Anal. Chem. 2004, 76, 1865-1870
Surface-Directed, Graft Polymerization within Microfluidic Channels Shuwen Hu,† Xueqin Ren,† Mark Bachman,‡,§ Christopher E. Sims,| G. P. Li,*,†,‡,§ and Nancy L. Allbritton*,†,|
Center for Biomedical Engineering, Integrated Nanosystems Research Facility, Department of Electrical and Computer Engineering, and Department of Physiology and Biophysics, University of California, Irvine, California 92697
We demonstrate a simple procedure to coat the surfaces of enclosed PDMS microchannels by UV-mediated graft polymerization. In prior applications, only disassembled channels could be coated by this method. This limited the utility of the method to coatings that could easily and tightly seal with themselves. By preadsorbing a photoinitiator onto the surface of PDMS microchannels, the rate of polymer formation at the surface was greatly accelerated compared to that in solution. Thus, a gel did not form in the lumen of enclosed microchannels. We demonstrate that the photoinitiator benzophenone remained on the surface of PDMS even after extensive washing. After addition of a variety of monomer solutions (acrylic acid, poly(ethylene glycol) monomethoxyl acrylate, or poly(ethylene glycol) diacrylate) and illumination with UV light, a stable, covalently attached surface coating formed in the microchannels. The electroosmotic mobility was stable in response to air exposure and to repeated cycles of hydration-dehydration of the coating. These surfaces also supported the electrophoretic separation of two model analytes. Placement of an opaque mask over a portion of the channel permitted photopatterning of the microchannels with a resolution of ∼100 µm. By using an appropriate mixture of monomers combined with masks, it should be possible to fabricate PDMS microfluidic devices with distinct surface properties in different regions or channels.
PDMS has several disadvantages including its extreme hydrophobicity, propensity to adsorb analytes, and porosity admitting molecules into the substrate itself. Consequently, there is a tremendous need for methods to rapidly and easily modify the surface properties of PDMS, particularly within the enclosed confines of a microfluidic channel. A number of approaches have been utilized to tailor the surface of PDMS microdevices, and a recent excellent review covers these strategies.2 Most of the coatings are difficult to micropattern (or even apply) within an intact microchannel. For example, chemical vapor deposition and UV-mediated grafting are capable of placing many different types of functional groups on the surface of a PDMS channel but require that the top and bottom pieces of the channel be coated prior to assembly.3,4 This limits the use of these methods to coatings that will readily seal with itself or another material. Other methods, for example, cerium(IV)-catalyzed polymerization, are also versatile in terms of placing different chemical groups on the surface of a PDMS microchannel but can be applied in intact, fully assembled channels.5 However, these coatings and others such as polyelectrolyte layers, lipid bilayers, and silanization are limited to patterning, which can be accomplished through the use of laminar flows.6-11 Alternative strategies to pattern molecules within an enclosed channel even at low resolution would be of great utility in the field of polymer microdevices. The use of light to initiate a chemical reaction has the potential to be used for surface patterning within a microchannel. A number
The use of poly(dimethylsiloxane) (PDMS) as a substrate for microfluidic devices has increased in popularity due to its straightforward and low fabrication cost.1 The durability and flexibility of PDMS make it an excellent material for the fabrication of valves, pumps, and other movable subcomponents of microfluidic devices. In addition, the compatibility of PDMS with living entities, such as mammalian cells, makes PDMS a material of choice for many biologic assays. Compared to other substrates,
(2) Makamba, H.; Kim, J. H.; Lim, K.; Park, N.; Hahn, J. H. Electrophoresis 2003, 24, 3607-3619. (3) Lahann, J.; Balcells, M.; Lu, H.; Rodon, T.; Jensen, K. F.; Langer, R. Anal. Chem. 2003, 75, 2117-2122. (4) Hu, S.; Ren, X.; Bachman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Anal. Chem. 2002, 74, 4117-4123. (5) Slentz, B. E.; Penner, N. A.; Regnier, F. E. J. Chromatogr., A 2002, 948, 225-233. (6) Kenis, P. J.; Ismagilov, R. F.; Takayama, S.; Whitesides, G. M.; Li, S.; White, H. S. Acc. Chem. Res. 2000, 33, 841-847. (7) Liu, Y.; Fanguy, J. C.; Bledsoe, J. M.; Henry, C. S. Anal. Chem. 2000, 72, 5939-5944. (8) Barker, S. L. R.; Ross, D.; Tarlov, M. J.; Gaitan, M.; Locascio, L. E. Anal. Chem. 2000, 72, 5925-5929. (9) Dou, Y. H.; Bao, N.; Xu, J. J.; Chen, H. Y. Electrophoresis 2002, 23, 35583566. (10) Linder, V.; Verpoorte, E.; Thormann, W.; de Rooij, N. F.; Sigrist, H. Anal. Chem. 2001, 73, 4181-4189. (11) Takayama, S.; Ostuni, E.; Qian, X.; McDonald, J. C.; Jiang, X.; LeDuc, P.; Wu, M. H.; Ingber, D. E.; Whitesides, G. M. Adv. Mater. 2001, 13, 570574.
* Corresponding authors. N.L.A.: (phone) 949-824-6493; (e-mail) nlallbri@ uci.edu; (fax) 949-824-8540. G.P.L.: (phone) 949-824-4194; (e-mail)
[email protected]; (fax) 949-824-3732. † Center for Biomedical Engineering. ‡ Integrated Nanosystems Research Facility. § Department of Electrical and Computer Engineering. | Department of Physiology and Biophysics. (1) Ng, J. M. K.; Gitlin, I.; Stroock, A. D.; Whitesides, G. M. Electrophoresis 2002, 23, 3461-3473. 10.1021/ac049937z CCC: $27.50 Published on Web 03/03/2004
© 2004 American Chemical Society
Analytical Chemistry, Vol. 76, No. 7, April 1, 2004 1865
of investigators have used UV-mediated grafting within enclosed microchannels.12-14 However, in all cases, UV-mediated grafting predominantly formed gel-based structures rather than surface coatings when applied to intact channels. Hu et al. did utilize UVmediated grafting to coat PDMS surfaces, but these channels had to be assembled after the coating process.4,15 When applied to an intact channel, this method also forms a gel rather than a surface coating. In addition, the small amount of free monomer within the channel is rapidly consumed by gel formation, leaving little to no monomer available for polymerization onto the surface. The gel also restricts diffusion of any residual monomer to the surface of the channels. Unless the channel can be disassembled and reassembled, the gel forms a plug in the channel restricting the movement of buffer and analytes. In this paper, UV-mediated grafting was directed toward the formation of a surface coating by adsorption of a photoinitiator onto the PDMS surface. Free photoinitiator was then removed from the luminal solution prior to graft polymerization. The adsorbed photoinitiator accelerated the rate of formation of the polymer on the surface relative to that in solution during the subsequent grafting step. Successful adsorption of benzophenone (the photoinitiator) to PDMS was demonstrated by measurement of the infrared absorption spectra of the PDMS surface. Multiple different monomers were successfully polymerized on the surface of intact PDMS microchannels. The properties of these covalently attached coatings were characterized by a variety of microscopic methods as well as by measurement of the electroosmotic mobility. The ability of a cross-linked coating to support the electrophoretic separation of two model peptides was also assessed. Last, low-resolution photopatterning of grafting within intact channels using a mask was demonstrated and characterized by microscopy. EXPERIMENTAL SECTION Reagents. Sylgard 184 was purchased from Dow Corning (Midland, MI), and silicon nitride-coated silicon wafers were obtained from Wafernet Inc. (San Jose, CA). Acrylic acid (AA), poly(ethylene glycol) monomethoxyl acrylate (PEG), 2-(methacryloxy)ethyltrimethylammonium chloride (MATC), poly(ethylene glycol) diacrylate (DiPEG), and benzyl alcohol were all obtained from Aldrich and used without further purification. All fluorescent reagents were purchased from Molecular Probes (Eugene, OR). Peptides were synthesized by the Beckman Peptide and Nucleic Acid Facility at Stanford University (Stanford, CA) and labeled with fluorescein as described previously.16 The peptide sequences were fluorescein-DLDVPIPGRFDRRVSVAAE (F-calc) and fluorescein-AEEEIYGEFEAKKKK (F-src).17,18 All other reagents and materials were purchased from Fisher Scientific (Pittsburgh, PA). (12) Seong, G. H.; Zhan, W.; Crooks, R. M. Anal. Chem. 2002, 74, 3372-3377. (13) Yu, C.; Xu, M.; Svec, F.; Fre´chet, J. M. J. J. Polym. Sci. Part A: Polym. Chem. 2002, 40, 755-769. (14) Moorthy, J.; Beebe, D. J. Lab Chip 2003, 3, 62-66. (15) Hu, S.; Ren, X.; Bachman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Electrophoresis 2003, 524, 3679-3688. (16) Lee, C. L.; Linton, J.; Soughayer, J. S.; Sims, C. E.; Allbritton, N. L. Nat. Biotechnol. 1999, 17, 759-762. (17) Nair, S. A.; Kim, M. H.; Warren, S. D.; Choi, S.; Songyang, Z.; Cantley, L. C.; Hangauer, D. G. J. Med. Chem. 1995, 38, 4276-4283. (18) Klee, C. B.; Ren, H.; Wang, X. J. Biol. Chem. 1998, 273, 13367-13370.
1866
Analytical Chemistry, Vol. 76, No. 7, April 1, 2004
Device Fabrication. Microfluidic channel patterns and the corresponding master were designed and fabricated as described previously.4 Sylgard 184 PDMS prepolymer was mixed thoroughly with its cross-linking catalyst at 10:1 (w/w) and degassed by vacuum. The polymer was cast against the silicon mold and polymerized at 70 °C for 1 h. After curing, the PDMS was peeled from the mold and holes (3.5-mm diameter) were punched into the polymer to create access ports and reservoirs. Flat PDMS substrates were obtained by casting the polymer mixture on a clean, flat surface. Final polymerization of the PDMS was performed by placing the pieces in a 65 °C oven overnight. The micromolded PDMS was sealed against a flat, PDMS substrate. UV-Graft Polymerization of PDMS Films. Films of PDMS with or without imprinted microchannels were grafted as described previously.15 The total monomer concentration was 10 wt %. After polymer grafting, the top and bottom portions of the microchannel were assembled by pressing them together manually. UV-Graft Polymerization of Microchannels without Adsorbed Benzophenone. Intact PDMS microchannels were filled with an aqueous solution containing NaIO4 (0.5 mM), benzyl alcohol (0.5 wt %), and monomer at the indicated concentration. The microchannel was placed in a custom-built irradiator (200-W mercury lamp) for the time indicated. The distance between the sample and the lamp was 5 cm. Uniform UV exposure was ensured by rotating the films under the UV source. UV-Graft Polymerization of Microchannels with Preadsorbed Benzophenone. An assembled microchannel was rinsed with benzophenone (10% in acetone) for 30 s. The channel was then washed with ∼100 channel volumes of water for 2 min at 22 °C. A solution of monomers (10% total monomer concentration in water), NaIO4 (0.5 mM), and benzyl alcohol (0.5 wt %) was loaded into the channel, which was then irradiated with ultraviolet light for 5 min. The channel was then washed extensively with water Toluidine Blue Staining. The top and bottom halves of a PDMS microdevice were separated. The pieces were washed in water (22 °C, 2 h) and dried under a vacuum. A double-edged razor blade was then used to slice the PDMS film perpendicular to the length of the channel. The cross-sectional piece of PDMS was then immersed in a 1% toluidine blue solution for 5 min. After rinsing with water, the PDMS was dried under a vacuum. Measurement of Infrared Absorption by Total Attenuated Reflection (ATR-IR). A flat piece of PDMS was immersed in acetone with or without benzophenone (10%) for 10 min. The PDMS film was washed in water at 80 °C for 10 h and then dried under a vacuum for 24 h. The ATR-IR spectra of PDMS films on a wedged germanium crystal were recorded using a single-beam spectrometer (Nicolet Magna-IR 860 spectrometer) equipped with a helium-neon laser, a triglycine sulfate detector, and a ZnSe reflection element. Spectra were recorded at 4-cm-1 resolution, and 4096 scans were collected per trace. A single-beam reference spectrum of a freshly cleaned germanium crystal was recorded before the measurements and used as the background spectrum. A water spectrum was also recorded, scaled empirically, and subtracted from the PDMS spectra to remove the water peaks in the region of 3500 cm-1.
Measurement of µeo. The current monitoring method was used to measure µeo in the microfabricated channels.4,19 The two buffers used were 20 mM phosphate (pH 7.0) and 18 mM phosphate (pH 7.0). Four measurements were performed on each device. The channel width and height was 60 µm. To measure the stability of µeo after exposure to air, the channels were flushed with water, dried under a vacuum, and then exposed to air at room temperature. At the times indicated, the channels were filled with aqueous buffer and µeo was measured. After the measurement, the channels were flushed with water, dried under a vacuum, and again exposed to air at room temperature until the next measurement of µeo. Electrophoresis of Peptides. Electrophoresis of peptides was performed on a simple “cross” chip with four reservoirs as previously described.15 The channels were 60 µm wide and 60 µm deep. The “injection” channel was 2 cm in length and was bisected by the “separation” channel, which was 3.5 cm in length. After movement of the sample through the entire “sample” channel, voltages were applied to the four reservoirs to move a “tightly pinched” sample plug into the separation channel.20 The sample plug was electrophoresed through the separation channel by application of 1000 V across the length of the channel. The fluorescence detection zone was located 2 cm downstream from the intersection of the injection and separation channels. Fluorescence was detected as described previously with a cooled CCD camera using an integration time of 100 ms.15 The electrophoretic buffer was Tris (25 mM) and glycine (192 mM) at pH 8.4. All solutions were degassed by sonication for 10 min immediately prior to use. Scanning Electron Microscopy. A Hitachi S-520 SEM was used to measure the morphology of the surfaces. The applied voltage was 1000 V. A thin layer of platinum was sputter-deposited onto the sample prior to imaging. RESULTS AND DISCUSSION UV-Grafting within an Enclosed Microchannel (“OneStep” Process). To determine whether the walls of a microchannel could be coated with polymer after the channel was fully assembled, the top and bottom halves of a PDMS microfluidic device were fabricated and sealed together (Figure 1A). Acrylic acid, NaIO4, and benzyl alcohol were loaded into the channel, which was then exposed to ultraviolet light for 3 h (Figure 1B). The top half of the PDMS microdevice, which contained the imprinted microchannel, was separated from the bottom flat portion and stained with toluidine blue. The channel always contained a gel that stained darkly with toluidine blue (n > 15) (Figure 1D). Since toluidine blue possesses a net positive charge, the gel was most likely composed of poly(acrylic acid), which is negatively charged. The hydrophobic PDMS substrate did not bind the dye. It was not possible to determine whether any surface coating also formed since the gel occupied the channel lumen. Varying the concentration of the acrylic acid, benzyl alcohol, or NaIO4 or the time of UV exposure did not result in the preferential formation of a surface coating rather than a gel throughout the lumen. Polymerization in the solution within the channel was faster than that at the channel wall. To remove the rapidly forming (19) Huang, X.; Gordon, M. J.; Zare, R. N. Anal. Chem. 1988, 60, 1837-1838. (20) Alarie, J. M.; Jacobson, S. C.; Culbertson, C. T.; Ramsey, J. M. Electrophoresis 2000, 21, 100-106
Figure 1. UV-grafting of microchannels. (A) Schematic of an intact microchannel composed of PDMS. (B) Schematic of the one-step photografting method in an enclosed microchannel. (C) Schematic of the two-step photografting method in an enclosed microchannel. (D) Transmitted light image of a cross section through the top half of a microchannel grafted by the one-step method and then stained with toluidine blue. The channel was exposed to UV light for 3 h. (E) Transmitted light image of a cross section through the top half of a microchannel grafted by the two-step method and then stained with toluidine blue. The channels in panels D and E were 60 µm wide and 60 µm deep.
polymer from the solution, acrylic acid (10%) was flowed under pressure through the microchannel while the channel was illuminated with ultraviolet light. After 5 h, a thin surface layer of poly(acrylic acid) was observable by light microscopy after staining with toluidine blue (data not shown). However, in addition to the very long irradiation time, the channel was prone to becoming clogged with the nascent polymers forming in the solution. It was not possible to achieve a reproducible surface coating by this method. UV-Grafting within an Enclosed Microchannel with Preadsorbed Photoinitiator (“Two-Step” Process). Ranby and colleagues reported a method to enhance the efficiency of polymerization on the surface of polyurethane.21 Benzophenone, a photoinitiator, was adsorbed onto polyurethane followed by photografting of a monomer. Localization of the photoinitiator to the surface of the material enhanced the rate of polymerization on the surface relative to that in the solution. To determine (21) Deng, J. P.; Yang, W. T.; Ranby, B. J. Appl. Polym. Sci. 2001, 80, 14261433.
Analytical Chemistry, Vol. 76, No. 7, April 1, 2004
1867
Figure 2. Localization of the grafted coating. (A) Schematic of a cross section through a microchannel and a transmitted light image of a cross section through the bottom portion of a microchannel. The channel was grafted with acrylic acid by the two-step method and then stained with toluidine blue. (B) Schematic of a top view of the bottom piece of a grafted channel and scanning electron micrograph of the bottom piece of a microchannel. The channel was also grafted by the two-step process but was not cross-sectioned or dye-stained. In both (A) and (B), the PDMS is green and the graft is dark blue.
whether benzophenone might also adsorb to PDMS, a flat piece of PDMS was incubated with benzophenone (10% in acetone) for 30 s and then washed in water at 80 °C for 10 h. The infrared absorption spectra of the surface was then measured by ATR-IR. The spectra of benzophenone-exposed PDMS showed a sharp peak at 1667 cm-1, which was not present in the spectra of PDMS films treated with only acetone (data not shown). The peak at 1667 cm-1 corresponds to the stretching vibration of the carbonyl group of benzophenone.22 Benzophenone was successfully adsorbed to the surface of the PDMS film. In addition, it was also possible that the benzophenone might have migrated into the PDMS substrate since hydrophobic molecules can diffuse into the bulk material of hydrophobic, porous polymers such as PDMS.21,23 To determine whether adsorbed benzophenone would enhance the rate of polymer formation on the surface of the PDMS microchannel relative to that in the solution, an assembled microchannel was rinsed with benzophenone and then washed with water (see methods, Figure 1A,C). A solution of acrylic acid was loaded into the channel, which was then irradiated with ultraviolet light for 5 min. The top half of the PDMS microdevice was separated from the bottom flat portion and stained with toluidine blue (Figure 1C). A surface coating that stained with toluidine blue was present on the inner wall of the channel (n > 20) (Figure 1E). No plug of gel was ever observed within the microchannel. The rate of polymer formation on the inner wall now greatly exceeded that forming within the solution. Since the benzophenone may have diffused into the bulk PDMS, the poly(acrylic acid) might be present predominantly within the PDMS bulk rather than as a surface layer extending into the channel lumen. To determine the location of the poly(acrylic acid), the lower half of the separated PDMS channel was also stained with toluidine blue. The width of the toluidine bluestained polymer strip shown in Figure 2A was 60 µm, the same width as the microfabricated channel. In addition, the poly(acrylic (22) Garcia, M. V.; Redondo, M. I. Spectrochim. Acta, Part A 1987, 43A, 879885. (23) Ocvirk, G.; Munroe, M.; Tang, T.; Oleschuk, R.; Westra, K.; Harrison, D. J. Electrophoresis 2000, 21, 107-115.
1868
Analytical Chemistry, Vol. 76, No. 7, April 1, 2004
acid) layer clearly extends out 5 µm above the PDMS substrate suggesting that, under these conditions, a surface coating is deposited onto the PDMS. When UV-grafted under the same conditions, channels without adsorbed benzophenone showed no toluidine blue staining, suggesting that little to no polymer formed on the surface. Shown in Figure 2B is a scanning electron micrograph of the bottom piece of a photografted microchannel with prior adsorption of benzophenone. The grafted strip was 60 µm in width (the same width as the microchannel) with sharp edges, suggesting that the polymer graft did not infiltrate between the two sealed pieces of PDMS or down into the PDMS bulk. Similar results were also obtained with other monomers, for example, PEG and MATC (data not shown). Electroosmotic Fluid Flow in Channels Grafted with the Two-Step Process. To assess the extent to which the two-step coating altered the surface properties of the enclosed microchannels, µeo was measured in microchannels grafted by the two-step” method with either AA or PEG alone or with a mixture of PEG, AA, and DiPEG in a ratio of 20:3:2. The total monomer concentration (10%) was identical for all three groups. µeo was (3.08 ( 0.14) × 10-4 (AA alone), (0.85 ( 0.06) × 10-4 (PEG alone), and (2.83 ( 0.16) × 10-4 cm2/V‚s (PEG/AA/DiPEG). As expected, the AAgrafted microchannel had the greatest electroosmotic fluid flow and all possessed electroosmotic fluid flows between that of native and oxidized PDMS.4 These results are similar to that obtained when the two halves of a microchannel are grafted separately by the one-step process and then assembled into a complete channel.4.15 To determine whether the electroosmotic properties of the grafted microchannels were stable over time, µeo was measured at varying times after exposure of two-step grafted channels to air. Microchannels were grafted with PEG or a mixture of PEG, AA, and DIPEG (20:3:2). The channels were filled with buffer, and µeo was measured. After 1 h, µeo was again measured in the buffer-filled channels. The channels were then filled with air. At varying subsequent times, the channels were refilled with buffer and µeo was measured again. After each of these measurements of µeo, the channels were refilled with air. µeo for the PEG-grafted and PEG/AA/DiPEG-grafted channels remained constant when exposed to air for long periods of time (Figure 3). These results are also similar to that obtained when the two halves of a microchannel are grafted separately by the one-step process and then assembled into a complete channel.4,15 The two-step process yields coatings with stability and properties similar to those obtained when channels are grafted in a single step and then assembled. Electrophoretic Separations on Microchannels Grafted by the Two-Step Process. To determine whether the grafted microchannels were suitable for electrophoretic applications, a microfluidic device with “cross” injector was constructed and grafted with PEG/AA/DiPEG (20:3:2).15 Two fluorescent peptides, commonly used as enzyme substrates in cell biology, F-calc and F-src, were loaded into the microdevice, injected into the separation channel, and electrophoresed within the device. Fluorescence in the separation channel was detected with a cooled CCD camera. No adsorption to the channel walls was visible for either peptide. Each peptide yielded a single peak when electrophoresed alone. When co-mixed, two widely separated peaks were always detected
Figure 3. Stability of µeo in microchannels grafted with the twostep method. Microchannels were grafted with either PEG or PEG/ AA/DiPEG. The magnitude of µeo was measured after exposure of PEG-grafted (open squares) or PEG/AA/DiPEG-grafted (open circles) microchannels to air. After grafting the channels, µeo was measured immediately (1 h time), and the devices were filled with water and stored in water. After the second measurement of µeo at 2 h, the channels were filled with air and stored in air. After all subsequent measurements of µeo, the channels were again filled with air.
Figure 4. Separation of peptides in microchannels grafted with the two-step method. PDMS channels were grafted with PEG, AA, and DiPEG. A mixture of F-src (50 µM) and F-calc (50 µM) was loaded into the device, injected, and separated. The fluorescence of the analytes in the separation channel was measured and plotted against time. The initial peak (52 s) is due to F-src and the second peak (92 s) to F-calc.
(Figure 4). F-src possessed a migration time of 52 ( 1 s with an efficiency of 9300 ( 700 theoretical plates (n ) 6). F-calc had a migration time of 92 ( 3 s and an efficiency of 5200 ( 400 theoretical plates (n ) 6). The migration times and separation efficiency of each peptide was the same irrespective of whether it was electrophoresed alone or as a mixture. The reproducibility of the migration time was similar to that achieved when these peptides were electrophoresed in PDMS devices coated with the one-step grafting method and then assembled.15 The efficiency of separation, however, was roughly half that reported on the PDMS devices coated with the one-step grafting method and then assembled. The devices described previously by Hu et al. had undergone extensive development targeted at separation of these enzyme-substrate peptides.15 It is likely that similar results could be achieved on the two-step grafted devices after a similar optimization of the coating procedure. Spatially Localized Photografting with the Two-Step Method. An advantage of ultraviolet-mediated grafting is the potential to pattern a surface using a mask. To determine whether
Figure 5. Spatially localized photografting of a microchannel. (A) An opaque mask was placed over the left portion of a channel that was coated with poly(acrylic acid) by the two-step method. Although the entire channel (60 µm wide and 60 µm deep) was coated with benzophenone and contained acrylic acid, the mask prevented the left half of the channel from being illuminated with UV light. The channel was dissembled and stained with toluidine blue. The upper portion of the microchannel containing the imprinted channel groove was photographed using transmitted light microscopy. (B) A channel (30 µm wide and 30 µm deep) was masked and coated with poly(acrylic acid) as in A). The channel was disassembled and the bottom piece of the channel was photographed under reflected light. The bottom piece was not stained with toluidine blue. Instead, the reflected light was used to highlight the edges of the protruding strip of polymer coating on the right half of the channel.
it might be possible to pattern a surface coating within an enclosed microchannel, benzophenone was preadsorbed throughout a microchannel. The channel was then filled with acrylic acid (10%). An opaque mask was placed over half of the channel, and the channel was illuminated with UV light for 5 min. The channel was disassembled, washed, and stained with toluidine blue. Shown in Figure 5A is an image acquired using transmitted light microscopy. The illuminated portion of the channel stained a deep blue while the masked side did not bind the dye. The transition region between the blue-stained and unstained channel sections was ∼100 µm (n ) 5). To further examine the length of the transition region, the lower portion of a channel was similarly masked and grafted with acrylic acid. The PDMS film was photographed by utilizing reflected light to highlight the coating (Figure 5B). The raised profile of the poly(acrylic acid) coating was clearly visible in the section of the channel that was not masked. No coating was present in the masked section. Again the transition zone at the interface between the coated and uncoated portions of the channel was ∼100 µm (n ) 6). The bottom portion of a similarly prepared channel was also examined by scanning electron microscopy. The elevated poly(acrylic acid) strip was apparent, and the transition region between the coated and uncoated channel regions was of dimensions similar to that in the transmitted and reflected light images. These data suggest that the spatial resolution of the mask combined with UV grafting was roughly 100 µm. However, improved resolution might be obtained using a collimated light source. CONCLUSIONS We demonstrated a simple method to selectively photograft a polymer onto the inner surface of an intact microchannel. The Analytical Chemistry, Vol. 76, No. 7, April 1, 2004
1869
formation of a gel within the microchannel was eliminated by adsorbing a photoinitiator onto the surface, thereby accelerating the rate of polymer formation on the surface relative to that in the solution. This coating process allows the PDMS microchannel to be sealed reversibly (native PDMS) or irreversibly (oxidized PDMS) and the coating then applied. The coating need not seal with itself as was the case in prior applications of UV-mediated grafting. The method was also very fast, requiring minutes to perform. In addition, the thickness of the coating should be variable by titrating either the photoinitiator concentration, the monomer concentration, or the UV-irradiation time. We demonstrated the feasibility of the grafting process with the single monomers, AA, PEG, and MATC, and with mixtures of monomers including cross-linkers. The coatings that are covalently linked to the PDMS were stable over time in response to repeated drying and rehydration. The polymeric coatings were of high quality in that they supported the separation of substrate peptides not separable on native or oxidized PDMS surfaces.15 In addition, the feasibility of coating only a portion of a channel with a polymer was demonstrated. The spatial resolution of the
1870
Analytical Chemistry, Vol. 76, No. 7, April 1, 2004
patterning method was ∼100 µm, which is sufficient for many different applications, in particular biological assays. For example, using this strategy in a sequential manner should yield channels with regions possessing different polymer surface coatings. Portions of a channel could be coated with a positive surface charge to enhance cell adhesion while other regions receive a material such as PEG to reject cell attachment. Alternatively portions of a channel may be coated with functional groups capable of reacting with and immobilizing proteins while a protein-resistant polymeric coating is applied the remainder of the channel. ACKNOWLEDGMENT This research was supported by the NIH (RR/CA14892, GM57015) and DARPA (N66001-01-C8014). The authors thank Gavin Meredith, Ruisheng Chang, Wei Wei, and Sharon Ma for technical advice and assistance.
Received for review January 10, 2004. January 17, 2004. AC049937Z
