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. 2020 Feb 17;10(3):123. doi: 10.1007/s13205-020-2111-z

Cell death induction by Ranunculus ternatus extract is independent of mitochondria and dependent on Caspase-7

Ming Fang 1, Takahisa Shinomiya 1, Yukitoshi Nagahara 1,✉
PMCID: PMC7026323  PMID: 32123647

Abstract

Ranunculus ternatus is a traditional Chinese medicine with an anticancer effect, but its underlying mechanism is unknown. In this study, we demonstrated by MTT assay that ethyl acetate extract (RTE) from R. ternatus exerts cytotoxic effects on human T cell lymphoma Jurkat cells. Then, to test the apoptosis induction ability of RTE to induce apoptosis, we analyzed phosphatidylserine exposure, DNA fragmentation, and caspase cleavage. RTE induced phosphatidylserine exposure and caspase-7 cleavage, but not caspase-3 cleavage. Sub-G1 cells were accumulated but DNA fragmentation was not observed. A pan-caspase inhibitor Z-Asp-CH2-DCB suppressed RTE-induced caspase cleavage and the above-described events. RTE also induced cell death in caspase-3 null human breast cancer MCF-7 cells, indicating that RTE-induced apoptotic-like cell death depends on the activation of one or more caspases, but not caspase-3. Moreover, RTE-induced cell death was not suppressed in Bcl-2 overexpressing Jurkat cells, suggesting that mitochondria were not involved in RTE-induced cell death. In conclusion, RTE-induced cell death was independent of mitochondria and dependent on caspase-7.

Keywords: Ranunculus ternatus, Cell death, Caspase, DNA fragmentation

Introduction

Cancer is a major public health problem worldwide (David and Zimmerman 2010), and the World Health Organization recently indicated that cancer deaths continue to increase (McGuire 2016). Cancer is a genetic disease; due to genetic mutation(s), cancer cells proliferate indefinitely, form malignant tumors, and invade surrounding healthy tissues (Ouyang et al. 2012). When a tumor's growth is localized, the cancer can usually be cured by the surgical removal of the tumor, but malignant tumors can form relocated lesions that easily become metastatic and cannot be treated with surgery. Chemotherapy and radiotherapy are usually administered for metastatic tumors (Nielsen et al. 2017), but these treatment modalities cannot selectively kill cancer cells without harming normal cells, and they often have serious side effects (Guo et al. 2014). Therapeutic methods that selectively kill cancer cells and present few side effects are needed.

Certain components extracted from traditional Chinese medicine have been reported to be injurious to cancer cells and to have less toxicity to normal cells (Karimian et al. 2014). This suggests that traditional Chinese medicine is less likely to damage normal cells and may still attack cancer cells and that such medicines could be used in new treatments for cancer cells. Components extracted from traditional Chinese medicines were reported to dampen malignant growth by arresting the cell cycle and inducing apoptosis in cancer cells (Karimian et al. 2014).

Apoptosis is a process of programmed cell death that occurs in multicellular organisms. Apoptosis is generally characterized by distinguishable morphological features such as cytoplasmic shrinkage, chromatin condensation, phosphatidylserine (PS) externalization, DNA fragmentation, and the formation of apoptotic bodies without affecting the plasma membrane integrity (Elmore 2007).

In the present study we focused on Ranunculus ternatus, which has been used as a traditional Chinese medicine (Pan and Sun 1986). R. ternatus is a perennial of the genus Ranunculaceae family and is distributed in wet regions in China and Japan (Zhang et al. 2007). The roots of R. ternatus are used as a medical herbal food, and the artificial cultivation of R. ternatus is widely performed. To date, traditional Chinese medicine containing R. ternatus has been reported to be effective against malignant lymphoma, leukemia, pulmonary tuberculosis, breast tumor, goiter, lung, gastric, esophageal tumor, and more (Zhang and Wan 1993; Chen et al. 2002; Tong et al. 2013). Both the organic form and the aqueous extract from R. ternatus have anticancer activity, and neither has any significant toxic effects (Wang et al. 2006). Phytochemical studies have confirmed that R. ternatus contains flavonoids, glycosides, benzine, organic acid, sterols, ester, amino acids, and constant and trace elements (Miao et al. 2014). Among the organic extracts, R. ternatus ethyl acetate extract (RTE) has been reported to contain sternbin, methylparaben, 3-[(4-O-d-glucopyranosyl)-phenyl]-2-propennoic acid, linocaffein, β-d-glucose, robustaflavone-4′-methylether, kayaflavone, podocarpusflavone A, bilobetin, isoginkgetin, amentoflavone, ternatoside A, ternatoside B and 4-O-d-glucopyranosyl-p-coumaric acid (Zhang and Tian 2006; Xiong et al. 2008; Tian et al. 2006). Amentoflavone and podocarpusflavone A have been reported to induce apoptosis (Pei et al. 2012; Yeh et al. 2012), but their apoptosis-inducing ability and its mechanism need further investigation. In addition, R. ternatus 70% ethanol extract can induce production of tumor necrosis factors (TNFs) by macrophages, suggesting that TNFs may be induced by RTE (Zhou et al. 1995). Herein we further examined the potential mechanism underlying RTE-induced apoptosis using the human T-cell lymphoma cell line Jurkat as a cancer model.

Materials and methods

Cell culture

Cells of the human lymphoid T-cell line Jurkat stably transfected with a human Bcl-2 expression plasmid (Bcl-2) and neomycin resistance plasmid (neo) were kindly provided by Dr. T. Miyashita of the National Research Institute for Child Health and Development (Tokyo, Japan). Jurkat (neo) cells and Jurkat (Bcl-2) cells were cultured in RPMI-1640 medium (Sigma-Aldrich, St. Louis, USA) supplemented with 3.5 µL/L 2-mercaptoethanol (Wako, Osaka, Japan), 75 mg/L kanamycin sulfate (Wako), 2 g/L NaHCO3 (Wako), heat-inactivated 10% (v/v) fetal bovine serum (FBS) (Biofill, Victoria, Australia). The human breast cancer MCF-7 cells were supplied by the Cell Resource Center for Biomedical Research, Tohoku University (Sendai, Japan), and cultured in Dulbecco's modified Eagle medium supplemented with 100 mg/L streptomycin, 100 U/mL penicillin, 2 g/L NaHCO3, and heat-inactivated 10% (v/v) FBS. All cell cultures were maintained at 37 °C in a humidified chamber under an atmosphere of 95% air and 5% CO2.

Organic layer collection protocol of R. ternatus

We used a previously reported protocol to collect the organic layer of R. ternatus (Zhuang et al. 2015). Briefly, 75 mL of 95% ethanol was added to 50 g of the root powder of R. ternatus and extracted by hot reflux at 100 °C for 3 h. The ethanol solution was recovered and then was removed by a rotary evaporator (Eyela, Tokyo, Japan). The concentrate was dispersed in Milli Q, and then ethyl acetate (Wako) and sodium chloride (Wako) were added and the organic layer was recovered. The organic layer was dehydrated with magnesium sulfate (Wako) and concentrated by a centrifugal evaporator (Thermo Fisher Scientific, Waltham, USA). The RTE concentrate was dissolved in dimethyl sulfoxide (DMSO; Wako) at the concentration of 100 mg/mL.

Cell viability assay

Jurkat cells (2 × 104) or MCF-7 cells (1 × 104), seeded overnight, were incubated in 96-well plates (BD Falcon, Franklin Lakes, USA) at 37 °C with or without RTE for 24 h. One hour prior to the end of the incubation period, 10 µL of 5 mg/mL 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT; Wako) was added to each well, and the plates were incubated at 37 °C for 1 h. Finally, the supernatants were discarded, and 100 μL of DMSO was added to dissolve MTT formazan. We used a microplate reader (ChroMate, Awareness Technology, Palm City, USA) to measure the absorbance of each well at 570 nm. The absorbance of the culture wells without test agents was set as 100%.

Assessment of sub-G1 cells and the cell cycle

Jurkat cells (2 × 106) were incubated with or without 50 µM of pan-caspase inhibitor Z-Asp-CH2-DCB for 1 h, then with or without RTE for 24 h. The cells were corrected, washed with phosphate-buffered saline (PBS) and suspended in permeabilizing buffer (0.1% Triton-X 100 in PBS). Thereafter, the cells were washed with PBS twice, resuspended in PBS containing 0.5 mg/mL RNase A and 1 mg/mL propidium iodide (PI; Wako), and analyzed by flow cytometry (FACSCalibur, Becton Dickinson, Mountain View, USA). We analyzed the results using Cell Quest software (Becton Dickinson).

Annexin-V binding

Annexin-V binding was assessed by flow cytometry. Jurkat cells (2 × 106) were incubated with or without 50 µM pan-caspase inhibitor Z-Asp-CH2-DCB for 1 h, then with or without RTE. The cells were corrected, washed with PBS and suspended in annexin V binding buffer (10 mM HEPES—NaOH, 140 mM NaCl, 2.5 mM CaCl2, pH 7.4). The pellet was dissolved for 10 min at room temperature in a dark room in 0.2 mL of annexin V binding buffer containing 2.5 µL of fluorescein isothiocyanate (FITC)-annexin V (Alexis Biochemicals, San Diego, USA). The cells were then washed with PBS, resuspended in PBS containing 1 mg/mL PI, and analyzed by flow cytometry. The data were analyzed with the Cell Quest software.

Hoechst 33342 staining

Jurkat cells (2 × 105) were incubated in 24-well plates (Sigma-Aldrich) at 37 °C with or without RTE for 24 h. The cells were then washed with PBS and resuspended in PBS containing 1 mg/mL PI and 200 μg/mL Hoechst 33342 (ImmunoChemistry Technologies, Bloomington, USA) for 10 min at 37 °C. Thereafter, the cells were analyzed by an EVOS® FL Cell Imaging System (Thermo Fisher Scientific) at × 400 magnification; the fluorescence was measured at 461-nm emission for Hoechst 33342 dye and 620-nm emission for PI.

TUNEL assay

We performed a terminal deoxynucleotidyl-transferase-mediated dUTP nick end labeling (TUNEL) assay using the MEBSTAIN Apoptosis TUNEL Kit Direct (Medical & Biological Laboratories, Woburn, USA) according to the manufacturer's instructions. Jurkat cells (2 × 106) were incubated with 0.2 mg/mL RTE for 24 h or 1 µM staurosporine for 4 h as a positive control. Then, the cells were washed with PBS containing 0.2% bovine serum albumin (BSA) and fixed in 4% paraformaldehyde (0.1 M NaH2PO4, pH 7.4) for 30 min at 4 °C. We then added 0.2 mL of 70% ethanol to the cell pellet and mixed it gently. The cell pellet was incubated for 30 min at − 20 °C for permeabilization. After the pellet was washed with PBS containing 0.2% BSA, 30 μL of terminal deoxynucleotidyl-transferase solution was added, followed by incubation for 1 h at 37 °C. Thereafter, the cells were washed with PBS containing 0.2% BSA and analyzed by flow cytometry. The data were analyzed using Cell Quest.

Western blot analysis

Jurkat cells (2 × 106), or MCF-7 cells (1 × 106), seeded overnight, were incubated with or without 50 µM pan-caspase inhibitor Z-Asp-CH2-DCB for 1 h, then with or without RTE for 24 h. The cells were corrected, and total protein from the cells was extracted by lysis buffer: 50 mM HEPES pH 7.5, 150 mM NaCl, 10% glycerol, 1% Triton X-100, 1.5 mM MgCl2, 1 mM EGTA, 1 mM sodium orthovanadate, and 1% protease inhibitor cocktail (Sigma-Aldrich). The protein concentration was determined by a bicinchoninic acid protein assay, and 20 μg of protein per sample was subjected to 15% polyacrylamide gel electrophoresis and then transferred to a polyvinylidene difluoride membrane. The membranes were blocked by 3% skim milk for 1 h and then incubated with the primary antibodies at 4 °C overnight. The primary antibodies used were: caspases-3 (Santa Cruz Biotechnology, Santa Cruz, USA), -7 (Cell Signaling Technology, Danvers, USA), -8 (Cell Signaling Technology), and -9 (Santa Cruz Biotechnology), PARP-1 (Santa Cruz Biotechnology), and β-actin (Cell Signaling Technology) (all antibodies were diluted 1:1000). After being washed with 0.1% Tween 20-PBS, the membranes were incubated with an HRP-labeled anti-mouse or anti-rabbit IgG secondary antibody (Cell Signaling Technology) at 1:2000 dilution for 1 h at room temperature. Positive bands were visualized with enhanced chemiluminescence reagents (Pierce ECL Western Blotting Substrate, Thermo Fisher Scientific) following exposure at 1 min, and then observed with an ImageQuant LAS-4000 camera system (GE Healthcare, Chicago, USA).

Determination of the mitochondrial membrane potential (ΔΨm)

Jurkat cells (2 × 105) were treated for 30 min with or without RTE at 0.2 mg/mL, or 10 μM mitochondria uncoupler, carbonyl cyanide m-chlorophenyl hydrazine (CCCP) for 30 min as a positive control. At 15 min before the end of the culture period, 100 nM of the fluorescent dye 3,3′-dihexyloxacarbocyanine iodide (DiOC6) (Molecular Probes, Eugene, USA) was added. After incubation, the cells were washed with PBS and then resuspended in PBS for the examination of the ∆Ψm by flow cytometry.

Statistical analyses

All statistical analyses were performed using Student's t-test. Significance was established at the p < 0.05 level.

Results

RTE suppressed the viability of Jurkat cells

To estimate the potency of the cytotoxicity of RTE, we performed an MTT assay. The treatment with RTE significantly reduced the viability of Jurkat cells in a dose-dependent manner, as shown in Fig. 1. The half maximal inhibitory concentration (IC50) was 0.20 mg/mL. This result indicates that the RTE treatment suppressed the Jurkat cells' viability.

Fig. 1.

Fig. 1

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RTE suppressed the viability of Jurkat cells. Jurkat cells were treated for 24 h with or without RTE for indicated concentrations were subjected to an MTT assay. The results are presented as the percentage of absorbance in untreated cultures. The data are the mean ± SD of the results of three independent experiments. *p < 0.05, ***p < 0.001 vs. untreated cells

RTE induced sub-G1 accumulation but did not arrest the cell cycle

Following our observation that RTE suppressed the cell viability of Jurkat cells, we investigated whether cell-cycle arrest occurred and whether RTE induced cytotoxicity. We confirmed the DNA content by performing a flow cytometry analysis staining with PI. Cells with a DNA content lower than that observed in the G1 phase are often referred to as “sub-G1 phase” cells. As shown in Fig. 2a, the 24-h treatment with RTE increased the proportion of sub-G1 cells from 2.76 to 50.0%.

Fig. 2.

Fig. 2

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RTE induced an accumulation of sub-G1 cells but did not arrest the cell cycle. a Jurkat cells were treated for 24 h with or without RTE. DNA contents were determined by flow cytometry as described in the “Material and methods”. Representative histograms of one of three independent measurements are shown. b The percentage of cells in each cell-cycle phase is shown in the chart graph. The data are the mean ± SD of the results of three independent experiments. Student’s t-test was used to determine differences between sub-G1 phase groups. **p < 0.01, ***p < 0.001 vs. untreated cells

We analyzed the percentages of cells in the G1, S, and G2 phases. The 24-h treatment with RTE decreased the percentage of G1-phase cells from 47.1 to 28.4%, the percentage of S-phase cells from 15.8 to 6.39%, and the percentage of G2-phase cells from 29.2 to 12.1% (Fig. 2b). These results indicate that RTE may induce DNA fragmentation in the cells but does not arrest their cell cycle.

RTE induced caspase-3 independent apoptotic cell death

We next investigated whether RTE induced cytotoxicity as a result of apoptosis. PS is exposed from the inner layer to the outer layer of the cell membrane during apoptosis, where it can be detected with annexin-V. Necrotic cell lost their membrane integrity and can be stained with cell-impermeable PI. Therefore, after inducing cytotoxicity by applying 0–0.2 mg/mL of RTE for 24 h, we stained the cells with both annexin-V and PI and examined them by flow cytometry. The rates of early apoptosis (i.e., cells externalizing PS but still maintaining their membrane integrity) and the rates of late-apoptosis/necrotic cells (i.e., cells that lost their membrane integrity and showed PI staining) increased dose-dependently (Fig. 3a).

Fig. 3.

Fig. 3

Fig. 3

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RTE did not induce typical apoptosis. a Jurkat cells were treated for 24 h with or without RTE at indicated concentrations. The cells were subjected to annexin-V and PI staining and analyzed by flow cytometry. A representative quadrant plot of one of three independent measurements is shown. The open bars represent the percentages of early-apoptotic cells (annexin-V+ /PI−) and the closed bars represents the percentages of late-apoptotic cells (annexin-V+ /PI+) detected. The data are the mean ± SD of the results of three independent experiments. *p < 0.05 vs. untreated cells. b Jurkat cells were treated with 0.2 mg/mL of RTE or without RTE for the indicated times. The open bars represent the percentages of early-apoptotic cells and the closed bars represent the percentages of late-apoptotic cells. The data are the mean ± SD of the results of three independent experiments. *p < 0.05, **p < 0.01 vs. untreated cells. c Jurkat cells were treated with or without RTE and stained with PI and Hoechst 33342, and the nuclear state was determined by fluorescent microscopy. The line and arrow indicate membrane-collapsed PI-stained swelling cells. Representative figures of one of three independent experiments are shown. d Jurkat cells were incubated with 0.2 mg/mL RTE for 24 h and 1 μM staurosporine for 4 h. DNA fragmentation was determined by flow cytometry with TUNEL assay as described in the “Materials and methods” section. Representative histograms of one of three independent measurements are shown. E: Jurkat cells were treated for 24 h with or without RTE at indicated concentrations. Western blots were performed with antibodies for PARP-1, caspase-7, caspase-3, and β-actin

Moreover, because RTE-induced significant upregulation of the rates of late-apoptosis/necrotic cells, we examined the effect of 0.2 mg/mL of RTE with shorter incubation time. Following treatment with 0.2 mg/mL of RTE for 0–8 h, the early-apoptosis cell rates, as indicated by the externalization of PS, were increased in somewhat a time-dependent manner (Fig. 3b).

The morphological characteristics of apoptosis include cell shrinkage, chromatin condensation, chromatin fragmentation, and fragmentation in multiple segregated bodies, plus the formation of apoptotic bodies (Elmore 2007). Fluorescent stains are often used to determine the state of the cell nucleus. We examined the state of the cell nuclei by conducting a fluorescent microscopy analysis staining with Hoechst 33342 (a cell-membrane permeable blue dye) and PI (a cell membrane non-permeable red dye) as shown in Fig. 3c. Following the applications of RTE for 24 h, fluorescent microscopy revealed that the crescents around the periphery of the cell nuclei or the entire chromatin were stained blue and had a bubbled appearance, suggesting that chromatin condensation occurred. A slight number of nuclei were stained red with cellular swelling occurred following RTE treatment at 0.2 mg/mL, suggesting that membrane integrity was lost and thus necrotic cell death had also occurred.

We investigated whether RTE induced DNA fragmentation by performing a TUNEL assay. The TUNEL method has enabled in situ visualizations of DNA fragmentation in nucleosomal units (Elmore 2007). Figure 3d shows that with the applications of RTE, a slight increase in dUTP-positive cells occurred. We also measured executioner caspase activity and nuclear enzyme PARP-1 cleavage in a western blotting analysis (Chaitanya et al. 2010). Figure 3e shows that, following the applications of RTE for 24 h, caspase-7 activation and PARP-1 cleavage were observed in Jurkat cells, but caspase-3 activation was not observed. These results demonstrated that RTE induced the externalization of PS, caspase-7 activation, and PARP-1 cleavage, but did not induce internucleosomal DNA fragmentation or caspase-3 activation. Our observations also indicate that RTE does not induce typical apoptosis involving executioner caspase-3 and frequent DNA fragmentation.

RTE-induced apoptotic cell death depends on caspase-7

RTE induced the activation of an execution caspase. There are two main pathways of caspase activation: extrinsic and intrinsic. Each pathway activates its own initiator caspases, i.e., caspase-8 and -9, and activated caspases-8 and -9 cleave downstream caspases such as caspase-3 and -7 (Elmore 2007). We further determined whether the RTE-induced cell death depends on one or more caspases using the pan-caspase inhibitor Z-Asp-CH2-DCB in a western blotting analysis and flow cytometry. Z-Asp-CH2-DCB is a cell-permeable broad-spectrum caspase inhibitor. The RTE-induced activations of caspase-7, -8 and -9 were suppressed by Z-Asp-CH2-DCB, and the cleavage of PARP-1 was significantly suppressed by this inhibitor (Fig. 4a).

Fig. 4.

Fig. 4

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RTE-induced cell death is dependent on caspase-7. a Jurkat cells were incubated with or without 50 μM Z-Asp-CH2-DCB for 1 h, and thereafter with or without 0.2 mg/mL RTE for 24 h. Western blots were performed with antibodies for PARP-1, caspases -8, -9, -7, and β-actin. b Jurkat cells were incubated with or without 50 μM Z-Asp-CH2-DCB for 1 h, and thereafter with or without 0.2 mg/mL RTE for 24 h. DNA contents were determined by flow cytometry. The chart graph shows the percentages of cells in the sub-G1 phase. The data are the mean ± SD of the results of three independent experiments. c Jurkat cells were incubated with or without 50 μM Z-Asp-CH2-DCB for 1 h, and thereafter with or without 0.2 mg/mL RTE for 24 h. Annexin-V and PI staining were performed. The open bars represent the percentages of early-apoptotic cells (annexin-V+ /PI−) and the closed bars represent the percentages of late-apoptotic cells (annexin-V+ /PI+) detected. The data are the mean ± SD of the results of three independent experiments. d MCF-7 cells were treated for 24 h with or without RTE at the indicated concentrations. The cells were subjected to an MTT assay. The results are presented as the percentage of absorbance in untreated cultures. The data are the mean ± SD of the results of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001 vs. untreated cells. E: MCF-7 cells were treated for 24 h with or without RTE at the indicated concentrations. Western blots were performed with antibodies for PARP-1, caspases -8, -9, and -7, and β-actin

Procaspase-7 is composed of a pro-domain, a large subunit (p20), a small subunit (p10), and a linker connecting the subunits. By initiator caspases, the pro-domain is cleaved to generate p20p10 caspase-7. Cleavage of the linker generates large subunits (p20) and small subunits (p10) to form activated heterotetramers (Lamkanfi and Kanneganti 2010). Recent studies revealed that p20p10 caspase-7, p41 caspase-8, and p35 caspase-9 also have enzymatic activity (Sawai 2013). A pan-caspase inhibitor can inhibit p20p10 caspase-7 activation but not p20 caspase-7 activation (Majerciak et al. 2010). In the present study, the RTE induced sub-G1 accumulation was suppressed with Z-Asp-CH2-DCB. DNA fragmentation decreased from 50.0% to 14.1% (Fig. 4b), and the RTE-induced externalization of PS was suppressed (Fig. 4c). The early-apoptosis cell rate decreased from 20.8 to 10.6%, and the late-apoptosis cell rates were unchanged.

Caspase-3 and caspase-7 coordinate the execution phase of apoptosis. Although caspase-3 and caspase-7 share 57% sequence identity throughout their catalytic domains, they are functionally distinct (Boucher et al. 2012). We therefore further examined whether RTE-induced cell death depends solely on caspase-7 using caspase-3-deficient human breast cancer MCF-7 cells. Treatment with RTE significantly reduced the viability of MCF-7 cells in a dose-dependent manner for 24 h (Fig. 4d). The IC50 was 0.32 mg/mL. This result suggests that RTE suppressed the MCF-7 cells' viability. Figure 4e shows that with the applications of RTE for 24 h, caspases -7, -8 and -9 were activated and PARP-1 was cleaved in the MCF-7 cells, suggesting that the RTE-induced cell death is dependent on caspase-7.

Mitochondria are not involved in RTE-induced apoptotic cell death

In the intrinsic or mitochondrial pathway of apoptosis, the ∆Ψm is reduced, and then cytochrome c is released and binds to apoptotic protease-activating factor 1, causing caspase-9 activation. Activated caspase-9 cleaves downstream caspases such as caspase-3 and -7, initiating the caspase cascade (Elmore 2007). The anti-apoptotic Bcl-2 protein resides on the outer mitochondrial membrane and prevent apoptosis (Elmore 2007); we thus investigated the perturbation of mitochondria during apoptosis by evaluating the ∆Ψm and mitochondria downstream caspases by the overexpression of mitochondria-localized Bcl-2 in Jurkat cells. To determine whether the RTE cytotoxicity was changed, we performed MTT assays to determine the cell viability of Jurkat (Bcl-2) cells. The results showed that the 24-h treatment with RTE significantly reduced the viability of Jurkat (Bcl-2) cells in a dose-dependent manner (Fig. 5a). The IC50 was 0.18 mg/mL. The cytotoxicity of RTE was unchanged by Jurkat (Bcl-2) cells.

Fig. 5.

Fig. 5

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Mitochondria were not involved in RTE-induced cell death. a Jurkat cells and Jurkat (Bcl-2) cells were treated for 24 h with or without RTE at indicated concentrations. The cells were subjected to an MTT assay. The results are presented as the percentage of absorbance in untreated cultures. The data are the mean ± SD of the results of three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001 vs. untreated cells. b Jurkat cells and Jurkat (Bcl-2) cells were treated for 30 min with or without RTE at 0.2 mg/mL, or 10 μM CCCP for 30 min. ∆Ψm was analyzed by flow cytometry using DiOC6. The bar graph indicates the quantified percentage of relative DiOC6 fluorescence. The data are the mean ± SD of the results of three independent experiments. *p < 0.05, **p < 0.01 vs. untreated cells. c Jurkat cells and Jurkat (Bcl-2) cells were incubated with or without 0.2 mg/mL RTE for 24 h. Western blots were performed with antibodies for caspase-7 and β-actin

We also examined the perturbation of mitochondria during apoptosis by determining the ∆Ψm. Cells were examined in a flow cytometry analysis using staining with DiOC6. As shown in Fig. 5b, 0.2 mg/mL RTE began reducing the ∆Ψm at 30 min after the treatment of Jurkat cells and Jurkat (Bcl-2) cells. We investigated the effects of mitochondria during apoptosis by examining the activity of caspase-7 downstream of the mitochondria. Figure 5c shows that with the application of 0.2 mg/mL RTE for 24 h, caspase-7 activation in Jurkat cells and Jurkat (Bcl-2) cells occurred. These results suggest that mitochondria are not involved in RTE-induced cell death.

Discussion

Our in vitro experiments revealed that RTE-induced cell death is followed by caspase activation and cleaving of PARP-1. A pan caspase inhibitor inhibited the RTE-induced cell death, suggesting that caspase activation plays an important role in RTE-induced cell death. RTE also induced cell death in MCF-7 cells, indicating that RTE-induced cell death depends on caspase-7, not caspase-3. Although MCF-7 cells are caspase-3 null adherent cell and tend to be less susceptible to death signals, their IC50 value was not so different from that of Jurkat cells in suspension, indicating that RTE-induced cell death does not involve caspase-3. Moreover, the overexpression of the mitochondria‐localized anti‐apoptotic protein Bcl‐2 did not inhibit RTE-induced cell death, suggesting that mitochondria are not involved in RTE-induced cell death.

DNA fragmentation is the hallmark of apoptosis. As shown in Figs. 2 and 3c, RTE induced chromatin condensation, increase of sub-G1 cells, and formation of micronuclei. We did not detect the formation of a large number of apoptotic bodies, however; these accounted for only a small part of the total formation of cells. In addition, we also found that necrotic cell death had occurred. We further determined the nucleosome units of DNA fragmentation by performing TUNEL assay and demonstrated that RTE treatment did not result in a significant increase of internucleosomal DNA fragmentation. These results indicate that RTE induces cell shrinkage and condensation of nuclei, but not the fragmentation of small DNA, further suggesting that RTE does not induce a typical form of apoptotic cell death and also has the ability to induce necrotic cell death. Mattes (2007) reported that necrotic cells appear to be in “sub-G1 phases” in the manner of apoptotic cells. Leicht et al. (2003) reported that they observed simultaneous apoptotic and necrotic cell death and detected caspase-7 activation; caspase-3 was not activated and DNA ladder-like fragmentation did not occur in their experiments. Those authors suggested that necrosis may have affected the apoptosis process so that caspase-3 activation and DNA ladder-like fragmentation did not occur. Their results resemble our present findings. We, therefore, speculate that RTE does not entirely follow typical form of apoptosis and that it may also induce necrosis-like cell death.

The aspartate-specific cysteine protease family of caspases is responsible for apoptosis (Elmore 2007). The cleavage of PARP-1 by caspases is a hallmark of apoptosis (Chaitanya et al. 2010). In the present study, RTE induced activations of caspase-8, caspase-9, and thereby cleavage of PARP-1 in Jurkat cells (Fig. 4a). These results suggested that both extrinsic and intrinsic pathways may be involved, and that caspase activation plays an important role in RTE-induced cell death. We examined the suppression of the intrinsic pathways by overexpression of Bcl-2 for its ability to block downstream mitochondria apoptotic events. However, cell death event was not suppressed, indicating that RTE-induced cell death was independent of mitochondria. Jiang et al. (2001) revealed that caspase-8 can activate caspase-9, suggesting that RTE-induced caspase-9 activation was induced by caspase-8. Usually, caspase-8 and caspase-9 may activate caspase-3 and caspase-7, but in our experiments caspase-3 was not activated (Fig. 3e), suggesting that the one of the components in RTE may function as a caspase-3 inhibitor, or may preferentially activate caspase-7. Boucher et al. (2012) suggested that the N-terminal domain of caspase-7 contains a transferable exosite to promote PARP-1 cleavage, and this allows caspase-7 to cleave PARP-1 more efficiently than caspase-3. Martini et al. (2017) further confirmed the importance and necessity of caspase-7’s N-terminal domain function. We thus hypothesized that RTE can induce cell death without interference from caspase-3. The results of our experiments demonstrated that RTE induced cell death through caspase-7 activation in MCF-7 cells (Fig. 4d, e), suggesting that RTE-induced cell death was dependent on caspase-7 activation. Collectively, these findings suggest that RTE may induce the N-terminal domain of capase-7 (which contains a transferable exosite) to promote PARP-1 cleavage, resulting in cell death. The relationship between RTE and caspase-7 thus merits further investigation.

There is another report about the unique phenomenon of caspase-7 activation. The traditional Chinese medicine extract Andrographolide was also found to be responsible for the induction of apoptosis via caspase-7 activation, but not caspase-3 activation (Yang et al. 2014). It is thus necessary to determine whether RTE can produce the same effects when applied to cancer cells other than those examined herein to lay the foundation for future widespread applications of RTE. Wang et al. (2006) indicated that RTE could reduce hepatoma volume in mice.

In conclusion, the present study demonstrated that RTE has an anticancer effect of inducing cell death that is dependent on caspase-7 through an extrinsic pathway. In the future, we will need to identify the ingredients in RTE that are responsible for this exclusive activation of caspase-7 and to determine the underlying mechanism.

Author contributions

FM, TS and YN conceived and designed the experiments. FM performed the experiments. FM and YN analyzed the data. FM, TS and YN wrote the manuscript.

Compliance with ethical standards

Conflict of interest

The authors declare no conflict of interest.

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