Cilostazol (OPC-13013) was generously donated by Otsuka Pharmaceutical (Tokushima, Japan). Protease inhibitor cocktail, trypan blue (0.4%), 5-bromo-4-chloro-3-indolyl β-D-galactoside (X-gal), glutaraldehyde, formaldehyde, potassium ferrocyanide, potassium ferricyanide, and etoposide were purchased from Sigma-Aldrich Chemical Co. (St. Louis, MO, USA). Fetal bovine serum (FBS), Dulbecco’s modified Eagle’s medium (DMEM) and other culture reagents were purchased from Hyclon (Logan, UT, USA). Anti-poly(ADP-ribose) polymerase (PARP), Bax, type II collagen, β-catenin, and β-actin antibodies were obtained from Santa Cruz Biotechnology (Santa Cruz, CA, USA). The secondary horseradish peroxidase (HRP)-conjugated antibody and the enhanced chemiluminescence (ECL) Western blotting kit were obtained from Amersham Pharmacia Biotech (Piscataway, NJ, USA).

Articular chondroctyes for primary culture were isolated from slices of knee joint cartilage of 5-week-old female Sprague-Dawley rats (Samtako BioKorea, Osan, Korea). Chondrocytes were isolated by enzymatic digestion for 1 h with 0.2% type II collagenase in DMEM. The chondrocytes were briefly centrifuged, and the cells were resuspended in DMEM supplemented with 10% heat-inactivated FBS and antibiotics (50 U/ml penicillin, 50 μg/ml streptomycin) at 37°C with 5% CO₂ in air atmosphere. Cells were plated on culture dishes at a density of 5×10⁴ cells/cm². The medium was replaced every 2 days, and cells reached confluence at ~4–5 days after culture; this was designated as passage 0 (P0). The P0 cells were serially subcultured up to P6 by plating cells at a density of 5×10⁴ cells/cm².

Cell viability was determined using the trypan blue exclusion assay. Chondrocytes were plated at 1×10⁵ cells per 6-well plates and incubated for 24 h. Cells were cultured for different times in the presence or absence of various concentrations of cilostazol in fresh DMEM medium. After incubation, the cells were washed with phosphate-buffered saline (PBS), and viable cells were scored by the trypan blue dye exclusion assay by a hemocytometer.

Cells were washed with 1% PBS/BSA and fixed in 4% paraformaldehyde for 15 min. Next, they were washed with PBS/BSA and permeabilized in 0.1% Triton-X 100 for 5 min on ice. Fluorescein isothiocyanate (FITC)-conjugated dUTP was used for the terminal deoxynucleotidyl transferase-mediated dUTP nick end-labeling (TUNEL) assay that was performed using the Apoptosis Detection System kit (Roche Molecular Biochemicals, Mannheim, Germany) according to the manufacturer’s instructions.

SA-β-gal staining assay was performed at pH 6.0 as described by Dimri et al, with a modification (21). Cells were washed in PBS, fixed for 5 min (room temperature) in 0.2% glutaraldehyde/2% formaldehyde, washed in PBS, and incubated with SA-β-gal stain solution (1 mg/ml X-Gal) 40 mM citrate/phosphate buffer (pH 6.0), 5 mM potassium ferrocyanide, 5 mM potassium ferricyanide, 150 mM NaCl, and 2 mM MgCl₂) in a chamber maintained at 37°C for 12 h. The degree of senescence-associated cells was calculated as a percentage of the total number of cells.

Chondrocytes (1×10⁶ cells/cm²) were grown in 60-mm culture dishes, and incubated for 24 h in fresh medium with or without cilostazol. Next, total RNA was isolated using TRIzol reagent and reverse transcription was performed using superscript reverse transcriptase (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instructions. Total RNA (2 μg) was used to prepare cDNA. The following primers were used in our study: type I collagen, forward, 5′-GACCCAAAGGTTCTCGTGGT-3′ and reverse, 5′-CTTTCTCCTCTCTGACCGGG-3′; type II collagen, forward 5′-GGTAAGTGGGGCAAGACCAT-3′ and reverse 5′-TTTTGCAGTCTGCCCAGTTC-3′; glyceraldehyde 3-phosphate dehydrogenase (GAPDH), forward, 5′-TGAAGGTCGGAGTCAACGGATTTGGT-3′ and reverse, 5′-CATGTGGGCCATGAGGTCCACCAC-3′. PCR reactions in 25 μl reaction volumes were performed as follows: 95°C for 5 min, 30 cycles of 94°C for 30 sec, either 52°C (type I collagen) or 56°C (type II collagen) for 30 sec, 72°C for 1 min, and 72°C for 5 min.

Equivalent amounts (20 μg) of total proteins were loaded onto 12% sodium dodecyl sulphate (SDS)-polyacrylamide gels for electrophoresis. The proteins were then transferred onto a nitrocellulose membrane by using an electroblotting apparatus (Bio-Rad, Richmond, CA), and the membranes were incubated with each primary antibody. The blots were washed with TBS-T and incubated with an HRP-conjugated secondary anti-rabbit antibody. The membranes were developed using the ECL reaction system and visualized using the LAS-3000 Luminescent Image Analyzer (FujiFilm, Japan). Image Gauge Ver. 3.0 software was used to calculate the changes in protein expression, and β-actin was used as an internal control to ensure equal protein sample loading.

Chondrocytes were cultured on collagen-coated 4-well glass chamber slides for 48 h in DMEM containing 10% FBS. Cells were fixed in 4% paraformaldehyde/PBS, followed by permeabilization in 1% Triton X-100/PBS. Appropriate primary antibodies were added to the cells for 30 min at 37°C. For secondary labeling, cells were incubated with FITC-conjugated secondary antibody (Invitrogen) for 30 min at 37°C. Nuclei were counterstained using propidium iodide (PI). Fluorescent images were observed and analyzed using a laser-scanning confocal microscope.

Each experiment was repeated at least 3 times. Data were expressed as the means ± SE from each independent experiment. The data for the experimental and control groups were tested for statistical significance by one-tailed Student’s t test, with P<0.05 accepted as the level of significance.

To determine the role of cilostazol in the maintenance of chondrocyte phenotypes, we first investigated the effect of cilostazol on the expression of type II collagen in chondrocytes. Primary rat articular chondrocytes were isolated, maintained, and treated with various concentrations (2, 10, and 50 μM) of cilostazol for 24 h. As shown in Fig. 1A and B, cilostazol significantly reduced the expression of type II collagen, which plays a crucial role in regulating chondrocyte functions by facilitating cell-matrix interactions. In addition, cilostazol stimulated the accumulation of β-catenin, a phenotypic marker for the differentiation and dedifferentiation of chondrocytes (Fig. 1C and D). These results suggested that cilostazol causes cellular dedifferentiation, i.e., loss of differentiated chondrocyte phenotypes, in primary articular chondrocytes.

Next, we assessed whether the cilostazol-induced dedifferentiation is associated with cellular senescence in primary chondrocytes. Cilostazol decreased cell proliferation without affecting cell viability (Fig. 2A). P0 cells were treated with various concentrations (2, 10, and 50 μM) of cilostazol for 48 h. Then, the induction of cellular senescence was confirmed by an increase in SA-β-gal-positive cells in cilostazol-treated chondrocytes compared to those in the control. Interestingly, cilostazol-treated chondrocytes showed significant increase in specific SA-β-gal staining (Fig. 2B).

To investigate whether cellular senescence is associated with the differentiation or dedifferentiation status of chondrocytes, we determined the effect of cilostazol on cellular senescence during subculture-induced dedifferentiation of chondrocytes. Primary chondrocytes were serially subcultured to induce cellular dedifferentiation. As shown in Fig. 3A, the expression of type I and type II collagens was significantly increased and decreased in P4 and P6 cells, respectively. The expression of β-catenin showed a similar pattern as type II collagen (data not shown). Importantly, serial subculture of chondrocytes also induced changes in senescent phenotypes, such as changes in cell morphology, decreases in cell proliferation, and increases in specific SA-β-gal staining (Fig. 3B). This suggests that chondrocyte senescence also increased during subculture-induced dedifferentiation.

Next, we observed cellular senescence in chondrocytes at different passages (P0, P2, P4, and P6) by subculture. Dedifferentiated chondrocytes showed significant cellular senescence that increased with passage number. This suggested that dedifferentiated chondrocytes slowly and spontaneously progressed to cellular senescence. Cilostazol significantly increased the SA-β-gal staining of P2 cells, and slightly increased the SA-β-gal staining of P4 and P6 cells (Fig. 3C). Cilostazol had a weak effect on senescence in dedifferentiated chondrocytes at high passage number. This is because with increasing passage number increases, senescence in cells increases. Taken together, these results suggest that senescence in chondrocytes is associated with the dedifferentiation status, which indicates the loss of differentiated chondrocyte phenotypes.

TUNEL assay was used to determine whether cilostazol-induced dedifferentiation and senescence changed the susceptibility of articular chondrocytes against apoptosis mediated by an apoptosis-inducing drug. Cells were pretreated with cilostazol for 48 h, and then stimulated with 200 μg/ml etoposide for an additional 24 h. The percentage of TUNEL-positive cells was markedly reduced when chondrocytes were treated with cilostazol and etoposide compared to when they were treated with etoposide alone (Fig. 4A). Etoposide is known to induce poly(ADP-ribose) polymerase (PARP) cleavage, and etoposide-induced PARP cleavage was reduced in cilostazol-treated cells (Fig. 4B). To show that senescent cells are more resistant than young cells, we used P0 and senescent P6 cells. As shown in Fig. 4C, when cells were stimulated with etoposide for 24 h, ~45 and 19% of P0 and P6 cells were TUNEL-positive cells, respectively. These results suggest that cilostazol renders chondrocytes resistant to the apoptosis inducer by inducing cellular senescence.