Bare-metal stents (diameter, 2 mm; length, 20 mm) were purchased from Dalian Yinyi Biomaterials Development Co., Ltd. (Dalian, China). Sunitinib was obtained from CST Biological Reagents Co., Ltd. (Shanghai, China) and sunitinib-eluting stents were produced in our laboratory as previously described by Chen et al (15). Sunitinib was dissolved in 100 µl ethanol and mixed with lubricating jelly (Jiangsu Province Jianerkang Medical Dressing Co., Ltd., Jintan, China) to a final concentration of 2 mg/g of jelly. Lubricating jelly is a commercially available product that consists of propylene glycol, a viscous liquid commonly used as an additive in food and medicine. Each balloon-mounted stent was individually dipped in sunitinib jelly to produce a thin coating weighing approximately 0.015 g; the amount of sunitinib eluted was determined from the additional weight of the coating. The stents were air-dried to completely evaporate the solvent. A coated stent was crimped onto a shrunken catheter balloon, and a pressure of 10 atm was applied to make the stent expand fully. The amount of sunitinib prior to and following expansion was analyzed.

Sunitinib release was measured using high performance liquid chromatography (HPLC; Agilent 1100; Agilent Technologies, Inc., Santa Clara, CA, USA), as previously described (16). The STES was immersed in a tube containing phosphate-buffered saline (PBS; pH=7.35–7.45). All tubes were incubated at 37°C throughout the release study. Samples were characterized at each predetermined time point.

All care and handling of the animals was in accordance with guidelines for the care and use of laboratory of animals, and the present study was approved by Nanjing University's ethical research committee (Nanjing, China). A total of 24 male New Zealand rabbits (2.5–3.5 kg) obtained from Jiangning Qinglongshan Animal Center (Nanjing, China), were used in the present study. Animals were maintained at 22°C and 65% humidity, and allowed ad libitum access to water and a basal diet. The rabbits were randomly assigned to two treatment groups: Bare-metal stent (BMS; n=12) and sunitinib-coated stents (STES; n=12). Under anesthesia with 5 mg droperidol and 100 mg ketamine (Fujian Fukang Pharmaceutical Co., Ltd., Fuzhou, China) administered intramuscularly, the carotid artery was accessed by dissection and isolated from the surrounding tissue. The balloon-expandable stent was inserted into the carotid artery through a small incision, and the stent was deployed by inflation with 10 atm. for 30 sec. Finally, the access site in the carotid artery was closed with 8–0 Prolene suture (Ethicon, Inc, Somerville, NJ, USA). Postoperatively, all rabbits received 25 mg aspirin prior to sacrifice, and a prophylactic antibiotic (300 mg cefuroxime) was administered for 7 days. The implanted stents were monitored by a Doppler handle probe (HP Sonos 4500; Philips Medical Systems, Inc., Bothell, WA, USA), and images of the blood flow were recorded every month. Diagnostic angiography was performed to confirm the position and patency rate of the stents.

At 1–3 months following stent implantation, rabbits were euthanized by direct intracardiac injection of potassium chloride and the stents and surrounding arteries were harvested. The arteries containing stents were divided into three segments. The proximal segment was opened longitudinally, flattened and fixed in 1.6% glutaraldehyde. It was then dehydrated, dried with CO₂, coated with gold and visualized under a scanning electron microscope (SEM; 3600N; Hitachi, Ltd., Tokyo, Japan). The percentage of the re-endothelialized area compared with the total luminal surface area was determined by examining SEM photomicrographs of each specimen. Protein was extracted from the ventral segment for western blot analysis. The distal segment was fixed in 10% neutral-buffered formalin for 24 h, embedded in methylmethacrylate, circumferentially cross-sectioned into 5-µm thick sections on a microtome fitted with a D-profile tungsten carbide knife (Delaware Diamond Knives, Inc., Wilmington, DE, USA) and stained with hematoxylin and eosin (17) for the quantification of histomorphological parameters. Tissue sections were visualized using a light microscope (BX51; Olympus Corporation, Tokyo, Japan), and the injury score (IS), average intimal thickness (AIT), areas of neointima (AN), media area, fibrin score (FI) and inflammation score (IMS) of each section was measured by digital morphometry using Image-Pro Plus software version 6.2 (Media Cybernetics, Inc., Rockville, MD, USA). IS was graded using the standardized protocol developed by Schwartz et al (18), which determined the degree of injury of the vessel wall in a semi-quantitative manner. The maximum score of 3 was applied when medial laceration extended through the external elastic lamina, and a score of 0 corresponded to an intact internal elastic lamina. Similarly, the extent of cellular infiltration was scored from 0–3 using a system described by Hong et al (19).

Rat smooth muscle cells (RSMCs) and rat endothelial cells (RECs) were purchased from the Cell Bank of the National Academy of Science (Shanghai, China). RSMCs were grown in Dulbecco's modified Eagle's medium (DMEM) (Sigma-Aldrich; Merck Millipore, Darmstadt, Germany) containing 10% fetal bovine serum (FBS; Sigma-Aldrich; Merck Millipore). RECs were cultured in Endothelial Basal Medium 2 (Sigma-Aldrich; Merck Millipore). Rat SMCs/ECs were chosen for the present preliminary study as rat antibodies are more commonly available. RSMCs and RECs were seeded in 96-well culture plates (7,500 cells/well). Sunitinib (0.1, 1 or 10 µM) or vehicle was added to the wells. Two days later, a 3-(4,5-dimethylthiazol)-2,5-diphenyltetrazolium (MTT) assay was performed as described previously (20) to determine the cytotoxicity of sunitinib.

To assess the direct effect of sunitinib on the proliferation of RSMCs, a 3H-thymidine incorporation assay was performed. (Methyl-3H)-thymidine (1 µCi/ml; PerkinElmer, Inc., Waltham, MA, USA) was added to wells 6 h prior to the termination of each experiment. Radioactivity was measured using a liquid scintillation counter.

To detect cell death, cells were stained with annexin V-fluorescein isothiocyanate (FITC) and propidium iodide (PI) according to the manufacturer's instructions (BD Biosciences, San Jose, CA, USA). RSMCs were cultured in a serum-free medium in the presence or absence of sunitinib for 24 h and then stimulated with 20 ng/ml PDGF-BB (CST Biological Reagents Co., Ltd.) homodimers for a further 24 h. The cells were subsequently trypsinized, collected and incubated with 5 µl annexin V-FITC and 5 µl PI (stock solution, 50 µg/ml) for 30 min at room temperature in the dark. Apoptotic (annexin V⁺/PI⁻) and necrotic (annexin V⁺/PI⁺) cells were detected on a FACSCalibur (BD Biosciences).

RSMCs were cultured in 6-well plates with BMS and STES for 12 h prior to examination. RSMCs attached to the stent struts were stained with 4,6-diamidino-2-phenylindole (DAPI; a blue fluorescent dye for cell nuclei) and observed under a fluorescent microscope. The numbers of attached SMCs were counted in three random high-power fields (magnification, ×200) per strut.

RSMCs were seeded in 35-mm dishes in DMEM and starved of FBS for 12 h, and cells were scratched with a 100-µl pipette tip. Cells were pretreated with 0, 0.1, 1 or 10 µM sunitinib and subsequently stimulated with 20 ng/ml PDGF-BB for 24 h. Cells were visualized under a light microscope and images were captured. Image-Pro Plus software version 6.2 (Media Cybernetics, Inc., Rockville, MD, USA) was used for quantification.

RSMCs were seeded in 6-well culture plates in DMEM and starved of FBS for 24 h, and treated with sunitinib or vehicle supplemented with 20 ng/ml PDGF-BB for a further 24 h. RSMCs were fixed with 4% paraformaldehyde for 10 min and permeabilized in 0.2% Triton X-100 for 10 min. The primary rabbit polyclonal anti-α-SMA antibody (1:100; catalog no. 14968; CST Biological Reagents Co., Ltd.) was applied to cells. Following washing with PBS three times, cells were incubated with the secondary antibody FITC-anti-rabbit IgG (1:400; catalog no. ab6717; Abcam, Cambridge, MA, USA) for 2 h at 37°C and observed under a fluorescent microscope.

For the in vitro experiment, protein was extracted using radioimmunoprecipitation assay buffer (Sigma-Aldrich; Merck Millipore) from RSMC cultured with various concentrations of sunitinib and 20 ng/ml PDGF-BB. Protein (20 µg) was loaded onto a 12% SDS-PAGE gel, separated by electrophoresis and transferred onto a polyvinylidene difluoride membrane. Membranes were blocked with 5% non-fat milk for 2 h, and incubated overnight at 4°C with the following primary antibodies: Rabbit phosphorylated (p)-PDGFR-β (1:1,000; catalog no. 3124), rabbit p-extracellular signal-regulated kinase (ERK; 1:1,000; catalog no. 9101) and rabbit glyceraldehyde-3-phosphate dehydrogenase (GAPDH; 1:2,000; catalog no. 5174), all purchased from CST Biological Reagents Co., Ltd. A horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (1:3,000; catalog no. ab6721; Abcam) was then added to the membranes at room temperature for 2 h. The signal was developed using Enhanced Chemiluminescence reagent (EMD Millipore, Bedford, MA) and visualized with the FlourChem FC2 Imaging System (ProteinSimple, San Jose, CA, USA). The protein bands were quantified using Image J software version 1.51b (National Institutes of Health, Bethesda, MD, USA). For the in vivo experiment, protein from neointimal tissue and media plus adventitia from BMS and STES was extracted and the protein expression levels of p-ERK and GAPDH were analyzed.

Data are expressed as the mean ± standard deviation. Unpaired Student's t-test was used to compare control and sunitinib-treated groups. Statistical analyses were performed in SPSS version 13.0 (SPSS, Inc., Chicago, IL, USA). P<0.05 was considered to indicate a statistically significant difference.

Incubation of bare-metal stents in the solution of sunitinib jelly resulted in coating with this compound in the present preliminary study; 32±5.4 µg sunitinib was bound to the stent. The amount of sunitinib on the stent did not alter following expansion. The results of the in vitro release of sunitinib are presented in Fig. 1A.

Stents were successfully implanted into the carotid arteries of 24 New Zealand rabbits (Fig. 1B), and the patency of the stent grafts was demonstrated by diagnostic angiography and Doppler ultrasound (Fig. 1C and D).

In-stent restenosis was examined by morphological analysis, as presented in Fig. 2. One month following implantation, the AIT was 0.12±0.03 mm in the BMS group and 0.07±0.04 mm in the STES group (P=0.06). Three months following implantation, clear neointimal formation was observed in BMS arteries; however, AIT was markedly reduced in STES (0.16±0.06 vs. 0.22±0.05 mm; P=0.03). In addition, the AN was 0.35±0.14 mm² in the BMS group and 0.21±0.12 mm² in the STES group (P=0.07) one month following implantation, and 0.64±0.20 mm² in the BMS group and 0.50±0.21 mm² in the STES group (P=0.09) three months following implantation (Fig. 2A and B).

No differences were observed in the IS or IMS between the two groups at one and three months following implantation, indicating that the observed effects on neointima formation did not result from structural differences or inflammation. IS were 1.17±0.41 (BMS group) and 1.00±0.63 (STES group; P=0.31), and 1.16±0.41 (BMS group) and 1.17±0.98 (STES group; P=0.50), at one and three months post-implantation, respectively. IMS were 0.33±0.52 (BMS group) and 0.17±0.41 (STES group; P=0.18), and 0.33±0.52 (BMS group) and 0.50±0.55 (STES group; P=0.31), at one and three months post-implantation, respectively.

Regarding the cytotoxicity of sunitinib, media area and FI were calculated. Media area was 0.42±0.03 in the BMS group and 0.40±0.04 in the STES group (P=0.21), and 0.42±0.06 in the BMS group and 0.39±0.04 in the STES group (P=0.25), at one and three months post-implantation, respectively. The FI values were 1.83±0.41 (BMS) and 2.00±0 (STES; P=0.18), and 2.00±0 (BMS) and 1.67±0.52 (STES; P=0.09), at one and three months post-implantation, respectively. These results suggest that local administration of sunitinib inhibited neointimal formation without damaging normal vascular media.

Re-endothelialization was analyzed by SEM. The re-endothelialized areas at one month following implantation were 91.7±3.7 and 89.5±6.8% in the BMS and STES groups, respectively (P=0.24; Fig. 2C), indicating that BMS and STES were efficient at inducing re-endothelialization. Notably, BMS and STES were observed to almost fully re-endothelialize areas, demonstrating that sunitinib did not delay endothelialization in the rabbit carotid model.

An MTT assay was performed in vitro with RSMCs and RECs to evaluate the effect of sunitinib on proliferation. Sunitinib treatment significantly suppressed the proliferation of RSMCs in a dose-dependent manner (0.1 µM, P=0.040; 1 µM, P=0.001; 10 µM, P<0.001); these suppressive effects were attenuated in RECs. No suppression was observed when RECs were treated with 0.1 or 1 µM sunitinib. The proliferation of RECs was slightly inhibited at 10 µM sunitinib (Fig. 3A). In addition, sunitinib treatment significantly suppressed DNA synthesis in RSMCs in a dose-dependent manner (0.1 µM, P=0.014; 1 µM, P=0.021; 10 µM, P=0.021). However, sunitinib had no significant effects on DNA synthesis in RECs.

As an antiproliferative effect is often accompanied by apoptosis or necrosis, the apoptotic effects of sunitinib on RSMCs were examined. Sunitinib significantly increased necrosis in a dose-dependent manner (0.1 µM, P=0.010; 1 µM, P=0.015; 10 µM, P=0.009), as assessed by PI staining, indicating that sunitinib may block RSMC proliferation by inducing necrosis (Fig. 3B). The in vitro effects of STES and BMS on RSMC adherence were then assessed. RSMC attachment to STES was reduced compared with BMS (21.8±9.3 vs. 11.2±5.4/strut; P=0.03; Fig. 3C).

Migration of SMCs from media to intima occurs following vascular injury, and PDGF is crucial for this migration. A scratch wound assay and cytoskeleton staining was therefore performed to investigate the effect of sunitinib on migration. The scratch wound assay is a classic two-dimensional migration model. As presented in Fig. 3D, stimulation with PDGF-BB induced migration was attenuated by sunitinib in a dose-dependent manner. Actin cytoskeleton polymerization is critical for cell migration; therefore, α-SMA in RSMCs was visualized with FITC. Actin polymerization was decreased following sunitinib treatment (Fig. 3E), suggesting that downregulated expression levels of α-SMA may represent one of the mechanisms underlying the inhibition of RSMC migration by sunitinib.

Various signaling pathways (particularly the ERK signaling pathway) are involved in SMC migration and growth. Therefore, ERK activation was analyzed by assessing phosphorylation, to further study the mechanisms underlying the inhibitory effects of sunitinib on proliferation and migration.

In the in vitro experiment, protein expression levels of p-PDGFR were decreased by sunitinib treatment in a dose-dependent manner, and PDGF-induced downstream p-ERK protein expression levels were suppressed (0.1 µM, P=0.021; 1 µM, P=0.024; 10 µM, P=0.031; Fig. 4A and B). This suggested that sunitinib specifically inhibited PDGF-induced ERK activation. In the in vivo experiment, the media, adventitia and neointima were harvested. The protein expression levels of p-ERK in the neointima were significantly decreased in STES compared with BMS (P=0.027; Fig. 4C and D).