Bispidinone and bispidinone analogs were obtained from the laboratory of Dr D.H. Park. Stock solution of the samples was dissolved in dimethyl sulfoxide (DMSO) as 20 mM and kept at 4°C. Further dilutions were made immediately prior to each experiment. Eagle’s minimal essential medium (EMEM), penicillin-streptomycin and trypsin-EDTA were purchased from HyClone (Logan, UT, USA). Fetal bovine serum (FBS) was obtained from Gibco-BRL (Carlsbad, CA, USA). The Cell Counting Kit-8 (CCK-8) was obtained from Dojindo (Osaka, Japan). The propidium iodide (PI)/RNase staining buffer and Annexin-FITC kit for apoptosis were from BD Pharmingen (San Diego, CA, USA).

HeLa cells were obtained from the American Type Culture Collection (ATCC; Manassas, VA, USA) and were cultured in EMEM supplemented with 10% FBS and 1% penicillin-streptomycin at 37°C in a humidified atmosphere with 5% CO₂.

HeLa cells were plated at 5×10³ cells/well in a 96-well microplate. After 24 h, the media were substituted with fresh media containing B16 at concentrations of 7.5, 15 and 30 μM. The plate was incubated for 48 h and cell viability was assessed using a WST-8 assay according to the manufacturer’s recommendations (24). CCK-8 solution (10 μl) was added to each well of the plate followed by 2-h incubation. The optical density for viable cells was read at 450 nm in a multimicroplate reader (Synergy HT; BioTek Instruments, Inc. Winooski, VT, USA). For the effect of B16 on HeLa cell proliferation, the cells were seeded at 5×10³ cells/ml media in 96-well plates and treated with or without B16 (25 μM) at various time points. Each experiment was repeated at least three times.

HeLa cells were distributed (2×10⁵ cells/well) into a 6-well plate and allowed to adhere overnight. The cells were treated with B16 (25 μM) for 24 and 48 h. Non-treated wells received an equivalent volume of DMSO (<0.1%) as a control. Optic phase-contrast images were captured with a Phase Contrast-2, ELWD 0.3 inverted microscope (Nikon, Japan).

Cells (3×10⁵) in a 60-mm Petri dish, treated with or without B16, were collected by trypsinization and washed with ice-cold phosphate buffered saline (PBS) via centrifugation at 2,500 g for 3 min. Cells (1×10⁵) were resuspended in 100 μl of binding buffer and stained with 5 μl of Annexin V-FITC and 10 μl of PI (50 μg/ml) for 15 min at room temperature, in the dark. Analysis was performed by FACSCalibur flow cytometer (Becton-Dickinson, San Jose, CA, USA) with 10,000 events/analysis. The data were analyzed using Cell Quest Pro software (Becton-Dickinson Instruments, Franklin Lakes, NJ, USA).

Cells (3×10⁵) in a 60-mm Petri dish, treated with or without B16, were collected by trypsinization and washed with ice-cold PBS via centrifugation as above. The cells were suspended in PBS and fixed with 70% ethanol (v/v). Samples were washed with ice-cold PBS and stained with PI/RNase staining buffer for 15 min at room temperature. The number of cells in different phases of the cell cycle was analyzed using a FACScan flow cytometer analysis system and 20,000 events were analyzed each time. The percentage of cells in different phases of the cell cycle was determined using Modifit software (Becton-Dickinson Instruments).

Briefly, the HeLa cells were applied to 12-well plates in growth medium (EMEM + 10% FBS + 1% penicillin-streptomycin). After the cells had grown to 70–80% confluence, they were rendered quiescent by incubation for 24 h in EMEM containing 2% FBS. The cells were treated with or without B16 in EMEM supplemented with 10% FBS and the cultures were incubated for 21 or 45 h. [³H]-dTTP was added at 1 μCi/ml (1 μCi=37 kBq) and incubated for an additional 3 h. Incorporated [³H]-dTTP was extracted in cell lysis buffer and measured in a liquid scintillation analyzer (Tris-Crab 2910TR; Perkin-Elmer Inc., Waltham, MA, USA) (25).

Generation of ROS was assessed using the fluorescent indicator 2,7-dichlorodihy-drofluorescein (H₂DCF-DA), a cell-permeable indicator for ROS shown to react with H₂O₂ (26). As previously described, H₂DCF-DA was oxidized to highly green fluorescent 2,7-dichlorofluorescein (DCF) by the generation of ROS. Cells (3×10⁵) in a 60-mm dish, treated with or without B16, were collected by trypsinization and centrifugation as describe above. The samples were washed with ice-cold PBS and stained with 2 μl H₂DCF-DA for 30 min at 37°C in the dark. Relative fluorescence intensities were monitored using the FACSCalibur flow cytometer and analyzed using CellQuest software with the FL-1 channel (green) set to 530 nm.

Rhodamine 123 was used as a fluorescent probe to detect the depolarization of mitochondrial-membrane potential. During apoptosis, the integrity of the mitochondrial membrane is disrupted, leading to depolarization of the membrane, opening of mitochondrial permeability transition pores and release of sequestered rhodamine 123. Cells (3×10⁵) in a 60-mm Petri dish), treated with or without B16, were collected by trypsinization and centrifugation as described above. The samples were washed with ice-cold PBS and stained with 5 μl rhodamine 123 for 30 min at 37°C in the dark. Relative fluorescence intensities were monitored using a FACSCalibur flow cytometer and analyzed using Cell Quest software with the FL-1 channel.

Following treatment of cells with or without B16, total cell lysates was prepared as previously described. Protein contents of the lysates were determined by the Bradford protein assay (Bio-Rad, Hercules, CA, USA). Proteins (8 μg) were resolved by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred onto nitrocellulose membranes (Schleicher & Schuell, BioScience, Inc., Keene, NH, USA) by western blotting. The results were quantified using Image J v. 1.43 software (National Institutes of Health, Bethesda, MD, USA). The following primary polyclonal antibodies were used: β-actin, Bcl-2, procaspase-9, procaspase-8 (1:1,000 dilution; Cell Signaling Technology Inc., Danvers, MA, USA), procaspase-3, p53 (1:300; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA) and Bax (1:1,000; BD Pharmingen, San Diego, CA, USA).

Results reported were obtained from at least three independent experiments with similar results. The results are presented as mean ± standard deviation (SD) in quantitative experiments. Statistical analysis was performed by one-way analysis of variance (ANOVA). ^*P<0.05 was considered to indicate a statistically significant difference. Microsoft Excel 2007 was used for statistical and graphical evaluations.

The cytotoxicity of B16 was determined using WST-8 assays. After incubation with increasing concentrations (7.5, 15, and 30 μM) for 48 h, B16 showed significant cytotoxicity in HeLa cells in a concentration-dependent manner, compared to the control (Fig. 2A). The IC₅₀ value of B16 was 22.66 μM. In addition, the proliferation of HeLa cells treated with 25 μM of B16 was measured at 12-h intervals for 60 h. As shown in Fig. 2B, the proliferation of HeLa cells treated with B16 continuously increased during the first 12 h, and then the proliferation rate observed at 12 h was maintained until 60 h. By contrast, the control cells maintained an exponential proliferation state throughout the 60-h incubation. These results suggested that B16 reduced the viability of HeLa cells in a dose- and time-dependent manner.

Morphological observation using light microscopy was performed to study B16-induced apoptosis in HeLa cells. As shown in Fig. 3A, the untreated cells spread regularly in culture plates and grew to near confluence. After 24 h of treatment with B16, some of the cells floated in the culture plates, while the majority of the attached cells maintained a shrunken shape. After 48 h of treatment, a significant proportion of the HeLa cells dislodged from the plates and the remaining adherent cells showed typical morphological changes, such as shrinkage, floating, and an increase in intercellular space. To assess the induction of apoptosis by B16 in HeLa cells, an Annexin V and PI double staining assay was performed. A 48-h treatment with B16 significantly increased the percentage of cells undergoing early apoptosis from 5.27 (0 h) to 25.11% (48 h) (Fig. 3B). In addition, the percentage of cells in late apoptosis (necrotic cells) was also increased by B16 treatment (Fig. 3B).

Cell growth and inhibition are tightly mediated through cell cycle control mechanisms (27). To examine the effects of B16 on the cell cycle, a DNA content assay was performed to analyze the cell cycle distribution of HeLa cells treated with 25 μM B16. In the cells undergoing apoptosis, a DNA peak that is characteristic of apoptosis (usually known as the sub-G0/G1 peak or apoptotic peak) appeared (Fig. 4). After 24 h of treatment, B16 induced a significant accumulation of cells in the S phase (21.44% in control cells vs. 29.21% in the B16-treated cells). Concomitantly, there was a significant decrease of cells in the G1 phase from 64.95 in the control cells to 48.88% in the B16-treated cells. The level of S-phase arrest induced by B16 in HeLa cells did not appear to be time dependent.

The results of the DNA content assay suggested that HeLa cells treated with B16 accumulated in the S phase. To confirm this hypothesis, we analyzed DNA replication in HeLa cells treated with B16 by determining the incorporation of [³H]-dTTP. In cells treated with B16 for 24 h and 48 h, [³H]-dTTP incorporation was reduced by 30 and 60%, respectively, compared to the controls (Fig. 5). This result suggested that B16 had an inhibitory effect on the process of DNA replication.

ROS plays an increasingly important role in apoptosis-induced cell death (28). Therefore, we determined the effect of B16 on ROS generation by measuring the fluorescence of the cell-permeable dye H₂DCF-DA in HeLa cells during a 6-h treatment with 25 μM B16. The mean H₂DCF-DA fluorescence value increased from 49.78 (0 h) to 101.98 (3 h) following treatment with B16, whereas the value of H₂DCF-DA fluorescence decreased after the 3-h time point (Fig. 6A). These results indicate that treatment with B16 induced changes in ROS generation.

The generation of ROS is associated with the disruption of the mitochondrial-membrane potential, and mitochondrial-membrane depolarization is considered to be a crucial hallmark of apoptosis. In our study, rhodamine 123 was used as a fluorescent probe to detect the loss of MMP. Fig. 6B shows that the mean rhodamine 123 fluorescence decreased from 86.97 (0 h) to 82.8 (12 h) and 74.83 (24 h) following treatment with 25 μM B16, suggesting that mitochondria were involved in the observed B16-induced apoptosis.

To investigate the underlying mechanism of apoptosis induced by B16, the expression levels of several apoptosis-associated proteins were determined by western blot analysis. As shown in Fig. 7A, a decrease in procaspase-3 and −9 expression levels was observed in cells treated with B16, demonstrating the activation of the caspase cascade in these cells. We also assessed the expression level of p53, a known tumor suppressor, and found that it increased following treatment with B16 (Fig. 7A). This result suggests that B16 suppresses tumor growth. B16 treatment also reduced the expression of procaspase-8 (Fig. 7B). As caspase-8 is known to be involved in the Fas signaling apoptotic pathway, this result suggested that B16-induced apoptosis is mediated by the Fas signaling pathway. The expression level of the pro-apoptotic protein Bax was significantly upregulated by B16 treatment in a time-dependent manner, whereas the expression level of the anti-apoptotic protein Bcl-2 showed a tendency for downregulation compared to the control. Correspondingly, a marked increase in the Bax/Bcl-2 ratio was observed (Fig. 7C). The changes in expression of Bcl-2 family members further confirmed that the mitochondrial pathway is involved in B16-induced apoptosis. Moreover, the caspase cascade is associated with the intrinsic and extrinsic apoptotic pathways. Thus, taken together, these results suggested that B16 induced apoptosis in HeLa cells via a caspase- and mitochondrial-dependent pathway.