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Self-Assembled, Mesoporous Polymeric Networks for Patterned Protein Arrays Dolly Batra,†,‡ Stefan Vogt,§ Phillip D. Laible,⊥ and Millicent A. Firestone*,†,‡ Materials Science Division, Center for Nanoscale Materials, Experimental Facilities, and Biosciences Division, Argonne National Laboratory, 9700 South Cass Avenue, Argonne, Illinois 60439 Received July 19, 2005. In Final Form: August 31, 2005 A facile, self-assembly approach to the fabrication of a robust, mesoporous, biocompatible polymeric network for the spatial organization of proteins is described. Surface-deposited poly(styrene) (PS) beads that assemble into a two-dimensional (2-D) hexagonal array are used to template cross-linked poly(vinyl alcohol) (PVA), yielding an inverse opal structure. The porous, water insoluble network is used to entrain a model, soluble protein, green fluorescent protein (GFP). The polymeric network is characterized by atomic force microscopy (AFM) and optical microscopy, and the spatial localization of the incorporated GFP is determined by fluorescence microscopy. The results demonstrate that this system may constitute a versatile platform for the lateral organization of biomolecules.
Introduction The synthesis of biocompatible, structured “soft” materials that organize biomolecules into periodic, functional arrays is an important issue in tissue engineering and fundamental studies of cell biology and is a necessary first step in the development of protein-based materials.1 The material requirements for a framework suitable for the immobilization of proteins are numerous and complex. Most importantly, the protein must be localized with high efficiency and without denaturation. Recently, the synthesis of ordered mesoporous solids (e.g., mesoporous silicas,2 anodized alumina,3,4 and inorganic or polymeric inverse opal structures5-8) and their application in bioseparations, biocatalysis, biosensors, and bioscaffolds have been reported. Many of these materials, however, suffer from various drawbacks that limit their utility. Transparent mesostructured silica films prepared by solvent evaporation using spin-coating or dip-coating, for example, often have randomly oriented mesochannels/pores that can only be oriented parallel to the substrate.9 Anodized alumina often suffers from poor optical transparency and cannot readily be made with pore sizes smaller than ∼20 nm. Porous polymeric networks adopting an inverse opal * To whom correspondence should be addressed. Millicent A. Firestone, Materials Science Division, Argonne National Laboratory, 9700 South Cass Avenue, Argonne, IL 60439. Phone: 630252-8298. Fax: 630-252-9151. E-mail:
[email protected]. † Materials Science Division. ‡ Center for Nanoscale Materials. § Experimental Facilities. ⊥ Biosciences Division. (1) Khademhosseini, A.; Jon, S.; Suh, K. Y.; Tran, T. N. T.; Eng, G.; Yeh, J.; Seong, J.; Langer, R. Adv. Mater. 2003, 15, 1995-2000. (2) Fan, J.; Lei, J.; Wang, L. M.; Yu, C. Z.; Tu, B.; Zhao, D. Y. Chem. Commun. 2003, 2140-2141. (3) Lee, S. B.; Mitchell, D. T.; Trofin, L.; Nevanen, T. K.; Soderlund, H.; Martin, C. R. Science 2002, 296, 2198-2200. (4) Wang, Z.; Haasch, R. T.; Lee, U. G. Langmuir 2005, 21, 11531157. (5) Lee, Y. J.; Braun, P. V. Adv. Mater. 2003, 15, 563-566. (6) Lee, Y. J.; Pruzinsky, S. A.; Braun, P. V. Langmuir 2004, 20, 3096-3106. (7) Kotov, N. A.; Liu, Y. F.; Wang, S. P.; Cumming, C.; Eghtedari, M.; Vargas, G.; Motamedi, M.; Nichols, J.; Cortiella, J. Langmuir 2004, 20, 7887-7892. (8) Zhang, Y.; Wang, S. P.; Eghtedari, M.; Motamedi, M.; Kotov, N. A. Adv. Funct. Mater. 2005, 15, 725-731. (9) Fukuoka, A.; Miyata, H.; Kuroda, K. Chem. Commun. 2003, 284285.
structure produced by colloidal crystallization techniques offer several clear advantages over other mesoporous matrixes, including biocompatibility and the ability to both introduce a wide variety of functional groups in the interior of the pore and control pore structure and connectivity (allowing for optimal diffusion through the matrix). In an effort to identify suitable polymeric mesoporous (defined here as pore dimensions between 100 and 1000 nm) frameworks, inverted colloidal crystalline frameworks composed of poly(N-isopropylacrylamide) (PNIPAM),10 poly(acrylates), poly(acrylic acid),11 and poly(2-hydroxyethyl methacrylate/styrene) copolymers12,13 were examined. Frameworks comprising elastomers such as poly(dimethylsiloxane) (PDMS)14 were also prepared. These materials (i.e., hydrogels) were chosen primarily because, by altering the environmental conditions (e.g., temperature, pH, solvent, and ligand binding), changes in their lattice constant can be induced, thereby giving rise to alterations in their optical properties (e.g., the color of the Bragg-diffracted light). Few systems have been reported, however, that specifically address the need for robust, mesoporous two-dimensional (2-D) polymeric frameworks that can be used to selectively partition and spatially organize proteins. Moreover, characterization of their suitability for biomolecule incorporation has been lacking,11 and the architectures are not suitable for the orientational ordering of the encapsulated proteins.15 In this communication, we present a facile, self-assembly approach for the fabrication of a robust, mesoporous, biocompatible polymeric network for the spatial organization of soluble proteins. This framework exhibits good optical transparency in the visible region of the spectrum and possesses favorable chemical and mechanical stability in aqueous buffer solutions. In addition, we demonstrate that the composition of the framework can be adjusted to resist protein adsorption, and the open wells allow for the spatial localization of a model soluble protein. This (10) Takeoka, Y.; Watanabe, M. Langmuir 2002, 18, 5977-5980. (11) Valsesia, A.; Colpo, P.; Silvan, M. M.; Meziani, T.; Ceccone, G.; Rossi, F. Nano Lett. 2004, 4, 1047-1050. (12) Jiang, P.; McFarland, M. J. J. Am. Chem. Soc. 2004, 126, 1377813786. (13) Chen, Y.; Ford, W. T.; Materer, N. F.; Teeters, D. Chem. Mater. 2001, 13, 2697-2704. (14) Yin, Y.; Li, Z. Y.; Xia, Y. Langmuir 2003, 19, 622-631. (15) Yap, F. L.; Zhang, Y. Langmuir 2005, 21, 5233-5236.
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polymeric framework offers significant potential as a versatile platform for the entrapment and organization of biomacromolecules. Experimental Section Protein Expression and Purification. Green fluorescent protein (GFP) from Aequoria victoria was overexpressed in the cytoplasm of an engineered strain of Rhodabacter sphaeroides. For optimal GFP production, a red-shifted variant that had been codon optimized for eukaryotic expression and modified for enhanced absorption cross-section and fluorescent yield (obtained from Quantum Biotechnologies Inc., Montreal, Quebec, Canada) was amplified and inserted in place of structural genes of the light-harvesting 1I antennae complex in the puf operon.16 Once incorporated into the vector, a seven-repeat histidine tag was fused in frame to the C-terminius. Culture and purification proceeded as previously described for membrane proteins produced in this manner with important modifications.17 In brief, cells were grown in chemoheterotrophic conditions (semi-aerobic, dark, 35 °C, 2 L of media in a 2.8-L Fernbach flask, 125 rpm) on YCC medium containing tetracycline (1 µg/mL).18 Using these conditions, the synthesis of proteins under the control of the puf operon is autoinduced as the cell density increases and the oxygen tension decreases. Cultures were harvested in the late log phase (Klett-Summerson calorimeter reading of 300; OD600 ) 3) and lysed in the presence of 100 mM NaCl and protease inhibitors (EDTA-free) using sonication and passage through a microfluidizer with the pressure set at 18 000 psi. After the cell debris was pelleted, the soluble fraction was separated from the membrane fraction by ultracentrifugation (250000g; 1.5 h). The supernatant from the final centrifugation step was passed through as a 0.45-µm filter prior to binding to Ni-NTA resin (Qiagen, Inc., Valencia, CA) for the purification of GFP. Standard immobilized metal affinity chromatography methods were employed with all buffers containing 100 mM NaCl for a maximal retention of GFP structural integrity and monodispersity. GFP was eluted with 100 mM imidazole. The eluant was removed during successive buffer washes as the protein was being concentrated in a centrifugal concentrator (over a 10-kDa MW cutoff filter; Vivascience, Hanover, Germany) or by dialysis (SnakeSkin tubing; 10-kDa MW cutoff; Pierce Scientific, Rockford, IL). Protein in 10 mM Tris (pH 7.8) and 100mM NaCl was either used immediately, stored at 4 °C, or flash frozen in liquid nitrogen for subsequent experiments. Synthesis of the Photoacid Generator (PAG). The watersoluble PAG, 2,4-(dihydroxyphenyl)dimethyl sulfonium triflate was synthesized using a two-step procedure developed by Willson and co-workers.19,20 Briefly, HCl was bubbled through a methanolic solution of dimethyl sulfoxide (DMSO) and resorcinol, producing a dimethylsulfonium chloride salt, which underwent an anion exchange with sodium triflate in water to afford the triflate salt in 33% yield. The final product was extracted into ethyl acetate, then reduced under vacuum. The product identity was confirmed by 1H NMR [3.4 (s, 6H, 2CH3) and aromatic frequencies at 7.6, 6.7, and 6.6 ppm (d, 3H, ArH)]. Synthesis of Porous Polymeric Networks and Protein Arrays. The polymer sphere template was formed from a 0.2% (w/v) aqueous suspension of carboxyl-terminated polystyrene spheres (500 nm, Interfacial Dynamics Corp., Tualatin, OR). A 0.1-0.3-mL sample of a dilute suspension (500 µL of a 2% (w/v) poly(styrene) (PS) sphere suspension in 5 mL of deionized water) was solvent-cast onto acid-cleaned glass slides held at 10 °C on a cold plate. The plate was slowly warmed to room temperature over 4 h, and the suspensions were allowed to dry at room (16) Scott, H. N.; Laible, P. D.; Hanson, D. K. Plasmid 2003, 50, 74-79. (17) Laible, P. D.; Scott, H. N.; Henry, L.; Hanson, D. K. J. Struct. Funct. Genomics 2003, 5, 167-172. (18) Taguchi, A. K. W.; Stocker, J. W.; Alden, R. G.; Causgrove, T. P.; Peloquin, J. M.; Boxer, S. M.; Woodbury, N. W. Biochemistry 1992, 31, 10345-10355. (19) Lin, Q. H.; Steinhausler, T.; Simpson, L.; Wilder, M.; Medeiros, D. R.; Willson, C. G.; Havard, J.; Frechet, J. M. J. Chem. Mater. 1997, 9, 1725-1730. (20) Havard, J. M.; Shim, S. Y.; Frechet, J. M. J.; Lin, Q. H.; Medeiros, D. R.; Willson, C. G.; Byers, J. D. Chem. Mater. 1999, 11, 719-725.
Letters temperature for 18 h. The dried PS spheres were then held at 35 °C for another 18 h to ensure the complete formation of the PS template. A solution of 2% (w/v) poly(vinyl alcohol) (PVA) [Mw ) 13-23 kDa; 87-89% hydrolyzed] containing 0.5% (w/v) 2,4-dihydroxyphenyl)dimethyl sulfonium triflate was cast onto the self-assembled PS template. The solution was allowed to dry for 4 h at 25 °C and then dried at 35 °C for another 18 h. After drying, we etched out the spheres by gently shaking the thin films in a solution of toluene for 18-36 h, replacing the toluene solution frequently. The resulting mesoporous network was photo-cross-linked via UV irradiation (254 nm) for 4 h. The PVA mesoporous network was incubated in a 1-2 µM solution of GFP for 30 min (25 mM potassium phosphate buffer; pH ) 6.5; [NaCl] ) 100 mM). The solution was washed with the same buffer 3 times, which was followed by 3 water washes. Physical Methods. Contact angle measurements were performed using a First Ten Angstroms FTA125 Dynamic Contact Angle Analyzer. The contact angle of deionized water (Milli-Q; 18 MΩ cm-1) was measured on a series of 2-mL sessile (freestanding) drops applied to the surface with a blunt cut syringe. Data were averaged over five spots for a given sample. Film thickness measurements were performed using an Ambios Technology XP-1 profilometer with a scan speed of 0.05 mm/sec and a stylus force of 49 µN. The porous cross-linked PVA networks were scored with a razor blade, and the thickness was measured over a length of 3 mm and averaged over four spots. 1H NMR spectra were obtained in (CD3)2CO using a 300 MHz Varian Unity INOVA spectrometer. UV spectra were obtained using a Varian Cary-5G UV-Vis-NIR spectrophotometer at a spectral resolution of 1 nm. Fluorescence spectra were collected on a Photon Technology International QM-3/2005 pulsed spectrofluorometer. Atomic Force Microscopy (AFM). Images were obtained using a Digital Instruments Dimension 3000 with a type IIA controller. The images were collected in tapping mode, which was operated with a 110-µm scanner with 10-nm microfabricated Si tips (DI TESP) with resonant frequencies between 315 and 340 kHz. A scan rate of 5-10 µm/s was used. Visible Light/Fluorescence Microscopy. Images were obtained using a Leica DMXRE light microscope (Leica Microsystems, Wetzlar, Germany) equipped with a high-resolution motorized XY stage (Ludl Bioprecision, Hawthorne, NY). Light micrographs were acquired using a 100× fluortar oil immersion objective with a 1.3 numerical aperture and a Leica DC 350 F CCD camera. GFP fluorescence images were taken with an FITC, Chroma filterset #41001 (Chroma, Rockingham, VT).
Results and Discussion Here, we describe the preparation of a periodic, mesostructured hydrogel comprising a porous, hexagonally ordered network of cross-linked PVA. This network is prepared by a facile, three-step process. First, a hexagonal array of polymer microspheres is formed by solvent-casting a dilute (0.2% w/v) aqueous suspension of monodisperse (e.g., 500 nm) PS beads onto a clean glass slide. The controlled (i.e., slow drying under ambient conditions) evaporation of water and the sedimentation of the particles produces a macroscopically (blue) opalescent (arising from the Bragg diffraction of visible light) film. Optical microscopy of the thin films reveals the presence of jagged lines separating irregularly shaped domains that arise during the drying process (data not shown). Tapping-mode AFM, which was used to further characterize the surface topography and the extent of the long-range ordering of the packed PS spheres, indicates (Figure 1A) that the procedure yields a well-ordered crystalline film comprising a close-packed hexagonal array of spheres over a large area (9 × 9 µm). However, the AFM image clearly shows various imperfections in the crystal packing, as evidenced by the presence of either a point (missing PS spheres) or line defect (slip dislocations). This colloidal crystal serves as a template onto which a porous, chemically and mechanically robust, PVA network is formed. PVA was selected because it is a highly
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Figure 2. AFM image (10 × 10 µm) of a porous PVA network produced after the removal of the PS template (toluene, 24 h) and a photo-cross-linking at 254 nm (4 h). Inset: an optical micrograph illustrating optical transparency and the slight yellow coloring of the macroscopic film.
Figure 1. AFM images showing: (A) The domain structure of a hexagonal-packed array (9 × 9 µm) of carboxy-terminated PS spheres (500 nm) produced by being solvent-cast onto a glass substrate. Region “i” highlights a missing PS sphere, and region “ii” highlights a line defect. (B) PVA/PAG-coated PS array produced by solvent-casting an aqueous solution of PVA/PAG onto the PS template (inset: 2 × 2 µm region of uncoated PS spheres). Scheme 1
polar, water-soluble, biocompatible polymer that has found numerous application in the biochemical and biomedical fields.21 To fabricate a robust PVA framework that is not susceptible to dissolution or pronounced swelling upon water immersion, it is necessary to render the PVA waterinsoluble via chemical cross-linking. In this work, a modification of an approach developed for water-processable resist materials was used.19,20 In brief, this approach (21) Hassan, C. M.; Peppas, N. A. Adv. Polym. Sci. 2000, 153, 3764.
employs a water-soluble photoactive catalyst, that is, a photoacid generator (PAG) (2,4-dihydroxyphenyl)dimethyl sulfonium triflate (1). The PAG is synthesized from resorcinol using the procedure of Willson and co-workers (Scheme 1).19 The introduction of the PAG to the PVA can be used to initiate photoinduced cross-linking (acidcatalyzed condensation of the alcohol groups), which gives rise to a cascade of solubility-modifying reactions.19,20 Specifically, here we employ a two-component aqueous mixture of 2% (w/v) PVA (87-89% hydrolyzed, Mw ) 1323 kDa) and 0.5% (w/v) PAG, which is solvent-cast on top of the PS colloidal crystalline template. A comparison of the tapping-mode AFM of a 500-nm PS 2-D crystal (Figure 1B inset) to those recorded after coating/infiltration and drying of the PVA/PAG mixture (Figure 1B) suggests that the polymer solution uniformly coats the PS spheres and does so without significantly disrupting the local packing structure of the template. After the PVA/PAG film is completely dried, the underlying PS template is removed by chemical etching with toluene (18-24 h). The remaining PVA/PAG film is irradiated with 254-nm UV light (4500 µW/cm2 at 1′′ for 4 h), initiating the cross-linking reactions that produce an insoluble porous polymer network. Attempts to remove the PS template after photo-cross-linking were unsuccessful, which is possibly due to the acid-catalyzed condensation of the carboxylate groups on the surface of the PS spheres with the alcohol groups of the PVA.22 The optical transparency of the polymer film remains good throughout the visible region (Figure 2 inset) but the film does exhibit a pale yellow color, which is presumably a result of the formation of a conjugated olefin network via the elimination of the alcohol groups.19,20 In addition, unlike the colloidal crystal template, these films do not appear iridescent to the naked eye. Moreover, the cross-linking causes a decrease in the wettability of the polymer, as (22) Brown, L.; Halling, P. J.; Johnston, G. A.; Suckling, C. J.; Valivety, R. H. J. Chem. Soc., Perkin Trans. 1 1990, 3349-3353.
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Figure 3. AFM images of a 2-D hexagonal array of mesopores before (A-C) and after (D-F) water incubation for 72 h. (A,D) High-magnification images of 3 × 3 µm areas. (B,E) Close-up images of a single pore. (C,F) Cross-sectional analyses of the pore features from B and E, respectively.
determined by water contact angle measurements made on thin films of solid un-cross-linked and cross-linked PVA thin films. These data show that, on average, the polymer surface goes from very hydrophilic (θ ) 12 ( 1°) to nonwetting (θ ) 52 ( 12°). The cross-linked inverse opal structure, as imaged by AFM, shows regions that adopt 2-D hexagonally ordered, hemispherical voids that are consistent with a replication of the PS opal template (Figure 2). However, the thin film suffers from regions of local defects that appear to most likely arise from PS beads that are not completely removed during the etching process. Closer examination of the hole morphology by AFM shows regions (3 × 3 µm) that contain well-ordered
2-D hexagonal arrays of hemispherical voids (Figure 3A). Figure 3B shows a more detailed view (1 × 1 µm) of a single pore and the corresponding line scan (Figure 3C), and shows a well-defined hemispherical structure composed of connected triangular features of polymer. The lateral dimensions (uncorrected for tip broadening) of the pore, measured as the in-plane width defined by the midpoint of the triangular features (peak-to-peak distance) are 489 ( 20 nm (averaged over 5 holes). Measurements from the pore center to the pore edge yield distances of 250 ( 20 nm, a value that corresponds well with the dimensions of the spherical particle template.
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Figure 4. Optical micrographs of a 500-nm porous, cross-linked PVA network. (A) DIC image of the 2-D hexagonal array. (B) Fluorescence image after incubation with GFP (image recorded using an FITC, Chroma #41001 filter set). Images were false colored to enhance regions of observed maximum fluorescence intensity. Images A and B were recorded in the same area. (C) 2-D FT of the fluorescence array observed in image B. (D) Digital overlay of the DIC image (panel A; false colored orange) with the fluorescence image (panel B; false colored green) showing the selective incorporation of GFP in the holes.
The effect of water on polymer swelling was evaluated by a comparison of the topographic AFM images of a given sample before and after it was soaked in water for 72 h. The large-area scan (3 × 3 µm), a close-up view of a single pore (1 × 1 µm), and the corresponding line scan are presented in Figure 3, panels D, E, and F, respectively. The large-area scan shows that the hexagonal ordering of the holes is not significantly altered by water exposure. The persistence of the periodic structure and the fact that the film remains firmly attached to the supporting glass slide suggest that the cross-linking chemistry employed here yields a mechanically robust thin film. Close examination of the pore structure, however, reveals some slight deformation in morphology, most likely the result of modest swelling of the polymer (hydrogel) network. These changes are not, however, significant enough to disrupt or destroy the periodic structure of the network. The influence of swelling can more easily be seen in the close-up image of the pore morphology, which no longer shows the well-resolved triangular features. Instead, a more diffuse image, indicative of an expanded polymer structure (i.e., a slight increase in the “wall” thickness of the pores or wells), is observed. The lateral dimensions of the pore and wall slightly expanded to 561 ( 26 nm, and the pore center-to-pore edge distance decreased to 241 ( 15 nm. These measurements indicate that, upon prolonged exposure to aqueous media, the network undergoes minor changes that involve a slight reduction in the open pore dimensions and a concomitant expansion in the wall thickness. Estimates of the cross-linked PVA film thickness, made by scoring the film and measuring the depth using either a profilometer or AFM, are typically 1.8-2.5 µm; thus, thicknesses corresponding to ∼3-5 times the dimension of the spherical template are routinely achieved. Because the hemispherical voids in the polymer film are oriented perpendicular to the plane of the glass substrate, they should be useful as “wells” into which proteins can be selectively partitioned and spatially localized. GFP isolated from jellyfish (Aequoria victoria) was selected to test this concept because it is a small cylindrical protein (30 × 40 Å) that spontaneously fluoresces upon illumination with blue light (here, a redshifted variant (λEX ) 474 nm) with peak emission at λ ) 508 nm). In nature, GFP transduces the blue chemiluminescence of aequorin into green light. Although it has been used primarily as an optical tag for studying living cells, there has been some recent interest in the use of GFP for optoelectronic devices or for pH or ion sensorss
applications that require it to be organized into a robust framework.23,24 The spatial localization of GFP into the porous, cross-linked PVA framework can be readily achieved by exploiting the greater hydrophilicity of the pore interior versus the surface of the polymeric network. Optical microscopy can be used to monitor the selective incorporation/spatial distribution of the GFP in the porous polymer network. Differential interference contrast (DIC) microscopy, an approach used to render contrast in transparent specimens, was employed to image the network and clearly shows a region of strong contrast between the pores and the cross-linked polymer (Figure 4A). Protein loading was achieved by short periods of incubation for the surface supported, mesoporous polymer network in GFP aqueous solutions followed by a rinse in a high-salt buffer. The spatial distribution of GFP into the network structure was imaged using fluorescence microscopy. Bright fluorescent regions patterned in a closepacked hexagonal array (corresponding to GFP localization in the holes) and dark, nonfluorescent regions (the crosslinked polymer) were observed (Figure 4B). Because the feature sizes are close to the diffraction limit of the focus of the optical microscope, the observed pattern is somewhat “fuzzy”.25 They are sharp enough, however, to clearly show a pattern of regions of high intensity versus regions of low intensity. Observation of the green fluorescence indicates that the GFP retains the tertiary structure of the native form.26 The lack of appreciable amounts (minimal fluorescence intensity observed) of GFP on the cross-linked PVA shows that the increased hydrophobicity is enough to render the polymer resistant to protein binding. The Fourier transform (FT) of the fluorescence array (in Figure 4B) produces an intensity pattern that clearly exhibits 6-fold (2-D hexagonal) symmetry (Figure 4C), further demonstrating the selective partitioning of the GFP into the pores. Samples that were not incubated in GFP revealed a nontextured/patterned weak background fluorescence signal (data not shown), which was evenly distributed throughout the thin film. A digital overlay of the DIC micrograph of the polymeric network and the (23) You, Y. J.; He, Y. K.; Borrows, P. E.; Forrest, S. R.; Petasis, N. A.; Thompson, M. E. Adv. Mater. 2000, 12, 1678-1681. (24) Cinelli, R. A.; Pellegrini, V.; Ferrari, A.; Faraci, P.; Nifosi, R.; Tyagi, M.; Giacca, M.; Beltram, F. Appl. Phys. Lett. 2001, 79, 33533355. (25) Matsushita, S. I.; Yagi, Y.; Miwa, T.; Tryk, D. A.; Koda, T.; Fujishima, A. Langmuir 2000, 16, 636-642. (26) Enoki, S.; Saeki, K.; Maki, K.; Kuwajima, K. Biochemistry 2004, 43, 14238-14248.
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fluorescence micrograph of the GFP-loaded material (Figure 4D) clearly demonstrate that the protein is preferentially found in the open holes, or hydrophilic regions, and is not retained strongly on the hydrophobic polymer, thus demonstrating that this approach is a simple and efficient means for patterning proteins. A prolonged storage of the samples yielded no evidence to suggest the diffusion of the protein into the polymeric framework structure. Most notably, a key advantage this approach has over microcontact printing is the ability to form closepacked arrays or high-density protein arrays. Conclusions In summary, we have developed a straightforward, selfassembly-based means to fabricate laterally patterned, ordered polymeric arrays of wells for the spatial localization of proteins. A biocompatible polymer, PVA, patterned into a hexagonal porous array, is rendered chemically and mechanically stable through the use of a PAG that catalyzes the chemical cross-linking of the polymer. The PVA cross-linked framework features good optical transparency and protein resistance, and its open, hydrophilic pores are available for protein incorporation. As a verification of the process, we demonstrated the localization of a soluble protein, GFP, into the wells. The localization is readily achieved by tuning the relative hydrophobicity of the cross-linked polymer relative to the
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pore interior. The lateral patterning may ultimately be tuned by controlling the size of the particle template and hence the pore dimensions and the spacing between the wells of localized protein. This approach offers the potential for producing high-density biomolecular arrays that are patterned on the length scales that are not readily achievable by photolithography and soft lithographic techniques (which typically have a characteristic feature size of 5-500 µm).27 Future work will be directed toward enhancing the binding of the proteins by implementing an affinity ligand derivatization of the surface of the underlying substrate or the interior of the polymer pore, studying (quantitatively) the loading levels of protein and determining the orientation of the spatially localized protein. Work exploring these opportunities and the utility of this approach for membrane proteins is now in progress. Acknowledgment. This work was performed under the auspices of the Office of Basic Energy Sciences, Division of Materials Sciences, United States Department of Energy, under contract No. W-31-109-ENG-38. We thank Adam Reynolds and Dawn Shaw for assistance in the production of the GFP. LA051948F (27) Xia, Y.; Whitesides, G. M. Angew. Chem., Int. Ed. 1998, 37, 551-575.
