Appl Biochem Biotechnol DOI 10.1007/s12010-014-1432-y
Rhizopus oryzae Lipase Immobilized on Hierarchical Mesoporous Silica Supports for Transesterification of Rice Bran Oil Prashanth Ramachandran & Guru Krupa Narayanan & Sakthivel Gandhi & Swaminathan Sethuraman & Uma Maheswari Krishnan
Received: 21 May 2014 / Accepted: 28 November 2014 # Springer Science+Business Media New York 2014
Abstract The tunable textural properties of self-oriented mesoporous silica were investigated for their suitability as enzyme immobilization matrices to support transesterification of rice bran oil. Different morphologies of mesoporous silica (rod-like, rice-like, and spherical) were synthesized and characterized by scanning electron microscopy (SEM), transmission electron microscopy (TEM), and nitrogen adsorption–desorption isotherms. The surface area, pore size, and ordered arrangement of the pores were found to influence the immobilization and activity of the enzyme in the mesopores. The immobilization in rod-like silica was highest with an immobilization efficiency of 63 % and exhibited minimal activity loss after immobilization. Functionalization of the mesoporous surface with ethyl groups further enhanced the enzyme immobilization. The free enzyme lost most of its activity at 50 °C while the immobilized enzyme showed activity even up to 60 °C. Transesterified product yield of nearly 82 % was obtained for 24 h of reaction with enzyme immobilized on ethyl-functionalized SBA-15 at an oil:methanol ratio of 1:3. Fourier transform infrared spectroscopy (FT–IR) and Gas chromatography–mass spectrometry (GC-MS) were used to characterize the transesterified product obtained. The reusability of the immobilized enzyme was studied for 3 cycles. Keywords SBA-15 . Rhizopus oryzae . Lipase . Immobilization . Mesoporous silica
Introduction Lipases are industrially important enzymes that catalyze the conversion of triacylglycerols to glycerol and fatty acid. The water-soluble lipases catalyze reactions involving water-insoluble substrates at the lipid-water interface, which has prompted investigators to seek appropriate milieu to retain the catalytic efficiency of the enzyme. Lipase-catalyzed transesterification of Prashanth Ramachandran and Guru Krupa Narayanan have equal contribution to this study.
P. Ramachandran : G. K. Narayanan : S. Gandhi : S. Sethuraman : U. M. Krishnan (*) Centre for Nanotechnology and Advanced Biomaterials, School of Chemical and Biotechnology, SASTRA University, Thanjavur 613401, India e-mail: [email protected]
Appl Biochem Biotechnol
triglycerides to fatty acid alkyl esters resulting in the formation of biodiesel is an important application that has gained momentum in the past decade. Immobilized lipases are more effective than their soluble counterparts in a number of ways. Immobilization improves the stability and reusability of enzymes. Factors such as immobilization method and the kind of support used play a major role in the stability of immobilized enzyme [1]. Hence, efforts are still on to develop an affordable immobilization matrix with high surface area and possessing adequate pores with sizes that exhibit negligible diffusional limitation of the substrate or the product [2, 3]. An important challenge in immobilizing lipase is the retention of active conformation of the enzyme during immobilization. The activation of Rhizopus oryzae lipase involves a shift from its closed to open confirmation. The catalytic site is buried under a helical segment called the lid (closed form) which rolls back at the oil-water interface thereby exposing the catalytic site (open form) [4]. The equilibrium between the closed and open conformation shifts towards the open form in the presence of a hydrophobic interface [5]. Immobilization of lipases on engineered supports could either restrict the free movement of the lid or may cause the immobilization of the enzyme in the open form itself so that it remains active. The affinity of lipase to hydrophobic supports has been investigated earlier [6], and its preference for such supports has been emphasized in literature. But in these cases, the inner pores of the support become inaccessible to the aqueous enzyme solution due to the highly hydrophobic character of the support surface [7]. Use of hydrophilic supports will be associated with the problem of clustering of enzymes and the presence of bound water molecules in the vicinity, which may be detrimental to the activity of lipase [8]. Hence, an ideal immobilization matrix is essential to realize the catalytic benefits of the enzyme. In case of lipase being immobilized on a porous solid then the enzyme inactivating factors like adsorption onto hydrophobic surface, aggregation and auto-proteolysis are mitigated to a great extent [9]. But this strategy suffers from diffusion problems (pH gradients, substrate, etc.). Reversible immobilization (non-covalent) protocols have also been suggested in cases where enzyme stabilization is not critical, and the support is reused after the enzyme is inactivated [10]. Epoxy, glutaraldehyde, glyoxal-activated supports have also been reported to increase the stability of the system and provide low steric hindrance [9, 11–13]. Cross-linked crystals and aggregates have also been extensively explored as immobilization matrices as they increase stability by arresting the mobility of the enzyme and do not require any separate support elements. However, the biocatalyst has a low mechanical resistance in these cases [14]. Mesoporous silica, especially SBA-15, with ordered pores, tunable pore sizes, and ease of surface functionalization has been a popular candidate for the immobilization of enzymes [15]. Several groups have worked on the use of mesoporous materials as matrices for immobilizing enzymes [14, 16]. One disadvantage associated with SBA15 matrices has been the leaching of enzymes from the silica support due to weak interactions between the support and the enzymes [17]. But this can be annulled by the use of suitably surface-functionalized silica, which could improve the interaction of the enzyme with the support. The present work aims to study the influence of the textural properties of mesoporous SBA-15 silica on immobilization and activity of lipase. The immobilized enzyme has been successfully employed for the transesterification of rice bran oil. The effect of surface functionalization on the enzyme loading and activity has also been investigated.
Appl Biochem Biotechnol
Materials and Methods Materials Commercially available rice bran oil was purchased from M/s. Kaleesuwari Refineries Private Limited, India. Methanol, sodium hydrogen phosphate, and sodium dihydrogen phosphate were purchased from Merck, India. Pluronic (P-123), tetra ethyl orthosilicate (TEOS), amino propyl triethoxy silane (APTES), triethoxy(ethyl)silane, cetyl trimethylammonium bromide (CTAB), tributyrin, and R. oryzae lipase in powdered form were purchased from Sigma– Aldrich, USA. All materials were used as such without further purification. Preparation of SBA-15 Three different morphologies of SBA-15 type of mesoporous silica were synthesized. These are schematically represented in Fig. 1. These were designated as rod-like, rice-like, and spherical SBA-15. Synthesis of rod-like SBA-15 was carried out using the procedure reported previously [18]. Briefly, 5 g of P-123 (tri-block copolymer) was taken in 60 mL of deionized water, and the pH was maintained below 1 by adding 2 M HCl. Nine grams of TEOS was added to the reaction mixture as silica source and stirred for 24 h followed by the aging process at 80 °C for 24 h. The white precipitate was filtered, washed with deionized water, and dried at 100 °C for 12 h under vacuum. The dried as-synthesized material was calcined at 550 °C for 6 h to remove the organic template (P-123). For obtaining rice-like SBA-15, 5 g of P-123 (tri-block copolymer) and 0.2 g NH4F were taken in 60 mL of deionized water and the pH was maintained at below 1 by adding 2 M HCl. Nine grams of TEOS was added to the reaction mixture as silica source and stirred for 24 h followed by the aging process at 80 °C for 24 h. The white precipitate was filtered, washed with deionized water, and dried at 100 °C for 12 h under vacuum. The dried as-synthesized material was calcined at 550 °C for 6 h to remove the organic template (P-123). For synthesis of spherical SBA-15, 2.5 g of P-123 (tri-block copolymer) was taken in 60 mL of deionized water and mixed with 2.5 g of CTAB dissolved in methanol. The pH was maintained below 1 by adding 2 M HCl. Nine grams of TEOS was added to the reaction mixture as silica source and stirred for 24 h followed by the aging process at 80 °C for 24 h. The white precipitate was filtered, washed with deionized water, and dried at 100 °C for 12 h under vacuum. The dried as-synthesized material was calcined at 550 °C for 6 h to remove the organic templates (P-123 & CTAB).
Fig. 1 Different morphologies of SBA-15 synthesized
Appl Biochem Biotechnol
Functionalization of SBA-15 The silica in which there was maximum immobilization was functionalized with hydrophobic and hydrophilic groups in an effort to increase the adsorption of enzyme. To make the surface of SBA-15 more hydrophilic, functionalization with amine group was carried out. Initially, 400 μL of APTES was mixed with 20 mL of toluene in which two grams of SBA-15 was added and refluxed at 80 °C for 20 h in N2 atmosphere. The precipitate was filtered, washed with acetone, and dried under vacuum. In order to introduce hydrophobic character to the mesoporous surface, functionalization with ethyl group was carried out. Four hundred microliters of triethoxy(ethyl)silane was mixed with 20 mL of toluene. Two grams of SBA-15 was added to the solution and refluxed at 95 °C for 20 h. The precipitate was filtered, washed with acetone, and dried under vacuum. Characterization of SBA-15 The mesoporous silica samples were analyzed using Fourier transform infrared spectroscopy (FT–IR). The analysis was carried out at a resolution of 4 cm−1 between 4000 and 450 cm−1 with an average of ten scans using the FT–IR spectrometer Spectrum 100, PerkinElmer, USA. The pore and surface morphology of SBA-15 were evaluated using field emission transmission electron microscopy (FE–TEM), JEM 2100F, JEOL, Japan and cold field emission scanning electron microscopy (FE– SEM), JSM 6701F, JEOL, Japan. The low-angle X-ray diffraction measurement (LA-XRD) was carried out on a D8 Focus, Bruker, Germany, over a 2θ range of 0.5 to 3° with a scan rate of 0.001°/s using CuKα radiation. The surface area, pore diameter, and pore volume were determined using N2 adsorption–desorption isotherms obtained using Accelerated Surface Area and Porosimetry analyzer (ASAP) 2020, Micromeritics, USA. Immobilization of Enzyme Forty milligrams of R. oryzae lipase was first dissolved in 100 mM phosphate buffer of pH 7.2 by stirring in a 30-mL vial using a magnetic stirrer. After obtaining a homogenous solution, 40 mg of immobilization support (rod-like, rice-like, or spherical SBA-15) was added and stirred using a magnetic stirrer for various time intervals ranging from 3 to 24 h. The solution was then centrifuged at 9610rpm for 10 min at 4 °C. The supernatant was collected, and the protein content was estimated using Lowry’s method [19]. By determining the amount of protein in the supernatant, the amount of enzyme adsorbed on to the support was calculated. The activity of the immobilized enzyme was measured using the procedure mentioned below. Activity Assay The activity of lipase was found by determining the extent of hydrolysis of tributyrin. The assay procedure involved the addition of 2 mL of tributyrin to a 20 mL of 100 mM phosphate buffer pH 7.2 containing the free or immobilized enzyme. The solution was stirred for 6 h and titrated against 0.1 N sodium hydroxide solution with phenolphthalein as an indicator. From the amount of sodium hydroxide consumed, the amount of butyric acid produced in 6 h of reaction was calculated. The activity of
Appl Biochem Biotechnol
the enzyme was represented as micromoles of butyric acid released per minute or unit. The specific activity was calculated per milligram of the enzyme immobilized in the support and was represented as U/mg enzyme immobilized. Optimal Temperature for Enzyme Activity The optimum temperature for activity of the free and immobilized enzymes was investigated for both 3 and 6 h in rod-like SBA-15. The above said assay procedure was carried out by exposing the reaction mixture to temperatures varying from 30 to 60 °C. The activity loss due to increase in temperature was found by calculating the amount of sodium hydroxide required to neutralize the butyric acid produced. Activity Loss The loss in activity during the reuse of enzyme was determined by using the tributyrin assay procedure after each use. The activity loss experiment was carried out for the optimally immobilized enzyme. Enzymatic Transesterification of Rice Bran Oil The procedure employed for the transesterification of rice bran oil is as follows. Methanol and rice bran oil were taken in 3:1 M ratio in a 30-mL glass vial. The pellet obtained after immobilization was re-suspended in 500 μL of phosphate buffer pH 7.2 and added to the reaction mixture. The mixture was stirred for 24 h. The solution was then left undisturbed to allow separation of the ester and glycerol layers. The ester layer was separated and heated. The two layers were then characterized using FT–IR and gas chromatography–mass spectrometry (GC–MS) to confirm their identity. Characterization of Transesterified Product The transesterification was confirmed by FT–IR analysis, and the spectrum was recorded in Attenuated Total Reflection (ATR) mode using Spectrum 100, PerkinElmer, USA. The data was used to confirm the long-chain, ester band in methyl esters and alcohol band in glycerol. The best sample was analyzed using GC–MS in order to confirm the conversion. GC–MS analysis was performed using Clarus 500, PerkinElmer, USA. The sample was introduced at a split ratio of 1:20 in the temperature range 50 to 300 °C in order to confirm the conversion percentage. Helium was used as carrier gas at a flow rate of 1 mL/min. Reuse of the Immobilized Enzyme The reusability of the immobilized enzyme (optimized enzyme preparation) that had been used for the transesterification reaction was to be ascertained. After the completion of the transesterification reaction, the two layers were allowed to separate, the immobilized enzyme partitioned to the glycerol layer. The glycerol layer was then centrifuged at 9610 rpm for 10 min to separate the immobilized enzyme. The pellet, obtained after centrifugation, was suspended in 1 mL phosphate buffer pH 7.2 and used again for transesterification reaction under the reaction conditions as mentioned above.
Appl Biochem Biotechnol
Results and Discussion Characterization of Supports The synthesized mesoporous silica samples were analyzed for morphological characteristics using field emission (FE)-SEM and field emission (FE)-TEM, and the images are shown in Fig. 2. The SEM micrographs confirm the formation of rod-like, rice-like, and spherical SBA-15. The TEM images reveal a highly ordered channel-like morphology for the rod-like SBA-15. The rice-like and spherical SBA-15 show a highly porous network, but no definite pore order is discernible. Figure 3 shows the FT–IR spectra of the mesoporous samples. The FT–IR spectra of all three silica samples showed characteristic vibration bands at 3459, 1086, and 800 cm−1 due to the presence of –OH, Si–O–Si, and Si–OH bonds, respectively. Pore size analysis of three mesoporous samples was done by Barrett–Joyner–Halenda (BJH) method of isotherm analysis (Fig. 4). The multipoint surface area of the mesoporous samples determined using the nitrogen adsorption–desorption isotherms (Fig. 5) was found to be 483, 518, and 743 m2/g for rod-like, rice-like, and spherical morphologies, respectively. The BJH isotherms show pore sizes of 7.35, 8.76, and 2.36 nm for rod, rice, and spherical morphologies. The pore volume analyzed at the highest relative pressure (P/P0 =1) was found to be 0.7602, 1.2032, and 0.3958 cm3/g for rod-like, rice-like, and spherical morphologies, respectively. An interesting observation in the BJH pore analysis is that the pore distribution for the rod-like SBA-15 is narrow while the rice-like SBA-15 exhibits a wide pore distribution. The pore sizes of the spherical SBA-15 are the lowest as observed from Fig. 4. The micropore area is large in the spherical SBA-15 when compared with the other two samples. It is evident that these differences in the textural properties of the mesoporous samples can influence the immobilization and catalytic efficiency of the enzyme. The surface functional groups in the immobilization supports significantly affect the binding of enzymes to the support. The surfaces of all the silica support have silanol groups (−SI–OH) as evident from the vibration frequencies observed in the FT–IR spectra (Fig. 3) which would make the surface hydrophilic. This in turn could influence the nature of interactions with the hydrophilic residues of the enzyme. The variation in the surface area of the different morphologies also direct the quantity of the enzyme immobilized in the matrix. The molecular weight of R. oryzae lipase is reported to be about 32 kDa [20]. The sizes of a majority of microbial lipases vary from around 4 to 5 nm; hence, the porosity and pore size of the immobilization support will have a major effect on its ability to load proteins as well as localization of the enzyme in the porous silica matrix [21]. Smaller pore sizes tend to localize the enzyme through weak associative forces on the outer surface while larger pores facilitate the accommodation of the enzyme inside the pore. Immobilization of Lipase Immobilization of lipase on different supports was carried out at room temperature with a support to enzyme ratio of 1:1 (weight ratio mg/mg; Table 1). The time for immobilization was varied from 3 to 24 h. For rod-like SBA-15, the immobilization was maximum when stirred for 6 h. The immobilization percentage reached 63.4 % by 6 h. The specific activity of the immobilized enzyme was decreased by 8.9 % when compared with free enzyme. When the immobilization was carried out for 3 h, the immobilization efficiency was reduced to 50 %, but the specific activity showed an increase by 47 % when compared with free enzyme. The
Appl Biochem Biotechnol
Fig. 2 Scanning electron micrographs (a, c, e) and transmission electron micrographs (b, d, f) of rod, rice, and spherical morphologies of SBA-15
decrease in activity with increased duration of stirring may be due to shear effects caused due to continuous stirring or constrainedness brought about by increased enzyme loading into the mesopores. For rice-like SBA-15, the immobilization was maximum at 77 % enzyme loading for 6 h of immobilization. The increased immobilization may be attributed to the greater pore size and higher surface area of the mesoporous sample. However, it was found that the enzyme activity
Appl Biochem Biotechnol
Fig. 3 FT–IR spectra of a rod-like, b rice-like, and c spherical SBA-15
was considerably reduced in spite of exhibiting higher immobilization. This may be due to the diffusional limitation of the support owing to the disordered arrangement of pores in the ricelike SBA-15 sample. The specific activity of the rice-like SBA-15 was between 0.2 and 0.4 U/ mg when compared to the free enzyme which exhibited a specific activity of 0.64 U/mg. When spherical SBA-15 was used for immobilization, the immobilization efficiency reached a maximum of 56 % when the immobilization procedure was carried out for 24 h. However, reproducibility in immobilization was not achieved which may be attributed to the non-uniformity of the particle sizes. The pore size of the spherical particles was found to be 2.1 nm. This may have also contributed to the retardation of the enzyme immobilization inside the pores and hence the low loading of the enzyme in to the support. It is also expected that most of the enzyme molecules are more likely held by weak interactive forces at the surface thereby facilitating rapid association and dissociation of the enzyme molecule from the surface. Based on the enzyme loading and enzyme activity data, the rod-like SBA-15 was found to be most conducive for lipase immobilization and hence was chosen for further trials. Characterization of Functionalized SBA-15 The rod-like SBA-15 was functionalized with ethyl and amine groups to introduce hydrophobic and hydrophilic character to the support, respectively. The functionalization was confirmed by FT–IR. Figure 6a, b shows the FT–IR spectra of the ethyl and amine-functionalized SBA15. Apart from the characteristic band between 1080 and 1100 cm−1 due to the −Si–O–Sistretching, the FT−IR spectrum of the ethyl-functionalized SBA-15 shows the appearance of a new band at 2926 cm−1 indicating the presence of −CH2− stretching which arises due to functionalization with ethyl group. In the case of amine-functionalized SBA-15, apart from the −Si–O–Si band at 1086 cm−1, the presence of a shoulder in the band at 3403 cm−1 indicates
Appl Biochem Biotechnol
Fig. 4 BJH isotherm spectra of a rod-like, b rice-like, and c spherical SBA-15
Appl Biochem Biotechnol Fig. 5 BET adsorption–desorption spectra of a rod-like, b ricelike, and c spherical SBA-15
Appl Biochem Biotechnol Table 1 Immobilization of lipase and its activity Sample
Rod-shaped SBA-15
Rice-shaped SBA-15
Spherical SBA-15
Time for immobilization (h)
Enzyme immobilized (%)
Activity (U)
Specific activity (U/mg)
24
62.90
14.80
0.58
12
55.75
14.00
0.63
6
63.37
14.75
0.58
3
49.43
24.00
1.21
24
77.35
8.34
0.26
12
64.00
4.61
0.17
6
77.11
7.86
0.25
3 24
54.25 50.00
8.80 8.75
0.41 0.39
12
39.50
12.40
0.78
6
34.80
11.80
0.84
3
41.10
10.47
0.63
presence of primary amine group. In addition, the band at 1559 cm−1 corresponding to N−H bending confirms the presence of amine group in the sample. The band at 2817 cm−1 may be attributed to −CH2− stretching because of the presence of propyl group which confirms APTES functionalization. Immobilization on Functionalized Supports Functionalization of silica surface with ethyl and amine groups imparts a hydrophobic and hydrophilic character to the silica support. This influences the hydrophobic and hydrophilic interactions between the support and the enzyme. The functionalized SBA-15 and enzyme were taken in weight ratios of 1:1 for immobilization. It was found that when immobilization
Fig. 6 FT–IR spectra of ethyl-functionalized (a) and amine-functionalized (b) SBA-15
Appl Biochem Biotechnol
was carried out on amine-functionalized silica, the amount of enzyme loaded onto the silica support was very less (Table 2). The immobilization percentage varied from 20.7 to 27.6 % when the time for immobilization was varied between 3 and 24 h. But it was found that the activity of the immobilized enzyme was equal to that of the free enzyme. When ethylfunctionalized silica was used for immobilization, the enzyme loading was high ranging from 0.67 to 0.7 mg enzyme/mg of support. This result is in agreement with earlier reports that have suggested a hydrophobic environment as the most suited for lipase immobilization [22]. The activity of the enzyme was found to remain intact after immobilization. Optimum Temperature for the Activity of Immobilized and Free Enzyme The optimum temperature of the immobilized and free enzyme was investigated and compared (Fig. 7). Lipase immobilized for 3 and 6 h in rod-like SBA-15 was taken for the studies. The activity of the immobilized and free enzyme was found by calculating the rate of hydrolysis of tributyrin at various temperatures. An incubation time of 6 h was provided to identify the effect of prolonged exposure to various temperatures on the enzyme. The enzyme immobilized in rod-like SBA-15 exhibited the maximum activity, and hence, it was chosen for this study. The optimum temperature for immobilized and free enzyme was found to be 30 °C. The specific activity of free enzyme reached 0.02 when exposed to a temperature of 50 °C and reduced to 0.00056 when exposed to a temperature of 60 °C. While for the enzyme which was immobilized for 6 h, the specific activity was 1.52 at 30 °C and reduced to 1.078 at 60 °C. The specific activity reduced by 29 % when the temperature was increased from 30 to 60 °C. This shows that the activity of the immobilized enzyme was higher at 60 °C when compared to that of free enzyme at the same temperature. The enzyme which was immobilized for 3 h showed similar denaturation pattern to that of the free enzyme. This indicates that when the immobilization procedure was carried out for 3 h, the enzyme gets adsorbed only on the outer surface and hence is not protected by the silica matrix. The higher temperature for immobilized enzyme can be attributed to the rigid silica framework that surrounds the enzyme which prevents the enzyme from undergoing changes and denaturation at higher temperatures. Activity Loss The loss in activity of immobilized enzyme was measured after each cycle. The activity in the first cycle was found to be 23.8 U. The enzyme activity in the subsequent cycles was found to decrease to 21 and 18.3 U. This may be due to desorption of some enzyme molecules from the mesopores, during repeated cycles. Table 2 Enzyme loading and activity on functionalized SBA-15 Functionalization Duration for immobilization (h)
Enzyme immobilized (%)
Activity of immobilized enzyme (U)
Specific activity of immobilized enzyme (U/mg)
Amine
20.7
24.014
2.95
26.5
6.72
3 6
Ethyl
9.88
24
27.6
21.542
2.08
3
67.4
24.458
0.90
6
50.0
21.944
1.11
24
70.2
17.778
0.63
Appl Biochem Biotechnol
Fig. 7 Represents the thermal stability of free and immobilized lipases
Enzymatic Transesterification of Rice Bran Oil For the transesterification reaction, rice bran oil and methanol were taken in ratios 1:3, 1:6, and 1:9. The ratio of 1:3, rice bran oil:methanol, was found to be optimum. The immobilized enzyme in rod-like SBA-15 was suspended in 2 mL of phosphate buffer pH 7.2 and added to the oil–methanol mixture and stirred at room temperature. The mixture was subjected to a reaction time of 24 h. After the reaction, the ester and glycerol layers were allowed to settle and the transesterified product layer separated. Lipase immobilized for 3 h, 6 h on rod-like SBA15, and for 3 h in ethyl-functionalized rod-like SBA-15 were used for the transesterification reaction. The yields obtained were 79.8, 79.6, and 81.7 %, respectively. The transesterified sample was heated to remove the glycerol and water contamination. The pure sample was analyzed using FT–IR. The prominent peak at 1742 cm−1 confirms the
Table 3 Composition of transesterified oil (biodiesel) obtained using Rhizopus oryzae lipase Constituent in transesterified oil fraction Octanoic acid, methyl ester
Peak area in GC–MS (%) 0.0255
5-Octadecenoic acid, methyl ester
0.0253
Nonanoic acid, 9-oxo-, methyl ester
0.0139
Tetradecanoic acid, methyl ester
0.0051
Pentadecanoic acid, methyl ester
0.0519
9-Hexadecenoic acid, methyl ester, (Z)-
0.3932
Hexadecanoic acid, methyl ester 9-Octadecenoic acid, methyl ester
19.3842 68.3299
Octadecanoic acid, methyl ester
3.0793
9,12-Octadecadienoic acid (Z,Z)-, methyl ester
0.1225
11-Eicosenoic acid, methyl ester
0.8863
Eicosanoic acid, methyl ester
0.7960
Docosanoic acid, methyl ester
1.1853
Appl Biochem Biotechnol
Fig. 8 GC–MS profile of the transesterified sample obtained by employing 3 h (immobilized) ethylfunctionalized rod-like SBA-15 for the transesterification reaction
presence of ester group in the sample. The peaks at 2922 and 2853 cm−1 confirm the presence of long-chain alkyl groups. The best sample obtained was subjected to GC–MS analysis in order to check for the presence of methyl esters, thus confirming the occurrence of transesterification reaction. It was found that the fatty acids that are present in the oil sample have been converted to their corresponding methyl esters (Table 3 and Fig. 8). Reuse of the Catalyst The reusability of immobilized enzyme increases its commercial viability. When lipase immobilized for 3 h was reused, the conversion efficiency dropped to 67.7 % after the three runs.
Conclusion Mesoporous SBA-15 with different morphology and textural properties has been successfully synthesized, characterized, and used as immobilization matrices for lipase. Rod-like morphology with highly ordered pores in the range 7–8 nm was found to exhibit high enzyme immobilization without compromising the enzyme activity. Functionalization with hydrophobic ethyl groups increased the enzyme immobilization without loss in enzyme activity. The immobilized lipase exhibited good transesterification efficiency and reusability up to 3 cycles. Our data suggests that mesoporous framework holds much promise as an immobilization matrix for lipase and for commercial biodiesel production. Acknowledgments The authors wish to acknowledge the characterization facilities established from the PG Teaching fund of the Nanomission council, Department of Science and Technology, and infrastructure support from SASTRA University.
References 1. Ye, P., Jiang, J., & Xu, Z. K. (2007). Adsorption and activity of lipase from Candida rugosa on the chitosanmodified poly(acrylonitrile-co-maleic acid) membrane surface. Colloids and Surfaces B: Biointerfaces, 60, 62–67. 2. Dizge, N., Aydiner, C., Imer, D. Y., Bayramoglu, M., Tanriseven, A., & Keskinler, B. (2009). Biodiesel production from sunflower, soybean, and waste cooking oils by transesterification using lipase immobilized onto a novel microporous polymer. Bioresource Technology, 100, 1983–1991.
Appl Biochem Biotechnol 3. Prlainovic, N. Z., Knezevic-Jugovic, Z. D., Mijin, D. Z., & Bezbradica, D. I. (2011). Immobilization of lipase from Candida rugosa on Sepabeads((R)): the effect of lipase oxidation by periodates. Bioprocess and Biosystems Engineering, 34, 803–810. 4. Bordes, F., Barbe, S., Escalier, P., Mourey, L., Andre, I., Marty, A., & Tranier, S. (2010). Exploring the conformational states and rearrangements of Yarrowia lipolytica Lipase. Biophysical Journal, 99, 2225– 2234. 5. Piamtongkam, R., Duquesne, S., Bordes, F., Barbe, S., Andre, I., Marty, A., & Chulalaksananukul, W. (2011). Enantioselectivity of Candida rugosa lipases (Lip1, Lip3, and Lip4) towards 2-bromo phenylacetic acid octyl esters controlled by a single amino acid. Biotechnology and Bioengineering, 108, 1749–1756. 6. Gao, S., Wang, Y., Diao, X., Luo, G., & Dai, Y. (2010). Effect of pore diameter and cross-linking method on the immobilization efficiency of Candida rugosa lipase in SBA-15. Bioresource Technology, 101, 3830– 3837. 7. Yin, P., Chen, W., Liu, W., Chen, H., Qu, R., Liu, X., Tang, Q., & Xu, Q. (2013). Efficient bifunctional catalyst lipase/organophosphonic acid-functionalized silica for biodiesel synthesis by esterification of oleic acid with ethanol. Bioresource Technology, 140, 146–151. 8. Grasset, L., Cordier, D., & Ville, A. (1977). Woven silk as a carrier for the immobilization of enzymes. Biotechnology and Bioengineering, 19, 611–618. 9. Mateo, C., Grazu, V., Pessela, B. C., Montes, T., Palomo, J. M., Torres, R., Lopez-Gallego, F., FernandezLafuente, R., & Guisan, J. M. (2007). Advances in the design of new epoxy supports for enzyme immobilization-stabilization. Biochemical Society Transactions, 35, 1593–1601. 10. Bayramoglu, G., Altintas, B., & Arica, M. Y. (2011). Reversible immobilization of uricase on conductive polyaniline brushes grafted on polyacrylonitrile film. Bioprocess and Biosystems Engineering, 34, 127–134. 11. Ducker, R. E., Montague, M. T., & Leggett, G. J. (2008). A comparative investigation of methods for protein immobilization on self-assembled monolayers using glutaraldehyde, carbodiimide, and anhydride reagents. Biointerphases, 3, 59–65. 12. Daglioglu, C., & Zihnioglu, F. (2012). Covalent immobilization of trypsin on glutaraldehyde-activated silica for protein fragmentation. Artificial Cells, Blood Substitutes, and Immobilization Biotechnology, 40, 378– 384. 13. Ferrarotti, S. A., Bolivar, J. M., Mateo, C., Wilson, L., Guisan, J. M., & Fernandez-Lafuente, R. (2006). Immobilization and stabilization of a cyclodextrin glycosyltransferase by covalent attachment on highly activated glyoxyl-agarose supports. Biotechnology Progress, 22, 1140–1145. 14. Vinoba, M., Bhagiyalakshmi, M., Jeong, S. K., Yoon, Y. I., & Nam, S. C. (2012). Immobilization of carbonic anhydrase on spherical SBA-15 for hydration and sequestration of CO2. Colloids and Surfaces B: Biointerfaces, 90, 91–96. 15. Li, S., Wu, Z., Lu, M., Wang, Z., & Li, Z. (2013). Improvement of the enzyme performance of trypsin via adsorption in mesoporous silica SBA-15: hydrolysis of BAPNA. Molecules, 18, 1138–1149. 16. Wan, M. M., Lin, W. G., Gao, L., Gu, H. C., & Zhu, J. H. (2012). Promoting immobilization and catalytic activity of horseradish peroxidase on mesoporous silica through template micelles. Journal of Colloid and Interface Science, 377, 497–503. 17. Magner, E. (2013). Immobilisation of enzymes on mesoporous silicate materials. Chemical Society Reviews, 42, 6213–6222. 18. Gandhi, S., Sethraman, S., & Krishnan, U. M. (2010). Influence of polyhydric solvents on the catalytic and adsorption properties of self-oriented mesoporous SBA-15 silica. Journal of Porous Materials, 18, 329–336. 19. Lowry, O. H., Rosebrough, N. J., Farr, A. L., & Randall, R. J. (1951). Protein measurement with the Folin phenol reagent. Journal of Biological Chemistry, 193, 265–275. 20. Hiol, A., Jonzo, M. D., Rugani, N., Druet, D., Sarda, L., & Comeau, L. C. (2000). Purification and characterization of an extracellular lipase from a thermophilic Rhizopus oryzae strain isolated from palm fruit. Enzyme and Microbial Technology, 26, 421–430. 21. Bayramoglu, G., Karagoz, B., Altintas, B., Arica, M. Y., & Bicak, N. (2011). Poly(styrene-divinylbenzene) beads surface functionalized with di-block polymer grafting and multi-modal ligand attachment: performance of reversibly immobilized lipase in ester synthesis. Bioprocess and Biosystems Engineering, 34, 735–746. 22. Balcao, V. M., & Malcata, F. X. (1998). On the performance of a hollow-fiber bioreactor for acidolysis catalyzed by immobilized lipase. Biotechnology and Bioengineering, 60, 114–123.
Rhizopus oryzae Lipase Immobilized on Hierarchical Mesoporous Silica Supports for Transesterification of Rice Bran Oil Prashanth Ramachandran & Guru Krupa Narayanan & Sakthivel Gandhi & Swaminathan Sethuraman & Uma Maheswari Krishnan
Received: 21 May 2014 / Accepted: 28 November 2014 # Springer Science+Business Media New York 2014
Abstract The tunable textural properties of self-oriented mesoporous silica were investigated for their suitability as enzyme immobilization matrices to support transesterification of rice bran oil. Different morphologies of mesoporous silica (rod-like, rice-like, and spherical) were synthesized and characterized by scanning electron microscopy (SEM), transmission electron microscopy (TEM), and nitrogen adsorption–desorption isotherms. The surface area, pore size, and ordered arrangement of the pores were found to influence the immobilization and activity of the enzyme in the mesopores. The immobilization in rod-like silica was highest with an immobilization efficiency of 63 % and exhibited minimal activity loss after immobilization. Functionalization of the mesoporous surface with ethyl groups further enhanced the enzyme immobilization. The free enzyme lost most of its activity at 50 °C while the immobilized enzyme showed activity even up to 60 °C. Transesterified product yield of nearly 82 % was obtained for 24 h of reaction with enzyme immobilized on ethyl-functionalized SBA-15 at an oil:methanol ratio of 1:3. Fourier transform infrared spectroscopy (FT–IR) and Gas chromatography–mass spectrometry (GC-MS) were used to characterize the transesterified product obtained. The reusability of the immobilized enzyme was studied for 3 cycles. Keywords SBA-15 . Rhizopus oryzae . Lipase . Immobilization . Mesoporous silica
Introduction Lipases are industrially important enzymes that catalyze the conversion of triacylglycerols to glycerol and fatty acid. The water-soluble lipases catalyze reactions involving water-insoluble substrates at the lipid-water interface, which has prompted investigators to seek appropriate milieu to retain the catalytic efficiency of the enzyme. Lipase-catalyzed transesterification of Prashanth Ramachandran and Guru Krupa Narayanan have equal contribution to this study.
P. Ramachandran : G. K. Narayanan : S. Gandhi : S. Sethuraman : U. M. Krishnan (*) Centre for Nanotechnology and Advanced Biomaterials, School of Chemical and Biotechnology, SASTRA University, Thanjavur 613401, India e-mail: [email protected]
Appl Biochem Biotechnol
triglycerides to fatty acid alkyl esters resulting in the formation of biodiesel is an important application that has gained momentum in the past decade. Immobilized lipases are more effective than their soluble counterparts in a number of ways. Immobilization improves the stability and reusability of enzymes. Factors such as immobilization method and the kind of support used play a major role in the stability of immobilized enzyme [1]. Hence, efforts are still on to develop an affordable immobilization matrix with high surface area and possessing adequate pores with sizes that exhibit negligible diffusional limitation of the substrate or the product [2, 3]. An important challenge in immobilizing lipase is the retention of active conformation of the enzyme during immobilization. The activation of Rhizopus oryzae lipase involves a shift from its closed to open confirmation. The catalytic site is buried under a helical segment called the lid (closed form) which rolls back at the oil-water interface thereby exposing the catalytic site (open form) [4]. The equilibrium between the closed and open conformation shifts towards the open form in the presence of a hydrophobic interface [5]. Immobilization of lipases on engineered supports could either restrict the free movement of the lid or may cause the immobilization of the enzyme in the open form itself so that it remains active. The affinity of lipase to hydrophobic supports has been investigated earlier [6], and its preference for such supports has been emphasized in literature. But in these cases, the inner pores of the support become inaccessible to the aqueous enzyme solution due to the highly hydrophobic character of the support surface [7]. Use of hydrophilic supports will be associated with the problem of clustering of enzymes and the presence of bound water molecules in the vicinity, which may be detrimental to the activity of lipase [8]. Hence, an ideal immobilization matrix is essential to realize the catalytic benefits of the enzyme. In case of lipase being immobilized on a porous solid then the enzyme inactivating factors like adsorption onto hydrophobic surface, aggregation and auto-proteolysis are mitigated to a great extent [9]. But this strategy suffers from diffusion problems (pH gradients, substrate, etc.). Reversible immobilization (non-covalent) protocols have also been suggested in cases where enzyme stabilization is not critical, and the support is reused after the enzyme is inactivated [10]. Epoxy, glutaraldehyde, glyoxal-activated supports have also been reported to increase the stability of the system and provide low steric hindrance [9, 11–13]. Cross-linked crystals and aggregates have also been extensively explored as immobilization matrices as they increase stability by arresting the mobility of the enzyme and do not require any separate support elements. However, the biocatalyst has a low mechanical resistance in these cases [14]. Mesoporous silica, especially SBA-15, with ordered pores, tunable pore sizes, and ease of surface functionalization has been a popular candidate for the immobilization of enzymes [15]. Several groups have worked on the use of mesoporous materials as matrices for immobilizing enzymes [14, 16]. One disadvantage associated with SBA15 matrices has been the leaching of enzymes from the silica support due to weak interactions between the support and the enzymes [17]. But this can be annulled by the use of suitably surface-functionalized silica, which could improve the interaction of the enzyme with the support. The present work aims to study the influence of the textural properties of mesoporous SBA-15 silica on immobilization and activity of lipase. The immobilized enzyme has been successfully employed for the transesterification of rice bran oil. The effect of surface functionalization on the enzyme loading and activity has also been investigated.
Appl Biochem Biotechnol
Materials and Methods Materials Commercially available rice bran oil was purchased from M/s. Kaleesuwari Refineries Private Limited, India. Methanol, sodium hydrogen phosphate, and sodium dihydrogen phosphate were purchased from Merck, India. Pluronic (P-123), tetra ethyl orthosilicate (TEOS), amino propyl triethoxy silane (APTES), triethoxy(ethyl)silane, cetyl trimethylammonium bromide (CTAB), tributyrin, and R. oryzae lipase in powdered form were purchased from Sigma– Aldrich, USA. All materials were used as such without further purification. Preparation of SBA-15 Three different morphologies of SBA-15 type of mesoporous silica were synthesized. These are schematically represented in Fig. 1. These were designated as rod-like, rice-like, and spherical SBA-15. Synthesis of rod-like SBA-15 was carried out using the procedure reported previously [18]. Briefly, 5 g of P-123 (tri-block copolymer) was taken in 60 mL of deionized water, and the pH was maintained below 1 by adding 2 M HCl. Nine grams of TEOS was added to the reaction mixture as silica source and stirred for 24 h followed by the aging process at 80 °C for 24 h. The white precipitate was filtered, washed with deionized water, and dried at 100 °C for 12 h under vacuum. The dried as-synthesized material was calcined at 550 °C for 6 h to remove the organic template (P-123). For obtaining rice-like SBA-15, 5 g of P-123 (tri-block copolymer) and 0.2 g NH4F were taken in 60 mL of deionized water and the pH was maintained at below 1 by adding 2 M HCl. Nine grams of TEOS was added to the reaction mixture as silica source and stirred for 24 h followed by the aging process at 80 °C for 24 h. The white precipitate was filtered, washed with deionized water, and dried at 100 °C for 12 h under vacuum. The dried as-synthesized material was calcined at 550 °C for 6 h to remove the organic template (P-123). For synthesis of spherical SBA-15, 2.5 g of P-123 (tri-block copolymer) was taken in 60 mL of deionized water and mixed with 2.5 g of CTAB dissolved in methanol. The pH was maintained below 1 by adding 2 M HCl. Nine grams of TEOS was added to the reaction mixture as silica source and stirred for 24 h followed by the aging process at 80 °C for 24 h. The white precipitate was filtered, washed with deionized water, and dried at 100 °C for 12 h under vacuum. The dried as-synthesized material was calcined at 550 °C for 6 h to remove the organic templates (P-123 & CTAB).
Fig. 1 Different morphologies of SBA-15 synthesized
Appl Biochem Biotechnol
Functionalization of SBA-15 The silica in which there was maximum immobilization was functionalized with hydrophobic and hydrophilic groups in an effort to increase the adsorption of enzyme. To make the surface of SBA-15 more hydrophilic, functionalization with amine group was carried out. Initially, 400 μL of APTES was mixed with 20 mL of toluene in which two grams of SBA-15 was added and refluxed at 80 °C for 20 h in N2 atmosphere. The precipitate was filtered, washed with acetone, and dried under vacuum. In order to introduce hydrophobic character to the mesoporous surface, functionalization with ethyl group was carried out. Four hundred microliters of triethoxy(ethyl)silane was mixed with 20 mL of toluene. Two grams of SBA-15 was added to the solution and refluxed at 95 °C for 20 h. The precipitate was filtered, washed with acetone, and dried under vacuum. Characterization of SBA-15 The mesoporous silica samples were analyzed using Fourier transform infrared spectroscopy (FT–IR). The analysis was carried out at a resolution of 4 cm−1 between 4000 and 450 cm−1 with an average of ten scans using the FT–IR spectrometer Spectrum 100, PerkinElmer, USA. The pore and surface morphology of SBA-15 were evaluated using field emission transmission electron microscopy (FE–TEM), JEM 2100F, JEOL, Japan and cold field emission scanning electron microscopy (FE– SEM), JSM 6701F, JEOL, Japan. The low-angle X-ray diffraction measurement (LA-XRD) was carried out on a D8 Focus, Bruker, Germany, over a 2θ range of 0.5 to 3° with a scan rate of 0.001°/s using CuKα radiation. The surface area, pore diameter, and pore volume were determined using N2 adsorption–desorption isotherms obtained using Accelerated Surface Area and Porosimetry analyzer (ASAP) 2020, Micromeritics, USA. Immobilization of Enzyme Forty milligrams of R. oryzae lipase was first dissolved in 100 mM phosphate buffer of pH 7.2 by stirring in a 30-mL vial using a magnetic stirrer. After obtaining a homogenous solution, 40 mg of immobilization support (rod-like, rice-like, or spherical SBA-15) was added and stirred using a magnetic stirrer for various time intervals ranging from 3 to 24 h. The solution was then centrifuged at 9610rpm for 10 min at 4 °C. The supernatant was collected, and the protein content was estimated using Lowry’s method [19]. By determining the amount of protein in the supernatant, the amount of enzyme adsorbed on to the support was calculated. The activity of the immobilized enzyme was measured using the procedure mentioned below. Activity Assay The activity of lipase was found by determining the extent of hydrolysis of tributyrin. The assay procedure involved the addition of 2 mL of tributyrin to a 20 mL of 100 mM phosphate buffer pH 7.2 containing the free or immobilized enzyme. The solution was stirred for 6 h and titrated against 0.1 N sodium hydroxide solution with phenolphthalein as an indicator. From the amount of sodium hydroxide consumed, the amount of butyric acid produced in 6 h of reaction was calculated. The activity of
Appl Biochem Biotechnol
the enzyme was represented as micromoles of butyric acid released per minute or unit. The specific activity was calculated per milligram of the enzyme immobilized in the support and was represented as U/mg enzyme immobilized. Optimal Temperature for Enzyme Activity The optimum temperature for activity of the free and immobilized enzymes was investigated for both 3 and 6 h in rod-like SBA-15. The above said assay procedure was carried out by exposing the reaction mixture to temperatures varying from 30 to 60 °C. The activity loss due to increase in temperature was found by calculating the amount of sodium hydroxide required to neutralize the butyric acid produced. Activity Loss The loss in activity during the reuse of enzyme was determined by using the tributyrin assay procedure after each use. The activity loss experiment was carried out for the optimally immobilized enzyme. Enzymatic Transesterification of Rice Bran Oil The procedure employed for the transesterification of rice bran oil is as follows. Methanol and rice bran oil were taken in 3:1 M ratio in a 30-mL glass vial. The pellet obtained after immobilization was re-suspended in 500 μL of phosphate buffer pH 7.2 and added to the reaction mixture. The mixture was stirred for 24 h. The solution was then left undisturbed to allow separation of the ester and glycerol layers. The ester layer was separated and heated. The two layers were then characterized using FT–IR and gas chromatography–mass spectrometry (GC–MS) to confirm their identity. Characterization of Transesterified Product The transesterification was confirmed by FT–IR analysis, and the spectrum was recorded in Attenuated Total Reflection (ATR) mode using Spectrum 100, PerkinElmer, USA. The data was used to confirm the long-chain, ester band in methyl esters and alcohol band in glycerol. The best sample was analyzed using GC–MS in order to confirm the conversion. GC–MS analysis was performed using Clarus 500, PerkinElmer, USA. The sample was introduced at a split ratio of 1:20 in the temperature range 50 to 300 °C in order to confirm the conversion percentage. Helium was used as carrier gas at a flow rate of 1 mL/min. Reuse of the Immobilized Enzyme The reusability of the immobilized enzyme (optimized enzyme preparation) that had been used for the transesterification reaction was to be ascertained. After the completion of the transesterification reaction, the two layers were allowed to separate, the immobilized enzyme partitioned to the glycerol layer. The glycerol layer was then centrifuged at 9610 rpm for 10 min to separate the immobilized enzyme. The pellet, obtained after centrifugation, was suspended in 1 mL phosphate buffer pH 7.2 and used again for transesterification reaction under the reaction conditions as mentioned above.
Appl Biochem Biotechnol
Results and Discussion Characterization of Supports The synthesized mesoporous silica samples were analyzed for morphological characteristics using field emission (FE)-SEM and field emission (FE)-TEM, and the images are shown in Fig. 2. The SEM micrographs confirm the formation of rod-like, rice-like, and spherical SBA-15. The TEM images reveal a highly ordered channel-like morphology for the rod-like SBA-15. The rice-like and spherical SBA-15 show a highly porous network, but no definite pore order is discernible. Figure 3 shows the FT–IR spectra of the mesoporous samples. The FT–IR spectra of all three silica samples showed characteristic vibration bands at 3459, 1086, and 800 cm−1 due to the presence of –OH, Si–O–Si, and Si–OH bonds, respectively. Pore size analysis of three mesoporous samples was done by Barrett–Joyner–Halenda (BJH) method of isotherm analysis (Fig. 4). The multipoint surface area of the mesoporous samples determined using the nitrogen adsorption–desorption isotherms (Fig. 5) was found to be 483, 518, and 743 m2/g for rod-like, rice-like, and spherical morphologies, respectively. The BJH isotherms show pore sizes of 7.35, 8.76, and 2.36 nm for rod, rice, and spherical morphologies. The pore volume analyzed at the highest relative pressure (P/P0 =1) was found to be 0.7602, 1.2032, and 0.3958 cm3/g for rod-like, rice-like, and spherical morphologies, respectively. An interesting observation in the BJH pore analysis is that the pore distribution for the rod-like SBA-15 is narrow while the rice-like SBA-15 exhibits a wide pore distribution. The pore sizes of the spherical SBA-15 are the lowest as observed from Fig. 4. The micropore area is large in the spherical SBA-15 when compared with the other two samples. It is evident that these differences in the textural properties of the mesoporous samples can influence the immobilization and catalytic efficiency of the enzyme. The surface functional groups in the immobilization supports significantly affect the binding of enzymes to the support. The surfaces of all the silica support have silanol groups (−SI–OH) as evident from the vibration frequencies observed in the FT–IR spectra (Fig. 3) which would make the surface hydrophilic. This in turn could influence the nature of interactions with the hydrophilic residues of the enzyme. The variation in the surface area of the different morphologies also direct the quantity of the enzyme immobilized in the matrix. The molecular weight of R. oryzae lipase is reported to be about 32 kDa [20]. The sizes of a majority of microbial lipases vary from around 4 to 5 nm; hence, the porosity and pore size of the immobilization support will have a major effect on its ability to load proteins as well as localization of the enzyme in the porous silica matrix [21]. Smaller pore sizes tend to localize the enzyme through weak associative forces on the outer surface while larger pores facilitate the accommodation of the enzyme inside the pore. Immobilization of Lipase Immobilization of lipase on different supports was carried out at room temperature with a support to enzyme ratio of 1:1 (weight ratio mg/mg; Table 1). The time for immobilization was varied from 3 to 24 h. For rod-like SBA-15, the immobilization was maximum when stirred for 6 h. The immobilization percentage reached 63.4 % by 6 h. The specific activity of the immobilized enzyme was decreased by 8.9 % when compared with free enzyme. When the immobilization was carried out for 3 h, the immobilization efficiency was reduced to 50 %, but the specific activity showed an increase by 47 % when compared with free enzyme. The
Appl Biochem Biotechnol
Fig. 2 Scanning electron micrographs (a, c, e) and transmission electron micrographs (b, d, f) of rod, rice, and spherical morphologies of SBA-15
decrease in activity with increased duration of stirring may be due to shear effects caused due to continuous stirring or constrainedness brought about by increased enzyme loading into the mesopores. For rice-like SBA-15, the immobilization was maximum at 77 % enzyme loading for 6 h of immobilization. The increased immobilization may be attributed to the greater pore size and higher surface area of the mesoporous sample. However, it was found that the enzyme activity
Appl Biochem Biotechnol
Fig. 3 FT–IR spectra of a rod-like, b rice-like, and c spherical SBA-15
was considerably reduced in spite of exhibiting higher immobilization. This may be due to the diffusional limitation of the support owing to the disordered arrangement of pores in the ricelike SBA-15 sample. The specific activity of the rice-like SBA-15 was between 0.2 and 0.4 U/ mg when compared to the free enzyme which exhibited a specific activity of 0.64 U/mg. When spherical SBA-15 was used for immobilization, the immobilization efficiency reached a maximum of 56 % when the immobilization procedure was carried out for 24 h. However, reproducibility in immobilization was not achieved which may be attributed to the non-uniformity of the particle sizes. The pore size of the spherical particles was found to be 2.1 nm. This may have also contributed to the retardation of the enzyme immobilization inside the pores and hence the low loading of the enzyme in to the support. It is also expected that most of the enzyme molecules are more likely held by weak interactive forces at the surface thereby facilitating rapid association and dissociation of the enzyme molecule from the surface. Based on the enzyme loading and enzyme activity data, the rod-like SBA-15 was found to be most conducive for lipase immobilization and hence was chosen for further trials. Characterization of Functionalized SBA-15 The rod-like SBA-15 was functionalized with ethyl and amine groups to introduce hydrophobic and hydrophilic character to the support, respectively. The functionalization was confirmed by FT–IR. Figure 6a, b shows the FT–IR spectra of the ethyl and amine-functionalized SBA15. Apart from the characteristic band between 1080 and 1100 cm−1 due to the −Si–O–Sistretching, the FT−IR spectrum of the ethyl-functionalized SBA-15 shows the appearance of a new band at 2926 cm−1 indicating the presence of −CH2− stretching which arises due to functionalization with ethyl group. In the case of amine-functionalized SBA-15, apart from the −Si–O–Si band at 1086 cm−1, the presence of a shoulder in the band at 3403 cm−1 indicates
Appl Biochem Biotechnol
Fig. 4 BJH isotherm spectra of a rod-like, b rice-like, and c spherical SBA-15
Appl Biochem Biotechnol Fig. 5 BET adsorption–desorption spectra of a rod-like, b ricelike, and c spherical SBA-15
Appl Biochem Biotechnol Table 1 Immobilization of lipase and its activity Sample
Rod-shaped SBA-15
Rice-shaped SBA-15
Spherical SBA-15
Time for immobilization (h)
Enzyme immobilized (%)
Activity (U)
Specific activity (U/mg)
24
62.90
14.80
0.58
12
55.75
14.00
0.63
6
63.37
14.75
0.58
3
49.43
24.00
1.21
24
77.35
8.34
0.26
12
64.00
4.61
0.17
6
77.11
7.86
0.25
3 24
54.25 50.00
8.80 8.75
0.41 0.39
12
39.50
12.40
0.78
6
34.80
11.80
0.84
3
41.10
10.47
0.63
presence of primary amine group. In addition, the band at 1559 cm−1 corresponding to N−H bending confirms the presence of amine group in the sample. The band at 2817 cm−1 may be attributed to −CH2− stretching because of the presence of propyl group which confirms APTES functionalization. Immobilization on Functionalized Supports Functionalization of silica surface with ethyl and amine groups imparts a hydrophobic and hydrophilic character to the silica support. This influences the hydrophobic and hydrophilic interactions between the support and the enzyme. The functionalized SBA-15 and enzyme were taken in weight ratios of 1:1 for immobilization. It was found that when immobilization
Fig. 6 FT–IR spectra of ethyl-functionalized (a) and amine-functionalized (b) SBA-15
Appl Biochem Biotechnol
was carried out on amine-functionalized silica, the amount of enzyme loaded onto the silica support was very less (Table 2). The immobilization percentage varied from 20.7 to 27.6 % when the time for immobilization was varied between 3 and 24 h. But it was found that the activity of the immobilized enzyme was equal to that of the free enzyme. When ethylfunctionalized silica was used for immobilization, the enzyme loading was high ranging from 0.67 to 0.7 mg enzyme/mg of support. This result is in agreement with earlier reports that have suggested a hydrophobic environment as the most suited for lipase immobilization [22]. The activity of the enzyme was found to remain intact after immobilization. Optimum Temperature for the Activity of Immobilized and Free Enzyme The optimum temperature of the immobilized and free enzyme was investigated and compared (Fig. 7). Lipase immobilized for 3 and 6 h in rod-like SBA-15 was taken for the studies. The activity of the immobilized and free enzyme was found by calculating the rate of hydrolysis of tributyrin at various temperatures. An incubation time of 6 h was provided to identify the effect of prolonged exposure to various temperatures on the enzyme. The enzyme immobilized in rod-like SBA-15 exhibited the maximum activity, and hence, it was chosen for this study. The optimum temperature for immobilized and free enzyme was found to be 30 °C. The specific activity of free enzyme reached 0.02 when exposed to a temperature of 50 °C and reduced to 0.00056 when exposed to a temperature of 60 °C. While for the enzyme which was immobilized for 6 h, the specific activity was 1.52 at 30 °C and reduced to 1.078 at 60 °C. The specific activity reduced by 29 % when the temperature was increased from 30 to 60 °C. This shows that the activity of the immobilized enzyme was higher at 60 °C when compared to that of free enzyme at the same temperature. The enzyme which was immobilized for 3 h showed similar denaturation pattern to that of the free enzyme. This indicates that when the immobilization procedure was carried out for 3 h, the enzyme gets adsorbed only on the outer surface and hence is not protected by the silica matrix. The higher temperature for immobilized enzyme can be attributed to the rigid silica framework that surrounds the enzyme which prevents the enzyme from undergoing changes and denaturation at higher temperatures. Activity Loss The loss in activity of immobilized enzyme was measured after each cycle. The activity in the first cycle was found to be 23.8 U. The enzyme activity in the subsequent cycles was found to decrease to 21 and 18.3 U. This may be due to desorption of some enzyme molecules from the mesopores, during repeated cycles. Table 2 Enzyme loading and activity on functionalized SBA-15 Functionalization Duration for immobilization (h)
Enzyme immobilized (%)
Activity of immobilized enzyme (U)
Specific activity of immobilized enzyme (U/mg)
Amine
20.7
24.014
2.95
26.5
6.72
3 6
Ethyl
9.88
24
27.6
21.542
2.08
3
67.4
24.458
0.90
6
50.0
21.944
1.11
24
70.2
17.778
0.63
Appl Biochem Biotechnol
Fig. 7 Represents the thermal stability of free and immobilized lipases
Enzymatic Transesterification of Rice Bran Oil For the transesterification reaction, rice bran oil and methanol were taken in ratios 1:3, 1:6, and 1:9. The ratio of 1:3, rice bran oil:methanol, was found to be optimum. The immobilized enzyme in rod-like SBA-15 was suspended in 2 mL of phosphate buffer pH 7.2 and added to the oil–methanol mixture and stirred at room temperature. The mixture was subjected to a reaction time of 24 h. After the reaction, the ester and glycerol layers were allowed to settle and the transesterified product layer separated. Lipase immobilized for 3 h, 6 h on rod-like SBA15, and for 3 h in ethyl-functionalized rod-like SBA-15 were used for the transesterification reaction. The yields obtained were 79.8, 79.6, and 81.7 %, respectively. The transesterified sample was heated to remove the glycerol and water contamination. The pure sample was analyzed using FT–IR. The prominent peak at 1742 cm−1 confirms the
Table 3 Composition of transesterified oil (biodiesel) obtained using Rhizopus oryzae lipase Constituent in transesterified oil fraction Octanoic acid, methyl ester
Peak area in GC–MS (%) 0.0255
5-Octadecenoic acid, methyl ester
0.0253
Nonanoic acid, 9-oxo-, methyl ester
0.0139
Tetradecanoic acid, methyl ester
0.0051
Pentadecanoic acid, methyl ester
0.0519
9-Hexadecenoic acid, methyl ester, (Z)-
0.3932
Hexadecanoic acid, methyl ester 9-Octadecenoic acid, methyl ester
19.3842 68.3299
Octadecanoic acid, methyl ester
3.0793
9,12-Octadecadienoic acid (Z,Z)-, methyl ester
0.1225
11-Eicosenoic acid, methyl ester
0.8863
Eicosanoic acid, methyl ester
0.7960
Docosanoic acid, methyl ester
1.1853
Appl Biochem Biotechnol
Fig. 8 GC–MS profile of the transesterified sample obtained by employing 3 h (immobilized) ethylfunctionalized rod-like SBA-15 for the transesterification reaction
presence of ester group in the sample. The peaks at 2922 and 2853 cm−1 confirm the presence of long-chain alkyl groups. The best sample obtained was subjected to GC–MS analysis in order to check for the presence of methyl esters, thus confirming the occurrence of transesterification reaction. It was found that the fatty acids that are present in the oil sample have been converted to their corresponding methyl esters (Table 3 and Fig. 8). Reuse of the Catalyst The reusability of immobilized enzyme increases its commercial viability. When lipase immobilized for 3 h was reused, the conversion efficiency dropped to 67.7 % after the three runs.
Conclusion Mesoporous SBA-15 with different morphology and textural properties has been successfully synthesized, characterized, and used as immobilization matrices for lipase. Rod-like morphology with highly ordered pores in the range 7–8 nm was found to exhibit high enzyme immobilization without compromising the enzyme activity. Functionalization with hydrophobic ethyl groups increased the enzyme immobilization without loss in enzyme activity. The immobilized lipase exhibited good transesterification efficiency and reusability up to 3 cycles. Our data suggests that mesoporous framework holds much promise as an immobilization matrix for lipase and for commercial biodiesel production. Acknowledgments The authors wish to acknowledge the characterization facilities established from the PG Teaching fund of the Nanomission council, Department of Science and Technology, and infrastructure support from SASTRA University.
References 1. Ye, P., Jiang, J., & Xu, Z. K. (2007). Adsorption and activity of lipase from Candida rugosa on the chitosanmodified poly(acrylonitrile-co-maleic acid) membrane surface. Colloids and Surfaces B: Biointerfaces, 60, 62–67. 2. Dizge, N., Aydiner, C., Imer, D. Y., Bayramoglu, M., Tanriseven, A., & Keskinler, B. (2009). Biodiesel production from sunflower, soybean, and waste cooking oils by transesterification using lipase immobilized onto a novel microporous polymer. Bioresource Technology, 100, 1983–1991.
Appl Biochem Biotechnol 3. Prlainovic, N. Z., Knezevic-Jugovic, Z. D., Mijin, D. Z., & Bezbradica, D. I. (2011). Immobilization of lipase from Candida rugosa on Sepabeads((R)): the effect of lipase oxidation by periodates. Bioprocess and Biosystems Engineering, 34, 803–810. 4. Bordes, F., Barbe, S., Escalier, P., Mourey, L., Andre, I., Marty, A., & Tranier, S. (2010). Exploring the conformational states and rearrangements of Yarrowia lipolytica Lipase. Biophysical Journal, 99, 2225– 2234. 5. Piamtongkam, R., Duquesne, S., Bordes, F., Barbe, S., Andre, I., Marty, A., & Chulalaksananukul, W. (2011). Enantioselectivity of Candida rugosa lipases (Lip1, Lip3, and Lip4) towards 2-bromo phenylacetic acid octyl esters controlled by a single amino acid. Biotechnology and Bioengineering, 108, 1749–1756. 6. Gao, S., Wang, Y., Diao, X., Luo, G., & Dai, Y. (2010). Effect of pore diameter and cross-linking method on the immobilization efficiency of Candida rugosa lipase in SBA-15. Bioresource Technology, 101, 3830– 3837. 7. Yin, P., Chen, W., Liu, W., Chen, H., Qu, R., Liu, X., Tang, Q., & Xu, Q. (2013). Efficient bifunctional catalyst lipase/organophosphonic acid-functionalized silica for biodiesel synthesis by esterification of oleic acid with ethanol. Bioresource Technology, 140, 146–151. 8. Grasset, L., Cordier, D., & Ville, A. (1977). Woven silk as a carrier for the immobilization of enzymes. Biotechnology and Bioengineering, 19, 611–618. 9. Mateo, C., Grazu, V., Pessela, B. C., Montes, T., Palomo, J. M., Torres, R., Lopez-Gallego, F., FernandezLafuente, R., & Guisan, J. M. (2007). Advances in the design of new epoxy supports for enzyme immobilization-stabilization. Biochemical Society Transactions, 35, 1593–1601. 10. Bayramoglu, G., Altintas, B., & Arica, M. Y. (2011). Reversible immobilization of uricase on conductive polyaniline brushes grafted on polyacrylonitrile film. Bioprocess and Biosystems Engineering, 34, 127–134. 11. Ducker, R. E., Montague, M. T., & Leggett, G. J. (2008). A comparative investigation of methods for protein immobilization on self-assembled monolayers using glutaraldehyde, carbodiimide, and anhydride reagents. Biointerphases, 3, 59–65. 12. Daglioglu, C., & Zihnioglu, F. (2012). Covalent immobilization of trypsin on glutaraldehyde-activated silica for protein fragmentation. Artificial Cells, Blood Substitutes, and Immobilization Biotechnology, 40, 378– 384. 13. Ferrarotti, S. A., Bolivar, J. M., Mateo, C., Wilson, L., Guisan, J. M., & Fernandez-Lafuente, R. (2006). Immobilization and stabilization of a cyclodextrin glycosyltransferase by covalent attachment on highly activated glyoxyl-agarose supports. Biotechnology Progress, 22, 1140–1145. 14. Vinoba, M., Bhagiyalakshmi, M., Jeong, S. K., Yoon, Y. I., & Nam, S. C. (2012). Immobilization of carbonic anhydrase on spherical SBA-15 for hydration and sequestration of CO2. Colloids and Surfaces B: Biointerfaces, 90, 91–96. 15. Li, S., Wu, Z., Lu, M., Wang, Z., & Li, Z. (2013). Improvement of the enzyme performance of trypsin via adsorption in mesoporous silica SBA-15: hydrolysis of BAPNA. Molecules, 18, 1138–1149. 16. Wan, M. M., Lin, W. G., Gao, L., Gu, H. C., & Zhu, J. H. (2012). Promoting immobilization and catalytic activity of horseradish peroxidase on mesoporous silica through template micelles. Journal of Colloid and Interface Science, 377, 497–503. 17. Magner, E. (2013). Immobilisation of enzymes on mesoporous silicate materials. Chemical Society Reviews, 42, 6213–6222. 18. Gandhi, S., Sethraman, S., & Krishnan, U. M. (2010). Influence of polyhydric solvents on the catalytic and adsorption properties of self-oriented mesoporous SBA-15 silica. Journal of Porous Materials, 18, 329–336. 19. Lowry, O. H., Rosebrough, N. J., Farr, A. L., & Randall, R. J. (1951). Protein measurement with the Folin phenol reagent. Journal of Biological Chemistry, 193, 265–275. 20. Hiol, A., Jonzo, M. D., Rugani, N., Druet, D., Sarda, L., & Comeau, L. C. (2000). Purification and characterization of an extracellular lipase from a thermophilic Rhizopus oryzae strain isolated from palm fruit. Enzyme and Microbial Technology, 26, 421–430. 21. Bayramoglu, G., Karagoz, B., Altintas, B., Arica, M. Y., & Bicak, N. (2011). Poly(styrene-divinylbenzene) beads surface functionalized with di-block polymer grafting and multi-modal ligand attachment: performance of reversibly immobilized lipase in ester synthesis. Bioprocess and Biosystems Engineering, 34, 735–746. 22. Balcao, V. M., & Malcata, F. X. (1998). On the performance of a hollow-fiber bioreactor for acidolysis catalyzed by immobilized lipase. Biotechnology and Bioengineering, 60, 114–123.
