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Article

Preparation of Immobilized Lipase on Silica Clay as a Potential Biocatalyst on Synthesis of Biodiesel

by
Ting Zou
1,
You-dan Duan
1,
Qiao-e Wang
2 and
Hai-ming Cheng
1,3,*
1
The Key Laboratory of Leather Chemistry and Engineering of Ministry of Education, Sichuan University, Chengdu 610065, China
2
Key Laboratory of Cosmetic, China National Light Industry, Beijing Technology and Business University, Beijing 100048, China
3
National Engineering Laboratory for Clean Technology of Leather Manufacture, Sichuan University, Chengdu 610065, China
*
Author to whom correspondence should be addressed.
Catalysts 2020, 10(11), 1266; https://doi.org/10.3390/catal10111266
Submission received: 3 October 2020 / Revised: 29 October 2020 / Accepted: 29 October 2020 / Published: 2 November 2020
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Abstract

Biodiesel offers an important alternative to fossil fuel. In this work, Eversa Transform 2.0 lipase was immobilized onto 3-aminopropyltriethoxysilane (APTES) modified silica clay (SC) by glutaraldehyde. The characteristics of the functionalized supports and the immobilized lipase were investigated by FTIR, TEM, BET, and XRD. The results show that the optimal conditions of lipase immobilization are as follows: 2% glutaraldehyde concentration, 15 mg/mL lipase concentration and incubating at 25 °C for 60 min. The immobilized lipase showed a high tolerance to temperature and pH variation in comparison to the free lipase. The immobilized lipase on SC was applied as a biocatalyst for the synthesis of biodiesel from methanol and canola oil. A biodiesel yield of 86% was obtained at a temperature of 45 °C via a three-step methanol addition. A conversion yield of 67% was maintained after reusing the immobilized lipase for five cycles. This work provides a strategy for the preparation of an efficient biocatalyst for the synthesis of biodiesel.

1. Introduction

Biodiesel is a clean alternative fuel which can be produced by renewable natural resources (oils and fats). In terms of its chemical composition, biodiesel is a series of long-chain fatty acid esters, which can be directly used in a compression ignition engine [1,2]. It has been reported that a variety of oil resources, such as vegetable oils, animal oils, waste cooking oils, and micro-algae oils, have been used for biodiesel production [3,4,5,6,7]. The transesterification of triglyceride is generally carried out by a catalyst, which can be acid, alkali, or enzyme catalysts [8]. Acid catalysts, such as sulfuric acid, hydrochloride, and phosphoric acid, are applied in biodiesel production due to their efficient catalytic performance. However, the acid catalysts suffer from problems of equipment corrosion, the emission of harmful gases, and harsh reaction conditions [9,10]. On the other hand, the alkali catalyzed process has fatal weaknesses such as saponification and the difficult separation of by-products [11]. More recently, enzyme-catalyzed transesterification has led to the development of recyclable and environmentally friendly catalysts. Such a catalyst has been considered to be the most promising prospective technology for biodiesel production [12].
Compared to free lipase, immobilized lipase has the following advantages [12,13,14]: 1. the immobilized lipase is more stable and more adaptable to environmental changes; 2. immobilized lipase could be recycled and the process operates continuously; 3. immobilized lipase improves the utilization efficiency of lipase and greatly reduces the production cost. Therefore, the development of lipase immobilization technology opens a new door for the development of biodiesel. The approaches in the use of immobilized lipases to produce biodiesel mainly focus on the selectivity of lipases, oil resources, and immobilization supports [15,16].
The enzyme support plays a critical role in the properties of the immobilized biocatalyst [17,18,19,20,21]. Inorganic materials such as carbon-based supports, silica-based supports, and magnetic particles exhibit good organic solvent stability, high surface area and pore size, and excellent mechanical properties, having been applied as supports for the immobilization of a wide range of lipases [22,23,24]. Silica clay (SC) is an industrial mineral raw material composed of silicon dioxide, metallic oxides and a few organic impurities. The wide distribution and low-cost provide the foundation for application as a potential support to prepare active and stable biocatalysts [25]. Moreover, SC is selected for its high specific surface area, facilitating the attachment of an increased number of lipases onto the surface of the support. The surface modification of SC by 3-aminopropyltriethoxysilane (APTES) can improve particle dispersion to prevent particle aggregation and increase the surface hydrophobicity to activate the active state of the enzyme [26,27].
In this work, a widely available material, silica clay, was used as the immobilization support of lipase. The silanized SC was prepared and used as the support for the immobilization of Eversa Transform 2.0 by covalent coupling with glutaraldehyde to synthetize biodiesel. It is shown that the immobilized biocatalyst retains a high activity and specificity for biodiesel conversion.

2. Results and Discussion

2.1. Characterization of the Modification and Immobilization

FT-IR spectra of silica clay (SC), 3-aminopropyltriethoxysilane (APTES)-modified SC (A-SC), and the immobilized lipase on glutaraldehyde-activated A-SC (Lip–Glu–A-SC) are shown in Figure 1. For the pure silica clay (Figure 1a), the bands at 3429 cm-1 and 1645 cm-1 were assigned to the Si–OH stretching and bending vibrations [28]. While the bands at 1091 cm-1 and 795 cm-1 were assigned to the Si–O stretching vibration and Si–O–Si symmetrical stretching vibration, respectively. For the A-SC (Figure 1b), the characteristic band at 2928 cm-1 is associated with the bending vibration of C–H from (APTES), thereby implying that the APTES was successfully coated onto the support [29]. For the spectrum of Lip–Glu–A-SC (Figure 1c), the band at 1645 cm-1 is assigned to the amide I band arising from the lipase protein. Furthermore, a new band that appeared at 1550 cm-1 could be due to the amide Ⅱ band [30]. The results confirm that the modification and immobilization processes were successful.
The N, H, C, and S contents of SC, A-SC, glutaraldehyde-activated A-SC (Glu–A-SC), and Lip–Glu–A-SC are shown in Table 1. Compared to pure SC, the nitrogen, hydrogen and carbon contents of A-SC were significantly increased, which is attributed to the successful introduction of APTES. For Glu–A-SC, with the increase in the carbon and hydrogen contents, the nitrogen content decreased from 1.85% to 1.23%, which confirms the successful bonding of glutaraldehyde. The change in sulfur content from 0% to 0.24% in Lip–Glu–A-SC indicates that the sulfur-containing protein was immobilized on the support.
The TEM image of the pure SC shows an almost homogeneous spherical or ellipsoidal shape and a smooth surface (Figure 2a). However, upon the successful bonding of APTES onto the SC, A-SC appears to have a rougher surface and a larger particle size (Figure 2b). As for the Glu–A-SC, the overlap between the support particles is so great that the size of the individual particles cannot be distinguished effectively (Figure 2c). The immobilization process slightly increased the average size of particles and the particle dispersion, confirming that the lipase molecules were well bonded onto the surface of support particles (Figure 2d).
The N2 adsorption–desorption isotherms obtained for the pure SC and Lip–Glu–A-SC are presented in Figure 3. The isotherm profiles were evaluated to be type-IV and have a type-H3 hysteresis loop, which is characteristic of mesoporous substances [30]. The pure SC had a BET surface area of 73.56 m2/g, while that of Lip–Glu–A-SC decreased to 42.97 m2/g, which was in good agreement with the previously reported values [31]. However, the average pore size and pore volume was the same for both. The results illustrate that the modification and immobilization processes occur only on the surface of the support, which is in good agreement with the TEM results. The XRD profiles of SC, A-SC, Glu–A-SC, and Lip–Glu–A-SC are shown in Figure 4. The main component of SC is silicon oxide, which is an amorphous material with characteristic wide peaks within the 2θ range of 15°~30° [22,32,33]. The XRD patterns are principally characterized by a wide peak centered at 21.65°. The peak centered at 26.57° can be ascribed to a small percentage of crystal impurities, with similar results having been reported in the previous literature. The XRD results indicate that the modification, crosslinking, and immobilization processes did not destroy the structure of the support, which also confirmed the successful immobilization of lipase.

2.2. Optimization of the Immobilization Process

The effects of glutaraldehyde concentration on the immobilization process were assessed for a lipase concentration of 10 mg/mL and various concentrations of glutaraldehyde in the range of 1% to 7% (v/v) at 30 °C and pH 7.5 for 60 min. As shown in Figure 5a, the optimal activity of immobilized lipase was achieved at a glutaraldehyde concentration of 1%. With a gradual increase in glutaraldehyde concentration above 1%, the activity of Lip–Glu–A-SC decreased. This implies that the introduction of a covalent bond might change the spatial conformation of immobilized lipase, leading to a reduction in the lipase activity. On the other hand, the immobilization efficiency was enhanced from 70% to 78% when the glutaraldehyde concentration was increased from 1% to 3%. However, the immobilization efficiency decreased for a glutaraldehyde concentration above 3%. The presence of excess glutaraldehyde may trigger self-crosslinking, resulting in a decrease in the number of free aldehyde groups on the surface of the support. In addition, the presence of excessive glutaraldehyde may cause excessive aldehyde groups to cross-link with the enzymes [31]. Therefore, as the concentration of glutaraldehyde further increases, the activity of the immobilized enzyme decreases. To achieve a compromise between the efficiency and cost, glutaraldehyde at a concentration of 2% was employed for the following immobilization process.
The effects of lipase concentration on the immobilization process were assessed for a glutaraldehyde concentration of 2% at 30 °C and pH 7.5 for 60 min by varying the lipase concentration within the range of 5–35 mg/mL. The amount of bound protein was enhanced with the increase in lipase concentration, reaching a maximum efficiency of bound protein at the highest lipase concentration (35 mg/mL) (Figure 5b). The highest activity of immobilized lipase was achieved for a lipase concentration of 20 mg/mL. As the lipase concentration increased above 20 mg/mL, the lipase activity decreased gradually. The reason for this may be that as the lipase concentration increases after binding, the considerable number of lipases will increase the steric hindrance between enzymes, leading to a decrease in lipase activity [26]. When the enzyme concentrations are 15 mg/mL and 20 mg/mL, respectively, there is almost no difference in enzyme activity. From an economic perspective, 15 mg/mL was employed for the following immobilization process.
The effects of contact time on the immobilization process were assessed with a glutaraldehyde concentration of 2% and a lipase concentration of 15 mg/mL at 30 °C and pH 7.5 by varying the contact time. The effect of contact time on the immobilization process is shown in Figure 5c. It can be seen that the relative activity reached the maximum in 60 min and maintained a fast growth rate before 60 min. The growth rate gradually slowed after 60 min. The amount of bound protein reached a relatively high value at 60 min. The effects of temperature on the immobilization process were assessed for a glutaraldehyde concentration of 2% and lipase concentration of 15 mg/mL at pH 7.5 for 60 min by varying the temperature from 20 to 50 °C. As illustrated in Figure 5d, the bound protein tended to increase with the increase in the temperature ranging from 20 to 30 °C, reaching the maximum immobilization efficiency at 30 °C. However, at a temperature higher than 30 °C, a relatively large decrease in bound protein was observed. The highest activity was acquired at 25 °C.

2.3. Immolization of Lipase on Modified Silica Clay

The application range and temperature tolerance are critical factors regarding the immobilized lipase for biodiesel production. Both free and immobilized lipase reach their maximum hydrolytic activity at 50 °C (Figure 6a). A dramatic decrease in free lipase activity was noticed within the temperature range from 60 to 70 °C, while a high activity of 90% was maintained, even when the Lip–Glu–A-SC was incubated at 70 °C. It should be noted that the activity of the immobilized lipase was stable at a wider range of temperatures than that of the free lipase. A similar result was obtained by Arica et al. [34].
Figure 6b shows the activities of free and immobilized lipases that were kept in phosphate buffer solution (0.1 mol/L) at 40 °C. To compare with the free lipase, the optimal pH value of immobilized biocatalyst shifted from 7.0 to 8.0. This change may be due to the covalent crosslinking technique. The relative activities of immobilized lipase remained at a high level within the pH range of 4.0 to 9.0, implying that the support expressed inertly, protecting lipase from inactivation or denaturation and remaining in its active form.
Figure 6c illustrates the activities of Lip–Glu–A-SC immobilized lipase kept in phosphate buffer solution (0.1 mol/L, pH 7.5) at 60 °C for 0~9 h. The relative activity of free lipase after 9 h is only 46% of its original activity, but conversely, that of the immobilized enzyme is about 90%. The results show that Lip–Glu–A-SC is more tolerant than the free lipase at a relatively high temperature. The increase in the thermal stability may be due to molecular rigidity, which is improved by immobilization. The remarkable improvement in thermal tolerance makes modified SC a popular support in biodiesel production, even in various industrial applications [27].

2.4. Application of Lip–Glu–A-SC for Synthesis of Biodiesel

The factors related to the conversion yield of biodiesel are shown in Figure 7. The conversion yield of biodiesel increased from 70.6% to 85.0 % when the dose of the immobilized lipase (11.5 mg protein per gram of immobilized lipase, see Figure S1) increased from 0.2 to 0.6 g (Figure 7a). As the dose of immobilized lipase was over 0.6 g, the conversion yield decreased slightly. This result may be due to the fact that an excess of immobilized lipase reduces the mass transfer efficiency of the reaction. This result may be due to the fact that an excess of immobilized lipase reduces the mass transfer efficiency [21]. Therefore, the dose of the immobilized lipase was set at 0.6 g in the following experiments.
The effect of the molar ratio of methanol/oil on the conversion yield of biodiesel is shown in Figure 7b. The conversion yield increased when the methanol/oil molar ratio was increased from 2:1 to 4:1. The highest conversion at 85.7% was achieved when the molar ratio of the methanol/oil was 4:1. A further increase in the molar ratio resulted in a decrease in the conversion yield. This may be due to the fact that excess methanol will reduce the activity of the immobilized lipase [35]. It has been reported that excess methanol is capable of driving the conversion reaction towards biodiesel [5]. The excessive short-chain alcohols could destroy the three-dimensional structure of lipase molecules, affecting the transesterification efficiency dramatically.
The effect of synthesis temperature on the yield of the biodiesel catalyzed by Lip–Glu–A-SC is shown in Figure 7c. The conversion yield of the biodiesel increased from 72% to 85.7% as the reaction temperature increased from 20 to 45 °C. However, a small decrease in the conversion yield could be observed as the temperature increased above 45 °C, which was probably due to the high temperature resulting in the inactivation of the immobilized lipase [36,37].
The effect of contact time on the conversion yield of biodiesel is shown in Figure 7d. For comparison, the conversion yield of the same amount of free lipase was at 75.2%. As observed, the conversion yield was at 86% when Lip–Glu–A-SC was added within 30 h of reaction. The conversion yield remained stable for a contact time between 30 and 45 h; therefore, 30 h was chosen as the optimal contact time in this work. The increase in the conversion yield after immobilization may due to the interfacial activation of lipases induced by capped-amino groups on the SC and the improved mass transfer efficiency of reactants around modified of silica clay [22].

2.5. Reusability of Lip–Glu–A-SC

The effect of recycling immobilized lipases on their activity is described in Figure 8. The catalytic efficiency of the immobilized lipase in the transesterification process only exhibited a small decrease and more than 67% of the activity can was retained after five cycles.

3. Materials and Methods

3.1. Materials and Reagents

Lipase Eversa Transform 2.0 (E.C. 3.1.1.3) was obtained from Novozymes China (Beijing, China). 3-aminopropyltriethoxysilane (APTES) and n-hexane (chromatographic grade) were purchased from Aladdin (Shanghai, China). Methyl heptadecanoate was purchased from Sigma-Aldrich China (Shanghai, China). Silica clay (SC) was obtained from Create New Material Company (Ningbo, China). The canola oil (99.9% fat, 0% free fatty acid, 0% protein, and 0% water) was purchased from a local market (Chengdu, China). All other reagents were of analytical grade and were used without further purification.

3.2. Modification of Silica Clay and Immobilization of Lipase

3.2.1. Modification of Silica Clay

A quantity of 4.0 g of SC was dispersed in 30 mL of ethanol for 30 min using ultrasonication. Then, 8 mL of 3-aminopropyltriethoxysilane (APTES) was injected into the ultrasonicated solution. The mixture was heated under reflux for 24 h under a nitrogen gas atmosphere with gentle magnetic stirring. Then, the modified support was filtered and rinsed with ethanol several times to remove the unreacted APTES. The APTES-modified SC was designated as A-SC.
Glutaraldehyde-activated support was synthesized by dispersing 1.0 g of A-SC in 10 mL glutaraldehyde solution. The mixture was incubated at 30 °C with gentle shaking. Then, the support was rinsed several times to eliminate the excess glutaraldehyde solution with 0.1 mol/L sodium phosphate buffer solution (pH 7.5). The activated support was dried at 60 °C to a constant weight and was designated as Glu–A–SC.

3.2.2. Immobilization Process

Ten milliliters of 0.1 mol/L sodium phosphate buffer (pH 7.5) containing 5–35 mg/mL of lipase and 1.0 g of Glu–A-SC was mixed with constant stirring for 20–180 min at 20–50 °C. Then, to remove the free lipase, the supports were washed three times with 0.1 mol/L sodium phosphate buffer (pH 7.5). The immobilized lipase was dried at 60 °C to a constant weight and was designated as Lip–Glu–A-SC. Scheme 1 illustrates the scheme of the preparation process.

3.3. Application of Lip–Glu–A-SC for Biodiesel Preparation

The silica clay immobilized lipase (Lip–Glu–A-SC) was applied as the biocatalyst for biodiesel synthesis. A typical transesterification reaction mixture consisted of 4.4 g of canola oil, a set amount of Lip–Glu–A-SC and 5 mL of n-hexane (with a pH of 6) at 45 °C. Methanol was added three times, with 0.16 g of methanol added dropwise each time. Once the reaction was completed, the immobilized lipase was separated by centrifugation and the supernatant was distilled under vacuum to remove excess methanol. An Agilent 7890 gas chromatographer (Agilent, Palo Alto, CA, USA) equipped with a flame ionization detector (FID) and a capillary column (EC-TM-WAX, 30 m × 0.32 mm ID × 0.25 μm film thickness) was used for the determination of the total fatty acid methyl esters content and N2 gas was chosen as the carrier gas for this process. The test conditions were as follows: the split ratio was 10:1, and the temperature of the injector and detector were both set at 300 °C. The column temperature was set at 160 °C for 3 min, and then heated to 240 °C at a rate of 10 °C/min and maintained at this temperature for 29 min. The injection volume was 1.0 μL. The yield of biodiesel (%) was calculated using Equation (1):
w t % = Σ Ai − A M H A M H × C M H V M H × 100 W
where wt% is the yield of biodiesel; ΣAi and AMH are the total peak area of fatty acid methyl esters and peak area of methyl heptadecanoate, respectively; CMH and VMH are the concentration (mg/mL) and volume (mL) of methyl heptadecanoate, respectively; W is the weight (mg) of the sample.

3.4. Hydrolytic Activity Assay

The hydrolytic activities of free or immobilized lipase were evaluated by an olive oil emulsion containing 4% (w/v) of polyvinyl alcohol 1750 (PVA 1750). The lipase sample was added to a mixture of 4 mL of olive oil emulsion and 5 mL of phosphate buffer solution (0.1 mol/L pH 7.5). The mixture was incubated at 40 °C for 15 min and titrated with 0.05 mol/L of sodium hydroxide solution, using 1% of phenolphthalein as the indicator. One unit of lipase activity was defined as the amount of enzyme which liberated 1 μmol of free fatty acid per min under assay conditions. The control, 15 mL of 95% ethanol, was added before the addition of enzyme.

3.5. Determination of Immobilization Efficiency

The protein content in the solution before and after immobilization was determined using the Lowry assay [26]. Bovine serum albumin (BSA) was used as the standard protein. The amount of bound protein on the support was calculated using Equation (S1). The immobilization efficiency (η) was calculated using Equation (2):
η % = ( C i − C f ) V i C i V f   × 100 %
where Ci (umol/mL) and Cf (umol/mL) are the initial and final lipase protein concentrations during the immobilization process, respectively. Vi and Vf are the initial and final volumes (mL) of the reaction solution, respectively. All samples were run in triplicate.

3.6. Characterization

The C, H, N and S contents were obtained by an elemental analyzer (Thermo Fisher Scientific, Boston, MA, USA). Transmission electron microscope (TEM) images of the samples were obtained using a Tecnai G2 F20 S-TWIN transmission electron microscope (FEI, Hillsboro, OR, USA), operating at 150 kV. The nitrogen adsorption isotherms of the samples were detected by a quanta chrome autosorb IQ-MP-MP-AG gas adsorption analyzer (Anton Paar, Shanghai, China). The specific surface area was determined using the Barrett–Emmett–Teller (BET) equation. The pore size and pore volume distribution were measured using the Barrett–Joyner–Halenda (BJH) model. X-ray diffraction (XRD) patterns were recorded on an X’ Pert Pro MPD DY129 X-ray diffractometer (Oxford, UK). Fourier transform infrared (FTIR) spectra were obtained using a Nicolet Is 10 spectrometer (Thermo Fisher Scientific, Boston, MA, USA) with a KBr pellet in a range from 400 to 4000 cm−1.

4. Conclusions

In this study, silica clay was modified by APTES and then crosslinked with glutaraldehyde for the immobilization of lipase (Lip–Glu–A-SC). The prepared Lip–Glu–A-SC was applied as a biocatalyst in the synthesis of biodiesel. The immobilization led to a remarkable improvement in the thermal and pH tolerance of Lip–Glu–A-SC in comparison to those of the free lipase. Lip–Glu–A-SC was proven to be an effective catalyst for biodiesel conversion under optimized conditions. The conversion yield of biodiesel was 67% after reusing the immobilized lipase for five cycles in an n-hexane system. The work shows that the cheap and easily obtained silica clay should be a potential support for the immobilization of enzymes.

Supplementary Materials

The following are available online at https://www.mdpi.com/2073-4344/10/11/1266/s1, Figure S1: Effect of immobilization conditions on the relative activity and the amount of bound protein, Figure S2: Gas chromatography of biodiesel synthesized by immobilized lipase.

Author Contributions

H.-m.C., Q.-eW. and T.Z. conceived and designed the experiments; T.Z. and Y.-d.D. performed the experiments; T.Z., Q.-eW. and H.-m.C. analyzed the data; T.Z. and H.-m.C. wrote the paper. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the Open Research Fund Program of Key Laboratory of Cosmetic, China National Light Industry, Beijing Technology and Business University, (grant number: KLC-2020-YB05).

Acknowledgments

The authors thank Xuekai Wang (Analytical & Testing Center, Sichuan University) for his help in GC detection.

Conflicts of Interest

The authors declare no conflict of interest.

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Catalysts 10 01266 g001
Figure 1. FTIR spectra of (a) silica clay (SC), (b) 3-aminopropyltriethoxysilane (APTES)-modified SC (A-SC) and (c) Lip–Glu–A-SC.
Figure 1. FTIR spectra of (a) silica clay (SC), (b) 3-aminopropyltriethoxysilane (APTES)-modified SC (A-SC) and (c) Lip–Glu–A-SC.
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Catalysts 10 01266 g002
Figure 2. TEM patterns of (a) SC, (b) A-SC, (c) Glu–A-SC, and (d) Lip–Glu–A-SC. Scale bar represents 50 nm.
Figure 2. TEM patterns of (a) SC, (b) A-SC, (c) Glu–A-SC, and (d) Lip–Glu–A-SC. Scale bar represents 50 nm.
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Catalysts 10 01266 g003
Figure 3. N2 adsorption–desorption isotherms of (a) SC and (b) Lip–Glu–A-SC. The insets to part (a,b) were the pore size distribution.
Figure 3. N2 adsorption–desorption isotherms of (a) SC and (b) Lip–Glu–A-SC. The insets to part (a,b) were the pore size distribution.
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Catalysts 10 01266 g004
Figure 4. XRD patterns of (a) SC, (b) A-SC, (c) Glu–A-SC, and (d) Lip–Glu–A-SC.
Figure 4. XRD patterns of (a) SC, (b) A-SC, (c) Glu–A-SC, and (d) Lip–Glu–A-SC.
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Catalysts 10 01266 g005
Figure 5. Effect of immobilization conditions on the relative activity and bound protein efficiency: (a) glutaraldehyde concentration, (conditions: lipase 10 mg/mL; temperature, 30 °C; time, 60 min); (b) lipase concentration, (glutaraldehyde 2%; temperature, 30 °C; time, 60 min); (c) time, (glutaraldehyde 2%; lipase 15 mg/mL; temperature, 30 °C); (d) temperature. (glutaraldehyde 2%; lipase 15 mg/mL; time, 60 min). The highest activity value for each group is set as 100%.
Figure 5. Effect of immobilization conditions on the relative activity and bound protein efficiency: (a) glutaraldehyde concentration, (conditions: lipase 10 mg/mL; temperature, 30 °C; time, 60 min); (b) lipase concentration, (glutaraldehyde 2%; temperature, 30 °C; time, 60 min); (c) time, (glutaraldehyde 2%; lipase 15 mg/mL; temperature, 30 °C); (d) temperature. (glutaraldehyde 2%; lipase 15 mg/mL; time, 60 min). The highest activity value for each group is set as 100%.
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Catalysts 10 01266 g006
Figure 6. Effects on the stability of immobilized and free enzymes. (a) temperature, incubated at pH 7.5 for 15 min; (b) pH, incubated at 40 °C for 15 min; and (c) time, incubated at pH 7.5 and 60 °C.
Figure 6. Effects on the stability of immobilized and free enzymes. (a) temperature, incubated at pH 7.5 for 15 min; (b) pH, incubated at 40 °C for 15 min; and (c) time, incubated at pH 7.5 and 60 °C.
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Catalysts 10 01266 g007
Figure 7. Effects on the transesterification yield: (a) immobilized lipase, (conditions: methanol/oil molar ratio, 4:1; temperature, 40 °C; time, 36 h); (b) methanol/oil molar ratio, (dose of the immobilized lipase, 0.6g, temperature, 40 °C; time, 36 h); (c) temperature, (dose of the immobilized lipase, 0.6 g; methanol/oil molar ratio, 4:1; time, 36 h); and (d) time, (dose of the immobilized lipase, 0.6 g, methanol/oil molar ratio, 4:1; temperature, 40 °C).
Figure 7. Effects on the transesterification yield: (a) immobilized lipase, (conditions: methanol/oil molar ratio, 4:1; temperature, 40 °C; time, 36 h); (b) methanol/oil molar ratio, (dose of the immobilized lipase, 0.6g, temperature, 40 °C; time, 36 h); (c) temperature, (dose of the immobilized lipase, 0.6 g; methanol/oil molar ratio, 4:1; time, 36 h); and (d) time, (dose of the immobilized lipase, 0.6 g, methanol/oil molar ratio, 4:1; temperature, 40 °C).
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Catalysts 10 01266 g008
Figure 8. Reusability of Lip–Glu–A–SC for biodiesel production. Reaction conditions: methanol/oil molar ratio,4:1; immobilized lipase dosage, 0.6 g; reaction time, 30 h; and reaction temperature, 45 °C.
Figure 8. Reusability of Lip–Glu–A–SC for biodiesel production. Reaction conditions: methanol/oil molar ratio,4:1; immobilized lipase dosage, 0.6 g; reaction time, 30 h; and reaction temperature, 45 °C.
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Scheme 1. Scheme of the preparation of immobilized lipase on APTES-modified silica clay.
Scheme 1. Scheme of the preparation of immobilized lipase on APTES-modified silica clay.
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Table 1. The nitrogen, hydrogen, carbon, and sulfur contents of the samples.
Table 1. The nitrogen, hydrogen, carbon, and sulfur contents of the samples.
SamplesN (%)C (%)H (%)S (%)
SC0.880.270.460
A-SC1.853.781.080
Glu–A-SC1.234.681.360
Lip–Glu–A-SC2.065.211.730.24
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Zou, T.; Duan, Y.-d.; Wang, Q.-e.; Cheng, H.-m. Preparation of Immobilized Lipase on Silica Clay as a Potential Biocatalyst on Synthesis of Biodiesel. Catalysts 2020, 10, 1266. https://doi.org/10.3390/catal10111266

AMA Style

Zou T, Duan Y-d, Wang Q-e, Cheng H-m. Preparation of Immobilized Lipase on Silica Clay as a Potential Biocatalyst on Synthesis of Biodiesel. Catalysts. 2020; 10(11):1266. https://doi.org/10.3390/catal10111266

Chicago/Turabian Style

Zou, Ting, You-dan Duan, Qiao-e Wang, and Hai-ming Cheng. 2020. "Preparation of Immobilized Lipase on Silica Clay as a Potential Biocatalyst on Synthesis of Biodiesel" Catalysts 10, no. 11: 1266. https://doi.org/10.3390/catal10111266

APA Style

Zou, T., Duan, Y.-d., Wang, Q.-e., & Cheng, H.-m. (2020). Preparation of Immobilized Lipase on Silica Clay as a Potential Biocatalyst on Synthesis of Biodiesel. Catalysts, 10(11), 1266. https://doi.org/10.3390/catal10111266

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