On-Chip Micropatterning of Plastic (Cylic Olefin Copolymer, COC

Langmuir 2007, 23, 1577-1583

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On-Chip Micropatterning of Plastic (Cylic Olefin Copolymer, COC) Microfluidic Channels for the Fabrication of Biomolecule Microarrays Using Photografting Methods Qiaosheng Pu,† Olufemi Oyesanya,† Bowlin Thompson,‡ Shantang Liu,† and Julio C. Alvarez*,† Department of Chemistry, Virginia Commonwealth UniVersity, P.O. Box 842006, Richmond, Virginia 23284-2006, and Department of Mechanical Engineering, Virginia Commonwealth UniVersity, P.O. Box 843015, Richmond, Virginia 23284-3015 ReceiVed September 11, 2006. In Final Form: NoVember 7, 2006 This paper reports on the surface modification of plastic microfluidic channels to prepare different biomolecule micropatterns using ultraviolet (UV) photografting methods. The linkage chemistry is based upon UV photopolymerization of acryl monomers to generate thin films (0.01-6 µm) chemically linked to the organic backbone of the plastic surface. The commodity thermoplastic, cyclic olefin copolymer (COC) was selected to build microfluidic chips because of its significant UV transparency and easiness for microfabrication by molding techniques. Once the polyacrylic films were grafted on the COC surface using photomasks, micropatterns of proteins, DNA, and biotinlated conjugates were readily obtained by surface chemical reactions in one or two subsequent steps. The thickness of the photografted films can be tuned from several nanometers up to several micrometers, depending on the reaction conditions. The micropatterned films can be prepared inside the microfluidic channel (on-chip) or on open COC surfaces (off-chip) with densities of functional groups about 10-7 mol/cm2. Characterization of these films was performed by attenuated-total-reflectance IR spectroscopy, fluorescence microscopy, profilometry, atomic force microscopy, and electrokinetic methods.

Introduction Herein we report the surface micropatterning of biomolecules inside plastic microfluidic chips using ultraviolet (UV) photoinduced methods. We selected cyclic olefin copolymer (COC) as building material because its significant UV transparency1,2 allows photolithographic interrogation of the COC surface under on-chip conditions. Furthermore, commodity thermoplastics like COC that permit easy replication of micrometer features by embossing or molding techniques hold the promise for the mass production of disposable microfluidic sensors.3-9 The patterning process relies on photoinduced polymerization of acryl monomers * To whom correspondence should be addressed. Voice: 804-828-3521; fax: 804-828-8599; e-mail: [email protected]. † Department of Chemistry. ‡ Department of Mechanical Engineering. (1) Rohr, T.; Ogletree, D. F.; Svec, F.; Frechet, J. M. J. Surface Functionalization of Thermoplastic Polymers for the Fabrication of Microfluidic Devices by Photoinitiated Grafting. AdV. Funct. Mater. 2003, 13, 264-270. (2) Khanarian, G. Optical Properties of Cyclic Olefin Copolymers. Opt. Eng. 2001, 40, 1024-1029. (3) Soper, A. S.; Ford, S. M.; McCarley, R. L.; Kelly, K.; Murphy, M. C. Polymetric Microelectromechanical Systems. Anal. Chem. 2000, 72, 643A651A. (4) Wang, Y.; Bikas, V.; Farquar, H. D.; Stryjewski, W.; Hammer, R. P.; McCarley, R. L.; Soper, A. S.; Cheng, Y.-W.; Barany, F. Microarrays Assembled in Microfluidic Chips Fabricated from Poly(methyl methacrylate) for the Detection of Low-Abundant DNA Mutations. Anal. Chem. 2003, 75, 1130-1140. (5) Liu, Y.; Ganser, D.; Schneider, A.; Liu, R.; Grodzinski, P.; Kroutchinina, N. Microfabricated Polycarbonate CE Devices for DNA Analysis. Anal. Chem. 2001, 73, 4196-4201. (6) Martynova, L.; Locascio, L. E.; Gaitan, M.; Kramer, G. W.; Christensen, R. G.; MacCrehan, W. A. Fabrication of Plastic Microfluidic Channels by Impriniting Methods. Anal. Chem. 1997, 69, 4783-4789. (7) McCormick, R. M.; Nelson, R. J.; Alonso-Amigo, M. G.; Benvegnu, D. J.; Hooper, H. H. Microchannel Electrophoretic Separations of DNA in InjectionMolded Plastic Substrates. Anal. Chem. 1997, 69, 2626-2630. (8) Xu, J.; Locascio, L. E.; Gaitan, M.; Lee, C. S. Room-Temperature Imprinting Method for Plastic Microchannel Fabrication. Anal. Chem. 2000, 72, 19301933. (9) Klintberg, L.; Svedberg, M.; Nikolajeff, F.; Thornell, G. Fabrication of a Paraffin Actuator Using Hot Embossing of Polycarbonate. Sens. Actuators, A 2003, 103, 307-316.

to produce cross-linked films covalently attached to the carbon atoms on the COC surface. Derivatization with target biomolecules such as DNA, proteins, and biotinlated conjugates is readily achieved in one or two subsequent steps. Despite the preliminary work on photografting of plastics,1,10-14 there are no reports on biomolecule immobilization on COC surfaces. Here, we describe the first example of immobilized biomolecules in COC microfludic channels. Experiments indicate that on-chip derivatization in selected areas of the COC microchannel can be easily done to produce films with sub-micrometer thickness but still maintaining a significant number of pendent groups for subsequent functionalization, that is, 10-7 mol/cm2. The compromise between these two parameters is very important for optimizing intrafilm diffusion in sensors on the basis of capture and release of analytes. In comparison to other immobilization strategies in microfluidic channels, such as hydrogel plugs or trapped coated microspheres,15,16 the method described here offers two advantages. First, the diffusion paths within these films are shorter than in (10) Ranby, B.; Yang, W. T.; Tretinnikov, O. Surface Photografting of Polymer Fibers, Films and Sheets. Nucl. Instrum. Methods Phys. Res., Sect. B 1999, 151, 301-305. (11) Deng, J. P.; Yang, W. T.; Ranby, B. Surface Photografting Polymerization of Vinyl Acetate (VAc), Maleic Anhydride, and Their Charge Transfer Complex. I. VAc(1). J. Appl. Polym. Sci. 2000, 77, 1513-1521. (12) Deng, J. P.; Yang, W. T.; Ranby, B. Surface Photografting Polymerization of Vinyl Acetate (VAc), Maleic Anhydride, and Their Charge Transfer Complex. II. VAc(2). J. Appl. Polym. Sci. 2000, 77, 1522-1531. (13) Yang, T. C. K.; Lin, S. S. Y.; Chuang, T. H. Kinetic Analysis of the Thermal Oxidation of Metallocene Cyclic Olefin Copolymer (mCOC)/TiO2 Composites by FTIR Microscopy and Thermogravimetry. Polym. Degrad. Stab. 2002, 78, 525-532. (14) Yang, W. T.; Ranby, B. Bulk Surface Photografting Process and its Applications. I. Reactions and Kinetics. J. Appl. Polym. Sci. 1996, 62, 533-543. (15) Zhan, W.; Seong, G. H.; Crooks, R. M. Hydrogel-Based Microreactors as a Functional Component of Microfluidic Systems. Anal. Chem. 2002, 74, 4647-4652. (16) Seong, G. H.; Zhan, W.; Crooks, R. M. Fabrication of Microchambers Defined by Photopolymerized Hydrogels and Weirs within Microfluidic Systems: Application to DNA Hybridization. Anal. Chem. 2002, 74, 33723377.

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a hydrogel plug which may occupy a significant portion of the channel cross section (several µm); second, pressure buildup caused by blockage of flow because of trapped microspheres or plugs is not present. Other advantages that stem from using a photografting method in COC channels are the following: the location, size, and shape of the modified area can be adjusted by using photomasks; functionalities such as carboxyl or amine groups that are useful for attaching biomolecules are directly obtainable from the photopatterning process; the film thickness can be controlled from 0.01 to a 6 µm by tuning the reaction conditions; the avidin-biotin linkage chemistry can be easily implemented allowing the attachment of virtually any biotinlated conjugate commercially available; and finally, the patterning can be performed both inside the COC microchannels (on-chip) and on open COC surfaces (off-chip). Our general goal in this project is to make microfluidic sensors that capitalize from these advantages and that can be used in low-volume and high-speed bioanalysis.17,18 Recently, we have reported the use of chemical coupling between electrochemical reactions as a way to enhance electrochemical sensitivity in microfluidic systems.19 The micropatterned areas inside the channel that we describe here will become chemical stations for the release or capture of analytical probes in coupled electrochemical detection schemes. The performance and versatility of a plastic material for microfluidic sensing applications is directly related to desirable features such as a surface chemical structure compatible with photoinduced linkage chemistries, good adhesion of metals for making electrochemical sensors, low background fluorescence for optical-based sensors, good UV transmittance to allow photolithographic interrogation of surface chemistry in on-chip conditions, high melting temperature (>90 °C) to allow polymerase chain reactions, resistant to chemicals, and finally, considerable stiffness so that high flow rates do not distort the channel’s shape. In this regard, COC exceeds or at least matches20 other thermoplastics such as polycarbonate (PC) or polymethylmethacrylate (PMMA) that are commonly used in microfluidic systems.3-9 UV-photoinitiated grafting is a well-documented method for modifying the surface of some thermoplastics10-14 including COC.1 Despite some patents21,22 and a few companies already commercializing COC microfluidic chips in customized designs, there are few reports with this material. Mair et al. and Craighead and co-workers have reported the fabrication of COC chips and their implementation for isoelectric focusing in electrophoresis,23-28 but their surface modification analysis is lacking thorough study. (17) Laurell, T.; Nilsson, J.; Marko-Vargo, G. The Quest for High-Speed and Low-Volume Bio-analysis. Anal. Chem. 2005, 77, 264A-272A. (18) Su, J.; Bringer, M. R.; Ismagilov, R. F.; Mrksich, M. Combining Microfluidic Networks and Peptide Arrays for Multi-Enzyme Assays. J. Am. Chem. Soc. 2005, 127, 7280-7281. (19) Khalid, I. M.; Pu, Q.; Alvarez, J. C. Thermodynamic and Kinetic Enhancement of Electrochemical Sensitivity by Chemical Coupling in Microfluidic Systems. Angew. Chem., Int. Ed. 2006, 45, 5829-5832. (20) Lamonte, R. R.; McNally, D. Uses and Processing of Cyclic Olefin Copolymers (COC). Plast. Eng. 2000, 56, 51-55. (21) Carvalho, B. L. Elastomeric Tools for the Fabrication of Elastomeric Devices and Uses Thereof. U.S. Patent Application 20040241049, 2004. (22) Jakobsen, M. H.; Kongsbak, L. Closed Substrate Platforms Suitable for Analysis of Biomolecules. U. S. Patent Application 200330152927, 2003. (23) Mair, D. A.; Geiger, E.; Pisano, A. P.; Frechet, J. M. J.; Svec, F. Injection Molded Microfluidic Chips Featuring Integrated Interconnects. Lab Chip 2006, 6, 1346-1354. (24) Gaudioso, J.; Craighead, H. G. Characterizing Electroosmotic Flow in Microfluidic Devices. J. Chromatogr., A 2002, 971, 249-253. (25) Kameoka, J.; Craighead, H. G.; Zhang, H.; Henion, J. A Polymeric Microfluidic Chip for CE/MS Determination of Small Molecules. Anal. Chem. 2001, 73, 1935-1941. (26) Li, C.; Yang, Y.; Craighead, H. G.; Lee, H. K. Isoelectric Focusing in Cyclic Olefin Copolymer Microfludic Channels Coated by Polyacrylamide Using a UV Photografting Method. Electrophoresis 2005, 26, 1800-1806.

Pu et al.

Lately, the interest for photopatterning of surfaces has been renewed with particular emphasis in micro- and nanopatterns,29 including materials such as PMMA,30 glass,31,32 and polydimethylsiloxane (PDMS).33,34 In some cases, the patterns were made inside microchannels using photomasks and in others only the open surface was modified. However, the underlying idea was to combine biomolecule patterning approaches with microfluidic platforms. This notion has been around for some time and is somehow descendent from the extensive work done with hydrogels to create biomolecule micropatches15,35-42 or functional structures inside microfluidic channels.40,43 The present results are, however, more directly related to the work of Ranby and co-workers and Yang et al. who developed efficient photografting techniques for plastics and who elucidated the mechanism of the process.10-14 In addition to conventional surface characterization using infrared (IR), atomic force microscopy (AFM), fluorescence microscopy, and profilometry, we also show electrokinetic measurements evaluating the surface charge of the films. Experimental Section Chemicals. Cyclic olefin copolymer resin (Topas 8007 × 10, glass-transition temperature 80 °C) was purchased from Ticona (Florence, KY). Benzophenone, methacrylamide, and sodium bicarbonate were available from Acros; sodium chloride, phosphoric acid (85%), methacrylic acid, N-[3-(dimethylamino)propyl]methacrylamide, acrylaminde, N-(3-dimethylaminopropyl)-N′-ethylcarbodi(27) Kai, J.; Sohn, Y. S.; Ahn, C. H. Protein Microarray on Cyclic Olefin Copolymer (COC) for Disposable Protein Lab-On-A-Chip. In 7th International Conference on Miniaturized Chemical and Biochemical Analysis Systems, Squaw Valley, CA, 2003. (28) Mela, P.; van den Berg, A.; Fintschenko, Y.; Cummings, E. B.; Simmons, B. A.; Kirby, B. J. The Zeta Potential of Cyclo-Olefin Polymer Microchannels and Its Effects on Insulative (Electrodeless) Dielectrophoresis Particle Trapping Devices. Electrophoresis 2005, 26, 1792-1799. (29) Brack, H. P.; Padeste, C.; Slaski, M.; Alkan, S.; Solak, H. H. Preparation of Micro- and Nanopatterns of Polymer Chains Grafted onto Flexible Polymer Substrates. J. Am. Chem. Soc. 2004, 126, 1004-1005. (30) MaCarley, R. L.; Vaidya, B.; Wei, S.; Smith, A. F.; Patel, A. B.; Feng, J.; Murphy, M. C.; Soper, S. A. Resist-Free Patterning of Surface Architectures in Polymer-Based Microanalytical Devices. J. Am. Chem. Soc. 2005, 127, 842843. (31) Ross, E. E.; Mansfield, E.; Huang, Y.; Aspinwall, C. A. In Situ Fabrication of Three-Dimensional Chemical Patterns in Fused Silica Separation Capillaries with Polymerized Phospholipids. J. Am. Chem. Soc. 2005, 127, 16756-16757. (32) Balakirev, M. Y.; Porte, S.; Vernaz-Gris, M.; Berger, M.; Arie, J.-P.; Fouque, B.; Chatelain, F. Photochemical Patterning of Biological Molecules Inside a Glass Capillary. Anal. Chem. 2005, 77, 5474-5479. (33) Wang, Y.; Lai, H. H.; Bacjman, M.; Sims, C. E.; Li, G. P.; Allbritton, N. L. Covalent Micropatterning of Poly(dimethylsiloxane) by Photografting through a Mask. Anal. Chem. 2005, 77, 7539-7546. (34) Chen, H. Y.; Lahann, J. Fabrication of Discontinuous Surface Patterns within Microfludic Channels Using Photodefinable Vapor-Based Polymer Coatings. Anal. Chem. 2005, 77, 6909-6914. (35) Heo, J.; Crooks, R. M. Microfluidic Biosensor Based on an Array of Hydrogel-Entrapped Enzymes. Anal. Chem. 2005, 77, 6843-6851. (36) Delamarche, E.; Bernard, A.; Schmid, H.; Bietsch, A.; Michel, B.; Biebuyck, H. Microfludic Networks for Chemical Patterning of Substrates: Design and Applications to Bioassays. J. Am. Chem. Soc. 1998, 120, 500-508. (37) Lenigk, R.; Liu, R. H.; Athavale, M.; Chen, Z.; Ganser, D.; Yang, J.; Rauch, C. B.; Liu, Y.; Chan, B.; Yu, H.; Ray, M.; Marrero, R.; Grodzinski, P. Plastic Biochannel Hybridization Devices: a New Concept for Microfluidic DNA Arrays. Anal. Biochem. 2002, 311, 40-49. (38) Liu, Y.; Rauch, C. B. DNA Probe Attachment on Plastic Surfaces and Microfluidic Hybridization Array Channel Devices with Sample Oscillation. Anal. Biochem. 2003, 317, 76-84. (39) Rehman, F. N.; Audeh, M.; Abrams, E. S.; Hammond, P. W.; Kenney, M.; Boles, T. C. Immobilization of Acrylamide-Modified Oligonucleotides by Co-Polymerization. Nucleic Acids Res. 1999, 27, 649-655. (40) Zangmeister, R. A.; Tarlov, M. J. UV Graft Polymerization of Polymerization of Polyacrylamide Hydrogel Plugs in Microfluidic Channels. Langmuir 2003, 19, 6901-6904. (41) Olsen, K. G.; Ross, D. J.; Tarlov, M. J. Immobilization of DNA Hydrogel Plugs in Microfluidic Channels. Anal. Chem. 2002, 74, 1436-1441. (42) Miller, G. A.; Belosludtsev, Y. Y.; Murphy, T. H.; Garner, H. R. Transparent Electronically Controlled DNA Chips. Biomed. MicrodeVices 2000, 2, 215-220. (43) Beebe, D. J.; Moore, J. S.; Bauer, J. M.; Yu, Q.; Liu, R. H.; Devadoss, C.; Jo, B. H. Functional Hydrogel Structures for Autonomous Flow Control Inside Microfluidic Channels. Nature 2000, 404, 588-590.

On-Chip Micropatterning of Microfluidic Channels imide hydrochloride (EDC), dimethylformamide (DMF), dimethyl sulfoxide (DMSO), avidin, bovine serium albumin (BSA), and tetraethyl orthosilicate were obtained from Sigma-Aldrich (Milwaukee, WI); sodium dodecylsulfate (SDS), N-hydroxysuccinimide (NHS), acrylic acid (AA), methaacrylic acid (MAA), and ethylenediamine were purchased from Fluka (Milwaukee, WI). Fluorescein-5-isothiocyanate (FITC), dapoxyl (2-aminoethyl)-sulfonamide, and biotin-4 fluorescein were obtained from Invitrogen (Carlsbad, CA), and phosphate-buffered saline (PBS) 10× solution was purchased from Fisher. The single-strand DNA 5′-H2N-(CH2)6GTT GAG GGG ACT TTC CCA GG-3′ (Am-Seq1), 5′-6fluorescein-CCT GGG AAA GTC CCC TCA AC-3′(FAM-Seq2), 5′-6-fluorescein-TAA CAC CCG TAT GAT AGT CT-3′ (5FAMSeq3) were synthesized and purified with high-performance liquid chromatrography (HPLC) by Integrated DNA Technologies, Inc (Coralville, IA). Equipment. A Super Spot MK III UV lamp equipped with a flexible lightguide (Lesco, Torrance, CA) was used as the light source for all photochemical modifications. The irradiation power was measured with a calibrated SuperSpot intensity meter (Lesco, Torrance, CA). Photomask used for the modification was made on quartz plates using photolithographic methods reported elsewhere.44, 45 Deionized (DI) water (18.2 MΩ) from a Milli Q system (Millipore, Billerica, MA) was used in all experiments. Fabrication of COC Plates and Microfluidic Chips. Round COC plates (63.3 mm in diameter and 1.05-mm thick) were made by injection molding of COC resin using a Toyo TI-556 injection molding machine (Rockaway, NJ). The plates were then cut into desired sizes for further use. Microfluidic chips were fabricated using a similar procedure reported in the literature.6,25 Half-cylindrical channel chips were fabricated by wire imprinting. A piece of COC plate and two glass microslides, Corning 2947 (Corning, NY), were cleaned with compressed air to remove dust particles. A wire stainless steel, 0.005 in. (127 µm) in diameter (Small Parts, Miami Lakes, FL, Catalog no. GWX-0050), was placed on the COC plate and was sandwiched between two glass microslides using four small binder clips. The assembly was put in an oven at 125 °C for 15 min. Then, the assembly was taken out from the oven and was allowed to cool down. The plastic plate and microslides usually detach spontaneously during the cooling process so that the wire can be pulled away to get the empty channel. The resulting channel plate and another COC plate with two holes previously drilled were sonicated in ethanol for 10 min. After drying, the two plates were assembled together with holes aligned along the channel and were sandwiched between two glass microslides clamped by binder clips. The assembly was put in the oven at 82 °C for 15 min to seal the chip. Trapezoidal channels were obtained by imprinting with silicon molds. Silicon molds were made by photolithographic method as described in the supporting material. Before imprinting, the surface of silicon mold was treated by spraying with 10% solution of tetraethyl orthosilicate in ethanol, was rinsed with ethanol, and was dried to facilitate mold release. The subsequent process of imprinting and bonding was the same as described for the half-cylindrical channels. Photochemical Modification. In all experiments, modification solutions were prepared by dissolving 10% of acryl monomers (w/v for solids and v/v for liquids) in DI water and by adding 0.1% photoinitiator benzophenone. Benzophenone was added in the form of acetone solution of 100 mg/mL. After vigorous mixing, the solutions were degassed for 20 min under vacuum (about 12-14 mmHg) and were filtered through 0.22-µm PTFE membrane. For all monomers used in the present study, there is precipitate of benzophenone formed during the mixing or degassing process, and thus benzophenone concentration was referred to as “saturated”. For COC plates, the modification solution was applied on the surface and was covered with a plain quartz plate or a gold photomask (44) Xia, Y. N.; Whitesides, G. M., Soft Lithography. Angew. Chem., Int. Ed. 1998, 37, 550-575. (45) Zhan, W.; Alvarez, J.; Crooks, R. M. Electrochemical Sensing in Microfluidic Systems Using Electrogenerated Chemiluminescence as a Photonic Reporter of Redox Reactions. J. Am. Chem. Soc. 2002, 124, 13265-13270.

Langmuir, Vol. 23, No. 3, 2007 1579 on quartz (patterned samples). To get clear patterns, the gold layer was placed upside down to avoid deflection of the UV light. Two wires of 0.127-mm diameter of steel were used as spacers to confine the distance between the mask and the COC surface. To avoid modification on the back of the plate, the backside was protected with Scotch tape. After that, the assembly was placed under the lightguide of the UV light at a selected distance (UV power varies depending on the distance) for a determined period of time in a black box (Caution: Strong UV light is harmful to unprotected eyes and skin, avoid direct exposure!). The irradiation power was determined with the intensity meter. After removing the Scotch tape, the plate was rinsed with frequently replenished DI water in an ultrasonic bath for 3 × 20 min. Afterward, the plates were ready for further derivatization or film characterization. For microchannel photografting, the modification solution was filled into the channel and was illuminated with UV light for a certain time as described above. To get a clear pattern, the inlets and outlets of the channel were sealed with cushions of PDMS to prevent flow inside the channel. The channel was flushed with DI water for 5 min with vacuum suction to remove free polymer and unreacted molecules. Surface Characterization. Attenuated total reflection Fourier transform infrared spectrometry (ATR-FTIR) spectra were recorded using a Nicolet Nexus 870-FTIR spectrometer at 4 cm-1 resolution with 1024 scans. Avatar Multi-Bounce HATR accessory was used for this purpose and the crystal was ZnSe with incidence angle of 45 °. Film thickness (>1 µm) was measured using Alpha-Step 500 profilometer (Tencor, Milpitas, CA) and film thickness below 1 µm was measured in tapping mode on a Nanoscope IIIA multimode atomic force microscopy (Digital Instruments, Veeco Metrology group, Santa Barbara, CA) using Ultrasharp silicon probes (MicroMasch, Wilsonville, OR). Fluorescent imaging was performed using an inverted fluorescence microscope (XDY-1, Microscopes Inc., St. Louis, MI) equipped with a 100 W mercury arc lamp. Filter sets suitable for fluorescein and Dapoxyl dye were obtained from Omega Optical (Brattleboro, VT). Pictures were taken using a Nikon Coolpix 5400 digital camera. Determination of Carboxylic Groups in Modified-COC Sheets.46 We used the method reported by Akamatsu et al.46 wherein selective adsorption and desortion of Ag+ from the sample is monitored by inductively coupled plasma atomic emission spectroscopy (ICP-AES). The complex 1:1 for COO--Ag+ allows the quantification of COOH groups on surface-bound films.46 Each modified sheet (2 × 2 cm) was immersed in 15 mL of AgNO3 solution (5.0 mM) and was magnetically stirred at room temperature overnight in the dark. The initial pH value of the AgNO3 solution was adjusted with NH4OH to 8.5 and after the adsorption process the sheets were rinsed with copious amounts of water three times. The pH of the rinsing water was adjusted with NH4OH to ∼8.5. Desorption of Ag+ from each COC sheet was done with 10 mL 1% HNO3 solution for 3 h with magnetic stirring at room temperature in the dark. The desorbed Ag+ was quantified by ICP-AES. Standard solutions of AgNO3 were prepared in 1% HNO3 and a blank solution was made only with 1% HNO3. The calibration curve is shown in Figure S2 (Supporting Information). Electroosmotic Flow (EOF) Measurements. The current monitoring method proposed by Huang47 et al. was used to measure EOF in the channels. Test solutions were 1 mM phosphate buffers of different pH’s (3.0-11.0) containing 8.0 mM (lower conductivity buffers) and 10 mM NaCl (higher conductivity buffer). In the measurement, platinum electrodes were placed into the reservoirs at both ends of the channel. A PS-350 high-voltage power supply (Stanford Research Systems, Sunnyvale, CA) was employed to provide high voltage over the electrodes. A 10 kΩ resistor was connected in series with the channel and the voltage drop cross was (46) Akamatsu, K.; Ikeda, S.; Nawafune, H. Site-Selective Direct Silver Metallization on Surface-Modified Polyimide Layers. Langmuir 2003, 19, 1036610371. (47) Huang, X.; Gordon, M. J.; Zare, R. N. Current-Monitoring Method for Measuring the Electroosmotic Flow Rate in Capillary Zone Electrophoresis. Anal. Chem. 1988, 60, 1837-1838.

1580 Langmuir, Vol. 23, No. 3, 2007 recorded by a Linear 1200 chart recorder (Linear Instruments Corp., Reno, NV). The channel was first filled with the lower conductivity buffer of certain pH. Just prior to the measurement, one reservoir was emptied and filled with the higher conductivity buffer of the same pH immediately. The amount was kept exactly the same as that in the other reservoir. We turned on the high voltage and marked the recorder simultaneously. The current increased when the higher conductivity buffer replaced the lower one and attained a higher stable level after the whole channel was replaced. The time (t) required for this was used to calculate the EOF mobility (µeof ) L2/(Vt), where L is the channel length, V is applied voltage). At least five measurements were made for each point. Subsequent Chemical Functionalization. NHS-Functionalized Carboxylic Groups. The poly(methacrylic acid) (or poly(acrylic acid)) modified COC plate was covered with 250 µL of DMF solution of 0.5 M NHS and 0.5 M EDC and was allowed to react at room temperature for 2 h. The resulting surface was rinsed with DI water and was dried with compressed air immediately after the rinsing process. The carboxylic group inside the channel was converted to succinimidyl ester by filling the channel with the same solution used for the plates. Then, the channel was flushed with DI water after 2 h and was dried with compressed air flow. Avidin Immobilization on the Modified Surface. Avidin was attached to the COC chip through the reaction of amine groups on avidin chain and the NHS activated carboxylic group. The solution of 0.5 mg avidin in 200 µL of 1× PBS (pH 7.4) was spread on the NHS functionalized plates and was allowed to react at room temperature overnight. The resulting plate was rinsed with 1× PBS and then was covered with 0.1 M ethanolamine in 1× PBS for 3 h. After that, it was rinsed with 1× PBS and was dried. The immobilized avidin was proved by incubation of the plate with 1.0 mM biotin-4-fluorescein solution in 1×PBS for 4 h and the fluorescent pattern was checked under the microscope after rinsing with 1× PBS thoroughly. Another NHS functionalized plate that was treated in the exact same way, except no avidin was applied, was used as a control. DNA Immobilization on the Modified Surface. A 20-mer DNA sequence with primary amine group at its 5′ terminal was attached to the modified surface. Thirty microliters of 1× PBS containing 1 mM Am-Seq1 was sandwiched between two COC plates with NHS activated carboxylic group pattern, face to face, and was allowed to react at room temperature in a sealed vessel overnight. Water was added inside the vessel to prevent DNA solution from drying because of evaporation. Then, the plates were rinsed with 1× PBS five times and were covered with 0.1 M ethanolamine in 1× PBS for 3 h at room temperature. The resulting surface was rinsed with copious DI water and 1× PBS and was dried with compressed air. The hybridization reaction was performed by sandwiching 30 µL of 0.2 mM complementary fluorescein labeled DNA (FAM-Seq2) or uncomplimentary one (FAM-Seq3) in 1× PBS (pH 7.4) and was kept in a sealed vessel overnight at room temperature. The resulting plates were rinsed with 1× PBS containing 0.1% SDS five times. The pattern was checked under the fluorescence microscope after they dried. DNA immobilization and hybridization inside the channels were performed by filling the channel with 5 µL of DNA solution following the same procedure as for COC plates. The chip reservoir was sealed with Scotch tape to prevent solution from drying inside the channel.

Results and Discussion Surface Modification and Spectroscopic Characterization. The surface modification of COC plates used here is based on a photografting method initially investigated by Ranby and coworkers and Yang et al. on some plastics10-14 and was later adapted by Rohr et al. to other thermoplastics with good grafting efficiency (i.e., >60%).1 The reaction mechanism occurs via hydrogen abstraction from the carbon chains on the plastic surface by a photochemically reduced initiator.10-14 Once carbon radical intermediates are generated on the plastic surface, vinyl or acryl

Pu et al. Scheme 1

monomers can react with them to form branched polymeric films anchored to the surface. The branching is a consequence of the presence of abstractable hydrogens on the grafted polymer and the formation of homopolymers usually not grafted can also happen simultaneously.10-14 The kinetics of the reaction, the morphology, and the thickness of the grafted film are highly dependent on factors such as solvent, UV irradiation time, monomer and photoinitiator used, and so forth. However, because the reaction is based on hydrogen abstraction, solid surfaces containing abstractable hydrogens like COC (Scheme 1) can be used for surface immobilization of molecular probes or control of surface properties. These two approaches are promising possibilities for the construction of microfluidic sensors. For example, immobilized probes in microfluidic channels can release and capture analytes in sensing schemes on the basis of bioaffinity.15,35,48 The modulation of surface properties such as the zeta potential can be used to control flow direction in electroosmotic pumps49,50 and also to improve the efficiency of some electrophoretic separations.51 Furthermore, because the photografting reaction is surface driven, there is a C-C covalent bond between the modified film and the surface thus improving the stability of the surface composite. In contrast, the preparation of hydrogel microstructures inside glass microchannels usually requires pretreatment of the glass surface with silane (Si-O bonds are not that stable at higher pH52) reagents to improve stability.15,35,48 Our goal in this work was to investigate the chemical flexibility of COC for the grafting of monomers that could afford immobilized functional groups (-NH2, -COOH) useful for further derivatization of analytical probes. Figure 1 shows the attenuated total reflectance FTIR spectrum of a series of films photografted on COC plates using different acrylic monomers and benzophenone as photoinitiator. For comparison, spectrum A in Figure 1 was obtained with an unmodified COC sheet. The observed IR bands in the range 2944-2866 cm-1 for CH stretches and the bending modes around 1458 cm-1 are in good agreement with previous reports.13,53 The other spectra in Figure 1 were taken after 12 min of UV irradiation (0.15 W/cm2) in the presence of acrylic monomers that had the following pendent groups: a secondary amine, N-[3-(dimethylaminopropyl]methacrylamide, B; an amide, methacrylamide, C; and a carboxylic acid group, (48) Seong, G. H.; Crooks, R. M. Fabrication of Microchambers within Microfluidic Systems using Photopolymerized Hydrogels: Application to DNA Hybridization. Anal. Chem. 2002, 74, 3372-3377. (49) McKnight, T. E.; Culbertson, C. T.; Jacobson, S. C.; Ramsey, J. M. Electroosmotically Induced Hydraulic Pumping with Integrated Electrodes on Microfluidic Devices. Anal. Chem. 2001, 73, 4045-4049. (50) Razunguzwa, T. T.; Timperman, A. T. Fabrication and Characterization of a Fritless Microfabricated Electrosomotic Pump with Reduced pH Dependence. Anal. Chem. 2004, 76, 1336-1341. (51) Palmer, C. P. Demonstrating Chemical and Analytical Concepts in the Undergraduate Laboratory Using Capillary Electrophoresis and Micellar Electrokinetic Chromatography. J. Chem. Educ. 1999, 76, 1542-1543. (52) McMurry, J. Organic Chemistry, 4th ed.; Brooks/Cole Publisching Company: Pacific Grove, CA, 1996; p 1243. (53) Forsyth, J.; Perena, J. M.; Benavente, R.; Perez, E.; Tritto, I.; Boggioni, L.; Britzinger, H. H. Influence of the Polymer Microstructure on the Thermal Properties of Cycloolefin Copolymers with High Norborene Contents. Macromol. Chem. Phys. 2001, 202, 614-620.

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Figure 2. Dependence of surface concentration of carboxylic groups on UV exposure time for the photografting of COC. Concentration of monomer is 10% (v/v), benzophenone is saturated, and UV power is 0.15 W/cm2. Figure 1. ATR-FTIR spectra for (A) COC and COC modified with (B) N-[3-(dimethylamino)propyl]methacrylamide, (C) methacrylamide, and (D) methacrylic acid. Photochemical reaction solutions contain 10% (w/v for solids and v/v for liquids) of monomers and saturated benzophenone as photoinitiator. UV irradiation time 12 min, UV power is 0.15 W/cm-2.

Scheme 2

Table 1. Film Thickness by AFM and Profilometry of Grafted Monomers on COC at Different UV Exposure Times time (min)

thickness for MAA (µm)

thickness for AA (µm)

5 10 15