Saikosaponin d induces cell death through caspase-3-dependent, caspase-3-independent and mitochondrial pathways in mammalian hepatic stellate cells

A human HSC cell line, LX-2 was purchased from MERCK MILLIPORE (SCC064). A rat HSC cell line, HSC-T6, immortalized with the large T antigen of the SV40. Both cell lines were cultured at 37 °C under a 5 % CO₂ atmosphere and in Dulbecco’s modified Eagle’s medium (DMEM; Gibco®, Life Technologies) supplemented with 100U/mL penicillin, 100 μg/mL streptomycin and 5 % heat-inactivated fetal bovine serum (FBS; Gibco®, Life Technologies). The HSC-T6 and LX-2 cells were seeded in 96-well plates at a density of 5 × 10³ cells/well in DMEM supplemented with 5 % FBS. Following an 18-h incubation at 37 °C under a 5 % CO₂ atmosphere, the old medium was replaced with fresh DMEM/1%FBS containing SSd (1 μM). After 0, 18, 24, 48 and 72 h of incubation, the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay (Invitrogen™, Life Technologies) was performed for cell proliferation detection as describe previous report [18]. The generated formazan products were solubilized with 100 μL of dimethyl sulfoxide (DMSO; Sigma-Aldrich), and the optical density was determined at 570 nm using an enzyme-linked immunosorbent (ELISA) reader (infinite M200PRO, TECAN).

Colony formation assay was performed according to Park et al.’s work [19, 20]. Both HSC-T6 and LX-2 cells were seeded (2 × 10³ cells/well) in a 6-well plate and incubated at 37 °C under a 5 % CO₂ atmosphere for 14 days. The colonies were fixed with 70 % ethanol at 4 °C and stained with 5 % Gentian Violet (Sigma) at room temperature.

Specific apoptosis was evaluated in both HSC-T6 and LX-2 cells by treating with SSd (1 μM) for 24 h. All living cells and cell debris were collected and fixed in 70 % ethanol/phosphate-buffered saline (PBS) at 4 °C, pelleted and resuspended in a buffer solution containing 200 μg/mL RNase A and 0.01 mg/mL propidium iodide (PI; Sigma-Aldrich). The cell cycle states of the HSC-T6 and LX-2 cells were determined by flow cytometry (Cytomic FC 500, BECKMAN COULTER).

Both HSC-T6 and LX-2 cells were seeded in a 6-well culture plate at a density of 2 × 10⁵ cells/well to determine the protein expression levels of collagen type I, collagen type III, caspase-3 and caspase-9. After 16-h incubation, the exhausted culture medium was discarded and replaced with fresh serum-free DMEM (Gibco®, Life Technologies) containing SSd at a working concentration of 1 μM). The culture medium and living cells were collected after subsequent 24-h incubation. The cell pellet was lysed by RIPA solution, and the total protein content was extracted. The protein expressions of collagen type I and III, caspase-3, and caspase-9 were measured by ELISA kits (Uscn Life Science Inc.). The activity of caspase-3/7 and caspase-9 was measured by the ApoTox-Glo™ Triplex assay kit (for caspase-3/7 activity detection; Promega) and Caspase-Glo® 9 assay kit (for caspase-9 activity detection; Promega) according to the manufacturer’s (protocol TRY instructions). The ApoTox-Glo™ Triplex assay kit also provides information on apoptosis. HSC-T6 and LX-2 cells were treated by SSd (0 μM, 0.1 μM, 0.25 μM, 0.5 μM, 1 μM, 1.5 μM, and 2 μM) for 24 h, and the results from 3 reactions at each SSd concentration were averaged. The fluorescence was measured by an ELISA reader (Fluoroskan Ascent FL, THERMO SCIENTIFIC), and the luminescence was measured by a luminometer (Centro LB 960, CBERTHOLD TECHNOLOGIES).

Total RNA was isolated by TriPure Isolation Reagent (Roche) based on the manufacturer’s protocol. Reverse transcriptional PCR was performed using the iScripe™ cDNA Synthesis kit (BIO-RAD). Quantitative polymerase chain reaction (qPCR) analysis and data collection were performed by an ABI 7500 FAST (Applied Biosystem). The following sequences of specific primers of target genes were adopted in qPCR: BAD-forward, 5′-CAGGCAGCCAATAACAGTCATC-3′; BAD-reverse, 5′-CCATCCCTTCATCTTCCTCAGT-3'; BAK-forward, 5′-AATGCCTACGAACTCTTCACCAA-3'; BAK-reverse, 5′-CAGTCAAACCACGCTGGTAGAC-3′; BAX-forward, 5′-TCATCCAGGATCGAGCAGAGA-3′; BAX-reverse, 5′-CCAATTCGCCGGAGACACT-3′; Bcl-2-forward, 5′-CATCTGCACACCTGGATCCA-3′; Bcl-2-reverse, 5′-TGAGCAGCGTCTTCAGAGACA-3′; Bcl-xL-forward, 5′-GATGGCCACCTACCTGAATGA-3′; Bcl-xL-reverse, 5′-CTCGGCTGCTGCATTGTTC-3′; PUMA-forward, 5′-ATGGCGGACGACCTCAAC-3'; PUMA-reverse, 5′-GGGAGGAGTCCCATGAAGAGA-3′.

HSC-T6 cells (1 × 10⁷ cells) were treated with SSd (1 μM) for 0, 0.25, 0.5, 1, 2, 4, and 8 h. Their mitochondrial and cytosolic fractions were then isolated by a Mitochondria/Cytosol Fractionation Kit (BioVision). Protein expression was detected by Western blotting using specific antibodies. The protein levels of COX3, GAPDH, BAX, BAK, apoptotic-protease-activating factor (Apaf)-1, cytochrome c (Cyt c), endonuclease G (EndoG) and apoptosis-inducing factor (AIF) were determined in isolated mitochondrial and cytosolic fractions (25 μg).

The cells were subsequently lysed in RIPA solution containing protease inhibitors (Roche). The total extracted protein content (25 μg) was separated by SDS-PAGE and transferred to polyvinyl difluoride (PVDF) membranes. The PVDF membranes were incubated by primary antibodies at a dilution of 1:500 or 1:1000 to detect procaspase-9, caspase-9, procaspase-3, caspase-3 (Cell Signalling), COX3, GAPDH (Santa Cruz), collagen I, collagen III, BAX, BAK, Bcl-2, Bcl-xL, Bcl-2-associated death promoter (BAD), p53 upregulated modulator of apoptosis (PUMA), β-actin, Apaf-1, Cyt c, EndoG, AIF (GeneTex) and α-SMA (abcam). The fold change in protein expression was expressed as a ratio calculated by dividing the specific protein band density by the β-actin band density.

Cellular ATP levels were detected by the Mitochondrial ToxGlo assay (Promega) according to the manufacturer’s protocol. Briefly, HSC-T6 cells were cultured at 1 × 10⁴ cells/well in a white, clear-bottom 96-well culture plate. The culture was incubated for 16 h, then the exhausted medium was discarded and replaced with a fresh medium containing 10 mM galactose instead of glucose. The cells were subsequently treated with SSd (0 μM, 0.1 μM, 0.25 μM, 0.5 μM, 0.75 μM, 1 μM, 1.5 μM, and 2 μM) for 24 h. An equal volume of the assay solution was added to each well, and the plate was subsequently incubated at room temperature for 30 min. Luminescence was measured by a luminometer (Centro LB 960, CBERTHOLD TECHNOLOGIES). The cellular oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) were measured by an XF24 bioenergetic assay (Seahorse Bioscience, Billerica, MA). Briefly, HSC-T6 cells were suspended in DMEM containing 5 % FBS and seeded on an XF24 microplate. The culture was incubated for 2 days, then the XF24 bioenergetic assay was initiated by removing the exhausted medium and replacing it with sodium-bicarbonate-free DMEM containing 5 % FBS. The extracellular flux changes in oxygen and in the pH of the medium surrounding the adherent cells were detected instantaneously by an XF24 extracellular Flux Analyzer (Seahorse Bioscience). The OCR and ECAR were detected at a steady state, then SSd (0.5 μM and 1 μM), oligomycin (1 μM), carbonyl cyanide 4-[trifluoromethoxy] phenylhydrazone (FCCP; 1 μM) and a mixture of rotenone (1 μM) and myxothiazol (1 μM) were injected sequentially into the wells to obtain the values of the maximal and non-mitochondrial respiration rate.

HSC-T6 cells were grown on coverslips in 24-well plates and incubated overnight. The cells were treated with SSd (1 μM) for 1 h, then stained with 100nM MitoTracker® Deep Red FM (Invitrogen™, Life Technologies) for 30 min at 37 °C. The cells were then fixed with 4 % cold paraformaldehyde for 20 min at 4 °C, and permeabilized with 0.1 % Triton X-100 for 1 min at room temperature. The cells were subsequently blocked with 1 % bovine serum albumin (BSA) for 30 min at room temperature and incubated with anti-Cyt c, anti-EndoG and anti-AIF antibody (GeneTex) overnight at 4 °C. The cells were then incubated with FITC-conjugated secondary antibody (Santa Cruz Biotechnology) for 60 min at room temperature, with DAPI (Molecular Probe) as a nuclear counterstain. The coverslips were mounted onto microscopy slides, and visualized under a confocal laser-scanning microscope (TCS SP5, LEICA).

All data are shown as the means of 3 independent experiments (mean ± S.D.). Statistical analysis was performed by the unpaired Student’s t-test, with p < 0.01 as significant.

To study the cytotoxic effects of SSd on HSCs of both HSC-T6 and LX-2, the MTT assay and colony formation assay were performed to examine the cell growth after exposure to SSd. SSd effectively inhibited cell proliferation of both HSC-T6 and LX-2 cells in a time-dependent manner (Fig. 1a). Additionally, SSd (0.5 μM and 1 μM) inhibited colony formation of both HSC-T6 and LX-2 cells (Fig. 1b). Numerous cell debris suddenly appeared when cells were treated with SSd (1 μM) at 72 h (Fig. 1c). Flow cytometry was analyzed to detect apoptotic signals (i.e., the sub-G1 phase of the cell cycle) after SSd treatment for 24 h. As indicated in Fig. 1d and e, the percentage of the sub-G1 phase and apoptotic bodies increased in both HSC-T6 and LX-2 cells treated with SSd. Activated HSCs are characterized by their expression of α-SMA and the synthesis of ECM constituents, particularly collagen I and III, during hepatic fibrosis [2, 21]. Our data indicate that SSd significantly reduced α-SMA expression in both HSC-T6 and LX-2 cells (Fig. 1f). Additionally, SSd markedly reduced collagen I and III expression within 72 h in both HSC-T6 and LX-2 cells by Western blotting and ELISA (Fig. 2a-d).

Caspase activity induction is involved in several ligand- and chemical-induced apoptotic processes. SSd treatment detected caspase-3/7 and caspase-9 activities in HSCs by using the ApoTox-Glo Triplex assay, Caspase-Glo 9 assay and Western blotting. The cytotoxicity, caspase-3/7 activity and caspase-9 activity of both HSC-T6 and LX-2 cells were increased, after treated with SSd for 24 h, as indicated in Fig. 3. Western blotting results also indicated that levels of caspase-3- and caspase-9-activated fragments rose after SSd treatment (Fig. 4a). To assess whether the total endogenous protein levels of caspase-3 and -9 were altered, the total forms of caspase-3 and -9 were measured by ELISA. Analytical results indicate that treatment with SSd for 24 h did not alter the total protein levels of caspase-3 and -9 compared to control groups in either HSC-T6 or LX-2 cells (Fig. 4b and c). A caspase-3 inhibitor, Z-DEVD-FMK, was adopted applied to investigate whether SSd-induced apoptosis was caspase-3-dependent. HSC-T6 and LX-2 cells were pre-incubated with Z-DEVD-FMK (100nM) for 1 h, and were subsequently co-treated with SSd for 24 h. Experimental results indicate that Z-DEVD-FMK partially inhibited the SSd-induced sub-G1 phase and cytotoxicity, as measured by flow cytometry and the MTT assay (Fig. 4d-f). These data indicate that SSd-induced apoptosis of both HSC-T6 and LX-2 cells may occur partially via caspase-3-dependent.

Mitochondrial fracture is common in apoptotic processes, and results in apoptotic factor release and caspase-9 activation. An experiment was performed to measure ATP production in HSC-T6 cells using the Mitochondrial ToxGlo Assay. Treatment with the indicated concentrations of SSd reduced cellular ATP production (Fig. 5a). To assess SSd-induced changes in the cells’ metabolic capacity and extracellular acidification, the cellular oxygen consumption and extracellular acidification were measured simultaneously by the Seahorse XF24 system. The steady-state oxygen consumption and extracellular acidification were measured, and then SSd (0.5 μM or 1 μM) was injected into the system at the fourth time point to stimulate the cells. Oligomycin was then injected to inhibit ATP synthase, and FCCP was added by injection to assess the maximal oxygen consumption. Finally, a mixture of rotenone and myxothiazol was injected to confirm that the respiration changes were mainly due to altered mitochondrial respiration. Experimental data show that SSd (1 μM) reduced levels of OCR and ECAR significantly (Fig. 5b and c). Injection of 0.5 μM of SSd has no effect on OCR at the fifth time point after, but it significantly inhibited ECAR (Fig. 5d and e). These results may indicate that SSd reduces oxygen consumption and the mitochondrial metabolic capacity of HSC-T6 cells.

After SSd treatment, the total protein content of HSC-T6 cells was extracted, and the expression of BAK, BAD, BAX, PUMA, Bcl-2 and Bcl-xL with specific antibodies was detected. SSd upregulated the BAK, BAD, and PUMA expressions within 8 h (Fig. 6a). Conversely, SSd downregulated the Bcl-2 expression, but did not affect the BAX or Bcl-xL expressions. The RNA expression levels of BAX, BAD, Bcl-2, and PUMA were consistent with the protein profiles (Fig. 6b). During apoptosis, BAX and BAK translocate from the cytoplasm to the mitochondria to form BAX/BAK pores. These BAX/BAK pores disturbed the membrane potential, leading to mitochondrial fracture and apoptotic factor release. To determine the effects experimentally, HSC-T6 cells were exposed to SSd (1 μM) exposure for 15, 30 and 60 min, and the mitochondrial and cytosolic fractions were then isolated for BAK and BAX detection. The organelle-specific marker COX3 expression was detected by Western blotting to ensure the purity of mitochondria [22]. As indicated in Fig. 7a, COX3 was present significantly in the mitochondrial fraction, while cytosol marker GAPDH was absent. BAK and BAX could be detected in the mitochondrial fraction of the 30- and 60-min SSd-treated cells (Fig. 7b). Additionally, GAPDH was present in the cytosolic fraction, but not in the mitochondrial fraction (Fig. 7c). SSd reduced BAK and BAX expressions in the cytosolic fraction within 60 min (Fig. 7d). The high purity of the mitochondria ensured that SSd increased BAK and BAX expression in mitochondria, while reducing it in cytoplasm. Moreover, the mitochondrial membrane potential and MitoTracker® Deep Red FM staining signal fell after SSd treatment (Fig. 7e and f). To further study the effect of SSd on apoptotic factor release, the mitochondrial and cytosolic fractions were isolated from HSC-T6 cells after SSd treatment. The purity of the mitochondrial and cytosolic fraction was also confirmed by the specific markers COX3 and GAPDH (Fig. 8a and b). Following SSd-induced mitochondrial function impairment, the mitochondial content of apoptotic factors, including Cyto c, EndoG, and AIF, fell while the cytoplasmic content of apoptotic factors rose (Fig. 8c and d). In addition, the apoptotic factor staining signal and mitochondrial staining signal fell after the 60-min SSd treatment, as revealed by fluorescent immunocytochemical staining and MitoTracker® Deep Red FM staining (Fig. 8e). These results suggest that SSd regulates pro- and anti-apoptotic protein expression and triggers BAX and BAK translocation, resulting in decrease of mitochondrial membrane potential, and apoptotic factor release.