Modified chitosan microspheres in non-aggregated amylase immobilization

International Journal of Biological Macromolecules 66 (2014) 46–51

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International Journal of Biological Macromolecules journal homepage: www.elsevier.com/locate/ijbiomac

Modified chitosan microspheres in non-aggregated amylase immobilization Medha Rana a , Amita Kumari b , Ghanshyam S. Chauhan c , Kalpana Chauhan a,∗ a b c

School of Chemistry, Shoolini University, Solan 173229, India School of Botany, Shoolini University, Solan 173229, India Department of Chemistry, Himachal Pradesh University, Shimla 171005, India

a r t i c l e

i n f o

Article history: Received 13 August 2013 Received in revised form 4 February 2014 Accepted 10 February 2014 Available online 18 February 2014 Keywords: Immobilization Amylase Microspheres Chitosan

a b s t r a c t Immobilized enzymes are useful as reusable catalysts in industrial processes. In this study, ␣-amylase was used as a model enzyme to evaluate the propensity of synthesized porous chitosan microspheres as immobilization matrix. Chitosan microspheres were synthesized by grafting and covalent gelation technique using acrylamide (AAm) and glutaraldehyde (GA) as chemical agents, respectively. The synthesized chitosan-cl-poly(AAm) demonstrated amylase immobilization capacity of 350 mg/g. Furthermore, SEM results supported the porous microsphere structure for chitosan-cl-poly(AAm) with non-aggregated amylase immobilization, which accounts for comparable activity of immobilized amylase (3.28 ␮mol/ml/min) in contrast to free amylase (3.46 ␮mol/ml/min). The immobilized ␣-amylase was characterized for optimal pH and temperature activity and showed better resistance to temperature and pH inactivation in contrast to free amylase. The immobilized amylase retained more than 60% of its initial activity when stored at 4 ◦ C for 30 days and retained 50% of its initial activity after seven successive repeated-use cycles. In conclusion, the study can be used as base for the immobilization of competent industrial biocatalysts in non-aggregated active structure. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Amylases (␣-amylase, ␤-amylase and glucoamylase) are among the most important industrial enzymes of technological promise in the food, paper and textile industries [1], detergents [2] and analytical chemistry [3]. But, the industrial use of such biocatalysts in enzyme reactors has certain practical problems i.e., high cost of isolation, purification of enzymes, structures instability once they are isolated from their natural environments and short operational lifetimes due to reaction conditions (organic solvents, elevated temperatures and pH can also cause the loss of activity due to aggregation or conformation loss) [4]. Three dimensional structure and conformation plays a crucial role in determining the catalytic efficiency of enzymes, thus eventually influencing their exploitability in biotechnological applications. The stability and good activity of the enzyme is an important concern in industrial biotechnology [5]. Immobilization of biocatalysts on solid support is very important to stabilize the structure of enzymes and hence their activities.

∗ Corresponding author. Tel.: +91 9459368088; fax: +91 1792 226364/308000. E-mail addresses: [email protected], [email protected] (K. Chauhan). http://dx.doi.org/10.1016/j.ijbiomac.2014.02.022 0141-8130/© 2014 Elsevier B.V. All rights reserved.

Moreover, the heterogeneity of the immobilized enzyme systems facilitates easy recovery of enzyme and product, repetitive use of enzymes, continuous operation of enzymatic processes and decreased cost of processing. Enzyme immobilization on solid carriers is an important way to improve enzyme performance in industrial processes [6]. The physical and chemical characteristics (Pore size, hydrophilic/hydrophobic balance and surface chemistry) of support have been reported accountable in enzyme immobilization and its catalytic properties [7,8]. The mosoporous materials have been evaluated as most promising carriers for enzyme immobilization [9–12]. But, the synthetic and processing cost of mesoporous structure limits the applicability in the encapsulation of biomolecules [13,14]. Furthermore, the encapsulation in mesoporous structure may cause the aggregation of enzyme and failure of catalytic activity [15]. Chen et al. [16,17] have reported the significance of particulate support (particle size) in immobilization applications. Luo and Zhang [18] have reported the enhanced immobilization on microporous cellulose microspheres. An egg shell microporous membrane has also been reported effective in enzyme immobilization [19]. The chitosan have been extensively used due to the advantage of a great compatibility with the enzymes [20,21]. In view of that, the aim of this study was to find a simple and efficient

M. Rana et al. / International Journal of Biological Macromolecules 66 (2014) 46–51

HOH2C

HOH2C O

O HO

O

CH2OH

O

O

NH2 HO H2N OH HO

O

NH2

Spacer group

O

HO

O

H2N NH2

O

NH2

O

2-aminoglucopyranose unit

CH2OH O

H2N HO OHH2N

O HOH2C

O

HO

OH

HOH2C

CH2OH O

O

HOH2C

Amylase

CH2OH

O

OH

pore structure

O

N N

O

47

O

O

O CH2OH

Scheme 1. Proposed structure of chitosan-cl-poly(AAm) with immobilized ␣-amylase.

method, to stabilize ␣-amylase with high yields and catalytic activity. Chitosan is the only cationic polysaccharide in nature derived from biomass. Its proteinaceous nature makes it worthy in enzyme immobilization [22]. However, chitosan potential as immobilization matrix can be enhanced via chemical modifications to realize the full potential of this versatile polysaccharide. Moreover, chitosan is amenable to chemical modifications due to versatility of chemical structure i.e., –OH, –CONH2 and –NH2 functional groups. Therefore, tailor made supports for enzyme immobilization can be produced with desired properties and uniform porosity by GA as spacer group.

stirring using a mechanical stirrer for 6 h. Furthermore, the reaction setup was kept undisturbed for the next 24 h for completion and homogeneity of product by condensation reactions. The crosslinked microspheres were isolated by centrifugation and dried by vacuum till constant weight. The characterization of functionalized chitosan microspheres and enzyme loaded microspheres was carried out by X-ray diffraction method (XRD), Fourier transform infrared spectroscopy (FTIR) and scanning electron micrograph (SEM). FTIR spectra were recorded using KBr pellets on Perkin Elmer, SEMs on Joel JSM 6100 and XRD on JOEL-8030 X-ray diffractometer. 2.3. Immobilization of ˛-amylase

2. Experimental 2.1. Material ␣-Amylase (EC 3.2.1.1) was obtained from the HiMedia Laboratories. The other chemicals used for the study were of analytical grades and used as received. The weights were measured on Denver Balance having minimum readability of 0.01 mg. 2.2. Modification and characterization

2.4. Enzyme activity and reusability

The commercial chitosan was copolymerized with acrylamide (AAm) in the molar ratio of 1:1 using ammonium persulphate (APS, 2.0% (w/w) of chitosan and AAm) as initiator at 65 ◦ C. After 2 h, the product was collected and washed with water and alcohol separately, respectively. The product was dried to a constant weight for further use. The efficiency of the product formation was evaluated as follows [23]. %Grafting Efficiency(%E) =

W  d

Wr

× 100

To the 10.0 ml buffered amylase (100 mg) solution (pH 7.0, 0.1 M) was added 0.250 g of modified chitosan microspheres. The mixture was stirred at 20 ◦ C in a rotatory shaker for 24 h. The supernatant was used for protein quantification by Lowry method [24]. Enzyme-supported chitosan microspheres were separated by centrifugation, and the unbound enzyme was removed by washing with phosphate buffer (pH 7.0, 0.1 M) solution. The immobilized enzyme was dried at 30 ◦ C for further use as biological catalyst.

(1)

where Wd is the weight of the synthesized product (Chitosang-poly(AAm), where -g- is for grafting) and Wr is the weight of all reacting species including chitosan, AAm and APS. Chitosang-poly(AAm) based microspheres were synthesized by a covalent gelation technique using GA (10.0% (w/w) of chitosan and AAm). GA acts as a bifunctional crosslinker and in the process of crosslinking it generates pores by bridging the two chains of the chitosan through the amino groups of the later as shown in Scheme 1 [4] Chitosang-poly(AAm) (2.0% w/v) was dissolved in acetic acid (pH 5.0–6.0) and GA was added drop wise at room temperature with constant

The enzyme activity for free and immobilized enzyme was assessed via colorimetric analysis using 3,5-dinitrosalicylic acid (DNSA) reagent [25]. 3.5 mg of enzyme and its immobilized equivalent (in 10 mg of support) was added to 1.0 ml of 2.0% (w/v) starch solution in phosphate buffer (pH 7.0, 0.1 M), separately. The test sets were incubated at 35 ◦ C for 30 min. To the incubated reaction set up 1.0 ml of DNSA was added and boiled for 10 min for color development. The optical density (OD) of the solution was determined at 550 nm in the UV–vis spectrophotometer (UV mini 1240 spectrophotometer). The effect of temperature (35 ◦ C to 75 ◦ C) on enzyme activity was studied at 30 min with 2.0% (w/v) starch in phosphate buffer (0.1 M, pH 7.0). The effect of pH (4.5 to 10) on enzyme activity in free and immobilized state was also studied at optimum 55 ◦ C. The activity of free and immobilized amylase was calculated by using earlier reported formula [26]. For each test, the activity of enzyme at the beginning of reaction was taken as 100% residual activity [27]. The reusability for immobilized amylase was studied at 55 ◦ C and for 30 min with 2.0% (w/v) of starch in phosphate buffer (pH 7.0, 0.1 M). After each activity measurement, the immobilized enzymes

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M. Rana et al. / International Journal of Biological Macromolecules 66 (2014) 46–51

were recovered by simple filtration and washed with buffer solution (pH 7.0, 0.1 M) for use in second cycle and so on. The activity of the first cycle for starch hydrolysis was taken as 100% and further activity for the repeated use was calculated as relative activity. All of the experiments of activity were performed in triplicate, and the results were expressed as averages. 3. Results and discussion 3.1. AAm graft copolymerization and covalent gelation The free radical initiator APS decomposes at 65 ◦ C via anion radical formation those abstract hydrogen from the –OH moieties present in the C-6 position of chitosan unit. The free radicals on the chitosan biopolymer serve as macro-initiators for the AAm monomer, which ultimately results copolymer of chitosan-g-poly(AAm). The percent grafting was 97% for chitosan-g-poly(AAm). Furthermore, the copolymer of chitosang-poly(AAm) was guided into uniform porous microspheres by covalent gelation. The percent efficiency for the microspheres formation was 93.24%. The ionotropic/covalent gelation method is very simple and mild process used for designing nano and micro-carriers as possible candidates for immobilization with low processing cost [28,29]. GA is a homo-bifunctional crosslinking agent, which reacts with –NH2 functional group of chitosan in covalent gelation by condensation reactions [30–32]. Aging (higher reaction time) serves to increase the mechanical strength of the gel by increasing the extent of crosslinking due to extensive condensation reactions between GA and chitosan. It is therefore important that the degree of condensation be carefully controlled to obtain gels of desired morphology and porosity. The porous structure provides the less interactive sites (as porous structure can provide site/void for immobilization, which results limited interaction between enzyme and support) for amylase immobilization, which results the conformation stability and high catalytic activity of the enzyme. 3.2. Immobilization characterization The immobilization results show the efficiency of the applied protocol with the immobilization capacity of 350 mg/g (≥85%) of chitosan based microspheres. The enhanced immobilization capacity can be accounted for porous structure of synthesized chitosan microspheres [6], which enhance the surface adsorption or partial entrapment with greater interaction by providing larger surface areas. The proposed structure for porous chitosan microspheres with immobilized amylase is shown in Scheme 1. The uniform porosity provides space for non-aggregated amylase immobilization and prevents the aggregation of the amylase via non-covalent forces (e.g., van der Waals attraction or electrostatic force) or strong covalent bonds (e.g., disulfide bridges). The non-aggregation of amylase is important for conformation stability or enzyme activity [33]. The physical entrapment of enzymes to the matrix is preferred because of its catalytic stability due to the retention in conformation. Furthermore, one free –CHO group in GA may react to –NH2 groups present in the enzyme molecule to produce imines or Schiffbases (covalent immobilization) [30]. The point of attachment is scarce due to porosity (less surface interaction, as porous structure provide confinement of interaction to particular site instead of whole enzyme molecule), which can accounts the conformational integrity of the enzyme for higher catalytic activity [34]. 3.2.1. FTIR characterization Chitosan-cl-poly(AAm) shows characteristic peaks at 3433.95 cm−1 (for O–H stretching, due to the polymeric association), 1641.22 and 1564.47 cm−1 (for –CONH2 stretching or

Fig. 1. FTIR Spectra for (a) chitosan-cl-poly(AAm) (b) immobilized ␣-amylase.

–NH2 bendings), 1412.39 cm−1 (for –C–H bending of –CH2 in GA) and 800–1200 cm−1 (C–O and C–C stretching vibrations of the hexopyranosyl moiety) (Fig. 1a). The characteristic peaks of reference chitosan-cl-poly(AAm) are at slightly modified cm−1 in contrast to the amylase immobilized spectrum. Furthermore, amylase immobilized chitosan-cl-poly(AAm) shows an intense peak at 1655.23 cm−1 as a characteristic band of peptide bond in ␣-amylase (Fig. 1b). FTIR also confirms about the covalent binding of the enzyme as Schiff-base function (C=N) has a characteristic absorption peak at 1640 cm−1 [35,36] Additionally, spectrum not shows any band at ∼1720 cm−1 , related to the free aldehyde group. Some changes can be observed in amylase immobilized chitosancl-poly(AAm) spectrum, the peaks are less sharp and with some differences in relative intensity than that of chitosan-cl-poly(AAm). 3.2.2. SEM and XRD characterization Fig. 2a and b show SEM images for pure chitosan and confirms irregular or approximately spherical shape with 100 ␮m size distribution. Furthermore, Fig. 2c evaluate approximately spherical shape (shown in inset) for modified chitosan microspheres of about 1–10 ␮m average size and uniform porosity. At high magnification, SEM images demonstrate small outgrowths on the surface for poly(AAm) modification (Fig. 2c and d). SEM image of ␣-amylase immobilized chitosan-cl-poly(AAm) microspheres illustrates the physical characterization for enzyme adsorption, and the image is worthy to show clear adherence of individual enzyme (nonaggregated) to the surface and within the pores of the support (Fig. 2d and e). It shows that the synthesized chitosan-cl-poly(AAm) porous microspheres is suitable for the partial entrapment of the non-aggregated amylase with conformational retention for their better catalytic activity. The porous matrix can certainly enhance the activity of the ␣-amylase, because it can partly entrap the enzyme without aggregation and can also hold it for longer duration with retention of conformation. The XRD measurements were carried out in order to investigate the structural change caused by modification and enzyme immobilization. Fig. 3 illustrate the XRD patterns for the native chitosan, synthesized microspheres of functionalized chitosan and amylase immobilized chitosan. The XRD spectrum of pure chitosan shows two peaks over 2 = 10–12◦ and 2 = 19–23◦ . It means that the

M. Rana et al. / International Journal of Biological Macromolecules 66 (2014) 46–51

49

Fig. 2. SEM images for ((a) and (b)) chitosan (c) chitosan-cl-poly(AAm) with spherical shape in inset (d) immobilized ␣-amylase at 2000 magnification (e) Immobilized ␣-amylase at 6000 magnification.

native chitosan essentially shows the micro-crystalinity (Fig. 3a), while after functionalization the peak pattern shows increased crystalinity of the particulate product (Fig. 3b). XRD spectra of crosslinked chitosan with GA which has two additional broad peaks at 2 = 19.88◦ and 2 = 26.89◦ , indicates that the sample is going from micro-crystalinity to crystalinity nature. The shift in the 2 values confirms the effective and uniform crosslinking would have taken place, which account for increased crystalinity. The rigid crystalline structure of pure chitosan is stabilized by intra and intermolecular hydrogen bonds. XRD results support the size of 0.001–1.0 ␮m for chitosancl-poly(AAm) microspheres by Scherrer equation [37]. XRD of chitosan-cl-poly(AAm) shows a new peak with modified intensity,

which can be accounted worthy for uniformity of modification. The XRD result for amylase immobilized gels do not exhibit any wellresolved peaks (Fig. 3c), which confirm the intercalation of the enzyme into the porous structure of chitosan-cl-poly(AAm). The covalent binding with ␣-amylase can also lead to enzyme intercalation. The XRD values of the diffraction peaks are listed in Table 1. Table 1 Pore structure and surface properties of the chitosan-cl-poly(AAm). Pos. [◦ 2Th.]

FWHM [◦ 2Th.]

d-Spacing [Å]

Area [cts*◦ 2Th.]

19.8863 26.8972

0.8699 0.1673

4.46476 3.31481

447.19 17.71

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intercalated into the support. It is the part of side chains of various amino acid groups that are responsible for intercalation [40].

3.3. Activity of the immobilized amylase

Fig. 3. XRD of (a) chitosan (b) chitosan-cl-poly(AAm) (c) immobilized ␣-amylase.

Amylase is highly polymeric species of 8 nm molecular size, hence possibility of attachment or entrapment within inter lamellar space can be ruled out. But a shift in the peak values is an evidence for intercalation [38,39]. Moreover, the inter-layer spacing ranges between 3.3 and 4.4 A˚ in chitosan-cl-poly(AAm), which accounts for non-aggregated amylase immobilization. Therefore it is evident from SEM and XRD results that the whole enzyme does not get Free amylase Immobilized amylase

100

(a)

The enzymatic starch hydrolysis is used to determine the efficiency of the applied immobilization protocol by DNSA method. The DNSA assay is based on the reduction of DNSA by glucose to 3amino-5-nitrosalicylic acid, which has red–brown color. The immobilized enzyme shows the maximum activity of 3.28 ␮mol/ml/min in comparison to free amylase (3.46 ␮mol/ml/min) at 35 ◦ C. This drop in the catalytic activity can be attributed to steric hindrance from support in the immediate vicinity of the enzyme molecules, which alter the natural molecular environment of enzymes and render their active sites less available to the substrate. The optimum catalytic activity shift to 55 ◦ C for immobilized amylase, while it decreases to 46% relative activity for free amylase at 75 ◦ C (Fig. 4a). The pH dependence of the activity of free and immobilized ␣-amylase was investigated at optimum 55 ◦ C. The pH results show the optimum activity for free amylase at 4.5 [41], while the immobilization improves the pH stability of the enzyme at the pH values i.e., 7.0 and 8.5 (Fig. 4b) as indicated elsewhere also [42]. The immobilized enzyme retained 91% and 82% of relative activity at pH 8.5 and 10, respectively. The increased activities of immobilized ␣-amylase with temperature indicate that porous microspheres retain the conformation (tertiary structure) by providing the equivalent space for non-aggregated enzyme immobilization. The activity of entrapped enzyme was assayed for seven cycles in order to find out the reuse of the entrapped enzyme. The results show good relative activity (80%) till third reuse cycle, while further reuse causes loss of activity to 80–49% value. The activity remains (49%) even in the seventh cycle of reuse (Fig. 4c). The decrease in activity can be credited to the leakage of the enzyme from the microspheres, due to the washing of microspheres at the end of each cycle. Further, the immobilized enzyme shows the constant activity pattern even after fourth cycle, which can be attributed to the covalent immobilization of ␣-amylase. The immobilized

Free amylase Immobilized amylase

100

(b)

90

% (Relative activity)

(c)

100

80

90 80

60

80 70 70

40

50

60

20

40

50

0

60

30

40

50

60

70 o

Temperature ( C)

80

4

5

6

7 pH

8

9

10

1

2

3

4

5

6

Cycle number

Fig. 4. (a) Effect of temperature (b) effect of pH on catalytic activity profiles for free and (c) reusability potential for immobilized ␣-amylase.

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amylase retains 60% of its initial appreciable activity even after 30 days of storage. 4. Conclusions In the present work, inexpensive porous chitosan microspheres have been synthesized for immobilizing ␣-amylase with ≥85% efficiency. The SEM results confirm the uniformity in porosity of synthesized microparticles, which ultimately support the competence of the applied protocol. The enzyme immobilization (350 mg/g) is based on the partial entrapment of protein molecules into pores of support generated in the course of covalent gelation. The uniform porosity provides the constructive interaction for physical and covalent immobilization of amylase with preservation of conformation for comparable catalytic activity (3.28 ␮mol/ml/min), thermal stability (65 ◦ C) and pH tolerance (pH 7.4 and 9.0). The results are of technological importance for detergent industry due to the enhanced thermal stability and tolerance to basic pH. In the repeated-use experiments, the immobilized amylase retained 60% of its initial activity when stored at 4 ◦ C for 30 days and retained 49% of its initial activity after seven consecutive repeated-use cycles. Altogether, these results provide useful information on how supports and/or enzymes have to be tailored to improve biocatalyst performance. References [1] P.H. Pandya, R.V. Jarsa, B.L. Newalkar, P.N. Bhalt, Microporous and Mesoporous Materials 77 (2005) 67–77. [2] Q. Shao, S.P.S. Chundawat, V. Balan, Biotechnology for Biofuels 12 (3) (2010) 1754–1764. [3] L. Kandra, Journal of Molecular Structure 667 (3) (2003) 487–498. [4] M.J. Moura, H. Faneca, M.P. Lima, M.H. Gil, M.M. Figueiredo, Biomacromolecules 12 (9) (2011) 3275–3284. [5] L. Cao, R.D. Schmid, Carrier-bound Immobilized Enzymes: Principles, Application and Design, WILEY-VCH Verlag GmbH & Co., Weinheim, 2005. [6] E.T. Hwang, M.B. Gu, Engineering in Life Sciences 13 (1) (2013) 49–61. [7] T. Boller, C. Meier, S. Menzler, Organic Process Research and Development 6 (4) (2002) 509–519. [8] L. Raj, G.S. Chauhan, W. Azmi, J.-H. Ahn, J. Manuel, Bioresource Technology 102 (3) (2011) 2177–2184. [9] M.I. Kim, J. Kim, J. Lee, H. Jia, H. Na Bin, J.K. Youn, J.H. Kwak, A. Dohnalkova, J.W. Grate, P. Wang, Biotechnology and Bioengineering 96 (2007) 210–218. [10] M.C. Rosales-Hernandez, J.E. Mendieta-Wejebe, J. Correa-Basurto, J.I. VazquezAlcantara, E. Terres-Rojas, J. Trujillo-Ferrara, International Journal of Biological Macromolecules 40 (5) (2007) 444–448. [11] A. Wang, H. Wang, S. Zhu, C. Zhou, Z. Du, S. Shen, Bioprocess and Biosystems Engineering 31 (5) (2008) 509–517.

51

[12] A.S.M. Chong, X.S. Zhao, Catalysis Today 93-95 (2004) 293–299. [13] H.R. Luckarift, J.C. Spain, R.R. Naik, M.O. Stone, Nature Biotechnology 22 (2) (2004) 211–213. [14] C-H. Lee, T-S. Lin, C-Y. Mou, Nano Today 4 (2009) 165–179. [15] N. Miletic, A. Nastasovic, K. Loos, Bioresource Technology 115 (2012) 126–135. [16] B. Chen, E.M. Miller, L. Miller, J.J. Maikner, R.A. Gross, Langmuir 23 (2007) 1381–1387. [17] B. Chen, M.E. Miller, R.A. Gross, Langmuir 23 (2007) 6467–6474. [18] X. Luo, L. Zhang, Biomacromolecules 11 (2010) 2896–2903. [19] C.S. Pundir, M. Bhambi, N.S. Chauhan, Talanta 77 (2009) 1688–1693. [20] B. Krajewska, Enzyme and Microbial Technology 35 (2-3) (2004) 126–139. [21] A.N. Singh, S. Singh, N. Suthar, V.K. Dubey, Journal of Agricultural and Food Chemistry 59 (11) (2011) 6256–6262. [22] V. Belessi, R. Zboril, J. Tucek, M. Mashlan, V. Tzitzios, D. Petridis, Chemistry of Materials 20 (2008) 3298–3305. [23] G.S. Chauhan, S. Chauhan, S. Kumar, A. Kumari, Bioresource Technology 99 (2008) 6464–6470. [24] O.H. Lowry, N.J. Rosebrough, A.L. Farr, R.L. Randau, Journal of Biological Chemistry 193 (1951) 265–273. [25] G.L. Miller, Analytical Chemistry 31 (1959) 426–428. [26] R. Kumar, A. Kumar, K. Chauhan, R. Gupta, J.H. Ahn, G.S. Chauhan, Bioresource Technology 100 (2009) 1474–1477. [27] A.B. Salleh, A. Ebrahimpour, R.N.Z.R.A. Rahman, M. Basri, Bioresource Technology 102 (2011) 6972–6981. [28] S. Racovita, S. Vasiliu, M. Popa, C. Luca, Revue Roumaine de Chimie 54 (9) (2009) 709–718. [29] E. Biro, A.S. Nemeth, C. Sisak, T. Feczko, J. Gyenis, Journal of Biochemical and Biophysical Methods 70 (2008) 1240–1246. [30] M.R. El-Aassar, Journal of Molecular Catalysis B: Enzymatic 85–86 (2013) 140–148. [31] L. Betancor, F. Lopez-Gallego, A. Hidalgo, N. Alonso-Morales, G. Mateo, R. Fernandez-Lafuente, J.M. Guisan, Enzyme and Microbial Technology 39 (2006) 877–882. [32] N. Alonso, F. Lıopez-Gallego, L. Betancor, A. Hidalgo, C. Mateo, J. Guisan, Journal of Molecular Catalysis B: Enzymatic 35 (2005) 57–61. [33] S. Colombie, A. Gaunand, B. Lindet, Journal of Molecular Catalysis B: Enzymatic 11 (2001) 559–565. [34] H-L. Liu, W-J. Chen, S-N. Chou, Colloids and Surfaces B: Biointerfaces 28 (2003) 215–225. [35] J. Knaul, S.M. Hudson, K.A.M. Creber, Journal of Polymer Science Part B: Polymer Physics 38 (1999) 1079–1094. [36] J.D. Schiffman, C.L. Schauer, Biomacromolecules 8 (2007) 594–601. [37] K. Prabakaran, T.V. Thamaraiselvi, S. Rajeswari, Trends in Biomaterials and Artificial Organs 19 (2006) 84–87. [38] W. Jia, E. Segal, D. Kornemandel, Y. Lamhot, M. Narkis, A. Siegmann, Synthetic Metals 128 (2002) 115–120. [39] D. Lee, K. Char, Polymer Degradation and Stability 75 (2002) 555–560. [40] R.D. Gougeon, M. Soulard, M. Reinholdt, J. Miehe-Brendle, J.M. Chezeau, R. LeDred, R. Marchal, P. Jeandet, European Journal of Inorganic Chemistry 1366 (2003). [41] O. Prakash, N. Jaiswal, Biotechnology and Applied Biochemistry 57 (2010) 105–110. [42] A. Kumari, A.M. Kayastha, Journal of Molecular Catalysis B: Enzymatic 69 (1-2) (2011) 8–14.