Communication

1341

DOI: 10.1002/marc.200600266

Summary: PANCMPCs containing phospholipid side moieties were electrospun into nanofibers with a mean diameter of 90 nm. Field emission SEM was used to characterize the morphologies of the nanofibers. These phospholipid-modified nanofibers were explored as supports for enzyme immobilization due to the characteristics of excellent biocompatibility, high surface/volume ratio, and porosity, which were beneficial to the catalytic efficiency and activity of immobilized enzymes. Lipase from Candida rugosa was immobilized on these nanofibers by adsorption. Preliminary results indicated that the properties of the immobilized lipase on these phospholipid-modified nanofibers were greatly promising.

Schematic representation of the structure and electrostatic properties of phospholipid-modified nanofibers.

Electrospun Nanofibers Modified with Phospholipid Moieties for Enzyme Immobilization Xiao-Jun Huang,1 Zhi-Kang Xu,*1 Ling-Shu Wan,1 Christophe Innocent,2 Patrick Seta2 1

Institute of Polymer Science, and Key Laboratory of Macromolecule Synthesis and Functionalization (Ministry of Education), Zhejiang University, Hangzhou 310027, China Fax: þ86 571 8795 1773; E-mail: [email protected] 2 Institute of Europe´e des Membranes, UMR CNRS no. 5635, 34293 Montpellier Cedex 05, France

Received: April 18, 2006; Revised: May 30, 2006; Accepted: June 1, 2006; DOI: 10.1002/marc.200600266 Keywords: biomimetic polymer; electrospinning; enzyme immobilization; lipase; nanofibers; phospholipids

Introduction In recent years, electrospinning has gained widespread attention since it is known to be an effective fabrication tool for preparing fibrous materials with diameter ranging from several micrometers down to tens of nanometers.[1–20] Among various polymers, acrylonitrile-based homopolymers and copolymers were most recently fabricated into nanofibrous materials with reinforcing superhydrophobic and/or catalytic properties.[6a,9–20] In our previous work, novel nanofibers possessing reactive carboxyl groups were fabricated from poly[acrylonitrile-co-(maleic acid)] and poly[acrylonitrile-co-(acrylic acid)] by the electrospinning process.[18,19] Lipase and catalase were covalently immobilized onto this nanofibers respectively via the activation of carboxyl groups. We found that the enzyme loading and the activity retention of immobilized enzymes on the nanofibers fabricated by electrospinning were much higher than those on the corresponding hollow fibers prepared by typical phase inversion process.[18] Thus, the process described in our work presents a convenient approach to

Macromol. Rapid Commun. 2006, 27, 1341–1345

fabricate nanofibers with reactive groups for enzyme immobilization. These enzyme-immobilized nanofibers with high enzyme loading and bioactivity may have great potentials in the fields of biocatalysts for polymer synthesis, in situ formation of nanofiber reinforcement composites, biosensors, and biocatalysis/separation.[21] However, to ensure the immobilized enzymes with both high activity and stability, surface biocompatibility for the nanofiber is also one important requirement, as the biocompatible surface of support can reduce some non-biospecific enzyme-support interactions, create a specific microenvironment for the enzyme, and thus benefit the enzyme activity.[18b] Phospholipids, as the principal components of natural biomembranes, have been proved to be inherently biocompatible with various proteins including enzymes.[22–30] Especially, polymer surfaces modified with phospholipid analogs have been shown to reduce protein adsorption significantly as these hydrated surfaces are able to interact with proteins without inducing conformational changes in their three-dimensional structures. Therefore, phospholipid moieties have often been incorporated into main chains or

ß 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

1342

X.-J. Huang, Z.-K. Xu, L.-S. Wan, C. Innocent, P. Seta

side groups to synthesize biomimetic polymers.[22–28] In our previous work,[28a] phospholipid-modified polyacrylonitriles (PAN) were synthesized by the copolymerization of acrylonitrile and 2-methacryloyloxyethyl phosphorylcholine (MPC). The zwitterionic phospholipid moieties could be rearranged on the polymer surface in an aqueous environment. Phospholipid moieties were also directly formed on the film/membrane surface of poly[acrylonitrileco-(2-hydroxyethyl methacrylate)] through chemical reactions.[28b,c] Results indicated that the surface biocompatibility of PAN films/membranes could be greatly improved by the incorporation of MPC into polymer chains or the tethering of phospholipid moieties on the surfaces.[28] In this work, electrospun nanofibers were fabricated from poly[acrylonitrile-co-(2-methacryloyloxyethyl phosphorylcholine)]s (PANCMPCs) with different contents of MPC. The phospholipid-modified nanofibers were expected to be attractive candidates for enzyme immobilization due to the potential advantages including: (1) the biocompatibility interface created by phospholipid moieties on the nanofiber surface will reduce some non-biospecific enzyme-support interactions, create a specific microenvironment for the enzyme, and thus benefit the enzyme activity; (2) the large specific surface will provide relatively high quantity of enzyme loading per unit mass; (3) the fine porous structure will entrust the accessibility of active sites and low diffusion

resistance necessary for high reaction rate and conversion; and (4) the easy recoverability from reaction media or applicability will be favorable for continuous operations.[18,19,21,31] Herein, one of the most exploited enzymes, lipase from Candida rugosa, was chosen as a model enzyme and immobilized on these nanofibers by physical adsorption (Figure 1).

Experimental Part MPC was synthesized according to the method of Ishihara.[32] PAN and PANCMPCs with different MPC contents, designated as PANCMPC05 and PANCMPC10 in the following text, were synthesized in our laboratory using a water-phase precipitation copolymerization process.[28] The designations 05 and 10 indicated that MPC content in the copolymers was 4.7 and 9.6 mol-%, respectively. Dimethyl sulfoxide (DMSO) was purified by vacuum distillation before use. Lipase (from Candida rugosa, protein concentration was 6.8 wt.-%), Bradford reagent, bovine serum albumin (BSA, M w ¼ 67 000 Da), and p-nitrophenyl palmitate (p-NPP) were purchased from Sigma and used as received. Other reagents were of analytical grade without further purification. To fabricate nanofibers with the electrospinning process, PAN and PANCMPCs were dissolved in DMF at room temperature with gentle stirring for 12 h to form 8 wt.-% homogeneous solutions. After air bubbles were removed completely, each solution was placed in a syringe (50 mL) bearing 1 mm

Figure 1. Schematic representation for the fabrication of phospholipid-modified nanofibers by electrospinning process for lipase immobilization. Macromol. Rapid Commun. 2006, 27, 1341–1345

www.mrc-journal.de

ß 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

1343

Electrospun Nanofibers Modified with Phospholipid Moieties for Enzyme Immobilization

inner diameter metal needles which was connected with a high voltage power supply (GDW-a, Tianjin Dongwen Highvoltage Power Supply Plant, China). The grounded counter electrode was connected to the aluminum foil collector. Typically, electrospinning was performed at 10 kV voltage, 15 cm distance between the needle tip and the collector. The flow rate of the solution was controlled by a microinfusion pump (WZ-50C2, Zhejiang University Medical Instrument Co., Ltd., China) to maintain at 2 mL
h1 from the needles outlet. It usually took half an hour to obtain a sufficiently thick sample that could be detached from the aluminum foil collector. The nanofibers on aluminum foil were dried under vacuum at 60 8C before it was detached. The morphology and fiber diameter of the electrospun nanofibers were obtained by sputter-coating with gold and examining under a field emission scanning electron microscope (FESEM). Lipase solution (5.0 mg
mL1) was prepared by adding appropriate amount of lipase powder to phosphate buffer (0.05 M, pH 7.0). A designed weight of nanofibers was submerged in 10 mL of lipase solution in a vertical orientation and shaken gently in a water bath at 30 8C for 3 h. Then, the nanofibers were taken out and rinsed with buffer until no soluble protein was detectable in washings. Protein concentrations in solutions were determined with Coomassie Brilliant Blue reagent following Bradford’s method.[33] BSA was used as standard to construct the calibration curve. The amount of adsorbed protein on the nanofibers was calculated from the protein mass balance among the initial and final lipase solution and washings. The lipase adsorption capacity of the nanofibers was defined as the amount of protein (mg) per gram of the nanofibers. Each reported value was the mean of at least three experiments, and the standard deviation was within ca. 5%. The activity of the immobilized lipase on the nanofibers in aqueous medium was determined according to the method reported previously.[18]

Results and Discussion Acrylonitrile-based homopolymers and copolymers were most recently fabricated into nanofibers with versatile properties for diverse applications.[9–20] Several factors such as the molecular weight of polymer, the concentration of solution, the flow rate of fluid, the strength of electric field, and the distance from the needle tip to the collector significantly influenced the size and morphology of the resultant fibers. In this work, PAN and PANCMPCs with different MPC contents were synthesized by a waterphase precipitation copolymerization process and then electrospun into nanofibers. When the MPC content in PANCMCPs was not very high, the characteristics of PANCMPCs solution were similar to that of PAN. Thus, phospholipid-modified nanofibers could be easily prepared by the electrospinning process. The morphology of the resulted nanofibers can be seen in Figure 2 and the diameters of the nanofibers are summarized in Table 1. As shown in Figure 2, when the molecular weight of the polymer was about 100 000 g
mol1 and the solution concenMacromol. Rapid Commun. 2006, 27, 1341–1345

www.mrc-journal.de

Figure 2. Electrospun nanofibers from 8 wt.-% PAN (a), PANCMPC05 (b), and PANCMPC10 (c) solutions.

tration was 8 wt.-%, an almost homogenous network of the electrospun fibers with diameter distribution between 70 and 120 nm was obtained [Figure 2(a–c)]. Following our previous results,[28] we reasonably deduced that these ß 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

1344

X.-J. Huang, Z.-K. Xu, L.-S. Wan, C. Innocent, P. Seta

Table 1.

Typical characteristics of the polymer and nanofibers.

Polymer

PAN PANCMPC05 PANCMPC10 a) b)

M w a)  104

MPC content in PANCMPCb)

Fiber diameter

g
mol1

mol-%

nm

10.3 9.6 9.3

– 4.7 9.6

100  20 90  20 90  20

The value of M w was derived from viscosity measurement. The content of MPC in PANCMPCs was calculated from 1 H-NMR.

phospholipid-modified nanofibers should show excellent biocompatibility and therefore might be used as enzyme immobilization carriers for biological catalysis/separation. Enzyme immobilization has been a popular strategy for most large-scale applications due to the ease in catalyst recycling, continuous operation, and product purification. Poor biocatalytic efficiency of immobilized enzymes, however, often limits the development of large-scale bioprocessing to compete with traditional chemical processes. Improvements of biocatalytic efficiency can be achieved by manipulating the structure of carrier materials for enzyme immobilization. Non-porous materials, to which enzymes are attached to the surfaces, are subjected to minimum diffusion limitation while enzyme loading per unit mass of support is usually low. On the other hand, porous materials can afford high enzyme loading, but suffer a much greater diffusional limitation of substrate. Reduction in the size of enzyme-carrier materials can generally improve the efficiency of immobilized enzymes. Electrospun nanofibers provide a large surface area for the attachment or entrapment of enzymes and the enzyme reaction. In the case of porous nanofibers, they can still reduce the diffusional path of substrate from the reaction medium to the enzyme active sites due to the reduced dimension in size. Electrospun nanofiber membranes are durable and easily separable, and can also be processed in a highly porous form for relieved mass-transfer of substrate through the membranes. Therefore, lipase from Candida rugosa was physically adsorbed

on the phospholipid-modified nanofibers. The properties of the free and immobilized lipases under the optimum conditions are shown in Table 2. It was found that the amount of adsorbed lipase on the nanofibers was up to about 22.0 mg
g1, corresponding to over ten times of that on the corresponding sheet membrane fabricated by casting.[28c] This result was ascribed to the large surface area/mass ratio of the nanofibers. On the other hand, the activity retention of the immobilized lipase on the PAN nanofibers was also higher than that on the PAN sheet membrane. The main reason for this could be the remarkable reduction of the diffusion resistance for the immobilized lipase on this nanofibers. Compared to that on the PAN nanofibers, the activity retention of lipase increased significantly from 56.4% on the PAN ones to 72.9 and 76.8%, respectively on the PANCMPC05 and PANCMPC10 nanofibers. This pronounced improvement of activity might be that the phospholipid moieties on the surface of nanofibers introduced a biofriendly microenvironment for the immobilized lipase, analogous to that in the natural biological membrane systems, could reduce some non-biospecific interaction between the nanofibers and immobilized lipase. During the lipase-immobilizing process, the nanofibers were thoroughly immersed in the phosphate buffer solutions of lipase. This highly hydrophilic environment with the ionic strength action of phosphate made the zwitterionic phospholipid moieties which were previously ‘‘buried’’ inside or beneath the surface layer to reorient on the upper surface of the nanofibers. With the increase of MPC content, a more stable, perfect hydrophilic and biocompatible external microenvironment was formed on the surface of nanofibers, which offered a more effective interfacial activation for the immobilized lipase. Therefore, the activity of the immobilized lipase increased with the increase of the phospholipid content in the PAN-based nanofibers. Table 2 lists the kinetic parameters Km and Vmax from a double reciprocal plot.[34] Results indicated that the Km values for the immobilized lipase on the nanofibers were obviously lower than that on the PAN sheet membrane. It demonstrated that the diffusion resistance was remarkably reduced by the reduction of the geometric size of the lipase support, in turn, the catalytic efficiency of the immobilized

Table 2. Adsorption capacity, activity, and kinetic parameters for immobilized lipases on the electrospun nanofibers and the PAN sheet membrane. Sample

Adsorbed protein mg
g

1

Free lipase Lipase immobilized on PAN sheet membranea) Lipase immobilized on PAN nanofibers Lipase immobilized on PANCMPC05 nanofibers Lipase immobilized on PANCMPC10 nanofibers a)

– 2.1  0.15 23.2  1.6 22.2  1.7 22.9  1.5

Specific activity U
mg

1

42.1 21.3 23.7 30.7 32.3

Activity retention % 100 50.6  0.9 56.4  0.7 72.9  0.8 76.8  0.6

Vmax U
mg

1

44.6 23.8 26.6 32.4 34.0

Km 103 M 0.44 1.64 0.79 0.88 0.86

PAN sheet membrane was prepared by casting method according to our previous work.[25]

Macromol. Rapid Commun. 2006, 27, 1341–1345

www.mrc-journal.de

ß 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

1345

Electrospun Nanofibers Modified with Phospholipid Moieties for Enzyme Immobilization

lipase was effectively improved. On the other hand, the Vmax values of the immobilized lipase on the PANCMPC nanofibers were obviously higher than that on the PAN ones. This result could also be ascribed to the biofriendly microenvironment for the immobilized lipase created by the phospholipid moieties on the upper surface of nanofibers.

[7] [8] [9] [10] [11]

Conclusion In this communication, we describe the preliminary preparation of phospholipid-modified nanofibers by an electrospinning process. PANCMPCs with different contents of MPC, as well as PAN, were fabricated into homogenous fibers with diameter in the nanometer range. In parallel, lipase from Candida rugosa was immobilized on these nanofibers and an effective enhancement on the efficiency of immobilized enzyme was realized. The nanofibers with excellent biocompatible phospholipid surfaces could create a suitable interface for the immobilized enzyme.

[12] [13] [14] [15]

[16] [17]

Acknowledgements: Financial supports from the National Natural Science Foundation of China (Grant no. 50273032) and the Programme Sino-Franc¸ais de Recherches Avance´es (No. PRA E03-05) are gratefully acknowledged.

[18]

[19] [20] [1] Y. Dzenis, Science 2004, 304, 1917. [2] D. Li, Y. Xia, Adv. Mater. 2004, 16, 1151. [3] Z. M. Huang, Y. Z. Zhang, M. Kotaki, S. Ramakrishna, Compos. Sci. Technol. 2003, 63, 2223. [4] [4a] L. M. Bellan, G. W. Coates, H. G. Craighead, Macromol. Rapid Commun. 2006, 27, 511; [4b] X. Lu, Q. Zhao, X. Liu, D. Wang, W. Zhang, C. Wang, Y. Wei, Macromol. Rapid Commun. 2006, 27, 430; [4c] Z. Li, H. Huang, C. Wang, Macromol. Rapid Commun. 2006, 27, 142; [4d] J. Li, A. He, C. C. Han, D. Fang, B. S. Hsiao, B. Chu, Macromol. Rapid Commun. 2006, 27, 114; [4e] H. S. Kim, H.-J. Jin, S. J. Myung, M. Kang, I.-J. Chin, Macromol. Rapid Commun. 2006, 27, 146. [5] [5a] S. Y. Gu, Q. L. Wu, J. Ren, G. J. Vancso, Macromol. Rapid Commun. 2005, 26, 716; [5b] G. M. Kim, R. Lach, G. H. Michler, Y. W. Chang, Macromol. Rapid Commun. 2005, 26, 728; [5c] M. Wei, J. Lee, B. Kang, J. Mead, Macromol. Rapid Commun. 2005, 26, 1127; [5d] X. Lu, Y. Zhao, C. Wang, Y. Wei, Macromol. Rapid Commun. 2005, 26, 1325; [5e] J. Zeng, H. Hou, J. H. Wendorff, A. Greiner, Macromol. Rapid Commun. 2005, 26, 1557; [5f] S. Nair, S. Natarajan, S. H. Kim, Macromol. Rapid Commun. 2005, 26, 1599. [6] [6a] F. Yi, Z.-X. Guo, P. Hu, Z.-X. Fang, J. Yu, Q. Li, Macromol. Rapid Commun. 2004, 25, 1038; [6b] K. Ohkawa, D. Cha, H. Kim, A. Nishida, H. Yamamoto, Macromol. Rapid Commun. 2004, 25, 1600; [6c] W. K. Son, J. H. Youk, T. S.

Macromol. Rapid Commun. 2006, 27, 1341–1345

www.mrc-journal.de

[21] [22] [23] [24] [25] [26] [27] [28]

[29] [30] [31]

[32] [33] [34]

Lee, W. H. Park, Macromol. Rapid Commun. 2004, 25, 1632. D. H. Reneker, A. L. Yarin, H. Fong, S. Koombhongse, J. Appl. Phys. 2000, 87, 4531. A. L. Yarin, S. Koombhongse, D. H. Reneker, J. Appl. Phys. 2001, 89, 3018. J. Liu, T. Wang, T. Uchida, S. Kumar, J. Appl. Polym. Sci. 2005, 96, 1992. F. Ko, Y. Gogotsi, A. Ali, N. Naguib, H. Ye, G. Yang, C. Li, P. Willis, Adv. Mater. 2003, 15, 1161. J. J. Mark, L. M. Viculis, A. Ali, R. Luoh, G. Yang, H. T. Hahn, F. K. Ko, R. B. Kaner, Adv. Mater. 2005, 17, 77. H. Q. Hou, J. J. Ge, J. Zeng, Q. Li, D. H. Reneker, A. Greiner, S. Z. D. Cheng, Chem. Mater. 2005, 17, 967. S. Kedem, J. Schmidt, Y. Paz, Y. Cohen, Langmuir 2005, 21, 5600. M. M. Demir, M. A. Gulgun, Y. Z. Menceloglu, B. Erman, S. S. Abramchuk, E. E. Makhaeva, A. R. Khokhlov, V. G. Matveeva, M. G. Sulman, Macromolecules 2004, 37, 1787. [15a] V. E. Kalayci, P. K. Patra, Y. K. Kim, S. C. Ugbolue, S. B. Warner, Polymer 2005, 46, 7191; [15b] S. F. Fennessey, R. J. Farris, Polymer 2004, 45, 4217; [15c] X. H. Qin, Y. Q. Wan, J. H. He, J. Zhang, J. Y. Yu, S. Y. Wang, Polymer 2004, 45, 6409. K. Acatay, E. Simsek, C. Ow-Yang, Y. Z. Menceloglu, Angew. Chem. Int. Ed. 2004, 43, 5210. K. Chakrabarti, P. M. G. Nambissan, C. D. Mukherjee, K. K. Bardhan, C. Kim, K. S. Yang, Carbon 2006, 44, 948. [18a] P. Ye, Z.-K. Xu, J. Wu, C. Innocent, P. Seta, Macromolecules 2006, 39, 1041; [18b] P. Ye, Z.-K. Xu, J. Wu, C. Innocent, P. Seta, Biomaterials 2006, 27, 4169; [18c] P. Ye, Z.-K. Xu, A.-F. Che, J. Wu, P. Seta, Biomaterials 2005, 26, 6394. Z.-G. Wang, Z.-K. Xu, L.-S. Wan, J. Wu, C. Innocent, P. Seta, Macromol. Rapid Commun. 2006, 27, 516. E. Zussman, A. L. Yarin, A. V. Bazilevsky, R. Avrahami, M. Feldman, Adv. Mater. 2006, 18, 348. J. Kim, J. W. Grate, P. Wang, Chem. Eng. Sci. 2006, 61, 1017. T. Nakaya, Y. J. Li, Prog. Polym. Sci. 1999, 24, 143. A. L. Lewis, Colloid. Surf. B 2000, 18, 261. E. F. Murphy, J. R. Lu, A. L. Lewis, J. Brewer, J. Russell, P. Stratford, Macromolecules 2000, 33, 4545. [25a] K. Ishihara, Sci. Technol. Adv. Mater. 2000, 1, 131; [25b] Y. Iwasaki, K. Ishihara, Anal. Bioanal. Chem. 2005, 381, 534. N. Nakabayashi, D. F. Williams, Biomaterials 2003, 24, 2431. [27a] Z.-K. Xu, Q.-W. Dai, J. Wu, X.-J. Huang, Q. Yang, Langmuir 2004, 20, 1481; [27b] H.-T. Deng, Z.-K. Xu, X.-J. Huang, J. Wu, P. Seta, Langmuir 2004, 20, 10168. [28a] X.-J. Huang, Z.-K. Xu, L.-S. Wan, Z.-G. Wang, J.-L. Wang, Macromol. Biosci. 2005, 5, 322; [28b] X.-J. Huang, Z.-K. Xu, L.-S. Wan, Z.-G. Wang, J.-L. Wang, Langmuir 2005, 21, 2941; [28c] X.-J. Huang, Z.-K. Xu, X.-D. Huang, Z.-G. Wang, K. Yao, Polymer 2006, 47, 3141. G. E. Lawson, Y. Lee, F. M. Raushel, A. Singh, Adv. Funct. Mater. 2005, 15, 267. J. Watanabe, K. Ishihara, Biomacromolecules 2006, 7, 171. [31a] H. F. Jia, G. Y. Zhu, B. Vugrinovich, W. Kataphinan, D. H. Reneker, P. Wang, Biotechnol. Prog. 2002, 18, 1027; [31b] B. C. Kim, S. Nair, J. Kim, J. H. Kwak, J. W. Grate, S. H. Kim, M. B. Gu, Nanotechnol. 2005, 16, 382. K. Ishihara, T. Ueda, N. Nakabayashi, Polym. J. 1990, 22, 355. M. Bradford, Anal. Biochem. 1976, 72, 248. S. H. Chiou, W. T. Wu, Biomaterials 2004, 25, 197.

ß 2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim