Photocatalyzed Surface Modification of Poly(dimethylsiloxane) with

Jul 8, 2010 - alginic acid (AA) was investigated. The PDMS substrates were first oxidized in a H2SO4/H2O2 solution to transform the SirCH3 groups on t...
0 downloads 0 Views 562KB Size
Download PDF
Anal. Chem. 2010, 82, 6430–6439

Photocatalyzed Surface Modification of Poly(dimethylsiloxane) with Polysaccharides and Assay of Their Protein Adsorption and Cytocompatibility Linyan Yang,† Li Li,† Qin Tu,† Li Ren,† Yanrong Zhang,† Xueqin Wang,† Zhiyun Zhang,† Wenming Liu,† Liangliang Xin,† and Jinyi Wang*,†,‡ Colleges of Science, Life Science, and Animal Medicine and Shaanxi Key Laboratory of Molecular Biology for Agriculture, Northwest A&F University, Yangling, Shaanxi 712100, People’s Republic of China An improved approach for the surface modification of poly(dimethylsiloxane) (PDMS) using carboxymethyl cellulose (CMC), carboxymethyl β-1,3-dextran (CMD), and alginic acid (AA) was investigated. The PDMS substrates were first oxidized in a H2SO4/H2O2 solution to transform the Si-CH3 groups on their surfaces into Si-OH groups. Then methacrylate groups were grafted onto the substrates through a silanization reaction using 3-(trimethoxysilyl)propyl methacrylate. Sequentially, cysteamine was conjugated onto the silanized surfaces by the reaction between the thiol and methacrylate groups under 254 nm UV exposure. Afterward, the amino-terminated PDMS substrates were sequentially reacted with CMC, CMD, and AA in the presence of N-hydroxysuccinimide and 1-ethyl-3-[3-(dimethylamino)propyl]carbodiimide, resulting in the grafting of polysaccharides onto PDMS surfaces. The composition and chemical state of the modified surfaces were characterized by X-ray photoelectron spectroscopy (XPS). In addition, the stability and dynamic characteristics of the polysaccharide-grafted PDMS substrates were investigated by XPS and temporal contact angle experiments. A protein adsorption assay using bovine serum albumin (BSA), chicken egg albumin, lysozyme, and RNase-A showed that the introduction of CMD and AA can reduce the adsorption of negatively charged BSA and chicken egg albumin, but increase the adsorption of the positively charged lysozyme and RNaseA. However, CMC-modified PDMS surfaces showed protein-repelling properties, regardless of whether the protein was positively or negatively charged. A cell culture and migration study of glioma C6, MKN-45, MCF-7, and HepG-2 cells revealed that the polysaccharide-modified PDMS greatly improved the cytocompatibility of native PDMS. Microfluidic systems provide a powerful platform for biological analysis and have been utilized for biological synthesis, sample * To whom correspondence should be addressed. Phone: + 86-29-870 825 20. Fax: + 86-29-870 825 20. E-mail: [email protected]. † Colleges of Science, Life Science, and Animal Medicine. ‡ Shaanxi Key Laboratory of Molecular Biology for Agriculture.

6430

Analytical Chemistry, Vol. 82, No. 15, August 1, 2010

purification, disease diagnostics, DNA sequencing, protein crystallization, and biosensor and single-cell analysis.1-3 In comparison with conventional benchtop systems, microfluidic systems often show significant advantages in their low reagents and sample volumes, better mimicry of the natural tissue environment, precise experimental control of the cellular microenvironment, and lowcost disposable devices.1,4,5 None of these features are easily available from conventional analytical technologies. To facilitate different purposes, a variety of polymeric materials, including poly(dimethylsiloxane) (PDMS),1-3 poly(methyl methacrylate) (PMMA), polycarbonate (PC), poly(ethylene terephthalate) (PET), polyurethane, poly(vinyl chloride) (PVC), and polyester, have been increasingly adopted in producing various microfluidic devices.5 Among these devices, PDMS-based microfluidic systems1,2,5,6 have been gaining popularity due to their distinct advantages,1,3,6 such as nontoxicity, easy fabrication, practical scalability, optical transparency, and gas permeability. In addition, the elasticity of PDMS matrixes enables the integration of pressure-driven valves and pumps7 with microfluidic channels, permitting the execution and automation of complex chemical8,9 and/or biological10 processes within a single microfluidic device. However, due to the hydrophobic nature of its surface, biological components from blood and body fluids interact strongly with the PDMS surface when it is present in a biological (1) Whitesides, G. M. Nature 2006, 442, 368–373. (2) Regehr, K. J.; Domenech, M.; Koepsel, J. T.; Carver, K. C.; Ellison-Zelski, S. J.; Murphy, W. L.; Schuler, L. A.; Alarid, E. T.; Beebe, D. J. Lab Chip 2009, 9, 2132–2139. (3) Hansen, C. L.; Sommer, M. O.; Quake, S. R. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 14431–14436. (4) Yin, H.; Pattrick, N.; Zhang, X.; Klauke, N.; Cordingley, H. C.; Haswell, S. J.; Cooper, J. M. Anal. Chem. 2008, 80, 179–185. (5) Cheng, X.; Gupta, A.; Chen, C.; Tompkins, R. G.; Rodriguez, W.; Toner, M. Lab Chip 2009, 9, 1357–1364. (6) Wong, I.; Ho, C. M. Microfluid. Nanofluid. 2009, 7, 291–306. (7) Unger, M. A.; Chou, H. P.; Thorsen, T.; Scherer, A.; Quake, S. R. Science 2000, 288, 113–116. (8) Lee, C. C.; Sui, G.; Elizarov, A.; Shu, C. J.; Shin, Y. S.; Dooley, A. N.; Huang, J.; Daridon, A.; Wyatt, P.; Stout, D.; Kolb, H. C.; Witte, O. N.; Satyamurthy, N.; Heath, J. R.; Phelps, M. E.; Quake, S. R.; Tseng, H. R. Science 2005, 310, 1793–1796. (9) Wang, J.; Sui, G.; Mocharla, V. P.; Lin, R. J.; Phelps, M. E.; Kolb, H. C.; Tseng, H. R. Angew. Chem., Int. Ed. 2006, 45, 5276–5281. (10) Hong, J. W.; Studer, V.; Hang, G.; Anderson, W. F.; Quake, S. R. Nat. Biotechnol. 2004, 22, 435–439. 10.1021/ac100544x  2010 American Chemical Society Published on Web 07/08/2010

environment.11,12 A significant amount of protein adsorption onto the PDMS surface caused by such hydrophobic interaction is the most important problem to overcome because it triggers many undesirable bioreactions and greatly decreases the experiment efficiency in many cases, such as sorting of cells, patterning of biological and nonbiological materials on substrates, and electrophoresis separation of biomolecules.13,14 Moreover, it always prevents the immediate use of PDMS-based microfluidic devices without any surface processing. To date, several approaches have been developed to confer hydrophilicity10,16 and biomolecule-repelling properties12,15-17 to PDMS surfaces. Numerous materials have also been employed for these purposes, such as hydroxylpropyl methyl cellulose, poly(vinylpyrrolidone), dextran, hyaluronic acid, polyacrylamide, poly(2-hydroxyethyl methacrylate), poly(vinyl alcohol), hydroxyethyl cellulose, poly(N-(hydroxyethyl)acrylamide), poly(acrylic acid), poly(2-(methacryloyloxy)ethyl phosphorylcholine), and poly(ethylene glycol).6 All these efforts have greatly improved the protein-repelling properties of PDMS substrates and expanded the applications of PDMS-based microfluidic systems. However, the majority of these works on surface modification of microfluidic devices are developed for electrophoresis applications due to the vast exploitation of microchips and demanding stringency on surface properties.6 Other microfluidic-related applications have also showed intensive demand on surface modification, such as microfluidics-based cellular study, a very popular field developed for the study of physiological processes in a miniaturized environment.18 In the field, the control and improvement of substrate surface properties of microfluidic systems is an indispensable prerequisite for the success of bioanalytical applications, since the culture environment is very important to maintain healthy cell conditions.2 Although several methods have been developed for cell-based studies,19,20 they have all focused on controlled cell immobilization and immunoassay. In fact, it is impossible to have just one material or approach that meets all the individual needs of microfluidic systems.15 Therefore, the development of novel methods and introduction of various biocompatible materials for the proper functioning of PDMS-based microfluidics remain largely out of reach. A study on biomaterials has proven that polysaccharides can be utilized as potential alternatives to poly(ethylene glycol), a wellknown material for preventing the nonspecific adsorption of (11) Sui, G.; Wang, J.; Lee, C. C.; Lu, W.; Lee, S. P.; Leyton, J. V.; Wu, A. M.; Tseng, H. R. Anal. Chem. 2006, 78, 5543–5551. (12) Zhou, J.; Yan, H.; Ren, K.; Dai, W.; Wu, H. Anal. Chem. 2009, 81, 6627– 6632. (13) Ocvirk, G.; Munroe, M.; Tang, T.; Oleschuk, R.; Westra, K.; Harrison, D. J. Electrophoresis 2000, 21, 107–115. (14) Xiao, D.; Zhang, H.; Wirth, M. J. Langmuir 2002, 18, 9971–9976. (15) Wu, Y.; Huang, Y.; Ma, H. J. Am. Chem. Soc. 2007, 129, 7226–7227. (16) Lahann, J.; Balcells, M.; Lu, H.; Rodon, T.; Jensen, K. F.; Langer, R. Anal. Chem. 2003, 75, 2117–2122. (17) Liu, Y.; Fanguy, J. C.; Bledsoe, J. M.; Henry, C. S. Anal. Chem. 2000, 72, 5939–5944. (18) Kamei, K.; Guo, S.; Yu, Z. T.; Takahashi, H.; Gschweng, E.; Suh, C.; Wang, X.; Tang, J.; McLaughlin, J.; Witte, O. N.; Lee, K. B.; Tseng, H. R. Lab Chip 2009, 9, 555–563. (19) Mandal, S.; Rouillard, J. M.; Srivannavit, O.; Gulari, E. Biotechnol. Prog. 2007, 23, 972–978. (20) Henares, T. G.; Mizutani, F.; Hisamoto, H. Anal. Chim. Acta 2008, 611, 17–30.

proteins, which also features biocompatibility and low toxicity.21 In addition, polysaccharides possess advantages over the routine materials utilized for PDMS modifications. For example, carboxymethyl cellulose (CMC) can inhibit postsurgical and postoperative adhesions,22 and carboxymethyl β-1,3-dextran (CMD) can significantly reduce the factor level of tumor necrosis in endotoxemic animals and increase granulocyte/macrophage colonystimulating activity, resulting in marrow-derived cell hyperplasia.23 Alginic acid (AA), on the other hand, possesses good biocompatibility and hydrophilicity.24 Recently, chitosan and dextran have been physically deposited or covalently grafted onto PDMS surfaces for biomolecule separation, immunoassay, and protein nonfouling.21,25,26 Few research works, however, have been performed for covalently coupling polysaccharides onto PDMS for application in cellular studies. In addition, most of the existing polysaccharide-based protocols are not feasible for modifying the microchannel surfaces that are embedded in PDMS matrixes. From the standpoint of device fabrication, surface-modified PDMS components often face the challenges of device assembly and microchannel sealing. These problems limit further development of PDMS-based microfluidic devices, especially for their applications in biological analysis. In this work, we describe an improved photocatalyzed approach for PDMS surface modification using the polysaccharides CMC, CMD, and AA. All the procedures were performed in aqueous solutions, resulting in a method suitable for polysaccharide-based microchannel modification. Using this approach, numerous PDMS substrates (wells and microchannels) were treated, and protein adsorption and cell growth viability of glioma C6, MKN-45, MCF7, and HepG-2 cells on the surfaces were investigated. The migration of these cells in modified microchannels was also analyzed. EXPERIMENTAL SECTION Materials and Reagents. RTV 615 PDMS prepolymer and a curing agent were purchased from GE Silicones (Minato-ku, Tokyo). Surface-oxidized silicon wafers were obtained from Shanghai Xiangjing Electronic Technology Ltd. (Shanghai, China). SU-8 2025 photoresist and developer were purchased from Microchem (Newton, MA). The compounds 3-(trimethoxysilyl)propyl methacrylate (MPTS) and cysteamine hydrochloride were obtained from Aladdin (Maryland). CMC and RNase-A were purchased from Sigma-Aldrich (Missouri). CMD was obtained from Zhuhai Tiantian Biotechnology Co., Ltd. (Zhuhai, China). AA was purchased from Qingdao Crystal Rock Biology Development Co., Ltd. (Qingdao, China). N-Hydroxysuccinimide (NHS) and 1-ethyl-3-(3-(dimethylamino)propyl)carbodiimide (EDC) were obtained from GL Biochem Ltd. (Shanghai, China). Chicken egg albumin and lysozyme were purchased from Dingguo Biotechnology Ltd. (Beijing, China). Bovine serum albumin (BSA) was (21) Martwiset, S.; Koh, A. E.; Chen, W. Langmuir 2006, 22, 8192–8196. (22) Ryan, C. K.; Sax, G. C. Am. J. Surg. 1995, 169, 154–160. (23) Vereschagin, E. I.; Lambalgen, A. A. V.; Dushkin, M. I.; Schwartz, Y. S.; Polyakov, L.; Heemskerk, A.; Huisman, E.; Thijs, L. G.; van den Bos, G. C. Folia Biol. (Prague) 1993, 39, 178–187. (24) Niino, H.; Kru ¨ ger, J.; Kautek, W. Appl. Phys. A: Mater. Sci. Process. 2001, 72, 53–57. (25) Hu, S. G.; Jou, C. H.; Yang, M. C. Biomaterials 2003, 24, 2685–2693. (26) Yu, L.; Li, C. M.; Liu, Y.; Gao, J.; Wang, W.; Gan, Y. Lab Chip 2009, 9, 1243–1247.

Analytical Chemistry, Vol. 82, No. 15, August 1, 2010

6431

obtained from Amresco Inc. (Solon, OH). The 2-(N-morpholino)ethanesulfonic acid (MES) used in the study was purchased from BBI (Ontario, Canada). The cell culture medium and fetal bovine serum (FBS) were obtained from Gibco Invitrogen Corp. (California). All solvents and other chemicals were purchased from local commercial suppliers and were of analytical reagent grade, unless otherwise specified. Deionized (DI) water (Milli-Q, Millipore, Bedford, MA) was used to prepare aqueous solutions. PDMS Device Fabrication. In this study, two types of PDMS devices (Supporting Information, devices 1 and 2) were utilized. To fabricate device 1, 24 g of well-mixed PDMS mixture (RTV 615 A and B in a 5:1 ratio) was poured onto a silicon wafer placed in a Petri dish. After curing in an 80 °C oven for 50 min, the PDMS layer was peeled off the silicon wafer and holes with a diameter of 6 mm were punched through the layer. It was then placed onto an appropriate glass slide with a thin PDMS film (RTV 615 A and B in a 20:1 ratio) and baked overnight prior to use. Device 2 was fabricated using a soft lithography method.1,3,7 One mold was first produced by photolithographic processes to create the fluidic components (channel width 200 µm, channel height 35 µm, channel length 6 mm) embedded into the layers of the PDMS matrix. To prepare the mold, a 35 µm thick negative photoresist (SU8-2025) was spin-coated onto a silicon wafer. After UV exposure and development, a mold with the required fluidic patterns was obtained. Before fabrication of the device, the mold was first exposed to trimethylchlorosilane ((TMS)Cl; Alfa Aesar, Lancs, England) vapor for 2-3 min. Then a well-mixed PDMS mixture (RTV 615 A and B in a 5:1 ratio) was poured into the mold to obtain a 2 mm thick fluidic layer containing the microchannels transferred from the mold. After curing in an 80 °C oven for 50 min, the PDMS layer with fluidic channels was peeled off the mold and through-holes were punched using a metal pin at the terminals of the inlet and outlet channels. The fluidic PDMS layer was then placed on top of a glass slide coated (2000 rpm, 60 s, ramp 15 s) with a PDMS mixture (RTV 615 A and B in a 20:1 ratio) and incubated for 45 min in the oven. Device 2 was ready for use after being baked for 48 h in an oven set at 80 °C. General Procedure for Surface Modification. PDMS substrates were first treated in H2SO4/H2O2 solution (3:1, v/v) for 30 s at 40 °C27 and washed with DI water until a pH of 7.0 was obtained. The substrates were then dipped in an aqueous solution with 0.4% (w/v) MPTS and 0.4% (v/v) acetic acid for 210 min. After rinsing with DI water five times, cysteamine hydrochloride solution (50 mg/mL, w/v) was introduced to the substrates, which were then exposed to 254 nm UV for 18 h.28 Before the polysaccharides were grafted onto the PDMS substrates, the cysteamine-grafted substrates were treated with an aqueous solution of triethylamine for 5 min to release the amino group from its hydrochloride salt state. After rinsing with DI water five times, the three polysaccharides CMC, CMD, and AA were conjugated onto the aminated PDMS substrates in the presence of EDC and NHS.29 To graft CMC onto the aminated substrates, CMC solution (2 mg/mL in 0.1 mol/L MES buffer, pH 6.0) was first converted to N-hydrox(27) Xu, C.; Taylor, P.; Ersoz, M.; Fletcher, P. D. I.; Paunov, V. N. J. Mater. Chem. 2003, 13, 3044–3048. (28) Roy, R.; Baek, M. G. Rev. Mol. Biotechnol. 2002, 90, 291–309. (29) Huck, C. W.; Stecher, G.; Bakry, R.; Bonn, G. K. Electrophoresis 2003, 24, 3977–3997.

6432

Analytical Chemistry, Vol. 82, No. 15, August 1, 2010

ysuccinimide esters by a sequential reaction with EDC (38.4 mg/mL in MES buffer, pH 6.0) for 15 min and NHS (23 mg/ mL in MES buffer, pH 6.0) for 60 min. The freshly aminated PDMS substrates were then reacted with the EDC- and NHSactivated CMC solution for 72 h at room temperature. After being washed five times with DI water to remove any excess reagents, the CMC-modified PDMS substrates were then stored in a refrigerator until use. The process of conjugating CMD onto the animated PDMS substrates was the same as above. Minor differences24 were observed in the conjugation of AA onto the aminated PDMS substrates. First, an AA solution (5 mg/mL in MES buffer, pH 6.0) was mixed with N,Ndimethylformamide (DMF; 3:1, v/v). Then the AA solution (3.75 mg/mL) was converted to N-hydroxysuccinimide esters by sequential reaction with EDC (36.3 mg/mL in MES buffer, pH 6.0) for 15 min and NHS (10.95 mg/mL in MES buffer, pH 6.0) for 60 min. The solution was finally introduced to the freshly aminated PDMS substrate and reacted for 28 h at room temperature. Modification of the microchannels in device 2 was similar to the procedure described above, except that the amination reaction, that is, the reaction of MPTS-grafted microchannels with cysteamine, took 21 h. All the reagents and solutions were introduced into the inlet connected to the microchannels and removed by a pipet after they were rinsed. X-ray Photoelectron Spectroscopy (XPS) Analysis. The XPS analyses were performed on an Axis Ultra X-ray photoelectron spectrometer with a Mg/Al X-ray source operating at 150 W (15 kV, 13 mA). The vacuum in the main chamber was kept above 3 × 10-6 Pa during XPS data acquisitions. Specimens were analyzed at an electron takeoff angle of 70°, measured with respect to the surface plane. For all modified PDMS substrates (1.0 cm × 1.0 cm), general survey scans (binding energy range 0-1100 eV, pass energy 160 eV) and high-resolution spectra (pass energy 80 eV) in the regions of C1s and N1s were recorded. Binding energies were referenced to the C1s binding energy at 284.8 eV. Contact Angle Measurements. Contact angle measurements were performed on PDMS specimens using Dropmeter 100 equipment (Maist Vision, Ningbo, China) via the sessile drops technique. DI water (18.4 MΩ) was used. Each data point given was based on 10 contact angle measurements at 5 different positions on the PDMS specimen. Advancing and receding contact angles were measured while the needle remained in the water droplet. Static contact angles were measured following the method previously reported.30 Stability Tests. To investigate the stability of polysaccharidemodified PDMS surfaces, the modified PDMS substrates were exposed to either air or DI water for 30 days. Their surfaces were characterized by XPS and contact angle measurements following the procedures described above. Protein Adsorption Analysis. Protein adsorption assay was performed following the method previously reported.31 Protein solutions were freshly prepared by dissolving BSA, chicken egg albumin, lysozyme, or RNase-A in phosphate-buffered saline (PBS; (30) Lam, C. N. C.; Kim, N.; Hui, D.; Kwok, D. Y.; Hair, M. L.; Neumann, A. W. Colloids Surf., A 2001, 189, 265–278. (31) Tangpasuthadol, V.; Pongchaisirikul, N.; Hoven, V. P. Carbohydr. Res. 2003, 338, 937–942.

pH 7.4) to give a final concentration of 0.5, 3, 5, 20, and 45 mg/ mL. After being rinsed with PBS, the polysaccharide-modified PDMS substrates (3 mm ×5 mm) were placed in 5 mL glass bottles, to which 2 mL of the freshly prepared BSA, chicken egg albumin, lysozyme, or RNase-A solution was added. Adsorption was allowed to proceed at 37 °C for 2 h under gentle shaking. After being washed with PBS five times and dried using N2, the PDMS substrates were sonicated in 2 mL of 1 wt % sodium dodecyl sulfate (SDS; Amerso Inc.) to remove reversibly adsorbed protein. The amount of protein adsorbed onto the substrates was determined by micro-BCA protein assay.32 The absorbance of the solution was measured by a microplate reader (Bio-Rad model 680) at 570 nm. Three repetitions were performed for all samples. Caution was used in handling all human biological material. Cell Culture. Glioma C6 cells, gastric cancer cells (MKN-45 cells), human breast adenocarcinoma cells (MCF-7 cells), and human hepatocellular liver carcinoma cells (HepG-2 cells) were all obtained from the Chinese Academy of Sciences (Shanghai, China). Four types of cells were routinely cultured using Dulbecco’s modified Eagle’s medium (DMEM; Invitrogen, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS; Invitrogen), 100 units/mL penicillin, and 100 µg/mL streptomycin in a humidified atmosphere of 5% CO2 at 37 °C. To maintain cells in the exponential growth phase, they normally passed at a ratio of 1:3 every three days. Before use, the cells were harvested through trypsinization with 0.25% trypsin (Invitrogen) in the Ca2+- and Mg2+-free Hanks balanced salt solution (CMF-HBSS) at 37 °C. Trypsinization was stopped by the addition of fresh supplemented DMEM, and the cell suspension was centrifuged at a rotational speed of 800 rpm for 3 min. The cells were then resuspended in supplemented DMEM (2 × 104 cells/mL) for use. Cell Growth on Polysaccharide-Modified Substrates. For each cell line, three wells of one device 1 were modified using the three polysaccharides. Another well was utilized for the control. Before cell culture, the PDMS devices were first sterilized under UV light for 1 h at room temperature and rinsed three times with a cell culture medium. The desired cell line was then seeded into the wells (2 × 103 cells/well) and cultured for one week. Cell growth curves on different surfaces were measured by counting the number of cells every 24 h using the hemocytometer method, and seven devices were simultaneously utilized for one round test. Each round test was repeated at least three times. Cell Migration in Polysaccharide-Modified Microchannels. After modification, sterilization, and rinsing following the procedures described above, the cells were seeded into the inlet reservoirs of device 2 (2 × 103 cells/reservoir) to establish a confluent monolayer and form a linearity edge along the interface between the reservoir and the channels. To keep a static and nongravitational environment for cell migration, culture medium levels in the inlet and outlet reservoirs were kept equal. The microfluidic devices were then kept in a humidified atmosphere of 5% CO2 at 37 °C for 7 days. Cell migration ability was quantified by measuring the distance (32) Smith, P. K.; Krohn, R. I.; Hermanson, G. T.; Mallia, A. K.; Gartner, F. H.; Provenzano, M. D.; Fujimoto, E. K.; Goeke, N. M.; Olson, B. J.; Klenk, D. C. Anal. Biochem. 1985, 150, 76–85.

traveled along the channels over the time taken. Each test was repeated at least three times. Microscopy and Image Analysis. An inverted microscope (Olympus, CKX41) with a CCD camera (QImaging, Micropublisher 5.0 RTV) and a mercury lamp (Olympus, U-RFLT50) was used to acquire phase contrast and fluorescence images. Time lapse images of cell growth and migration were obtained every 24 h. The software Image-Pro Plus 6.0 (Media Cyternetics, Silver Spring, MD) and SPSS 12.0 (SPSS Inc.) were employed to perform image analysis and data statistical analysis, respectively. Data are presented as means ± SD for the measured contact angles, cell growth curves, and cell migration ability.

RESULTS AND DISCUSSION Surface Modifications. Modification (Scheme 1) of the working surfaces (Supporting Information, wells of device 1 and microchannels of device 2) of these devices starts from the solution-phase oxidation reaction of PDMS surfaces, which was carried out by treating the PDMS surfaces with H2SO4/H2O2 solution.27 Then methacrylate groups were grafted onto the silanol-covered PDMS wells or microchannels through a silanization reaction using MPTS. Afterward, the methacrylate groups were converted to amino groups by the reaction between cysteamine and the MPTS-grafted surface under 254 nm UV exposure.28 It should be emphasized that the use of a long amination reaction time is critical for MPTS-grafted microchannel modification since the introduction of methacrylate groups and the thickness of the fluidic layer may affect the intensity of UV light that reaches the microchannel surfaces. Studies on the UV transmittance of MPTS-grafted PDMS substrates, as well as the relationship between the density of amino groups grafted onto the PDMS slide-covered PDMS surfaces and the reaction time, showed that, after 21 h, the density of the amino group grafted onto the microchannel surfaces was close to that of the UV directly exposed substrates (Supporting Information). Finally, the amino-grafted PDMS surfaces were subjected to reactions with the three polysaccharides CMC, CMD, and AA to produce CMC-grafted PDMS, CMDgrafted PDMS, and AA-grafted PDMS surfaces, respectively. Low contact angle means more polysaccharide was grafted onto the PDMS substrates. To ensure the maximum amount of the polysaccharides was grafted onto the PDMS substrates, the optimized reaction time for CMC, CMD, and AA was 72, 72, and 28 h, respectively, since the contact angles of the polysaccharidemodified PDMS surfaces were the minimum after these reaction periods. The stabilized static contact angles in air after these reaction periods were 46.5 ± 0.81°, 42.5 ± 0.78°, and 44.2 ± 0.83° for CMC-, CMD-, and AA-modified PDMS surfaces, respectively. These reaction conditions were employed to treat the devices utilized for the next studies. XPS Analysis. A number of wafer-molded PDMS substrates with dimensions of 1.0 cm ×1.0 cm were treated according to the same approach described above to give the MPTS-grafted, aminografted, and polysaccharide-grafted substrates for subsequent surface XPS characterizations. The properties of these silanolcovered surfaces are, however, dynamic. As a result, the progresAnalytical Chemistry, Vol. 82, No. 15, August 1, 2010

6433

Scheme 1. Schematic Presentation of the PDMS Modification Process Using the Polysaccharides CMC, CMD, and AA under UV Catalysisa

a Reagents and conditions: (i) H2SO4/H2O2, 40° C, 30 s; (ii) 0.4% (v/v) acetic acid solution; (iii) 254 nm UV; TEA, 5 min; (iv) EDC/NHS, 72 h; (v) EDC/NHS, 72 h; (vi) EDC/NHS, 28 h.

sive restoration of hydrophobicity occurs within a few minutes.33 Therefore, silanization reactions on the freshly prepared silanolcovered PDMS surfaces were performed immediately, and the surface properties were characterized on the subsequent silanization surfaces. Figure 1A shows the low-resolution XPS survey spectra of these surfaces, all of which are semiquantitative because this analysis is unlikely to separate the photoemission contributions of the modified layers from the background of bulk PDMS.11 The XPS spectrum of native PDMS substrates exhibited a ratio of 0.61 between the O1s and C1s photoemissions. A different O1s: C1s ratio of 0.53 was observed for MPTS-grafted substrates, indicating that the introduction of MPTS changed the surface chemical compositions of native PDMS substrates and more carbon was introduced. It also implies that MPTS was grafted onto silanol-covered surfaces (for the elemental composition of each step of the modification process and the ratios of O1s to C1s, see the Supporting Information). Further evidence of MPTS-grafted surfaces is supported by high-resolution XPS spectra (Figure 1B). High-resolution C1s XPS spectra of the MPTS-grafted substrates have peaks at 282.65 eV (C-C), 283.05 eV (Cs(CdO)sO-), 284.15 eV (C-O), and 286.65 eV (OsCdO).34,35 A comparison with the high-resolution C1s XPS spectra of native PDMS [281.79

eV (C-Si), 282.44 eV (C-H)]36 indicates that MPTS was grafted onto the surface. In particular, the appearance of the carbonyl peak at 286.65 eV and the R-carbon at 283.05 eV provided obvious evidence that MPTS was successfully conjugated onto the silanolcovered surfaces. For the cysteamine-grafted surface (NH2PDMS), the appearance of the N1s peak in low-resolution XPS survey spectra (Figure 1A) implies the introduction of amino groups. The high-resolution N1s XPS spectra (Figure 1C) of the NH2-grafted surfaces have peaks at 397.35 eV (C-NH2) and 400.0 eV (protonated amine).37 In addition, C1s XPS spectra of the NH2-grafted surface (Figure 1B) also show that similar peaks appeared for the MPTS-PDMS surfaces. All the results above indicate that cysteamine was grafted onto MPTS-grafted surfaces. In the high-resolution C1s XPS spectra of polysaccharidegrafted surfaces (Figure 1B), the appearance of a peak at 282.85 eV (Cs(CdO)sNH-) and its disappearance at 283.05 eV (Cs(CdO)sO-) from the surface of NH2-PDMS demonstrate that the formation of an amide bond between the carboxyl group of the polysaccharides and the terminal amino groups on the NH2-PDMS surface occurred. High-resolution N1s XPS spectra (Figure 1C) of the polysaccharide-grafted surfaces all have a single peak at 397.9 eV (-NH(CdO)-).38 All this information

(33) Delamarche, E.; Geissler, M.; Bernard, A.; Wolf, H.; Michel, B.; Hilborn, J.; Donzel, C. Adv. Mater. 2001, 13, 1164–1167. (34) Siddiquey, I. A.; Ukaji, E.; Furusawa, T.; Sato, M.; Suzuki, N. Mater. Chem. Phys. 2007, 105, 162–168. (35) Beamson, G. J. Electron Spectrosc. Relat. Phenom. 2001, 121, 163–181.

(36) EI-Ashgar, N. M.; EI-Nahhal, I. M.; Chehimi, M. M.; Delamar, M.; Babonneau, F.; Livage, J. Monatsh. Chem. 2006, 137, 263–275. (37) Sahoo, R. R.; Patnaik, A. J. Colloid Interface Sci. 2003, 268, 43–49. (38) Niu, Z.; Gao, F.; Jia, X.; Zhang, W.; Chen, W.; Qian, K. Y. Colloids Surf., A 2006, 272, 170–175.

6434

Analytical Chemistry, Vol. 82, No. 15, August 1, 2010

Figure 1. (A) XPS wide scan spectra of the native PDMS, MPTS-grafted PDMS (MPTS-PDMS), cysteamine-grafted PDMS (NH2-PDMS), and polysaccharide-grafted PDMS (CMC-PDMS, CMD-PDMS, and AA-PDMS). (B) High-resolution XPS C1s spectra of the native PDMS, MPTS-PDMS, NH2-PDMS, CMC-PDMS, CMD-PDMS, and AA-PDMS. (C) High-resolution XPS N1s spectra of NH2-PDMS, CMC-PDMS, CMD-PDMS, and AA-PDMS.

demonstrates that the polysaccharides were covalently grafted onto the PDMS surfaces. This was also confirmed by their ATRFT-IR spectra (Supporting Information), in which amide I and amide II peakes were at 1641 and 1565 cm-1, respectively.39 Stability Tests. To investigate the stability of polysaccharidemodified PDMS surfaces in aqueous and ambient environments, numerous modified PDMS substrates were exposed to either air or DI water for 30 days. Dynamic surface characteristics of the modified substrates were monitored by temporal contact angle experiments. The results (Supporting Information) reveal that the introduction of polysaccharides onto PDMS surfaces greatly improved the hydrophilicity of native PDMS substrates (the advancing contact angle of the native PDMS substrate is 119.1 ± 2.38°). The stability of these surfaces, whether in the water phase or in air, tends to remain constant after two days (the stabilized advancing contact angles in air for CMC-, CMD-, and AA-modified PDMS are 56.6 ± 0.71°, 53.2 ± 0.64°, and 53.9 ± 0.76°, respectively). All these results suggest that robust cross-linked polysaccharide-silane layers were created on the PDMS surfaces, thus exhibiting long-lasting hydrophilicity. The long-lasting hydrophilicity of the surface is attributed to the use of chemical covalent coupling. To track changes in the composition and the chemical state of the modified surfaces before and after treatment during stability tests, polysaccharide-modified PDMS substrates treated for 30 days under aqueous and ambient environments were all (39) Ma, J.; Xu, Y.; Fan, B.; Liang, B. Eur. Polym. J. 2007, 43, 2221–2228.

characterized by XPS (Supporting Information). In comparison to the fresh polysacchide-modified PDMS substrates, the results show that the treatments did not appear to affect the composition and chemical state of the polysaccharide-modified PDMS surfaces. The XPS spectra of the polysaccharide-modified PDMS substrates before and after treatment also provided supporting evidence that the long-lasting hydrophilicity and stability of the modified surface are due to covalent coupling. Protein Adsorption Analysis. More recently, studies on polysaccharide coatings have demonstrated that different polysaccharide structures possess different protein adsorption abilities, depending on their electrostatic and steric-entropic interactions.40 In physiological media, most proteins carry a net charge, with the sign and magnitude of the net charge depending on the isoelectric point of the protein (pI). Electrostatic interactions between proteins and charged surfaces, therefore, often play a major role in the adsorption behavior of proteins.31 The assay of polysaccharide-modified PDMS surfaces using two methods previously reported41-43 showed that there are residual carboxyl (40) McArthur, S. L.; McLean, K. M.; Kingshott, P.; St John, H. A. W.; Chatelier, R. C.; Griesser, H. J. Colloids Surf., B 2000, 17, 37–48. (41) Hu, S.; Ren, X.; Bachman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Anal. Chem. 2004, 76, 1865–1870. (42) Wang, Y.; Bachman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Langmuir 2006, 22, 2719–2725. (43) Wang, Y.; Lai, H.; Bachman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Anal. Chem. 2005, 77, 7539–7546.

Analytical Chemistry, Vol. 82, No. 15, August 1, 2010

6435

Figure 2. Assay of positively and negatively charged protein adsorption on the native and polysaccharide-modified PDMS. Prior to use, all the polysaccharide-modified PDMS substrates were stabilized at ambient temperature for at least 2 days: (A) adsorption of BSA on the native and polysaccharide-modified PDMS, (B) adsorption of chicken egg albumin on the native and polysaccharide-modified PDMS, (C) adsorption of lysozyme on the native and polysaccharide-modified PDMS; (D) adsorption of RNase-A on the native and polysaccharide-modified PDMS.

groups on the surfaces (Supporting Information). Therefore, BSA, chicken egg albumin, lysozyme, and RNase-A, four globular proteins varying in charge,31 were selected for protein adsorption assay in the current study. The amounts of protein adsorbed onto native PDMS and polysaccharide-modified PDMS surfaces are shown in Figure 2, which indicates that polysaccharide-modified PDMS surfaces tend to reduce BSA and chicken egg albumin adsorption (Figure 2A,B) but increase lysozyme and RNase-A adsorption (Figure 2C,D), regardless of high or low protein concentrations (except for CMCmodified PDMS). As mentioned above, polysaccharide-modified PDMS surfaces maintained residual carboxylic acid groups on their surfaces. At pH 7.4, the carboxylic acid was converted to a negatively charged carboxylate ion.31 Therefore, BSA (pI 4.7) and chicken egg albumin (pI 4.7), two carboxylic acid-rich proteins, were significantly less adsorbed onto the carboxylic acid-rich surface, possibly due to charge-charge repulsion. On the other hand, lysozyme (pI 11) and RNase-A (pI 9.4) contain a large number of -OH and -NH2 groups.31 Therefore, they are positively charged at pH 7.4. Hydrogen-bonding and chargecharge attraction could be responsible for the increased adsorption of lysozyme and RNase-A. The results from this study seem to fit well with a previous report on the study of the charge interaction of protein and polyelectrolyte films.31 In comparison with the amount of protein adsorbed onto different polysaccharide-modified PDMS substrates, CMC-modified PDMS surfaces exhibited repelling actions to both positively and negatively charged proteins, matching well with actual applications in vivo for reducing the 6436

Analytical Chemistry, Vol. 82, No. 15, August 1, 2010

“fouling” of surfaces by biological species.22 The repelling mechanism has not yet been addressed. Cell Culture and Vitality Evaluation. To investigate the cytocompatibility of polysaccharide-modified PDMS surfaces, C6, MKN-45, MCF-7, and HepG-2 cells were all cultured onto them using device 1. The cell modality and proliferation were then analyzed. The cell shape was selected as an important factor for determining the cell behavior. One set of inverted microscope images of cell growth is presented in Figure 3. As the images show, on the native PDMS surface, C6 cells do not fully adhere. These cells also show a round shape and aggregate together to grow. Furthermore, only a small number of cells with short neurites was determined. This type of cell took on poor proliferation abilities. The cells were well-spread through the polysaccharide-modified surfaces and showed shuttle- or ellipse-like shapes. Many C6 cells had tubby or slight neurites. The other three cell lines also showed similar phenomena. They were not fully adhered onto the native PDMS surfaces and exhibited round shapes. All cell lines showed completely spread shapes on the modified surfaces, which fractionally indicates good growth capability. To further determine the cell proliferation vitality as cultured on different surfaces, the speed of cell proliferation was also quantified by counting the number of cells every 24 h using the hemocytometer method. The results are shown in Figure 4. The figure obviously shows that the proliferation vitality of cells cultured on polysaccharide-modified PDMS surfaces was higher than that of cells cultured on native PDMS surfaces. The results suggest that the introduction of polysaccharides ameliorated the

Figure 3. Microscopic images of glioma C6 cells, MKN-45 cells, MCF-7 cells, and HepG-2 cells cultured on the native or polysaccharidemodified PDMS for 4 days. Prior to use, all the polysaccharide-modified PDMS devices were stabilized at ambient temperature for at least 2 days.

Figure 4. Growth curves of different cell lines on the native or polysaccharide-modified PDMS: (A) glioma C6 cells, (B) MKN-45 cells, (C) MCF-7 cells, (D) HepG-2 cells. Prior to use, all the polysaccharide-modified PDMS devices were stabilized at ambient temperature for at least 2 days.

properties of PDMS for cell growth. The difference in cell proliferation vitality for homotypical cells on different surfaces and heterotypical cells on the same surfaces might be due to the fact that different modified surfaces may have different interaction abilities with the cell member ligands. On the other hand, it may also be because different tissue-derived cell membranes possess

different functional groups. As such, they may also exhibit different interactions with modified surfaces.44,45 (44) Gumbiner, B. M. Cell 1996, 84, 345–357. (45) McBeath, R.; Pirone, D. M.; Nelson, C. M.; Bhadriraju, K.; Chen, C. S. Cell 2004, 6, 483–495.

Analytical Chemistry, Vol. 82, No. 15, August 1, 2010

6437

Figure 5. Migration distances of (A) glioma C6 cells, (B) MKN-45 cells, (C) MCF-7 cells, and (D) HepG-2 cells in the native and polysaccharidemodified PDMS microchannels after culturing in device 2 for 7 days. Prior to use, all the polysaccharide-modified PDMS microchannels were stabilized at ambient temperature for at least 2 days.

Cell Migration in Microchannels. To further study the cytocompatibility of polysaccharide-modified PDMS surfaces, cell migration experiments were also performed in polysaccharidemodified microchannels using device 2. The main reason for this is that cell migration plays a crucial role in various biological processes, including embryogenesis, wound healing, immune responses, and tissue development.46,47 Understanding the mechanisms underlying cell migration will also be important for emerging areas of biotechnology which focus on cellular transplantation and the manufacture of artificial tissues, as well as for the development of new therapeutic strategies for controlling invasive tumor cells. To a certain extent, the cell migration ability reflects the cell vitality. The mean migration distances of C6, MKN-45, MCF-7, and HepG-2 cells in the modified microchannels are displayed in Figure 5 (one set of propidium iodide-stained fluorescence images of cell migration over 7 days is presented in the Supporting Information). As the results show, the polysaccharide-modified surface improved the cell migration ability compared to that in native PDMS microchannels. In native PDMS microchannels, all cells migrated short distances, whereas in polysaccharide-modified PDMS microchannels, cell migration was significantly promoted. These results were in agreement with the results from the cell proliferation vitality assay presented above. (46) Lo´pez-Bendito, G.; Cautinat, A.; Sa´nchez, J. A.; Bielle, F.; Flames, N.; Garratt, A. N.; Talmage, D. A.; Role, L. W.; Charnay, P.; Marı´n, O.; Garel, S. Cell 2006, 125, 127–142. (47) Gurtner, G. C.; Werner, S.; Barrandon, Y.; Longaker, M. T. Nature 2008, 453, 314–321.

6438

Analytical Chemistry, Vol. 82, No. 15, August 1, 2010

CONCLUSIONS In this work, three polysaccharides, CMC, CMD, and AA, were successfully covalently grafted onto PDMS substrates using a photocatalyzed surface modification method. Although long-term reaction steps were employed in the process, the method greatly extended the biocompatibile and low toxic material polysaccharide application in microfluidics. Also, this method is suitable for the modification of microchannels embedded in PDMS matrixes. Contact angle measurements showed that the introduction of the three polysaccharides greatly improved the hydrophilicity of native PDMS substrates. The modified surfaces also have long-lasting stability. Protein adsorption assay exhibited that CMD- and AAmodified PDMS surfaces can reduce the adsorption of negatively charged BSA and chicken egg albumin, but increase that of positively charged lysozyme and RNase-A. However, CMCmodified PDMS surfaces exhibited protein-repelling properties for both types of proteins. The cell culture and migration study revealed that the polysaccharide-modified PDMS greatly ameliorated the cytocompatibility of native PDMS. Compared with the native PDMS substrates, the introduction of the polysaccharides on the PDMS surfaces greatly improves the viability of the cells. This work demonstrates the potential of applying polysaccharides in a PDMS microfluidics-based cellular study. It also expands the application of PDMS-based microfluidics to a certain extent. In further experiments, when the amidation reaction is performed, residual carboxyl groups on the modified surfaces may be used for biomolecule immobilization. The polysaccharides may also be used as bifunctional polymeric surfaces, that is, for specific

biomolecule immobilization and nonspecific fouling resistance in the microfluidic system. ACKNOWLEDGMENT We acknowledge the funding provided by the National Natural Science Foundation of China (Grant Nos. 209 750 82 and 207 750 59), the Ministry of Education of the People’s Republic of China (Grant NCET-08-0464), the State Forestry Administration of the People’s Republic of China (Grant No. 200904004), the Scientific Research Foundation for the Returned Overseas Chinese Scholars,

State Education Ministry, and Northwest A&F University. The first two authors contributed equally to this work. SUPPORTING INFORMATION AVAILABLE Additional information as noted in the text. This material is available free of charge via the Internet at http://pubs.acs.org. Received for review March 1, 2010. Accepted June 29, 2010. AC100544X

Analytical Chemistry, Vol. 82, No. 15, August 1, 2010

6439