Toward a Rational Design of Surface Textures Promoting

Jan 16, 2014 - The safe integration of cardiovascular devices requires the sustainable coverage of their luminal surface by endothelial cells (ECs). T...
4 downloads 12 Views 2MB Size
Download PDF
Letter pubs.acs.org/NanoLett

Toward a Rational Design of Surface Textures Promoting Endothelialization Eva Potthoff,§,⊥ Davide Franco,†,⊥,‡ Valentina D’Alessandro,† Christoph Starck,‡ Volkmar Falk,‡ Tomaso Zambelli,∥ Julia A. Vorholt,§ Dimos Poulikakos,*,† and Aldo Ferrari*,† †

Laboratory of Thermodynamics in Emerging Technologies, Department of Mechanical and Process Engineering, ETH Zurich, Sonneggstrasse 3, CH-8092 Zurich, Switzerland ‡ Clinic of Cardiac and Vascular Surgery, University Hospital Zurich, Raemistrasse 100, 8091 Zurich, Switzerland § Institute of Microbiology, ETH Zurich, Wolfgang-Pauli-Strasse 10, 8093 Zurich, Switzerland ∥ Laboratory of Biosensors and Bioelectronics, Institute for Biomedical Engineering, ETH Zurich, Gloriastrasse 35, 8092 Zurich, Switzerland S Supporting Information *

ABSTRACT: The safe integration of cardiovascular devices requires the sustainable coverage of their luminal surface by endothelial cells (ECs). The engineering of active surface textures has the potential to coordinate cellular adhesion and migration under the action of hemodynamic forces. We define a paradigm to rationally design textures maximizing EC activities as a function of the applied stresses. This is based on harnessing the adhesions established by ECs through fine-tuning of the vertical extend of the underlying surface nanotopography. KEYWORDS: Endothelial cell adhesion, FluidFM, topography, focal adhesion kinase, paxillin, migration

A

onstrates that luminal endothelialization of cardiovascular implants is hampered in regions characterized by disturbed flow or altered material properties.1 This observation indicates that the local hemodynamic stress profile directly affects the stability and behavior of ECs and must be considered when engineering substrates aimed at promoting endothelialization in vivo. The implementation of micrometer or nanoscale surface modifications to known biomaterials is a promising strategy to control the cellular processes, which contribute to the development and maintenance of a functional endothelium.7,8 Substrate topography strongly modulates ECs functions contributing to endothelialization, that is, adhesion and migration, providing a facile and cost-effective approach to the surface engineering of cardiovascular implants.8−10

dverse responses to implants are a major obstacle to the sustainable treatment of cardiovascular diseases.1,2 At the interface between foreign body and blood, inflammatory reactions can be elicited by metallic or polymeric materials. In the case of stents, this may lead to acute thrombosis or chronic stenosis (i.e., the narrowing of the luminal cavity). In mechanical circulatory assist systems, thromboembolism is often caused by thrombus formation at the level of the inflow cannular or within the system.3,4 A fundamental and unmet need is thus to enable a stable interfacing cellular layer both preventing the direct contact between implant and blood and locally regulating tissue homeostasis.1,5 This function in physiological conditions in vivo is ensured by the endothelium, a specialized tissue composed by endothelial cells (ECs).6 Hence, the fast and stable coverage of target substrates by ECs (altogether defined as the process of endothelialization) is considered as the optimal solution to avoid thrombus formation and thromboembolic death in patients with cardiovascular implants.5 Importantly, clinical evidence dem© 2014 American Chemical Society

Received: December 20, 2013 Revised: January 13, 2014 Published: January 16, 2014 1069

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

Antibodies. Rabbit anti-FAK (no. 3285), rabbit antiphospho-FAK (no. 3283), rabbit anti-Paxillin (no. 2542), and rabbit anti phospho-Paxillin (no. 2541) were purchased from Cell Signaling Technology (Beverly, MA). Goat anti-VEC (vascular endothelial cadherin; no. 6458) was purchased from Santa Cruz Biotechnology Inc. (U.S.A.). Mouse antivinculin (V4505) and mouse antitubulin (T5168) were purchased from Sigma Aldrich (U.S.A.). Secondary donkey antirabbit-alexa-555 (A31572), donkey antigoat-alexa-488 (A11055), and antimouse-alexa-488 (A-21202) antibodies were from Invitrogen (Carlsbad, CA). Cell Culture. Human umbilical vein endothelial cells (HUVECs; Invitrogen, U.S.A.) were grown in medium 200PRF supplemented with fetal bovine serum 2% v/v, hydrocortisone 1 μg/mL, human epidermal growth factor 10 ng/mL, basic fibroblast growth factor 3 ng/mL, gentamicin 10 μg/mL, amphotericin 0.25 μg/mL, and heparin 10 μg/mL (all reagents from Invitrogen) and were maintained at 37 °C and 5% CO2. All reported experiments were performed using cells with less than seven passages in vitro. The substrates were then coated with gelatin according to the protocol by Lampugnani et al.23 The substrates were stored at 4 °C until the seeding of the cells. To perform the colocalization analysis and the single cell experiments (Figures 3 and 4) cells were detached from a subconfluent cell culture and resuspended in prewarmed medium. The cells were then counted, diluted to reach a concentration of 6.5 × 103 cells in 1 mL of medium and then seeded onto the substrates. This protocol reproducibly yielded isolated individual cells. For the normal adhesion force and for the cell migration under flow measurements, cells were incubated under cell culture conditions for 45 min with 5chloromethylfluorescein diacetate (CellTracker Green CMFDA, Invitrogen) at the final concentration of 1 μM in serum-free medium. To obtain fully confluent monolayers (Supporting Information Figure 4) cells were seeded on COC substrates at high density (3.5−5 × 104 cell/cm2) and cultured for three days. Adhesion Force Measurements-Sample Preparation. HUVECs were trypsinized in 0.25% trypsin-EDTA (Invitrogen) for 1−2 min to detach the cells from the culture dish. After centrifugation for 7 min at 800 rpm, cells were washed three times with prewarmed filtered (0.22 μm pore size) CO2independent medium (Invitrogen) supplemented with 10% FBS (Invitrogen), 1% pen/strep (Invitrogen), and 2 mM Lglutamine (Sigma-Aldrich) and subsequently seeded onto the desired substrate. The cell concentration was adjusted to obtain sufficiently high numbers of single cells on the desired substrate at the start of the experiment to about 5−10 cells/mm2. Before starting the adhesion strength measurements, cells were allowed to attach and spread on the substrate for 25 min at 37 °C. For contractility inhibition experiments, HUVECs were incubated in CO2-independent medium containing 50 μM blebbistatin (Sigma-Aldrich) dissolved in DMSO for 30 min after the centrifugation step. The adhesion measurements were started after additional 25 min incubation time of the cells on the substrate at 37 °C. Adhesion Force Measurements- Single-Cell Force Spectroscopy (SCFS) Procedures Using FluidFM. A FluidFM24 (Cytosurge AG, Zurich and Nanosurf AG, Liestal, Switzerland) mounted on top of an Axio Observer Z1 inverted microscope (Carl Zeiss, Jena, Germany) was used for the SCFS

However, most reports to date provided only empirical correlations between selected surface geometries and the resulting cell coverage.7,11 A rational approach to fine-tune the interaction at the interface between ECs and structured materials is still missing. EC adhesion to a substrate is mediated by specialized cellular machineries named focal adhesions (FAs).12 The establishment of a FA is initiated by the binding of membrane receptors of the integrin family to their respective ligands on the substrate.13 This early recognition triggers the intracellular recruitment of a number of cytoplasmic proteins yielding the assembly of a hierarchical and ordered protein structure (i.e., the adhesion plaque).14 The adhesion plaque is initially endowed with a signaling function, to which a structural function can be then added by the interaction with the cell cytoskeleton.15 This maturation process is mediated by two pivotal proteins: the focal adhesion kinase (FAK) and paxillin (PAX).16 Early upon integrin binding, FAK is activated through autophosphorylation at specific tyrosine residues.17 Phosphorylated (p)FAK can in turn catalytically activate cytoplasmic PAX. This active form of PAX (pPAX) is able to dock at the adhesion site, corecruiting the adaptor protein vinculin.16 The resulting enrichment of vinculin at the adhesion plaque eventually mediates its interaction with actin filaments.18 Altogether, the interplay between FAK and PAX provides a molecular relay between the signaling and structural function of FAs.16 Importantly, only through mature FAs, ECs can apply force to the substrate in the form of myosin-II mediated contractility of actin filaments.12,16,19,20 This force ultimately regulates the strength of cell-to-substrate interaction. Indeed, substrates allowing the formation of mature FAs promote stable cell adhesion, while the prevalence of immature FAs is typical of migrating cells.21 The engineering strategy presented here aims at generating facile surface structures able to interfere with the maturation of FAs established by ECs. In particular, we show that substrate geometry critically modulates the structural relay between FAK and PAX. On the basis of this paradigm, we define substrates that selectively maximize fundamental cell activities contributing to endothelialization. We demonstrate to this end that the contributions to the migration of the ECs, to the resistance to flow-mediated shear, and to the resistance to normal traction can be selectively and locally maximized by nanometric variations in the vertical surface topography. Materials and Methods. Substrate Fabrication. The gratings were imprinted on 180 μm thick untreated cyclic olefin copolymer (COC) foils (Ibidi, Germany) using nanoimprint lithography (NIL) as previously reported.22 Three molds were fabricated with ridge and groove widths of 1000 nm and ridge height of 100, 400, and 1000 nm. The depth of the grooves was adjusted by tuning the etching time while the side wall steepness was kept constant. The COC substrates were placed on top of the mold and softened by raising the temperature up to 160−180 °C. A pressure of 50 bar was then applied for 10 min before cooling down to 40 °C. Finally, the pressure was released and the mold was detached from the substrate with a scalpel. This procedure generated squared patterns of 1 cm side length. Samples were then treated with oxygen plasma (100 W for 30 s) to increase the hydrophilicity of the surface and to promote cell adhesion. The water static contact angle of COC before treatment was 94.3 ± 0.4° and was reduced to 27.8 ± 1.3° after treatment. The imprinted gratings were systematically characterized by scanning-electron microscopy before cell culturing. 1070

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

(WSS) to individual HUVECs adhering to the different substrates. Briefly the shear stress applied on the cells can be expressed as function of the channel dimensions, medium viscosity, and volumetric flow rate. While channel dimensions and medium properties were fixed in our experimental setup (w = 20 mm, h = 0.3 mm, μ = 8.4 × 10−4 Pa s), the flow rate was controlled using a peristaltic roller pump (Model 66, Harvard Apparatus, U.S.A.) to apply WSS of 6 Pa to the cells. After seeding the cells on the COC substrate, the sample was incubated for 25 min into the flow chamber. Thereafter, the flow chamber was mounted under an inverted wide-field microscope (Nikon, Japan) in order to record the migration of HUVECs under constant flow. Once the flow chamber was connected to the flow system, the pump was activated and a shear stress of 6 Pa was applied to the cells. The flow experiment was performed within an incubated chamber (Life Imaging Services, Switzerland) at 37 °C and 5% CO 2 concentration. The same settings were applied to expose HUVEC monolayers to flow. In this case, cells were fixed after 18 h of constant flow. Wide-Field Microscopy. Wide-field imaging was performed with a 40×, 1.3 NA oil immersion objective (PlanApo, Nikon) using an inverted Nikon-Ti wide-field microscope (Nikon, Japan) equipped with an Orca R-2 CCD camera (Hamamatsu Photonics, Japan). Images of HUVECs stained with TRICTphalloidin, or tubulin, and vinculin or pFAK antibodies were acquired using the TRICT and FITC filter sets. For each sample, 15 individual cells were imaged, collecting threechannels Z-stacks with lateral resolution of 323 nm/px and vertical resolution of 300 nm/image. Live cell imaging of cell migrating under flow was performed within an incubated chamber (Life Imaging Services, Basel, Switzerland) at 37 °C and 5% CO2 concentration. Images were collected with a 20×, 0.45 NA long distance objective (Plan Fluor, Nikon). The total acquisition time was 16−18 h and images were acquired every 15 min. Focal drift during the experiments was eliminated using the microscope PFS autofocus system. At the end of the experiment, the resulting time-lapses were converted into a single 8 bit movie for each imaging field under analysis. Confocal Microscopy. Confocal images of HUVECs interacting with different substrates were collected with a Leica SP2-FCS (Leica, Germany) using a 63×, 1.4 NA, oil immersion objective (Plan-Apo, Leica). Emission from DAPI was excited with the 405 nm wavelength of a Solid State laser and collected in the 450−480 nm optical window. The Alexa488 emission was excited with the 488 nm wavelength of a HeNe laser and collected in the 500−540 nm optical window. Finally, the Alexa555 emission was excited with the 555 nm wavelength of a HeNe laser and collected in the 600− 800 nm optical window. For each sample, 15 individual cells were imaged, collecting three-channels with lateral resolution of 116 nm/px. Image Analysis. The value of Pearson’s coefficient was extracted from each confocal image stack using the colocalization plug-in of Imaris (Bitplane, Switzerland). During the colocalization analysis, calculated threshold values6 were imposed for both the pFAK or pPAX and the vinculin channels. Cell migration tracks were obtained using the “Manual Tracking” plug-in of ImageJ. Each track was analyzed using MATLAB to extract the counterflow, streamwise, and stationary components. A counterflow migration step was detected when within two subsequent frames (ΔT = 15

measurements applying an adapted protocol from Potthoff et al.25 Rectangular, tipless, 200 μm long and 36 μm wide silicon nitride probes with an 8 μm aperture were chosen for the study of HUVECs25 (Cytosurge AG). Before each experiment, cantilever sensitivity and spring constant were calibrated using software-implemented scripts based on the formalism described by Sader et al.26 Cantilevers exhibit a spring constant of about 1.8 N/m. Prior to the deposition of an antifouling coating, cantilevers were plasma-cleaned at 18 W for 30 s (Plasma Cleaner PDC-32G, Harrick Plasma, U.S.A.). Activated cantilevers together with 0.5 mL Sigmacote (Sigma-Aldrich) were placed in a vacuum desiccator (Duran group, Germany) overnight in order to deposit a hydrophobic film on the cantilevers, minimizing protein adsorption. The combination with a 100 μm piezoelectric Z-stage fixed to the optical microscope allowed for mammalian cell experiments requiring a long detachment distance. Force measurements were recorded at 37 °C within an incubation chamber. A selected HUVEC was approached with a set point of 50 nN, followed by a pause of 5 s (at constant height) during which an underpressure of 700 mbar was applied to grasp the cell. The probe was then retracted at a given piezo velocity, maintaining the underpressure while recording the deflection signal (i.e., the force). An approach and retraction speed of 1 μm/s was applied over a pulling range of 80 μm. After an optical control where the cell was detached from the substrate, an overpressure pulse was applied to expel the cell such that the following cell could be optically targeted and approached. In the case that a cell was sticking to the cantilever after an SCFS, the cantilever was washed in 0.25% trypsin-EDTA (Invitrogen) for a few minutes and then washed in PBS, 0.2 g of KCl, 1.44 g of Na2HPO4, and 0.24 g of KH2PO4 (all from Fluka) in 1 L of distilled water (pH 7.4) with 1000 mbar overpressure applied through the cantilever before it was used for the next cell. One force−distance (F−d) curve was recorded per each measured cell, while 16−38 cells per substrate within at least three independent biological replicates were measured. Representative example curves are shown in Supporting Information Figure 2. Adhesion forces were extracted from the F−d curves using a SPIP 6.0.10 version (Image Metrology, Denmark). Immunostaining. HUVECs were fixed and permeabilized for 3 min with 3% paraformaldehyde (PFA) and 0.5% TritonX100 in PBS at room temperature (RT). The cells were then postfixed with 3% PFA in PBS for 15 min. After washing the samples three times for 5 min with PBS, they were incubated with 5% bovine serum albumin (BSA) for 1 h at RT. The samples were then incubated with antivinculin primary antibody together with antiphospho-FAK or antiphosphoPAX primary antibody overnight at 4 °C. Alternatively, the samples were incubated with antivinculin or antiphospho-FAK primary antibody together with TRITC-phalloidin (Sigma, U.S.A.), with antitubulin together with antiphospho-FAK primary antibodies, or with anti-VEC primary antibody overnight at 4 °C. Subsequently, the samples were rinsed four times for 1 h with 5% BSA in PBS and then incubated at room temperature with antimouse-alexa-488 secondary antibody and donkey antirabbit-alexa-555 antibodies for 45 min at RT. Finally, the samples were washed three times (1 h each) in PBS, postfixed for 2 min in 3% PFA, briefly washed again with PBS, mounted with DAPI-containing Vectashield (Vector Laboratories Inc., U.S.A.) and immediately imaged. Cell Migration under Flow. A custom designed parallel plate flow chamber was used to apply a constant wall shear stress 1071

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

Figure 1. Focal adhesions maturation profile on substrates with different topographies. (a) Distribution of pFAK (red channel; upper row), vinculin (green channel; lower row), and DAPI (nuclei, blue channel; lower row) fluorescent signal on FLAT substrates or gratings with increasing groove depth and (b) distribution of pPAX (red channel; upper row), vinculin (green channel; lower row) and DAPI (nuclei, blue channel; lower row) fluorescent signal on FLAT substrates and gratings with increasing groove depth. The maximum projection of confocal Z-stacks are reported. The direction of the gratings is indicated in the lower left corner of each panel (lower row). Scale bars correspond to 10 μm. The white boxes in the red channel (pFAK and pPAX in panels a and b, respectively) identify regions of interests at the cell edge reported as magnified, inverted view in the middle panel. (c) Biochemical analysis of FAK and Paxillin phosphorylation in HUVECs on substrates with different topographies. The panels show the Western Blots of total and activated FAK (tFAK and pFAK, respectively) and Paxillin (tPAX and pPAX, respectively) protein levels of cells directly harvested 25 min after seeding. (d,e) Western blot quantification of independent experiments. The signal ratio between the phosphorylated 1072

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

Figure 1. continued and total protein forms is reported in panel d. The signal ratio between the total protein form and a loading control (tubulin) is reported in panel e. The horizontal bars indicate the differences between the HUVECs in suspension (Susp.) or interacting with gratings with respect to the FLAT substrate. Error bars represent the measured s.e.m. Significant differences between the population means are reported.

Figure 2. Maximal normal adhesion force of HUVECs on substrates with different topographies measured by FluidFM. (a) Schematic view of the experimental principle showing the initial step of a backward single-cell force spectroscopy by retracting the hollow cantilever after cell targeting and immobilization on the cantilever. The blue arrows indicate the underpressure applied in the channel for cell immobilization. (b) Comparison of the maximal normal adhesion force of HUVECs adhering to FLAT substrates, gratings with increasing groove depth (100, 400, and 1000 nm), or HUVECs treated with Blebbistatin (Blebb). The histograms report the mean measured value. Error bars correspond to the standard error of the mean. The total number of measured cells is reported in the upper right corner. Significant differences between the population means are reported (** indicate p < 0.01).

min), the cell moved 3 μm or more in the direction opposite to flow. A streamwise migration step was counted when displacement of more than 3 μm was measured in the direction of flow. Cell movements of less than 3 μm were defined as stationary. Thus, each time point was associated with a specific migration behavior as described above. Finally, the average permanence time (i.e., the number of subsequent frames) in each state was calculated and plotted (Figure 4). Western Blot. The Western blot analysis was performed as previously reported.9 Briefly, nine COC microstructured substrates were assembled within an incubation chamber casted in polydimethylsiloxane (PDMS), yielding a total surface of approximately 9 cm2. After sterilization with ethanol, freshly detached HUVECs (120 000) were seeded in the PDMS chamber. After 25 min of incubation, the medium was removed and lysis buffer supplemented with a phosphatase inhibitor cocktail (Sigma Aldrich) was directly added to the cells. Lysed cells were then collected by scraping. Cell lysates were analyzed by standard Western blot using antibodies against total FAK and PAX and their phosphorylated isoforms (in position Try397 and Try118 respectively). As loading control, an antibody against tubulin was used. The bands intensity was quantified with the “Gel Analysis” tool of ImageJ. Statistical Analysis. Statistical comparison of the force measurements, Pearson’s coefficients, and migration contribution were performed using a nonparametric Mann−Whitney test (α = 0.05). All quantitative measurements reported are expressed as average values ± the standard error of the mean. The total number of events counted is reported in the upper right corner of the presented graphs. Results. Focal Adhesion Maturation on Structured Substrates. It has been shown that ECs respond maximally to anisotropic topographies (gratings) with lateral feature size of 1000 nm.9,11 The contribution of vertical topographical

features depends of the ability of cells to interact and limit their contacts to the top of the ridges (thus bridging over grooves).9 We tested a set of topographies with identical lateral feature size (ridges and grooves of 1000 nm) and increasing groove depth (flat control, 100, 400, and 1000 nm). This vertical range was chosen since mammalian cells do not sense grooves shallower than 50 nm,27 while on the other end the bridging effect on topography is saturated on grooves deeper than 1000 nm.9 To identify the effect of topography on FA maturation we determined the localization and expression levels of FAK and its active phosphorylated (Tyr397-pFAK) form, of PAX and its active phosphorylated (Tyr118-pPAX) form, of vinculin, and of the cell cytoskeleton16 on grooved substrates or on chemically identical flat substrates (Figure 1 and Supporting Information Figure 1). pFAK, pPAX, and vinculin-rich adhesions were visualized at the cell-substrate interface of ECs. FAs were distributed along the cell periphery (Figure 1a,b) on all tested substrates. Specifically, FAs established on flat substrates appeared to be immature and of reduced size (Figure 1a). Accordingly, the pFAK and pPAX (Figure 1a,b) staining exhibited low signal at the adhesion points while vinculin showed a diffuse localization in the cytoplasm. The contact with gratings increased the concentration of pFAK at adhesions that appeared larger and well oriented. These effects became prominent with increasing groove depth with adhesions established on 1000 nm deep gratings showing larger size and higher content of pFAK. Interestingly, the effect of topography on FA maturation, evaluated in terms of pPAX and vinculin enrichment, did not correlate with the activation of FAK. Adhesions established on shallow gratings (100 nm) displayed the highest pPAX content and a corresponding strong localization of vinculin and actin at the adhesion sites (Figure 1b and Supporting Information 1073

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

Figure 3. Focal adhesion maturation profile on different topographies under directional flow. (a) Distribution of pFAK (red channel; upper row), vinculin (Vin, green channel; middle row) and DAPI (nuclei, blue channel; middle row) fluorescent signal and the corresponding colocalization signal (pFAK/Vin; lower row) on FLAT substrates or gratings with increasing groove depth. The maximum projection of confocal Z-stacks are reported. The direction of the gratings is indicated in the lower left corner of each panel (lower row). Scale bars correspond to 10 μm. The blue arrows indicate the direction of the flow. The white boxes in the colocalization channel identify regions of interests reported as (b) magnified view of 1074

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

Figure 3. continued colocalization signal (pFAK/Vin) in the front (upstream) and rear (downstream) regions of the cells. (c) The Pearson’s correlation coefficient (pFAK/Vin) is shown as a function of vertical feature size under both static and directional flow conditions. Significant differences between the population means are reported (* for p < 0.05, ** for p < 0.01). The connecting lines are intended as a guide to the eye. (d) Distribution of pPAX (red channel; upper row), vinculin (green channel; lower row) and DAPI (nuclei, blue channel; lower row) fluorescent signal on FLAT substrates and gratings with increasing groove depth. The maximum projections of confocal Z-stacks are reported. The direction of the gratings is indicated in the lower left corner of each panel (lower row). Scale bars correspond to 10 μm. The white boxes in the colocalization channel identify regions of interests reported as (e) magnified view of colocalization signal (pPAX/Vin) in the front (upstream) and rear (downstream) regions of the cells. (f) The Pearson’s correlation coefficient (pPAX/Vin) are shown as a function of vertical feature size under both static and directional flow conditions. Significant differences between the population means are reported (** for p < 0.01). The connecting lines are intended as a guide to the eye.

adhesion force were measured for ECs adhering to deeper gratings (598 ± 123 nN on 1000 nm deep gratings). Depending on the grating depth, the cell contact area changed slightly. Apparently, the normal adhesion force and the contact area did not correlate, as shown in previous studies.30 In order to test whether the measured mean adhesion force increase on shallow gratings depended on cell-mediated contractility, cells were pretreated with Blebbistatin. Treatment with the myosin-II inhibitor, reduced the adhesion force to 50% of control levels (295 ± 44 nN). These results demonstrate that the contact to shallow gratings yielded a significant increase in the resistance to vertical traction and thus normal adhesion force is maximized on topographies allowing the establishment of mature FAs (Figure 1). Additionally they demonstrate that this effect depends on cell-mediated contractility. Effect of WSS of Focal Adhesion Maturation. Next we evaluated the effect of flow-mediated walls shear stress (WSS) on the maturation of FAs established by ECs. To this end, we used a custom-designed flow bioreactor and exposed ECs to a steady state flow resulting in a constant WSS of 6 Pa.31 In these experiments, gratings were oriented parallel to the direction of flow. The distribution of pFAK, pPAX, and vinculin (Figure 3) before and after flow-conditioning describes the changes in the FA maturation profile. The size and intensity of pFAK clusters at the adhesion sites (along the cell edge) significantly increased in ECs contacting gratings (Figure 3a), while no evident changes could be observed in cells interacting with flat substrates (Figure 3a). The corresponding recruitment of vinculin to FAs can be appreciated in the colocalization channel (Figure 3a,b). In particular, adhesions established at the upstream edge of cells on gratings displayed higher colocalization signals (Figure 3b) than the adhesions formed at the downstream edge indicating that cells polarized in the direction opposite to the flow. An inverse polarization was typical of cells interacting with flat substrates, which showed a higher number of small pFAK/pPAX/vinculin adhesions at the rear edge. The quantitative measurement of the colocalization signal before and after exposure to flow supports the above observations (Figure 3c). Upon WSS stimulation, pFAK and vinculin colocalization significantly increased in ECs interacting with 100 and 400 nm deep gratings. No significant changes were detected on flat substrates, while cells interacting with 1000 nm deep gratings displayed a reduced colocalization indicating an overall decrease in FA maturation (Figure 3b). Congruent results were obtained when visualizing the distribution and colocalization of pPAX and vinculin. On gratings, the adhesions formed at the front edge were larger and showed a more intense colocalization than at the rear edge, while an opposite polarization was detected in most of ECs interacting with flat substrates (Figure 3d,e). Colocalization

Figure 1). On deeper gratings pPAX and vinculin localization at FAs was less efficient, scaling down to control levels on 1000 nm deep gratings (Figure 1b and Supporting Information Figure 1). A quantitative evaluation of the expression levels of FAK and PAX and of their active, phosphorylated forms was obtained through Western blot analysis (Figure 1c−e). The blot quantification confirmed a marked increase of the pFAK expression in cells adhering to 1000 nm deep gratings while no significant variations from control were detected on shallower substrates (Figure 1d). pPAX levels showed a different trend with a small expression increase on 400 nm deep gratings and a similar decrease on 1000 nm deep ones. As expected, cells maintained in suspension, thus not establishing FAs, showed a clear reduction of pFAK levels. Finally, the total FAK levels were mostly comparable on all tested substrates with only a detectable increase on 100 nm deep gratings. On the other hand, the total PAX level was strongly reduced in cells in suspension and increased in cells contacting 1000 nm deep gratings. This observation suggests a fast turnover of PAX (Figure 1e).28 Taken together, these results indicate that substrate topography variations have a significant impact on the FAs maturation profile. In particular, adhesions established on 1000 nm deep gratings are characterized by a stronger signaling activity mediated by pFAK, while an increase in structural components (pPAX, vinculin, and actin) was observed in adhesion formed on 100 and 400 nm deep gratings. ECs Resistance to Vertical Traction. In order to evaluate EC resistance to normally applied tractive force we measured the maximal normal adhesion force in individual cells adhering to flat substrates or gratings (Figure 2 and Supporting Information Figure 2). For this, we applied FluidFM-based single cell force spectroscopy (SCFS) according to the protocol established in ref 25 (Figure 2a). With conventional AFM only the adhesion of round cells already immobilized on the cantilever can be investigated during the initial stage of the cell-substrate interaction.29 The FluidFM approach, however, allowed measuring forces with which cells adhered after growth on the gratings until they were fully spread without chemical or mechanical perturbations, expanding the range of measurable forces. It also enabled serial SCFS measurements with the same cantilever and thus allowed one to obtain statistically relevant data by measuring 16 or more cells per substrate. The mean adhesion force measured for ECs contacting control flat substrates was 619 ± 70 nN, close to the force measured for other cell types on similar substrates.25 Importantly, cells adhering to shallow gratings were able to resist forces which were 1.8 fold (1113 ± 86 nN) and 1.4 fold (860 ± 59 nN) higher (on 100 and 400 nm deep gratings; respectively, Figure 2b). No changes in the maximal normal 1075

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

Figure 4. HUVECs migration under directional flow. (a) Tracks extracted from time-lapses (for a total of 18 h) of HUVECs migrating on FLAT substrates or gratings under steady state flow yielding a shear stress of 6 Pa. The blue line in the upper left corner indicates the direction of the flow. The orientation of gratings is reported in the lower left corner. The total number of measured tracks is reported in the upper right corner of each panel. (b−d) The histograms report the average contributions to migration on the different substrates. In particular, the fraction of counterflow (b), the stationary (c), and the streamwise movement events (d) are reported. Significant differences between the population means are reported (* for p < 0.05, ** for p < 0.01). The total number of counted events is indicated in the upper right corner of each panel.

analysis (Figure 3f) showed that flow increased the overall colocalization between pPAX and vinculin on flat substrates. A similar change was measured on 100 nm deep gratings, while the maximal increase was detected on 400 nm deep gratings (Figure 3f). Interestingly, no colocalization changes were measured in ECs interacting with 1000 nm deep gratings. Altogether these data indicate that upon exposure to flow, cells on gratings acquired a typical polarization in the direction opposite to flow. Additionally, the WSS induced an overall maturation of FAs on 100 and particularly on 400 nm deep gratings while the maturation of adhesions established on 1000 nm deep gratings was reduced. Effect of WSS of ECs Migration. In order to link the FA maturation changes induced by WSS, to the resulting EC behavior we acquired long-term time-lapse movies of individual cells under flow (Supporting Information Movie 1). The resulting cell migrations tracks are reported in Figure 4a. On flat substrates a large fraction of cell movement was directed along the flow while only a small migration component in the counterflow direction was measured (Figure 4b,d). This yielded an overall streamwise movement of cells (52% of cells; Figure 4a) while no cell displayed a net counterflow migration. Counterflow migration was significantly increased on all gratings with the highest increase detected on 1000 nm deep gratings resulting in an overall cell movement against the flow in 27% of the cells (Figure 4b). Static resistance to flow (i.e., no migration) was similarly enhanced on gratings with highest

stationary behavior on 400 nm deep gratings (Figure 4c). Here, the large majority of cells (80%) did not move significantly during the recording. In all, these data demonstrate that the contact with gratings reinforced the resistance to flow and enabled counterflow migration with maximal effects detected on 400 nm deep and 1000 nm deep gratings, respectively. Discussion. The set of data presented in this work defines a new model for the interaction between substrate topography and ECs. Endothelial adhesion is modulated by the vertical size of topographical features in the range between 100 and 1000 nm. While deep gratings (1000 nm) promote the activation of focal adhesion kinase, thus enhancing its signaling activity (Figures 1 and 5), shallower gratings (100 and 400 nm) act as to reinforce the relay to structural components, through the activation of paxillin, causing the recruitment of vinculin and actin (Figures 1 and 5 and Supporting Information Figure 1) at the adhesion sites. This effect on the focal adhesion maturation profile confers specific and substrate-dependent properties to endothelial cells, which are demonstrated upon exposure to externally applied stresses (Figures 2 and 4). The application of tractive stimuli demonstrates that the interaction with shallow gratings (100 and 400 nm deep; Figure 2) dramatically improves the maximal normal adhesion force of endothelial cells. This effect can be partially explained by an enhanced structural maturation of focal adhesions (Figure 1). A cell interacting with a substrate exerts both in-plane (shear) and 1076

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

Figure 5. Model of topographic control of cell adhesion forces. (a) Relay between signaling and structural components in focal adhesions as a function of the substrate topography. (b) Summary of endothelial cell performance in terms of maximal vertical adhesion force (solid black squares), resistance to flow (solid red circles), and migration in the direction opposite to flow (solid blue triangles) as a function of the substrate topography. Results are taken from the experiments reported in Figures 2 and 4 and plotted as normalized values. The connecting lines are intended as a guide to the eye.

When cells are exposed to flow-mediated shear, the substratedependent changes in the focal adhesion profile (Figure 3) are linked to the appearance of specific migration phenotypes (Figure 4). Here, the interaction with topography enables cells to polarize and migrate in the direction opposite to the flow (Figure 4). This behavior is maximized on substrates triggering focal adhesion kinase activity at the adhesion sites (Figures 1, 3, and 4). The resulting signaling is essential for the generation and maintenance of membrane protrusions that drive cell polarization upon spreading and migration.8,35 In this scenario, the interaction with deep gratings may result in the establishment of adhesions that provide sufficient resistance to shear while maintaining a dynamic state typical of migrating cells.21 This effect may prove beneficial in fully differentiated endothelia exposed to shear, allowing the maintenance of a confluent monolayer under flow (Supporting Information Figure 4). Consistently, when the structural component of focal adhesions is reinforced by the interaction with shallower

out-of-plane (normal) tractions, which are associated with the focal adhesions.32,33 When adhering to shallow topographies, cells contact both the top of ridges and the bottom of grooves thus adapting their membrane to the underlying structures.9 Here, focal adhesions are preferentially formed along the ridge edges thus in the regions of highest membrane curvature.9 On deeper gratings (800 nm or deeper), cells contact exclusively the top of ridges, and membrane arcs bridge over grooves.8,34 Interestingly, the normal adhesion force diminishes as a function of the groove depth (Figure 2). This exponential decay correlates well with the cell groove-bridging efficiency (Supporting Information Figure 3). On the basis of these observations and the lack of correlation between the normal adhesion force and the contact area we speculate that membrane curvature, and the resulting spatial confinement of focal adhesions, contributes to reinforce the normal component of acto-myosin traction exerted by endothelial cells on the substrate (Figure 2). 1077

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

(7) Dreier, B.; Gasiorowski, J. Z.; Morgan, J. T.; Nealey, P. F.; Russell, P.; Murphy, C. J. Am. J. Physiol. 2013, 305 (3), C290−8. (8) Franco, D.; Milde, F.; Klingauf, M.; Orsenigo, F.; Dejana, E.; Poulikakos, D.; Cecchini, M.; Koumoutsakos, P.; Ferrari, A.; Kurtcuoglu, V. Biomaterials 2013, 34 (5), 1488−97. (9) Franco, D.; Klingauf, M.; Bednarzik, M.; Cecchini, M.; Kurtcuoglu, V.; Gobrecht, J.; Poulikakos, D.; Ferrari, A. Soft Matter 2011, 7 (16), 7313−7324. (10) Liliensiek, S. J.; Wood, J. A.; Yong, J.; Auerbach, R.; Nealey, P. F.; Murphy, C. J. Biomaterials 2010, 31 (20), 5418−26. (11) Biela, S. A.; Su, Y.; Spatz, J. P.; Kemkemer, R. Acta Biomater. 2009, 5 (7), 2460−6. (12) Spatz, J. P.; Geiger, B. Methods Cell Biol. 2007, 83, 89−111. (13) Zaidel-Bar, R.; Itzkovitz, S.; Ma’ayan, A.; Iyengar, R.; Geiger, B. Nat. Cell Biol. 2007, 9 (8), 858−67. (14) Kanchanawong, P.; Shtengel, G.; Pasapera, A. M.; Ramko, E. B.; Davidson, M. W.; Hess, H. F.; Waterman, C. M. Nature 2010, 468 (7323), 580−4. (15) Cohen, M.; Joester, D.; Geiger, B.; Addadi, L. ChemBioChem 2004, 5 (10), 1393−9. (16) Pasapera, A. M.; Schneider, I. C.; Rericha, E.; Schlaepfer, D. D.; Waterman, C. M. J. Cell Biol. 2010, 188 (6), 877−90. (17) Mitra, S. K.; Hanson, D. A.; Schlaepfer, D. D. Nat. Rev. Mol. Cell Biol. 2005, 6 (1), 56−68. (18) Humphries, J. D.; Wang, P.; Streuli, C.; Geiger, B.; Humphries, M. J.; Ballestrem, C. J. Cell Biol. 2007, 179 (5), 1043−57. (19) Balaban, N. Q.; Schwarz, U. S.; Riveline, D.; Goichberg, P.; Tzur, G.; Sabanay, I.; Mahalu, D.; Safran, S.; Bershadsky, A.; Addadi, L.; Geiger, B. Nat. Cell Biol. 2001, 3 (5), 466−72. (20) Zimerman, B.; Arnold, M.; Ulmer, J.; Blummel, J.; Besser, A.; Spatz, J. P.; Geiger, B. IEE Proc.: Nanobiotechnol. 2004, 151 (2), 62−6. (21) Cavalcanti-Adam, E. A.; Volberg, T.; Micoulet, A.; Kessler, H.; Geiger, B.; Spatz, J. P. Biophys. J. 2007, 92 (8), 2964−74. (22) Ferrari, A.; Cecchini, M.; Dhawan, A.; Micera, S.; Tonazzini, I.; Stabile, R.; Pisignano, D.; Beltram, F. Nano Lett. 2011, 11 (2), 505− 11. (23) Lampugnani, M. G.; Corada, M.; Andriopoulou, P.; Esser, S.; Risau, W.; Dejana, E. J. Cell Sci. 1997, 110 (Pt 17), 2065−77. (24) Meister, A.; Gabi, M.; Behr, P.; Studer, P.; Voros, J.; Niedermann, P.; Bitterli, J.; Polesel-Maris, J.; Liley, M.; Heinzelmann, H.; Zambelli, T. Nano Lett. 2009, 9 (6), 2501−7. (25) Potthoff, E.; Guillaume-Gentil, O.; Ossola, D.; Polesel-Maris, J.; LeibundGut-Landmann, S.; Zambelli, T.; Vorholt, J. A. PloS One 2012, 7 (12), e52712. (26) Sader, J. E.; Sanelli, J. A.; Adamson, B. D.; Monty, J. P.; Wei, X.; Crawford, S. A.; Friend, J. R.; Marusic, I.; Mulvaney, P.; Bieske, E. J. Rev. Sci. Instrum. 2012, 83 (10), 103705. (27) Loesberg, W. A.; te Riet, J.; van Delft, F. C.; Schon, P.; Figdor, C. G.; Speller, S.; van Loon, J. J.; Walboomers, X. F.; Jansen, J. A. Biomaterials 2007, 28 (27), 3944−51. (28) Abou Zeid, N.; Valles, A. M.; Boyer, B. Cell Commun. Signaling 2006, 4, 8. (29) Hosseini, B. H.; Louban, I.; Djandji, D.; Wabnitz, G. H.; Deeg, J.; Bulbuc, N.; Samstag, Y.; Gunzer, M.; Spatz, J. P.; Hammerling, G. J. Proc. Natl. Acad. Sci. U.S.A. 2009, 106 (42), 17852−7. (30) Dao, L.; Gonnermann, C.; Franz, C. M. J. Mol. Recognit. 2013, 26 (11), 578−89. (31) Orsenigo, F.; Giampietro, C.; Ferrari, A.; Corada, M.; Galaup, A.; Sigismund, S.; Ristagno, G.; Maddaluno, L.; Koh, G. Y.; Franco, D.; Kurtcuoglu, V.; Poulikakos, D.; Baluk, P.; McDonald, D.; Grazia Lampugnani, M.; Dejana, E. Nat. Comm. 2012, 3, 1208. (32) Legant, W. R.; Choi, C. K.; Miller, J. S.; Shao, L.; Gao, L.; Betzig, E.; Chen, C. S. Proc. Natl. Acad. Sci. U.S.A. 2013, 110 (3), 881− 6. (33) Maskarinec, S. A.; Franck, C.; Tirrell, D. A.; Ravichandran, G. Proc. Natl. Acad. Sci. U.S.A. 2009, 106 (52), 22108−13. (34) Rossier, O. M.; Gauthier, N.; Biais, N.; Vonnegut, W.; Fardin, M. A.; Avigan, P.; Heller, E. R.; Mathur, A.; Ghassemi, S.; Koeckert, M. S.; Hone, J. C.; Sheetz, M. P. EMBO J. 2010, 29 (6), 1055−68.

gratings (Figure 3), endothelial cells acquire a stronger stationary behavior, which correlates with stronger normal adhesion force (Figures 2 and 4). In this configuration, the coupling to the cytoskeleton (Figures 1 and 3 and Supporting Information Figure 1) may stabilize focal adhesions thus improving adhesion strength and demoting cell migration. Taken together, the results of the model described here define a new paradigm for the development of surface modifications for cardiovascular implants. When aiming at obtaining a stable endothelial coating at the interface with the bloodstream, the local variations in the hemodynamic stress profile must be considered.36 Our results imply that, owing to the local differences in the stresses generated by the blood flow, a single, uniform surface topography may prove insufficient to induce a complete and functional coating by endothelial cells. Therefore, topographies with a range of overlaid feature sizes may be preferable. The contact between endothelial cells and basal substrates featuring vertical topographies with multiple sizes (in the range between 100 and 1000 nm), tailored according to a map of hemodynamic stresses at the implant luminal surface, is bound to yield a combination of maximal normal adhesion strength, resistance to shear, and counterflow migration.

■

ASSOCIATED CONTENT

S Supporting Information *

Further details regarding the phenotype of endothelial cells interacting with the substrates, the force measurements, and the effect on cell monolayers are provided. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. Author Contributions ⊥

E.P. and D.F. equally contributed to this work.

Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This work is part of the Zurich Heart Project and supported by Hochschulmedizin Zürich and by a grant of the Mäxi Foundation, by the Swiss innovation promotion agency KTICTI (11722.1 PFNM-NM) (to T.Z. and J.A.V.), as well as by the European Union Seventh Framework Programme (FP7/ 2007-2013) under Grant Agreement NMP4-LA-2009-229289 NanoII and Grant Agreement NMP3-SL-2009-229294 NanoCARD. The authors gratefully acknowledge Dr. Gaia Restivo for her technical support.

■

REFERENCES

(1) Otsuka, F.; Finn, A. V.; Yazdani, S. K.; Nakano, M.; Kolodgie, F. D.; Virmani, R. Nat. Rev. Cardiol. 2012, 9 (8), 439−53. (2) Patel, H. J.; Su, S. H.; Patterson, C.; Nguyen, K. T. Biotechnol. Prog. 2006, 22 (1), 38−44. (3) Inoue, T.; Croce, K.; Morooka, T.; Sakuma, M.; Node, K.; Simon, D. I. JACC Cardiovasc. Interv. 2011, 4 (10), 1057−66. (4) Komatsu, R.; Ueda, M.; Naruko, T.; Kojima, A.; Becker, A. E. Circulation 1998, 98 (3), 224−33. (5) Baiguera, S.; Ribatti, D. Angiogenesis 2013, 16 (1), 1−14. (6) Dejana, E. Nat. Rev. Mol. Cell Biol. 2004, 5 (4), 261−70. 1078

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079

Nano Letters

Letter

(35) Zhang, X.; Jiang, G.; Cai, Y.; Monkley, S. J.; Critchley, D. R.; Sheetz, M. P. Nat. Cell Biol. 2008, 10 (9), 1062−8. (36) Rikhtegar, F.; Pacheco, F.; Wyss, C.; Stok, K. S.; Ge, H.; Choo, R. J.; Ferrari, A.; Poulikakos, D.; Muller, R.; Kurtcuoglu, V. PloS One 2013, 8 (3), e58147.

1079

dx.doi.org/10.1021/nl4047398 | Nano Lett. 2014, 14, 1069−1079