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Nanotopographic Control of Cytoskeletal Organization Heather M. Powell,† Douglas A. Kniss,‡ and John J. Lannutti*,† Department of Materials Science and Engineering, and College of Medicine, Department of Obstetrics and Gynecology, Laboratory of Perinatal Research and Center for Biomedical Engineering, The Ohio State UniVersity, Columbus, Ohio 43210 ReceiVed NoVember 7, 2005. In Final Form: January 25, 2006 Growth of 3T3-L1 preadipocytes on a nanoscalar poly(ethylene terephthalate) (PET) surface produced an absence of the intracellular stress fibers characteristic of cell growth on “normal” planar surfaces. This phenomenon was consistently observed from time zero throughout 3 days of culture and was accompanied by changes in paxillin expression along with an approximately 50% decrease in the number of adherent cells in response to 500 dynes/cm2 of shear stress. This suggests that the cytoskeleton in cells adherent to nanofibrillar surfaces does indeed form, but at a smaller, more difficult to observe scale. We propose a novel mechanism by which the growth and clustering of integrin-associated focal adhesions on surface nanofibrils regulates cytoskeletal development. The width of the extracellular matrix contacts is constrained by the width of the nanofibrils and the absence of any surface between them. The limited dimensions of these point contacts then constrain receptor polymerization and the associated aggregation of actin filaments. The existence of a topographic mechanism leading to growth-limited integrin clustering is hypothesized.
Introduction A unifying design consideration in tissue engineering involves seeding cells onto scaffolds for the purpose of generating threedimensional (3D) tissue-like structures. An important challenge involves the promotion of specific levels of cellular adhesion, motility, orientation, and cell-matrix and cell-cell communication sufficient to encourage the formation of a neotissue that adequately mimics the properties of the desired endogenous tissue. Many investigations have studied the effect of scaffold properties on cell behavior in an effort to determine those characteristics that elicit a desired response. Surface topography has been extensively studied as one of the classic factors known to control cell behavior.1-9 However, its ability to alter either adhesion to * Corresponding author. Mailing address: 477 W Hall, 2041 College Rd., The Ohio State University, Columbus, OH 43210; tel: 614-292-3926; fax: 613-292-1537; e-mail:
[email protected]. † Department of Materials Science and Engineering. ‡ Laboratory of Perinatal Research and Center for Biomedical Engineering. (1) Brunette, D. M. Fibroblasts on micromachined substrata orient hierarchically to grooves of different dimensions. Exp. Cell Res. 1986, 164, 11-26. (2) Brunette, D. M. Spreading and orientation of epithelial cells on grooved substrata. Exp. C. Res. 1986, 167, 203-217. (3) Chehroudi, B.; Gould, T. R. L.; Brunette, D. M. Effects of grooved epoxy substratum on epithelial cell behaviour in vitro and in vivo. J. Biomed. Mater. Res. 1988, 22, 459-473. (4) Chehroudi, B. B. M.; Brunette, D. M. The effects of micromachined surfaces on formation of bonelike tissue on subcutaneous implants as assessed by radiography and computer image processing. J. Biomed. Mater. Res. 1997, 34, 279-290. (5) Chehroudi, E.; Gould, T. R. L.; Brunette, D. M. A light and electron microscopic study of the effects of surface topography on the behavior of cells attracted to titanium-coated percutaneous implants. J. Biomed. Mater. Res. 1991, 25, 387-405. (6) Perizzolo, D.; Lacefield, W. R.; Brunette, D. M. Interaction between topography and coating in the formation of bone nodules in culture for hydroxyapatite- and titanium-coated micromachined surfaces. J. Biomed. Mater. Res. 2001, 56, 494-503. (7) den Braber, E. T.; de Ruijter, J. E.; Smits, H. T. J.; Ginsel, L. A.; von Recum, A. F.; Jansen, J. A. Effect of parallel surface microgrooves and surface energy on cell growth. J. Biomed. Mater. Res. 1995, 29, 511-518. (8) den Braber, E. T.; de Ruijter, J. E.; Ginsel, L. A.; von Recum, A. F.; Jansen, J. A. Quantitative analysis of fibroblast morphology on microgrooved surfaces with various groove and ridge dimension. Biomaterials 1996, 17, 2037-2044. (9) Chesmel, K. D.; Clark, C. C.; Brighton, C. T.; Black, J. Cellular response to chemical and morphologic aspects of biomaterials surfaces. II. The biosynthetic and migratory response of bone cell populations. J. Biomed. Mater. Res. 1995, 29, 1101-1110.
a scaffold surface or cell-substrate signaling remains poorly defined. An evolving parallel focus has suggested that cells are sensitive to the modulus of the surface.10-12 A comprehensive understanding of cell-substrate interactions requires a thorough investigation of cellular adhesion. Cellular adhesion provides environmental cues that influence cellular decisions to proliferate, differentiate, or migrate13 and is a critical initial step in the formation of cellular communities. Adhesion is mediated by specialized transmembrane proteins (integrins) with surface-adsorbed extracellular matrix (ECM) proteins. At these sites of attachment to a surface, integrin receptors cluster, leading to the formation of focal adhesions (FAs) and the recruitment of several intracellular signaling molecules, including the cytoskeletal proteins vinculin and paxillin.14 Localization of these cytoskeletal-associated proteins to the cytoplasmic face of the FAs triggers a physical linkage between the ECM and the actin cytoskeleton.15-17 FAs are believed to regulate adhesion and locomotion in response to an environmental stimuli and act as sites of force generation and transmission.18-20 (10) Lo, C. M.; Wang, H. B.; Dembo, M.; Wang, Y. L. Cell movement is guided by the rigidity of the substrate. Biophys. J. 2000, 79, 144-152. (11) Wong, J. Y.; Velasco, A.; Rajagopalan, P.; Pham, Q. Directed movement of vascular smooth muscle cells on gradient-compliant hydrogels. Langmuir 2003, 19, 1908-1913. (12) Genes, N. G.; Rowley, J. A.; Mooney, D. J.; Bonassar, L. J. Effect of substrate mechanics on chondrocyte adhesion to modified alginate surfaces. Arch. Biochem. Biophys. 2004, 422, 161-167. (13) Turner, A. M. P.; Dowell, N.; Turner, S.; Kam, L.; Isaacson, M.; Craighead, H. G.; Shain, W. Attachment of astroglial cells to microfabricated pillar arrays of different geometries. J. Biomed. Mater. Res. 2000, 51, 430-441. (14) Miyamoto, S.; Teramoto, H.; Coso, O. A.; Gutkind, J. S.; Burbelo, P. D.; Akiyama, S. K.; Yamada, K. M. Integrin function - molecular hierarchies of cytoskeletal and signaling molecules. J. Cell Biol. 1995, 131, 791-805. (15) Nikolopoulos, S. N.; Turner, C. E. Actopaxin, a new focal adhesion protein that binds paxillin LD motifs and actin and regulates cell adhesion. J. Cell Biol. 2000, 151, 1435-1447. (16) Turner, C. E.; Kramarcy, N.; Sealock, R.; Burridge, K. Localization of paxillin, a focal adhesion protein, to smooth-muscle dense plaques, and the myotendinous and neuromuscular-junctions of skeletal-muscle. Exp. Cell Res. 1991, 192, 651-655. (17) Jockusch, B. M.; Bubeck, P.; Giehl, K.; Kroemker, M.; Moschner, J.; Rothkegel, M.; Rudiger, M.; Schluter, K.; Stanke, G.; Winkler, J. The molecular architecture of focal adhesions. Annu. ReV. Cell DeV. Biol. 1995, 11, 379-416. (18) Sawhney, R. K.; Howard, J. Molecular dissection of the fibroblast-traction machinery. Cell Motil. Cytoskeleton 2004, 58, 175-185.
10.1021/la052993q CCC: $33.50 © 2006 American Chemical Society Published on Web 04/27/2006
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Paxillin, often referred to as an FA adapter protein, is a small 559 amino acid protein localized to FAs.21 Paxillin contains a number of motifs, including LD motifs, LIM motifs, an SH3 domain binding site, and SH2 domain binding sites, all of which mediate protein-protein interactions.22 These modules allow the binding of a variety of structural and signaling molecules, including focal adhesion kinase (FAK) and vinculin.23 The phosphorylation of paxillin following integrin-dependent cell adhesion24 has long been associated with the coordinate formation of FAs and stress fibers24,25 and plays an important role in the regulatory dynamics of FAs.26 Because paxillin and the phosphorylation of paxillin are linked to many cellular processes, such as adhesion, migration, signaling, and cell survival, the native ability of any surface to modulate paxillin expression and activation would provide more in-depth control of cellular behavior. In this study, the ability of nanoscalar surfaces to alter cellular adhesion, morphology, cytoskeletal arrangement, and paxillin expression and localization was investigated. Nanofibrillar surfaces, comprised of fibrils roughly 20 nm in diameter and 200 nm in length, were generated on widely used poly(ethylene terephthalate) (PET) tissue culture inserts via oxygen plasma etching.23 High-resolution scanning electron microscopy (SEM) and atomic force microscopy (AFM) were used to characterize both untreated and treated/nanofibrillar versions of these PET surfaces prior to culture. We investigated the ability of these nanoscalar textures to alter cellular adhesion, morphology, cytoskeletal rearrangement, and paxillin expression and localization. Cells were cultured on these surfaces and analyzed for general morphology via SEM, actin organization via phalloidin staining, and cell-substrate interaction through the immunofluorescence of paxillin. In this context, the widespread assumption that nanoscalar features are completely immune to influence by the media merited additional attention. The data suggests that nanoscale modification of surface topography can influence cell adhesion and the assembly of FAs. Materials and Methods Cell Culture. Mouse 3T3-L1 preadipocytes were obtained from the American Type Culture Collection (ATCC). The cells were maintained in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS, Gibco), 2 mM l-glutamine, 1 mM sodium pyruvate, and penicillin-streptomycin. Cultures were incubated in a humidified (19) Zaidel-Bar, R.; Ballestrem, C.; Kam, Z.; Geiger, B. Early molecular events in the assembly of matrix adhesions at the leading edge of migrating cells. J. Cell Sci. 2003, 116, 4605-4613. (20) Pelham, R. J.; Wang, Y. L. Cell locomotion and focal adhesions are regulated by substrate flexibility. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 1366113665. (21) Shaw, L. M.; Turner, C. E.; Mercurio, A. M. The alpha-6a-beta-1-integrin and alpha-6b-beta-1-integrin variants signal differences in the tyrosine phosphorylation of paxillin and other proteins. J. Biol. Chem. 1995, 270, 2364823652. (22) Schaller, M. D.; Schaefer, E. M. Multiple stimuli induce tyrosine phosphorylation of the Crk-binding sites of paxillin. Biochem. J. 2001, 360, 5766. (23) Brown, M. C.; Curtis, M. S.; Turner, C. E. Paxillin LD motifs may define a new family of protein recognition domains. Nat. Struct. Biol. 1998, 5, 677-678. (24) Burridge, K.; Turner, C. E.; Romer, L. H. Tyrosine phosphorylation of paxillin and Pp125(Fak) accompanies cell-adhesion to extracellular-matrix - A role in cytoskeletal assembly. J. Cell Biol. 1992, 119, 893-903. (25) Seufferlein, T.; Rozengurt, E. Sphingosylphosphorylcholine rapidly induces tyrosine phosphorylation of P125(Fak) and paxillin, rearrangement of the actin cytoskeleton and focal contact assembly - Requirement of P21(Rho) in the signaling pathway. J. Biol. Chem. 1995, 270, 24343-24351. (26) Richardson, A.; Malik, R. K.; Hildebrand, J. D.; Parsons, J. T. Inhibition of cell spreading by expression of the C-terminal domain of focal adhesion kinase (FAK) is rescued by coexpression of Src or catalytically inactive FAK: A role for paxillin tyrosine phosphorylation. Mol. Cell. Biol. 1997, 17, 6906-6914.
Powell et al. chamber with 5% CO2/95% air at 37 °C. The medium was changed every 2 days. Surface Modification via Reactive Ion Etching. PET Thermanox plastic coverslips (Nalgene Nunc International, Rochester, NY) were used in this study. The thickness and diameter of the PET disks were 200 µm and 13 mm, respectively. For the adhesion studies, rectangular (60 × 20 mm) Thermanox coverslips were used. PET substrates were modified utilizing reactive ion etching following a method described previously27 using a Technics Micro-RIE Series 800-II (Technics, Concord, CA) operated at 50 W and 160 mTorr of oxygen supplied to the system at a rate of 25 cm3 m. PET coverslips were etched continuously for 30 min with oxygen plasma then placed upright in a multiwell cell culture plate to prevent subsequent contact of these delicate nanoscalar features with any other surface. Scanning Electron Microscopy. The topography of the etched and nonetched surfaces was qualitatively examined using SEM (FEI Sirion). Both etched and untreated films were sputter coated with a 10 nm layer of gold-palladium and analyzed in ultrahigh-resolution mode with a 5 kV accelerating voltage. Atomic Force Microscopy. The surface topography of the films was quantified using AFM (Digital Instruments Nanoscope IIIa, Veeco Metrology, Tucson, AZ) in tapping mode using an LTESP probe (Veeco Metrology, Tuscon, AZ). The PET films were examined in room-temperature phosphate-buffered saline (PBS) at a scan rate of 0.500 Hz. Several areas 25 µm2 in size were imaged and analyzed with Digital Instruments software version 4.22r2. Topographical roughness parameters Ra (average deviation from arithmetic centerline), Rmax (maximum peak height), and Rq (root-mean-square deviation of the surface) were determined. X-ray Photoelectron Spectroscopy (XPS). XPS analysis was performed on untreated and treated disks after rinsing consecutively with PBS and distilled water. Disks were analyzed (Kratos AXIS Ultra) for the percentage of C, N, and O using a mono Al source operated at 13 kV and 10 mA in hybrid lens mode. Cellular Proliferation and Morphology. Plasma-etched and untreated PET substrates were placed in a 24-well cell culture plate (containing 13-mm diameter wells) and sterilized by immersion in 70% alcohol for 30 min followed by washing with PBS followed by culture medium. Cells were seeded onto sterilized untreated and treated disks at a density of 1.5 × 104 cells/well and cultured for 7 days. On days 1, 2, 3, 5, and 7, cell proliferation was determined utilizing the MTS assay (CellTiter 96 AQueous non-radioactive cell proliferation assay, Promega, Madison WI). For each time point, duplicate disks were used, and experiments were repeated three times. Cell morphology on the surfaces was studied using SEM. Cells on PET disks were fixed with 3% glutaraldehyde in a 0.1 M phosphate buffer containing 0.1 M sucrose (pH 7.4) for 2 h. They were then washed with the sucrose-phosphate buffer for 30 min. The samples were osmicated in 1% OsO4 for 1 h and washed again. After dehydration in a graded ethanol series, the samples were then subjected to a graded ethanol-hexamethyldisilazane (HMDS, Ted Pella, Redding, CA) series (3:1 ETOH/HMDS to 100% HMDS). The specimens were immersed in 100% HMDS, dried overnight by evaporation and then sputter coated with Au-Pd and examined using SEM at 5 kV (Sirion, FEI). Transmission Electron Microscopy (TEM). To examine the cell-material interface, the 3T3-L1 fibroblasts were seeded onto untreated and treated disks as before. Following the 24 h culture period, the disks were rinsed with PBS and fixed for 30 min in 2% glutaraldehyde in 0.1 M phosphate buffer containing 0.1 M sucrose. The cell-seeded disks were then rinsed, post fixed in 1% osmium tetroxide for 30 min, and exposed to a graded ethanol series. Hydroxymethyl methacrylate (HPMA) was then used to rinse the disks prior to infiltration with Polybed 812 resin (Polysciences Inc., Warrington, PA). Polybed 812 resin was added to the wells, and the plate was slowly rocked for 1.5 h. The disks were then embedded in the Polybed by overnight polymerization at 60 °C. Cell-seeded disks were then trimmed and sectioned on a Reichert Ultracut E (27) Powell, H.; Lannutti, J. Nanofibrillar surfaces via reactive ion etching. Langmuir 2003, 19, 9071-9078.
Control of Cytoskeleton Organization ultramicrotome using a diamond knife cutting at 70 nm. Sections were stained for 15 min in 2% uranyl acetate followed by 5 min in Reynolds lead citrate, and examined with a Philips CM 12 TEM operated at 60 kV. Immunocytochemistry (ICC). The expression of filamentous or F-actin and the FA-associated protein paxillin were examined by phalloidin staining and ICC, respectively. Briefly, untreated and treated PET disks were seeded with cells (1.5 × 104 cells/13 mm diameter well) and cultured for 0-72 h with the zero point considered to be the time at which approximately 30% of the cells had attached to the untreated surface. This “zero hour” time point was established to assist in the analysis of cells at the onset of attachment and spreading. Cell attachment to the treated surface appeared to be slower but not markedly so. This time was determined to be 45 min after initial seeding; all subsequent experiments were performed with the zero point set to 45 min. The cell-seeded disks were then rinsed with PBS and fixed with 4% paraformaldehyde in PBS for 1 h. After fixation, the samples were rinsed gently with PBS and permeabilized with 0.5% Triton X-100 PBS for 15 min. Samples were pretreated for 1 h with 5% horse serum (HS) PBS followed by incubation overnight at 4 °C with the primary antibody for paxillin (Upstate Cell Signaling, Lake Placid, NY). The disks were thoroughly rinsed with PBS then stained with the appropriate secondary antibody for 1 h at room temperature. Disks were subsequently incubated with phalloidin (1:40, AlexaFluor phalloidin, Molecular Probes, Eugene, OR) for 1 h at room temperature. After rinsing with PBS, the double-labeled disks were examined via confocal microscopy (Zeiss 510 META). Negative controls were prepared following the same procedures but omitting the primary antibodies. Image Analysis. Fluorescence images were quantitatively analyzed for paxillin spot size at the 24 h time point. Briefly, paxillin images were reduced to 8-bit files then thresholded to reduce the amount of noise. Image J software was then used to calculate the area (µm2) of each paxillin spot. For each sample treatment, a minimum of 10 images were taken from three different experiments. Results were reported as a histogram of the spots larger than a specified area. Adhesion Studies.The relative adhesion strength of cells grown on both untreated and treated PET rectangular coverslips was tested using a parallel plate device constructed following Kawamoto et al.28 To ensure that this arrangement did not create turbulent flow conditions, a tracking dye was added to the inlet port prior to the initial media flow. The flow of dye was tracked visually, and the flow pattern was clearly laminar. For adhesion experiments, cells were seeded at a density of 2.4 × 103 cells/mm2 on the nanofibrillar and untreated PET coverslips for 24 h. This seeding density, lower than that used for ICC and proliferation assays, was chosen to ensure that cells did not reach confluence. If cells reach confluency, the media could flow primarily above the cells decreasing the amount of shear stress.28 Seeded PET coverslips were then placed into the parallel plate device with the cell side oriented toward the gasket and exposed to a laminar flow of DMEM supplemented with 25 mM HEPES. The apparatus was maintained at 37 °C in a water bath. Cells were exposed to 0-500 dynes/cm2 of shear stress for 5 min. Following exposure to shear, those cells remaining were removed using trypsin and counted (Coulter Multisizer II, Beckman). Each experiment was repeated six times and expressed as average ( SEM. Statistical Analysis. For cell growth and adhesion, a one-way analysis of variance (ANOVA), followed by Tukey-Kramer multiple comparison analysis, was performed. The data were presented as either mean ( standard deviation or mean ( SEM, and p < 0.05 was considered significant.
Results Surface Characteristics of Modified PET Substrates. As shown previously,27 reactive ion etching of as-received, relatively (28) Kawamoto, Y.; Nakao, A.; Ito, Y.; Wada, N.; Kaibara, M. Endothelial cells on plasma-treated segmented-polyurethane - Adhesion strength, antithrombogenicity and cultivation in tubes. J. Mater. Sci.: Mater. Med. 1997, 8, 551-557.
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Figure 1. As-received PET tissue culture coverslips.
dense PET (Figure 1) with oxygen for 30 min produced a nanofibrillar surface topography with fibrils roughly 300 nm in length and 20 nm in diameter (Figure 2A). However, upon exposure to media for 24 h at 37 °C, these surface fibrils become unstable, as is evident by the appearance of rafts of fractured nanofibrils and their remaining bases (Figure 2B,C). To produce thicker, more stable nanostructures, “partially cooled” structures27,29 were used. This technique generated shorter (∼200 nm) clustered fibrils: this clustering provided additional strength (Figure 2D), while retaining a distinctive nanoscalar character. Examination after exposure to media at 37 °C for 24 h revealed that these thermally bonded nanofibers were physically stable, as neither raft formation nor fractured fibrils were observed. This surface was then characterized and quantified under PBS utilizing wet-mode AFM. The average roughness of the nanofibrillar surface was 51 ( 6 nm and the maximum peak height was 402 ( 156 nm (Figure 3). The average roughness of the untreated surface was 4 nm. Reactive plasmas have been shown to produce an increased affinity for water through the addition of polar groups to macromolecules; however, this enhanced hydrophilicity is not stable with time.30-32 The wetting angle of etched PET returns to its original value following exposure to air for 24 hours.31 This “hydrophobic recovery” is believed to result from the movement of surface species into the bulk, restoring the original chemical nature of the surface. XPS analysis for oxygen, nitrogen, carbon, and sulfur revealed that oxygen plasma appeared to result in a slight oxygen enrichment (Table 1). However, the accuracy of chemical quantification by XPS is typically reported as 5-10%; it can range from 1% for spectra in which the signal strength is high, up to 20% for spectra containing a large amount of noise and produced using a low signal strength.33 Given the relatively low signal strength (due to the nanoscaled nature of the surface33), these differences in mass concentration are not statistically significant. (29) Derns, B. C.; Rodriguez, F. The role of heat transfer during reactive-ion etching of polymer films. J. Vac. Sci. Technol. 1990, B8, 1985-1989. (30) Nakamatsu, J.; Delgado-Aparicio, L. F.; Da Silva, R.; Soberon, F. Aging of plasma-treated poly(tetrafluoroethylene) surfaces. J. Adhes. Sci. Technol. 1999, 13, 753-761. (31) Beake, B. D.; Ling, J. S. G.; Leggett, G. J. Scanning force microscopy investigation of poly(ethylene terephthalate) modified by argon plasma treatment. J. Mater. Chem. 1998, 8, 1735-1742. (32) Bourceanu, G.; Gheorghiu, M. D.; Moisa, C. Effect of the gas nature and etching time of poly(ethylene terephthalate) (PET) film on the high-frequency plasma. ReV. Roum. Phys. 1985, 30, 145-150. (33) Tougaard, S. Quantification of nano-structures by electron spectroscopy. In Surface Analysis by Auger and X-Ray Photoelectron Spectroscopy; Briggs, D., Grant, J. T., Eds.; IM Publications: West Sussex, U.K., 2003.
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Figure 2. SEM images of the PET coverslip surface (A) after etching for 30 min in oxygen plasma, (B) after etching followed by soaking in media at 37 °C for 24 h (low magnification), (C) after etching followed by soaking in media at 37 °C for 24 h (high magnification), and (D) after “thermal bonding” followed by soaking in media at 37 °C for 24 h (high magnification). The latter structure was used for all subsequent experiments. The image shown in panel C reveals that small “rafts” of broken fibrils (white arrow) are present following the exposure. Table 1. XPS wt % Analysis of As-Received and Oxygen Plasma-Etched PET Disksa sample
O
N
C
PET as-is PET etched
33.9 40.2
0.48 0.62
65.6 59.2
a Although the mean oxygen and nitrogen content of the two disks varies by as much as 6.4%, this disparity is within reported values for error on samples with low signal strength.
Cellular Proliferation. To assess cell growth on untreated and treated PET, nonradioactive MTS assays were performed at various times (1-7 days) after plating. We detected a statistically significant difference in 3T3-L1 growth on treated versus untreated surfaces for day 2 (p < 0.001) and day 5 (p < 0.05) (Figure 4), with the total cell number on the unmodified surface being greater than the modified surface. The differences in cell number were not substantial, however, and were not significant for all time points. Nanofeatures Alter Cell Morphology and Cytoskeletal Organization. Cell morphology was qualitatively evaluated via SEM at time points ranging from 0 to 72 h. Cells were categorized as either round (spherically shaped and had not begun to spread), spread (exhibits spreading but retains equiaxed shape), or elongated (has spread and was elongated). After 4 h in culture, 78 ( 21% of the cells on smooth surfaces were rounded in appearance, whereas only 6.9 ( 6.7% of the cells on the nanoscalar surfaces remained rounded. After 24 h in vitro, all cells were spread onto the growth surface and elongated (Figure 5A-D). When grown on the nanoscalar surfaces, the cells appear to be slightly larger and extend a greater number of nanoscalar lamellapodia to the surface (Figure 5D,E). Interestingly, these lamellapodia appear to subdivide and closely interact with the nanofibrils (Figure 5E) at a similar nanometric scale. The inset shows that these much smaller filopodia extend from the original cell boundary. Using an AlexaFluor phalloidin stain, we detected substantial differences in the organization of the actin cytoskeleton on cells
grown on nanofibrillar versus untreated PET surfaces. Cells on untreated surfaces formed a cortical ring of actin as the cells began to attach to the substrate, as evidenced from actin staining at 0 h (Figure 6A). Cells on treated surfaces had a dense diffuse stain of actin throughout the cell (Figure 6D). As the cells began to spread on the unmodified surface, stress fibers became visible (Figure 6B). In contrast, cells grown on treated substrates exhibited far less F-actin organization and did not contain visible stress fibers (Figure 6E). After 72 h in vitro, 3T3-L1 cells seeded on nanofibrillar surfaces showed no evidence of either clearly F-actin or stress fibers (Figure 6F). Beyond 72 h in vitro, these cells begin to achieve confluency; at this point cell-surface interactions become less dominant, complicating these observations. Substrate Nanofibrils Alter Paxillin Expression and Aggregation. The expression and localization of paxillin was determined via ICC on cells seeded for 0-72 h on both untreated and nanofibrillar surfaces. At early time points (t < 4 h), paxillin expression on both surfaces is comparable (data not shown). With further culture time, paxillin expression on untreated surfaces begins to differ from that on the treated surfaces (Figure 7). Cells on untreated surfaces are characterized by well-defined aggregates of paxillin localized at stress fiber caps (Figure 7A). In contrast, fibroblasts on treated surfaces display a relatively diffuse pattern and only a few regions of paxillin aggregation (Figure 7B). Image analysis was used to quantify this difference in paxillin aggregate size. Paxillin aggregates to a higher degree on untreated surfaces, with the mean aggregate size averaging 1 µm2; the majority of paxillin aggregates on treated surfaces are less than 0.4 µm2 in size (Figure 8). Nanofibrillar Surfaces and Cellular Adhesion. Quantification of the cells remaining on either surface following exposure to shear stress revealed a trend of decreasing cell numbers as shear stress increased. Interestingly, however, the average percentage of cells remaining on the untreated surfaces is greater than that on the nanofibrillar surfaces for all time points (Figure
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Figure 5. SEM micrographs of fibroblasts grown on control (A,B) and treated (C,D) disks. At higher magnifications (E), the filopodia appears to subdivide to interact with the nanofibrils at the same nanoscalar level via generation of successively smaller extensions.
point does the cellular membrane make contact with the “valleys” between these peaks.
Discussion
Figure 3. Wet-mode AFM conducted in PBS. (A) two-dimensional (2D) and (B) 3D images of the thermally bonded PET structure.
Figure 4. MTS assay of 3T3-L1 cells grown on both as-is and plasma-etched PET. Data presented as mean ( std.
9). Throughout much of this range, cells on untreated surfaces are approximately twice as likely to remain adherent. TEM of the cell-surface interface (Figure 10) shows that these cells contact only the uppermost points of the nanofibrils/ groups of nanofibrils. In cross-section, the surface itself shows nanofibrils originating from larger “peaks” in the surface. At no
Chemical and physical characterization of the as-received and treated surfaces reveals that the plasma etching process produces no significant alteration in surface chemistry (Table 1), but creates a noteworthy transformation in the surface topography (Figure 2). The form and particularly the stability of these nanoscaled surfaces is of obvious importance to the application of nanotopography to cell culture. The apparent destruction of the initial nanoscalar morphology is, to our knowledge, the first observation of nanostructural instability in this context. Brownian motion of the individual nanofibrils likely leads to oscillation and then localized agglomeration followed by fracture, producing nanofibril “rafts”, even before these surfaces are exposed to the action of adherent mammalian cells. This is an unexpected consequence of attempting to make a fully synthetic surface adopt a more biological appearance at realistic length scales. The robust nature of the “thermally bonded” surface allows cell culture to take place without altering topography while maintaining an “ECMlike” surface that better mimics the body’s endogenous scaffolding. Topographic enrichment of a surface has been generally regarded as a means of enhancing attachment in a variety of cell types. Thus, our observations of weaker cell adhesion accompanied by more limited actin reorganization when a nanoscale topography was present are clearly counterintuitive. Cells on nanofibrillar surfaces extend dendritic filopodia to these synthetic nanofibrils (Figure 5E), indicating that they mimic the 3D morphology of the natural ECM in such a way as to promote a strong filopodial response. Similar dimensional control over cell spreading has been seen in hepatocytes grown on 3D collagen foams with a pore size of 81.6 versus 9.9 µm.34 Hepatocytes within these large pore foams exhibited a spread phenotype in
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Figure 6. Inverted, grayscaled F-actin staining of cells grown on untreated disks at (A) 0 h, (B) 24 h, and (C) 72 h. Cells grown on treated disks at (D) 0 h, (E) 24 h, and (F) 72 h. Scale bar ) 20 µm (panels A and C) and 50 µm (panels B, C, E, and F).
which filopodia extended to the pore walls. In contrast, hepatocytes grown on small pore foams do not enter the scaffold and instead populate the surface of the construct while maintaining a cuboidal morphology with no extended filopodia. The paxillin ICC and phalloidin staining of cells cultured on untreated surfaces reveal large aggregates of paxillin at the end of each stress fiber. Cells grown on treated surfaces display a diffuse pattern of paxillin and no visible stress fibers. These data suggest that the lack of easily visible stress fibers in cells grown on treated surfaces is a consequence of alterations in FA formation. Similar instances involving spatial control of cellular behavior can be found in prior work by Ingber and others who showed that cell shape, attachment, and organization can be controlled by locally decorating planar surfaces with adhesion proteins.35 In this study, bacteriological dishes were coated with fibronectin (FN) at varying densities. Endothelial cells plated on dishes coated with high densities of FN (>500 ng/cm2) attached and spread, but did not form capillary tubes, in contrast to intermediate density coatings (100-500 ng/cm2). Cells cultured on surfaces with less than 100 ng FN/cm2 attached but remained rounded. Subsequent investigations indicated that endothelial cells exhibit increased attachment, spreading, and enhanced proliferation on surfaces with higher FN densities. Cells grown on substrates with greater than 2000 molecules per µm2 increased in cell number, whereas cells cultured on densities lower than 235 molecules/µm2 lost viability over time.36 Intermediate densities exhibited intermediate growth rates. Capillary endothelial-cell spreading and DNA synthesis have also been shown to increase with higher FN density surfaces.37 Massia and Hubbell38 observed that, at low densities (34) Ranucci, C. S.; Kumar, A.; Batra, S. P.; Moghe, P. V. Control of hepatocyte function on collagen foams: sizing matrix pores toward selective induction of 2-D and 3-D cellular morphogenesis. Biomaterials 2000, 21, 783-793. (35) Ingber, D. E.; Folkman, J. How does extracellular-matrix control capillary morphogenesis? Cell 1989, 58, 803-805. (36) Ingber, D. E. Fibronectin controls capillary endothelial-cell growth by modulating cell-shape. Proc. Natl. Acad. Sci. U.S.A. 1990, 87, 3579-3583. (37) Ingber, D. E.; Prusty, D.; Frangioni, J. V.; Cragoe, E. J.; Lechene, C.; Schwartz, M. A. Control of pH intracellular and growth by fibronectin in capillary endothelial-cells. J. Cell Biol. 1990, 110, 1803-1811. (38) Massia, S. P.; Hubbell, J. A. Human endothelial-cell interactions with surface-coupled adhesion peptides on a nonadhesive glass substrate and 2 polymeric biomaterials. J. Biomed. Mater. Res. 1991, 25, 223-242.
Figure 7. Confocal images of 3T3L-1 cells grown for 24 h following seeding onto untreated (A) and treated (B) substrates. Cells on both substrates were stained for actin (magenta) and paxillin (green). Note the diffuse actin and paxillin staining pattern in cells on the nanoscalar surfaces compared to the well-defined paxillin spots at the end of the stress fibers (see arrow) in cells on the smooth surfaces.
Control of Cytoskeleton Organization
Figure 8. Histogram of paxillin spot size on both untreated and treated PET substrates. All cells were cultured for 24 h.
Figure 9. Relative strength of adhesion of cells grown on treated and untreated surfaces for 24 h prior to exposure to a given shear stress for 5 min. Data reported as mean + SEM.
Figure 10. TEM micrograph of the cell-surface interface. In the cross-section, the nanofibrils are visible originating from the “peaks” of the PET surface supporting them. Some damage to the nanofibrils appears to be evident, and this is likely an artifact of the sample preparation process. The cell appears to be making contact only with the tips of the fibrils. The scale bar is 200 nm.
of surface-bound RGD peptides, cells attach but do not form visible stress fibers. However, at higher densities of RGD peptides (a spacing of ∼140 nm or less), spreading is accompanied by the development of FAs and stress fibers.38 A critical number of interacting integrins must be clustered for visible FAs and stress fibers to form. These and other examples39-43 concern intracellular control by the careful extracellular presentation of ligands on a flat substrate. In contrast, the system under investigation presents only deliberate variations in nanotopography. By using the novel
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Figure 11. Model for integrin-ligand binding on smooth versus nanofibrillar surfaces. (A) While, on nanofibrillar surfaces, surfaceadsorbed ligands are present both on the fibrils and in the valleys, cells are believed to bind only at tips of the fibrils. Thus the spatial restriction of the ligands results in restriction of integrin accumulation. F-actin forms, but additional F-actin clustering cannot occur because of the limited availability of nanofibrils. (B) In contrast, surfaceadsorbed ligands on smooth surfaces are continuous; this promotes the formation of F-actin, leading to bundling into stress fibers.
intersection of the concept of nanotopography with integrin assembly and stress fiber development, we can generate possible explanations for the observed effects. In formulating mechanistic insights and general principles regarding the influence of nanoscalar topography, however, it is important to first compare the scale of the nanofibrils to that of the cell itself. The width of these nanofibrils is approximately 1/500 of the cell diameter; the spacing between them is approximately 200 nm or about 15 integrins wide. Thus, only a finite number of integrins can be theoretically bound to the tip of a given nanofibril. In addition, the density of the nanofibrils must affect the spacing of the integrin clusters within a cell wall. With these caveats, Figure 11 (adapted from ref 44) shows our proposed model for cellular attachment to both nanofibrillar and untreated surfaces. The cells contact only the tops of the nanofibrils (as point contacts), and the proteins adsorbed there. The spacing of these nanofibrils is at least 200 nm, a distance greater than the 140 nm ligand density previously seen to facilitate FA formation.38 This low ligand density would then inhibit the formation of easily visible paxillin aggregation and stress fiber formation. The physical adhesion data shows that cells cultured on the nanostructured surfaces maintain ∼50% of the untreated anchorage, suggesting that the anchorage is weaker, but is certainly not absent. Because cell adhesion is not completely impaired on treated surfaces, the presence of nanofibrils does not inhibit FA formation, but evidently impedes the aggregation of FA proteins. FA’s form on both surfaces, but the spatial inhibition of larger scale integrin clustering on the nanofibrillar surfaces prevents visible stress fiber formation and limits the size of paxillin aggregates. In contrast to the purely chemical alterations explored previously, the mechanism in Figure 11 utilizes a physical separation between the extracellular contact points to control intracellular cytoskeletal development. Ligands adsorbed to portions of the topography inaccessible to the cell cannot participate. (39) Katz, B. Z.; Zamir, E.; Bershadsky, A.; Kam, Z.; Yamada, K. M.; Geiger, B. Physical state of the extracellular matrix regulates the structure and molecular composition of cell-matrix adhesions. Mol. Biol. Cell 2000, 11, 1047-1060. (40) Maheshwari, G.; Brown, G.; Lauffenburger, D. A.; Wells, A.; Griffith, L. G. Cell adhesion and motility depend on nanoscale RGD clustering. J. Cell Sci. 2000, 113, 1677-1686. (41) Houseman, B. T.; Mrksich, M. The microenvironment of immobilized Arg-Gly-Asp peptides is an important determinant of cell adhesion. Biomaterials 2001, 22, 943-955. (42) Neff, J. A.; Tresco, P. A.; Caldwell, K. D. Surface modification for controlled studies of cell-ligand interactions. Biomaterials 1999, 20, 2377-2393. (43) Adams, J. C. Characterization of cell-matrix adhesion requirements for the formation of fascin microspikes. Mol. Biol. Cell 1997, 8, 2345-2363. (44) Burridge, K.; Chrzanowska-Wodnicka, M. Focal adhesions, contractility, and signaling. Annu. ReV. Cell DeV. Biol. 1996, 12, 463-518.
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These observations appear to conflict with those previously observed,45 in which specific nanoscalar topography had no obvious effects on cellular morphology. Both sets of data are in agreement concerning the lack of topographic effects on cell growth kinetics versus “flat” surfaces. Note, however, that in this prior study, the majority of the peaks were only 50 nm in height. Referring to Figure 11, it seems likely that peak heights below a certain value will be insufficient to keep the cell from contacting the “valleys” between them, and stress fiber development will be unaffected. The associated effects of the stable nanofibrils on cytoskeletal morphology are noteworthy both for the magnitude of the effects and the fact that this effect is retained even after 3 days of cellular ECM deposition. Rajagopalan et al.46 reported weak cytoskeletal development and cellular contractility in balb/c 3T3 fibroblasts cultured on RGD-modified substrata. Similar results were seen by Streeter and Rees,47 who reported a lack of stress fiber formation in fibroblasts on GRGDS-coated substrates; TEM showed that the cell made only “point contacts” with the substrate versus more extensive adhesion to FN. We also utilized TEM and observed a very similar network of point contacts (Figure 10), as opposed to the uniform interaction with the non-nanoscaled surface. We can utilize a recent view48 of receptor polymerization as either nucleation or growth to develop a picture of cell response to these different surfaces during the initial stages of adhesion on the other side of the cell membrane from these point contacts. Nucleation involves the aggregation of two or more sets of mobile receptors into a cluster that does not dissociate. Growth describes the addition of either single receptors or small groups into preexisting clusters. Nucleation and adhesion are spatially unfettered on untreated surfaces, and cluster growth eventually dominates. Nanofibrillar surfaces facilitate adhesion and spreading
(see Figure 5E) by maximizing the density of locations (see Figure 7) upon which small individual clusters nucleate to form early focal complexes.49 Limitations to growth occur, as ligands below the plane of interaction defined by the tops of the nanofibrils are not accessible. Conversely, the lifetime of a ligand-receptor complex becomes longer, and cluster diffusion is hindered to the point that FA’s cannot grow to allow the intracellular formation of visible stress fibers. The number of adhesion sites/cell is larger and the cell spreads more effectively.50 At the same time, however, the cluster-surface bonds are smaller, and the cell can more rapidly rearrange/turnover these contacts allowing improved cell motility.51 This provides a reasonable explanation for the ability of a nanoscaled surface to invoke the formation of nanoscaled intracellular and extracellular features. Our observations advance the possibility that the nanoscale can be used to alter cell behavior in a predictable manner on a given substrate without considerable change in chemical composition. At first blush, such ability provides an opportunity to modify tissue engineering scaffolds in a manner that can tailor subsequent levels of cell adhesion, cell shape and phenotype. Although our focus has been on the cytoskeleton, given the burgeoning understanding of paxillin’s central role in growth factor modulation and signal transduction,52 these surface effects are important across a range of biological systems.
(45) Xie, Y.; Sproule, T.; Li, Y.; Powell, H.; Lannutti, J. J.; Kniss, D. A. Nanoscale modifications of PET polymer surfaces via oxygen-plasma discharge yield minimal changes in attachment and growth of mammalian epithelial and mesenchymal cells in vitro. J. Biomed. Mat. Res. 2002, 61, 234-245. (46) Rajagopalan, P.; Marganski, W. A.; Brown, X. Q.; Wong, J. Y. Direct comparison of the spread area, contractility, and migration of balb/c 3T3 fibroblasts adhered to fibronectin- and RGD-modified substrata. Biophys. J. 2004, 87, 28182827. (47) Streeter, H. B.; Rees, D. A. Fibroblast adhesion to RGDS shows novel features compared with fibronectin. J. Cell Biol. 1987, 105, 507-515. (48) Kato, M.; Mrksich, M. Using model substrates to study the dependence of focal adhesion formation on the affinity of integrin-ligand complexes. Biochemistry 2004, 43, 2699-2707.
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Acknowledgment. H.M.P. would like to thank the National Defense Science and Engineering Graduate Fellowship for financial support. The authors would also like to thank the Campus Electron Optics Facility, Campus Microscopy and Imaging Facility, and the Ohio MicroMD Laboratory at the Ohio State University for use of their facilities.
(49) DeMali, K. A.; Wennerberg, K.; Burridge, K. Integrin signaling to the actin cytoskeleton. Curr. Opin. Cell Biol. 2003, 15, 572-582. (50) Gaudet, C.; Marganski, W. A.; Kim, S.; Brown, C. T.; Gunderia, V.; Dembo, M.; Wong, J. Y. Influence of type I collagen surface density on fibroblast spreading, motility, and contractility. Biophys. J. 2003, 85, 3329-3335. (51) Ballestrem, C.; Hinz, B.; Imhof, B. A.; Wehrle-Haller, B. Marching at the front and dragging behind: differential alpha-V beta 3-integrin turnover regulates focal adhesion behavior. J. Cell Biol. 2001, 155, 1319-1332. (52) Brown, M. C.; Turner, C. E. Paxillin: Adapting to change. Physiol. ReV. 2004, 84, 1315-1339.
