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Immobilization studies and biochemical properties of free and immobilized Rhizopus oryzae lipase onto CaCO3: A comparative study
Biochemical Engineering Journal 37 (2007) 34–41
Immobilization studies and biochemical properties of free and immobilized Rhizopus oryzae lipase onto CaCO3: A comparative study Hanen Ghamgui, Nabil Miled, Maha Karra-chaˆabouni, Youssef Gargouri ∗ Laboratoire de Biochimie et de G´enie Enzymatique des Lipases, ENIS route de Soukra, 3038 Sfax, Tunisia Received 12 April 2006; received in revised form 13 March 2007; accepted 16 March 2007
Abstract Lipase from Rhizopus oryzae (ROL) was immobilized by physical adsorption onto CaCO3 . The immobilization yield was more than 95% during 30 min and corresponds to the loading of 2570 IU/g support. The optimum temperature for both free and immobilized lipase activities was 37 ◦ C. After 24 h of incubation at 50 ◦ C, the immobilized ROL maintained 67% of its initial activity, while the free enzyme was completely inactivated. Therefore, the immobilization seems to improve highly the lipase thermal stability. Besides, the immobilized lipase showed a higher stability than free lipase when stored at 4 ◦ C. The kinetics of the olive oil hydrolysis by the immobilized lipase showed that the hydrolysis rate reached the maximum within 15 min of incubation with the substrate. The hydrolytic activity of the immobilized lipase on olive oil used as substrate was higher than that of the free lipase form, as shown by a higher amount of released free oleic acid. We studied the ethyl oleate ester (biofuel) synthesis by immobilized and free ROL. The conversion yield of this ester was also found to be higher with the immobilized lipase than with the free lipase form (83% versus 6%). Furthermore, electron microscopy allowed us to observe that the morphology of the surface of CaCO3 after the adsorption of ROL showed a large contact area of multipoint attachment with the enzyme. © 2007 Elsevier B.V. All rights reserved. Keywords: CaCO3 ; Rhizopus oryzae lipase; Immobilization; Hydrolysis; Synthesis; Electron microscopy
1. Introduction During the last decade, lipases have been extensively used as biocatalysts for a variety of reactions in organic solvent systems such as the synthesis of esters and the modification of lipids [1–3]. In an aqueous system, these biocatalysts are able to catalyze the hydrolysis of triacylglycerols to give fatty acids and glycerol, both essential in the oleochemical industry [4]. Besides, the lipase-catalyzed hydrolysis of triolein was also used for the production of mono- and di-glycerides, which are widely used as emulsifiers for food and cosmetics [5]. Different techniques for the immobilization of lipase, such as physical adsorption [6], covalent bonding to a solid support [7] and physical entrapment within a polymer matrix support [8] Abbreviations: ROL, Rhizopus oryzae lipase; MO, mono-olein; DO, diolein; OA, oleic acid; TO, triolein; TLC, thin layer chromatography; IU, international unit; TU, total units ∗ Corresponding author. Tel.: +216 74 675055; fax: +216 74 675055. E-mail address: [email protected] (Y. Gargouri). 1369-703X/$ – see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.bej.2007.03.006
have been used to protect the enzyme from the nonpolar solvent environment and enable its reuse. Among the immobilization techniques, the adsorption is one of the simplest methods with a higher commercial potential than other methods. Although the immobilization of lipases has several advantages, the selection of the support is a prominent factor influencing the enzymatic reactions. Comparative studies indicated that dramatic differences in activity are observed among lipases adsorbed onto different materials [9,10]. Furthermore, most studies have observed that one or more properties of immobilized enzymes are different from those of the soluble form [11,12]. In a previous report from our laboratory [13], a crude lipase preparation from Rhizopus oryzae (ROL) was adsorbed on different supports. The CaCO3 was able to adsorb more lipase than the Celite. In addition to its low cost, the CaCO3 has several advantages for use as a matrix including its lack of both toxicity and chemical reactivity with the enzyme. Hence, we determined the best conditions for lipase immobilization on CaCO3 . To explain the difference between the immobilization yields obtained with two supports, the morphology of the CaCO3
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and the Celite 545 before and after enzyme immobilization was examined using electron microscopy and compared to the Celite 545 support. The biochemical properties of the immobilized ROL onto CaCO3 were checked and compared to those obtained with the free one. Therefore, the enzyme performance in the hydrolysis of olive oil and the synthesis of ethyl oleate were established.
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2.4. Temperature activity profile The optimum temperature for the free and the immobilized lipase forms were determined by measuring the hydrolytic activity using olive oil emulsion as substrate [15] at different temperatures (25–55 ◦ C) and at pH 8.0. 2.5. pH activity profile
2. Materials and methods 2.1. Materials Carbonate of calcium (CaCO3 ) was obtained from Pharmacia (Uppsala, Sweden). Celite 545 was purchased from Fluka (Buchs, Switzerland). Virgin olive oil was purchased from the local market. Oleic acid was from Prolabo (Paris, France). All other substances were of analytical grade. The pH-Stat was from Metrohm (Herisau, Switzerland). The shaker (Certomat H/HK) was from B. Braun Biotech (Germany, Melsungen). The freeze-dry (Christ Alpha 1-2, France) was from Bioblock Scientific. 2.2. Production and lyophilization of Rhizopus oryzae lipase Rhizopus oryzae lipase was produced, in our laboratory, as described by Ben Salah et al. [14]. After 24 h of culture, cells were removed by centrifugation and the lipase in the supernatant was precipitated by the addition of ammonium sulphate up to 70% of saturation followed by centrifugation at 8000 rpm at 4 ◦ C for 30 min. The pellet was dissolved in 20 mM sodium acetate buffer pH 5.4 containing 20 mM NaCl and 2 mM benzamidine. Then, the solution was centrifuged at 8000 rpm for 10 min and the supernatant containing the lipase was used for the lyophilization or the immobilization. The lyophilization was carried out for 8 h at −36 ◦ C and 0.37 mbar. The loss of the activity due to the lyophilization was determined as being 2.35% (based on hydrolysis activity before and after the lyophilization). This preparation was stored at 4 ◦ C and used directly as free lipase. The enzyme immobilization was made onto CaCO3 and Celite 545 as described by [13]. 2.3. Lipase hydrolytic activity The activities of the free and the immobilized lipases were measured with a pH-Stat, under the standard assay conditions, using olive oil emulsion as substrate [15]. The reaction mixture contains 10 ml of olive oil emulsion (1 ml of olive oil and 9 ml of Arabic gum at 10%), 20 ml of distilled water and 100 l of bovine serum albumin 12.5%. The activity was expressed as units per ml of enzymatic solution. One international unit (IU) of lipase activity was defined as the amount of enzyme that catalyzes the liberation of 1 mol of fatty acid from olive oil per min at pH 8.0 and at 37 ◦ C.
The optimum pH for the free and the immobilized lipase forms were determined by measuring the hydrolytic activity using olive oil emulsion as substrate [15]. The assays were carried out by incubating the enzyme in 50 mM buffer at different pH values (3–11) for 30 min at 25 ◦ C. Buffers used were glycin, pH 3–4, acetate, pH 5–6, Tris–HCl, pH 7–9, and boric acid, pH 10–11. 2.6. Thermal stability of free and immobilized lipase The free and the immobilized lipase forms were each incubated at various temperatures (30–70 ◦ C) for 24 h. The residual activity after incubation was determined using the standard assay method [15]. The hydrolytic activity of the not incubated enzyme was taken as 100%. 2.7. Hydrolysis of olive oil by free and immobilized lipases The hydrolysis reaction of olive oil was performed using the free and the immobilized lipases as biocatalysts. The reaction was carried out in a reactor maintained at 37 ◦ C by a water jacket with stirring at pH 8.0 and 7.0 for the free and the immobilized lipase, respectively. The reaction mixture contains 5 ml of olive oil emulsion (0.5 ml of olive oil and 4.5 ml of arabic gum at 10%), 10 ml of distilled water and 50 l of bovine serum albumin 12.5%. Reaction was started by adding 1500 IU of either free or immobilized lipase form. The fatty acids released were titrated, at various incubation times with 0.2 N of sodium hydroxide, with keeping constant pH. The hydrolysis percentage corresponds to the ratio free fatty acid × 100/total fatty acid. 2.8. Esterification of ethyl oleate in a solvent-free system The esterification reactions were carried out in screw-capped flasks containing 2 g of total weight taken at oleic acid to ethanol molar ratio of 1, 300 IU of the free or the immobilized lipase, stirred at 200 rpm and at 37 ◦ C. A reaction in the same conditions without adding enzyme was realized in parallel. Aliquots of the reaction mixture were withdrawn at different times. The immobilized enzyme was removed by centrifugation at 8000 rpm for 5 min and the residual acid content was assayed by titration with 0.2 N sodium hydroxide using phenolphthalein as an indicator and 3 ml of ethanol as a quenching agent. The conversion (%) of ester synthesis was calculated based on the conversion of the oleic acid to ester after a given time.
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2.9. Qualitative analysis of reaction products We added 100 l of concentrated HCl to reaction samples in order to neutralize the fatty acids liberated. The neutral lipid classes were extracted using 30 ml of chloroform:methanol (2/1, v/v). The organic phase was retained, the solvent was evaporated on a rotary evaporator and the samples were dissolved in 2 ml of chloroform. The different lipid classes were analyzed by thin layer chromatography (TLC) on Silica 60 F254 previously activated at 60 ◦ C for 30 min. The developing solvent was a mixture of hexane:diethyl ether:methanol:acetic acid (78/17/2/3, v/v/v/v). The lipid spots were visualized with iodine vapor.
lipase was loaded onto the support to reach a maximum value at 2570 TU. At low enzyme loading, the lipase molecules seemed to maximise the contact with the surface, which may result in a loss of conformation and, consequently in reduced activities. However, for a loading amount higher than 2570 TU, multilayer adsorption might have occurred and effectively inhibited access to the enzyme active sites. Minovska et al. [16] reported that the same support displayed a similar immobilization yield (92%) using the commercial Candida rugosa type VII lipase.
3.1.1. The effect of the lipase loading on the immobilization yield To determine the best conditions of lipase immobilization, we studied the influence of the enzyme amount [in the range of 856–10286 TU] to be adsorbed to 1 g of CaCO3 used as support. As shown in Fig. 1, the yield of immobilization increased as more
3.1.2. The effect of the time course on the immobilization yield The immobilization yield was measured at different incubation times of an enzymatic solution (2570 TU) with 1 g of support at 4 ◦ C. The immobilization was found to proceed very rapidly, since it was almost complete within 30 min (Fig. 2). However, we observed a loss of enzymatic activity after 120 min of incubation time. This fact seems to suggest that the desorption of the enzyme could occur easily after 120 min of incubation time with the support probably due to disruption of the weak physical forces linking the enzyme to its support. As a result, a 30 min incubation time of the enzymatic solution with the support was considered ideal to achieve the maximum adsorption of the enzyme onto CaCO3 . Likewise, an incubation of 30 min was needed to achieve the best immobilization yield of Candida rugosa lipase onto AmberliteXAD7 [17]. The immobilization on CaCO3 was carried out with 2570 TU/g support during 30 min and gave rise to the highest immobilization yield (95%). Therefore, these conditions were employed for further experiments. Rosu et al. [18] have studied the adsorption of Pseudomonas SP KWI 56 lipase on the same support. They showed that the immobilization yield of
Fig. 1. Yields of immobilized ROL using different initial activity of soluble lipase. The activity was measured at the pH-stat on olive oil emulsion at pH 8 at 37 ◦ C. Bars indicate standard deviation.
Fig. 2. Yields of immobilized ROL using different incubation time of enzymatic solution with the CaCO3 . The activity was measured at the pH-stat on olive oil emulsion at pH 8 at 37 ◦ C. Bars indicate standard deviation.
2.10. Scanning electron microscopy (SEM) A Baltec Critical Point Dryer 30 field emission scanning electron microscope (Balzers Union, Germany) was used to examine the surface morphology of the carrier before and after the immobilization of ROL. The samples were sputter-coated with gold prior to analysis. 3. Results and discussion 3.1. Factors influencing enzyme leakage
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this enzyme reached only 57.8%. The covalent immobilization of C. rugosa on agarose was checked by Arroyo et al. [19], the method required several chemical steps, the percentage of adsorbed enzyme was very low (3.5%) and the cost was higher. After covalent immobilization, the life time of the immobilized C. rugosa was significantly extended. 3.2. Physicochemical properties of free and immobilized lipases The immobilization procedure was expected to bring changes in the original physicochemical properties as well as in the kinetic behaviour of the enzyme. 3.2.1. Effect of temperature on lipase activity The temperature dependence of the hydrolysis reaction with the free and the immobilized lipases was studied. It was observed that increasing the temperature from 25 to 37 ◦ C increased concomitantly the activity of both free and immobilized lipases (Fig. 3). At 37 ◦ C, this activity reached its optimal value of 31 and 28 IU/mg with the immobilized and the free lipases, respectively (which corresponds to 100%). Therefore, the optimum temperature for both free and immobilized lipases was found to be 37 ◦ C. Beyond 37 ◦ C, the immobilized lipase seemed interestingly to display higher activities than the free one. Indeed, the former retained 65 and 30% of its activity at 45 and 55 ◦ C, respectively. Meanwhile, the free lipase retained only 10% of its initial activity at 45 ◦ C and was completely inactivated at 55 ◦ C. These results seem to differ from previous findings. Hiol et al. [20] reported that the optimum temperature for the immobilized R. oryzae on Amberlite IRC 50 was 35 ◦ C while Montero et al.
Fig. 3. Effect of temperature on free () and immobilized () ROL activities. Enzymes were assayed with olive oil emulsion as substrate at pH 8.0. Bars indicate standard deviation.
Fig. 4. Thermal stability of free () and immobilized () ROL. Enzymes were assayed with olive oil emulsion as substrate at pH 8.0 and 37 ◦ C. Bars indicate standard deviation.
[21] observed a slight shift in the optimal temperature from 45 to 50 ◦ C after immobilization of C. rugosa lipase on Accurel. 3.2.2. Thermal stability of the free and the immobilized lipases To confirm the tolerance of the immobilized lipase towards high temperatures, we carried out experiments on the thermal stability of the immobilized lipase. This can be useful to explore its potential applications. Free and immobilized lipases were incubated at various temperatures for 24 h and the residual activ-
Fig. 5. Residual activity of free and CaCO3 -immobilized lipase as affected by incubation at 4 ◦ C from day 1 to 235. Bars indicate standard deviation.
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Fig. 6. The effect of pH on free () and immobilized () R. oryzae lipase activities. Bars indicate standard deviation.
ity was measured at 37 ◦ C. When compared to the free lipase, the immobilized lipase exhibited a better thermal stability at all temperatures tested (Fig. 4). We observed that upon incubation for 24 h at 50 ◦ C, the free lipase was completely inactivated while the immobilized lipase displayed a residual activity of about 67%. At 60 ◦ C after 24 h of heat treatment, the immobilized lipase retained 32% of its original activity while the free enzyme showed thermal inactivation. These data indicated that the thermal stability of lipase could be enhanced by the immobilization process. De Oliveira et al. [12] have reported that after only 1 h of heat treatment at 60 ◦ C, the Candida rugosa lipase immo-
Fig. 8. Synthesis of ethyl oleate in a solvent-free system by the free () and the immobilized () ROL. Reaction conditions were 300 UI of lipase, an oleic acid/ethanol molar ratio of 1 at 37 ◦ C and stirred at 200 rpm. Bars indicate standard deviation.
bilized on styrene-divinylbenzene retained 50% of its original activity. Under the same conditions, the free enzyme lost 94% of its activity [12]. Here, we show that the immobilization of Rhizopus oryzae lipase onto the CaCO3 increased remarkably its stability towards high temperatures.
Fig. 7. (a) Olive oil hydrolysis by the free () and the immobilized () ROL. Reaction conditions were 1500 UI of lipase, a temperature of 37 ◦ C, with stirring (200 rpm) and a pH of 8.0 and 7.0 for free and immobilized enzymes, respectively. Bars indicate standard deviation. (b) TLC analysis of the hydrolysis product with free (lane 2) and immobilized (lane 3) lipase. A reaction in the same conditions without adding enzyme was used as standard (lane 1).
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Fig. 9. Photomicrographs of the CaCO3 particles (a) before and (b) after the immobilization of ROL and the Celite 545 before (c) and (d) after the immobilization of ROL.
3.3. Effect of storage temperature on activity of lipase The possibility of period of time storing immobilized lipase for a period of time at a certain temperature is one of the key factors to be considered. Generally, enzymes remain active when kept at a low temperature probably because lipases tend to keep their original structure. CaCO3 -immobilized lipase retained 100% of its catalytic activity when stored at 4 ◦ C but the activity of the free lipase gradually decreased to 50% over a period of 151 days (Fig. 5). The stability achieved was most likely due to multiple attachment of the enzyme to the support, preventing any intermolecular process such as proteolysis and aggregation and therefore, creating a more rigid enzyme molecule [22]. 3.4. Effect of pH on lipase activity The effect of pH on the activity of both free and the immobilized ROL was checked in the pH range of 3–11 at 37 ◦ C. As you can see from Fig. 6, the maximum activity was measured at pH 8 with free ROL. In contrast when ROL was immobilized,
the optimum pH was shifted to pH 7. Fig. 6 also shows that the immobilized lipase presented high activities at acidic pH values. A shift of the optimum pH value from 6 to 7 upon immobilization of C. rugosa lipase onto porous chitosan beads was also observed by Pereira et al. [23]. A change of the optimum pH for porcine pancreatic lipase entrapped in Carragenan beads was reported by Desai et al. [24]. 3.5. Performance of free and immobilized lipase in hydrolysis and synthesis reactions 3.5.1. Hydrolysis of olive oil emulsion by free and immobilized lipase The capacities of the immobilized and the free lipase forms to hydrolyze olive oil emulsion were checked. The percentage of the olive oil hydrolysis by the immobilized lipase reached 69% within 15 min of incubation time with the substrate (Fig. 7a). This value was only 29% with the free lipase. Moreover, the TLC analysis showed that the immobilized lipase enhanced the formation of mono-olein and oleic acid leading to a practically
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complete hydrolysis of triolein (Fig. 7b). A similar increase in the lipase activity upon immobilization was described for C. rugosa lipase, using palm oil as substrate, and SGE-A294 copolymer as support [25]. The reason for the increased activity after the immobilization may include protection of the enzyme inside the micropores of the support from the alterations of the microenvironment. Furthermore, the lipase form multiple interactions with CaCO3 through hydrogen or hydrophobic interaction, and enzyme stabilization through multipoint attachment to the support can be postulated. Hence, the hydrolysis of triacylglycerols using ROL immobilized onto CaCO3 can be an attractive method to produce mono-olein that can be used as food emulsifier.
enzyme. This immobilized lipase was applied successfully in the hydrolysis of olive oil and the synthesis of ethyl oleate ester. Furthermore, using a scanning electron micrograph, it was observed that the CaCO3 has a large contact area for multipoint attachment with the enzyme. This may be evidence for a high amount of lipase loading. Besides the simplicity of this immobilization method and the availability of the CaCO3 , the immobilized Rhizopus oryzae lipase showed high activities to perform hydrolysis and esterification reactions. Therefore, natural CaCO3 showed a promising future for the application of natural resource as a support for biocatalysts in various organic syntheses. These would allow the easy immobilization of ROL through a simple and inexpensive method.
3.5.2. Synthesis of ethyl oleate ester by free and immobilized lipases The ability of the immobilized and the free ROL to catalyze synthesis reaction was investigated. We take as example the synthesis of ethyl oleate ester which can be used as a biological additive, a PVC plasticizer and as an additive for hydraulic fluid [26]. As shown in Fig. 8, a higher conversion yield of 83% was obtained with the immobilized lipase. However, the conversion percentage does not exceed 6% using the free lipase. Several studies have described the production of ethyl esters using various immobilized lipases but the conversion yield varied considerably with strain source [27–29]. Therefore, the immobilized lipase was an attractive biocatalyst to carry out the synthesis in a solvent-free system and the hydrolysis reactions.
Acknowledgments
3.6. Morphology by scanning electron micrograph (SEM) The morphology of the CaCO3 before and after the immobilization was examined. We observed that the support particles have a large surface area (Fig. 9a). After the lipase immobilization onto CaCO3 , the support surface area was filled with rounded structures, which were likely to be protein aggregates (Fig. 9b). The flat CaCO3 surface seems to provide a large contact area for a multipoint attachment with the enzyme. The free surface for lipase adsorption seems to be lower in the case of the Celite 545 support than that of the CaCO3 (Fig. 9a and c). Indeed, in a previous work, we obtained an immobilization yield of 93% using the CaCO3 as a support [13]. This yield was only 66% with the Celite 545. This difference was explained in previous work by the preferential binding of the lipase to a CaCO3 support [13]. However, the surface area, as shown by the electronic microscopy, may offer another explanation for this difference in the yield of immobilization. 4. Conclusions The effectiveness of an immobilization process depends on the support used. The increase in stability and the high activity shown by CaCO3 -immobilized lipase would be encouraging for its choice in industrial applications. The immobilization on CaCO3 carried out at 4 ◦ C with 2570 IU/g support during 30 min gave rise to the highest immobilization yield (95%). The immobilized lipase displayed a better thermal stability than the free
We are grateful to Pr. Fakhfakh Z. (FSS, Sfax) for his kind help in the SEM analysis. We are very grateful to Mr. Hajji A. (ENIS, Sfax) for his kind help with the English. We also thank Pr. Gharsallah N`eji (FSS, Sfax) for his help. This work received financial support from «Minist`ere de la recherche scientifique, de la technologie et du d´eveloppement des comp´etences, Tunisie» granted to the Laboratoire de Biochimie et de G´enie Enzymatique des Lipases (ENIS, Sfax). References [1] O.M. Laia, H.M. Ghazalia, F. Cho, C.L. Chong, Physical and textural properties of an experimental table margarine prepared from lipase-catalysed transesterified palm stearin: palm kernel olein mixture during storage, Food Chem. 71 (2000) 173. [2] M. Iso, B. Chen, M. Eguchi, T. Kudo, S. Shrestha, Production of biodiesel fuel from triglycerides and alcohol using immobilized lipase, J. Mol. Catal. B: Enzym. 16 (2001) 53. ˇ Knez, [3] M.D. Romero, L. Calvo, C. Alba, M. Habulin, M. Primoˇziˇc, Z. Enzymatic synthesis of isoamyl acetate with immobilized Candida antarctica lipase in supercritical carbon dioxide, J. Supercrit. Fluids 33 (2005) 77. [4] K. Bagi, L.M. Simon, B. Szaj´ani, Immobilization and characterization of porcine pancreas lipase, Enzyme Microb. Technol. 20 (1997) 531. [5] F.J. Plou, M. Barandiaran, M.V. Calvo, A. Ballesteros, E. Pastor, High-yield production of mono- and di-oleylglycerol by lipase-catalyzed hydrolysis of triolein, Enzyme Microb. Technol. 18 (1996) 66. [6] M. Kamori, T. Hori, M. Yamashita, T. Hori, Y. Yamashita, Y. Hirose, N. Naoshinobu, Immobilization of lipase on a new inorganic ceramics support, toyonite and the reactivity and enantioselectivity of the immobilized lipase, J. Mol. Catal. B: Enzym. 9 (2000) 269. [7] D.R. Walt, V. Agayn, The chemistry of enzyme and protein immobilization with glutardehyde, Trends Anal. Chem. 13 (1994) 425. [8] C. Pizarro, M. Fernandez-Torroba, C. Benito, J. Gonzalez-Saiz, Optimization by experimental design of polyacrylamide gel composition as support for enzyme immobilization by entrapment, Biotechnol. Bioeng. 53 (1997) 497. [9] M. Reslow, P. Adlercreutz, B. Mattiason, On the importance of the support material for bioorganic synthesis influence of material partition between solvent, Enzyme Microb. Technol. 73 (1999) 384. [10] F.J. Plou, M.A. Cruces, M. Ferrer, G. Fuentes, E. Pastor, M. Bernab´e, M. Christensen, F. Comelles, J.L. Parra, A. Ballesteros, Enzymatic acylation of di- and trisaccharides with fatty acids: choosing the appropriate enzyme, support and solvent, J. Biotechnol. 96 (2002) 55. [11] G. Pencreac’h, M. Leullier, J. Baratti, Properties of free and immobilized lipases from Pseudomonas cepacia, Biotechnol. Bioeng. 56 (1997) 181.
H. Ghamgui et al. / Biochemical Engineering Journal 37 (2007) 34–41 [12] P.C. De Oliveira, G.M. Alves, H.F. De Castro, Immobilisation studies and catalytic properties of microbial lipase onto styrene-divinylbenzene copolymer, Biochem. Eng. J. 5 (2000) 63. [13] H. Ghamgui, M. Karra chˆaabouni, Y. Gargouri, 1-Butyl oleate synthesis by immobilized lipase from Rhizopus oryzae: a comparative study between n-hexane and solvent-free system, Enzyme Microb. Technol. 35 (2004) 355. [14] R. Ben Salah, K. Fendri, Y. Gargouri, La lipase de Rhizopus oryzae: production, purification et caract´eristiques biochimiques, Rev. Fr. Corps Gras. 41 (1994) 133. [15] Y. Gargouri, R. Julien, A. Sugihara, L. Sarda, R. Verger, Inhibition of pancreatic and microbial lipases by proteins, Biochim. Biophys. Acta 795 (1984) 326. [16] V. Minovska, E. Winkelhausen, S. Kuzmanova, Lipase immobilized by different techniques on various support materials applied in oil hydrolysis, J. Serb. Chem. Soc. 70 (2005) 609. [17] M. Basri, W.M. Zin Wan Yunus, W.S. Yoong, K. Ampon, C.N.A. Razak, A.B. Salleh, Immobilization of lipase from Candida rugosa on synthetic polymer beads for use in the synthesis of fatty esters, J. Chem. Technol. Biotechnol. 66 (1996) 169. [18] R. Rosu, Y. Uozaki, Y. Iwasaki, T. Yamane, Repeated use of immobilized lipase for monoacylglycerol production by solid-phase glycerolysis of olive oil, J. Am. Oi1 Chem. Soc. 74 (1997) 445. [19] M. Arroyo, J.A. Moreno, J.V. Sinisterra, Immobilisation/stabilisation on different hydroxilic supports of lipase from Candida rugosa, J. Mol. Catal. B: Enzym. 83 (1993) 261. [20] A. Hiol, M.D. Jonzo, N. Rugani, D. Druet, L. Sarda, L.C. Comeau, Purification and characterization of an extracellular lipase from a thermophilic Rhizopus oryzae strain isolated from palm fruit, Enzyme Microb. Technol. 26 (2000) 421.
41
[21] S. Montero, A. Blanco, M.D. Virto, L.C. Landeta, I. Agud, R. Solozabal, Immobilization of Candida rugosa lipase and some properties of the immobilized enzyme, Enzyme Microb. Technol. 15 (1993) 239. [22] M. Basri, K. Ampon, W.M.Z. Wan Yunus, C.N.A. Razak, A.B. Salleh, Immobilization of hydrophobic lipase derivation on to organic polymer beads, J. Chem. Technol. Biotechnol. 59 (1994) 37. [23] E.B. Pereira, H.F. Castro, F.F. Moraes, G.M. Zanin, Kinetic studies of lipase from Candida rugosa: a comparative study between free and immobilized enzyme onto porous chitosan beads, Appl. Biochem. Biotechnol. 91 (2001) 739. [24] P.D. Desai, A.M. Dave, S. Devi, Entrapment of lipase into K-carrageenan beads and its use in hydrolysis of olive oil in biphasic system, J. Mol. Catal. B: Enzym. 31 (2004) 143. [25] L. Mojovic, Z. Knezevic, R. Popadic, S. Jovanovic, Immobilization of lipase from Candida rugosa on a polymer support, Appl. Microbiol. Biotechnol. 50 (1998) 676. [26] S. Hazarika, P. Goswami, N.N. Dutta, A.K. Hazarika, Ethyl oleate synthesis by Porcine pancreatic lipase in organic solvents, Chem. Eng. J. 85 (2002) 61. [27] Li Deng, T. Tan, F. ang, X. Xu, Enzymatic production of fatty acid alkyl esters with a lipase preparation from Candida sp. 99-25, Eur. J. Lipid Sci. Technol. 105 (2003) 727. [28] M. Kaieda, T. Samukawa, T. Matsumoto, K. Ban, A. Kondo, Y. Shimada, H. Noda, F. Nomto, K. Ohtsuka, E. Izumoto, H. Fukuda, Biodiesel fuel production from plant oil catalyzed by Rhizopus oryzae lipase in a watercontaining system without an organic solvent, J. Biosci. Bioeng. 88 (1999) 627. [29] M. Kaieda, T. Samukawa, A. Kondo, H. Fukuda, Effect of methanol and water contents on production of biodiesel fuel from plant oil catalyzed by various lipases in a solvent-free system, J. Biosci. Bioeng. 91 (2001) 12.
Immobilization studies and biochemical properties of free and immobilized Rhizopus oryzae lipase onto CaCO3: A comparative study Hanen Ghamgui, Nabil Miled, Maha Karra-chaˆabouni, Youssef Gargouri ∗ Laboratoire de Biochimie et de G´enie Enzymatique des Lipases, ENIS route de Soukra, 3038 Sfax, Tunisia Received 12 April 2006; received in revised form 13 March 2007; accepted 16 March 2007
Abstract Lipase from Rhizopus oryzae (ROL) was immobilized by physical adsorption onto CaCO3 . The immobilization yield was more than 95% during 30 min and corresponds to the loading of 2570 IU/g support. The optimum temperature for both free and immobilized lipase activities was 37 ◦ C. After 24 h of incubation at 50 ◦ C, the immobilized ROL maintained 67% of its initial activity, while the free enzyme was completely inactivated. Therefore, the immobilization seems to improve highly the lipase thermal stability. Besides, the immobilized lipase showed a higher stability than free lipase when stored at 4 ◦ C. The kinetics of the olive oil hydrolysis by the immobilized lipase showed that the hydrolysis rate reached the maximum within 15 min of incubation with the substrate. The hydrolytic activity of the immobilized lipase on olive oil used as substrate was higher than that of the free lipase form, as shown by a higher amount of released free oleic acid. We studied the ethyl oleate ester (biofuel) synthesis by immobilized and free ROL. The conversion yield of this ester was also found to be higher with the immobilized lipase than with the free lipase form (83% versus 6%). Furthermore, electron microscopy allowed us to observe that the morphology of the surface of CaCO3 after the adsorption of ROL showed a large contact area of multipoint attachment with the enzyme. © 2007 Elsevier B.V. All rights reserved. Keywords: CaCO3 ; Rhizopus oryzae lipase; Immobilization; Hydrolysis; Synthesis; Electron microscopy
1. Introduction During the last decade, lipases have been extensively used as biocatalysts for a variety of reactions in organic solvent systems such as the synthesis of esters and the modification of lipids [1–3]. In an aqueous system, these biocatalysts are able to catalyze the hydrolysis of triacylglycerols to give fatty acids and glycerol, both essential in the oleochemical industry [4]. Besides, the lipase-catalyzed hydrolysis of triolein was also used for the production of mono- and di-glycerides, which are widely used as emulsifiers for food and cosmetics [5]. Different techniques for the immobilization of lipase, such as physical adsorption [6], covalent bonding to a solid support [7] and physical entrapment within a polymer matrix support [8] Abbreviations: ROL, Rhizopus oryzae lipase; MO, mono-olein; DO, diolein; OA, oleic acid; TO, triolein; TLC, thin layer chromatography; IU, international unit; TU, total units ∗ Corresponding author. Tel.: +216 74 675055; fax: +216 74 675055. E-mail address: [email protected] (Y. Gargouri). 1369-703X/$ – see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.bej.2007.03.006
have been used to protect the enzyme from the nonpolar solvent environment and enable its reuse. Among the immobilization techniques, the adsorption is one of the simplest methods with a higher commercial potential than other methods. Although the immobilization of lipases has several advantages, the selection of the support is a prominent factor influencing the enzymatic reactions. Comparative studies indicated that dramatic differences in activity are observed among lipases adsorbed onto different materials [9,10]. Furthermore, most studies have observed that one or more properties of immobilized enzymes are different from those of the soluble form [11,12]. In a previous report from our laboratory [13], a crude lipase preparation from Rhizopus oryzae (ROL) was adsorbed on different supports. The CaCO3 was able to adsorb more lipase than the Celite. In addition to its low cost, the CaCO3 has several advantages for use as a matrix including its lack of both toxicity and chemical reactivity with the enzyme. Hence, we determined the best conditions for lipase immobilization on CaCO3 . To explain the difference between the immobilization yields obtained with two supports, the morphology of the CaCO3
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and the Celite 545 before and after enzyme immobilization was examined using electron microscopy and compared to the Celite 545 support. The biochemical properties of the immobilized ROL onto CaCO3 were checked and compared to those obtained with the free one. Therefore, the enzyme performance in the hydrolysis of olive oil and the synthesis of ethyl oleate were established.
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2.4. Temperature activity profile The optimum temperature for the free and the immobilized lipase forms were determined by measuring the hydrolytic activity using olive oil emulsion as substrate [15] at different temperatures (25–55 ◦ C) and at pH 8.0. 2.5. pH activity profile
2. Materials and methods 2.1. Materials Carbonate of calcium (CaCO3 ) was obtained from Pharmacia (Uppsala, Sweden). Celite 545 was purchased from Fluka (Buchs, Switzerland). Virgin olive oil was purchased from the local market. Oleic acid was from Prolabo (Paris, France). All other substances were of analytical grade. The pH-Stat was from Metrohm (Herisau, Switzerland). The shaker (Certomat H/HK) was from B. Braun Biotech (Germany, Melsungen). The freeze-dry (Christ Alpha 1-2, France) was from Bioblock Scientific. 2.2. Production and lyophilization of Rhizopus oryzae lipase Rhizopus oryzae lipase was produced, in our laboratory, as described by Ben Salah et al. [14]. After 24 h of culture, cells were removed by centrifugation and the lipase in the supernatant was precipitated by the addition of ammonium sulphate up to 70% of saturation followed by centrifugation at 8000 rpm at 4 ◦ C for 30 min. The pellet was dissolved in 20 mM sodium acetate buffer pH 5.4 containing 20 mM NaCl and 2 mM benzamidine. Then, the solution was centrifuged at 8000 rpm for 10 min and the supernatant containing the lipase was used for the lyophilization or the immobilization. The lyophilization was carried out for 8 h at −36 ◦ C and 0.37 mbar. The loss of the activity due to the lyophilization was determined as being 2.35% (based on hydrolysis activity before and after the lyophilization). This preparation was stored at 4 ◦ C and used directly as free lipase. The enzyme immobilization was made onto CaCO3 and Celite 545 as described by [13]. 2.3. Lipase hydrolytic activity The activities of the free and the immobilized lipases were measured with a pH-Stat, under the standard assay conditions, using olive oil emulsion as substrate [15]. The reaction mixture contains 10 ml of olive oil emulsion (1 ml of olive oil and 9 ml of Arabic gum at 10%), 20 ml of distilled water and 100 l of bovine serum albumin 12.5%. The activity was expressed as units per ml of enzymatic solution. One international unit (IU) of lipase activity was defined as the amount of enzyme that catalyzes the liberation of 1 mol of fatty acid from olive oil per min at pH 8.0 and at 37 ◦ C.
The optimum pH for the free and the immobilized lipase forms were determined by measuring the hydrolytic activity using olive oil emulsion as substrate [15]. The assays were carried out by incubating the enzyme in 50 mM buffer at different pH values (3–11) for 30 min at 25 ◦ C. Buffers used were glycin, pH 3–4, acetate, pH 5–6, Tris–HCl, pH 7–9, and boric acid, pH 10–11. 2.6. Thermal stability of free and immobilized lipase The free and the immobilized lipase forms were each incubated at various temperatures (30–70 ◦ C) for 24 h. The residual activity after incubation was determined using the standard assay method [15]. The hydrolytic activity of the not incubated enzyme was taken as 100%. 2.7. Hydrolysis of olive oil by free and immobilized lipases The hydrolysis reaction of olive oil was performed using the free and the immobilized lipases as biocatalysts. The reaction was carried out in a reactor maintained at 37 ◦ C by a water jacket with stirring at pH 8.0 and 7.0 for the free and the immobilized lipase, respectively. The reaction mixture contains 5 ml of olive oil emulsion (0.5 ml of olive oil and 4.5 ml of arabic gum at 10%), 10 ml of distilled water and 50 l of bovine serum albumin 12.5%. Reaction was started by adding 1500 IU of either free or immobilized lipase form. The fatty acids released were titrated, at various incubation times with 0.2 N of sodium hydroxide, with keeping constant pH. The hydrolysis percentage corresponds to the ratio free fatty acid × 100/total fatty acid. 2.8. Esterification of ethyl oleate in a solvent-free system The esterification reactions were carried out in screw-capped flasks containing 2 g of total weight taken at oleic acid to ethanol molar ratio of 1, 300 IU of the free or the immobilized lipase, stirred at 200 rpm and at 37 ◦ C. A reaction in the same conditions without adding enzyme was realized in parallel. Aliquots of the reaction mixture were withdrawn at different times. The immobilized enzyme was removed by centrifugation at 8000 rpm for 5 min and the residual acid content was assayed by titration with 0.2 N sodium hydroxide using phenolphthalein as an indicator and 3 ml of ethanol as a quenching agent. The conversion (%) of ester synthesis was calculated based on the conversion of the oleic acid to ester after a given time.
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2.9. Qualitative analysis of reaction products We added 100 l of concentrated HCl to reaction samples in order to neutralize the fatty acids liberated. The neutral lipid classes were extracted using 30 ml of chloroform:methanol (2/1, v/v). The organic phase was retained, the solvent was evaporated on a rotary evaporator and the samples were dissolved in 2 ml of chloroform. The different lipid classes were analyzed by thin layer chromatography (TLC) on Silica 60 F254 previously activated at 60 ◦ C for 30 min. The developing solvent was a mixture of hexane:diethyl ether:methanol:acetic acid (78/17/2/3, v/v/v/v). The lipid spots were visualized with iodine vapor.
lipase was loaded onto the support to reach a maximum value at 2570 TU. At low enzyme loading, the lipase molecules seemed to maximise the contact with the surface, which may result in a loss of conformation and, consequently in reduced activities. However, for a loading amount higher than 2570 TU, multilayer adsorption might have occurred and effectively inhibited access to the enzyme active sites. Minovska et al. [16] reported that the same support displayed a similar immobilization yield (92%) using the commercial Candida rugosa type VII lipase.
3.1.1. The effect of the lipase loading on the immobilization yield To determine the best conditions of lipase immobilization, we studied the influence of the enzyme amount [in the range of 856–10286 TU] to be adsorbed to 1 g of CaCO3 used as support. As shown in Fig. 1, the yield of immobilization increased as more
3.1.2. The effect of the time course on the immobilization yield The immobilization yield was measured at different incubation times of an enzymatic solution (2570 TU) with 1 g of support at 4 ◦ C. The immobilization was found to proceed very rapidly, since it was almost complete within 30 min (Fig. 2). However, we observed a loss of enzymatic activity after 120 min of incubation time. This fact seems to suggest that the desorption of the enzyme could occur easily after 120 min of incubation time with the support probably due to disruption of the weak physical forces linking the enzyme to its support. As a result, a 30 min incubation time of the enzymatic solution with the support was considered ideal to achieve the maximum adsorption of the enzyme onto CaCO3 . Likewise, an incubation of 30 min was needed to achieve the best immobilization yield of Candida rugosa lipase onto AmberliteXAD7 [17]. The immobilization on CaCO3 was carried out with 2570 TU/g support during 30 min and gave rise to the highest immobilization yield (95%). Therefore, these conditions were employed for further experiments. Rosu et al. [18] have studied the adsorption of Pseudomonas SP KWI 56 lipase on the same support. They showed that the immobilization yield of
Fig. 1. Yields of immobilized ROL using different initial activity of soluble lipase. The activity was measured at the pH-stat on olive oil emulsion at pH 8 at 37 ◦ C. Bars indicate standard deviation.
Fig. 2. Yields of immobilized ROL using different incubation time of enzymatic solution with the CaCO3 . The activity was measured at the pH-stat on olive oil emulsion at pH 8 at 37 ◦ C. Bars indicate standard deviation.
2.10. Scanning electron microscopy (SEM) A Baltec Critical Point Dryer 30 field emission scanning electron microscope (Balzers Union, Germany) was used to examine the surface morphology of the carrier before and after the immobilization of ROL. The samples were sputter-coated with gold prior to analysis. 3. Results and discussion 3.1. Factors influencing enzyme leakage
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this enzyme reached only 57.8%. The covalent immobilization of C. rugosa on agarose was checked by Arroyo et al. [19], the method required several chemical steps, the percentage of adsorbed enzyme was very low (3.5%) and the cost was higher. After covalent immobilization, the life time of the immobilized C. rugosa was significantly extended. 3.2. Physicochemical properties of free and immobilized lipases The immobilization procedure was expected to bring changes in the original physicochemical properties as well as in the kinetic behaviour of the enzyme. 3.2.1. Effect of temperature on lipase activity The temperature dependence of the hydrolysis reaction with the free and the immobilized lipases was studied. It was observed that increasing the temperature from 25 to 37 ◦ C increased concomitantly the activity of both free and immobilized lipases (Fig. 3). At 37 ◦ C, this activity reached its optimal value of 31 and 28 IU/mg with the immobilized and the free lipases, respectively (which corresponds to 100%). Therefore, the optimum temperature for both free and immobilized lipases was found to be 37 ◦ C. Beyond 37 ◦ C, the immobilized lipase seemed interestingly to display higher activities than the free one. Indeed, the former retained 65 and 30% of its activity at 45 and 55 ◦ C, respectively. Meanwhile, the free lipase retained only 10% of its initial activity at 45 ◦ C and was completely inactivated at 55 ◦ C. These results seem to differ from previous findings. Hiol et al. [20] reported that the optimum temperature for the immobilized R. oryzae on Amberlite IRC 50 was 35 ◦ C while Montero et al.
Fig. 3. Effect of temperature on free () and immobilized () ROL activities. Enzymes were assayed with olive oil emulsion as substrate at pH 8.0. Bars indicate standard deviation.
Fig. 4. Thermal stability of free () and immobilized () ROL. Enzymes were assayed with olive oil emulsion as substrate at pH 8.0 and 37 ◦ C. Bars indicate standard deviation.
[21] observed a slight shift in the optimal temperature from 45 to 50 ◦ C after immobilization of C. rugosa lipase on Accurel. 3.2.2. Thermal stability of the free and the immobilized lipases To confirm the tolerance of the immobilized lipase towards high temperatures, we carried out experiments on the thermal stability of the immobilized lipase. This can be useful to explore its potential applications. Free and immobilized lipases were incubated at various temperatures for 24 h and the residual activ-
Fig. 5. Residual activity of free and CaCO3 -immobilized lipase as affected by incubation at 4 ◦ C from day 1 to 235. Bars indicate standard deviation.
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Fig. 6. The effect of pH on free () and immobilized () R. oryzae lipase activities. Bars indicate standard deviation.
ity was measured at 37 ◦ C. When compared to the free lipase, the immobilized lipase exhibited a better thermal stability at all temperatures tested (Fig. 4). We observed that upon incubation for 24 h at 50 ◦ C, the free lipase was completely inactivated while the immobilized lipase displayed a residual activity of about 67%. At 60 ◦ C after 24 h of heat treatment, the immobilized lipase retained 32% of its original activity while the free enzyme showed thermal inactivation. These data indicated that the thermal stability of lipase could be enhanced by the immobilization process. De Oliveira et al. [12] have reported that after only 1 h of heat treatment at 60 ◦ C, the Candida rugosa lipase immo-
Fig. 8. Synthesis of ethyl oleate in a solvent-free system by the free () and the immobilized () ROL. Reaction conditions were 300 UI of lipase, an oleic acid/ethanol molar ratio of 1 at 37 ◦ C and stirred at 200 rpm. Bars indicate standard deviation.
bilized on styrene-divinylbenzene retained 50% of its original activity. Under the same conditions, the free enzyme lost 94% of its activity [12]. Here, we show that the immobilization of Rhizopus oryzae lipase onto the CaCO3 increased remarkably its stability towards high temperatures.
Fig. 7. (a) Olive oil hydrolysis by the free () and the immobilized () ROL. Reaction conditions were 1500 UI of lipase, a temperature of 37 ◦ C, with stirring (200 rpm) and a pH of 8.0 and 7.0 for free and immobilized enzymes, respectively. Bars indicate standard deviation. (b) TLC analysis of the hydrolysis product with free (lane 2) and immobilized (lane 3) lipase. A reaction in the same conditions without adding enzyme was used as standard (lane 1).
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Fig. 9. Photomicrographs of the CaCO3 particles (a) before and (b) after the immobilization of ROL and the Celite 545 before (c) and (d) after the immobilization of ROL.
3.3. Effect of storage temperature on activity of lipase The possibility of period of time storing immobilized lipase for a period of time at a certain temperature is one of the key factors to be considered. Generally, enzymes remain active when kept at a low temperature probably because lipases tend to keep their original structure. CaCO3 -immobilized lipase retained 100% of its catalytic activity when stored at 4 ◦ C but the activity of the free lipase gradually decreased to 50% over a period of 151 days (Fig. 5). The stability achieved was most likely due to multiple attachment of the enzyme to the support, preventing any intermolecular process such as proteolysis and aggregation and therefore, creating a more rigid enzyme molecule [22]. 3.4. Effect of pH on lipase activity The effect of pH on the activity of both free and the immobilized ROL was checked in the pH range of 3–11 at 37 ◦ C. As you can see from Fig. 6, the maximum activity was measured at pH 8 with free ROL. In contrast when ROL was immobilized,
the optimum pH was shifted to pH 7. Fig. 6 also shows that the immobilized lipase presented high activities at acidic pH values. A shift of the optimum pH value from 6 to 7 upon immobilization of C. rugosa lipase onto porous chitosan beads was also observed by Pereira et al. [23]. A change of the optimum pH for porcine pancreatic lipase entrapped in Carragenan beads was reported by Desai et al. [24]. 3.5. Performance of free and immobilized lipase in hydrolysis and synthesis reactions 3.5.1. Hydrolysis of olive oil emulsion by free and immobilized lipase The capacities of the immobilized and the free lipase forms to hydrolyze olive oil emulsion were checked. The percentage of the olive oil hydrolysis by the immobilized lipase reached 69% within 15 min of incubation time with the substrate (Fig. 7a). This value was only 29% with the free lipase. Moreover, the TLC analysis showed that the immobilized lipase enhanced the formation of mono-olein and oleic acid leading to a practically
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complete hydrolysis of triolein (Fig. 7b). A similar increase in the lipase activity upon immobilization was described for C. rugosa lipase, using palm oil as substrate, and SGE-A294 copolymer as support [25]. The reason for the increased activity after the immobilization may include protection of the enzyme inside the micropores of the support from the alterations of the microenvironment. Furthermore, the lipase form multiple interactions with CaCO3 through hydrogen or hydrophobic interaction, and enzyme stabilization through multipoint attachment to the support can be postulated. Hence, the hydrolysis of triacylglycerols using ROL immobilized onto CaCO3 can be an attractive method to produce mono-olein that can be used as food emulsifier.
enzyme. This immobilized lipase was applied successfully in the hydrolysis of olive oil and the synthesis of ethyl oleate ester. Furthermore, using a scanning electron micrograph, it was observed that the CaCO3 has a large contact area for multipoint attachment with the enzyme. This may be evidence for a high amount of lipase loading. Besides the simplicity of this immobilization method and the availability of the CaCO3 , the immobilized Rhizopus oryzae lipase showed high activities to perform hydrolysis and esterification reactions. Therefore, natural CaCO3 showed a promising future for the application of natural resource as a support for biocatalysts in various organic syntheses. These would allow the easy immobilization of ROL through a simple and inexpensive method.
3.5.2. Synthesis of ethyl oleate ester by free and immobilized lipases The ability of the immobilized and the free ROL to catalyze synthesis reaction was investigated. We take as example the synthesis of ethyl oleate ester which can be used as a biological additive, a PVC plasticizer and as an additive for hydraulic fluid [26]. As shown in Fig. 8, a higher conversion yield of 83% was obtained with the immobilized lipase. However, the conversion percentage does not exceed 6% using the free lipase. Several studies have described the production of ethyl esters using various immobilized lipases but the conversion yield varied considerably with strain source [27–29]. Therefore, the immobilized lipase was an attractive biocatalyst to carry out the synthesis in a solvent-free system and the hydrolysis reactions.
Acknowledgments
3.6. Morphology by scanning electron micrograph (SEM) The morphology of the CaCO3 before and after the immobilization was examined. We observed that the support particles have a large surface area (Fig. 9a). After the lipase immobilization onto CaCO3 , the support surface area was filled with rounded structures, which were likely to be protein aggregates (Fig. 9b). The flat CaCO3 surface seems to provide a large contact area for a multipoint attachment with the enzyme. The free surface for lipase adsorption seems to be lower in the case of the Celite 545 support than that of the CaCO3 (Fig. 9a and c). Indeed, in a previous work, we obtained an immobilization yield of 93% using the CaCO3 as a support [13]. This yield was only 66% with the Celite 545. This difference was explained in previous work by the preferential binding of the lipase to a CaCO3 support [13]. However, the surface area, as shown by the electronic microscopy, may offer another explanation for this difference in the yield of immobilization. 4. Conclusions The effectiveness of an immobilization process depends on the support used. The increase in stability and the high activity shown by CaCO3 -immobilized lipase would be encouraging for its choice in industrial applications. The immobilization on CaCO3 carried out at 4 ◦ C with 2570 IU/g support during 30 min gave rise to the highest immobilization yield (95%). The immobilized lipase displayed a better thermal stability than the free
We are grateful to Pr. Fakhfakh Z. (FSS, Sfax) for his kind help in the SEM analysis. We are very grateful to Mr. Hajji A. (ENIS, Sfax) for his kind help with the English. We also thank Pr. Gharsallah N`eji (FSS, Sfax) for his help. This work received financial support from «Minist`ere de la recherche scientifique, de la technologie et du d´eveloppement des comp´etences, Tunisie» granted to the Laboratoire de Biochimie et de G´enie Enzymatique des Lipases (ENIS, Sfax). References [1] O.M. Laia, H.M. Ghazalia, F. Cho, C.L. Chong, Physical and textural properties of an experimental table margarine prepared from lipase-catalysed transesterified palm stearin: palm kernel olein mixture during storage, Food Chem. 71 (2000) 173. [2] M. Iso, B. Chen, M. Eguchi, T. Kudo, S. Shrestha, Production of biodiesel fuel from triglycerides and alcohol using immobilized lipase, J. Mol. Catal. B: Enzym. 16 (2001) 53. ˇ Knez, [3] M.D. Romero, L. Calvo, C. Alba, M. Habulin, M. Primoˇziˇc, Z. Enzymatic synthesis of isoamyl acetate with immobilized Candida antarctica lipase in supercritical carbon dioxide, J. Supercrit. Fluids 33 (2005) 77. [4] K. Bagi, L.M. Simon, B. Szaj´ani, Immobilization and characterization of porcine pancreas lipase, Enzyme Microb. Technol. 20 (1997) 531. [5] F.J. Plou, M. Barandiaran, M.V. Calvo, A. Ballesteros, E. Pastor, High-yield production of mono- and di-oleylglycerol by lipase-catalyzed hydrolysis of triolein, Enzyme Microb. Technol. 18 (1996) 66. [6] M. Kamori, T. Hori, M. Yamashita, T. Hori, Y. Yamashita, Y. Hirose, N. Naoshinobu, Immobilization of lipase on a new inorganic ceramics support, toyonite and the reactivity and enantioselectivity of the immobilized lipase, J. Mol. Catal. B: Enzym. 9 (2000) 269. [7] D.R. Walt, V. Agayn, The chemistry of enzyme and protein immobilization with glutardehyde, Trends Anal. Chem. 13 (1994) 425. [8] C. Pizarro, M. Fernandez-Torroba, C. Benito, J. Gonzalez-Saiz, Optimization by experimental design of polyacrylamide gel composition as support for enzyme immobilization by entrapment, Biotechnol. Bioeng. 53 (1997) 497. [9] M. Reslow, P. Adlercreutz, B. Mattiason, On the importance of the support material for bioorganic synthesis influence of material partition between solvent, Enzyme Microb. Technol. 73 (1999) 384. [10] F.J. Plou, M.A. Cruces, M. Ferrer, G. Fuentes, E. Pastor, M. Bernab´e, M. Christensen, F. Comelles, J.L. Parra, A. Ballesteros, Enzymatic acylation of di- and trisaccharides with fatty acids: choosing the appropriate enzyme, support and solvent, J. Biotechnol. 96 (2002) 55. [11] G. Pencreac’h, M. Leullier, J. Baratti, Properties of free and immobilized lipases from Pseudomonas cepacia, Biotechnol. Bioeng. 56 (1997) 181.
H. Ghamgui et al. / Biochemical Engineering Journal 37 (2007) 34–41 [12] P.C. De Oliveira, G.M. Alves, H.F. De Castro, Immobilisation studies and catalytic properties of microbial lipase onto styrene-divinylbenzene copolymer, Biochem. Eng. J. 5 (2000) 63. [13] H. Ghamgui, M. Karra chˆaabouni, Y. Gargouri, 1-Butyl oleate synthesis by immobilized lipase from Rhizopus oryzae: a comparative study between n-hexane and solvent-free system, Enzyme Microb. Technol. 35 (2004) 355. [14] R. Ben Salah, K. Fendri, Y. Gargouri, La lipase de Rhizopus oryzae: production, purification et caract´eristiques biochimiques, Rev. Fr. Corps Gras. 41 (1994) 133. [15] Y. Gargouri, R. Julien, A. Sugihara, L. Sarda, R. Verger, Inhibition of pancreatic and microbial lipases by proteins, Biochim. Biophys. Acta 795 (1984) 326. [16] V. Minovska, E. Winkelhausen, S. Kuzmanova, Lipase immobilized by different techniques on various support materials applied in oil hydrolysis, J. Serb. Chem. Soc. 70 (2005) 609. [17] M. Basri, W.M. Zin Wan Yunus, W.S. Yoong, K. Ampon, C.N.A. Razak, A.B. Salleh, Immobilization of lipase from Candida rugosa on synthetic polymer beads for use in the synthesis of fatty esters, J. Chem. Technol. Biotechnol. 66 (1996) 169. [18] R. Rosu, Y. Uozaki, Y. Iwasaki, T. Yamane, Repeated use of immobilized lipase for monoacylglycerol production by solid-phase glycerolysis of olive oil, J. Am. Oi1 Chem. Soc. 74 (1997) 445. [19] M. Arroyo, J.A. Moreno, J.V. Sinisterra, Immobilisation/stabilisation on different hydroxilic supports of lipase from Candida rugosa, J. Mol. Catal. B: Enzym. 83 (1993) 261. [20] A. Hiol, M.D. Jonzo, N. Rugani, D. Druet, L. Sarda, L.C. Comeau, Purification and characterization of an extracellular lipase from a thermophilic Rhizopus oryzae strain isolated from palm fruit, Enzyme Microb. Technol. 26 (2000) 421.
41
[21] S. Montero, A. Blanco, M.D. Virto, L.C. Landeta, I. Agud, R. Solozabal, Immobilization of Candida rugosa lipase and some properties of the immobilized enzyme, Enzyme Microb. Technol. 15 (1993) 239. [22] M. Basri, K. Ampon, W.M.Z. Wan Yunus, C.N.A. Razak, A.B. Salleh, Immobilization of hydrophobic lipase derivation on to organic polymer beads, J. Chem. Technol. Biotechnol. 59 (1994) 37. [23] E.B. Pereira, H.F. Castro, F.F. Moraes, G.M. Zanin, Kinetic studies of lipase from Candida rugosa: a comparative study between free and immobilized enzyme onto porous chitosan beads, Appl. Biochem. Biotechnol. 91 (2001) 739. [24] P.D. Desai, A.M. Dave, S. Devi, Entrapment of lipase into K-carrageenan beads and its use in hydrolysis of olive oil in biphasic system, J. Mol. Catal. B: Enzym. 31 (2004) 143. [25] L. Mojovic, Z. Knezevic, R. Popadic, S. Jovanovic, Immobilization of lipase from Candida rugosa on a polymer support, Appl. Microbiol. Biotechnol. 50 (1998) 676. [26] S. Hazarika, P. Goswami, N.N. Dutta, A.K. Hazarika, Ethyl oleate synthesis by Porcine pancreatic lipase in organic solvents, Chem. Eng. J. 85 (2002) 61. [27] Li Deng, T. Tan, F. ang, X. Xu, Enzymatic production of fatty acid alkyl esters with a lipase preparation from Candida sp. 99-25, Eur. J. Lipid Sci. Technol. 105 (2003) 727. [28] M. Kaieda, T. Samukawa, T. Matsumoto, K. Ban, A. Kondo, Y. Shimada, H. Noda, F. Nomto, K. Ohtsuka, E. Izumoto, H. Fukuda, Biodiesel fuel production from plant oil catalyzed by Rhizopus oryzae lipase in a watercontaining system without an organic solvent, J. Biosci. Bioeng. 88 (1999) 627. [29] M. Kaieda, T. Samukawa, A. Kondo, H. Fukuda, Effect of methanol and water contents on production of biodiesel fuel from plant oil catalyzed by various lipases in a solvent-free system, J. Biosci. Bioeng. 91 (2001) 12.