Veratramine (with a purity ≥98%) was extracted and isolated from the roots and rhizomes of Veratrum nigrum L., and provided by Nanjing Spring & Autumn Biological Engineering Co., Ltd. The chemical structure of veratramine is presented in Fig. 1A. The human liver cancer HepG2 cell line was purchased from the American Type Culture Collection. FBS, penicillin streptomycin and cell culture media were purchased from Hangzhou Sijiqing Biological Engineering Materials Co., Ltd. The Cell Counting Kit-8 (CCK-8) assay kit was obtained from Nanjing KeyGen Biotech Co., Ltd. The apoptosis detection kit and antibodies used for western blotting were purchased from Beyotime Institute of Biotechnology. The reverse transcription kit and PCR kit were purchased from Vazyme Biotech Co., Ltd. Acridine orange (AO) and 3-methyladenine (3-MA) were obtained from Sigma-Aldrich; Merck KGaA. All other reagents and solvents (AR grade) were purchased from Sinopharm Chemical Reagent Co., Ltd. Ten BALB/c nude mice, aged 4–6 weeks (16–18 g), were obtained from Shanghai Slack Laboratory Animals Co., Ltd. These mice were housed in specialized mouse cages at ~20°C, humidity 50–60%, specific pathogen-free, dark conditions, and fed with normal food and clean water.

The human liver cancer HepG2 cells were cultured in RPMI-1640 media containing 10% FBS, penicillin (100 units/ml) and streptomycin (100 units/ml). The cells were maintained in a humid incubator (Sanyo XD-101; Sanyo) with 5% CO₂ at 37°C.

Cell viability was assessed using a CCK-8 assay. HepG2 cells (5×10³ cells/well) were seeded in 96 well plates for 24 h, and then treated with various concentrations of veratramine (0, 2.5, 5, 10, 20, 40 and 80 µM) for an additional 24, 48 or 72 h. Subsequently, 10 µl of CCK-8 solution was added to each well and the cells were further incubated for 4 h. The absorbance was measured at 450 nm using a microplate reader (Bio-Rad Laboratories, Inc.). IC₅₀ values were calculated using the cell survival data curve.

The effect of veratramine on HepG2 cell migration and invasion was assessed using a wound healing assay and Transwell assay, respectively. For the wound healing assay, HepG2 cells were seeded in 6 well plates and cultured to 90% confluence. The cell monolayer was scratched using a 10-µl pipette tip. Subsequently, the cells were treated with veratramine (40 µM) for 48 h. After drug treatment, the cells were observed through an inverted microscope (Olympus Corporation), and the width of the scratch was measured.

For the Transwell assay, Transwell culture inserts were placed into the wells of 6-well plates and coated with a layer of Matrigel™. Subsequently, RPMI-1640 medium containing 10% FBS was added to the lower chamber and HepG2 cells were seeded in the upper chamber. After veratramine (40 µM) treatment for 48 h, the invaded cells in the lower chamber were fixed with 1% formaldehyde for 10 min at 25°C and stained with 0.5% crystal violet for another 5 min. Finally, the cells were observed under an inverted microscope (magnification, ×200).

Cell apoptosis was evaluated using the Annexin V-FITC/7AAD double staining assay. Briefly, HepG2 cells (3×10⁵ cells/well) were seeded in 6-well plates for 24 h. Subsequently, the cells were treated with increasing concentrations of veratramine (10, 20 and 40 µM) for 48 h. After incubation, the cells were harvested and stained with Annexin V-FITC and 7AAD according to the manufacturer's instructions. Apoptotic cells were quantified using flow cytometry (FACSCalibur; BD Biosciences) and the data were analyzed using FlowJo software (FlowJo V10; Tree Star, Inc.).

RT-qPCR was used to evaluate the expression levels of the key apoptotic genes, BCL2 and BAX, and AP-1 downstream gene ETS1. After phosphate-buffered saline (PBS) or veratramine treatment for 48 h, HepG2 cells were subjected to total RNA isolation using TRIzol^® reagent. Subsequently, cDNA was synthesized using the HiScript III 1st Strand cDNA Synthesis kit and qPCR was performed using the ChamQ SYBR qPCR Master Mix according to the manufacturer's instructions. Thermocycling parameters were optimized as 5 min at 95°C, followed by 40 cycles at 95°C (15 sec), 60°C (30 sec); melt curve: 95°C (15 sec), 60°C (30 sec) and 95°C (15 sec). PCR primers were provided by Guangzhou RiboBio Co., Ltd., and β-actin was used as the internal reference. Quantitative analysis of relative mRNA expression levels was performed according to the 2^−ΔΔCq method (36). Primer sequences were as follows: BCL2 forward, 5′GTGGAGGAGCTCTTCAGGGA3′ and reverse, 5′AGGCACCCAGGGTGATGCAA3′); BAX forward, 5′GGCCCACCAGCTCTGAGCAGA3′ and reverse, 5′GCCACGTGGGCGTCCCAAAGT3′; ETS1 forward, 5′GCGCGCTAGCAACTTGCTACCATCCCGT3′ and reverse, 5′GCGCAAGCTTTGCCATCACTCGTCGGC3′; β-actin forward, 5′ACAAAGTGGTCATTGAGGGC3′and reverse, 5′GCCGTCAGGCAGCTCGTAGC3′.

Formation of acidic vesicular organelles (AVO) is a hallmark of autophagic cells. Acridine orange (AO) dye readily penetrates cell membranes and visualizes the occurrence of the acidic vesicular organelles (AVOs). Therefore, AO staining was used to detect autophagy. HepG2 cells were cultured overnight and treated with varying concentrations of veratramine (10, 20 and 40 µM) for 48 h. Subsequently, the cells were stained with 10 µg/ml of AO dye for 20 min at 37°C in the dark. AVO formation (Red) was observed using fluorescence microscopy (magnification, ×200). The percentage of autophagic cells was measured using flow cytometry.

HepG2 cells were cultured overnight and then treated with varying concentrations of veratramine (10, 20 and 40 µM) for 48 h. An equal volume of PBS was used as a control. After drug treatment, the cells were harvested and lysed in RIPA lysis buffer supplemented with a protease inhibitor cocktail. Cell debris were removed by centrifugation at 14,000 × g at 4°C for 15 min and the protein concentration was determined using BCA assays. Subsequently, 30 µg of total protein was separated on a 10% SDS-PAGE gel and electroblotted onto PVDF membranes (EMD Millipore). The membranes were blocked with 5% skim milk in TBS-Tween-20 (TBST; 20 mmol/l Tris-HCl, 150 mmol/l NaCl and 0.1% Tween-20) for 1 h at room temperature (RT), and then incubated with anti-Beclin1 (product no. 3495), anti-LC3 (product no. 3868), anti-PI3K (product no. 4249), anti-phospho-PI3K (product no. 17366), anti-Akt (product no. dilution 4685), anti-phospho-Akt (product no. 9614), anti-mTOR (product no. 2983), anti-phospho-mTOR (product no. 5536) or anti-GAPDH (product no. 5174; all with a dilution 1:1,000; all from Cell Signaling Technology, Inc.) primary antibodies at 4°C, overnight. After washing 3×10 min with TBST, the membranes were incubated with horseradish peroxidase-conjugated secondary antibodies (product no. 7074; dilution 1:2,000; Cell Signaling Technology, Inc.) for 1 h at RT. The protein bands were visualized using ECL reagent and the G:BOX chemiXR5 Imaging System (Syngene). Band intensities were quantified using Image-Pro Plus 6.0 software, with GAPDH used as an internal control.

The liver cancer xenograft model was established by subcutaneous injection of human HepG2 cells (1×10⁶ cells suspended in 0.1 ml PBS for each mouse) into the armpits of nude female BALB/c mice (6–8 weeks old). Once the tumors reached a volume of approximately 75–100 mm³, the mice were randomly divided into two groups, with five mice per group, a PBS group and veratramine group. PBS (200 µl) or veratramine (2 mg/kg) were injected into the mice via the tail vein. Based on the IC₅₀ value of veratramine on HepG2 cells in vitro, and previous studies (37–39), 2 mg/kg of veratramine was used for the animal experiment. The mice were treated three times a week for 4 weeks. During the treatment, the weights of the mice and the volume of the tumors were measured every week. The volume of the tumors was calculated according to the following formula: V = 0.5 × length × width². Three days after the last treatment, blood was collected from the mice and the mice were sacrificed. Subcutaneous tumors from the mice were excised and weighed. The number of white blood cells, neutrophils and platelets, as well as the concentrations of alanine aminotransferase, aspartate aminotransferase and creatinine were examined. The heart, liver, spleen, lungs and kidneys were excised and then fixed with 4% paraformaldehyde. The paraffin sections of the organs were performed for pathological evaluation with hematoxylin-eosin (H&E) staining (hematoxylin, 5 min; eosin, 2 min) at room temperature. The results of H&E staining were evaluated by pathologist in a blinded fashion. The animal experiments were approved by the Animal Care and Use Committee of Yijishan Hospital of Wannan Medical College.

The experiments were performed in triplicate and the data are presented as the mean ± SD. Data were analyzed using GraphPad Prism 6.0 software (GraphPad Software, Inc.). Student t-tests or ANOVAs with Tukey's post hoc test were used to evaluate the data, and P<0.05 was considered to indicate a statistically significant difference.

CCK-8, wound healing and Transwell assays were performed to examine the effects of veratramine on HepG2 cell proliferation, migration and invasion, respectively. As revealed in Fig. 1B, veratramine significantly inhibited HepG2 cell proliferation in a time- and dose-dependent manner, and the IC₅₀ values of veratramine on HepG2 cells for 24, 48, and 72 h were 26.54 µM, 19.81 and 9.12 µM, respectively. Furthermore, the data revealed that 40 µM of veratramine treatment reduced the cell migration rate compared to the PBS control (Fig. 1C and D). Consistently, the number of invasive cells was also decreased after veratramine treatment for 48 h. Collectively, these results indicated that veratramine not only suppressed cell growth but also reduced cell migration and invasion in HepG2 cells. This demonstrated the potential for veratramine to treat liver cancer.

The effect of veratramine on apoptosis was investigated using Annexin V-FITC/7AAD double staining and RT-qPCR. The cells were treated with the indicated dose of veratramine for 48 h and the same volume of PBS was used as a negative control. As indicated in Fig. 2A and B, flow cytometric data revealed that veratramine induced apoptosis in HepG2 cells, and the proportion of apoptotic cells (Annexin V⁺) increased from 8.41% (PBS control) to 51.86% (40 µM of veratramine). Although multiple signaling pathways are involved in apoptosis, the majority of them are regulated by the Bcl-2 family (40). BCL2 and BAX are representative genes in the Bcl-2 family. Among them, BCL2 is an anti-apoptotic gene, whereas BAX is a pro-apoptotic gene. Studies have indicated that the ratio of BCL2 to BAX is considered to be a decisive factor as to whether a cell will commit to apoptosis. Therefore, in order to further verify whether veratramine can induce apoptosis, the mRNA expression levels of BCL2 and BAX were detected using RT-qPCR. RT-qPCR results revealed that veratramine significantly upregulated the expression level of BAX and downregulated the expression level of BCL2, which further demonstrated that veratramine induced apoptosis in HepG2 cells (Fig. 2C and D).

To explore the mechanism of action behind the antitumor activity, the effect of veratramine on autophagy in HepG2 cells was investigated. AO staining was performed to label autophagosomes. Flow cytometric results revealed that the number of autophagic cells (AO⁺) increased with increasing veratramine concentrations. After 48 h of veratramine treatment (40 µM), the proportion of autophagic cells reached 45.3% (Fig. 3A and B). Moreover, fluorescence microscopy images further confirmed that veratramine induced autophagy in HepG2 cells (Fig. 3C). Autophagy is strictly regulated by gene expression, with Beclin-1 and LC3 proteins being markers of autophagy (41,42). During autophagy, LC3-I is converted to LC3-II and then localized to the autophagosome membrane (41). The expression levels of LC3-II are positively correlated with the degree of autophagy, and the ratio of LC3-II/I can be used to estimate the level of autophagy (41,42). In the present study, western blot analysis revealed that Beclin-1 and LC3-II proteins were significantly upregulated, while LC3-I protein was significantly downregulated, in the veratramine-treated cells. The Beclin-1 protein expression levels and the ratio of LC3-II/I were the highest in the high concentration group (40 µM) (Fig. 3D-F). Overall, the aforementioned results demonstrated that veratramine treatment could induce autophagy in HepG2 cells.

The PI3K/Akt/mTOR signaling pathway is an intracellular signaling pathway that plays an important role in the pathogenesis of cancer (30). As a downstream protein of PI3K and Akt, phosphorylated mTOR protein promotes cancer cell proliferation and angiogenesis (28,29). A previous study has demonstrated that the PI3K/Akt/mTOR pathway is associated with autophagy (43). Therefore, it was hypothesized that veratramine may inhibit the PI3K/Akt/mTOR pathway to promote autophagic cell death. In order to verify this hypothesis, the phosphorylation levels of PI3K, Akt and mTOR proteins were assessed using western blotting. The results revealed that the levels of p-PI3K, p-Akt and p-mTOR proteins were significantly lower in veratramine-treated cells than in the control cells, and the changes were achieved in a dose-dependent manner (Fig. 4A and B). Moreover, a previous study demonstrated that veratramine modulated AP-1-dependent gene transcription by directly binding to programmable DNA (35). ETS proto-oncogene 1, transcription factor (ETS1), an AP-1 downstream gene (35), has been revealed to be predominantly expressed in various tumor subtypes and can mediate activation of the PI3K/Akt/mTOR pathway (44,45). As anticipated, the mRNA level of ETS1 was significantly downregulated in veratramine-treated HepG2 cells (Fig. 4C). Based on these results, it was concluded that veratramine induced autophagy by blocking the PI3K/Akt/mTOR signaling pathway in HepG2 cells.

To evaluate the effect of autophagy on veratramine-induced cell death, autophagy inhibitor 3-MA was used and the proportion of apoptotic cells and the cell viability was examined. Cells were co-incubated with veratramine (40 µM) for 48 h in the presence or absence of 3-MA (2.5 mM). As revealed in Fig. 5A and B, after the addition of 3-MA, the number of autophagosomes was significantly decreased, demonstrating that autophagy induced by veratramine was significantly inhibited. Fluorescence microscopy results further confirmed that autophagic vacuoles were reduced after 3-MA co-treatment (Fig. 5C). Moreover, cell viability in cells treated with a combination of veratramine and 3-MA, was higher than that in cells treated with veratramine alone, which indicated that 3-MA attenuated the cytotoxicity of veratramine on HepG2 cells (Fig. 5D). Similarly, the proportion of apoptotic cells in the co-treatment group was lower compared with that of the veratramine alone treatment group (Fig. 5E and F). Collectively, these results indicated that veratramine induced-autophagy promoted apoptosis in HepG2 cells.

Based on the findings of the present in vitro experiments, the antitumor effect of veratramine was further verified in the HepG2 subcutaneous xenograft mouse model. No mice died or acted abnormally before the end of the animal experiment. By monitoring the tumor volume, it was observed that veratramine treatment significantly inhibited tumor growth compared with the PBS control (Fig. 6A-B). Consistent with the tumor volume, tumor weight in the veratramine group was significantly lower than the tumor weight in the PBS group (Fig. 6C). In addition, the systemic toxicity of veratramine was also evaluated in vivo. The weight of the mice in the veratramine-treated group was not significantly different from that in the PBS group (Fig. 6D). Furthermore, veratramine treatment had no significant effect on the number of blood cells as well as the liver and kidney function compared with the PBS control (Fig. 6E-H). Moreover, the results of H&E staining revealed that no obvious tissue damage, such as inflammation (infiltration of inflammatory cells) and/or necrosis (tissue structural destruction), was observed in tissue sections of the heart, liver, spleen, lung and kidney (Fig. 6I), which indicated that veratramine had no obvious organ toxicity. Collectively, these results demonstrated that veratramine not only suppressed tumor growth in vivo, but also had no obvious systemic toxicity.