Article pubs.acs.org/Biomac
Lipase Immobilization on Differently Functionalized Vinyl-Based Amphiphilic Polymers: Influence of Phase Segregation on the Enzyme Hydrolytic Activity Mariangela Bellusci,† Iolanda Francolini,‡ Andrea Martinelli,‡ Lucio D’Ilario,‡ and Antonella Piozzi*,‡ †
Department of Chemistry and Material Technology, ENEA CR Casaccia, Via Anguillarese 301, 00123, Rome, Italy Department of Chemistry, Sapienza University of Rome, P.le Aldo Moro 5, 00185 Rome, Italy
‡
ABSTRACT: Microbial lipase from Candida rugosa was immobilized by physical adsorption onto an ethylene−vinyl alcohol polymer (EVAL) functionalized with acyl chlorides. To evaluate the influence of the reagent chain-length on the amount and activity of immobilized lipase, three differently long aliphatic fatty acids were employed (C8, C12, C18), obtaining EVAL functionalization degrees ranging from 5% to 65%. The enzyme−polymer affinity increased with both the length of the alkyl chain and the matrix hydrophobicity. In particular, the esterified polymers showed a tendency to give segregated hydrophilic and hydrophobic domains. It was observed the formation of an enzyme multilayer at both low and high protein concentrations. Desorption experiments showed that Candida rugosa lipase may be adsorbed in a closed form on the polymer hydrophilic domains and in an open, active structure on the hydrophobic ones. The best results were found for the EVAL-C18 13% matrix that showed hyperactivation with both the soluble and unsoluble substrate after enzyme desorption. In addition, this supported biocatalyst retained its activity for repetitive cycles.
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INTRODUCTION Lipases (triacylglycerol ester hydrolase, EC 3.1.1.3) are ubiquitous enzymes of considerable physiological significance and industrial potential. Indeed, they catalyze not only hydrolysis but also various reverse reactions, such as esterification, transesterification and thiotransesterification, in anhydrous organic solvents, biphasic systems, and micellar solutions, with chiral specificity.1−4 Examples of lipase applications are resolution of racemic mixtures,5,6 synthesis of new surfactants and drugs,7,8 bioconversion of oils, fats, detergent formulations,9−11 and, more recently, for biodiesel production.12,13 Enzyme immobilization onto solid carriers is a way to overcome some of the drawbacks associated with the use of enzymes such as their high cost of production and purification. In addition, enzyme immobilization improves the stability of the biocatalyst, facilitates its recovery and reuse, offers better reaction control, and increases product purity and yield.14 The main immobilization techniques are covalent binding, ionic or hydrophobic adsorption, and enzyme entrapment. The multipoint covalent attachment is considered one of the most efficacious techniques for thermal stabilization of immobilized enzymes.15,16 © 2012 American Chemical Society
Immobilization by ionic or hydrophobic interactions has the advantage of being reversible, low cost and allows the support to be reused after enzyme inactivation. In addition, the immobilized enzyme can be more active than the native one if the interactions between the biocatalyst and the support stabilize the form with higher activity.17 This is the case of the lipases that can exist in two forms: a closed form (inactive) with the catalytic site protected by the lid and an open form (active) in which the catalytic site is accessible to the substrate due to the lid displacement. In aqueous media, the equilibrium between the two forms is shifted toward the closed one. In the presence of drops of an insoluble substrate, the lipases are activated by hydrophobic interfaces (“interfacial activation”) that bring about a lipase conformational change in which a part of the enzyme molecule (lid) opens uncovering the catalytic site.18 Several types of hydrophobic matrices have been used for lipase immobilization.19−22 In fact, it has been proposed that Received: December 2, 2011 Revised: January 31, 2012 Published: February 1, 2012 805
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the interaction between the lipase and hydrophobic supports are similar to those established with an insoluble substrate. Particularly, polymers have gained significant attention due to their easy-to-tailor chemical and mechanical properties.23 Sugiura and Isobe24,25 first demonstrated that the lipase from different sources may be adsorbed onto several hydrophobic interfaces. This selective adsorption was used for lipase purification from lyophilized crude enzyme preparations.26−30 In addition, the lipases immobilized onto suitable hydrophobic supports can exhibit an enhanced activity (hyperactivation)14 also in the absence of external interfaces as they can be fixed in a hyperactivated structure.31−34 More recently, a series of different hydrophilic supports to immobilize the lipases were also experimented.35−38 However, only few studies have dealt with the issue of lipase immobilization by a systematic approach. Particularly, the use of a hydrophilic support improves the efficiency of enzyme immobilization, while the accessibility of the substrate to the lipase active site is promoted by employing hydrophobic supports. Therefore, the use of amphiphilic matrices should allow a proper hydrophobic/hydrophilic balance needed for the development of a supported biocatalyst possessing both a high hydrolytic rate and a good stability of lipase activity. In this paper, a vinyl-based amphiphilic polymer EVAL (ethylene−vinyl alcohol molar ratio 40/60), both pristine and functionalized with long alkyl chains, was employed to immobilize Candida rugosa lipase (CRL) to evaluate the effect of hydrophobic/hydrophilic balance on enzyme loading and activity. In previous studies, this polymer was differently functionalized by our group for drug adsorption39 and fumarase immobilization.40 In this work, alkyl chains at different length were introduced into EVAL by partial reaction of its hydroxy groups with capryloyl chloride (CpCl, C8), lauroyl chloride (LauCl, C12), and stearoyl chloride (StCl, C18). EVAL alkyl derivatives with different esterification degrees were obtained by varying reagent/polymer ratios. The synthesized polymers were characterized by 1H NMR spectroscopy, thermal analysis, dynamic contact angle, and water swelling measurements. Data relevant to catalytic properties of the free and immobilized enzyme were included as well as the operational and storage stability of some of the developed systems was studied.
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Scheme 1. Repetitive Unit of EVAL (a) and Reaction Scheme for the Synthesis of EVAL Derivatives
extraction with CHCl3 in a Soxhlet and finally dried under vacuum for three days at 50 °C. Polymer Characterization. 1H NMR spectra were recorded in DMF-d7 solution (Aldrich), employing a Varian XL 200 instrument. The chemical shifts are quoted in ppm, using tetramethylsilane as internal standard. The esterification degree obtained by functionalization of EVAL with acyl chlorides was determined by 1H NMR measurement.41 Differential scanning calorimetry (DSC) measurements were carried out by a Mettler TA 3000 calorimeter provided with a TC 10 A processor, by keeping the cell (DSC30) under N2 flow. The explored temperature range was −50−250 °C and the employed heating rate was 10 °C/min. The thermogravimetric analysis (TGA) was carried out employing a Mettler TG 50 thermobalance, provided with the same TC 10 A processor. The explored temperature range was 25−600 °C, and the heating rate was 10°/min, under N2 flow. The dynamic contact angle (DCA) measurements were performed by using a computerized CAHN DCA 312 dynamic contact angle analyzer. Samples were prepared by coating glass slides with DMF polymer solutions. The contact angles in advancing were measured in the immersion phase of the sample. Each measurement was made in triplicate at 25 °C employing a stage speed of 50 μm/s. The swelling experiments were performed in water by dipping polymer films at room temperature for different times. The extent of swelling Gw was calculated by the following eq 1:
EXPERIMENTAL SECTION
Gw =
Materials. Candida rugosa Type VII lipase (CRL, 724 units/mg solid) was purchased from Sigma. Tributyrin (TB, Fluka) and ethyl butyrate (EB, Aldrich) for the hydrolysis assay and the nonionic surfactant Triton X-100 (polyoxyethylene octyl phenyl ether, Fluka) were used as received. An ethylene−vinyl alcohol polymer (EVAL, DAJAC), containing ethylene and vinyl alcohol units in a 40/60 molar ratio, was employed as starting material. Its hydroxy groups were functionalized with capryloyl chloride (CpCl, Fluka), lauroyl chloride (LauCl, Fluka), or stearoyl chloride (StCl, Fluka). Dimethylformamide (DMF, Fluka) and tributylamine (TBA, Fluka) were distilled before use. All other organic solvents were analytical grade and used without further purification. Polymer Functionalization. The partial esterification reaction of EVAL hydroxy groups was carried out, under gentle stirring for 3 h at 50−60 °C in N2 atmosphere, by adding equimolar amounts of TBA and acyl chloride (CpCl, LauCl, or StCl) to a solution of 1 wt % EVAL in DMF. Different stoichiometric ratios, that is, 0.1, 0.3, 0.5, and 0.8, between mol of acyl chloride and EVAL hydroxy groups were used (Scheme 1). After the reaction, the solution was treated with a water− chloroform mixture, and the polymer was recovered at the CHCl3− H2O interface. The functionalized polymer was further purified by
G(t )‐G 0 × 100 G0
(1)
where G0 is the initial weight of the film, G is the weight of the swollen film (dried by a smooth treatment with filter paper), and t is the swelling time. Swelling kinetics was obtained by plotting Gw versus time. To evidence the surface and bulk porosity of polymer films, the samples were observed by scanning electron microscopy (SEM, LEO1450VP). In particular, for bulk measurements the films were hardened for immersion into liquid nitrogen and fractured. Then the films were fixed onto SEM supports and, after gold shading, the fractured surfaces were examined. Preparation of the Immobilized CRL. Polymer films (diameter about 2 cm, thickness about 80 μm) were obtained by deposition, on Teflon plates, of a 5 wt % solution of EVAL or EVAL derivatives in DMF, followed by solvent evaporation in a vacuum oven at 60 °C. CRL immobilization on polymer films was performed by employing different concentrations of enzyme (0.2−1 mg/mL) in 15 mL of 10 mM phosphate buffer (PBS, KH2PO4/K2HPO4, pH = 7.4). The adsorption was carried out in 50 mL flask under mild magnetic stirring (400 rpm) at room temperature for about 24 h. For EVAL-C18 13%, the adsorption was also carried out for 30 min. 806
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Enzymatic Assay. The hydrolytic activity of free CRL was assayed on a tributyrin emulsion at 37 °C in PBS. Parameters influencing the hydrolytic reaction of enzyme, including reaction time, amount of substrate, phosphate buffer, surfactant, and stirring speed, were investigated. In particular, the reaction mixture containing 225 μL of tributyrin, 525 μL of Triton X-100 and 1100 μL of CRL in PBS was incubated for 2 h under mild stirring (400 rpm). An enzyme concentration ranging from 0.05 mg/mL to 0.5 mg/mL was used. The reaction was stopped by the addition of 225 μL of a 1:1 vol % acetone−ethanol solution. The liberated butyric acid was titrated with 0.1 N NaOH solution using phenolphthalein as indicator. The adsorbed enzyme was determined by the difference between the butyric acid content produced by the used enzyme solutions, before and after the adsorption, plus that of the washing solutions, such a butyric acid content being proportional to the CRL adsorbed amount. The calibration curve necessary for determining the amount of immobilized enzyme and its efficiency was achieved employing the above-mentioned conditions, considered as optimal for the free CRL hydrolytic reaction. As far as the enzyme immobilized onto polymer films is concerned, the hydrolysis reaction was carried out in the absence of detergent because it was hypothesized that CRL can recognize the hydrophobic polymer surface as solid interface. Thus, the films were only incubated with 225 μL of tributyrin and 15 mL of PBS for 2 h under stirring (400 rpm). a total of 1 unit (U) of enzyme activity was defined as the amount of enzyme that catalyzed the production of 1 μmoL of free acid per min under the experimental conditions. The efficiency (η) of the immobilized CRL was determined as the ratio between the activity of the immobilized enzyme and that of the same amount of free enzyme in buffer solution. To investigate the nature and strength of the enzyme interaction with the matrices the hydrolytic activity of the immobilized derivatives was also assayed by using a soluble substrate, ethyl butyrate (EB), whose critical micellar concentration (CMC), determined by using CRL in solution, was 330 mM (data not shown). Moreover, desorption experiments were performed by employing Triton X-100 at 1 and 3% concentrations for 1 h. Reusability and Storage Stability. The operational stability of the immobilized enzyme was assayed by using EVAL-C18 13% in successive batches (5 cycles) performed for 1 h at 25 °C. Moreover, the storage stability of free and immobilized lipase was carried out at 25 °C by immersing free or immobilized enzyme in a PBS solution (10 mM) for 30 days. In both cases, the residual hydrolytic activity was determined by using the experimental conditions described in the previous section.
Table 1. Data Relevant to the Synthesized Polymers polymer
molar ratioa reagent/OH
integral ratiob OH/CH
Tg (°C)c
EVAL EVAL-C8 20% EVAL-C8 30% EVAL-C8 65% EVAL-C12 12% EVAL-C12 18% EVAL-C12 35% EVAL-C18 5% EVAL-C18 13% EVAL-C18 41%
0.3 0.5 0.8 0.3 0.5 0.8 0.1 0.3 0.5
1 0.80 0.70 0.35 0.88 0.82 0.65 0.95 0.87 0.59
60 44 35 30 40 25 20 51 50 48
θadv (°)d 79 80 80 90 83 82 92 89 96 99
± ± ± ± ± ± ± ± ± ±
1 2 1 1 2 1 1 1 1 1
a
Ratios between mol of acyl chloride and equivalent of EVAL hydroxy groups. bIntegral ratios calculated from 1H NMR data. cGlass transition temperature determined in second scan. dAdvancing contact angles measured in water.
increase of the solubility of the esterified polymer in the reaction medium. This higher solubility with respect to the unreacted EVAL presumably favored the progress of the reaction between polymer hydroxy groups and StCl. DSC and TGA Data. Amphiphilic polymers can exhibit phase segregation due to the different chemical nature of hydrophilic and hydrophobic segments. Although there can be some degree of mixing between these segments, a phase separation, responsible for their physical and mechanical properties, occurs. The extent of this phenomenon depends on the presence of functional groups or side chains in the polymer structure. The thermal transitions, particularly the glass transition temperature (Tg), are affected by possible interactions between hydrophilic and hydrophobic segments. Indeed, if phase mixing occurs, the mobility of the segments decreases with a consequent increase of Tg. In Figure 1, as an example, DSC curves of EVAL-C18 polymers compared to EVAL are reported. As for EVAL, the split
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RESULTS AND DISCUSSION H NMR Data. 1H NMR measurements were used to determine the esterification degree of the polymers. In particular, we considered the ratio between the value of the integral relevant to the OH groups and that corresponding to the methylene groups, as previously reported.41 It was checked that this ratio was equal to 1 for the unreacted EVAL. In Table 1, the data obtained at different alkyl chain content are reported. When the ratios between mol of acyl chloride and equivalents of EVAL hydroxy groups were of 0.3, 0.5, and 0.8, the esterification degrees were 20, 30, and 65% for capryloyl chloride (C8 chain length) and 12, 18, and 35% for lauroyl chloride (C12 chain length). Therefore, the chain length plays a crucial role in the esterification reaction causing a higher yield for the C8 alkyl chain-containing polymer. As for stearoyl chloride (C18 chain length), we employed reagent/OH molar ratios of 0.1, 0.3, and 0.5, achieving esterification degrees of 5, 13, and 41%, respectively. The surprisingly high esterification degree obtained with the 0.5 molar ratio was explained by the 1
Figure 1. DSC thermograms of EVAL-C18 polymers (the EVAL thermogram, for the sake of clarity, is also reported).
melt peak in the 160−180 °C range was attributed to the presence of crystalline regions due to OH−OH interactions. In addition, at about 60 °C, a flex attributable to the glass transition temperature (Tg) was observed. In the thermograms of the 807
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cycle, the advancing angle decreased by about 4° for EVAL and 7° both for EVAL-C8 65% and EVAL-C12 35% (DCA not shown). These findings indicate that EVAL, being partially hydrophilic, was able to expose its hydroxyl groups after contact with water. Instead, the increase in surface wettability observed in the other two samples was due to the higher amorphous content of these polymers favoring the rearrangements of the alkyl chains. Water swelling measurements showed that all samples possessed considerable bulk hydrophobicity (Figure 2). By fitting
esterified polymers, a decrease both of the melting enthalpy and temperature besides an endothermic transition in the temperature range of 60−80 °C was observed. Similar results were found for EVAL-C8 and EVAL-C12 (data not shown). The melting temperature of polymers decreased progressively with increasing of the functionalization degree. This can be explained by the reduction of hydrogen bond interactions between the polymer hydroxy groups. The disappearance of the melting peak at high esterification degree observed for EVALC8 and EVAL-C12 indicates a greater amorphous character of these samples (DSC not shown) than EVAL-C18. Moreover, the transition at lower temperature was attributed to the melting of the semicrystalline domains due to van der Waals interactions between the alkyl chains. Indeed, the esterified polymers, particularly EVAL-C18, are inclined to give segregated hydrophilic and hydrophobic domains consisting of alcoholic and alkyl chain regions. Therefore, the extent of these domains depends on both the length of the introduced alkyl chains and the esterification degree. Because, in the first DSC scan, the glass transition was not clearly observed, the samples were subjected to a second heating. In this case, the disappearance of the lower temperature transition allowed the Tg determination, whose values are reported in table 1. As for EVAL-C8 and EVAL-C12, the significant Tg decreasing with the increase of the functionalization degree can be explained in terms of a plasticizing effect exerted by the alkyl chains. This finding can be related to a minor phase mixing of these esterified polymers with respect to EVAL. Thermogravimetric analysis was carried out to investigate the thermal stability of the different matrices (data not shown). For all the functionalized polymers, a loss in weight in the 220− 370 °C range was observed and attributed to the acid release. Moreover, experimental data showed that the degradation onset temperature decreased with the increase of the esterification degree of the polymers. DCA and Swelling Data. Contact angle, θ, is a quantitative measure of the wetting of a solid by a liquid. It is defined geometrically as the angle formed by a liquid at the three phase boundary where a liquid, gas, and solid intersect. A low value of contact angle (θ) indicates that the liquid wets well, while a high contact angle indicates a poor wetting of the solid. If the angle θ is less than 90°, the liquid is said to wet the solid. If it is greater than 90°, it is said to be nonwetting. A zero contact angle represents complete wetting. If the three phase (liquid/ solid/vapor) boundary is in actual motion, the angles produced are called dynamic contact angles and are referred to as “advancing” and “receding” angles. The advancing contact angle is obtained from data generated as the solid advances into the liquid. The sample is immersed to a set depth, and the process is reversed. As the solid retreats from the liquid, the data collected are used to calculate the receding contact angle. Therefore, to evaluate the surface hydrophobicity of the samples, measurements of dynamic contact angle were performed. The data related to the advancing contact angle are reported in Table 1. As for EVAL-C8 and EVAL-C12, it may be noted that only at the highest esterification degrees (65 and 35%, respectively) the wettability is poor. On the contrary, for the EVAL-C18 polymers, the wettability was poor already at low content of alkyl chains. The highest value of contact angle was found for EVAL-C18 41%. To evaluate a possible effect due to kinetic hysteresis, two immersion cycles of samples were performed. In the second
Figure 2. Kinetics curves relevant to the swelling in water of polymers: EVAL-C8 (a), EVAL-C12 (b), and EVAL-C18 (c). The EVAL curve is reported in all figures for the sake of clarity.
the experimental data versus the root square of time, a different behavior was found for the EVAL-C18 derivatives. 808
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than the limit of the BET analysis (Brunauer, Emmett and Teller analysis), approximately of 1 m2. Therefore, to evidence the bulk and surface porosity, polymer samples were observed by SEM. As can be noted in Figure 7, EVAL showed a good porosity both in surface and bulk (pore size roughly 400 nm). On the contrary, EVAL derivatives possessed a low porosity. Indeed, both the film surface and fractured side of EVAL-C18 13% (reported as an example in Figure 7) were rather smooth also when observed at high magnification (Figure 7c). Particularly, the higher polymer hydrophobicity the smoother the surface. Taking into account that the surface offered by a not porous polymer film with a diameter of 2 cm is about 6.3 × 10−4 m2, we can hypothesize that, given the high enzyme loaded (Figure 4), a multilayer adsorption occurred particularly for EVAL derivatives that possessed a poor porosity. Instead for EVAL, the penetration of the enzyme into support pores can better explain the high enzyme loading. As for the hydrolytic activity vs tributyrin, the low efficiency found for EVAL is then probably due to diffusion limitations of the heterogeneous substrate into the film pores. To verify this hypothesis, a soluble substrate (ethyl butyrate, EB) was employed. By using EB at concentration lower than CMC (40 mM), the EVAL matrix containing 6.9 mgCRL/gpolymer anyway showed a poor hydrolytic activity (η < 3%), evidencing that, besides mass transfer limitations, also an immobilization of CRL aggregates occurred. Instead, the efficiency significantly increased up to η ≅ 50% when an ethyl butyrate concentration of 400 mM (concentration above CMC) was employed. This improvement in catalytic activity with respect to that found with tributyrin (η ≅ 10%) was probably due to a partition effect of the soluble substrate that brings about an increase of its concentration in the proximity of the support (where the environment is hydrophobic) and the activation of the adsorbed enzyme. As for EVAL derivatives, it is worth noticing that EVALC8 65% and EVAL-C12 35% achieved the saturation loading (Figure 4) already at low enzyme concentration showing, however, a poor hydrolytic activity (Figure 6). This behavior could be attributed to the higher swelling ability of these polymers that causes the flattening on the polymer surface of the alkyl chains not compacted in hydrophobic domains (see DCA discussion). Therefore, in presence of segregated matrices possessing a reduced extension of hydrophobic and hydrophilic domains that can rearrange in an aqueous environment, the enzyme probably maximizes its contact with the surface, establishing interactions that bring about enzyme conformational changes and loss of catalytic activity. Otherwise, the low CRL activity found for the polymers showing a moderate but steady level of phase segregation (the other EVAL-C8 and EVAL-C12 matrices) could be due to a different adsorption of the enzyme: a fraction interfacially bonded on the hydrophobic domains and another fraction in an aggregate fashion on the hydrophilic ones. This latter fraction, probably adsorbed as a multilayer, depressed the activity of the CRL interfacially bonded. As for EVAL-C18 polymers, EVAL-C18 13% that displayed a remarkable bulk and surface hydrophobicity as well as a good phase segregation (Figure 1) showed the best hydrolytic activity at low enzyme loading (Figure 6c). In this case, the formation of large hydrophobic domains allowed the enzyme to establish interactions with the matrix involving the hydrophobic area surrounding the active site and the internal face of the lid.
Indeed, notwithstanding, all polymers showed a Fickian mechanism for water uptake (case I diffusion mechanism), and different diffusion coefficients were determined. In fact, average values ranged from 1 × 10−5 to 3 × 10−5 mm2/min for the EVAL-C8 and EVAL-C12 systems, whereas a water diffusion coefficient of 1 order of magnitude lower was found for the EVAL-C18 matrices (average value of 2 × 10−6 mm2/min). This finding could be attributed to the lower amorphous content of the EVAL-C18 derivates (Figure 1), which limits water uptake. Enzymatic Assay of the Immobilized Lipase. The effect of Triton X-100 on the free CRL activity, reported in Figure 3,
Figure 3. Effect of the Triton X-100 concentration on the hydrolytic activity of the free lipase. The experimental conditions were 225 μL of tributyrin and 1100 μL of CRL (2 mg/mL) in PBS, 2 h incubation time, and 400 rpm stirring.
showed that ∼200 mg/mL surfactant is required to minimize the interfacial tension between the substrate and the enzyme. In Figure 4, the data relevant to CRL immobilization onto EVAL and its derivatives are reported. It can be observed that the loaded amount increased with the increase of the enzyme concentration while the hydrolytic activity profile, reported in figure 5 and 6, was inversely proportional to the amount of bonded lipase. Bosley and Peilow42 studied the influence of enzyme loading on the immobilization of several lipases on porous polypropylene. The experimental data showed that the efficiency fell as the loading increased. The authors hypothesized that at high protein loading the systems suffered from mass transfer limitations which hindered the diffusion of the heterogeneous substrate. Similar results were obtained by de Oliveira et al.22 using CRL immobilized onto a styrene−divinylbenzene copolymer. Also, O’Connell and Varley43 hypothesized that at high protein concentration CRL could be adsorbed in the form of a multilayer on colloid gas aphrons. The same hypothesis could be advanced by us. Indeed, by assuming the geometry of the protein as a sphere (⌀ = 7 nm) and taking into account an average molecular weight of about 60 KDa,44 it can be roughly calculated that the minimum amount of CRL bound in our experiments would require about a 0.040 m2 polymer surface. Unfortunately, it was not possible to evaluate the real film surface area because its value was lower 809
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Figure 4. Effect of CRL concentration on the enzyme loading for EVAL (A), EVAL-C8 (B), EVAL-C12 (C), and EVAL-C18 (D).
The greater CRL affinity for the EVAL-C18 13% matrix can also be evidenced by plotting the efficiency vs the length of the alkyl chain, under similar amount of adsorbed enzyme and functionalization degree (Figure 8). As expected, the enzyme efficiency increased with the alkyl chain length. These results are in agreement with findings of Shaw et al.20 who studied the effect of chain length on the lipase-coupling yield for different materials. The authors showed that an alkyl chain longer than six carbons favored the immobilization of active enzyme. Besides the enzyme conformation (closed or open form), the matrix hydrophobicity may also effect the type of adsorbed isoenzyme. Indeed, the commercial CRL used in this study is a crude product constituted by several CRL isoenzymes and other contaminant enzymes. Among the isoenzymes, the two main fractions are Lip1 and Lip3, as reported in the literature.45,46 It was evidenced that, differently from Lip1, Lip3 can exist in solution both in monomeric and dimeric form. The ability of Lip3 to self-associated is due to both the higher flexibility of the
Figure 5. Effect of lipase loading on efficiency (hydrolytic activity/ loading) for lipase immobilized onto EVAL.
Therefore, CRL results mainly immobilized in an open structure at least in the first layers close to the surface. 810
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Figure 7. SEM micrographs of the EVAL (A) and EVAL-C18 13% (B) fractured films. (C) EVAL-C18 13% surface at higher magnification.
Figure 6. Effect of lipase loading on efficiency (hydrolytic activity/ loading) for lipase immobilized onto EVAL-C8 (a), EVAL-C12 (b), and EVAL-C18 (c).
lid and the greater hydrophobicity of the catalytic site.47,48 This dimeric form possesses a good affinity for soluble substrates also below their CMC, suggesting the presence of an accessible active site. The Lip1 and Lip3 isoforms are catalytically different in solution.45 Indeed, it was found that the transition between closed and open conformations is slower for Lip1 than Lip3. This finding determines a difference in the minimum concentrations of soluble substrate needed to obtain the interfacial activation of the two isoenzymes. In particular, the Lip3 closed−open equilibrium is shifted toward the open form at lower soluble substrate concentration than Lip1. Starting from these considerations, the polymer surface could also have a different affinity for the isoenzymes (chromatographic effect). This could bring about the obtainment of a
Figure 8. Effect of the alkyl chain length on efficiency of lipase immobilized onto the functionalized polymers. The comparison between matrices was performed under similar amount of both adsorbed lipase and esterification degree (EVAL-C8 20%, EVAL-C12 12%, and EVAL-C18 13%). 811
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could be related to the presence of greater Lip3 fraction in monomer and dimer form. Finally, in an attempt to load a very small enzyme amount, the CRL adsorption on the EVAL-C18 13% was also performed for 30 min. The adsorbed enzyme, corresponding to 0.7 mgCRL/gpolymer, showed hyperactivation also toward tributyrin (4-fold increase in catalytic activity). Reusability and Storage Stability. The operational stability of the immobilized CRL was evaluated in a repeated batch process for the most active matrix after desorption experiments. Figure 10 shows the effect of repeated use on the
supported biocatalyst with a catalytic activity related to the Lip1/Lip3 ratio present in the adsorbed lipase multilayer. Indeed, the higher efficiency value found in EVAL-C18 13% with respect to EVAL C-18 5% could be explained by the formation of larger hydrophobic domains that probably promote a higher adsorption of the most active CRL isoenzyme determining a replica effect in the above layers up to the outer one. It is worth noticing that both the EVAL-C18 5% and EVAL-C18 13% matrices adsorbed the same CRL amount and showed similar matrix phase segregation. Recently, Lorente and co-workers48 showed that Pseudomonas fluorescens lipase aggregates can be dissociated by detergent addition. To evidence the adsorption of different isoenzymes onto polymer surfaces, desorption experiments of CRL adsorbed on the EVAL, EVAL-C18 5% and EVAL-C18 13% matrices were performed by using Triton X-100 at 1 and 3% (w/w) concentrations. At 1% detergent concentration, an enzyme release of about 70% for EVAL and 80% for the other two matrices was observed, reaching in all cases a value of adsorbed enzyme of approximately 2 mgCRL/gpolymer corresponding to just a few enzyme layers, whereas, for the higher detergent concentration (3%), an enzyme complete release and its inactivation were obtained. By testing the efficiency of the matrices treated with 1% Triton X-100 toward EB at a concentration lower than CMC, it was found that only EVAL-C18 polymers showed hyperactivation toward this soluble substrate (from 4- to 20-fold increase in catalytic activity for EVAL-C18 5% and EVAL-C18 13%, respectively). This finding suggests the adsorption of CRL in an open conformation particularly for EVAL-C18 13% having larger in size hydrophobic domains, as previously hypothesized. As for the tributyrin test, a lower efficiency with respect to the one determined before desorption experiments was observed for both matrices (Figure 9). This demonstrates that, although a
Figure 10. Effect of repeated use on the activity of the lipase immobilized on EVAL-C18 13%. Cycles were performed for 1 h at 25 °C.
activity of the enzyme immobilized on EVAL-C18 13%. Immobilized biocatalyst, under the adopted experimental conditions, retained an almost complete activity for at least five cycles. Also, for the storage stability of free and immobilized CRL in PBS solution, polymer-immobilized enzyme did not exhibit any loss in activity up to 30 days at 25 °C (data not shown).
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CONCLUSION Microbial lipase from Candida rugosa was immobilized by physical adsorption onto an ethylene−vinyl alcohol polymer (EVAL) functionalized with acyl chlorides of different chain lengths (C8, C12, C18). It was observed that the chain length plays a crucial role in determining the esterification yield that resulted in more elevated yields for capryloyl chloride with respect to the other two acyl chlorides. The esterified polymers showed a good tendency to give phase segregation as a consequence of the presence of the alkyl chains. Results showed that the enzyme−polymer affinity increased both with the length of the alkyl chain and the matrix hydrophobicity. Instead, the enzyme efficiency (η) toward tributyrin decreased with increasing amounts of the bonded enzyme. In the presence of matrices possessing hydrophilic (EVAL) and hydrophobic domains that can rearrange in an aqueous environment (i.e., EVAL-C8 65% and EVAL-C12 35% derivatives), the enzyme is mainly adsorbed by means of protein aggregates and in closed form. Otherwise, the polymers showing a moderate but steady level of phase segregation (the other EVAL-C8 and EVAL-C12 matrices) adsorbed CRL in active form on the hydrophobic domains and in aggregate fashion on the hydrophilic ones. On the contrary, the formation of large hydrophobic domains allows the interfacial activation of enzyme promoting the lid opening. In fact, EVAL-C18 13% matrix showed a
Figure 9. Polymer efficiency toward tributyrin and ethyl butyrate after desorption experiments with 1% Triton X-100.
remarkable reduction of enzyme layers by desorption experiments was obtained, diffusion limitations still exist probably due to a moderate film porosity. In addition, considering that, after desorption, the two EVAL-C18 polymers possessed the same amount of adsorbed CRL, the higher activity of EVAL-C18 13% toward tributyrin 812
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Biomacromolecules
Article
(22) De Oliveira, P. C.; Alves, G. M.; De Castro, H. F. Biochem. Eng. J. 2000, 5, 63−71. (23) Fang, Y.; Huang, X. J.; Chen, P. C.; Xu, Z. K. BMB Rep. 2011, 44, 87−95. (24) Sugiura, M.; Isobe, M. Chem. Pharm. Bull. (Tokyo) 1976, 24, 72−78. (25) Sugiura, M.; Isobe, M. Biochim. Biophys. Acta 1975, 397, 412− 417. (26) Montero, S.; Blanco, A.; Virto, M. D.; Landeta, L. C.; Agud, I.; Solozabal, R.; Lascaray, J. M.; de Renobales, M.; Llama, M. J.; Serra, J. L. Enzyme Microb. Technol. 1993, 15, 239−247. (27) Rùa, M. L.; Ballesteros, A. Biotechnol. Tech. 1994, 8, 21−26. (28) Bornscheuer, U.; Reif, O. W.; Lausch, R.; Freitag, R.; Scheper, T.; Kolisis, F. N.; Menge, U. Biochim. Biophys. Acta 1994, 1201, 55−60. (29) Sarda, L.; Desnuelle, P. Biochim. Biophys. Acta 1958, 30, 513− 521. (30) Isobe, M.; Sugiura, M. Chem. Pharm. Bull. (Tokyo) 1977, 25, 1987−1991. (31) Reetz, M. T.; Zonta, A.; Simpelkamp, J. Biotechnol. Bioeng. 1996, 49, 527−534. (32) Reetz, M. T. Adv. Mater. 1997, 9, 943−954. (33) Bastida, A.; Sabuquillo, P.; Armisen, P.; Fernandez-Lafuente, R.; Huguet, J.; Guisan, J. M. Biotechnol. Bioeng. 1998, 58, 486−493. (34) Fernandez-Lafuente, R.; Armisen, P.; Sabuquillo, P.; FernandezLorente, G.; Guisan, J. M. Chem. Phys. Lipids 1998, 93, 185−197. (35) Miletic, N.; Vukovic, Z.; Nastasovic, A.; Loos, K. J. Mol. Catal. B: Enzyme 2009, 56, 196−201. (36) Buthe, A. Methods Mol. Biol. 2011, 743, 223−237. (37) Kosaka, P. M.; Kawano, Y.; El Seoud, O. A.; Petri, D. F. Langmuir 2007, 23, 12167−12173. (38) Ren, Y.; Rivera, J. G.; He, L.; Kulkarni, H.; Lee, D. K.; Messersmith, P. B. BMC. Biotechnol. 2011, 11, 63. (39) Marconi, W.; Monopoli, G.; Piozzi, A.; Di Rosa, R. J. Bioact. Compat. Polym. 2001, 6, 294−306. (40) Marconi, W.; Faiola, F.; Piozzi, A. J. Mol. Catal. B: Enzyme 2001, 15, 93−99. (41) Marconi, W.; Cordelli, S.; Napoli, A.; Piozzi, A. Macromol. Chem. Phys. 1999, 200, 1191−1199. (42) Bosley, J. A.; Peilow, A. D. J. Am. Oil Chem. Soc. 1997, 74, 107− 111. (43) O’Connell, P. J.; Varley, J. Biotechnol. Bioeng. 2001, 74, 264− 269. (44) Kazlauskas, R. J. Trends Biotechnol. 1994, 12, 464−472. (45) Pernas, M. A.; Lopez, C.; Prada, A.; Hermoso, J.; Rùa, M. L. Colloids Surf., B 2002, 26, 67−74. (46) Pernas, M. A.; Lopez, C.; Rùa, M. L.; Hermoso, J. FEBS Lett. 2001, 501, 87−91. (47) Ghosh, D.; Wawrzak, Z.; Pletnev, V. Z.; Li, N.; Kaiser, R.; Pangborn, W.; Jornvall, H.; Erman, M.; Duax, W. L. Structure 1995, 3, 279−288. (48) Fernandez-Lorente, G.; Palomo, J. M.; Fuentes, M.; Mateo, C.; Guisan, J. M.; Fernandez-Lafuente, R. Biotechnol. Bioeng. 2003, 82, 232−237.
good catalytic activity with the insoluble substrate and hyperactivation with the soluble one. This study demonstrated for the first time that not only the matrix hydrophobicity, but also its hydrophilic features need to be taken into consideration to develop polymer supports suitable for CRL immobilization. In fact, only the amphiphilic polymers possessing a proper hydrophobic/hydrophilic balance as well as a good and steady phase segregation are able to adsorb CRL in active form.
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AUTHOR INFORMATION
Corresponding Author
*Fax: (+39)-06-49913692. E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS The authors are grateful to University “La Sapienza” (Ateneo Funds) and Italian Ministry of Education, University and Research for financial support.
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REFERENCES
(1) Brink, L. E. S.; Tramper, J.; Luyben, K.; Ch., A. M.; Van’t Riet, K. Enzyme Microb. Technol. 1988, 10, 736−743. (2) de Zeote, M. C.; Kock-van Dalen, A. C.; van Rantwijk, F.; Sheldon, R. A. J. Mol. Catal. B: Enzyme 1996, 1, 109−113. (3) Gupta, M. N. Eur. J. Biochem. 1992, 203, 25−32. (4) Martinek, K.; Levashov, A. V.; Klyachko, N.; Khmelnitski, Y. L.; Berezin, I. V. Eur. J. Biochem. 1986, 155, 453−468. (5) Jaeger, K. E.; Eggert, T. Curr. Opin. Biotechnol. 2004, 15, 305− 313. (6) Ikeda, I.; Klibanov, A. M. Biotechnol. Bioeng. 1993, 42, 788−791. (7) Kuboki, A.; Ishihara, T.; Kobayashi, E.; Ohta, H.; Ishii, T.; Inoue, A.; Mitsuda, S.; Miyazaki, T.; Kajihara, Y.; Sugai, T. Biosci. Biotechnol. Biochem. 2000, 64, 363−368. (8) Martin, B. D.; Linhardt, R. J.; Dordick, J. S. Biomaterials 1998, 19, 69−76. (9) Jaeger, K. E.; Ransac, S.; Dijkstra, B. W.; Colson, C.; van, H. M.; Misset, O. FEMS Microbiol. Rev. 1994, 15, 29−63. (10) Balcao, V. M.; Paiva, A. L.; Malcata, F. X. Enzyme Microb. Technol. 1996, 18, 392−416. (11) Villeneuve, P. Eur. J. Lipid Sci. Technol. 2003, 105, 308−317. (12) De Diego, T.; Manjon, A.; Lozano, P.; Iborra, J. L. Bioresour. Technol. 2011, 102, 6336−6339. (13) Wang, L.; Du, W.; Liu, D.; Li, L.; Dai, N. J. Mol. Catal. B: Enzyme 2006, 43, 29−32. (14) Mateo, C.; Palomo, J. M.; Fernandez-Lorente, G.; Guisan, J. M.; Fernandez-Lafuente, R. Enzyme Microb. Technol. 2007, 40, 1451− 1463. (15) Betancor, L.; Luckarift, H. R. Trends Biotechnol. 2008, 26, 566− 572. (16) Hernandez, K.; Fernandez-Lafuente, R. Enzyme Microb. Technol. 2011, 48, 107−122. (17) Fernandez-Lorente, G.; Fernandez-Lafuente, R.; Palomo, J. M.; Mateo, C.; Bastida, A.; Coca, J.; Haramboure, T.; Hernandez-Justiz, O.; Terreni, M.; Guisan, J. M. J. Mol. Catal. B: Enzyme 2001, 11, 649− 656. (18) Brzozowski, A. M.; Derewenda, U.; Derewenda, Z. S.; Dodson, G. G.; Lawson, D. M.; Turkenburg, J. P.; Bjorkling, F.; Huge-Jensen, B.; Patrk, S. A.; Thim, L. . Nature 1991, 351, 491−494. (19) Bosley, J. Biochem. Soc. Trans. 1997, 25, 174−178. (20) Shaw, J. F.; Chang, R. C.; Wang, F. F.; Wang, Y. J. Biotechnol. Bioeng. 1990, 35, 132−137. (21) Palomo, J. M.; Munoz, G.; Fernandez-Lorente, G.; Mateo, C.; Fernandez-Lafuente, R.; Guisan, J. M. J. Mol. Catal. B: Enzyme 2002, 19−20, 279−286. 813
dx.doi.org/10.1021/bm2017228 | Biomacromolecules 2012, 13, 805−813
