Isolation, Identification, and Immobilization of Thermally Stable Lipase

Sep 26, 2013 - Centre of Scientific Excellence - Group of Encapsulation & Nanobiotechnology, National Research Center, El-Behooth St., Dokki,. Cairo, ...
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Biocatalysts: Isolation, Identification, and Immobilization of Thermally Stable Lipase onto Three Novel Biopolymeric Supports Magdy M. M. Elnashar,*,†,

∥,⊥

Hanan Mostafa,‡ Nagy A. Morsy,

§, ∥,#

and Ghada E. A. Awad‡,

∥

†

Polymers Department, ‡Chemistry of Natural and Microbial Products Department, §Chemistry of Natural Compounds Department, ∥ Centre of Scientific Excellence - Group of Encapsulation & Nanobiotechnology, National Research Center, El-Behooth St., Dokki, Cairo, Egypt ⊥ College of Medicine, Biochemistry Department, Taif University, Hawiya Taif, KSA # Faculty of Sciences & Arts, Chemistry Department, King Abdulaziz University, Khulais, KSA ABSTRACT: Lipase, one of the most important and versatile industrial enzymes, has been isolated, identified, and immobilized onto three novel supports prepared based on our US patent (US20110076737). Nine fungal isolates were cultivated, and maximum lipase activity of 285 U/mL was achieved from the fungal isolate identified as Rhizopus oryzae GF1. The enzyme was shown to be thermally stable at 50 °C for 210 min. Three different environmentally friendly biopolymers prepared according to our US patent have been used to immobilize covalently the lipase from Rhizopus oryzae GF1. The structures of the gel beads; grafted alginate, carrageenan and alginate-carrageenan; have been proved by the FTIR. The best formulation, alginatecarrageenan, covalently immobilized 183.5 U/g lipase and was further optimized to load 223 U/g lipase. The immobilization process increased the operational temperature from 30 to 50 °C compared to the free enzyme. The hydrolysis of oil using the free and the immobilized lipase was achieved at the same time, 90 min, which reflects no diffusion limitation. The shelf stability showed that the immobilized enzyme retained full activity for over 9 weeks at 4 °C, whereas the free enzyme lost 80% of its initial activity after 4 weeks. The reusability test proved the durability of the grafted beads for 20 cycles with a retention of 97% of the immobilized enzyme activity compared to 23% by other authors after the 10th use. hydrophilic, and biocompatible. For example,7 they used alginate and carrageenan to entrap tannase and then crosslinked the gel beads with chitosan followed by glutaraldehyde. Unfortunately, the entrapment technique limits their industrial use as supports for enzyme immobilization due to enzyme leakage. That was the motivator in our research group to modify these biopolymers to covalently immobilize enzymes to avoid enzyme’s leakage. In this work, we first, isolated and identified lipase from local strains, Rhizopus oryzae GF1, and to reduce the price cost, the resulting crude lipase was covalently immobilized onto three different biopolymers formulations prepared for the first time in our laboratory according to our USPTO8 immobilization onto I. modified carrageenan gel beads; II. modified alginate gel beads; III. modified alginate-carrageenan gel beads. The novel gel formulations were prepared in bead shapes using the Encapsulator to enable uniform gel beads production on the semipilot scale and to increase the gel’s surface area. The grafted formulations were illustrated using a schematic diagram, and the chemical modification was proved using the FTIR. On one hand, the three gel formulations were first studied for their thermal stability, and second, they were used to immobilize lipase and the best gel formulation was further used to optimize the enzyme loading capacity (E.L.C.). On the other hand, the free and immobilized enzymes were first characterized for their

1. INTRODUCTION One of the most versatile enzymes that have a broad variety of industrial application is lipases. Lipases can catalyze many reactions, which lead to multiple industrial applications in detergent formulations, organic chemical processing, synthesis of biosurfactants, the dairy industry, the oleochemical industry, the agrochemical industry, paper manufacture, cosmetics, nutrition, and pharmaceutical processing.1 However, lipase itself is expensive. Accordingly, there is a market needed to produce cheap lipase and/or cheap immobilized lipase that could be achieved by preparing low-cost enzyme in addition to a cheap carrier. Immobilization of enzymes on solid supports facilitates the development of continuous, large scale commercial processes as opposed to the small scale operations which employ soluble enzymes.2 Another advantage for immobilization, it often enhances the thermal and chemical stability of the immobilized enzyme as it can impart resistance to the denaturing effect of various solvents, which lead to predictable decay rates.3 Moreover, for many applications enzymes are preferably used in an immobilized state in order to easily separate the catalyst from the product steam, reuse, continuous operation, the possibility of better control of reactions, and hence more favorable economical factors can be expected. Unfortunately, good enzyme supports are expensive (silica-based carriers and synthetic polymers such as Eupergit C) and that caused many to search for a cheaper substitute such as CaCO3,4 rice husk and rice straw,5 or chitin and chitosan.6 Alginate and carrageenan are natural polymers that could be good candidates for immobilization of enzymes as they are abundant in nature, © 2013 American Chemical Society

Received: September 7, 2013 Accepted: September 26, 2013 Published: September 26, 2013 14760

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mixed with dry KBr. The mixture was ground into a fine powder using an agate mortar before it was compressed into a KBr disk under a hydraulic press at 10,000 psi. Each KBr disk was scanned 16 times at 4 mm/s at a resolution of 2/cm over a wavenumber range of 400−4000/cm, using Happ-Genzel apodization. The characteristic peaks were recorded. 3.3. Isolation, Production, and Identification of Thermostable Lipase Producing Bacteria. 3.3.1. Isolation of Lipase Producing Microorganisms. Isolation of fungi with lipolytic activity was carried out according to the method adopted in ref 10. Five grams of contaminated clarified butter from different sources was suspended in 50 mL of sterile distilled water. One milliliter of the butter suspension was inoculated in a medium consisting of 0.025% yeast extract and 0.1% commercial cooking oil emulsified in aqueous 10% (w/v) Arabic gum.11 The inoculated flasks were incubated for 10 days with shaking at 45 °C. One milliliter of the culture was plated onto agar plates of the same medium. Single colonies were selected and inoculated many times in potato dextrose medium (PDA) slants. 3.3.2. Lipase Production. Production of lipase was carried out using a medium consisted of the following: Peptone, 3.0; KH2PO4, 0.2; KCl, 0.05; MgSO4·7H2O, Glucose, 1.0; and commercial cocking oil 1.0 g %.12 The pH of the medium was adjusted to pH 7. Cultivation was carried out in 250 mL Erlenmeyer flasks containing 50 mL of production medium. The flasks were inoculated with 1.0 mL containing (5 × 106 spores/mL) of 5 days old culture prepared by inoculating a slant of the fungus in 50 mL of isolation medium. The inoculated flasks were incubated at 30 °C for 72 h in a rotary shaker adjusted at 200 rpm. At the end of the incubation period, the fungal culture was centrifuged at 5000 rpm for 20 min under cooling. The culture filtrate was used in further investigation. 3.3.3. Identification of the Lipase Producing Fungal Isolate. Nine fungal isolates were studied, and the most potent one that produced the highest lipase productivity was chosen for further investigations and has been identified. Identification was carried out by microscopic examination, and the mycelium and conidiophores were observed after having been cultured on a concavity slide with PDA medium at 28 °C and then colored by a fungus staining solution (20.0 g of phenol, 20.0 mL of lactic acid, 40.0 mL of glycerin, 20 mL of distilled water, and 0.05 g of cotton blue). Furthermore, the 18S rDNA was amplified by a polymerase chain reaction (PCR) using primers designed to amplify a 1500 bp fragment of the 18S rDNA region. This PCR reaction used DNA thermal cycler (Whatmann Biometra UNO, Gottingen, Germany). The forward primer was 18SF149:5′-GGAAGGG (G/A) TGTATTTATTAG-3, and the reverse primer was 18SR 701: 5′GTAAAAGTCCTGGTTCCC-3′. The fungal identification was carried out at City for Scientific Research, Borg el Arab, Alexandria, Egypt. 3.3.4. Cell Banking. This strain was subcultured in agar medium, and the arisen colonies were harvested by glycerol solution (20%) and put in a series of 2 mL Cryogen vials (Nalgene, USA). The tubes were frozen immediately at −20 °C for 24 h followed by storage as a working cell bank at −80 °C for further use. This was an important step to ensure that the starter culture of each experiment was of the same generation number. 3.4. Determination of Lipase Activity. Lipase enzyme activity was carried out according to Parry et al., 2001.13 One

activities at different pHs and temperatures, and second, they were studied for their thermal stability at different temperatures. The optimum gel formulation among the three carriers was used with the optimum enzyme conditions for substrate conversion, and the results were compared with the free enzyme. Finally, the most important experiments for evaluation of any immobilized enzymes, the shelf and the operational stability, were studied.

2. MATERIALS k-Carrageenan (Mr: 154,000; sulfate ester ∼25%) and alginic acid sodium salt from brown algae (CAS # 9005-38-3) were obtained from Fluka. Polyethyleneimine (MW: 423), Cat # 468533, was obtained from Aldrich. The triglyceride is a locally produced extra pure olive oil. Other chemicals were of AnalR or equivalent quality. The Encapsulator, model IE-50 was purchased from Innotech Encapsulator in Switzerland. 3. METHODS 3.1. Preparation and Grafting of k-Carrageenan, Alginate, and Carrageenan-Alginate Gel Beads. Three gel formulations were prepared using 2% (w/v) k-carrageenan, 2% (w/v) alginate, and 2% (w/v) carrageenan-alginate (ratio 1:1) dissolved in distilled water. In the 3 formulations, all gel beads are having the same water content so that the comparison would be feasible. The solutions were mixed thoroughly using an overhead mechanical stirrer until complete dispersion had occurred. The gels were prepared in uniform beads using the Inotech Encapsulator. The three gel solutions were grafted according to our USPTO,8 where the above three gel solutions were dropped through a nozzle of 300 μm using the Innotech Encapsulator in hardening solutions of (a) 2.25% (w/v) KCl (K+) for k-carrageenan and (b) 2% (w/v) CaCl2 (Ca2+) for the alginate and carrageenan-alginate for 2 h as a control gel. Gel beads were further modified using polyelectrolytes followed by a mediator to bind lipase enzyme covalently. The polyelectrolyte used was polyethylenimine (PE), and the mediator was glutaraldehyde. The role of (PE) was to improve the gels thermal stability, whereas the glutaraldehyde was to incorporate new functionalities to the gel beads (aldehyde groups). The control formulations (unmodified gel beads) were prepared by dropping the above beads in a solution of 4% (v/v) polyethylenimine (PE) at pH 9.5 for 3 h. Whereas, the treated gels beads were prepared by soaking the control formulations in a solution of 2.5% (v/v) glutaraldehyde (GA) for 2 h to crosslink the gel surface and moreover to incorporate free aldehyde groups as follows:9 a) Alg/Ca2+/PE/GA b) Carr/K+/PE/GA c) Alg-Carr/Ca2+/PE/GA 3.2. Elucidation of the Modified Gels Using ATR-FTIR. The attenuated total reflectance Fourier transform infrared (ATR-FTIR) has been used to identify the new functionalities on the grafted gels. IR transmission spectra were obtained using a FTIR spectrophotometer (FTIR-8300, Shimadzu, Japan). The test is aiming to prove the presence of the new functional group, carbonyl group, after carrageenan and alginate or alginate-carrageenan gel modification with PEI followed by glutaraldehyde. A total of 2% (w/w) of the sample, with respect to the potassium bromide (KBr; S. D. Fine Chem) disk, was 14761

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milliliter of culture filtrate was mixed with 2.5 mL of deionized water and 1 mL of 0.1 M of tris−HCl buffer (pH 7.5), 3 mL of 10% (v/v) triglyceride emulsion. The mixture was incubated for 2 h at 37 °C in a shaking water bath. At the end of the incubation time 10 mL of 99% ethyl alcohol was added, and the resulting mixture was then titrated against 0.05 N NaOH using thymolphthalein as indicator. Boiled enzyme samples were used as blanks. One unit of lipase activity was defined as the amount of enzyme that liberates 1 μmol of free fatty acids under the assayed conditions. 3.5. Immobilization of Lipase onto Grated Gel Beads. In this experiment, the three gel formulations a) Alg/Ca2+/PE/GA b) Carr/K+/PE/GA c) Alg-Carr/Ca2+/PE/GA were incubated with lipase solution of 1:5 dilution for 24 h at 37 °C and at pH 7.5. The best formulation was used for further optimization. 3.5.1. Optimization of the Enzyme Loading Capacity Using Grafted Alginate-Carrageenan Beads. In this experiment, 1 g of the gel formulation of Alg-Carr/Ca2+/PE/GA was incubated in 5 mL of 1:20, 1:10, 1:5, 1:3, and 1:1 dil enzyme in 0.1 M acetate phosphate buffer at 37 °C, pH 7.5 for 24 h. The amount of the initial enzyme solutions per 5 mL were 71.25, 142.5, 285, 475, and 1425 U, respectively. The E.L.C. or the amount of enzymes’ units immobilized onto gel beads was calculated as follows E.L.C. = (Mo − M f )/W

3.5.2.2. Temperature Profile. The optimum temperatures for the free and immobilized lipase were examined. The same units (225 U) of free and immobilized enzymes were incubated for 2 h at pH 6 into the assay mixture as in Section 3.5 at 20− 70 °C. The concentration of the formed product was estimated by measuring the resulting amount of the titrated 0.05 N NaOH using thymolphthalein as indicator, which is equivalent to the produced free fatty acid. The data were normalized to 100% activity. The highest enzyme activity is expressed as 100%, and each pH is expressed relatively as a percentage of the 100% activity. 3.5.2.3. Enzyme’s Thermal Stability. To prove the stability of the immobilized enzyme at high temperatures, the enzymes were incubated in the enzyme’s buffer solution for a period of 2 h at 50 and 60 °C, and then they were examined for enzyme activity as above. The data were normalized to 100% activity. The highest enzyme activity is expressed as 100%, and each temperature is expressed relatively as a percentage of the 100% activity. 3.5.4. Oil Hydrolysis, Shelf, and Operational Stability of Immobilized Lipase. To evaluate the efficiency of the immobilized enzyme, three main experiments were carried out. The oil hydrolysis using the optimum conditions was obtained from the above optimization for the free and immobilized enzyme and the shelf and reusability of the immobilized enzyme. 3.5.4.1. Oil Hydrolysis. The same units (225 U) of free and immobilized enzymes were incubated at 45 °C for 2.5 h at pH 6 into the assay mixture as in Section 3.4. Samples were withdrawn at interval times from 30 min to 2.5 h and analyzed for oil hydrolysis. 3.5.4.2. Shelf Stability. The shelf stability of immobilized and free lipase was studied for the free and the immobilized enzyme over a period of 9 weeks at 4 °C. Ten grams of the immobilized enzyme containing (225 U/g beads) and their equivalent of the free enzyme (225 U/mL) were stored in 0.1 M of tris-HCl buffer (pH 7.5) at 4 °C. The samples were covered to avoid dehydration and loss of the solvent. A sample of the free enzyme (1 mL) or the immobilized enzyme (1 g gel beads) has been withdrawn every week and assayed for enzyme activity. One gram of the grafted gel beads was added to 2.5 mL of deionized water and 1 mL of 0.1 M of tris-HCl buffer (pH 6), 3 mL of 10% (v/v) triglyceride emulsion. The mixture was incubated for 1.5 h at 50 °C in a shaking water bath, and the substrate solution was assayed as above. The starting operational activity was considered as 100% relative activity, and data were normalized to 100% activity. 3.5.4.3. Operational Stability. The reusability of immobilized lipase was studied using the novel grafted gel beads. The best conditions obtained from the optimum pH, temperature, and hydrolysis of oil were taken. One gram of the grafted gel beads was added to 2.5 mL of deionized water and 1 mL of 0.1 M of tris-HCl buffer (pH 6), 3 mL of 10% (v/v) triglyceride emulsion. The mixture was incubated for 1.5 h at 50 °C in a shaking water bath, and the substrate solution was assayed as above. The same gel disks were then washed with distilled water and reincubated with another substrate solution; this procedure was repeated 20 times, and the initial activity was considered as 100%. The relative activity was expressed as a percentage of the starting operational activity.

(1)

where Mo is the initial enzyme activity (U), Mf is the enzyme activity of the filtrate (U) after immobilization, and W is the weight of wet gel beads (g). The best formulation was used for the lipase’s catalytic experiments. On the other hand, the immobilization efficiency (IE) has been also calculated by using the following equation I.E.% = (Mi /Mo)*100

(2)

where Mo is the initial enzyme activity (U), and Mi is the enzyme activity of the immobilized enzyme per gram gel beads (U). Moreover, the immobilization yield (I.Y.) has been calculated from the following equation I.Y.% = (C /A − B)*100

(3)

where A is the activity of free enzyme added, and B is the activity of remaining enzyme, whereas C is the activity of immobilized enzyme. 3.5.2. Evaluation of Lipase Catalytic Activity. Various factors were studied to evaluate the catalytic activity of both the free and immobilized lipase such as effect of pH, temperature of reaction, and thermal stability of the enzymes, as follows: 3.5.2.1. pH Profile. The optimum pHs for the free and immobilized lipase were examined. Same units (225 U) of free and immobilized enzymes were incubated for 2 h at 37 °C into assay mixture as in Section 3.5. at pH 5−9. The concentration of the formed product was estimated by measuring the resulting amount of the titrated 0.05 N NaOH using thymolphthalein as indicator, which is equivalent to the produced free fatty acid. The data were normalized to 100% activity. The highest enzyme activity is expressed as 100%, and each pH is expressed relatively as a percentage of the 100% activity. 14762

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4. RESULTS AND DISCUSSION 4.1. Screening of Different Bacterial Isolates for Production of Lipase Enzyme. Nine fungal isolates were cultivated in the production medium in order to know the most potent one able to produce the highest activity of lipase enzyme (Figure 1) showing that all fungal isolates produce a

carrageenan blend. Second, we used different concentrations of the enzyme’s solution with the best gel beads’ formulation. 4.4.1. Immobilization of Crude Lipase onto the Three Gel Beads Formulations. According to the results shown in Figure 3a and in Table 1, it is obvious that the enzyme loading capacity

Figure 1. Screening of fungal isolates for their ability to produce lipase enzyme.

considerable amount of lipase enzyme, but maximum lipase activity was achieved from the fungal isolate coded GF1, which was able to produce about 285 U/mL of lipase enzyme. Thus, fungal isolate GF1 was chosen for further investigations. 4.2. Fungal Isolate Identification. The lipase producing fungus isolate was identified as Rhizopus oryzae by both microscopic examination and DNA sequence, Therefore it was designated as Rhizopus oryzae GF1. 4.3. IR of the Three Gel Formulations. As shown in Figure 2, the intensity of the functional groups, −OH and

Figure 3. a. Immobilized lipase onto grafted alginate, carrageenan, and alginate-carrageenan gel beads. Enzyme was soaked for 4 h at 37 °C and at pH 7.5. b. Optimization of enzyme loading capacity onto carrageenan gel beads.

(ELC) using the alginate-carrageenan blend (183.5 U/g gel) is the highest and almost equivalent to the sum of both alginate (76.7 U/g gel) and carrageenan (115.8 U/g gel) separately. The high ELC of the Alg-Carr formulation could be due to this blend combining the functional groups of both alginate (−OH, −COOH) and carrageenan (−OH, −OSO 3 H), which increased the chances to attract and bind more enzyme to the modified blend surface via hydrogen bonding or hydrophilic−hydrophilic interaction. Another hypothesis could be that the anionic characters of the formed blend attracted more PE to the gel beads surface via anionic-cationic complex formation as previously described by the authors,20 which bound consequently to more aldehyde groups and as a result to more enzyme molecules as illustrated in Scheme 1. In other words, as shown in Scheme 1, more enzymes were bound to the Alg-Carr formulation as a result of more free aldehydic groups being present than that in the formulations of Alg or Carr. On the other hand, the immobilization efficiency (I.E.%) has shown that the Alg-Carr formulation has immobilized an amount of enzymes (79%), which is almost the sum of that of Alg (33%) and Carr (50%) formulations separately. However, it could not verify if some of the immobilized enzymes were blocked or became inactive during the immobilization or not. To verify the former, we calculated the immobilization yield (I.Y.) and it was almost 100% in the case of Carr and Alg-Carr

Figure 2. IR of the three gel formulations: modified alginate, carrageenan, and alginate-carrageenan gel beads.

−CHO, at 3100−3500 and 1700 cm−1, respectively, of the gel formulations of Alg-Carr/PE/GA is almost the sum of that of Alg/PE/GA and Carr/PE/GA. Thus we expect more enzymes to be loaded onto the Alg-Carr/PE/GA than that for the Alg/ PE/GA or the Carr/PE/GA. 4.4. Immobilization and Optimization of the Enzyme Loading Capacity (ELC). In this part, we carried 2 experiments starting with using different types of polymeric gel beads based on alginate and carrageenan and alginate14763

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Table 1. Immobilized Lipase onto Alg, Carr, and Carr-Alg Gel Beads gel beads formulation

total units (enz. soaking soln)

U/g beads

immobilization efficiency, % (I.E.%)

immobilization yield, % (I.Y.%)

Alg Carr Alg-Carr

285 285 285

94 ± 5.7 142 ± 17 225 ± 2.8

33 ± 2.0 50 ± 6.0 79 ± 1

66 ± 4.0 95 ± 5.3 100 ± 1.6

Scheme 1. Schematic Diagram To Describe the Modification of the Gel Beads and the Enzyme Loading Capacity: (a) Alg/PE/ GA/Enz, (b) Carr/PE/GA/Enz, and (c) Alg-Carr/PE/GA/Enz

same ELC of 222.5 U/g beads but with a much higher immobilization efficiency of 46.8%. We could also tell from Table 1 that at low enzyme concentration, there is almost no loss of enzyme activity after immobilization; for example, at enzyme dilution of 1:20 (71.5 U), the immobilization efficiency was almost 100%. Optimization of pH and Temperature of the Immobilized Lipase. The optimum pH values for the free and the immobilized enzyme were very close as they were at pH 7 and pH 6−6.5, respectively, as shown in Figure 4a. However,

formulations, whereas it was only 66% in the case of Alg gel beads. In other words, some of the enzymes were blocked or inactivated during immobilization of enzymes onto the Alg gel beads and almost 100% of the enzymes were still active after immobilization onto Alg-Carr and Carr gel beads. 4.4.3. Optimization of the Immobilized Lipase onto AlgCarr/PE/GA Gel Beads. As shown in Figure 3b, the ELC increased gradually from 72.5 U/g gel to reach 222.5 U/g gel by increasing the enzyme concentration to 1:3 dilution. However, at no enzyme dilution, there is almost no increase in ELC. This could be regarded to saturation of the glutaraldehyde molecules with the enzyme at 1:3 enzyme dilutions so that no more enzymes could be loaded to the gel beads.14 Data of the ELC as well as the immobilization efficiency were tabulated in Table 2. Table 2. Immobilized Lipase onto Carr-Alg Gel Beads enz. concn

total units (soaking soln)

U/g beads

immobilization efficien cy, %

1:20 1:10 1:05 1:03 1:01

71.25 142.5 285 475 1425

72.5 ± 3.5 132 ± 2.8 183.5 ± 4.9 222.5 ± 3.5 223 ± 1.4

101.8 ± 5.0 92.6 ± 2.0 64.4 ± 1.7 46.8 ± 0.7 15.6 ± 0.1

According to the results in Table 2, the maximum ELC of 223 U/g beads was obtained when we used enzyme dilution of 1:01 (1425 U). Unfortunately, the immobilization efficiency of this formulation was the lowest, 15.6%, and the loss of enzymes was quite high as only 223 U were immobilized out of 1425 U. Accordingly, we have chosen the next higher concentration of enzyme dilution of 1:03 (475 U), which showed almost the

Figure 4. a. pH profile of the immobilized crude enzyme. b. Temperature profile of the immobilized crude enzyme. 14764

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thermal stability at high 50−60 °C compared to that obtained by ref where the free lipase at 50 °C lost all its initial activity after 120 min of heat treatment; the immobilized enzyme retained about 63% of its initial activity. At 60 °C, the free lipase lost all its initial activity after 45 min heat treatment, while the immobilized counterpart retained only 45% of its initial activity. In general, the decrease of enzymes’ activity might be due to the disturbance of globular structure of the protein by heat. On the other hand, the immobilization process stabilized more of the 3D of the enzyme; in addition the polymeric material for immobilization forms a cage surrounding the enzyme to protect it from the outside heat. In brief, the immobilized preparations were more stable than the soluble enzymes at higher temperatures and that is favorable for industrial use. 4.6. Oil Hydrolysis Using the Free and Immobilized Lipase. The results for the hydrolysis of olive oil using the free and immobilized lipase were shown in Figure 6. Both the free

the immobilized enzyme showed a better pH profile at low pHs, pH 5.5−6.5, whereas the free enzyme showed a better pH profile at high pHs, pH 7−9. In general, both trends were close to each other at most pHs as shown in Figure 4a. In contrast, Arica et al., 200015 immobilized lipase onto poly(2-hydroxyethylmethacrylate-co-methacrylamidophenylalanine). They found that the free enzyme was more stable at pH 4−7, whereas the immobilized enzyme was at pH 7.5−9. This behavior could be regarded to the nature of the carriers as they used copolymers of different functional groups, microenvironment, and morphology other than the one we used, and as a result they had different enzyme-polymer interactions. Alternatively, the imine groups in our polymer could be labile at acidic medium and that could affect the immobilized enzymes. One of the main goals of this article was to improve the enzyme’s thermal stability to be suitable for industrial use as previously mentioned in the Introduction. As shown in Figure 4b, the thermal stability of the enzyme covalently immobilized to the grafted gel beads revealed a higher and outstanding thermal stability over the free enzyme where the optimum temperature of the free enzyme was at 30 °C and that of the immobilized enzyme was at 50 °C. A similar optimum temperature of 50 °C for the immobilized lipase was obtained in refs 16 and 17. In another study, some authors achieved lower optimum temperatures for immobilized lipase; for example, in ref it was found that immobilization of lipase from C. rugosa on chitosan showed an optimum reaction temperature of 30 °C, while in ref 19 it was found that immobilization of lipase from the same organism on kaolin showed higher activity at 40 °C. By comparing our results to other authors’ results, we could conclude that the immobilized enzyme’s thermal stability is highly affected by the type of the carrier. 4.5. Thermal Stability of the Free and Immobilized Lipase. The thermal stability of the free and immobilized enzymes was shown in Figure 5. The temperature stability of

Figure 6. Hydrolysis of oil using the free and the immobilized lipase.

and the immobilized enzyme were following the same trend at most times. For example, at 30 min, the immobilized enzyme reached 65% conversion compared to 55% for the free enzyme. At 60 min, the free enzyme reached a plateau and conversion of 100% of olive oil, whereas the immobilized enzyme reached 82%. At 90 min and onward, both enzyme forms had the same plateau of 100% conversion. This result could be regarded to little to no diffusion limitation has encountered the productsubstrate conversion using the immobilized enzyme. However, the little delay for the immobilized enzyme to reach the plateau at 60 min could be regarded to experimental error as the immobilized enzyme was faster than the free one at 30 min, and it was equal to the free enzyme at 90 min. Another explanation, the acidic medium for the reaction at pH 6, could have a negative effect on the imine groups of the immobilized enzyme and made it less stable. 4.7. Operational and Shelf Stability of Immobilized Lipase. The shelf stability of the free and immobilized enzymes was studied at 4 °C for 9 weeks as shown in Figure 7a. The free enzyme retained full activity for 2 weeks, after that it declined to 80% after 3 weeks and dramatically to 10% after 4 weeks and lost completely its activity after 5 weeks. On the other hand, the immobilized enzyme retained full activity for over 9 weeks, which is very useful for marketing the immobilized enzyme. On the other hand,21 immobilized lipase on chitosan nanofibrous membrane and the results were as follows: the immobilized lipase retained 56.2% of its initial activity after 4 weeks when stored in phosphate buffer (pH = 7.0) at 4 °C activity, while the free lipase retained only 36.6% after 10 days and lost most of its initial activity after 4 weeks. The extended stability of the immobilized form could be attributed to prevention of the 3D structure of the enzyme

Figure 5. Temperature-stability profile of free and immobilized lipase.

the free and immobilized enzymes at 50−60 °C revealed that the immobilized form is more stable for a longer time than the free enzymes. For example, at 50 °C, the immobilized enzymes retained 100% of its relative activity for an incubation period of 210 min, whereas that of the free enzymes decreased gradually till it reached 85%. At 60 °C, the temperature at which the enzymes are preferably used in industries to avoid microbial contamination, both enzymes’ form retained 100% of their relative activity at 90 min. However, at 150 min, the free enzyme lost 20% of its relative activity, whereas the immobilized enzyme still remained 100% of its relative activity. At a longer incubation time at 210 min, the free enzyme retained 64% of its relative activity compared to 86% for the immobilized enzyme. Both enzyme forms showed better 14765

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AUTHOR INFORMATION

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*Fax: 202-3370931. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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REFERENCES

(1) Sharma, R.; Chisti, Y.; Banerjee, U. C. Production, Purification, Characterization, and Applications of Lipases. A Review Article. Biotechnol. Adv. 2001, 19, 627. (2) Malcata, F. X.; Reyes, H. R.; Garcia, S. G.; Hill, C. G.; Amundson, C. H. Immobilized Lipase Reactors for Modification of Fats and Oils − A Review. J. Am. Oil Chem. Soc. 1990, 67, 890. (3) Khmelnitsky, Y. L.; Levashov, A. V.; Klyachko, N. L.; Martinek, K. Engineering Biocatalytic Systems in Organic Media with Low Water Content. Enzyme Microb. Technol. 1988, 10, 710. (4) Rosu, R.; Iwasaki, Y.; Shimizu, N.; Doisaki, N.; Yamane, T. Intesification of Lipase Performance in a Transesterification Reaction by Immobilization on CaCO3 Powder. J. Biotechnol. 1998, 66, 51. (5) Tantrakulsiri, J.; Jeyashoke, N.; Krisanangkura, K. Utilization of Rice Hull Ash as Support Material for Immobilization of Candida cylindracea Lipase. J. Am. Oil Chem. Soc. 1997, 74, 173. (6) Felse, P. A.; Panda, T. Studies in Applications of Chitin and its Derivatives. Bioprocess Eng. 1999, 20, 505. (7) Nakane, k.; Ogihara, T.; Ogata, N.; Kurokawa, Y. EntrapImmobilization of Invertase on Composite Gel Fiber of Cellulose Acetate and Zirconium Alkoxide by Solgel Process. J. Appl. Polym. 2001, 81, 2084. (8) Elnashar, M. M. Carriers for Immobilization and Reusability of Enzymes. USPTO, U.S. Patent, 20110076737 A1, 2010. (9) Elnashar, M. M.; Yassin, A. M. Covalent Immobilization of βGalactosidase on Carrageenan Coated Chitosan. J. Appl. Polym. Sci. 2009, 114, 17. (10) Gowland, P.; Kernick, M.; Sundaram, T. K. Thermophilic Bacterial Isolates Producing Lipase. FEMS Microbiol. Lett. 1987, 48, 339. (11) Abdel-Fattaha, Y. R.; Gaball, A. A. Identification and OverExpression of a Thermostable Lipase from Geobacillus Thermoleovorans Toshki in Escherichia coli. Microbiol. Res. 2008, 163, 13. (12) Akhtar, M. W.; Mirza, A. Q.; Chughtai, M. I. D. Lipase Induction in Mucor hiemalis. Appl. Environ. Microbiol. 1980, 40, 257. (13) Parry, R. M.; Chandan, R. C.; Shahani, K. M. Rapid and Sensitive Assay for Milk Lipase. J. Dairy Sci. 1966, 49, 356. (14) Elnashar, M. M. Preparation of Carrageenan Treated with Synthetic Polymer as a Carrier for Biotechnological Applications. Egyptian Patent, 24600, 2009. (15) Arica, M. Y.; Kacar, Y.; Ergene, A.; Denizli, A. Reversible Immobilization of Lipase on Phenylalanine Containing Hydrogel Membranes. Process Biochem. 2001, 36, 847. (16) Roy, N.; Ray, L.; Chattopadhyay, P. Production of Lipase in a Fermentor using a Mutant Strain of Corynebacterium sp.: Its Partial Purification and Immobilization. Indian J. Exp. Biol. 2004, 42, 202. (17) Kanwar, S. S.; Verma, H. K.; Kaushal, R. K.; Gupta, R.; Chimni, S. S.; Kumar, Y.; Chauhan, G. S. Effect of Solvents and Kinetic Parameters on Synthesis of Ethyl Propionate Catalysed by Poly (AAcco- HPMA-cl-MBAm)-Matrix-Immobilized Lipase of Pseudomonas aeruginosa BTS-2. World J. Microbiol. Biotechnol. 2005, 21, 1037. (18) Hung, T. C.; Giridhar, R.; Chiou, S. H.; Wu, W. T. Binary Immobilization of Candida rugosa Lipase on Chitosan. J. Mol. Catal. B: Enzym. 2003, 26, 69. (19) Rahman, M. B. A.; Tajudin, S. M.; Hussein, M. Z.; Rahman, R. N. R.; Salleh, A. B.; Basri, M. Immobilization Application of Natural Kaolin as Support for of Lipase from Candida rugosa as Biocatalyst for Effective Esterification. Appl. Clay Sci. 2005, 29, 111. (20) Bayramoglu, G.; Kacar, Y.; Denizli, A.; Arıca, M. Y. Covalent Immobilization of Lipase onto Hydrophobic Group Incorporated Poly(2-hydroxyethyl methacrylate) Based Hydrophilic Membrane Matrix. J. Food Eng. 2002, 52, 367.

Figure 7. a. Shelf stability of the immobilized crude enzyme. b. Reusability of the immobilized enzyme.

from denaturation as a result of the covalent bonding of lipase biomacromolecules onto the biopolymeric beads. The main advantage of immobilization of enzymes is the easy separation and reusability. The data shown in Figure 7b indicated that the immobilized lipase retained over 97% of its activity after 20 reuses. The retention of the enzyme activity for almost 20 cycles with no loss of activity is proof that the enzymes were immobilized to the gel beads via covalent bonds. Otherwise, if some enzymes were immobilized via physical bonds, they will not tolerate the reusability test and they would be lost. These results were far better than the one by Garlapati et al., 2013.21 They succeeded in retaining 95% of the initial activity of immobilized lipase from Rhizopus oryzae after 6 cycles; however, the enzyme activity dropped to 23% by the 10th cycle. The slight decrease in the immobilized enzyme activity by the 20th use using the grafted Alg-Carr gel might be attributed to inactivation of enzyme due to continuous use.22

5. CONCLUSION In this report, we succeeded to produce thermally stable lipase from screening of seven fungal isolates. The best organism producing lipase, Rhizopus oryzae GF1, has been identified, and its activity was 285 U/mL. The enzyme was thermally stable at 50 °C for up to 2.5 h. Three formulations of gel beads were prepared with the size of 300 μm using the Encapsulator. The beads contained free aldehyde groups and were used to covalently immobilize the enzyme through its free −NH2 groups to form Schiff’s base. The FTIR showed a clear band at 1700 cm−1 for the free aldehyde groups in the three gel beads formulations. The intensity of the −CHO band was stronger for the Alg-Carr/PE/GA formulation than that of Alg/PE/GA and Carr/PE/GA. This could be the main reason why the enzyme loading capacity of the formulation of Alg-Carr/PE/ GA (183.5 U/g) is higher than that of the Alg/PE/GA (76.7 U/g) and the Carr/PE/GA (115.8 U/g). This interpretation has been illustrated in Scheme 1 as freer aldehyde groups were found in the Alg-Carr/PE/GA formulation. This result was highly encouraging to immobilize the semipurified lipase in our next manuscript to see if this behavior is going to repeat or not! 14766

dx.doi.org/10.1021/ie402960d | Ind. Eng. Chem. Res. 2013, 52, 14760−14767

Industrial & Engineering Chemistry Research

Article

(21) Garlapati, V. K.; Kant, R.; Kumari, A.; Mahapatra, P.; Das, P.; Banerjee, R. Lipase Mediated Transesterification of Simarouba Glauca Oil: A New Feedstock for Biodiesel Production. Sustainable Chem. Processes 2013, 1, 11. (22) Elnashar, M.; Yassin, M. Lactose Hydrolysis by β-Galactosidase Covalently Immobilized to Thermally Stable Biopolymers. Appl. Biochem. Biotechnol. 2009, 159, 426.

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dx.doi.org/10.1021/ie402960d | Ind. Eng. Chem. Res. 2013, 52, 14760−14767