The 5,637 urothelial cell line, a grade II carcinoma, was purchased from the Bioresource Collection and Research Center (Hsinchu, Taiwan). The 5,637 cells were maintained in RPMI-1640 medium (Gibco Life Technologies, Grand Island, NY, USA) supplemented with 10% fetal bovine serum (FBS; Biological Industries, M.P. Ashrat, Israel), 1.5 g/l sodium bicarbonate, 4.5 g/l D-glucose, 1% penicillin-streptomycin (Gibco Life Technologies), 1 mM sodium pyruvate and 10 mM HEPES. Cells were plated on 100-mm plastic dishes and incubated in a CO₂ incubator at 37°C, with 5% CO₂ and 95% filtered air.

Cell viability was determined using a colorimetric 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. The cells were seeded in 96-well plates at a density of 4×10³ cells/well for 24 h, and then were incubated with various concentrations of test agents for another 12–36 h. MTT was added into the medium at 37°C for 2 h, and the medium was then discarded and dimethyl sulfoxide (DMSO) was added to dissolve the formazan product. Each well was measured by light absorbance at 540 nm. The result was expressed as the percentage of the normal saline-treated control group.

Firstly, 1×10⁶ cells were seeded in 100-mm dishes for a 24-h incubation. Afterwards, TSA or sterile alcohol was added for another 12 and 24 h of treatment. The cells were then harvested, centrifuged at 800 × g for 5 min and fixed with ice-cold 75% ethanol overnight at 4°C. Following removal of the ethanol, the cells were stained with a DNA staining solution [containing 1 mg/ml propidium iodide (PI) and 10 mg/ml RNase A dissolved in phosphate-buffered saline (PBS)] for 30 min at room temperature. The relative DNA content of the stained cells was measured using a FACScan flow cytometer (BD Biosciences, San Jose, CA, USA). The cell doublets were removed by gating the left area of the FL2-W vs. FL2-A dot plot for analysis. The cell cycle data from flow cytometry were analyzed using ModFit LT™ software (Verity Inc., Sunnyvale, CA, USA).

Apoptotic cells were also detected using Annexin V labeling. Cells were treated as above and harvested, and then stained with 2 µl Annexin V-FITC and 2 µl PI staining solution (Bio-Genesis Technologies, Inc., Taipei, Taiwan) in the dark at room temperature for 15 min. The cell samples were immediately analyzed by the same flow cytometry and software program as previously mentioned.

The fluorescence intensity of Rhodamine 123 (Sigma-Aldrich, St. Louis, MO, USA), which is permeable to the mitochondrial membrane then specifically quenched in the mitochondria due to the MMP, was used as a measure of membrane damage. In brief, 5×10⁵ cells were incubated with 5 µM Rhodamine 123 for 10 min at 37°C. The cells were then centrifuged at 200 × g for 6 min, resuspended in PBS, and kept on ice in the dark. The staining intensity was determined using the FACS analysis (Ex/Em = 485/535 nm) as previously mentioned. Negative staining for Rhodamine 123 represented the loss of ΔΨm.

Briefly, the TSA-treated and control cells were washed with PBS and incubated for 20 min in 150 µl of lysis buffer containing PRO-PREP™ protein extraction solution (iNtRON Biotechnology, Gyeonggi-do, Korea), 0.1 M NaF and 0.5 M vanadate. The mixture was centrifuged at 12,000 × g for 5 min, and then the total protein extract in the supernatant fluid was collected and aliquoted for 30 µg for further analysis. Protein concentrations were determined using the Pierce BCA assay kit (Thermo Scientific, Rockford, IL, USA). For immunoblotting, proteins on the analytical 10% SDS-PAGE gels were transferred to a polyvinylidene difluoride membrane using a trans-blot apparatus. Antibodies against cleaved caspase-3, caspase-9, PARP, survivin (Cell Signaling Technology, Inc., Danvers, USA), α-tubulin, GAPDH, β-actin (GeneTex, Irvine, CA, USA), p-Akt, cyclin D1, p21 (Santa Cruz Biotechnology, Paso Robles, CA, USA) and Sp1 (Upstate Biotechnology, Lake Placid, NY, USA) were used as the primary antibodies. Mouse, rabbit or goat IgG antibodies conjugated to horseradish peroxidase were used as the secondary antibodies. An enhanced chemiluminescence kit and Chemi-Smart 3000 system (Vilber Lourmat Corporation, Torcy, France) were used for detection, and the quantity of each band was determined using MultiGauge software (Fujifilm, Tokyo, Japan).

Numerical data are expressed as the mean ± standard error from triplicate experiments. Statistical differences were analyzed using one-way analysis of variance analysis followed by Student's t-test. All statistics were calculated using SigmaPlot version 12.5 (Systat Software, Inc., San Jose, CA, USA).

TSA is a very toxic chemical in 5,637 cells. As shown in Fig. 1A, the 5,637 cells in the normal culture exhibited a spindle-shaped morphology. After exposure to 500 nM TSA for 12 h, some cells rounded up. After 24 h, >70% of the cells rounded up and the attached cells showed marked change to an elongated shape with filamentous protrusions. The viability of the 5,637 cells was then assessed using MTT assay. The 5,637 cells treated with TSA displayed decreased cell viability in a dose-dependent manner after 24 h (Fig. 1B). Furthermore, it was found that most of the cells died after co-culture with 500 nM TSA for 36 h, which was also well reflected in the markedly low cell viability presented here. Since the IC₅₀ value of TSA was determined between 250 and 500 nM after a 24-h treatment, these concentrations were used in the following experiments.

HDAC inhibitors have been reported to cause growth arrest at the G1 and G2/M phases in a wide variety of tumor cells (23). To elucidate whether TSA induces growth arrest in 5,637 cells, flow cytometric analysis was used. We measured cell cycle distribution at 12 and 24 h, respectively, after treatment with varying concentrations of TSA (125, 250 and 500 nM). The results showed that G2/M arrest and the sub-G1 portion were formatted along with the rising concentration of TSA at 12 h. At 24 h, there was a moderate increase in G1 phase portion at a lower dosage (125 and 250 nM), while a large degree of cell death was correlated with enhanced sub-G1 phase cells observed under a high dosage (500 nM) of TSA (Fig. 2A). Since p21 is the principal cyclin-dependent kinase inhibitor (CDKI) involved in cell cycle arrest upon DNA damage (24), we analyzed p21 expression by western blotting following TSA treatments. As shown in Fig. 2B, p21 expression was significantly increased in a dose-dependent manner following a 12-h treatment. In contrast, cyclin D1 was correspondingly found downregulated following both a 12- and 24-h treatment (Fig. 2B). These findings demonstrated that TSA may regulate G2/M and G1 phase arrest by modulating both p21 and cyclin D1 expression in 5,637 cells. Due to the rationale that HDAC inhibitors may regulate gene expression, the p21 mRNA expression level was then detected to verify whether TSA has an effect on p21 mRNA. However, as shown in Fig. 2C, the amount of p21 mRNA did not change following TSA treatment, suggesting that an alternative pathway was attributable to the enhanced p21 protein accumulation rather than the direct transcriptional activation of TSA.

Based on the evidence from the cell cycle assay that TSA induced severe death of 5,637 cells, we next examined the apoptosis status using Annexin V-PI staining in cells exposed again to four varying concentrations of TSA for 12 and 24 h. As shown in Fig. 3A, TSA caused dose-dependent apoptosis based on an increase in the number of Annexin V-positive cells at 12 h and at 24 h (UR, late apoptosis). In addition, apoptosis was further confirmed by the evidence of cleaved PARP and cleaved caspase-3 as detected by western blotting (Fig. 3B). In essence, the cell viability was restored when the pan-caspase inhibitor Z-VAD-FMK was applied (Fig. 3C), suggesting that TSA-induced cell death was mediated by the caspase-dependent apoptotic pathway.

The progression of apoptosis has been closely related to the injury suffered from MMP collapse (25). Since the TSA-induced apoptotic clues come from the activation of intrinsic pathway, subsequently we measured the change in MMP to assess whether the apoptosis process was associated with irreversible mitochondrial depolarization. In the present study, the loss of MMP in TSA-treated 5,637 cells was determined by use of Rhodamine 123 dye via flow cytometric assay. As shown in Fig. 4A, the loss of MMP was noted in the histograms, in which a clear peak shift was shown along with the increasing TSA concentration, indicating that TSA could lead to depolarization of the inner mitochondrial membrane in a dose-dependent manner. The percentage of the cells with mitochondrial depolarization was 13.6, 14.9 and 16.1% in response to 125, 250 and 500 nM of TSA treatment at 12 h, and 27.4, 50.8 and 77.8% at 24 h, respectively. Furthermore, the downstream apoptotic effector caspase-9 was also demonstrated to be activated accompanied by the MMP loss (Fig. 4B). Collectively, these observations strongly support the notion that TSA-induced apoptosis in 5,637 cells occurred through the intrinsic mitochondrial pathway.

Previously published data explored in various cancer cell studies have indicated a role for PI3K/Akt in the cytotoxocity of TSA (26,27). We next examined whether the PI3k-Akt signaling pathway may also contribute to TSA-induced apoptosis in 5,637 cells. As shown in Fig. 5A, TSA dose-dependently reduced Akt phosphorylation (ser473) after 12 h of treatment. While at 24 h, only 500 nM TSA suppressed Akt phosphorylation. This evidence suggested that TSA-induced suppression of the PI3K/Akt pathway was more effective at early time points. Moreover, when cells were treated with PI3K inhibitor LY294002 (10 µM) alone, pro-apoptotic molecule caspase-9 was also subsequently activated (Fig. 5B), which was consistent with a previous study showing that LY294022 attenuates cell viability in 5,637 cells (28). Given these observations, we suggest that the enhanced apoptosis in 5,637 cells induced by TSA is mediated, at least in part, through the PI3K-Akt pathway.

Reports have previously noted that TSA suppresses Sp1 binding to the promoter of survivin (BIRC5), which belongs to a member of the inhibitor of apoptosis (IAP) family that inhibits caspase activation thereby leading to induction of apoptosis (29,30). Thus, in the present study, we examined whether TSA affects Sp1 and survivin expression in 5,637 cells. Results from Fig. 6A demonstrated that TSA dose-dependently downregulated Sp1 expression after a 24-h treatment, accompanied by a decrease in the survivin protein level (Fig. 6B). However, to further make clear the essential effect exerted by Sp1, we used mithramycin A, which is a prototypic Sp1 inhibitor possessing capacity to displace Sp1 from its transcription sites (31), to block Sp1 activity. As shown in Fig. 6C, the cell growth was markedly suppressed by treatment of mithramycin A. However, pre-treatment of pan-caspase inhibitor for 1 h antagonized the influence of mithramycin A and effectively restored the cell viability (Fig. 6D). This evidence indicated that TSA counteracted Sp1 transcriptional activity, which at least led to downregulation of survivin protein expression following apoptosis at 24 h in 5,637 cells.