Superior activities of lipase immobilized on pure and hydrophobic clay supports: Characterization and catalytic activity studies

Superior activities of lipase immobilized on pure and hydrophobic clay supports: Characterization and catalytic activity studies

Journal of Molecular Catalysis B: Enzymatic 97 (2013) 36–44 Contents lists available at ScienceDirect Journal of Molecular Catalysis B: Enzymatic jo...

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Journal of Molecular Catalysis B: Enzymatic 97 (2013) 36–44

Contents lists available at ScienceDirect

Journal of Molecular Catalysis B: Enzymatic journal homepage: www.elsevier.com/locate/molcatb

Superior activities of lipase immobilized on pure and hydrophobic clay supports: Characterization and catalytic activity studies R. Reshmi, S. Sugunan ∗ Department of Applied Chemistry, Cochin University of Science and Technology, Kochi 682022, Kerala, India

a r t i c l e

i n f o

Article history: Received 26 December 2012 Received in revised form 4 April 2013 Accepted 6 April 2013 Available online 31 May 2013 Keywords: Montmorillonite K-10 Candida rugosa lipase Immobilization Hydrolysis Kinetics

a b s t r a c t Lipase from Candida rugosa was immobilized on 3-aminopropyltriethoxysilane-modified montmorillonite K-10 (Mt–S) support via glutaraldehyde spacer (Mt–G). Simple physical adsorption technique was also employed to immobilize lipase on Mt. The supports as well as the novel immobilized biocatalysts were characterized by a combination of techniques, namely X-ray diffraction (XRD), nitrogen adsorption studies and Scanning Electron Microscopy (SEM). Using the hydrolysis reaction of p-nitrophenyl palmitate in aqueous and organic media, the properties of the immobilized lipases were assayed and compared with those of the free enzyme. The effects of reaction temperature, pH, thermal and storage stabilities of the immobilized lipases were evaluated. The optimum reaction temperature rose from 35 ◦ C to 50 ◦ C for Mt–I and to 55 ◦ C for Mt–G. The covalently immobilized conjugate, Mt–G retained almost 90% activity at 50 ◦ C, while the free enzyme retained only 20% activity after 120 min heat treatment. Immobilized lipases exhibited enhanced storage stability than the native lipase (up to 40 days). The kinetic parameters of the free and immobilized lipases, Km and Vmax were also assayed. The activity of the free lipase in heptane (0.213 U/mg) was 0.51% of that in the aqueous medium (41.6 U/mg). The Km value for the free lipase was nearly 50-fold higher in organic media. The free lipase was 195-fold more active in water compared to that in organic solvent. © 2013 Published by Elsevier B.V.

1. Introduction Of all the clay minerals, smectites are the most chemically susceptible for modification and application. Nanoclays, have emerged as alternative supports for enzyme immobilization, since they can enhance the biocatalytic efficiency by reducing diffusional limitations as well as by increasing enzyme loading because of superior specific surface area per weight unit [1,2]. Clays and related materials have potentially interesting properties such as hydrophobic/hydrophilic behavior, electrostatic interactions, layered structure, swelling capability, ion exchange ability, mechanical and thermal stability and bacterial resistance [3]. Another advantage is the presence of silanol groups that, after activation by different functional groups [4] act as attachment sites for bioactive species. Clays possess high specific surface area available between 200 and 800 m2 /g. The facility of water dispersion-recuperation has three kinds of entities: (i) the neutral siloxane surface, (ii) reactive OH groups and (iii) permanent charged sites resulting from isomorphic cation substitutions. The

∗ Corresponding author. Tel.: +91 484 2575804; fax: +91 484 2577595. E-mail address: [email protected] (S. Sugunan). 1381-1177/$ – see front matter © 2013 Published by Elsevier B.V. http://dx.doi.org/10.1016/j.molcatb.2013.04.003

non-polar portion of larger biological molecules can efficiently be bound to this type of surface through van der Waals forces. Immobilization of enzymes on clays has been studied for a long time and it is still of great interest [1,5,6]. Moreover, new methods of immobilization have been developed, such as immobilization within a phyllosilicate sol–gel matrix [7] or composite chitosan-clays [8]. The hydrophilic nature of the parent clays can be converted into hydrophobic ones by replacing the natural exchangeable cations with cationic organic molecules forming the so-called “organoclays”. Hydrophobic modification of the clay intrasurface allows many hydrophobic guest molecules to be readily intercalated. The study of clays became even more interesting because of its cheapness. Nanoscale zerovalent iron (NZVI) supported on poly (hydroxo Al (III)) cations-pillared bentonite (Al-bent) resulted in the enhanced nitrate removal [9], while NZVI supported on hydrophobic organobentonite showed enhanced removal of organic contaminants [10,11]. Clays may constitute half or more of the solid fraction of soils and are known to catalyze numerous transformations of organic compounds when those are sorbed on their surfaces [12,13]. Fusi et al. [14] reported that montmorillonite may adsorb enzyme molecules on both external and internal surfaces. Naidja and Huang [15] found that the large molecules of aspartase (MW

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180,000) were intercalated within the montmorillonite layers. On the other hand, Harter and Stotzky [16] reported that the adsorption of catalase (MW 238,000) onto Ca-montmorillonite did not result in expansion of the mineral structure. In the work carried out by Gopinath et al. [17], it was proposed from surface area measurements, that the enzymes ␣-amylase, glucoamylase and invertase are situated at the periphery of the clay mineral particles whereas the side chains of different amino acid residues penetrated between the layers. The covalently bound invertase on montmorillonite K-10 resisted leaching even after 15 cycles due to the stronger bond between the grafted groups and the enzyme, while the adsorbed systems were prone to leaching [18]. The enzyme activity and the activity recovery of lipase immobilized on PAPTES were found to be the highest when lipase was immobilized onto three different modified palygorskite supports [19]. Lipases (triacylglycerol ester hydrolases, E.C.3.1.1.3) have been classified as enzymes that hydrolyze fats and oils with subsequent release of free fatty acids, diacylglycerols, monoacylglycerols and glycerol [20]. At present, the main disadvantage for the use of lipase enzyme in industrial production is the cost of the enzyme. The use of immobilized enzyme is thus a valid approach because it would allow the reutilization of the enzyme. Candida rugosa stands out due to its wide application in oil hydrolysis, transesterification, esterification and enantioselective biotransformation [21]. The demand for lipase in industry is due to its high activity and low price [22,23]. The reaction catalyzed in organic media (esterification or transesterification) is usually different from the one in aqueous media (hydrolysis). This fact makes these comparisons difficult because the transfer reaction was carried out in an organic solvent. Direct comparisons of enzyme activity in aqueous and organic media are scarce in the literature [24]. A simple colorimetric assay for the determination of lipase activity in aqueous and organic media using hydrolysis of p-nitrophenyl fatty acid ester was adopted in this study and this makes the activity comparison much easier. A relatively hydrophobic solvent (n-heptane) (i.e., with a high value of log P) was selected to maintain the maximum amount of water in the enzyme solid phase and to obtain a good solubilization of the substrate. In the present work, organosilanes were covalently bonded to the clay surfaces via condensation reaction with the surface silanol groups (Si OH) which resulted in a more tight interactions between organic components and clay. To prevent enzyme leaching, the silanized derivative was stabilized by cross linking with glutaraldehyde. Immobilization of lipase from C. rugosa was carried out by adsorption on pure montmorillonite (Mt) and on the above organically modified clays (Mt–G). The covalently bound systems showed remarkable activity and stability compared to physically adsorbed biocatalysts. Currently, strong interests in such supports are due to ecofriendly demands in many modern industrial applications. To the best of our knowledge, there are no reports on the immobilization of lipases on modified montmorillonite and its detailed physicochemical characterization. The lipolytic activity, kinetic characteristics and storage stability of the montmorillonite-immobilized lipases were investigated. The properties of the free and immobilized lipase preparations were compared in aqueous and organic media respectively.

2. Experimental 2.1. Materials Lipase (from C. rugosa, lyophilized powder) and montmorillonite K-10 were supplied by the Sigma Chemical Co. and used as received. Bovine serum albumin 99% (BSA), Gum arabic, Triton X-100, p-nitrophenyl palmitate (p-NPP) and

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glutaraldehyde were purchased from Sigma-Chemical Co. Heptane was procured from Merck. 2.2. Characterization of support Powder XRD of the immobilized systems and the pure clay support were taken on a Rigaku D/Max-C system with ˚ within the 2 range Ni filtered CuK␣ radiation ( = 1.5406 A) 2◦ –15◦ at a scanning rate of 0.5◦ /min at room temperature. A Micromeritics 2000 surface area analyzer was used to measure the nitrogen adsorption isotherms of the samples at liquid nitrogen temperature. Prior to the measurement, the clay samples were degassed at 200 ◦ C and the immobilized samples were degassed at room temperature overnight in nitrogen flow. The scanning electron microscopy of the samples were carried out on a JOEL JSM 840 A (Oxford make) model 16,211 SEM analyzer with a resolution of 1.3 eV. The samples were dusted on a metal stab and coated with a layer of gold to minimize charge effects. 2.3. Preparation of clay supports 2.3.1. Functionalization of the supports The surface of clay sample was amino functionalized by reacting 1 g of solid with 50 ml of a 2.2 mmol solution of Amino propyltriethoxy silane (APTES) in molecular sieve 4A dried toluene (v/v) at 373 K for 6 h with stirring [25]. Products were separated by filtration, washed with dry toluene and methanol. It was also washed with dichloromethane by Soxhlet extraction for 18 h and dried at ambient temperature. Following this, 1 g of amino functionalized support was reacted with 25 ml of a 2.5% solution of glutaraldehyde in phosphate buffer (pH = 6.62) for 4 h at room temperature. The product was washed exhaustively with distilled water till all excess glutaraldehyde was removed which was tested by Tollen’s reagent method and then dried at room temperature. 2.3.2. Immobilization procedure In aqueous environment, the enzyme was dissolved in 100 mM sodium phosphate buffer (4 mg/ml, pH 7.0) and mixed with 100 mg of the support. The immobilization process was carried out at 4 ◦ C in a shaking water bath for 16 h under low stirring at room temperature. After this, the supports were thoroughly washed with phosphate buffer (0.1 M, pH 7), dried at room temperature and held at 4 ◦ C for further use. 2.3.2.1. Protein assay. Protein was determined according to Lowry et al. [26] using bovine serum albumin (BSA) as a standard. The amount of bound protein was determined indirectly by the difference between the amount of protein introduced into the coupling reaction mixture and the amount of protein in the filtrates. The immobilization capacity of the protein on the support was defined as the amount of protein (mg) per gram of the support. 2.4. Substrate preparation and lipase assay 2.4.1. Activity assay of lipase in aqueous media A modified version of the procedure by Winkler and Stuckmann [27] based on the hydrolysis of p-NPP was used throughout this study to measure lipase activity. The stock substrate solution was prepared as per the method of Kordel et al. [28] with slight modifications, by dissolving p-NPP in 2-propanol to obtain a 16.5 mM stock solution. One milliliter of the stock substrate solution was added to 9 ml of 0.01 M phosphate buffer, containing 0.4% (w/v) Triton X-100 and 0.1% (w/v) arabic gum, pH 7.0. Lipase activity was assayed spectrophotometrically by measuring the rate of hydrolysis of p-NPP at 410 nm and 37 ◦ C in a UV spectrophotometer. The reaction mixture composed of 900 ␮l of

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freshly prepared substrate solution and 100 ␮l of the enzyme extract. Enzyme activity was expressed as unit/ml (U/ml), where one unit of activity was defined as the amount of enzyme that catalyzed the release of 1 ␮mol of p-nitrophenol (p-NP) per min under the assay conditions. The extinction coefficient of p-NP at pH 8.0 was determined as 17,500 M−1 cm−1 /l. 2.4.2. Activity assay of lipase in the organic medium The reaction rate of the free and immobilized lipase preparations in heptane was determined according to the process described by Pencreac’h and Baratti [29]. In the standard conditions, the reaction mixture was composed of 2 ml of n-heptane containing 10 mM p-NPP in an Erlenmeyer flask. The reaction was started by the addition of 10 mg free lipase preparation (or 50 mg immobilized lipase preparation). The mixture was incubated at 37 ◦ C. After 5 min of reaction, agitation was stopped, the lipase powder was allowed to settle for 30 s, and the clear supernatant was withdrawn. Fifty microlitres of supernatant was immediately mixed with 1.0 microlitres, 10 mM NaOH, directly in 1.0 ml cuvette of the spectrophotometer. The p-NP was extracted by the aqueous alkaline phase. It displayed a yellow color because of the alkaline pH. The absorbance was read at 410 nm against a blank without enzyme and treated in parallel. Molar extinction coefficient of 17.4 × 103 M−1 cm−1 for p-NP was estimated using the standard solutions of p-NP in n-heptane. 2.5. Biochemical characterization of the immobilized supports 2.5.1. Immobilization yield, activity yield, catalytic efficiency and effectiveness factor Immobilization yield (IY) (%) was calculated according to the following equation: IY (%) =

A−B A × 100

where A is the total activity of enzyme added in the initial immobilization solution; B, the activity of the residual enzyme in the immobilization and washing solutions after the immobilization procedure. The amount of protein in the enzyme solution and in the elution solutions was determined by the Lowry method [26], and the amount of protein (p) bound on the carriers was calculated from the formula p=

(Ci − Ct )V W

where p is the amount of bound enzyme onto carriers (mg/g), Ci and Cf are the concentrations of the enzyme protein initial and final in the reaction medium (mg/ml), V is the volume of the reaction medium (ml), W is the weight of the carriers (g). The catalytic efficiency of an enzyme is determined by maximum specific constant Kcat /Km (M−1 s−1 ). The effectiveness factor (EF) was used as a comparison parameter for the immobilized system. The formula for ‘EF’ is given by EF =

Vmax(immobilized enzyme) Vmax(free enzyme)

The activity yield (AY%), is defined as follows: AY% = Uads /Ueq × 100; where Ueq are the equilibrated enzyme units, Uads are the adsorbed units measured as the difference between Ueq and units remaining in the supernatant, and Uact is the active adsorbed activity.

The relative activity (Ra ) of the free and immobilized lipase was calculated according to the formula: Ra (%) =

A × 100 Am

where Ra is the relative activity of CRL (%); A is the activity of the immobilized enzyme (U g−1 ); Am is the activity of the free enzyme in solution (U g−1 ). 2.5.2. Effect of temperature on enzyme activity The hydrolytic activities of the lipases were determined at different temperatures (in the range between 30 and 70 ◦ C). Residual hydrolytic activity was measured using the method described above. The thermal stability assays were performed by incubation of the immobilized and free forms of CRL lipase at different temperatures (50 and 55 ◦ C) for different time intervals, cooled down to room temperature and the activity of enzyme was measured as described above. The initial activity of freshly prepared immobilized lipase in the first run was defined as 100% activity. The relative activities were determined as percentage yield of activities at different temperature compared to the activity of reaction at 30 ◦ C as % activity at different temperature × 100 maximum % activity (30 ◦ C)

2.5.3. Kinetic parameters The kinetic parameters, Km and Vmax , of free and immobilized lipase were calculated from the Michaelis–Menten models via Lineweaver–Burk and linear regression with Sigma plot software (version 10; Systat Software Inc., Richmond, CA, USA) using varying concentrations of p-NPP (0.2–1.5 mM) in the aqueous medium, and from 1 mM to 50 mM in heptane. The catalytic efficiency (Vmax /Km ) for the hydrolysis of the p-NPP substrate was also determined. 2.5.4. Storage stability of free and immobilized lipase The activity of free and immobilized lipase was measured as fresh and after storage in phosphate buffer (50 mM, pH 7.0) at 4 ◦ C during 40 days in a batch operation mode with the experimental conditions given above. The relative activities of the enzyme were compared to the activity at day 1 as Relative activity (%) =

% activity at different incubation period × 100 maximum % activity (storage temperature)

3. Results and discussions 3.1. X-ray diffraction (XRD) analysis The parent montmorillonite clay gave a distinct diffraction peak around 2 equal to 8.88◦ , which corresponds to a basal spacing of 9.95 A˚ (Fig. 1(a)). After grafting with 3-aminopropyltriethoxysilane and glutaraldehyde (Fig. 2(b) and (c)) there was not much change in the d value (Table 1). Upon immobilization of lipase (Fig. 1(b) the d value remained unaltered which confirmed that the texture of Mt clay is maintained and the modification takes place only at the external surface, further supported by surface area measurements. Thus, the enzyme did not enter into the interlayer space and mostly inhabited on the external surfaces and at the edges of the interlayer sheets through hydrogen bonding, van der Waals and electrostatic forces of interactions. Similar results were obtained with lipase on modified bentonite with monolayer surfactant (BMS) and on bentonite with bilayer surfactant (BBS) and alkaline phosphatase on Na-sepiolite and modified sepiolite [30,31].

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Fig. 3. Nitrogen adsorption isotherms of (a) Mt and (b) MtI. Fig. 1. X-ray diffraction patterns of (a) Mt and (b) MtI.

Table 1 d values of the pure, functionalized and immobilized samples of clay. Sample

˚ d values (A)

Mt MtI Mt–S Mt–G

9.95 9.95 9.97 9.94

3.1.1. Nitrogen adsorption/desorption analysis Adsorption isotherm of montmorillonite K-10 clay belongs to the type II in the Brunauer, Deming, Deming and Teller (BDDT) classification, characteristic of nitrogen adsorption on macroporous adsorbents (with less or no porosity) (Fig. 3(a) and (b)). Furthermore, the hysteresis loops of these isotherms are assigned to type H4 in the IUPAC classification [32], which is representative of the slit-shaped pores in layered materials. A wider pore size distribution is depicted in Figs. 4 and 5. After adsorption of lipase, the amount of N2 adsorbed decreased while there was not much change in the P/P0 value (Fig. 3(b)), which confirmed that the adsorption was entirely external and no intercalation took place in the clay which was also evident form the XRD results (Table 2). There was not much shift in the pore size distribution curves after immobilization (Fig. 5(b)). After functionalization with silane and glutaraldehyde, the surface area decreased with no appreciable

decrease in the pore diameter and pore volume (Fig. 6(a)–(c)). As there was no intercalation upon functionalization it is clear that the binding of organic groups is entirely external. Therefore it can be concluded that the texture of the montmorillonite was maintained and 3-aminopropyltriethoxysilane was connected only with the surface of clay.

Fig. 2. X-ray diffraction patterns of (a) Mt, (b) Mt–S and (c) Mt–G.

Fig. 5. Pore size distribution of (a) Mt and (b) MtI.

Fig. 4. Nitrogen adsorption isotherms of (a) Mt, (b) Mt–S and (c) Mt–G.

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Table 2 Textural parameters of immobilized and the functionalized clay samples. Sample

Surface area (m2 /g)

˚ Pore diameter (A)

Pore volume (cm3 /g)

Mt MtI Mt–S Mt–G

246 73 186 133

36 34.7 36 35

0.42 0.17 0.36 0.30

Fig. 8. Effect of temperature on the activity of free and immobilized lipases () free lipase, () MtI, (䊉) Mt–G.

3.2. Biochemical characterization of free and immobilized conjugates

Fig. 6. Pore size distribution of (a) Mt, (b) Mt–S and (c) Mt–G.

3.1.2. Scanning electron microscopy The morphologies of montmorillonite K-10 clay and the immobilized conjugate (Mt–I) are displayed in SEM photographs (Fig. 7). Generally, both morphologies shown were uniformly layered structures with a flaky aspect on a smooth surface. The appearance of boulder like structures may be due to the presence of enzyme and there was an increase in particle size after immobilization. A slight porous nature was observed after functionalization with glutaraldehyde.

3.2.1. Effect of temperature on the activity and stability of immobilized enzyme The optimum reaction temperature of the free enzyme was at 35 ◦ C and that of the MtI were heightened to 50 ◦ C while that of MtG; the optimum temperature was increased to 55 ◦ C (Fig. 8). This is consistent with previous findings, indicating that enzymes immobilized on carriers by covalent bonding could lead to higher stability [33]. A similar enhancement in thermal stability was observed when lipase was immobilized onto silanized palygorskite [34]. The proteins expose buried hydrophobic amino acid residues on the surface. Thus the contact area between the protein and the hydrophobic groups of the matrix should increase, resulting in an increase in the hydrophobic interaction of the proteins for the adsorbent at higher temperature. The free lipase lost almost

Fig. 7. Scanning electron micrographs of Mt, MtI and Mt–G.

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Fig. 9. Thermal stabilities of free lipase and immobilized lipases at (a) 50 ◦ C and (b) 55 ◦ C () free lipase, () MtI, (䊉) Mt–G.

85% its initial activity within around 120 min, at 50 ◦ C and 55 ◦ C while the adsorbed systems MtI hold over 80% of their initial activities after 2 h heat treatment (Fig. 9). On the other hand, the covalently bound systems could retain almost 90% activity after 120 min heat treatment and this enhanced thermal stability observed for Mt–G is in consistent with other’s conclusion [35]. Although heat considerably reduces conformational flexibility of native and immobilized lipase, immobilized lipases are still capable of performing its vibrational and more complex movement required for efficient catalytic activity [36]. The adsorbed enzyme also exhibits a reduction in mobility, but as time precedes this reduction in mobility is eliminated due to the weak nature of the enzyme support bond and therefore the protein unfolds leading to a loss of activity. The covalently bound lipase, Mt–G presented an improved thermal stability and thus may be attractive biocatalysts for industrial purposes. Lipase immobilized onto layered double hydroxide added with sodium dodecyl sulphate (SDS) exhibited the highest thermal stability by retaining 60% of its activity at 70 ◦ C [36]. This behavior is similar to that was reported for other support types [37,38]. 3.2.2. Storage stability The free enzyme lost more than 70% of its activity after 25 days storage (Fig. 10). The decrease in activity of the free lipase might be due to its susceptible autolysis during the storage time. Under the same storage conditions, the activity of the immobilized lipase decreased more slowly than that of the free

lipase. The activity of the covalently bound systems did not change significantly even after 40 days and they could maintain more than 85% activity compared to the adsorbed ones, MtI which could maintain more than 75% activity. The extended stability of the covalently bound supports could be attributed to the prevention of structural denaturation as a result of the covalent bonding of lipase molecules onto the modified supports. A similar retention in activity of 90% was reported [39] in the immobilization of C. rugosa lipase on layered double hydroxides (LDHs) and Ni/Al–sodium dodecyl sulfonate layered double hydroxide nanocomposites (Ni–SDS–LDHs) after 30 days storage at 4 ◦ C while native lipase could retain only 20% activity. 3.2.3. Kinetic parameters and activities of free and immobilized lipases in aqueous and organic media The Lineweaver plots of free and immobilized lipases in organic and aqueous media are shown in Figs. 11 and 12. The kinetic parameters obtained in both the media are compared in Table 3. The Km value for the immobilized lipases was higher than that for the free lipase which indicated that the immobilized enzyme had an apparent lower affinity for its substrate than that of free enzyme, which might be caused by the steric hindrance of the active site by the support, the loss of enzyme flexibility necessary for substrate binding, or diffusional resistance to solute transport near the particles of the support. The Vmax value of the immobilized lipases was smaller than that of the free enzyme. The reason why the immobilized lipase had a larger Km value could be due to multipoint covalent immobilization [40]; thus the structure of enzymes could be rigidified on the carriers. As part of the lipase being immobilized onto the clay surface by multipoint covalent bond, the blocking of active site of lipase would result in the loss of enzyme activity (or a smaller Vmax ) and also could cause physical obstacle for the adsorption of the substrate on lipase [41], thus increasing the diffusion resistance of ester to covalently bound Table 3 Kinetic parameters of the free and the immobilized lipases.

Fig. 10. Effect of storage stability on the activity of free and immobilized lipases () free lipase, () MtI, (䊉) Mt–G.

Sample

Medium

Vmax (U/mg)

Km (mM)

R2 a

Free lipase

Aqueous Organic

44.4 8.68

0.17 0.27

0.9831 0.9970

MtI

Aqueous Organic

35.2 0.26

0.26 0.28

0.9799 0.9931

Mt–G

Aqueous Organic

25.3 3.93

0.27 0.29

0.9348 0.9937

a

Correlation coefficient.

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Fig. 11. Lineweaver–Burk plots of the free and immobilized lipases in organic medium.

Fig. 12. Lineweaver–Burk plots of the free and immobilized lipases in aqueous medium.

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Table 4 Activity parameters for the free and immobilized lipases. Sample

Medium

Bound protein (mg/g) support)

Specific activity (U/mg)

Immobilization yield (%)

Catalytic effieciency (M−1 s−1 )Vmax /Km (*103 )

Effectiveness factor ()

Activity yield (%)

Free lipase

Aqueous Organic

– –

41.6 0.213

– –

258.1 3.1

– –

100 –

MtI

Aqueous Organic

50.3

29.2 0.24

87

135.6 7.2

0.79

83.7 87.3

Mt–G

Aqueous Organic

89.6

25.7 0.27

83.8

92.4 7.5

0.56

86.5 105.2

lipase. Km increased and Vmax value decreased after immobilization of Saccharomyces cerevisiae lipase on Mg–Al hydrotalcite [42] and Burkholderia lipase on celite carriers via covalent bonding [43]. Km value for the immobilized lipase by covalent binding was higher than that by adsorption which showed the easiness with which the enzyme-substrate complex is formed in the case of adsorbed systems. Vmax values of the immobilized lipases by adsorption were higher than that by covalent binding. The Km value of free lipase preparation was lower in the aqueous medium than in heptane and a reverse trend was observed in the Vmax value. A similar trend in the kinetic properties in aqueous and organic media was observed when C. rugosa lipase was immobilized on PACNMA membrane [44]. The Km value for the free lipase was nearly 50-fold higher in organic media. The activity yield of lipase by chemical bonding was higher than that by adsorption while the immobilization yield showed the reverse trend (Table 4). The specific activity of Mt–G was lower than MtI as chemical bonding is a rigorous method which causes substantial decrease of enzyme activity. However, the amount of immobilized protein by adsorption was lower than that by chemical bonding. The amount of lipase immobilized was increased by grafting the acid activated mica with different functional groups due to the strong interaction of the enzyme with the attached hydrophobic groups [45]. Organically modified smectite nanoclays (ORKUN, ORSWy2 and ORLAP) in which the intercalation of organic surfactants between layers imparts a more hydrophobic character to their surface, retained approximately 10% higher amounts of lipase than their unmodified precursors [46]. Catalytic efficiency was greater for the covalently bound systems than the adsorbed ones probably due to the interfacial activation of the lipase in organic solvents. The effectiveness factor was lower than 1 for the immobilized systems in aqueous medium which depicts that immobilization has some effect on substrate and product diffusion. Activities in organic media were found to be lower than those in aqueous media, which was attributed to the low amount of water available, the lower structural flexibility of the enzymes in organic media [47] and the diffusional effects since it is a heterogeneous catalysis system yielding to a narrow range of activity. This effect is especially important with lipases for which the opening of the lid covering the active site is necessary for good activity. A similar assay with nitrophenyl esters as substrate has been recently reported for subtilisin in organic media using amino acids [48]. The increase in activity was higher in the case of covalently bound systems in organic medium which could be explained by several hypotheses. Firstly, free lipase aggregated because it was insoluble in the organic medium, while immobilized lipase scattered on a large surface area could easily contact with substrates [49]. Secondly, the formation of covalent bonds between the enzyme and the support surface increased the stability of the enzyme conformation against the organic medium. Thirdly, some properties of the modified supports could benefit the activity of the immobilized lipase. The hydrophobic interaction can stabilize the “open state” conformation of lipase and favor the active site

accessibility to substrates and this substrate partitioning may also affect the activity of the covalently bound systems [50]. 4. Conclusions Lipase from C. rugosa was immobilized onto montmorillonite via two techniques i.e., adsorption and covalent binding. Montmorillonite, the functionalized materials as well as the immobilized counterparts were characterized by powder-XRD, SEM and BET surface area. The hydrolytic activity of the free and immobilized lipases in water (emulsified substrate) and in heptane (insoluble enzyme powder) was assayed using p-nitrophenyl palmitate as substrate. A very promising result of this work was the observation that the activity of the immobilized enzyme becomes less sensitive to reaction conditions than that of its free counterpart. The immobilized enzymes displayed enhanced catalytic efficiency and exhibited a better storage stability. The Km value of the covalently immobilized lipase was higher than that by adsorption. The activity of the free lipase in heptane (0.213 U/mg) was 0.51% of that in the aqueous medium (41.6 U/mg). The properties of the organomodified nanoclays and stability of immobilized lipases exhibited interesting characteristics that would be suitable for industrial biotransformations. Acknowledgements The authors thank Sophisticated Instruments Facility, Indian Institute of Science, Bangalore and Sophisticated Test and Instrumentation Centre (STIC), CUSAT, Cochin for providing SEM and XRD data. References [1] I.E. Fuentes, C.A. Viseras, D. Ubiali, M. Terreni, A.R. Alcantara, J. Mol. Catal. B: Enzym. 11 (2001) 657–663. [2] E. Serefoglou, K. Litina, D. Gournis, E. Kalogeris, A.A. Tzialla, I.V. Pavlidis, H. Stamatis, E. Maccallini, M. Lubomska, P. Rudolf, Chem. Mater. 20 (2008) 4106–4115. [3] F. Secundo, J. Miehe-Brendle, C. Chelaru, E.E. Ferrandi, E. Dumitriu, Microporous Mesoporous Mater. 109 (2008) 350–361. [4] J.C. Dai, J.T. Huang, Appl. Clay Sci. 15 (1999) 51–65. [5] M.S. Carrasco, J.C. Rad, S. Gonzales-Carcedo, Bioresour. Technol. 51 (1995) 175–181. [6] M.B.A. Rahman, S.M. Tajudin, M.Z. Hussein, Appl. Clay Sci. 29 (2005) 111–116. [7] A.F. Hsu, K. Jones, W.N. Marmer, T.A. Foglia, Biotechnol. Lett. 25 (2003) 1713–1716. [8] M.Y. Chang, R.S. Juang, Process Biochem. 39 (9) (2004) 1087–1091. [9] Y. Zhang, Y. Li, H. Dong, J. Li, X. Zheng, Chem. Eng. J. 171 (2011) 526–531. [10] Y. Zhang, Y. Li, X. Zheng, Sci. Total Environ. 409 (2011) 625–630. [11] Y. Li, Y. Zhang, J. Li, X. Zheng, Environ. Pollut. 159 (2011) 3744–3749. [12] U. Mingelgrin, S. Saltzman, B. Yaron, Soil Sci. Soc. Am. J. 41 (1977) 519–523. [13] M.C. Wang, P.M. Huang, Geoderma 112 (2003) 31–50. [14] P. Fusi, G.G. Ristori, L. Calamai, G. Stotzky, Soil Biol. Biochem. 21 (1989) 911–920. [15] A.P. Naidja, M. Huang, J. Mol. Catal. A: Chem. 106 (1996) 255–265. [16] R.D. Harter, G. Stotzky, Soil Sci. Soc. Am. Proc. 37 (1973) 116–123. [17] S. Gopinath, S. Sugunan, Appl. Clay Sci. 35 (2007) 67–75. [18] G. Sanjay, S. Sugunan, Catal. Commun. 6 (2005) 81–86. [19] C. Huang, Y. Liu, X. Wang, J. Mol. Catal. B: Enzym. 55 (2008) 49–54.

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