Adsorption and Activity of Candida rugosa Lipase on Polypropylene

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Adsorption and Activity of Candida rugosa Lipase on Polypropylene Hollow Fiber Membrane Modified with Phospholipid Analogous Polymers Hong-Tao Deng,† Zhi-Kang Xu,*,† Xiao-Jun Huang,† Jian Wu,‡ and Patrick Seta§ Institute of Polymer Science, Zhejiang University, Hangzhou 310027, P. R. China, Department of Chemistry, Zhejiang University, Hangzhou 310027, P. R. China, and Institut Europe´ en des Membranes, UMR CNRS no. 5635, 34293 Montpellier Cedex 05, France Received June 22, 2004. In Final Form: August 25, 2004 Efforts have recently been made toward the study of interactions of phospholipid with various enzymes. It seems that phospholipids may be directly involved in regulating the enzyme activity. In this work, three phospholipid analogous polymers (PAPs), containing hydrophobic octyloxy, dodecyloxy, and octadecyloxy groups (abbreviated as 8-PAP, 12-PAP, and 18-PAP, respectively), were tethered on polypropylene hollow fiber microfiltration membrane (PPHFMM) to create a biocompatible interface for lipase immobilization. Lipase from Candida rugosa was immobilized on these PPHFMMs by adsorption. The adsorption capacity, activity, and thermal stability of enzyme on the PAP-modified PPHFMMs were compared with those of enzyme on the nascent ones. It was found that, as for the PAP-modified PPHFMMs, the adsorption capacities of lipase are lower than that of the nascent ones, while the activity retention of immobilized lipase increases from 57.5% to 74.1%, 77.5%, and 83.2% respectively for the 8-PAP-, 12-PAP-, and 18-PAP-modified PPHFMMs. In addition, the experimental results of thermal stability show that the residual activity of the immobilized lipase at 50 °C for 2 h is 62% for the 8-PAP-modified PPHFMM, 59% for the 12-PAPmodified PPHFMM, and 66% for the 18-PAP-modified PPHFMM, which are also higher than that of the nascent ones.

Introduction Lipases (EC 3.1.1.3) have gained considerable importance as versatile biocatalysts for the hydrolysis/synthesis of a wide range of esters and amides.1-3 A promising property of lipases is their activation in the presence of hydrophobic interface, which was first reported by Sarda and Desnuelle.4 Up to now, this lipase activation at interfaces has been recognized as a common feature. In the absence of interfaces, lipases have some elements of secondary structure (termed the “lid”) covering their active sites and making them inaccessible to substrates. However, in the presence of hydrophobic interfaces important conformational rearrangements take place, yielding the “open state” of lipases. These rearrangements result in the exposure of hydrophobic surfaces, the interaction with the hydrophobic interface, and the corresponding functionality on the enzyme. In this case, lipases seem to become strongly adsorbed to hydrophobic interfaces through a large hydrophobic surface that surrounds the catalytic site. This large hydrophobic surface involves residues from the internal face of the lid as well as from other protein chains. Therefore, in recent years, special emphasis has been paid to the selective adsorption of lipases on tailor-made strongly hydrophobic support surfaces.5-13 This immobilization procedure is based on the assumption that the large hydrophobic area that * Corresponding author: Fax ++ 86 571 8795 1773; e-mail [email protected]. † Institute of Polymer Science, Zhejiang University. ‡ Department of Chemistry, Zhejiang University. § Institut Europe ´ en des Membranes, UMR CNRS no. 5635. (1) Balca˜o, V. M.; Paiva, A. L.; Malcata, F. X. Enzyme Microb. Technol. 1996, 18, 392-416. (2) Jaeger, K.-E.; Reetz, M. T. TIBTECH 1998, 16, 396-403. (3) Paiva, A. L.; Balca˜o, V. M.; Malcata, F. X. Enzyme Microb. Technol. 2000, 27, 187-204. (4) Sarda, L.; Desnuelle, P. Biochim. Biophys. Acta 1958, 30, 513521.

surrounds the active site of lipases is the one mainly involved in their adsorption on strongly hydrophobic solid surfaces. Thus, lipases recognize these surfaces similarly to those of their natural substrates, and they suffer interfacial activation during immobilization. Main advantages for this immobilization method include (a) promoting a dramatic activation of lipases after their immobilization (that is, adsorbed lipases show very enhanced esterase activity in the absence of additional hydrophobic interfaces), (b) promoting a possibility to associate the immobilization with the purification of lipases, and (c) promoting a strong but reversible immobilization that enables us to recover these expensive supports after inactivation of immobilized lipases. Artificial membranes have been applied in biotechnology due to their interesting properties of high specific surface area and the possibility to combine separation with chemical reaction.14 The fact that lipases are activated in the presence of aqueous/ hydrophobic interfaces has interested membrane scientists and biotechnologists in their attempt to find polymeric membrane as an efficient (5) Fernandez-Lafuente, R.; Armise´n, P.; Sabuquillo, P.; Ferna´ndezLorente, G.; Guisa´n, J. M. Chem. Phys. Lipids 1998, 93, 185-197. (6) Wannerberger, K.; Arnebrant, T. Langmuir 1997, 13, 3488-3493. (7) Gunnlaugsdottir, H.; Wannerberger, K.; Sivik, B. Enzyme Microb. Technol. 1998, 22, 360-367. (8) Guisan, J. M.; Sabuquillo, P.; Fernandez-Lafuente, R.; FernandezLorente, G.; Mateo, C.; Halling, P. J.; Kennedy, D.; Miyata, E.; Re, D. J. Mol. Catal. B: Enzym. 2001, 11, 817-824. (9) Gill, I.; Pastor, E.; Ballesteros, A. J. Am. Chem. Soc. 1999, 121, 9487-9496. (10) Palomo, J. M.; Mun˜oz, G.; Ferna´ndez-Lorente, G.; Mateo, C.; Fernandez-Lafuente, R.; Guisa´n, J. M. J. Mol. Catal. B: Enzym. 2002, 19, 279-286. (11) Palomo, J. M.; Segura, R. L.; Fernandez-Lorente, G.; Pernas, M.; Rua, M. L.; Guisan, J. M.; Fernandez-Lafuente, R. Biotechnol. Prog. 2004, 20, 630-635. (12) Tsai, S.-W.; Shaw, S.-S. J. Membr. Sci. 1998, 146, 1-8. (13) Balcao, V. M.; Vieira, C.; Malcata, F. X. Biotechnol. Prog. 1996, 12, 164-172. (14) Gekas, V. C. Enzyme Microb. Technol. 1986, 8, 450-460.

10.1021/la0484624 CCC: $27.50 © 2004 American Chemical Society Published on Web 10/12/2004

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carrier for the immobilization of enzyme. Various membrane materials from hydrophilics to hydrophobics were reported in the literature.12,13,15-21 Among them, polypropylene membrane is more interesting15,16 due to its hydrophobicity, well-controlled porosity, and chemical inertness as well as high potentials for comprehensive applications. However, the poor biocompatibility of this membrane may cause nonbiospecific interaction, protein denaturation, and enzyme activity loss.22 Therefore, one can envisage that it is possible to introduce a biofriendly interface on the membrane surface for lipase immobilization through surface modification technologies, which may reduce some nonbiospecific enzyme-support interaction, create a specific microenvironment for the lipase, and benefit the enzyme activity. However, typical surface modification normally leads to the enhancement of both biocompatibility and hydrophilicity on the membrane surface. Simultaneous improvement of biocompatibility and hydrophobicity for the membrane surface were rarely described.23,24 It is well-known that various lipids, such as phospholipids and glycolipids, are the main components of cell membrane. Up to now, considerable interests have been paid to the study of lipid-protein interactions, which are of fundamental importance for both the structural integrity and the various functions of all biological membranes. In particular, the chemical composition and physical properties of the host lipid bilayer can markedly influence the activity, thermal stability, and the location and disposition of a large number of integral membrane proteins in biological membrane systems.25,26 Phospholipids, as an important category of lipid, have been proved to be inherently biocompatible with various proteins including enzymes.27-32 Recently, efforts have been made toward the study of interactions of phospholipid with various enzymes like phospholipase,28 protein kinase,29 glucose oxidase,30 and lipase.31,32 In some cases it has been shown that phospholipids may be directly involved in regulating the enzyme activity. In our previous work,33 a novel method for the surface modification of a polypro(15) Garcia, H. S.; Malcata, F. X.; Hill, C. G., Jr.; Amundson, C. H. Enzyme Microb. Technol. 1992, 14, 535-545. (16) Indlekofer, M.; Funke, M.; Claassen, W.; Reuss, M. Biotechnol. Prog. 1995, 11, 436-442. (17) Goto, M.; Goto, M.; Nakashio, F.; Yoshizuka, K.; Inoue, K. J. Membr. Sci. 1992, 74, 207-214. (18) Zbigniew, S. J. Membr. Sci. 1994, 97, 209-214. (19) Lozano, P.; Pe´rez-Marin, A. B.; De Diegoa, T.; Go´mezb, D.; Paolucci-Jeanjean, D.; Belleville, M. P.; Rios, G. M.; Iborra, J. L. J. Membr. Sci. 2002, 201, 55-64. (20) Sakaki, K.; Giorno, L.; Drioli, E. J. Membr. Sci. 2001, 184, 2738. (21) Arica, M. Y.; Kac¸ ar, Y.; Ergene, A.; Denizli, A. Process Biochem. 2001, 36, 847-854. (22) Kasemo, B. Surf. Sci. 2002, 500, 656-677. (23) Liu, Z.-M.; Xu, Z.-K.; Wang, J.-Q.; Yang, Q.; Wu, J.; Seta, P. Eur. Polym. J. 2003, 39, 2291-2299. (24) Deng, H.-T.; Xu, Z.-K.; Liu, Z.-M.; Wu, J.; Ye, P. Enzyme Microb. Technol., in press. (25) Sandermann, H. Acta 1978, 515, 209-237. (26) Lewis, R. N. A. H.; Zhang, Y. P.; Liu, F.; McElhaney, R. N. Bioelectrochemistry 2002, 56, 135-140. (27) Bakas, L. S.; Chazalet, M. S.-P.; Bernik, D. L.; Disalvo, E. A. Colloids Surf., B: Biointerfaces 1998, 12, 77-87. (28) Pete, M. J.; Exton, J. H. Biochim. Biophys. Acta 1995, 1256, 367-373. (29) Yu, J. S.; Chan, W. H.; Yang, S. D. Biochem. Biophys. Res. Commun. 1997, 237, 331-335. (30) Snejdarkova, M.; Rehak, M.; Babincova, M.; Sargent, D. F.; Hianik, T. Bioelectrochem. Bioenerg. 1997, 42, 35-42. (31) Wickham, M.; Wilde, P.; Fillery-Travis, A. Biochim. Biophys. Acta 2002, 1580, 110-122. (32) Cajal, Y.; Svendsenb, A.; Bolo´sa, J. De.; Patkarb, S. A.; Ascuncion, A. M. Biochimie 2000, 82, 1053-1061. (33) Xu, Z.-K.; Dai, Q.-W.; Wu, J.; Huang, X.-J.; Yang, Q. Langmuir 2004, 20, 1481-1488.

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pylene microfiltration membrane by tethering phospholipid analogous polymers (PAPs) was given, which includes the photoinduced graft polymerization of N,N-dimethylaminoethyl methacrylate (DMAEMA) and the ring-opening reaction of grafted poly(DMAEMA) with 2-alkyloxy2-oxo-1,3,2-dioxaphospholanes. It was found that these PAP-modified polypropylene microfiltration membranes exhibit excellent biocompatibility, and meanwhile some hydrophobic properties on the membrane surface were also retained after modification. In the present work, three PAPs, containing hydrophobic octyloxy, dodecyloxy, and octadecyloxy groups (respectively abbreviated as 8-PAP, 12-PAP, and 18-PAP), were tethered on a polypropylene hollow fiber microfiltration membrane (PPHFMM) to create a biocompatible interface for lipase immobilization (Figure 1). Lipase from Candida rugosa was immobilized on these membranes by adsorption. The adsorption capacity, activity, and thermal stability of immobilized lipases were investigated. It was envisaged that the comparison between the different supports for lipases immobilization may help to learn the influence of phospholipid layer on the membrane surface, especially the length of hydrophobic alkyloxy groups, and on the activity and stability of immobilized lipases. Experimental Section Materials. PPHFMM was prepared with melt-extruded/coldstretched (MECS) method in our lab.34,35 The inner and outer diameters of this hollow fiber were 240 and 290 µm, respectively, with porosity of 50% and an average pore diameter of 0.10 µm. Lipase (from Candida rugosa), Bradford reagent,36 and bovine serum albumin (BSA) were purchased from Sigma and used as received. Other reagents were AR grade and purified following normal procedures before use. Preparation and Characterization of PAP-Modified Membranes. The preparation and characterization of PAPmodified PPHFMMs followed the process reported in our previous paper.33 Polypropylene microfiltration membranes with almost the same grafting degree of PAP were also fabricated for the measurements of water contact angles. Static Water Contact Angle Measurements. The static water contact angles of the membrane surface were measured by the sessile drop method and the captive bubble method at 25 °C with a contact angle goniometer (KRUSS DSA10-MK, Germany) equipped with video capture. At least 10 contact angles were averaged to get reliable data, and the standard error for the result was below 5% of the average value. Immobilization of Lipase by Adsorption. The adsorption immobilization of lipase refers to the literature.37 Lipase solutions (0.25-2.50 mg/mL) were prepared by adding appropriate amounts of lipase powder to phosphate buffer (0.05 M, pH 7.0). A bundle of hollow fiber membranes (145 fibers, 7.5 cm long) was submerged in 10 mL of lipase solution in a vertical orientation and shaken gently in a water bath at 30 °C for 3 h. The ratio of the area of membranes to the volume of lipase solution is 10 cm2/cm3, which can ensure the significant changes in the concentration of lipase solution after adsorption process and make all of the membrane surface contact solution completely during the adsorption process. Finally, the membranes were taken out and rinsed with buffer until no soluble protein was detectable in washings. Protein concentration in solutions was determined with Coomassie Brilliant Blue reagent following Bradford’s method.36 Bovine serum albumin (BSA) was used as standard to construct the calibration curve. The amount of adsorbed protein on the membranes was calculated from the protein mass balance among the initial and final lipase solution and washings. The (34) Xu, Z.-K.; Wang, J.-L.; Shen, L.-Q.; Meng, D.-F.; Xu, Y.-Y. J. Membr. Sci. 2002, 196, 221-229. (35) Xu, Z.-K.; Dai, Q.-W.; Liu, Z.-M.; Kou, R.-Q.; Xu, Y.-Y. J. Membr. Sci. 2003, 214, 71-81. (36) Bradford, M. Anal. Biochem. 1976, 72, 248-254. (37) Malcata, F. X.; Garcia, H. S.; Jr, C. G. H. Biotechnol. Bioeng. 1992, 39, 647-657.

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Figure 1. Schematic representative for the preparation of phospholipid analogous polymer-tethered PPHFMMs. lipase adsorption capacity of the membranes was defined as the amount of protein (mg) per square meter of surface areas of the hollow fiber membranes (including outer and inner surface). The reported values were the mean of at least three experiments, and the standard deviation was within ca. (5%. Desorption Experiments. The lipase-immobilized PPHFMMs (145 fibers, 7.5 cm long) were incubated in 10 mL of phosphate buffer (0.05 M, pH 7.0) and shaken in a water bath at 30 °C for 24 h. Supernatant samples were removed at intervals, and protein content was analyzed as described above. The desorption ratios of lipases was calculated by the following expression:

desorption ratio ) [(amount of lipases released) × 100]/ [amount of lipases adsorbed on the membranes] The reported values were the mean of at least three experiments, and the standard deviation was within ca. (5%. Activity Assay of Free and Immobilized Lipases. The pH stat method with olive oil titrimetric assay was used in this work. The lipase-immobilized PPHFMMs were cut into short segments to make them disperse well in the emulsion during assay process. The substrate emulsion was prepared by thoroughly mixing 130 mL of olive oil with 400 mL of gum arabic solution (11% gum arabic powder and 1.25% CaCl2‚2H2O, m/v) and stored at 4 °C. 24 mL of substrate emulsion was mixed with 9 mL of deionized water and 2 mL of sodium taurocholate solution (0.5%, m/v). The emulsion was incubated in a water bath at a certain temperature for several minutes, and then pH was adjusted to the desired

value with NaOH solution. One milliliter of lipase solution (1 mg/mL) or suitable amounts of the lipase-immobilized membranes was added into the emulsion. The pH was held constant for 10 min by continuously adding 0.01 M NaOH standard solution. The consumed volume of NaOH standard solution was recorded. The blank value in the absence of lipase-immobilized PPHFMMs was measured by the same way. One lipase unit corresponded to the release of 1 µmol of fatty acid per minute under the assay conditions. Specific activity was defined as the number of lipase unit per milligram of protein. Activity retention value was the ratio of specific activity of immobilized lipase with that of free lipase. Each data was the average of three parallel experiments at least. The standard error for the result was below 5% of the average value. Thermal Stability. Free and immobilized lipase preparations were stored in phosphate buffer (0.05 M, pH 7.0) at 50 °C for 2 h. Samples were periodically withdrawn for activity assay. The residual activities were determined as above.

Results and Discussion Adsorption/Desorption Behaviors of Lipase on the PAP-Modified PPHFMMs. Figure 2 shows the adsorption isotherm of lipase on the nascent and PAP-modified PPHFMMs. It can be seen that, for each membrane support, the increment of protein concentration enhances the driving force for the adsorption, which, in turn, increases the adsorbed amount of protein until a plateau value was observed. This plateau value of each membrane

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Table 1. Comparison of Adsorption Capacity and Activity of Lipases on the Nascent, 8-PAP-Modified, 12-PAP-Modified, and 18-PAP-Modified PPHFMMS water contact angle (deg)a membrane

grafting degree (wt %)

sessile-drop method

captive-bubble method

adsorbed protein (mg/m2)

specific activity (U/mg protein)

activity retention (%)

nascent PPHFMM 8-PAP-modified PPHFMM 12-PAP-modified PPHFMM 18-PAP-modified PPHFMM

0 10.6 10.9 10.8

108.4 ( 1.8 53.2 ( 1.3 92.4 ( 1.5 96.2 ( 1.1

105.1 ( 2.4 38.1 ( 0.9 39.3 ( 1.1 38.5 ( 0.8

45.3 ( 1.1 41.3 ( 1.4 41.8 ( 1.3 39.5 ( 1.5

69.9 90.1 94.3 101.2

57.5 ( 2.8 74.1 ( 3.2 77.5 ( 3.7 83.2 ( 3.3

a

Water contact angle was measured on sheet membrane with almost the same grafting degree of PAP. Table 2. Effect of Grafting Degree of 18-PAP on the Adsorption Capacity and Activity of Immobilized Lipases

Figure 2. Effect of initial protein concentration on the adsorbed amount of protein: (2) nascent PPHFMM; (O) 10.6 wt % 8-PAPmodified PPHFMM; (4) 10.9 wt % 12-PAP-modified PPHFMM; (b) 10.8 wt % 18-PAP-modified PPHFMM.

Figure 3. Effect of time on desorption ratio: (2) nascent PPHFMM; (O) 10.6 wt % 8-PAP-modified PPHFMM; (4) 10.9 wt % 12-PAP-modified PPHFMM; (b) 10.8 wt % 18-PAPmodified PPHFMM.

is listed in Table 1. The data indicate that the modifying process decreases the adsorbed amount of protein to some extent. The reduced adsorption strength after modification suggested by the observed results needs to be confirmed through desorption experiments. The plot of desorption ratio as a function of time is shown in Figure 3. The data were obtained from the protein concentration in phosphate buffer after the lipase-immobilized membranes were incubated in this buffer solution at 30 °C. In that case, only the lipase adsorbed weakly on the membranes can be desorbed. It was found that the time needed to reach about 6% desorption ratio is 8 h for the nascent PPHFMM, while the period prolongs to about 12 h for the PAPmodified PPHFMMs. Furthermore, the desorption ratio at desorption/adsorption equilibrium is about 10.0%, 8.6%, 8.8%, and 8.1% for the nascent PPHFMM, the 8-PAP-

grafting degree (wt %)

adsorbed protein (mg/m2)

specific activity (U/mg protein)

activity retention (%)

4.2 6.7 8.5 10.8

43.3 ( 1.6 41.1 ( 1.4 41.4 ( 1.5 39.5 ( 1.5

91.5 96.9 98.4 101.2

75.3 ( 3.9 79.7 ( 3.9 80.9 ( 4.1 83.2 ( 3.3

modified PPHFMM, the 12-PAP-modified PPHFMM, and the 18-PAP-modified PPHFMM, respectively. On the basis of these results, it can be concluded that the introduction of PAP on the membrane surface significantly enhances the adsorption strength for lipase. This adsorption strength can be attributed to the electrostatic and hydrophobic interaction of lipase protein respectively with the zwitterion and the long alkyloxy moieties of PAP. As for the decreased adsorption capacity after modification, it can be ascribed to the partial block of the membrane pores and the reduction of the available adsorption area on the membrane surface. In fact, similar results were also observed in our previous surface-modified PPHFMMs.38 In addition, with increasing the grafting degree of 18PAP, the decreasing trend of adsorption capacity can also be observed in Table 2. This phenomenon to some extent confirms the above interpretation. Effect of Surface Modification on the Activity of Lipase. The activities of lipases immobilized on the nascent and the PAP-modified PPHFMMs are compared in Table 1. It can be seen that the activity retention of lipase immobilized on the nascent PPHFMM is 57.5%. This can be ascribed to the hydrophobic characteristic of the membrane surface. The importance of hydrophobic interface in promoting lipase adsorption and inducing conformational change, which opens the active site of the enzyme, is well recognized.4-13 However, the activity retention of lipase increases significantly from 57.5% for the nascent PPHFMM to 74.1%, 77.5%, and 83.2% respectively for the 8-PAP-, 12-PAP-, and 18-PAP-modified PPHFMMs. Some activity data of immobilized lipases were reported by other researchers.39,40 Among them, on account of the diversity of enzyme sources and the activity assay methods, the specific activity (U/mg protein) is hard to be comparable, but the value of activity retention can effectively reflect the capacity of support for enzyme immobilization. Compared with previously reported values,39,40 the activity retention of 83.2% in this case should be a rather encouraging result. One possible interpretation for this pronounced improvement of activity may be that the layer formed by the phospholipid analogues introduces a biofriendly microenvironment for the immobilized (38) Deng, H.-T.; Xu, Z.-K.; Wu, J.; Ye, P.; Liu, Z.-M.; Seta, P. J. Mol. Catal. B: Enzym. 2004, 28, 95-100. (39) Montero, S.; Blanco, A.; Virto, M. D.; Landeta, L. C.; Agud, I.; Solozabal, R.; Lascaray, J. M.; Renobales, M.; Llama, M. J.; Serra, J. L. Enzyme Microb. Technol. 1993, 15, 239. (40) Trusek-Holownia, A.; Noworyta, A. Desalination 2002, 144, 427432.

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lipases, which is analogous to that in the natural biological membrane systems. In addition, it is interestingly observed that, at basically the same grafting degree of PAP, the more carbon atoms in the long alkyloxy group of PAP, the higher activity increase can be obtained, which indicates the important participation of the hydrophobic alkyloxy moieties in the interfacial activation of lipase. The long hydrophobic alkyloxy groups seem to offer strong hydrophobic interaction with the hydrophobic domain around the active center of lipase, which stabilizes the “open state” conformation of lipase and favors the active site’s accessibility to substrates. Furthermore, the data in Table 2 show that the activity of immobilized lipases increases with increasing the grafting degree of 18-PAP. It may be due to the high density of octadecyloxy groups dispersing on the membrane surface, reinforcing the interfacial activation of lipase. As for the sequence of activity increase of the three PAP-modified PPHFMMs, it may be elucidated by taking into account the conformational property of PAP on the membrane surface. As reported in our previous paper,33 when the PAP-modified membrane was washed with THF and dried after the surface-modifying process, the long alkyloxy groups fingered out, and a hydrophobic upper surface was formed spontaneously on the membrane surface. Afterward, in the enzyme-immobilizing process, the membrane was thoroughly immersed in the phosphate buffer solution of lipase. This highly hydrophilic environment with the ionic strength action of phosphate, to some extent, can induce the conformational turning-over between the moieties of zwitterion and the moieties of long alkyloxy group, which made parts of the zwitterion moieties reach the upper surface of the membrane and caused the imperfection of the hydrophobic alkyloxy groups surface on the membrane. Comparing the three PAP-modified PPHFMMs, the PAP with short alkyloxy groups seems to be more prone to this conformational change due to the low steric hindrance during conformational turning-over. It means that the long alkyloxy groups surface on the membrane is more stable and perfect than the short one in the hydrophilic external environment, which can offer a more effective interfacial activation for the immobilized lipase. In an attempt to prove this assumption, the water contact angle of the PAP-modified polypropylene microfiltration membrane (PPMM) in sheet form was measured by both the sessile drop method and the captive bubble method, and the results are listed in Table 1. It can be seen that, for the PAP-modified membrane, the contact angle measured by the captive bubble method in water is much lower than that measured by the sessile drop method in air, but the difference for the nascent membrane can be omitted. This difference between the two measuring methods is a typical phenomenon caused by the conformation alteration of the hydrophilic macromolecular chains.41,42 It means that in the captive-bubble method the long-time contact of PAP-modified membrane with water results in a conformational turning-over between the moieties of zwitterion and the moieties of long alkyloxy group of PAP. Furthermore, as can be seen from Table 1, the water contact angle measured by sessile drop method increased in the sequence of 18-PAP-modified PPMM > 12-PAP-modified PPMM > 8-PAP-modified PPMM, consistent with the sequence of activity increase, which seems to support the above assumption. Therefore, on the basis of the above discussion, it is conceivable that the different capability of conformational (41) Yasuda, H.; Sharma, A. K.; Yasuda, T. J. Polym. Sci., Polym. Phys. Ed. 1981, 19, 1285-1291. (42) Tretinnikov, O. N.; Ikada, Y. Langmuir 1994, 10, 1606-1614.

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Figure 4. Effect of pH on relative activity: (9) free lipase; (2) nascent PPHFMM; (O) 10.6 wt % 8-PAP-modified PPHFMM; (4) 10.9 wt % 12-PAP-modified PPHFMM; (b) 10.8 wt % 18PAP-modified PPHFMM.

change of three PAPs is the major cause for the sequence of activity increase. In addition, the hydrophobic alkyloxy groups of PAP with different length can somehow induce hydrophobic substrates diffusing to support through hydrophobic interaction, which seems to contribute to the above sequence of activity increase too. Furthermore, the long and stretched alkyloxy moieties can function as a “spacer” to some extent and hereby extend the distance from lipase to support, which may be another reason. It is necessary to mention that, although many literatures verified the hydrophobic interfacial activation of immobilized lipases, a more hydrophobic interface does not always means higher activity obtained for the enzyme. In other words, the moderate hydrophilicity/hydrophobicity of support surface will benefit the improvement of lipases’ activity.6,7,43 In this work, with respect to the data of water contact angle, despite the decreased surface hydrophobicity caused by surface modification, the pronounced improvement of lipases’ activity was still observed for PAP-modified PPHFMMs. In addition to the effective participation of the stretched-out hydrophobic alkyloxy moieties in the interfacial activation of lipase mentioned above, another possible interpretation for this result may be that the layer by the phospholipid analogues introduces a biofriendly microenvironment for the immobilized lipases, analogous to that in the natural biological membrane systems, which contributed to the reduction of some nonbiospecific interaction between supports and immobilized enzymes. It is the major reason for the fact that a large number of materials modified by phospholipid bilayer and lipid vesicle are used extensively as supports for enzyme immobilization.30-32,44,45 On the other hand, some researchers think that electrostatic interaction also favors the stabilization of “open state” conformation of lipases to some extent.32 In this case, the possibility of partial participation of electrostatic interaction in interfacial activation should not be excluded, although it is secondary to the hydrophobic interaction on account of the interfacial activation mechanism of lipases. Effect of pH and Temperature on the Lipase Activity. Figure 4 shows the effect of pH on the activity of the free and the immobilized lipases. It is found that the optimum pH value for the free lipase is about 7.7, (43) Wannerberger, K.; Welin-Klintstrom, S.; Arnebrant, T. Langmuir 1997, 13, 784-790. (44) Phadtare, S.; Parekh, P.; Gole, A.; Patil, M.; Pundle, A.; Prabhune, A.; Sastry, M. Biotechnol. Prog. 2002, 18, 483-488. (45) Walde, P.; Ichikawa, S. Biomol. Eng. 2001, 18, 143-177.

Candida rugosa Lipase

Figure 5. Effect of temperature on relative activity: (9) free lipase; (2) nascent PPHFMM; (O) 10.6 wt % 8-PAP-modified PPHFMM; (4) 10.9 wt % 12-PAP-modified PPHFMM; (b) 10.8 wt % 18-PAP-modified PPHFMM.

whereas those for the immobilized lipases shift to the alkaline region at about 8.5, 8.3, 8.7, and 8.5 respectively for the nascent, 8-PAP-modified, 12-PAP-modified, and 18-PAP-modified PPHFMMs. It can be explained as that upon immobilization the active site becomes more exposed to solvent than that in the globular, folded, dissolved lipase form; therefore, proton transfer to the amino acid residues at the active site becomes less hindered.46 As shown in Figure 5, the immobilization makes the optimum temperature for lipase activity shifts from about 35 °C of the free enzyme to 40, 43, 43, and 45 °C of the enzymes immobilized on the nascent, 8-PAP-modified, 12PAP-modified, and 18-PAP-modified PPHFMM, respectively. These results can be attributed to a low restriction in the diffusion of the substrate and products at higher reaction temperature. In addition, the improved resistance of protein to thermal denaturation is also an important factor. Thermal Stability. Figure 6 shows the thermal stability of free and immobilized lipases studied in this work. It can be seen that free lipase lost all its initial activity within about 100 min. Lipase adsorbed on the 18-PAP-modified PPHFMM preserves initial activity about 66% in 2 h, which is higher than that of lipase immobilized on the nascent membrane (37%). In addition, the residual activity of 62% and 59% can also be achieved respectively by 8-PAP- and 12-PAP-modified PPHFMM under the same conditions. The improved thermal stability of lipases on PAP-modified PPHFMMs can be explained reasonably as that the interaction of lipases with PAPs stabilizes the conformation of enzyme protein and im(46) Duinhoven, S.; Poort, R.; Van der Voet, D.; Agterof, W. G. M.; Rorde, W.; Lyklema, J. J. Colloid Interface Sci. 1995, 170, 340-350.

Langmuir, Vol. 20, No. 23, 2004 10173

Figure 6. Effect of time on residual activity: (9) free lipase; (2) nascent PPHFMM; (O) 10.6 wt % 8-PAP-modified PPHFMM; (4) 10.9 wt % 12-PAP-modified PPHFMM; (b) 10.8 wt % 18PAP-modified PPHFMM.

proves the resistance of protein to thermal denaturation. However, with respect to the sequence of activity increase, no similar law is observed in the comparison of the thermal stability of three modified membranes. It may be due to the fact that not only hydrophobic but also electrostatic interaction between lipases and PAPs is involved in the stabilization of enzyme protein conformation. Conclusions Phospholipid analogous polymers, containing octyloxy, dodecyloxy, and octadecyloxy groups, were tethered on a PPHFMM to create a biocompatible interface for lipase immobilization. The partial block of the membrane pores caused by surface modification results in the decreased adsorption capacity. In addition, as a result of the stabilizing action of hydrophobic alkyloxy groups to the “open state” conformation of lipase and the excellent biocompatibility of phospholipid analogues layer, tethering PAP on the membrane surface increases the activity retention of immobilized lipase from 57.5% to 74.1%, 77.5%, and 83.2% respectively for 8-PAP-, 12-PAP-, and 18-PAP-modified PPHFMM. Furthermore, due to the fact that the electrostatic and hydrophobic interaction between lipase and PAP stabilized the conformation of enzyme protein, improved thermal stability was also obtained by the PAP-modified PPHFMMs. Acknowledgment. The authors are grateful for the financial support from the National Natural Science Foundation of China (Grants 20074033 and 50273032), the National Basic Research Program of China (Grant 2003CB15705), and the Programme Sino-Franc¸ ais de Recherches Avance´es (Grant PRA E03-04). LA0484624