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Article Cite This: ACS Omega 2018, 3, 18203−18213
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Synthesis of Rhizopus arrhizus Lipase Nanoparticles for Biodiesel Production Rohit Kumar Sharma,†,‡ Manoj Saxena,†,‡,∥ Crystal A. O’Neill,†,§ Hector A. R. Ramos,†,‡ and Kai Griebenow*,†,‡,§
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†
Molecular Sciences Research Center, University of Puerto Rico, 1390 Ponce de Leon Avenue, Suite 2, San Juan, Puerto Rico 00931-3346, United States ‡ Department of Environmental Science, University of Puerto Rico-Río Piedras, P.O. Box 70377, San Juan, Puerto Rico 00936-8377, United States § Department of Chemistry, University of Puerto Rico, Río Piedras Campus, 17 Ave. Universidad STE 1701, San Juan, Puerto Rico 00925-2537, United States S Supporting Information *
ABSTRACT: We developed a nanoparticulate Rhizopus arrhizus lipase formulation to enhance its activity and to increase the conversion yield of lipids into fatty acid methyl esters (FAME, a.k.a., biodiesel). More than 95% purity of the lipase was achieved in a two-step purification. Nanoparticle formulation was afforded by co-lyophilization of the lipase with methyl-βcyclodextrin (MβCD), an established lyoprotectant. Scanning electron microscopy and dynamic light scattering measurements showed a size of 75− 200 nm for the nanoparticles depending on the ratio of lipase-to-MβCD employed during co-lyophilization. Fourier transform infrared spectroscopic analysis by Gaussian curve fitting of the resolution-enhanced amide I region of lyophilized and nanoparticulate lipase indicated a more native-like secondary structure in the latter. A 98% substrate-to-FAME conversion was achieved in 10 h in n-hexane by lipase nanoparticles, whereas the crude and lyophilized enzyme showed 65 and 70% conversion in 18 h, respectively. In this aspect, the lipase nanoparticles were superior to all other reported systems. Operational stability after 5 catalytic conversions of nanoparticles was found to be >81%. In summary, we herein developed a novel lipase formulation for efficient catalysis in lipid-to-biodiesel conversion.
1. INTRODUCTION In recent decades, biodiesel has emerged as an alternative fuel with reduced carbon footprint, in particular, when using renewable sources of lipids. The two important factors to assure commercial success in biodiesel production are the choice of economically sustainable raw materials and employment of an economic and efficient catalyst. For that reason, optimization of both, the biomass as well as the (bio)catalyst. has been at the heart of biodiesel research in recent years.1−5 Lipases offer advantages over conventional catalysts in biodiesel production including the production of only negligible amounts of wastewater, better product yield and quality, reduced energy costs because of its ability to catalyze at low (ambient) temperatures, being a renewable catalyst, and displaying prominent stability in nonconventional environments.3,6 To improve the operational stability of the biocatalyst, immobilization of lipases is frequently being performed, which also facilitates efficient recovery of the enzyme after the reaction.2,3,6,7 However, immobilization can be detrimental to enzyme activity, i.e., by causing steric hindrance for substrate binding. The support material chosen for immobilization may © 2018 American Chemical Society
also lead to interfacial inactivation during synthesis or use. Poor mass transfer due to high loading of the enzyme on the support material or incorrect choice of support material are a few examples of conditions that can lead to reduced catalytic efficiency.6 Additionally, inactivation by methanol and glycerol accumulating in pores of the support material is another problem specific to biodiesel production, which to some extent could be controlled by using a solvent of higher polarity, which assists in solvation of glycerol and reduce the destabilizing impact of methanol on the enzyme.8 In this study, we extend efforts to improve the activity and stability of a lipase catalyst in biodiesel production. However, in contrast to immobilization of the lipase on a designed nanomaterial, we formulated nanoparticles of the lipase itself by the phase separation method using MβCD as an excipient. We previously developed this method to improve the stability and function of proteins in pharmaceutical and nonaqueous enzymology applications.9−11 Furthermore, it has been Received: August 16, 2018 Accepted: December 10, 2018 Published: December 24, 2018 18203
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Table 1. Comparative Data of Ester Conversion by Lipases in Different Solvent Systems substrate/alcohol soybean oil and methanol acetic acid and butanol oleic acid and butanol soybean oil and methanol soybean oil and methanol olive oil and methanol
lipase R. R. R. R. R. R.
oryzae oryzae oryzae oryzae cells oryzae cells arrhizus
system
time (h)
% conversion
reference
solvent-free hexane and heptane hexane aqueous aqueous hexane
>45 24 >50 50 70 12
80−90 76−80 75−83 90% 90 99
41 42 44 49 60 this study
Figure 1. (a) Effect of the pH on the activity of purified R. arrhizus lipase determined by olive oil titration. Lipase was incubated in buffers at various pH for 10−12 h before titration. (b) Effect of the reaction temperature on lipase activity at pH 6.5 determined by using p-nitrophenyl laurate as substrate. The purified enzyme was incubated at different temperatures (20−50 °C) in a water bath for 30 min before using it in the reaction. The condition with the highest activity was defined as 100%. Error bars represent the average ± standard deviation of three replicates (****p ≤ 0.0001, **p ≤ 0.001, *p ≤ 0.05, ns p > 0.05 calculated by one-way analysis of variance (ANOVA) mutliple comparison test).
lyophilization or other dehydration methods cannot remove all protein-bound water. Optimum enzyme activity depends on a minimum amount of water in the system, which varies from enzyme to enzyme and strongly depends on the solvent system. High conversion can only be accomplished when the right combination of the biocatalyst, water, and the solvent is chosen.14,15,21 Halling and co-workers found that this phenomenon is best described by the water activity in the system.22,23 Lipase from Rhizopus spp. has been utilized to produce biodiesel from the various combinations of oils and alcohols but mostly immobilized lipase was employed in these studies (Table 1). We selected olive oil as our model lipid source because olive oil-derived biodiesel studies are rare.24 Besides, the fatty acid composition of typically used cooking oil and brown grease usually have high percentage of oleic, linoleic, and palmitic acids, which are the main constituents of olive oil. Thus, a good FAME yield using olive oil will be an encouraging step in guiding production of biodiesel from waste cooking oils.25,26 The lipase nanoparticles were used to produce biodiesel, which demonstrated a faster and more complete catalytic conversion of olive oil-to-FAME when compared to crude and purified lyophilized lipase. In absence of the supporting materials necessary for immobilization, this method provides a superior catalyst that has an advantage in better mass transfer and fewer opportunities of being deactivated by glycerol during the reaction.
established that MβCD leads to improved activities of serine proteases12 as well as lipases in organic solvents.13 For this work, lipase from R. arrhizus was selected as our model enzyme, which was purified to more than 95% purity prior to nanoparticle synthesis. Several process parameters were optimized. Determination of the optimum pH is imperative to achieve the most efficient lipase nanoparticles for biodiesel production since our procedure involves lyophilization. Under such conditions, enzymes have a so-called pH-memory in organic solvents14,15 because the ionization state of the active site residues is maintained during lyophilization and is critical for optimum activity in organic solvents.16 The enzyme will only work efficiently in the nonaqueous environment when the protonation of the side chains is the same as required in the aqueous system for optimum activity. Similarly, employing additives prior to the freeze-drying step can improve the activity of the dehydrated enzymes in organic solvents, probably by various mechanisms ranging from structural preservation of the enzyme to improved structural mobility, active site, and substrate solvation properties, among others.17 In several studies, the possible mechanism(s) of such activity enhancement was experimentally evaluated.9,10,17−19 The optimum ratio of lipase-to-MβCD was experimentally evaluated based on previous findings that showed that MβCD works only at an optimum concentration, after which enzyme activity starts decreasing.9,11,12,20 Subsequently, we also optimized the water amount in the biodiesel reaction because water is one of the key factors for the catalysis in nonaqueous media. A minimum amount of water is always present because 18204
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methyl acetate even after incubating it for several hours.33,34 At ratios of 1:4 resulted in the formation of polydisperse and large nanoparticles, which might be due to interference during the phase separation as the solution became too concentrated (Figure 2e,f).11 In the case of both crude and lyophilized enzymes, heterogeneity decreased upon an increase in the MβCD content (Figure 2, and Additional Information, Figure S6). An inverse dependency of the specific activity on the size of the protein particles was observed. For example, the activity of the lyophilized lipase increased 7, 32, and 48 fold compared to the crude lipase, for the various lipase-MβCD co-lyophilizates, whereas the size of the nanoparticles decreased in the same order up the 1-to-4 ratio. At the 1:6 ratio, the activity decreased compared to 1:4 ratio, and an increment in the size of the nanoparticles was observed accordingly (Table 2). It has to be pointed out, however, that the size is only one factor contributing to the activity increase in nanoparticles compared to the lyophilized enzyme. Other factors identified include reduced lipase structural rigidity in the presence of MβCD, which in turn help enzyme to attain more active conformation in the organic solvent.9 Additionally, colyophilization of enzymes with MβCD stabilizes the protein structure and prevents structural changes during lyophilization, which contributes to increased activity in organic solvents.9,11 Finally, the nanoparticulate nature of the catalyst reduces potential substrate diffusional limitations that can occur in solid-phase catalysis.21 The lower activity of enzymes in nonaqueous systems has been a drawback in applications, and development of strategies overcoming this issue has been an area of ongoing research during the last few decades. To verify the effect of MβCD on the lipase substrate-to-FAME conversion without co-lyophilization, a control experiment was performed where MβCD was added directly at the 1:4 mass ratio to the reaction mixture. No effect of MβCD was observed in the FAME conversion. This result is in the agreement with previous findings,9 which revealed that MβCD has to be co-lyophilized with an enzyme to obtain highly active formulations, presumably to prevent dehydration-induced structural changes. 2.3. FTIR Spectroscopy of Lyophilized and Nanoparticles in Solid Form and Secondary Structure Prediction by Curve Fitting of the Amide I Band. Being sensitive toward the secondary structure of proteins, the amide I vibrational mode provides quantitative information on protein secondary structure.35−37 Purified samples of lyophilized and nanoparticulate lipase were analyzed in solid form as KBr pellets, and the inverted calculated second derivative spectra are shown in Figure 3a,b. There were around seven spectral components visible in both samples, but spectra of the nanoparticles were better resolved as indicated by better band separation. The curve fitting was performed with seven bands and starting parameters of the fitting guided by the second derivative spectra. Results of the curve fitting are shown in (c) and (d) and are summarized in detail in Table 3. Frequencies of the secondary structure components α-helix, β-sheets, βturn, and unordered structures were compared between second derivatization and curve fitting spectra and percentage of each component was calculated as shown in Table 3. For most of the spectral component frequencies, differences between second derivative and a curve fit were not more than 3
2. RESULTS AND DISCUSSION 2.1. Effects of pH and Temperature. We found the optimum pH of the lipase to be 6.5 (Figure 1a), but no significant reduction of relative activity was observed at pH 7 compared to 6.5, which is in agreement with other reports on similar lipases or closely related species.27−29 The temperature optimum of activity was found to be 40 °C (Figure 1b), which is also close to that found for a similar species.30 2.2. Nanoparticle Activity, Optimization, and Characterization. At 0.15 M salt concentration, we obtained the maximum activity of the lyophilized enzyme in the transesterification model reaction of 4-nitrophenyl laurate with ethanol in n-hexane (Additional Information, Figure S4). This concentration was selected for all subsequent experiments. The improved activity of the lyophilized lipase triggered by salts in organic solvents could be due to several factors. First, due to the presence of a salt matrix, the chances of protein deactivation by organic solvents get minimized because salts reduce the direct contact of protein with the surrounding solvent layer. Second, salts increase the catalytic turnover by providing better hydration of the protein. Thus, at an optimum concentration of salts, the enzyme may have a more native-like structure in organic solvents.17,31 Nanoparticles were formed by phase separation as shown in Scheme 1 during the freezing process as described by Morita et Scheme 1. Formation of Lipase Nanoparticles by the Phase Separation Method
al.,32 which depends on the right concentrations of protein and excipient. In our study, the optimum ratio of protein-toexcipient was 1:4 (w/w) because the nanoparticles formed were the smallest (Figure 2d) and had the highest activity (Table 2). To obtain the nanoparticles, the lipase-MβCD colyophilizate was washed with ethyl acetate (Figure 2c). We selected ethyl acetate for washing of the dehydrated lyophilized powder because ethyl acetate does not affect the activity of many proteins in such procedures.11 Similarly, others reported that the activity of the lipase was not affected by ethyl or 18205
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Figure 2. Scanning electron microscopy (left panel) and dynamic light scattering (right panel) results of the nanoparticles formed by different weight ratios of lipase-to-MβCD (w:w). (a, b) 1:2, (c, d) 1:4, (e, f) 1:6. More information is given in Table 2.
cm−1. The percentage of α-helix content increased from 23.33 to 32.76%, whereas the percentage of β-sheets decreased from 53.62 to 34.41% from lyophilized enzyme to nanoparticles. The percentage of β-turns/other secondary structure did not change significantly and remained 23.03 and 22.01%,
respectively, but some unordered components of 10.81% at 1645 cm−1 were observed in nanoparticles. Other reports on the percentage of α helix of lipases from Rhizopus niveus (homologous to Rhizopus oryzae), Rhizomucor meihei and pancreatic lipase revealed the high percentage of α helix 18206
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Table 2. Estimation of Particle Size, Polydispersity Index (PDI), and Activity of Nanoparticles Formed Using Different Mass Ratios of Lipase-to-MβCD lipase/MβCD ratio (w/w)
particle radius (nm)a
polydispersity indexb (PDI)
crude lyophilized 1:2 1:4 1:6
175 ± 50 150 ± 25 75 ± 25 200 ± 25
0.47 0.68 0.70 1.00
activity in n-hexanec (μmol min−1 mg−1)
% fold activity enhancementd
± ± ± ± ±
1.00 7.00 32.00 48.00 31.00
0.17 1.205 5.41 8.23 5.21
0.084 0.034 0.259 0.640 0.229
a Determined by DLS (see Section 4.8. for details). Crude lipase was too heterogeneous to be measured. bPDI is a degree of the homogeneity of the sample. Values of less than 0.7 are an indication of a monodisperse sample. cSee Section 4.5 for details. dFor % fold activity enhancement, crude lipase activity was considered 1.00.
Figure 3. Amide I FTIR spectra after second derivatization. (a) Lyophilized enzyme. (b) Nanoparticles. FSD FTIR spectra and Gaussian curve fitting. The solid black lines represent the superimposed FSD spectra and the results of the curve fitting; individual Gaussian bands are shown in different color lines and frequency of each signal is mentioned. (c) Lyophilized enzyme. (d) Nanoparticles.
(>33%) in aqueous solution,38−40 which is very close to the percentage of α-helix in nanoparticles we found in this study. These findings agree with previous reports that co-lyophilization of proteins with MβCD reduces structural changes occurring during lyophilization.9,11 From the above quantitative data, we can conclude that like in other proteins,9,11 MβCD plays an important role in protecting the secondary structure of the lipase during lyophilization, which is also evident from the higher activity of the nanoparticles. 2.4. Effect of Water on FAME Conversion Percentage. Previous work by Kaieda et al. on R. oryzae lipase, which is a closely related species of R. arrhizus, reported that the lipase
catalyzed methanolysis efficiently in up to 30% of water.41 Similarly, Salah et al. found that the esterification reaction for the synthesis of butyl acetate ester increased dramatically when reaction conditions changed from solvent-free (no water added) to 45% (w/w) initially added water and then finally to an organic solvent, which gave the highest ester yield.42 These results emphasize that lipases from Rhizopus species can perform well in both aqueous and nonaqueous systems. However, efficient conversion in organic solvents is only possible when different parameters are precisely optimized. To achieve the high conversion of lipid-to-FAME, we optimized the initial water amount in our system. As shown in Figure 4, the reaction was performed in the presence of variable amounts 18207
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in very apolar solvent systems, such as hexane.43 A similar trend was observed by Ghamgui el al.44 for the synthesis of 1butyl oleate in both n-hexane and a solvent-free system when the reaction was performed in the absence of water. Wehtje et al.43 studied the effect of the water activity (aw) on the enzyme activity of three different lipases, including R. arrhizus. Their findings revealed that lipases from R. arrhizus have the ability to attain optimum catalytic activity at relatively low water activity, which means that less residual water is required to reach optimum catalysis in a nonaqueous environment. 2.5. Time Course Study of FAME Synthesis. Figure 5 shows a time course study of FAME synthesis catalyzed by
Table 3. Amide I Band Analysis for the Secondary Derivatives and Curve Fitting of Lyophilized and Nanoparticle Lipase in Solid Form band position protein lyophilized
nanoparticles
second derivative 1655 1616, 1629, 1640, 1675 1661, 1687 1653 1626, 1636, 1683 1662, 1672 1645
curve fitting
area %
assignmenta
1653 1615, 1628, 1640, 1675 1664, 1686 1653 1623, 1635, 1682 1663, 1672 1644
23.33 53.62
α-helix β-sheet
23.03 32.76 34.41
β-turn/other α-helix β-sheet
22.01 10.81
β-turn/other unordered
a
Band assignment according to commonly reported assignments in the amide I region.37,55
Figure 5. Kinetics of FAME synthesis by different formulations of lipase from R. arrhizus. Reaction conditions: oil-to-methanol ratio 1:3 (mol/mol), oil-to-n-hexane ratio 1:0.5 (v/v), 1% of water (wt % of the oil), and 10 mg of lipase. All reactions were performed at 40 °C and 200 rpm for 20 h on a reciprocal shaker. Error bars represent the average ± standard deviation of three replicates (****p ≤ 0.0001 and **p ≤ 0.001 calculated by two-way ANOVA Tukey’s multiple comparisons test).
Figure 4. Effect of water added to n-hexane on the lipid-to-FAME conversion on different formulations of lipase from R. arrhizus. Reaction conditions: oil-to-methanol ratio 1:3 (mol/mol), oil-to-nhexane ratio 1:0.5 (v/v); 0, 1, 2, 4, 6, 8, and 10 wt % water of the oil, and 10 mg of lipase was added to each reaction mixture. All reactions were performed at 40 °C and 200 rpm for 18 h on a reciprocal shaker. Error bars represent the average ± standard deviation of three replicates (****p ≤ 0.0001 and ***p ≤ 0.001 calculated by two-way ANOVA Tukey’s multiple comparisons test).
crude, lyophilized, and lipase nanoparticles in n-hexane. The fastest conversion was observed by the nanoparticles in the initial phase of the reaction: nearly 18% FAME conversion was obtained in the first 2 h. Only 2.4 and 8.3% of FAME synthesis were achieved by crude and lyophilized lipase preparations during the same time. After 10 h, almost 98% conversion was achieved by the lipase nanoparticle catalyst. FAME synthesis by crude and lyophilized lipases reached only around 64 and 70% conversion yield after 18 h. Increasing the reaction time further did not improve the FAME conversion in these cases due to inactivation of the lipase. In other studies, when a liquid formulation of R. oryzae was used in the methyl ester formation using soybean oil as a substrate, 90.5 wt % ester was produced in 67 h.41 In a different work, a yeast expression system was used to produce intracellular active R. oryzae lipase as a wholecell biocatalyst system to produce methyl ester from the soybean oil. However, only 71 wt% yield of methyl ester was obtained during a 165 h long cycle under aqueous conditions.45 Similarly, immobilized R. oryzae in butyl acetate synthesis took 20−25 h to reach 60% conversion,42 whereas synthesis of 1-butyl oleate took 30 h and maximum conversion reached around 75% only.44 It is obvious that with our lipase nanoparticles, a higher conversion in less time (10 h) was
of water in the reaction medium (0, 1, 2, 4, 6, 8, and 10%, w/ w). At 1% of water content, we obtained the maximum substrate-to-FAME conversion, which can probably be explained with the very nonpolar solvent system used. Only small amounts of water are needed in apolar organic solvents to achieve near maximum water activity. Accordingly, a drastic decrease in FAME conversion was observed when the water amount was increased from 1 to 2%. Subsequent further increases in the amount of water caused additional decreases in lipase activity. Nanoparticles had the highest activity under all conditions employed, which demonstrates that this preparation is per se more resistant to denaturation stress, which is a result of increasing water concentration.36 The activity decrease is likely due to increasing lipase denaturation at increasing water content. In addition, a high amount of water can favor hydrolysis over esterification.21 It is important to point out that even without the addition of water to the reaction, there is residual water bound to the enzyme after the dehydration step, which might be sufficient to achieve a reasonably active lipase 18208
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Figure 6. Effects of the methanol and ethanol concentrations on the lipid conversion to biodiesel by different formulations of lipase from R. arrhizus. (a) Methanol. (b) Ethanol. Reaction conditions are as follows: oil-to-alcohol ratio 1:3, 1:6, and 1:9 (mol/mol), oil-to-n-hexane ratio 1:0.5 (v/v), 1% of water (wt % of the oil), and 10 mg of lipase. All reactions were performed at 40 °C and 200 rpm for 18 h on a reciprocal shaker. Error bars represent the average ± standard deviation of three replicates (****p ≤ 0.0001 calculated by two-way ANOVA Tukey’s multiple comparisons test).
matrix used to immobilize the enzyme and can reduce the mass transfer rate. Ghamgui el al.44 achieved 75−83% percentage of conversion of the ester in n-hexane when they used immobilized lipase of R. oryzae, whereas we observe nearly 98% conversion by using lipase nanoparticles. 2.7. Operational Stability of Lipase Nanoparticles. Hama et al. used a packed-bed reactor and immobilized R. oryzae cells on polyurethane foam biomass particles and thus achieved 81.4% methyl ester conversion after five catalytic cycles.49 In another report, 73.8% synthesis of 1-butyl-oleate was achieved after six catalytic cycles.44 Figure 7 shows the reusability profile of all three formulations of the enzyme of
accomplished than with crude, lyophilized enzyme, and also compared with other formulations, which is of great commercial significance. 2.6. Methanol and Ethanol Effects on FAME/Ethyl Ester (EE) Conversion for Crude, Lyophilized, and Nanoparticulate Lipases. FAME/EE conversion in each reaction was determined, as shown in Figure 6. In the case of both, methanol and ethanol, nanoparticles were found to be superior to lyophilized lipase as demonstrated by a higher conversion of lipids to the esters even when high stoichiometry of alcohol was used. Although the lipid conversion in both, methanol and ethanol, decreased significantly at increasing alcohol concentration, ethanol caused somewhat less enzyme deactivation. These findings are in agreement with literature data where it has been shown that long chain alcohols cause less inactivation of the enzyme, but also give lower conversion yields.2,44 An excess amount of alcohol in biodiesel production leads to the inactivation of lipases.8,46 Although this detrimental phenomenon can be minimized by careful selection of the solvent and lipase formulation, some solvent- and alcoholinduced inactivation cannot be ruled out completely.47 Although the exact mechanism of inactivation of lipases by alcohols is not precisely known, some recent studies suggest that high concentrations of alcohol can trigger protein unfolding and irreversible inactivation.46 Enzyme structural dynamics is another factor affecting the conversion of the substrate to product. Solvents impact the structural dynamics based on their effect on water activity of the system.14,15 This interaction is related to the dielectric constant of the solvent and reflects the stability and activity of a particular enzyme in that particular environment.15,23,48 Therefore, using n-hexane as the solvent has advantages. First, it helps in dissolving the substrate, which increases the mass transfer and eventually contributes to a higher conversion percentage of esters. Second, its log P value is >3.5, which lessens the inactivation of the enzyme as it strips less essential bound water from the enzyme and increases the durability of the enzyme.21 In addition, using the enzyme in free form (mobilized) is a valuable strategy because glycerol, which is the byproduct of the reaction, can accumulate in the pores of the
Figure 7. Operational stability of biodiesel formation by different formulations of lipase from R. arrhizus. The enzyme was pelleted by centrifugation, washed 4 times with n-hexane, and dried. Reaction conditions are as follows: oil-to-methanol ratio 1:3 (mol/mol), oil-ton-hexane ratio 1:0.5 (v/v), 1% of water (wt % of the oil), and 10 mg of lipase. All reactions were performed at 40 °C and 200 rpm for 18 h on a reciprocal shaker. Error bars represent the average ± standard deviation of three replicates (****p ≤ 0.0001 and **p ≤ 0.001 calculated by two-way ANOVA Tukey’s multiple comparisons test). 18209
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deviation in the activity versus pH curve was noted when constructing the graph, and this has been described in the literature before.52 The pH at which best activity was obtained was selected for all experiments. 4.2.2. Activity at Optimum Temperature of the Purified Lipase. For determination of the optimum temperature, purified enzyme was incubated at different temperatures. A temperature ranges from 20 to 50 °C was selected, and at every 5 °C relative activity was monitored by taking a total of 5 points. Lipase samples were incubated in a water bath for 30 min,29 and activity was measured using a UV-2450 (Shimadzu, Japan) spectrophotometer equipped with a temperaturecontrolled cell holder. Briefly, a standard curve of concentration vs. optical density of p-nitrophenol was prepared in 0.1 M Tris-Cl buffer at pH 8.2. For lipase activity, an equal volume of 0.1 M Tris buffer and 420 μM of p-nitrophenyl laurate substrate solutions was mixed, and the reaction was started by adding 1 mL of purified lipase solution. Absorbance was recorded at 410 nm every 20 s for up to 15 min. From the standard curve, the concentration of hydrolyzed substrate was calculated.51 For specific activity calculations, the amount of protein in each sample was calculated, as described in Section 4.3. 4.3. Protein Concentration Determinations of Crude, Lyophilized, and Nanoparticulate Lipase. Protein concentration was determined by using the Pierce BCA Protein Assay Kit (Pierce) as per the manufacturer’s instructions. Briefly, a standard curve with bovine serum albumin (0−250 μg/mL) was constructed. From the standard curve equation, the concentrations of crude, purified, and lyophilized lipase as well as redissolved nanoparticle samples were calculated. All reactions were performed in 96-well microplate, and the absorbance at 562 nm was measured by using the Multiskan FC Microplate reader (ThermoFisher, Waltham, MA). 4.4. Formation and Optimizations of Lyophilized and Nanoparticulate Lipase. To prepare the lyophilized lipase, first the optimum concentration of buffer was experimentally evaluated. For that lipase was dissolved in five different concentrations of sodium phosphate buffer (0.025, 0.050, 0.100, 0.150, and 0.200 M at pH 6.5) and lyophilized. All lyophilized samples were tested for activity in a transesterification model reaction (see Section 4.5), and the relative activity was determined (Additional Information, Figure S4). The buffer concentration at which maximum activity was achieved was selected and used as a control enzyme for all experiments. Nanoparticles of lipase were formed by co-lyophilization from different protein-to-MβCD ratios (1:2, 1:4, and 1:6; w/ w) in 0.15 M sodium phosphate buffer at pH 6.5. Activity and size of the nanoparticles were measured, as described in Sections 4.5. and 4.8. The nanoparticles which provided the best activity were selected for all subsequent experiments. The nanoparticles were obtained as described11 with some minor modifications (Scheme 1). Briefly, lipase was dialyzed against 0.15 M sodium phosphate buffer at pH 6.5 overnight using a 10 kDa cutoff dialysis membrane. Next, 10 mg/mL of lipase protein solution was co-lyophilized with the excipient MβCD at a 1:4 ratio (w/w, protein-to-excipient) because this ratio provided lipase nanoparticles with the best activity. The amount of protein in the solution was estimated, as mentioned in Section 4.3. The resulting solution was flash-frozen in liquid nitrogen and lyophilized for 48 h using a 5 L Modulyo D freeze dryer (Thermo Savant, Waltham, MA) at a condenser
this study, which was determined by lipid-to-FAME conversion by each enzyme in five subsequent conversion batches. Recovery of the enzyme was performed by centrifugation and washing of the pellets four times with n-hexane before using them in the next batch. Nanoparticles showed excellent operational stability as still up to 81% conversion was achieved after the 5th catalytic cycle. In contrast, both crude and lyophilized enzymes showed reduced operational stability and conversions were only 16 and 43%, respectively, after the 2nd catalytic cycle and only 8 and 22% after the 3rd cycle. After the 5th batch conversion, both crude and lyophilized lipases were practically inactive. These findings clearly indicate that formulation as nanoparticles vastly improved the operational stability of the lipase in biodiesel production.
3. CONCLUSIONS In this study, we report a two-step purification strategy of R. arrhizus lipase and show that activity increased 50 fold after purification in aqueous conditions and 48 fold after nanoparticle formation in an organic solvent. Comparative studies of nanoparticles with crude and lyophilized enzymes revealed a high percentage of FAME conversion by nanoparticles and excellent operational stability. We further demonstrated that nanoparticles could be used for five conversion cycles without encountering significant loss in catalytic efficiency, which shows superior performance of nanoparticles over crude and lyophilized samples. Previous studies also confirmed that lipases from Rhizopus spp. have good stability in organic solvents and they are a good choice for ester synthesis in nonaqueous systems. Although, to achieve high product yield, optimization of catalyst, solvent system, water amount, and other parameters is necessary. 4. EXPERIMENTAL SECTION 4.1. Chemicals. Crude lipase from R. arrhizus (EC 3.1.1.3), para-nitrophenol, para-nitrophenyl palmitate, MβCD, methyl heptadecanoate, and olive oil (highly refined, low acidity grade) were purchased from Sigma-Aldrich (St. Louis, MO). The molar amount of the oil was calculated from its saponification value.50 All solvents were purchased in the anhydrous form (Sure/Seal bottles: water content below 0.005%) from Fisher Scientific (Waltham, MA) and used without further drying. 4.2. Lipase Activity in Aqueous System. The specific activities of crude and purified lipases (Additional Information, Figure S1d) were determined by the titrimetric method using olive oil/gum arabic emulsion as substrate, as described by Pinsirodom et al.51 The reaction was performed for 30 min in a 40 °C water bath. For specific activity calculations, the amount of protein in each sample was calculated, as described in Section 4.3. 4.2.1. Activity at Optimum pH of the Purified Lipase. The pH at which purified lipase displayed optimum activity was determined in the same manner as mentioned above. The only difference was that purified lipase was incubated in buffers of varying pH for 10−12 h. A pH ranges from 6.0 to 9.0 was selected, and value for every half unit was monitored by taking total 7 points. Two different buffers, 50 mM sodium phosphate pH 6.0−8.0 (pH buffering capacity within the range of pH 5.8−8.0) and 50 mM Tris-Cl pH 8.0−9.0 (pH buffering capacity within the range of 7.2−9.0), were prepared, and each sample was titrated with 0.05 N NaOH. No unexpected 18210
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ACS Omega
Article
temperature of −45 °C and a pressure of
