Nanoscale Matrix Topography Influences Microscale Cell Motility

Article pubs.acs.org/journal/abseba

Nanoscale Matrix Topography Influences Microscale Cell Motility through Adhesions, Actin Organization, and Cell Shape Samila Nasrollahi, Sriya Banerjee, Beenish Qayum, Parag Banerjee, and Amit Pathak* Department of Mechanical Engineering and Materials Science, Washington University, Saint Louis, Missouri 63130, United States ABSTRACT: Mammalian cells are exposed to complex microenvironments of varying micro- and nanoscale structural features. These multiscale extracellular cues dictate important aspects of cell behavior, including migration, proliferation and differentiation. In this study, we fabricated anodized aluminum oxide (AAO) membranes of either 80 or 40 nm pore diameters. We utilized these membranes as extracellular matrix scaffolds to culture NIH-3T3 fibroblast cells and investigated how the surface nanotopography might regulate their motility. We observed faster and more persistent fibroblast migration on AAO membranes with larger pores. Through various cell−matrix interaction markers, we found that the surfaces with higher nanoporosity enhance motility through larger focal adhesions, aligned actin fibers, and polarized cell morphology. Our findings reveal the importance of nanoscale topographical cues present in the matrix environment in regulating submicrometer-scale subcellular mechanisms of stress fiber organization and adhesion formation, micrometer-scale cell−matrix interactions, and cell motility over hundreds of micrometers. KEYWORDS: nanotopography, AAO membranes, cell migration

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INTRODUCTION Tissues are composed of living cells surrounded by physically diverse microenvironments that vary in stiffness, geometry, and topography, all of which control cell behavior through varying cell−matrix interactions.1 The ultrastructural analysis of the extracellular matrix (ECM) of various tissues, including bone, nerve, cartilage, or blood vessels, has revealed their complex topographic patterns that vary over nano- and microscales.2 In the modern tissue engineering approaches,3−5 the biocompatibility and biological plasticity of the synthetic matrix scaffolds are being enhanced by incorporating micro- and nanostructured features that better mimic the topography of native tissues. Over the past decade, several studies have revealed that matrix size, shape, and organization impact various aspects of cell behavior including cell adhesion, polarity, motility, and differentiation.6−10 Among other topographical cues, surface feature size plays a pivotal role in regulating cell functions. For example, surface roughness beyond a critical value led to reduced cell proliferation.11 The MG63 and 3T3 fibroblasts exhibit bigger adhesions and larger spreading on substrates coated with smaller nanoparticles.12,13 The human mesenchymal stem cells (hMSCs) modify their morphology according to the matrix topography and elongate with the grating features, which in turn influences the fate decisions of hMSCs.14,15 Also, hMSCs cultured on larger TiO2 nanotubes become more elongated and exhibit osteogenic differentiation.16 Fibroblasts cultured on nanoislands of greater height form more filopodial extensions but spread less, compared to those on shorter islands.17 Although the effects of micrometer-scale matrix properties on cell migration have been widely studied, our understanding of © XXXX American Chemical Society

how nanoscale matrix topography impacts cell migration remains limited mainly because of the lack of matrix platforms that allow precise control over surface nanostructure in a reproducible manner. Among the nanostructured synthetic materials developed thus far, the mechanical and material characteristics of AAO membranes make it an ideal material for various applications in the field of tissue engineering.18−21 Briefly, the AAO membranes are developed on an aluminum layer during the anodic oxidation process inside acidic electrolytes.20,22 During fabrication, the size of nanopores is controlled by optimizing the applied voltage. After fabrication, the aluminum layer is separated to obtain self-supporting membranes. Due to their tunable nanotopographical features, high porosity, and biocompatibility, the AAO membranes have been recognized as suitable substrates for cultivation of different cell types, such as osteoblasts, neurons, liver cells, and stem cells.23−27 It has already been established that the pore size of the membrane influences cell proliferation and differentiation. For example, a long-term cultivation of human ostroblast-like (HOB) cells on nanoporous membranes resulted in increased alkaline phosphatase (ALP) activity, which in turn influenced cellular adhesion and differentiation.28 Additionally, AAO membranes have been used for studying the influence of surface porosity on proliferation and migratory behavior of keratinocytes and fibroblast cells.23,29 Special Issue: Multiscale Biological Materials and Systems: Integration of Experiment, Modeling, and Theory Received: September 15, 2016 Accepted: November 4, 2016 Published: November 4, 2016 A

DOI: 10.1021/acsbiomaterials.6b00554 ACS Biomater. Sci. Eng. XXXX, XXX, XXX−XXX

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ACS Biomaterials Science & Engineering

Figure 1. Fabrication of AAO membranes. (A) Top view SEM image of AAO membrane shows hexagonally ordered pores, seen as dark dots in the image. (B) As-grown AAO membranes have a pore diameter of 40 nm, as measured from the diameter of the dark dots. (C) Upon chemical etching in a 5 wt % phosphoric acid solution, the pore diameter increases to 80 nm. (D) An SEM image of the tilted AAO membrane shows the cross-section of the nanopores. extensive wash with PBS. Subsequently, membranes were dried under UV irradiation for 2 h. For single-cell analysis, NIH 3T3 cells were seeded on top of the membranes at cell density of 2500 cells/cm2 and cultured for 3 days. Time-Lapse Microscopy of Live Cells. Live cell imaging was conducted using the Zeiss Cell Observer microscope, equipped with an incubator chamber for controlled temperature, humidity, and CO2 and a motorized, programmable stage. To obtain time-lapse movies, phase contrast images of NIH 3T3 cells were acquired with a 10× objective for at least 36 h at 30 min intervals. For cell migration analysis, individual movies were imported to ImageJ software (National Institutes of Health), and cell migration tracks were extracted using the Manual Tracking plugin. Cell migration tracks were subsequently analyzed to obtain migration speed, directionality, direction autocorrelation (DA), and mean square displacement (MSD). Migration speed was calculated as a ratio between the sum of distances traveled by the cell between each time point and the total time. Directionality was calculated as a ratio of total distance and net displacement traversed by a cell over a given time period. To evaluate the distance traversed by cells in each time interval, we performed MSD analysis.33 By obtaining the position vector on the cell trajectory at time t, MSD is calculated using the following equation, in which brackets denote average over all times and for all N cells.

Taken together, numerous previous studies have now demonstrated that AAO membranes are suitable substrates for culturing a variety of cell types and the nanoscale topographical features of the AAO membranes influence different aspects of cell behavior. However, it remains unknown how the pore size of these membranes regulates the migratory behavior of single cells. Further, it is not yet clear which subcellular mechanisms could mediate the nanotopography-dependent cell polarization and migration. In this study, we therefore aim to investigate how nanoporosity of the AAO membrane influences the behavior of NIH 3T3 fibroblasts on multiple length scales, from cell migration and morphology (micron-scale) to actin fiber alignment and focal adhesion formation (submicron-scale). Here, we fabricated AAO membranes of 40 and 80 nm pore diameters, cultured fibroblasts, and compared their migratory behavior on the two membrane types. We also measured fiber alignment and focal adhesion sizes of these cells to understand the subcellular mechanisms that link the cell’s ability to detect nanoscale matrix cues and accordingly adapt its microscale migratory responses.

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MSD(Δt ) = [x(t + t0) − x(t0)]2 + [y(t + t0) − y(t0)]2

EXPERIMENTAL SECTION

t ,N

We also calculated the DA parameter, which measures the alignment of velocity vectors overtime. DA parameter is defined using the following equation, in which v(t0) is the vector at the starting time t0, and the v(t0 + t) is the vector at time (t0+ t). Brackets indicate that all calculated cosines are averaged for all possible starting times t0 over all cells (N).

Fabrication of AAO Membranes with Different Sizes of Nanopores. The nanoporous AAO membrane was synthesized from 250 μm thick Al foils (Alfa Aesar, 99.999% purity). The Al foil was degreased in ethanol and deionized water. Next, the foil was electropolished using a perchloric acid and ethanol solution (1:5 vol %) at 15 V for 3 min at 4 °C. The electropolished Al foils underwent a 2step anodization process30 in 0.3 M oxalic acid solution at 40 V and 8 °C. The first anodization was run for a period of 6 h. The growth rate of the pores is 72 nm/min. Thus, pores ∼25.9 μm deep were formed. This porous oxide layer was then etched away using a phosphoric acid (6%) and chromic acid (1.8%) solution, leaving behind a textured Al foil which would lead to a well ordered porous oxide in the second anodization step. The second anodization was run for a fixed time of ∼11 h, which yields pores ∼47.5 μm deep. To prepare the membranes, we stripped the oxide layer from one side of the foil using a 3 M sodium hydroxide solution. The remaining Al beneath the oxide layer was then removed using a solution31 of 6 g of copper chloride in 75 mL of 38% hydrochloric acid and 75 mL of deionized water. The as-prepared AAO membranes have a pore diameter of 40 nm. The pore diameters were adjusted to 80 nm by a timed 16 min etch, in 5 wt % phosphoric acid solution,32 maintained at 38 °C using a water bath. Cell Culture on AAO Membranes. NIH 3T3 fibroblasts were cultured in growth media containing low-glucose DMEM (Invitrogen) supplemented with 1% L-glutamine, 1% penicillin/streptomycin, and 10% fetal bovine serum (Invitrogen), and maintained at 37 °C and 5% CO2 during culture. Prior to cell seeding, AAO membranes were sterilized by soaking in 70% ethanol for at least 24 h followed by

DA = (v(t0)v(t0 + t ) t0, N = cos θ(t0 , t0 + t )

t0, N

Immunofluorescence, Confocal Microscopy, and Image Analysis. After at least 2 days of culture on the AAO membranes, cells were fixed with 3.7% paraformaldehyde solution for 10 min, washed with 1× phosphate buffered saline (PBS), and permeabilized with 0.2% Triton X-100 in PBS for 5 min. Cells were blocked using 1% BSA solution prior to treatment with a primary monoclonal mouse paxillin antibody (1:100; Abcam). Before adding the secondary antibody (1:500, goat antimouse Alexa Fluor 647; Invitrogen), phalloidin (1:200; Invitrogen), and DAPI (1:250; Santa Cruz), the primary antibody was removed and rinsed three times with 1X PBS. After removing the secondary antibody solution, samples were extensively washed with 1X PBS and stored in 1X PBS at 4 °C until imaging. Immunofluorescence microscopy was conducted using a laser-scanning confocal microscope (Zeiss LSM 730; Carl Zeiss MicroImaging, Germany). Confocal stacks were acquired with 1 μm intervals at 20X objective. Captured z-stacks were imported to ImageJ (NIH), and the stacks were projected with the maximum intensity. To analyze actin fiber orientation, z-projected images of phalloidin were analyzed using the OrientationJ plugin in ImageJ. For individual cells, the degree of stress fiber alignment was B


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Figure 2. Effect of AAO membrane nanoporosity on migratory behavior of fibroblasts. (A) Migration speed of cells cultured on membranes with 40 and 80 nm pores. (B) Directionality ratio of cells cultured on membranes of different pore sizes; plot indicates more persistent movement for cells cultured on membranes of larger pores. N > 30 cells per condition from at least two separate experiments. Square brackets denote statistical significance. (C) Time lapse series of phase contrast images for cells migrating on membranes with 40 nm (top panel) and 80 nm (bottom panel) pores. Scale bar = 50 μm.

Figure 3. Cells explored a larger space on more porous membranes. Plot indicates single-cell migration trajectories for 10 cells randomly selected among cells seeded on membranes with (A) 80 nm and (B) 40 nm pores. (C) Average MSD analysis over time, which is computed at each time interval by averaging the MSD values for at least 20 cells. (D) Direction Autocorrelation (DA) calculated for every time interval and averaged for the entire cell population selected for analysis. Error bars = SE. N > 30 cells per condition from at least two separate experiments. measured in terms of coherency varying between 0 (isotropic distribution) and 1 (highly aligned fibers). To quantify the size of focal adhesions and their distribution throughout the cells, punctate paxillin sites were manually traced and the size of each adhesion complex was obtained using the Analyze Particle tool in ImageJ. Cell area was

measured using the Shape Descriptors tool in ImageJ, after manually tracing the cell outline. Aspect ratios were defined as the ratio of major axis/minor axis length. Statistical Analysis. Results are reported as the mean ± Standard Error (SE), unless stated otherwise. A Student’s t-test was used to C


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ACS Biomaterials Science & Engineering identify the significant differences between experimental conditions. A p value of 0.05 or smaller was used to identify the statistically significant differences.

additional insight into the mechanism by which nonporous membranes regulate cell migration, we analyzed cell morphology and cytoskeletal structure. First, we measured cell spreading in terms of area and aspect ratio. We found that individual cells occupied larger area and attained a more elongated shape (higher aspect ratio) on membranes with the bigger 80 nm pores, as plotted in Figure 4A, B. Next, we used confocal microscopy to

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RESULTS Fabrication and Characterization of AAO Membranes. The AAO forms cylindrical pores which are hexagonally ordered and have an as-grown pore diameter of 40 nm, as can be seen from Figure 1B (top view SEM image of an AAO membrane is shown in Figure 1A). As stated before, the pores grow 47.5 μm deep, making it a nanostructure with an aspect ratio (pore length/pore diameter) of 1187. The pore density is estimated to be 1010 cm−2. The pore diameters can be increased by isotropic chemical etching of the membrane in a 5 wt % phosphoric acid solution. An AAO membrane with pore diameter of 80 nm is shown in Figure 1C. This structure has an aspect ratio of 594. An SEM image of the tilted AAO membrane shows the partial side view of the pores (Figure 1D). It can be seen that the pores are well-ordered and run parallel to each other. Faster and More Directed Cell Migration on AAO Membranes with Larger Nanopores. To test if the migratory behavior of fibroblast cells responds to the nanotopography of the ECM, we measured migration speed, directionality, mean square displacement (MSD), and direction autocorrelation (DA) for cells grown on AAO membranes of either 40 or 80 nm pore diameters. We allowed the cells to attach to the membrane surfaces for 5 h before capturing time-lapse movies of their movement, as described above. We found that the cells migrated with a speed of ∼0.40 μm/min on membranes with 80 nm pores, which was approximately 60% faster than the cells on membranes with 40 nm pores (Figure 2A). We also evaluated the effect of nanoporosity on the directionality of cellular movement, defined as a ratio of total distance and net displacement traversed by a cell over a given time period. We found that cells on 80 nm pores migrated with a greater directionality, ∼ 0.75, compared to those on the 40 nm pores, ∼ 0.55,(Figure 2B). Thus, on surfaces with larger nanopores, cells not only migrate faster but also exhibit an enhanced directionality during their movement. Next, we plotted migration trajectories of several representative cells (Figure 3A, B) and found that the cells explored a wider space with intersecting paths on the membranes with 80 nm pores. In contrast, the cells on the smaller pores stayed within the vicinity of their original location and did not collide their paths with other cells. We also calculated the mean square displacement, which gives a measure of the area explored by cells for any given time interval, and confirmed that the cells migrating on the more porous membranes explored wider territory (Figure 3C). Importantly, the increased spatial exploration (Figure 3) by the migratory cells on the 80 nm membranes was not only due to their increased speed (Figure 2A), but also to the more persistent and directional movement (Figure 2B). To evaluate directionality of cellular movement over time, we calculated the direction autocorrelation for each time interval, which shows the alignment of velocity vectors over time. As plotted in Figure 3D, the lower DA value for cells on 40 nm membranes indicates that the velocity vector at any time interval is less aligned with the previous velocity vectors, when compared to the cells on more porous surface. Collectively, these findings indicate that the nanoporosity of the AAO membranes fundamentally alters the migratory behavior of cells, with faster and more persistent migration on large pores. Greater Cell Spreading, Elongation, and Stress Fiber Alignment on More Porous Nanomembranes. To gain

Figure 4. Cell morphology and stress fiber alignment on membranes of varying pore diameter. (A) Spreading area and (B) aspect ratio of cells on membranes with 40 and 80 nm wide pores. (C) Representative confocal images demonstrating bipolar, multipolar, and nonpolar morphologies (left to right). Images were obtained by merging images of phalloidin (green) and DAPI (blue). (D) Percent emergence of each morphology phenotype and (E) migration speed associated with each morphology on membranes of different pore diameters. (F) Representative immunofluorescence images of F-actin distribution, stained for phalloidin (green), and nuclei stained with DAPI (blue) in fibroblasts cultured on membranes of larger pores (top) and small pores (bottom). (G) Quantification of actin fiber alignment as a function of membrane pore diameter. Horizontal brackets denote statistical significant difference (p < 0.05). N > 30 cells per condition from at least two separate experiments. Scale bar = 50 μm.

visualize the alignment of actin fibers and cell polarization (Figure 4A,C). We encountered three different types of cell morphologies across both types of membranes: (1) nonpolar cells, with no distinguishable front or rear, (2) bipolar cells, with distinct front and rear regions, or (3) multipolar cells, with more than two distinct poles, as shown in Figure 4C. Here, we define poles as prominent cellular extensions that protrude at least 5 μm outside of the main cell body. On membranes with large pores, the bipolar morphology was the dominant phenotype accounting for more than 70% of the cells, followed by multipolar phenotype accounting only for 25% of cell population. Interestingly, almost D


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Figure 5. Larger focal adhesions on membranes with larger nanopores. (A) Representative immunofluorescence images of paxillin (red) and DAPI (blue) in fibroblasts cultured on membranes of 80 nm (top) and 40 nm (bottom) pores. (B) Scatter plot compares the mean focal adhesion area on membranes of different pore sizes. Histograms of focal adhesion sizes in cells on membranes with (C) 80 nm and (D) 40 nm pores. N > 30 cells per condition from at least two separate experiments. Statistically different pairs (p < 0.05) are indicated by horizontal square brackets. Scale bar = 50 μm.

obtained from the analysis of at least 20 cells (Figure 5C, D) for each membrane type (40 and 80 nm pores). Strikingly, more than 65% of the adhesion complexes on 80 nm membranes were bigger than 8 μm2, whereas only less than 20% of adhesions were that size on the membranes with smaller pores. Thus, single fibroblasts migrating on membranes with bigger nanopores formed larger adhesion complexes.

no cells were found with nonpolar shapes (Figure 4D). However, on membranes with smaller pores, we observed all three types of cell shapes with nonpolar morphology being the dominant one (Figure 4D). We also found that the cells with bipolar morphology migrated faster than those with either multipolar or nonpolar morphologies, regardless of the nanoporosity of membranes (Figure 4E). On the basis of these results, we concluded that increasing the pore diameter caused a bipolar morphology with distinct front and rear of the cell (Figure 4D), which facilitates a more streamlined migration (Figure 4E). By performing confocal microscopy of F-actin expression, we found greater alignment of actin bundles in the cells migrating on the 80 nm membranes (Figure 4F). However, on the membranes with smaller nanopores, actin fibers in the cells were somewhat uniformly distributed with very few discernible actin bundles (Figure 4F). We performed quantitative image analysis of F-actin distribution by calculating a coherency parameter, which revealed that the cells cultured on membranes with larger nanopores had more aligned actin fibers compared to the cells on smaller pores (Figure 4G). Thus, higher stress fiber alignment on substrates with larger nanopores predicts faster cell motility. Nanoporosity of the Surface Influences Focal Adhesions. To understand the effect of nanopore size on focal adhesion complexes, we stained the cells for paxillin, a focal adhesion protein, performed image analysis, and measured the size of the adhesion complexes in cells on both types of membranes (Figure 5). On average, cells formed larger adhesion complexes on membranes with 80 nm pores. In contrast, the adhesion complexes were smaller and more diffused on the 40 nm membranes (Figure 5A). The quantitative analysis of these paxillin images revealed that the average focal adhesion area on 80 nm membrane was ∼7.5 μm2, compared to the smaller average adhesion size of ∼4.5 μm2 on the 40 nm membranes (Figure 5B). To better understand the variation of adhesion sizes across samples, we plotted histograms of focal adhesion sizes

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DISCUSSION The physical interaction of living cells with their extracellular microenvironments of varying micro- and nanoscale features34−43 regulates fundamental cellular responses such as migration, proliferation, and differentiation. Matrices with defined nanometer scale topography are now receiving more attention due to their unique capabilities, including higher surface-to-volume ratio and their greater biological plasticity.24,44 Among synthetic materials with controlled nanotopographical surface features, AAO membranes have been widely used as suitable cell culture substrates for investigating cell proliferation and differentiation. For example, the membranes with smaller pores allow relatively less proliferation in different cell types, including fetal human osteoblasts, epithelial cells, and fibroblasts.23,26,27,29 In general, cells alter their morphology according to matrix topography of varying length scales.38,45 For instance, cellular elongation and polarization of epithelial cells inside micrometer-scale confined regions leads to faster cell migration and malignant transformation of epithelial cells.9,46 Further, it has been shown that the human mesenchymal stem cells exhibit greater osteogenic differentiation and higher cytoskeletal tension on substrates with larger pores.16 In agreement with previous observations of cellular response to nanoscale matrix features, we have found that the 3T3 fibroblasts adopt different morphologies on AAO membranes with different sizes of nanopores. Our measurements show that cells on membranes with larger nanopores undergo greater E


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under Subcontract DE-AC36-08GO28308 to the National Renewable Energy Laboratory, Golden, Colorado) and the Government of India, through the Department of Science and Technology under Subcontract IUSSTF/JCERDC-SERIIUS/ 2012. Electron microscopy facilities were provided by the Institute of Materials Science & Engineering (IMSE) at Washington University in St. Louis.

spreading and attain more elongated and bipolar morphologies. We argue that the membranes with smaller pores (40 nm) present a larger contact area for attachment, compared to those with 80 nm pores, and allow the cells to easily form more dynamic adhesions and protrusions in any direction. Thus, smaller pores lead to multipolar and nonpolar cell shapes that are not ideal for efficient migration (Figure 4E). Because of the extensive availability of attachment areas, cells on these membranes demonstrate less efficient motility with lower directionality and a meandering movement. However, on membranes with larger pores, the reduction in surface area could make it harder for cells to stabilize many protrusions in multiple directions. Thus, it is likely that the cells maintain a bipolar morphology, with front and rear regions, which is the minimum possible number of poles and extensions a welladhered cell must sustain. According to our measurements, this elongated morphology is also associated with more aligned stress fibers and stable focal adhesions. Note that the larger adhesions on more porous membranes do not work against cellular translocation; instead, the cells on these surfaces benefit from the associated stress fiber alignment and cell elongation, both of which are known to aid faster cell motility.46−48 Because of the smaller contact area on more porous membranes, cells might have to work harder and explore larger substrate territory to find stable adhesion points, which in turn might aid their motility.

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(1) Kim, D.-H.; Provenzano, P. P.; Smith, C. L.; Levchenko, A. Matrix nanotopography as a regulator of cell function. J. Cell Biol. 2012, 197 (3), 351−360. (2) Nguyen, A. T.; Sathe, S. R.; Yim, E. K. From nano to micro: topographical scale and its impact on cell adhesion, morphology and contact guidance. J. Phys.: Condens. Matter 2016, 28 (18), 183001. (3) Langer, R.; Tirrell, D. A. T. D., Designing materials for biology and medicine. Nature 2004, 428 (6982), 487−492. (4) Liu, C.; Xia, Z.; Czernuszka, J. T. Design and Development of Three-Dimensional Scaffolds for Tissue Engineering. Chem. Eng. Res. Des. 2007, 85 (7), 1051−1064. (5) Ratner, B. D.; Bryant, S. J. Biomaterials: where we have been and where we are going. Annu. Rev. Biomed. Eng. 2004, 6, 41−75. (6) Di Cio, S.; Gautrot, J. E. Cell sensing of physical properties at the nanoscale: Mechanisms and control of cell adhesion and phenotype. Acta Biomater. 2016, 30, 26−48. (7) Pathak, A.; Kumar, S. Biophysical regulation of tumor cell invasion: moving beyond matrix stiffness. Integr. Biol. 2011, 3 (4), 267−278. (8) Pathak, A.; Kumar, S. Transforming potential and matrix stiffness co-regulate confinement sensitivity of tumor cell migration. Integr. Biol. 2013, 5 (8), 1067−1075. (9) Nasrollahi, S.; Pathak, A. Topographic confinement of epithelial clusters induces epithelial-to-mesenchymal transition in compliant matrices. Sci. Rep. 2016, 6, 18831. (10) Nelson, C. M.; VanDuijn, M. M.; Inman, J. L.; Fletcher, D. A.; Bissell, M. J. Tissue Geometry Determines Sites of Mammary Branching Morphogenesis in Organotypic Cultures. Science 2006, 314 (5797), 298−300. (11) Washburn, N. R.; Yamada, K. M.; Simon, C. G.; Kennedy, S. B.; Amis, E. J. High-throughput investigation of osteoblast response to polymer crystallinity: influence of nanometer-scale roughness on proliferation. Biomaterials 2004, 25 (7−8), 1215−1224. (12) Arnold, M.; Cavalcanti-Adam, E. A.; Glass, R.; Blummel, J.; Eck, W.; Kantlehner, M.; Kessler, H.; Spatz, J. P. Activation of integrin function by nanopatterned adhesive interfaces. ChemPhysChem 2004, 5 (3), 383−8. (13) Goreham, R. V.; Mierczynska, A.; Smith, L. E.; Sedev, R.; Vasilev, K. Small surface nanotopography encourages fibroblast and osteoblast cell adhesion. RSC Adv. 2013, 3 (26), 10309. (14) Dalby, M. J.; Gadegaard, N.; Oreffo, R. O. Harnessing nanotopography and integrin-matrix interactions to influence stem cell fate. Nat. Mater. 2014, 13 (6), 558−69. (15) Wong, S. T.; Teo, S. K.; Park, S.; Chiam, K. H.; Yim, E. K. Anisotropic rigidity sensing on grating topography directs human mesenchymal stem cell elongation. Biomech. Model. Mechanobiol. 2014, 13 (1), 27−39. (16) Oh, S.; Brammer, K. S.; Li, Y. S.; Teng, D.; Engler, A. J.; Chien, S.; Jin, S. Stem cell fate dictated solely by altered nanotube dimension. Proc. Natl. Acad. Sci. U. S. A. 2009, 106 (7), 2130−5. (17) Dalby, M. J.; Riehle, M. O.; Johnstone, H. J.; Affrossman, S.; Curtis, A. S. Polymer-demixed nanotopography: control of fibroblast spreading and proliferation. Tissue Eng. 2002, 8 (6), 1099−108. (18) Kim, Y.; Jung, B.; Lee, H.; Kim, H.; Lee, K.; Park, H. Capacitive humidity sensor design based on anodic aluminum oxide. Sens. Actuators, B 2009, 141 (2), 441−446. (19) Poinern, G. E.; Fawcett, D.; Ng, Y. J.; Ali, N.; Brundavanam, R. K.; Jiang, Z. T. Nanoengineering a biocompatible inorganic scaffold for skin wound healing. J. Biomed. Nanotechnol. 2010, 6 (5), 497−510.

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CONCLUSIONS Our findings demonstrate that subcellular adhesions and cytoskeletal structure are able to detect nanometer scale changes in surface topography and accordingly alter cell morphology and migration−a micron-scale outcome. Despite the vast improvement in the development of materials used for tissue engineering applications, our understanding of the cell−matrix interaction at the nanoscale still remains incomplete largely due to the absence of platforms mimicking the nanostructure of the native ECM. The technique used in this study for the fabrication of AAO membranes enables a precise and reproducible tuning of the surface nanostructure, which increases the potential of providing additional insights into the biophysical regulation of cell migration by nanoscale matrix cues. Such knowledge could inspire the development of novel surface topographies for a desired level of cell motility, which in turn could enable better control over cellular function in the tissue-engineering-based reconstruction of damaged tissues.

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REFERENCES

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Phone: (314) 935-7585. Address: One Brookings Dr., CB 1185, St. Louis, MO 63130. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was in part supported by grants to A.P. from the National Science Foundation (CAREER Award 1454016) and the Edward Mallinckrodt, Jr. Foundation (New Investigator Award). S.B. was fully supported by the US−India Partnership to Advance Clean Energy-Research (PACE-R) for the Solar Energy Research Institute for India and the United States (SERIIUS), funded jointly by the U.S. Department of Energy (Office of Science, Office of Basic Energy Sciences, and Energy Efficiency and Renewable Energy, Solar Energy Technology Program, F


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