Polyclonal antibodies against cyclin E (sc-481), CDK2 (sc-163) and CDK4 (sc-260), p21WAF1 (sc-756), p53 (sc-126), p27 (sc-528) and GAPDH (sc-20357) were all obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Monoclonal antibody against cyclin D1 (04-221) was obtained from Millipore (Temecula, CA, USA). Polyclonal antibodies against ERK (9102), phosphorylated (p)-ERK (9101), p38 MAPK (9212), p-p38 MAPK (9211), JNK (9258), p-JNK (9251), AKT (9272) and p-AKT (9271) were all purchased from Cell Signaling Technology (Lakewood, NJ, USA). Polyclonal MMP-9 antibody (AB19016) was purchased from Millipore. HRP-conjugated secondary antibodies against goat anti-rabbit IgG-HRP (sc-2004), goat anti-mouse IgG-HRP (sc-2005) and donkey anti-goat IgG-HRP (sc-2020) were purchased from Santa Cruz Biotechnology, Inc. All procedures and protocols used in the present study were approved by the Ethics Committee of Chung-Ang University (Anseong, Korea).

We used a total of 100 g of air-dried Rosa Hybrida flowers (purchased from Jincheon Agricultural Technology Center, Jincheon-gun, Korea), which were finely crushed and soaked in water, followed by heating at 100°C. The extract was concentrated via a rotary evaporator (Rotavapor-R; Brinkmann Instruments, Westbury, NY, USA), lyophilized, and freeze-dried. The yield of the final extract was approximately 10% w/w, and the extract was diluted in saline buffer.

VSMCs were obtained from the aortas of 2 young male Sprague-Dawley rats (8 weeks old, weighing 200–250 g; DHbiolink, Eumseong, Korea) using enzymatic digestion, as described previ ously (23). Briefly, the rats were sacrificed by CO₂ inhalation, the vessels were obtained under sterile conditions and were washed several times in Hanks' balanced salt solution (HBSS). Following removal of the adventitia from the aortas, digestion of the aortas in 5 ml of digestion solution (0.125 mg/ml elastase, 0.25 mg/ml soybean trypsin inhibitor, 10 mg/ml collagenase I, 2.0 mg/ml crystallized bovine albumin and 15 mM HEPES; all from Sigma-Aldrich; St. Louis, MO, USA) was performed at 37°C for 45 min. The digested cells were filtered with a sterile 100-µm nylon mesh (BD Biosciences, Franklin Lakes, NJ, USA), centrifuged at 1,000 rpm for 10 min and rinsed twice in Dulbecco's modified Eagle's medium (DMEM; Corning, Inc., Corning, NY, USA) containing 10% fetal calf serum. They were then cultured in DMEM. To characterize the cells as VSMCs, immunofluorescence staining was performed with a monoclonal antibody against smooth muscle-α-actin (Sigma-Aldrich). The explants were incubated in DMEM containing 10% FBS (Corning, Inc.), 2 mM glutamine (Sigma-Aldrich), 50 µg/ml gentamicin (Sigma-Aldrich), and 50 µl/ml amphotericin B (Sigma-Aldrich) at 37°C in an atmosphere with 5% CO₂. The cells were used at passages five to eight. For use in the experiments, the cells were grown to 80–90% confluence and rendered quiescent by serum-starvation for at least 24 h in DMEM without FBS.

The cells were cultured to 80–90% confluence, and then starved while being cultured in serum-free DMEM for 24 h. The cells were incubated with various concentrations of WERH (100, 200, 400 and 600 µg/ml) for 40 min and then cultured with or without PDGF (20 ng/ml; Millipore) for a further 24 h. A modified 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT; Sigma-Aldrich) assay was used to determine cell viability, as described previously (12,23).

After 24 h starvation, the cells were treated with the indicated concentrations of WERH and then cultured with or without PDGF (20 ng/ml) for a further 24 h. This was followed by the addition of 1 µCi/ml [methyl-³H]thymidine (PerkinElmer NEN., Boston, MA, USA) for 4 h. Incorporated thymidine was precipitated with cold 10% trichloroacetic acid, solubilized in 0.2 M NaOH, and counted using a scintillation counter (PerkinElmer NEN, Inc., Waltham, MA, USA).

The cells were collected and fixed in 70% ethanol (Millipore) for 2 h at 4°C. After washing with ice-cold phosphate-buffered saline (PBS; Sigma-Aldrich), the cells were incubated with RNase (0.1 mg/ml; Sigma-Aldrich) and propidium iodide (50 µg/ml; Sigma-Aldrich) for 30 min. Cell cycle analysis was performed using a Becton-Dickinson FACStar flow cytometer equipped with Becton-Dickinson Cell Fit software (both from BD Biosciences), as previously described (12,23).

The cells were lysed using 250 µl lysis buffer [containing, in mmol/l, HEPES (pH 7.5) 50, NaCl 150, EDTA 1, EGTA 2.5, DTT 1, β-glycerophosphate 10, NaF 1, Na₃VO₄ 0.1, and phenylmethylsulfonylfluoride 0.1 and 10% glycerol, 0.1% Tween-20, 10 µg/ml of leupeptin, and 2 µg/ml aprotinin; all from Sigma-Andrich]. After centrifugation at 12,000 rpm for 20 min at 4°C, the protein concentrations of the cell lysates were determined by Bradford assay. Equal amounts of the samples were resolved via electrophoresis on SDS-polyacrylamide (10%) gels, and then electroblotted onto nitrocellulose membranes (Hybond; Amersham Corp., Arlington Heights, IL, USA). The membranes were incubated overnight with specific primary antibodies at 4°C followed by peroxidase-conjugated secondary antibodies for 1 h. Immunocomplexes were visualized using an ECLPlus™ Western Blot Detection system (Amersham Biosciences, Piscataway NJ, USA). Experiments were performed at least 3 times.

For the immunoprecipitation assay, equal amounts of cell lysates were incubated with specific antibodies overnight at 4°C. Immunocomplexes were added using protein A-sepharose beads (Santa Cruz Biotechnology, Inc.), followed by incubation at 4°C for 2 h. The immunoprecipitates were washed with lysis buffer 3 times, and then resuspended in SDS-PAGE sample buffer containing β-mercaptoethanol (Bio-Rad, Richmond, CA, USA). The samples were analyzed by immunoblot analysis. For immune complex kinase assays, the immunoprecipitated samples were washed 3 times with lysis buffer and twice with kinase buffer (containing, in mM/l, HEPES 50, MgCl₂ 10, DTT 1, β-glycerophosphate 10, NaF 1, and sodium orthovanadate 0.1). The pellet was resuspended in 25 µl kinase buffer containing either 1 µg glutathione S-transferase (GST)-pRb C-terminal (pRb amino acids 769–921 fusion protein; Santa Cruz Biotechnology, Inc.) or 5 µg histone H₁ (Life Technologies, Grand Island, NY, USA), 20 µM/l ATP, and 5 µCi [γ³²P]ATP (4,500 µCi/mmol; ICN, Costa Mesa, CA, USA), and incubated for 20 min at 30°C. The reaction was then stopped by the addition of 25 µl of 2X concentrated Laemmli sample buffer (Santa Cruz Biotechnology, Inc.). Electrophoresis was performed with the samples using an SDS-polyacrylamide (10%) gel. The phosphorylated forms of pRb and histone H₁ were visualized with a BAS 2000 bioimaging analyzer (Fujifilm, Tokyo, Japan).

Serum-starved cells were grown to 90% confluence, and then damaged with a pipette tip. The medium was replaced with fresh medium containing different concentrations of WERH in the presence or absence of PDGF (20 ng/ml). After 24 h incubation, a phase-contrast microscope (Optika, Ponteranica, Italy) was use to capture images and to evaluate the wound closure effected by cell migration.

An invasion assay was then performed using modified Boyden chambers with a polycarbonate nucleopore membrane (Corning, Inc.). The serum-starved cells were cultured in DMEM containing WERH with or without PDGF and they were then poured into each of the upper chamber plates for 24 h to allow for cell invasion through the membrane. Non-invading cells on the upper surface were removed using a cotton swab, and the cells that had invaded the membrane were stained with 0.1% crystal violet solution (containing 2.5 mM crystal violet, 1% MeOH, 4% paraformaldehyde) for 20 min. Invasive cells were then quantified by counting the cells, using a microscope, in 6 independent areas at ×20 magnification/well.

Conditioned medium was resolved via electrophoresis of polyacrylamide gels supplemented with 1 mg/ml gelatin. After washing the gels twice with 2.5% Triton X-100 at room temperature for 2 h, they were reacted with a buffer (10 mM CaCl₂, 150 mM NaCl, and 50 mM Tris-HCl, pH 7.5) at 37°C overnight. The gels were then stained with 0.2% Coomassie blue, followed by destaining with 50% methanol and 5% acetic acid. Areas of gelatinase activity were visualized as clear bands against a dark blue field.

Nuclear extracts and EMSA were performed as described previously (12,18). The cultured cells were harvested by centrifugation, washed and suspended in a buffer [10 mM HEPES (pH 7.9), 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM DTT and 0.5 mM PMSF] for 15 min on ice. After vortexing the cells with 0.5% Nonidet P-40, a nuclear pellet was then assembled by centrifugation at 12,000 rpm for 1–2 min at 4°C and extracted in buffer (20 mM HEPES pH 7.9, 0.4 M NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM DTT and 1 mM PMSF) for 15 min at 4°C. The nuclear extract (10–20 µg) was reacted at 4°C for 30 min using the 100-fold excess of an unlabeled oligonucleotide containing the -79 MMP-9 cis element of interest (Bioneer, Daejeon, Korea). The sequences used were as follows: AP-1, CTGACCCCTGAGTCAGCACTT; NF-κB, CAGTGGAATTCCCCAGCC; and Sp1, GCCCATTCCTTCCGCCCCCAGATGAAGCAG. The binding reaction mixture was then incubated at 4°C for 20 min in a buffer (25 mM HEPES buffer pH 7.9, 0.5 mM EDTA, 0.5 mM DTT, 0.05 M NaCl, and 2.5% glycerol) with 2 µg poly(dI/dC) (Bioneer) and 5 fmol (2×10⁴ cpm) of a Klenow end-labeled (³²P-ATP) 30-mer oligonucleotide (Bioneer), which spanned the DNA binding site in the MMP-9 promoter. The reaction mixture was electrophoresed at 4°C in a 6% polyacrylamide gel with TBE (89 mM Tris, 89 mM boric acid and 1 mM EDTA) running buffer. The gel was washed with water, dried and exposed to X-ray film overnight. The shifted bands were detected and evaluated using ImageQuant software (GE Healthcare Life Sciences, Pittsburgh, PA, USA).

In the present study, the results are represented as the means ± SE. Statistical comparisons between groups were analyzed using factorial ANOVA analysis and Fisher's least significant difference test. A P-value <0.05 was considered to indicate a statistically significant difference.

To evaluate the effect of WERH on cell proliferation, VSMCs were treated with WERH, and this was followed by the addition of PDGF. VMSC proliferation was induced by PDGF, and this was verified by MTT assay and a thymidine incorporation assay. The PDGF-induced proliferative effects were reversed to a control level (non-treatment) at a dose of 400 µg/ml WERH and cell toxicity was not noted (Fig. 1A and B). Cell death was not observed in the vehicle-treated group (Fig. 1A). The survival of VSMCs was significantly reduced at a 600 µg/ml concentration of WERH (Fig. 1A). Similar results were obtained in a thymidine incorporation experiment (Fig. 1B). Subsequent experiments were performed using a non-toxic cellular concentration of WERH (below 400 µg/ml). These results demonstrated that WERH inhibited VSMC proliferation induced by PDGF, without causing cell death.

We then investigated whether WERH induced changes in cell cycle distribution in PDGF-stimulated VSMCs. As shown in Fig. 1C, flow cytometric analysis indicated that in PDGF-stimulated VSMCs WERH treatment induced an accumulation of cells in the G1-phase cell cycle and reduced the number of cells in the S-phase cell cycle, compared with PDGF-stimulated VSMCs which were not treated with WERH. These results suggest that growth inhibition promoted by WERH in VSMCs was caused by G1-phase cell cycle arrest. Subsequent experiments examined the cell cycle-regulatory proteins that are closely associated with the G1-phase cell cycle. The expression levels of cyclin D1, cyclin E, CDK2 and CDK4 were upregulated in PDGF-stimulated VMSCs (Fig. 2A). The treatment of VSMCs with WERH resulted in a significant reduction in the PDGF-stimulated expression levels of cyclin D1, cyclin E, CDK2 and CDK4 (Fig. 2A). The kinase activity of CDKs, as cyclin-CDK complexes, is known to be essential to G1- to S-phase cell cycle progression (6,7). Thus, the following experiment investigated the effect of WERH on the kinase activity of CDKs in PDGF-stimulated VSMCs. PDGF stimulation increased the kinase activity of CDK2 and CDK4 in VSMCs. The increased kinase activity of CDK2 and CDK4 induced by PDGF was suppressed in the presence of WERH (Fig. 2B).

As CDKIs are known to be negative regulators that control the transition from the G1- to S-phases of the cell cycle (9,10), the expression levels of the CDKIs p21WAF1 and p27KIP1 were examined. PDGF treatment induced the expression of p21WAF1 and decreased the expression of p27KIP1 in VSMCs (Fig. 2C). Stimulation of VSMCs to PDGF had almost no effect on the expression level of tumor suppressor protein p53. In addition, WERH treatment increased p21WAF1 expression in PDGF-stimulated VSMCs. However, WERH had no effect on the PDGF-induced inhibition of p27KIP1 expression in VSMCs. The level of p53 expression was not markedly affected by the addition of WERH in the presence of PDGF (Fig. 2C). To further investigate whether the observed inhibitory effect on the CDKs was due to interactions with p21WAF1, immunoprecipitation analysis was performed using cell lysates in the presence of WERH. An increase in the association of p21WAF1 with CDK2 and CDK4 was detected in the PDGF-stimulated VSMCs (Fig. 2D). WERH treatment increased the levels of p21WAF1/CDK2 and p21WAF1/CDK4 in VSMCs following stimulation with PDGF (Fig. 2D). These results demonstrated that WERH suppressed the PDGF-stimulated proliferation of VSMCs, through p21WAF1-mediated G1-phase cell cycle arrest, which involved inhibition of the kinase activity of CDKs.

To identify the signaling pathways involved in the PDGF responses in VSMCs, we examined the phosphorylation of MAPKs and AKT in VSMCs. Stimulation of VSMCs with PDGF led to an increase in the phosphorylation of ERK1/2, JNK, p38 MAPK, and AKT (Fig. 3). We next investigated whether WERH is capable of inhibiting the phosphorylation of ERK1/2, JNK, p38 MAPK and AKT in PDGF-stimulated VSMCs. As shown in Fig. 3, treatment with WERH significantly suppressed the PDGF-induced phosphorylation of ERK1/2 and AKT in VSMCs. However, WERH treatment did not markedly affect the phosphorylation of JNK and p38 MAPK induced by PDGF in VSMCs. These results indicated that WERH blocks the proliferation of PDGF-stimulated VSMCs by suppressing ERK1/2 and AKT signaling.

The migration and invasion of VSMCs are critical steps in the formation of vascular lesions that lead to the development of atherosclerosis (13,14). An in vitro wound healing assay and an invasion assay were performed to determine the effects of WERH on the PDGF-induced migration and invasion of VSMCs. PDGF treatment significantly increased the motility and invasiveness of VSMCs within 24 h (Fig. 4). Treatment of VSMCs with WERH resulted in decreased migration of VSMCs (Fig. 4A). In addition, the invasiveness of PDGF-stimulated VSMCs was significantly obstructed in the presence of WERH (Fig. 4B). These results clearly suggest that WERH plays an inhibitory role in the migration and invasion of VSMCs induced by PDGF.

MMP-9 expression is involved in the migration and invasion of VSMCs during the development of vascular lesions (13,14). Thus, the next experiment examined whether WERH inhibited the expression of MMP-9 in PDGF-stimulated VSMCs using a gelatin zymographic assay and immunoblot analysis. The expression of MMP-9 was stimulated by exposure to PDGF as demonstrated by zymographic assay (Fig. 5A). Treatment with WERH significantly abolished PDGF-stimulated MMP-9 expression in VSMCs. The immunoblot analysis results were similar (Fig. 5A). To further investigate the exact molecular mechanism of action of the inhibitory effect of WERH, an EMSA experiment was employed using three motifs, NF-κB, AP-1 and Sp-1 cis-elements, that are responsible for MMP-9 expression (18,19). PDGF treatment significantly stimulated the binding activity of the NF-κB, AP-1 and Sp-1 motifs in VSMCs. The addition of WERH markedly reduced the increased binding ability of NF-κB, AP-1 and Sp-1 motifs in PDGF-stimulated VSMCs (Fig. 5B). These results demonstrated that the transcription factors NF-κB, AP-1 and Sp-1 are involved in the WERH-induced inhibition of MMP-9 expression in PDGF-stimulated VSMCs.