Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres

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Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres Juliana Bergamasco a, Marcelo V. de Araujo a, Adriano de Vasconcellos a, Roberto A. Luizon Filho a, Rafael R. Hatanaka b, Marcus V. Giotto c, Donato A.G. Aranda d, Jose´ G. Nery a,* a

Departamento de Fı´sica, Instituto de Biocieˆncias, Letras e Cieˆncias Exatas, UNESP e Universidade Estadual Paulista, Campus de Sa˜o Jose´ do Rio Preto, Sa˜o Paulo 15054-000, Brazil b Universidade Estadual Paulista Ju´lio de Mesquita Filho, Instituto de Quı´mica de Araraquara, Departamento de Quı´mica Orgaˆnica, Rua Prof. Francisco Degni, s/n, Quitandinha, 14800-900 Araraquara, SP, Brazil c Institute of Materials Science, University of Connecticut, 97 North Eagleville Road, Storrs, CT 06269-3136, USA d Laborato´rio Greentec, Escola de Quı´mica/UFRJ, Caixa Postal 68572, Rio de Janeiro-RJ, Cep 21941972, Brazil

article info

abstract

Article history:

Polyvinyl alcohol (PVA) microspheres with different degree of crystallinity were used as

Received 11 February 2013

solid supports for Rhizomucor miehei lipase immobilization, and the enzyme-PVA complexes

Received in revised form

were used as biocatalysts for the transesterification of soybean oil to fatty acid ethyl esters

9 September 2013

(FAEE). The amounts of immobilized enzyme on the polymeric supports were similar for

Accepted 13 September 2013

both the amorphous microspheres (PVA4) and the high crystalline microspheres (PVA25).

Available online xxx

However, the enzymatic activity of the immobilized enzymes was depended on the crystallinity degree of the PVA microspheres: enzymes immobilized on the PVA4 microspheres

Keywords:

have shown low enzymatic activity (6.13 U mg
1), in comparison with enzymes immobi-

Biomass

lized on the high crystalline PVA25 microspheres (149.15 U mg
1). A synergistic effect was

Fatty acid ethyl esters (FAEE)

observed for the enzyme-PVA25 complex during the transesterification reaction of soybean

Enzyme immobilization

oil to FAEE: transesterification reactions with free enzyme with the equivalent amount of

Polymeric support

enzyme that were immobilized onto the PVA25 microspheres (5.4 U) have yielded only 20%

Poly(vinyl) alcohol

of FAEE, reactions with the pure highly crystalline microsphere PVA25 have not yielded FAEE, however reactions with the enzyme-PVA25 complexes have yielded 66.3% of FAEE. This synergistic effect of an immobilized enzyme on a polymeric support has not been observed before for transesterification reaction of triacylglycerides into FAEE. Based on ATR-FTIR, 23Na- and 13C-NMR-MAS spectroscopic data and the interaction of the polymeric network intermolecular hydrogen bonds with the lipases residual amino acids a possible explanation for this synergistic effect is provided. ª 2013 Elsevier Ltd. All rights reserved.

* Corresponding author. Tel.: þ55 17 3221 2490; fax: þ55 17 3221 2247. E-mail addresses: [email protected], [email protected] (J.G. Nery). 0961-9534/$ e see front matter ª 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biombioe.2013.09.006

Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

2

1.

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Introduction

Chemically biodiesel is a mixture of fatty acid methyl or ethyl esters (FAME or FAEE) formed by the direct transesterification of vegetable oils or animals fat. In this reaction, triglycerides molecules react with an excess of short-chain alcohol (methanol or ethanol) in the presence of an appropriate catalyst, undergoing a sequence of three consecutive reversible reactions, which include conversion of triglycerides to diglycerides, followed by the conversion of diglycerides to monoglycerides [1,2]. Biodiesel can be produced using different types of catalysts, and each one of them has its advantages and disadvantages, and excellent reviews dealing with the pros and cons of using these catalysts for biodiesel production in large scale have been published in the recent years [3e5]. Among these catalysts, enzymes are very attractive for biodiesel production due to their specificity, mild reaction conditions and easy purification of the subproduct glycerol [6,7]. Lipases (triacylglycerol ester hydrolases, EC 3.1.1.3) are high specific used enzymes employed in the transesterification and esterification of triglycerides into FAME or FAEE, since through specific mechanisms they can naturally catalyze the breakdown of fats and oils with subsequent release of free fatty acids, diacylglycerides, monoglycerides and glycerides at a watereoil interface [8,9]. However, there are several drawbacks for using enzymes for the biodiesel production in large industrial scale, such as their elevated economic cost, difficult recovery of the catalyst after the reaction and high possibility of deactivation or denaturation of the enzymes [7,10]. One possible and viable alternative to overcome some of these drawbacks is the immobilization of the enzymes on different kind of inorganic [11,12] and organic solid supports [2,13e15]. In general the enzyme immobilization process is followed by an enhancement of their thermal stability, enzymatic activity and in some few cases even a synergistic effect between the enzymes and the inorganic supports has been observed [11]. Among several available biodegradable polymers that can be employed for the preparation of polymeric matrixes to used as solid supports for the immobilization of cells, enzymes and proteins, poly(vinyl alcohol) (PVA) is one of the most versatiles due to its easy availability, chemical structure, non-toxicity and low price. In general the immobilization of cells, proteins and enzymes on PVA supports is done through covalent bonding, adsorption, chemical cross-linking, photo-crosslinking, freezing and thawing [15]. Recently high crystalline PVA microspheres specially designed for biomedical applications, more specifically for embolization and chemoembolization medical procedures were reported in the literature [16,17]. One of the main structural features of these highly crystalline PVA microspheres is formation of large percentage of intermolecular hydrogen bonds formed by the chemical bonding of the CH carbon of the polymer interchains with hydroxyl groups in the crosslinked region [16]. The presence of the these intermolecular hydrogen bonds could in principle allow the use of

these PVA microspheres as a solid support for enzymes immobilization, therefore in the present work we report the immobilization of Rhizomucor miehei lipase onto highly crystalline microspheres and the use of these enzyme-PVA complexes as catalysts in the transesterification of soybean oil into FAEE. Several parameters such as the amount of immobilized enzyme, enzyme activity of immobilized and free enzyme, the yield of the fatty acid ethyl esters (FAEE) obtained using the enzyme-PVA complexes as catalysts and the occurrence synergetic effects for enzyme-PVA complexes were investigated.

2.

Experimental section

2.1.

Materials

2.1.1.

Chemicals

In the preparation of PVA microspheres the following chemicals were used: polyvinyl alcohol (molecular weight of 78,000 Da and degree of hydrolysis 86.5e89.5%) and benzoyl peroxide (minimum purity of 99%) were obtained from Vetec Quı´mica Fina (Brazil); terc-butanol, sodium hydroxide and vinyl acetate (minimum purity of 99%) were obtained from Merck and sodium sulfate 99% was obtained from Sigma Aldrich. The immobilization of lipase onto PVA microspheres was performed using phosphate acetate buffer prepared from sodium phosphate monobasic and sodium phosphate dibasic (minimum purity of 99%) obtained from Sigma Aldrich. For the determination of unbound protein was used Bradford reagent obtained from Sigma Aldrich. 4-Nitrophenyl palmitate (Sigma Aldrich), triton X-100 and isopropyl alcohol were used for the esters hydrolysis reactions. Refined soybean oil cultivated from Glycine max (L.) Merr containing 0.5% of free fatty acid and 1% of water obtained from Cargill Oil Company (Brazil) and anhydrous ethanol (Sigma-Aldrich, 99.8%) was used for syntheses of the FAEE.

2.1.2.

Enzyme

The enzyme used was the commercial PALATASE 20,000 L (molecular size 31,600 Da, pI 3.8, obtained from SigmaeAldrich), which is a purified lipase from Rhizomucor miehei 1.3 specific (EC 3.1.1.3), produced by submerged fermentation of genetically modified Aspergillus oryzae microorganism.

2.2.

Experimental procedure

2.2.1.

Polyvinyl alcohol (PVA) microspheres syntheses

PVA microspheres were synthesized according to a procedure previously described in the literature [16]. A typical synthesis was prepared in two steps. Step 1: Preparation of the PVAc microspheres: In a homemade reactor (250 cm3) fitted with condenser and under nitrogen atmosphere, a solution A (70 mg of polyvinyl alcohol (suspending agent) dissolved in 100 cm3 of distilled water) was slowly warmed up until 70  C, and then kept at this temperature for 60 min with constant stirring. After 60 min, 1 g of benzoyl peroxide (initiator) was added to the solution A, and 2 min later a solution B (25 cm3 of vinyl acetate monomer dissolved in 25 cm3 of alcohol terc-butyl) was also added to the

Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

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solution A. The reaction was allowed to continue for 4 h resulting in poly(vinyl acetate) (PVAc) microspheres. Step 2: Saponification of the PVAc microspheres to PVA microspheres: The reaction performed in step 1 was cooled down to 35  C. Then to this reaction it was added 10 cm3 of a saline solution of Na2SO4 (100 kg m
3 concentration). Immediately after the addition of the saline solution, NaOH solutions with different concentrations (4 mol m
3, 12 mol m
3 and 25 mol m
3) were slowly added to the reaction. The saponification reactions were conducted with continuous stirring for 12 h at 35  C. Afterwards the final products were filtrated and washed several times with distilled water until the pH of the residual water has reached the pH range of 6e8. The microspheres were then poured into cold distilled water (10  C) and frozen (
20  C) for 12 h. Microspheres saponificated with NaOH (4 mol m
3) were amorphous ones and were named PVA4, while microspheres saponificated with NaOH (12 mol m
3 and 25 mol m
3) were crystalline ones and were named PVA12, and PVA25 respectively. The final products were lyophilized (Liobras, Sa˜o Carlos, Brazil) before the physicochemical characterization by XRD, SEM, ATR-FTIR, 13C CPMAS-NMR, 23Na MAS-NMR.

2.2.2.

Immobilization of lipase onto PVA microspheres

The immobilization of the enzyme on the PVA microspheres was carried out according with a methodology described by Lee et al. [18] and Dizge et al. [2]. The experiments were performed in the following manner: In a homemade reactor a solution consisting of 1.5 mg of enzyme was dissolved into 20 cm
3 of phosphate buffer (0.05 mol m
3, pH 7.0) and to this solution it was added 400 mg of soybean oil. This mixture was incubated at 37  C for 30 min under constant stirring, afterwards 1 g of PVA microspheres (100e500 mm diameter range) were added to it. The experiment was kept at room temperature for 25 h with gentle stirring, afterwards the microspheres were collected by centrifugation, and then washed with the phosphate buffer in order to remove all the unbound protein and the enzyme-polymer complexes were lyophilized and stored at low temperature (
20  C) until the use. The amount of the immobilized enzymes was determined according to the Bradford method [19] using the supernatant phase. The detailed Bradford experiments are described in Section 2.2.4.

2.2.3.

Physical characterization of PVA microspheres

All the pure and immobilized PVA microspheres were characterized by XRD, SEM, ATR-FT-IR, 13C CPMAS NMR, 23Na NMR and zeta potential. XRD data were collected with a Rigaku RotaFlex RU200B (Tokyo, Japan) on a rotating anode source with a flat-plate BraggeBrentano geometry, operating with Cu Ka radiation (wavelength ¼ 1.5418  A) at 50 kV and 100 mA, and equipped with a graphite monochromator. The powder diffraction patterns were recorded in the range 2a ¼ 5e80 with a step scan of 0.02 and at a rate of 10 s/step. SEM images were recorded using an XL30 FEG instrument, and before the analysis, a thin coating of gold was deposited onto the samples. ATR-FTIR spectra were collected using a Smart Orbit ATR (Thermo Scientific) diamond accessory and a Nicolet 6700 spectrometer (Thermo Scientific) equipped with a DTGS detector. All samples were scanned for 64 times at the spectral resolution of 4 cm
1 between 4000 cm
1 and 500 cm
1.

3

Solid-state magic angle spinning (MAS) NMR spectra were acquired with a Bruker DMX 300 NMR spectrometer operating at field strength of 7.05 T, hence a resonance frequency of 75.4 MHz for 13C and 79.38 MHz for 23Na (300 MHz for 1H) by using a double resonance 4 mm MAS probe. The magic angle of the double resonance 4 mm MAS probe was set using the 79Br resonance of KBr. Approximately 100 mg of each sample was packed in a Zirconia rotor with Kel-F caps and spun at 10 kHz at room temperature. 13C NMR: the solid-state cross polarization 13C CPMAS NMR spectra were acquired with two pulse phase modulation (TPPM) proton decoupling with 75 kHz of field strength. The 90 pulse width for 1H was 3.6 ls with contact time of 1 ms, delay time of 5 s, 20,000 scans, 4096 data points, and 30 kHz of spectral width. Chemical shifts were given with respect to tetramethyl silane (TMS) by using an external sample of solid glycine (carbonyl at 176.5 ppm) for 13 C as the secondary reference. 23 Na NMR: the short selective pulse width was 1.6 ms (p/6) with a delay time of 2 s applied to allow thermal equilibrium. There were 50,000 scans performed yielding 8192 data points and 50 kHz of spectral width. A Lorentzian line broadening of 20 Hz was applied to the data. Chemical shifts were given with respect to a 0.1 M NaCl solution as the secondary reference. The zeta potential of the samples was measured to determine the surface properties with pure and immobilized PVA samples using Nano Zetasizer ZS 90 (Malvern Instruments, Worcester shire, UK). For measurement, the pure and immobilized supports were dissolved in distilled water at a concentration of 1 mg cm
3.

2.2.4. Determination of the unbound proteineBradford reaction The concentration of the immobilized enzyme was determined through the method described by Bradford which uses bovine serum albumin as a standard [19]. The concentration of enzyme adsorbed on the PVA microsphere, Pg (g kg
1), was calculated using the following equation (1): Pg ¼

Co Vo
Cf Vf ; w

(1)

where C0 is initial protein concentration (mg cm
3), Cf the protein concentration of the filtrate (mg cm
3), V0 the initial volume of lipase solution (cm
3), Vf the volume of filtrate (cm
3) and w is the weight of PVA support used (kg) [12]. In a typical experiment, 950 mm3 of Bradford Reagent was added into an eppendorf, and this solution was mixed with 50 mm3 of the supernatant phase (see Section 2.2.2). After 20 min, the samples were measured in a spectrophotometer using a wavelength at 595 nm.

2.2.5.

Determination lipase activity

The enzymatic activities of the free lipase and the lipase immobilized on PVA microspheres were determined by the hydrolysis of p-4-nitrophenyl palmitate (p-NPP) following the formation of 4-nitrophenol by the method of Winkler and Stuckmann [20] and Krieger et al. [21]. According to this method a stock solution of the p-NPP (3 mg cm
3) was prepared using HPLC grade isopropyl alcohol (solution A). Another solution (solution B) comprised of (2%) triton X-100, diluted in phosphate buffer (pH 7; 0.05 M) was prepared, and

Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

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then 1000 mm3 of solution A was added to 9000 mm3 of solution B. This mixture or final solution was named substrate. The activity of free enzyme was measured according to the following procedure: In 900 mm3 of the substrate solution was added to 100 mm3 of free enzyme diluted in the phosphate buffer, and then the mixture was incubated for 1 min at 37  C in a water bath. The 4-nitrophenol released was determined photometrically (410 nm) and the assays were performed in triplicate and one unit of lipase activity was defined as the amount of enzyme required to produce 1 mmol of 4nitrophenol released from 4-NPP per minute under the assay conditions. The lipolytic activity of free enzyme, U, was calculated according to equation (2).   Abs  Vt  103  D ε  Xe  T

2.2.7.

 U¼

(2)

U ¼ Unit of enzyme activity is expressed as U/mL, where one unit of activity is defined as the release of 1 mmol p-nitrophenol released from the p-NPP/mL min within the assay conditions; Abs ¼ Absorbance of sample at 410 nm; Vt ¼ Total volume of reaction (mL); Xe ¼ Volume of enzyme solution (mL) or mass of the immobilized enzyme; ε ¼ Coefficient of molar extinction (L mol
1 cm
1); T ¼ Incubation time; 103 ¼ Factor correction of coefficient (ε); D ¼ Sample dilution, if necessary. The activities of pure PVA and enzyme-PVA complexes were measured according to the following procedure: 1 mg of the samples were mixed with 100 mm3 of phosphate buffer (pH 7) and 900 mm3 of substrate, and then this mixture incubated for 1 min in a water bath kept at 37  C. The reaction was interrupted by adding 150 mm3 of 0.1 mol m
3 sodium carbonate with 5% triton X-100 to the previous mixture and cooling it down with the help of an ice bath. Absorbance measurements at 410 nm were immediately performed against a blank reaction without enzyme.

2.2.6.

reactions was followed by thin layer chromatography (TLC) based on a procedure described by Yang et al. [23]. At predetermined time intervals, a small volume (100 mm3) of the reaction mixture was collected, and mixed with 500 mm3 of hexane for 2 min. After separation by centrifugation, 3 mm3 of the upper layer was applied to a silica gel plate. A solution of hexane/ethyl acetate/acetic acid (90:10:1) was used as a developing solvent and a solution of methanol/sulfuric acid (1:1) was used as a color reagent. After spraying the color reagent over silica gel plate, followed by the development solvent, the silica gel plate was heated at high temperature and then analyzed.

FAEE production using the enzyme-PVA complexes

The transesterification reactions of the soybean oil to FAEE were performed with the two different catalysts groups: free enzyme and the enzyme-PVA complexes (enzyme-PVA4, enzyme-PVA12 and enzyme-PVA25). Transesterification reactions catalyzed by the free enzymes were performed with an oil:ethanol ratio of 1:6 and 0.08 mg of enzyme (equivalent to 5.4 U which were the total amount of enzymes immobilized on the polymeric supports) at 37  C and ethanol was slowly added in order to preclude the inactivation of the enzyme. After 72 h of reaction the products (FAEE, glycerol) were separated by centrifugation at 7280 g. Transesterification reactions with the enzyme-PVA complexes (enzyme-PVA4, enzyme-PVA12 and enzyme-PVA25) were also performed in the same experimental conditions and the amount of, the biocatalysts used were of 4% relative to mass of oil [22]. The progress of the transesterification

Gas chromatography

The quantifications of ester content in FAEE samples were performed, in triplicates, according to EN 14103 (Fat and oil derivatives e Fatty Acid Methyl Esters (FAME) e Determination of Ester and linolenic acid methyl esters contents) in a Shimadzu GC 2010 gas chromatograph with flame ionization detector (GC-FID). The chromatograph was configured with injector in split mode coupled to auto-sampler AOC 5000 for liquid samples. EN 14103 establishes the chromatographic conditions used in the quantification of fatty acid esters: sample injection volume ¼ 1 cm3, split ¼ 1:20, injector and detector temperatures ¼ 250  C, isothermal oven temperature ¼ 210  C, pressure of helium carrier gas ¼ 83 kPa due to “split” or adjusted to visualize clearly the peak of the methyl standard C24:1. The quantification of the percentage of ethyl esters in the samples was performed as following: 20 mg cm
3 stock solutions of methyl myristate (C14:0) and methyl nervonate (C24:1) standards were prepared, while methylic and ethylic heptadecanoate standards were prepared in 10 mg cm
3 stock solutions. Methyl esters standards C14:0 and C24:1 were used to identify the range of integration. In addition, we used the methylic (C17:0) ester as an internal standard (IS).

2.2.8.

Thermodynamic parameters

Preliminary thermodynamic parameters such as energy of activation (Ea), energy of deactivation (Ed), enthalpy ðDHoa Þ and entropy ðDSoa Þ of activation were calculated following the standard procedures reported in the literature [24e30]. According to these studies the thermal activation of the lipase in its free and immobilized form obey the following Arrhenius relation:
Ea AT ¼ Aexpð RT Þ ;

(3)

where AT is the enzyme activity, A is the pre-exponential factor, Ea is the activation energy, R is the gas constant and T is the absolute temperature in Kelvin [24]. One of the mean reasons to immobilize enzymes on solid supports is to decrease their elevated economic cost, therefore it is important to determine the economic feasibility of the immobilized enzyme process [7,10]. In this case it is important not only to determine the activation energy (Ea) of the enzymes, but also the potential for loss of the enzymatic activity during use. Usually the primary cause of the loss of enzyme activity is thermal deactivation of the enzyme which is correlated to the energy of deactivation (Ed). Several studies

Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

b i o m a s s a n d b i o e n e r g y x x x ( 2 0 1 3 ) 1 e1 6

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reported in the literature [24e27] treat the thermal deactivation of enzyme as a first order reaction and as a consequence of this assumption the deactivation rate constant can be obtained following an adapted Arrhenius type of reaction as the one described by the equation (4): ln DAT ¼


Ed þ lnDA RT

(4)

where DAT is the enzyme activity in deactivation state, DA is the initial enzyme activity in deactivation state and Ed is the inactivation energy [24,26,27]. In this study the Ea was determined using the equation (3) by taking into consideration the dependence of the lipase activity (AT) in the 30  Ce44  C temperature range for both the free and the immobilized lipase. The determination of the Ed for the free and immobilized enzyme was calculated using the equation (4) by taking also into consideration the dependence of the lipase activity (AT) in 44  Ce60  C temperature range. The enthalpy ðDHoa Þ and entropy ðDSoa Þ of activation were calculated from the plot of ln k/T versus 1/T according to the procedures described by Yan et al. [28], and other authors [29,30].

3.

Results and discussion

Fig. 1 e XRD patterns of the as made PVA microspheres: a) amorphous PVA4 microspheres, b) crystalline PVA12 microspheres and c) crystalline PVA25 microspheres.

3.1. XRD and SEM characterization of the as made PVA microspheres and the enzyme-PVA complexes XRD data provide strong evidences that the crystalline structures of synthesized polymeric particles change with the concentration of NaOH used during the saponification process. The saponification reactions performed with NaOH 4 mol m
3 concentration have resulted in polymeric particles with low crystallinity and their XDR patterns are characterized by very broad Bragg diffraction peaks in the regions 2q ¼ 7e30 as shown in Fig. 1a, and the absence of well-defined and intensive Bragg diffraction peaks for these samples confirms their amorphous nature. On the other hand, it was noticed that increasing the NaOH concentration to 12 mol m
3 and 25 mol m
3 has caused the formation of polymeric particles with higher crystallinity as shown in the XDR patterns of these samples (Fig. 1b, c). The most striking case was observed for the samples treated with NaOH concentration of 25 mol m
3, since the typical XDR patterns of these samples have displayed four distinct Bragg diffraction peaks at the 2q region of 11.5, 19.5, 23.0 and 40.5 as shown in Fig. 1c. The width and the peak shape of the XRD pattern of the PVA25 microspheres in comparison with the other ones (PVA12 and PVA4) clearly indicate that the PVA25 microspheres are more crystalline than the other ones. Fig. 2 shows SEM micrographs of the as made PVA4, PVA12 and PVA25 microspheres before immobilization. PVA4 is characterized by a smooth and irregular surface (Fig. 2a), while for PVA12 and PVA25 there are the formation of scaffolds (Fig. 2b, c). SEM of the enzyme-PVA25 complex (Fig. 3) shows an apparently homogeneous distribution throughout the PVA microsphere, where all the previous free scaffolds present in the sample were completely filled up with the enzyme.

3.2. Effect of the immobilization on the optimal temperature and thermal stability of the enzymes In order to determine the optimum temperature for Rhizomucor miehei enzyme, several enzymatic activities assays were carried out with the free enzyme at temperature ranges varying from 36  C to 52  C and its optimal temperature was found at 44  C (Fig. 4). Increasing the temperature above the optimum temperature (44  C) had caused a decrease of enzyme activity which could be attributed to the deactivation and denaturation of the enzyme. These results are consistent with previous studies reported in the literature for the Rhizomucor miehei’s optimum temperature, which were also found to be in the temperature range of 30  Ce50  C at a pH of 8.0 [31e33]. In order to verify if the enzyme immobilization onto PVA microspheres had affected the original enzyme’s optimum temperature (44  C), several enzymatic assays were performed with the enzyme-polymer complexes. The results have shown that there were no significant variations for enzyme-PVA12 and enzyme-PVA25 complexes, and the optimum temperature was found to be the same for both the free enzyme and the immobilized enzyme. On the other hand, immobilization of the enzyme on PVA4 microspheres has resulted in significant decrease of the enzyme activity, and no optimum temperature was determined for this enzymepolymer complex due to the fact that enzyme activity was kept constant in the 36  Ce52  C temperature range (see Fig. 4). The thermal stability of the Rhizomucor miehei lipase was also studied for free enzyme and the polymer-enzyme complexes at 44  C which was the experimental optimal temperature found for the free Rhizomucor miehei lipase in this investigation, and the experiments were performed at six

Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

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Fig. 2 e SEM micrographs of the polymeric supports: a) PVA4, b) PVA12 and c) PVA25.

different time intervals (0, 60, 120, 180, 240 and 300 min) for the free enzyme and the enzymes-polymer complexes. The thermal stability profiles of the free enzyme and of the polymer-enzyme complexes (enzyme-PVA4, enzymePVA12, enzyme-PVA25) are shown in Fig. 5. It can be noticed that for the free enzyme after 60 min reaction at 44  C there was an enzymatic activity reduction of 10% in comparison

with the original value, and it was kept constant through the lasting 300 min. On the other hand, the polymer-enzyme complexes (enzyme-PVA12, enzyme-PVA25) have shown no reduction in their original enzymatic activities, since the enzymatic activities were kept constant from the beginning until the end of the experiments. It means that the immobilization process has increased the thermal stability of the

Fig. 3 e a) SEM micrograph of as made PVA25 support, b) SEM micrograph of the enzyme-PVA25 complex. Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

b i o m a s s a n d b i o e n e r g y x x x ( 2 0 1 3 ) 1 e1 6

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Fig. 4 e Effect of temperature on the enzymatic activity of free and immobilized Rhizomucor miehei lipase on the different PVA supports.

enzyme compared to its thermal stability in the free form, which has suffered a reduction of 10% in the first 60 min of the reaction. An exception was noticed for the enzyme-PVA4 complex, because it has exhibited from the beginning until the end of the experiment low enzymatic activity. These results are in agreements with other studies reported in the literature [34,35].

3.3. ATR-FTIR analyses of the PVA microspheres and enzyme-PVA complexes The main possible interactions involved in the adsorption of proteins on solid surfaces are hydrophobic and electrostatic interactions and also hydrogen bonds [36]. In order to understand the interactions of the immobilized enzymes onto the PVA supports, ATR-FTIR studies were performed for the as made PVA microspheres and enzyme-PVA complexes. ATRFTIR has been extensively used to examine the structure of proteins not only in solution, but also the structural features of adsorbed proteins on diverse classes of solids [37e39]. According to the literature [40,41] the main characteristic absorption bands for PVAc and their respective chemical assignment are: 2925 cm
1 (CeH stretching), 1730 cm
1 (C]O stretching), 1376 cm
1 ((CH)eReCH3 bending) and 1130 cm
1(CeOeC stretching), while for PVA [40,42] the absorption bands and their respective assignments are: 3550e3200 cm
1 (OH.OH stretching), 2900e2700 cm
1 (C]H

stretching), 1650e1630 cm
1 (OH.OH bending),
1 ((CH)eCH2 bending), and 1141 cm
1 (CeO 1461e1417 cm stretching). Fig. 6 shows the experimental spectra for the as made PVA4, PVA12 and PVA25 microspheres. In the case of PVA4 (Fig. 6a) the main observed absorption bands 2937 cm
1, 1730 cm
1, 1424 cm
1, 1373 cm
1, 1223 cm
1 and 1017 cm
1. All the observed absorption bands are in agreement with the data reported in the literature [40,41]. The presence of an intense absorption band at 1730 cm
1 was assigned to the acetyl groups, and it was clear indication that the saponification process was an inefficient in the preparation of the PVA4 microspheres. On the other hand, this absorption band (1730 cm
1) has completely disappeared in the spectra of PVA12 and PVA25 samples as shown in Fig. 6b and c. This was a clear indication that the saponification processes were very efficient in the case of these samples. The ATR-FTIR spectra of PVA12 and PVA25 were quite similar, except for two additional bands observed for PVA25 at 1650 cm
1 and 1560 cm
1 as shown in the see expanded Fig. 6c. The absorption band at 1650 cm
1 was assigned to the OHeOH bend, while the 1560 cm
1 was attributed to the interaction of the sodium ions and the carboxyl group (COO
), which was a direct consequence of the high concentration of the sodium hydroxide used in the preparation of this sample [43,44]. ATR-FIT can also provide useful insights into the secondary structure changes of the enzymes caused by the immobilization process. ATR-FTIR spectra of proteins have an

Fig. 5 e Enzymatic stability at 44  C of free and immobilized Rhizomucor miehei lipase on the different PVA supports. Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

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Fig. 6 e FTIR-ATR of the as made PVA microspheres: a) PVA4, b) PVA12 and c) PVA25.

important “finger print” in the region 1200 cm
1e1700 cm
1, since this region covers three majors group of infrared absorption bands related to proteins. The first one is related to the amide I band (C]O stretch weakly coupled with CeN stretch and the NeH bending) and it is located in the 1610 cm
1e1700 cm
1 region. The second one is related to the amide II band (CeN stretch strongly coupled with the NeH bending) and it is found in the region 1530 cm
1e1550 cm
1. The third one is related to the amide III (NeH in plane bending coupled to CeN stretching; CeH and NeH deformation) and it is mainly observed between 1450 cm
1 and 1200 cm
1 regions [42,45,46]. The experimental ATR-FTIR data for the free and the enzyme-PVA complexes are shown in Fig. 7. A detailed ATRFTIR spectrum of the free Rhizomucor miehei lipase is shown in Fig. 7d and it shows the maximum of the amide I band occurs at 1650 cm
1 and this band consists of a group of overlapped signals, which contains information on secondary protein structure of the enzymes. The characteristic bands of amide III (1413 cm
1, 1310 cm
1, 1220 cm
1) were also observed for the free enzymes [47]. Analyses of the ATR-FTIR for enzyme-PVA12 (Fig. 7b) and enzyme-PVA25 (Fig. 7c) complexes show that the adsorbed Rhizomucor miehei lipase has kept most of its b-sheet structure in comparison with the free enzyme (Fig. 7d). The fact that there was no difference in the positions, bandwidths,

intensity and area ratio of the amide I, strongly suggests that the lipase was adsorbed on the enzyme-PVA12 and enzymePVA25 microspheres without drastic changes of the b-sheet structure. The characteristic bands of the amide III were also kept unchanged for these complexes. However, the same did not occur for enzyme-PVA4 complex (Fig. 7a). Analyses of the adsorption bands at the amide I region will clearly show that there were drastic changes in the position, bandwidths and intensity of the adsorption band related to the b-sheet structure, indicating that the enzyme was not immobilized in its active form. The low yield of FAEE with the enzyme-PVA4 in comparison with the other enzyme-PVA complexes (see Section 3.7) corroborates this observation.

13 3.4. C-CPMAS-NMR and 23Na-MAS-NMR of the pure PVA microspheres and the enzyme-PVA complexes 13

C CPMAS NMR spectrum of the pure PVA microspheres (PVA4, PVA12 and PVA25) is shown in Fig. 8. The 13C CPMAS NMR spectra were similar to the ones reported by Semenzim et al. [16]. There are five chemical shifts associated to the PVA4 microsphere shifts (Fig. 8c): 170.53, 129.72, 66.75, 39.64 and 20.99 ppm. Chemical shifts at 170.53 and 20.99 ppm were assigned to the PVAc, while chemical shifts at 39.69, 66.75 and 20.9 were associated to the PVA. Therefore, on the basis of

Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

b i o m a s s a n d b i o e n e r g y x x x ( 2 0 1 3 ) 1 e1 6

9

Fig. 7 e FTIR-ATR of the enzyme-PVA complexes: a) enzyme-PVA4, b) enzyme-PVA12 and c) enzyme-PVA25, d) free enzyme.

these values reported in the literature, it is clear that PVA4 microspheres are a blend of the two polymers. PVA12 and PVA25 spectra (Fig. 8b, a) show four distinct chemical shifts at 77.33, 71.32, 65.03 and 45.9 ppm which is indication of pure PVA microspheres. The chemical shifts located at 77.33, 71.32, and 65.03 ppm were associated with the intramolecular and intermolecular hydrogen bonding of the CH carbons (methine carbons) to the hydroxyl groups, while 45.9 ppm is associated with the methylene carbon resonance [16,48]. The immobilizations of the enzymes on the PVA4, PVA12 and PVA25 have induced no significant structural modification in the polymeric network, since the 13C-CPMAS NMR spectra of the enzyme-PVA complexes were similar to the 13CCPMAS NMR spectra of the pure polymer. One possible explanation for this behavior might be due to the fact that the 13C-CPMAS NMR technique is not sensitive enough to detect the small amount of immobilized enzymes onto the polymeric supports. On the other hand, significant difference between the 23NaMAS-NMR spectra of the pure polymeric supports (PVA4, PVA12 and PVA25) and the enzyme-PVA complexes (enzymePVA4, enzyme-PVA12 and enzyme-PVA25) have been observed as shown in Figs. 9 and 10. Chemical shifts observed for pure PVA4 (7.30 ppm,
7.55 ppm), PVA12 (0.50 ppm,
7.80 ppm) and PVA25 (
2.57 ppm,
10.44 ppm) are associated with two different sodium environments. According to

the literature three species of sodium ions have been identified in polymeric networks: isolated (5.0e7.5 ppm), fully hydrated (0 to
2.5 ppm) and aggregated sodium ions (
12 to
23 ppm) [49,50]. Therefore, based on the literature the two different chemical shifts observed for PVA4 can be assigned to isolated sodium ions (7.3 ppm) and to the aggregated sodium ions (
7.5 ppm). It is interesting to note that the chemical shift associated with hydrated sodium ions is absent in the 23NaMAS-NMR spectrum of PVA4. Chemical shifts 0.5 ppm (PVA12) and
2.5 ppm (PVA25) can be assigned to the hydrated sodium ions, while
7.8 ppm (PVA12) and
10.4 ppm (PVA25) are associated with aggregated sodium ions. Chemical shifts associated with the presence of isolated sodium ions were not observed for the samples PVA4 and PVA25. 23 Na-MAS-NMR data of the enzyme-PVA complexes clearly indicates that the immobilization of the Rhizomucor miehei enzyme onto the PVA surfaces has caused major changes of the sodium ions environments in the polymeric network. Fig. 10c shows the 23Na-MAS-NMR of the enzyme-PVA4 complex and it reveals now four distinct chemical shifts at 4.78,
1.00,
5.54 and
13.85 ppm. It should be noticed that the isolated sodium ions observed for the PVA4 have completely disappeared, indicating that somehow the enzyme has interacted with the isolated sodium ions. It is possible that now all the sodium ions existing in the enzyme-PVA4 are in the form of hydrated and aggregated sodium ions. Enzyme-PVA12

Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

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b i o m a s s a n d b i o e n e r g y x x x ( 2 0 1 3 ) 1 e1 6

Fig. 8 e

13

C-CPMAS-NMR of the as made PVA microspheres: a) PVA25, b) PVA12 and c) PVA4.

revealed eight chemical shifts of 0.95,
3.58,
6.02,
15.79,
47.86,
55.28,
74.42,
90.45 ppm, whereas enzyme-PVA25 reveals seven chemical shifts at 0.49,
6.23,
12.96,
21.19,
25.18,
74.06,
90.76 ppm (Fig. 10b, a). These new chemical shifts indicate that the enzyme interacts with the sodium ions of the polymeric network, therefore creating new complex environments for the sodium ions. Although the correct assignment of all chemical shifts observed for enzyme-PVA12 and enzyme-PVA25 complexes is not a straight forward task, it is reasonable to affirm that the only two possible environments for the existence of sodium ions in these two complexes, are either in the form of hydrated or aggregated species [49,50]. To the best of our knowledge this is the first 23 Na-MAS-NMR data that clearly shows the influence of an enzyme in the reorganization of the sodium ions environment in a polymeric network.

3.5.

Zeta potential

Electrostatic potential of the pure PVA microspheres was measured and the data are shown in Fig. 11. As it was expected all the microspheres have presented a negative charged surface, however the electrostatic potential varies according to the concentration of NaOH used in the saponification reaction. The experimental electrostatic potential for all microspheres without enzyme immobilization was the following: PVA4 (
23.2 mV), PVA12 (
20.4 mV) and PVA25

(
8.84 mV). One possible explanation for the smallest electrostatic potential observed for PVA25 microspheres compared to the others, is probably due to the fact that the large amount of Naþ present in the polymeric network counterbalances the OH
of the polymeric network, therefore decreasing the surface electrostatic potential. After the immobilization there was an increase of the negative charges on the surface of all enzyme-polymer complexes in comparison with the pure microspheres: (enzyme-PVA4 (
38.70 mV), enzyme-PVA12 (
31 mV), enzyme-PVA25 (
28.90 mV)). In principle the increasing of the negative charge of the enzymecomplexes could positively affect the lipase activity according to the mechanism of “electrostatic catapult” proposed by Peterson et al. [51]. According to this mechanism after the cleavage of an ester bond by a lipase, a free fatty acid is released. This fatty acid will become deprotonated, thus carrying a negative charged molecule that easily causes the acylation of the enzyme, therefore decreasing its activity. Therefore, according to this electrostatic catapult mechanism, the high negative potential existing at the bottom of active site would contribute to a more efficient repulsion of the negatively charged free fatty acid, therefore avoiding the acylation and the decrease of the lipase’s active. However the fact that enzyme-PVA4 complex has presented the lowest enzymatic activity and FAEE yields compared to the other complexes indicates this mechanism cannot straightforward applied to these complex systems.

Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

b i o m a s s a n d b i o e n e r g y x x x ( 2 0 1 3 ) 1 e1 6

Fig. 9 e

Fig. 10 e

11

23

Na-MAS-NMR of the as made PVA microspheres: a) PVA25, b) PVA12 and c) PVA4.

23

Na-MAS-NMR of the enzyme-PVA complexes: a) enzyme-PVA25, b) enzyme-PVA12 and c) enzyme-PVA4.

Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

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Fig. 11 e Zeta potential for the pure polymeric supports and the enzyme-PVA complexes.

3.6. Lipase immobilization efficiency and enzymatic activity of immobilized enzymes The amount of lipase immobilized onto the polymeric supports and enzymatic activity of free and immobilized enzymes were investigated. The amount of immobilized enzyme was 30 g kg
1 for all the three microspheres, however enzyme and their enzymatic activity were quite different from each other. The enzymatic activity measured for enzyme-PVA4 complex was very low (6.13 U mg
1) in comparison with the enzyme-PVA12 complex (67.23 U mg
1) and enzyme-PVA25 complex (149.15 U mg
1). Since all the three supports have immobilized the same amount of enzyme (30 g kg
1) enzymatic activity assays were performed with the free enzyme using the equivalent amount of the immobilized enzymes. In this case the measured enzymatic activity was of (11.6 U mg
1) (see Fig. 12). The correlation between these data and the ATR-FTIR data strongly suggested that the enzyme immobilized onto the PVA4 was not immobilized in its active form, therefore showing a low enzymatic activity in comparison to the others. The high enzymatic catalytic observed for enzyme-PVA25 complex has influenced the transesterification reaction of soybean to FAEE as shown in the next section.

3.7.

FAEE production

Previous to the transesterification reactions, all the lyophilized enzyme-polymer complexes (enzyme-PVA4, enzymePVA12 and enzyme-PVA25) were treated with the addition of phosphate buffer, according to a procedure described by Klibanov [52]. The addition of a small amount of phosphate buffer is required to preserve the conformation of the enzyme. The transesterification reactions with the free enzyme, enzyme-PVA4, enzyme-PVA12 and enzyme-PVA25 complexes have resulted in ethyl esters yields of 20.0%, 1.7%, 40.7% and 66.3% respectively (Fig. 13). These results clearly indicate an interesting synergistic effect between the enzyme and the polymeric supports, especially for the enzyme-PVA12 and enzyme-PVA25 complexes. This effect is due to the fact the amount of ethyl ester formed by the enzyme-PVA25 complex (66.3%) and enzyme-PVA12 (40.7%) is higher than the amount formed by the free enzyme (20%). It should be noticed that the pure PVA4, PVA12 and PVA25 microspheres were not able to perform the transesterification of the vegetable oil into FAEE. This synergistic effect of the enzyme and the solid support has already been observed for zeolitic supports as reported by Vasconcellos et al. [11], however a survey of the available

Fig. 12 e Enzymatic activities of free enzyme, enzyme-PVA4, enzyme-PVA12 and enzyme-PVA25. Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

b i o m a s s a n d b i o e n e r g y x x x ( 2 0 1 3 ) 1 e1 6

13

Fig. 13 e Ethyl esters produced by free enzyme, enzyme-PVA4, enzyme-PVA12 and enzyme-PVA25.

literature dealing with enzymes immobilization onto polymeric supports aiming the production of biodiesel [11e13,22,33] will show that there is no reference to this particular synergistic effect between the enzymes and the polymeric supports.

3.8. Preliminary thermodynamic studies of the free enzyme and enzyme-PVA25 complexes Preliminary thermodynamics parameters were calculated only for the free enzyme and the enzyme-PVA25 complexes due to the fact the most significant catalytic results were obtained for this system as described in the previous sections. Determination of the thermodynamics parameters started with the measurement of the relative activity for both the free and immobilized enzyme and the results are shown in Fig. 14. The thermodynamics parameters were calculated using the Arrhenius equations discussed in Section 2.2.8 and the results are summarized in Table 1 (see Supplementary materials section). It is clear from these that no solid conclusions can be withdrawn from these data due to the weakness of the experimental statistic data. Statistically the determination of activation energies, and pre-exponential factors would require a much wider range of rates than the measured ones. Due to the fact that there is also a shift in statistical weight

when a variable is transformed to logarithmic form, the so called “Arrhenius plot” is probably not statistically valid e and no conclusions can be drawn from it. The fact the experimental data are composed of five points and in such regression the slope and intercept consume 2 of the 5 degree of freedom will cause a change in the weighting and will lead in false of accuracy [53,54], therefore we cannot compare our experimental obtained for Ea, Ed, DHoa , and DSoa with the other results that have been reported in literature [24,26,28,55e60]. In order to determine more precise and accurate thermodynamic constants for this system under investigation more detailed experiments will be performed in the future.

4. Possible explanations for the observed synergistic effect We tried to find a possible explanation for the synergistic effect observed for the enzyme-PVA25 complex in the transesterification reaction of vegetable oil into FAEE. Our interpretation was based on the ATR-FTIR and the 23Na-NMRMAS data. As stated before, the interactions involving the adsorption of proteins on solid surfaces are only possible through hydrophobic, hydrogen bonds and electrostatic

Fig. 14 e Effect of assay temperature (30 Ce60  C) on free enzyme and enzyme-PVA25 lipase activity. Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006

14

b i o m a s s a n d b i o e n e r g y x x x ( 2 0 1 3 ) 1 e1 6

interactions. Therefore, the increase of enzyme activity observed for enzyme-PVA25 complex can be attributed to the interaction of the amino acid residues of the enzyme with the hydroxyl groups present on the surface of the microspheres. From the crystallographic point of view Herrgard et al. [61] have identified a functionally important electrostatic network which includes the amino acids residues S144, D203, H257, Y260, H143, Y28, R80, and D91. This electrostatic network consists of residues belonging to the catalytic triad (S144, D203, H257), of residual amino acids located in proximity to the active site (Y260), and of residual amino acids that stabilize the geometry of the active site (Y28, H143), and of residual amino acids located in the lid (D91) or close to the first hinge (R80). According to their studies [61] the lid (D91) and the first hinge (R80) are associated with the interfacial activation of lipases, which occurs when an a-helical lid opens up by rotating around two hinge regions, while the stabilization of the enzyme in its open conformation is related to the siteesite electrostatic interaction energies between the residues R86 and D61, D113 and E117. Therefore, it is possible that the positively charged arginine (R86) will interact with the hydroxyls groups present on the surface of the PVA25 polymer through an acidebase binding mechanism as reported by Macario et al. [62] and the enzymes would be immobilized in its open and active confirmation. The ATR-FTIR data and the high yields of FAEE obtained with enzyme-PVA25 complex strongly support this affirmation. Meanwhile, it is possible that due to the fact the surface of PVA4 is dominated by acetate groups, the interaction of this polymeric support with the enzyme would occur with residual amino acids that do not stabilize the enzyme in its open and active conformation [11,61,63] and the ATR-FTIR data also support this affirmation. 23Na-MAS-NMR data for the pure PVA-25 polymer clearly indicate the presence hydrated sodium species at
2.57 ppm and aggregate sodium species at
10.44 ppm, however once the enzyme is immobilized onto the polymer surface a drastic change in the sodium environment is observed due to the complex interaction between the sodium ions and the enzyme. Daniel [56] and Vieille et al. [57] have reported that the presence sodium salt bridge can contribute to proteins stabilization. The complex 23Na-MASNMR data observed for the enzyme-PVA25 complex suggest that this type of interaction might be occurring in this system, therefore contributing for the immobilization of the enzyme in its opened and active form [56,57].

5.

Conclusions

Highly crystalline PVA microspheres were found to be suitable solid materials for enzyme immobilization, aiming the production of FAEE. The highest yield of FAEE was obtained using the enzyme-PVA25 complexes (66.3%) and a synergistic effect was observed for this enzyme-polymer complex. ATR-FTIR data indicate that the structural feature of the enzyme is preserved during the immobilization process on the high crystalline microspheres, but not for the low crystalline. 23NaMAS-NMR spectra indicate that the immobilized enzyme interacts with the sodium ions of the polymeric network, and

the presence of isolated, fully hydrated and aggregated sodium ions species were detected.

Acknowledgment This ongoing research project is financially supported by The State of Sa˜o Paulo Research Foundation (FAPESP) under Program The Young Investigator Award 05/54703-6 (J.G.N.), Award 11/10092-4 and Award 11/51851-5. We also thank CAPES and the National Council for Scientific and Technological Development (CNPq) for financial support under the Awards 07/478104-3 and 558880-2010-0.

Appendix A. Supplementary data Supplementary data related to this article can be found at http://dx.doi.org/10.1016/j.biombioe.2013.09.006.

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Please cite this article in press as: Bergamasco J, et al., Enzymatic transesterification of soybean oil with ethanol using lipases immobilized on highly crystalline PVA microspheres, Biomass and Bioenergy (2013), http://dx.doi.org/10.1016/ j.biombioe.2013.09.006