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Specificity of a Lipase in Ester Synthesis: Effect of Alcohol Neena N. Gandhi, Sudhirprakash B. Sawant,’ and Jyeshtharaj B. Joshi Department of Chemical Technology, University of Bombay, Matunga, Bombay-400 019, India
Ester synthesis by the Mucor miehei lipase has been studied for various alcohol substrates: n-propanol, n-butanol, isoamyl alcohol, n-hexanol, n-octanol, 2-ethylhexanol, n-decanol, and lauryl alcohol. The effects of temperature, the nature of the acid, and immobilization of the lipase on its substrate specficity have been elucidated by carrying out esterifications a t 29 and 50 “C with lauric and oleic acids and by using both the soluble and immobilized (resin-adsorbed) forms of the lipase as catalysts. Higher synthesis rates were obtained with oleic acid than with lauric acid. A bimodal distribution pattern was observed for the reaction rate as a function of alcohol chain length. Two superimposed “bells” were obtained with maxima a t C4 (butanol) and C10 (decanol) a t 29 “C. Whereas immobilization of the lipase did not influence this substrate specificity, an increase in temperature to 50 “C caused a shift in the first peak from C4 to C6 (hexanol), while the second peak position was not affected. The minimum, in all cases, was found to be a t C8 (octanol).
Introduction Esters of fatty acids and alcohols have applications in industries such as foods, cosmetics, emulsifiers and lubricants, and flavors and fragrances, (Bauer et al., 1990; Kao Corporation, 1987). For instance, butyl laurate is used as a plasticizer (Hawley, 1981)and in flavor compositions, mainly for apricot and peach flavors, usually accompanied by other low-boilingesters (Arctander, 1969). Butyl oleate has applications in flavors, as a plasticizer, particularly for PVC as a solvent, as a lubricant, in waterproofings, and in polishes (Hawley, 1981,1991). Such esters can be produced by the reaction of an alcohol with a fatty acid using ion-exchange resins (Bhagade and Nageshwar, 1978; Dassy et al., 19941, inorganic acids (Newman, 1941),p-toluenesulfonic acid, and enzymes (Barros et al., 1989; Ishii et al., 1990) as catalysts. Common practice for the manufacture of these esters involves catalytic hydrolysis or esterification directed by alkali metals at high temperatures (100-300 “C) and/or extremes of pressure (Benson, 1967; Swern, 1964). This not only leads to high operating costs but also results in poor selectivity, undesirable side reactions, such as fatty acid polymerization and autoxidation, and formation of impurities like color bodies. The latter results in the need for elaborate refining procedures. The use of enzymes to catalyze esterification/alcoholysis has become a more promoting method than acid- or base-catalyzed reactions for ester production (Pecnik and Knez, 1992). Enzyme-catalyzed reactions are superior to conventional chemical methods due to mild reaction conditions, high catalytic efficiency, and inherent selectivity of natural catalysts (Linfield et al., 1984; Sonnet, 1988), which results in much purer products. Lipases (E.C. 3.1.1.3)belong to the enzymatic class ofhydrolases and are convenient enzymes for ester production. Although the natural reaction they catalyze is that of triacylglycerol hydrolysis, they are found to catalyze the reverse reaction of synthesis just as efficiently. Their substrate specificity is broader than that of many other known enzymes, ranging from the natural substrate of * Author to whom correspondence should be addressed.
glycerides to aliphatic and aromatic acids and alcohols to their esters to thiols, amines, sugars, and amides (Bhirud et al., 1991a,b; Ikeda et al., 1991). Hence, a single lipase can be utilized for the synthesis of a variety of compounds. The broad substrate spectrum of a number of lipases has been documented by various workers. Rees et al. (1991) studied the affinity of the Chromobacterium viscosum lipase (entrapped in microemulsion-based organogels) for primary, secondary, and aromatic alcohols and acids. Pecnik and Knez (1992)compared the transesterification of C4-C8 alcohols with vinyl acrylate using Candida cylindracea. Langrand et al. (1990)investigated the catalytic potential of lipases from Aspergillus species, Rhizopus arrhizus, Mucor miehei, and Candida rugosa in n-heptane. They covered Cl-C6 alcohols, geraniol, and citronellol with C2-C6 acids. The substrate specificity of lipases (about 9-10) for various alcohols, sugars, thiols, and amines was studied in an aqueous medium by Lazar et al. (1986). A similar study on four other microbial lipases was reported by Okumura et al. (1979). It was noted that, in all of the preceding cases, the yields after a specified reaction period have been considered. It is likely that equilibrium may have been attained in some of them. Hence, a comparison of yields may prove erroneous. A better indicator of the substrate affinity of the lipase would be the initial activity. Another observation we made during our literature survey was that while the effect of acyl and alcohol chain length on ester synthesis by several lipases has been studied (Takahashi et al., 1988; Ishii et al., 19901, not much information is available on the alcohol specificity of the M . miehei lipase. Gatfield (1984), Miller et al. (1988),and Manjon et al. (1991)studied the effect of acid chain length. Knez et al. (1990) worked with the lower alcohols (Cl-C5). Both studies are reported for the immobilized lipase, lipozyme. To our knowledge, there has been no study on the substrate specificity of the free M . miehei lipase. Also, in most cases, a solvent has been used. This has the effect of reducing the substrate concentration “seen” by the enzyme, so that a clear picture of the exclusive effect of the nature of the substrate on enzyme activity may not emerge. The solvent can also affect the polarity of the medium, which
0 1995 American Chemical Society and American Institute of Chemical Engineers 8756-7938/95/3011-0282$09.00/0
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in turn affects the reaction rate. It may also cause the partitioning of water from the microenvironment of the lipase. The effect of temperature on lipase selectivity also has not been investigated. A detailed study of the effects of temperature and immobilization on lipase substrate selectivity,in conjunction with supporting physical data, could prove useful toward elucidating the structurefunction relationship of the lipase molecule. The present communication is an endeavor toward bridging this gap. A variety of medium-chain laurates and oleates have been synthesized in solvent-free medium using both soluble and immobilized forms of the M . miehei lipase, lipozyme.
Experimental Procedures Materials. Lipozyme-10.000L is a M. miehei lipase from Novo Industries (Bagsvaerd, Denmark). The dark brown liquid has a reported activity of 10 000 LU/g (LU, lipase unit with respect to micromoles of butyric acid liberated by tributyrin hydrolysis per minute at 30 "C and pH 7). Lipozyme-TM, a M . miehei-derived lipase, was obtained from Novo Industries, immobilized onto a macroporous anion-exchangeresin of particle size 300-600 pm, with a reported activity of 25 BIU/g [BIU, batch interesterification unit, defined as micromoles of palmitic acid incorporated into triolein per minute at 40 "C, in a reaction system consisting of 12 mL of an equimolar mixture of triolein and palmitic acid (56.6 mM each) using 275 mg of lipozymel (Novo Industries Ah,1986). Lauric acid wgs obtained from Loba Chemie (Bombay, India), and oleic acid was from Bombay Oil Industries Ltd. (Bombay). Isoamyl alcohol, 2-ethylhexanol, n-butanol, sodium hydroxide, and methanol were procured from S. d Fine-chem Ltd., and lauryl alcohol was from SRL (Bombay, India). n-propanol, n-hexanol, and noctanol were from CDH (Bombay, India). n-Decanol was obtained from Merck-Schuchardt(Schuchardt,Germany). All chemicals were of LR grade. All reagents and enzymes were used without further purification. Apparatus. The glass reactor had a capacity of 100 mL and a diameter of 40 mm. It had a close-fitting, fournecked lid and was fitted with a pitched-blade turbine impeller consisting of six blades. Four baffles of diameter 3 mm each were also provided. The entire reactor assembly was mounted in a thermostatic water bath, which was maintained at the reaction temperature with an accuracy of f l "C. Methods. 1. Using Soluble Lipozyme. Acid (0.1 mol) and alcohol (0.2 mol) were mixed and equilibrated at the reaction temperature and placed in the reaction vessel. Esterification was initiated with 0.5 mL of enzyme, diluted to 5 mL with distilled water. The speed of agitation was 33.3 revis (rev, revolutions). This procedure, along with the specified substrate concentrations, temperature, etc., was followed in all cases. Any variations in this procedure will be stated in the text when necessary. 2. Using Lipozyme-TM Resin Beads. In this case, a mixture of 0.1 and 0.2 mol of acid and alcohol, respectively, was placed in the reactor and equilibrated at the reaction temperature. Lipozyme beads (250 mg) were added to initiate the reaction. The stirring speed was 2.5 reds. No water was added in this case. Variations in any of the experimental conditions will be specifically mentioned. Analysis. Aliquots (0.5 mL) of the reaction mixture were withdrawn periodically. Each sample was dissolved in 10 mL of methanol and analyzed titrimetrically for
60
t
TI ME (SIx IO-] Figure 1. Progress curves for various alcohols at 29 "C.
Reaction conditions: 0.2 mol of alcohol; 0.1 mol of lauric acid; 0.5 mL of soluble lipozyme diluted t o 5 mL with distilled water; 33.3 reds; 29 "C. Symbols used are as follows: (+) n-propanol; (*) n-butanol; (A) isoamyl alcohol; (0)n-hexanol; ( 0 )2-hethylhexanol; ( x ) n-octanol; (e) n-decanol; (H) lauryl alcohol.
the residual acid content using sodium hydroxide with phenolphthalein as an indicator. The percentage conversion and/or the moles of acid reacted were calculated by comparing the obtained titers with that at the commencement of the reaction.
Results and Discussion 1. Esterification Using Soluble Lipozyme. Experiments were conducted at 29 and 50 "C. Results are shown in Figures 1-4. It may be noted that since the reaction medium is a water-in-oil dispersion the rates and/or conversions obtained would be determined by the interfacial area, which, in turn, is a function of the speed of agitation and the ratio of the volumes of the aqueous and organic phases. For a given ratio of the latter, the interfacial area is dependent on the speed of agitation. As this area increases, a greater number of enzyme molecules will move out of the interior and become adsorbed onto these interfacial sites. Once all the molecules have occupied sites at the interface, however, any increase in interfacial area due to the increase in agitation speed will have no effect. All of the experiments have, therefore, been conducted within this zone. The following observationswere made: (a) Overall, the degree of conversion was found to vary with the substrate (Figures 1and 2). This is a reflection of the interplay of the influence of various parameters, such as the molecular size, hydrophobicity, and solubility of the alcohol in question, in the enzyme-containing aqueous phase. (b)A bimodal distribution pattern was obtained for the plots of initial rate versus aliphatic carbon chain length of the alcohols at both temperatures: the first phase for C < 8 and bhe second one for C > 8. Each phase displayed a bell-shaped distribution (Figure 3). Langrand et al. (1990) observed a similar pattern for immobilized lipozyme (as a function of acyl chain length), as well as for other lipases. For acids with M . miehei lipase, they observed two superimposed bell-shaped distributions with maxima at C2 and C5. For Candida cylindrucea lipase, Engel et al. (1991) obtained maxima for acids at C4 and C8. According to these authors, immobilization as well as temperature can shift these maxima. This was observed in the present case (for alcohols) too. At 29 "C, the maximum was at C4 in the first phase, which shifted to C6 at 50 "C. The peak for
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. -
TIME (SIx
lo-'
Figure 2. Progress curves for various alcohols at 50 "C. Reaction conditions: 0.2 mol of alcohol; 0.1 mol of lauric acid; 0.5 mL of soluble lipozyme diluted to 5 mL with distilled water; 33.3 reds; 50 "C. Symbols used are as follows: (+) n-propanol; (*) n-butanol; (a)isoamyl alcohol; (0) n-hexanol; (0)2-ethylhexanol; ( x ) n-octanol; ( 0 )n-decanol; (m) lauryl alcohol.
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z
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8w 3
8V w
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a
CARBON CHAIN LENGTH
Figure 3. Initial reaction rate as a function of alcohol chain length. Reaction conditions: 0.2 mol of alcohol; 0.1 mol of lauric acid; 0.5 mL of soluble lipozyme diluted to 5 mL with distilled water; 33.3 reds; 29 and 50 "C. Symbols used are as follows: (*) 29 "C; ( 0 )50 "C; (2-ET) 2-ethylhexanol.
the second bell was found to be at C10 (for both temperatures, however). Ergan et al. (1990) reported a definite optimum chain length(s) for a particular lipase. This different affinity of a lipase molecule for varying substrates can be understood in terms of the binding energy (Malcata et al., 1992) that is released when a substrate binds at the active site. Only a few of the many substrates that bind at the active site can release a sufficient amount of binding energy required for effecting a change in conformation of the lipase to a form that is a more efficient catalyst (induced-fitmodel for enzyme action) (Dixon and Webb, 1979; Koshland, 1959; Laidler, 1951). Substrates such as propanol, which is too small, are not able to release enough energy, so that the change in conformation of the native lipase to the desired catalytically active form does not occur or is at best incomplete. Hence, the reaction will proceed slowly. On the other hand, substrates that are too long (for instance, lauryl alcohol) are able t o release an amount of binding energy that ought
to be sufficient to effect the desired conformational change. However, some of this energy is required to change the conformation of the substrate so as t o make it fit into the active site. Hence, only a small fraction of the energy released by the binding process will actually be made available to derive the conformational change of the enzyme. Consequently, optimum activity will not be achieved. Malcata et al. (1992)had proposed a similar explanation for the occurrence of bimodal distribution curves. (c) No reaction with n-octanol and 2-ethylhexanol occurred at either temperature (Figures 1 and 2). The nonreactivity of 2-ethylhexanol can be explained by taking into account the steric considerations. However, the fact that n-octanol is also a C8 alcohol may mean that the enzyme has a very low affinity for a C8 substrate. This may be understood by considering the existence of two alcohol-binding sites on the lipase molecule, along the same lines as Parida and Dordick's (1993) postulation of two acyl-binding pockets in the active site of the Candida cylindracea lipase molecule. Parida and Dordick (1993) obtained curves similar to [where V, is the those in Figure 3 for plots of V,,lK, maximum initial reaction rate obtained (with respect to substrate concentration) and K, is the Michaelis constant] against the chain length of 2-hydroxy acids. In their case, a minimum was obtained at C6 for different experimental conditions. The authors attributed this result to the existence of a small pocket for acids with chain length C < 6 (an esterase site, acting mainly on soluble substrates) and of a larger pocket for acids with C > 6 (typical lipase active site, acting preferentially on insoluble substrates). This hypothesis was supported by enantioselective data (which emphasizes the importance of substrate orientation in catalysis). We therefore, hypothesize a similar two-alcohol binding site model, with a C8 alcohol not fitting into either site properly. The orientation of 2-ethylhexanol also may not be favorable. Esterification of octanol by immobilized M. miehei lipase has been reported previously by Bhirud et al. (1991b) and Kirk et al. (1992). However, upon careful analysis of their work, it was found that the quantities of enzyme used in most cases were much higher than those used in our experiments. To confirm this, a reaction was carried out using 5 mL of undiluted lipase (as opposed to the usual 0.5 mL diluted to 5 mL with distilled water). Oleic acid and n-octanol were taken in the usual 0.1:0.2 molar ratio, and the reaction mixture was stirred magnetically at 30 "C. In 7 h, about 7% conversion was obtained, and it kept increasing thereafter. Thus, what appeared to be nonreaction was actually reaction limited by the quantity of enzyme employed (implying that the specific activity of the enzyme with respect to esterification with n-octanol is very low). Hence, it may be stressed that whenever the low reactivity of C8 alcohols is mentioned in this communication, it should be regarded in the context of the experimental conditions. (d) While lower polar alcohols (C3-C5) showed lower rates at 50 "C than at 29 "C due to deactivation of the enzyme, the converse trend was observed for the higher alcohols (C10, C12). This result is explained in detail in the following. An increase in temperature would be liable t o change the lipase activity distribution toward longer carbon chain length, because the solubility of the lower alcohols (C I5) in the aqueous enzyme-containingphase would increase with temperature. Therefore, the lipase would be exposed to higher alcohol concentrations, resulting in increased denaturation (and, thus, reduced activity).
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Lipozyme deactivation by butanol has been reported (for the immobilized form) by Mukesh et al. (1993a,b). This effect will be expected to intensify with increasing chain length and increased hydrophobicity of the alcohol, resulting in deactivation due to the solvation of internal hydrophobic amino acid residues in the protein molecule and the attendant disruption of the enzyme's threedimensional structure (Wong and Wong, 1992). However, the higher alcohols (C > 6) have little or no solubility in the aqueous phase. Any slight increase in temperature may only serve to enhance the accessibility of the substrate to the active site of the lipase. Hence, a shift of the peak of the first bell to the right with an increase in temperature from 29 to 50 "C is expected. This shift was observed. (e) For butyl laurate synthesis, it was observed that, at 50 "C, for up to 40 min, 13% conversion was obtained at a rate similar to that at 29 "C. Beyond this conversion, however, there was no further reaction. The aqueous phase with the enzyme was separated and contacted under otherwise identical conditions with a fresh mixture of substrates at 29 "C. No esterification was observed, indicating deactivation of the enzyme. It was found that the deactivation was thermally irreversible. On the basis of this observation, the laurates and oleates of other alcohols were also synthesized at 50 "C, so as to elucidate the behavior of the enzyme in the presence of other alcohols at that temperature. Similar results were obtained with n-propanol, which is a good substrate for the enzyme at 29 "C, but at 50 "C, irreversible thermal deactivation is found to occur, and again no effect of water addition was observed. However, in this case, the deactivation was instantaneous (unlike butanol, where reaction was observed to take place for the first 40 min). The intermediate concentrations (Fink, 1987)of watermiscible alcohols are known to facilitate protein denaturation by solvating out internally placed hydrophobic amino acid residues, i.e., they stabilize the denatured as compared to the native configuration. The degree of protein denaturation by lower molecular weight alcohols increases with increasing carbon length (up to C5) and the capacity to form hydrogen bonds, for example, C4 > C3 > C2 > C1. [For higher homologues, a decrease in water miscibility or solubility results in a decline in the denaturation effect with increasing carbon chain length.] The higher deactivating effect with propanol than with butanol appears to be in contradiction with this observation. However, the considerably high miscibility of propanol relative to butanol may prove a weightier factor than the slight difference in hydrophobicity. This deactivation theory can be used to explain the leveling off of the curves for the lower alcohols in Figures 2 and 4. (f) When oleic acid was reacted with butanol, a trend was observed similar to that for lauric acid at the given temperature, with the inhibitory effect of butanol at 50 "C, also seen in the case of oleic acid (Figure 4). There was a slight increase in rate and conversion with an increase in chain length and degree of unsaturation of the acid. At 50 "C, the rates obtained with both acids were similar. 2. Esterification Using Immobilized Lipozyme. As before, 0.1 and 0.2 mol of lauridoleic acid and alcohol, respectively, were used. Enzyme beads (250 mg) were used to initiate the reaction. All experiments were carried out at 30 "C at an agitation speed of 2.5 revls. Results are shown in Figures 5-7. (a) For alcohols up to C8, a slight increase in reaction rate was observed for oleic acid as compared to lauric
285
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Figure 4. Progress curves for different acids at 29 and 50 "C. Reaction conditions: 0.2 mol of butanol; 0.1 mol of acid; 0.5 mL of soluble lipozyme diluted to 5 mL with distilled water; 33.3 reds. Symbols used are as follows: (+) lauric acid (29 "C); (A) lauric acid (50 "C); (0)oleic acid (29 "C); (*) oleic acid (50 "C).
/
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Figure 5. Progress curves for various alcohols with lauric acid. Reaction conditions: 0.2 mol of alcohol; 0.1 mol of lauric acid; 250 mg of lipozyme beads; 2.5 reds; 30 "C. Symbols used are as follows: (+) n-propanol; (*) n-butanol; (A) isoamyl alcohol; ( 0 )n-hexanol; ( x ) n-octanol; (0)n-decanol; (m) lauryl alcohol.
acid. Thus, oleic acid appears to be a better substrate for lipozyme than lauric acid. The reportedly higher selectivity of this lipase for long-chain acids (Langrand et al., 1990; Marlot et al., 19851, thus, is evident in our case as well. This observation was similar to that observed for the acids using the soluble enzyme. (b) With n-octanol both acids gave very low conversions. However, unlike the soluble lipozyme case, there was some reaction. Upon separation of the enzyme beads from the preceding reaction mixture and addition of the same to a 1:2 standard lauric acidhutanol mixture, the activity of the enzyme was the same as that obtained previously using fresh enzyme (beads) with a lauric acid/ butanol mixture. Again, this points to the low affinity of the M . miehei lipase for this alcohol. Miller et al. (1988) obtained high synthesis rates for octyl ester synthesis with acids of carbon chain length C5-C10. Kirk et al. (1992) carried out the synthesis of the octyl esters of C7-Cl8 fatty acids. Bhirud et al. (1991b) also carried out the alkylation of stearyl amine with C8-Cl4 alcohols in isopropyl alcohol as solvent,
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I
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Conclusions The activity distribution of the M . miehei lipase with respect to its alcohol substrates was found to be a bimodal one. A shift in the maxima loci was observed upon
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raising the temperature, while immobilization appeared to have no effect. The minimum activity was obtained with C8 alcohols in all cases. Higher activity was obtained with oleic acid than with lauric acid.
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(e) Degrees of conversion varied with different reactants as expected, which may reflect their varied hydrophobicities, molecular sizes (and hence), diffusion rates, and kinetic rates.
Acknowledgment N.N.G. is grateful to CSIR, India, for the research fellowship provided.
702 TIME (5)
10')
Figure 6. Progress curves for various alcohols with oleic acid. Reaction conditions: 0.2 mol of alcohol; 0.1 mol of oleic acid; 250 mg of lipozyme beads; 2.5 reds; 30 "C. Symbols used are as follows: (+) n-propanol; (*) n-butanol; (A) isoamyl alcohol; (0) n-hexanol; ( x 1 n-octanol; ( 0 )n-decanol; (B) lauryl alcohol.
CARBON CHAIN LENGTH
Figure 7. Initial reaction rate as a function of alcohol chain length. Reaction conditions: 0.2 mol of alcohol; 0.1 mol of acid; 250 mg of lipozyme beads; 2.5 reds; 30 "C. Symbols used are as follows: (*) lauric acid; ( 0 )oleic acid.
with good yields. All of these studies were carried out using lipozyme beads. As mentioned for the soluble lipozyme case, an explanation for this may be found in the fact that the enzyme quantities used were considerably higher than those used by us. (c) There appears to be no effect of branching in the alcohol carbon chain or secondary nature of alcoholic functionality in this case (as also for the soluble lipozyme), since activity with isoamyl alcohol falls well in line with that of other alcohols. (d) The lower conversion rate of lauryl alcohol compared to n-butanol and n-hexanol is probably due to the much longer carbon chain and, hence, the higher hydrophobicity (resulting in lower concentrations of the substrate in the vicinity of the enzyme). A similar observation was made for soluble lipozyme. A bimodal distribution pattern for activity with respect to the aliphatic carbon chain length of alcohols was observed with the maximum of the first bell at C4 and that of the second at C10,similar to that of the soluble enzyme. Thus, immoblization appeared to have no effect on the activity distribution of lipozyme.
Literature Cited Arctander, S. Perfume Flavor Chemicals; Montclair, NJ, 1969; Vols. I and 11. Barros, M. R. A.; Cabral, J. M. S.; Willson, B. C.; Hamel, J.-F. P.; Cooney, C. L. Esterification-coupled Extraction of Organic Acids: Partition Enhancement and Underlying Reaction and Distribution Equilibria. Biotechnol. Bioeng. 1989,34,900915. Bauer, K.; Garbe, D.; Surburg, H. Common Fragrance and Flavor Materials, 2nd revised ed.; VCH Publishers: New York, 1990. Benson, F. R. Poly01 Surfactants. In Nonionic Surfactants: Surfactant Science Series; Schick, M. J., Ed.; Marcel Dekker: New York, 1967; Vol. 1, pp 247-299. Bhagade, S. S.; Nageshwar, G. D. Catalysts by Ion Exchange Resins. Esterification. Chem. Petro-Chem. J . 1978,9,3-12. Bhirud, V. S.; Subrahmanyam, V. V. R.; Vaidya, S. D. Influence of Reaction Media on Esterification Catalyzed by Mucor miehei Lipase. J . Oil Technol. Assoc. India 1991a,44-47. Bhirud, V. S.; Subrahmanyam, V. V. R.; Vaidya, S. D. Alkylation of Primary Amines with Fatty Alcohols using Immobilized Mucor miehei Lipase as Catalyst. J.Oil Technol. Assoc. India 1991b,47-48. Dassy, S.; Wiame, H.; Thyrion, F. C. Kinetics of the Liquid Phase Synthesis and Hydrolysis of Butyl Lactate Catalyzed by Cation Exchange Resin. J . Chem. Technol. Biotechnol. 1994, 59, 149-156. Dixon, M.; Webb, E. C. The Enzymes; Longman: London, 1979; p 267. Engel, K.-H.; Bohnen, M.; Dobe, M. Lipase-catalyzed Reactions of Chiral Hydroxyacid Esters: Competition of Esterification and Transesterification. Enzyme Microb. Technol. 1991,13, 655-660. Ergan, F.; Trani, M.; Andre, G. Production of Glycerides from Glycerol and Fatty Acid by Immobilized Lipase in Nonaqueous Media. Biotechnol. Bioeng. 1990,35,195-200. Fink, A. L. Acyl Group Transfer-The Serine Proteinases. In Enzyme mechanisms; Page, M. I., Williams, A., Eds.; Royal Society of Chemistry: London, 1987; pp 159-177. Gatfield, I. L. The Enzymatic Synthesis of Esters in Nonaqueous Systems. Ann. N.Y. Acad. Sci. 1984,434,569-572. Hawley, G. G., Ed. The Condensed Chemical Dictionary; Van Nostrand Reinhold Co.: New York, 1981. Ikeda, I.; Tanaka, J.; Sukuki, K. Synthesis of Acrylic Esters by Lipase. Tetrahedron Lett. 1991,32,6865-6866. Ishii, T.; Mori, T.; Chen, J.; Itoh, Y.; Shimura, S.; Kirimura, K.; Usami, S. Ester Synthesis by a Crude Lipase of Rhizopus oligosporus in an Aqueous System. J . Ferment. Bioeng. 1990, 70, 188-189. Kao, Corporation. Preparation of Lower Alcohol Fatty Acid Ester by Alcoholysis of Oil and Lower Alcohols in Presence of Thermoresistant Lipase. Jap. Patent JP 1,010,094, 1987. Kirk, 0.;Bjorkling, F.; Godtfredsen, S. E.; Larsen, T. 0. Fatty Acid Specificity in Lipase-catalyzed Synthesis of Glucoside Esters. Biocatalysis 1992,6, 127-134.
207
Biotechnol. Prog., 1995, Vol. 11, No. 3 Knez, Z.; Leitgeb, M.; Zavrsnik, D.; Lavric, B. Synthesis of Oleic Acid Esters with Immobilized Lipase. Fat Sci. Technol. 1990, 92,169-172. Koshland, D. E., Jr. Mechanisms of Transfer Enzymes. In The Enzymes, 2nd ed.; Boyer, P. D., Lardy, H., Myrback, K., Eds.; Academic Press: New York, 1959;Vol. 1, pp 305-346. Laidler, K. J. The Influence of Pressure on the Rates of Biological Reactions. Arch. Biochem. 1961,30,226-236. Langrand, G.; Rondot, N.; Triantaphylides, C.; Baratti, J. Short Chain Flavor Esters Synthesis by Microbial Lipases. Biotechnol. Lett. 1990,12,581-586. Lazar, G.; Weiss, A.; Schmid, R. D. Synthesis of Esters by Lipases. In Proceedings of the World Conference on Emerging Technologies in the Fats and Oils Industry; Baldwin, A. R., Ed.; American Oil Chemists’ Society: Champaign, IL, 1986; pp 346-354. Linfield, W. M.; Barauskas, R. A.; Sivieri, L.; Serota, S.; Stevenson, R. W., Sr. Enzymatic Fat Hydrolysis and Synthesis. J . Am. Oil Chem. SOC.1984,61,191-195. Malcata, F. X.; Reyes, H. R.; Garcia, H. S.; Hill, C. G., Jr.; Amundson, C. H. Kinetics and Mechanisms of Reactions Catalyzed by Immobilized Lipases. Enzyme Microb. Technol. 1992,14,426-446. Manjon, A.; Iborra, J. L.; Arocas, A. Short-chain Flavor Ester Synthesis by Immobilized Lipase in Organic Media. Biotechnol. Lett. 1991,13,339-344. Marlot, C.; Langrand, G.; Triantaphylides, C.; Baratti, J. Ester synthesis in organic solvent catalyzed by lipases immobilized on hydrophilic supports. Biotechnol. Lett. 1985,7,647-650. Miller, C.; Austin, H.; Posorske, L.; Gonzlez, J. Characteristics of an Immobilized Lipase for the Commercial Synthesis of Esters. J . Am. Oil Chem. Soc. 1988,65,927-931. Mukesh, D.; Banerji, A. A.; Newadkar, R.; Bavinakatti, H. S. Lipase Catalyzed Transesterification of Vegetable Oils-A Comparative Study in Batch and Tubular Reactors. Biotechnol. Lett. 1993a,15,77-82.
Mukesh, D.; Iyer, R. S.; Wagh, J. S.; Mokashi, A. A.; Banerji, A. A.; Newadkar, R.; Bevinakatti, H. S. Lipase Catalyzed Transesterification of Castor Oil. Biotechnol. Lett. 1993b,15, 251-256. Newman, M. S. A New Method for the Esterification of Certain Sterically Hindered Acids. J . Am. Chem. Soc. 1941, 63, 2431-2435. Novo Industries Ms. NOVO Method for the Determination of Lipase Batch Interesterification Activity; Novo Industries: Bagsvaerd, Denmark, 1986. Okumura, S.; Iwai, M.; Isujisaka, Y. Synthesis ofvarious Kinds of Esters by Four Microbial Lipases. Biochim. Biophys. Acta 1979,575,156-165. Parida, S.;Dordick, J. S. Tailoring Lipase Specificity by Solvent and Substrate Chemistries. J . Org. Chem. 1993,58,32383244. Pecnik, S.;Knez, 2. Enzymatic Fatty Ester Synthsis. J . Am. Oil Chem. Soc. 1992,69,261-265. Rees, G. D.; Nascimento, M. G.; Jenta, T. R. J.; Robinson, B. H. Reverse Enzyme Synthesis in Microemulsion-based Organogels. Biochim. Biophys. Acta 1991,1073,493-501. Sonnet, P. E. Lipase Selectivities. J . Am. Oil Chem. SOC.1988, 65,900-904. Swern, D.Bailey’s Industrial Oil and Fat Products; J. Wiley: New York, 1964. Takahashi, K.; Saito, Y.; Inada, Y. Lipases made Active in Hydrophobic Media. J . Am. Oil. Chem. SOC.1988,65,911916. Wong, S. S.; Wong, L.-J. C. Chemical crosslinking and the stabilization of proteins and enzymes. Enzyme Microb. Technol. 1992,14,866-874. Accepted October 3, 1994.@ BP940089A *Abstract published in Advance ACS Abstracts, February 1, 1995.
