ARTICLE pubs.acs.org/Biomac
Surface Functionalization of TiO2 Nanotubes with Bone Morphogenetic Protein 2 and Its Synergistic Effect on the Differentiation of Mesenchymal Stem Cells Min Lai,† Kaiyong Cai,*,† Li Zhao,‡ Xiuyong Chen,† Yanhua Hou,† and Zaixiang Yang† †
Key Laboratory of Biorheological Science and Technology (Chongqing University), Ministry of Education, College of Bioengineering, Chongqing University, Chongqing 400044, P. R. China ‡ China National Centre for Biotechnology Development, No. 16, Xi Si Huan Zhong Lu, Haidian District, Beijing 100036, P. R. China ABSTRACT: To investigate the influence of surface-functionalized substrates with nanostructures on the behaviors of mesenchymal stem cells, we conjugated bone morphogenetic protein 2 (BMP2) onto TiO2 nanotubes with different diameter sizes of 30, 60, and 100 nm for in vitro study. Polydopamine was employed as the intermediate layer for the conjugation of BMP2. The successful conjugation of BMP2 onto TiO2 nanotubes was revealed by field-emission scanning electron microscopy (FE-SEM), X-ray photoelectron spectroscopy (XPS), and contact angle measurements. Immunofluorescence staining of vinculin, osteocalcin (OCN), and osteopontin (OPN) revealed that BMP2-functionalized TiO2 nanotubes was favorable for cell growth. More importantly, MSCs cultured onto BMP2-functionalized TiO2 nanotubes displayed significantly higher (p < 0.05 or p < 0.01) differentiation levels of ALP and mineralization after 7 and 14 day cultures, respectively. The results suggested that surface functionalization of TiO2 nanotubes with BMP2 was beneficial for cell proliferation and differentiation. The approach presented here has potential application for the development of titanium-based implants for enhanced bone osseointegration.
’ INTRODUCTION The investigation of the interactions between cells and biomaterials is one of the key issues for the development of biomaterials. The biological behaviors of cells are highly regulated by the local microenvironment of chemistry and topography on mesoscale, microscale, and nanoscale sizes.1 Recently, how to control the cell behavior within microenvironments on the nanoscale attracted more and more attention. Cells were verified to respond to material surface that has nanoscale features in a controlled manner. For instance, cells demonstrated distinct responses to nanoscale patterns with disorder and order geometry.2,3 The pillar-structured surface was reported to control the cell architecture and cell function.4 Controlled arrays of high aspect ratio substrate-bound nanowires were also proven to regulate the growth and maintain the differentiated states of neuronal cells.5 In another study, nanoimprinted pattern was reported to guide the morphology and motility of smooth muscle cells.6 Titanium (Ti) and its alloys have been widely used as orthopedic implants in clinical applications because of their bioinert and good mechanical properties.7 However, how to improve the osseointegration between an implant and surrounding nature bone tissue, in turn to extend the lifetime of implant, still remain unresolved.8,9 Nature bone is generally composed of organic extracellular matrix components (collagen, proteoglycan, etc.) and inorganic components with nanoscale features.1 To mimic closely the nanoscale architecture of human bone, the construction of an implant presenting both local r 2011 American Chemical Society
chemistry and nanoscale topography becomes one of the hot topics in the related field. Recent studies indicated that TiO2 nanotubes formed onto Ti substrates could be employed as an ideal platform to present nanoscale topography. The diameters and shape of TiO2 nanotubes were tunable in a controlled manner by optimizing fabrication parameters.10-12 Previous studies demonstrated that TiO2 nanotubes with different diameters could affect the proliferation and differentiation of mesenchymal stem cells (MSCs).13,14 Nevertheless, no study investigated the influence of combined nanotopography and local chemistry on the biological behaviors of MSCs. Except for topography, local chemistry of a substrate is another important factor that contributes to cellular microenvironment. It thus directly regulates the interactions between cells and materials. Surface modification is a feasible approach to change the surface chemistry of substrates. Growth factors, hormones, and chemicals could be directly immobilized onto the surface of a substrate via either chemical or physical strategy. Previously, we successfully modified titanium substrates with GFP-BMP encoding plasmid DNA via layer-by-layer technique.15 Bone morphogenetic proteins (BMPs) play important roles in bone and cartilage formation. Among the members of BMP Received: November 30, 2010 Revised: January 28, 2011 Published: March 07, 2011 1097
dx.doi.org/10.1021/bm1014365 | Biomacromolecules 2011, 12, 1097–1105
Biomacromolecules family, BMP2 has been demonstrated to promote bone formation in vivo.16,17 The growth factor of BMP2 was proven to stimulate the differentiation of osteogenic cells. For instance, BMP2 promoted the osteogenesis when incorporated into hyaluronic acid hydrogels along with MSCs.18 Carstens et al. demonstrated in situ osteogenesis in a craniofacial mandibular defect using absorbable collagen sponges as delivery vehicles for recombinant human BMP2 (rhBMP2).19 The purpose of this study was to fabricate BMP2-functionalized TiO2 nanotubes with different diameters and to investigate the synergetic effect of nanotopography and chemical cue (BMP2) of TiO2 nanotubes on the regulation of proliferation and differentiation of MSCs.
’ EXPERIMENTAL SECTION Materials. Titanium foils (0.25 mm thickness, 99.5%) were provided by Alfa Aesar (Tianjin, China). 3-(4,5-Dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide (MTT), alkaline phosphatase (ALP), 3,4dihydroxyphenylalanine (dopamine), and bicinchoninic acid (BCA) assay kit were purchased from Sigma Chemical. Recombinant human BMP2 was purchased from PeproTech. TiO2 Nanotube Fabrication. TiO2 nanotubes were prepared according to a previous study.10 Titanium foils were sequentially rinsed with acetone, isopropanol, ethanol, and distilled water each for 20 min with sonication, respectively. For anodization, platinum foil was employed as negative electrode, whereas titanium foils were used as positive electrode. A mixture solution of glycerol and water (1:1, vol) containing 0.27 M NH4F was used as electrolyte. TiO2 nanotubes with different diameters were formed at potential of 10, 20, and 25 V at room temperature for 3 h, respectively. After electrochemical treatment, all samples were rinsed with distilled water. Polydopamine Coating and BMP2 Conjugation. TiO2 nanotube substrates were immersed in solution of 50 mL of dopamine (2 mg/ mL) with 10 mM Tris buffer (pH 8.5) for overnight in the dark. Treated samples were rinsed with distilled water to remove the residual dopamine for further treatment. The polydopamine-coated TiO2 nanotube substrates (20 pieces) were then immersed in 50 mL of BMP2 solution (80 ng/mL) with 10 mM Tris buffer (pH 8.5) and incubated for overnight at room temperature. Then, samples were rinsed with PBS to remove the physically adsorbed BMP2 and dried at ambient temperature for following cell experiments. The polydopamine-coated TiO2 nanotubes were denoted as PDOP-TiO2 nanotubes, whereas the BMP2 grafted polydopamine-coated TiO2 nanotubes were denoted as BMP2-PDOP-TiO2 nanotubes. Surface Characterization. The morphology of the substrates was observed by field-emission scanning electron microscopy (FEINova 400 Nano SEM, Phillips, Holland). The chemical composition of the substrates was determined by X-ray Photoelectron Spectroscopy (XPS) using the model PHI 5600 system (Perkin-Elmer) with a Mg KR source (1253.6 eV). Contact angles of the substrates were measured by model 200 video-based optical system (Future Scientific Tai Wan, China). The images of the water drops on the sample surface were recorded by a camera and analyzed with the software supplied by the manufacturer. The contact angles were calculated from the mean value of five different substrates. Cell Culture. MSCs were isolated from bone marrow of rats’ femur and tibia (Wistar rat, with weight of 150-200 g) according to a previous study.20 Cells were cultured in DMEM supplemented with 10% bovine serum (FBS, Gibco) under 5% CO2 atmosphere at 37 °C. Medium was changed on the first day, then every 2 days. After confluence, cells were detached with 0.25% trypsin in 1 mM tetrasodium EDTA, centrifuged, and resuspended in the complete medium for reseeding in new culture flasks. MSCs at the third passage were used for cellular experiments. The initial cell seeding density was 4 104 cells/well (24-well plate) in this study.
ARTICLE
Immunofluorescence Staining. After 2 days of culture, MSCs adhered to substrates were fixed with 2% glutaraldehyde at 4 °C for 20 min. Samples were then washed with PBS and permeablized with 0.2% Triton X-100 at 4 °C for 2 min. Next, the treated samples were washed three times with PBS and incubated with 1% bovine serum albumin (BSA)/PBS at 37 °C for 1 h. Subsequently, goat monoclonal antibody against vinculin (1:200) (Santa Cruz Biotechnology) was added at 4 °C for overnight. After that, mouse-antigoat fluorescein isothiocyanate (FITC)-conjugated secondary antibody (1:100) (ZSGB-BIO, Beijing, China) was added at 37 °C for another 1 h. Samples were rinsed with PBS for further treatment. Finally, treated samples were stained with 5 U/mL rhodamine-phalloidin (Invitrogen) at 4 °C for overnight and counterstained with 10 μg/mL Hoechst 33258 (Sigma-Aldrich, St. Louis, MO) at room temperature for 5 min. The procedure for osteocalcin (OCN) and osteopontin (OPN) staining was similar to that of vinculin, however, with different primary antibodies. The staining was performed after culture for 21 days. The stained samples were finally mounted with 90% glycerinum and observed with CLSM (TCS SP5, Leica, Germany). Cell Viability. Cell viability was determined by MTT assay. In brief, MSCs were cultured on unmodified substrates, modified substrates, and tissue culture polystyrene (TCPS) for 7 and 14 days, respectively. Next, 100 μL of MTT (5 mg/mL) was added to each well and incubated at 37 °C for another 4 h. MTT-containing medium was removed, and 0.5 mL of dimethyl sulfoxide (DMSO) was added to each well to dissolve formazan crystal. The optical density of the solution was measured at 490 nm with a spectrophotometric microplate reader (Bio-Rad 680). Each treatment was performed five times. The mean value was used as the final result. Total Intracellular Protein Content Assay. The assay was performed according to a previous study.21 MSCs were cultured on unmodified substrates, modified substrates, and TCPS for 7 and 14 days, respectively. MSCs were lysed by 1% Triton X-100 with three freezethaw cycles. Total protein content in the cell lysates was measured with a BCA assay kit. The absorbance was measured at a wavelength of 570 nm with a spectrophotometric microplate reader (Bio-Rad 680). The total intracellular proteins (expressed as milligrams) expressed by MSCs were determined from a standard absorbance curve versus known concentration of albumin run in the parallel experiment. Alkaline Phosphatase Activity Assay. The ALP activity assay was performed according to a previous study.22 Cell lysate was prepared as above-mentioned. Paranitrophenyl phosphate was employed as a reference substrate to determine the ALP activity of MSCs cultured on different substrates. The absorbance at wavelength of 405 nm was measured with a spectrophotometric microplate reader (Bio-Rad 680). The ALP activity (expressed as μmol of converted p-nitro-phenol/min) was normalized by the total intracellular protein production. The ALP activity was thus expressed as μmol p-nitrophenol/min/mg protein. Mineralization Assay. After 21 days of culture, mineralization was measured after the elution of alizarin red-S (ARS) (Sigma-Aldrich) staining according to a previous study.13 Cells adhered to different substrates were washed with PBS and then fixed with 2% glutaraldehyde at 4 °C for 20 min. Next, samples were washed three times with distilled water and stained with 40 mM ARS (pH 4.1) at room temperature for 20 min. The surface layer on the substrate was collected with acetic acid (10%, v/v) and transferred to a microcentrifuge tube. The microcentrifuge tubes were heated to 85 °C for 10 min and then centrifuged at 20 000 g min-1 for 15 min. The supernatant was transferred to a new microcentrifuge tube and neutralized by the addition of 10% v/v ammonium hydroxide. The absorbance of supernatant was measured at wavelength of 405 nm with a spectrophotometric microplate reader (Bio-Rad 680). Statistical Analysis. All data were expressed as means ( standard deviation (SD) with n = 5. The statistical analysis was performed with OriginPro (version 6.1) at confidence levels of 95 and 99%. 1098
dx.doi.org/10.1021/bm1014365 |Biomacromolecules 2011, 12, 1097–1105
Biomacromolecules
’ RESULTS Figure 1 shows the scheme of the conjugation of BMP2 onto the surfaces of TiO2 nanotubes. TiO2 nanotubes were fabricated via an anodization approach. Prepared TiO2 nanotubes substrates were directly dipped into dopamine solution with an adjusted pH value of 8.5. Self-polymerization occurred to form a polydopamine film onto surfaces of TiO2 nanotubes substrates.23 The catechol and quinone functional groups of polydopamine film were then utilized to conjugate BMP2 onto TiO2 nanotubes. Surface Characterization. The surface morphologies of native and modified TiO2 nanotubes were characterized by SEM (Figure 2). TiO2 nanotubes with average diameters of 30, 60, and 100 nm were formed on the top layers of Ti substrates when applying anodization potential of 10, 20, and 25 V, respectively (Figure 2 a). Figure 2 b displays the representative SEM images of TiO2 nanotubes with different diameters after being treated with dopamine coating and further conjugation of
Figure 1. Schematic illustration of the conjugation of BMP2 onto TiO2 nanotubes.
ARTICLE
BMP2, respectively. TiO2 nanotubes were partially covered by a layer, which might be derived from the formation of polydopamine film. To investigate the successful formation of polydopamine film and conjugation of BMP2 onto TiO2 nanotubes, XPS measurement was employed in this study. Figure 3 shows the representative XPS spectra of native TiO2 nanotubes, PDOP-TiO2 nanotubes, and BMP2-PDOP-TiO2 nanotubes (30 nm). The surface chemical composition of each sample was listed in Table 1. Native TiO2 nanotubes demonstrate three elements of C, Ti, and O (Figure 3 a). The high content of carbon was derived from the environmental contamination.24 After being coated with dopamine, an additional peak at 399.2 eV was observed (Figure 3 b). The presence of N element was derived from amino groups of dopamine molecules. The high content of 7% (Table 1) for N directly suggests that polydopamine film was formed onto TiO2 nanotubes. After conjugation of BMP2 onto PDOP-TiO2 nanotubes surfaces, the N content further increased to 8.6% (Table 1), which was mainly due to the higher content of N in BMP2 molecules compared with that in PDOP molecules. This result was consistent with a previous study.25 The result confirmed that BMP2 was successfully conjugated onto the surfaces of TiO2 nanotubes. To confirm further whether BMP2 was conjugated onto TiO2 nanotubes, a water contact angle measurement was performed to monitor each treatment. Water contact angle measurement is a method that indicates the hydrophobic/hydrophilic properties of the measured substrates. Native TiO2 nanotubes displayed superhydrophilic properties with contact angle 0.05) was observed. However, MSCs cultured onto 100 nm Ti nanotubes showed lower (p < 0.05) mineralization than those on native Ti. Taken together, all results provided direct evidence that surface functionalization of TiO2 nanotubes with BMP promoted the osteogenic differentiation of MSCs.
’ DISCUSSION In this study, the TiO2 nanotubes with various diameters were surface-functionalized with BMP2. We demonstrated that nanotopography and local chemistry synergistically stimulated the proliferation and differentiation of MSCs. It is well known that cellular microenvironments play an important role in the regulation of cells behaviors. Cellular responses (adhesion, migration, proliferation, and differentiation, etc.) to a material were highly directed by the surface characteristics, such as
local chemistry and surface topography.27 The investigation of the synergistic effect of surface topography and chemical cues on cell behaviors is of crucial importance for the development of biomaterials. As depicted in Figure 1, we used a simple method to conjugate BMP2 onto TiO2 nanotube surfaces via an intermediate layer of polydopamine. TiO2 nanotubes provided desirable nanoscale topography for cell adhesion and spreading; on the other hand, the conjugated BMP2 afforded biochemical stimulus for the proliferation and differentiation of MSCs. Therefore, the design of incorporation of nanostructures of TiO2 nanotubes and osteoinductive factor (BMP2) provided an instructive extracellular microenvironment for MSCs. Different diameters of titanium nanotubes with regular structures were successfully fabricated (Figure 2a). The diameters of titanium nanotubes were directly related to the anodization potentials.11,12 Surface functionalization of TiO2 nanotubes was conducted by initially introducing polydopamine as an intermediate layer. After polydopamine coating and BMP2 conjugation, a thin layer was formed onto the surface of titanium substrate and partially covered the TiO2 nanotubes (Figure 2b). XPS and contact angle were further employed to investigate the conjugation of BMP2. Except for titanium and oxygen elements, titanium substrate with nanotubes displayed carbon signals (32.8%) on its surface (Figure 3 a and Table 1). It was derived 1102
dx.doi.org/10.1021/bm1014365 |Biomacromolecules 2011, 12, 1097–1105
Biomacromolecules
ARTICLE
Figure 8. Osteopontin expression of MSCs seeded onto (a) Ti, (b) BMP2-PDOP-Ti, (c) 30 nm TiO2 nanotubes, (d) BMP2-PDOP-30 nm TiO2 nanotubes, (e) 100 nm TiO2 nanotubes, and (f) BMP2-PDOP-100 nm TiO2 nanotube (scale bar, 200 μm). Actin filaments of the cells stain (red), cell nuclei (blue), and osteopontin (green).
Figure 9. Quantification of cell mineralization on different substrates after 21 days of culture. Error bars represent means ( SD for n = 5, **p < 0.01.
from the contamination. Titanium is a chemically active material, which is easily contaminated by adventitious, unavoidable hydrocarbon adsorption from ambient air. It was also observed in our
previous study24 and other reports.28,29 No fluorine was observed, which indicated that there was no chemical residual after anodization. The result directly suggested that the carbon contamination was contributed to the physical adsorption from air. Dopamine is the analogue of adhesive proteins in mussel with good biocompatibility. It was proven to be an efficient agent for multifunctional coatings of any substrates.25,26,30 In this study, we coated TiO2 nanotubes by dipping substrates in alkaline dopamine solution. Self-polymerized, surface-adhesive polydopamine thin film onto TiO2 nanotubes was then utilized for further conjugation of BMP2. Although the deposited polydopamine coating is chemically heterogeneous, catechol and quinone functional groups presented in the polydopamine coating, in particular, the latter of which could be used for covalent coupling of biomacromolecules,25,30 such as trypsin and vascular endothelial growth factor (VEGF), under the alkaline condition. In this study, we employed to conjugate BMP2. Around 7% nitrogen was observed after dopamine coating, which suggested the successful coating. After BMP2 conjugation, the content of nitrogen increased to 8.6% (Table 1). The nitrogen content of BMP2 was calculated around 16.9% based on its peptide structure. Therefore, we could estimate that the coverage of BMP2 onto the TiO2 nanotubes was ∼9.5%. Water contact angle measurements further revealed that the conjugation occurred. All 1103
dx.doi.org/10.1021/bm1014365 |Biomacromolecules 2011, 12, 1097–1105
Biomacromolecules of the above data confirmed that BMP2 was successfully conjugated to titanium substrates (Figures 2 and 3, Table 1). Cell adhesion is the first event when cells come into contact with a material. It is essentially important to regulate subsequent cellular behaviors, such as cell proliferation, differentiation, protein production, as well as gene expressions.27,31 Cell adhesion is normally mediated by the transmembrane integrin protein, which is located at the site of adhesion. Vinculin is an intracellular protein involved in the linkage among cell adhesion membranous molecules, integrins, and actin filaments.32 It plays a key role in initiating and establishing cell adhesion and cytoskeletal development. The focal adhesion of a cell is highly sensitive to its microenvironment, which is composed of biochemical signals from the extracellular neighborhood cells and surface characteristics (chemistry, topography, etc.) of the materials.33 MSCs grown on BMP2-functionalized TiO2 nanotubes displayed higher expression of vinculin than those grown on native TiO2 nanotubes (Figure 4). The result clearly demonstrated the stimulation of BMP2 on the adhesion of MSCs. Previous study revealed that mechanism was that BMP2 was helpful for the ECM organization and enhanced expression of fibronection, R5 and β1 integrin subunit, and focal adhesion kinase (p125FAK).34 The comparatively high expression of vinculin promoted actin formation and cytoskeleton organization. This phenomenon was clearly demonstrated by MSCs adhered to substrates of BMP2-PDOP-30 nm TiO2 nanotubes and BMP2-PDOP-60 nm TiO2 nanotubes rather than BMP2-PDOP100 nm TiO2 nanotubes. Compared with MSCs grown on native TiO2 nanotubes, cells grown on BMP2-functionalized TiO2 nanotubes displayed higher (p < 0.01) cell viabilities after 7 and 14 days of culture. PDOP coating had little influence on the proliferation of MSCs. This result proved that cell adhesion via sensing extracellular matrix (ECM) greatly improved the cell proliferation (Figures 4 and 5). Previous studies also observed that BMP2 had a positive effect on the proliferation of MSCs and osteoblasts via a pathway of activation of receptor-regulated Smads, respectively.35-37 Although grafted BMP2 had an effect on cell proliferation, the nanoscale surface topography still influences the cell growth. MSCs grown on native TiO2 nanotubes with diameters of 30 and 60 nm displayed higher proliferation tendency than those grown on titanium substrate and 100 nm TiO2 nanotubes. Previous studies have demonstrated that vitality, proliferation, and differentiation of the MSCs could be critically influenced by the nanotopography of TiO2 nanotubes.13,14 The result was consistent with previous studies.14,38 The reason was that PODA coatings only partially blocked the nanotubes. The mechanism proposed by a previous study was that spacing >70 nm did not support focal adhesion formation and cell signaling and thus induction of the cell apoptosis.14 As an immature marker of osteoblast and an essential element of ossification,39 ALP activity was quantitatively measured to indicate the early osteogenic tendency (Figure 6). Compared with MSCs grown on native TiO2 nanotubes, cells grown on BMP2-functionalized TiO2 nanotubes displayed higher (p < 0.05 or p < 0.01) ALP activity after 7 and 14 days of culture. PDOP coating had little influence on the proliferation of MSCs. The result clearly demonstrated that surface functionalization with BMP2 promoted the differentiation of MSCs. BMP2-functionalized 30 nm TiO2 nanotubes displayed the highest ALP activity among groups after 7 days of culture. It was attributed to the synergetic effect of surface topography and functionalization with BMP2. A noticeable phenomenon was that the ALP activity of MSCs cultured for 14 days was relatively lower than those after 7 days culture in each group. It was related to the fact that MSCs differentiated into mature
ARTICLE
osteoblasts in the later stage of differentiation,39 whereas ALP activity suggested the early stage of differentiation. The same trend was also observed in our previous study.15 To investigate the potential of surface functionalization of BMP2 to induce the differentiation of MSCs, the expressions of OCN and OPN, which are used as the markers to indicate the osteogenic tendency,40 were visualized via immunocytochemical staining. It was performed on different substrates after 21 days of culture. MSCs grown on BMP2-PDOP-30 nm TiO2 nanotubes expressed much higher levels of OCN and OPN than those on samples of BMP2PDOP-Ti and BMP2-PDOP-100 nm TiO2 nanotubes. Moreover, for native samples, 30 nm TiO2 nanotubes displayed higher potential of differentiation induction for MSCs than native titanium and 100 nm TiO2 nanotubes. OCN and OPN are major noncollagenous bone matrix proteins.41 OCN, as a bone extracellular matrix vesicle, is related to the formation of mineralized tissue. It provides a protected environment for mineralization when connected to Ca2þ and hydroxyapatite. OPN, as a phosphorylated bone matrix sialoprotein, is believed to play an important role in cell adhesion and biomineralization.42 The biomineralization of MSCs was quantitatively measured by ARS staining (Figure 9). The result once again suggested that surface functionalization with BMP2 promoted the differentiation of MSCs. BMP2 is a member of BMP family, which is related to the transforming growth factor-β (TNF-β) superfamily.37 It was extensively reported to be a strong bone inducer during new bone formation and bone repair.43,44 The cell signaling cascade starts from the activation of BMP receptors, which is similar to that of TNF-β receptors.45 BMP receptors are composed of cell transmembrane serine and threonine protein kinases. When MSCs adhered to BMP2conjugated TiO2 nanotubes, the dimerization of the two serine/ threonine protein kinases was promoted. The threonine protein kinases phosphorylated the serine protein kinases. The activation of the serine protein kinases initialized phosphorylation of downstream effector proteins of receptor-regulated Smads (R-Smads), resulting in signal transduction.46,47 However, R-Smads’ affinity for DNA was relatively low. It commonly formed complexes with transcription factors to allow strong binding.46 A previous study confirmed that Runt-related transcription factor 2 (Runx2) formed a complex with R-Smads in vivo when activated by BMP2.48 Previous studies confirmed that Runx2 was critical for osteoblast differentiation.49,50 Therefore, surfaces presenting conjugated BMP2 of TiO2 nanotubes induced the differentiation of MSCs into osteoblasts.
’ CONCLUSIONS In this study, we fabricated surface-functionalized TiO2 nanotubes with BMP2 through the intermediate layer of polydopamine. XPS, SEM, and contact angle measurements demonstrated that BMP2 was successfully conjugated onto the TiO2 nanotubes. Carbon contamination was found regarding the prepared substrates with TiO2 nanotubes. The results of in vitro tests confirmed that surface-functionalized TiO2 nanotubes with BMP2 synergistically promoted the differentiation of MSCs. Furthermore, the study provided a platform to investigate synergistic effect of local chemistry and topography on the biological behaviors of cells. ’ AUTHOR INFORMATION Corresponding Author
*Tel: þ86-23-65102507. Fax: þ86-23-65102877. E-mail:
[email protected]. 1104
dx.doi.org/10.1021/bm1014365 |Biomacromolecules 2011, 12, 1097–1105
Biomacromolecules
’ ACKNOWLEDGMENT This work was financially supported by China Ministry of Science and Technology (973 project no. 2009CB930000), Natural Science Foundation of China (11032012), Fok Ying Tung Education Foundation (121035), Natural Science Foundation of Chongqing Municipal Government (CSTC, 2010AB5116, and 2008AB5129), MOE Key Laboratory of Macromolecular Synthesis and Functionalization, Zhejiang University (2010MSF01), “111 project”(B06023), and Fundamental Research Funds for the Central Universities (project no. CDJXS10 23 22 11). ’ REFERENCES (1) Stevens, M. M.; George, J. H. Science 2005, 318, 1135–1138. (2) Dalby, M. J.; Gadegaard, N.; Tare, R.; Andar, A.; Riehle, M. O.; Herzyk, P. Nat. Mater. 2007, 6, 997–1003. (3) Huang, J. H.; Graeter, S. V.; Corbellini, F.; Rinck, S.; Bock, E.; Kemkemer, R.; Kessler, H.; Ding, J. D.; Spatz, J. P. Nano Lett. 2009, 9, 1111–1116. (4) Matschegewski, C.; Staehlke, S.; Loeffler, R.; Lange, R.; Chai, F.; Kern, D. P. Biomaterials 2010, 31, 5729–5740. (5) Bechara, S. L.; Judson, A.; Popat, K. C. Biomaterials 2010, 31, 3492–3501. (6) Yim, E. K.; Reano, R. M.; Pang, S. W.; Yee, A. F.; Chen, C. S.; Leong, K. W. Biomaterials 2005, 26, 5405–5413. (7) St Pierre, C. A.; Chan, M.; Iwakura, Y.; Ayers, D. C.; Kurt-Jones, E. A.; Finberg, R. W. J. Orthop. Res. 2010, 29, 1418–1424. (8) Linder, L.; Carlsson, A.; Marsal, L.; Bjursten, L. M.; Branemark, P. I. J. Bone Jt. Surg., Br. Vol. 1988, 70, 550–555. (9) Azevedo, C. R. F. Eng. Failure Anal. 2003, 10, 153–164. (10) Balaur, E.; Macak, J. M.; Taveira, L.; Schmuki, P. Electrochem. Commun. 2005, 7, 1066–1070. (11) Zwilling, V.; Aucouturier, M.; Darque-Ceretti, E.; BoutryForveille, A. Electrochim. Acta 1999, 45, 921–929. (12) Macak, J. M.; Hildebrand, H.; Marten-Jahns, U.; Schmuki, P. J. Electroanal. Chem. 2008, 621, 254–266. (13) Park, J.; Bauer, S.; Schlegel, K. A.; Neukam, F. W.; von der Mark, K.; Schmuki, P. Small 2009, 5, 666–671. (14) Park, J.; Bauer, S.; von der Mark, K.; Schmuki, P. Nano Lett. 2007, 7, 1686–1691. (15) Hu, Y.; Cai, K. Y.; Zhang, R.; Luo, Z.; Yang, L.; Jandt, K. D.; Deng, L. H. Biomaterials 2009, 30, 3626–3635. (16) Karageorgiou, V.; Meinel, L.; Hofmann, S.; Malhotra, A.; Volloch, V.; Kaplan, D. J. Biomed. Mater. Res. 2004, 71(A), 528–537. (17) Benoit, D. S.; Collins, S. D.; Anseth, K. S. Adv. Func. Mater. 2007, 17, 2085–2093. (18) Kim, J. A.; Kim, I. S.; Cho, T. H.; Lee, K. B.; Hwang, S. J.; Tae, G. Biomaterials 2007, 28, 1830–1836. (19) Carstens, M. H.; Chin, M.; Li, X. J. J. Craniofacial Surg. 2005, 16, 1033–1042. (20) Pountos, I.; Corscadden, D.; Emery, P.; Giannoudis, P. V. Injury 2007, 38, 23–33. (21) Cai, K. Y.; Yao, K. D.; Lin, S. B.; Yang, Z. M.; Xie, H. Q.; Li, X. Q. Biomaterials 2002, 23, 1153–1160. (22) Cai, K. Y.; Yao, K. D.; Lin, S. B.; Yang, Z. M.; Xie, H. Q.; Li, X. Q. J. Biomed. Mater. Res. 2002, 60, 398–404. (23) Waite, J. H. Nat. Mater. 2008, 7, 8–9. (24) Cai, K. Y.; M€uller, M.; Bossert, J.; Rechtenbach, A.; Jandt, K. D. Appl. Surf. Sci. 2005, 250, 252–267. (25) Poh, C. K.; Shi, Z.; Lim, T. Y.; Neoh, K. G.; Wang, W. Biomaterials 2010, 31, 1578–1585. (26) Lee, H.; Dellatore, S. M.; Miller, W. M.; Messersmith, P. B. Science 2007, 318, 426–430. (27) Dalby, M. J.; Di Silvio, L.; Harper, E. J.; Bonfield, W. J. Mater. Sci.: Mater. Med. 2002, 13, 311–314. (28) Winkelmann, M.; Gold, J.; Hauert, R.; Kasemo, B.; Spencer, N. D.; Brunette, D. M.; Textor, M. Biomaterials 2003, 24, 1133–1145.
ARTICLE
(29) Jang, H. K.; Whangbo, S. W.; Kim, H. B. J. Vac. Sci. Technol., A 2002, 18, 917–921. (30) Lee, H.; Rho, J.; Messersmith, P. B. Adv. Mater. 2009, 21, 431–434. (31) Grigoriou, V.; Shapiro, I. M.; Cavalcanti-Adam, E. A.; Composto, R. J.; Ducheyne, P.; Adams, C. S. J. Biol. Chem. 2005, 280, 1733–1739. (32) Humphries, J. D.; Wang, P.; Streuli, C.; Geiger, B.; Humphries, M. J.; Ballestrem, C. J. Cell Biol. 2007, 179, 1043–1057. (33) Wen, K. K.; Rubenstein, P. A.; Demali, K. A. J. Biol. Chem. 2009, 284, 30463–30473. (34) Shah, A. K.; Lazatin, J.; Sinha, R. K.; Lennox, T.; Hickok, N. J.; Tuan, R. S. Biol. Cell. 1999, 91, 131–142. (35) Rai, B.; Teoh, S. H.; Ho, K. H.; Hutmacher, D. W.; Cao, T.; Chen, F.; Yacob, K. Biomaterials 2004, 25, 5499–5506. (36) Akino, K.; Mineta, T.; Fukui, M.; Fujii, T.; Akita, S. Wound Repair Regener. 2003, 11, 354–360. (37) ten Dijke, P. Curr. Med. Res. Opin. 2006, 22, S7–S11. (38) Oh, S.; Brammer, K. S.; Li, Y. S. J.; Teng, D.; Engler, A. J.; Chien, S.; Jin, S. Proc. Natl. Acad. Sci. U.S.A. 2009, 106, 2130–2135. (39) Yamamoto, Y.; Ohsaki, Y.; Goto, T.; Nakasima, A.; Iijima, T. J. Dent. Res. 2003, 82, 962–966. (40) Stein, G. S.; Lian, J. B. Endocr. Rev. 1993, 14, 424–442. (41) Ayukawa, Y.; Takeshita, F.; Inoue, T.; Yoshinari, M.; Shimono, M.; Suetsugu, T. J. Biomed. Mater. Res. 1998, 41, 111–119. (42) Boskey, A. L. Clin. Orthop. Relat. Res. 1992, 281, 244–274. (43) Yu, Y. Y.; Lieu, S.; Lu, C.; Colnot, C. Bone 2010, 47, 65–73. (44) Liu, T.; Gao, Y.; Sakamoto, K.; Minamizato, T.; Furukawa, K.; Tsukazaki, T.; Shibata, Y.; Bessho, K.; Komori, T.; Yamaguchi, A. J. Cell Physiol. 2007, 211, 728–735. (45) Wrana, J. L.; Attisano, L.; Wieser, R. Nature 1994, 370, 341–347. (46) ten Dijke, P.; Fu, J.; Schaap, P. J. Bone Jt. Surg., Am. Vol. 2003, 85-A, 34–38. (47) Miyazono, K. Bone 1999, 25, 91–93. (48) Hanai, J.; Chen, L. F.; Kanno, T. J. Biol. Chem. 1999, 274, 31577–31582. (49) Pungchanchaikul, P.; Bloch-Zupan, A.; Ferretti, P. Cells Tissues Organs 2010, 192, 283–291. (50) Lee, J. S.; Lee, J. M.; Im, G. I. Biomaterials 2011, 32, 760–768.
1105
dx.doi.org/10.1021/bm1014365 |Biomacromolecules 2011, 12, 1097–1105
