobilized lipase. Complete conversions were obtained using this design. In the latter case, a membrane physically separated the enzyme and salt solution: the immobilized enzyme being placed on the shell side (outside) while the salt solution was circulated on the lumen side (inside). Salt solution (diluted by the water of reaction) was resaturated by passing it through a bed of salt. This, in principle, is an extension of the idea suggested by Wehtje et al.[182] The limitation of the reactor developed by Rosell et al.[194] was that aw control at low levels could be rendered difficult by the high viscosity of the saturated salt solutions. This problem, however, can be partially overcome by properly selecting hollow-fiber membranes. The reactor design was amenable to scale up.

IV.

LIPASE-CATALYZED ESTERIFICATION REACTIONS

Research on lipase-catalyzed production of various kinds of esters has increased tremendously in the recent past. Esters are present in fats and oils, and natural and synthetic polymers. They are useful intermediates or end products in the chemical industry. Although the enzymatic synthesis of esters has been studied for many years with crude preparations of pancreatic and ricinus lipases, whether one enzyme catalyzes both the hydrolytic and synthetic reactions remained unconfirmed until the early 1960s. Fukumoto et al.[196] for the first time purified the lipase of A. niger to homogeneity and demonstrated that glyceride synthesis was effected by the reversal of hydrolysis. The same group later reported glyceride[33] and aliphatic



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ester[197] synthesis by lipases to further prove the occurrence of the reverse reaction. Lipases from A. niger, R. delemar, G. candidum, and Penicillium cyclopium were found to synthesize esters of oleic acid and various primary alcohols. However, only G. candidum lipase synthesized esters of secondary alcohols. Esters of tertiary alcohols, phenols, or sugar alcohols were not synthesized by any lipase. These reactions were performed without any solvent. However, in the later stages, research was directed towards use of organic solvents for ester synthesis reactions.[5] The application of lipases for synthetic purposes is now well documented. Enzymatic production of esters can be achieved either by reaction between free acid and hydroxyl groups of alcohol or by ester exchange or transesterification (includes alcoholysis, acidolysis, and interesterification).[77] Lipases in two different physical states, solution or solid, were used in nonaqueous media for ester synthesis. As a solution, they were used in the native form or after modification with amphiphilic molecules to improve their solubility in organic media. Solid enzymes were used in the form of precipitates, lyophilizates, crosslinked crystals, or immobilized particles. Lipase-catalyzed esterification reactions have been actively pursued to produce various kinds of commercially important esters. Esters of short-chain alcohols and short-chain fatty acids are extremely important aroma compounds.[198] Esters of short-chain alcohols and long-chain fatty acids are valuable oleochemicals that may be used as lubricants, diesel fuel, and antistatic reagents.[199] Esters of long-chain fatty acids and polyhydric alcohols like glycerol, sorbitol, and other carbohydrates (called—emulsifiers/surfactants) find immense application in food and pharmaceutical industries.[15] Some of the important categories of lipase-catalyzed esterification reactions are discussed below.

A. Modification of Fats and Oils Lipases have many possible uses in the treatment and modification of fats and oils.[200] Some of the important esters are the glycerides present in the fats and oils used in food and other industries. Many lipases exhibit sn-1,3 specificity and can hence be used to regioselectively (inter)esterify natural triglyceride. However, the acyl migration from sn-2 to the sn-1 or sn-3 positions must be suppressed. By exchanging some fatty acids (totally or in part) of the triglycerides of fats and oils with those from a different oil, it is possible to modify the physico-chemical and nutritional properties and palatability of the starting material.[201] Processes employing solvent-free media[202] as well as organic solvents[203] have been studied. Three types of modified triglycerides have gained commercial importance: (i) cocoa butter equivalents, (ii) fats with improved spreadability, and (iii) structured lipids or highly digestive triglycerides.


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1. Cocoa Butter Equivalents Cocoa butter has a melting point around human body temperature (378C) and therefore is well suited as a matrix for suppositories. It also has a major application in the production of chocolates, for which rapid melting is desirable for mouth feel. Several tons of cocoa butter were imported from tropical countries like Kenya and Malaysia. Major efforts have been made to produce a substitute for cocoa butter using cheaper vegetable oils.[115] In this regard, the selectivity of most lipases for the 1 and 3 positions of glycerol is advantageous, because most triglycerides present in cocoa butter have stearic or palmitic acid at positions 1 and 3 and oleic acid strictly at position 2 (SOS or SOP), and chemical interesterification is not selective.[204] Cocoa butter equivalents can be produced through lipase-catalyzed interesterification of suitable natural triglycerides, such as middle fraction of palm oil (containing palmitic acid moiety at 1 and 3 positions and oleic acid at 2, POP) or sunflower oil (having a high content of oleic acid, OOO) with stearic acid or tristearin (SSS). The enzyme’s specificity can be hampered by acyl migration and it is possible to esterify glycerol and free fatty acids using a 1,3-specific lipase.[179] Uniqema (a former subsidiary of the Unilever group and recently acquired by ICI) produces 100 ton of chocolate fat (SOS) by R. miehei lipase-catalyzed transesterification of high oleic sunflower oil with stearic acid in a solvent-free packed-bed reactor.[205] Fuji Oil Co. of Japan has also established similar process using Rhizopus lipase with a production capacity of several thousand tons per annum.[206]

2. Fats with Improved Spreadability The melting point of an oil can be modulated by the degree of catalytic hydrogenation of double bonds present in unsaturated fatty acids. This is carried out on a large scale for the preparation of margarines and shortenings from plant oils.[207] Alternatively, the desired melting behavior can be achieved via sn-1,3specific lipase-catalyzed interesterification of suitable triglyceride mixtures or a combination of both procedures.[208,209]

3. Structured Lipids (Highly Digestive Triglycerides) Another interesting class of products synthesized by lipase catalysis is the structured lipids[209] that can serve diverse purposes viz.: AAA, ABA, AAB, and ABC types. Among these, AAA type can be rather easily synthesized by (chemical or enzymatic) esterification of glycerol and fatty acid, which usually results in almost complete conversions even at stoichiometric proportions of the substrates. Other types of triglycerides require regiospecific reactions, for which positionally



531

specific lipases are essential. ABA-type structured lipids can be synthesized by sn-1,3-specific lipase-catalyzed reactions: (i) interesterification of two different triglycerides, (ii) acyl exchange between triglyceride and fatty acid or with fatty acid ethyl ester, or (iii) acylation of glycerol with fatty acid giving symmetric 1,3-diacyl-sn-glycerol, followed by chemical acylation at sn-2 position (since there is no sn-2-specific lipase available, this step cannot be performed with ordinary lipases). AAB-type structured lipids are prepared by mono-substitution at either sn-1 or sn-3 positions of triglyceride with fatty acid or its ester, avoiding the formation of di-substituted by-product. Stereopreference of certain lipases to sn-1 over sn-3 position enables the syntheses of chiral AAB- and ABC-type triglycerides. One example of ABA-type structured lipids are the ones in which medium-chain fatty acids (M) are present at 1 and 3, and long-chain saturated or unsaturated ones (L) at 2. These are easier to digest and can still provide essential fatty acids and hence especially suited for infant food formulations and for patients with absorption (in the body) problems. Triglycerides of MLM type provide rapid delivery of energy through enhanced hydrolysis and absorption; pancreatic lipase hydrolyzes medium-chain triglycerides preferentially resulting in monoglycerides that are efficiently absorbed from intestine. Several products of this type are commercially available, e.g., Caprenin (C6 – C22:0 – C8) of Procter & Gamble (Cincinnati, OH, USA), but are chemically synthesized. A lipase-derived commercial product of this kind, Betapol (1,3oleoyl-2-palmitoyl glycerol, OPO or mimic of human milk fat), is marketed by Loders Croklaan for infant diet. OPO is produced in industry by lipasecatalyzed acidolysis of tripalmitin with oleic acid and mixed triglycerides are separated by physical methods.[210] In order to achieve a high yield of OPO, Schmid et al.[211] investigated a two-step process involving (i) alcoholysis of tripalmitin using sn-1,3 lipase resulting in 2-monopalmitin (2-MP), and (ii) esterification of 2-MP with oleic acid using the same lipase. The best results were obtained with immobilized lipases from R. miehei and R. delemar. The OPO produced by this two-step process has been observed to be much purer (, 96% palmitic acid at sn-2 position) compared to commercial Betapol that contains only 65% palmitic acid at sn-2 position. A drawback of this method is that it requires recrystallization of 2-MP at low temperature, and hence cannot be applied to polyunsaturated fatty acids (PUFA) containing 2-monoglyceride, which possess very low melting points. Triglycerides of MLM type enriched in essential long-chain fatty acids of fish oils (eicosapentaenoic or docosahexaenoic acid) are helpful in preventing coronary heart diseases and inflammatory diseases.[212] In these reactions, lipase specificity towards (especially) long-chain v-3-polyunsaturated ones is beneficial. Triglycerides of this composition are preferentially produced by means of sn-1,3specific lipases since chemical interesterification of highly unsaturated triglycerides promotes side reactions like oxidation, cis– trans isomerization, or migration of the double bond.


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Other possible applications of lipase interesterification are in facilitating an easy handling of palm oil by reacting with canola or soybean oil to decrease the low temperature viscosity of the oil.[213] Lipase-catalyzed esterification can also be applied to enzymatic refining of hyperacid oils by reacting their free fatty acids with or without added glycerol.[214] Enzymatic modification of vegetable oils has already been implemented at the industrial scale: Loders Croklaan has built a plant for the purpose at Wormeveer, Netherlands.[210] This subject has been reviewed extensively by Adlercreutz,[215] Akoh,[216] Bornscheuer,[217,218] Iwasaki and Yamane,[209] Malcata et al.,[219] and Quinlan and Moore.[210]

B. Wax Esters Methyl and ethyl esters of long-chain acids provide valuable oleochemical species that may function as diesel fuel.[199] On the other hand, long-chain esters (those derived from alcohols and acids with a carbon chain C12 or more), which are typically referred to as waxes, have potential applications from lubricants to cosmetics. Natural wax esters presently in use are obtained only from sources such as jojoba seed (Simmondsia chinensis ), carnauba (Copernicia cerifera ), and sperm whale. Attempts to produce substitutes of these premium wax esters have been reported in the literature.[220 – 222] It is possible to synthesize wax esters that can be used in personal care products through direct synthesis from free fatty acids and long-chain alcohols or alcoholysis of triglycerides in the reaction mixtures composed of substrates alone.[220] Research on the production of biodegradable, environmentally acceptable esters for biodiesel, lubricants, solvents, surface-active agents from vegetable oils through lipase catalysis has markedly increased during the recent past.[223] For example, butyl oleate may be used as biodiesel additive, PVC plasticizer, water resisting agent, and in hydraulic fluids.[224] Rapeseed oil fatty acid esters of 2-ethyl-1-hexanol can be a viable replacement to conventional organic solvents in a number of detergent applications (e.g., in shampoos for car cleaning) and as a solvent for printing ink.[225] Biodegradable lubricants, developed in the 1980s for 2-stroke engines, consisted of neopentylpolyol esters of branched chain fatty acids as the main base fluid. Eychenne et al.[226] recently reviewed the developments in the production of lubricating oils based on neopentylpolyols such as neopentylglycol, pentacrythritol, and trimethylolpropane. In the mid 1980s, biodegradable chain-saw oils based on natural esters of rapeseed oils were introduced into the market.[225] Biodegradable trimethylolpropane esters of fatty acids from sunflower oil or rapeseed oil can be utilized for the production of hydraulic fluids. Further, these esters were also developed as lubricants for jet turbine, motorcar, and gas turbine engines.[225] Lipase-catalyzed esterification and transesterification have been employed for the production of biodegradable solvents[227] and polymers.[223] Schuch and



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Mukherjee[228] showed that lipase-catalyzed alcoholysis of triacylglycerols with long-chain alcohols yielding wax esters is by far the fastest of the interesterification reactions. Further, Mukherjee and Kiewitt[229] reported lipasecatalyzed reactions for the preparation of natural resembling wax esters.

C.

Surfactant Esters

This group mainly contains esters of sugar alcohols. Monoesters of fatty acids and sugars or sugar alcohols are a very important group of nonionic surfactants with potential applications in many industries. These amphiphilic molecules have very good emulsifying, stabilizing, or conditioning effects. Traditional high-temperature chemical esterification to produce these esters leads to coloration of the final product and, dehydration and cyclization in the case of sugar alcohols. These problems can be overcome by the use of enzymes under milder conditions for the synthesis of these esters.[134] Janssen et al.[133] reported the synthesis of sugar esters in aqueous media, but could achieve only a low yield. Klibanov’s group synthesized carbohydrate esters in organic solvents (such as pyridine, which however, are toxic and may not be compatible with food application) in which both sugars and fatty acids are soluble.[121] The surfactant industry, with an annual production of 3 million tons, worth US$ 4 million, is facing increased consumer demand for biocompatible and biodegradable products.[230,231] Saccharide-fatty acid esters, produced from inexpensive agricultural feedstock meet these conditions and also have excellent surface-active properties.[230] Biosurfactants have been produced by enzyme catalysis from a variety of monosaccharides (e.g., fructose, galactose, xylose, and glucose) and sugar alcohols (e.g., sorbitol) ([231] and references cited therein). In recent times, disaccharide esters (e.g., those of sucrose) have received immense attention due to their higher solubility in water compared to monosaccharide esters.[232] But, successful synthesis of sucrose esters is not yet reported in the literature. Engineering of the reaction medium is especially challenging for reactions involving di- and polysaccharide substrates due to their low solubility in organic solvents and high hygroscopicity. However, in most of the solvents that yield high enzymatic activity (i.e., hydrophobic), solubility of saccharides is very poor. Various strategies (disadvantages in parentheses) employed to increase saccharide solubility in hydrophobic solvents include:[231,233] use of aqueous– nonaqueous two-phase liquid systems (reaction and mass transfer rates are slow); use of saccharide ethers or glycopyranosides (ester properties are changed by the presence of an ether group); derivatization with isopropylidene, a known ZOH group blocking agent, or complex formation using phenylboronic acid (expensive and requires tedious downstream processing to remove derivatives); suspension of silica gel-containing adsorbed fructose in acyl donor-containing liquid phase (difficult to separate silica gel and enzyme);[183] and


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use of polar or polar –nonpolar cosolvent mixtures at reflux (refluxing cosolvents at large scale may pose safety problems). Recently, the importance of controlling or engineering the esterification reaction conditions in such systems has been investigated. Cao et al.[234] demonstrated the accelerated esterification of saccharide-fatty acid by crystallizing the monoester product at subambient temperature by employing a solvent that enhanced or hindered the solubilization of saccharides and monoesters, respectively. Alternatively, the composition of the reaction medium can be altered during the progress of reaction to enhance the rate and extent of esterification. Moreover, polyol solubility often increases greatly during the reaction even under solvent-free conditions due to the formation of mono- and diesters.[235] Zhang and Hayes[233] investigated this important aspect for enhancement of rate of saccharide esterification by medium engineering and observed 10 – 100 times higher conversions compared to the earlier reported yields. Chemically prepared alkyl glucosides, which are now being produced on an industrial scale for nonfood applications, show properties similar to the sugar esters and hence are major competitors for the same. Uniqema has commercialized lipase-catalyzed production of monoesters of ethylglycoside on a 100-kg scale.

D. Organic Synthesis Chirality plays a crucial role in nature. The synthesis of chirally pure materials is an important and challenging task and the preparation of enantiomerically pure compounds has received as much attention, in the last two decades. Enantiomers of a given molecule are different entities in terms of their biological action, especially pharmacological properties. Chiral purity of drug molecules is therefore warranted essential. The FDA has issued guidelines concerning the development of drugs having chiral centers.[236] The FDA does not prohibit the marketing of racemates, but the choice of a racemic synthesis over the development of a chirally pure drug must be justified. It also requires investigations on the bioavailability and pharmacological data of the chiral drug and the final approval is based on complete background information on each enantiomer and the racemic mixture. In this scenario, the fate of already marketed racemic drugs and those under development seems to be uncertain. There is an increasing demand for preparing optically active pure compounds in pharmaceutical industry and also considerable interest in remarketing the former racemic drugs in enantiopure form. Although many nonenzymatic methods have emerged for enantioselective synthesis, biotransformations are likely to play an increasingly important role to meet the challenge posed to organic chemists. For almost every kind of chemical reaction, there exists an enzyme-catalyzed equivalent. Enzymes are known to mediate a variety of reactions (e.g., oxidation, reduction, hydrolysis, condensation, and isomerization) including CZC and CZX (hetero atom) bond formation.



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Among various methods available, kinetic resolution of racemates or asymmetric synthesis from prochiral precursors (both enzymatic) are attractive.[237] Lipases have found major attention of chemists as the most versatile class of biocatalysts for this purpose, because they can accommodate a wide variety of substrates, natural or synthetic, and still exhibit chiral recognition or selectivity. Lipases, by nature, are evolved to withstand denaturing interface and also organic solvents. Due to the reason that acyl-lipase formed in the first step of the reaction can be considered as an acylating agent, a wide range of substrate specificity is observed for lipases. Therefore, hydroperoxides, amines, and thiols also become nucleophilic substrates for lipases in addition to those containing the hydroxyl group. As a result of several unique properties, a plethora of synthetic reactions have been carried out and about one-third of all biotransformations are performed with lipases.[24] Lipases are known to exhibit regio- and/or stereoselectivity while catalyzing reactions of polyfunctional compounds, isomers, or racemic or prostereogenic substrates. Regio- and stereoselective reactions involving triglycerides have been outlined under Section IV.A. Therefore, only reactions involving nonglyceride substrates have been described in this section. Most asymmetrization substrates bear common functionalities such as alcohols, ketones, and esters. As natural substrates of lipases are esters of a prochiral alcohol with an achiral acid, the main substrates of lipase catalysis would be esters of chiral alcohols (Type I substrates as per Faber’s denomination).[24] Nevertheless, some lipases can also recognize the chirality in the acid moiety of the ester (Type II substrates). However, in order to maximize the use of lipases as tools in synthetic chemistry, it is essential to broaden the choice of functional classes as effective substrates. Two main strategies, hydrolysis or acyl transfer, can be used for the resolution of racemic mixtures. Generally lipases operate at the same prochiral center regardless of media, and hence it is possible that one can choose between pathways of enantiospecific hydrolysis or synthesis. A typical regioselective reaction (catalyzed by porcine pancreatic lipase) and an enantioselective transesterification (catalyzed by R. miehei lipase) are depicted in Schs. 3 and 4, respectively, both involving nonglyceride substrates. 1. Kinetic Resolution of Chiral Compounds Kinetic resolution of racemates can be carried out using lipases with a theoretical yield of 50%. The first generation processes were based on the hydrolysis of derivatives such as acylated amines or essential alcohols and acids.

Scheme 3. Regioselective esterification by porcine pancreatic lipase (83% yield, 99% monoacetate is obtained).


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Scheme 4. Enantioselective transesterification by R. miehei. E ¼ 206 (acylation), E ¼ 8 (hydrolysis) (from Ref. [43]).

In recent years, however, enantioselective synthesis and transesterification reactions in organic media have been extensively studied[122] as they offer many advantages including: (i) solubility problems can be overcome more easily than in aqueous solution, (ii) the number of steps can be reduced (since prior derivatization is not necessary). Racemic forms of 2-aryl propionic acid, commonly referred to as profens (an important group of nonsteroidal anti-inflammatory drugs, NSAIDs), are best examples where the racemate produced side effects. The anti-inflammatory and analgesic effects of the profens are attributed almost exclusively to the S-enantiomer. Thus, it is necessary to resolve the racemic mixture to avoid any possible side effects. Lipases have been successfully used in the resolution of profens.[238] The profens were either converted to esters chemically and then enantioselectively hydrolyzed to the active S-acid through biocatalysis or the active S-form was converted to S-ester in organic media using lipases followed by chemical hydrolysis of S-ester to S-acid. Duan et al.[238] investigated enantioselective esterification of ketoprofen with n-propanol catalyzed by an immobilized lipase from C. antarctica (Novozyme 435 of Novo Nordisk, Denmark). They have reported a single step process for the same by esterifying the R-isomer of ketoprofen faster than that of the S-isomer resulting in the formation of R-ester and S-acid in the product stream. However, it is a known fact that lipase-catalyzed reactions are usually reversible and the reverse reaction should be suppressed to avoid racemization. It is also important to drive the reaction equilibrium in the desired direction. Thus, water generated during esterification should be removed by vacuum or by using adsorbents. Alternatively, a large excess of acyl donor (e.g., ethyl acetate) can be employed instead of a solvent (solvent-free system). Trifluoro- or trichloroethyl acetate and cyclic anhydrides (representing good leaving groups) have been employed successfully in such systems. Irreversible transesterification reagents such as oxime esters may also drive the equilibrium in the desired direction (i.e., ester formation). The most widely accepted choice is the use of enol esters (vinyl



537

Scheme 5. Irreversible acylation (transesterification) with enol esters (from Ref. [43]).

or isopropenyl ester) and the primary alcohol formed during hydrolysis tautomerizes to result in a carbonyl compound, and thus becoming unavailable for the reverse reaction (Sch. 5).[239,240] Theoretical yields are limited to 50% in the kinetic resolutions, since the other racemic form is not utilized in the reaction. By using the meso-diesters or diol reagents, which undergo enantioselective hydrolysis or transesterification through enantiotropic group differentiation, yields up to 100% are attainable; this phenomenon is termed the “Meso Trick” (Sch. 6).

Type I Substrates (Chiral Esters Possessing Stereocentered Alcohol Moiety ) Several examples are available for this type of substrates in the literature. However, the practical use of this kind of reactions can be reflected very well in the processes to obtain precursors of homochiral drugs such as the antiviral agent Lamivudine[241] or the antifungal Coriolic acid,[242] or the production of enantiopure C3 compounds.[243] Better results can be obtained if the substrate is prochiral as in case of synthesizing cyclopentanoids having a variety of biological activities as analogs of prostaglandins, prostacyclins, and thromboxanes.[244]

Type II Substrates (Chiral Esters Possessing Stereocentered Acid Moiety ) The biocatalytic kinetic resolution of chiral racemic acids is generally carried out in hydrolytic mode, because the rate of hydrolytic reaction in water is generally faster than that of the esterification in organic media. Most employed lipases for enantiomeric hydrolysis of carboxylic acid esters and/or lactones are those from C. rugosa and porcine pancreas. The resolution of several chiral acids

Scheme 6.

Application of the Meso Trick (from Ref. [43]).


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has been performed by hydrolytic approach, but good E-values are obtained except for the resolution of ketorolac, an anti-inflammatory agent.[245]

2. Selective Acylation The serine residue of the active site is acylated in the first step of lipase catalysis followed by nucleophilic attack. Nucleophile may be water (hydrolysis) or an alcohol (esterification). Apart from these, several other groups of compounds also act as nucleophiles. Several reactions of industrial relevance have been investigated using lipase-catalysis which are discussed as follows:

Resolution of Racemic Alcohols and Acids Majority of the publications on lipase-catalyzed reactions deal with this kind of reactions.[24,43] Although there are several examples described in the literature, only few of these processes have found industrial applications. There are several important reasons for this: (i) In a number of cases (e.g., pyrethroids, b-blockers), no undesirable side effects have been found for the wrong enantiomer, and hence the racemic product can be used as such. (ii) Chemical methods such as preferential crystallization (naproxen, L -menthol), catalytic asymmetric synthesis (Diltiazem), or fermentative procedure or synthons from the chirality pool (aphenoxy-propionic acid herbicides) are more economical. (iii) The need to racemize the wrong enantiomer is a major drawback. While this is feasible in most cases, it will be an economic burden on the process.[43] While the use of prostereogenic or meso precursors is a principal solution to this problem, it is often not possible to define such precursors for a given synthetic problem. Despite these hurdles, there are at least two industrial lipase-catalyzed processes: the preparation of Diltiazem by Tanabe Seiyaku Co. (Tokyo, Japan) and the preparation of several chiral amines by BASF. Some examples of this category have been described below under Type I and Type II categories:

Type I Substrates Several lipases catalyze the acylation of prochiral diols or racemic alcohols. The resolution can be carried out using either a carboxylic acid or an acid derivative (ester or anhydride) as an acyl donor. Kinetic resolution of several primary and secondary alcohols using hexanoic and octanoic acids as acyl donors is well reported in the literature.[24] To avoid the reversibility of the esterification and concomitant depletion of optical purity of the remaining substrate, the best solution is the use of special acyl donors that ensure a more or less irreversible type of reaction. Although it is



539

helpful in some cases to employ excess acyl donor, it would not be economically feasible and it is not always compatible with the high enzyme activity. The most common methodologies to shift the equilibrium towards the desired product are to employ enol esters (vinyl esters, particularly vinyl acetate) and anhydrides, and secondly the use of activated esters such as 20 -haloethyl, cyanomethyl, and oxime esters (resulting in reduced nucleophilicity of the leaving groups due to the presence of electron withdrawing substituents). Bevinakatti and Newadkar[246] described the resolution of N-ethoxy carbonyl-2-amino-1-butanol (precursor of Ethambutol, an antitubercular drug) using a lactone (2-phenyl oxazolin-5-one), which is a rarely used acyl transfer agent. It is also not usual to perform enantioselective acylation of racemic alcohols using uncommon vinyl cinnamate or cinnamic anhydride,[247] and this example is also peculiar because it constitutes the only case of failure of the empirical “Kazlauskas rule.”[248] Compounds possessing hydroxyl groups at specific positions (e.g., sobrerol, chloramphenicol, or oligopeptide esters) have been selectively acylated using porcine pancreatic lipase.[24] Sugars are the important examples of polyhydroxy compounds and numerous studies have been directed to their regioselective acylation or deacylation by lipases.

Type II Substrates There are numerous examples of lipase-catalyzed resolution of chiral acids via enantioselective esterification or transesterification.[24] R. miehei lipasecatalyzed resolution of methyl trans-b-phenyl glycidate results in the (2S, 3R ) compound with better yields and selectivities compared to other lipases.[249] The authors used a more sterically hindered alcohol (isobutanol) and increased the alcohol concentration to render the reverse reaction virtually negligible. The most employed substrates for resolution are 2-arylpropionic acids (the profen family), an important group of NSAIDs. Best resolutions are described by transesterification of racemic ketoprofen with n-butanol or 2-pyridyl ethanol[250] and by esterification of racemic ibuprofen with n-butanol.[251]

Acylation of Stereoisomers Lipases (e.g., porcine pancreatic lipase) exhibited a preference to E isomer of allylic alcohols in which E/Z stereoisomers are present. In most cases, unfortunately, selectivities are only modest. Morgan et al.[252] reported the separation of a mixture of (E )-geraniol and (Z )-nerol by acylation with trifluoroethyl butyrate or hexanoic anhydride in diethyl ether in presence of porcine pancreatic lipase, in which geraniol was acylated four times faster. The E isomers of secondary allylic alcohols such as 3-undecen-2-ol or 4-phenyl-3-buten2-ol, however, reacted 20 – 40 times faster than the Z isomers under similar


540


Scheme 7. Asymmetrization of prostereogenic diester [1,4-dihydro-4-(3-nitrophenyl)-2,6dimethyl-pyridine dicarboxylate ðR2 ¼ CH2 OCOCH3 Þ]: (a) R1 ¼ CH2 OCH3 ; Lipase AK, 63% (S )-monoester, 95% ee; R1 ¼ CH2 Ph; Candida OF-360, 24% (R )-monoester, 73% ee (from Ref. [253]).

conditions. In addition, the S enantiomer of the E isomer does not participate in the reaction, due to high enantioselective nature of this enzymatic transesterification ðE . 100Þ:

Asymmetrization of Meso-diols and Prostereogenic Alcohols and Esters An example for asymmetric reaction (formation of enantiopure compounds) is depicted in Sch. 7. Similar reaction of diesters with lipase has also been used for the synthesis of enantio-pure lactones (Sch. 8). Asymmetrization of meso-diols (Sch. 9) usually lead to quantitative conversions of the substrate preventing racemization of the wrong enantiomer and extensive research has been done in this area.

Acylation of Other Nucleophiles and Unusual Acyl Groups Lipases were also shown to acylate a wide range of nucleophiles such as ROOH or RZNH2. Besides, several unusual acyl acceptors (nucleophiles) and acyl donors have also been reported. Bulky acylating reagents usually react more slowly as slim fatty acids are natural acylating agents of lipase. Depending on the configuration of their substrate binding sites, however, different lipases may show

Scheme 8. Asymmetrization of diester with lipase for the formation of the g-lactone. 100% Conversion (. 98% ee ) with R ¼ Et; Porcine pancreatic lipase, hexane; 100% conversion (32% ee ) with R ¼ Et; P. fluorescens lipase, hexane; 100% conversion (. 95% ee ) with R ¼ Bn; Porcine pancreatic lipase, hexane were observed (from Ref. [43]).



Scheme 9. diesters.

541

Asymmetric products obtained from lipase-catalyzed asymmetrization of meso-

quite different enantioselectivities towards bulky acyl groups. Figure 2 shows example that represents the versatility of lipases as biocatalysts.

3. Resolution of Sterically Hindered Compounds Carboxyl esters bearing a fully substituted chiral center adjacent to the ester moiety (e.g., esters of tert-alcohols and of a,a-disubstituted carboxylates) are usually not accepted as substrates by lipases. Several proteases and few esterases are capable of hydrolyzing these substrates despite their steric bulkiness. But the number of these highly useful enzymes is rather limited. Two strategies are developed to circumvent this limitation. (i) The use of “activated esters” bearing electron-withdrawing groups enhances the electrophilic properties of the ester moiety, thereby increasing the enzymatic reaction rate, may help to overcome slow reaction rates. (ii) Spatial separation of the bulky quarternary carbon atom bearing the chiral center from the ester group to be hydrolyzed by a spacer moiety led to readily acceptable modified, but nonactivated, substrates. The nonactivated, activated, and spacer separated substrates are depicted in Fig. 3. An excellent review has recently discussed the use of these strategies with examples.[254]

E. Biopolymer Production Biopolymers are gaining tremendous attention in the recent years. Of the biotechnological productions (microbial/enzymatic), enzymatic synthesis is shown to be superior for the purpose because of the high selectivity of enzymes to produce polymers with high degree of chemical, regio- or optical-


542


Figure 2.

Unusual acyl donors or acceptors of lipase catalysis (from Ref. [43]).

selectivity.[255] Enzymatic polymerizations are receiving considerable research interest as a new route for polymer production since biocatalysis is expected to generate environmentally acceptable properties like biodegradability and biocompatibility. Both enzymatic and chemo-enzymatic routes have been utilized for the synthesis of a variety of novel polymers for applications as water



543

Figure 3. Sterically hindered substrate types. “G” is an electron-withdrawing group and “X” is a spacer moiety (from Ref. [254]).

adsorbents, hydrogels, biodegradable materials, chiral resolving matrices, and liquid crystals.[255] Lipases have been used to synthesize optically active polymers from a racemic mixture of monomers.[256] Lipase-catalyzed esterification of various monosaccharides with vinyl acrylate produced 6-acryloyl esters, which gave, upon chemical polymerization, water soluble polyacrylates that could be cross-linked to obtain insoluble material with the capacity to absorb up to 50-fold its weight of water.[257] Lipases are effective catalysts for polyesterification[258] and oligomerization[256] reactions in organic solvents. Use of chemical catalysis for these reactions is limited due to undesirable effects on the subsequent polymerization reactions. Recently, Iglesias et al.[259] reported C. antarctica lipase-catalyzed regioselective polyesterification of glycerol and adipic acid. UV-matrix-assisted laser desorption ionization time of flight mass spectroscopy was used for the analysis and low molecular weight polyesters ð1314 – 1716Þ with very narrow polydispersities ð1:0 – 1:2Þ were obtained. Several hydrolytic enzymes can be utilized in organic solvents to carry out certain condensation reactions that are difficult or impossible in aqueous media. Fermentative production and chemical synthesis of aliphatic polyesters have been widely studied,[260] however, these studies are outside the framework of this review.

1. Polycondensation Lipase-catalyzed polyester synthesis was first performed by dehydration between a hydroxyl group and a carboxyl group.[261] The polyesters thus obtained had molecular weights in the range 600 – 1300: Although, dehydration reaction is normally performed in nonaqueous media, recently lipase-catalyzed dehydrative polymerization was performed in aqueous media.[262] Similar reaction was also reported in a solvent-free heterogeneous system.[263]


544


The reaction employing an activated carboxylic acid ester as monomer proceeded more readily via polycondensation than dehydration. Activating groups are often 2,2,2-trichloroethyl- and 2,2,2-trifluoroethyl-units.[264] An enantioselective polymerization of a chiral, epoxide-substituted diester with 1,4-butanediol was also performed using porcine pancreatic lipase.[264] Enantioselectivity achieved was high as reported by the enantiomeric purity of the unreacted epoxy monomer (.97%). It was also reported that only primary alcoholic groups of sucrose were reacted during the synthesis of sucrose-containing linear polyesters from sucrose and bis(2,2,2-trifluoroethyl) dicarboxylates.[16] As the transesterification is an equilibrium reaction, polymerization via transesterification is sometimes disturbed by the liberated alcohol. Lipase-catalyzed polymerization of divinyl adipate with glycols afforded polyesters with a high molecular weight (up to 6700), where the liberated vinyloxy group forms acetaldehyde but not an alcohol.[265] This mechanism was later employed as a single-step process for polyester synthesis.[266]

2. Ring-Opening Polymerization of Lactones Lipases have also been found to induce the ring-opening polymerization of nonsubstituted lactones with various ring sizes. 1-Caprolactone (7-membered) polymerization was the first and most extensively studied.[267] A small sized lactone, b-propiolactone (4-membered) was polymerized using Pseudomonas lipase to a mixture of linear and cyclic oligomers with molecular weight of several hundreds.[268] Further, enzymatic polymerization of substituted 4-membered lactones was also reported.[269] Apart from these small-sized lactones, ringopening polymerization of medium-sized lactones (d-valerolactone, 6-membered) and 1-caprolactone using lipases from C. rugosa, P. fluorescens, and porcine pancreatic lipase have been reported.[270] However, the molecular weight was not significantly high (,2000) in case of d-valerolactone polymerization, while molecular weights obtained for 1-caprolactone polymerization were strongly dependent upon the origin of lipase.[271] Polymerization of 9-membered lactone (8octanolide) was demonstrated by Kobayashi et al.[270,271] through lipase catalysis. Four macrolides: 11-undecanolide (12-membered), 12-dodecanolide (13-membered), 15-pentadecanolide (16-membered), and 16-hexadecanolide (17-membered) were also polymerized using lipases.[272] The highest molecular weight (,25,000) was achieved with 11-undecanolide polymerization. In all the cases, the polymer contained carboxylic acid (at one end) and alcohol (at the other end) as terminal groups. Immobilized C. antarctica lipase (CAL) showed high catalytic activity compared to P. fluorescens (PFL) and P. cepacia (PCL) lipases for polymerization. CAL effectively catalyzed polymerization of 8-octanolide,[270] macrolides,[273] and lactones possessing a methyl substituent such as a-methyl-dvalerolactone, a-methyl-1-caprolactone, and b-methyl-d-valerolactone.[274]



545

3. Ring-Opening Copolymerization of Lactones The CAL was also found to induce the copolymerization of lactones. Kobayashi et al.[270,271] demonstrated the copolymerization of 8-octanolide and two other lactones (1-caprolactone, 12-dodecanolide) using CAL. 8-Octanolide was observed to give low polymerization yield compared to the other two. The formation of random copolymer was also confirmed by 13C-NMR spectroscopy. Formation of random copolymer is, in fact a unique feature of enzymatic polymerization. Conversely, conventional methods of copolymerization (anionic process) normally result in a block-type copolymer due to large polymerizability difference of lactones with a different ring size.

4. Control of Polymer Terminal Structure/End-Functionalization Facile control of terminal structure is essential as the terminal-functionalized polymers (macromonomers and telechelics) are often utilized as prepolymers for the production of functional polymers. A single-step production of methacryl-type polyester macromonomers by the initiator and terminator methods was reported. CAL was found to catalyze initiation of the polymerization of lactones by introducing an alcohol moiety at the polymer terminal (initiator method).[275] A methacryloyl group was introduced successfully at the terminal position of 12-dodecanolide polymer using 2-hydroxyethyl methacrylate as an initiator to result in a methacryl-type polyester macromonomer. This methodology was also employed for the production of v-alkenyl- and alkynyl-type macromonomers using 5-hexen-1-ol and 5-hexyn-1-ol as initiators. Alkyl glucopyranosides were also reported to initiate CAL-mediated polymerization of 1-caprolactone to result in a polymer bearing sugar moiety at the terminal position.[267] End-functionalized polymers were also produced via lipase-catalyzed polymerization of 12-dodecanolide in presence of vinyl esters.[276] Vinyl methacrylate and divinyl sebacate were employed as terminators to produce a methacryl-type macromonomer and a polyester telechelic with carboxylic group at either ends. Recently, Kobayashi[262] reviewed various aspects of polymerization reactions including polysaccharide synthesis and enzymatic polymerization including lipase- and oxidoreductase-catalyzed reactions.

F. Flavor Esters Low molecular weight esters are the principle components responsible for the aroma of fruits like pear, pineapple, strawberry, apple, and banana, etc. In the 1800s, reports began to appear about the isolation of microorganisms that produced different fruity aromas.[114] The production of aromas in cultures of yeast, molds, and bacteria was attributed to ester formation caused by the reaction


546


between organic acids and ethanol.[116,277,278] Later on, research was concentrated on finding the basic mechanism involved for the formation of fruity esters. Lipase is the enzyme principally involved in the release of free fatty acids from triglycerides. These free fatty acids then react with alcohol to form esters, which are responsible for the fruity odor. Pseudomonas are not considered as desirable organisms to be utilized for generating food flavors, since many of these bacteria are pathogens. Therefore, research was directed to find organisms that are accepted for use in foods. Several lactic acid bacteria (Streptococcus lactis, S. lactis subsp. diacetylactis, S. cremoris, and several Lactobacilli ) were useful in this regard.[114] In addition to producing flavor compounds, many microorganisms serve as sources of enzymes that can be utilized to alter/produce the food flavors. Flavor compound synthesis by biotechnological processes plays an increasing role in food industry.[279] This is the result of scientific advances in biological processes, making use of microorganisms or enzymes as an alternative to chemical synthesis, combined with developments in advanced analytical techniques such as HPLC, GC, IR, or mass spectroscopy. Enzymes as biocatalysts offer a variety of possibilities over microorganisms: enzymes are specific, stereo-selective, by-product formation is minimized, and product recovery becomes easy. A great deal of research has been directed towards the use of lipid-related enzymes—mainly lipases.[280 – 284] Flavor ester synthesis by means of lipases is an interesting alternative considering that there are many well-known flavor esters in the natural aroma of fruits, traditionally obtained by extraction or by chemical synthesis. Lipases offer several advantages and the areas where lipases can have a great impact on the flavor industry include: (i) production of natural flavoring ingredients, (ii) production of part or complete flavor, and (iii) increased yields of essential oils, oleoresin, and flavor components in natural material processing. Existing technologies of flavor production are predominantly based on natural plant materials. However, agricultural production of flavors is subject to seasonal and regional variations/fluctuations. Chemical synthesis may produce unwanted by-products, and the products obtain “synthetic” label. Recent consumer trend for “natural” products has altered the preferences away from chemically synthesized flavors. Flavors obtained through biocatalysis can be considered “natural” by regulating agencies and thus can command premium even when priced substantially high. For instance, the naturally produced cis-3-hexanol is priced at $4000 per kg while the synthetic counterpart costs only about $80 per kg.[285] According to the International Organization of the Flavor Industry (IOFI) notation, the definition for natural flavors is “those obtained by appropriate physical, enzymatic, or microbiological processes from materials of vegetable or animal origin, either in raw state or after processing for human consumption.”[286] Therefore, several food manufacturers, at present, are keen in sources of natural



547

ingredients. This is more so for flavoring in the food, dairy, and beverage industries. Some of the initial reports on the synthesis of flavor esters via lipase catalysis include those of Gatfield,[287] Inada et al.,[288] Iwai et al.,[31] and Langrand et al.[198,289] Apart from using isolated enzymes, few attempts have also been made to employ enzymes present in microbes directly without isolation. R. arrhizus mycelium has been used directly to catalyze the conversion of octanol and palmitic or oleic acid to the corresponding esters.[290] However, most of the work at present is related to the use of enzymes isolated from microorganisms. Some arguments have been raised regarding the “natural” label of flavor esters produced through lipase catalysis because the solvents used as a reaction medium may not be natural and sometimes be toxic. Despite these hurdles, recent advances in the use of supercritical fluids and solvent-free systems (where only substrates are present with no added solvent) appear to be suited ideally to the perfume and flavor industry. Some of the important esterification reactions used for the synthesis of various kinds of esters are listed in Table 6.

V. KINETICS AND PROCESS OPTIMIZATION A. Kinetics of Lipase-Catalyzed Esterification The application of lipases for lipolysis, esterification, transesterification, and lactonization reactions in organic solvents has increased significantly during the last decade.[341] For the design of suitable reactors, kinetic information on the rate of product formation and the effects of changes in system conditions is needed. Developing rate expressions to characterize several lipase-catalyzed reactions and deciphering the mechanisms involved has been found to be a challenging task.[342] Investigations on reaction kinetics would provide such information and facilitate the prediction of reactor performance under a wide range of system conditions. In contrast to the wealth of information pertaining to the kinetics of lipasecatalyzed hydrolysis reactions, relatively little is reported on the kinetics of esterification and transesterification reactions. Most of the models reported to date are based on the application of simple Michaelis– Menten kinetics, which seem to be valid for the most simple enzymatic reactions. Some researchers have proposed kinetic models for lipase-catalyzed esterification and transesterification reactions based on Ping-Pong mechanism.[343 – 345] It is generally agreed that the reaction occurs via the formation of an active complex between an acyl group and the active site of the lipase.[346,347] Despite the fact that several kinetic studies are reported, the information needed for industrial scale design and analysis continues to be rather limited.[348] In addition, most of the literature reports deal with the esterification of short-chain alcohols and medium- or long-chain acids, in which


548

Table 6. Lipase-Catalyzed Esterification Reactions: A Survey Lipase Sourcea (State)b ANL, RhDL, GCL, PCL (free)

Solvent (s) Casein solution

Acyl Donor Oleic acid

ANL, GCL, PcPL, RhDL Aqueous (free)

C2 – C6, tigric acids

CRL, ANL (free)

Oleic acid

Water

Stearic, oleic, and linoleic acids Oleic acid

CRL (IM)

Heptane

Butyric acid

RmML (IM)

Various solvents

PsFGL (free)

Isooctane

RmML (IM)

—

Various carboxylic acids Aliphatic acids Oleic acid

Oleic, erucic acids

Salient Features

Esterification efficiency: RhDL ¼ GCL . PCL . ANL. All 18 alcohols esterified easily. Only GCL esterified 28 alcohols and diols. No lipase esterified 38 alcohols. Stearic hindrance effect of substrates Geraniol, farnesol ANL showed activity for all C3 – C5 acids. Others showed no activity for most acids tested. Acetic and tigric acids not esterified with geraniol Glycerol Slow reactions at room temperature. Low yields. Equilibrium shifted using 50% excess oleic acid and removal of water by distillation Sucrose, glucose, Of all lipases, CRL was most active. Low yields fructose, and of stearic acid esters due to poor dispersion of solid sorbitol stearic acid in the reaction medium Glycerol RmML and ChVL showed highest yields. Stability of enzymes and product compositions were studied Alcohols, sugars, Synthesis of various kinds of esters using lipases amines, thiols from different sources. No esterification was observed with sugars, thiols, etc. Ethanol Periodic hydration of immobilized lipase was found essential for its long-term use Various alcohols Study on characteristics of the immobilized lipase. High activity in hydrophobic solvents and very low activities in hydrophilic solvents Racemic alcohols Enantioselective esterification using surfactantcoated, solvent-soluble lipase Acetone – Acetone –glycerol acryl ester synthesis glycerol C1 – C12 1, 2, and 38 alcohols, diols, phenols, surgar alcohols

References [197]

[31]

[291]

[34]

[292] [293]

[27] [102]

[294] [295]


ANL, CRL, EL, PsXL, Aqueous PeXL,RhXL, RmML, RmXL (IM) ChVL, RhDL,RmML, Water PsFL (free) ANL, CRL, RmXL, HLL, Buffer RhAL, PsXL (IM)

Alcohol


— Isooctane reverse micelles (RMs) —

Oleic acid Lauric acid

Glycerol Butanol

Oleic acid

RmML (sol) CRL (free) AXL (free) RhAL (my)

Heptane

C2 – C6 acids

C1 – C5 1, 28 alcohols C1 – C6 alcohols, citronellol, geraniol

RmML (IM)

—

Oleic acid

Butanol

CRL (free) PsFL (free) RmML (IM) RmML (IM)

Hexane, octane, decane, CH2Cl2 Isopropyl alcohol

C2, C4 acids Stearyl amine

C2, C4, C5 alcohols C8 – C14 alcohols

CRL (IM)

Hexane

Propionic acid

Ethanol

RhDL (RM)

Isooctane RMs

Lauric acid

Glycerol

RmML (IM)

Solvent-free (SF)

Glycerol

RmML (free)

C10, oleic acids Butyric acid

n-Hexane, biphasic (aqueous-hexane) Heptane C2, C8, C16 acids

RmML (IM)

ChVL (MBG)

Butanol

C2, C8, C14 alcohols

High glyceride yields with effective water removal Rapid loss of activity. No alteration in pH profile or substrate specificity observed Lipase activity lower with isoalcohols compared to primary ones Esterification efficiency: RmML . RhAL . CRL . AXL. Yields of 35 short-chain flavor esters using various lipases were compared. Lipases of different origins show different selectivities Effect of water content on activity and equilibrium conversion was evaluated. Initial water content did not affect esterification at low pressures (0.32 bar) Vmax and Km determined. Effect of solvent hydrophobicity on ester yield investigated Low reaction rates for C8 alcohol; rates increased with increasing alcohol chain length Covalent attachment to nylon. Improved activity and temperature stability upon immobilization. Inactivation by high ethanol concentrations Optimization of reverse micellar media for monoglyceride synthesis Medium-chain glyceride synthesis in SF system

[15] [296]

Water removal using Dean – Stark apparatus had little effect on final yield. Complete dehydration caused inactivation Lipase immobilized in micro-emulsion based organogels to serve as novel solid-phase catalysts. They were active at , 2 208C and adapted for continuous operation

[302]

[297] [198]

[173]

[298] [299] [300]

[301] [191]


RmML (IM) CRL, RhDL (RM)

[303]

549

(continued )


Lipase Sourcea (State)b

Solvent (s)

Acyl Donor

550

Table 6.

Continued

Alcohol

Salient Features

References

Oleyl alcohol

Esterification with or without water removal

[304]

Heptane

C18, C22 Unsaturated Butyric acid

Citronellol

[305]

RmML (IM)

Hexane, SCCO2

C14 acid

Ethanol

RmML (IM)

—

Oleic acid

RmML (IM)

Various organic solvents

Decanoic acid

Isopropylidene glycol Dodecanol

Water content of bulk solvent decreased as reaction progressed. Activity recovered upon dehydration of used enzyme Vmax, Km, and Ki(alcohol) were determined in both solvents Kinetic analysis of the system

CRL, HXL, PcPL, RmML, RhNL (IM)

Hexane

Decanoic acid

Dodecanol

RmML (IM)

Solvent-free, n-heptane

Butyric acid

Geraniol

ChVL (free)

C10 acid

Glycerol

ANL, CRL, PsFL (IM)

Hydrocarbons, ethers, ketones n-Hexane

Lauric acid

PcPL (IM)

Hexane

Oleic acid

Methanol, ethanol, amyl alcohol Ethanol

RmML (IM)

SCCO2

RmML (IM)

Hexane is a better immobilization medium compared to water. Optimum pH for CRL ¼ 7.0, ANL ¼ 6.5, and PsFL ¼ 7.0 Increased stability of immobilized lipase in the presence of an additive. Additives used: albumin, gelatin, casein, and PEG

[307] [308]

[309]

[310]

[311] [312]

[313]


Reaction rate is dependent on aw with the optimal aw ¼ 0.5 for RmML immobilized on anion-exchange resin At aw ¼ 0.12, RmML and RhNL are active while HXL, CRL, and PcPL require high aw. Immobilization onto anion-exchange resin gives similar behavior to immobilization onto macroporous polypropylene Lipozyme was unsuitable for solvent-free ester synthesis. Water was adsorbed to the hydrophilic support material Product composition studies in different solvents

[306]


n-Hexane

Fatty acid esters Acetic acid

Citronellol, geraniol Citronellol

CAL (IM)

n-Hexane, various organic solvents

CAL (IM)

n-Hexane, various organic solvents

Acetic acid

Geraniol

CAL (IM)

Solvent-free

Acetic acid

Citronellol

ANL, CRL, PPL, PsXL, RhXL, RmJL (free)


Lauric acid

Benzyl alcohol

PcPL, PsFGL, PsFL (free)

Benzene

Geraniol

PPL (free and IM)

SCCO2

Farnesyl acetic acid Butyric acid

Racemic glycidol

GCL (free and IM)

Hydrocarbons

CRL, RhDL (free)

AOT-Isooctane

Oleic and stearic acids Lauric acid

Ethanol, butanol, hexanol, octanol 1,3-Propanediol

PsXL, RmML(IM)


Hexanoic acid

Benzyl alcohol

. 95% molar conversions obtained for short-chain fatty acid esters of both the alcohols 98% conversion after 14 hr at 208C. Enzyme inhibition by acetic acid but not by citronellol. Higher log P did not necessarily mean higher enzyme activity Enzyme inhibition by acetic acid but not by geraniol. Conversion was independent of initial solvent’s water content. Solvent log P ¼ 0.85 led to .80% conversion Optimum alcohol/acid molar ratio ¼ 1.56, solvent log P ¼ 1.63. Desiccants improved yield by 10% Enzyme inhibition by benzyl alcohol. Activity is dependent on type of coating-surfactant, solvent, lipase preparation, aqueous pH, and origin of lipase Lipases, chemically modified with PEG derivatives, preferentially catalyze (R )-isomer of secondary alcohols Enantioselectivity of enzyme is unaffected by enzyme hydration in SCCO2 Selectivity unaffected upon immobilization onto silica gel Substrate inhibition by lauric acid at above 70 mM. Kinetic analysis of the esterification reaction Exceptions to general trend of enzyme activity with respect to solvent log P value. Importance of compatibility of solvents with substrates/products

[314] [315]

[316]

[317] [318]

[319]

[320] [321]


CAL (IM)

[322] [323]

(continued ) 551


552

Table 6. Lipase Sourcea (State)b

Solvent (s)

Acyl Donor

None

Lauric acid

RmML (IM)

Lauric acid

PsXL (free)

Aqueous, biphasic (isooctane – water) Hexane

Palmitic acid

ANL, CRL, PPL, PsXL, RmJL, RhXL (free)

Aromatic and aliphatic hydrocarbons

Various chain length fatty acids

UML (free and IM)

Hexane

RmML (IM)

Isooctane

Various fatty acids or ethyl esters Oleic acid

RmML(free and IM)


Butyric acid

CAL, RmML(IM)

None

C2 – C4 acids

CRL (IM)

Toluene

Various fatty acids

Alcohol Butanol

Salient Features

Aggregation of soluble RmML dispersed by the addition of water. Higher stability of immobilized RmML Butanol, lauryl Thermo-deactivation of RmML aggravated by alcohol exposure to butanol. Higher reaction rates in biphasic systems Cetyl alcohol Repeated use of lipase –surfactant complex in a membrane reactor Menthol Nonionic-surfactant coated CRL showed increased enantioselectivity and reaction rate over native CRL. Isooctane gave the highest rate and enantioselectivity compared to other solvents tested 1,2: 3,4-di-oImproved activity and stability upon immobilization isopropylidene onto Amberlite XAD-2. Substrate inhibition by galactopyranose linoleic and linolenic acids n-Butanol Increased ester yield due to removal of water by pervaporation from the reaction mixture Geraniol Reaction was better in n-heptane compared to n-hexane, isooctane, and cyclohexane. Regeneration of enzyme by Na2SO4 Geraniol Unlike RmML, CAL showed stable activity during four consecutive runs and low sensitivity for water content Sulcatol Kinetic studies on fatty acid esterification catalyzed by immobilized CRL. Reaction follows Ping-Pong mechanism with competitive alcohol inhibition

References [324]

[325]

[326] [327]

[328]

[190] [329]

[330]

[331]


RmML(sol and IM)

Continued


Isooctane

Lauric acid

Menthol

RhAL (free), RmML (IM) None CRL (free) n-Hexane

Butyric acid Tributyrin

Geraniol Geraniol

CRL (free)

Isooctane

Racemic naproxen

Trimethylsilyl methanol

RmML (IM)

None

Oleic acid

Octanol

CRL (free)

Hexane

a-Lipoic acid

Primary alcohols

CRL (free)

Isooctane

Lauric acid

Menthol

RmML (IM)

None

Oleic acid

RmML (IM), PPL

n-Heptane/solventfree

Butyric acid

Lauryl alcohol, isosorbide, sorbitol Butanol

Nature of cosolvents used in preparing surfactantcoated lipase affects both the yield and the activity. Hydrophilic cosolvents were most suitable Unlike RhAL, RmML tolerates high substrate levels Use of RSM for optimizing TE reaction. Variables selected: reaction time, temperature, enzyme concentration, substrate molar ratio and added water Improved enantioselectivity of lipase in presence of bis(2-ethylhexyl) sodium sulfosuccinate (AOT). CRL lipase is inhibited competitively by alcohol and noncompetitively by AOT Esterification reaction catalyzed by lipase immobilized on a hydrophobic support follows the Ping-Pong bi– bi mechanism Optimum enantioselective esterification with n-hexanol. Model for enzyme active site suggested Surfactant-coated lipase catalyzed esterification followed a Ping-Pong Bi – Bi mechanism with dead-end inhibition by alcohol Comparison of tubular recycle reactor with stirred tank reactor. Comparison of various enzyme kinetic models Description of an assay method for determination of esterification activity of lipases

[332]

[333] [334]

[335]

[336]

[337] [338]

[339]


CRL (free)

[340]

a

553

Abbreviations used: ANL: A. niger, AXL: Aspergillus sp., CAL: C. antarctica, CLL: Candida lipolytica, CPL: C. parapsilosis, CRL: C. rugosa, CXL: Candida sp., ChVL: Ch. viscosum, EL: Enterbacterium, GCL: G. candidum, HLL: H. lanuginosa, HXL: Humicola sp., PCL: P. cyclopium, PXL: Penicillium sp., PsCPL: P. cepacia, PsFGL: P. fragi, PsFL: P. fluorescens, PsXL: Pseudomonas sp., PPL: Porcine pancreas, RhAL: R. arrhizus, RhDL: R. delemar, RhNL: R. niveus, RhXL: Rhizopus sp., RmJL: Rhizomucor javanicus, RmML: R. miehei, RmXL: Rhizomucor sp., UML: Utsilago maydis. b Abbreviations used in parentheses: State of the enzyme used. (sol)—Soluble, (IM)—Immobilized, (my)—Mycelium, (RM)—Reverse micelles, (ME)— Microemulsion, (MBG)—Microemulsion based organogels, and (free)—Native enzyme.


554


alcohol inhibition is well established while acid effects are unclear. We have proposed a kinetic model for the lipase-catalyzed esterification of short-chain substrates (isoamyl alcohol and butyric acid).[349] Several authors recommended the use of thermodynamic activities instead of concentrations in kinetic equations particularly when the contribution of solvent interaction on substrate availability to the enzyme is considered.[350] Lipasecatalyzed esterification has been shown to follow Ping-Pong kinetic model.[174,331,344] Competitive inhibition by the alcohol has usually been found to be significant, resulting in the following equation for the rate of the forward reaction:[351] v¼

h

V max

i KA KB 1 þ ½A 1 þ K½BiB þ ½B

ðV:1Þ

where v is the initial reaction rate, Vmax is the maximum reaction rate, KA and KB are the binding constants for the fatty acid (A) and the alcohol (B), and KiB is the inhibition constant for the alcohol. Valivety et al.[174] and Janssen et al.[352] have suggested incorporation of another inhibition term in the rate equation for water (product), if the amount of water present is considerable. This term will only be significant if the concentrations of both A and B are low. However, at high concentrations of A or B or both A and B, this term will become negligible, leading to Eq. (V.1).[352] In our kinetic study, we have employed substantially higher concentrations of substrates and low conversions were utilized for initial rate determination to avoid product inhibition. We have used a model system in which both alcohol and acid are of short-chain length observing a Ping-Pong Bi – Bi mechanism with inhibition by both the substrates. A general expression for the forward rate according to this mechanism is given by:[351] v¼

V ½A½B h max i h i ½B ½A ½A½B þ K A ½B 1 þ K iB þ K B ½A 1 þ K iA

ðV:2Þ

where KiA is the inhibition constant for the acid, the other notations are same as in Eq. (V.1).

B. Mass Transfer Influences on Kinetics The significant difference between aqueous and nonaqueous enzymology is that immobilized enzymes, which catalyze reactions in organic solvents are heterogeneous catalysts and hence are subject to different diffusional limitations compared to their solubilized counterparts. For enzyme particles suspended in organic solvents, there will be two diffusional barriers that a substrate molecule



555

must cross before catalysis. Firstly, the substrate must diffuse from the bulk solvent, through a boundary layer, to the enzyme surface (external mass transfer). Secondly, the substrate must diffuse through the particle (may be consisting of enzyme, salt, and water) to the enzyme active site (internal mass transfer). Varying speeds on the shaker can give an indication of the effect of external mass transfer while internal diffusional limitations can be gauged by changing the particle sizes. When the enzyme activities are compared in various organic solvents, it is important to ensure that the roles of external and internal mass transfer do not change for different solvents. Few researchers have investigated mass transfer effects to understand the enzyme behavior.[349,353 – 355] We have assessed mass transfer limitations in a solvent-free system (unpublished data) and have found that mass transfer limitations were significant for long-chain substrates while short-chain substrates (water soluble) pose reasonably no difficulties. This could be due to the highly hydrophilic nature of the substrates. While hydrophobic substrates (which exhibit low solubility in the aqueous microenvironment) require vigorous agitation, the hydrophilic substrates preferably enter the microaqueous layer surrounding the enzyme. Both the substrates employed in our case (isoamyl alcohol and butyric acid) have sufficient polar nature to enter microaqueous interface. Goldberg et al.[356] have also observed that diffusional limitations in highly hydrated enzyme preparations were less important for hydrophilic than hydrophobic substrates because of their higher solubility in the hydrated solid phase. While simply increasing agitation speed can minimize external diffusion, internal diffusion limitations can be overcome by reducing the particle size as well as by low enzyme loading on matrix.

C. Process Optimization Response surface methodology (RSM)-based on various experimental designs for the optimization of lipase-catalyzed synthesis of various esters has been reported.[28,29,334,357] Response surface methodology is an efficient statistical tool for optimization of multiple variables using minimum number of experiments.[358] It is superior to the traditional approach of optimization by varying one parameter at a time while keeping others constant. The relationships between important esterification variables (substrate and enzyme concentrations, temperature, water activity, reaction time, etc.) and ester yield, and optimum conditions for ester synthesis have been investigated. The designs utilized for the optimization include 3-variable 3-level Box– Behnken design[29] and central composite rotatable design (CCRD).[28] The experimental data obtained as per the design can be fitted to a second order polynomial equation


556


to obtain coefficients that can be utilized for creating response surfaces, predictive correlations, and generating optimum conditions.

VI. STABILITY OF LIPASES Enzymes inactivate at high temperatures in aqueous media due to both partial unfolding and covalent alterations in the primary structure.[38] Water is required for both these mechanisms and hence, it should be possible to enhance the enzyme thermostability in nonaqueous environments, mainly as a consequence of protein rigidity in these systems. The thermostability observed is quite impressive. The porcine pancreatic lipase remained stable at 1008C for .12 hr.[37] Such dramatic thermostabilization is seldom possible by using other approaches like chemical cross-linking, immobilization, or even protein engineering. Apart from rigidity, another reason for enhanced thermostability is that a number of covalent processes involved in irreversible or reversible inactivation of proteins such as deamidation, peptide hydrolysis, and cystein decomposition require water, but are extremely slow in nonaqueous systems. Ueda et al.[359] observed that the lipase from Rhodococcus equi is highly active at 708C in alcoholic solvents as long as the solvents are dry, while in wet solvents stability is very poor. Several of these experiments are restricted to relatively nonpolar solvents. Biocatalyst stability in polar solvents is quite low. However, Reslow et al.[360] examined the thermostability of Celite immobilized chymotrypsin at 508C and had shown that the solvents having log P . 0:7 support the thermostability. This kind of information is quite useful and need to be collected for other enzyme systems also. Most of the previous work had been restricted to relatively nonpolar substrates. Stability studies involving polar and water soluble substrates are also essential to bring out a clearer picture. Enzymatic catalysis has recently been extended to both gas phase and supercritical solvent systems. The enhanced thermostability of enzymes in nonaqueous media is important for these systems since it extends the temperature range in which the enzymes are active. Despite the advantage of high thermostability of enzyme in nonaqueous systems, the rates of reaction in fact decrease beyond optimum temperature. In many instances, thermostabilization of enzymes also results in stabilization towards other denaturing conditions. Arnold[144] pointed out that there might be correlations between enhanced thermostability and stability in nonaqueous systems. In support of this, malic enzyme and alcohol dehydrogenase from Sulfolobus solfataricus (an extremophile) have been found to show a correlation between thermostability and organic solvent resistance.[361] In this respect, proteins from extremophiles do not differ significantly from their mesophilic counterparts. Their adaptation to extreme conditions, either intrinsic or through interaction with extrinsic factors, is accompanied by only marginal increases in the free energy of stabilization. No general rule or strategy of stabilization has yet been framed.[362]



557

Adverse Influence of Water of Reaction on Stability During synthetic reactions like esterification, complications arise because water is a product. Several researchers[282,290,308] observed that in addition to a decrease in equilibrium yield, accumulation of water results in a decrease in the enzyme activity. Moreover, accumulated water is also shown to affect adversely the long-term stability of the enzyme.[282,300,363] The reduction in water content of the enzymes usually results in an increase in their half-lives.

A. Enzyme Stabilization in Nonaqueous Media In recent years, well-defined thermostabilities have been elucidated by analyzing ultrastable proteins and verifying their specific anomalies by rational design. Taking thermostability as an example, several experimental approaches have been employed to assign specific structural alterations to changes in stability: selection of temperature-sensitive mutants; systematic variations of amino acid residues in the core or in the periphery of model proteins; fragmentation of domain proteins or modifications of connecting peptides between domains, and alteration of subunit interactions by mutagenesis or solvent perturbation.[364] Jaenicke and Bo¨hm reviewed various strategies of stabilization of enzymes.[362] The approaches followed to stabilize enzymes are mainly two: (i) medium engineering and (ii) biocatalyst engineering.

B. Medium Engineering It is generally agreed that enzymes in nonaqueous systems can be active provided that the essential water layer around them is not stripped off.[5,142] Medium engineering in the context of biocatalysis in nonaqueous media involves the modification of the immediate vicinity of the biocatalyst. First rule is that nonpolar solvents are better than polar ones since the former provide a better microenvironment for the enzyme. Thus, the solvents with log P . 4:0 are most suitable while those with log P , 2:0 would constitute a poor choice. Solvents in the intermediate range of log P ð2:0 – 4:0Þ are unpredictable and are likely candidates for medium engineering. The properties of the substrates and products should also be considered for medium engineering. If the immediate microenvironment of the enzyme favors substrate solubility and has low product solubility, the reaction rates would be high. Laane,[365] in an excellent review on medium engineering, discussed how the microenvironment of the enzyme is controlled by the nature of matrices to which the biocatalyst may be linked. While the solvent effect appears to hold true for most enzymes, there are few exceptions. Porcine pancreatic lipase is catalytically active in anhydrous pyridine[35] suggesting that the enzyme can retain its bound water even in


558


the presence of a water-miscible solvents. This property is thought to be evolutionary. Lipases are activated by water-insoluble fatty acid esters at above critical micelle concentration. Hence, their natural environment is nonpolar. The ability to bind essential water tightly might have evolved as a prerequisite condition for catalysis in hydrophobic environments. Several aspects have to be considered in choosing an appropriate solvent for a given reaction: (i) the compatibility of the solvent with the selected reaction (substrates and products), (ii) solvent must be inert, (iii) solvent density should be low so as to minimize mass transfer limitations, and (iv) other solvent properties like surface tension, toxicity, flammability, waste disposal, and cost. Halling[363,366] presented a detailed account on predictions that can be made to elucidate the influence of solvent selection on the equilibrium.

C. Biocatalyst Engineering Immobilization of enzymes and protein engineering have been known for improving biocatalyst efficiency/stability in aqueous media. Similar efforts to improve biocatalyst performance in organic solvents have been made. Several recent developments have taken place in this area including the generation of active and stable homogeneous (soluble) biocatalysts. Solubilization of enzymes in hydrophobic media requires the covalent or noncovalent modification of the native enzyme. Covalent techniques are well described in the literature (e.g., attachment of PEG chains to enzymes).[101] Noncovalent modifications are much less common, although they are able to provide highly active and soluble enzyme forms. For example, protein –lipid complexes soluble in organic solvent have been produced using nonionic lipids.[367] Khmelnitsky et al.[99] provided an excellent overview of the earlier research in the area of biocatalyst engineering. Recently, Villeneuve et al.[368] reviewed various lipase stabilization strategies.

D. Microwave Treatment External field assisted enzymatic reactions are newly reported and are shown to provide high enzyme stability. While the accelerating effect of microwave treatment on chemical reactions is well documented,[369] only recently has this technique been extended to biocatalysis. Enzymatic reactions under microwave treatment can be carried out in solvent-free systems at temperatures up to 1108C employing enzyme and substrates impregnated onto a solid support. Microwave treatment under dry conditions was also found to enhance the lipase enantioselectivity during the acylation of 1-phenyl ethanol.[370]



559

VII. MODIFIED LIPASES Efforts have been made for improving enzyme efficiency/stability/recyclability in organic solvents through various forms/modifications including immobilization of the enzyme. Laane[365] used the term “Biocatalyst Engineering” for this. Various forms of enzymes employed in nonaqueous systems are: (i) enzymes dissolved in substrate solutions,[371] (ii) solid enzyme powder suspended in organic solvents,[372] (iii) solid enzyme adsorbed on support,[360] (iv) PEGmodified enzymes soluble in organic solvents,[373] (v) enzyme entrapped within a gel,[374] and (vi) immobilized enzyme suspended in organic solvents.[375] The insolubility of enzymes in monophasic organic systems pose difficulties such as mass transfer limitations (intraparticle and external) that can mask the true intrinsic kinetics.[18] Methods used to eliminate mass transfer limitations in conventional heterogeneous chemical systems have been effectively applied to enzyme catalysis in organic media. One such method to minimize external mass transfer limitation is to spread the enzyme onto materials with large surface areas like glass beads.[376] As the enzymes are insoluble in organic media, the bound enzyme will have less chance to desorb from the immobilization surface. While no general guidelines can yet be provided for choosing the most appropriate form of the enzyme for a specific purpose, some recent approaches of modifying enzymes for catalysis in organic solvents are given as follows.

A. Immobilization Researchers have shown immobilization as a tool for enhancing the function of biocatalysts in organic solvents.[377] The traditional types of immobilization techniques applied to lipases include adsorption onto solid supports such as silica gel, covalent attachment to solid supports, and entrapment in organic polymers or microemulsions. In general, adsorption techniques are easy to perform, but the bond is often weak. Such biocatalysts sometimes lack the degree of stabilization achieved by covalent attachment or entrapment. On the other hand, techniques based on covalent attachment may be relatively tedious and often require several chemical steps. Thus, entrapment seems to be the method of choice. However, ease of performance, increased activity, and long-term stability are also important goals.[378] Halling[379] and Welsh et al.[298] reported that immobilized lipases give faster reaction rates as compared to their native counterparts. Use of dehydrated biocatalysts in organic solvents often results in decreased activity in dry systems. Immobilization of enzymes on silica gel, followed by dehydration by rinsing with dry n-propanol, resulted in a 2-fold more active enzyme preparations compared to lyophilized powder.[380] Tanaka and Kawamoto[375] have dealt with the topic of immobilized enzymes in organic solvents. Unfortunately, in a number of studies on enzyme immobilization, only one-time performance is reported.


560


Avnir[381] described a novel approach to enzyme entrapment using inorganic matrices such as silica gel. Accordingly, the so-called sol – gel process, initiated by the hydrolysis of Si(OR)4, is performed in the presence of enzyme. Hydrolysis and condensation of Si-monomers in the presence of an acid or base catalyst trigger the cross-linking process with the formation of amorphous SiO2, a porous inorganic matrix that grows around the enzyme in a three-dimensional manner. Enzyme activities of 25– 100% relative to the nonimmobilized counterparts were observed. Relative activities of less than 5% were observed with lipase (e.g., P. cepacia ) immobilized in this kind of gels. It was concluded that the polar microenvironment of SiO2-matrix is not ideal for lipases and hydrophobic sol– gel materials derived from organically modified silane precursors were speculated to provide a better microenvironment.[382] Thus, monomers of the type RSi(OCH3)3 or mixtures of these and Si(OCH3)4 tetramethoxysilane (TMOS) were used in the sol– gel process in presence of lipases. Gels with relatively low quantities of MTMS content exhibited low activities, while those having .75% MTMS were 13 times more active than nonimmobilized enzyme. A number of other precursors such as RSi(OCH3)3 were also examined, the organic R-groups being C2H5, n-C3H7, n-C4H9, and n-C18H37. Even higher enzyme activities were observed in these studies and disilanes such as (CH3O)3 Si(CH2)6 Si(OCH3)3 were also well studied including the investigations on the use of beneficial additives such as PEG.[382] Various lipases (e.g., C. antarctica, A. niger, H. lanuginosa, R. arrhizus, P. cepacia, and P. aeruginosa ) were entrapped using this process.[383] An increase in activity by factors of 5– 20 is common and in some cases factors up to 88 were observed.[384] Major attractive features of the lipase-containing gels are: they exhibit pronounced chemical and thermal stabilities, easy recovery from reaction mixture, high reusability efficiency, and higher enantioselectivity. For these reasons, the gels have been commercialized (Reagent of the Year 1997 from Fluka, Switzerland). Although the reasons for the increased enzyme activities have not been unambiguously elucidated, it is clear that the sol–gel materials need to contain hydrophobic alkyl groups. In the absence of such hydrophobic groups in the matrix (e.g., use of Si(OCH3)4 as the sol–gel precursor), only ,5% activity is observed.

B. Chemical Modification 1. Cross-Linking Use of glutaraldehyde to prepare insoluble cross-linked enzyme crystal (CLEC) has received significant research attention for application in organic solvents. P. cepacia lipase CLEC-catalyzed resolution of phenyl acetate on a pilot scale was demonstrated by Collins et al.[385] Although chemical cross-linking has been shown to be a facile and inexpensive method to enhance thermostability and tolerance to organic solvents, the extent and exact location of the modifications are often not clear. It is also difficult to conclude which changes resulted in the enhanced stabilization. In addition, since heterogeneous mixtures are generated



561

and the number of residual viable enzyme active sites remaining after modification was mostly not reported, it is difficult to assess and compare the data from different publications.

2. Modification with Polyethylene Glycol Chemical modification with monofunctional reagents permits the binding of specific monomeric or polymeric functionalities. In particular, covalent modification with amphiphilic polymer PEG has been successful in solubilizing proteins for applications in organic solvents as well as in the preparation of therapeutically active proteins with reduced antigenicity and enhanced in vivo stability.[386] Inada and colleagues[101] have modified several enzymes including horse radish peroxidase, catalase, lipase, and chymotrypsin with a PEG derivative. The modified enzymes were stable and active in a variety of polar and apolar solvents. Polyethylene glycol chains are thought to form a protective shell around the solubilized enzymes and entrap water, similar to that observed in reverse micelles.[387] Solubilized enzymes in organic media were discouraged for use in continuous processes due to enzyme recovery problems. Takahashi et al.[388] addressed this issue by converting the soluble PEG-enzymes into magnetic derivatives through treatment with Feþ2 and Feþ3 ions and recovering the enzymes with complete activity in a magnetic field. The issue of carrying out preparative scale biotransformations with solubilized enzymes to overcome internal diffusion limitation still remains a subject of debate. Hernaíz et al.[389] demonstrated enhanced stability of C. rugosa lipase in isooctane by treating the enzyme with pnitro-phenyl chloroformate and cyanuric acid chloride activated PEG. The former was more active than the latter. In both the cases, the stability enhancement was accompanied by a decreased lipolytic activity although transesterification activity was improved. An alternative to PEG is to use the nonionic amphiphilic polyoxyethylene lauryl ether (Brij 35) as a covalent modifier, which has been successfully applied to catalase.[390]

3. Modification with Polymers Ito et al.[391] described a new enzyme modification procedure in which various vinyl monomers were graft-polymerized using enzyme containing aliphatic azo groups as initiators. The modification enabled lipase to be soluble in organic solvents. The modified lipase-catalyzed esterification was performed in chloroform. Hydrophobic polymers (such as polystyrene and polymethyl methacrylate) as well as amphiphilic polymers (such as poly N-vinylpyrrolidone) were employed for lipase modification. It is believed that these grafted polymers provide a waterlike microenvironment for the enzymes. Improved thermostability


562


of enzymes (including lipases) by modification with synthetic polymers has been reported ([391] and references cited therein).

4. Modification of Amino Acids Chemical modification of enzymes is an excellent tool to investigate the active sites of lipases as well as to modify or improve the activities of the enzymes. Kawase and Tanaka[392,393] modified several amino acid residues of C. rugosa lipase and examined the activities of the modified enzyme in organic solvent as well as aqueous systems. Treatment of lipase with dithiothreitol significantly enhanced not only the esterification activity but also operational stability. Chemical introduction of small molecular functionalities has also been proven beneficial.[394,395] Fadnavis et al.[395] have modified C. rugosa lipase with diethyl-p-nitro-phenyl-phosphate (DPNP) to alter the lipase specificity. Although incorporation of unnatural amino acids using combined site-directed mutagenesis and chemical modification approach was performed for several enzymes, this approach has not been applied to lipases. Chemical modification of enzymes has recently been reviewed.[396]

5. Lipid- and Surfactant-Coated Enzymes One of the successful techniques employed to overcome the enzyme stability problem involves the complexation of enzymes with lipids and surfactants, thereby rendering the enzymes stable and soluble in nonaqueous systems.[397 – 399] Okahata and colleagues[397,398] employed this technique for glycosidases to carry out transglycosylation reactions. Okahata and Ijiro[400] first reported the preparation of lipid-coated lipase. Lipases from R. delemar, Pseudomonas fragi, and R. niveus were coated with nonionic synthetic dialkyl amphiphiles. The same group also reported an enantioselective esterification of racemic alcohols and aliphatic acids in isooctane employing lipid-coated P. fragi lipase.[294] Noda et al.[399] employed surfactant-coated enzymes for the ring-opening polymerization of lactones and found that the rate of polymerization of pentadecalactone catalyzed by the surfactant-coated Lipase-PS was 100-fold higher than that obtained using native enzyme. Lipase coated with sorbitan monostearate (Span 60) has been reported to improve the interesterification rate of tripalmitin and stearic acid in n-hexane.[401] Lipases overloaded with surfactant molecules remained active, indicating that surfactant molecules interact with specific sites on the enzyme. Isono et al.[326] employed Span 60 in synthesizing a lipase – surfactant complex of Pseudomonas sp. lipase to catalyze esterification of cetyl alcohol and palmitic acid in n-hexane. Higher reaction rates in organic media have also been observed with Mucor javanicus lipase coated with a nonionic surfactant, glutamic acid dioleylester ribitol



563

amide compared to native lipase.[402] Modification with various surfactants showed that the activity was greater when the enzymes were coated with surfactants having a branch or a double bond compared to those having straight chains, provided that the number of carbons in the surfactants was the same. Investigations to provide insight into the surfactant – lipase interactions and structure– function relationship have been carried out by Hermoso et al.[403] and Folmer et al.[404] Co-solvents employed during the preparation of the lipase– surfactant complex affect the three-dimensional structure of lipases. Nonetheless, Kamiya et al.[332] found no correlation between the log P of co-solvents and the performance of surfactant-coated C. rugosa lipase in the esterification of (R )-menthol with lauric acid, although hydrophobic cosolvents were generally suitable. Several reports indicate that the effect of surfactants on the enzyme activity and stability are mainly dependent on the origin of the lipase and the type and structure of the surfactant. There are no reports pertaining to the use of surfactants on immobilized lipase or the use of surfactant– lipase complex in solvent-free systems to date. Although surfactants have been shown to activate the CLECs of lipases,[405] the mechanism is yet unclear because the cross-linking presumably prevents significant conformational changes.

C. Modifications by Lyophilization Enzymes as biocatalysts are often prepared from a suitable aqueous or buffer solution via lyophilization. It has been reported that the lyophilization procedure could lead to undesirable changes in protein secondary structure[406] and about 40% of the active sites may suffer denaturation.[407] Several methods have been reported to enhance enzyme activity during this stage. The addition of specific small molecules (excipients) in the freeze-drying stage often results in improved catalytic activity and in some cases altered substrate specificities of the enzymes. Addition of an inorganic salt has been shown to dramatically enhance the activity of enzymes in organic solvents.[354] Initial reaction rate studies revealed that the activation is basically intrinsic and not just the relaxation of diffusional limitations.[408] Activation has also been achieved by the addition of crown ethers.[409] Dabulis and Klibanov[410] observed dramatic enhancement of enzyme activity in organic solvents with enzymes lyophilized from aqueous solution containing lyoprotectants (a ligand). This ligand-induced activation was observed regardless of the structural similarity of the ligand and substrates. Further, nonligand lyoprotectants (sorbitol, sugars, and PEG) also enhanced enzyme activity in organic solvents when present in aqueous solution prior to enzyme lyophilization. Addition of excipients to suspensions of native enzymes in organic solvents had no appreciable effect on enzyme activity indicating that the activation is based on the ability of the excipient to alleviate denaturation of enzymes during lyophilization. It has also been reported that adding surfactants[411] or hydrophobic sol– gel


564


materials[412] before lyophilization enhanced the lipase activity in organic solvents up to 100-fold.

D. Ion-Pairing Dordick and colleagues[413] described ion-pairing of biocatalysts in the presence of very low concentrations of ionic surfactants, which resulted in remarkably active ion-paired enzymes. The activities of subtilisin and a-chymotrypsin were .1000-fold higher than their native counterparts for the synthesis of peptides.[414] The high catalytic activity of ion-paired enzymes in organic solvents also paved the way for their incorporation into plastic materials for preparing “Biocatalytic Plastics.”[415]

E. Molecular Imprinting Molecular imprinting involves complex formation between a macromolecule and low-molecular weight ligand in solution, followed by drying and washing the solid complex with a selective solvent that removes the ligand but does not dissolve the polymer. Since the solid state offers only a very low molecular mobility of the polymer chain, the macromolecule retains the conformation induced by the ligand even after the ligand has been released from the complex. Russell and Klibanov[146] demonstrated the first example for molecular imprinting of subtilisin. Lyophilization of this enzyme from aqueous solution containing a competitive inhibitor, followed by the removal of inhibitor, created an enzyme with 100-fold more activity compared to that of native enzyme. This phenomenon was subsequently extended to a related protease, a-chymotrypsin.[416,417] In addition, ligand imprinting was also demonstrated with nonenzymatic proteins,[410,418] resulting in new adsorbents in organic media. Examples of successful molecular imprinting of a nonenzymatic protein with its transition state analogs to convert it into an enzymelike catalyst have been reported.[419,420] The proteins acquired the ability to catalyze b-elimination reaction in organic solvent, although the reaction rates were only modest. Rich and Dordick[421] demonstrated the activity of molecular imprinting to control the substrate selectivity of subtilisin-catalyzed acylation of nucleosides in organic solvents.

F. Trapping in Presence of Interfaces With the background of earlier molecular imprinting studies, Braco and coworkers ([422] and references cited therein) arrived at a rational strategy to enhance lipase activity in nonaqueous media. Most of the lipases activate at interfaces as a result of important rearrangements including the displacement of



565

the lid covering the active site and/or the adjustment of the catalytic machinery by means of forming an “oxyanion hole.” Braco employed a strategy named trapping in presence of interfaces (TPI), which is based on the following rationale: inducing activating conformational change in lipase (probably lid opening and/or fitting the machinery) in aqueous solution by binding to amphiphile interfaces, trapping the activated enzyme form by rapid freeze-drying, and further retaining the active form in the lyophilized powder by virtue of the conformational rigidity of proteins in anhydrous solvents. The outcome of this treatment is a higher reaction rate by the activated lipase compared to that of nonactivated counterpart.[422] While excipient addition was highly successful in achieving high activities in case of proteases, lipase activation was minimal in comparison.[422] When salts and sugars were incorporated in the lyophilization buffer, only a slight increase in activity was observed compared to TPI treatment. Given that in the closed conformation of lipase the lid covers the active site, it could be less susceptible to denaturation during lyophilization and hence lyoprotectants will have no major role in improving the lipase activity. Therefore, in case of lipases, the protection methods are insufficient to provide full activation. The TPI strategy seems to provide “interface memory” necessary to obtain the activated lipase.

G. Protein Engineering Despite hard competition from chemical modification and related techniques, protein engineering has become an increasingly important strategy.[423] The importance of protein engineering in industry continues to grow with the expanding range of protein applications. Extremophilic proteins isolated from organisms from extreme environments are emerging as an important source of new backbones for engineering proteins to attain new properties. The current strategies of protein engineering include rational design and directed evolution. The Lipase Engineering Database (accessible at: http:// www.led.uni-stuttgart.de), a useful resource on sequence– structure– function relationships of microbial lipases, includes information of 92 microbial lipase and homologus serine hydrolases (assigned into 15 superfamilies and 32 homologous families).[424] This database would be of aid to understand the functional role of individual amino acids and also to devise effective strategies for protein engineering of lipases.

1. Rational Design This was the earliest approach to protein engineering and is still widely employed to introduce desired characteristics into a target protein. This strategy hinges on relating structure to function, frequently via molecular modeling. The growing understanding of how to engineer certain basic enzyme properties (e.g.,


566


stability, activity, and surface properties) is beginning to make rational design more efficient. Advances in rational design depend on the progress made in structure determination, improved modeling protocols, and significant new insights into structure– function relationships. A new promising technique for obtaining structural details of the proteins that are difficult to crystallize is the twodimensional crystallization, which utilizes metal ion coordination to surface histidines in order to crystallize proteins at interfaces.[425] Dolder et al.[426] demonstrated the importance of the micelle – vesicle phase transition in the mechanism of two-dimensional crystallization for membrane proteins. Advances in modeling of free energy perturbation methods[427] and molecular dynamics calculations[428] form the basis for this area. Moult[429] discussed the state-of-theart in comparative modeling and ab initio protein structure predictions.

2. Directed Evolution Although improving enzymes by site-directed mutagenesis continues to be important, an alternate method—directed evolution—gained increasing attention from academic and industrial laboratories to modify and improve important biocatalysts.[430] Directed evolution combines random mutagenesis with screening or selection for the desired property and is especially useful for cases like solvent tolerance or thermostability where available theories are inadequate to predict which structural changes will give improvement. The goals currently envisaged are to efficiently modify either the enzyme or the reaction conditions to improve activity, stability, or selectivity. The major topic of current research is directed toward increased enzyme stability. Rational design, in this context, is inferior because the molecular basis for increased stability is ill defined. There are several examples to support this. Kidd et al.[431] mutated a surface lysine to tyrosine in subtilisin assuming that surface charge removal stabilizes the enzyme. The stability increased, but the crystal structure indicated that the increase was due to ˚ away. Arnold and an unidentified change in the weak calcium-binding site 12 A colleagues utilized random mutagenesis to create improved subtilisin E[432] and p-nitrobenzyl esterase having increased activities in DMF.[433] In both cases, the mutated amino acids were far from the active site and could not have been predicted using a rational design approach. Thermostability is also difficult to improve rationally and hence is a good target for directed evolution. Random mutagenesis and screening resulted in more thermostable subtilisin[434] and lipase.[435] Contrary to these successes, a rational approach to increase the thermostability of Penicillium camembertii lipase by introducing a disulfide link failed.[436] Random mutagenesis (using UV light or chemical mutagens) is long known to microbiologists for improving microbial strains. However, the excitement of directed evolution comes from other mutagenesis techniques such as error-prone



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PCR that targets a single gene or region of a gene ensuring that the improvement occurs only in the target protein but not in the genome. Directed evolution (also called molecular evolution, asexual PCR, or in vitro evolution) involves using PCR,[437] expressing the protein and then selecting or screening for those with improved properties. The DNA shuffling technique, introduced by Stemmer,[430,438] places the directed evolution approach apart from earlier random mutagenesis and screening efforts. Two significantly different ways of implementing directed evolution have been published: screening through single and then higher multiples of variants to cover sequence space systematically[432,433] and examination of large numbers of variants with multiple mutations using high definition screens or selection to find the best one.[430,437 – 439] The preferred approach may depend on the nature of the target and the type of selection or screens that can be developed. Enhanced enantioselectivity of a lipase[440] and an esterase[441] has been reported using directed evolution. The development of high-throughput screening methods that accurately reflect the target property is critical for success of directed evolution.[442] Not many publications on screening methods are available. Recently, Venekei et al.[443] described a simple and effective screening method for proteases. Janes et al.[444] developed a fast method for measuring the enantioselectivity of hydrolases, which might be useful for screening mutants with improved enantioselectivity. Reetz[445] provided an informative account on currently available and useful combinatorial and high-throughput screening systems. Bornscheuer’s group developed directed evolution strategy[441,446,447] as well as new methods to improve enantioselectivity.[448 – 450] Directed evolution was concentrated on the use of various recombinant esterases from P. fluorescens[451,452] and from Streptomyces diastatochromogenes.[453] Escherichia coli, Bacillus subtilis, and Saccharomyces cerevisiae have all been used successfully as host organisms for creating and expressing large and functional enzyme mutant libraries. Several choices are available for creating DNA libraries, e.g., error-prone PCR, combinatorial oligonucleotide mutagenesis, DNA shuffling, random-priming recombination (RPR), random chimeragenesis on transient templates (RACHITT), staggered extension process (StEP recombination), recombined extension on truncated templates (RETT), in vivo recombination. Whether mutations should be targeted to specific regions or distributed throughout the gene and for identifying improved variants (screens vs. selections) are the issues of debate. There is no agreement as to a single best approach and there may never be one, since they all address different needs and situations. The important problems tackled in the recent past include enhancing activity,[454,455] altering substrate specificity[456,457] and enantioselectivity,[440,441] and improving stability.[458,459] A variety of other features already exhibited by the parent enzyme at some (probably low) level were also improved.[460,461] Many reviews on several aspects of directed evolution have appeared in recent times.[445,462 – 466]


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3. Modeling of Lipase Selectivity To identify the amino acid residues that are functionally relevant or mediate stereoselectivity, x-ray structure determination, computer-aided molecular modeling, and protein engineering have been applied. Crystallographers have solved most lipase structures and paved the way for researchers to use computer modeling to rationalize the enantioselectivities of lipases.[467,468] Extensive research has also been directed to identify the binding regions of the acyl and alcohol substrates in various lipases and to rationalize the observed enantioselectivity. The current goals of modeling are directed mainly on three aspects: (i) to explain the known behavior of an enzyme at the molecular level, (ii) to suggest how to change the selectivity of a reaction by modifying the enzyme, substrate or reaction conditions, and (iii) to predict quantitatively the degree of selectivity of an enzyme-catalyzed reaction. Most lipases, certainly those approved for food applications, do not discriminate well between natural substrates containing saturated vs. polyunsaturated or long vs. short/medium chain carboxylic acids. It would be advantageous, from a practical viewpoint, to develop a biocatalyst that can selectively convert the substrates with acyl groups of up to a certain chain length or discriminate between cis- and trans-configurations of fatty acids. Towards this objective, Klein et al.[469] utilized molecular dynamic simulations of substrate binding to design mutations to change the acyl chain length selectivity. They predicted that replacement of Val209 and Phe112 with bulkier tryptophan residues in the acyl-binding groove of R. delemar lipase would sterically hinder the docking of long-chain (.C4) fatty acids. This prediction was confirmed in sitedirected mutagenesis experiments, where the double mutant (Val209Trp þ Phe112Trp) showed increased (.80-fold) selectivity for short-chain substrate. However, several other predictions did not give the desired results (e.g., replacement of Val94 with tryptophan, predicted to block the hydrolysis of substrates longer than caprylates, did not alter the chain length specificity). Similarly, Jorger and Haas[470] observed only modest (2– 3-fold) improvement in the activity toward tricaprylin in comparison to triolein, upon a single (Phe95Asp) and a double (Phe95Asp þ Phe214Arg) mutation. Gaskin et al.[471] evaluated the feasibility of altering the chain length specificity of R. miehei lipase by randomly mutating Phe94 (responsible for accommodating the acyl chain of the substrate) in the protein groove. They observed that the substitution of Phe94 with glycine resulted in an enzyme that displays about six times less activity against resorufin ester but exhibits 3 –4 times higher activity with short-chain (C4) ester substrates. At present, modeling cannot predict enantioselectivity quantitatively. The failure could possibly be due to omitting entropy contributions to enantioselectivity (most modeling programs calculate only enthalpy) and incomplete understanding of the origin of enantioselectivity. Mutagenesis trials with dehydrogenases suggest that changing substrate specificity requires not only mutations in the binding site, but also more distant ones that fine-tune the positions



569

of the main chain.[472] Instead of modeling, organic chemists usually predict selectivity qualitatively using empirical rules, which summarize the observed selectivity of lipases. One such useful rule predicts the fast reacting enantiomer of a secondary alcohol based on the sizes of the substituents at the stereocenter.[473] This rule has also been extended to primary amines (NH2ZCHZRZR1) that possess the same shape as secondary alcohols.[474] Parameters other than the size (e.g., electronic effects) also affect enantioselectivity.[475] However, this rule is not applicable for the primary alcohols with oxygen at the stereocenter. The hydroxyl group in the fast-reacting enantiomer of secondary alcohols protrudes out while in fast-reacting enantiomer of primary alcohol, the CH2OH group points away. Two reports[476,477] described modeling to reason out this apparent contradiction and both these models differ on the location of the large substituent. Zuegg et al.[476] suggested that the large substituent binds in the same place for both primary and secondary alcohol and the CH2OH group points away from the active site histidine to avoid disrupting the catalytic action. The second group,[477] on the other hand, suggested the presence of an “alternate hydrophobic pocket” (different part of the alcohol binding pocket, in other words an extension of the medium sized pocket) in which the large substituent binds. According to this model, the CH2OH group points in a different direction due to different kinds of binding of the two types of alcohols. This model strengthens the observation that the lipases from P. cepacia and porcine pancreas (both of which contain the alternate binding pocket ) are the main candidates that exhibit highest enantioselectivity toward primary alcohols. It also explains the case of primary alcohols having oxygen at the stereocenter, which do not obey the rule. The alternate binding pocket contains numerous tyrosines and one of them, Try29, may form a hydrogen bond with such substrates. The effects of this hydrogen bond are unpredictable. Lang et al.[478] determined the x-ray crystal structure of a transition state analog bound to the active site of P. cepacia lipase. This analog mimics ester hydrolysis at the primary ZOH group of a triglyceride analog. This structure indeed showed one substituent bound to the alternate hydrophobic pocket, which the authors called “HH pocket.” The x-ray structures of (a) R. miehei lipase complexed with a C6 phosphonate inhibitor, (b) C. rugosa lipase complexed with a long sulfonyl chain, of the human pancreatic lipase – colipase complex covalently inhibited by the two enantiomers of a C11 alkyl chain phosphonate, and (c) porcine pancreatic lipase covalently inhibited by ethyleneglycol monooctyl ether represent the breakthroughs in mimicking the natural tetrahedral intermediates. However, none of these compounds resemble a natural triglyceride. Cygler et al.[479] presented a first structural view of lipase stereoselectivity toward secondary alcohols by complexing (R )- and (S )-methyl ester hexylphosphonate transition state analog to C. rugosa lipase. In fast-reacting (R )-enantiomer, a hydrogen bond is present between the alcohol oxygen of the substrate and the active site histidine, while this bond is absent in slow reacting (S )-enantiomer. Based on these results, the authors proposed that this hydrogen bond is responsible for the stereoselectivity of


570


Scheme 10. Structure of transition state analog used for the x-ray crystallographic studies of the active site of P. cepacia lipase.

the enzyme. On the other hand, Uppenberg et al.[480] obtained similar results with C. antarctica lipase B complexed with a long-chain polyoxyethylene detergent and suggested that the hydrogen bond is present in both fast- and slow-reacting enantiomers. They proposed that the enzyme’s enantioselectivity could not be explained simply by the presence or absence of this hydrogen bond. Other factors such as the size of alcohol-binding pocket are suggested to play key roles as well. Longhi et al.[481] prepared a complex of cutinase with an enantiopue triglyceride analog. Unfortunately, this study, expected to reveal the stereoselective substrate interactions with protein, was a failure because the inhibitor was bound in the active site in an exposed position with alkyl chains. Lang et al.[478] provided a breakthrough by determining the x-ray structure of P. cepacia lipase complexed with an analog of medium alkyl chain length, RC-(RP,SP)-1,2-dioctyl carbamoylglycero-3-O-p-nitrophenyl octylphosphonate (TC8) (Sch. 10). The enzyme is in the open conformation with the lid displaced to allow access of the substrate analog and four binding pockets were detected: an oxyanion hole and three pockets for accommodating the sn-1, sn-2, and sn-3 fatty acid chains. The boomerang-shaped active site is divided into a large hydrophobic groove, in which the sn-3 acyl chain snugly fits, and a part that embeds the inhibitor’s alcohol moiety. The alcohol-binding pocket can be subdivided into a mixed hydrophilic/hydrophobic cleft for the sn-2 moiety of the substrate and a smaller hydrophobic groove for the sn-1 chain. Van der Waals interactions are the main forces that hold the radyl groups of the analog in position. A hydrogen bond between the ester oxygen atom of the sn-2 chain and the active site histidine contributes to guide the position of the inhibitor. This hydrogen bond is similar to the one observed by Cygler et al.[479] in case of C. rugosa lipase. The bound lipid analog assumes the bent-tuning-fork conformation preferred by lipids at an interface.[482] As the sn-2 pocket provides the true and intimate interactions with the substrate, this pocket is presumably the determinant of the enzyme’s stereopreferences. Zhang and Kazlauskas[483] modeled an enantio- and regioselective hydrolysis of a chiral bisphenol using Chromobacterium viscosum lipase. Although the stereocenter is situated far from the reaction site, the reaction is highly selective. It was suggested that the appropriate filling of the large alcoholbinding pocket is essential. This pocket tilts to one side in the reaction, the direction of which suits the orientation of fast-reacting enantiomer. Both smaller



571

and larger acyl groups would bind poorly because they either cannot fill the pocket or would extend out of the pocket. Another empirical rule based on modeling has been successful in predicting which enantiomer reacts faster in the case of carboxylic acids having stereocenter at the a-position. C. rugosa lipase shows high enantioselectivity for such substrates.[467,484] Modeling suggests that large substituents bind in a tunnel while the stereocenter (present at the a-position) lies near the mouth of this tunnel. Further modeling explained why addition of long-chain alcohols (which bind in the tunnel and prevent the normal productive orientation of fast reacting enantiomer) lowers the enantioselectivity.[467] Similarly, substrates containing branched, large substituents cannot fit in the tunnel and hence do not follow this model. The large, branched substituent lies outside the tunnel in the hydrophobic pocket. In such orientation, modeling predicted a preference for the opposite enantiomer, which was confirmed experimentally.[485] Holmquist and Berglund[486] further examined the importance of amino acid residues within the tunnel for the enantioselectivity based on site-directed mutagenesis experiments involving a related lipase from G. candidum, which also possess a tunnel. P. cepacia lipase is one of the most widely employed enzymes for the enantiomeric resolution of esters of secondary alcohols. This lipase shows preference for R over S compounds, but both enantiomers can be converted indicating that the less preferred enantiomer can also be productively bound to the active site. Modeling indicated that the acyl chain bound to the primary hydroxyl group of the glycerol moiety clashes with the hydrophobic side chains of Leu287 and Ile290. Hirose et al.[487] investigated, using site-directed mutagenesis, the importance of amino acid residues for stereoselectivity of P. cepacia lipase. They succeeded in altering the enantioselectivity from R to S specificity by introducing a combination of three mutations, Val266Leu, Leu287Ile, and Phe221Leu. Val266 is located at the entrance of the acyl (sn-3) pocket whereas Leu287 is at the beginning of the sn-2 pocket. Phe221 is present at the surface of the enzyme, ˚ away from the inhibitor. While the Val266Leu and Leu287Ile substitutions ,20 A are envisaged to affect the size and width of the respective pockets, Phe221 seems to be far away to directly influence the stereoselectivity of the enzyme. Nevertheless, the Phe221Leu mutation on its own was observed to decrease enantioselectivity slightly indicating the need for further probe to elucidate the role of Phe221 as an enantioselectivity-determinant factor. Moderate enantioselectivity towards triglycerides and their analogs is exhibited by R. oryzae lipase. Surprisingly, the favored enantiomer reverses upon changing the substituent at sn-2 position.[488] These authors employed computeraided modeling and used docking calculations to explain the reversals in enantioselectivity. They have grouped the substrates as per the flexibility of the b-bond in the sn-2 substituent. The R. oryzae lipase favors hydrolysis at the sn-1 position when the sn-2 substituent possess flexible bond (e.g., ether). But when a rigid bond (e.g., amide or phenyl) containing sn-2 substituent is present, hydrolysis is favored at sn-3 position. Modeling indicated that in all cases, the sn-2


572


substituent binds in a “hydrophobic dent,” which corresponds to an extension of the medium-sized binding pocket and is similar to the alternate hydrophobic pocket mentioned in the case of P. cepacia lipase. The sn-2 substituent is positioned near Leu258. When the substituent has a flexible b-bond, it can avoid clashes with Leu258; but when it has a rigid b-bond it can avoid clashes only by turning the substrate so that the sn-3 group is in the hydrolysis site. To test their hypothesis that enantioselectivity is based mainly on steric interactions with Leu258, the authors mutated Leu to Phe to increase the steric interactions. In accordance with the hypothesis, the enantioselectivity for the substrates with the rigid b-bond increased significantly.[489] However, no reversal is observed in case of R. miehei lipase, which prefers sn-1 position for all substrates.[490] A loop has been identified that makes the hydrophobic dent shallower, which in turn restricts the steric interactions and the rigid substrates cannot reverse enantioselectivity. As a major determinant of stereoselectivity, a structural element (the His gap ) has been identified. In Mucorales lipases, it consists of the catalytic His257 and its C-terminal Leu258 and is assumed to closely interact with the sn-2 substituent of the substrate. The central role of the His gap in determining stereoselectivty was further confirmed by mutating Leu258 in R. oryzae lipase as explained earlier. Varying size and physico-chemical properties of the His gap led to variants with distinct stereoselectivites. Pleiss et al.[491] investigated the role of His gap motif in other microbial lipases with known three-dimensional structures (C. rugosa lipase and C. antarctica lipase B). While stereoselectivity of C. rugosa lipase toward trioctanoin and sn-2 substituted analog is similar to R. miehei lipase, C. antarctica lipase B displayed opposite stereopreference toward the hydrolysis of trioctanoin. Based on these results, a general set of rules have been proposed to predict stereopreference of the lipases from R. oryzae, R. miehei, C. rugosa, and C. antarctica B as well as mutated R. oryzae. For all lipases tested, a single torsion angle (FO3 – C3) was found to correlate to the experimentally determined stereopreference of ester hydrolysis and allows predictions. From these investigations, it is clear that stereoselectivity is governed by three important factors: (i) flexibility of the sn-2 functional group of the triglycerides and analogs, (ii) shape and width of the His gap, and (iii) shape of the substrate binding site. In the “unnatural” environment, both activity and selectivity of lipases are strongly influenced by a number of process variables such as water activity, addition of various salts, formation of counterions, base effects, catalyst preparation, and the nature of immobilization support. Halling[492] provided an informative review on these factors summarizing the latest studies by various research groups on modeling of solvent effects. Hæffner and Norin[493] provided an account of molecular modeling of lipase-catalyzed reactions as applied to predicting enantioselectivities. Although use of modeling to predict the enantioselectivity is being used by numerous research groups, much of the available data is empirical and not easily interpreted at the molecular level. Modeling predicts which enantiomer reacts



573

faster in most cases. It is very difficult to predict the degree of enantioselectivity due to number of reasons: (i) difficulties in mimicking and modeling the transition state, (ii) finding the global minimum, (iii) need to find improved computational tools to calculate free energy, and (iv) difficulties in mimicking solvation. Kazlauskas[494] recently reviewed molecular modeling and limitations.

4. Combinatorial Biosynthesis There are also reports on metabolic pathway engineering by directed evolution. One significant example is the construction of an arsenate detoxification pathway.[495] The ability of directed evolution to simultaneously transfer a large number of genes has been utilized to engineer “unnatural natural products” and forms the basis of combinatorial biosynthesis. In this process, the alteration of gene sequences and gene products could be made in a combinatorial fashion to exploit the nature’s inherent diversity.[496] Of late, combinatorial biocatalysis is emerging as an important area. This employs iterative reactions catalyzed by isolated enzymes or whole cells, in a natural or unnatural environment, on substrates in solution or on a solid phase. Combinatorial biocatalysis harnesses the natural diversity of enzymatic reactions for iterative synthesis of organic compound libraries.[497,498] Use of lipases has led to the generation of a library of derivatives of dibenzyl-1,2-phenylenedioxy diacetate.[499] Lipases and proteases have also been employed in nonaqueous media to generate a library of acylated flavonoid derivatives ([498] and references cited therein).

VIII.

CONCLUDING REMARKS

Enzymatic reactions are no longer restricted to aqueous solutions and biocatalysis in nonaqueous media has significantly influenced the view of biochemists, chemists, and engineers towards enzymology. Chemists can now take advantage of enzyme specificities under mild conditions to catalyze reactions that were earlier limited to the use of tedious chemical catalysis. However, despite significant progress and expanding range of industrial applications, lipase catalysis is not yet very common in the industrial scene compared to carbohydrases and proteases. Biocatalysis is still not viewed as a first-line alternative, but as a course of last resort and attempted when other possible synthetic schemes fail. As a result, biocatalysis has not reached its full potential. Nevertheless, there are several industrially successful examples of biocatalytic processes and industry’s perception on biocatalysis is rapidly changing towards positive side. Biocatalysis has always been a key focus area in biotechnology and new approaches for the utilization of biocatalysts have been sparked by remarkable new insights into protein structure and function and from advances in genetic engineering. Despite much advancement, there is probably no clear scientific basis


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yet for a rational use of nonaqueous biocatalysis, due to lack of fundamental understanding. Several basic issues have yet to be resolved. The parameters that govern enzyme activity in nonaqueous media must be addressed to form a quantitative point of view. Diffusional limitations must be addressed to correlate intrinsic enzyme kinetics in various solvents. The role of water and the nature of the organic solvent on biocatalysis must be understood fully. While several lipases prefer hydrophobic solvents, there are exceptions. While water activity (aw) as a parameter instead of water concentration is well known, in our view a more detailed characterization of the effects of substrates, products and solvents, and the free energy relationships of these components may be beneficial for a better understanding of lipase catalysis in nonaqueous media. The role of pH in nonaqueous biocatalysis is rather a fuzzy concept, which clearly needs a better understanding. Lipases hold considerable promise in synthetic organic chemistry and have already found practical applications in detergents, oleochemistry, cheese production, medical therapy, and industrial production of specialty chemicals. Lipases from various (,30) sources have been cloned, sequenced, and expressed in host organisms. Pure recombinant lipases are already commercially available in free or immobilized form, or as a screening set (Chirazyme screening set from Roche). Even cross-linked crystals of lipases from C. rugosa and P. cepacia (ChiroCLEC) are commercially available (Altus, Cambridge, MA). Tertiary structures of .12 lipases have been resolved and this number will continue to grow due to their industrial importance. Consequently, rational approaches to modify lipases for specific application will flourish. Protein engineering is emerging as a major thrust area in improving enzyme activities and in finding novel applications. Recent successes in the directed evolution have brought this approach into limelight. On the other hand, chemical modification approach is also actively pursued by several researchers to gain insights into the activity or stability enhancement. While this approach is reemerging as a superior strategy, a need for comparative studies exists. Research progress on extremophilic proteins is also showing influence on the biocatalysis in general. Comparing mesophilic and thermophilic enzymes suggested that thermostability is evolutionary adaptation by increased conformational rigidity in thermophiles. The structural knowledge of extremophilic proteins would aid protein engineering to target specific amino acids of mesophilic proteins for modification in order to achieve activity or stability enhancement. It is also clear that the fundamental science is driving the biocatalysis forward. The field is getting more molecular and structural as researchers dwell to understand the basis for specificity to fine-tune the catalysts. Biocatalyst optimization could witness profound impact from the developments in gene shuffling and combinatorial screening. The broad field of biocatalysis is truly at the interface between chemistry and biology. Advances in both the fields through improved techniques in molecular biology or development of chemo-biocatalytic synthetic strategies are



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revolutionizing the chemical and pharmaceutical industries. The increased awareness of chemists of the synthetic power of biocatalysis and the increased interest of biologists and biochemical engineers in synthetic chemistry will surely enhance the importance of biocatalysis significantly. The contribution of the chemist is to probe the specificity of the available enzymes by designing substrates to generate useful products. The current market for chiral drugs is about $115 billion worldwide. It is estimated to grow up to $146 billion by 2003.[500] The global market for industrial and specialty enzymes is about $1.5 billion in 1998 and is ever increasing with a predicted 5 – 10% growth per annum [http://www.jic. bbsrc.ac.uk/exhibitions/bio-future/industry.htm]. Fewer than 30 enzymes account for more than 90% of the industrial enzymes currently in use. It is expected that in the years to come, the exchange of knowledge and experience between crystallographers, chemists, biochemists, geneticists, and enzymologists on one hand and food, chemical and biochemical engineers on the other would create an indepth understanding of lipase catalysis. Biocatalysis is beginning to have an impact on chemical process development in the pharmaceutical and agricultural industries. Industry has played a crucial role in applying the biocatalysis from the laboratory to the industry. This trend will almost certainly continue with increasing vigor. It is hoped that this review would help to achieve this broader goal. ACKNOWLEDGMENTS Authors thank the Director, CFTRI, Mysore for encouragement and providing facilities. One of the authors, S.H.K. is grateful to the Council of Scientific and Industrial Research, New Delhi, India for the award of Senior Research Fellowship. Mr. K. C. Sekhar is thanked for help in preparation of the manuscript.

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