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Covalent Micropatterning of Poly(dimethylsiloxane) by Photografting through a Mask Yuli Wang,†,‡,§ Hsuan-Hong Lai,| Mark Bachman,†,‡ Christopher E. Sims,§ G. P. Li,*,†,‡ and Nancy L. Allbritton*,§
Integrated Nanosystems Research Facility, Department of Electrical Engineering and Computer Science, Department of Physiology and Biophysics, and Department of Chemical Engineering and Materials Science, University of California, Irvine, California 92697
A new photografting method to micropattern a covalent surface modification on poly(dimethylsiloxane) (PDMS) provides advantages in simplicity and efficiency. To accomplish the entire process on the benchtop, the PDMS was initially treated with benzophenone dissolved in a water/acetone mixture. This process permitted limited diffusion of the photoinitiator into the PDMS surface. Polymerization of acrylic acid was initiated by exposure of the benzophenone-implanted PDMS to UV radiation through a photomask with a thin aqueous layer of acrylic acid sandwiched between the PDMS and photomask. This procedure resulted in patterned poly(acrylic acid) (PAA) on the PDMS surface. In the modified regions, PAA and PDMS formed an interpenetrating polymer network extending 50 µm into the PDMS with an X-Y spatial resolution of 5 µm. The carboxyl groups of the PAA graft could be derivatized to covalently bond other molecules to the patterned PAA. Two bioanalytical applications of this micropatterned surface were demonstrated: (1) a guide for cell attachment and growth and (2) a substrate for immunoassays. 3T3 cells were shown to selectively localize to modified surface regions where they could be cultured for up to 7 days. Additionally, the micropatterned surface was used to immobilize either protein A or antibody for heterogeneous immunoassays. Polymers are of growing importance in microfabricated devices especially those utilized in the fabrication of biosensors and labon-chips. While the first such devices were fabricated in glass or silicon, polymers have proven to be a viable alternative principally due to the ability to rapidly and inexpensively prototype devices on the benchtop. Of the polymers employed in fabricating these devices, poly(dimethylsiloxane) (PDMS), is one of the most utilized.1 The attractive properties of PDMS include low cost, ease * Corresponding authors. E-mail:
[email protected]. Fax: 949-824-9137. Phone: 949-824-6493. E-mail:
[email protected]. Fax: 949-824-3732. Phone: 949-8244194. † Integrated Nanosystems Research Facility. ‡ Department of Electrical Engineering and Computer Science. § Department of Physiology and Biophysics. | Department of Chemical Engineering and Materials Science. (1) McDonald, J. C.; Whitesides, G. M. Acc. Chem. Res. 2002, 35, 491-499. 10.1021/ac0509915 CCC: $30.25 Published on Web 11/03/2005
© 2005 American Chemical Society
of fabrication, self-sealing capability, chemical stability, high gas permeability, biological compatibility, and optical transparency. These characteristics have led to the widespread use of PDMS in microfabricated tools designed for biological applications, such as microfluidic systems. PDMS-based microfluidic devices have proven of value for ultra-small-volume reagent delivery, microscale chemical reactions, electrophoretic separations, and cell manipulations.2-7 The mechanical properties of PDMS make it ideal for fabrication of integrated devices. Components such as valves, pumps, switches, optical lenses, waveguides, and others can be fabricated directly on the device.1,8-10 Thus, PDMS has found wide utility in the fabrication of microdevices. The intrinsic properties of PDMS can be both beneficial and disadvantageous, depending on the intended purpose of the device. The mechanical and optical properties of PDMS lend themselves to a variety of uses particularly in the areas of biochemical and cellular assays. Due to the detrimental effects of nonpolar organic solvents, such as swelling and delamination, PDMS has been primarily used with aqueous solutions, but this does not present a significant drawback for most bioanalytical applications. The native hydrophobicity of the PDMS surface has been one of the biggest limitations in biological applications, as this property enhances nonspecific adsorption of biomolecules, limits cell attachment, and presents challenges in surface wetting and channel filling. Fortunately, the surface of PDMS can be chemically modified or physically masked by adsorption to meet the specific requirements of many applications. For example, the channel walls of PDMS microfluidic chips have been modified to (2) Bruin, G. J. M. Electrophoresis 2000, 21, 3931-3951. (3) Sia, S. K.; Whitesides, G. M. Electrophoresis 2003, 24, 3563-3576. (4) Whitesides, G. M.; Ostuni, E.; Shuichi, T.; Jiang, X.; Ingber, D. E. Annu. Rev. Biomed. Eng. 2001, 3, 335-373. (5) Effenhauser, C. S.; Bruin, G. J. M.; Paulus, A.; Ehrat, M. Anal. Chem. 1997, 69, 3451-3457. (6) Chiu, D. T.; Jeon, N. L.; Huang, S.; Kane, R. S.; Wargo, C. J.; Choi, I. S.; Ingber, D. E.; Whitesides, G. M. Proc. Nat. Acad. Sci. U.S.A. 2000, 97, 2408-2413. (7) Leclerc, E.; Sakai, Y.; Fujii, T. Biomed. Microdevices 2003, 5, 109-114. (8) Ng, J. M. K.; Gitlin, I.; Stroock, A. D.; Whitesides, G. M. Electrophoresis 2002, 23, 3461-3473. (9) Unger, M. A.; Chou, H. P.; Thorsen, T.; Scherer, A.; Quake, S. R. Science 2000, 288, 113-116. (10) Camou, S.; Fujita, H.; Fujii, T. Lab Chip 2003, 3, 40-45.
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enhance electrophoretic separations.11,12 Surface modification brings about several advantages that enhance the applicability of PDMS for microfluidic-based separations: generation of stable electroosmotic flow, enhanced surface hydrophilicity for aqueous samples, decreased adsorption of biomolecules, and improved peak resolution. A number of surface modification schemes have been reported to tailor the PDMS microchannels for this purpose.11,13-20 Another example of the beneficial value of surface modifications for PDMS is the performance of bioassays on a solid substrate. The use of a PDMS-based device for DNA assays has been reported in which the PDMS surface was modified by oxygen plasma followed by silanization.21 This modification generates reactive chemical groups that can be used to immobilize oligonucleotides on the PDMS surface. One of the most widespread uses for PDMS surface modification has been to generate favorable substrates for cell attachment and growth.22-26 Patterning of cell attachment sites on PDMS devices has found relevance for studies of cell growth as well as for high-throughput cell-based assays.26-31 As mentioned above, the surface of PDMS can be modified by both covalent and physical modification and a variety of methods are available for tailoring the surface properties of PDMS. Simple physical adsorption of a substance is often used to impart the desired surface property. This method has been employed to modify the inner surfaces of assembled microfluidic devices as well as for developing surfaces for cell attachment.11 Since adsorption has limitations, particularly in the stability of the coating, a variety of covalent modification schemes have been developed.11,12 A simple and direct method is to oxidize the PDMS (11) Makamba, H.; Kim, J. H.; Kwanseop, L.; Park, N.; Hahn, J. H. Electrophoresis 2003, 24, 3607-3619. (12) Dolnick, V. Electrophoresis 2004, 25, 3589-3601. (13) Hu, S.; Ren, X.; Bachman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Anal. Chem. 2002, 74, 4117-4123. (14) Hu, S.; Ren, X.; Bachman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Anal. Chem. 2004, 76, 1865-1870. (15) Xiao, D.; Le, T. V.; Wirth, M. J. Anal. Chem. 2004, 76, 2055-2061. (16) Roman, G. T.; Hlaus, T.; Bass, K. J.; Seelhammer, T. G.; Culbertson, C. T. Anal. Chem. 2005, 77, 1414-1422. (17) Chen, L.; Ren, J.; Bi, R.; Chen, D. Electrophoresis 2004, 25, 914-921. (18) Slentz, B. E.; Penner, N. A.; Regnier, F. E. J. Chromatogr., A 2002, 948, 225-233. (19) Berdichevsky, Y.; Khandurina, J.; Guttman, A.; Lo, Y. H. Sens. Actuators, B 2004, 97, 402-408. (20) Dou, Y. H.; Bao, N.; Xu, J. J.; Chen, H. Y. Electrophoresis 2002, 23, 35583566. (21) Liu, D.; Perdue, R. K.; Sun, L.; Crooks, R. M. Langmuir 2004, 20, 59055910. (22) Lee, J. N.; Jiang, X.; Ryan, D.; Whitesides, G. M. Langmuir 2004, 20, 1168411691. (23) Reyes, D. R.; Perruccio, E. M.; Becerra, S. P.; Locascio, L. E.; Gaitan, M. Langmuir 2004, 20, 8805-8811. (24) De Silva, M. N.; Desai, R.; Odde, D. J. Biomed. Microdevices 2004, 6, 219222. (25) Jiang, X. Y.; Takayama, S.; Qian, X. P.; Ostuni, E.; Wu, H. K.; Bowden, N.; LeDuc, P.; Ingber, D. E.; Whitesides, G. M. Langmuir 2002, 18, 32733280. (26) Toworfe, G. K.; Composto, R J.; Adams, C. S.; Shapiro, I. M.; Ducheyne, P. J. Biomed. Mater. Res. 2004, 71A, 449-461. (27) Li, N.; Tourovskaia, A.; Folch, A. Crit. Rev. Biomed. Eng. 2003, 31, 423488. (28) Taylor, A. M.; Rhee, S. W.; Tu, C. H.; Cribbs, D. H.; Cotman, C. W.; Jeon, N. L. Langmuir 2003, 19, 1551-1556. (29) Tan, J. L.; Liu, W.; Nelson, C. M.; Raghavan, S.; Chen, C. S. Tissue Eng. 2004, 10, 865-872. (30) Raghavan, S.; Chen, C. S. Adv. Mater. 2004, 16, 1303-1313. (31) Tourovskaia, A.; Figueroa-Masot, X.; Folch, A. Lab Chip 2005, 5, 14-19.
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surface with ultraviolet (UV) radiation, a UV/ozone treatment, or an oxygen plasma treatment to generate a hydrophilic silicone oxide layer.32-34 However, the hydrophilic nature of oxidized PDMS is only temporary as the migration of PDMS chains leads to recovery of the native hydrophobic state.35-37 The oxidized PDMS surface can be further modified with alkylsilanes via a silanization reaction of surface silanol groups.38 Multiple other strategies for the covalent, chemical modification of PDMS are now available.39 To generate coated PDMS surfaces with longterm stability, we have modified PDMS by two approaches, both of which employ UV-induced graft polymerization.13,14 Using a onestep method, PDMS is dipped into an aqueous solution containing sodium periodate, benzyl alcohol, and monomer(s) and then exposed to UV radiation for 1-4 h. UV radiation generates free radicals, which induce polymerization in the solution as well as on the surface. The resultant surface polymerization leads to a covalent chemical modification of the surface.13 The surface properties of PDMS can be tailored by combining monomers with different properties, for example, neutral and charged monomers.40 More recently, an efficient two-step process employing a photoinitiator adsorbed to the PDMS surface prior to UV-induced graft polymerization was demonstrated. This method enhances the rate of polymerization at the PDMS surface relative to that in the aqueous solution, permitting surface polymerization with only 5 min of UV exposure. Both methods have been used to tailor the surface properties of PDMS microfluidic devices for electrophoretic separations.13,14,40,41 In bioanalytical applications such as DNA assays and immunoassays, it is desirable to covalently modify PDMS on the microscale in order to place different chemical moieties on the same surface. Such selective patterning often requires a surface modification process conducted together with a micropatterning technique. A variety of techniques have been described for the deposition of materials on PDMS in microscale geometries, including microcontact printing (µCP), microfluidic-flow patterning, and physical masking.28,42-47 In almost all instances, the deposition involves adsorption of molecules to the PDMS surface (32) Huck, T. S. W.; Bowden, N.; Onck, P.; Pardoen, T.; Hutchinson, J. W.; Whitesides, G. M. Langmuir 2000, 16, 3497-3501. (33) Ouyang, M.; Yuan, C.; Muisener, R. J.; Boulares, A.; Koberstein, J. T. Chem. Mater. 2000, 12, 1591-1596. (34) Lai, J. Y.; Lin, Y. Y.; Denq, Y. L.; Chen, J. K. J. Adhes. Sci. Technol. 1996, 10, 231-242. (35) Fritz, J. L.; Owen, M. J. J. Adhes. 1995, 54, 33-45. (36) Hillborg, H.; Tomczak, N.; Olah, A.; Schonherr, H.; Vancso, G. J. Langmuir 2004, 20, 785-794. (37) Olah, A.; Hillborg, H.; Vancso, G. J. Appl. Surf. Sci. 2005, 239, 410-423. (38) Ferguson, G. S.; Chaudry, M. K.; Biebuyck, H. A.; Whitesides, G. M. Macromolecules 1993, 26, 5870-5875. (39) Uyama, Y.; Kato, K.; Ikada, Y. Adv. Polym. Sci. 1998, 137, 1-39. (40) Hu, S.; Ren, X.; Bachman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Langmuir 2004, 20, 5569-5574. (41) Hu, S.; Ren, X.; Bachman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Electrophoresis 2003, 24, 3679-3688. (42) Xia, Y. N.; Whitesides, G. M. Angew. Chem,, Int. Ed. Engl. 1998, 37, 551575. (43) Quist, A. P.; Pavlovic, E.; Oscarsson, S. Anal. Bioanal. Chem. 2005, 381, 591-600. (44) Bernard, A.; Renault, J. P.; Michel, B.; Bosshard, H. R.; Delamarche, E. Adv. Mater. 2000, 12, 1067-1070. (45) Bernard, A.; Michel, B.; Delamarche, E. Anal. Chem. 2001, 73, 8-12. (46) Dertinger, S. K. W.; Jiang, X. Y.; Li, Z. Y.; Murthy, V. N.; Whitesides, G. M. Proc. Nat. Acad. Sci. U.S.A. 2002, 99, 12542-12547. (47) Folch, A.; Jo, B. H.; Hurtado, O.; Beebe, D. J.; Toner, M. J. Biomed. Mater. Res. 2000, 52, 346-353.
rather than covalent attachment. Patterned, covalent surface modifications of inorganic substrates such as gold, silicon, and glass have been accomplished using µCP and a self-assembly monolayer process to pattern polymers, proteins, DNA, cells, and other entities.30,42,44,48-52 The success of this technique relies on the reactive surface property of the inorganic substrates; however, covalent modification of native PDMS using µCP has not found widespread use due to the inertness and hydrophobicity of PDMS. Diaz-Quijada and Wayner have demonstrated patterned, covalent modification of a PDMS surface.53 In this study, the PDMS surface was oxidized by initial photolithographic patterning with aluminum followed by exposure to ozone treatment; the oxidized regions were then further derivatized with (3-aminopropyl)triethoxysilane. This treatment generated a micropatterned surface to which biomolecules such as DNA could be coupled.53 Drawbacks to this elegant approach were the need for multiple steps and access to a cleanroom facility. In this paper, a simple benchtop method was developed for the patterned, covalent modification of PDMS surfaces. This method extends our previously published two-step, UV-induced surface graft polymerization by providing a scheme to pattern the surface on the micrometer scale without the need for cleanroom facilities.14 Use of a photomask to direct graft polymerization at selected locations on the PDMS surface permits simultaneous micropatterning and surface modification. Modification of the surface was confirmed by microscopic observation after staining with visible and fluorescent dyes, as well as measurement of the topography by atomic force microscopy (AFM). The efficacy of the micropatterned PDMS surface was demonstrated for two bioanalytical applications: patterning cell growth and antibody immobilization for heterogeneous immunoassays. EXPERIMENTAL SECTION Materials. The Sylgard 184 silicone elastomer kit was purchased from Dow Corning (Midland, MI). Precleaned glass slides (75 mm × 25 mm × 1 mm) were purchased from Corning Glass Works (Corning, NY). Iron oxide photoplates (Ferroxoplate, 3 in. × 3 in. × 0.060 in.) were from Towne Technologies Inc. (Somerville, NJ). 2-Methacryloxyethyltrimethylammonium chloride, poly(ethylene glycol) monomethoxyl acrylate (average molecular weight, 454), acrylic acid, benzophenone, benzyl alcohol, sodium periodate, 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide (EDC), toluidine blue, 6-aminofluorescein, rhodamine B, ethylenediamine, poly(L-lysine)-FITC labeled (MW 15 000-30 000), protein A, and protein A-fluorescein isothiocyanate conjugate staphylococcus aureus (protein A-FITC) were all obtained from Sigma-Aldrich (St. Louis, MO). Anti-green fluorescent protein (GFP) antibody was from Molecular Probes (Eugene, OR), and the purified recombinant enhanced GFP protein (rEGFP) was obtained from Clontech (Palo Alto, CA). All the reagents were (48) Xia, Y. N.; Mrksich, M.; Kim, E.; Whitesides, G. M. J. Am. Chem. Soc. 1995, 117, 9576-9577. (49) Kumar, A.; Whitesides, G. M. Appl. Phys. Lett. 1993, 63, 2002-2004. (50) Lange, S. A.; Benes, V.; Kern, D. P.; Horber, J. K. H.; Bernard, A. Anal. Chem. 2004, 76, 1641-1647. (51) Kane, R. S.; Takayama, S.; Ostuni, E.; Ingber, D. E.; Whitesides, G. M. Biomaterials 1999, 20, 2363-2376. (52) Husemann, M.; Mercerreyes, D.; Hawker, C. J.; Hedrick, J. L.; Shah, R.; Abbott, N. L. Angew. Chem,, Int. Ed. Engl. 1999, 38, 647-649. (53) Diaz-Quijada, G. A.; Wayner, D. D. M. Langmuir 2004, 20, 9607-9611.
Figure 1. Schematic of the two-step process for covalent micropatterning of PDMS.
used without further purification, except that acrylic acid was distilled under reduced pressure to remove the inhibitor (monomethyl ether of hydroquinone) and stored in a -20 °C freezer until use. Photomask Fabrication. Iron oxide photomasks with various micropatterns were fabricated according to the traditional microfabrication process.54 Micropatterning of PDMS by Photografting through a Mask. Glass-supported planar films of PDMS were made as follows. PDMS prepolymer (10:1 mixture of base/curing agent of Sylgard 184 kit) was mixed with hexane at a ratio of 50:50 w/w, and 2 mL of this mixture was spread on a glass slide. The slide was placed in a fume hood overnight to evaporate hexane after which the PDMS was cured in an 80 °C oven for 1 h. The PDMS together with the attached glass surface was cut into 25 × 25 mm pieces. The PDMS film thickness was estimated to be 500 µm. The surface modification of PDMS was carried out by a two-step process depicted in Figure 1. In the first step of this process, PDMS was immersed in 10 wt % benzophenone solution in varying ratios of water/acetone mixtures for 1 min, then immediately rinsed in methanol, and dried with a stream of nitrogen. During this treatment, benzophenone diffused into the PDMS surface. In the second step of this process, a photomask was used to direct the modification at the selected areas of the benzophenoneimplanted PDMS. Acrylic acid (40 µL of 10 wt % in water) with benzyl alcohol (0.5 wt %) and sodium periodate (0.5 mM) was added to the center of the PDMS. An iron oxide photomask was then slowly lowered over the PDMS surface. This PDMS/mask assembly was then placed inside a Loctite Zeta 7411 UV Flood System equipped with a 400-W metal halide lamp, with a distance between the lamp and photomask of 16 cm. The assembly was exposed to UV radiation for 25 min. The PDMS was detached (54) Rai-Choudhury, P., Ed. Handbook of Microlithography, Micromachining, and Microfabrication; SPIE Optical Engineering Press: Bellingham, WA, 1997.
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from the assembly, rinsed with deionized water, and immersed in acetone for 5 min. The goal of the acetone immersion was to remove unreacted benzophenone from the PDMS slab. The PDMS was then rinsed with water and incubated in sodium phosphate buffer (0.1 M, pH 8) overnight to remove poly(acrylic acid) (PAA), which was not covalently attached to the PDMS. Attempts at photografting other monomers (2-methacryloxyethyltrimethylammonium chloride or poly(ethylene glycol) monomethoxyl acrylate) were performed under the same conditions as that used for acrylic acid. Staining PDMS with a Dye. The grafted PDMS surface was observed by microscopy after staining with a visible dye (toluidine blue) or a fluorescent dye (6-aminofluorescein). Both toluidine blue and 6-aminofluorescein have positively charged amine groups and adsorb to the negatively charged carboxylate groups of PAA, but not to PDMS. The samples were immersed in 0.1% dye (wt %) in sodium phosphate buffer (0.1 M, pH 8) for 5 min, rinsed with water, and then dried in a stream of nitrogen. The dried samples were then imaged under transillumination on an inverted microscope (Nikon TE 300). Covalent Coupling of Molecules to PAA-Grafted PDMS Surfaces. Molecules possessing free amino groups were covalently linked to the carboxyl groups of grafted PAA via carbodiimide-mediated amide formation. A chamber was constructed by using Sylgard 184 to attach a silicon O-ring (24-mm outer diameter) to the modified PDMS sample. The chamber was filled with sodium phosphate buffer (500 µL of 0.02 M, pH 4.5) for 5 min. The buffer was removed, and EDC (2 wt %) in sodium phosphate buffer (0.02 M, pH 4.5, 500 µL) was added for 4 h to activate the carboxylate groups of PAA on the surface. After rinsing with phosphate-buffered saline (PBS) (138 mM NaCl, 27 mM KCl, 10 mM PO4, pH 7.4), the chamber was filled with a solution of the amine-containing molecule in PBS (200-500 µL). The concentration of the amine-containing molecule was 10 mg/ mL ethylenediamine, 1 mg/mL poly(L-lysine)-FITC labeled, and 0.1 mg/mL protein A-FITC. The reaction mixture was incubated overnight at room temperature with a glass slide covering the top of the chamber to prevent evaporation. The chamber was then rinsed with PBS, and the entire sample (including the chamber) was immersed in PBS and stored at 4 °C. Cell Culture. NIH 3T3 cells were grown at 37 °C in a humidified 5% CO2 atmosphere in Dulbecco’s modified eagle medium supplemented with fetal bovine serum (10%), and Lglutamine (584 mg/L). Penicillin (100 units/mL) and streptomycin (100 µg/mL) were added to the medium to inhibit bacterial growth. Cells were plated at concentrations determined empirically to produce approximately one cell per 1000× field of view on the day of the experiment and allowed to recover for 24 h. Immediately prior to use, the growth medium was removed from the cell chamber and replaced with PBS. Cells cultured on the PDMS surfaces were imaged under transillumination on an inverted microscope (Nikon TE 300). Antibody Immobilization. Antibody was immobilized to PAA grafted regions via two strategies. In the first strategy, anti-GFP antibody was directly coupled to the PAA surface via the carbodiimide-mediated reaction (described above) using a concentration of anti-GFP of 180 µg/mL in PBS. In the second strategy, protein A was first covalently coupled to PAA by the carbodiimide7542
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mediated reaction, using a protein A concentration of 500 µg/ mL. Anti-GFP (180 µg/mL) was then bound to surface-immobilized protein A by overnight treatment using 200 µL of the antiGFP solution. Samples prepared with both strategies were then rinsed with PBS and treated with 200 µL of rEGFP (166 µg/mL) for 2 h. The samples were then rinsed with PBS and observed by fluorescence microscopy (excitation 480 ( 20 nm, emission 535 ( 25 nm, Nikon TE300 microscope). AFM. The surface topography of the surface-modifed PDMS sample was acquired in air with a Nano-R AFM (Pacific Nanotechnologies Inc., Santa Clara, CA) in material sensing mode. RESULTS AND DISCUSSION Implantation of Benzophenone into the Surface of PDMS. Previous studies have shown that entrapment of the photoinitiator benzophenone at or just beneath a polymeric surface can enhance the efficiency of UV-induced graft polymerization at that surface.14,55 In those studies, acetone-induced swelling of PDMS led to the diffusion of benzophenone into the PDMS walls of a microchannel.14 When the microchannel was exposed to UV light, polymerization occurred primarily on the surfaces of the channel walls. UV exposure of the channel through a mask yielded a pattern with very poor spatial resolution (∼100 µm). Two reasons are likely to explain this poor resolution. Extensive entrapment of high levels of benzophenone occurs within the device walls. Activation of benzophenone and diffusion of that activated benzophenone into nonilluminated regions of the channel could then occur throughout the illumination time of the device. Second, release of benzophenone from the PDMS walls into the channel lumen may occur where the activated benzophenone could react with monomers. The activated monomers and growing polymer would then be free to diffuse to nearby nonilluminated channel regions and react with the PDMS walls. To improve the spatial resolution of patterning, the depth of entrapment of benzophenone was varied by dissolving the benzophenone (10 wt %) in a water/acetone mixture and titrating the ratio of water to acetone. Since water alone does not swell PDMS, increasing the percentage of water relative to that of acetone led to decreased PDMS expansion and, therefore, reduced the diffusion of benzophenone into the PDMS. The ratio of water/ acetone was varied over a range of 40:60 to 0:100 and the incubation time in the benzophenone was 1 min. The benzophenone concentration was fixed at 10 wt % since benzophenone becomes insoluble when the ratio of water/acetone is higher than 40:60. To visualize the extent of diffusion of a small hydrophobic molecule such as benzophenone into the PDMS surface, rhodamine B (MW 479), a hydrophobic fluorescent dye, was used in place of benzophenone (MW 182). PDMS was incubated with rhodamine B (10 wt %) in varying water/acetone mixtures in a manner identical to that used for benzophenone entrapment. Figure 2A shows the fluorescence image of the cross section of the PDMS surface implanted with rhodamine B. The fluorescence is visible to a depth of ∼100 µm and is at higher intensity near the PDMS surface relative to that deeper into the PDMS (Figure 2B). Rhodamine B enrichment in the PDMS was found to be stable after vigorous washing with water for several hours, although the (55) Deng, J. P.; Yang, W. T.; Ranby, B. J. Appl. Polym. Sci. 2001, 80, 14261433.
Figure 2. Implantation of small molecules within PDMS. (A) Fluorescence image of a cross section of the PDMS surface after treatment with 10% rhodamine B in a water/acetone (35:65 w/w) solution for 1 min. The dotted white line marks the air-PDMS interface. The fluorescence is seen to extend into the substance of the PDMS at the exposed surface. (B) Fluorescence intensity profile across the PDMS image in (A) showing infiltration of the dye into the PDMS to a depth of ∼100 µm.
rhodamine migrated out of the PDMS after immersion (
