Synthesis of ethyl oleate by esterification in a solvent-free system using lipase immobilized on PDMS-modified nonwoven viscose fabrics

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Synthesis of ethyl oleate by esterification in a solvent-free system using lipase immobilized on PDMS-modified nonwoven viscose fabrics Weina Li a,b , Huaqing Shen a , Miaomiao Ma a , Luo Liu a , Caixia Cui a , Biqiang Chen a,∗ , Daidi Fan b , Tianwei Tan a,∗ a b

College of Life Science and Technology, Beijing University of Chemical Technology, Beisanhuan East Road 15, Beijing 100029, China Department of Chemical Engineering, Northwest University, Taibaibeilu 229, Xi’an 710069, China

a r t i c l e

i n f o

Article history: Received 9 April 2015 Received in revised form 1 July 2015 Accepted 16 July 2015 Available online xxx Keywords: Batch stirred tank reactor Esterification Lipase immobilization Nonwoven viscose fabric

a b s t r a c t A lipase from the yeast Yarrowia lipolytica was immobilized on a PDMS-modified nonwoven viscose fabric and used in the synthesis of ethyl oleate. The efficiency of immobilization improved to 75.2% when the concentrated lipase slurry (180 mg mL−1 ) was used as the source of lipase. When compared with the original immobilized lipase, the lipase immobilized on the PDMS-modified fabric exhibited more stable catalytic activity over 35 batches; exhibited a 20-fold lower affinity for oleic acid, a 42-fold less ethanol-induced inhibition; could be reused in 10 iterative 5L batch stirred tank reactor processes. When compared to that of free lipase, the pH stability range of immobilized lipase was narrower (pH 6–7 vs pH 5–8); the optimum reaction temperature was higher (40 ◦ C vs 37 ◦ C); and the thermally more stable (70% vs 5% of activity was retained after pre-incubation for 4 h at 45 ◦ C). The variation in the activity exhibited in an organic solvent could be correlated to the log P. Catalytic efficiency was ∼13-fold lower upon excessive lipase immobilization. XPS/ATR-FTIR confirmed the introduction of PDMS onto lipaseimmobilized viscose. The simple enzyme immobilization method could potentially be useful for the production of ethyl oleate at an industrial scale. © 2015 Elsevier Ltd. All rights reserved.

1. Introduction With the rapid developments in enzyme-based technologies, microbial lipases have received much attention. The yeast Yarrowia lipolytica (YLL) has found applications in several fields, such as bioconversion of hydrophobic substrates [1], biosynthesis of gold nanoparticles [2], and production of specialty lipids [3]. In

Abbreviations: YLL, yeast Yarrowia lipolytica; YlLip2, Lip2 lipase from yeast Yarrowia lipolytica; ATR, attenuated total reflectance; BSA, bovine serum albumin; BSTR, batch stirred tank reactor; FTIR, Fourier transform infrared spectroscopy; GC, gas chromatography; KBr, potassium bromide; Km , Michaelis constant; PDMS, polydimethylsiloxane; p-NP, p-nitrophenol; p-NPP, p-nitrophenyl palmitate; PVC, polyvinyl chloride; Vmax , maximum reaction rate; XPS, X-ray photoelectron spectroscopy; V+ , maximum velocity of forward reaction; KmA , Michaelis constant of OA; KmB , Michaelis constant of EtOH; Kib , inhibition constant of EtOH. ∗ Corresponding authors. E-mail addresses: [email protected] (W. Li), [email protected] (H. Shen), [email protected] (M. Ma), [email protected] (L. Liu), [email protected] (C. Cui), [email protected] (B. Chen), [email protected] (D. Fan), [email protected] (T. Tan).

contrast to the soluble enzymes used in industrial processes, immobilized enzymes often offer advantages in terms of stability, volume-specific biocatalyst loading, recyclability, and simplified downstream processing [4,5]. For example, Lip2 lipase from yeast Y. lipolytica (YlLip2), immobilized on macro-porous resin, exhibits enhanced catalytic properties and can be used for the enrichment of polyunsaturated fatty acids [6]. Similarly, YlLip2, immobilized in Accurel MP 1000, efficiently catalyzes the incorporation of medium-chain length fatty acid into olive oil [7]. Although several methods for enzyme immobilization have been described, only a few successful commercial processes employ immobilized enzymes [8,9]. Fickers and Marty [10] have reviewed the utility of lipase obtained from YLL in genetics, regulation, biochemical characterization, and biotechnological applications. Our group has focused its research efforts towards improving enzyme thermostability [11], purifying enzymes by chromatography [12], preventing thermal inactivation, and characterizing the conformational changes [13]. Immobilized lipases have been used to carry out lipase-catalyzed reactions, such as the synthesis of 2-ethylhexyl palmitate [14], fatty acid methyl ester [15–17], diglycerides [18], and ethyl oleate [19].

http://dx.doi.org/10.1016/j.procbio.2015.07.012 1359-5113/© 2015 Elsevier Ltd. All rights reserved.

Please cite this article in press as: W. Li, et al., Synthesis of ethyl oleate by esterification in a solvent-free system using lipase immobilized on PDMS-modified nonwoven viscose fabrics, Process Biochem (2015), http://dx.doi.org/10.1016/j.procbio.2015.07.012

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We have investigated the utility of fibrous materials, i.e., based on textiles, for enzyme immobilization. When compared to porous materials, fibrous materials possess excellent properties suitable for the immobilization of enzymes, such as a high specific surface area, good mechanical strength, chemical stability, and lower diffusion resistance of the substrates or products [20,21]. In 2007, immobilized YlLip2 lipase was used for the enzymatic production of commercial biodiesel commercialization with a capacity of 10,000 tons in Shanghai [22]. However, further improvements in immobilization techniques that can afford higher enzymatic activity and stability need to be explored. Previously, we developed an entire inexpensive esterification process for the production of ethyl oleate (as biodiesel), including a straightforward protocol for the immobilization of lipase and optimizing yields of the extended esterification batch reaction. Lipase from a fermentation broth, when immobilized on a non-woven fabric, afforded ∼19 reuse numbers [23]. When compared to lipase immobilized on inexpensive nonwoven fabric membranes, YlLip2 immobilized on a hydrophobically treated viscose fiber (primarily composed of cellulose as determined by SEM and FTIR) showed better stability [24]. In this study, methods for immobilization of lipase via fermentation broth or crude lipase powder solution were evaluated. Synthesis of ethyl oleate by esterification in a solvent-free system was used as the model reaction. Kinetic parameters for esterification reaction carried out by hydrophobic immobilized lipase, original immobilized lipase, and crude lipase powder were determined. These parameters—maximum velocity (V+ ), the affinity between oleic acid and the lipase (KmA ), and the degree of substrate inhibition (Kib )—were then used to evaluate the influence of the relatively hydrophobic matrix on the lipase activity. In the 5-L Batch stirred tank reactors (BSTR), the most widely used reactor for enzymatic processes, shear stress from stirring eventually disrupts the enzyme-loaded membrane [25]. In this study, the operational stability and the detachment of lipase from the fabrics when the esterification reaction is carried using immobilized lipases out in 5-L BSTR was evaluated. Finally, characteristics of the immobilized lipase, such as optimal pH, optimal temperature, solvent tolerance, kinetics of p-NPP hydrolysis, and composition (XPS/ATR-FTIR analysis) were analyzed.

2. Materials and methods 2.1. Materials Crude YlLip2 lipase powder was prepared in our lab according to a previously published procedure, which involves dehydration by cold acetone [13]. MALDI-TOF mass spectral analysis showed that the lipase was encoded by gene LIP2 (GenBank accession no. AJ012632) [12]. Olive oil was purchased from the Beijing Fangcao Chemical Factory (Beijing, China). The amino-functionalized silicone polymer, polydimethylsiloxane (PDMS, Fig. S1) used to modify the membranes (mechanism of immobilization is shown in Scheme S1), was purchased from Dow Corning Corporation (Midland, MI), and the micro-emulsified PDMS (PM6058) was purchased from Beijing Hanlin Chemical Co., Ltd. (Beijing, China). Bovine serum albumin (BSA) was used as the standard protein. All other organic solvents were purchased from the Beijing Chemical Factory and were of analytical grade. Nonwoven fabric samples made of viscose were donated by the Beijing Jie Xiang Co., Ltd. (Beijing, China). The viscose used was composed of rayon (65 g m−2 ) produced from cellulose fibers that had been extracted and purified from cotton lint, wood, or other plant materials.

2.2. Immobilization of lipase from the fermentation broth and solution of the powder Protocol for the immobilization of lipase for use in BSTR is described here. The cells from the fermentation broth were removed by centrifugation (6792 × g, 10 min, 4 ◦ C). The supernatant fermentation broth was used for immobilizing the enzyme on the fabric membrane. The immobilized lipase (3 cm × 3 cm) was prepared as follows: a piece of nonwoven fabric was placed in each lipase solution for 2 h. This ensured that the lipase was adsorbed and, consequently, immobilized on the fabric. Subsequently, the fabric membrane was removed and dried at room temperature. The adsorption protocol described above was repeated several times (in the same piece of fabric) to obtain the desired lipase loading as described previously [23]. Samples of immobilized lipase used for characterization: for the preparation of PDMS-treated viscose samples, viscose fabric with treated with the PDMS micro-emulsion (to increase the hydrophobicity of the viscose membrane) as described previously for “VI. Fk5 micro-emulsion” [24]. Support membranes were immersed in PDMS micro-emulsion (400× dilution) for 1 h at room temperature and then dried at 70 ◦ C for 1 h. Crude lipase solution (5 mL; 30 mg mL−1 ) were placed on a pieces of the dried support membrane and the adsorption facilitated by shaking the same at 120 rpm for 2 h at 25 ◦ C. Subsequently, the immobilized lipase support membranes were dried at room temperature (25 ◦ C). For the preparation of PDMS-treated immobilized lipase sample, the immobilized lipase (1 g) was treated with a solution of an amino silicone polymer (1%, w/v; 0.8–5 Pa s) in n-hexane (20 mL). The immobilized lipase was immersed in the amino silicone polymer solvent for 2 h and then baked at 120 ◦ C for 2 min. Assays were performed to ensure that the enzyme was not inactivated under the drastic treatment conditions, Fig. S2. 2.3. Measurement of hydrolytic activity of lipase Initial studies were performed using olive oil as the hydrolysis substrate. A method was developed (described below) to quantify an amount of lipase as a “lipase unit” (U). Samples of lipase for the measurement of catalytic activity were prepared. A solution of the crude lipase powder (30–60 mg mL−1 ) was diluted in KH2 PO4 /K2 HPO4 buffer (100 mmol L−1 , pH 8) to obtain solution containing desired units of the enzyme. An aliquot (1 mL) of the supernatant of the centrifuged fermentation broth was diluted in water (50 mL) and the pellet (0.5 g) was then dissolved in water (100 mL). Hydrolytic activities of the lipase samples were measured using titrimetric assays based on an olive oil emulsion method, but with a few modifications [26]. For use in the assay, olive oil [5%, v/v] was emulsified in distilled water containing 2% (w/v) PVA in a homogenizer for 6 min at maximum speed. The assay mixture consisting of emulsion (5 mL), phosphate buffer (4 mL, 100 mmol L−1 , pH 8), and free enzyme (1 mL) was assembled and the reaction was incubated at 35 ◦ C for 10 min at 120 rpm. Ethanol (20 mL) was added to stop the hydrolysis reaction. The activity of the enzyme was determined by titration of the released fatty acid with NaOH (0.05 M). One unit (IU) of lipase activity was defined as the amount of enzyme that catalyzes the liberation of 1 ␮mol of fatty acid from the olive oil substrate per min at 35 ◦ C. The lipase activity from the different fractions, constituting cell-bound lipase and intracellular lipase, was expressed as units per milliliter of lipase solution or per gram of sediment. The immobilization efficiency and proteincoupled content were calculated as previously described [24]. The compound p-nitrophenyl palmitate (p-NPP) was also used as the substrate for hydrolysis. Lipase activity was measured using a spectrophotometric assay with p-NPP as the substrate. The

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substrate solution (1.65 mM) was prepared by dissolving p-NPP in isopropanol. A 20 mM phosphate buffer (pH 8, containing 0.1% Triton-100) was used as the reaction buffer. An aliquot (200 ␮L) of lipase solution, a portion (70.2 mg) of the membrane loaded with lipase, or a blank solution (200 ␮L; 20 mM phosphate buffer, pH 8, containing 0.1% Triton-100) was added to the reaction buffer (2.16 mL), following which, the substrate solution (0.24 mL) was added to the reaction medium. The enzyme/substrate or blank/substrate mixtures were incubated for 1 min. The molar extinction coefficient of p-nitrophenol (p-NP) was estimated to be 14,528 cm−1 M−1 from the absorbance of standard p-NP solutions measured at 405 nm. The slope of the absorbance versus time curve gave the reaction rate. One unit of enzyme activity corresponded to the release of 1 ␮mol p-NP per minute under the standard assay conditions. Olive oil was used as the substrate in all cases except where mentioned otherwise, e.g., in Section 3.3.3. 2.4. Measurement of the effect of pH, temperature, and organic solvents on the activity and stability of immobilized lipase Effect of pH and temperature on the activity of immobilized lipase: The rates of hydrolytic reactions catalyzed by crude lipase solution and immobilized lipase were measured in different buffers (50 mM each) covering the pH ranged from pH 4 to pH 10. The buffers used include citric acid-sodium citrate buffer (pH 4–6), Tris–HCl buffer (pH 7–9), and glycine-NaOH (pH 10). The hydrolytic activity of lipase samples was measured over the 25–50 ◦ C temperature range. Effect of pH and temperature on the stability of immobilized lipase: After initially storing the lipase samples in buffers with different pH values (4–10) for 16 h in room temperature, the hydrolytic activities of the lipase samples were measured at 35 ◦ C. The activities of these samples were reported relative to the activity at pH 7, which was set as 100%. The lipase samples were incubated at 35–45 ◦ C for 4 h. At different time points (1, 2, and 4 h) aliquots were withdrawn and the activity measured. The activity of the enzyme determined under optimal conditions (optimal pH and temperature) was set as 100%; other activities were reported relative to this activity. Effect of solvent on the activity of immobilized lipase: immobilized lipase (15 mg), or crude lipase powder (10 mg) was immersed in organic solvent (1 mL) for 24 h at 15 ◦ C. Subsequently, the residual activity was measured and reported relative to the activity of the enzyme in 0.1 M PBS buffer pH 8. 2.5. Esterification of oleic acid with ethanol in batch stirred tank reactor (BSTR) Initially, esterification reactions were conducted in a 50 mL conical flask. Equimolar quantities of oleic acid (2.82 g) and ethanol (584 ␮L), a piece of immobilized lipase (3 cm × 3 cm) were incubated at 30 ◦ C and the flask was shaken at 190 rpm. The fatty acid conversion rate was determined by titration with NaOH, as described previously [23,24]. At the end of reaction, using 0.2 M NaOH titration to determine the conversation of oleic acid after adding 20 mL ethanol to mix with the catalyzed system. The activity units of the immobilized lipase samples prepared either from the fermentation broth or lipase powder solution were determined under these conditions. For determining the kinetic parameters of the esterification reaction, varying concentrations of oleic acid (2–20 mmol) were independently mixed with ethanol (5 mmol), 20 mg crude lipase powder, or 30 mg immobilized lipase and incubated at 30 ◦ C for 3 or 4 h while being shaken at 500 rpm. Initial velocities of the reactions were measured. Subsequently, similar sets of reactions were

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performed at different concentrations of ethanol (7.5, 10, 15, and 20 mmol). Next the reactions were conducted in BSTR. For this, immobilized lipase (139.5 g), water absorbent (silica gel/molecular sieve mixtures; 200 g), and oleic acid (3.14 mol; 887.5 g) were taken in the BSTR. Ethanol was added in a stepwise manner. In the 1st batch, ethanol (3.54 mol; 207 mL) was added in one-lot. In the subsequent 9 batches (batch 2–10), equal portions of ethanol were added in 4 stages for each batch addition (at 0, 1, 2, and 3 h time-points). After the theoretical yield was realized with each added batch, the immobilized lipase was removed using the pipe holder, the new substrate was added and mixed with the reaction liquid in the BSTR, and finally, the lipase on the pipe holder was re-inserted into the reaction mix. The esterification reactions was carried out at 30 ◦ C and 1000 rpm. 2.6. Attenuated total reflectance–Fourier transform infrared spectroscopy (ATR–FTIR) measurements A Nicolet Nexus 670 FTIR spectrometer equipped with an attenuated total reflectance (ATR) detector was used to record attenuated total reflection. FTIR spectroscopy in attenuated total reflectance mode allows enzymes and proteins in a solid phase to be studied in situ. ATR–FTIR was used to evaluate the effects of membrane treatment (hydrophobically treated vs untreated membrane) on the conformation of immobilized lipase. Moreover, the conformation of the crude lipase was also examined by ATR–FTIR. Frozen dried lipase samples in powder form were diluted with potassium bromide (KBr) and pressed into a conventional wafer for FTIR analysis. 2.7. Reactor setup The 5-L BSTR reactor was built by modifying a fermenting tank with a standard turbine agitator. A total of 12 hollow cylindrical polyvinyl chloride (PVC) tubes were fixed at the bottom of a metallic holder placed in the reactor. The PVC tubes (with 8 mm diameter holes in the tube walls) were lined with 4 A˚ molecular sieves. Membranes bearing the immobilized lipase were wound around the outer walls of the tubes, covering the mesh-filled holes (Scheme 1). 2.8. Measurement of water content The amount of water dissolved in the liquid reaction mixture and the amount of water in the immobilized lipase were measured using a Carl Fisher water measuring device KLS-4119 (Shanghai Precision & Scientific Instrument Co., Ltd., Shanghai, China). For the analysis, samples (100–200 mg) were taken from the reaction mixture. 2.9. Measurement of ethanol content An aliquot (500 ␮L) of the sample solution was dissolved in water (27 mL) and the resulting mixture was used for determining the ethanol content using gas chromatography (GC, Shimadzu GC-2010, Japan). The determination of ethanol using the GC was performed according to a method described previously [27]. Both the injector and detector were maintained at 230 ◦ C. The temperature of the column was programmed as follows: step 1, 120 ◦ C for initial 0.5 min; ramp 1, increase to 180 ◦ C (20 ◦ C/min); step 2, hold at 180 ◦ C for 3 min; ramp 2, increase to 230 ◦ C (30 ◦ C/min); and, finally, step 3, hold at 230 ◦ C for 10 min. We used the external standard method for creating a calibration curve, wherein the calibration curve was established using media mixed with various concentrations of ethanol concentrations (1–9 g L−1 ).

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Oleic acid conversion (%)

100.0

A

80.0

60.0

-1

1900U mL (4 times adsorption) -1 7800U mL (lipase powder) -1 6400U mL -1 4500U mL -1 3200U mL -1 900U mL -1 500U mL -1 20U mL

40.0

20.0

0.0 0

5

10

15

20

25

Reaction time (h)

2.10. Measurement of protein leakage from matrices and operational stability in the esterification of lauric acid and dodecanol

B

80.0

Oleic acid conversion (%)

Scheme 1. Schematic of the 5-L batch stirred tank reactor (BSTR). Immobilized lipase (139.5 g), water absorbent (200 g; silica gel/molecular sieve mixture), oleic acid (887.5 g; 3.14 mol), and stepwise addition of 3.54 mol ethanol (207 mL, a onestage stepwise addition in the 1st batch; a four-stage stepwise addition in the subsequent 9, i.e., 2 through 10, batches). After the yield from each batch reached its theoretically predicted value, the immobilized lipase was withdrawn from the reaction mixture, new components were added and mixed together in the reaction liquid for 5 min, and the immobilized lipase, contained in the pipe holder, was replaced back into the reaction mixture. The reactions were performed at 30 ◦ C and 1000 rpm as described (see Section 2 for additional details).

70.0 60.0 50.0 40.0 30.0

original immobilized lipase PDMS treated

20.0 10.0 0

Leakage of lipase from the nonwoven viscose fabric into the aqueous solution was examined. To quantify this, immobilized lipase (3 cm × 3 cm of the membrane) was added to 200 mL of water. The solution was shaken at 150 rpm and 55 ◦ C. At periodic intervals, aliquots (1 mL) of the solution were removed and the absorbances of the samples were measured at 595 nm using the Bradford method with BSA as the standard [28]. Furthermore, the operational stability of the immobilized lipase (two pieces of 22 cm × 8 cm) was similarly evaluated in a 500 mL round-bottom flask under mechanical stirring. The lipase was immersed in a reaction liquid (150 mL; 0.25 M lauric acid and 0.25 M dodecanol in hexane) at 55 ◦ C, 150 rpm, and 1 day batch−1 .

5

10

15

20

25

30

35

40

Reuse numbers (3h/batch) Fig. 1. (A) Esterification by immobilized lipase prepared from fermentation broth (the lipase powder solution) at different activity units. Unless specifically mentioned, the lipase was immobilized from supernatant of the fermentation broth with a one-time adsorption. Note that the following deviations in the protocol: adsorption was carried out 4 times for immobilizing the lipase from fermentation broth with an activity of 1900 U mL−1 ; the lipase was immobilized from solution of the lipase powder with an activity of 7800 U mL−1 . (B) Operational stability of hydrophobic immobilized lipase during the esterification of oleic acid with ethanol in presence of 8% (v/v) water, two pieces of viscose-immobilized lipase (3 cm × 3 cm) prepared from 1900 U mL−1 fermentation broth (4× adsorption; see Section 2 for details). The standard reaction was performed in a 50 mL conical flask containing 8% (v/v) water (added initially), and equimolar quantities of oleic acid and ethanol (three-stage stepwise addition of 10 mmol) at 30 ◦ C for 3 h with shaking at 190 rpm.

3. Results and discussion 3.1. Immobilized lipase preparations derived from fermentation broth and crude lipase powder solution During the cultivation of YLL, the lipase activity from different cellular fractions (cell-bound lipase, intracellular lipase, and extracellular lipase) has been found to be distributed according to cell growth [29]. During the initial growth phase, the production of lipase occurred predominantly in the cell-bound regions. In the late stationary phase, highest extracellular levels of lipase were observed in the culture medium due to the release of lipase. Previously [23], we noticed that highest rates of hydrolysis (using olive oil as the substrate) with the immobilized enzyme were observed when the enzyme was immobilized using a fermentation broth with higher activity. We observed a very similar trend in our time course when the activity of the fermentation broth was 3000–4500 U mL−1 . However, when the lipase was immobilized by four repeated adsorptions using a fermentation broth with an activity of 1900 U mL−1 , the activity of the resulting immobilized lipase was equal to that of the immobilized lipase samples prepared by

a single adsorption using a fermentation broth with an activity of 4500 U mL−1 . After 3 h, the oleic acid conversion by the immobilized lipase (prepared by 4× adsorption from fermentation broth with an activity of 1900 U mL−1 ) ∼80%, and an equilibrium of 86% was reached within 1 day (Fig. 1A). Fig. 1B shows the operational stability of hydrophobic immobilized lipase (4×, 1900 U mL−1 fermentation broth). The hydrophobic immobilized lipase was more stable and exhibited higher catalytic activity in the reaction system within 35 batches; by comparison, under optimal conditions, lipase immobilized on a cotton silk fabric afforded >19 reuse numbers at 5 h/batch, and an esterification rate >80%) [23]. The relatively hydrophobic matrix provided a proper microenvironment for the immobilized lipase by preventing the accumulation of excess water during long-term reuse. Next, we used centrifugation to investigate the distribution of lipase activity in the original fermentation broth (Table 1). We found that a supernatant and a pellet could be separated by centrifugation of the fermentation broth at 425 × g for 10 min. When compared to the pellet formed on centrifugation at 6792 × g, the

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Table 1 Distribution of the lipase activity in the fermentation broth. Centrifugal

106 × g, 5–10 min 239 × g, 5 min 425 × g, 10 min 6792 × g, 10 min

Phenomenon

Hydrolytic activity

No delaminating, no precipitation. A thin layer of pellet is formed along the wall of centrifugal tube Solid–liquid separation, soft cell pellet Solid–liquid separation

relatively softer pellet obtained at 425 × g was much easier to handle. Therefore, in addition to the supernatant, the wet pellet (with a lipase activity of 1000–3000 U g−1 ) can also be potentially used for the immobilization of the lipase, provided proper methods are followed (such as homogenization of the pellet). We also used dried crude lipase powder in our study because it is convenient, stable, and readily stored. Because the lipase powder was prepared by spray-drying fermentation broth with protective additives, such as milk powder and maltodextrin [30], the main difference between the lipase powder and the lipase in the fermentation broth was in their composition. When adsorbed to membranes at comparable activities, the lipase immobilized from the fermentation broth (6400 U mL−1 ) e xhibited higher activity than that immobilized from the solution of lipase powder (7800 U mL−1 ) (Fig. 1). After 3 h, the conversions of oleic acid by lipase immobilized from fermentation broth (6400 U mL−1) and the solution of the powder (7800 U mL−1 ) were 74.9% and 41.2%, respectively. The activity of free lipase, the composition of the lipase solution (e.g., from the fermentation broth or powder solution), and the loading of the lipase on the carrier (influenced by adsorption time)—all factors affected the final activity of the immobilized lipase. When compared to the efficiency of lipase adsorption onto the hydrophobic membrane from the crude lipase solution (immobilization efficiency of 23.9% and a protein-coupled content of 47.3%) [24], the immobilization of the lipase from the slurry of fermentation broth (18 mg mL−1 ) was higher (immobilization efficiency of 75.2% and protein-coupled content of 82.6%; Table 2). The percentage of lipase immobilized and the activity recovered were monitored for the large-scale (2.4 L) immobilization of lipase (Table S1). Table S1 shows that the lipase solution was totally consumed after four-adsorptions and the weight of the membrane carrying in the lipase increased to 307 g. Therefore, the use of a concentrated solution of lipase is recommended for industrial applications because it improves the activity utilization ratio of the lipase solution. 3.2. Synthesis of ethyl oleate by esterification in a solvent-free system 3.2.1. Kinetic study The presence of excess ethanol in the lipase-catalyzed esterification reaction can lead to two consequences, enzyme denaturation [12] and/or inhibition of the lipase-catalyzed reaction [23]. The mechanism for the deactivation of lipase in excess methanol Table 2 Immobilization parameters from concentrated slurry lipase solution.

Immobilization efficiency (%) Protein-coupled content (%)

Original viscose

Hydrophobic viscose

67.9 71.6

75.2 82.6

Lipase (58,000 U, 7.57 mg) was adsorbed onto original or hydrophobic viscose (0.6 g) from a concentrated solution of lipase (10 mL; 180 mg/mL). The slurry was nearly completely adsorbed in several minutes. Then, the immobilized lipase was freeze dried.

Supernatant (U mL−1 )

Sediment (U g−1 )

/ / 3200 ± 0 2500 ± 100

/ / 1429 ± 107 3268 ± 454

(widely used in the production of biodiesel) was recently reported by Li et al. [31]. Furthermore, the study by Yu et al. revealed that YlLip2 was more sensitive to ethanol [12]. Sandoval et al. [32] reported the solvent-free synthesis of ethyl oleate with thermodynamic activity-based enzyme kinetics approach. Therefore, to enhance the degree of esterification, the concentration of ethanol was controlled by the stepwise addition of the same. In our study, mass-transfer limitations (both external and internal) were not observed; the rate of the reaction was not influenced either by a change in the agitation speed or by variations in the amount of the immobilized enzyme (Fig. S3). The inhibition of the reaction by excess ethanol was attributed to the formation of a dead-end complex between enzyme and excess ethanol. As elucidated in Fig. 2, substrate A (oleic acid) binds to E; A is cleaved into product P (water) that dissociates and X (fatty acid) that remains conjugated to E (complex EX); substrate B (EtOH) binds to EX complex and gives EXB intermediate; X and B are fused together giving product Q (biodiesel) that dissociates. The free enzyme can also bind EtOH (substrate B), leading to the formation of an unproductive complex EB. Kinetics data were fitted to the rate-equation corresponding to the Ping-Pong mechanism, adjusting for the inhibition by EtOH (B), and are shown in the scheme in Fig. 2. None of the datasets could be approximated by the unconstrained fit, where all parameters (V+ , KmA , KmB , Kib ) were subjected to optimization. Under these circumstances, some parameters tended to either infinity or zero. By selectively varying two parameters (V+ and one of the Michaelis-Menten/dissociation constants), a plausible interval could be determined for all the parameters. They are shown in Table S2, together with the values of the constrained variables. The estimates of the substrate concentrations (required to maintain the maximum velocity) are also shown; note that these velocities are the initial rates of conversion. As listed in Table 3, maximum velocities of the immobilized lipase samples were lower than that of the crude lipase powder, more so in the case of hydrophobically treated immobilized lipase. The maximum velocity of the hydrophobic immobilized lipase was half that of the original immobilized lipase and 1/3rd that of the crude lipase powder. The KmA (35.4 M) of hydrophobic immobilized lipase was 10- and 20-fold higher than that of crude lipase powder and original immobilized lipase, respectively. Therefore, after modification with a hydrophobic group, the affinity of the immobilized lipase for the oleic acid was clearly reduced. Kib represents the concentration of substrate when the half the activity of the enzyme is inhibited. The Kib (8.5 M) of reaction catalyzed by the hydrophobic immobilized lipase was 17- and 42-fold higher than that of crude lipase powder and original immobilized lipase, respectively, and accounted for the lowest inhibition effect of ethanol on the hydrophobic immobilized lipase. 3.2.2. Hydrophobic immobilized lipase applied in 5L-BSTR 3.2.2.1. Accumulation of water during the single-batch esterification reaction catalyzed by immobilized lipase. Water plays a significant role in the mechanism of enzyme-catalyzed esterification. First, the water produced in the reaction affects the reaction

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Fig. 2. Schemes describing the ordinary Ping-Pong mechanism, supplemented by substrate inhibition. Here A = oleic acid; B = EtOH; P = water; Q = biodiesel; X = fatty acid; V+ = maximum velocity for the forward reaction; KmA = Michaelis constant of oleic acid; KmB = Michaelis constant of EtOH; and Kib = inhibition constant of EtOH. Schematic representation of the results (circles) in 3 D coordinates fitted by the equation of the ordinary Ping-Pong mechanism with EtOH-induced inhibition for the esterification reaction with crude lipase powder (left panel), immobilized lipase (middle panel), and hydrophobic immobilized lipase (right panel; serious loss of activity can be attributed to over time-baking).

Table 3 Estimates of the substrate concentrations required to maintain the maximum velocity. Crude lipase powder

Hydrophobic immobilized lipase

Selected

Plausible

Selected

Plausible

Selected

170–300 2–8 M 2–4 M 0.2–1 M

250 4.27 2.99 0.5

100–250 0.3–2.7 M 1.7–6 M 0.05–4 M

200 2.12 4.89 0.2

100–300 15–70 M 0.5–2 M 3–10 M

150 35.44 1 8.49

v = 7–10 a = 2.5–3 M b = 0.75–2.5 M

80.0

100.0

60.0

75.0

40.0

1.5 1.2 0.9

50.0

water content(%, w/w) ethanol conversion(%) oleic acid conversion(%)

20.0

0.0

0

2

4 6 8 Reaction time (h)

25.0

10

0.0

0.6 0.3

Water content(%, w/w)

equilibrium. Second, in non-aqueous systems, the level of water content can be critical to maintain the conformation of the catalytic site in the enzyme. Water’s influence as an important factor contributing to the enzyme activity was considered [33–35]. The role of water in the solvent-free enzymatic esterification reaction was primarily studied by the adding water initially to the immobilized lipase, as well as by using a water adsorbent in reaction liquid [23]. In this study, the trends in the distribution of water in the reaction liquid and the water accumulated in the immobilized lipase were primarily investigated. The conventional BSTR was used as the model reactor to determine the performance of the hydrophobic immobilized lipase in a single-batch reaction without the stepwise addition of ethanol. As evident from Fig. 3, we found no obvious trend in the substrate concentration with change in the water content (1.19–0.53%; corresponding to an introduced water amount of 9.89–16.35 g) of the reaction liquid. Findings from this study are consistent with

v = 18–20 a = 2.5–3 M b = 0.75–1.5 M

Ethanol conversion(%)

v = 20–30 a = 2–3 M b = 0.75–2 M

Oleic acid conversion(%)

V+ = KmA = KmB ≥ Kib ≤ Velocity estimates (maximum)

Immobilized lipase

Plausible

0.0

Fig. 3. Time course of the reaction between oleic acid and ethanol in a single batch reaction carried out in a BSTR (equimolar amounts, 3.54 mol, of reactions, with no stepwise ethanol addition). The reaction conditions are described in detail in Reaction in BSTR of Section 2.

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30

40

50

75.0

6060

40

50.0 20 25.0

lauric acid conversation (%) protein content ( /mL)

0

0.0 0

2

4

6

8

/mL)

20

Protein content (

Lauric acid conversion (%)

10

10 12 14 16 18 20 22

Reuse numbers(day/batch) Fig. 4. Protein detachment and catalytic operational stability of the immobilized lipase. The amount of lipase that had detached from the membrane was present in the surrounding aqueous environment and could be monitored over time. The operational stability of the immobilized lipase (two pieces, 22 cm × 8 cm each) was evaluated in a 500 mL round-bottom flask under mechanical stirring. Reaction conditions: 150 mL reaction liquid (0.25 M lauric acid and 0.25 M dodecanol in hexane), 55 ◦ C, 150 rpm, and 1 day batch−1 .

those an earlier study which revealed that the water content in the biocatalyst solution is approximately independent of the substrate concentration [32]. When ethanol was added in a single lot, the trend in ethanol conversion was similar to that of oleic acid conversion. Water was a contaminant of the commercially procured reactants. Ethanol and oleic acid carried 0.94 and 10.28 g of water, respectively, into the reaction system. The immobilized lipase, before being immersed in the reaction mixture, contained 7.37% (w/w) of water. We monitored the water content of the reaction system over time to understand the water distribution. The conversion of oleic acid reached 72.9% at 11 h, which accounted for a theoretical yield of 41.17 g of water (Fig. 3). Experimental results revealed that at the end of 11 h, the corresponding water content in the immobilized lipase increased to 8.66% from its initial value of 1.77% (16.35 g) of water and accounted for 39.70% of the total theoretical water content. During esterification under batch conditions, the water that accumulates in the immobilized lipase tends to increase. Water accumulation in the biocatalyst becomes more significant at higher conversion levels because more water is formed, and the solution’s polarity also decreases as acid and alcohol substrates are converted to their less polar ester products [36]. However, the activity of the immobilized lipase can rebound if the enzyme-bound membrane is dried overnight [23]. In this study, we found that the lipase activity was restored when the membrane was dried to a water content of 4.39% (w/w). The water content of the biocatalyst is considered to be the main factor affecting reaction equilibrium, enzymatic activity, and the long-term stability of enzymes [33,34]; however, simple air-drying techniques and other measurements can help to restore the activity of immobilized enzymes. Detachment of proteins from an immobilized protein membrane over time has been examined by several researchers as a criterion to compare different immobilization methods [37,38]. We investigated the detachment of lipase from the hydrophobic membrane in water by monitoring the protein content in solution over time and compared the result with that of lipase leakage when the immobilized lipase was placed in n-hexane (Fig. 4). During the 5day incubation in water, the lipase rapidly desorbed from the fabric in the first 2 h and was totally desorbed by day 4. After the fabric with the immobilized lipase was dried, almost no loaded enzyme remained in the fabric. In n-hexane, only a small amount of lipase that was loosely bound to the membrane’s surface fibers escaped when incubation started; there was hardly any detachment of the

Oleic acid conversion(%)

100.0

Incubation time (h) 100.0 0

7

80.0

60.0

40.0

20.0

0.0 0

20

40

60

80

100

120

140

160

Cumulative operation time(h) Fig. 5. Iterative batches with the reuse of the immobilized lipase and stepwise addition of ethanol (a one-stage stepwise addition in the 1st batch; a four-stage stepwise addition in the batches 2–10). The reaction conditions are described in detail in Reaction in BSTR of Section 2.

lipase subsequently. Enzymes that are physically immobilized on the surface of the carrier sometimes detach from the carriers, which can be problematic for industrial applications [37]. The hydrophobic fabric immobilized enzyme could be recycled for more than 80 times with no significant decrease in esterification activity (3 h/batch) [39]. In our study, by the 12th batch, the mechanical stirring had sheared the immobilized lipase to floccule strips; however, the subsequent esterification rate was not affected. By the 20th batch, in which the water content made up 20% (v/v) of the immobilized lipase, the esterification rate was reduced by 50%. Therefore, it can be speculated that when employing an immobilized lipase in BSTRs, the detachment of lipase from the fabrics as a result of shearing has a significantly lesser effect on the catalyst function that due to an increase in water content. 3.2.2.2. Operational stability of the hydrophobic immobilized lipase. In Nie’s study, the final conversion of the fatty acid methyl ester from waste oil could reach 92% in a fix bed reactor, with more than 20 day-life of immobilized lipase [40]. As proteins are generally unstable in short-chain alcohols (such as methanol and ethanol), with the presence of ethanol being more detrimental for the function of the type of lipase used in this study [12], we added ethanol in a four-stage stepwise fashion to prevent enzyme inhibition. Because of the protection offered by the substrate/product to the enzyme activity [41], the immobilized membrane was immersed in the reaction mixture at all times during the batch operation, except when ethanol was added. Fig. 5 shows the reaction time course of fatty acid conversion in each cycle. For the 1st batch (without stepwise ethanol addition), the esterification rate first increased quickly during the first 5 h (60% conversion) and then decreased slowly, reaching the reaction equilibrium (84% conversion) in 12 h. The reaction time courses for the batches 2–4 were identical; the reaction equilibrium (∼90% conversion, comparable to that by the lipase immobilized on a silk membrane, Fig. S4) was reached within 10 h. Our results agree with the findings that addition of less than stoichiometric amounts of ethanol at each step prevents substrate-mediated enzyme inhibition [42]. The final reaction conversion rate using the four-stage stepwise addition of ethanol improved by ∼10% when compared to that of the reaction involving addition of ethanol in a single lot. After the 5th batch, the reaction slowed significantly, and the equilibrium was reached after 12 h (88.3%), 19 h (87.2%), and 36 h (81%) of ethanol addition for the 6th, 8th, and 10th batches, respectively. These results indicate that the lipase immobilized on nonwoven viscose fabric membrane can be reused for at least 10 batches

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Fig. 6. The effect of pH and temperature on the activity (A, C) and stability (B, D) of the immobilized lipase.

3.3. Characterization of the application-oriented immobilized lipase 3.3.1. The effect of pH, temperature on the activity and stability of immobilized lipase The optimum pH for the esterification by the immobilized lipase was 7; the optimal pH for the activity of immobilized lipase is slightly acidic when compared to that of the free lipase in solution (pH 8; see Fig. 6A) [12,19]. The hydrolytic activity by the immobilized lipase was relatively high (85%) only in the pH 6–7 range. Meanwhile, the free lipase retains 85% of its optimal activity across a wider range of pH values (pH 5–8) even after a 16 h pre-incubation at room temperature. Results from the evaluation of the optimum temperature and thermal stability are shown in Fig. 6(C and D). In contrast to the results from Yu’s study (optimal temperature for the activity of purified lipase was 40 ◦ C) [12], the optimum temperature for free lipase in this study was 30–35 ◦ C. Through site-directed mutations, Wen et al. [11] improved the thermal stability of YlLip2 (wild-type) from the optimum temperature 37 ◦ C to ∼40 ◦ C (thermostable variant of Y/Lip2). Interestingly, immobilization of lipase also increased the optimal reaction temperature to 40 ◦ C (Fig. 6C); the same results were observed when lipase was immobilized on PDMS-fibers (using silk woven fabric) [19]. In their investigations into the thermal stability of the enzyme, Yu et al. [12] found that when incubated at 30 ◦ C and 35 ◦ C for 4 h, the purified lipase retained 95% and 83%, respectively, of its original activity. However, only 32% and 5% of the activity was retained when the lipase was incubated for 4 h at 40 ◦ C and 45 ◦ C, respectively. Similarly in our study, we observed that the free lipase retained over 75% of the original activity after being incubated for 4 h at 25 ◦ C. However, the activity dropped sharply at pre-incubation temperatures higher than 35 ◦ C, with more than

half activity being lost after 2 h pre-incubation (Fig. 6D). The thermal stability of the immobilized lipase in 35–45 ◦ C range was significantly higher, with over 70% of the activity being retained after 4 h incubation at 45 ◦ C. Therefore, immobilization significantly improved the thermal stability of lipase. 3.3.2. Tolerance of immobilized lipase to organic solvents Tolerance of immobilized lipase towards organic solvents was evaluated by measuring the activity of the enzyme after immersing the immobilized lipase in different organic solvents (Fig. 7). Lu et al. revealed that in more hydrophobic solvents, such as n-hexane, the yield of the ester was high, demonstrating their suitability as solvents for transesterification [17]. Similarly, lipase powder and immobilized lipase retain over 60% of their activity in nonpolar solvents, such as methylbenzene and n-hexane; however, they nearly completely deactivated in strong polar organic solvents, such as DMSO and methanol. It is likely that in polar solvents, the water molecules retained by the lipase are effective in allowing 100.0

Relative activity(%)

(150 h) under suitable operation conditions, where the reaction rate decreases as the total time of reaction increases.

crude lipase powder original immobilized lipase hydrophobic immobilized lipase

80.0 60.0

N,N-Dimethyl acetamide acetone ethanol

40.0 20.0

DMSO methanol

0.0 -2

-1

0

1

isopropyl alcohol

2 toluene 3 4 chloroform n-hexane

solvent(Log P) Fig. 7. Tolerance of immobilized lipase to organic solvents.

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W. Li et al. / Process Biochemistry xxx (2015) xxx–xxx Table 4 Activities, kinetic parameters, and relative content of structures estimated from Fourier transform infrared spectroscopy (FTIR) for both the free and immobilized enzymes.

3.3.4. XPS and ATR-FTIR analysis The polyhydroxy compound viscose is made up of C and O elements. The presence of a Si2p peak in the X-ray photoelectron spectrum (XPS) of PDMS-treated immobilized lipase sample indicated that the Si (Fig. 8) was introduced into viscose by physical adsorption of PDMS (Fig. 9A); this observation was consistent with

20000 0 1000

800

600

400

200

0

Binding Energy (eV) Fig. 8. X-ray photoelectron spectroscopy (XPS) of the viscose fabric before and after the treatment with polydimethylsiloxane (PDMS).

the new peaks observed in the IR spectrum related to the −CH3 , Si−O–Si, and Si−C bonds at ∼1260, ∼1021, and ∼800 cm−1 , respectively. Furthermore, after the PDMS treatment, both the bare and lipase-immobilized viscose showed the same peaks, indicating that PDMS was physically adsorbed (Fig. 9A). The amide I band, corresponding to the v (CO) carbonyl stretching mode of the peptide was present in the 1700–1600 cm−1 region [44]. Because of the high degree of overlap in the amide 100

-1

800cm Si-C

80

-1

original immobilized lipase PDMS treated viscose viscose-PDMS lipase powder

1021cm Si-O-Si

A

60

40

20 1800

1600

1400

1200

1000

800

-1

Wavenumber(cm ) B 0.05

0.8

original immobilized lipase PDMS treated

lipase powder

0.04 0.6 0.03

0.02

0.4

0.01

Abs of lipase powder

3.3.3. Kinetics of p-NPP hydrolysis Both the free and immobilized lipase showed enhanced activity with an increase in the concentration of substrate (0.165–1.65 mM). The apparent kinetic parameters calculated using the Lineweaver–Burk plot is listed in Table 4. The kcat (kcat = vmax /[E]T ) corresponds to the maximum number of substrate molecules converted to product per molecule of enzyme. The kcat of lipase decreased 31-fold upon the immobilization. This indicates that when excess lipase was immobilized on the membrane, the lipase lost most of its activity. In general, Km values are affected by diffusional limitations, steric effects, and ionic strength; the Km of lipase decreased 2.45-fold upon immobilization, indicating that physically binding the enzyme to the membrane can reduce the diffusion barrier that limits the substrate’s ability to reach the enzyme, thereby facilitating the substrate’s access to the active site on the enzyme. Compared to the catalytic efficiency (kcat /Km ) of the free lipase, catalytic efficiency of activity lipase immobilized on the hydrophobically treated and untreated membranes were ∼13and ∼15-fold lower, respectively. These results show that there were no obvious differences in the kinetic parameters of enzymes immobilized on either the hydrophobically treated membrane or the untreated membrane.

40000

-1

the enzyme to retained its catalytically active structure, while in non-polar solvents, the water bound to the surface of the lipases are easily removed by the solvent, resulting in the loss of lipase activity due to disruptions in the structure. However, the activity of immobilized enzymes was not improved in strong polar solvents, such as methanol and ethanol, as previous observed with enzyme immobilized on silk fiber [39]. In general, the activity of hydrophobic treated immobilized lipase changed marginally with the change in log P of different organic solvents. However, the activity of the immobilized lipase in methylbenzene and DMSO were significantly different. An attempt to correlate the measured optimal yield of the ester with the properties of the solvents revealed a reasonable correlation between the highest yield and log P values, however, with a few outliers [17]. While pretreating lipase in a solution of methanol (10–20%) in water did enhance enzyme activity and methanol tolerance [43].

Si(2p)

60000

1099cm C-F

The aliquot (200 ␮L) of crude lipase solution contained 1.66 mg protein. The original (untreated) membrane (70.2 mg) was loaded with 5.42 mg protein. The PDMStreated (hydrophobic) membrane (70.2 mg) was loaded with 6.26 mg protein. a The surface structure of the crude lipase preparations in the amide I spectroscopic region were estimated by pure protein secondary structure analysis method (Table S3).

80000

-1

131.8◦ 34.04 0.50 0.67 0.7 35.2 19.0

1260cm 3 CH

122◦ 32.82 0.55 0.64 0.9 35.4 22.9

Counts/s

/ 311.74 17.13 1.57 10.9 39.5 20.5

100000

Transmittance(%)

Contact angle Vmax (␮M min−1 ) kcat (min−1 ) Km (mM) kcat /Km (mM−1 min−1 ) a ˛-Helix a ˇ-Sheet

PDMS treated

C(1s)

120000

Immobilized lipase Original

original viscose viscose-PDMS O(1s)

140000

Abs of immobilized lipase

Crude lipase powder

9

0.2 0.00 1750 1725 1700 1675 1650 1625 1600 1575 1550 1525 1500 -1

Wavenumber(cm ) Fig. 9. (A) Fourier transform infrared (FTIR) spectra of the crude lipase powder, lipase immobilized on the PDMS-treated (hydrophobic) membrane, and lipase immobilized on the original (untreated) membrane; (B) Attenuated total reflectance (ATR) of the crude lipase powder and immobilized lipase.

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I band, a reduced-bandwidth analysis was performed using the second derivative of each spectrum. Fig. 9B shows the spectra (1750–1500 cm−1 ) of the investigated samples—the crude lipase powder, lipase immobilized on the hydrophobic (PDMS-treated) membrane, and lipase immobilized on the untreated (original) membrane. Through the analysis of the second derivative spectra (Fig. S5), and the bands’ position assignment (Table S3), it was revealed that the structure of the crude lipase preparation in the amide I spectroscopic region was mostly preserved after the immobilization (consisting of ∼20% ˇ sheets and ∼35% ˛ helices). 4. Conclusions In summary, we demonstrated the use of a hydrophobic nonwoven viscose fabric containing immobilized lipase for catalyzing the esterification reaction in a BSTR. The immobilized lipase was reused in 10 iterative batches, for a total reaction time of 150 h. Use of a concentrated lipase solution for immobilization improves the activity utilization ratio. The hydrophobic immobilized lipase (in the fermentation broth of 1900 U mL−1 for 4×) exhibited more stable catalytic activity within 35 batches. The mechanism of the esterification reaction catalyzed by the immobilization for the synthesis of ethyl oleate in a solvent-free system followed the Ping-Pong bi-bi mechanism, where ethanol was responsible for substrate-mediated enzyme inhibition. When compared with that of the original immobilized lipase, the hydrophobic immobilized lipase exhibited a 20-fold lower affinity for oleic acid, and a 42fold lower ethanol-induced inhibition. There were no significant differences in the optimum pH/pH stability, the optimum temperature/thermal stability, and the organic solvent tolerance between the two immobilized lipases. Analysis of the amide I region in the spectrum revealed that the surface structure of the crude lipase was largely preserved. Acknowledgements This study was supported by the National Basic Research Program of China (973 program) (2013CB733600 and 2012CB72520), the National Nature Science Foundation of China (21436002), and the National High-Tech R&D Program of China (863 Program) (2014AA022100). Also thank Sergey Fedosov in Department of Engineering, Faculty of Science and Technology, Aarhus University for his help in enzyme kinetic calculations. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:10.1016/j.procbio.2015.07.012 References [1] P. Fickers, P.H. Benetti, Y. Wache, et al., Hydrophobic substrate utilisation by the yeast Yarrowia lipolytica, and its potential applications, FEMS Yeast Res. 5 (2005) 527–543. [2] M. Agnihotri, S. Joshi, A.R. Kumar, et al., Biosynthesis of gold nanoparticles by the tropical marine yeast Yarrowia lipolytica NCIM 3589, Mater. Lett. 63 (2009) 1231–1234. [3] A.V. Bankar, A.R. Kumar, S.S. Zinjarde, Environmental and industrial applications of Yarrowia lipolytica, Appl. Microbial. Biot. 84 (2009) 847–865. [4] A. Liese, L. Hilterhaus, Evaluation of immobilized enzymes for industrial applications, Chem. Soc. Rev. 42 (2013) 6236–6249. [5] R.A. Sheldon, S. van Pelt, Enzyme immobilisation in biocatalysis: why, what and how, Chem. Soc. Rev. 42 (2013) 6223–6235. [6] Y. Yan, X. Zhang, D. Chen, Enhanced catalysis of Yarrowia lipolytica lipase LIP2 immobilized on macroporous resin and its application in enrichment of polyunsaturated fatty acids, Bioresour. Technol. 131 (2013) 179–187. [7] L. Casas-Godoy, A. Marty, G. Sandoval, et al., Optimization of medium chain length fatty acid incorporation into olive oil catalyzed by immobilized Lip2 from Yarrowia lipolytica, Biochem. Eng. J. 77 (2013) 20–27.

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Please cite this article in press as: W. Li, et al., Synthesis of ethyl oleate by esterification in a solvent-free system using lipase immobilized on PDMS-modified nonwoven viscose fabrics, Process Biochem (2015), http://dx.doi.org/10.1016/j.procbio.2015.07.012