Preparation of Biodiesel by Lipase-Catalyzed Transesterification of

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Energy & Fuels 2007, 21, 368-372

Preparation of Biodiesel by Lipase-Catalyzed Transesterification of High Free Fatty Acid Containing Oil from Madhuca indica Veena Kumari, Shweta Shah, and Munishwar N. Gupta* Department of Chemistry, Indian Institute of TechnologysDelhi, Hauz Khas, New Delhi 110 016, India ReceiVed May 14, 2006. ReVised Manuscript ReceiVed NoVember 3, 2006

The oil from Madhuca indica has a high free fatty acid content, and hence, it is difficult to convert it into biodiesel by using a chemical catalyst. In this work, a commercial preparation of lipase from Pseudomonas cepacia was used as a catalyst for converting this oil into ethyl esters. Solvent-free media were used for this process. The process optimization consisted of varying parameters, such as water content in the reaction medium and salt present during the drying of lipase prior to its use as powder. The lipase immobilized on accurel gave 96% conversion in 6 h. The best results were obtained using modified biocatalyst formulations, which are called cross-linked enzyme aggregates (CLEAs) and protein-coated microcrystals (PCMCs). While free enzyme powder after process optimization gave 98% conversion in 6 h using 50 mg of lipase, CLEAs gave 92% conversion in 2.5 h (using an equivalent of 50 mg of enzyme) and PCMCs gave 99% conversion in 2.5 h using the same amount of enzyme. In the case of PCMCs, 99% conversion could be obtained just by using a biocatalyst equivalent to 25 mg of enzyme but with an extended time period of 16 h.

Introduction Biodiesel is a nontoxic, biodegradable, and renewable source of energy.1 Environmental benefits of its use (in place of diesel) include lower exhaust emissions of particulate matter and greenhouse gases, such as CO, CO2, and SOx. Biodiesel consists of monoalkyl esters of long-chain fatty acids. It is produced from vegetable oils or fats by either acid/alkali or lipasecatalyzed transesterification with methanol or ethanol.1 In the Indian context, the bulk of the efforts have been directed toward obtaining biodiesel by chemical transesterification of Jatropha curcas oil.2 However, it is felt that alternative starting oils also need to be studied.3 Madhuca indica (Mahua) constitutes one such potential alternative source for biodiesel.4,5 Mahua is a forest tree, which grows in several parts of India. The flowers find use in the production of country liquor. The seed kernels are rich in fat with a rather high melting point. Thus, at 25 °C, it is in the form of a semisolid fat. Hydrogenation of this fat (after refining) is carried out for use in products for human consumption.6 The fatty acid composition of the oil reveals the presence of oleic acid (46.3%) and linoleic acid (17.9%) as the major unsaturated fatty acids and palmitic acid (17.8%) and stearic acid (14.0%) as the major saturated fatty acids. In view of its availability in large amounts, use of its semisolid fat as starting triglycerides for the production of biodiesel has been suggested and explored.4,5 All of these efforts have used a * To whom corresopndence should be addressed: Chemistry Department, Indian Institute of TechnologysDelhi, Hauz Khas, New Delhi 110 016, India. Telephone: 91-11-2659-1503 and/or 91-11-2659-6568. Fax: 91-112658-1073. E-mail: [email protected]. (1) Ma, F.; Hanna, M. A. Bioresour. Technol. 1999, 70, 1-15. (2) Francis, G.; Edinger, R.; Becker, K. Nat. Resour. Forum 2005, 29, 12-24. (3) Srivastava, A.; Prasad, R. Renewable Sustainable Energy ReV. 2000, 4, 111-133. (4) Ghadge, S. V.; Raheman, H. Biomass Bioenergy 2005, 28, 601605. (5) Puhan, S.; Vedaraman, N.; Ram, B. V. B.; Sankarnarayanan, G.; Jeychandran, K. Biomass Bioenergy 2005, 28, 87-93. (6) Rukmini, C. Food Chem. Toxicol. 1990, 28, 601-605.

chemical catalyst-based transesterification approach. Mahua oil contains a high amount of free fatty acids (FFAs) (about 20%), and thus, a pretreatment step is necessary to avoid soap formation.4 In such cases, enzyme-based transesterification offers an important advantage. Several attempts have been made to use lipase for the production of biodiesel.7-9 There are some other advantages of using an enzyme-based route as well:1,10,11 The reaction is carried out at moderate temperature. Thus, the catalyst and process temperature do not degrade the reactor material. Downstream processing of biodiesel and glycerol (as a byproduct) is much easier. Also, unlike chemical catalysis, which works better with methanol,10 enzymes seem to prefer ethanol. In the case of chemical catalysis, the high temperature necessary in the process improves the miscibility between oil and methanol, while in the case of biocatalysis, the reaction is carried out at lower temperatures at which the miscibility of methanol in oil is very poor.12,13 Methanol is also known to cause enzyme inactivation more than ethanol.12,13 Hence, ethanol is generally preferred for carrying out lipase-catalyzed transesterification for the preparation of biodiesel.7,12,13 Ethanol as such is a renewable starting material for plant feedstock. Thus, an enzyme-based route fits better as a part of developing sustainable technology for biofuels. All of this has generated immense interest in using lipase for the production of biodiesel from a variety of oils/fats, such as soybean,14 sunflower,8,15,16 (7) Nelson, L. A.; Foglia, T. A.; Marmer, W. N. J. Am. Oil Chem. Soc. 1996, 73, 1191-1195. (8) Dossat, V.; Combes, D.; Marty, A. Enzyme Microb. Technol. 2002, 30, 90-94. (9) Shah, S.; Sharma, S.; Gupta, M. N. Jatropha oil. Energy Fuels 2004, 18, 154-159. (10) Fukuda, H.; Kondo, A.; Noda, H. J. Biosci. Bioeng. 2001, 5, 405416. (11) Shah, S.; Sharma, S.; Gupta, M. N. Indian J. Biochem. Biophys. 2003, 40, 392-399. (12) Shimada, Y.; Watanabe, Y.; Sugihara, A.; Tominaga, Y. J. Mol. Catal. B: Enzym. 2002, 17, 133-142. (13) Salis, A.; Pinna, M.; Monduzzi, M.; Solinas, V. J. Biotechnol. 2005, 119, 291-299.

10.1021/ef0602168 CCC: $37.00 © 2007 American Chemical Society Published on Web 12/06/2006

Biodiesel of High FFA Containing Oil from Mahua

cottonseed,17 rapeseed,7,18 palm oil,19,20 mango kernel,21 Jatropha oil,9 and beef tallow22 (lipase-catalyzed transesterification of mahua oil has not been attempted thus far). This is the case, despite the current high cost of the biocatalyst. It is hoped that efficient downstream processing techniques would make enzyme production cost much cheaper.23,24 Also, if the enzyme-based transesterification is adopted at a large scale, a large demand would induce large-scale production of the enzyme and would result in lowering the market price of lipases. The existing usage of enzymes in several areas, such as detergents,25 dairy products,26 and textile and leather processing27 reflect the soundness of such a strategic approach. The present work shows that the preparation of ethyl esters of oils present in mahua could be carried out in solvent-free media (i.e., substrate alone constituted media)8 by using lipase from Pseudomonas cepacia. It is also shown that the adaptation of current knowledge about biocatalyst design for efficient catalysis in low water media (as such media are sometime called28,29) leads to an increase in the efficiency of conversion. Experimental Section Materials. Mahua oil was purchased from local market. Lipases from P. cepacia (PS), Pseudomonas fluorescens (AK), and Mucor jaVanicus (M) were kind gifts from Amano Enzyme Inc., Nagoya, Japan. Rhizomucor miehei lipase immobilized on anion-exchange resin (Lipozyme RM IM) was a kind gift from Dr. J. S. Rao, Novozymes, South Asia Pvt. Ltd., Bangalore, India. Accurel MP 100 was a kind gift from Membrane Gmbh, Obernburg, Germany. The absolute ethanol [purity by gas chromatography (GC) > 99.9%] (Merck, Germany) used had a water content of 0.1% (v/v). All of the solvents used in this work were of low water grade and were further dried by gentle shaking with 3 Å molecular sieves (E. Merck, Mumbai, India). Methods: Preparation of pH-Tuned Enzymes. Lipases (50 mg) from P. cepacia, P. fluorescens, and M. jaVanicus were dissolved in 0.5 mL of 0.02 M phosphate buffer at pH 7.0, 8.0, and 7.5, respectively (these are the pH optima of the lipases as reported by the vendors and other workers).30-32 All enzyme solutions were immediately frozen at -20 °C and lyophilized for 24 h. These were referred to as “pH-tuned” enzyme preparations.9,33 (14) Schwab, A. W.; Dykstra, G. J.; Selke, E.; Sorenson, S. C.; Pryde, E. H. J. Am. Oil Chem. Soc. 1988, 65, 1781-1786. (15) Mittelbach, M. J. Am. Oil Chem. Soc.1990, 67, 168-170. (16) Antolin, G.; Tinaut, F. V.; Briceno, Y.; Castano, V.; Perez, C.; Ramirez, A. I. Bioresour. Technol. 2002, 83, 111-114. (17) O ¨ znur, K.; Tu¨ter, M.; Aksoy, H. A. Bioresour. Technol. 2002, 83, 125-129. (18) De, B. K.; Bhattacharya, P. K.; Bandhu, C. J. Am. Oil Chem. Soc. 1999, 76, 451-453. (19) Abigor, R. P.; Vadia, P.; Foglia, T.; Hass, M.; Jones, K.; Okefa, E.; Obibuzor, J.; Bator, M. Biochem. Soc. Trans. 2000, 28, 979-981. (20) Crabbe, E.; Nolasco-Hipolito, C.; Kobayashi, G.; Sonomoto, K.; Ishizaki, A. Proc. Biochem. 2001, 37, 65-71. (21) Linko, Y. Y.; La¨msa¨, M.; Wu, X.; Vosukainen, W.; Sappa¨la¨, J.; Linko, P.; J. Biotechnol. 1998, 66, 41-50. (22) Hsu, A-F.; Jones, K.; Marmer, W. N.; Foglia, T. A. J. Am. Oil Chem. Soc. 2001, 78, 585-588. (23) Przybycien, T. P.; Pujar, N. S.; Steele, L. M. Curr. Opin. Biotechnol. 2004, 15, 469-478. (24) Gupta, M. In Methods in Affinity-Based Separations of Enzymes and Proteins; Gupta, M. N., Ed.; Brikhauser Verlag: Basel, Switzerland, 2002; pp 1-15. (25) Arbige, M. V.; Pitcher, W. H. Trends Biotechnol. 1989, 7, 330335. (26) John, V. T.; Abraham, G. In Biocatalysts for Industry; Dordick, J. S., Ed.; Plenum Press: New York, 1990; Vol. 10, pp 193-197. (27) Houde, A.; Kademi, A.; Leblanc, D. Appl. Biochem. Biotechnol. 2004, 118, 155-170. (28) Gupta, M. N.; Roy I. Eur. J. Biochem. 2004, 271, 2575-2583. (29) Hudson, E. P.; Kppler, R. K.; Clark, D. S. Curr. Opin. Biotechnol. 2005, 16, 637-643. (30) Ishihara, H.; Okuyama, H.; Ikezawa, H.; Tejima, S. Biochem. Biophys. Acta 1975, 388, 413-422.

Energy & Fuels, Vol. 21, No. 1, 2007 369 Enzyme Immobilization. The polypropylene support, Accurel MP 100 (50 mg), was placed in 5 mL capped vials and wetted with 150 µL of 95% ethanol followed by the addition of 4 mL of P. cepacia lipase solutions in 20 mM potassium phosphate buffer at pH 6.0. The vials were incubated at 25 °C with constant shaking at 100 rpm for overnight. The solutions were then withdrawn from each vial and stored, while the MP100 particles were washed twice with 1 mL of phosphate buffer.34 The lipase activity and protein were determined in immobilization solution and washings. The immobilized lipase preparations were dried by a speed vacuum system (UVS4004 Universal Vacuum system, Thermo Savant). P. cepacia Lipase Cross-Linked Enzyme Aggregates (CLEAs). P. cepacia lipase preparation (50 mg) was dissolved in 0.05 M phosphate buffer at pH 7.0 (1 mL), followed by centrifugation for 10 min at 8000g. The supernatant was collected and, to this, was added bovine serum albumin. This aqueous mixture was then added dropwise to a shaking centrifuge tube (150 rpm) containing 4 mL of the acetone. After 30 min, glutaraldehyde (25% v/v in water) was added so that the final concentration of glutaraldehyde was 25 mM. The tube was shaken continuously during this addition. The mixture was kept at 4 °C for 4 h with constant shaking at 300 rpm. The same organic solvent (1 mL) was added to this mixture, which was then centrifuged at 8000g. The supernatant was decanted, and the residue was washed 3 times with the same organic solvent (5 mL), centrifuged, and decanted. The final enzyme preparation was kept in the acetone at 4 °C. Prior to use, CLEAs kept in the acetone were centrifuged and the supernatant was decanted.35 Preparation of Protein-Coated Microcrystals (PCMCs) of P. cepacia Lipase. The P. cepacia lipase preparation (50 mg) was dissolved in 0.05 M phosphate buffer at pH 7.0 (1 mL), followed by centrifugation for 10 min at 8000g. The supernatant was collected (which now contains clear lipase solution) and was further concentrated by a speed vacuum system to 0.2 mL. A saturated solution of potassium sulfate (300 µL) was added to the varying amounts of concentrated lipase solution. This combined solution was then added dropwise to a shaking vial (150 rpm) containing 6 mL of acetone. The precipitate obtained was then centrifuged at 2200g for 5 min to remove acetone. The precipitate was further washed three times with 2 mL of acetone.36 Enzymatic Transesterification Reaction. Mahua oil (0.5 g) and ethanol were taken at a ratio of 1:4 in different screw-capped vials. Different pH-tuned lipase preparations (50 mg) were added to these different vials, and the mixtures were incubated at 40 °C with a constant shaking at 200 rpm.9 The progress of the reaction was monitored by removing aliquots (40 µL) at various time intervals. The ethyl esters formed were analyzed by GC. GC Analysis. The ethyl esters were analyzed on a Nucon 5700 gas chromatograph with a flame ionization detector. The capillary column (length, 30 m; internal diameter, 0.25 mm) was packed with 70% phenyl polysilphenylenesiloxane. Nitrogen was used as a carrier gas at a constant pressure of 4 kg cm-2. The column oven temperature was programmed from 150 to 250 °C min-1 (at the rate of 10 °C min-1) with injector and detector temperatures at 240 and 250 °C, respectively.9 The percent yield was calculated in terms of the weight of alkyl esters obtained per unit weight of oil. A standard curve was plotted for the weight of alkyl esters verses the GC peak area. For plotting the standard curve, alkyl esters were extracted with a mixture containing 4 volumes of hexane/ether (1:1, v/v) and 2 volumes of (31) Kojima, Y.; Yokoe, M.; Mase, T. Biosci., Biotechnol., Biochem. 1994, 58, 1564-1568. (32) Takeda, Y.; Aono, R.; Doukyu, N. Extremophiles 2006, 10, 14334909. (33) Mattiasson, B.; Adlercreutz, P. Trends Biotechnol. 1991, 9, 394398. (34) Salis, A.; Sanjust, E.; Solinas, V.; Monduzzi, M. Biocatal. Biotransform. 2005, 23, 381-386. (35) Shah, S.; Sharma, A.; Gupta, M. N. Anal. Biochem. 2006, 18, 154159. (36) Kreiner, M.; Moore, B. D.; Parker, M. C. Chem. Commun. 2001, 1096-1097.

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Figure 1. Screening of lipases for biodiesel production from Mahua oil. Mahua oil (0.5 g) and ethanol were taken at a ratio of 1:4. pHtuned lipase preparations were added and incubated at 40 °C with constant shaking at 200 rpm. PS, P. cepcia lipase; IM, immobilized R. miehei lipase; AK, P. fluorescens lipase; M, M. jaVanicus lipase.

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Figure 2. Effect of various solvents on biodiesel production from Mahua oil using P. cepacia lipase. Mahua oil (0.5 g) and ethanol were taken at a ratio of 1:4. pH-tuned P. cepacia lipase was added (50 mg) followed by the addition of different solvents (2 mL) viz., hexane, octane, and acetonitrile. A control was also run where no solvent was added. These solutions were incubated at 40 °C with a constant shaking at 200 rpm for 24 h.

saturated NaCl as described by Abigor et al.19 The compositions of products were identified by comparing retention times with standards of a known composition. Protein Estimation. The protein concentration was determined according to the procedure described by Bradford,37 using bovine serum albumin as the standard protein. Assay of Lipase Activity. The hydrolytic activity of lipase was monitored by following the rate of hydrolysis of p-nitrophenylpalmitate spectrophotometrically at 410 nm.38

Results and Discussion Screening of Lipases for Biodiesel Preparation. Figure 1 shows the results of the screening of four different commercial preparations of lipases for their transesterification efficiency with mahua oil. Ethanol was used for the transesterification of mahua oil in solvent-free media. M. jaVanicus lipase was nearly inactive for catalyzing transesterification. Immobilized R. miehei lipase and free P. cepacia lipase were found to give appreciable yields at 80% (w/w), followed by P. fluorescens lipase at 62% (w/w). The conversion time was 24 h, and the reaction temperature was 40 °C. Further work was done with P. cepacia lipase. Effect of Solvents on Transesterification. There are several advantages associated with the solvent-free approach, with an important one being the elimination of the step of solvent evaporation.8 Use of anhydrous organic solvent media, however, is occasionally reported to give higher transesterification as compared to solvent-free media.15 The reaction was carried out by using octane, hexane, and acetoniltrile (as somewhat randomly chosen solvents of different polarity) as the reaction media. Only octane gave an 84% higher yield as compared to solvent-free media (Figure 2). In view of our earlier experience of the possibility of improving the percent conversion by process optimization in a solvent-free media approach,9 above-mentioned advantages associated with a solvent-free approach, and only marginal improvement in conversion by using octane, further work was carried out with solvent-free media. Effect of Buffer Molarity during Lyophilization on Transesterification. It has been shown that the amount of buffer salt present during lyophilization (while doing pH tuning of the (37) Bradford, M. M. Anal. Biochem. 1976, 72, 248-254. (38) Kilcantey, K. N.; Wilkinson, M. G.; Fox, P. F. Enzyme Microb. Technol. 2002, 31, 310-320.

Figure 3. Effect of varying the concentration of buffer on the transesterification reaction. P. cepacia lipase (50 mg) was lyophilized in 0.5 mL of Tris-HCl buffer of varying concentrations (0.5, 0.8, 1.0, and 1.2 M) at pH 7.0. The lyophilized enzyme powders were added to the reaction media containing Mahua oil and ethanol at a ratio of 1:4. These mixtures were incubated at 40 °C with a constant shaking at 200 rpm for 24 h. The experiments were performed in triplicate, and the error bars represent the percentage error in each set of readings.

enzyme) also affects the reaction rates.39,40 The mechanism of salt activation has been studied earlier.39,40 It is believed that the enhanced activity with an increasing buffer content results from a better dispersion of the enzyme in the reaction medium. This prevents intermolecular interactions, which are likely to result in mass-transfer limitations and changes in the enzyme conformation. Figure 3 shows the effect of varying concentrations of Tris-HCl buffer on the transesterification of Mahua oil catalyzed by P. cepacia lipase. The maximum yield [99% (w/ w) in 24 h] was obtained when the concentration of Tris-HCl buffer during lyophilization was 0.8 M. Effect of Water on Transesterification. Amount of water present in the media is another critical parameter, which is known to influence biotransformations in nonaqueous media.28,41,42 Figure 4 shows the effect of different amounts of water (39) Triantafyllou, A. O.; Wehhtje, E.; Adlercreutz, P.; Mattiasson, B. Biotechnol. Bioeng. 1997, 54, 67-76. (40) Ru, M. T.; Hirokane, S. Y.; Lo, A. S.; Dordick, J. S.; Reimer, J. A.; Clark, D. S. J. Am. Chem. Soc. 2000, 122, 1565-1571.

Biodiesel of High FFA Containing Oil from Mahua

Figure 4. Effect of water on biodiesel preparation using P. cepacia lipase. P. cepacia lipase (50 mg) was lyophilized in 0.5 mL of 0.8 M Tris-HCl buffer at pH 7.0. The lyophilized enzyme powders were added to the reaction media containing Mahua oil and ethanol at a ratio of 1:4. Different amounts of water were added to the reaction media viz., 2, 4, 6, 8, and 10% (w/w, enzyme). These mixtures were incubated at 40 °C with a constant shaking at 200 rpm for 6 h. The experiments were performed in triplicate, and the error bars represent the percentage error in each set of readings.

present in the reaction mixture for biodiesel preparation. The highest conversion of biodiesel (98% in 6 h) was obtained at a 6% (w/w) water level. The trend is in agreement with the general experience in carrying out enzymatic reactions in low water media.28,42 Initially, as water is added, it adds onto polar amino acids present on the enzyme surface. This breaks hydrogen bonds among these and imparts conformational flexibility to the enzyme molecule, which is required for catalysis. After an optimum water level, the hydrolytic reaction become significant and competes with the desired transesterification reaction. Hence, the reaction yield of the alkyl ester diminishes. Biodiesel Preparation from Lipase Immobilized on Accurel. Immobilization of the enzyme has often been found to improve its performance in nonaqueous media (as well).28,43 Accurel has been one of the most frequently used commercially available matrixes for lipase immobilization.43,44 About 50 mg of lipase (containing 5 µg of protein/mg of lipase powder) could be immobilized on 50 mg of support with an adsorption efficiency of 99%. About a 96% conversion level could be achieved in 6 h at 40 °C. This was comparable to the conversion obtained with free enzyme. Another approach was to explore whether less enzyme could be used to obtain an adequate conversion level. Figure 5 shows that 50 mg of enzyme was necessary to obtain a 96% conversion yield in 6 h. Biodiesel Preparation from High-Activity Biocatalyst Preparations. The last few years have witnessed the design of a few high-activity biocatalyst preparations for use in nonaqueous enzymology.28 CLEAs35,45 and PCMCs36 represents two such exciting possible enzyme formats. Superiority of such biocatalyst designs has been shown with model systems and (to a much limited extent) in the synthesis of fine chemicals. In both cases, lyophilization is avoided. It is now well(41) Triantafyllou, A. O.; Wehtje, E.; Adlercreutz, P.; Mattiasson, B. Biotechnol. Bioeng. 1995, 45, 406-414. (42) Anthonsen, T.; Sjursens, B. J. In Methods in Nonaqueous Enzymology; Gupta, M. N., Ed.; Birkhauser Verlag: Basel, Switzerland, 2000; pp 14-35. (43) Al-Duri, B.; Yong, Y. P. J. Mol. Catal. B: Enzym. 1997, 3, 177188. (44) Bosley, L.; Peilow, A. D. J. Am. Oil Chem. Soc. 1997, 74, 107111. (45) Lo´pez, P.; Cao, L.; van Rantwijk, F.; Sheldon, R. A. Biotechnol. Lett. 2002, 24, 1379-1383.

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Figure 5. Effect of immobilization of lipase on accurel for biodiesel preparation from Mahua oil. The immobilized lipase preparations (50 mg of lipase on 50 mg of accurel and 25 mg of lipase on 50 mg of accurel) were respectively added to the reaction media containing Mahua oil and ethanol at a ratio of 1:4. These mixtures were incubated at 40 °C with a constant shaking at 200 rpm for 24 h. Aliquots were taken out at 3, 6, 12, and 24 h and were analyzed by GC.

Figure 6. Biodiesel production from Mahua oil catalyzed from CLEAs and PCMCs of P. cepacia lipase. CLEAs and PCMCs prepared from different amount of lipases viz., 50, 25, 12.5, and 6.25 mg were respectively added to the reaction media containing Mahua oil and ethanol at a ratio of 1:4. These mixtures were incubated for 2.5 h at 40 °C with a constant shaking at 200 rpm. The experiments were performed in triplicate, and the error bars represent the percentage error in each set of readings.

established that lyophilization causes structural changes that lower the catalytic efficiency of the enzymes in low water media.46,47 CLEAs are matrix-free immobilized preparations. CLEAs are formed in two steps. The first step of the precipitation leads to the formation of physical aggregates. These physical aggregates are held together by noncovalent bonding and redissolve when dispersed in water. Cross-linking of these physical aggregates produce CLEAs, which are structurally more robust. Even in water, CLEAs withstand the denaturing condi(46) Lee, Y.-Y.; Dordick, J. S. Curr. Opin. Biotechnol. 2002, 13, 376384. (47) Roy, I.; Gupta, M. N. Biotechnol. Appl. Biochem. 2004, 39, 165177.

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tions much better than corresponding free enzymes.35,45,48 Hence, CLEAs per se have a greater stability than free enzymes.35,45,48 In the case of PCMCs, protein molecules are present as a uniform layer on the surface of the crystal.36 The substrate is easily accessible to the enzyme. Hence, PCMCs as heterogeneous catalysts are believed to have less mass-transfer limitations as compared to lyophilized powders. Also, even in CLEAs, the active site would not be on the biocatalyst surface, and this design is expected to have somewhat of a higher mass-transfer limitation as compared to PCMCs. Applications of these relatively new biocatalyst designs for the production of biodiesel have not been explored thus far. Figure 6 shows that the use of CLEAs and PCMCs prepared from P. cepacia lipase did indeed accelerate the rate of transesterification. About 92 and 99% conversions could be obtained with 50 mg of enzyme in 2.5 h only by using 50 mg of enzyme. Exploratory experiments (48) Schoevaart, R.; Wolbers, M. W.; Golubovic, M.; Ottens, M.; Kieboom, A. P. G.; van Rantwijk, F.; van der Wielen, L. A. M.; Sheldon, R. A. Biotechnol. Bioeng. 2004, 87, 754-762.

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suggest that, by using a larger reaction time of 16 h, it is possible to obtain 96% conversion using PCMCs prepared from 25 mg of P. cepacia lipase. It seems that upon further optimization with PCMCs, a more economical and efficient process design would be possible. These further studies are in progress. The current work shows that, by the judicious choice of biocatalyst design and process optimization, the lipase-based route can be successfully used to obtain biodiesel from oils, such as mahua (with high FFA content), which, at present, are difficult to transesterify by chemical routes. Acknowledgment. The funds provided by the Council of Scientific and Industrial Research (Extramural and Technology Mission on Oilseeds, Pulses, and Maize), Department of Science and Technology (DST), and Department of Biotechnology (DBT), all of which are Government of India Organizations, are gratefully acknowledged. EF0602168