LCA (purity ≥98%) was purchased from Zhongxin Pharmaceutical Group Corp., Ltd. (Tianjin, China). Roswell Park Memorial Institute (RPMI) 1640 medium was obtained from Gibco (Thermo Fisher Scientific, Inc., Waltham, MA, USA). Super neonatal bovine serum (NBS) was purchased from Hangzhou Sijiqing Biological Engineering Materials Co., Ltd. (Hangzhou, China). Annexin V/propidium iodide (PI) apoptosis kit was purchased from Nanjing KeyGen Biotech. Co. Ltd. (Nanjing, China). Molecular Probes 2′,7′-dichlorodihydrofluorescein diacetate (H₂DCFDA; Thermo Fisher Scientific, Inc.,), ethylene glycol tetraacetic acid (EGTA), Fluo-3 AM ester, and Hoechst 33258 were purchased from Sigma-Aldrich (St. Louis, MO, USA). BAPTA-AM was purchased from AAT Bioquest, Inc. (Sunnyvale, CA, USA).

The T24 human bladder cancer cells were purchased from the Cell Bank of the Committee on Type Culture Collection of the Chinese Academy of Sciences (Shanghai, China). The cells were maintained in RPMI 1640 medium supplemented with 10% NBS (v/v), 100 U/ml penicillin and 100 μg/ml streptomycin at 37°C in a humidified atmosphere containing 5% CO₂. Cells were allowed to attach for 24 h prior to treatment. LCA was dissolved in dimethyl sulfoxide (DMSO) and diluted with fresh medium to achieve the desired concentration. The final concentration of DMSO did not exceed 0.2% in the fresh medium, and DMSO at this concentration is known to have no significant effect on cell viability.

Cell viability was measured using the sulforhodamine B (SRB) assay (24). Briefly, T24 cells were trypsinized and seeded into 96-well plates at 1.0×10⁵ cells/ml. Subsequently, the cells were exposed to LCA (0, 20, 40 and 80 μM) for 24 h, followed by a further 24 h incubation in fresh medium. The T24 cells were then fixed with trichloroacetic acid and stained for 30 min with 0.4% (wt/vol) SRB (Sigma-Aldrich) dissolved in 1% acetic acid. Unbound dye was removed by four washes with 1% acetic acid, and protein-bound dye was extracted with 150 ml DMSO for determination of optical density was detected at a wavelength of 490 nm using a Varioskan Flash 3001 plate reader (Thermo Fisher Scientific, Inc.).

Apoptosis was assessed using Annexin V-fluorescein isothiocyanate (FITC) and PI labeling, as previously described (25). Cells were treated with 0, 20, 40 and 80 μM LCA for 24 h. Subsequently, the cells were washed twice with phosphate-buffered saline (PBS), and were resuspended in staining buffer containing 5 μl (1 mg/ml) PI and 5 μl Annexin V-FITC. Double-labeling was performed at room temperature for 10 min in the dark prior to flow cytometric analysis. Cell staining was detected using a FACStar flow cytometer (BD Biosciences, Franklin Lakes, NJ, USA).

Alterations in the nuclear morphology of apoptotic cells were observed by labeling the cells with the nuclear stain Hoechst 33258, and examining them under a fluorescent microscope. After treatment with 0, 20, 40 and 80 μM LCA for 24 h, the cells were fixed in formaldehyde (40 g/l) in PBS for 20 min, followed by Hoechst 33258 (10 mg/l) staining for 30 min in the dark at 37°C. Nuclear morphology was subsequently observed under a fluorescence microscope (MIC00266; Zeiss, Oberkochen, Germany) (26).

Ca²⁺ concentration was measured by Fluo-3 AM staining and microscopy. Briefly, T24 cells were incubated with or without Ca²⁺ chelators (200 μM EGTA and 10 μM BAPTA-AM) for 1 h prior to 5 μM LCA treatment for 24 h. Subsequently, the cells were harvested and washed twice, and were resuspended in Fluo 3 AM (5 μM) at 37°C for 30 min. After washing three times, the stained cells were observed under a computer-assisted microscope (MIC00266; Carl Zeiss AG, Oberkochen, Germany) at an excitation wavelength of 488 nm and an emission wavelength of 525 nm.

The intracellular levels of ROS in T24 cells were assessed using H₂DCFDA (27). Briefly, cells were incubated with 55 μM LCA or with 10 μM BAPTA-AM (a chelator of cytosolic Ca²⁺) for 1 h prior to LCA treatment. The treated cells were then washed in PBS and incubated with 30 μM H₂DCFDA for 30 min at 37°C. Fluorescence was detected using a fluorescent plate reader at 485/525 nm excitation/emission wavelengths (Varioskan Flash 3001; Thermo Fisher Scientific, Inc.) and data were expressed as median fluorescence intensity. The stained cells were then observed under a computer-assisted microscope (MIC00266; Carl Zeiss AG).

The MMP was assessed using a dual-emission potential-sensitive probe, 5,5′,6, 6′-tetra-chloro-1, 1′,3,3′-tetraethyl-imidacarbocyanine iodide (JC-1; KeyGen Biotech Co., Ltd., Nanjing, China). The ratio of red to green fluorescence of JC-1 depends solely on membrane potential, with a decrease being indicative of membrane depolarization (28). Briefly, the cells were exposed to LCA (0, 20, 40 or 80 μM) for 4, 8, 16 or 24 h. Subsequently, the cells were loaded with 2 mg/l JC-1 at 37°C for 20 min, and were analyzed using a plate reader (Varioskan Flash 3001; Thermo Fisher Scientific, Inc.).

Total RNA was extracted from T24 cells using TRIzol (Sangon Biotech Co., Ltd., Shanghai, China). RNA quality was determined using the A260/A280 ratio and 1.5% agarose gel electrophoresis. cDNA synthesis was performed using Moloney murine leukemia virus reverse transcriptase with a First Strand cDNA Synthesis kit (Fermentas; Thermo Fisher Scientific, Inc.). The synthesized cDNA was amplified by Riboloek Nase-free ddH₂O (8 μl), template RNA (3 μl), Oligo(dT)18 (1 μl) to a final volume of 12 μl. Tubes were placed into the C1000 Thermal Cycler (Bio-Rad Laboratories, Inc., Hercules, CA, USA) at 70°C for 5 min, following which 5X reaction buffer (4 μl), Riboloek Nase Indihitor (1 μl), dNTP M-M RT RTase (1 μl) and RNase DEPC-treated water to a final volume of 20 μl were added and incubated at 42°C 1 h for denaturation and 70°C 5 min for annealing. The synthesized cDNA was amplified by Oligo(dT)₁₈, according to the instructions of a PCR Amplification kit (Fermentas; Thermo Fisher Scientific, Inc.). The PCR primers (synthesized by Sangon Biotech Co., Ltd.; presented in Table I) and their cycling conditions were set as indicated. The PCR reaction volume consisted of 12.5 μl 2X PCR Master (Sangon Biotech Co., Ltd.), 3 μl cDNA template and 0.5 μl of each primer. The cycling conditions were as follows: Pre-denaturation at 94°C for 3 min, with 30–35 cycles of denaturation at 94°C for 30 sec, annealing for 30 sec, extension at 72°C and a final extension at 72°C for 10 min, using a C1000 Thermal Cycler (Bio-Rad Laboratories, Inc.). The gene products were quantified using agarose gel electrophoresis (Biodee Biotechnology Co., Ltd., Beijing, China) and a Bio-Rad gel imaging system (Bio-Rad Laboratories, Inc.).

The quantitative PCR analysis was carried out according to the manufacturer's protocol of the Taqman One-Step PCR Master Mix (Applied Biosystems; Thermo Fisher Scientific, Inc.). Total cDNA (2 μl) was added per 25 μl reaction, alongside 0.5 μl sequence-specific primers and 12.5 μl SYBR Premix Ex Taq. All target gene primers and probes were purchased commercially (Sangon Biotech Co., Ltd.). Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as an internal control. The cycling conditions were as follows: Pre-denaturation at 95°C for 4 min, denaturation at 95°C for 5 sec, annealing for 30 sec, extension at 72°C and a final extension at 72°C for 10 min, using a Rotor-Gene Q Real time PCR machine (Qiagen China Co., Ltd, Shanghai, China). Relative expression levels of the target genes were calculated based on the 2^−ΔΔCq method of relative quantification (29), according to the following equation: Relative expression level = 2(^{Cq value of GAPDH - Cq value gene of interest}). The primer sequences are presented in Table I.

Data are presented as the mean ± standard from at least three independent experiments. Data were evaluated by one-way analysis of variance followed by Student-Newman-Keuls test, uding Origin software, version 8.0 (OriginLab, Northampton, MA, USA). In all cases, P<0.05 was considered to indicate a statistically significant difference.

To examine cell viability in vitro, the SRB assay was used to determine the inhibitory effects of LCA on proliferation. A total of 24 h post-LCA treatment, LCA reduced the proliferation of T24 cells in a dose-dependent manner; the half maximal inhibitory concentration was ~55 μM (P=4.3×10⁻¹³; Fig. 1A). Subsequently, it was investigated whether LCA was able to induce cell death through an apoptotic mechanism. Annexin V-FITC and PI double-labeling was used for the detection of phosphatidylserine externalization, a hallmark of early phase apoptosis. Compared with the control group, a high proportion of Annexin V⁺ labeling was detected in cells treated with LCA, thus indicating that they were in the early phase of apoptosis (P=1.3×10⁻⁵; Fig. 1B and C). These results indicate that LCA significantly induced apoptosis in T24 cells.

Typical apoptotic morphological alterations, as indicated by condensed nuclei and nuclear fragmentation, were apparent after exposure to 40 μM LCA. Apoptotic nuclear alterations were markedly increased in the cells pretreated with 80 μM LCA (Fig. 2).

Following treatment of T24 cells with LCA, Ca²⁺ levels were significantly increased compared with the control group. These results indicate that LCA promoted Ca²⁺ release in a time-dependent manner (30 min, P= 4.3×10⁻¹³; 2 h, P=1.2×10⁻⁵; 4 h, P=0.0084; Fig. 3A). In order to investigate whether LCA could induce an increase in cytosolic Ca²⁺ through extracellular or intracellular Ca²⁺ pools, the T24 cells were pretreated with EGTA (an extracellular Ca²⁺ chelator) and BAPTA-AM (an intracellular Ca²⁺ chelator). EGTA and BAPTA-AM significantly suppressed LCA-induced Ca²⁺ release (P=1.9×10⁻⁶ and P=2.2×10⁻⁶, respectively; Fig. 3B). Furthermore, BAPTA-AM could attenuate LCA-induced apoptosis, whereas EGTA had no inhibitory effects on apoptosis (Fig. 3C). Collectively, these results indicate that LCA mainly induced release of intracellular Ca²⁺ to promote apoptosis.

To determine whether cytosolic Ca²⁺ release is a signal leading to ROS accumulation in LCA-induced apoptosis, T24 cells were pretreated with BAPTA-AM, then treated with LCA for 1 h, or the cells were treated with LCA alone. The levels of ROS generation were lower in the LCA and BAPTA-AM treated group, as compared with in the LCA-treated group (apoptotic rates, P= 0.0241; ROS, P= 0.0128; Fig. 3D). These results suggest that BAPTA-AM may decrease LCA-induced apoptosis and ROS generation, thus indicating that cytosolic Ca²⁺ release may act upstream of ROS generation in LCA-treated T24 cells.

As shown in Fig. 4A, treatment of T24 cells with LCA for 4, 8, 16 and 24 h induced a time-dependent reduction in MMP (P=6.0×10⁻⁸, P=9.5×10⁻⁷, P=0.0793, P=1.1×10⁻⁵, respectively), thus suggesting that LCA-induced cell apoptosis may be associated with mitochondrial dysfunction.

To further explore whether LCA induces apoptosis via the regulation of mitochondrial apoptosis-associated genes, B-cell lymphoma (Bcl)-2-associated X protein (Bax), Bcl-2-interacting mediator of cell death (Bim), Bcl-extra large (xL), apoptotic protease activating factor-1 (Apaf-1), caspase-9 and caspase-3 were detected in T24 cells. The cells were treated with various concentrations of LCA for 24 h. As shown in Fig. 4B and C, the mRNA expression levels of Bcl-xL were downregulated (P=0.052), whereas Bax (P= 4.1×10⁻¹²), Bim (P=2.4×10⁻⁸), Apaf-1 (P=1.8×10⁻¹⁴), caspase-9 (P= 0.0107) and caspase-3 (P= 0.0001) expression levels were upregulated in a concentration-dependent manner.

To determine whether intracellular Ca²⁺ levels exert a critical role on LCA-induced mitochondrial apoptosis, BAPTA-AM, an intracellular Ca²⁺ chelator, was used. T24 cells were treated with or without BAPTA-AM for 1 h prior to LCA treatment for 24 h. As shown in Fig. 5, compared with the LCA-treated group, Bax (P=0.0374), Apaf-1 (P=5.7×10⁻⁵), caspase-9 (P=0.0211) and caspase-3 (P=0.0095) expression were markedly downregulated. However, there was no significant difference in Bcl-xL mRNA expression between the LCA-treated and LCA + BAPTA-AM-treated groups. These data suggest that intracellular Ca²⁺ has a critical role in mitochondrial apoptosis.

The involvement of ER stress signaling in the responses triggered by LCA-induced apoptosis was evaluated based on the expression of calpain 2 and caspase-4. As shown in Fig. 6, cells were treated with LCA for 24 h, and calpain 2 and caspase-4 expression levels were increased in a concentration-dependent manner. Subsequently, the T24 cells were treated with or without BAPTA-AM for 1 h prior to LCA treatment for 24 h,. As shown in Fig. 7, compared with the LCA-treated group, calpain 2 (P=0.0418) and caspase-4 (P=3.8×10⁻⁵) expression levels were downregulated in response to BAPTA-AM treatment. These results suggest that Ca²⁺ is involved in the ER stress-related apoptotic pathway.