Synthesis of an Antioxidant n-propyl Gallate by Magnetic Iron Nanoparticles-bound Lipolase

Abstract

Citrate capped silver nanoparticles have been synthesized through chemical reduction method (Lee-Meisel method) whereas tri-sodium citrate acts as both reducing and stabilizing agents. The influence of citrate concentration, reaction time and reaction temperature during the synthesis of nanoparticles were evaluated. The properties of Ag NPs functionalized with tri-sodium citrate in varying concentrations have been investigated through UV-Vis and FTIR spectroscopy. UV spectra showed the SPR band at 418nm and conformed that the Ag NPs are spherical in shape through SEM. The size and structure of AgNps are determined through X-ray diffraction. FT-IR studies showed the presence of organic components of tri-sodium citrate which is responsible for reduction of silver nanoparticles. SERS spectra conformed the bonding site between the silver surface and oxygen atom of tri-sodium citrat.

Keywords

Magnetite iron nanoparticles, Immobilization, Lipolase 100L, Enzyme stability, Esterification, n-propyl gallate.

Introduction

A variety of nano-carriers, nano-particles, nano-fibers, nano-tubes etc. have displayed greater competence to mimic nanoscale environment for the enzyme as compared to micro- and macro-environments provided by traditional support carriers that have been commonly employed for enzyme immobilization. The porous structured beads (silica, alginate etc.), membranes and fibers (>100 μm) have some inherent drawbacks that include lower stability, enzyme leaching and poor dispersibility. Among various types of nano-particles; the iron based magnetic nano-particles have been used for enzyme immobilization due to their non-toxicity and biocompatibility. Magnetic behavior of iron nano-particles results in the simple and speedy recovery of nanobiocatalyst from reaction mixture and reusability of biocatalyst. Surface modification of magnetic NPs is an important issue that needs consideration prior to immobilization of enzyme onto NPs. Functionalization of NPs with silane is a prerequisite for covalent attachment of enzyme onto modified nano-particles, which prevents enzyme leaching and results in enhanced stability and longer shelf life of enzyme. Monomeric stabilizers, inorganic materials like silica [1,2], gold [3] or gadolinium (III) [4,5] and polymer stabilizers like dextran, carboxymethylated dextran, carboxydextran, starch, arabinogalactan, glycosaminoglycan, sulfonated styrenedivinylbenzene, polyethylene glycol, polyvinyl alcohol, poloxamers, and polyoxamines [6,7] for carrying out surface modification and functionalization of nano-carriers have been reported. Lipases (EC 3.1.1.1) occupy a place of prominence among biocatalysts in a rapidly growing biotechnological and pharmaceutical market owing to their novel and multiple applications in synthetic chemistry.

Besides bacteria, yeast and fungi are also good sources for lipase production. The lipase hydrolyzes preferentially long chain of fatty acids [8,9] besides other reactions like esterification, amidation, alcoholysis and aminolysis. However, to achieve biocatalysis at industrial scale, it is essential to employ a stable and cheaper biocatalyst, which is non-toxic, stable, easily separable/ recoverable and could be used repetitively in aqueous or organic media. The Lipolase 100L sourced from a thermophilic fungus Thermomyces langinosus was chosen for immobilization on to iron magnetic NPs because of its inherent thermostability (up to 60oC), alkalophilic nature (up to pH 9) and its known potential as an industrial biocatalyst [10]. The n-popyl gallate (E310), a propyl ester of gallic acid is a well-known antioxidant used in foods containing oils and fats, food packaging materials, non-food items like hair products, cosmetics, lubricants etc. Propyl gallate is considered a safe antioxidant to protect oil & fat-containing foods from rancidity and spoilage by preventing the formation of peroxides. The biological effects of n-propyl gallate include antimicrobial activity, ultraviolet (UV) radiation protection, chemoprotection, antimutagenesis, antitumorigenesis, antiteratogenesis and anticarcinogenesis (Safety assessment sheet of propyl gallate, 2007). In the present study, Fe3O4/ γFe2O3 nano-particles were prepared, characterized for enzymatic activity and subsequently employed to achieve esterification of n-propanol and gallic acid to synthesize n-propyl gallate in an organic medium.

Experimental

Materials and methods

The chemicals that included FeCl₃.6H₂0, NaCl, K₂HPO₄ and KCl were purchased from S.D. Fine-Chem. Ltd., Hyderabad, India; tetraethoxy silane (TEOS), FeCl₂.4H₂O, Bovine Serum Albumin (BSA), p-nitrophenyl palmitate (p-NPP) and p-nitrophenol (p- NP) were purchased from Alfa Aesar, Manchaster, England. NaH₂PO₄ and methanol were purchased from HIMEDIA Laboratory Ltd., Mumbai, India. Ammonia solution, ethanol, n-hexane, n-heptane, n-nonane, cyclohexane, DMSO, n-propanol, iso-propanol, iso-butanol, and Tris buffer were purchased from Merck Darmstadt, Germany. All the chemicals used in the present study were of high analytical grade. A commercial Lipolase 100L of Thermomyces lanuginosus was supplied as a gift by Novozymes A/S (Bagsvaerd, Denmark). The Lipolase 100L contained 8,202 U/ml lipase activity and 505.6 mg protein/ml representing a specific enzyme activity of 16.2 U/mg protein.

Synthesis of iron magnetic nano-particles

Co-precipitation [11] a widely used method for synthesis of magnetic nano-particles (Fe₃O₄ or γFe₂O₃) is based on stoichiometric molar ratio of ferrous and ferric salts in the mixture and may be represented by following chemical reaction;

Fe²⁺ + 2Fe³⁺ + 8OH^-                     Fe₃O₄ + H₂O

At a molar ratio of 2:1 (Fe³⁺/Fe²⁺), pH 8 to 14 and in the presence of non-oxidizing environment, the precipitate of Fe₃O₄ was obtained [12]. However, Fe₃O₄ (magnetite) form is unstable as it is readily oxidized to γFe₂O₃ (maghemite) form in the presence of oxygen.

Fe₃O₄ + 2H⁺                        γFe₂O₃ + Fe²⁺ + H₂O

The aqueous solutions of ferric (III) chloride hexahydrate (FeCl₃.6H₂O) (0.6M) and ferrous (II) chloride tetrahydrate (FeCl₂.4H₂O) (0.3M) were taken in molar ratio 2:1. Aqueous ammonia was drop-wise added to the mixture with continuous stirring and passage of N₂ in the reaction cocktail. The reaction was performed at a pH 10-12. Thereafter, the solution was stirred at 40°C for 30 min, 150 rpm followed by heating at 80°C for another 30 min under shaking. The magnetic NPs were retained followed by extensive washing with distilled water. The NPs were completely dried by freeze drying.

Surface modification of magnetic NPs

In brief, the NPs (Fe₃O₄ or γ Fe₂O₃) were set in the form of a suspension by dropping 1.5 g of NPs in 150 ml of ethanol. The suspension was ultrasonicated (35% amplitude for 20 min at room temperature). Deoxygenated water (1 ml) and 750 μl of TEOS were added to above NPs suspension. The suspension was stirred for 12 h at 37°C and NPs were collected by using a magnet, The NPs were ethanol washed to remove traces of unbound silane. These NPs dried in a rotary-evaporator were stored under N₂ to get finer particles with reduced particle size [13].

Binding of lipolase on silane-coated NPs

A commercial Lipolase 100L (Novozymes, Denmark) was covalently immobilized on to silane-coated modified magnetic NPs. For lipase immobilization [14], 30 mg of modified NPs were suspended in sodium phosphate buffered saline (PBS, pH 7.4) and the suspension was sonicated for 5 min at room temperature. Lipase-PBS solution (5 ml of certain concentration) was added to it in the presence of N2 and suspension was kept under shaking for 24 h at 37°C for efficient immobilization of lipase onto magnetic NPs. Lipase-immobilized NPs were separated by a magnet and supernatant was collected. These NPs were given several PBS washing and were finally dispersed into PBS for further use.

Lipase activity assay

Lipase activity of supernatant and immobilized biocatalyst was measured by a previously reported method [15] using p-NPP as a substrate. The absorbance of p-nitrophenol released during the reaction was measured at 410 nm. One unit (U) of Lipolase activity was defined as μmol(s) of p-nitrophenol released per minute by hydrolysis of p-NPP by 1 ml of soluble enzyme or 1 mg of immobilized biocatalyst (weight of NPs included) at 45°C under standard assay conditions.

Loading capacity of NPs for lipolase and determination of protein binding efficiency

To investigate the total loading capacity and protein binding efficiency of magnetic NPs, various dilutions of Lipolase in PBS (pH 7.4) were prepared (Figure 1). The 5 ml of each of the Lipolase-PBS stock (dilution) was loaded onto 5 ml of NPs suspension (30 mg NPs in 5 ml PBS) and at 37°C, overnight with gentle stirring. Thereafter, supernatant from each of the sample(s) was recovered and used to determine protein binding on NPs, which was calculated by subtracting the total protein used for binding from that of total protein recovered in the supernatant [16].

Figure 1: Amount of Lipolase immobilized onto magnetic iron nano-particles.

The appropriately diluted Lipolase-PBS stock mixed with 5 ml of NPs suspension (30 mg NPs in 5 ml PBS pH 7.4) was kept at 37°C overnight under gentle stirring. The supernatant from each of the preparations was recovered and assayed to determine protein binding on NPs, which was calculated by subtracting the amount of protein recovered in the supernatant from the total protein used for binding on to NPs.

Characterization of synthesized and NPs-bound lipolase

The size and morphology of synthesized and NPs-bound Lipolase were observed by Scanning Electron Microscope (SEM) with an accelerating voltage of 15 kV.

Thermo-tolerance of NPs-bound lipolase

To determine the maximum activity of NP-Lipolase at different temperatures, NPs-bound Lipolase was assayed at temperature range 40-70°C in separate reaction mixtures after 10 min incubation under shaking. The obtained enzyme activities were recorded thereafter.

Effect of different solvents on NPs-bound lipolase

Ideally, biocatalyst must be able to work in different organic solvents in order to carry out various synthetic reactions effectively. NPs-bound Lipolase was pre-exposed to different pure solvents i.e. n-heptane, n-nonane, cyclohexane, DMSO, iso-propanol and iso-butanol at 45°C for 10 min. The NPs-bound biocatalyst was washed with 1X PBS (pH 7.4) and checked for lipase activity.

Reusability of NPs-bound lipolase

Reusability and recovery become very important parameters when we use immobilized enzymes. Magnetic NPs have turned out as an excellent lipase carriers resulting in stability of enzyme molecules. Reusability of the biocatalyst was examined by recycling NPs-bound Lipolase up to 10 cycles of hydrolysis of p-NPP conducted at 45°C for 10 min, each. After each cycle, NPsbound Lipolase was separated using magnet and was given thrice PBS (pH 7.4) washes to prepare it for fresh cycle of enzymatic assay with p-NPP.

Enzymatic esterification of n-propanol and gallic acid

Esterification reaction involving the synthesis of propyl gallate in solvent-mediated system was carried out by varying the molar ratios of n-propanol/ gallic acid (100-500 mM) catalyzed by 50 μl homogeneous suspension of NPs-bound Lipolase. Final reaction volume was made to 2 ml by DMSO taken in Teflon-capped glass-vial (5 ml capacity). The reaction was performed at 55°C for 14 h along with respective controls (with no enzyme). Samples were analyzed by High performance liquid chromatography (HPLC) for presence of n-propyl gallate.

HPLC analysis

A C18 column (250 × 4.6 mm, 5 m; Intersil, GL Sciences Inc) was employed to determine the amount of n-propyl gallate synthesized in the reaction cocktail by the action of NPs-bound Lipolase. The sample size was 20 μl and all assays were done in duplicate and mean values were presented. An isocratic mobile phase comprising formic acid (0.1%, v/v) and acetonitrile (80%, v/v) taken in ratio 50: 50 was used at a flow rate of 1 ml/min. A reference curve was calibrated with n-propyl gallate (0.1 to 0.5 mM).

Result

The results of the various experiments are presented in the following sections

Loading capacity of lipolase and determination of protein binding efficiency on magnetite nano-particles

Different dilutions of Lipolase (conc. <5%, ~ 41.5 mg/ml) with PBS (1X, pH 7.4) were prepared to determine the loading capacity and protein binding efficiency of Lipolase onto magnetite nano-particles. The loading of enzyme on to NPs reached a saturated amount of about 6.1 mg Lipolase per milligram magnetite nanoparticles with 97.4% binding efficiency (Figure 1). However, it was found that with any further increase in the loaded protein, % binding of Lipolase onto nano-particles thereby decreased.

Characterization of lipolase bound NPs

The size and morphology of the synthesized nano-particles and NPs bound Lipolase were observed using SEM (Figure 2). The average size of distinct particles was found to be in the range of 30-90 nm. The SEM images of NPs-bound Lipolase revealed that the resulting nano-biocatalyst comprises of agglomerated structures of NPs-Lipolase.

Figure 2: Surface morphology of (A) naked Fe₃O₄/ γFe₂0₃ nano-particles; (B) Lipolase bound to NPs [Magnification X 8,500], respectively.

Thermo-tolerance of NPs-bound lipolase

The effect of temperature and thermos-tolerance on the NP-immobilized Lipolase was evaluated (Figure 3). The results showed that maximum NP-Lipolase activity was obtained at temperature 45°C (148 U/ml). However, it was found that NPs-bound lipase was active even at temperature of 70°C (135 U/ml) and thus the NPs-bound biocatalyst was capable of working at broader temperature range i.e. 40 to 70°C.

Figure 3: Thermo-tolerance of NPs-bound Lipolase.

The NPs-bound Lipolase was incubated at 40 to 70°C in separate reaction mixtures for 10 min under shaking and the residual lipase activity was assayed thereafter.

Effect of different solvents on the activity of NPs-bound lipolase

NPs-bound Lipolase showed greater activity in the presence of alkanes as compared to alcohols when used as solvents. The highest activity of the NPs-bound biocatalyst was found after exposure to n-heptane (158.7 U/ml) followed by n-hexane (151.1 U/ml), n-nonane (128.5 U/ml) and cyclohexane (121.4 U/ml). It was observed that with an increasing C-chain length of alcohols, the hydrolytic activity of NPs-bound biocatalyst decreased (Figure 4).

Figure 4: Effect of various organic solvents on hydrolytic activity of NPs-bound Lipolase.

Reusability of NPs-bound biocatalyst

The hydrolytic activity of NPs-bound Lipolase more or less (~99% of original Lipolase activity) was retained up to 10 cycles of repetitive hydrolysis (Figure 5). Moreover, the NPs-bound biocatalyst was easy to wash and retained by magnetic attraction.

Figure 5: Reusability of NPs-bound Lipolase in continuous cycles of hydrolysis of p-nitrophenyl palmitate.

NPs-bound Lipolase was pre-exposed to each of the pure solvents at 45°C for 10 min. The biocatalyst washed with PBS (pH 7.4) after each use was assayed for residual lipase activity.

Synthesis of n-propyl gallate by using NP-bound lipolase

The ability of NPs-bound Lipolase to carry out esterification reaction was tested by HPLC analysis of the reaction products. The results (Table 1) showed the presence of n-propyl gallate in the reaction mixture. In order to determine optimum ratio of reactants to obtain the maximum yield of product, different concentration of reactants were prepared by varying the concentration of one reactant at a time and keeping the other constant, making the final volume of reaction mix to 2 ml with DMSO. At equimolar (500 mM) concentartion of n-propanol and gallic acid, the maximum synthesis n-propyl gallate with a conversion of ~81.4% was achieved. On the other hand, an excess of either n-propanol or gallic acid reduced the formation of n-propyl gallate in the reaction system. It seems that excess of acid or alcohol might be responsible for denauration of protein/ enzyme ledaing to a decline in the catalytic activity of the NP-bound Lipolase.

Table 1: Synthesis of n-propyl gallate at different molar concentrations of reactants.

The reaction mixture containing varying ratio of n-propanol and gallic acid in DMSO was incubated at 55°C for 14 h in the presence of the NPsbound biocatalyst.

Discussion

Recent advances in nano-technology have led to the development of various nano-structured materials which for immobilization of enzymes and biocatalysts. Surface chemistry [17] and different physiochemical properties like surface area to volume ratios, aquaphilicity, hydrophobicity, particle size-shape, topography and chemical reactivity coupled with inherent chemical, electrical and magnetic properties of the nano-particles make them suitable host for immobilization of some industrially important enzymes [18]. In the light beneficial properties of nano-particles, successful attempt was made to synthesize the magnetic nano-particles of iron that efficiently bound to commercial enzyme Lipolase, the NPs possessed remarkable stability and reusability [19,20]. In an another experiment with immobilization of Candida rugosa lipase on alkyl silane coated Fe₃O₄ nanoparticles, amount of immobilized lipase was found to be about 0.20 mg lipase per mg of nanoparticles [14]. In the present study, the NPs-bound enzyme reached a saturated amount of about 6.06 mg Lipolase per milligram magnetite nano-particles with 97.4% binding efficiency, which was noticeably higher than previous reports. A decrease of protein % binding that was observed while increasing the protein loading is quite likely on account of the saturation of binding of lipase/ protein molecules on to surface of NPs. Further, it might also be because of obscuring of enzyme molecules placed beneath the top layer of enzyme molecules or stearic hindrance of substrate molecules to the enzyme catalytic site(s).

A Mucor javanicus lipase, which was effectively immobilized on silica nanoparticles [21] showed optimized pH at 9.0 with working range 8 to 10 as compared to free enzyme optimized at 8.0 pH. Immobilized lipase onto the EDA-GA or EDA-NCS activated nano-particles at 60oC retained 62% or 68% to the initial activity after 120 min incubation, respectively. In an another study, [22] a bacterial lipase from Arthrobacter sp. (ABL: IIIM Jammu, India strain, MTCC No. 5125) immobilized on non-magnetic (Type A) and magnetic (Type B) supports derived from copolymerization of 3-aminopropyltriethoxysilane and TEOS showed enhanced stability at pH 5-9 retaining 65-95% activity after 48 h incubation as compared to free enzyme, which was not stable under these conditions [23]. The non-magnetic and magnetic sol-gel composites retained 60-70% activity after 1 h incubation at 70°C. However, in the current work, NPs-bound Lipolase was capable working in much broader pH and temperature range, with pH 6.5 to 9.5 with good activity at extremes pH as well and showed temperature tolerance in the range 40°C to 70°C. Moreover, NPs-bound Lipolase retained almost 80-85% of its initial activity even after 28 h of incubation at 70°C much higher than previous reports [14,22,24]. Reusability and recovery of immobilized enzyme are amongst greatly desired and important parameters that have to be considered in order to ensure efficient and effective immobilization. The immobilized lipase on alkyl trimethoxysilane-coated Fe₃O₄ nano-particles (C18-Fe₃O₄) retained 65% of its initial activity after 7 cycles [14]. The decrease of activity was attributed to possible denaturation or leaching (detachment) of lipase molecules from nano-particles. In case of porcine pancreas lipase covalently immobilized on the surface of silica-coated modified magnetite nanoparticles, immobilized lipase retained 63.5% of its initial activity after 6 cycles [24]. However, NP-Lipolase showed enormous stability and repetitive use. It was active and retained 99% of its activity even after 10^th cycle. This confirmed that the lipase bound to silane-coated NPs are highly stable and overcomes enzyme-leaching problem reported by earlier workers.

DMSO is miscible with water and has the property to dissolve the enzyme, which invariably results in the inactivation of protein of interest [25]. The tolerance of NPs-bound Lipolase in different solvents suggested its likely use in organic synthesis, enantioselectivity and associated applications. The NPs-bound Lipolase showed enhanced activity in case of alkanes and DMSO. It showed the highest activity towards p-NPP (C16 acyl group) among various tested in the presence of n-heptane. When the NPsbound Lipolase was employed for the synthesis of an ester namely ‘n-propyl gallate’ the maximum synthesis of n-propyl gallate in DMSO with a conversion of ~81.4% could be achieved. However, an excess of either n-propanol or gallic acid reduced the formation of n-propyl gallate in the reaction system possiblly because of denauration of protein/ enzyme ledaing to a decline in the catalytic activity of the iron magnetic NPs-bound Lipolase.

Conclusion

Magnetite/maghemite nanoparticles (Fe₃O₄/ γFe₂O₃) synthesized by using FeCl₃ and FeCl₂ in a molar ratio 2:1 were modified with TEOS. A commercial lipase from Thermomyces lanuginosus i.e. ‘Lipolase 100L’ was used for immobilization. Nano-biocatalyst was highly stable in broader temperature range and showed increased activity in the presence of alkanes. NP-Lipolase was highly stable and enzyme-leaching problem as reported previously was resolved. NPs-bound Lipolase remained almost active over 10 cycles of hydrolysis and retained ~99% of its initial hydrolytic activity. n-Propyl gallate was effectively synthesized by NPs-bound Lipolase at 55°C. An equimolar (500 mM) reaction mixture of n-propanol and gallic acid in DMSO gave the maximum yield of the product. The study showed remarkable stability and reusability of NPs-bound biocatalyst not only in aqueous and organic solvents but also employed successfully the NPs-bould Lipolase for the synthesis of n-propyl gallate with a conversion rate of 81.4% in a reasonably short period of time indicating suitability of NPs-bound Lipolase for achieving biocatalysis in an organic solvent system. n-Propyl gallate is an antioxidant commonly used in foods containing oils & fats, food packaging materials, non-food items like hair products, cosmetics, lubricants etc.

Acknowledgement

The financial support from Department of Biotechnology, Ministry of Science and Technology, Government of India to Department of Biotechnology, Himachal Pradesh University, Shimla (India) is thankfully acknowledged. Moreover, the authors do not have any conflict of interest at their place of work.

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