Immobilization of lipase on porous monodisperse chitosan microspheres
Yang Chen ∗ Junteng Liu Chunjie Xia Chenxi Zhao Zhongqi Ren Weidong Zhang
Beijing Key Laboratory of Membrane Science and Technology, Beijing University of Chemical Technology, Beijing, People’s Republic of China
Abstract Porous monodisperse chitosan microspheres were synthesized for enzyme immobilization. The microspheres were prepared using microchannels and modified with glutaraldehyde. The microspheres had a mean diameter of 495 µm; the polydispersity indices were less than 0.08, and the specific surface area was between 121 and 173 m2 /g. Candida sp. 1619 lipase was selected as a model lipase. Immobilization conditions such as enzyme loading, glutaraldehyde
concentration, and immobilization time were optimized. The temperature, pH, and storage stability of the free and immobilized enzymes were also investigated. The immobilized enzyme had broad-ranging pH and temperature optima as compared with free enzyme. The storage stability of the immobilized enzyme was higher than that of the free enzyme. C 2014 International Union of Biochemistry and Molecular Biology, Inc. Volume 00, Number 0, Pages 1–6, 2014
Keywords: chitosan, immobilized enzyme, lipase activity, monodisperse, porous microspheres, protein loading
1. Introduction The specificity and catalytic performance of enzymes such as their hydrolysis, esterification, and aminolysis [1, 2] make them particular industrial biocatalysts in biochemical, biomedical, and food industrial processes [3]. However, natural enzymes rarely have the characteristics of industrial catalysts. Enzymes are often difficult to recover and reuse as catalysts from reaction solutions or from substrates and products [4]. Enzyme immobilization is the most useful way to improve the catalytic performance of enzymes in nonnatural environments, providing many important advantages over the use of natural enzymes; these advantages include enzyme activity, stability, reusability, continuous operation, and process economics [5]. Enzyme immobilization has been researched using
∗ Address for correspondence: Junteng Liu, PhD, Associate Professor, Beijing Key Laboratory of Membrane Science and Technology, Beijing University of Chemical Technology, PO Box 1, No. 15, N. 3rd Ring Rd. East, Beijing 100029, People’s Republic of China. Tel.: +86-10-6442-3628; Fax: +86-10-6443-6781; e-mail: [email protected]. Received 24 December 2013; accepted 7 May 2014
DOI: 10.1002/bab.1242 Published online in Wiley Online Library (wileyonlinelibrary.com)
various carriers, such as gel [6], inorganic materials [7], and polymer [8]. Chitosan is a natural acetylated glucosamine biopolymer. Given its hydrophilicity and biocompatibility, chitosan has attracted considerable attention in the development of enzyme immobilization [9]. Chitosan also has many hydroxyl and amino groups, which produce benefits when linked to enzymes [10]. As an enzyme carrier, chitosan has various molecular forms, such as fibers, membranes, and beads. Because of their high porosity and large specific surface area, porous chitosan microspheres offer low mass transfer resistance and small steric hindrance to enzymes within their three-dimensional polymer networks. When enzymes are immobilized by coupling to porous particles, diffusion limitations and catalytic activity decrease caused by immobilization are principally dependent on their structure, such as the sphere diameter and pore size [11]. The catalytic activity of immobilized enzymes increases as the sphere diameter decreases [12]. However, the diameter of particles is not always “the smaller, the better” under different situations; especially in large-scale industrial manufacturing, the immobilized enzyme is filled into columns or packed-bed reactors [8]. The immobilized enzyme is difficult to separate from products and to reuse if the particle size is too small. Therefore, micrometer-sized particles are especially suited for industrial applications with their many active sites and large specific surface area to fix enzyme molecules [13, 14].
1
Biotechnology and Applied Biochemistry Several methods to prepare chitosan microparticles have been reported, including emulsion techniques [15], spraydrying [16], and coacervation [17]. However, the size and size distribution of microspheres are difficult to control by these methods. The weakness of nonuniform sphere sizes with wide size distribution may limit practical applications. Thus, a reproducible technique for producing monodisperse chitosan microspheres should be developed to ensure that every batch of immobilized enzymes has consistent application value. The microfluidic chip is an emerging tool for manufacturing microparticles with precisely controlled and monodisperse size distributions [18–21]. Microfluidic methods have been developed as novel approaches for the controllable synthesis of monodisperse microspheres. Several research groups [22–24] have utilized such techniques to produce chitosan microspheres. Monodisperse chitosan microspheres with controlled sizes (100–500 µm in diameter) and narrow size distribution (a variation of less than 10%) have been successfully synthesized. Pore structure considerably influences immobilization and sphere size. In the current study, cross-linking was used to control pore structure. Enzyme immobilization on such cross-linked support does not require chemical activation because the cross-linker, normally a bifunctional reagent, has two functions: cross-linking and activation. Glutaraldehyde is often used as a cross-linking and surface-activating agent because of its reliability and ease of use and, more importantly, because of the availability of amino groups for the reaction with glutaraldehyde not only on enzymes but also on chitosan. In the present work, chitosan microspheres were prepared by microchannel and modified with glutaraldehyde. The characteristics of porous chitosan microspheres were studied. Candida sp. 1619 lipase was selected as a model lipase for immobilization on microparticles. To maximize the adsorption and expression of the enzymes, immobilization conditions were optimized in terms of immobilization time, enzyme loading, and glutaraldehyde concentration. In addition, the thermal, pH, and storage stability of the free and immobilized lipases were evaluated.
2. Experiment 2.1. Materials Candida sp. 1619 lipase was obtained from CTA New Century Biotechnology Co. (Beijing, People’s Republic of China). Bovine serum albumin was obtained from Beijing Jingke Chemical Reagent Corporation (Beijing, People’s Republic of China). Chitosan (degree of deacetylation = 90%) was obtained from Sinopharn Chemical Reagent Co. (Shanghai, People’s Republic of China). Liquid paraffin, glutaraldehyde, Span80, and Tween20 were obtained from Fuchen Chemical Reagent Co. (Tianjin, People’s Republic of China). Oleic acid and olive oil were obtained from Beijing Chemical Reagents Company (Beijing, People’s Republic of China). Coomassie brilliant blue G-250 was obtained from Fluka Corporation (Buchs,
2
Switzerland). All other reagents were commercially available and of the highest grade.
2.2. Preparation of chitosan microspheres Chitosan solution was prepared by initially dissolving 0.5 g of chitosan in 50 mL of an aqueous 1% (v/v) acetic acid solution. Liquid paraffin was used as the oil phase. These fluids were injected into a microchannel at fixed flow rates by syringe pumps (LSP02-1B, Longer Precision Pump Co., Hebei, People’s Republic of China). The chitosan aqueous solution (the disperse phase) was sheared by the liquid paraffin (the continuous phase), developing a series of monodisperse spherical structures called water-in-oil emulsions. When these emulsions were introduced to a 1 mol/L NaOH solution through a Teflon tube, they reacted with OH− at the interface and instantly formed chitosan microparticles. Then, these microparticles underwent cross-linking and modification when they were introduced to a glutaraldehyde solution. After 4 H of hardening, chitosan hydrogels were observed.
2.3. Lipase immobilization Candida sp. 1619 lipase was immobilized on chitosan microparticles by adsorption and glutaraldehyde modification. A certain quantity of lipase was added to sodium phosphate buffer (0.2 mol/L, pH 7.0) to prepare the lipase solution. The solution was then stored at 4 ◦ C for about 6 H until the lipase was completely dissolved. Dried chitosan particles (0.5 g) were suspended in lipase solution for a preset time at room temperature. Then immobilized microparticles were filtered and washed by sodium phosphate buffer (0.2 mol/L, pH 7).
2.4. Analytical methods 2.4.1. Determination of protein loading Protein loading was calculated according to difference in protein content between the original and filtered lipase solution. The protein concentration was estimated by Bradford’s method, using Coomassie brilliant blue G-250 [25]. Bovine serum albumin was used to establish the standard curve.
2.4.2. Determination of immobilized lipase activity Lipase activity was assayed by modified copper soap colorimetry [26]. One unit of lipase activity is defined as 1 µmol of free fatty acid released per minute per gram of immobilized lipase [27]. The substrate, which comprised 3 mL of phosphate buffer (pH 7.0) and 1 mL of olive oil, was pretreated for 5 Min at 37 ◦ C. Afterwards, immobilized lipase was added to the substrate, and reaction was started. Eight milliliters of methylbenzene was used to terminate the reaction after 15 Min. Lipase activity was analyzed by a spectrophotometer at 710 nm after 1 mL of copper reagent [(CH3 COO)2 Cu] was added to the methylbenzene phase. Residual activity was defined as the ratio of the activity of immobilized lipase to the activity at optimum reaction conditions. Each data was the average of at least three parallel experiments, and the standard deviation was within ±5%. The value for the average activity of the placebo version of the substrate and chitosan microspheres was consistently
Immobilization of Lipase
Particle size distributions.
FIG. 1
FIG. 3
low but was subtracted from the resulting values for each measurement of the free and immobilized lipase.
3. Results and Discussion 3.1. Characteristics of porous chitosan microspheres Porous monodisperse chitosan microspheres were prepared by microchannel and cross-linked with glutaraldehyde. The mean diameter of the prepared microspheres was 495 µm, and the polydispersity indices were less than 0.08 (Fig. 1). As shown in Fig. 2a, the chitosan microparticles were porous particles. The specific surface area was measured using the Brunauer–Emmett–Teller method of adsorption of nitrogen gas. The specific surface area was between 121 and 173 m2 /g. The degree of lipase immobilization is determined by the specific surface area of the microspheres. In addition, the porosity of microparticles controls the rate of enzymatic bioconversions by controlling the diffusion of substrates and products through pores. A large specific surface area can reduce diffu-
FIG. 2
Effect of glutaraldehyde concentration on specific surface area of chitosan microspheres.
sional limitations and promote the expression of lipase activity. The concentration of the cross-linker determines the porous structure of microspheres and requires optimization. As shown in Fig. 3, the specific surface area of the chitosan microspheres increased as glutaraldehyde concentration was increased from 1% to 9% (v/v). The surface of a chitosan microsphere with lipase immobilization is shown in Fig. 2b, in contrast with the surface of a chitosan microsphere without lipase immobilization (Fig. 2a). Some nanoparticles overspread the surface of the chitosan microsphere. Candida lipase has a molecular volume of 5 × 4.2 × 3 nm3 and a molecular mass of 45,000 to 60,000 Da [28]. The lipase is adsorbed onto the porous scaffolds.
3.2. Effect of immobilization time on lipase immobilization The time required for immobilizing lipase was optimized for the chitosan microparticles. As shown in Fig. 4, the activity of
Scanning electron microscope images: (a) surface of freeze-dried chitosan microsphere and (b) surface of freeze-dried chitosan microsphere with lipase loaded.
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FIG. 5 FIG. 4
Effect of immobilizing time on the activity of immobilized lipase (lipase concentration was 15 g/L, lipase solution was 20 mL, and glutaraldehyde concentration was 7%).
the immobilized lipase increased as the immobilized time was increased. Because the cross-linked enzyme creates a monolayer of enzymes on the microsphere surface, activity increased when immobilization time was increased. The immobilized enzyme exhibited maximal activity after 9 H of immobilization, and extended immobilization time reduced the lipase activities. With extended immobilization time, more enzyme molecules were immobilized on the microspheres, and the excessively immobilized enzyme can make the aperture of the microspheres relatively thin, lowering the accessibility of the substrate to the active sites [29]. Thus, the optimal immobilizing time was found to be 9 H.
3.3. Effect of enzyme loading on lipase immobilization Chitosan microparticles were loaded with lipase solution (20 mL) at different lipase concentrations, and protein loading and lipase activity was evaluated. As shown in Fig. 5, protein loading increased when lipase concentration increased from 5 to 30 g/L. However, lipase activity decreased when the lipase concentration exceeded 15 g/L. Presumably, more enzyme molecules went inside the microspheres after the active sites of the carriers were saturated. The enzyme molecules cannot efficiently perform catalysis for the limitation of substrate diffusion. Meanwhile, enzymes can be hidden and some active sites can be damaged in a high-immobilization enzyme system [29]. To maintain a balance of the protein loading and enzyme activity, the optimal enzyme concentration was 15 g/L.
4
FIG. 6
Effect of enzyme concentration on the enzyme loading amount and the activity of immobilized lipase (immobilizing time was 9 H, lipase solution was 20 mL, and glutaraldehyde concentration was 7%).
Effect of glutaraldehyde concentration on the enzyme loading amount and the activity of immobilized lipase (immobilizing time was 9 H, lipase concentration was 15 g/L, and lipase solution was 20 mL).
3.4. Effect of glutaraldehyde concentration on lipase immobilization Glutaraldehyde is not only the cross-linker but also the denaturant. Thus, glutaraldehyde concentration can directly affect the specific surface area of chitosan microspheres as well as the activity of immobilized lipase. As shown in Fig. 6, when the glutaraldehyde concentration was lower than 7%, the activity of immobilized enzyme kept pace with the glutaraldehyde concentration. However, when the concentration was higher than 7%, lipase activity decreased, because not enough glutaraldehyde participates in
Immobilization of Lipase
FIG. 7
Effect of temperature on the activity of free and immobilized enzymes.
FIG. 8
Effect of pH on the activity of free and immobilized enzymes.
FIG. 9
Effect of storage on the activity of free and immobilized enzymes.
the cross-linking reaction when the glutaraldehyde concentration is lower than 7%. However, when the concentration is too high, glutaraldehyde is involved in aldol condensation, which can affect the formation of pores on the surface of microspheres and thus make immobilization harder. In addition, as glutaraldehyde concentration exceeds 7%, the extensive interaction of individual enzymes with aldehyde groups on the surface of the microspheres possibly changes the enzyme conformation and lowers the enzyme activity [30]. Thus, the optimal concentration of glutaraldehyde was found to be 7%.
3.5. Thermal stability The effect of temperature on the activity of free and immobilized lipase was studied by measuring residual enzyme activity from 27 to 67 ◦ C. As shown in Fig. 7, the optimal temperature of free and immobilized lipase was 37 ◦ C. The influence of temperature on immobilized lipase was much smaller than that on free lipase. At 67 ◦ C, the free lipase retained only 18.9% of residual activity, whereas the immobilized lipase retained 63.3% of its initial activity. The immobilization step increases enzyme’s rigidity, generally reflected by an increase in stability toward denaturation by raising the temperature [31]. Thus, this immobilization method enhanced the thermal stability of the lipase.
immobilized lipase still retained 71.2% of its initial activity. Thus, this immobilization method optimized the pH stability of the lipase.
3.6. pH stability
3.7. Storage stability
The effect of pH on the residual activity of free and immobilized lipase was studied at pH values between 5 and 10 at 37 ◦ C (Fig. 8). The optimal pH for free lipase was 6, and that for mobilized enzymes was 7. As expected, the immobilized lipase exhibited a wider pH range of activity than the free lipase. The immobilization process advanced the pH stability of lipase activity, especially in the alkaline condition. At pH 10, the free lipase retained only 12.8% of residual activity, whereas the
Storage stability for the immobilized enzyme is a significant index for evaluating the properties of the enzyme, making the immobilized enzyme more advantageous than the free one. Free and immobilized enzymes were stored in sodium phosphate buffer (0.2 mol/L, pH 7.0) at 4 ◦ C for 30 days. Aliquots of these lipases were periodically withdrawn for assay. Figure 9 shows the effect of storage on the activity of free and immobilized enzymes. After 7 days, the activity of the free
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Biotechnology and Applied Biochemistry enzyme decreased to less than 20% of the initial value, whereas the immobilized enzyme still retained about 80% of its initial activity. The immobilized enzyme exhibited 50% of its initial activity only after about 30 days. However, the activity of the free enzyme decreased to less than 10% at that time.
[5] [6] [7] [8] [9] [10]
4. Conclusion Chitosan microspheres were prepared by microchannel and modified with glutaraldehyde. The mean diameter of the prepared microspheres was 495 µm, and the polydispersity indices were less than 0.08. The specific surface area of the microsphere was between 121 and 173 m2 /g. Candida sp. 1619 lipase was selected as the model lipase to study immobilization on the microspheres. Immobilization conditions were optimized in terms of immobilization time, enzyme loading, and glutaraldehyde concentration. The optimal immobilizing time, enzyme loading, and glutaraldehyde concentration were 9 H, 15 g/L, and 7%, respectively. The immobilized lipase had a wide range of pH and temperature optima. The storage stability of the immobilized enzyme was higher than that of the free enzyme. This method can improve controlled-release delivery systems, and the preparation of monodisperse chitosan microparticles will provide many potential biomedical applications.
5. References [1] Brady, D., Steenkamp, L., Skein, E., Chaplin, J. A., and Reddy, S. (2004) Enzyme Microb. Technol. 34, 283–291. [2] Liu, C. H., and Chang, J. S. (2008) Bioresour. Technol. 99, 1616–1622. [3] Hung, T. C., Giridhar, R., Chiou, S. H., and Wu, W. T. (2003) J. Mol. Catal. B Enzym. 26, 69–78. [4] Zhang, Y., Xu, J., Yuan, Z., Xu, H., and Yu, Q. (2010) Bioresour. Technol. 101, 3153–3158.
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[11] [12] [13] [14] [15] [16] [17] [18] [19] [20] [21] [22] [23] [24] [25] [26] [27] [28] [29] [30] [31]
Dosanjh, N. S., and Kaur, J. (2002) Biotechnol. Appl. Bioc. 36(1), 7–12. Prasad, M., and Palanivelu, P. (2014) Biotechnol. Appl. Biochem., in press. Wang, Y. J, and Caruso, F. (2005) Chem. Mater. 17, 953–961. Hu, J., Li, S. J., and Liu, B. (2006) Biotechnol. J. 1, 75–79. Su, Z., Yu, Y., Liang, C., Li, L., and Yu, S. (2013) Biotechnol. Appl. Biochem. 60, 231–235. Adriano, W. S., Edilson Filho, H. C., Silva, J. A., and Gonc¸alves, L. R. (2005) Biotechnol. Appl. Bioc. 41(3), 201–207. Vaidya, B. K., Ingavle, G. C., Ponrathnam S., and Kulkarni, B. D. (2008) Bioresour. Technol. 99, 3623–3629. Bailey, J. E., and Cho, Y. K.(1983) Biotechnol. Bioeng. 25, 1923–1935. Guo, Z., Bai, S., and Sun, Y. (2003) Enzyme Microb. Technol. 32, 776–782. Budriene, S., Gorochovceva, N., Romaskevic, T., Yugova, L. V., and Miezeliene, A. (2005) Cent. Eur. J. Chem. 3, 95–105. Genita, I., Perugini, P., and Conti, B. (1997) Int. J. Pharm. 152, 237–246. Shi, X. Y., and Tan, T. W. (2002) Biomaterials 23, 4469–4473. Bayomi, M. A., Al-Suwayeh, S. A., and El-Helw, A. M. (1998) Pharm. Acta Helv. 73, 187–192. Becker, H., and Locascio, L. E. (2002) Talanta 56, 267–287. Eddington, D. T., and Beebe, D. J. (2004) Adv. Drug Deliv. Rev. 56, 199–210. Christopher, G. F., and Anna, S. L. (2007) J. Phys. D Appl. Phys. 40, 319–336. Teh, S. Y., Lin, R., Hung, L. H. and Lee, A. P. (2008) Lab Chip 8, 198–220. Yang, C. H., Lin, Y. S., Huang, K. S., Huang, Y. C., and Wang, E. (2009) Lab Chip 9, 145–150. Xu, J. H., Zhao, H., Lan, W. J., and Luo, G. S. (2012) Adv. Healthc. Mater. 1, 106–111. Wang, W., Yang, C., Cui, X. Q., Bao, Q. L., and Li, C. M. (2010) Microfluid. Nanofluid. 9,1175–1183. Bradford, M. M. (1976) Anal. Biochem. 72, 248–254. Saisuburamaniyan, N., Krithika, L., Dileena, K. P., Sivasubramanian, S., and Puvanakrishnan, P. (2004) Anal. Biochem. 330, 70–73. Wang, S. G., Zhang, W. D., Li, Z., Ren, Z. Q., and Liu, H. X. (2010) Appl. Biochem. Biotechnol. 162(7), 2015–2026. Gao, S. L., Wang, Y. J., Diao, X., Luo, G. S., and Dai, Y. Y. (2010) Bioresour. Technol. 101, 3830–3837. Jiang, D. S., Long, S. Y., Huang, J., Xiao, H. Y., and Zhou, J. Y. (2005) Biochem. Eng. J. 25, 15–23. Changa, M. Y., and Juang, R. S. (2007) Biochem. Eng. J. 35, 93–98. Abdel-Naby, M. A. (1993) Appl. Biochem. Biotechnol. 38, 69–81.
Immobilization of Lipase
Yang Chen ∗ Junteng Liu Chunjie Xia Chenxi Zhao Zhongqi Ren Weidong Zhang
Beijing Key Laboratory of Membrane Science and Technology, Beijing University of Chemical Technology, Beijing, People’s Republic of China
Abstract Porous monodisperse chitosan microspheres were synthesized for enzyme immobilization. The microspheres were prepared using microchannels and modified with glutaraldehyde. The microspheres had a mean diameter of 495 µm; the polydispersity indices were less than 0.08, and the specific surface area was between 121 and 173 m2 /g. Candida sp. 1619 lipase was selected as a model lipase. Immobilization conditions such as enzyme loading, glutaraldehyde
concentration, and immobilization time were optimized. The temperature, pH, and storage stability of the free and immobilized enzymes were also investigated. The immobilized enzyme had broad-ranging pH and temperature optima as compared with free enzyme. The storage stability of the immobilized enzyme was higher than that of the free enzyme. C 2014 International Union of Biochemistry and Molecular Biology, Inc. Volume 00, Number 0, Pages 1–6, 2014
Keywords: chitosan, immobilized enzyme, lipase activity, monodisperse, porous microspheres, protein loading
1. Introduction The specificity and catalytic performance of enzymes such as their hydrolysis, esterification, and aminolysis [1, 2] make them particular industrial biocatalysts in biochemical, biomedical, and food industrial processes [3]. However, natural enzymes rarely have the characteristics of industrial catalysts. Enzymes are often difficult to recover and reuse as catalysts from reaction solutions or from substrates and products [4]. Enzyme immobilization is the most useful way to improve the catalytic performance of enzymes in nonnatural environments, providing many important advantages over the use of natural enzymes; these advantages include enzyme activity, stability, reusability, continuous operation, and process economics [5]. Enzyme immobilization has been researched using
∗ Address for correspondence: Junteng Liu, PhD, Associate Professor, Beijing Key Laboratory of Membrane Science and Technology, Beijing University of Chemical Technology, PO Box 1, No. 15, N. 3rd Ring Rd. East, Beijing 100029, People’s Republic of China. Tel.: +86-10-6442-3628; Fax: +86-10-6443-6781; e-mail: [email protected]. Received 24 December 2013; accepted 7 May 2014
DOI: 10.1002/bab.1242 Published online in Wiley Online Library (wileyonlinelibrary.com)
various carriers, such as gel [6], inorganic materials [7], and polymer [8]. Chitosan is a natural acetylated glucosamine biopolymer. Given its hydrophilicity and biocompatibility, chitosan has attracted considerable attention in the development of enzyme immobilization [9]. Chitosan also has many hydroxyl and amino groups, which produce benefits when linked to enzymes [10]. As an enzyme carrier, chitosan has various molecular forms, such as fibers, membranes, and beads. Because of their high porosity and large specific surface area, porous chitosan microspheres offer low mass transfer resistance and small steric hindrance to enzymes within their three-dimensional polymer networks. When enzymes are immobilized by coupling to porous particles, diffusion limitations and catalytic activity decrease caused by immobilization are principally dependent on their structure, such as the sphere diameter and pore size [11]. The catalytic activity of immobilized enzymes increases as the sphere diameter decreases [12]. However, the diameter of particles is not always “the smaller, the better” under different situations; especially in large-scale industrial manufacturing, the immobilized enzyme is filled into columns or packed-bed reactors [8]. The immobilized enzyme is difficult to separate from products and to reuse if the particle size is too small. Therefore, micrometer-sized particles are especially suited for industrial applications with their many active sites and large specific surface area to fix enzyme molecules [13, 14].
1
Biotechnology and Applied Biochemistry Several methods to prepare chitosan microparticles have been reported, including emulsion techniques [15], spraydrying [16], and coacervation [17]. However, the size and size distribution of microspheres are difficult to control by these methods. The weakness of nonuniform sphere sizes with wide size distribution may limit practical applications. Thus, a reproducible technique for producing monodisperse chitosan microspheres should be developed to ensure that every batch of immobilized enzymes has consistent application value. The microfluidic chip is an emerging tool for manufacturing microparticles with precisely controlled and monodisperse size distributions [18–21]. Microfluidic methods have been developed as novel approaches for the controllable synthesis of monodisperse microspheres. Several research groups [22–24] have utilized such techniques to produce chitosan microspheres. Monodisperse chitosan microspheres with controlled sizes (100–500 µm in diameter) and narrow size distribution (a variation of less than 10%) have been successfully synthesized. Pore structure considerably influences immobilization and sphere size. In the current study, cross-linking was used to control pore structure. Enzyme immobilization on such cross-linked support does not require chemical activation because the cross-linker, normally a bifunctional reagent, has two functions: cross-linking and activation. Glutaraldehyde is often used as a cross-linking and surface-activating agent because of its reliability and ease of use and, more importantly, because of the availability of amino groups for the reaction with glutaraldehyde not only on enzymes but also on chitosan. In the present work, chitosan microspheres were prepared by microchannel and modified with glutaraldehyde. The characteristics of porous chitosan microspheres were studied. Candida sp. 1619 lipase was selected as a model lipase for immobilization on microparticles. To maximize the adsorption and expression of the enzymes, immobilization conditions were optimized in terms of immobilization time, enzyme loading, and glutaraldehyde concentration. In addition, the thermal, pH, and storage stability of the free and immobilized lipases were evaluated.
2. Experiment 2.1. Materials Candida sp. 1619 lipase was obtained from CTA New Century Biotechnology Co. (Beijing, People’s Republic of China). Bovine serum albumin was obtained from Beijing Jingke Chemical Reagent Corporation (Beijing, People’s Republic of China). Chitosan (degree of deacetylation = 90%) was obtained from Sinopharn Chemical Reagent Co. (Shanghai, People’s Republic of China). Liquid paraffin, glutaraldehyde, Span80, and Tween20 were obtained from Fuchen Chemical Reagent Co. (Tianjin, People’s Republic of China). Oleic acid and olive oil were obtained from Beijing Chemical Reagents Company (Beijing, People’s Republic of China). Coomassie brilliant blue G-250 was obtained from Fluka Corporation (Buchs,
2
Switzerland). All other reagents were commercially available and of the highest grade.
2.2. Preparation of chitosan microspheres Chitosan solution was prepared by initially dissolving 0.5 g of chitosan in 50 mL of an aqueous 1% (v/v) acetic acid solution. Liquid paraffin was used as the oil phase. These fluids were injected into a microchannel at fixed flow rates by syringe pumps (LSP02-1B, Longer Precision Pump Co., Hebei, People’s Republic of China). The chitosan aqueous solution (the disperse phase) was sheared by the liquid paraffin (the continuous phase), developing a series of monodisperse spherical structures called water-in-oil emulsions. When these emulsions were introduced to a 1 mol/L NaOH solution through a Teflon tube, they reacted with OH− at the interface and instantly formed chitosan microparticles. Then, these microparticles underwent cross-linking and modification when they were introduced to a glutaraldehyde solution. After 4 H of hardening, chitosan hydrogels were observed.
2.3. Lipase immobilization Candida sp. 1619 lipase was immobilized on chitosan microparticles by adsorption and glutaraldehyde modification. A certain quantity of lipase was added to sodium phosphate buffer (0.2 mol/L, pH 7.0) to prepare the lipase solution. The solution was then stored at 4 ◦ C for about 6 H until the lipase was completely dissolved. Dried chitosan particles (0.5 g) were suspended in lipase solution for a preset time at room temperature. Then immobilized microparticles were filtered and washed by sodium phosphate buffer (0.2 mol/L, pH 7).
2.4. Analytical methods 2.4.1. Determination of protein loading Protein loading was calculated according to difference in protein content between the original and filtered lipase solution. The protein concentration was estimated by Bradford’s method, using Coomassie brilliant blue G-250 [25]. Bovine serum albumin was used to establish the standard curve.
2.4.2. Determination of immobilized lipase activity Lipase activity was assayed by modified copper soap colorimetry [26]. One unit of lipase activity is defined as 1 µmol of free fatty acid released per minute per gram of immobilized lipase [27]. The substrate, which comprised 3 mL of phosphate buffer (pH 7.0) and 1 mL of olive oil, was pretreated for 5 Min at 37 ◦ C. Afterwards, immobilized lipase was added to the substrate, and reaction was started. Eight milliliters of methylbenzene was used to terminate the reaction after 15 Min. Lipase activity was analyzed by a spectrophotometer at 710 nm after 1 mL of copper reagent [(CH3 COO)2 Cu] was added to the methylbenzene phase. Residual activity was defined as the ratio of the activity of immobilized lipase to the activity at optimum reaction conditions. Each data was the average of at least three parallel experiments, and the standard deviation was within ±5%. The value for the average activity of the placebo version of the substrate and chitosan microspheres was consistently
Immobilization of Lipase
Particle size distributions.
FIG. 1
FIG. 3
low but was subtracted from the resulting values for each measurement of the free and immobilized lipase.
3. Results and Discussion 3.1. Characteristics of porous chitosan microspheres Porous monodisperse chitosan microspheres were prepared by microchannel and cross-linked with glutaraldehyde. The mean diameter of the prepared microspheres was 495 µm, and the polydispersity indices were less than 0.08 (Fig. 1). As shown in Fig. 2a, the chitosan microparticles were porous particles. The specific surface area was measured using the Brunauer–Emmett–Teller method of adsorption of nitrogen gas. The specific surface area was between 121 and 173 m2 /g. The degree of lipase immobilization is determined by the specific surface area of the microspheres. In addition, the porosity of microparticles controls the rate of enzymatic bioconversions by controlling the diffusion of substrates and products through pores. A large specific surface area can reduce diffu-
FIG. 2
Effect of glutaraldehyde concentration on specific surface area of chitosan microspheres.
sional limitations and promote the expression of lipase activity. The concentration of the cross-linker determines the porous structure of microspheres and requires optimization. As shown in Fig. 3, the specific surface area of the chitosan microspheres increased as glutaraldehyde concentration was increased from 1% to 9% (v/v). The surface of a chitosan microsphere with lipase immobilization is shown in Fig. 2b, in contrast with the surface of a chitosan microsphere without lipase immobilization (Fig. 2a). Some nanoparticles overspread the surface of the chitosan microsphere. Candida lipase has a molecular volume of 5 × 4.2 × 3 nm3 and a molecular mass of 45,000 to 60,000 Da [28]. The lipase is adsorbed onto the porous scaffolds.
3.2. Effect of immobilization time on lipase immobilization The time required for immobilizing lipase was optimized for the chitosan microparticles. As shown in Fig. 4, the activity of
Scanning electron microscope images: (a) surface of freeze-dried chitosan microsphere and (b) surface of freeze-dried chitosan microsphere with lipase loaded.
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FIG. 5 FIG. 4
Effect of immobilizing time on the activity of immobilized lipase (lipase concentration was 15 g/L, lipase solution was 20 mL, and glutaraldehyde concentration was 7%).
the immobilized lipase increased as the immobilized time was increased. Because the cross-linked enzyme creates a monolayer of enzymes on the microsphere surface, activity increased when immobilization time was increased. The immobilized enzyme exhibited maximal activity after 9 H of immobilization, and extended immobilization time reduced the lipase activities. With extended immobilization time, more enzyme molecules were immobilized on the microspheres, and the excessively immobilized enzyme can make the aperture of the microspheres relatively thin, lowering the accessibility of the substrate to the active sites [29]. Thus, the optimal immobilizing time was found to be 9 H.
3.3. Effect of enzyme loading on lipase immobilization Chitosan microparticles were loaded with lipase solution (20 mL) at different lipase concentrations, and protein loading and lipase activity was evaluated. As shown in Fig. 5, protein loading increased when lipase concentration increased from 5 to 30 g/L. However, lipase activity decreased when the lipase concentration exceeded 15 g/L. Presumably, more enzyme molecules went inside the microspheres after the active sites of the carriers were saturated. The enzyme molecules cannot efficiently perform catalysis for the limitation of substrate diffusion. Meanwhile, enzymes can be hidden and some active sites can be damaged in a high-immobilization enzyme system [29]. To maintain a balance of the protein loading and enzyme activity, the optimal enzyme concentration was 15 g/L.
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FIG. 6
Effect of enzyme concentration on the enzyme loading amount and the activity of immobilized lipase (immobilizing time was 9 H, lipase solution was 20 mL, and glutaraldehyde concentration was 7%).
Effect of glutaraldehyde concentration on the enzyme loading amount and the activity of immobilized lipase (immobilizing time was 9 H, lipase concentration was 15 g/L, and lipase solution was 20 mL).
3.4. Effect of glutaraldehyde concentration on lipase immobilization Glutaraldehyde is not only the cross-linker but also the denaturant. Thus, glutaraldehyde concentration can directly affect the specific surface area of chitosan microspheres as well as the activity of immobilized lipase. As shown in Fig. 6, when the glutaraldehyde concentration was lower than 7%, the activity of immobilized enzyme kept pace with the glutaraldehyde concentration. However, when the concentration was higher than 7%, lipase activity decreased, because not enough glutaraldehyde participates in
Immobilization of Lipase
FIG. 7
Effect of temperature on the activity of free and immobilized enzymes.
FIG. 8
Effect of pH on the activity of free and immobilized enzymes.
FIG. 9
Effect of storage on the activity of free and immobilized enzymes.
the cross-linking reaction when the glutaraldehyde concentration is lower than 7%. However, when the concentration is too high, glutaraldehyde is involved in aldol condensation, which can affect the formation of pores on the surface of microspheres and thus make immobilization harder. In addition, as glutaraldehyde concentration exceeds 7%, the extensive interaction of individual enzymes with aldehyde groups on the surface of the microspheres possibly changes the enzyme conformation and lowers the enzyme activity [30]. Thus, the optimal concentration of glutaraldehyde was found to be 7%.
3.5. Thermal stability The effect of temperature on the activity of free and immobilized lipase was studied by measuring residual enzyme activity from 27 to 67 ◦ C. As shown in Fig. 7, the optimal temperature of free and immobilized lipase was 37 ◦ C. The influence of temperature on immobilized lipase was much smaller than that on free lipase. At 67 ◦ C, the free lipase retained only 18.9% of residual activity, whereas the immobilized lipase retained 63.3% of its initial activity. The immobilization step increases enzyme’s rigidity, generally reflected by an increase in stability toward denaturation by raising the temperature [31]. Thus, this immobilization method enhanced the thermal stability of the lipase.
immobilized lipase still retained 71.2% of its initial activity. Thus, this immobilization method optimized the pH stability of the lipase.
3.6. pH stability
3.7. Storage stability
The effect of pH on the residual activity of free and immobilized lipase was studied at pH values between 5 and 10 at 37 ◦ C (Fig. 8). The optimal pH for free lipase was 6, and that for mobilized enzymes was 7. As expected, the immobilized lipase exhibited a wider pH range of activity than the free lipase. The immobilization process advanced the pH stability of lipase activity, especially in the alkaline condition. At pH 10, the free lipase retained only 12.8% of residual activity, whereas the
Storage stability for the immobilized enzyme is a significant index for evaluating the properties of the enzyme, making the immobilized enzyme more advantageous than the free one. Free and immobilized enzymes were stored in sodium phosphate buffer (0.2 mol/L, pH 7.0) at 4 ◦ C for 30 days. Aliquots of these lipases were periodically withdrawn for assay. Figure 9 shows the effect of storage on the activity of free and immobilized enzymes. After 7 days, the activity of the free
Biotechnology and Applied Biochemistry
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Biotechnology and Applied Biochemistry enzyme decreased to less than 20% of the initial value, whereas the immobilized enzyme still retained about 80% of its initial activity. The immobilized enzyme exhibited 50% of its initial activity only after about 30 days. However, the activity of the free enzyme decreased to less than 10% at that time.
[5] [6] [7] [8] [9] [10]
4. Conclusion Chitosan microspheres were prepared by microchannel and modified with glutaraldehyde. The mean diameter of the prepared microspheres was 495 µm, and the polydispersity indices were less than 0.08. The specific surface area of the microsphere was between 121 and 173 m2 /g. Candida sp. 1619 lipase was selected as the model lipase to study immobilization on the microspheres. Immobilization conditions were optimized in terms of immobilization time, enzyme loading, and glutaraldehyde concentration. The optimal immobilizing time, enzyme loading, and glutaraldehyde concentration were 9 H, 15 g/L, and 7%, respectively. The immobilized lipase had a wide range of pH and temperature optima. The storage stability of the immobilized enzyme was higher than that of the free enzyme. This method can improve controlled-release delivery systems, and the preparation of monodisperse chitosan microparticles will provide many potential biomedical applications.
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Immobilization of Lipase
