The human bladder cell line T24 was purchased from the American Type Culture Collection (Manassas, VA, USA). Cells were maintained in RPMI-1640 medium (Hyclone, Logan, UT, USA) supplemented with 10% FBS (Hyclone) and antibiotics under a humidified atmosphere with 5% CO₂.

Casticin was purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA), dissolved in sterile dimethyl sulfoxide (DMSO) as a 200-mM stock solution, and diluted in medium to the indicated concentration before use. z-LEHD-fmk (z-Leu-Glu(OMe)-His-Asp-(OMe)-fluoremethylketone), z-DEVD-fmk (N-benzyloxycarbonyl-Asp-Glu-Val-Asp-fluoromethylketone), z-VAD-fmk (N-benzyloxycarbonyl-Ile-Glu-Thr-Asp-fluoromethyl ketone), SB203580, and SP600125 were purchased from Calbiochem (San Diego, CA, USA). N-acetylcysteine (NAC) was obtained from Sigma-Aldrich (St. Louis, MO, USA). Necrostatin-1 was purchased from Selleckchem (Houston, TX, USA).

The percentage of T24 cells undergoing apoptosis was determined by flow cytometry using FITC-labeled Annexin-V (BD Biosciences, San Diego, CA, USA) and 7-aminoactinomycin D (7-AAD) (BD Biosciences). To determine optimal conditions, experiments were performed using different concentrations of casticin (0, 10, 20, 50, 100 and 200 µM) and different periods of incubation (2, 4, 8, 24 and 48 h). DMSO (0.05%) was used as a vehicle control. To examine the role of caspases, cells were pretreated with z-LEHD-fmk (20 µM, caspase-9 inhibitor), z-DEVD-fmk (20 µM, caspase-3 inhibitor) or z-VAD-fmk (20 µM, pan-caspase inhibitor) for 2 h before casticin treatment. Cells were harvested, rinsed with PBS, and resuspended in 100 µl of 1X Annexin-V binding buffer [10 mM HEPES/NaOH (pH 7.4), 140 mM NaCl, 2.5 mM CaCl₂]. After addition of 3 µl of Annexin-V-FITC and 3 µl of 7-AAD the cells were incubated at room temperature for 15 min in the dark with gentle vortexing. Stained cells were analyzed using a FACSCalibur flow cytometer (BD Biosciences) equipped with CellQuestpro software (BD Biosciences).

Changes in mitochondrial membrane potential were measured using DiOC₆ (3,3′-dihexyloxacarboxyanine iodide; Molecular Probes, Eugene, OR, USA). Cells were treated with casticin or DMSO for 24 h, washed twice in PBS, resuspended in PBS supplemented with DiOC₆ (20 nM), incubated at 37°C for 15 min in the dark, and immediately analyzed with a flow cytometer using the FL-1 filter.

Intracellular accumulation of ROS was monitored by flow cytometry after staining with the fluorescent probe DCFH-DA (10 µM, 2′,7′-dichlorodihydro-fluorescein diacetate; Molecular Probes) as previously described (15) with slight modification. DCFH-DA is deacetylated in cells by esterases to yield a non-fluorescent compound, DCFH, which remains trapped within the cell and is cleaved and oxidized by ROS in the presence of endogenous peroxidase to yield a highly fluorescent compound, DCF (2′,7′-dichlorofluorescein). Cells were preincubated with 10 µM DCFH-DA for 30 min at 37°C and then seeded in 6-well plates (5×10⁵ cells/ml) and treated with or without casticin for 24 h. Cells were washed, resuspended in PBS, and ROS levels were determined using a FACSCalibur flow cytometer.

Cells were washed in PBS and lysed in NP-40 buffer (Elpis Biotech, Daejeon, Korea) supplemented with a protease inhibitor cocktail (Sigma-Aldrich). To address phosphorylation events, an additional set of phosphatase inhibitors (Cocktail II, Sigma-Aldrich) was added to the NP-40 buffer. Protein concentration was determined using a BCA assay kit (Pierce, Rockford, IL, USA). Proteins (10 µg/sample) were resolved by SDS-PAGE and transferred to nitrocellulose (Millipore Corp., Billerica, MA, USA). The membranes were blocked with 5% skim milk and western blot analysis was performed using a standard commercial method. Immunoreactivity was detected using an enhanced chemiluminescence (ECL) kit (Advansta Corp., Menlo Park, CA, USA) and Multiple Gel DOC system (Fujifilm). Primary antibodies (Abs) against the following proteins were used: caspase-8, caspase-3, caspase-9, PARP, β-actin, Bcl-2, Bax, Mcl-1, Bak, p53, XIAP, RIP, phospho-JNK (Thr¹⁸³/Tyr¹⁸⁵), JNK, phospho-p38-MAPK (Thr¹⁸⁰/Tyr¹⁸²), p38-MAPK, phospho-ERK1/2 (Thr²⁰²/Tyr²⁰⁴), and ERK1/2 (Cell Signaling Technology, Beverly, MA, USA); XAF1 and TAp73 (Abcam, Cambridge, UK); TAp63 (BioLegend, San Diego, CA, USA).

Experimentally verified human TAp73-siRNA duplex and negative control-siRNA were obtained from Bioneer (Daejeon, Korea). Cells were transiently transfected under optimized conditions. Briefly, cells were seeded at a concentration of 1×10⁵ cells per well in a 6-well plate and grown overnight. Cells in each well were transfected with 200 nM siRNA using Lipofectamine RNAiMAX Reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions. Cells were used for further experiments 48 h after transfection.

Data were expressed as mean ± standard deviation (SD). Statistical analysis was conducted using one-way analysis of the variance (ANOVA). P-values <0.05 were considered statistically significant.

To evaluate whether casticin affects cell survival through regulation of the mitochondria membrane potential, the percentage of apoptotic cells in casticin-treated T24 cells was determined by flow cytometry using FITC-labeled Annexin-V and 7-AAD and the mitochondria membrane potential was measured by staining with DiOC₆. Casticin treatment of T24 cells resulted in a dose- and time-dependent increase in the Annexin-V⁺/7-AAD⁺ population (Fig. 1A and B). Casticin also significantly disrupted the integrity of the mitochondria membrane after 8 h of treatment (Fig. 1A and B). We further analyzed the activity of caspases by immunoblotting. Levels of cleaved forms of caspase-8, caspase-9 and caspase-3 were increased in T24 cells after treatment with casticin. Cleavage of PARP was also detected in the presence of casticin (Fig. 1C). Treatment with specific caspase inhibitors z-LEHD-fmk (caspase-9 inhibitor) or z-DEVD-fmk (caspase-3 inhibitor) or the pan-caspases inhibitor z-VAD, effectively blocked casticin-mediated apoptosis (Fig. 1D). These results suggest that casticin-induced apoptosis of bladder cancer cells involves interruption of the mitochondria membrane potential and activation of caspases.

On the basis of previous results we investigated the expression of XAF1, apoptosis-related proteins, and p53 family proteins to elucidate the underlying mechanism of casticin-induced T24 cell apoptosis. Although casticin treatment had no influence on the expression level of p53 and induction of TAp63, the level of TAp73 was markedly elevated in casticin-exposed T24 cells. XAF1 expression was also upregulated, whereas XIAP was downregulated (Fig. 2A). The levels of antiapoptotic proteins, including Mcl-2 and Bcl-2, were decreased, whereas expression of proapoptotic proteins (Bak and Bax) was increased (Fig. 2B). Because JNK and p38 MAPK regulate phosphorylation and activation of TAp73 (18,19), we next examined whether activation of mitogen-activated protein kinases (MAPKs) is correlated with the induction of TAp73 in casticin-treated T24 cells. Casticin significantly induced the phosphorylation of JNK and p38 MAPK, whereas activation of ERK was blocked (Fig. 2C). Pharmacological inhibition of JNK and p38 with SP600125 and SB203580, respectively, not only suppressed the activation of caspase-9 and caspase-3 (Fig. 2D), but also attenuated the casticin-induced expression of XAF1, TAp73, Bak and Bax (Fig. 2E and F). These results suggest that MAPKs are involved in the upregulation of XAF1 and TAp73 in T24 cells after treatment with casticin.

We next investigated whether the apoptotic effect of XAF1 is regulated by the expression and activation of TAp73 in casticin-exposed T24 cells. To define the relationship between TAp73 activation and induction of XAF1, we analyzed the expression of XAF1 and activation of MAPKs in casticin-treated T24 cells after silencing TAp73 with siRNA. Knockdown of TAp73 with siRNA resulted in downregulation of XAF1 but had no effect on the activation of JNK and p38 MAPK (Fig. 3A). Gene silencing of TAp73 effectively blocked the cleavage of caspase-9, caspase-3, and PARP in casticin-treated T24 cells (Fig. 3B). In addition, the level of proapoptotic proteins (Bak and Bax) was suppressed in TAp73-knockdown T24 cells after treatment with casticin (Fig. 3C). As ROS production is an early signal mediator for apoptosis induction after treatment with casticin (3,6,20) we examined the effect of ROS on XAF1 and TAp73 activation in casticin-treated T24 cells. Casticin upregulated intracellular ROS levels within 1 h (Fig. 4A). Moreover, NAC (a ROS scavenger) not only efficiently inhibited ROS production (Fig. 4B), but also blocked generation of the cleaved forms of caspase-9, caspase-3, and PARP (Fig. 4C). Furthermore, expression of XAF1, TAp73, phosphorylated p38, and phosphorylated JNK was suppressed by NAC treatment (Fig. 4D). These results suggest that ROS-mediated TAp73 expression is one of the critical factors for activation of XAF1 and induction of apoptosis in casticin-treated T24 cells.

RIP kinases mediate apoptosis or necrosis induced by tumor necrosis factor (TNF)-α or genotoxic stresses through the accumulation of ROS (21,22). ROS are also closely associated with the caspase cascade and activation of MAPKs (23). Our previous results showed that casticin promoted ROS-induced MAPK phosphorylation to activate caspases and induce apoptosis of T24 cells. Therefore, we examined whether RIP kinases are involved in ROS production for activation of the TAp73/XAF1 signaling pathway in casticin-treated T24 cells. Casticin elicited RIP kinase expression (Fig. 5A). Treatment with necrostatin-1 (a RIP kinase inhibitor) blocked the cleavage of caspases (Fig. 5B), attenuated ROS generation (Fig. 5C), and suppressed MAPK-mediated TAp73/XAF1 activation (Fig. 5D) in T24 cells after treatment with casticin. However, gene silencing of TAp73 with siRNA and treatment with MAPK inhibitors (SP600125 and SB203580) failed to block the expression of RIP kinase in casticin-treated T24 cells (Fig. 5E and F). In addition, NAC treatment of casticin-exposed T24 cells had no influence on the activation of RIP kinase (Fig. 5G). These results suggest that the activation of RIP kinase plays a critical role in casticin-induced apoptosis through ROS production.