Controlling Cell Adhesion on Human Tissue by Soft Lithography

To organize a single layer of cells of uniform phenotype, therefore, we explore the use of soft lithography to control RPE cell adhesion and morpholog...
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Langmuir 2004, 20, 4155-4161

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Controlling Cell Adhesion on Human Tissue by Soft Lithography Christina J. Lee,† Mark S. Blumenkranz,‡ Harvey A. Fishman,‡ and Stacey F. Bent*,† Department of Chemical Engineering, Stanford University, Stanford, California 94305-5025, and Department of Ophthalmology, Stanford University School of Medicine, Stanford, California 94305-5308 Received August 11, 2003. In Final Form: February 9, 2004 Soft lithographic techniques are widely used for fundamental biological applications. This study investigates the extension of soft lithography for use on human tissue to create a biological implant by systematically studying the effect of pattern size on cellular morphology. We focus on mimicking a key layer of the physiological retina with an organized monolayer of epithelial cells to act as a new treatment for age-related macular degeneration. We show that epithelial cells can be confined to cytophilic islands defined on lens capsule by the inhibitory polymer poly(vinyl alcohol). In addition, as the size of the cytophilic islands grows, both the fraction of islands with cells attached and the number of cells adhered to each island increase. High densities of cell adhesion and single cell attachment per island were achieved with a 25 µm pattern size. Over time, the cells spread over the 5 µm wide barriers to form a confluent monolayer that may eventually serve as a functional retinal implant. With the ability to apply soft lithography to tissue samples, human tissue may become a universal membrane substrate for other ocular diseases or in tissue engineering applications elsewhere in the body.

Introduction Over the past decade, the techniques of soft lithography have gained widespread use. Soft lithographic techniques, which include microcontact printing, microfluidic networking, micromolding in capillaries, and replica molding, offer many advantages over conventional lithography, including their low cost, ready accessibility to general users, and compatibility with a range of sample geometries. Because of these advantages, soft lithographic methods have been studied extensively for a wide variety of biological applications. They are being utilized, for example, in the direction of cell adhesion, the control of apoptosis, and the fabrication of biosensors and cell sorters.1-3 Other potential applications include high-throughput screening of drug candidates, cell-based assays for monitoring changes in intracellular calcium, and DNA separations.4 In most of these examples, the soft lithographic techniques have been applied to flat and rigid materials, such as glass, plastic, and silicon.5-11 Restriction of soft lithography to these traditional substrates, however, may ultimately limit the range of biological applications. Although glass, silicon, and plastic are easily modified for selective cell adhesion, they may not be biocompatible or have the †

Stanford University. Stanford University School of Medicine. * To whom correspondence should be addressed.

‡

(1) Ito, Y. Biomaterials 1999, 20, 2333-2342. (2) Kane, R.; Takayama, S.; Ostuni, E.; Ingber, D.; Whitesides, G. Biomaterials 1999, 20, 2363-2376. (3) Quake, S. R.; Scherer, A. Science 2000, 290, 1536-1540. (4) Mitchell, P. Nat. Biotechnol. 2001, 19, 717-721. (5) Folch, A.; Toner, M. Annu. Rev. Biomed. Eng. 2000, 2, 227-256. (6) Wilbur, J.; Kumar, A.; Biebuyck, H.; Kim, E.; Whitesides, G. Nanotechnology 1996, 7, 452-457. (7) Luk, Y.; Kato, M.; Mrksich, M. Langmuir 2000, 16, 9604-9608. (8) Lu, L.; Kam, L.; Hasenbein, M.; Nyalakonda, K.; Bizios, R.; Gopferich, A.; Young, J.; Mikos, A. Biomaterials 1999, 20, 2351-2361. (9) Branch, D.; Wheeler, B.; Brewer, G.; Leckband, D. IEEE Trans. Biomed. Eng. 2000, 47, 290-300. (10) St. John, P. M. J. Neurosci. Methods 1997, 75. (11) Wheeler, B.; Corey, J.; Brewer, G.; Branch, D. J. Biomech. Eng.s Trans. ASME 1999, 121, 73-78.

proper permeability or flexibility to function appropriately in the body. The range of applications can be extended considerably by using soft materials. Soft materials possess the flexibility to conform to the shape of the tissues and the organs in the body and may minimize damage due to friction if the implants shift against neighboring tissue. Further optimization of the substrate, particularly in applications of transplantation, may ultimately lead to the selection of human tissue as the soft material of choice. This selection is expected to be driven primarily by concerns of biocompatibility, which in the case of synthetic materials is still inadequate for permanent implants.12 Human tissue extracted from the patient, on the other hand, is autologous and may circumvent the immune response, which is the major hurdle in all implants. It is expected that in order to make functional implants using tissue, one will likely need to modify the surface properties of the tissue, particularly with respect to cell adhesion. This can potentially be done using soft lithography. Yet although microcontact printing (µCP) and microfluidic networking (µFN) have been extended to form patterns on synthetic soft materials such as biodegradable polymer surfaces, the use of these techniques on tissue is largely unexplored. We propose that the methods of soft lithography will provide a powerful means to spatially modify the surfaces of tissue just as it has with nonbiological substrates. We previously have reported initial studies showing that soft lithographic techniques can be adapted for use on human tissue.13 Here we focus on systematically investigating the effect of pattern size on cellular morphology and density. We demonstrate the application of these techniques to organize retinal pigment epithelial cells on human tissue for the development of a retinal implant to treat age-related macular degeneration. (12) Ikada, Y.; Tsuji, H. Macromol. Rapid Commun. 2000, 21, 117132. (13) Lee, C.; Huie, P.; Leng, T.; Peterman, M.; Marmor, M.; Blumenkranz, M.; Bent, S.; Fishman, H. Arch. Ophthalmol. 2002, 120, 1714-1718.

10.1021/la035467c CCC: $27.50 © 2004 American Chemical Society Published on Web 04/09/2004

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Figure 1. Human retinal pigment epithelial cells. Scanning electron micrograph of RPE cells in a human retina obtained from an eye bank. The cells have retained their uniform shape and organization but have lost structural features, such as microvilli. The scale bar is 10 µm.

Age-related macular degeneration (AMD) is the most prevalent form of blindness among individuals in industrialized nations over 65.14 The earliest manifestations of AMD are the deterioration of retinal pigment epithelial (RPE) cells and a change in the underlying basement membrane, resulting in lowered permeability and modification of the metabolism of the membrane. RPE cells are vital to the function of the retina as each RPE cell supports approximately 40 photoreceptors (the specialized neurons that convert light to a chemical signal) by regulating their flow of nutrients and by metabolizing their wastes.15 A promising treatment for AMD is to replace the diseased RPE cells with new, healthy cells. In the human retina, the healthy RPE cells form a highly organized, close-packed layer (Figure 1). In a transplant, the implanted RPE cells should mimic the size and morphology with retention of tight junctions and other essential metabolic functions. Different approaches to creating a functional implant have been explored. One approach is the injection of a suspension of epithelial cells into the subretinal space, which has been shown to rescue some photoreceptors for a limited period of time.16 However, simply injecting the pigmented cells into the subretinal space is unlikely to resolve AMD because the cells fail to form well-ordered monolayers, and they may arrange themselves into multilayers with random orientations and phenotypic variability.17 In contrast, the function of the RPE cells (14) Klein, R. In Age-Related Macular Degeneration; Maguire, M. G., Ed.; Mosby: St. Louis, MO, 1999; pp 31-55. (15) Marmor, M. F. In The Retinal Pigment Epithelium; Marmor, M. F., Wolfensberger, T. J., Eds.; Oxford University Press: New York, 1998; p 6. (16) Thumann, G.; Aisenbrey, S.; Schraermeyer, U.; Lafaut, B.; Esser, P.; Walter, P.; Bartz-Schmidt, K. U. Arch. Ophthalmol. 2000, 118, 13501355.

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Figure 2. RPE cells cultured on human lens capsule. Hoffman modulation contrast image of RPE cells cultured on human lens capsule for 1 day. The scale bar is 100 µm.

may be highly dependent on the orientation and morphology of the cells.18 We are pursuing a second approach in which epithelial cells are cultured on a substrate to create a single organized layer. The success of this approach, however, will depend on two issues: biocompatibility of the substrate and the phenotype of RPE cells on the substrate. To address the concerns of biocompatibility, we have focused on tissue. The tissue that we examined as a substrate for RPE culture and transplantation is lens capsule. The lens capsule is an extracellular matrix membrane, composed mainly of proteins, such as collagen IV, heparan sulfate proteoglycan, and fibronectin. It is easily obtainable, as it is routinely removed in cataract surgery. The anterior lens capsule is approximately 5 mm in diameter and 9-14 µm thick. This membrane is attracting much interest because in addition to being biocompatible, it may offer the possibility of an autologous transplant. Thus, lens capsule may serve as a good substitute for Bruch’s membrane, which is only slightly thinner at 2-4 µm and has a similar compositional makeup. Figure 2 shows that lens capsule encourages indiscriminate epithelial cell attachment, leading to a disorganized cell layer. The disorganized cells exhibit a range of morphologies and phenotypes, which in turn may lead to diminished cellular function. To organize a single layer of cells of uniform phenotype, therefore, we explore the use of soft lithography to control RPE cell adhesion and morphology on this tissue. Experimental Section Materials. Human lens capsules extracted during cataract surgery (IRB protocol ID # 74930) were stored in phosphate (17) Zarbin, M. A.; Sugino, I. K.; Castellarin, A. A. In Age-Related Macular Degeneration; Maguire, M. G., Ed.; Mosby: St. Louis, MO, 1999; pp 363-382. (18) Singhvi, R.; Kumar, A.; Lopez, G. P.; Stephanopoulos, G. N.; Wang, D. I. C.; Whitesides, G. M.; Ingber, D. E. Science 1994, 264, 696-698.

Cell Adhesion on Human Tissue buffered saline (PBS) at 4 °C. Following sterilization in PBS under UV (254 nm) for 3 h, the lens capsules were treated with 0.05% trypsin-EDTA for 10 min at 37 °C. In vivo, the lens capsule is covered with native epithelial cells. The epithelial cells are removed with trypsin to provide a clean, initial substrate. Lens capsule curls and floats in solution, requiring it to be spread onto a surface before culturing cells on it. In addition, lens capsule is extremely delicate; it must be handled carefully to prevent tearing. With the aid of tweezers and a flexible, plastic spatula, the lens capsule was removed from solution and, under a stereoscope, was spread out in a single layer onto a plastic coverslip (Thermanox, EM Sciences, Fort Washington, PA) to give it structural integrity and then allowed to dry in a laminar flow hood. (All reagents were obtained from Invitrogen, Carlsbad, CA.) Human retinal pigment epithelial cells (ARPE-19, ATCC, Manassas, VA) were maintained in D-MEM/F-12 supplemented with 10% fetal bovine serum at 37 °C, 6.5% CO2. The cells were separated from 100 mm tissue culture dishes by treatment with 0.05% trypsin-EDTA for 5 min at 37 °C, using physical dislodgment, followed by resuspension in culture media. The cells were passaged weekly at a 1:10 ratio. Lens capsules were seeded at an approximate density of 103 cells/mm2. Cells were counted in a Bright-line hemacytometer (Hausser Scientific, Horsham, PA). The cells were allowed to settle and adhere onto the lens capsule in culture media in a 37 °C, 6.5% CO2 environment. Soft Lithography. Poly(dimethylsiloxane) (PDMS) stamps were prepared as described in the literature.19 Briefly, a chrome mask with the desired micron-sized patterns was fabricated at the Stanford Nanofabrication Facility using conventional photolithography. The mask was used to pattern a 7 µm positive photoresist-coated silicon wafer (SPR220-7, Shipley Co., L.L.C., Marlborough, MA). PDMS in a 10:1 mixture of elastomer to curing agent (Sylgard 184, Dow Corning Corp., Midland, MI) was poured onto the patterned silicon wafer, degassed, and cured at 100 °C. After 1 h, the PDMS stamp (mold) was removed from the patterned silicon wafer. PDMS stamps (0.5 cm2 ) were exposed to an air plasma (Harrick PDC-32G, Ossining, NY) for 1 min at 100 W to obtain a hydrophilic surface. The soft lithographic techniques were modified to allow for the printing of PVA as described previously.13 In particular, current methods of µCP rely on the evaporation of the solvent to form a thin, physisorbed layer on the PDMS stamp. This layer is then transferred from the PDMS stamp to the substrate. In the case of PVA, we were unable to pattern the tissue in this manner. Since PVA has good adhesive properties, if only a thin layer using a small amount of PVA is formed on the PDMS stamp, minimal PVA is found to transfer. The strong adhesion to the elastomer was verified by examining the PDMS stamp under a microscope after printing; the images show clearly that the PVA remains on the stamp. Thus, we have employed a “wet transfer” method to deposit PVA on tissue. To stamp the lens capsule surface, the PDMS stamp was placed carefully onto a thin layer of 2% poly(vinyl alcohol) (PVA) (MW 70 000-100 000, Sigma, St. Louis, MO) in DI water (18.2 MΩ cm-1) with 0.1 mg/mL fluorescein (Sigma, St. Louis, MO). Immediately after contact, the PDMS stamp was removed from the thin layer of solution and placed in contact with the lens capsule to “wet transfer” the solution. A 40 g weight was placed on top of the stamp for 30 min. After removal of the elastomer stamp, the lens capsule remained dry and adhered to the plastic coverslip. The microprinted lens capsule was then sterilized under a UV lamp (254 nm) for 3 h. RPE cells were subsequently cultured on the sterilized lens capsule. For microfluidic patterning (µFN), the same chrome mask was used to pattern a negative photoresist (SU-8, MicroChem Corp, Newton, MA) that was spun onto a silicon wafer. The resulting PDMS stamp (0.5 cm2) has honeycomb channels that are 30 µm tall. For flow of the aqueous PVA solution, the PDMS stamp was first plasma cleaned (1 min at 100 W in air plasma) to form hydrophilic walls. The stamp was then placed face down on the lens capsule and sandwiched by a second plastic coverslip. The (19) Whitesides, G.; Ostuni, E.; Takayama, S.; Jiang, X.; Ingber, D. Annu. Rev. Biomed. Eng. 2001, 3, 335-373.

Langmuir, Vol. 20, No. 10, 2004 4157 lens capsule and stamp were submerged in PVA for 5 min, allowing the channels to fill with solution. A 40 g weight was applied to ensure conformal contact between the stamp and the lens capsule and to prevent the channels from leaking. Following submersion, the lens capsule and stamp were removed from the solution and allowed to dry in the dark to prevent photobleaching of the fluorescein. After 48 h, the weight was removed, and the lens capsule was examined under a fluorescence microscope. Finally, the lens capsule was sterilized under UV at 254 nm for 3 h. Microscopy. Lens capsule was imaged with a digital camera (Nikon Cool Pix 900) attached to an inverted microscope. All fluorescently stamped lens capsules were imaged using the inverted microscope (Nikon Eclipse TE300) with a xenon source (75 W) through a FITC filter (Chroma Technology Corp., Brattleboro, VT) connected to a digital camera (Hamamatsu OrcaER). Both patterned and unpatterned cells were imaged on the inverted microscope using a Hoffman modulation contrast condenser. Scanning electron microscopy images were obtained by fixing cells in 5% gluteraldehyde/2% paraformaldehyde in 0.1 M sodium cacodylate buffer, pH 7.4 for 3 h. They were washed in sodium cacodylate (0.1 M, pH 7.4) and postfixed in 1% osmium tetroxide for 3 h at 4 °C. Finally, the cells were dehydrated in a graded series of alcohols before they underwent critical point drying and were coated with gold. All electron microscopy materials were from EM Science, Fort Washington, PA. The viability of patterned RPE cells was tested with the LIVE/ DEAD Viability/Cytotoxicity Assay Kit (Molecular Probes, Eugene, OR). The assay is a 2 µM calcein acetoxymethyl AM ester and a 4 µM ethidium homodimer-1 solution in RPE buffer (14.2 mM NaCl, 5.6 mM KCl, 2.2 mM CaCl2 H2O, 3.6 mM NaHCO3, 1 mM MgCl2‚6H2O, 30 mM Hepes Buffer, pH 7.4). The cells were washed with RPE buffer before being incubated in the LIVE/DEAD solution for 30 min at room temperature in the dark. After loading, the cells were rinsed in prewarmed buffer (37 °C) to remove excess stain and observed in this buffer. The calcein ester, which can be observed with a fluorescein filter, permeates cells with intact plasma membranes. It becomes more intensely fluorescent upon activity of an ubiquitous intracellular esterase, indicating that the patterned cells are alive. Ethidium homodimer-1, which can be observed with the Texas Red filter, binds nucleic acids. It is not cell membrane permeable, and thus, its fluorescence is only enhanced for cells with damaged membranes.

Results and Discussion Stamps were fabricated for both µCP and µFN approaches. A Hoffman modulation contrast image of a typical poly(dimethylsiloxane) (PDMS) stamp used in the µCP experiments is shown in Figure 3A. Each stamp is 0.5 cm × 1 cm in size. The fluid deposits wherever the raised honeycomb lines of the PDMS stamp contact the lens capsule. For the example shown in Figure 3A, the width of the raised honeycomb lines is 2 µm, and each recessed hexagon is 25 µm in diameter. The feature height is 7 µm. For µFN, a stamp with the opposite features is required. The scanning electron micrograph (SEM) in Figure 3B is of a PDMS stamp used to define the channel walls for µFN. The channels, 10 µm in width, surround 50 µm hexagonal pillars. In this sample, the walls are vertical and smooth, reach 30 µm in height, and retain their hexagonal shape. The hexagonal pillars protect the underlying lens capsule while the fluid flows through the channels. Because lens capsule encourages cell attachment, we must deposit molecules that prevent cell adhesion so that the cells adhere only to unmodified regions of lens capsule. Soft lithography can then be used to confine the RPE cells to a specific size. The choice of molecular “ink” was determined by examining several known inhibitory polymers, including bovine serum albumin, octadecyltricholorosilane, poly(2-hydroxyethyl methacrylate), and poly(vinyl alcohol). We found the best results with poly-

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Figure 3. PDMS stamps. (a) Hoffman modulation contrast image of PDMS stamp used in microcontact printing. The raised honeycomb lines are 2 µm in width and 7 µm in height. The recessed hexagons are 25 µm in diameter. The scale bar is 25 µm. (b) Scanning electron micrograph of PDMS stamp used in microfluidic networking. The 30 µm tall hexagon pillars are 50 µm in diameter separated by 10 µm. The scale bar is 25 µm.

(vinyl alcohol) (PVA), a biocompatible polymer which is currently used in many biomedical applications.20 For example, it has U.S. Food and Drug Administration approval in application as embolization particles for reducing blood loss in certain types of brain surgery. Thus, we believe that PVA will likely be well tolerated in other systems and not cause an immune response in the subretinal space. As described below, we have determined that PVA deters RPE cell adhesion, consistent with literature reports on the inhibitory properties of PVA toward bovine endothelial cells,21 and exhibits excellent stability on the tissue. The human lens capsule patterned by µCP is shown in the fluorescence image of Figure 4A. Although the lens capsule autofluoresces, the PVA/fluorescein mixture fluoresces more brightly than does the unmodified tissue and is clearly visible in a fluorescence micrograph. The PVA/ fluorescein mixture can be identified in Figure 4A as the 10 µm wide honeycomb pattern, and the unmodified tissue, corresponding to recessed regions of the PDMS stamp, appears as an array of dark hexagons 50 µm in diameter. These unmodified regions are designed to act as the cytophilic islands. Figure 4B is a Hoffman modulation contrast image of RPE cells after 9 h of culture on the PVA-patterned lens capsule, while Figure 4C is a fluorescence image of RPE cells after 1 day in culture using the LIVE/DEAD stain. Shown in Figure 4C is the excitation of live dye; no (20) Giusti, P.; Lazzeri, L.; Barbani, N.; Narducci, P.; Bonaretti, A.; Palla, M.; Lelli, L. J. Mater. Sci.: Mater. Med. 1993, 4, 538-542. (21) Sugawara, T.; Matsuda, T. J. Biomed. Mater. Res. 1995, 29, 1047-1052.

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fluorescence from the dead cell dye was observed. Figure 4D is a high-magnification Hoffman modulation contrast image of the patterned RPE cells after 3 days in culture. We can make several observations from Figure 4. First, we observe in Figure 4B that the RPE cells selectively attach to the lens capsule, avoiding the PVA-coated regions. The PVA pattern is hence successfully reproduced in the cellular pattern. Second, we detect that on average three to four cells adhere to each 50 µm diameter cytophilic island. Third, we observe in Figure 4C that all the cells remain viable. They fluoresce intensely upon excitation with UV (ex/em 495/515 nm) after loading with the live stain, illustrating that each patterned cell is alive. The corresponding image of the cells upon excitation of the dead stain (ex/em 495/635 nm) is not shown because all present cells are viable. The only signal seen at 635 nm is a background signal from the autofluorescence of the lens capsule. The RPE cells confined to cytophilic islands also show structural characteristics of normal RPE cells. Examination under electron microscopy (not shown) exhibits the presence of microvilli.13 In addition, although the coverage is inhomogeneous, the patterning on lens capsule is quite widespread, covering almost the entire sample. Under the low magnification of Figure 4C, the borders of the lens capsule can be seen; the region on the left is the underlying plastic coverslip. This low-magnification view illustrates that we are able to process lens capsule while retaining its integrity. We have also observed excellent stability for the pattern transferred in PVA. For example, the particular sample shown in Figure 4 was reseeded with RPE cells 7 days after the initial patterned cells were removed in a harsh sterilization process to remove bacterial contamination. The sterilization procedure consisted of multiple, vigorous rinse steps and incubation for 30 min in RPE media supplemented with 1 mg/mL gentamicin. Following sterilization, the newly added human RPE cells continued to adhere only to regions of unmodified tissue, indicating that the PVA retained its inhibitory properties. On the other hand, while the PVA performs well in inhibiting cell adhesion, it does not completely block the spreading of cells once they have adhered to the attachment sites. For example, we observe in Figure 4D that after 3 days the cells spread over PVA barriers and contact neighboring cells. This ability of the cells to spread over the PVA barriers after initial organization by patterning is beneficial to the creation of a viable RPE transplant, since a close-packed layer is desired as the final product. Hence, this series of measurements illustrates that we are able to (1) achieve cell patterns that follow the PVA template over large areas of lens capsule and (2) create a layer of RPE cells that is initially organized. It also allows us to verify the viability of the cells and confirm PVA stability. To explore the dependence of the RPE cell confinement on the pattern size, we have used PDMS stamps of various sizes to apply PVA to lens capsule and monitored the resulting adhesion and spreading of the cells. Figure 5 shows the results of a pattern incorporating 15 µm cytophilic islands, a scale which is closer to that present in the native macula, separated by 4 µm lines. A fluorescence image of human lens capsule patterned in PVA by µFN using this 15 µm stamp is shown in Figure 5A. The pattern is widespread, although there are a few defects where the borders of the cytophilic islands are poorly defined. Figure 5B is a Hoffman modulation contrast image of RPE cells cultured for 6 h on the patterned lens capsule. It can be seen that the cells preferentially attach to the unmodified areas of the lens

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Figure 4. Cell adhesion to 50 µm cytophilic islands. (a) Fluorescence image of human lens capsule patterned by µCP. The PVA honeycomb lines are 10 µm in width and separate 50 µm regions of hexagonal shape of unmodified lens capsule. The unmodified lens capsule appears dark in the image. The scale bar is 100 µm. (b) Hoffman modulation contrast image of human RPE cells on the PVA-patterned lens capsule after 9 h in culture (seeded at 2 × 103 cells/mm2). The scale bar is 100 µm. (c) Fluorescence image of viable human RPE cells patterned on human lens capsule after 1 day, measured by LIVE/DEAD stain. This image was taken after sterilization in 1 mg/mL gentamicin. The scale bar is 1 mm. (d) High-magnification Hoffman modulation contrast image of human RPE cells cultured for 3 days on PVA-patterned human lens capsule (second seeding 7 days later at 103 cells/mm2). The scale bar is 50 µm.

capsule and not to the PVA lines, consistent with the results of the 50 µm study. However, unlike on the 50 µm hexagonal pattern, in which multiple cells attached to each hexagon, we observe that only a single cell adheres to each island on the 15 µm pattern. On the other hand, the lens capsule is now very sparsely populated by cells, leaving many attachment sites devoid of cells. Figure 5C shows that after 24 h the cells start to spread and adhere onto neighboring attachment sites. They also exhibit clumping behavior, showing a preference for neighboring cells over the lens capsule. We postulate that this is due to the migratory behavior of the cells. With time, the cells that have attached to the lens capsule will migrate toward other cells nearby and will adhere to these, forming cell clumps. We do not believe that the defects in the pattern are the cause of this clumping behavior since the appearance of clumping is more widespread than the occurrence of defects. Thus, with the 15 µm hexagonal pattern, we are able to achieve initial single cell adhesion to each cytophilic island, but the attachment is too sparse to create the desired packed layer of RPE cells. To compare with the 15 and 50 µm attachment sites, we also investigated a PDMS stamp with an intermediate sized pattern: 25 µm in diameter separated by 2 µm. The human lens capsule patterned with PVA by µCP using this stamp is shown in the fluorescence image

of Figure 6A. Figure 6B is a Hoffman modulation contrast image of the patterned lens capsule that has been seeded with human RPE cells. It is apparent that the RPE cells have adhered only to the cytophilic islands created on the lens capsule, as was observed in the 50 and 15 µm samples. The RPE cells have also been confined to a diameter no larger than 25 µm, with each cytophilic island occupied by a single cell. Figure 6C shows a fluorescence image of RPE cells after 24 h of culture on patterned lens capsule. The cells, loaded with the LIVE/ DEAD stain, fluoresce from the live stain and show evidence in this image of retaining both the pattern and cell viability. Over longer periods of time, the RPE cells begin to spread over the PVA barriers and contact each other to form a monolayer, as was seen in the 50 µm patterns. It is evident from Figures 4-6 that pattern size has an effect on RPE cell adhesion. We have systematically explored this effect for a range of pattern sizes and have summarized the results in Table 1. From our studies, we observed several clear trends. First, the occupation fraction (the fraction of islands with at least one cell adhered to it) though exhibiting scatter, generally grows with increasing diameter of the cytophilic island. A large jump occurs between 15 and 20 µm, which is a possible indication that the cytophilic area defined by the 15 µm pattern is too small for cell adhesion. We believe this leads to the

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Figure 5. Cell adhesion to 15 µm cytophilic islands. (a) Fluorescence micrograph of human lens capsule patterned by µFN. The cytophilic islands are 15 µm in diameter separated by PVA lines of 4 µm. The scale bar is 50 µm. (b) Hoffman modulation contrast image of human RPE cells cultured for 6 h on the patterned lens capsule (seeded at 103 cells/mm2). The scale bar is 50 µm. (c) Hoffman modulation contrast image of human RPE cells cultured for 24 h on the patterned lens capsule. The scale bar is 50 µm.

clumping behavior that is observed in cells confined to the 15 µm islands but is not seen at larger pattern sizes. Second, as expected, the occupation number (the average number of cells attached to each cytophilic island) increases with pattern size. As the island increases in size, there is a higher probability that multiple cells may adhere to one island. Finally, we note that although the occupation fraction and the occupation number both

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Figure 6. Cell adhesion to 25 µm cytophilic islands. (a) Fluorescence image of human lens capsule patterned by µCP. The cytophilic islands are 25 µm in diameter and are separated by 2 µm PVA lines. The scale bar is 100 µm. (b) Hoffman modulation contrast image of human RPE cells (seeded at 5 ×103 cells/mm2) after 2.5 h in culture on the patterned lens capsule. LC indicates the areas that are lens capsule surface. P indicates the regions that are plastic. The scale bar is 100 µm. (c) Fluorescence micrograph of human RPE cells patterned on human lens capsule after 24 h. The cells were stained with LIVE/DEAD stain. Viable cells appear bright in this image. LC indicates the areas that are lens capsule surface. P indicates the regions that are plastic. The scale bar is 100 µm.

increase with island size, the island density drops by virtue of the pattern. Therefore, the overall cell density, which is the product of these three values (occupation fraction, occupation number, and island density), remains relatively constant over all pattern sizes. We do observe

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Table 1. Summary of Cell Density as a Function of Pattern Sizea island diameter (µm)

occupation fraction

island density (mm-2)

occupation number

cell density (mm-2)

15 20 25 30 35 40 50

0.15 0.57 0.46 0.62 0.59 0.55 0.81

4234 3127 2079 1482 1110 863 564

1.0 1.0 1.4 1.5 1.6 1.6 3.0

578 1704 1161 1048 1030 772 1254

a Island diameter refers to the diameter of the patterned island. Occupation fraction is the fraction of islands with at least one cell adhered to it. Island density is the number of islands patterned per square mm. Occupation number is the average number of cells attached to each cytophilic island. Cell density is the number of cells adhered in each square mm.

a peak in cell density at 20 µm, and thus, this pattern size may be the best candidate for creating a functional RPE layer. First, with a high cell density, there are potentially enough cells in the layer to be able to support the photoreceptors, and second, with an occupation number of one, we have more control over the morphology and size of every single cell. Soft lithography is clearly effective in confining RPE cells to a range of sizes on lens capsule. The methods of soft lithography, therefore, do extend to tissue. There are several modifications and challenges that were found to be necessary in adapting to biological samples. First, the lens capsule, at only 15 µm in thickness, requires careful manipulation to prevent tearing. A supporting material to give it structural integrity was required. By spreading lens capsule onto plastic before processing, we were able to deposit PVA via soft lithography onto the tissue. Second, although the application of pressure in soft lithography has previously been studied extensively because of concerns of deformation in the PDMS stamp,22 a critical issue with tissue samples is deformation of and damage to the substrate itself. Hence, control of pressure needs to be reevaluated in this application. The use of excessive pressure on the stamp can distort the lens capsule before deformation of the elastomer occurs. This distortion can cause topographical changes that may affect the cell adhesion. Historically, microcontact printing has most commonly utilized self-assembled monolayers. Tissue, however, has a complex surface with unknown chemical functionalities, and the molecular specificity of the linkages needed to attach self-assembled monolayers would be difficult to achieve on such samples. A thin polymer film, on the other hand, performs well without the need for specific surface sites. We have found that a thin polymer layer of PVA adheres to the tissue, retains pattern integrity, and deters cell adhesion. The optimal method for depositing this polymer film on tissue was found to be a wet-transfer variant of microcontact printing. Whereas current methods typically involve evaporation of the solvent prior to the transfer of material, PVA is transferred most successfully and reproducibly to the tissue when the solvent is retained. Since PVA is often used in adhesives, it follows (22) Hui, C.; Jagota, A.; Lin, Y.; Kramer, E. Langmuir 2002, 18, 1394-1407.

logically that PVA remains on the PDMS stamp even after attempts are made to transfer the PVA to the tissue and that a modification to transferring the PVA while solvent remains is necessary. However, this modification leads to some cases of inhomogeneity being introduced into the patterns. Because the PVA is still fluid when it is transferred, it is possible with excessive pressure to push some of the material to the edge of the pattern. As a result, more material lies just to the edge of the honeycomb lines than in the lines itself, shrinking the cytophilic area. This is the case shown in Figure 4A. PVA in the honeycomb lines appears less intense than the region just outside the lines. With optimization of the pressure applied to the elastomer, however, this wet-transfer variation is a reliable and reproducible method to pattern the tissue with homogeneous lines. We also explored µFN and were able to obtain good results with this approach, as it shares many similarities with the modified wet transfer method of µCP. The biggest advantage of µFN lies in its potential for the simple formation of complex patterns of different functional groups. For example, different solutions can be flowed down separate channels, without disrupting existing patterns, or alternatively, laminar flow under controlled conditions allows the transfer of multiple solutions without mixing in single channels. Since this complex patterning is not required for the creation of the RPE layer, we have chosen to use the wet transfer variant of µCP in the studies reported here, except for those shown in Figure 5. Conclusion The soft lithographic techniques, µCP and µFN, have been modified in this study to extend beyond silicon, glass, and biodegradable polymers to human tissue. Adapting these techniques to tissue extends the range of possible biological applications. We have successfully patterned the lens capsule with a honeycomb network of inhibitory PVA, creating cytophilic islands of unmodified lens capsule. RPE cells selectively adhered to these islands and were able to spread across PVA barriers after several days to contact their neighbors. The confinement helped to define the morphology in terms of size. It is hoped that this control will help retain the function of the cell; studies are now ongoing to explore this question. Culturing RPE cells onto lens capsule brings us one step closer to creating an autologous implant that can restore function to the retina. Finally, microprinted human tissue such as lens capsule or other biological membranes may prove applicable as a universal membrane substrate for other ocular diseases or in tissue engineering applications elsewhere in the body. Acknowledgment. We thank Aqueelah C. McKinley for assistance with the size dependence study. We also thank Roopa Dalal for her assistance with scanning electron microscopy. We thank Dr. Martin L. Fishman, Dr. Anne Fung, Dr. Joanne Giaconi, and Dr. Christopher Engelman for obtaining lens capsule specimens. This work was funded by a generous grant from VISX Inc. and the Stanford University Bio-X Interdisciplinary Initiatives Program. C.J.L. thanks the NIH Biotechnology Training Grant for support. LA035467C