Narrow Window in Nanoscale Dependent Activation of Endothelial

To assess the role of residual fluoride content in amorphous nanotubes in cell behavior including cell adhesion and proliferation, MSC were plated on ...
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NANO LETTERS

Narrow Window in Nanoscale Dependent Activation of Endothelial Cell Growth and Differentiation on TiO2 Nanotube Surfaces

2009 Vol. 9, No. 9 3157-3164

Jung Park,†,§ Sebastian Bauer,‡,§ Patrik Schmuki,‡ and Klaus von der Mark*,† Department of Experimental Medicine I, Nikolaus-Fiebiger-Center of Molecular Medicine, Friedrich-Alexander-UniVersity of Erlangen-Nuremberg, 91054 Erlangen, Germany, and Department of Materials Science, Institute for Surface Science and Corrosion (LKO), UniVersity of Erlangen-Nuremberg, Martensstrasse 7, 91058 Erlangen, Germany Received April 28, 2009; Revised Manuscript Received July 7, 2009

ABSTRACT Critical features of biomimetic materials used for vascular grafts and stents are surface structure and chemical features of the implant material supporting adhesion, proliferation, and differentiation of endothelial cells and smooth muscle cells, the major cell types of blood vessels. Recently, experimental evidence from several laboratories have indicated a strong stimulation of cellular activities on vertically aligned TiO2 nanotube surfaces in comparison to amorphous TiO2 surfaces. Conflicting reports exist, however, concerning the nanoscale dimension, and the role of the chemistry and crystallinity of the nanotubes in eliciting cell responses. Here we demonstrate that 15 nm nanotubes provide a substantially stronger stimulation of differentiation of mesenchymal cells to endothelial cells and smooth muscle cells than 70-100 nm nanotubes, while high rates of apoptosis were seen on 100 nm nanotubes. Also endothelial cell adhesion, proliferation, and motility were several-fold higher on 15 nm than on 100 nm nanotubes. Furthermore, our data indicate a clear dominance of the nanoscale geometry on endothelial cell behavior over surface chemistry and crystallinity of the TiO2 nanotube surface. These findings indicate that fine-tuning of TiO2 surfaces at nanoscale will be an essential parameter in optimizing endothelial cell and smooth muscle cell responses to vascular implants.

Recently, increasing experimental evidence has accumulated showing that nanoscale topography is an important factor for cellular recognition of the biological microenvironment and biomimetic material used for vascular grafts, stents, or bone implants. The sensitivity of cellular responses to surface topography was demonstrated using a lateral spacing model made of self-organizing nanoporous Al and Si surfaces,1 or a nanoscale surface protrusion model based on polymer demixing2 and ordered gold cluster arrays.3 Modifying topography, chemistry, and surface energies on titanium surfaces had a strong impact on different cellular responses and modulated the in vivo success rate of implantations.4,5 Several recent publications have shown that cell activations on fabricated TiO2 nanotubes are directed by topographical cues of the surface at nanoscale level and differ significantly from a unstructured, flat TiO2 surfaces or polystyrene culture dish surfaces,6-8 but the results are in part conflicting. While some groups showed enhanced cell behavior using anatase * Corresponding author. Tel. +49-9131-8529104. E-mail: kvdmark@ molmed.uni-erlangen.de. † Friedrich-Alexander-University of Erlangen-Nuremberg. ‡ University of Erlangen-Nuremberg. § Both authors contributed equally to the presented work. 10.1021/nl9013502 CCC: $40.75 Published on Web 08/04/2009

 2009 American Chemical Society

phase of nanotubes within 70-100 nm diameters compared to flat Ti surfaces,6,7 our previous study showed significantly enhanced cellular activity of mesenchymal stem cells (MSC) on 15 nm amorphous nanotubes in comparison to 70-100 nm nanotubes.9,10 Yet in another study, proliferation of smooth muscle cells was higher on flat TiO2 surfaces than on 30 nm nanotubes, while endothelial cells showed the opposite reaction.11 Part of the conflicts may be due to different methods used to generate TiO2 surfaces, resulting in different surface topology, crystalline status, and chemical composition. For example, several reports indicate superior cell support by TiO2 nanotubes in a crystalline anatase form as compared to amorphous TiO2,12 and anatase coating has been shown to improve implant osseointegration.13,14 Direct comparative experiments, however, have not been performed yet on TiO2 nanotube surfaces. Further explanations for discordant results from different laboratories may be due to the use of different cell types, including primary cells versus cell lines, and different methods employed to assess cellular activities. Thus, the dominant factors determining specific cell activities and cell fate on TiO2 nanotubular microenvironments remain to be defined for each cell type with quantitative and specific assays.

Figure 1. Topography, crystallographic structure, and chemistry of TiO2 nanotubes. (a) SEM top-views of the nanotubular layers of different tube diameters formed by anodization of titanium in 1 M H3PO4/0.12 M HF at potentials between 1 and 20 V. (b) XRD patterns recorded for an as-formed sample and after annealing in air at 450 °C for 1 h show the transformation of as-formed amorphous TiO2 nanotubes to crystalline anatase phase. (c) X-ray photoelectron spectroscopy showing fluoride (F 1s) peaks of three different 100 nm nanotube samples: high F- in as-formed nanotube surface, low F- after 3 days soaking in water, and no F- after annealing at 450 °C for 1 h (anatase). (d) Cell adhesion to high fluoride, 15 and 100 nm TiO2 nanotubes (8.38 and 8.49 at % F-, respectively) and low F- content (2 at %). (e) At 3 days GFP labeled mesenchymal stem cells plated on 15 nm nanotubular sheets were fully confluent, in contrast to 100 nm nanotube surfaces of both high fluoride and low fluoride TiO2 nanotubes.

Here we report on the behavior of three different cell types relevant for blood vessel formation (endothelial cell line mlEND, bone marrow mesenchymal cells (MSC), and human cord blood endothelial progenitor cells) on TiO2 nanotube surfaces with respect to (i) nanoscale size of tube diameter, (ii) the crystalline structure of TiO2 as anatase versus amorphous material, and (iii) the content of remaining fluoride in the nanotubes resulting from the preparation procedure. The cellular activities analyzed on these surfaces included cell adhesion, proliferation, differentiation, migration and apoptosis. 3158

We generated self-assembled, amorphous, and heat-treated, crystallized anatase phase TiO2 nanotubes by anodizing Ti sheets in a phosphate--fluoride electrolyte at different voltages ranging from 1 to 20 V with six different diameters between 15, 20, 30, 50, 70, and 100 nm (Figure 1a) as described previously.15-17 As-formed TiO2 nanotubes typically have an amorphous structure; the anatase phase was produced following the nanotube formation by annealing in air at 450 °C for 1 h (Figure 1b). For the analysis of residual fluoride remaining in the nanotubes following the annealing process, both amorphous Nano Lett., Vol. 9, No. 9, 2009

and anatase nanotube surfaces were analyzed by X-ray photoelectron spectroscopy (XPS). The XPS measurements showed that residual fluoride content in the nanotubes was less than 0.1 at % after annealing at 450 °C for 1 h used for anatase nanotubes, while in as-formed samples usual fluoride contents after soaking in water were about 4 at % (Figure 1c). Therefore it was possible to evaluate the direct effect of crystalline, anatase phase surfaces without an influence of residual fluoride content on cell behavior. To assess the role of residual fluoride content in amorphous nanotubes in cell behavior including cell adhesion and proliferation, MSC were plated on amorphous TiO2 nanotubes produced in an electrolyte of 1 M H3PO4/0.30 M HF (“high HF”, 8.38 and 8.49 at % fluoride on 100 and 15 nm nanotubes, respectively) or 0.12 M HF (“low HF, 2 at %). Our results show that the high fluoride concentration did not significantly change the number of adherent MSC cells 1 day after plating, irrespective of the nanotube diameter (Figure 1d). After 3 days, the cell numbers strongly increased on 15 nm nanotubes and were equal on both high and low HF conditions. On 100 nm nanotubes, however, the increase in cell number was much lower than on 15 nm nanotubes, while high HF nanotubes doubled that on low HF nanotubes (Figure 1d). Figure 1e confirms that cell adhesion to 15 nm nanotubes exceeded by far that on 100 nm nanotubes on both high and low HF surfaces, while more cells attached and survived on high fluoride than on low fluoride 100 nm nanotubes. These findings indicate that a high fluoride content in the TiO2 nanotubes does not affect initial cell adhesion but may support cell proliferation after 3 days. In osteoblast cultures, however, fluoride-treated, grit-blasted TiO2 surfaces supported greater proliferation and increased bone sialoprotein and BMP-2 expression.18 Also RUNX-2 and Osterix levels were significantly increased on treated titanium implant surfaces.19,20 Other ions introduced into the tubular structure were found to be at a similarly low concentration level independent of the tube diameter or anodization time (Figure S1, Supporting Information). Therefore those background ions cannot be responsible for the observed size effects. TiO2 nanotubes smaller than 15 nm diameter were not tested as they turned out to be structurally unstable. As shown previously for mesenchymal stem cells (MSC),9,10 both adhesion and proliferation rates of endothelial cells on flat amorphous TiO2 surfaces were about 20% lower than on 15 nm nanotubes (Figure S2a,b, Supporting Information). To assess the role of the crystalline structure of TiO2 in supporting endothelial cell functions, TiO2 nanotubes prepared in a size range of 15-100 nm diameter as F--free anatase phase and amorphous surfaces were incubated with the mouse endothelial cell line mlEND21 (Figure 2) and rat bone marrow stem cells (MSC)9 (Figure S3, Supporting Information). Altogether, the pattern of cell responses was strongly dependent on the tube diameter, but rather similar on amorphous versus anatase nanotube samples, with some exceptions (see below). Adhesion and proliferation rates of mlEND cells equally increased with decreasing diameter of Nano Lett., Vol. 9, No. 9, 2009

the nanotubes, on both amorphous and anatase nanotube surfaces at 1, 3, and 7 days after cell plating (Figure 2a-c; Figure S3a-c, Supporting Information). Only on 50-70 nm nanotubes, but not on smaller nanotubes, 3 days after plating cell numbers were significantly lower on anatase than on amorphous nanotubes, indicating higher rates of cell death (Figure 2b), since the proliferation rates measured by a colorimetric WST assay were not different on both surfaces (Figure 2c). Adhesion and proliferation rates of MSC cells, however, were somewhat higher on amorphous than on anatase nanotubes (Figure S3a,c, Supporting Information). Three days after cell plating, mlEND endothelial cells developed abundant focal contacts on both anatase and amorphous 15 nm nanotubes, but not on 100 nm nanotubes, as shown by immunostaining for paxillin (Figure 2d). Most important, immunostaining for vWF (von Willebrand factor), a marker for mature endothelial cells, demonstrated that endothelial cell differentiation was strongly supported on both amorphous and anatase 15 nm but not on 100 nm nanotubes (Figure 2e). Synthesis of vWF was also considerably higher on 15 nm nanotubes than on smooth TiO2 surfaces (Figure S2c, Supporting Information). In contrast, on 100 nm nanotubes a large number of endothelial cells (36.7%) underwent apoptosis, compared to 8.5% on 15 nm surfaces (Figure 2f). This was detected by flow cytometry after staining with FITC-annexin-V at 2 days after plating. For comparison, apoptosis rates on polystyrene culture dishes used as control were 9.8% (data not shown). These results indicate that for adhesion and growth of endothelial cells and MSC, the nanotopography of the microenvironment is a dominant factor in comparison to the crystalline structure or fluoride content of the TiO2 nanotubes. Furthermore, the data show that endothelial cells respond to the same nanosize range 15-30 nm as osteoblasts, osteoclasts,10 or mesenchymal stem cells, as shown in our previous studies,9 while spacings between 50-100 nm show less support for these activities, and instead induce apoptosis. To confirm the stimulatory effect of 15 nm nanotubes on differentiation of the murine endothelial cell line mlEND reported above with primary endothelial cells, we analyzed the ability of amorphous and anatase nanotubes to support differentiation of bone marrow MSC to endothelial cells. Differentiation was induced in culture medium containing a VEGF and FGF growth factor cocktail, which directs stem cell differentiation specifically to endothelial cells22,26 (Figure S4, Supporting Information). Immunofluorescence analysis of the endothelial cell markers vWF23 and VE-cadherin24 6 days after incubation in differentiation medium revealed strong induction of differentiation on 15 nm but not on 100 nm nanotubes, on both amorphous and anatase surfaces (Figure 3a,b,c). Most importantly, the deposition of VEcadherin, an endothelial-cell-specific cell adhesion molecule aligned at the borders of neighboring endothelial cells on 15 nm nanotubes (Figure 3a,b) indicated the formation of intense cell-cell contacts required for the formation of functional endothelial cell layers. Also the differentiation of CD133 positive human cord blood stem cells25 to endothelial cells is enhanced on 15 nm nanotubes but does not take place 3159

Figure 2. Nanosize-dependence of endothelial cell behavior on amorphous versus crystalline TiO2 nanotubes. Mouse endothelial cells mlEND were plated on amorphous and anatase nanotubes. Cell adhesion increased with decreasing nanotube diameter with a maximum at 15 nm, irrespective of the crystalline structure of the nanotubes on diameters (a). At 3 days cell numbers were higher proliferation on amorphous 50-70 nm than on anatase nanotubes of the same size (b). At 7 days proliferation rates measured after 7 days in culture were the same on amorphous and anatase nanotubes (c). Immunostaining for paxillin (red) revealed abundant focal contact formation on 15 nm but not on 100 nm nanotubes (d). Extensive staining for vWF, a marker for mature endothelial cell surface was seen at 7 days after plating on 15 nm nanotubes of both amorphous and anatase nanotubes indicating enhanced endothelial cell differentiation. (von Willebrand factor: red; F-actin: green; nuclear DAPI staining: blue) (e). On 100 nm nanotubes a large number of endothelial cells (36.7%) underwent apoptosis (programmed cell death) compared to 8.5% on 15 nm shown by apoptosis assay using annexin V-FITC (green: unstained control) (f).

on 100 nm nanotubes, as shown by staining for the endothelial cell marker PECAM (CD31) (Figure 3c). By culturing MSC in a different medium designed to drive differentiation into smooth muscle cells,22,26 cell differentiation of MSC cells was directed toward smooth muscle cells. As observed with endothelial cells, differentiation to smooth muscle cells as indicated by expression of smooth muscle actin was also highly stimulated on 15 nm but not on 100 nm nanotubes, irrespective of the crystalline structure (Figure 3d). 3160

The elongated cell shape of endothelial cells seen on 15 but not 100 nm nanotubes (Figure 3c) suggested enhanced migration activity on 15 nm nanotubes. To asses the nanosize dependent cell motility in a quantitative manner, cell tracks of fluorescence labeled MSC cells and mlEND endothelial cells migrating during 24 h on 15 and 100 nm nanotube surfaces were recorded by video microscopy and depicted as rosette with starting points aligned in the center to assess their relative motility.27 (Figure 3e). The data show significantly enhanced cell motility of both MSC and endothelial Nano Lett., Vol. 9, No. 9, 2009

Figure 3. Endothelial differentiation on amorphous vs anatase phase nanotubes. Differentiation of MSC (a,b) to endothelial cells on different TiO2 nanotube surfaces was induced with specific differentiation medium. Immunofluorescence analysis of vWF (red) and VE-cadherin (green) revealed a strong expression of endothelial markers and formation of VE-cadherin positive cell-cell contacts on both amorphous (a) and anatase phase 15 nm nanotubes (b) but not on 100 nm nanotubes (a, b). (c) Similarly, the differentiation of human cord blood endothelial progenitor was enhanced on 15 nm amorphous and anatase nanotubes but not on 100 nm nanotubes, as seen by immunostaining for PECAM (red) (F-actin; green, nuclear staining in blue). On both amorphous and anatase 15 nm, but not 100 nm nanotubes, cells assumed migratory and elongated cell shape. (d) Also smooth muscle cell differentiation (c) from GFP-labeled MSC was highly stimulated on 15 nm nanotubes irrespective of the crystalline structure as shown by immunostaining for smooth muscle actin (red) (green: green fluorescent protein in MSC cytoplasm). (d) Further cell migration activity was significantly different on different nanotube sizes (e). Cell trackings of GFP labeled MSC cells and DiI-labeled mlEND cells on amorphous TiO2 nanotubes by fluorescence-video microscopy showed strongly enhanced cell motility on 15 nm nanotubes a compared to 100 nm nanotubes. (right panel in e, * ) p < 0.001, ** ) p < 0.005, unpaired t test).

cells on 15 nm nanotubes as compared to 100 nm nanotubes. Video microscopy of MSC and endothelial cells revealed that cells rapidly extend and retract filipodia on 100 nm surface, indicating a frustraneous attempt to migrate, but in Nano Lett., Vol. 9, No. 9, 2009

fact cells do not move, in contrast to 15 nm surfaces (see videos, Supporting Information). Resettling of vascular grafts with cells and capillary formation during wound healing including bone regeneration 3161

following a TiO2 implantation require active endothelial proliferation migration and differentiation, a process referred to as neo-angiogenesis, which involves the recruitment of endothelial cells to the site of injury. Two common mechanisms of neo-angiogenesis, i.e., endothelial migration and sprouting from preexisting endothelial cells, as well as recruitment of endothelial precursor cells from circulation are critical for targeting cells for successful bone-implant integration.28 Endothelial progenitor cells expanded in vitro from CD133(+) stem cells by stimulation with VEGF (vascular endothelial growth factor) have shown to contribute to neo-angiogenesis after cell transplantation in heart stroke animal model or ischemic limb animal model.28,29 Here we have demonstrated that TiO2 nanotube surfaces in the range of 15-30 nm diameter provide specific topographical cues to support adhesion, proliferation, migration, and differentiation of endothelial cells, which proposes the utilization of such nanostructured surfaces as suitable biomimetic materials for the use of vascular grafts. The responses of endothelial cells and MSC to TiO2 nanotube surfaces are largely independent from the fluoride content and from the crystalline structure of TiO2 but underlie a stringent control by the nanoscale topography. The findings are to some extent consistent with previous studies on the role of nanoscale dimensions of cellular responses to structured surfaces. Enhanced cell adhesion and focal contact formation in distinct spacings of 98%. Isolated cells were cultivated in medium 199 (M199; GIBCO-BRL) supplemented with 20% fetal bovine serum for 7 days, plated on different nanotubes and stimulated for endothelial cell differentiation with medium 199 containing 5 ng/mL FGF-2 (human recombinant fibroblast growth factor-basic; SIGMA) and 10 ng/mL VEGF (vascular endothelial growth factor) for 7 days. Cell Adhesion, Proliferation Assay, and Apoptosis Assay. Endothelial cell line (mlEND) and GFP-labeled mesenchymal stem cells and were plated on titanium surfaces with cell densities of 5000/cm2. For cell adhesion experiment 24 h after cell plating nonadherent cells were washed off with PBS and adherent cells were counted at 6 different areas (1280 × 1024 pixels) under a fluorescent microscope (5× magnification). Cell growth was analyzed by counting cells 3 days after cell plating. Cell proliferation rates were quantified at 7 days using the cell proliferation assay WST-1 (Roche) according to the manufacturer’s instructions. Cell apoptosis assays were performed 2 days after plating mLENDS at a cell density of 10 000/cm2 on 15 and 100 nm nanotubes of 9 cm2 titanium plates. Polystyrene culture dishes served as a control. After resuspending the harvested cells in 100 µL of binding buffer (10 mM HEPES, pH 7.4, 140 mM NaCl, 2.5 mM CaCl2), we incubated cells with 5 µL of annexin V conjugated with FITC (a kind gift of Dr. Ernst Po¨schl, Erlangen) for 15 min at room temperature in the dark. Annexin V-positive cells were counted using a fluorescenceactivated cell sorter (Becton-Dickinson, FACs Calibur). Immunocytochemistry. Cell preparation and fixation for immunocytochemical staining was done as described previously.9,10 For visualization of focal contacts on nanotubes, Nano Lett., Vol. 9, No. 9, 2009

endothelial cells grown for 3 days were fixed, permeabilized, and incubated with antibodies of mouse monoclonal antipaxillin (Signal Transduction) followed by a polyclonal goat antimouse Cy5-conjugated secondary antibody (Chemicon). Cell nuclei were stained blue with DAPI (Roth). Endothelial cell differentiation was evaluated by staining with antibodies against vWF (Chemicon), anti VE-cadherin (Santa Cruz), and anti PECAM (Cell Biochem). Smooth muscle cell differentiation of MSC was followed by staining with a mouse monoclonal antibody for smooth muscle actin (Sigma). Bound antibodies were labeled with antimouse Cy5- and antirabbit Cy5-conjugated secondary antibodies (Chemicon) and analyzed by fluorescence microscopy using an Axioplan 2 microscope (Zeiss). Cell nuclei were stained blue with DAPI (Roth). For cytoplasmic actin staining cells were stained with antibodies of Alexa488-labeled phalloidin (Biosource). Cell Tracking for Quantification of Cell Migration. To analyze endothelial cell motility quantitatively on different sizes of nanotubes, DiI-labeled endothelial cell mlEND and GFP-labeled MSC were plated on 15 and 100 nm nanotubes at cell density of 5000/cm2 and incubated with endothelial differentiation medium. One day after cell plating, cell migration was monitored by time-lapse video microscopy in intervals of 15 min during 24 h and analyzed using Openlab software (Improvision). For quantification of cell motility, migration tracks of each 10 cells of MSC and mlEND cells were collected and plotted as rosettes with common starting points in the center.27 The migration velocity of each cell track during 24 h was calculated from the number of pixels/track; for statistical analysis the unpaired t test was used. Acknowledgment. We gratefully thank Mrs. Hildebrand for XPS investigations, Mrs. Marten-Jahns for XRD investigations, Mrs. Friedrich for SEM investigations, and Andreas Pittrof, Department of Materials Science, University of Erlangen-Nuremberg. This work was supported by the Deutsche Forschungsgemeinschaft (SCHM1597/ 9-1 and MA534/20-1). Supporting Information Available: Figures showing XPS investigations on ion uptake during TiO2 nanotube formation, cell count and proliferation rates of endothelial cells on 15 nm nanotubes and smooth surfaces and furthermore of mesenchymal stem cells on amorphous and anatase TiO2 nanotubes. Videos of DiI-labeled endothelial cells and GFP-labeled MSC plated on 15 nm (video 1) and 100 nm (video 2) nanotubes with endothelial differentiation medium. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Popat, K. C.; Chatvanichkul, K. I.; Barnes, G. L.; Latempa, T. J., Jr.; Grimes, C. A.; Desai, T. A. J Biomed. Mater. Res. A 2006, 955-964, 6. (2) Dalby, M. J.; Riehle, M. O.; Yarwood, S. J.; Wilkinson, C. D.; Curtis, A. S. Exp. Cell Res. 2003, 284 (2), 274–82. (3) Boyen, H. G.; Kastle, G.; Weigl, F.; Koslowski, B.; Dietrich, C.; Ziemann, P.; Spatz, J. P.; Riethmuller, S.; Hartmann, C.; Moller, M.; Schmid, G.; Garnier, M. G.; Oelhafen, P. Science 2002, 297 (5586), 1533–6. 3163

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Nano Lett., Vol. 9, No. 9, 2009