Materials Science and Engineering C 37 (2014) 28–36

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Materials Science and Engineering C journal homepage: www.elsevier.com/locate/msec

Electrical stimulation to promote osteogenesis using conductive polypyrrole films Wei-Wen Hu a,⁎, Yi-Ting Hsu a, Yu-Che Cheng b,c, Chuan Li d, Ruoh-Chyu Ruaan a,b, Chih-Cheng Chien c,e,f, Chih-Ang Chung d, Chia-Wen Tsao d a

Department of Chemical and Materials Engineering, National Central University, Jhongli City, Taiwan Institute of Biomedical Engineering, National Central University, Jhongli, Taiwan Department of Medical Research, Cathay General Hospital, Taipei, Taiwan d Department of Mechanical Engineering, National Central University, Jhongli, Taiwan e School of Medicine, Fu Jen Catholic University, Taipei, Taiwan f Department of Anesthesiology, Sijhih Cathay General Hospital, Sijhih City, Taipei, Taiwan b c

a r t i c l e

i n f o

Article history: Received 19 August 2013 Received in revised form 8 November 2013 Accepted 6 December 2013 Available online 19 December 2013 Keywords: Polypyrrole Osteogenesis Electrical stimulation Bone marrow stromal cells Mineralization Direct current field

a b s t r a c t In this study, we developed an electrical cell culture and monitoring device. Polypyrrole (PPy) films with different resistances were fabricated as conductive surfaces to investigate the effect of substrate-mediated electrical stimulation. The physical and chemical properties of the devices, as well as their biocompatibilities, were thoroughly evaluated. These PPy films had a dark but transparent appearance, on which the surface cells could be easily observed. After treating with the osteogenic medium, rat bone marrow stromal cells cultured on the PPy films differentiated into osteoblasts. The cells grown on the PPy films had up-regulated osteogenic markers, and an alkaline phosphatase activity assay showed that the PPy films accelerated cell differentiation. Alizarin red staining and calcium analysis suggested that the PPy films promoted osteogenesis. Finally, PPy films were subjected to a constant electric field to elucidate the effect of electrical stimulation on osteogenesis. Compared with the untreated group, electrical stimulation improved calcium deposition in the extracellular matrix. Furthermore, PPy films with lower resistances allowed larger currents to stimulate the surface cells, which resulted in higher levels of mineralization. Overall, these results indicated that this system exhibited superior electroactivity with controllable electrical resistance and that it can be coated directly to produce medical devices with a transparent appearance, which should be beneficial for research on electrical stimulation for tissue regeneration. © 2013 Elsevier B.V. All rights reserved.

1. Introduction Various cells have been proven to be susceptible to electrical stimulation and electrically assisted therapy has been applied to different tissues, such as nerves and muscles, to promote their regeneration [1–4]. Direct current field (DCF), capacitive coupling electrical field (CCEF), and electromagnetic field (EMF) are three frequently used forms of electric stimulation [5]. Of these, CCEF and EMF are noninvasive and have been studied widely. However, these treatments are not spatially specific so both normal and pathological tissues would be stimulated [6]. By contrast, DCF can be applied directly to a wound site and the range of stimulation can be confined to avoid unwanted side-effects [7]. Stimulation using electrodes mediated by a fluid is the simplest method for DCF treatment [8–13]. However, electrolysis may occur to change the local pH values [14]. In addition, the electric current may trigger a gradient ion distribution, which could affect the cell physiology ⁎ Corresponding author at: Department of Chemical and Materials Engineering, National Central University, No.300, Jhongda Rd., Jhongli City, Taoyuan County, Taiwan. Tel.: +886 3 422 7151x34243; fax: +886 3 422 5258. E-mail address: [email protected] (W.-W. Hu). 0928-4931/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.msec.2013.12.019

and increase the difficulty of clarifying the effects of DCF on cells [15]. These drawbacks all limit the mechanistic studies of the effects of DCF on cell differentiation. Substrate-mediated DCF is an alternative approach where cells are cultured on conductive material surfaces for in situ treatment. The electric current passes through the cell substrate; thus, electrolysis and ion movement can be avoided. To simulate an in vivo environment, the electric resistance of conductive materials should be similar to those of real tissues. Different materials have been used for substrate-mediated DCF treatments, but metal is the most popular because of its good conductivity [16–18]. However, compared with normal tissues, the extremely low resistance of metallic materials leads to the passage of a high electric current during stimulation, which may negatively affect cells. The corrosion of metal may also occur during DCF treatment, which can release cytotoxic debris [19]. To overcome these difficulties, composite materials have been developed to produce biocompatible surfaces with appropriate resistivity. Multiwalled carbon nanotubes have been embedded in a polylactide matrix to electrospin conductive nanofibers [20]. Polypyrrole (PPy) has also been doped into biodegradable polymers to prepare conductive films [21,22]. These materials have good conductivity and can be used for substrate-mediated DCF treatment,

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but they are all opaque so the surface cells cannot be observed directly. Thus, fluorescent staining and scanning electron microscopy are required for visualization, whereas the cell morphology is probably affected during fixation. In addition, end-point observations of these systems are impossible to continuously monitor surface cells. These problems suggest that the preparation of transparent conductive materials with adjustable resistivity would be beneficial for the research on the effects of DCF on cell differentiation. Therefore, we aimed to develop a convenient device that can be used easily to investigate the effect of electrical stimulation on surface cells. Conductive PPy films were polymerized via chemical oxidation to deposit them on substrate surfaces. The electrical resistance of PPy films was regulated by controlling the monomer concentrations. These films were biocompatible and had a transparent appearance so the surface cells could be observed easily. Rat bone marrow stromal cells (rBMSCs) were used as a model system to test our devices. The differentiation of rBMSCs was accelerated on these electroactive surfaces. Next, the PPy films were applied to a homemade device before DCF treatment. The mineralization of rBMSCs was improved greatly after subjecting them to electric fields, which suggests that this device can promote osteogenesis. These results support that this substrate-mediated DCF system is suitable for performing osteogenesis and it has potential applications in other areas of tissue differentiation research. 2. Materials and methods 2.1. Materials Pyrrole and ammonium persulfate were purchased from Acros (Geel, Belgium) and Showa (Tokyo, Japan), respectively. Fetal bovine serum (FBS), Dulbecco's modified Eagle medium (D-MEM), and trypsin-EDTA were obtained from Biowest (Nuaillé, France). Dexamethasone, 2phospho-L-ascorbic acid trisodium salt, β-glycerophosphate disodium salt hydrate, Triton X-100, and glutaraldehyde were purchased from Sigma-Aldrich (St Louis, MO, USA). 2.2. The preparation of PPy films Polystyrene (PS) Petri dishes with a diameter of 35 mm (Nunc, Penfield, NY, USA) were used as the substrate for PPy film deposition. Pyrrole was dissolved in water at concentrations of 0.1, 0.3, and 0.5 M. Aqueous solutions of oxidant ammonium persulfate were prepared using the corresponding pyrrole solutions so the molar ratio of pyrrole relative to ammonium persulfate was always 5:1 (i.e. 0.02, 0.06, and 0.1 M). Next, 2 ml of pyrrole and ammonium persulfate solutions were added to Petri dishes, with gently mixing for 15 min at room temperature. Finally, these films were washed with deionized (DI) water and dried in an oven. 2.3. Scanning electron microscopy (SEM) The morphologies of the PPy films were visualized by SEM (3500N, Hitachi, Japan) after gold sputtered-coating. To determine the film thickness, the films were cryofractured in liquid nitrogen to produce cross-sections. 2.4. Characterization of PPy films To characterize the deposited PPy films, Fourier transform infrared (ATR-FTIR) spectroscopy (Spectrum 100, PerkinElmer, Waltham, MA, USA) was used to obtain the IR spectra at a resolution of 1 cm−1. In addition, elemental analysis of the PPy films was performed using X-ray photon-electron spectrometry (XPS, K-alpha, Thermo) where the binding energy was measured from 0 to 1100 eV.

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2.5. Four-point probe analysis Four-point probe (EverBeing, Hsinchu, Taiwan) analysis was used to determine the sheet resistance of PPy films. Twenty points in different regions were examined in each film to confirm the spatial stability of the sheet resistance. Furthermore, the resistivities of PPy films were converted using the thickness measurements derived from the SEM results. 2.6. In vitro cell culture of rBMSCs Rat bone marrow stromal cells (rBMSCs) harvested from the 8week-old Sprague–Dawley rats were used in this study to simulate osteogenesis. These cells were maintained in regular medium of DMEM with 10% FBS. Osteogenic supplements (100 μm ascorbic-2-phosphate, 10 mM β-glycerophosphate, and 100 nM dexamethasone) were added to the regular medium during osteogenesis experiments. 2.7. Biocompatibility of PPy films Polypropylene rings were glued onto PPy films to create wells. To evaluate the biocompatibility of PPy films, rBMSCs were seeded onto PPy films at a density of 17,000 cells/cm2, and an (4,5-cimethylthiazol2-yl)-2,5-diphenyl tetrazolium bromide (MTT) assay was performed to quantify cell viability. After culture for 2 days, 20 μl of MTT solution (5 mg/ml in phosphate-buffered saline (PBS)) and 180 μl of medium were added to each well for 3 h at 37 °C. The supernatant was removed and 200 μl of dimethyl sulfoxide (DMSO) was added to dissolve the formazan, which was analyzed spectrophotometrically at a wavelength of 550 nm. A lactate dehydrogenase (LDH) assay was used to quantify the cell numbers with CytoTox 96 Non-Radioactive Cytotoxicity Assay (Promega). Briefly, 100 μl of fresh medium was replaced in each well before the assay and 15 μl of lysis buffer was added to sample for 1 h at 37 °C to release LDH from live cells. After transferring 50 μl of the LDH-released medium to 96-well multiplates, 50 μl of LDH reagent was added, which was followed by incubation for 30 min at room temperature. Finally, 50 μl of stop solution was added to each well and they were analyzed spectrophotometrically at a wavelength of 490 nm. A standard curve was also produced by lysing known cell amounts, which was used to convert the absorbance into a total cell number. 2.8. Quantitative and qualitative analyses of calcium deposition in the extracellular matrix (ECM) To quantify the level of calcium deposition, a colorimetric assay was performed using the calcium-(o-cresolphthalein complexone) (Ca-OCPC) complex method [23]. Before the assay, the osteogenic medium was removed from the well using two washes of PBS. Next, 100 μl per well of 0.5 N acetic acid was used to release calcium. Ten microliters of calcium-released sample was added to 200 μl of calcium-binding reagent (0.1 g/l of o-cresolphthalein complexone and 1 g/l of 8-hydroxyquinoline) and 200 μl of buffer reagent (1.6 M of 2-amino-2-methyl-1-propanol, pH 10.7). After 15 min incubation at room temperature, 100 μl of purple-colored Ca-OCPC complex was transferred to 96-well multiplates, which was quantified based on the absorbance at a wavelength of 575 nm. The amount of calcium in cell lysate was converted based on the linear calibration results obtained using calcium chloride standard solutions. Before alizarin red S staining, the cultures were rinsed with PBS and fixed (1% glutaraldehyde in PBS) for 30 min at 37 °C, then stained with 2% alizarin red S solution for 20 min at room temperature. After washing with PBS, the stained samples were observed using an inverted microscope (Eclipse Ti-U, Nikon, Japan).

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2.9. Alkaline phosphatase activity (ALP) analysis Before the ALP assay, the osteogenic medium was removed from the well with two washes of PBS. Cell lysis buffer (0.2% of Triton X-100 in 1 M diethanolamine buffer) was added at 50 μl/well. Repeated cycles of freezing and thawing were applied to achieve cell lysis. The substrate p-nitrophenyl phosphate (pNPP) was added at 50 μl/well. After 30 min incubation at 37 °C, 50 μl of 0.3 M NaOH was added to stop the reaction and the absorbance was measured at 405 nm. 2.10. Real-time polymerase chain reaction (RT-PCR) Total RNA was isolated from cells using an RNeasy Mini Kit (Qiagen, Hilden, Germany) according to the manufacturer's instructions. Approximately 2 μg of total RNA was reverse-transcribed using the Superscript III First-strand system (Invitrogen, Carlsbad, CA, USA). The transcribed cDNA was amplified by TaqMan-polymerase chain reaction (ABI, Foster city, CA, USA) and monitored using an ABI 7300 Real-Time PCR System. The following primer pairs related to the targeted RNA were used in this study: β-actin, 5′-CTGGCTCCTAGCACCATGA-3′ and 5′-TAGAGCCA CCAATCCACACA-3′; core binding factor 1 (Cbfa 1), 5′-CCACAGAGCTAT TAAAGTGACAGTG-3′ and 5′-AACAAACTAGGTTTAGAGTCATCAAGC-3′; osteocalcin (OCN), 5′-ATAGACTCCGGCGCTACCTC-3′ and 5′-CCAGGGGA TCTGGGTAGG-3′. Roche Universal ProbeLibrary Probes were used in this study (β-actin: UPL #63; Cbfa1: UPL #98; OCN: UPL #125). cDNA was analyzed to determine the genes of interest and a housekeeping gene (β-actin) in independent reactions, and the cDNA levels were normalized relative to that of β-actin. Briefly, ΔCt was the differences in the cycle number at the threshold fluorescence level between the genes of interest and β-actin, which was applied to normalize the transcription levels (2−ΔCt). These results were compared with the corresponding results harvested from cells cultured on tissue culture polystyrene (TCPS) in regular medium without osteogenic supplements [24]. 2.11. Electrical stimulation To apply the DCF to rBMSCs on PPy films, a homemade electrical cell culture and monitoring device was developed. PPy films deposited in 35 mm dishes were trimmed to produce rectangular specimens that measured 3.0 cm × 1.8 cm. Three polypropylene rings with diameters of 6 mm and height of 8 mm were glued to the PPy films. Two stainless steel electrodes were placed at two ends of the PPy films and several electrodes were connected in parallel to an external power resource (Regulated DC power supply, Hola). One day before the DCF treatment, the rBMSCs were seeded onto the PPy films. The culture medium was replaced with the osteogenic medium and constant electric fields were applied to the PPy films in the homemade devices. To evaluate the effect of electrical stimulation on cell viability, a MTT assay and morphological observations were performed 1 day after the DCF treatment. The cells were kept in culture until the analysis and the osteogenic medium was refreshed every other day. The mineralization levels were determined on Day 14 using the Ca-OCPC complex method and alizarin red S staining. 2.12. Statistical analysis The statistical analyses were performed with SPSS (Chicago, IL, USA). A two-tailed Student's t-test was performed to make comparison and the errors were reported as standard deviations. 3. Results and discussions 3.1. Fabrication of PPy films Different concentrations of pyrrole, i.e. 0.1 M, 0.3 M, and 0.5 M, were used to fabricate PPy films, which were denoted as PPy0.1, PPy0.3, and

PPy0.5, respectively. The films that produced by more pyrrole had a darker appearance. All of the films were dark but transparent (Fig. 1a). Compared with most conductive materials with opaque surfaces, these transparent films can be easily applied for direct observations of surface cell morphology. Cross-sections of the PPy films were visualized by SEM. All of the films were dense and their thickness increased with the pyrrole concentrations used during polymerization (Fig. 1b). To characterize the chemical structures of PPy films, ATR–FTIR and XPS analyses were performed (Fig. 2). In the IR spectra, there were two specific peaks at 1690 and 1360 cm− 1, which were ascribed to the C_N and the C\N groups, respectively [25] (Fig. 2a). The peak at 1160 cm−1 was attributed to its O_N vibration [25]. The absorptions at 1475 and 1550 cm−1 were due to the stretching vibrations of C_C and pyrrole ring, respectively [26,27]. In addition, the absorption of S_O at 1050 cm−1 was attributed to the initiator ammonium persulfate [28]. In the XPS analysis, the peaks with binding energies of 280– 290, 370–400, and 540–550 eV corresponded to the presence of carbon, nitrogen, and oxygen elements, respectively (Fig. 2b). These results demonstrated that the PPy films were fabricated successfully. 3.2. Electrical properties of PPy films To facilitate substrate-mediated DCF treatment, the resistance of a biomaterial should be as similar as possible to native tissues to mimic the in vivo environment. In this study, bone was used as the model system to test the developed device. Faes et al. reported that the resistivity of tibial cancellous bone is about 4.75 Ω-m [29]. Therefore, the fabricated PPy films needed to be at a similar level. To manipulate the resistance of PPy films, different concentrations of pyrrole were used for polymerization. A four-point probe analysis was performed to determine the sheet resistance of PPy films, which was the reciprocal of the sheet conductance (Fig. 3a). The thickness of PPy films (Fig. 1b) was used to convert the results into resistivities and conductivities (Fig. 3b). Higher concentrations of pyrrole produced PPy films with lower resistivities and higher conductivities. Most of the conductive materials used for electrical stimulation have a sheet resistance of 1–1000 kΩ and a conductivity of 1–10−3 S/cm; hence, our fabricated PPy films should be appropriate for physiological requirements [22,30–35]. 3.3. Biocompatibilities of PPy films Conductive polymers have been proved that electroactive surfaces may increase intercellular communication to promote cell proliferation and adhesion [36–39]. Therefore, we evaluated whether our synthetic PPy films were biocompatible. The cell number and cell viability of rBMSCs on PPy films were evaluated using LDH and MTT assays, respectively (Fig. 4a). The numbers of cell on PPy films were slightly less than that on TCPS, but the differences were not significant (p N 0.1). Similarly, the MTT results suggested that cells grown on PPy films maintained their relative viabilities of N80%, and there were no significant differences between PPy films. The PPy films were transparent; therefore, the surface cell morphologies could be easily observed, which showed that they were almost not different from those on TCPS (Fig. 4b). These results suggested that PPy films may maintain cell viability without cytotoxicity. 3.4. Effect of conductive surfaces on the osteogenesis of rBMSCs Various studies have indicated that electroactive surfaces may promote cell differentiation [36,40,41]. Our results also suggested that PPy films were biocompatible (Fig. 4). It should be interesting to evaluate their effects on cell differentiation. Therefore, rBMSCs were seeded onto PPy films for 1 day, and the osteogenic medium was used for the subsequent culture. Because osteoblasts that differentiated from rBMSCs can deposit calcium in the ECM, the level of calcium in the

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(a) PPy0.1

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Fig. 1. PPy film deposition. (a) PPy films were fabricated using different concentrations of pyrrole. All of the PPy films were dark but transparent. (b) SEM pictures showing cross-sections of PPy films (scale bar = 50 μm).

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Binding Energy (eV) Fig. 2. Characteristics of PPy films. (a) ATR-FTIR spectra of PPy films. (b) XPS analysis results.

ECM is an important index of osteogenesis [42]. In the present study, the Ca-OCPC complex method and alizarin red S staining were used for quantitative and qualitative analyses of mineralization. For the Ca-OCPC assay, the deposited calcium was analyzed based on the absorbance of a purple-colored complex (Fig. 5a). Calcium was undetectable in all groups on Day 7. However, there was a significant increase in calcium deposition in all PPy films compared with the control TCPS group on Day 9. On Day 14, the trend was more obvious and the calcium level increased with increasing pyrrole concentrations. The highest calcium deposition in the PPy films (PPy0.5) on Day 14 was the three times that found in the TCPS group. The calcium level continued to increase in the TCPS group on Day 28, but the increase was saturated in the other PPy groups. However, all of the PPy groups still contained more calcium in ECM than that of the TCPS group. The calcium deposition rate was also normalized relative to the surface cell number to elucidate the relationship between cell proliferation and mineralization (Fig. 5b). The trends in the normalized calcium deposition were similar to those of the total deposition on Days 7, 9, and 14, which suggest that improvements in the calcium levels were not affected by cell numbers. Interestingly, because the mature bone matrix resulted in differentiated cell death, a lower cell number led to a higher normalized calcium deposition on Day 28. Alizarin red S staining was applied on Day 14 to confirm the Ca-OCPC assay results (Fig. 5c). Films fabricated by more pyrrole had higher calcium deposition levels, and all of the PPy films exhibited denser staining than that of the TCPS group. These results were similar to those obtained with the Ca-OCPC assay, supporting that these electroactive surfaces improved osteogenesis. To elucidate the improvement of the conductive surfaces for osteogenesis, we analyzed the mRNA and protein expression levels of osteogenic-specific markers. The early marker alkaline phosphatase (ALP) was quantified using substrate pNPP to determine its expression by differentiated rBMSCs (Fig. 6a). The highest ALP expressions of rBMSCs on PPy films all occurred earlier than those on the TCPS, which suggests that these electroactive surfaces accelerated cell differentiation.

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Fig. 3. Electrical properties of PPy films. (a) The sheet resistances of PPy films were analyzed by four-point probe analysis, and their reciprocals were the sheet conductance. (b) The resistivity and conductivity of PPy films were determined based on the corresponding film thickness measured by SEM.

Cbfa1 and OCN are two osteo-related markers which are expressed during early-middle and late periods of osteogenesis, respectively [43]. The transcription levels of these two genes were quantified using RT-PCR on Day 7 and Day 14 (Fig. 6b). There was a higher level of Cbfa1 on Day 7 than on Day 14 because of its early function during osteogenesis [44]. In contrast to the TCPS group, where there was a slight increase, Cbfa1 mRNA was greatly upregulated in all PPy groups on Day 7. Cbfa1 is a transcriptional factor that regulates osteoblast differentiation; thus, the fact that the rBMSCs had high levels of Cbfa1 suggests that the PPy films promoted osteogenesis. By contrast, OCN was not transcribed on Day 7, but it was upregulated on Day 14. Because OCN is regulated by Cbfa1, the high levels of Cbfa1 in the PPy groups led to the subsequent upregulation of OCN [45]. The trends in the mRNA levels of Cbfa1 and

OCN were similar to the calcium deposition results, which showed that the PPy films were superior to the TCPS surfaces and their higher conductivity led to greater mRNA transcription. These results suggest that these electroactive PPy films are suitable for eliciting the cell differentiation of osteoblast lineages. 3.5. The electrical stimulation of rBMSCs on PPy films Wolff's law is the gold standard for bone regeneration and it states that mineralization can be induced to resist a mechanical load. This presents the interesting question of how osteoblasts and osteoclasts sense the mechanical force. Collagen is the main matrix protein in mineralized tissue. It has been shown that stress-induced strain can

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Fig. 4. Biocompatibility of PPy films. (a) An MTT assay was applied to evaluate the bioactivity of rBMSCs cultured on PPy films and the results obtained by TCPS were defined as the control group to determine the relative viability. In addition, the surface cell numbers were evaluated using a LDH assay. (*: p b 0.05, **: p b 0.01 compared with TCPS). (b) The morphologies of rBMSCs were observed 1 day after seeding on PPy films (scale bar = 100 μm).

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Fig. 5. Calcium depositions by rBMSCs on PPy films. (a) After treatment with the osteogenic medium, the rBMSCs cultured on different PPy films were analyzed to determine the calcium deposition levels in the ECM using the Ca-OCPC complex method. (b) The deposited calcium was normalized against the surface cell number, which was determined using a LDH assay. (c) Alizarin red S staining was used to determine the level and distribution of mineralization. (*: p b 0.05, **: p b 0.01 compared with TCPS) (scale bar = 100 μm).

generate potential gradients along collagen fibers to provide a local stimulus for bone-generating cells [46]. This piezoelectric effect suggests that an electrical cue may be the stimulus for mechanically loaded bone. Thus, electric stimulation has been studied as a physical cue for promoting bone regeneration [47]. Therefore, DCF treatment has been shown to accelerate bone regeneration in various osseous defects, such as callus formation in delayed union and spine fusion [48–50]. In this study, osteogenesis was used as the model to test our system. We developed a homemade device where polypropylene rings were glued onto PPy films to create wells for cell seeding and rBMSCs were placed 1 day before the experiments (Fig. 7). After changing to the osteogenic medium, two electrodes were applied to two ends of the PPy films with a distance of 2.86 cm and they were connected to a power supply to apply the DCF treatment. Before the in vitro experiments, PBS was added to the wells and 10 V DCF was applied for 12 h to ensure that the electrical properties of the PPy films were satisfactory at physiological pH. There were no significant differences in the resistance of PPy films before and after DCF treatment, suggesting that the film conductivity was stable throughout electrical stimulation (data not shown). To determine the appropriate parameters for electrical stimulation, PPy0.5 was used as a platform for DCF treatment. Voltages of 10, 1, and 0.1 V were applied to create electrical fields of 3.5, 0.35, and 0.035 V/cm, respectively, and the durations of DCF treatment were 2, 4, and 12 h. A MTT assay was performed after 1 day of the treatment to confirm whether the DCF treatment damaged the surface cells, and the results were normalized based on the untreated group to determine their relative viabilities (Fig. 8a). The DCF treatment using 0.035 and 0.35 V/cm for 2 h and 4 h produced similar MTT results to the untreated group, whereas the viabilities at 12 h decreased to 75–80%, thereby suggesting that long-term DCF treatment appeared to damage the rBMSCs. In addition, the cells treated with 3.5 V/cm were greatly reduced based on their MTT results. The cell morphologies also had a similar trend that cells were normal after treatment with 0.035 and 0.35 V/cm DCF for 2 h

and 4 h (Fig. 8b). However, cell extension was slightly reduced when the treatment was performed for 12 h. For the 3.5 V/cm groups, numerous dead cells were found on the films and the residual cells were aggregated as clumps even for 2 h treatment. Furthermore, some bubbles were found with the 3.5 V/cm treatment, which suggests that this electric field may have been too high to allow cell survival. After 14 day of culture, the mineralization level was analyzed using the Ca-OCPC complex method and alizarin red S staining. Compared with the rBMSCs grown on PPy films without DCF treatment, the calcium deposition levels were improved using electrical fields of 0.35 and 0.035 V/cm (Fig. 8c). However, calcium was almost undetectable when 3.5 V/cm was applied. With the 2 h DCF treatment, the calcium level was slightly increased but the difference was not significantly compared with the control group. By contrast, the level of calcium in the ECM was greatly enhanced when the DCF treatment was increased to 4 h. However, the calcium deposition rate was decreased when the duration of DCF treatment was increased to 12 h. The cell morphology after the 12-h treatment had shrunk to some extent (Fig. 8b), which demonstrates that long-term DCF treatment probably has a negative effect on rBMSCs. The corresponding alizarin red S staining also detected similar trends where the deep-stained calcium-containing tissues were found at the 4-h DCF treatment (Fig. 8d). Because rBMSCs treated with 0.35 V/cm for 4 h had the highest calcium deposition, these electrical parameters were also used to test the electrical stimulation effects of PPy films with different resistivities. After DCF treatment, the rBMSCs cultured on PPy films all had higher calcium levels in the ECM compared with the untreated groups (Fig. 8e). The percentage improvement increased with the film conductivity where the levels of calcium deposited on the PPy films prepared with 0.1, 0.3, and 0.5 M pyrrole were 151%, 175%, and 196% of the levels of the untreated groups, respectively. This may have been because the PPy films with lower resistances allowed higher currents to pass through the membrane, which enhanced the extent of mineralization.

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Fig. 6. Evaluation of the osteogenic markers of rBMSCs on PPy films. (a) The expression of alkaline phosphate (ALP) was measured using substrate pNPP. (b) Real-time PCR was applied to quantify the mRNA levels of Cbfa1 and OCN in rBMSCs at Days 7 and 14. The results obtained with rBMSCs without the osteogenic medium treatment (w/o OM) were applied as the baseline to determine the increases in the intracellular mRNA levels. (*: p b 0.05, **: p b 0.01 compared with the TCPS group treated with the osteogenic medium).

Fig. 7. Homemade electrical cell culture device used to apply electrical stimulation via PPy films. PPy films prepared on polystyrene surfaces were trimmed to produce rectangular specimens of 3.0 × 1.8 cm and three polypropylene rings were glued to create wells for cell seeding. Two stainless steel electrodes were placed at two ends of the PPy films and several electrodes were connected in parallel to an external power source.

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PPy0.3

PPy0.5

Fig. 8. Effect of the application of a DCF on osteogenesis. Different DCFs were applied to PPy films for various durations. (a) An MTT assay was conducted at 1 day after the DCF treatment and the result obtained using PPy0.5 without DCF treatment was defined as the control group to determine the relative viability. (b) Surface cell morphologies after DCF treatment. (c) After DCF treatment, rBMSCs were cultured for 14 days and analyzed to determine their calcium deposition levels in the ECM using the Ca-OCPC complex method. (d) Alizarin red S staining was used to visualize the level and distribution of mineralization. (e) rBMSCs cultured on different PPy films were treated with DCF (0.35 V/cm) for 4 h. After 14 days of culture, the Ca-OCPC complex method was applied to analyze the levels of calcium deposition in the ECM. (*: p b 0.05, **: p b 0.01 compared with PPy films without DCF treatment) (scale bar = 100 μm).

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Therefore, the level of calcium can be regulated by controlling the resistance of the PPy films. In this study, the highest mineralization increase induced by the DCF treatment was significantly greater than that in the TCPS group (fivefold higher on Day 14, compared with Fig. 5a). Previous studies suggest that DCF administration can facilitate bone tissue regeneration [16,17,20–22]. It has also been reported that DCF treatment may increase the intracellular concentration of calcium ion [15,51]. Increased levels of free calcium ions can activate calmodulin to trigger protein kinase and calcineurin; thus, the differentiation of mesenchymal stem cells may be enhanced to facilitate the formation of mineralized tissue [52,53]. Although the improvement of osteogenesis via electrical stimulation has been demonstrated, optimizing the treatment remains a challenge. In this study, we developed an electrical culture system where appropriate DCF conditions can be specified and the conductivity of the PPy films could be adjusted according to the physiological requirements. This easily monitored device should be a useful tool for elucidating the effects of DCF treatment on cell differentiation. 4. Conclusions In this study, we developed a practical substrate-mediated DCF in vitro cell culture system. Using oxidant chemical polymerization, biocompatible PPy films were fabricated with a dark but transparent appearance which can be easily applied to access the morphology of surface cell. Bone marrow stromal cells were grown on the conductive PPy films, and they exhibited a superior osteogenic capacity. The DCF treatment also greatly promoted mineralization, and the optimal DCF treatment was 0.35 V/cm for 4 h. The osteo-differentiation of rBMSCs can be improved significantly by controlling the preparation of PPy films and level of electrical stimulation,. Furthermore, our developed PPy film fabrication could be easily deposited to different medical devices, and their resistances can be adjusted according to the requirements. These properties suggested that PPy-mediated electrical stimulation has potential applications in tissue regeneration. Acknowledgment This research was supported by the grant of NSC 101-2221-E-008090-MY2 from the National Science Council of Taiwan, and 101CGHNCU-A2 from the National Central University and the Cathy General Hospital Joint Research Center. References [1] L. Ghasemi-Mobarakeh, M.P. Prabhakaran, M. Morshed, M.H. Nasr-Esfahani, H. Baharvand, S. Kiani, S. Al-Deyab, S. Ramakrishna, J. Tissue Eng. Regen. Med. 5 (2011) E17–E35. [2] K.T. Ragnarsson, Spinal Cord 46 (2008) 255–274. [3] P. Decherchi, E. Dousset, T. Marqueste, F. Berthelin, F. Hug, Y. Jammes, L. Grelot, Sci. Sports 18 (2003) 253–263. [4] P.H. Peckham, J.S. Knutson, Annu. Rev. Biomed. Eng. 7 (2005) 327–360. [5] M. Griffin, A. Bayat, Eplasty 11 (2011) e34. [6] J.C. Gan, P.A. Glazer, Eur. Spine J. 15 (2006) 1301–1311. [7] C.T. Brighton, Z. Friedenberg, E.I. Mitchell, R.E. Booth, Clin. Orthop. Relat. Res. (1977) 106–123. [8] I.S. Kim, J.K. Song, Y.M. Song, T.H. Cho, T.H. Lee, S.S. Lim, S.J. Kim, S.J. Hwang, Tissue Eng. Part A 15 (2009) 2411–2422. [9] I.S. Kim, J.K. Song, Y.L. Zhang, T.H. Lee, T.H. Cho, Y.M. Song, D.K. Kim, S.J. Kim, S.J. Hwang, Biochim. Biophys. Acta, Mol. Cell. Res. 1763 (2006) 907–916.

[10] K.E. Hammerick, A.W. James, Z.B. Huang, F.B. Prinz, M.T. Longaker, Tissue Eng. Part A 16 (2010) 917–931. [11] N. Ozkucur, T.K. Monsees, S. Perike, H.Q. Do, R.H.W. Funk, PLoS One 4 (2009). [12] A.K. Dubey, S.D. Gupta, B. Basu, J. Biomed. Mater. Res. Part B 98 (2011) 18–29. [13] B. Ercan, T.J. Webster, Biomaterials 31 (2010) 3684–3693. [14] A.N. Zengo, C.A. Bassett, G. Prountzos, R.J. Pawluk, A. Pilla, J. Dent. Res. 55 (1976) 383–390. [15] Q. Wang, S.Z. Zhong, O.Y. Jun, L.X. Jiang, Z.K. Zhang, Y. Xie, S.Q. Luo, Clin. Orthop. Relat. Res. (1998) 259–268. [16] S. Bodhak, S. Bose, W.C. Kinsel, A. Bandyopadhyay, Mater. Sci. Eng. C Mater. Biol. Appl. 32 (2012) 2163–2168. [17] B.M. Isaacson, L.B. Brunker, A.A. Brown, J.P. Beck, G.L. Burns, R.D. Bloebaum, J. Biomed. Mater. Res. B 97B (2011) 190–200. [18] S.D. McCullen, J.P. McQuilling, R.M. Grossfeld, J.L. Lubischer, L.I. Clarke, E.G. Loboa, Tissue Eng. C 16 (2010) 1377–1386. [19] R.A. Gittens, R. Olivares-Navarrete, R. Tannenbaum, B.D. Boyan, Z. Schwartz, J. Dent. Res. 90 (2011) 1389–1397. [20] S.J. Shao, S.B. Zhou, L. Li, J.R. Li, C. Luo, J.X. Wang, X.H. Li, J. Weng, Biomaterials 32 (2011) 2821–2833. [21] J. Zhang, K.G. Neoh, X. Hu, E.T. Kang, W. Wang, Biotechnol. Bioeng. 110 (2013) 1466–1475. [22] S. Meng, Z. Zhang, M. Rouabhia, J. Bone Miner. Metab. 29 (2011) 535–544. [23] H. Castano, E.A. O'Rear, P.S. McFetridge, V.I. Sikavitsas, Macromol. Biosci. 4 (2004) 785–794. [24] S. Kim, S.S. Kim, S.H. Lee, S.E. Ahn, S.J. Gwak, J.H. Song, B.S. Kim, H.M. Chung, Biomaterials 29 (2008) 1043–1053. [25] K. Arora, A. Chaubey, R. Singhal, R.P. Singh, M.K. Pandey, S.B. Samanta, B.D. Malhotra, S. Chand, Biosens. Bioelectron. 21 (2006) 1777–1783. [26] M.A. Chougule, S.G. Pawar, P.R. Godse, R.N. Mulik, S. Sen, V.B. Patil, Soft Nanosci. Lett. 1 (2011) 6–10. [27] H.J. Kharat, K.P. Kakde, P.A. Savale, K. Datta, P. Ghosh, M.D. Shirsat, Polym. Adv. Technol. 18 (2007) 397–402. [28] V. Shaktawat, K. Sharma, N.S. Saxena, J. Ovonic Res. 6 (2010) 239–245. [29] T.J. Faes, H.A. van der Meij, J.C. de Munck, R.M. Heethaar, Physiol. Meas. 20 (1999) R1–R10. [30] G.X. Shi, M. Rouabhia, S.Y. Meng, Z. Zhang, J. Biomed. Mater. Res. Part A 84A (2008) 1026–1037. [31] G.X. Shi, M. Rouabhia, Z.X. Wang, L.H. Dao, Z. Zhang, Biomaterials 25 (2004) 2477–2488. [32] G.X. Shi, Z. Zhang, M. Rouabhia, Biomaterials 29 (2008) 3792–3798. [33] P. Moroder, M.B. Runge, H.A. Wang, T. Ruesink, L.C. Lu, R.J. Spinner, A.J. Windebank, M.J. Yaszemski, Acta Biomater. 7 (2011) 944–953. [34] A.S. Rowlands, J.J. Cooper-White, Biomaterials 29 (2008) 4510–4520. [35] A. Kotwal, C.E. Schmidt, Biomaterials 22 (2001) 1055–1064. [36] P.R. Bidez, S.X. Li, A.G. MacDiarmid, E.C. Venancio, Y. Wei, P.I. Lelkes, J. Biomater. Sci. Polym. Ed. 17 (2006) 199–212. [37] M. Onoda, Y. Abe, K. Tada, Thin Solid Films 519 (2010) 1230–1234. [38] H.J. Wang, L.W. Ji, D.F. Li, J.Y. Wang, J. Phys. Chem. B 112 (2008) 2671–2677. [39] Q.S. Zhang, Y.H. Yan, S.P. Li, T. Feng, Mater. Sci. Eng. C Mater. Biol. Appl. 30 (2010) 160–166. [40] V. Lundin, A. Herland, M. Berggren, E.W. Jager, A.I. Teixeira, PLoS One 6 (2011) e18624. [41] E. Mooney, P. Dockery, U. Greiser, M. Murphy, V. Barron, Nano Lett. 8 (2008) 2137–2143. [42] G.S. Stein, J.B. Lian, J.L. Stein, A.J. Van Wijnen, M. Montecino, Physiol. Rev. 76 (1996) 593–629. [43] F.J. Hughes, W. Turner, G. Belibasakis, G. Martuscelli, Periodontology 2000 (41) (2006) 48–72. [44] H. Perinpanayagam, T. Martin, V. Mithal, M. Dahman, N. Marzec, J. Lampasso, R. Dziak, Arch. Oral Biol. 51 (2006) 406–415. [45] R.T. Franceschi, G.Z. Xiao, D. Jiang, R. Gopalakrishnan, S.Y. Yang, E. Reith, Connect. Tissue Res. 44 (2003) 109–116. [46] E. Fukada, I. Yasuda, J. Phys. Soc. Jpn. 12 (1957) 1158–1162. [47] A.C. Ahn, A.J. Grodzinsky, Med. Eng. Phys. 31 (2009) 733–741. [48] S. Inoue, T. Ohashi, R. Imai, M. Ichida, I. Yasuda, Clin. Orthop. Relat. Res. (1977) 92–96. [49] D.C. Paterson, G.N. Lewis, C.A. Cass, Clin. Orthop. Relat. Res. (1980) 117–128. [50] W.J. Kane, Spine 13 (1988) 363–365. [51] G.J. Bourguignon, W. Jy, L.Y.W. Bourguignon, J. Cell. Physiol. 140 (1989) 379–385. [52] L. Sun, H.C. Blair, Y.Z. Peng, N. Zaidi, O.A. Adebanjo, X.B. Wu, X.Y. Wu, J. Iqbal, S. Epstein, E. Abe, B.S. Moonga, M. Zaidi, Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 17130–17135. [53] M. Zayzafoon, K. Fulzele, J.M. McDonald, J. Biol. Chem. 280 (2005) 7049–7059.

Electrical stimulation to promote osteogenesis using conductive polypyrrole films.

In this study, we developed an electrical cell culture and monitoring device. Polypyrrole (PPy) films with different resistances were fabricated as co...
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