Ginsenoside Rh2 represses autophagy to promote cervical cancer cell apoptosis during starvation

HEK293T (ATCC, CRL-11268), HeLa (ATCC, CCL-2), and C-33 A (ATCC, HTB-31) cells were cultured in DMEM supplemented with 10% fetal bovine serum (FBS) at 37 °C in a 5% CO₂ environment. Lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA) was used for transient plasmid and siRNA transfections. The sequences of the siRNA targeting Atg7 were 5′-GACAUUAAGGGUUAUUACUTT-3′and 5′-AGUAAUAACCCUUAAUGUCTT-3′. The control siRNA sequences were 5′-UUCUCCGAACGUGUCACGUTT-3′ and 5′-ACGUGACACGUUCGGAGAATT-3′.

Cells were seeded into 96-well culture plates. The following day, culture medium was replaced with 100 µl of serum-free medium or normal medium with dimethylsulfoxide (DMSO) or different compounds and incubated for 24 h. Before detection, CCK8 substrate was added into the plates and incubated for 1 h at 37 °C. Absorbance was measured at 450 nm using a microplate reader (Tecan Infinite 200PRO, Tecan, Mannedorf, Switzerland).

The primary antibodies used in this study were as follows: anti-Tubulin mAb (BioLegend, San Diego, CA, USA), anti-p62 mAb, anti-LC3A/B mAb (Cell Signaling Technology, Danvers, MA, USA), anti-Cytochrome C mAb, anti-COX IV pAb, anti-Lamin B1 mAb, anti-AIF mAb, anti-CTSB pAb (Beyotime, shanghai, China). HRP-conjugated goat anti-mouse IgG and goat anti-rabbit IgG (Jackson Immunoresearch, West Grove, PA, USA) and Alexa Fluor 488-conjugated goat anti-rabbit IgG antibody (Molecular Probes, Waltham, MA, USA) were used for detection.

Ginsenoside 20 (S)-Rh2 was purchased from Chengdu Must Biotech (Chengdu, China). The Annexin V-FITC/PI apoptosis detection kit was purchased from BD Biosciences (Franklin Lakes, NJ, USA). Rhodamine 123, Ac-DEVD-CHO, DAPI, the nuclear and cytoplasmic protein extraction kit and cell mitochondria isolation kit were purchased from Beyotime. CCK-8 was purchased from BOSTER Biotech (Wuhan, China). Rapamycin and Bafilomycin A1 (BA1) were purchased from InvivoGen (San Diego, CA, USA).

The LC3B gene was obtained by PCR from cDNA of HeLa cells and subsequently subcloned using standard molecular biology procedures into the pEGFP-C3. The primers: forward 5′-GGAATTCTATGCCGTCGGAGAAGAC-3′ and reverse 5′-CGGGATCCTTACACTGACAATTTCAT-3′. Annexin A2 fragments were amplified from HeLa cDNA by PCR and subcloned into the VR1012 vector with an HA tag added in frame at its N-terminus. The primers: forward 5′-ATGTCTACTGTTCACGAAATCC-3′ and reverse 5′-GTCATCTCCACCACACAGG-3′. HA tag primers: forward 5′-CGTCTAGAGCCACCATGTCTACTGTTCACGAA-3′ and reverse 5′-CGGGATCCTCAAGCATAGTCTGGGACGTCGTATGGGTAGTCATCTCCACCACA-3′. GFP-mRFP-LC3, CD317 IHA, and VR1012 and pEGFP-N3 were described previously [22, 23].

Cells were double-stained with propidium iodide (PI) and Annexin V-FITC using an apoptosis detection kit for flow cytometry. The samples were analyzed using a FlowSight^® Imaging Flow Cytometer (Merck Millipore, Burlington, MA, USA) and the software IDEAS Application V6.1.

Cells were stained with 1 µM rhodamine 123 for 30 min at 37 °C. After washing with ice-cold phosphate-buffered saline (PBS), the cells were acquired and analyzed by flow cytometry.

Cells were then fixed with 4% formaldehyde, permeabilized with 0.5% Triton X-100, and then blocked with 10% FBS. Immunostaining was performed by incubating cells with primary antibody and Alexa Fluor 488-conjugated IgG antibody. After DAPI staining, cells were observed using a fluorescence microscope with a 20× objective lens.

Cells were harvested and resuspend with lysis buffer, boiled for 15 min, separated by SDS- PAGE, and then transferred onto nitrocellulose membranes (Whatman, Maidstone, UK). After blocking in 5% nonfat milk, membranes were probed with primary and secondary antibodies. Immunoreactivity was visualized by chemiluminescence. ImageJ software was used for densitometric analysis.

Cells were treated as indicated and fixed in 4% glutaraldehyde at 4 °C overnight. After dehydration, ultrathin sections were embedded and stained with uranyl actate/lead citrate. Images were captured under a transmission electron microscope (H-7650, Hitachi, Japan).

After rinsing in PBS and fixing with 4% paraformaldehyde for 10 min, the cells were stained for acidic vesicular organelles in the dark for 30 min. Then, the cells were observed by fluorescence microscopy at 488 nm and imaged.

All data represent at least 3 independent experiments and are expressed as mean ± SEM. Statistical comparisons were made by one-way analysis of variance and Dunnett’s post hoc test. *p < 0.05; **p < 0.01; ***p < 0.001; ns > 0.05.

G-Rh2 has been shown to have antitumor effect on a variety of cancer cells, including cervical cancer cells. Serum deprivation is often used as a means of emulating the tumor microenvironment. Here, we investigated the cytostatic activity of G-Rh2 during serum deprivation and normal conditions. As shown in Fig. 1, in normal conditions, G-Rh2 (0–10 µM) had no effect on cell proliferation. 5, 7.5 and 10 µM G-Rh2 significant inhibited cancer cells proliferation under serum-free conditions (Fig. 1a and b, Additional file 1: Figure S1B and C). Meanwhile, the cell viability of normal cells (293T and HUVEC) treated with 5 µM G-Rh2 were no obvious difference compared with control group (Fig. 1c and d). These data suggested that low concentration G-Rh2 exerted strong cytostatic activity under serum-free conditions.

To determine whether the decrease in cell viability was caused by apoptosis, we used Hoechst 33342 staining and Annexin V/PI analysis. Figure 2a shows chromatin condensation and nuclear fragmentation in HeLa and C-33A cells treated with G-Rh2. As showed in Fig. 2b (Additional file 2: Table S1), G-Rh2 did not affect apoptosis in cancer cells cultured in normal medium, but markedly enhanced apoptosis during 24 h serum deprivation. Moreover, in serum-free conditions, 7.5 µM G-Rh2 caused a decrease in the percentage of S and G0/G1 phase cells (Additional file 1: Fig. S1D, Additional file 3).

To investigate the apoptosis pathway, we employed Ac-DEVD-CHO, a specificity caspase 3 inhibitor. The inhibitor reduced serum deprivation-induced apoptosis but no significant effect on the apoptosis induced by G-Rh2 (Fig. 3a, Additional file 3: Table S1). Mitochondria are the primary organelles that induce apoptosis [24]. We compared the levels of mitochondrial membrane potential (MMP) between DMSO- and G-Rh2-treated using by Rhodamine 123. Flow cytometric analysis showed that G-Rh2 had little effect on mitochondrial function in normal conditions but prominently reduced MMP in serum-deprived cells (Fig. 3b). Next, we detected the distribution of apoptosis-inducing proteins (AIF) and cytochrome c. As shown in Fig. 3c and d, the level of nuclear AIF was significantly higher in G-Rh2-treated cells from serum-deprived conditions than in the DMSO controls. This nuclear accumulation was accompanied by a decline in mitochondrial distribution, suggesting that G-Rh2 enhanced AIF release and nuclear translocation in serum-free cells. Meanwhile, the distribution of cytochrome c in the mitochondria and cytoplasm had no visible change in any conditions. Together, these data suggested that G-Rh2 promotes serum deprivation-induced apoptosis through the AIF pathway.

According to recent report, an autophagy modulator scoring system (AMSS) was designed to evaluate methodological integrity [25, 26]. To verify whether G-Rh2 participates in autophagy, we examined the effect of G-Rh2 on autophagy by measuring GFP-LC3B puncta formation. The results revealed that G-Rh2 obviously increased the number of LC3B puncta in normal and starving HeLa cells and C-33A cells (Fig. 4a). Transmission electron microscopy was used to observe the accumulation of autophagic vacuoles in G-Rh2-treated cells, which revealed an increased number of autophagic vacuoles compared with control cells (Fig. 4b).

An increase in LC3 II in response to a drug may represent either increased generation of autophagosomes or a block in autophagosomal maturation and degradation. Meanwhile, a reduction of substrate protein p62 can be regarded as a marker for increased autophagic flux. Western blot results showed that LC3 II and p62 protein levels were increased following G-Rh2 treatment (Fig. 5a, lanes 2 and 6). Rapamycin strongly induced autophagy leading to significant degradation of LC3 and p62 (Fig. 5a, lanes 3 and 7). The autophagy inhibitor BA1 prevented maturation of autophagic vacuoles and increased LC3 and p62 levels (Fig. 5a, lanes 4 and 8). The effect of G-Rh2 was similar to autophagy inhibitor BA1. In addition, we used HeLa and C-33A cells transfected with the tandem fluorescent-tagged LC3B (mRFP-GFP-LC3B) plasmid. RFP and GFP exhibit differing pH sensitivity, wherein GFP is quantitatively quenched in acidic compartments while RFP is stable. Therefore this probe capitalizes on the pH difference between the acidic autolysosome and the neutral autophagosome to monitor flux of LC3B from autophagosomes (RFP-positive GFP-positive; yellow dots) to autolysosomes (RFP-positive GFP-negative; red dots). Nutrient starvation treatment only resulted in the production of large amounts of red puncta. In the cells treated with G-Rh2, we observed the enhanced formation of yellow puncta. The results suggested that upon G-Rh2 treatment the autophagosomes do not fuse with lysosomes or that lysosomal function is impaired (Fig. 5b).

Because the autophagic flux depends on low pH, we analyzed the lysosomal pH by an acridine orange (AO) assay. AO is a fluorescent nucleic acid dye that accumulates in acidic spaces such as lysosomes. Under the low pH conditions in lysosomes, the dye emits red light when excited by blue light. Compared to the control group with serum, the DMSO group without serum showed an increased red signal, indicating that the pH value was decreased and autophagy was promoted, whereas the addition of G-Rh2 reduced the red signal, indicating inhibition of autophagy (Fig. 5c). The lysosome enzyme like Cathepsin B (CTSB) was used to examine the lysosome function. Western blotting assay demonstrated that G-Rh2 remarkedly downregulated the total and mature from protein level of CSTB in HeLa and C-33A cells (Fig. 5d). These data suggested that in Rh2-treated cells, the increased LC3 II levels and formation of GFP-LC3 puncta were not due to enhanced autophagy, but due to reducing lysosomal activity, inhibiting the fusion of autophagosome and lysosome, leading to a block of autophagic flux.

To determine whether autophagy regulation was responsible for G-Rh2-induced apoptosis, we designed a siRNA to knockdown the expression of Atg7. As shown in Fig. 6a, the expression of Atg7 was downregulated in siAtg7 group and the sicontrol has no effect on the expression of Atg7. Meanwhile, the expression level of LC3 II was significantly reduced in the siAtg7 group. These results suggested the autophagy was inhibited in the cells transfected siAtg7. Then, for examine whether autophagy inhibition in serum free condition will trigger AIF-induced apoptosis, we detected the apoptosis rate and expression level of AIF in nuclear in the cells transfected siAtg7. As shown in Fig. 6b (Additional file 3: Table S1), knockdown Atg7 remarkably promoted apoptosis in serum free condition. According to the western blotting results, the level of nuclear AIF was higher in cells transfected siAtg from serum-deprived conditions than in the cells transfected sicontrol (Fig. 6c). In addition, autophagy inhibitors and G-Rh2 had additive effects on HeLa cell apoptosis during serum deprivation. Treatment with the autophagy agonist rapamycin significantly rescued G-Rh2-induced apoptosis in serum-free conditions (Additional file 4: Fig. S2). These data suggested that autophagy plays an important role in G-Rh2-induce apoptosis under serum-deprived conditions.

Annexin A2, is a member of the Annexin family that plays multiple cellular biological roles [27, 28]. Recently report claimed that 20(S) G-Rh2 directly bound Annexin A2 and promoting apoptosis in tumor cells [29]. To address whether Annexin A2 suppressed apoptosis following G-Rh2 treatment and serum deprivation, we forced Annexin A2 expression in HEK293T and detected its influence on apoptosis. CD317 is thought to protect cancer cells against serum deprivation-induced apoptosis, and was used as a positive control in this experiment [30]. The results showed that Annexin A2 overexpression did not affect apoptosis rates in serum-starved DMSO treated cells, but significantly suppressed apoptosis in G-Rh2-treated cells that were serum-deprived (Fig. 7 and Additional file 4: Figure S2B). This may be due to an excess of Annexin A2 binding to G-Rh2 and hindering G-Rh2 activity, or that expression of the protein improves starvation- induced autophagy and reduces apoptosis.