Langmuir 1998, 14, 741-744
741
Patterned Protein Layers on Solid Substrates by Thin Stamp Microcontact Printing C. D. James,*,† R. C. Davis,† L. Kam,‡,§ H. G. Craighead,† M. Isaacson,† J. N. Turner,§,| and W. Shain§,| School of Applied and Engineering Physics, Cornell University, Ithaca, New York 14853, Department of Biomedical Engineering, Rensselear Polytechnic Institute, Troy, New York 12180, Wadsworth Center, New York State Department of Health, Albany, New York 12208, and Department of Biomedical Sciences, School of Public Health, University of Albany, Albany, New York 12208 Received September 18, 1997. In Final Form: December 11, 1997 Microcontact printing (µCP) is a new method of molecularly patterning surfaces on a micrometer scale. In this paper, we present the extension of microcontact printing to producing patterned layers of proteins on solid substrates. µCP avoids the use of strong acids and bases necessary in photolithographic patterning, allowing its use for patterning of proteins and other biological layers. We also describe the methods of thin stamp microcontact printing that allow printing of isolated features previously unattainable by microcontact printing. A solution of polylysine in borate-buffered saline was printed onto a glass coverslip, yielding micrometer scale features over an area of 4 cm2.
Introduction Molecular patterning of solid substrates has many biological applications including biosensing, medical implants, and control of cell adhesion and growth.1-6 Localization of a cell to a desired position on a sample surface has been realized by microscopic molecular patterning of surfaces.3,4,6,7 High-resolution photolithographic and focused laser methods have been used to pattern surfaces with molecular layers.2,7 However, photolithography requires the use of harsh solvents and bases, making it incompatible with many biological molecules. The laser method uses an interference technique that does not allow generation of patterns of arbitrary complexity. Here we introduce a new method of producing patterned protein layers which has specific importance to a number of emerging technologies including advanced tissue engineering, biomineralization, DNA computing, and cultured neural networks.8-11 Microcontact printing (µCP) is a new method of chemically and molecularly patterning surfaces on a submicrometer scale. It has been used to pattern self-assembled monolayers (SAMs) of compounds such as hexadecanethiol and octadecyltrichlorosilane †
Cornell University. Rensselear Polytechnic Institute. § New York State Department of Health. | University of Albany. ‡
(1) Corn, R. M.; Jordan, C. E.; Frey, B. L.; Kornguth, S. Langmuir 1994, 10, 3642. (2) Dontha, N.; Nowall, W. B.; Kuhr, W. G. Anal. Chem. 1997, 69, 2619. (3) Chen, C. S.; Mrksich, M.; Huang, S.; Whitesides, G. M.; Ingber, D. E. Science 1997, 276, 1425. (4) Morgan, H.; Pritchard, D. J.; Cooper, J. M. Biosens. Bioelectron. 1995, 10, 841. (5) St John, P. M.; Kam, L.; Turner, S. W.; Craighead, H. G.; Issacson, M.; Turner, J. N.; Shain, W. J. Neurosci. Methods 1997, 75, 171. (6) Clark, P.; Connolly, P.; Curtis, A. S. G.; Dow, J. A. T.; Wilkinson, C. D. W. Development 1990, 108, 635. (7) Kleinfeld, D.; Kahler, K. H.; Hockberger, P. E. J. Neurosci. 1988, 8, 4098. (8) Healy, K.; Thomas, C.; Rezania, A.; et al. Biomaterials 1996, 17, 195. (9) Mooney, J. F.; Hunt, A. J.; et al. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 12287. (10) Corn, R. M.; Frutos, A. G.; Liu, Q.; Thiel, A. J.; et al. Nucleic Acids Res. 1997, 25, 4748. (11) Stenger, D. A., McKenna, T. M., Eds. Enabling Technologies for Cultured Neural Networks; Academic Press, Inc.: San Diego, CA, 1994.
(OTS) respectively on gold12 and SiO213,14 surfaces. µCP has an advantage over conventional photolithographic patterning methods in that it requires no harsh chemicals, making it suitable for patterning biologically active layers. Additionally, it should be possible to pattern multiple molecular layers by repeated application of µCP. However, two problems in µCP as presently used hinder its use in printing arbitrary patterns of biological molecules. First, the poly(dimethylsiloxane) (PDMS) elastomer used for the stamp in µCP is hydrophobic. When printing with waterbased, biological solutions (such as proteins in saline solution), the poor wetting characteristics of the hydrophobic stamp yield extremely nonuniform patterns. The second problem is that the soft elastomer stamp lacks the rigidity necessary for precision alignment and geometrical control of the pattern. The large compliance of the stamp also complicates printing small isolated features due to sagging between features. We describe here two developments in µCP which resolve these problems. The first is a method for printing protein layers. Here we describe the printing of polylysine layers on glass coverslips. Polylysine was chosen because of its utility in promoting selective attachment of cells to surfaces. The second development is the use of a thin elastomeric stamp on a rigid glass backing in order to facilitate printing of small isolated features and improve alignment and geometrical control of the patterns. A fluorescent tag was conjugated to the polylysine used in these experiments to enable visualization of the printed polylysine on the substrate surface. This allows for a quick verification of the quality of a given printing procedure. Atomic force microscopy (AFM) measurements were performed on the printed layers to determine layer thickness and coverage. Stamping with Aqueous Protein Solutions. µCP has been applied in the field of microlithography to achieve chemical modification of substrate surfaces, but patterning with polarized compounds such as aqueous protein solutions has not been investigated. The elastomer typically (12) Kumar, A.; Biebuyck, H. A.; Whitesides, G. M. Langmuir 1994, 10, 1498. (13) Xia, Y.; Mrksich, M.; Kim, E.; Whitesides, G. M. J. Am. Chem. Soc. 1995, 117, 9576. (14) St. John, P. M.; Craighead, H. G. Appl. Phys. Lett. 1996, 68, 1022.
S0743-7463(97)01048-2 CCC: $15.00 © 1998 American Chemical Society Published on Web 01/28/1998
742 Langmuir, Vol. 14, No. 4, 1998
Letters
Figure 1. (a) Printing isolated features using conventional thick stamp µCP. Stamp sagging causes printing in undesired locations. (b) Printing isolated features using a thin stamp with a rigid back support; stamp sagging is eliminated.
used for µCP, poly(dimethylsiloxane) (PDMS), is hydrophobic (due to the nonpolar methyl sidegroups along the siloxane backbone of the polymer chain). This presents a problem when attempting to use water-based, biological solutions as the “ink” in µCP. Some molecules such as polylysine, which contain positively charged amine side chains, present a similar problem of being difficult to deposit on PDMS for transfer to a substrate. However, studies have shown that when PDMS is treated in a lowtemperature plasma cleaner under oxygen, hydrophilicity and protein adsorption are temporarily increased on the surface of the polymer.15,16 X-ray photoelectron spectroscopy studies17 have shown that the oxygen plasma creates species which attack the silicon-carbon bonds, separating the methyl groups from the siloxane backbone, thus increasing the amount of oxygen on the surface of the
Figure 2. (a) Thin stamp fabrication: a master is placed onto PDMS. (b) Pressure distributes the PDMS into a thin layer. (c) Final thin stamp after curing, cooling, and rinsing.
PDMS. This induces hydrophilicity since hydrogen bonding is now able to occur between the surface of the PDMS and water molecules (as well as amine groups). However, hydrophobicity returns with time due to several factors such as diffusion of polarized groups into the bulk of the polymer.17 Here we use a low-temperature plasma treatment of PDMS to enable the patterning of polylysine on glass coverslips. Thin Stamp Microcontact Printing. µCP has been applied primarily in cases where a large percentage of the elastomer surface is in contact with the substrate during stamping. We attempted to apply conventional µCP to produce isolated features on a glass substrate. During
Figure 3. SEM micrographs of isolated features printed with thin stamp µCP. All scale bars are 50 mm. SEM images were taken at a 75° angle. (a) SEM micrograph of a thin stamp containing an array of 20 mm dots centered on 10 mm wide lines. Lines are 240 mm in length. (b) Corresponding fluorescent image of a printed polylysine line. (c) SEM micrograph of an isolated 25 mm diameter dot on a thin stamp. (d) Corresponding fluorescent image of a printed polylysine dot.
Letters
Langmuir, Vol. 14, No. 4, 1998 743
Figure 4. (a) Polylysine pattern intended to be produced on a glass substrate using conventional thick stamp µCP. (b) Resulting pattern due to sagging of the PDMS stamp in a recessed region. Light regions indicate polylysine.
stamping, a weight is placed on the back of the stamp, forcing the stamp to make intimate contact with the surface being printed on. Under the applied force, the elasticity of the stamp allows sagging to occur in large recessed areas of the stamp surface. These recessed regions of the stamp make contact with the surface resulting in chemical modification of the surface in undesired locations (Figure 1a). This renders the conventional method of “thick” stamp µCP inadequate for printing isolated features. The thin stamp design described here utilizes a glass slide as a rigid support for a thin PDMS stamp with a base thickness of approximately 10 µm and features of approximately 10 µm in height (Figure 1b). The height of the features allows for stamping to occur normally, while the rigid back support prevents sagging in larger recessed areas on the stamp surface while the stamp is under pressure. Experimental Section Thin Stamp Fabrication. Using photolithographic techniques, 10 mm thick photoresist (AZ Hoechst 4903) on silicon wafers is patterned to create masters for the µCP procedure. Our (15) Garbassi, F.; Morra, M.; Occhiello, E. Polymer Surfaces: From Physics to Technology; John Wiley & Sons Ltd.: England, 1994; p 427. (16) Holly, F. J.; Owen, M. J. Physicochemical Aspects of Polymer Surfaces; Mittal, K., Ed.; Plenum Press: New York, 1981; Vol. 2, p 625. (17) Morra, M.; Ochiello, E.; Marola, R.; Garbassi, F.; Johnson, D. J. Colloid Interface Sci. 1990, 137, 11.
master patterns consist of rectangular arrays of lines and dots in the resist. The lines are 240 mm in length of varying widths from 2 to 20 mm on a 250 mm pitch spacing. The dots vary in diameters from 2 to 20 mm also on a 250 mm pitch spacing. Large rectangles (5 × 2.5 mm) were placed on both sides of the line and dot patterns to keep an excess of pressure from being exerted on the fine line and dot features. To prevent the PDMS from bonding to exposed silicon on the masters, the wafers are fluorinated with perfluorodecyltrichlorosilane vapor under a nitrogen environment for 2 h. The stamps are fabricated from 2 × 2 cm silicon masters cleaved from the master wafer. A drop of Sylgard 184 silicone elastomer (Dow Corning Corp.) mixed with curing agent in a 10:1 ratio, is placed onto a microscope slide. A 2 × 2 cm section of the master is then placed on top of the drop of PDMS (Figure 2a). A 50 g weight is placed on top of the master, leaving a thin layer of elastomer between the master and the glass backing (Figure 2b). The weighted stack is placed in a vacuum desiccator for 12 h to remove bubbles from the elastomer. This results in a base height (not including the ∼10 mm feature height) of the PDMS stamp of approximately 10 mm. The elastomer is cured for 1.5 h at 60 °C, during which the PDMS forms a strong bond to the glass backing. After the stamp is allowed to cool, the master is carefully removed from the PDMS stamp and the stamp is rinsed in acetone to remove photoresist from the stamp surface (Figure 2c). Scanning electron microscopy (SEM) micrographs shown in parts a and c of Figure 3 display the resulting thin stamp features. µCP Polylysine Solution. Polylysine hydrobromide, (C6H12N2OHBr)n, with an average Mw ) 37 600 g/mol, was obtained from Sigma Chemical Co. The fluorescent aminereactive dye 5,6-carboxytetramethylrhodamine, succinimidyl ester (TAMRA,SE; Molecular Probes Inc., Catalog no. C-1171) is dissolved in dimethyl sulfoxide (DMSO) and then conjugated to the polylysine at a 10:1 dye-to-protein molar ratio. The conjugation is carried out in borax-boric acid buffer (0.1 M, pH ∼8.5) using deionized, Millipore-filtered water, and a Slide-ALyzer 2K MWCO Dialysis Cassette Kit (Pierce, Prod. # 66207) is used to separate unbound dye from dye-conjugated polylysine. The final µCP “ink” consists of 50 mL of dye-conjugated polylysine, 150 mL of DMSO, and 1 mL of borax-boric acid buffer. Prior to stamping, this solution was filter sterilized with 0.22 mm syringe filters (Fisher Scientific). Substrate Patterning Using Microcontact Printing. Immediately before beginning the microcontact printing (µCP) procedure, a temporary increase in hydrophilicity and protein adsorption of the stamps is achieved through treatment in a low-temperature plasma cleaner/sterilizer (Harrick Scientific) evacuated with a mechanical roughing pump. After this treatment, the polylysine solution is placed on the surface of the patterned stamp for 15 min to allow for protein adsorption and then blown dry with a nitrogen blow-off gun for 2 min with compressed nitrogen regulated to 50 psi. The stamp is placed in contact with a glass coverslip (Carolina Biological Supply, Catalog no. 63-3033) for 15 min under a weight of 50 g to promote electrostatic bonding between the polylysine molecules and the glass substrate. The coverslips are then rinsed in borate buffer and Millipore filtered water to remove unbonded polylysine. Characterization. Fluorescence Microscopy. A mercury arc lamp, which has a principal line at 546 nm, was used to induce fluorescence of the TAMRA, SE dye (absorption peak 547 nm, emission peak 575 nm) conjugated to the polylysine. A Zeiss Axiotron optical microscope with an Omega Optical XF101 fluorescence filter set was used to observe the protein patterns on the glass substrate. AFM. A Digital Instruments Nanoscope3a multimode atomic force microscope (AFM) was used in tapping mode to acquire the images. A drive voltage of 25 mV was used to generate a 2 V modulation signal with a Digital instruments TESP probe. This corresponds to a dither amplitude of approximately 20 nm. A set point of 70% of the maximum voltage for stable feedback was used. SEM. SEM micrographs of the µCP stamps were acquired on a Hitachi S800 SEM at 10 kV. The PDMS stamps were coated with Au-Pd prior to imaging.
744 Langmuir, Vol. 14, No. 4, 1998
Letters
Figure 5. Tapping-mode AFM image of the edge of a printed polylysine feature. An average profile of the edge is shown in the insert. The profile was generated by averaging the line scans in the boxed region in the image. A transition length of less than 100 nm is shown.
Results A conventional thick stamp was used to stamp polylysine on a glass surface. Figure 4a is a sketch of the desired printed pattern. Figure 4b is a fluorescence image of the actual stamped pattern. The TAMRA tagged polylysine shows up as light regions. Stamp sagging caused polylysine to be transferred from recessed regions of the stamp. The sagging of the stamp does not allow small isolated features to be printed by conventional µCP. This difficulty with stamp sagging can arise whenever the span of the recessed region becomes much larger than the recess depth of the stamp. In turn, if the recess depth is increased to alleviate sagging, small stamp features become overly compliant. Isolated polylysine features were printed with a thin elastomer stamp. The stamp pattern consists of adjacent arrays of isolated 5-25 mm wide lines and 5-25 mm diameter dots. A fluorescence image of a 20 mm dot centered on an isolated 10 mm line is shown in Figure 3b. The dark regions surrounding the line show that no stamp sagging has occurred. A fluorescence image of an isolated 25 mm dot is shown in Figure 3d. Complete polylysine coverage is seen on the dot and no polylysine transfer is detected in undesired regions. The coverage and thickness of the stamped polylysine layer were determined by AFM. However, due to the change in chemical composition seen by the probe as it scans over the edge, the tapping mode AFM profile does not give an accurate measurement of the polylysine layer thickness. Figure 5 shows an AFM image of the edge of a polylysine line printed with thick stamp µCP. The image indicates good coverage and yields a thickness measurement of 0.5 nm. The resolution of the printing process is excellent. The transition length at the polylysine edge is less than 100 nm. These measurements correspond to the printing procedure described above. The thickness of the polylysine layer is dependent on the compressed
nitrogen flow rate and drying time as well as the pressure applied to the stamp by the weight. The sharpness of patterns on stamped coverslips appears to degrade with successive use of a stamp and occasionally the raised features on the stamps themselves would be torn from their position upon separation from the glass coverslip. For use in cell culture work the printed pattern must be robust in saline solution environments. Fluorescence images indicate the pattern did not degrade with immersion in boric acid-borax buffer for 48 h. Fluorescence images taken after storage of the patterned coverslips under refrigeration in air for 5 weeks indicated no diffusion of polylysine to originally unstamped regions. Conclusions High-resolution patterned polylysine layers have been produced by µCP. Hydrophobicity of PDMS elastomers was successfully averted through low-temperature plasma treatment. A process for fabricating thin elastomer stamps on a stiff glass backing has been developed allowing for high-resolution micrometer-sized, isolated polylysine features to be printed on glass substrates. This results in a relatively inexpensive, fast, and reproducible method for modifying glass surfaces with proteins and should be compatible with a wide variety of biological molecules. The ability to chemically pattern surfaces with biological proteins provides a necessary step toward the control of adhesion and growth of cells on surfaces. Acknowledgment. This work was partially supported by a grant from DARPA/ITO. The authors acknowledge the use of the facilities at the Cornell Nanofabrication Facility, which is one node of the National Nanofabrication Users Network (NNUN) funded by the National Science Foundation, and the Cornell Materials Science Center funded by DMR of the NSF. C.D.J. was supported by a NSF fellowship. LA9710482
