- Email: [email protected]
Effect of pretreatment by different organic solvents on esterification activity and conformation of immobilized Pseudomonas cepacia lipase
Process Biochemistry 45 (2010) 1176–1180
Contents lists available at ScienceDirect
Process Biochemistry journal homepage: www.elsevier.com/locate/procbio
Short communication
Effect of pretreatment by different organic solvents on esterification activity and conformation of immobilized Pseudomonas cepacia lipase Yun Liu a,∗ , Xuan Zhang b , Hui Tan c , Yunjun Yan a , B.H. Hameed d a
College of Life Science and Technology, Huazhong University of Science and Technology, No. 1037 Luoyu Road, Wuhan 430074, Hubei, PR China Melbourne Graduate School of Science, University of Melbourne, Victoria 3010, Australia College of Life Science and Technology, Central South University of Nationalities, Wuhan 430072, PR China d School of chemical Engineering, Engineering Campus, Universiti Sains Malaysia, 14300 Nibong, Tebal, Penang, Malaysia b c
a r t i c l e
i n f o
Article history: Received 1 December 2009 Received in revised form 16 March 2010 Accepted 16 March 2010 Keywords: Pseudomonas cepacia lipase Esterification activity Fourier transform-infrared spectrometry Secondary structure
a b s t r a c t Experiments were carried out to investigate the effect of pretreatment by organic solvents with different hydrophobicities, functional groups and molecular constitutions as activation agents on initial esterification activity and secondary structure of immobilized Pseudomonas cepacia lipase. The results showed that esterification activity of immobilized P. cepacia lipase treated with organic solvents containing –C O– and –C N– functional groups was highest, followed by the one treated with –C–C– functional groups but the lowest with –OH and aromatics functional groups. An organic solvent with a branched structure was more favorable compared with a straight chain in terms of enhancing enzyme activity. Conformational studies via Fourier transform-infrared spectroscopy indicated that the catalytic activity variance was attributed to the secondary structure changes for immobilized P. cepacia lipase treated with organic solvents. Moreover, the effects of moisture, pH and temperature on the esterification activity of immobilized P. cepacia lipase were also addressed. © 2010 Elsevier Ltd. All rights reserved.
1. Introduction Lipases (glycerol ester hydrolases EC 3.1.1.3) constitute a special group of enzymes which biological function is to catalyze the hydrolysis of triglycerides into diglycerides, monoglycerides, free fatty acids (FFA) and glycerol [1]. Numerous scientific researches showed that marked differences in esterification activity were observed in terms of lipase source, degree of purity, state (free or immobilized), substrate, and reaction media (solvent-free or biphasic). The application of enzymes in organic solvents media has become one of the most exciting fields of enzymology in recent years [2–5]. The usage of organic solvents is especially advantageous to transform substrates that are unstable or poorly soluble in water. Furthermore, at low moisture content, many water-dependent side reactions are avoided, and the denaturation of lipases is also minimized. However, lipase catalytic efficiency in organic media is, in most cases, in the orders of magnitude lower than in aqueous systems [6]. This behavior is ascribed to different causes, such as diffusion limitations, high saturating substrate concentrations, restricted protein flexibility, low stabilization of the enzyme substrate intermediate and even partial
∗ Corresponding author. Tel.: +86 27 87792214; fax: +86 27 87792213. E-mail addresses: [email protected] (Y. Liu), [email protected] (Y. Yan). 1359-5113/$ – see front matter © 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.procbio.2010.03.023
enzyme denaturation which become irreversible in anhydrous solvents. Nevertheless, upon placing a lipase in an organic medium as an interfacial activation agent, the lipase can be altered its native structure and function [7]. Therefore, the correlation between conformational structure and esterification activity requires direct measurement of active-site structure and the effect of the reaction medium on the transition state of the reaction [8]. Using Fourier transform-infrared (FT-IR) spectroscopy, it is very easy to know the changes of the non-covalent forces (hydrogen bonding, ionic, hydrophobic and van der Waals interactions) that maintain the native secondary and tertiary structures of lipases, as a result of the addition of organic solvents to an aqueous enzyme solution [9]. Pseudomonas cepacia lipase (now called Burkholderia cepacia lipase, also known under its commercial trade name as lipase PS) has been recently proven to be among the most versatile lipases. Although the application of lipase PS in organic solvents medium had intensively been studied [3,10–12], the relationship of esterification activity and conformation behavior for lipase PS treated with organic solvents has rarely been comprehensively explored so far. Therefore, the main purposes of this work are: (1) to investigate the esterification activity of lipase PS treated with organic solvents (including ethanol, acetone, isopropanol, acetonitrile, n-hexane, nheptane and toluene) with different solvent hydrophobicity (log P), functional groups (including –OH, C N, C O, C–C, and phenyl group) and molecular constitution (including straight chain and
Y. Liu et al. / Process Biochemistry 45 (2010) 1176–1180
branch chain). (2) To elucidate the secondary structure changes of lipase PS using FT-IR spectroscopy after it is treated with the above-mentioned organic solvents.
1177
a mixture of 0.40 g lauric acid (0.2 M), 0.37 g dodecanol (0.2 M) in 10 mL isooctane solvent and 0.01 mL buffer solution. The vial was placed in a controlled temperature shaker for 30 min at 37 ◦ C and shaken at the rate of 200 rpm. The reaction was stopped after 30 min by adding 10 mL mixture solutions of ethanol and acetone (1:1, v/v), and the mixture was immediately titrated the unreacted FFA with 0.05 M alcoholic NaOH solution using phenolphthalein as indicator. One unit is the amount of the enzyme which catalyzes the formation of 1 mol dodecanol laurate in 1 min at 37 ◦ C. Therefore, initial esterification activity (U min−1 g−1 ) was expressed as mol of ester formed per min per gram protein. After immobilization, the esterification activity of lipase PS increased 40.72-folds (free lipase PS power 3032 ± 2.57 U min−1 g−1 and immobilized lipase PS 127,530 ± 1.44 U min−1 g−1 ). Protein content determination of lipase PS was detailed by Petkar et al. [14]. The protein content of free and immobilized lipase PS was 0.09 wt% and 0.04 wt%, respectively.
2. Materials and methods 2.1. Materials Novozym 435 (lipase B from Candida antarctica), and Lipozyme TLIM (lipase from Thermomyces lanuginosus) were from Novo Nordisk, Denmark. Lipase PS powder (lipase from P. cepacia) was from Amano Pharmaceutical Co. Ltd. (Nagoya, Japan). Macroporous resin NKA with particle size of 0.3–1.25 mm was from Tianjin Nankai Sci. & Tech. Co. Ltd. Tianjin, China. Ethanol, acetone, isopropanol, acetonitrile, n-hexane, n-heptane and toluene were of analytical grade and from Sinopharm Chemical Reagent Co. Ltd. Shanghai, China. All other reagents made in China were of analytical grade.
2.5. Conformation analysis of immobilized lipase PS by FT-IR spectrometry The measurement of the lipases secondary structure by FT-IR spectrometry was detailed by Quiroga et al. [15]. IR spectra was measured at 25 ◦ C by a Vextex 70 FTIR spectrometer (Bruker Optik GMBH, Germany) equipped with a nitrogen-cooled, mercury–cadmium–tellurium (MCT) detector, in the region of 4000–400 cm−1 . Conditions were 4 cm−1 spectral resolution, 20 kHz scan speed, 128 scan co-additions, and triangular apodization. By setting the microscope square diaphragm aperture to 100 m × 100 m, an excellent spectrum was collected within a few minutes. The spectrum acquisition (all samples were overlaid on a zinc selenide attenuated total reflectance (ATR) accessory) is from IR spectra, and the secondary structure elements based on the information of amide I region and the band assignment were manipulated using WinSpec software (LISE-Faculteˇis Universitaires Notre-Dame de la Paix, Namur, Belgium).
2.2. Lipase PS immobilized on macroporous resin NKA To screen a suitable resin support for lipase PS immobilization, four types of macroporous resins (AB-8, NKA, Amberlite XAD 7HP and Diaion HP2MGL resins) were tested. It was demonstrated that lipase PS immobilized on resin NKA exhibited the highest specific activity, immobilization efficiency and activity recovery. In addition, macroporous resin NKA is an inert support and stable in organic solvents. So, resin NKA is chosen as the support matrix for lipase PS immobilization in this work. The immobilization procedures are as follow: 10 g resin NKA was immersed in 50 mL absolute ethanol for 4 h. After evaporating ethanol, resin NKA was eluted 6 times with distilled water and placed in a beaker filled with 50 mL 0.05 M phosphate buffer (pH 7) for overnight at 4 ◦ C, and then the remaining buffer was eliminated so that no more liquid phase was present in the tested tube. 50 mL of 0.05 M phosphate buffer (pH 7) containing 8.0 g free lipase PS was added into the tube and the mixture was stirred in a rotary shaker at 200 rpm and 37 ◦ C for 2 h. The suspension was filtered through a Buchner funnel and the immobilized lipase PS was washed on the filter paper with 50 mL of 0.05 M phosphate buffer (pH 7) to remove soluble enzyme for 5 times. The immobilized lipase PS was stored at 4 ◦ C for usage after it was dried in a FD-1D-50 vacuum desiccator. The immobilization efficiency of immobilized lipase PS was estimated to be 94% (total initial activity minus total activity remaining in solution after immobilization).
3. Results and discussion 3.1. Effect of organic solvents with different functional groups The effect of pretreatment by seven different organic solvents on initial esterification activity of the immobilized lipase PS was investigated and the results are shown in Table 1. Comparing with the control, the esterification activity of the lipase PS treated with all solvents except for ethanol and toluene are significantly higher. The esterification activity is highest for acetone with –C O– functional group and isopropanol with branch chain of –OH functional group. On the contrary, the esterification activity is lowest for ethanol with –OH functional group and toluene with aromatics functional group. Analogous trends are observed for Novozym 435 and Lipozyme TLIM (Table 1). The variance of esterification activity is observed after the immobilized lipase PS is treated with ethanol (–OH), acetone (–C O–) and acetonitrile (–C N–), which display similar log P but different functional groups. It is suggested that functional group of organic solvents is an important factor affecting the esterification activity of lipases. A probable explanation is ascribed to the variation of water retained in the microenvironment of the catalytic active site, which is necessary to maintain the dynamical properties of the enzyme [16]. This explanation was addressed in many literatures. For example, Secundo and Carrea [11] demonstrated that Novozym 435 showed differ-
2.3. Immobilized lipase PS treated with organic solvents To elucidate the effect of organic solvents as activation agents on the esterification activity of immobilized lipase PS, organic solvents with different hydrophobicity (log P), functional group and molecular constitution were used to treat the immobilized lipase PS. The tested organic solvents were ethanol, acetone, acetonitrile, isopropanol, n-heptane, n-hexane, and toluene. 0.2 g of the immobilized lipase PS was placed in different test tubes, and then 2 mL of various solvents (i.e., actone, isopropanol, and n-hexane) were added to the tubes to swell the enzyme particles. Each treatment was immersed and stirred in a rotary shaker at 20 ◦ C for 2 h. Then, the solvent was filtered through a Buchner funnel by vacuum. The treated immobilized lipase was dried to remove the organic solvents in a vacuum desiccator before esterification activity assay. Another experiment containing the enzyme without organic solvent treatment was carried out as a control. 2.4. Assay of enzyme initial esterification activity Esterification activity was assayed for the change of free fatty acid (FFA) concentration before and after an esterification reaction. The determination procedures of lipase PS initial activity were modified according to the method described by Talukder et al. [13]. 40 mg lipases were added into a screw capped vial containing
Table 1 Effects of pretreatment by organic solvents with different functional groups on the initial esterification activities of the three immobilized lipases (mean ± STDV, n = 3). Treatment methodsb
log P
Functional group
a
Control Ethanol Acetone Acetonitrile Isopropanol n-hexane n-heptane Toluene a
−0.24 −0.23 −0.33 0.8 5.7 5.5 2.9
–OH –C O– –C N– –OH –C–C– –C–C– C6 H5 –
Immobilized PS (U min−1 g−1 ) 8,875.00 7,968.75 16,668.75 13,231.25 13,856.25 14,531.25 11,562.50 8,800.00
± ± ± ± ± ± ± ±
a
1.40 1.00 1.76* 0.76* 1.04* 0.75* 2.18 0.52
Novozym 435 (U min−1 g−1 ) 9,893.75 8,906.25 20,625 14,687.5 23,856.25 16,456.25 10,731.25 9,793.75
± ± ± ± ± ± ± ±
a
2.02 0.25 0.50* 0.87* 0.29* 0.63* 4.48 3.75
Lipozyme TLIM (U min−1 g−1 ) 3,543.75 3,125.00 8,850.00 6,356.25 10,731.25 7,656.25 4,862.50 2,187.50
± ± ± ± ± ± ± ±
0.29a 1.04 0.76* 1.15* 0.76* 1.95* 0.50 1.32
Esterification activities of the immobilized lipase PS without organic solvent treatment were measured as control reference. Substrate: a mixture of 0.40 g lauric acid (0.2 M), 0.37 g dodecanol (0.2 M) in 10 mL isooctane solvent and 0.01 mL buffer solution, reaction time 30 min, temperature 37 ◦ C, and shaking rate 200 rpm. * P < 0.001 level significant. b
1178
Y. Liu et al. / Process Biochemistry 45 (2010) 1176–1180 Table 2 Quantitative estimation (%) of the secondary structure elements of treated immobilized lipase PS calculated by FT-IR analysis in amide I region.
Fig. 1. Effect of pretreatment by organic solvents with different hydrophobicity and molecular constitution on the immobilized lipase PS. Substrate: a mixture of 0.40 g lauric acid (0.2 M), 0.37 g dodecanol (0.2 M) in 10 mL isooctane solvent and 0.01 mL buffer solution, reaction time 30 min, temperature 37 ◦ C, pH = 6.8, shaking rate 200 rpm.
ent activity when treated with organic solvents with different functional groups. Su and Wei [5] reported that higher substrate conversion rate was obtained for t-butanol, t-pentanol, acetonitrile and 1,4-dioxane when Novozym 435 was treated with different pure polar solvents. Generally, solvents with a log P value below 2 are not suitably used to treat the lipases for enhancement of the catalytic activity [17]. However, high catalytic activity was observed when Novozym 435 and Lipozyme TLIM were treated with acetonitrile and 1,4-dioxane which both have very low log P values (log P = −0.33) [5]. It has been proven that the activity of lipases treated with organic solvent is not only dependent on the hydrophobicity (log P), but also the functional group of organic solvents.
3.2. Effect of organic solvent with different molecular constitution In order to further demonstrate the effect of molecular constitution of organic solvent on the esterification activity of immobilized lipase PS, a comparison experiment was conducted with three organic solvents sharing the same hydroxyl group, among which two were ethanol and butanol with straight chain (–OH group located at the first carbon atom) but with different log P, and another one was isopropanol with branch chain (–OH group located at the second carbon atom), whose log P is equal to butanol. It is observed that, with straight chain molecular constitution, lipase PS exhibits different esterification activity after treatment with ethanol and butanol which are of different log P (Fig. 1). Comparing to the activity before treatment (8875 ± 1.40 U min−1 g−1 ), the activity of lipase PS is slightly higher (10,206.25 ± 0.29 U min−1 g−1 ) after treatment with butanol but a little lower (7968.75 ± 1.00 U min−1 g−1 ) with ethanol. The reason is due to the variant log P between butanol (log P = 0.8) and ethanol (log P = −0.24), which affects the microenvironment moisture around the catalytic active site. However, butanol and isopropanol share the same log P (log P = 0.8), but immobilized lipase PS shows much higher activity after treatment with isopropanol than with butanol. This observation suggests that the molecular constitution of organic solvent also plays an important role, and a solvent with a branched chain is more contributions to enhancing enzymatic activity, which agrees well with that reported by Wu et al. [18], who demonstrated that the activity of lipase from Candida rugosa treated with isooctane was much higher than that with octane, although the two organic solvents are of the same log P (log P = 4.5). In conclusions, molecular constitution also plays an important role in affecting lipase PS activity apart from the hydrophobicity and group function of organic solvents.
Organic solvents
␣-Helix (%)
-Sheet (%)
-Turns (%)
Random coil (%)
Controla Ethanol Toluene n-Heptane Butanol Acetone Acetonitrile Isopropanol n-Hexane
33.4 28.7 26.3 18.2 17.1 16.8 19.1 17.5 17.2
28.2 34.6 39.1 43.2 44.2 46.7 45.2 46.8 44.9
30.8 24.1 23.2 26.7 24.1 28.9 27.4 26.8 27.5
7.6 11.9 12.0 10.9 11.5 8.7 7.1 8.6 9.7
a The immobilized lipase PS without organic solvent treatment were measured as control.
3.3. The secondary structure of immobilized lipase PS by FT-IR spectrometry It is logical to consider conformational structure change of the treated immobilized lipase PS as a probable cause for the esterification activity variance. To verify this hypothesis, FT-IR experiments were conducted to analyze the secondary structure variation of the treated immobilized lipase PS. FT-IR technique renders feasible the study of a protein’s secondary structure because proteins absorb in infra in the amide regions, due to the peptide bond vibrations. The amide I region (mainly due to the C O stretching vibration) found at approximately 1600–1700 cm−1 is mostly used in protein secondary structure determination due to its sensitivity in conformational changes and the significantly higher signal intensity than in other amide bands. The secondary structure element variations of treatment immobilized lipase PS are shown in Table 2. The ATR spectra of the second derivative and deconvoluted amide I spectra (1600–1700 cm−1 ) of immobilized lipase PS treated with organic solvents have been shown for supporting information (Fig. SI-1). As it can be seen from Table 2, the immobilized lipase PS treated with organic solvents undergoes some alterations in its secondary structure. In general, treated immobilized lipase PS exhibits a decrease in ␣-helix and -turns content and an increase in sheet and random coil except for the case of acetonitrile which random coil decreased. The decrease in ␣-helix content of immobilized lipase PS treated with organic solvents probably affects the active site of lipase PS leading to a crucial effect on its activity. This observation is in accordance with the initial esterification activity observed in Sections 3.1 and 3.2. However, an increase in -sheet content was observed in all cases when treated with organic solvents. This increase can be attributed to a loss of hydrogen-bonding interactions between the water molecules and the surface of the protein resulting in a more rigid structure for the treated lipase PS. In conclusions, the secondary structure change of lipase PS is contributed to the variations of its esterification activity when it is treated with organic solvents. 3.4. Basic enzymatic properties of treated immobilized lipase PS In order to investigate the basic enzymatic properties of immobilized lipase PS treated with organic solvents, the routine experiments on the effects of pH (from 4.0 to 8.0), moisture content (from 0 wt% to 2 wt%) and temperature (from 25 ◦ C to 55 ◦ C) on catalytic activity were carried out. The results are shown in Fig. 2. Comparing with the untreated lipase PS, the behavior of treated lipase PS activity changed markedly at all tested pH ranges. From Fig. 2A, the maximum esterification activity of the treated lipase PS is 12,918.75 ± 0.38 U min−1 g−1 at the optimal pH 6.0. Beyond pH 6.0, the esterification activity of the treated lipase PS decreases. Specifically, the low activity at lower pH (below pH 6.0) is attributed to a change in lipase PS micro-structure, which is caused by the
Y. Liu et al. / Process Biochemistry 45 (2010) 1176–1180
1179
At 45 ◦ C, the esterification activity of the treated lipase PS ranges from 9893.75 ± 1.04 to 116,125.50 ± 0.88 U min−1 g−1 . A further increase of temperature beyond 45 ◦ C leads to protein denaturation of lipase PS, resulting in a decrease of enzyme activity [18,20]. 4. Conclusions To investigate the effect of pretreatment by different organic solvents on enzyme activity and conformational structure, eight organic solvents, such as acetone, isopropanol, n-hexane, acetonitrile, butanol, ethanol, toluene, and n-heptane, are employed to treat the immobilized lipase PS. It is demonstrated that hydrophobicity, functional group and molecular constitution of organic solvents are all crucial factors to affect esterification activity. FTIR studies show that secondary structure of the immobilized lipase PS is altered to a significant extent, which is responsible for the variance of esterification activity. It is concluded that there is a close correlation between secondary structure and esterification activity of immobilized lipase PS, which is of significance for its further application. Acknowledgements The authors are deeply indebted to the Analytical and Testing Center of Huazhong University of Science and Technology for FT-IR analysis, and the financial support from the National High Technology Research and Development Program of China (863 Program) (2009AA03Z232 and 2007AA100703), Program for New Century Excellent Talents in University (NCET-07-0336), Doctoral Education Fund for New Teachers (20090142120090), and the Natural Science Foundation of Hubei Province (2008CDB359 and 2009CDA046). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.procbio.2010.03.023.
Fig. 2. Effect of pH, moisture and temperature on the activity of the organic solvent treated immobilized lipase PS. Substrate: a mixture of 0.40 g lauric acid (0.2 M), 0.37 g dodecanol (0.2 M) in 10 mL isooctane solvent and 0.01 mL buffer solution, reaction time 30 min, temperature 37 ◦ C, shaking rate 200 rpm.
absorption of excessive H+ leading to an irreversible protein denaturation of lipase PS. The enzyme activity decreases rapidly due to a more severe deactivation of the enzyme at higher pH (above pH 6.0). In addition, pH effect can also be manifested due to partitioning of hydrogen ions between the solution and the support surface [19]. Similar trends of effect of pH on the activities were also observed in lipases from C. rugosa, C. antarctica, Rhizopus delemar [18–21]. Effect of moisture on esterification activity of the immobilized lipases had been intensively reported [7,10,21]. It is demonstrated (Fig. 2B) that the optimum moisture content is 0.1 wt% for the immobilized lipase PS after treatment with all tested organic solvents. Above 0.1 wt% moisture content, an obvious decrease of esterification activity is observed. The probable reason is attributed to a shift in the chemical equilibrium in the higher water content range. Similar observations were also reported by Petkar et al. [14] and Naoe et al. [21], who both found that Humicola lanuginosa lipase, C. antarctica lipase B and Rhizomucor miehei lipase exhibited high activities at extremely low water content, as well as lipase from R. delemar. From Fig. 2C, it is noted that the optimum temperature is 45 ◦ C for the immobilized lipase PS after treatment with organic solvents.
References [1] Liu Yun, Yan Yunjun, Zhang Xuan, Tan Hui, Xin Honglin, Yao An’na. Biodiesel production by combined lipases catalyzed transesterification: optimization and kinetics [J]. AIChE Journal, 2009, doi:10 1002/aic.12090. [2] Fjerbaek L, Christensen KV, Norddahl B. A review of the current state of biodiesel production using enzymatic transesterification. Biotechnol Bioeng 2009;102:1298–315. [3] Hara P, Hanefeld U, Kanerv LT. Immobilised Burkholderia cepacia lipase in dry organin solvents and ionic liquids: a comparison. Green Chem 2009;11:250–6. [4] Liu Yun, Tan Hui, Zhang Xuan, Yan Yunjun, Hameed BH. Effect of monohydric alcohols on enzymatic transesterification for biodiesel production. Chem Eng J 2010;157:223–9. [5] Su E, Wei D. Improvement in lipase-catalyzed methanolysis of triacylglycerols for biodiesel production using a solvent engineering method. J Mol Catal BEnzym 2008;55:118–25. [6] Priego J, Ortíz-Nava C, Carrillo-Morales M, López-Munguía A, Escalante J, Castillo E. Solvent engineering: an effective tool to direct chemoselectivity in a lipase-catalyzed michael addition. Tetrahedron 2009;65:536–9. [7] Klibanov AM. Improving enzymes by using them in organic solvents. Nature 2001;409:241–6. [8] Hiramatsu H, Kitagawa T. FT-IR approaches on amyloid fibril structure. Biochim Biophys Acta 2005;1753:100–7. [9] Natalello A, Ami D, Brocca S, Lotti M, Doglia SM. Secondary structure, conformational stability and glycosylation of a recombinant Candida rugosa lipase studied by Fourier-transform infrared spectroscopy. Biochem J 2005;385:511–7. [10] Pencreach G, Baratti JC. Activity of Pseudomonas cepacia lipase in organic media is greatly enhanced after immobilization on a polypropylene support. Appl Microbiol Biotechnol 1997;47:630–5. [11] Secundo F, Carrea G. Lipase activity and conformation in neat organic solvents. J Mol Catal B-Enzym 2002;19:93–102. [12] Secundo F, Spadaro S, Carrea G, Overbeeke PLA. Optimization of Pseudomonas cepacia lipase preparations for catalysis in organic solvents. Biotechnol Bioeng 1999;62:554–61.
1180
Y. Liu et al. / Process Biochemistry 45 (2010) 1176–1180
[13] Talukder MMR, Tamalampudy S, Li CJ, Le YL, Wu JC, Kondo A, et al. An improved method of lipase preparation incorporating both solvent treatment and immobilization onto matrix. Biochem Eng J 2007;33:60–5. [14] Petkar M, Lali A, Caimi P, Daminati M. Immobilization of lipases for nonaqueous synthesis. J Mol Catal B-Enzym 2006;39:83–90. [15] Quiroga E, Camí G, Marchese J, Barberis S. Organic solvents effect on the secondary structure of araujiain hI, in different media. Biochem Eng J 2007;35:198–202. [16] Koskinen AMP, Klibanov AM. Enzymatic reactions in organic media. London: Blackie Academic & Professional; 1996. [17] Laane C, Boeren S, Vos K, Veeger C. Rules for optimization of biocatalysis in organic solvents. Biotechnol Bioeng 1987;30:81–7.
[18] Wu JC, Song BD, Xing AH, Hayashi Y, Talukder MMR, Wang SC. Esterification reactions catalyzed by surfactant-coated Candida rugosa lipase in organic solvents. Proc Biochem 2002;37:1229–33. [19] Klibanov AM. Why are enzymes less active in organic solvents than in water. Trends Biotechnol 1997;15:97–101. [20] Abdullah AZ, Sulaiman NS, Kamaruddin AH. Biocatalytic esterification of citronellol with lauric acid by immobilized lipase on aminopropyl-grafted mesoporous SBA-15. Biochem Eng J 2009;44:263–70. [21] Naoe K, Ohsa T, Kawagoe M, Imai M. Esterification by Rhizopus delemar lipase in organic solvent using sugar ester reverse micelles. Biochem Eng J 2001;9:67–72.
Contents lists available at ScienceDirect
Process Biochemistry journal homepage: www.elsevier.com/locate/procbio
Short communication
Effect of pretreatment by different organic solvents on esterification activity and conformation of immobilized Pseudomonas cepacia lipase Yun Liu a,∗ , Xuan Zhang b , Hui Tan c , Yunjun Yan a , B.H. Hameed d a
College of Life Science and Technology, Huazhong University of Science and Technology, No. 1037 Luoyu Road, Wuhan 430074, Hubei, PR China Melbourne Graduate School of Science, University of Melbourne, Victoria 3010, Australia College of Life Science and Technology, Central South University of Nationalities, Wuhan 430072, PR China d School of chemical Engineering, Engineering Campus, Universiti Sains Malaysia, 14300 Nibong, Tebal, Penang, Malaysia b c
a r t i c l e
i n f o
Article history: Received 1 December 2009 Received in revised form 16 March 2010 Accepted 16 March 2010 Keywords: Pseudomonas cepacia lipase Esterification activity Fourier transform-infrared spectrometry Secondary structure
a b s t r a c t Experiments were carried out to investigate the effect of pretreatment by organic solvents with different hydrophobicities, functional groups and molecular constitutions as activation agents on initial esterification activity and secondary structure of immobilized Pseudomonas cepacia lipase. The results showed that esterification activity of immobilized P. cepacia lipase treated with organic solvents containing –C O– and –C N– functional groups was highest, followed by the one treated with –C–C– functional groups but the lowest with –OH and aromatics functional groups. An organic solvent with a branched structure was more favorable compared with a straight chain in terms of enhancing enzyme activity. Conformational studies via Fourier transform-infrared spectroscopy indicated that the catalytic activity variance was attributed to the secondary structure changes for immobilized P. cepacia lipase treated with organic solvents. Moreover, the effects of moisture, pH and temperature on the esterification activity of immobilized P. cepacia lipase were also addressed. © 2010 Elsevier Ltd. All rights reserved.
1. Introduction Lipases (glycerol ester hydrolases EC 3.1.1.3) constitute a special group of enzymes which biological function is to catalyze the hydrolysis of triglycerides into diglycerides, monoglycerides, free fatty acids (FFA) and glycerol [1]. Numerous scientific researches showed that marked differences in esterification activity were observed in terms of lipase source, degree of purity, state (free or immobilized), substrate, and reaction media (solvent-free or biphasic). The application of enzymes in organic solvents media has become one of the most exciting fields of enzymology in recent years [2–5]. The usage of organic solvents is especially advantageous to transform substrates that are unstable or poorly soluble in water. Furthermore, at low moisture content, many water-dependent side reactions are avoided, and the denaturation of lipases is also minimized. However, lipase catalytic efficiency in organic media is, in most cases, in the orders of magnitude lower than in aqueous systems [6]. This behavior is ascribed to different causes, such as diffusion limitations, high saturating substrate concentrations, restricted protein flexibility, low stabilization of the enzyme substrate intermediate and even partial
∗ Corresponding author. Tel.: +86 27 87792214; fax: +86 27 87792213. E-mail addresses: [email protected] (Y. Liu), [email protected] (Y. Yan). 1359-5113/$ – see front matter © 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.procbio.2010.03.023
enzyme denaturation which become irreversible in anhydrous solvents. Nevertheless, upon placing a lipase in an organic medium as an interfacial activation agent, the lipase can be altered its native structure and function [7]. Therefore, the correlation between conformational structure and esterification activity requires direct measurement of active-site structure and the effect of the reaction medium on the transition state of the reaction [8]. Using Fourier transform-infrared (FT-IR) spectroscopy, it is very easy to know the changes of the non-covalent forces (hydrogen bonding, ionic, hydrophobic and van der Waals interactions) that maintain the native secondary and tertiary structures of lipases, as a result of the addition of organic solvents to an aqueous enzyme solution [9]. Pseudomonas cepacia lipase (now called Burkholderia cepacia lipase, also known under its commercial trade name as lipase PS) has been recently proven to be among the most versatile lipases. Although the application of lipase PS in organic solvents medium had intensively been studied [3,10–12], the relationship of esterification activity and conformation behavior for lipase PS treated with organic solvents has rarely been comprehensively explored so far. Therefore, the main purposes of this work are: (1) to investigate the esterification activity of lipase PS treated with organic solvents (including ethanol, acetone, isopropanol, acetonitrile, n-hexane, nheptane and toluene) with different solvent hydrophobicity (log P), functional groups (including –OH, C N, C O, C–C, and phenyl group) and molecular constitution (including straight chain and
Y. Liu et al. / Process Biochemistry 45 (2010) 1176–1180
branch chain). (2) To elucidate the secondary structure changes of lipase PS using FT-IR spectroscopy after it is treated with the above-mentioned organic solvents.
1177
a mixture of 0.40 g lauric acid (0.2 M), 0.37 g dodecanol (0.2 M) in 10 mL isooctane solvent and 0.01 mL buffer solution. The vial was placed in a controlled temperature shaker for 30 min at 37 ◦ C and shaken at the rate of 200 rpm. The reaction was stopped after 30 min by adding 10 mL mixture solutions of ethanol and acetone (1:1, v/v), and the mixture was immediately titrated the unreacted FFA with 0.05 M alcoholic NaOH solution using phenolphthalein as indicator. One unit is the amount of the enzyme which catalyzes the formation of 1 mol dodecanol laurate in 1 min at 37 ◦ C. Therefore, initial esterification activity (U min−1 g−1 ) was expressed as mol of ester formed per min per gram protein. After immobilization, the esterification activity of lipase PS increased 40.72-folds (free lipase PS power 3032 ± 2.57 U min−1 g−1 and immobilized lipase PS 127,530 ± 1.44 U min−1 g−1 ). Protein content determination of lipase PS was detailed by Petkar et al. [14]. The protein content of free and immobilized lipase PS was 0.09 wt% and 0.04 wt%, respectively.
2. Materials and methods 2.1. Materials Novozym 435 (lipase B from Candida antarctica), and Lipozyme TLIM (lipase from Thermomyces lanuginosus) were from Novo Nordisk, Denmark. Lipase PS powder (lipase from P. cepacia) was from Amano Pharmaceutical Co. Ltd. (Nagoya, Japan). Macroporous resin NKA with particle size of 0.3–1.25 mm was from Tianjin Nankai Sci. & Tech. Co. Ltd. Tianjin, China. Ethanol, acetone, isopropanol, acetonitrile, n-hexane, n-heptane and toluene were of analytical grade and from Sinopharm Chemical Reagent Co. Ltd. Shanghai, China. All other reagents made in China were of analytical grade.
2.5. Conformation analysis of immobilized lipase PS by FT-IR spectrometry The measurement of the lipases secondary structure by FT-IR spectrometry was detailed by Quiroga et al. [15]. IR spectra was measured at 25 ◦ C by a Vextex 70 FTIR spectrometer (Bruker Optik GMBH, Germany) equipped with a nitrogen-cooled, mercury–cadmium–tellurium (MCT) detector, in the region of 4000–400 cm−1 . Conditions were 4 cm−1 spectral resolution, 20 kHz scan speed, 128 scan co-additions, and triangular apodization. By setting the microscope square diaphragm aperture to 100 m × 100 m, an excellent spectrum was collected within a few minutes. The spectrum acquisition (all samples were overlaid on a zinc selenide attenuated total reflectance (ATR) accessory) is from IR spectra, and the secondary structure elements based on the information of amide I region and the band assignment were manipulated using WinSpec software (LISE-Faculteˇis Universitaires Notre-Dame de la Paix, Namur, Belgium).
2.2. Lipase PS immobilized on macroporous resin NKA To screen a suitable resin support for lipase PS immobilization, four types of macroporous resins (AB-8, NKA, Amberlite XAD 7HP and Diaion HP2MGL resins) were tested. It was demonstrated that lipase PS immobilized on resin NKA exhibited the highest specific activity, immobilization efficiency and activity recovery. In addition, macroporous resin NKA is an inert support and stable in organic solvents. So, resin NKA is chosen as the support matrix for lipase PS immobilization in this work. The immobilization procedures are as follow: 10 g resin NKA was immersed in 50 mL absolute ethanol for 4 h. After evaporating ethanol, resin NKA was eluted 6 times with distilled water and placed in a beaker filled with 50 mL 0.05 M phosphate buffer (pH 7) for overnight at 4 ◦ C, and then the remaining buffer was eliminated so that no more liquid phase was present in the tested tube. 50 mL of 0.05 M phosphate buffer (pH 7) containing 8.0 g free lipase PS was added into the tube and the mixture was stirred in a rotary shaker at 200 rpm and 37 ◦ C for 2 h. The suspension was filtered through a Buchner funnel and the immobilized lipase PS was washed on the filter paper with 50 mL of 0.05 M phosphate buffer (pH 7) to remove soluble enzyme for 5 times. The immobilized lipase PS was stored at 4 ◦ C for usage after it was dried in a FD-1D-50 vacuum desiccator. The immobilization efficiency of immobilized lipase PS was estimated to be 94% (total initial activity minus total activity remaining in solution after immobilization).
3. Results and discussion 3.1. Effect of organic solvents with different functional groups The effect of pretreatment by seven different organic solvents on initial esterification activity of the immobilized lipase PS was investigated and the results are shown in Table 1. Comparing with the control, the esterification activity of the lipase PS treated with all solvents except for ethanol and toluene are significantly higher. The esterification activity is highest for acetone with –C O– functional group and isopropanol with branch chain of –OH functional group. On the contrary, the esterification activity is lowest for ethanol with –OH functional group and toluene with aromatics functional group. Analogous trends are observed for Novozym 435 and Lipozyme TLIM (Table 1). The variance of esterification activity is observed after the immobilized lipase PS is treated with ethanol (–OH), acetone (–C O–) and acetonitrile (–C N–), which display similar log P but different functional groups. It is suggested that functional group of organic solvents is an important factor affecting the esterification activity of lipases. A probable explanation is ascribed to the variation of water retained in the microenvironment of the catalytic active site, which is necessary to maintain the dynamical properties of the enzyme [16]. This explanation was addressed in many literatures. For example, Secundo and Carrea [11] demonstrated that Novozym 435 showed differ-
2.3. Immobilized lipase PS treated with organic solvents To elucidate the effect of organic solvents as activation agents on the esterification activity of immobilized lipase PS, organic solvents with different hydrophobicity (log P), functional group and molecular constitution were used to treat the immobilized lipase PS. The tested organic solvents were ethanol, acetone, acetonitrile, isopropanol, n-heptane, n-hexane, and toluene. 0.2 g of the immobilized lipase PS was placed in different test tubes, and then 2 mL of various solvents (i.e., actone, isopropanol, and n-hexane) were added to the tubes to swell the enzyme particles. Each treatment was immersed and stirred in a rotary shaker at 20 ◦ C for 2 h. Then, the solvent was filtered through a Buchner funnel by vacuum. The treated immobilized lipase was dried to remove the organic solvents in a vacuum desiccator before esterification activity assay. Another experiment containing the enzyme without organic solvent treatment was carried out as a control. 2.4. Assay of enzyme initial esterification activity Esterification activity was assayed for the change of free fatty acid (FFA) concentration before and after an esterification reaction. The determination procedures of lipase PS initial activity were modified according to the method described by Talukder et al. [13]. 40 mg lipases were added into a screw capped vial containing
Table 1 Effects of pretreatment by organic solvents with different functional groups on the initial esterification activities of the three immobilized lipases (mean ± STDV, n = 3). Treatment methodsb
log P
Functional group
a
Control Ethanol Acetone Acetonitrile Isopropanol n-hexane n-heptane Toluene a
−0.24 −0.23 −0.33 0.8 5.7 5.5 2.9
–OH –C O– –C N– –OH –C–C– –C–C– C6 H5 –
Immobilized PS (U min−1 g−1 ) 8,875.00 7,968.75 16,668.75 13,231.25 13,856.25 14,531.25 11,562.50 8,800.00
± ± ± ± ± ± ± ±
a
1.40 1.00 1.76* 0.76* 1.04* 0.75* 2.18 0.52
Novozym 435 (U min−1 g−1 ) 9,893.75 8,906.25 20,625 14,687.5 23,856.25 16,456.25 10,731.25 9,793.75
± ± ± ± ± ± ± ±
a
2.02 0.25 0.50* 0.87* 0.29* 0.63* 4.48 3.75
Lipozyme TLIM (U min−1 g−1 ) 3,543.75 3,125.00 8,850.00 6,356.25 10,731.25 7,656.25 4,862.50 2,187.50
± ± ± ± ± ± ± ±
0.29a 1.04 0.76* 1.15* 0.76* 1.95* 0.50 1.32
Esterification activities of the immobilized lipase PS without organic solvent treatment were measured as control reference. Substrate: a mixture of 0.40 g lauric acid (0.2 M), 0.37 g dodecanol (0.2 M) in 10 mL isooctane solvent and 0.01 mL buffer solution, reaction time 30 min, temperature 37 ◦ C, and shaking rate 200 rpm. * P < 0.001 level significant. b
1178
Y. Liu et al. / Process Biochemistry 45 (2010) 1176–1180 Table 2 Quantitative estimation (%) of the secondary structure elements of treated immobilized lipase PS calculated by FT-IR analysis in amide I region.
Fig. 1. Effect of pretreatment by organic solvents with different hydrophobicity and molecular constitution on the immobilized lipase PS. Substrate: a mixture of 0.40 g lauric acid (0.2 M), 0.37 g dodecanol (0.2 M) in 10 mL isooctane solvent and 0.01 mL buffer solution, reaction time 30 min, temperature 37 ◦ C, pH = 6.8, shaking rate 200 rpm.
ent activity when treated with organic solvents with different functional groups. Su and Wei [5] reported that higher substrate conversion rate was obtained for t-butanol, t-pentanol, acetonitrile and 1,4-dioxane when Novozym 435 was treated with different pure polar solvents. Generally, solvents with a log P value below 2 are not suitably used to treat the lipases for enhancement of the catalytic activity [17]. However, high catalytic activity was observed when Novozym 435 and Lipozyme TLIM were treated with acetonitrile and 1,4-dioxane which both have very low log P values (log P = −0.33) [5]. It has been proven that the activity of lipases treated with organic solvent is not only dependent on the hydrophobicity (log P), but also the functional group of organic solvents.
3.2. Effect of organic solvent with different molecular constitution In order to further demonstrate the effect of molecular constitution of organic solvent on the esterification activity of immobilized lipase PS, a comparison experiment was conducted with three organic solvents sharing the same hydroxyl group, among which two were ethanol and butanol with straight chain (–OH group located at the first carbon atom) but with different log P, and another one was isopropanol with branch chain (–OH group located at the second carbon atom), whose log P is equal to butanol. It is observed that, with straight chain molecular constitution, lipase PS exhibits different esterification activity after treatment with ethanol and butanol which are of different log P (Fig. 1). Comparing to the activity before treatment (8875 ± 1.40 U min−1 g−1 ), the activity of lipase PS is slightly higher (10,206.25 ± 0.29 U min−1 g−1 ) after treatment with butanol but a little lower (7968.75 ± 1.00 U min−1 g−1 ) with ethanol. The reason is due to the variant log P between butanol (log P = 0.8) and ethanol (log P = −0.24), which affects the microenvironment moisture around the catalytic active site. However, butanol and isopropanol share the same log P (log P = 0.8), but immobilized lipase PS shows much higher activity after treatment with isopropanol than with butanol. This observation suggests that the molecular constitution of organic solvent also plays an important role, and a solvent with a branched chain is more contributions to enhancing enzymatic activity, which agrees well with that reported by Wu et al. [18], who demonstrated that the activity of lipase from Candida rugosa treated with isooctane was much higher than that with octane, although the two organic solvents are of the same log P (log P = 4.5). In conclusions, molecular constitution also plays an important role in affecting lipase PS activity apart from the hydrophobicity and group function of organic solvents.
Organic solvents
␣-Helix (%)
-Sheet (%)
-Turns (%)
Random coil (%)
Controla Ethanol Toluene n-Heptane Butanol Acetone Acetonitrile Isopropanol n-Hexane
33.4 28.7 26.3 18.2 17.1 16.8 19.1 17.5 17.2
28.2 34.6 39.1 43.2 44.2 46.7 45.2 46.8 44.9
30.8 24.1 23.2 26.7 24.1 28.9 27.4 26.8 27.5
7.6 11.9 12.0 10.9 11.5 8.7 7.1 8.6 9.7
a The immobilized lipase PS without organic solvent treatment were measured as control.
3.3. The secondary structure of immobilized lipase PS by FT-IR spectrometry It is logical to consider conformational structure change of the treated immobilized lipase PS as a probable cause for the esterification activity variance. To verify this hypothesis, FT-IR experiments were conducted to analyze the secondary structure variation of the treated immobilized lipase PS. FT-IR technique renders feasible the study of a protein’s secondary structure because proteins absorb in infra in the amide regions, due to the peptide bond vibrations. The amide I region (mainly due to the C O stretching vibration) found at approximately 1600–1700 cm−1 is mostly used in protein secondary structure determination due to its sensitivity in conformational changes and the significantly higher signal intensity than in other amide bands. The secondary structure element variations of treatment immobilized lipase PS are shown in Table 2. The ATR spectra of the second derivative and deconvoluted amide I spectra (1600–1700 cm−1 ) of immobilized lipase PS treated with organic solvents have been shown for supporting information (Fig. SI-1). As it can be seen from Table 2, the immobilized lipase PS treated with organic solvents undergoes some alterations in its secondary structure. In general, treated immobilized lipase PS exhibits a decrease in ␣-helix and -turns content and an increase in sheet and random coil except for the case of acetonitrile which random coil decreased. The decrease in ␣-helix content of immobilized lipase PS treated with organic solvents probably affects the active site of lipase PS leading to a crucial effect on its activity. This observation is in accordance with the initial esterification activity observed in Sections 3.1 and 3.2. However, an increase in -sheet content was observed in all cases when treated with organic solvents. This increase can be attributed to a loss of hydrogen-bonding interactions between the water molecules and the surface of the protein resulting in a more rigid structure for the treated lipase PS. In conclusions, the secondary structure change of lipase PS is contributed to the variations of its esterification activity when it is treated with organic solvents. 3.4. Basic enzymatic properties of treated immobilized lipase PS In order to investigate the basic enzymatic properties of immobilized lipase PS treated with organic solvents, the routine experiments on the effects of pH (from 4.0 to 8.0), moisture content (from 0 wt% to 2 wt%) and temperature (from 25 ◦ C to 55 ◦ C) on catalytic activity were carried out. The results are shown in Fig. 2. Comparing with the untreated lipase PS, the behavior of treated lipase PS activity changed markedly at all tested pH ranges. From Fig. 2A, the maximum esterification activity of the treated lipase PS is 12,918.75 ± 0.38 U min−1 g−1 at the optimal pH 6.0. Beyond pH 6.0, the esterification activity of the treated lipase PS decreases. Specifically, the low activity at lower pH (below pH 6.0) is attributed to a change in lipase PS micro-structure, which is caused by the
Y. Liu et al. / Process Biochemistry 45 (2010) 1176–1180
1179
At 45 ◦ C, the esterification activity of the treated lipase PS ranges from 9893.75 ± 1.04 to 116,125.50 ± 0.88 U min−1 g−1 . A further increase of temperature beyond 45 ◦ C leads to protein denaturation of lipase PS, resulting in a decrease of enzyme activity [18,20]. 4. Conclusions To investigate the effect of pretreatment by different organic solvents on enzyme activity and conformational structure, eight organic solvents, such as acetone, isopropanol, n-hexane, acetonitrile, butanol, ethanol, toluene, and n-heptane, are employed to treat the immobilized lipase PS. It is demonstrated that hydrophobicity, functional group and molecular constitution of organic solvents are all crucial factors to affect esterification activity. FTIR studies show that secondary structure of the immobilized lipase PS is altered to a significant extent, which is responsible for the variance of esterification activity. It is concluded that there is a close correlation between secondary structure and esterification activity of immobilized lipase PS, which is of significance for its further application. Acknowledgements The authors are deeply indebted to the Analytical and Testing Center of Huazhong University of Science and Technology for FT-IR analysis, and the financial support from the National High Technology Research and Development Program of China (863 Program) (2009AA03Z232 and 2007AA100703), Program for New Century Excellent Talents in University (NCET-07-0336), Doctoral Education Fund for New Teachers (20090142120090), and the Natural Science Foundation of Hubei Province (2008CDB359 and 2009CDA046). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.procbio.2010.03.023.
Fig. 2. Effect of pH, moisture and temperature on the activity of the organic solvent treated immobilized lipase PS. Substrate: a mixture of 0.40 g lauric acid (0.2 M), 0.37 g dodecanol (0.2 M) in 10 mL isooctane solvent and 0.01 mL buffer solution, reaction time 30 min, temperature 37 ◦ C, shaking rate 200 rpm.
absorption of excessive H+ leading to an irreversible protein denaturation of lipase PS. The enzyme activity decreases rapidly due to a more severe deactivation of the enzyme at higher pH (above pH 6.0). In addition, pH effect can also be manifested due to partitioning of hydrogen ions between the solution and the support surface [19]. Similar trends of effect of pH on the activities were also observed in lipases from C. rugosa, C. antarctica, Rhizopus delemar [18–21]. Effect of moisture on esterification activity of the immobilized lipases had been intensively reported [7,10,21]. It is demonstrated (Fig. 2B) that the optimum moisture content is 0.1 wt% for the immobilized lipase PS after treatment with all tested organic solvents. Above 0.1 wt% moisture content, an obvious decrease of esterification activity is observed. The probable reason is attributed to a shift in the chemical equilibrium in the higher water content range. Similar observations were also reported by Petkar et al. [14] and Naoe et al. [21], who both found that Humicola lanuginosa lipase, C. antarctica lipase B and Rhizomucor miehei lipase exhibited high activities at extremely low water content, as well as lipase from R. delemar. From Fig. 2C, it is noted that the optimum temperature is 45 ◦ C for the immobilized lipase PS after treatment with organic solvents.
References [1] Liu Yun, Yan Yunjun, Zhang Xuan, Tan Hui, Xin Honglin, Yao An’na. Biodiesel production by combined lipases catalyzed transesterification: optimization and kinetics [J]. AIChE Journal, 2009, doi:10 1002/aic.12090. [2] Fjerbaek L, Christensen KV, Norddahl B. A review of the current state of biodiesel production using enzymatic transesterification. Biotechnol Bioeng 2009;102:1298–315. [3] Hara P, Hanefeld U, Kanerv LT. Immobilised Burkholderia cepacia lipase in dry organin solvents and ionic liquids: a comparison. Green Chem 2009;11:250–6. [4] Liu Yun, Tan Hui, Zhang Xuan, Yan Yunjun, Hameed BH. Effect of monohydric alcohols on enzymatic transesterification for biodiesel production. Chem Eng J 2010;157:223–9. [5] Su E, Wei D. Improvement in lipase-catalyzed methanolysis of triacylglycerols for biodiesel production using a solvent engineering method. J Mol Catal BEnzym 2008;55:118–25. [6] Priego J, Ortíz-Nava C, Carrillo-Morales M, López-Munguía A, Escalante J, Castillo E. Solvent engineering: an effective tool to direct chemoselectivity in a lipase-catalyzed michael addition. Tetrahedron 2009;65:536–9. [7] Klibanov AM. Improving enzymes by using them in organic solvents. Nature 2001;409:241–6. [8] Hiramatsu H, Kitagawa T. FT-IR approaches on amyloid fibril structure. Biochim Biophys Acta 2005;1753:100–7. [9] Natalello A, Ami D, Brocca S, Lotti M, Doglia SM. Secondary structure, conformational stability and glycosylation of a recombinant Candida rugosa lipase studied by Fourier-transform infrared spectroscopy. Biochem J 2005;385:511–7. [10] Pencreach G, Baratti JC. Activity of Pseudomonas cepacia lipase in organic media is greatly enhanced after immobilization on a polypropylene support. Appl Microbiol Biotechnol 1997;47:630–5. [11] Secundo F, Carrea G. Lipase activity and conformation in neat organic solvents. J Mol Catal B-Enzym 2002;19:93–102. [12] Secundo F, Spadaro S, Carrea G, Overbeeke PLA. Optimization of Pseudomonas cepacia lipase preparations for catalysis in organic solvents. Biotechnol Bioeng 1999;62:554–61.
1180
Y. Liu et al. / Process Biochemistry 45 (2010) 1176–1180
[13] Talukder MMR, Tamalampudy S, Li CJ, Le YL, Wu JC, Kondo A, et al. An improved method of lipase preparation incorporating both solvent treatment and immobilization onto matrix. Biochem Eng J 2007;33:60–5. [14] Petkar M, Lali A, Caimi P, Daminati M. Immobilization of lipases for nonaqueous synthesis. J Mol Catal B-Enzym 2006;39:83–90. [15] Quiroga E, Camí G, Marchese J, Barberis S. Organic solvents effect on the secondary structure of araujiain hI, in different media. Biochem Eng J 2007;35:198–202. [16] Koskinen AMP, Klibanov AM. Enzymatic reactions in organic media. London: Blackie Academic & Professional; 1996. [17] Laane C, Boeren S, Vos K, Veeger C. Rules for optimization of biocatalysis in organic solvents. Biotechnol Bioeng 1987;30:81–7.
[18] Wu JC, Song BD, Xing AH, Hayashi Y, Talukder MMR, Wang SC. Esterification reactions catalyzed by surfactant-coated Candida rugosa lipase in organic solvents. Proc Biochem 2002;37:1229–33. [19] Klibanov AM. Why are enzymes less active in organic solvents than in water. Trends Biotechnol 1997;15:97–101. [20] Abdullah AZ, Sulaiman NS, Kamaruddin AH. Biocatalytic esterification of citronellol with lauric acid by immobilized lipase on aminopropyl-grafted mesoporous SBA-15. Biochem Eng J 2009;44:263–70. [21] Naoe K, Ohsa T, Kawagoe M, Imai M. Esterification by Rhizopus delemar lipase in organic solvent using sugar ester reverse micelles. Biochem Eng J 2001;9:67–72.