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Effect of support surface chemistry on lipase adsorption and activity
Journal of Molecular Catalysis B: Enzymatic 94 (2013) 69–76
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Journal of Molecular Catalysis B: Enzymatic journal homepage: www.elsevier.com/locate/molcatb
Effect of support surface chemistry on lipase adsorption and activity Ye Peng ∗ , Han Zhu-Ping, Xu Yong-Juan, Hu Peng-Cheng, Tong Ji-Jun Key Laboratory of Advanced Textile Materials and Manufacturing Technology of Education Ministry, Department of Chemistry, Zhejiang Sci-Tech University, Hangzhou 310018, PR China
a r t i c l e
i n f o
Article history: Received 7 January 2013 Received in revised form 30 March 2013 Accepted 26 April 2013 Available online 4 May 2013 Keywords: Support surface chemistry Lipase Enzyme immobilization Functional group-ended polymer
a b s t r a c t The functional group controlled surfaces (S-F, functional groups including NH2 , COOH, OH, CH3 and CF3 ) were fabricated by mixing polystyrene (PS) respectively with functional group-ended polystyrenes (PS-Fs). These surfaces were used as model supports to investigate the effect of surface chemistry on lipase adsorption and activity. The order of the amount of adsorbed lipase on the surface with similar functional group density was S-CH3 > PS > S-CF3 > S-NH2 > S-COOH > S-OH. It could be found that on the surface containing hydrophobic group, lipase could take more side-on orientations with larger spreading, while on the surface containing hydrophilic group, lipase could take more end-on orientations with smaller spreading. Lipase immobilized on the surface containing OH showed the highest activity. The adsorption of substrates and products on the functional group controlled surfaces was also measured. The surface containing NH2 showed a higher activity which might be ascribed to the fact that there was a high substrate concentration on the lipase/support interface. The order of the activity retention of the immobilized lipase was S-OH > S-NH2 > S-COOH > S-CF3 > S-CH3 > PS. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Lipases, as versatile biocatalysts, have gained considerable importance and the potential applications of lipases in biotransformations have been fully documented with a plethora of examples, on both hydrolyzation in aqueous media and synthesis in organic media [1–4]. Due to an opposite polarity between the enzyme (hydrophilic) and their substrate (lipophilic), lipase reaction occurs at the interface between the aqueous and the oil phase. Hence, interfaces are the key spots for lipase biocatalysis [5–8]. There are three types of interfaces: air–water, oil–water and solid–water, which could activate lipase [9]. The solid–water interface provides the possibility to immobilize enzyme and to tailor a surface with which the lipase interacts. In fact, enzymes have always been immobilized onto insoluble or solid supports for industrial applications, which are regarded as a useful tool to enhance their thermal and operational stabilities, and reduce the cost [10–13]. Special emphasis has been attached to the selective adsorption of lipases on tailor-made strongly hydrophobic support surfaces [14–17]. This immobilization procedure is based on the assumption that the large hydrophobic area that surrounds the active site of lipases is the one mainly involved in their adsorption on strongly hydrophobic solid surfaces. However, some
∗ Corresponding author. Tel.: +86 571 86843691; fax: +86 571 86843653. E-mail address: [email protected] (P. Ye). 1381-1177/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.molcatb.2013.04.015
authors reported that lipase could also retain higher activity on the hydrophilic support [9,18]. In lipase catalysis interface composition, which is described by the term “interface quality” [19–23], plays a vital role. There are at least five kinds of materials on the interface, including lipase, support, substrate, product and solution. Recent studies demonstrate that lipase activity as a function of interface composition is more attributed to substrate inaccessibility rather than to enzyme denaturation or inactivation, as it is often hypothesized. Holmberg and co-workers measured the adsorption of two substrates capric acid (decanoic acid) and monocaprin on the lipase-covered surface, and found that lipase adsorbed at the hydrophilic surface favored hydrolysis and capric acid was the main product when monocaprin was used as substrate [9]. However, knowledge about the effect of the interface composition in lipase catalysis is still limited and complete understanding of enzyme–substrate or product–substrate interaction is necessary. To control support surface properties, the modification of polymer end groups has emerged as a practical means [24–26]. In this study, three hydrophilic groups ( NH2 , COOH and OH) and two hydrophobic groups ( CH3 and CF3 ) were quantitatively introduced onto the support surfaces by mixing PS solution with PS-Fs, respectively. Lipase (from Candida rugosa) [27], was immobilized on the supports as a model enzyme. The adsorption of two substrates (p-nitrophenyl palmitate (p-NPP) and olive oil) and three products (palmitic acid (PA), glycerol and p-nitrophenol (p-NP)) was also measured. The effects of the support surface chemistry on the adsorption and activity of lipase were investigated.
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PS-COOH + NH2-CH2CH2-F DCC PS-CONH-CH2CH2-F Scheme 1. Synthesis of functional group-ended polystyrene (PS-F, functional groups including NH2 , OH, CH3 and CF3 ).
PS-CONH-CH2CH2-F + H2O HCL
PS-COOH + NH2-CH2CH2-F
Scheme 2. Hydrolyzation of functional group-ended polystyrene (PS-F, functional groups including NH2 , OH, CH3 and CF3 ) to carboxyl-ended polystyrene (PSCOOH) in HCL solution.
2. Materials and methods 2.1. Materials Lipase (from C. rugosa), Bradford reagent, bovine serum albumin (BSA) and p-nitrophenyl palmitate (p-NPP) were purchased from Sigma and used as received. Polystyrene (PS, Mn 92,100, PDI 2.6) was purchased from Aldrich Co. All the other chemicals were of analytical grade and used without further purification. 2.2. Synthesis of the functional group-ended polystyrenes Carboxyl-ended polystyrene (PS-COOH, Mn 4400, PDI 1.2) was prepared as previously reported [28]. Hydroxyl-ended polystyrene (PS-OH), trifluoromethyl-ended polystyrene (PS-CF3 ), methyl-ended polystyrene (PS-CH3 ) and amino-ended polystyrene (PS-NH2 ) were synthesized by the reaction between PS-COOH with hydroxyl-ethane-amine, trifluoromethyl-ethane-amine, methylethane-amine or ethylenediamine in the presence of dicyclocarbondiamide (DCC) in dehydrated tetrahydrofuran (THF) solution in a three-neck round-bottom flask equipped with a stopcock and a magnetic stirring bar, respectively. Prior to use, the flask was vacuumed and backfilled with dry nitrogen several times. The reaction mixture was then kept at room temperature for 48 h under a nitrogen atmosphere. After reaction, the mixture was filtered to remove 1,3-dicyclohexylurea and then precipitated into water, and the resulting product was dried in vacuum for 24 h. The product was then dissolved into THF, and the resulting solution was passed through an alumina column to remove the impurity (Scheme 1). 2.3. Preparation of the functional group controlled surfaces Prescribed amounts of PS were dissolved in xylene at a concentration of 5 wt% for the preparation of casting solutions. The functional group controlled surfaces were prepared by mixing PS solution (95 wt%) with functional groups-ended polystyrenes solution (5 wt%) (PS-F, functional groups such as CH3 , CF3 , NH2 , COOH and OH), then casting the solution (2 mL) on rimmed glass plates (7.6 cm × 2.6 cm) and drying at 25 ◦ C in dried air, finally drying at 150 ◦ C for 10 h, respectively. Each support taken off the glass plate had two surfaces that were formed in different environments. One surface of the support contacting with the dried air was designated as the air-side surface, and the other surface contacting with the glass plate was designated as the glass-side surface. The surface density of functional group might be different on the two side surfaces. In order to eliminate the possible error owing to this difference, the supports were poured with pure PS solution (0.2 mL) on their one side surfaces, the air-side surface for the support containing COOH, NH2 or OH, and the glass-side surface for the support containing CH3 or CF3 , and dry them again to form one side of pure PS surfaces. 2.4. Characterization of the functional group controlled surfaces X-ray photoelectron spectroscopy (XPS) experiments were performed on a PHI-5000C ESCA system (Perkin–Elmer) with Al K␣ radiation (h = 1486.6 eV). Contact angles of water on the support surfaces were measured by the Sessile drop method using a DSA10 drop sharp analysis (Kruss Co., Germany) at 25 ◦ C. The volume of the water drops used was always 3 L. All the reported values were averages of at least eight measurements taken at different
locations on the support surface and had a typical error of the mean of ±1◦ . The surface free energy was calculated according Owens and Wendt’s theory from the measured contact angles of water and diiodomethane on the samples [29]. The surface densities of COOH were obtained by measuring the load and release of methylene blue (MB) [30]. In order to measure the surface densities of functional group, such as CH3 , NH2 , OH and CF3 , the amide bond of PS-Fs were hydrolyzed to carboxyl group in HCL solution (20 wt%) at 50 ◦ C for 24 h, respectively (Scheme 2). 2.5. Lipase immobilization The support (about 10 cm × 10 cm) was cut into 5 mm × 5 mm and thoroughly washed with deionized water, and then rinsed with phosphate buffered solution (PBS) (25 mM, pH 7.0). After that, the support was submerged in 10 mL lipase solution (0.5 mg/mL) at room temperature in a shaking water bath for 8 h. Finally, the supports were removed, thoroughly rinsed with PBS and then rinsed with de-ionized water. The amount of adsorbed protein on the support was determined by measuring the initial and final concentrations of protein within the protein solutions and washings using Coomassie Brilliant Blue reagent, following Bradford’s method. As mention above, the support had two surfaces, one was the functional group controlled surface and the other was the pure PS surface. The value of the amount of adsorbed protein on the functional group controlled surface was calculated by subtracting the amount of adsorbed protein on the pure PS surface of the same size in the same experiment conditions. The surface morphology of these supports after lipase adsorption was characterized with Transmission electron microscopy (TEM, JEM-2000) and Silver nitrate was used to stain protein. 2.6. Activity assay of lipase The reaction rate of the free and immobilized lipase preparations was determined according to the Hydrolysis reaction of p-NPP reported by Chiou and Wu [31]. One enzyme unit was the amount of biocatalyst liberating 1.0 mol of p-NP per minute in these conditions. The pH stat method with olive oil titrimetric assay was also used in this work [32]. One enzyme unit was the amount of biocatalyst liberating 1.0 mol of fatty acid per minute in these conditions. Activity retention was defined as the ratio of the activity of the amount of the enzyme immobilized on the support to the activity of the same amount of free enzyme. 2.7. Sum frequency generation spectroscopy (SFG) The sum frequency generation (SFG) vibrational spectra were obtained by using a custom-designed EKSPLA SFG spectrometer. Briefly, the visible input beam at 532 nm was generated by frequency doubling the fundamental output pulses with wavelength of 1064 nm from an EKSPLA Nd:YAG laser. Both the visible and infrared beams were focused at the sample surface with diameters of 0.5 mm with a pulse width of ∼30 ps and a repetition rate of 50 Hz. The incident angles of the visible beam and the IR beam were respectively 60◦ and 53◦ . In this study, the SFG spectra with an ssp polarization combination (i.e., an s-polarized sum frequency
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output, an s-polarized visible input, and a p-polarized infrared input) were collected at the surfaces of the functional supports [33]. 2.8. Adsorption of substrate and products The support (about 7.6 cm × 2.6 cm) was cut into 5 mm × 5 mm and submerged into 3.5 mL p-NPP solution (7.0 mM), p-NP solution (1 × 10−2 mM) or PA solution (1 × 10−2 mM), respectively. The adsorption process was carried out at room temperature in a shaking water bath for 5 min. Finally, the supports were removed, thoroughly rinsed with de-ionized water. The amount of adsorbed p-NPP or p-NP on the support was determined by measuring the initial and final concentrations of p-NPP or p-NP within the solutions and washings with UV–vis spectrophotometer. The amount of adsorbed PA on the support was determined by measuring the initial and final concentrations of PA within the solutions and washings by Gas Chromatography (Tianmei GC7900, China). Contact angles of olive oil or glycerol on the support surfaces were measured by the Sessile drop method using a DSA-10 drop sharp analysis (Kruss Co., Germany) at 25 ◦ C. The volume of the olive oil or glycerol drops used was always 3 L. All the reported values were averages of at least eight measurements and had a typical error of the mean of ±1◦ . 3. Results and discussion 3.1. Characterization of the functional group controlled surfaces To investigate enzyme adsorption, one important approach is to accurately control the surface chemistry of support, including the kind and density of functional group on the support surface. In this experiment, the functional group ended PSs were mixed with normal PS to fabricate the functional group controlled surfaces. Contact angle measurement is one of the most sensitive and effective methods for probing surface structure of polymers with 0.5–1 nm of surface sensitivity [34]. The contact angle of water on the support surface containing PS-F was determined and the results are shown in Table 1. Compare to PS, the contact angle of water on the hydrophilic group controlled surface (S-OH, S-NH2 and S-COOH) decreased from 86◦ to 64◦ , 70◦ and 60◦ , while that on the hydrophobic group controlled surface (PS-CH3 and PS-CF3 ) increased to 99◦ and 104◦ , respectively. This phenomenon could be explained by the fact that the hydrophilic functional group ( COOH, NH2 or OH) could segregate onto the glass-side surface, while the hydrophobic functional group ( CH3 or CF3 ) could segregate onto the air-side surface in order to decrease interface free energy [35]. Accordingly, compared to PS, the surface free energy and the polar surface free energy of the surfaces containing hydrophilic groups ( COOH, NH2 or OH) obviously increased. While, for the surfaces containing hydrophobic groups ( CH3 or CF3 ), the polar surface free energies decreased. This phenomenon was corresponding with the variation of the contact angle of water on the surfaces. The surface chemistry of the support was characterized by XPS analysis (shown in Fig. 1). For the pure PS surface, there was only a little oxygen peak, which might be due to oxidization during thermal treatment. On the XPS spectra of the surface containing PS-COOH, the oxygen peak increased obviously, which indicated carboxyl group existed on the surface. For the surface containing OH, nitride peak appeared, which came from the amide bond of PS-OH. For the surface containing CF3 , nitride and fluoride peaks appeared. As for surface containing NH2 , nitride peak became most obviously. For the surface containing CH3 , nitride and oxygen peaks existed. According to the results above,
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functional groups ( COOH, OH, CF3 , NH2 and CH3 ) were quantitatively introduced onto the support surfaces and these supports were employed to immobilize lipase in the next sections. 3.2. Lipase immobilization on the functional group controlled surfaces Table 2 lists the amount, activity and kinetic parameters of the immobilized lipases on the support surfaces. The number of immobilized protein on the surface containing CH3 was largest among these supports. CH3 is a typical hydrophobic group which could form hydrophobic interaction with protein to decrease the protein/support interface energy. Similar result could be found for the pure PS surface, which also showed a larger amount of immobilized protein due to its hydrophobic property. As for the surface containing CF3 , which showed available resistance for protein [36], the amount of adsorbed protein was smaller than that on PS. On the surfaces containing hydrophilic group, COOH, NH2 or OH, the amount of adsorbed protein was relative low. In protein adsorption, water plays a vital role, which complexes to the protein and support surface and then releases to bulk solution to form a new protein/support interface [37]. As hydrophilic groups could form stable hydrogen bond with water, the release of water from these groups could be rather difficult [38,39]. Thus, the introduction of COOH, NH2 or OH on the surfaces could effectively depress the protein adsorption [40,41]. The interaction between the enzyme and the supports weaken by introducing the hydrophilic groups, but the enzyme binding mechanism was still hydrophobic. The amount of adsorbed protein on the OH controlled surface was lowest. Lipase (its isoelectric point is 5.2) had a negative net charge under these conditions (the solution pH was 7.0), so it could form an attractive electrostatic interaction with NH2 on the support surface [42]. As there still were some positive groups on lipase surface, it was possible to form an electrostatic interaction between lipase and COOH on the support surface [43,44]. The electrostatic interaction could reinforce the interaction between the surface and protein and lead to a higher amount of adsorbed protein. Thus, the order of the amount of adsorbed protein on the surfaces with similar functional group density was S-CH3 > PS > S-CF3 > S-NH2 > SCOOH > S-OH. Furthermore, TEM was employed to observe the adsorbed lipase (shown in Fig. 2). In Fig. 2a, it could be found that lipase (the dark particle) was adsorbed on the PS surface (the gray background) with a diameter of 20–30 nm. Similar results were also found for the surfaces containing hydrophobic groups ( CH3 or CF3 ). For the surfaces containing hydrophilic groups ( NH2 , COOH or OH), there were the decreases of the surface area occupied by protein molecule, which was corresponding to the decreases of the amount of adsorbed protein. Especially, in Fig. 2f, for the surface containing OH, the amount of adsorbed protein was lowest and its diameter was about 15 nm. Lipase appears its shape (6.5 nm × 9.7 nm × 17.5 nm) [27], and adsorbs with two orientations of end-on and side-on. The difference of the diameter of the proteins adsorbed on the supports might be evidence that indicate the adsorbed proteins taking different orientation. On the surface containing hydrophobic group, lipase could take more side-on orientations with larger spreading due to the strong hydrophobic interaction between lipase and the surface, while on the surface containing hydrophilic group, the interaction was weaken and lipase could take more end-on orientations with smaller spreading. Additionally, C. rugosa lipase tends to aggregate through its very hydrophobic lid [45], so there were some large lipase particles with the diameter of 50–60 nm on the surfaces. Fig. 3 shows the SFG spectra of lipases on the support surfaces in the infrared frequency ranging from 2800 to 3100 cm−1 , which corresponds to the C H stretching vibrations. The SFG resonant signals
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Table 1 The contact angle of water, surface energy, polar surface energy and functional group surface density on the support surfaces (S, the pure PS surface; S-F, the surface containing functional group, such as CH3 , CF3 , OH, NH2 and COOH). Sample
S
S-CH3
Contact angle of water (◦ ) Surface energy (mN/m) Polar surface energy (mN/m) Functional group surface density (×10−4 mmol/m2 )
86 ± 1 42.7 ± 1.8 0.80 ± 0.02 –
99 39.5 0.52 15.8
of the C H stretching vibrations can provide clues to conformation and orientation of alkyl chains on the interface [46–50]. In the ssp spectrum of lipase (Fig. 3a), two intense resonances at 2875 and 2940 cm−1 belong respectively to the methyl symmetric stretching and its Fermi resonance (a coupling between the
± ± ± ±
S-CF3 1 2.0 0.02 0.5
104 37.8 0.01 16.2
S-OH ± ± ± ±
1 1.6 0.01 0.5
64 49.3 9.2 14.1
S-NH2 ± ± ± ±
1 2.1 0.20 0.4
70 48.1 8.2 15.0
± ± ± ±
S-COOH 1 1.8 0.16 0.4
60 50.2 10.1 14.6
± ± ± ±
1 2.1 0.22 0.5
fundamental stretching and overtone of methyl bending mode). In the spectrum of PS (Fig. 3h), a intense resonance at 3060 cm−1 could be attributed to the benzene stretching. For the spectra of lipases adsorbed on the functional supports (Fig. 3b–g), there were obvious increases of resonances at 2875 and 2940 cm−1 and an
Fig. 1. XPS spectra for the pure PS surface (a), the surfaces containing functional group
COOH (b),
OH (c),
CF3 (d),
NH2 (e) and
CH3 (f), respectively.
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Table 2 The amount, activity and kinetic parameters of the immobilized lipases on the support surfaces. Sample
S
Amount of adsorbed protein (ng/cm2 ) Activity retention (%) (olive oil) Activity retention (%) (p-NPP) Vmax (U/mg) Km (mM)
126 20 18 852 10.9
S-CH3 ± ± ± ± ±
6 1 1 54 0.6
134 24 26 1165 10.1
± ± ± ± ±
S-CF3 6 1 1 62 0.5
101 23 22 1034 9.2
S-OH ± ± ± ± ±
5 1 1 57 0.6
Fig. 2. TEM photographs of the pure PS surface (a) and the surfaces containing functional group ( CH3 (b),
decrease of resonance at 3060 cm−1 . These results indicated that lipase completely occupied the support surfaces. And two new resonances at 2850 and 2910 cm−1 , which belong respectively to the methylene symmetric stretching (ss) and methylene asymmetric stretching (as), could also be recognized. These phenomena were due to the variation of lipase conformation. Compared to the supports containing hydrophilic groups, the increase of resonances at 2850 and 2910 cm−1 was more obviously in the spectra of lipase adsorbed on the supports containing hydrophobic groups, which might also support that there was a more remarkable variation of lipase conformation on the hydrophobic supports. Especially, for the support containing CH3 (Fig. 3f), there was an intense resonances at 3050 cm−1 , which should belong to the benzene stretching of
62 61 62 2783 7.0
CF3 (c),
S-NH2 ± ± ± ± ±
OH (d)
4 2 2 96 0.5
75 47 60 2569 5.8
NH2 (e) and
± ± ± ± ±
S-COOH 4 2 2 102 0.4
68 43 42 1814 7.7
± ± ± ± ±
4 2 2 87 0.5
COOH (f)) after lipase adsorption.
the residues of styrene acrylic acid and tyrosine which come from lipase. 3.3. Activity of immobilized lipase In our experiment, the adsorption of the substrate and product were measured (shown in Figs. 4 and 5). As p-NPP contains a long hydrophobic carbon chain, this molecule could easily adsorb on the hydrophobic surface. The amount of adsorbed p-NPP on the surface containing CH3 was largest. For the surface containing OH and COOH, the amount of adsorbed p-NPP were rather low. Interestingly, the amount of adsorbed p-NPP on the surface containing NH2 was rather high, which might be due to the fact that
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Fig. 3. SFG spectra of lipase for the nascent (a) and adsorbed lipase on the pure PS surface (g) and the surfaces containing functional group ( OH (b), NH2 (c), COOH (d), CF3 (e) and CH3 (f)), respectively. SFG spectrum of PS (h).
Fig. 4. The amount of adsorbed p-NPP and PA on the functional controlled surfaces.
Fig. 5. The contact angle of olive oil and glycerol on the functional controlled surfaces.
NH2 could attract nitro group of p-NPP. The hydrolysis products of p-NPP are PA and p-NP. Considering the tiny concentration of product in early stage of the catalysis reaction, a low concentration of product solution was chosen for the adsorption experiment. PA also has a long carbon chain, so it adsorption behavior was similar to p-NPP. p-NP shows strong water solubility, so the amount of adsorbed p-NP was rather small (these data are not shown). For glycerol and olive oil, it is difficult to directly measure their adsorption amount. Thus, contact angle was used to characterize the affinity of the two molecules to these surfaces (shown in Fig. 5). The value of contact angle of olive oil on the surface containing CH3 was smallest, which indicated that the affinity of olive oil to the surface was highest among these surfaces. For the surfaces containing hydrophilic groups, the contact angle of olive oil was higher than that of PS. Since the most part of the surfaces was still occupied by PS, these surfaces also appeared to have a high affinity to the hydrophobic olive oil. Olive oil hydrolyzes two products, glycerol and fatty acids. PA could be regarded as a model of fatty acids. Compared to the hydrophobic surfaces, the angle of glycerol on the hydrophilic surfaces was low, since glycerol is a trihydric alcohol with high polarity. The hydrolysis reaction of lipid produces two kinds of products, fatty acids (such as PA) and alcohols (such as p-NP and glycerol). The alcohol molecules have strong water solubility and rarely adsorb on the support surfaces. Fatty acid with long carbon chain could easily adsorb on these support surfaces. However, considering the tiny concentration of productions in the reaction system, especially at the beginning stage, the effect of adsorbed fatty acid on the lipase activity should be rather limited compared to the adsorbed substrate. Now it has been recognized as a common feature that lipases could be activated in the presence of hydrophobic interface [5,7]. Special emphasis has been paid to the selective adsorption of lipases on tailor-made strongly hydrophobic support surfaces [14]. In our experiment, after adsorption on PS, lipase retained only 20% activity. There existed strong hydrophobic interaction between PS surface and lipase, which lead to the obvious spreading of lipase. Thus, lipase’ conformation changed evidently and it activity lost largely. Lipase adsorbed on the surface containing CH3 showed a higher activity retention than that on the PS surface. The increase of the amount of adsorbed substrate on the surface containing CH3 could also help to enhance lipase activity. For the surface containing CF3 , the open-state conformation of lipase could also be induced by the hydrophobic support surface. However, the amount of adsorbed substrate on the surface containing CF3 decreased. Thus, the activity retention on the surface containing CF3 was lower than that on the surface containing CH3 . On the surface containing OH, the adsorbed lipase showed the highest activity. After the introduction of this hydrophilic group, there still existed a large number of PS chain on this surface. During lipase adsorption, the hydrophobic area on the support surface could form hydrophobic interaction with lipase’s lid and help to yield open-state conformation of lipase. At the same time, the hydrophilic group on the support surface could form hydration layer with water, which effectively inhibited the spreading of adsorbed lipase and induced an end-on orientation of adsorbed lipase. From Fig. 2, it could be seen that the diameter of adsorbed lipase was about 10–15 nm on this surface. Besides the short diameter of lipase observed with TEM, the high activity of the lipase adsorbed on the surface containing OH might also be evidence that support the conclusion that the adsorbed lipase could take the end-on orientation. The end-on orientation of lipase could favor the surface activation of lipase’ lid and avoid the unnecessary hydrophobic interaction between the support surface and the hydrophobic residues from other lipase chain (shown in Fig. 6).
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Fig. 6. The adsorbed lipase on the functional controlled surface.
Deng et al. reported that the activity of adsorbed lipase increased by tethering phospholipid on polypropylene to create a biocompatible interface [32]. Therefore, a surface with hydrophobic area and hydrophilic group might be a promising support surface to keep lipase activity. As discussed above, there existed electrostatic interaction between lipase and the surface containing COOH or NH2 . This interaction could increase the amount of adsorbed lipase and change lipase native conformation. Thus, the activity of lipase adsorbed on the surface containing COOH or NH2 was lower than that on the surface containing OH. Interestingly, for the surface containing NH2 , the lipase activity retention of hydrolysis of p-NPP was obviously higher than that of olive oil, which might be ascribed to the fact that there is a high p-NPP concentration on the support surface due to the attraction between NH2 and nitro group of p-NPP. Kinetic parameters for the hydrolytic activity of the free and immobilized lipases, Km and Vmax , were assayed (shown in Table 2). Vmax showed the same trend with the activity retention, SOH > S-NH2 > S-COOH > S-CF3 > S-CH3 > PS. Lipases immobilized on the surface containing hydrophilic group, OH, NH2 or COOH, showed higher catalysis activity, which might be due to the hydration layer which created by the hydrophilic group on the support surface could inhibit the spreading of lipase and keep the nature conformation of lipase. The order of Km value was PS > SCH3 > S-CF3 > S-COOH > S-OH > S-NH2 . Km is defined as the substrate concentration that gives a reaction velocity of 1/2 Vmax . This parameter reflects the effective characteristics of the enzyme and depends upon both partition and diffusion effects. The surface containing NH2 showed the lowest Km value, which might be owing to the fact that NH2 could attract the nitro group of p-NPP. 4. Conclusions In this work, the functional group controlled surfaces were fabricated and used as model supports to investigate the effect of surface chemistry on lipase adsorption and activity. The hydrophilic groups, COOH, NH2 and OH, could form hydration layer with water and thus effectively reduce the amount of adsorbed protein. On the surface containing hydrophobic group, lipase could take more side-on orientations with larger spreading due to the strong hydrophobic interaction, while on the surface containing hydrophilic group, these interaction was weakened and lipase could take more end-on orientations with smaller spreading. Lipase immobilized on the surface containing OH showed the highest catalysis activity, which might be due to the less spreading and the end-on orientation of absorbed lipase on this surface. Therefore, a surface with hydrophobic area and hydrophilic group might be a promising support surface to keep lipase activity. The
surface containing NH2 appeared to have a higher activity, which might be ascribed to the fact that there was a high substrate concentration on the surface due to the attraction between NH2 and the nitro group of p-NPP. This phenomenon indicated that it might be an advisable method to design the support surface with specific group which could effectively adsorb substrate to increase enzyme activity. Thus, the support surface chemistry could also affect the activity of immobilized lipase by the substrate adsorption on the support interface. This work might be useful for the understanding of lipase adsorption and its activity with more quantitative information and the design and preparation of support for enzyme immobilization. Acknowledgements Authors are grateful for the financial support from the National Natural Science Foundation of China (Grant Nos. 50803059 and 31200746), the Natural Science Foundation of Zhejiang Province (Grant No. Y407275). References [1] M. Svedendahl, K. Hult, P. Berglund, J. Am. Chem. Soc. 127 (2005) 17988–17989. [2] C.E. Humphrey, N.J. Turner, M.A.M. Easson, S.L. Flitsch, R.V. Ulijn, J. Am. Chem. Soc. 125 (2003) 13952–13953. [3] P.M. Kosaka, Y. Kawano, O.A. El Seoud, D.F.S. Petri, Langmuir 23 (2007) 12167–12173. [4] P.J. Wilde, B.S. Chu, Adv. Colloid Interface Sci. 165 (2011) 14–22. [5] A.W. Sonesson, H. Brismar, T.H. Callisen, U.M. Elofsson, Langmuir 23 (2007) 2706–2713. [6] B.S. Chu, G.T. Rich, M.J. Ridout, R.M. Faulks, M.S.J. Wickham, P.J. Wilde, Langmuir 25 (2009) 9352–9360. [7] P. Ye, Z.K. Xu, J. Wu, C. Innocent, P. Seta, Macromolecules 39 (2006) 1041–1045. [8] P. Reis, K. Holmberg, H. Watzke, M.E. Leser, R. Miller, Adv. Colloid Interface Sci. 148 (2009) 237–250. [9] P. Reis, K. Holmberg, T. Debeche, B. Folmer, L. Fauconnot, H. Watzke, Langmuir 22 (2006) 8169–8177. [10] P.V. Iyer, L. Ananthanarayan, Process Biochem. 43 (2008) 1019–1032. [11] A. Dyal, K. Loos, M. Noto, S.W. Chang, C. Spagnoli, K.V.P.M. Shafi, J. Am. Chem. Soc. 125 (2003) 1684–1685. [12] H. Jia, G. Zhu, B. Vugrinovich, W. Kataphinan, D.H. Reneker, P. Wang, Biotechnol. Progr. 18 (2002) 1027–1032. [13] K. Wannerberger, S. Welin-Klintström, T. Arnebrant, Langmuir 13 (1997) 784–790. [14] Q. Jin, G. Jia, Y. Zhang, Q. Yang, C. Li, Langmuir 27 (2011) 12016–12024. [15] K. Wannerberger, T. Arnebrant, Langmuir 13 (1997) 3488–3493. [16] I. Gill, E. Pastor, A. Ballesteros, J. Am. Chem. Soc. 121 (1999) 9487–9496. [17] J.M. Palomo, R.L. Segura, G. Fernandez-Lorente, M. Pernas, M.L. Rua, J.M. Guisan, R. Fernandez-Lafuente, Biotechnol. Progr. 20 (2004) 630–635. [18] P. Ye, Z.K. Xu, J. Wu, C. Innocent, P. Seta, Biomaterials 27 (2006) 4169–4176. [19] M. Furuhashi, G. Tuncman, C.Z. Görgün, L. Makowski, G. Atsumi, E. Vaillancourt, K. Kono, V.R. Babaev, S. Fazio, M.F. Linton, R. Sulsky, J.A. Robl, R.A. Parker, G.S. Hotamisligil, Nature 447 (2007) 959–965. [20] J. Muderhwa, H.J. Brockman, Biol. Chem. 267 (1992) 24184–24192. [21] A. Alouou, J. Rodriguez, S. Fernandez, D. Oosterhout, D. Puccinelli, F. Carrière, Biochim. Biophys. Acta 1761 (2006) 995–1013. [22] H. Stamatis, A. Xenakis, F.N. Klosis, Biotechnol. Adv. 17 (1999) 293–318.
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Contents lists available at SciVerse ScienceDirect
Journal of Molecular Catalysis B: Enzymatic journal homepage: www.elsevier.com/locate/molcatb
Effect of support surface chemistry on lipase adsorption and activity Ye Peng ∗ , Han Zhu-Ping, Xu Yong-Juan, Hu Peng-Cheng, Tong Ji-Jun Key Laboratory of Advanced Textile Materials and Manufacturing Technology of Education Ministry, Department of Chemistry, Zhejiang Sci-Tech University, Hangzhou 310018, PR China
a r t i c l e
i n f o
Article history: Received 7 January 2013 Received in revised form 30 March 2013 Accepted 26 April 2013 Available online 4 May 2013 Keywords: Support surface chemistry Lipase Enzyme immobilization Functional group-ended polymer
a b s t r a c t The functional group controlled surfaces (S-F, functional groups including NH2 , COOH, OH, CH3 and CF3 ) were fabricated by mixing polystyrene (PS) respectively with functional group-ended polystyrenes (PS-Fs). These surfaces were used as model supports to investigate the effect of surface chemistry on lipase adsorption and activity. The order of the amount of adsorbed lipase on the surface with similar functional group density was S-CH3 > PS > S-CF3 > S-NH2 > S-COOH > S-OH. It could be found that on the surface containing hydrophobic group, lipase could take more side-on orientations with larger spreading, while on the surface containing hydrophilic group, lipase could take more end-on orientations with smaller spreading. Lipase immobilized on the surface containing OH showed the highest activity. The adsorption of substrates and products on the functional group controlled surfaces was also measured. The surface containing NH2 showed a higher activity which might be ascribed to the fact that there was a high substrate concentration on the lipase/support interface. The order of the activity retention of the immobilized lipase was S-OH > S-NH2 > S-COOH > S-CF3 > S-CH3 > PS. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Lipases, as versatile biocatalysts, have gained considerable importance and the potential applications of lipases in biotransformations have been fully documented with a plethora of examples, on both hydrolyzation in aqueous media and synthesis in organic media [1–4]. Due to an opposite polarity between the enzyme (hydrophilic) and their substrate (lipophilic), lipase reaction occurs at the interface between the aqueous and the oil phase. Hence, interfaces are the key spots for lipase biocatalysis [5–8]. There are three types of interfaces: air–water, oil–water and solid–water, which could activate lipase [9]. The solid–water interface provides the possibility to immobilize enzyme and to tailor a surface with which the lipase interacts. In fact, enzymes have always been immobilized onto insoluble or solid supports for industrial applications, which are regarded as a useful tool to enhance their thermal and operational stabilities, and reduce the cost [10–13]. Special emphasis has been attached to the selective adsorption of lipases on tailor-made strongly hydrophobic support surfaces [14–17]. This immobilization procedure is based on the assumption that the large hydrophobic area that surrounds the active site of lipases is the one mainly involved in their adsorption on strongly hydrophobic solid surfaces. However, some
∗ Corresponding author. Tel.: +86 571 86843691; fax: +86 571 86843653. E-mail address: [email protected] (P. Ye). 1381-1177/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.molcatb.2013.04.015
authors reported that lipase could also retain higher activity on the hydrophilic support [9,18]. In lipase catalysis interface composition, which is described by the term “interface quality” [19–23], plays a vital role. There are at least five kinds of materials on the interface, including lipase, support, substrate, product and solution. Recent studies demonstrate that lipase activity as a function of interface composition is more attributed to substrate inaccessibility rather than to enzyme denaturation or inactivation, as it is often hypothesized. Holmberg and co-workers measured the adsorption of two substrates capric acid (decanoic acid) and monocaprin on the lipase-covered surface, and found that lipase adsorbed at the hydrophilic surface favored hydrolysis and capric acid was the main product when monocaprin was used as substrate [9]. However, knowledge about the effect of the interface composition in lipase catalysis is still limited and complete understanding of enzyme–substrate or product–substrate interaction is necessary. To control support surface properties, the modification of polymer end groups has emerged as a practical means [24–26]. In this study, three hydrophilic groups ( NH2 , COOH and OH) and two hydrophobic groups ( CH3 and CF3 ) were quantitatively introduced onto the support surfaces by mixing PS solution with PS-Fs, respectively. Lipase (from Candida rugosa) [27], was immobilized on the supports as a model enzyme. The adsorption of two substrates (p-nitrophenyl palmitate (p-NPP) and olive oil) and three products (palmitic acid (PA), glycerol and p-nitrophenol (p-NP)) was also measured. The effects of the support surface chemistry on the adsorption and activity of lipase were investigated.
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PS-COOH + NH2-CH2CH2-F DCC PS-CONH-CH2CH2-F Scheme 1. Synthesis of functional group-ended polystyrene (PS-F, functional groups including NH2 , OH, CH3 and CF3 ).
PS-CONH-CH2CH2-F + H2O HCL
PS-COOH + NH2-CH2CH2-F
Scheme 2. Hydrolyzation of functional group-ended polystyrene (PS-F, functional groups including NH2 , OH, CH3 and CF3 ) to carboxyl-ended polystyrene (PSCOOH) in HCL solution.
2. Materials and methods 2.1. Materials Lipase (from C. rugosa), Bradford reagent, bovine serum albumin (BSA) and p-nitrophenyl palmitate (p-NPP) were purchased from Sigma and used as received. Polystyrene (PS, Mn 92,100, PDI 2.6) was purchased from Aldrich Co. All the other chemicals were of analytical grade and used without further purification. 2.2. Synthesis of the functional group-ended polystyrenes Carboxyl-ended polystyrene (PS-COOH, Mn 4400, PDI 1.2) was prepared as previously reported [28]. Hydroxyl-ended polystyrene (PS-OH), trifluoromethyl-ended polystyrene (PS-CF3 ), methyl-ended polystyrene (PS-CH3 ) and amino-ended polystyrene (PS-NH2 ) were synthesized by the reaction between PS-COOH with hydroxyl-ethane-amine, trifluoromethyl-ethane-amine, methylethane-amine or ethylenediamine in the presence of dicyclocarbondiamide (DCC) in dehydrated tetrahydrofuran (THF) solution in a three-neck round-bottom flask equipped with a stopcock and a magnetic stirring bar, respectively. Prior to use, the flask was vacuumed and backfilled with dry nitrogen several times. The reaction mixture was then kept at room temperature for 48 h under a nitrogen atmosphere. After reaction, the mixture was filtered to remove 1,3-dicyclohexylurea and then precipitated into water, and the resulting product was dried in vacuum for 24 h. The product was then dissolved into THF, and the resulting solution was passed through an alumina column to remove the impurity (Scheme 1). 2.3. Preparation of the functional group controlled surfaces Prescribed amounts of PS were dissolved in xylene at a concentration of 5 wt% for the preparation of casting solutions. The functional group controlled surfaces were prepared by mixing PS solution (95 wt%) with functional groups-ended polystyrenes solution (5 wt%) (PS-F, functional groups such as CH3 , CF3 , NH2 , COOH and OH), then casting the solution (2 mL) on rimmed glass plates (7.6 cm × 2.6 cm) and drying at 25 ◦ C in dried air, finally drying at 150 ◦ C for 10 h, respectively. Each support taken off the glass plate had two surfaces that were formed in different environments. One surface of the support contacting with the dried air was designated as the air-side surface, and the other surface contacting with the glass plate was designated as the glass-side surface. The surface density of functional group might be different on the two side surfaces. In order to eliminate the possible error owing to this difference, the supports were poured with pure PS solution (0.2 mL) on their one side surfaces, the air-side surface for the support containing COOH, NH2 or OH, and the glass-side surface for the support containing CH3 or CF3 , and dry them again to form one side of pure PS surfaces. 2.4. Characterization of the functional group controlled surfaces X-ray photoelectron spectroscopy (XPS) experiments were performed on a PHI-5000C ESCA system (Perkin–Elmer) with Al K␣ radiation (h = 1486.6 eV). Contact angles of water on the support surfaces were measured by the Sessile drop method using a DSA10 drop sharp analysis (Kruss Co., Germany) at 25 ◦ C. The volume of the water drops used was always 3 L. All the reported values were averages of at least eight measurements taken at different
locations on the support surface and had a typical error of the mean of ±1◦ . The surface free energy was calculated according Owens and Wendt’s theory from the measured contact angles of water and diiodomethane on the samples [29]. The surface densities of COOH were obtained by measuring the load and release of methylene blue (MB) [30]. In order to measure the surface densities of functional group, such as CH3 , NH2 , OH and CF3 , the amide bond of PS-Fs were hydrolyzed to carboxyl group in HCL solution (20 wt%) at 50 ◦ C for 24 h, respectively (Scheme 2). 2.5. Lipase immobilization The support (about 10 cm × 10 cm) was cut into 5 mm × 5 mm and thoroughly washed with deionized water, and then rinsed with phosphate buffered solution (PBS) (25 mM, pH 7.0). After that, the support was submerged in 10 mL lipase solution (0.5 mg/mL) at room temperature in a shaking water bath for 8 h. Finally, the supports were removed, thoroughly rinsed with PBS and then rinsed with de-ionized water. The amount of adsorbed protein on the support was determined by measuring the initial and final concentrations of protein within the protein solutions and washings using Coomassie Brilliant Blue reagent, following Bradford’s method. As mention above, the support had two surfaces, one was the functional group controlled surface and the other was the pure PS surface. The value of the amount of adsorbed protein on the functional group controlled surface was calculated by subtracting the amount of adsorbed protein on the pure PS surface of the same size in the same experiment conditions. The surface morphology of these supports after lipase adsorption was characterized with Transmission electron microscopy (TEM, JEM-2000) and Silver nitrate was used to stain protein. 2.6. Activity assay of lipase The reaction rate of the free and immobilized lipase preparations was determined according to the Hydrolysis reaction of p-NPP reported by Chiou and Wu [31]. One enzyme unit was the amount of biocatalyst liberating 1.0 mol of p-NP per minute in these conditions. The pH stat method with olive oil titrimetric assay was also used in this work [32]. One enzyme unit was the amount of biocatalyst liberating 1.0 mol of fatty acid per minute in these conditions. Activity retention was defined as the ratio of the activity of the amount of the enzyme immobilized on the support to the activity of the same amount of free enzyme. 2.7. Sum frequency generation spectroscopy (SFG) The sum frequency generation (SFG) vibrational spectra were obtained by using a custom-designed EKSPLA SFG spectrometer. Briefly, the visible input beam at 532 nm was generated by frequency doubling the fundamental output pulses with wavelength of 1064 nm from an EKSPLA Nd:YAG laser. Both the visible and infrared beams were focused at the sample surface with diameters of 0.5 mm with a pulse width of ∼30 ps and a repetition rate of 50 Hz. The incident angles of the visible beam and the IR beam were respectively 60◦ and 53◦ . In this study, the SFG spectra with an ssp polarization combination (i.e., an s-polarized sum frequency
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output, an s-polarized visible input, and a p-polarized infrared input) were collected at the surfaces of the functional supports [33]. 2.8. Adsorption of substrate and products The support (about 7.6 cm × 2.6 cm) was cut into 5 mm × 5 mm and submerged into 3.5 mL p-NPP solution (7.0 mM), p-NP solution (1 × 10−2 mM) or PA solution (1 × 10−2 mM), respectively. The adsorption process was carried out at room temperature in a shaking water bath for 5 min. Finally, the supports were removed, thoroughly rinsed with de-ionized water. The amount of adsorbed p-NPP or p-NP on the support was determined by measuring the initial and final concentrations of p-NPP or p-NP within the solutions and washings with UV–vis spectrophotometer. The amount of adsorbed PA on the support was determined by measuring the initial and final concentrations of PA within the solutions and washings by Gas Chromatography (Tianmei GC7900, China). Contact angles of olive oil or glycerol on the support surfaces were measured by the Sessile drop method using a DSA-10 drop sharp analysis (Kruss Co., Germany) at 25 ◦ C. The volume of the olive oil or glycerol drops used was always 3 L. All the reported values were averages of at least eight measurements and had a typical error of the mean of ±1◦ . 3. Results and discussion 3.1. Characterization of the functional group controlled surfaces To investigate enzyme adsorption, one important approach is to accurately control the surface chemistry of support, including the kind and density of functional group on the support surface. In this experiment, the functional group ended PSs were mixed with normal PS to fabricate the functional group controlled surfaces. Contact angle measurement is one of the most sensitive and effective methods for probing surface structure of polymers with 0.5–1 nm of surface sensitivity [34]. The contact angle of water on the support surface containing PS-F was determined and the results are shown in Table 1. Compare to PS, the contact angle of water on the hydrophilic group controlled surface (S-OH, S-NH2 and S-COOH) decreased from 86◦ to 64◦ , 70◦ and 60◦ , while that on the hydrophobic group controlled surface (PS-CH3 and PS-CF3 ) increased to 99◦ and 104◦ , respectively. This phenomenon could be explained by the fact that the hydrophilic functional group ( COOH, NH2 or OH) could segregate onto the glass-side surface, while the hydrophobic functional group ( CH3 or CF3 ) could segregate onto the air-side surface in order to decrease interface free energy [35]. Accordingly, compared to PS, the surface free energy and the polar surface free energy of the surfaces containing hydrophilic groups ( COOH, NH2 or OH) obviously increased. While, for the surfaces containing hydrophobic groups ( CH3 or CF3 ), the polar surface free energies decreased. This phenomenon was corresponding with the variation of the contact angle of water on the surfaces. The surface chemistry of the support was characterized by XPS analysis (shown in Fig. 1). For the pure PS surface, there was only a little oxygen peak, which might be due to oxidization during thermal treatment. On the XPS spectra of the surface containing PS-COOH, the oxygen peak increased obviously, which indicated carboxyl group existed on the surface. For the surface containing OH, nitride peak appeared, which came from the amide bond of PS-OH. For the surface containing CF3 , nitride and fluoride peaks appeared. As for surface containing NH2 , nitride peak became most obviously. For the surface containing CH3 , nitride and oxygen peaks existed. According to the results above,
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functional groups ( COOH, OH, CF3 , NH2 and CH3 ) were quantitatively introduced onto the support surfaces and these supports were employed to immobilize lipase in the next sections. 3.2. Lipase immobilization on the functional group controlled surfaces Table 2 lists the amount, activity and kinetic parameters of the immobilized lipases on the support surfaces. The number of immobilized protein on the surface containing CH3 was largest among these supports. CH3 is a typical hydrophobic group which could form hydrophobic interaction with protein to decrease the protein/support interface energy. Similar result could be found for the pure PS surface, which also showed a larger amount of immobilized protein due to its hydrophobic property. As for the surface containing CF3 , which showed available resistance for protein [36], the amount of adsorbed protein was smaller than that on PS. On the surfaces containing hydrophilic group, COOH, NH2 or OH, the amount of adsorbed protein was relative low. In protein adsorption, water plays a vital role, which complexes to the protein and support surface and then releases to bulk solution to form a new protein/support interface [37]. As hydrophilic groups could form stable hydrogen bond with water, the release of water from these groups could be rather difficult [38,39]. Thus, the introduction of COOH, NH2 or OH on the surfaces could effectively depress the protein adsorption [40,41]. The interaction between the enzyme and the supports weaken by introducing the hydrophilic groups, but the enzyme binding mechanism was still hydrophobic. The amount of adsorbed protein on the OH controlled surface was lowest. Lipase (its isoelectric point is 5.2) had a negative net charge under these conditions (the solution pH was 7.0), so it could form an attractive electrostatic interaction with NH2 on the support surface [42]. As there still were some positive groups on lipase surface, it was possible to form an electrostatic interaction between lipase and COOH on the support surface [43,44]. The electrostatic interaction could reinforce the interaction between the surface and protein and lead to a higher amount of adsorbed protein. Thus, the order of the amount of adsorbed protein on the surfaces with similar functional group density was S-CH3 > PS > S-CF3 > S-NH2 > SCOOH > S-OH. Furthermore, TEM was employed to observe the adsorbed lipase (shown in Fig. 2). In Fig. 2a, it could be found that lipase (the dark particle) was adsorbed on the PS surface (the gray background) with a diameter of 20–30 nm. Similar results were also found for the surfaces containing hydrophobic groups ( CH3 or CF3 ). For the surfaces containing hydrophilic groups ( NH2 , COOH or OH), there were the decreases of the surface area occupied by protein molecule, which was corresponding to the decreases of the amount of adsorbed protein. Especially, in Fig. 2f, for the surface containing OH, the amount of adsorbed protein was lowest and its diameter was about 15 nm. Lipase appears its shape (6.5 nm × 9.7 nm × 17.5 nm) [27], and adsorbs with two orientations of end-on and side-on. The difference of the diameter of the proteins adsorbed on the supports might be evidence that indicate the adsorbed proteins taking different orientation. On the surface containing hydrophobic group, lipase could take more side-on orientations with larger spreading due to the strong hydrophobic interaction between lipase and the surface, while on the surface containing hydrophilic group, the interaction was weaken and lipase could take more end-on orientations with smaller spreading. Additionally, C. rugosa lipase tends to aggregate through its very hydrophobic lid [45], so there were some large lipase particles with the diameter of 50–60 nm on the surfaces. Fig. 3 shows the SFG spectra of lipases on the support surfaces in the infrared frequency ranging from 2800 to 3100 cm−1 , which corresponds to the C H stretching vibrations. The SFG resonant signals
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Table 1 The contact angle of water, surface energy, polar surface energy and functional group surface density on the support surfaces (S, the pure PS surface; S-F, the surface containing functional group, such as CH3 , CF3 , OH, NH2 and COOH). Sample
S
S-CH3
Contact angle of water (◦ ) Surface energy (mN/m) Polar surface energy (mN/m) Functional group surface density (×10−4 mmol/m2 )
86 ± 1 42.7 ± 1.8 0.80 ± 0.02 –
99 39.5 0.52 15.8
of the C H stretching vibrations can provide clues to conformation and orientation of alkyl chains on the interface [46–50]. In the ssp spectrum of lipase (Fig. 3a), two intense resonances at 2875 and 2940 cm−1 belong respectively to the methyl symmetric stretching and its Fermi resonance (a coupling between the
± ± ± ±
S-CF3 1 2.0 0.02 0.5
104 37.8 0.01 16.2
S-OH ± ± ± ±
1 1.6 0.01 0.5
64 49.3 9.2 14.1
S-NH2 ± ± ± ±
1 2.1 0.20 0.4
70 48.1 8.2 15.0
± ± ± ±
S-COOH 1 1.8 0.16 0.4
60 50.2 10.1 14.6
± ± ± ±
1 2.1 0.22 0.5
fundamental stretching and overtone of methyl bending mode). In the spectrum of PS (Fig. 3h), a intense resonance at 3060 cm−1 could be attributed to the benzene stretching. For the spectra of lipases adsorbed on the functional supports (Fig. 3b–g), there were obvious increases of resonances at 2875 and 2940 cm−1 and an
Fig. 1. XPS spectra for the pure PS surface (a), the surfaces containing functional group
COOH (b),
OH (c),
CF3 (d),
NH2 (e) and
CH3 (f), respectively.
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Table 2 The amount, activity and kinetic parameters of the immobilized lipases on the support surfaces. Sample
S
Amount of adsorbed protein (ng/cm2 ) Activity retention (%) (olive oil) Activity retention (%) (p-NPP) Vmax (U/mg) Km (mM)
126 20 18 852 10.9
S-CH3 ± ± ± ± ±
6 1 1 54 0.6
134 24 26 1165 10.1
± ± ± ± ±
S-CF3 6 1 1 62 0.5
101 23 22 1034 9.2
S-OH ± ± ± ± ±
5 1 1 57 0.6
Fig. 2. TEM photographs of the pure PS surface (a) and the surfaces containing functional group ( CH3 (b),
decrease of resonance at 3060 cm−1 . These results indicated that lipase completely occupied the support surfaces. And two new resonances at 2850 and 2910 cm−1 , which belong respectively to the methylene symmetric stretching (ss) and methylene asymmetric stretching (as), could also be recognized. These phenomena were due to the variation of lipase conformation. Compared to the supports containing hydrophilic groups, the increase of resonances at 2850 and 2910 cm−1 was more obviously in the spectra of lipase adsorbed on the supports containing hydrophobic groups, which might also support that there was a more remarkable variation of lipase conformation on the hydrophobic supports. Especially, for the support containing CH3 (Fig. 3f), there was an intense resonances at 3050 cm−1 , which should belong to the benzene stretching of
62 61 62 2783 7.0
CF3 (c),
S-NH2 ± ± ± ± ±
OH (d)
4 2 2 96 0.5
75 47 60 2569 5.8
NH2 (e) and
± ± ± ± ±
S-COOH 4 2 2 102 0.4
68 43 42 1814 7.7
± ± ± ± ±
4 2 2 87 0.5
COOH (f)) after lipase adsorption.
the residues of styrene acrylic acid and tyrosine which come from lipase. 3.3. Activity of immobilized lipase In our experiment, the adsorption of the substrate and product were measured (shown in Figs. 4 and 5). As p-NPP contains a long hydrophobic carbon chain, this molecule could easily adsorb on the hydrophobic surface. The amount of adsorbed p-NPP on the surface containing CH3 was largest. For the surface containing OH and COOH, the amount of adsorbed p-NPP were rather low. Interestingly, the amount of adsorbed p-NPP on the surface containing NH2 was rather high, which might be due to the fact that
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Fig. 3. SFG spectra of lipase for the nascent (a) and adsorbed lipase on the pure PS surface (g) and the surfaces containing functional group ( OH (b), NH2 (c), COOH (d), CF3 (e) and CH3 (f)), respectively. SFG spectrum of PS (h).
Fig. 4. The amount of adsorbed p-NPP and PA on the functional controlled surfaces.
Fig. 5. The contact angle of olive oil and glycerol on the functional controlled surfaces.
NH2 could attract nitro group of p-NPP. The hydrolysis products of p-NPP are PA and p-NP. Considering the tiny concentration of product in early stage of the catalysis reaction, a low concentration of product solution was chosen for the adsorption experiment. PA also has a long carbon chain, so it adsorption behavior was similar to p-NPP. p-NP shows strong water solubility, so the amount of adsorbed p-NP was rather small (these data are not shown). For glycerol and olive oil, it is difficult to directly measure their adsorption amount. Thus, contact angle was used to characterize the affinity of the two molecules to these surfaces (shown in Fig. 5). The value of contact angle of olive oil on the surface containing CH3 was smallest, which indicated that the affinity of olive oil to the surface was highest among these surfaces. For the surfaces containing hydrophilic groups, the contact angle of olive oil was higher than that of PS. Since the most part of the surfaces was still occupied by PS, these surfaces also appeared to have a high affinity to the hydrophobic olive oil. Olive oil hydrolyzes two products, glycerol and fatty acids. PA could be regarded as a model of fatty acids. Compared to the hydrophobic surfaces, the angle of glycerol on the hydrophilic surfaces was low, since glycerol is a trihydric alcohol with high polarity. The hydrolysis reaction of lipid produces two kinds of products, fatty acids (such as PA) and alcohols (such as p-NP and glycerol). The alcohol molecules have strong water solubility and rarely adsorb on the support surfaces. Fatty acid with long carbon chain could easily adsorb on these support surfaces. However, considering the tiny concentration of productions in the reaction system, especially at the beginning stage, the effect of adsorbed fatty acid on the lipase activity should be rather limited compared to the adsorbed substrate. Now it has been recognized as a common feature that lipases could be activated in the presence of hydrophobic interface [5,7]. Special emphasis has been paid to the selective adsorption of lipases on tailor-made strongly hydrophobic support surfaces [14]. In our experiment, after adsorption on PS, lipase retained only 20% activity. There existed strong hydrophobic interaction between PS surface and lipase, which lead to the obvious spreading of lipase. Thus, lipase’ conformation changed evidently and it activity lost largely. Lipase adsorbed on the surface containing CH3 showed a higher activity retention than that on the PS surface. The increase of the amount of adsorbed substrate on the surface containing CH3 could also help to enhance lipase activity. For the surface containing CF3 , the open-state conformation of lipase could also be induced by the hydrophobic support surface. However, the amount of adsorbed substrate on the surface containing CF3 decreased. Thus, the activity retention on the surface containing CF3 was lower than that on the surface containing CH3 . On the surface containing OH, the adsorbed lipase showed the highest activity. After the introduction of this hydrophilic group, there still existed a large number of PS chain on this surface. During lipase adsorption, the hydrophobic area on the support surface could form hydrophobic interaction with lipase’s lid and help to yield open-state conformation of lipase. At the same time, the hydrophilic group on the support surface could form hydration layer with water, which effectively inhibited the spreading of adsorbed lipase and induced an end-on orientation of adsorbed lipase. From Fig. 2, it could be seen that the diameter of adsorbed lipase was about 10–15 nm on this surface. Besides the short diameter of lipase observed with TEM, the high activity of the lipase adsorbed on the surface containing OH might also be evidence that support the conclusion that the adsorbed lipase could take the end-on orientation. The end-on orientation of lipase could favor the surface activation of lipase’ lid and avoid the unnecessary hydrophobic interaction between the support surface and the hydrophobic residues from other lipase chain (shown in Fig. 6).
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Fig. 6. The adsorbed lipase on the functional controlled surface.
Deng et al. reported that the activity of adsorbed lipase increased by tethering phospholipid on polypropylene to create a biocompatible interface [32]. Therefore, a surface with hydrophobic area and hydrophilic group might be a promising support surface to keep lipase activity. As discussed above, there existed electrostatic interaction between lipase and the surface containing COOH or NH2 . This interaction could increase the amount of adsorbed lipase and change lipase native conformation. Thus, the activity of lipase adsorbed on the surface containing COOH or NH2 was lower than that on the surface containing OH. Interestingly, for the surface containing NH2 , the lipase activity retention of hydrolysis of p-NPP was obviously higher than that of olive oil, which might be ascribed to the fact that there is a high p-NPP concentration on the support surface due to the attraction between NH2 and nitro group of p-NPP. Kinetic parameters for the hydrolytic activity of the free and immobilized lipases, Km and Vmax , were assayed (shown in Table 2). Vmax showed the same trend with the activity retention, SOH > S-NH2 > S-COOH > S-CF3 > S-CH3 > PS. Lipases immobilized on the surface containing hydrophilic group, OH, NH2 or COOH, showed higher catalysis activity, which might be due to the hydration layer which created by the hydrophilic group on the support surface could inhibit the spreading of lipase and keep the nature conformation of lipase. The order of Km value was PS > SCH3 > S-CF3 > S-COOH > S-OH > S-NH2 . Km is defined as the substrate concentration that gives a reaction velocity of 1/2 Vmax . This parameter reflects the effective characteristics of the enzyme and depends upon both partition and diffusion effects. The surface containing NH2 showed the lowest Km value, which might be owing to the fact that NH2 could attract the nitro group of p-NPP. 4. Conclusions In this work, the functional group controlled surfaces were fabricated and used as model supports to investigate the effect of surface chemistry on lipase adsorption and activity. The hydrophilic groups, COOH, NH2 and OH, could form hydration layer with water and thus effectively reduce the amount of adsorbed protein. On the surface containing hydrophobic group, lipase could take more side-on orientations with larger spreading due to the strong hydrophobic interaction, while on the surface containing hydrophilic group, these interaction was weakened and lipase could take more end-on orientations with smaller spreading. Lipase immobilized on the surface containing OH showed the highest catalysis activity, which might be due to the less spreading and the end-on orientation of absorbed lipase on this surface. Therefore, a surface with hydrophobic area and hydrophilic group might be a promising support surface to keep lipase activity. The
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