Colorectal cancer is a highly prevalent cancer, and the second leading cause of cancer-related death worldwide (Sung
Podophyllotoxin (PT) is a natural lignan derived from the roots and rhizomes of Mayapple
Apoptosis, which is programmed cell death, is essential for normal tissue homeostasis (Qiao and Wong, 2009). Two signaling pathways are involved in activating apoptosis, the extrinsic and intrinsic pathways (Khan
Reactive oxygen species (ROS) including superoxide, the hydroxyl radical, and hydrogen peroxide are highly reactive molecules derived from oxygen (Martindale and Holbrook, 2002). ROS induce the activation of the mitogen-activated protein kinase (MAPK) signaling pathway involved in cellular processes, such as stress adaptation, differentiation, cell growth, proliferation, and apoptosis (Martindale and Holbrook, 2002; Ko
The molecular mechanism of the anticancer effects of PT on colorectal cancer cells has not been reported. This study was conducted to evaluate the anticancer effect of PT on colorectal cancer and determine its apoptotic mechanism related to ROS and p38 MAPK.
Roswell Park Memorial Institute (RPMI)-1640 medium, fetal bovine serum (FBS), trypsin, penicillin/streptomycin (p/s), and phosphate-buffered saline (PBS), were from Hyclone (Logan, UT, USA). podophyllotoxin (PT; purity greater than 98%), dimethyl sulfoxide (DMSO), 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT), N-acetylcysteine (NAC), basal medium eagle (BME), carbobenzoxy-valyl-alanyl-aspartyl-(O-methyl)-fluoromethylketone (Z-VAD-FMK), and p38 inhibitor (SB203580) were procured from Sigma Aldrich (St. Louis, MO, USA). Antibodies specific to cyclin B1, cdc2, β-actin, p21, p27, 78-kDa glucose-regulated protein (GRP78), C/EBP homologous protein (CHOP), death receptor (DR)4, DR5, apoptotic protease-activating factor-1 (Apaf-1), Bid, Bcl-xl, Mcl-1, Bad, caspase 3 and horseradish peroxidase-conjugated secondary antibodies (goat-anti-rabbit, goat-anti-mouse) were obtained from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Antibodies against phosphor (p)-p38 MAPK (Thr180/Tyr182) and p38 were purchased from Cell Signaling Technology (Beverly, MA, USA).
Human colon cancer HCT116 cells were obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA), and maintained in RPMI-1640 medium containing 10% FBS and 100 U/mL penicillin/streptomycin. The cells were incubated at 37°C in a 5% CO2 humidified incubator. To determine the effect of PT on HCT116 cells, the cells were treated with varying concentrations of PT (0, 0.1, 0.2, and 0.3 μM) for 24 or 48 h. DMSO was used as the vehicle.
The MTT assay was used to determine cell viability. HCT116 cells were seeded in a 96-well plate (5,000 cells/well) and treated with PT the next day. After 24 or 48 h of treatment with PT, MTT reagent was added to each well (0.5 mg/mL final concentration) for 1 h incubation at 37°C. Then, the formazan crystals were dissolved by adding 100 μL of DMSO to each well. The absorbance of the samples was measured at 570 nm using a Multiskan GO spectrophotometer (Thermo Scientific, Vantaa, Finland) to assess cell viability. The relative cell viability was calculated compared to the negative controls.
The soft agar assay was used to evaluate anchorage-independent growth. For the bottom layer, 1 mL of 0.6% agar was poured into each well of a 6-well plate with culture medium and PT in DMSO, and allowed to solidify. Then, the upper layers (8×103 cells/well) were seeded with culture medium containing 0.3% agar and PT in DMSO. Seven days later, microscopic observation was done to measure the size and number of colonies (Leica Microsystems, Wetzlar, Germany).
To detect HCT116 cell apoptosis, the Muse™ Annexin V & Dead Cell Kit (MCH100105, EMD Millipore, Billerica, MA, USA) was used following the manufacturer’s instructions. Briefly, cells treated with PT for 48 h were harvested and washed in 1×phosphate-buffered saline (PBS), and stained with the Muse™ Annexin V & Dead Cell reagents (EMD Millipore). The stained cells were incubated at room temperature (RT) for 20 min in the dark. Fluorescence intensity was measured by flow cytometry using a Muse™ Cell Analyzer (EMD Millipore), and the cells were sorted into live (annexin V-/7-AAD-), early apoptotic (annexin V+/7-AAD-), late apoptotic (annexin V+/7-AAD+), and necrotic (annexin V-/7-AAD+) cells. The sum of the early and late apoptotic cells was calculated as the total number of apoptotic cells.
For cell cycle analysis, cells treated with PT were harvested, washed with ice-cold 1×PBS, and 70% ethanol was added for fixing. The cells were incubated overnight at –20°C, and then the fixed cells were collected by centrifugation at 4,000 rpm for 10 min and rinsed with 1×PBS. The cell pellet was resuspended in Muse™ Cell Cycle Reagent (MCH100106, EMD Millipore), and the cells were incubated for 30 min at RT in the dark. The DNA content was assessed by measuring the fluorescence intensity by flow cytometry (Muse™ Cell Analyzer).
The Muse™ Oxidative Stress kit (EMD Millipore) was used to measure intracellular ROS levels. In brief, cells treated with PT were harvested and washed with 1×assay buffer, and then incubated with Muse™ Oxidative Stress Reagent working solution (MCH100111, EMD Millipore) at 37°C for 30 min. The ROS levels were determined by flow cytometry (Muse™ Cell Analyzer).
To assess the changes in the mitochondrial membrane potential (MMP) in PT-treated cells, the cells were stained with MitoPotential dye and 7-AAD using the Muse™ MitoPotential Kit (MCH100110, EMD Millipore). Briefly, cells treated with PT were harvested and washed with 1×assay buffer. The washed cells were resuspended in Muse™ MitoPotential working solution and incubated for 20 min at 37°C. Muse™ MitoPotential 7-AAD reagent was added for 5 min of incubation at RT, then changes in the MMP were determined by flow cytometry (Muse™ Cell Analyzer).
To analyze the activation of multiple caspases including caspase-1, -3, -4, -5, -6, -7, -8, and -9, the Muse™ Multi-caspase Kit (MCH100109, EMD Millipore) was used according to the manufacturer’s instruction. Briefly, cells treated with PT were harvested and washed with 1×caspase buffer. The washed cells were resuspended in Muse™ Multi-caspase Reagent working solution for 30 min at 37°C. Then, the cells were incubated for 5 min after adding Muse™ Caspase 7-AAD working solution. The activation of multiple caspases was analyzed by flow cytometry using a Muse™ Cell Analyzer.
To lyse the cells, the harvested cells were washed with cold 1×PBS, and then resuspended for sonication in RIPA buffer (iNtRON Biotechnology, Seongnam, Korea). A Bio-Rad DC Protein Assay kit was used to determine the protein concentration of the cell lysates (Bio-Rad, Hercules, CA, USA). Then, equal amounts of protein samples were resolved by 6-15% sodium dodecyl sulfate-polyacrylamide gel electrophoresis. The resolved proteins were transferred to poly-vinylidene fluoride membranes (EMD Millipore). The membranes were blocked in 3% or 5% skim milk in 1×PBST (PBS containing 0.1% Tween 20) for 2 h at RT and washed with PBST. The respective primary antibodies were used to probe the membranes. The dilution of the primary antibodies was 1:1,000 in 1×PBST, and the incubation was either 2 h at RT, or overnight at 4°C. Next, the membranes were washed with PBST three times, then incubated with horseradish peroxidase-conjugated secondary antibodies. The dilution of antibodies was 1:4,000-6,000, and the incubation was 1 h at RT. Visualization of Western blots was done using Western Blotting Luminol Reagent (Santa Cruz Biotechnology), and detected with ImageQuant LAS 500 (GE Healthcare, Uppsala, Sweden).
All data are presented as the mean ± standard deviation (SD), of three independent triplicate experiments. One-way ANOVA was used to assess the statistical significance of variation among multiple groups.
To analyze the inhibition of HCT116 cell proliferation by PT (Fig. 1A), the MTT assay was performed. Fig. 1B shows that PT significantly reduced cell viability in dose- and time-dependent manners. The IC50 value was 0.23 μM for 48 h of treatment (Fig. 1B). The soft agar assay results indicated that colony formation was inhibited by treatment with increasing concentrations of PT. HCT116 cells treated with 0.1 μM PT produced 94.9% fewer colonies and 87.24% smaller colonies compared to the untreated control cells (Fig. 1C-1E). We further conducted annexin V-FITC/7-AAD staining, and the cells were analyzed using a Muse™ Cell Analyzer. As the PT concentration increased from 0 to 0.1, 0.2, and 0.3 μM, the percentage of apoptotic cells increased from 2.59% to 18.47%, 28.87%, and 65.90%, respectively (Fig. 1F, 1G). These results suggested that HCT116 cell apoptosis was induced by PT treatment.
We examined the effect of PT on the cell cycle phase populations of HCT116 cells using the Muse™ Cell Analyzer. The percentage of cells in the sub-G1 phase increased from 5.33% in the DMSO-treated group to 19.93%, 34.67%, and 49.27% after treatment with 0.1, 0.2, and 0.3 μM PT, respectively (Fig. 2A, 2B). Compared to the control cells, the percentage of HCT116 cells in the G2/M phase increased as the treatment concentration of PT increased, whereas the G0/G1 phases were decreased, and the S phases were unaffected by PT treatment (Fig. 2A, 2C) Furthermore, we conducted Western blots to assess the expression of G2/M phase-related proteins in HCT116 cells. After PT treatment, cyclin B1 and cdc2 were downregulated in HCT116 cells, and simultaneously p27 and p21 were upregulated in a concentration-dependent manner compared to the control group (Fig. 2D). These observations suggest that the PT-mediated decrease in cell viability were accompanied by G2/M cell cycle arrest.
Our next goal was to investigate whether the PT-induced apoptosis was mediated by the generation of intracellular ROS. We measured the production of ROS in HCT116 cells after treatment with PT for 48 h using a Muse™ Oxidative Stress Kit. The percentage of ROS+ HCT116 cells increased from 5.88% to 42.59%, after treatment with 0 µM to 0.3 µM PT (Fig. 3A, 3B). HCT116 cells were co-treated with 0.3 μM PT and 4 mM ROS scavenger (NAC) and the cell viability was analyzed by the MTT assay. As shown in Fig. 3C, the reduction in cell viability induced by treatment with PT alone was significantly prevented by addition of NAC, indicating that ROS production was essential for PT-induced HCT116 cell death. To determine if the increase in ROS generation induced ER stress, we examined the expression of ER stress markers by Western blot analysis. In PT-treated HCT116 cells, GRP78 and CHOP were up-regulated compared to the controls (Fig. 3D). We further evaluated the levels of TRAIL receptor-associated proteins in HCT116 cells following PT treatment. Treatment with PT resulted in the up-regulation of DR5 and DR4 in concentration-dependent manners (Fig. 3D). These results demonstrated that PT induced ER stress-mediated apoptosis in HCT116 cells via the upregulation of ROS.
We performed Western blotting of HCT116 cells to examine whether PT could regulate activation of the p38 signaling pathway. The treatment of HCT116 cells with 0.3 μM PT increased p38 MAPK phosphorylation (Fig. 4A). To confirm the role of p38 MAPK in mediating PT-induced apoptosis, HCT116 cells were pretreated with 5 μM SB203580 (p38 MAPK inhibitor) for 3 h before treatment with 0.3 μM of PT for 48 h. As shown in Fig. 4B, the apoptotic effect of PT was prevented by pretreatment HCT116 cells with the p38 inhibitor. Taken together, these results suggest that the p38 signaling pathway was a key regulator of PT-induced apoptosis in HCT116 cells.
To see whether PT treatment resulted in the loss of MMP, we monitored the membrane potential of mitochondria after treatment with PT. We observed a concentration-dependent increase in the percentage of all depolarized cells from 6.03 ± 0.30% in the controls to 7.28 ± 0.28%, 26.00 ± 0.72%, and 44.65 ± 1.01% in following treatment with 0.1, 0.2, and 0.3 µM PT, respectively (Fig. 5A, 5B). To investigate the expression levels of apoptotic pathway-related proteins after PT treatment, we performed Western blot assay. When HCT116 cells were incubated with PT, the expression of Bid, Bcl-xl and Mcl-1 was reduced in a dose-dependent manner, whereas Bad expression was increased in a dose-dependent manner. Also, treatment with PT resulted in the increased expression levels of Apaf-1 and the cleavage of pro-caspase 3 in HCT116 cells (Fig. 5C). To determine whether the PT-induced apoptosis could be associated with multi-caspases, caspase activity was analyzed by the Muse™ Multi-caspase Kit. Incubation of HCT116 cells with 0.1, 0.2, and 0.3 µM PT for 48 h increased the percentage of cells with caspase activity in the untreated group from 6.15 ± 0.38% to 8.62 ± 1.12%, 31.61 ± 2.56, and 48.04 ± 1.56%, respectively (Fig. 5D, 5E). Furthermore, HCT116 cells were pretreated with Z-VAD-FMK (4 μM), a pan-caspase inhibitor, for 3 h, and then exposed to 0.3 μM PT for 48 h. Cell viability decreases were significantly prevented in PT and Z-VAD-FMK treatment groups, compared to the viability in the PT-treated groups of HCT116 cells (Fig. 5F) These results suggested that PT induced the apoptosis of colon cancer cells via a caspase-dependent pathway.
PT is a phytochemical exerting various biological activities, and multiple studies have demonstrated the anticancer activity of PT and its derivatives in cancer cell lines and animal experiments. PT inhibited the growth of a human breast cancer cell line by 50% at 1 nM (Chattopadhyay
Apoptosis is characterized by morphological and biological changes including chromatin condensation followed by DNA degradation (Khan
ROS are involved with diverse cellular functions, and low to moderate levels of ROS seem essential for cell viability (Liu
In response to oxidative stress in the ER, the MAPK signaling pathway can be activated (Darling and Cook, 2014). JNK and p38 belong to the MAPK pathway and can be activated through a series of phosphorylations in the signaling pathway when ROS generation increases. The activation of MAPK in this context seems to be dependent upon elevated levels of ROS, as an antioxidant such as NAC could inhibit activation of the MAPK signaling pathway (Son
ER stress can promote the opening of mitochondrial permeability transition pore and induce the loss of MMP, leading to the intrinsic apoptotic pathway (Zeeshan
In conclusion, PT induced G2/M phase cell cycle arrest, the loss of MMP, the activation of multi-caspases, the p38 MAPK signaling pathway, and apoptosis in HCT116 cells. Considering that the anticancer effect of PT was prevented by NAC, increases in ROS generation mediated the anticancer activity of PT. Our results demonstrated that PT induced the apoptosis of HCT116 cells through the upregulation of ROS. More molecular targets of PT are emerging, and the structural versatility of PT derivatives could lead to the development of potential anticancer therapeutics.
We greatly appreciated using the Convergence Research Laboratory (established by the MNU Innovation Support Project in 2019) to conduct this research. This research was funded by the Basic Science Research Program of National Research Foundation Korea, grant number 2019R1A2C1005899.
The authors have no financial conflicts of interest to declare.