Lipase Catalyzed-transesterification of Vegetable Oils by Lipolytic Bacteria
Lipase-catalyzed transesterification of vegetable oils is an important reaction that produces fatty acid alkyl esters which are valuable intermediates in oleo chemistry and are excellent substitutes to diesel fuel. In present work lipase producing bacteria were isolated from oil spilled soil samples collected from different areas of Raipur, India by serial dilution method. Lipase activity of extracellular lipase was determined by titrimetric method. The sodium alginate entrapment was carried out to immobilize lipase according to the standard method. Out of 15 bacterial isolates (LPB1-LPB15), LPB1 exhibited the maximum extracellular lipase activity on lipase assay medium. Thus, it was selected for further study. Olive oil was found to be the best substrate for lipase production (0.0070 μg/mL /min) among the substrates tested. This isolate exhibited further increase in activity with value of 0.0099 μg/mL/min using olive oil as substrate in production medium supplemented with lecithin as emulsifier at pH 7.2 after 3 days of incubation at 30°C (160 rpm). The transesterification capability of the crude extracellular lipase from LPB1 was assessed using thin layer chromatography by using hexane/diethyl ether/acetic acid as solvent system in the ratio of 90:10:1 (v/v/v). The free extracellular lipase exhibited the formation of methyl esters with the vegetable oils tested such as karanja (Rf 0.59), neem (Rf 0.59), castor (Rf 0.6) and olive oil (Rf 0.62). Both the soluble and immobilized lipase of this isolate demonstrated the methanolysis of non edible oil of Karanja (Pongamia pinnata) within 1-3 h.
April 09, 2010; Accepted: October 13, 2010;
Published: November 04, 2010
Lipases (Triacylglycerol acylhydrolases EC 22.214.171.124) are a class of hydrolases
which catalyze the hydrolysis of triglycerides to glycerol and free fatty acids
on an oil-water interface (Kamini et al., 2000).
In addition they are carboxylesterases that catalyze the hydrolysis and transesterification
of esters. The syntheses of esters can exhibit enantioselective properties (Varma
et al., 2007; Bezbradica et al., 2007;
Abbas et al., 2002). Apart from its prevalent
use in biosurfactants, aroma and flavor compounds, lubricant and in polyesters,
it has also find use in biodiesel preparation (Edmundo et
al., 1998; Athawale et al., 2003; Hills,
2003; Kumar and Gross, 2000; Jaeger
and Eggert, 2002). Lipase has been found in many species like Gryllus
campestris (Ozlem et al., 2007) and Cocos
nucifera Linn. (Ejedegba et al., 2007).
Lipolytic bacteria can be isolated from soil, raw milk and waste water
(Mohan et al., 2008; Abdou
et al., 2003; Bhumibhamon et al., 2002).
Several bacterial species producing extracellular lipases have been reported
(Bora and Kalita, 2009; Amoozegar
et al., 2008; Zeng et al., 2004).
Effective transesterification reactions using s everal lipases from P. aeruginosa
MTCC 5113, E. aerogenes, P. fluroscens MTCC 103, have been also reported
by several researchers (Singh et al., 2008;
Kumari et al., 2009; Devanesan
et al., 2007).
Transesterification of vegetable oils received considerable attention in past
few years, which produces fatty acid and alkyl esters that are valuable intermediates
in oleo chemistry and ethyl esters which are excellent substitute for diesel
fuels (Freedman et al., 1986; Schuchardt
et al., 1998). Enzymes perform very specific transesterification
reaction (bio-transformations) (Stamatis et al.,
2000); have made them increasingly popular in industries where less specific
chemical processes produce unwanted by-products. The disadvantage associated
with enzymatic transesterification is cost of enzyme preparation. Immobilization
generally increases the reusability of the enzyme (Roy et
al., 2003) and hence lowers the cost as well as helps the biocatalyst
to become efficient in nonaquous media (Shah et al.,
Furthermore, more than hundred types of tropical plants that produce oil-bearing
materials have been identified. Among them are rubber seed oil, fenugreek oil,
Neem oil, karanja oil, castor oil, olive oil and jatropha oil. They are good
sources of oils with special fatty acid compositions and therefore have to be
exploited by much research as excellent substrate for transesterification reaction
(Shah et al., 2004; Kumari
et al., 2009).
The present study reports the isolation of lipase producing bacteria from soil and to assess both the free and immobilized lipase preparations for transesterificaion of vegetable oils in a solvent-free system to produce methyl esters.
MATERIALS AND METHODS
Sample collection: Soil samples were aseptically collected from Raipur city with the help of soil auger in sterile sealed plastic bags for isolation of lipase producing bacteria under laboratory conditions in the year 2009. The soil samples from oil spilled areas included Kitchen garden waste, dairy farm compost (Sanchi dairy industry), compost yard (Kedia distillery limited) and Jatropha oil mill waste.
Isolation of lipolytic bacteria: The lipolytic bacteria were isolated
from collected soil samples by serial dilution method. For this 1.0 g of soil
sample was dissolved in 100 mL of sterile distilled water. This was serially
diluted (10-1 to 10-5) and diluted samples were plated
on solid agar medium containing olive oil in enrichment media (0.5% w/v (NH4)2
SO4, 0.05% w/v K2HPO4, 0.03% w/v MgSO4.7H2O,
2% v/v olive oil and 1.5% w/v agar, pH 7.0). Colony capable for utilizing olive
oil as sole source of carbon was isolated and individually streaked on modified
lipase assay media containing 1.5% w/v peptone, 0.5% w/v NaCl, 0.1% w/v CaCl2,
1% v/v Tween 80, 1.5% w/v agar at pH 7.2 as described by Shukla
and Gupta (2007). The plates were incubated at 30°C for 72 h. The formation
of white precipitate of calcium monostearate around the colony was considered
as positive colony for extracellular lipase secretion and used for further study.
Gram staining was carried out as given by Brucker (1986).
The stock cultures were maintained routinely on nutrient agar media.
Lipase production: The pure culture of lipolytic bacteria was maintained
on nutrient agar medium (0.5% w/v peptone, 5% w/v beef extract, 0.15% w/v yeast
extract, 0.5% w/v NaCl and 1.5% w/v agar pH 7.5). The extra cellular lipase
production was carried out in a medium composed of 3% w/v yeast extract, 1%
w/v KH2PO4, 0.1% w/v MgSO4.7H2O,
0.5% w/v maltose and 0.2% v/v olive oil at pH 7.2. Medium was sterilized and
inoculated with 1% seed inoculum prepared in nutrient broth followed by incubation
at 37°C for 48 h at 160 rpm in a shaker incubator. By using different substrate
sources such as olive oil, olive oil and lecithin (emulsifier), tween 80 and
castor oil, their effect on lipase production by selected bacterial isolate
was assessed at pH of 7.2. The cells were then centrifuged at 10,000 rpm for
15 min and the supernatant was used directly as crude preparation of lipase
for further studies.
Lipase assay: The lipase activity in the culture supernatant was determined
by titrimetric method (Sadasivam and Manickam, 1991).
Lipase activity (E.C. 126.96.36.199) was measured by titration the fatty acid released
with 0.1 M NaOH using 0.1% alcoholic phenolphthalein as indicator.
One unit of lipase activity was defined as the amount of enzyme releasing one mole of free fatty acid in 1 min under standard assay condition.
Immobilization of extracellular lipase by entrapment: The sodium alginate
entrapment of crude lipase was carried out according to the standard method
(Bhushan et al., 2008). Alginate with a concentration
range of 4-10 % and LB1 lipase preparation were mixed together and stirred for
15 min at 4°C to obtain a homogenous mixture. Alginate-lipase homogenous
mixture was extruded drop by drop into a cold CaCl2 solution (0.4
M) through pipette. The beads were stirred in CaCl2 solution for
45 min and then thoroughly washed with buffer (Tris acetate 0.107 M, pH 7.2).
Alginate immobilized lipase beads were stored at 4°C in Tris acetate buffer
till further use.
Set up for transesterification reaction: Transesterification reaction was carried out for different oil samples such as Olive, Neem, Karanja and Castor oil separately with a short chain alcohol, i.e., methanol. Oil and methanol were taken in the ratio of 1:4 (mol/mol) in a screw capped vial. To this oil: methanol mixture 2 mL of crude lipase was added and incubated at 40°C with constant stirring at 200 rpm for 3 h in a shaker incubator. Process of methanolysis was performed with both free and immobilized lipase preparation.
Analysis of esters: The formation of methyl esters of vegetable oil
in the reaction mixture was analyzed by thin-layer chromatography on silica
gel H (E. Merck, Mumbai, India) plates. The coated silica gel plates were spotted
with transesterified oil samples and bio-diesel. The chromatogram was developed
in chromatographic chamber using hexane/diethyl ether/acetic acid as solvent
system in the ratio of 90:10:1 (v/v/v). The spots were detected in the iodine
chamber and Rf values were calculated and compared with authentic standards
(Gordon et al., 1994).
RESULT AND DISCUSSION
Isolation and screening of lipolytic bacteria: The lipase producing
bacteria were isolated from different soil samples. Enrichment culture technique
and lipase assay media enable the isolation of extracellular lipase producing
bacteria. In total, 15 isolates were isolated by enrichment culture technique
from the soil samples and among them; three isolates (LPB1, LPB4 and LPB7) showed
measurable lipolytic activity as shown in Fig. 1a-d.
The precipitation zone for strain LPB1 (Fig. 1) was found
to be appreciable among the three isolates and hence was further explored for
lipase production and transesterification process. The LPB1 was identified as
gram negative and rod shaped on gram staining.
||Photo images of zone of precipitation generated by isolates
of lipase producing bacteria on Tween 80 agar medium. (a) LPB12, negative
isolate from compost yard the positive isolates (b) LPB7, (c) LPB4 and (d)
LPB1 from kitchen garden soil, Jatropha oil mill waste and dairy farm compost,
Lipase production: The lipase production efficiency of lipolytic bacteria
was assayed in presence of different substrates i.e., Olive oil, Olive oil and
Lecithin (emulsifier), Castor oil and Tween 80. The assay was also performed
at various pH ranges i.e., from 5.6 to 8.0. Optimization of extracellular lipase
production was carried out in bacterial (Babu and Rao, 2007)
and fungal (Iftikhar and Hussain, 2002) lipase. It was
observed that strain LPB1 produces lipase at all pH range tested. With an increase
in the pH from 5.6 to 7.2 the lipase production was increased in isolate LPB1
from 0.006 to 0.0077 (μg/mL/min). Beyond pH 7.2 a fall in lipase production
was observed (Fig. 2). Out of the six different values of
pH of production medium the pH optima for maximum production of lipase in strain
LPB1 was found to be 7.2 (0.0099 μg/mL/min). The production of lipase decreased
in slightly acidic (pH 5.6) or alkaline (pH 8.0) whereas, at pH 7.2 the highest
production was reported. The production of extracellular lipase can be induced
by using the lipidic substrate (Akhtar et al., 1980).
Hence, the lipase production was found measurable for all tested lipidic substrate
(Fig. 3). Olive oil was found to be the best substrate for
lipase production among the substrates tested. The lipase activity was found
comparable in presence of Olive oil and Tween 80 (0.0070 μg/mL/min and
0.0077 μg/mL/min, respectively). An increase in lipase activity was observed
from 0.007 to 0.009 (μg/mL/min) in presence of lecithin as emulsifier in
production media containing Olive oil as substrate at pH 7.2. The incorporation
of surface active substances can increase the availability of lipidic substrate
by emulsifying the lipid and could further increase the extracellular lipase
activity (Naka et al., 1986). The value reported
by us are almost similar to that obtained for lipase production from Bacillus
sp. (Mohan et al., 2008; Achamma
et al., 2003). Achamma et al. (2003)
has also reported maximum production of lipase when olive oil has been used
||Effect of pH on lipase production by isolate LPB1 in production
media. The result is a mean of four observations
||Effect of various substrates on lipase production by isolate
LPB1 at pH 7.2. The result is a mean of four repeats
Similar result indicating olive oil to be the best substrate of lipase production
in various strains of fungi has been reported by Annibale
et al. (2006).
Transesterification: Formation of methyl ester of various oil samples
(Karanja, Castor, Neem and Olive) has been monitored by thin layer chromatography
on silica gel plate employing hexane/diethyl ether/acetic acid as solvent system.
The spots were detected in the iodine chamber and identified by comparison with
the Rf values of authentic standards form literature for methyl oleate (Gordon
et al., 1994). Figure 4a shows the methyl ester
spots obtain and developed by TLC of oil samples transesterified by lipase obtained
from LPB1. Transesterification of different oil samples such as karanja (Rf
0.59), neem (Rf 0.59), castor (Rf 0.6) and olive oil (Rf 0.62), respectively
could be confirmed since the Rf values of samples calculated is comparable with
the Rf value of standard methyl oleate ester (Rf 0.63).
||(a) Standard methyl oleate and methyl esters of oil samples
tested when observed from left to right in the silica gel TLC plates after
transesterification reaction (Solvent system: hexane/diethyl ether/acetic
acid, 90:10:1 (v/v/v)). TLC of methanolyzed samples using crude LPB1 lipase.
Lanes1, Standard methyl esters (Rf 0.63), lanes 2-5 reaction mixture of
Karanja oil (Rf 0.59), Neem oil (Rf 0.59), Castor Oil (Rf 0.6), Olive oil
(0.62) (Left to right), (b) TLC analysis of reaction mixture after methanolysis
of karanja oil using immobilized LPB1 lipase. Lanes1, Standard methyl esters
(Rf 0.63), Lanes 2-4 reaction mixture of Karanja oil after 1-3 hr (Rf 0.6,
Hence we report the methonlysis of oil samples catalyzed by LPB1 crude lipase
isolated from dairy compost (Sanchi, Raipur).
Karanja oil has been chosen for the time dependent tranesterification studies
since it is cheap, non edible and present abundantly in India. Hence, it can
serve as a good source of raw material for bio fuel generation. Since, immobilization
impart stability to enzyme system and also increases the shelf life and reusability
of enzyme. Hence an attempt has been made to check out the efficiency of LPB1
lipase in immobilized condition for transesterification reaction. Figure
4b shows the transesterification of Karanja oil at various interval of time
(1-3 h) by alginate immobilized LPB1 crude lipase. Transesterification started
within first hours of incubation of Karanja oil with immobilized lipase. Transesterification
has been worked out in aqueous and non aqueous system using lipase for bio-diesel
production by many workers (Singh et al., 2008).
In order to utilize lipases in non aqueous environment the enzyme has frequently
been immobilized on various support matrix (Jagannathan,
et al., 2008; Shieh et al., 2003; Hus
et al., 2002). Alginate has been used as immobilization matrix for
lipase immobilization by many workers (Bhushan et al.,
2008; Devanesan et al., 2007).
From the present study it has been demonstrated that the LPB1 strain shows the efficiency of methanolysis of non edible oil from Karanja. The crude lipase of LPB1 both in soluble form as well as alginate immobilized form exhibited the transesterification of this oil within 3 h of incubation of reaction mixture. In comparison with the highly purified commercial lipase, the inexpensive crude lipase preparation obtained from LPB1 had a distinct rate of tranesterification and at the same time is comparatively cheaper. However,,the further research on various parameters affecting the methyl ester yields namely temperature, pH, reaction time and number of bead and molecular ratio of oil to methanol is to be studied.
Abbas, H., A. Hiol, V. Deyris and L. Comeau, 2002. Isolation and characterization of an extracellular lipase from Mucor sp. strain isolated from palm fruit. Enzyme Microbial Technol., 31: 968-975.
Abdou, A.M., 2003. Purification and partial characterization of psychrotrophic Serratia marcescens lipase. J. Dairy Sci., 86: 127-132.
CrossRef | Direct Link |
Achamma, T.M., K. Manoj, A.Valsa, S. Mohan and R. Manjula, 2003. Optimization of growth condition for the production of extra cellular lipase by Bacillus mycoides. Indian J. Microbiol., 43: 67-69.
Akhtar, M.W., A.Q. Miraz and M.D.I. Chughtai, 1980. Lipase induction in Mucor hiemalis. Applied Environ. Microbiol., 18: 257-263.
Direct Link |
Amoozegar, M.A., E. Salehghamari, K. Khajeh, M. Kabiri and S. Naddaf, 2008. Production of extracellular thermohalophilic lipase from a moderately halophilic bacterium, Salinivibrio sp. Strain SA-2. J. Basic Microbiol., 48: 160-167.
Athawale, V., N. Manjrekar and M. Athawale, 2003. Effect of reaction parameter on synthesis of citronellyl methacrylate by lipase-catalyzed transesterification. Biotechnol. Prog., 19: 298-302.
Babu, I.S. and G.H. Rao, 2007. Optimization of process parameters for production of lipase in submerged fermentation by Yarrowia lipolytica NCIM 3589. Res. J. Microbiol., 2: 88-93.
Bezbradica, D., D. Mijin, S. Siler-Marinkovic and Z. Knezevic, 2007. The effect of substrate polarity on the lipase-catalyzed synthesis of aroma esters in solvent-free systems. J. Mol. Catalysis B. Enzyme, 45: 97-101.
Bhumibhamon, O., K. Achara and F. Suptawee, 2002. Biotreatment of high fat and oil wastewater by lipase producing microorganisms. Kasetsart J. Nat. Sci., 36: 261-267.
Direct Link |
Bhushan, I., R. Parshad, G.N. Qazi and V.K. Gupta, 2008. Immobilzation of lipase by entrapment in Ca-alginate beads. J. Bioactive Compatible Polymers, 23: 552-562.
Bora, L. and M. Kalita, 2009. Production of extracellular lipase from Bacillus sp. LBN4 by solid state fermentation. Internet J. Bioeng., Vol. 3, No. 2.
Brucker, M.C., 1986. Gram staining a useful laboratory technique. J. Nurse-Midwifery, 31: 156-158.
D'Annibale, A., G.G. Sermanni, F. Federici and M. Petruccioli, 2006. Olive-mill wastewaters: A promising substrate for microbial lipase production. Bioresour. Technol., 97: 1828-1833.
Devanesan, N.G., T. Viruthagiri and N. Sugumar, 2007. Transesterification of jatropha oil using immobilized Pseudomonas fluorescens. Afr. J. Biotechnol., 6: 2497-2501.
Direct Link |
Edmundo, C., D. Valerie, C. Didier and M. Alain, 1998. Efficient lipase-catalyzed production of tailor-made emulsifier using solvent engineering coupled to extractive processing. J. Am. Chem. Soc., 75: 309-313.
Ejedegba, B.O., E.C. Onyeneke and P.O. Oviasogie, 2007. Characteristics of lipase isolated from coconut (Cocos nucifera Linn) seed under different nutrient treatments. Afr. J. Biotechnol., 6: 723-727.
Direct Link |
Freedman, B., R.O. Butterfield and E.H. Pryde, 1986. Transesterification kinetics of soyabean oil. J. Am. Chem. Soc., 63: 1375-1386.
Gordon, J.A., S.K. Heller, T.L. Kaduce and A.A. Spector, 1994. Formation and release of a peroxisome-dependent arachidonic acid metabolite by human skin fibroblasts. J. Biol. Chem., 269: 4103-4109.
Hills, G., 2003. Industrial use of lipases to produce fatty acid esters. Eur. J. Lipid Sci. Technol., 105: 601-607.
Direct Link |
Hus, A.F., K. Jones, T.A. Fogolia and W.N. Marmer, 2002. Immobilized enzymes: Temperature indicators for dielectric pasteurization process. Applied Biochem. Biotechnol., 36: 181-181.
Iftikhar, T. and A. Hussain, 2002. Effect of nutrients on extracellular lipase production by mutant strain of Rhizopus oligosporus Tuv-31. Biotechnology, 1: 15-20.
Direct Link |
Jaeger, K.E. and T. Eggert, 2002. Lipases for biotechnology. Curr. Opin. Biotechnol., 13: 390-397.
CrossRef | PubMed | Direct Link |
Jegannathan, K.R., S. Abang, D. Poncelet, E.S. Chan and P. Ravindra, 2008. Production of biodiesel using immobilised lipase: A critical review. Crit. Rev. Biotechnol., 28: 253-264.
CrossRef | PubMed |
Kamini, N.R., T. Fuijii, T. Kurosu and H. Lefuji, 2000. Lipase catalysed methanolysis of vegetable oils in aqueous medium by Cryptococcus sp. S-2. Process Biochem., 36: 317-324.
Kumar, A. and R.A. Gross, 2000. Candida antarctica lipase B catalyzed transesterification: New synthetic routes to copolymers. J. Am. Chem. Soc., 122: 11767-11770.
Kumari, A., P. Mahapatra, V.K. Garlapati and R. Banerjee, 2009. Enzymatic transesterification of Jatropha oil. Biotechnol. Biofuels, 2: 1-7.
CrossRef | PubMed | Direct Link |
Mohan, T.S., A. Palavesam and G. Immanvel, 2008. Isolation and characterization of lipase-producing Bacillus strains from oil mill waste. Afr. J. Biotechnol., 7: 2728-2735.
Direct Link |
Naka, Y., S. Amano and K. Yamashita, 1986. Effect of surface active agents on the production of Pseudomonas lipase. Yukagaku, 35: 459-462.
Ozlem, O., A.A. Mehmet and G. Salih, 2007. Partial purification of total body lipase from Gryllus campestris L. (Orthoptera: Gryllidae). Fen Bilimleri Dergisi Cilt 28 Sayı 2, http://eskiweb.cumhuriyet.edu.tr/edergi/makale/1544.pdf.
Roy, I., A. Gupta, S.K. Khare, V.S. Bisaria and M.N. Gupta, 2003. Immobilization of xylan-degrading enzymes from Melanocarpus albomyces IIS 68 on the smart polymer Eudragit L-100. Applied Microbiol. Biotechnol., 61: 309-313.
Sadasivam, S. and A. Manickam, 1991. Biochemical methods for Agricultural Sciences. Wiley Eastern Limited and Tamilnadu Agricultural University, New Delhi.
Schuchardt, U., R. Sercheli and R.M. Vargas, 1998. Transesterfication of vegetable oils: A review. J. Braz. Chem. Soc., 9: 199-210.
Shah, S., K. Solanki and M.N. Gupta, 2007. Enhancement of lipase activity in nonaqueous media upon immobilization on multiwalled carbon nanotubes. Chem. Central J., 1: 30-30.
Direct Link |
Shah, S., S. Sharma and M.N. Gupta, 2004. Biodiesel preparation by lipase catalyzed transesterification of jatropha oil. Energy Fuels, 18: 154-159.
Shieh, C.J., H.F. Liao and C.C. Lee, 2003. Optimization of lipase-catalyzed biodiesel by response surface methodology. Bioresour. Technol., 88: 103-106.
CrossRef | Direct Link |
Shukla, P. and K. Gupta, 2007. Ecological screening for lipolytic molds and process optimization for lipase production from Rhizopus Oryzae KG-5. J. Applied Sci. Environ. Sanitation, 2: 35-42.
Direct Link |
Singh, M., S. Singh, R.S. Singh, Y. Chisti and U.C. Banerjee, 2008. Transesterification of primary and secondary alcohols using Pseudomonas aeruginosa lipase. Bioresour. Technol., 99: 2116-2120.
Stamatis, H., E.C. Voutsas, C. Delimitsou, F.N. Kolisis and D. Tassios, 2000. Enzymatic production of alkyl esters through lipase cataysed transesterification reaction in organic solvents: Solvent effects and prediction capabilities of equilibrium conversions. Biocatalysis Biotransformations, 18: 259-269.
Direct Link |
Varma, R., S.M. Kasture, B.G. Gaikwad, S. Nene and U.R. Kalkote, 2007. Lipase catalysed ennantioselective amidation of α phenyl-ethylamine. Asian J. Biochem., 2: 279-283.
Direct Link |
Zeng, Z., X. Xiao, P. Wang and F. Wang, 2004. Screening and characterization of psychrotrophic, lipolytic bacteria from deep-sea sediments. J. Microbiol. Biotechnol., 14: 952-958.
Direct Link |