Introduction
Primary liver cancer (PLC) is a prevalent form of cancer, ranking as the fourth most deadly tumor and the sixth most common in terms of new cases globally.1 Hepatocellular carcinoma (HCC) accounts for around 90% of all PLC cases, with an annual incidence of 850,000 new cases.2,3 Approximately 80% of HCC patients are diagnosed at an advanced stage due to the absence of early symptoms and rapid disease progression.4 Although drugs like sorafenib are accessible, the five-year survival rate for patients with advanced liver cancer remains below 5%.5,6 Consequently, it is crucial and of great significance to investigate novel treatment alternatives for HCC.
Oncolytic viruses (OVs) are viral agents that specifically target and eliminate cancer cells while sparing healthy tissues.7 OVs encompass both naturally occurring and genetically engineered viruses, with the latter showing enhanced efficacy and improved targeting capabilities.8 OVs can reproduce within tumor cells, generating additional offspring virions that destroy neighboring tumor cells through various mechanisms, including direct lysis of tumor cells, stimulation of local or systemic immune responses against the tumor, and modification of the tumor microenvironment.9 Several studies have demonstrated significant antitumor effects of oncolytic vaccinia virus containing Aphrocallistes vastus lectin in HCC cells and mouse models with transplanted human tumors.10,11 Additionally, the wild-type Newcastle disease virus HK84 strain has been reported to specifically suppress HCC, with its anticancer activity linked to the activation of interferon (IFN) type I.12
Senecavirus A (SVA) is a natural oncolytic virus.13 It is a non-segmented RNA virus with a single positive-stranded genome and lacks a vesicle membrane. SVA belongs to the genus Senecavirus within the Picornaviridae family.14 The viral particles exhibit a characteristic symmetrical icosahedral shape, measuring approximately 27 nm in diameter and weighing around 30 kDa.15 In 2015, the International Committee on Taxonomy of Viruses renamed the original strain of Seneca Valley virus (SVV), known as SVV-001, to SVA. Since SVA reproduces using RNA intermediates and does not go through a DNA stage or integrate into the host genome, it is considered relatively safe. Research indicates that SVA has a strong capability to target tumors with neuroendocrine characteristics.8 A solitary intravenous administration of the oncolytic picornavirus SVV-001 eradicated medulloblastomas in primary tumor-derived orthotopic xenograft mouse models.16 Surprisingly, the administration of 1014 vp/kg of the oncolytic virus SVV-001 successfully eradicated tumors in two out of six heterotransplant mouse models of a specific small cell lung cancer variant, with long-lasting effects.17 Nevertheless, there have been no documented studies on the potential application of SVA for treating HCC.
The objective of this study was to examine the therapeutic impact of SVA on HCC through both in vitro and in vivo experiments. The MIHA cell line, a normal immortalized human hepatocyte cell line, was utilized as a normal control in the study.18 Our investigation revealed that SVA suppresses the growth of HepG2 cells by triggering apoptosis and halting the cell cycle. Administering SVA through direct injection into the tumor effectively suppresses HCC growth in a mouse model while maintaining a favorable safety profile. The results indicate that SVA has the potential to be a new and effective treatment approach for HCC.
Methods
Cells and virus
Hepatocellular carcinoma cell line HepG2 was cryopreserved in our laboratory. The MIHA cell line, a normal immortalized human hepatocyte cell line, was generously provided by Professor Juan Chen from Chongqing Medical University, China, and used as a normal control. The BHK-21 (baby hamster kidney cells) and SVA (GenBank: MH716015.1) were acquired as a generous donation from Professor Xiao Liu of Southwest University, China. BHK-21, MIHA, and HepG2 cells were cultivated in Dulbecco’s modified Eagle’s medium (DMEM) from VivaCell, China, supplemented with 1% mycoplasma inhibitor from Trans, China, and 10% (V/V) fetal bovine serum (FBS) from Biological Industries, Israel. The cells were cultured in a cell incubator at 37°C with 5% CO2 concentration.
Virus amplification and purification
BHK-21 cells were seeded in 10 cm culture dishes until they reached approximately 80% coverage, then the medium was replaced with 10 mL of serum-free medium. The SVA virus stock was introduced and incubated for 4 h at 37°C to facilitate virus adsorption. Subsequently, FBS was introduced to achieve a final concentration of 2%. When over 80% of the cells displayed cytopathic effects, the virus was collected by subjecting the cells to two cycles of freezing and thawing, followed by centrifugation at 12,000 ×g for 30 m to remove solid particles. The supernatant was then centrifuged at 120,000 ×g for 2 h at 4°C to concentrate it. The precipitate was resuspended in phosphate-buffered saline (PBS) to a final volume that was 2% of the original supernatant volume. The viral suspension was then aliquoted and stored at −80°C until needed.
TCID50 assay
Viral titers were determined by TCID50 assays as previously described.19 HepG2 cells were distributed at a density of 1×104 cells per well in a 96-well plate with DMEM supplemented with 10% FBS. Following the attachment of cells, the virus stock was diluted in serum-free DMEM from 10−1 to 10−10 in serial steps. The diluted virus was then applied to the cells for 4 h, with eight replicates for each dilution. Subsequently, the inoculum was substituted with 200 µL of DMEM supplemented with 2% FBS. Cellular morphology was observed daily to document the number of cytopathic effects. The TCID50 (tissue culture infectious dose 50) was determined using the Reed-Muench method.20
Cell viability assay
HepG2 and MIHA cells were distributed in a 96-well plate at densities of 2×103 and 3×103 cells per well, respectively. The cells were subjected to SVA at specific multiplicities of infection (MOI) as specified in various assays for 4 h at 37°C. After adsorption, the medium was replaced with 200 µL of DMEM containing 10% FBS. The cells were then incubated for 24, 48, and 72 h, with three replicates for each time point. The CCK-8 assay, using the Biosharp kit from China, was conducted following the methodology previously outlined.21 In summary, a combination of 100 µL of serum-free medium and 10 µL of CCK-8 reagent was introduced into each well. Following a 2-h incubation period at 37°C protected from light, the absorbance at a wavelength of 450 nm was quantified using a spectrophotometer from PerkinElmer, United States. Cell viability was calculated using the following formula:
Cell survival rate (%)=ODexperiment−ODblankODcontrol−ODblank×100%.
Colony formation assay
HepG2 and MIHA cells were placed in a 6-well plate at densities of 1×103 and 2×103 cells per well, respectively. The cells were exposed to either PBS or SVA at a MOI of 0.1 or 0.5. They were cultured until each clone contained more than 50 cells (two weeks). The cells were then fixed with 4% paraformaldehyde (Solarbio, China) for 30 m, stained with 0.1% crystal violet (Solarbio, China) for 15 m at room temperature, and imaged.22 Colonies were quantified using ImageJ software.
Cell cycle analysis
Cells were infected with SVA for 24 h. The cell cycle distribution was determined using the DNA Content Quantitation Assay kit (Cell Cycle) from Solarbio, China, following the previously described method.23 Specifically, 1 million cells were collected and preserved in pre-cooled 70% ethanol for 4 h at 4°C. The cells were then exposed to 100 µL of RNase A at 37°C for 30 m. Afterward, 400 µL of PI staining solution was added. The mixture was incubated at 4°C for 30 m in the dark and analyzed using flow cytometry (FACSCanto™, BD, USA). The data were analyzed with FlowJo software.
Detection of apoptotic cells by flow cytometry
The evaluation of apoptosis was conducted via the implementation of Annexin V/PI double staining, utilizing an Annexin V-fluorescein (FITC)/PI apoptosis kit (BD, USA) as previously described.24 The cells were cultured and exposed to SVA for a duration of 24 h. A total of 500,000 cells were collected, and five microliters of Annexin V and five microliters of PI staining solution were introduced. Subsequently, the cells were placed in a dark environment and maintained at room temperature for a period of 30 m. The mixture underwent flow cytometry to evaluate cell apoptosis. The data were subsequently processed using the FlowJo software. The different quadrants in the flow cytometry analysis are defined as follows: Annexin V-FITC (−)/PI (+), indicating the number of deceased cells; Annexin V-FITC (+)/PI (+), representing the number of cells in the late apoptotic/necrotic state; Annexin V-FITC (+)/PI (−), signifying the number of cells in the early apoptotic state; and Annexin V-FITC (−)/PI (−), indicating the quantity of viable cells.
Nuclear staining by Hoechst 33258
The identification of altered nuclear morphology and apoptotic body formation was conducted via Hoechst 33258 staining, in accordance with the previously described method.25 The cells were cultured and exposed to SVA for a duration of 24 h, after which they were fixed for 30 m using 4% paraformaldehyde fixative (Solarbio, China) at room temperature. Subsequently, the cells were exposed to Hoechst 33258 staining solution (Solarbio, China) and incubated for 30 m at 37°C in the dark. Following two washes with PBS, the nuclei stained with Hoechst 33258 were visualized using an inverted fluorescence microscope manufactured by Olympus in Japan. Apoptotic cells exhibit a number of distinctive characteristics, including nuclear condensation, fragmentation, the formation of apoptotic bodies, and an increased fluorescence intensity.
Mitochondrial membrane potential (MMP) assay
The cells were infected with SVA for a duration of 24 h. The MMP was measured using a mitochondrial membrane potential assay kit with JC-1 (Beyotime, China), as previously described.26 In conclusion, a total of one million cells were gathered and subjected to centrifugation at a speed of 1,000 revolutions per minute for a duration of 5 m. The cell precipitate should be retrieved and a 500 µL solution of JC-1 staining (20 µg/mL) should be applied to it. The mixture should be incubated at a temperature of 37°C in a light-free environment for a duration of 20 m. Subsequently, the cells underwent two washes with the JC-1 staining buffer prior to being subjected to flow cytometry analysis. The data were analyzed using the FlowJo software.
In vivo xenograft tumor model
Male BALB/c nude mice, aged four to six weeks and weighing 18–21 g, were purchased from Chengdu Ensiweier Biotechnology Co., Ltd. HepG2 cells in the logarithmic growth phase were harvested and diluted to a concentration of 5×107 cells per milliliter. Then, 0.1 mL of the cell suspension was injected subcutaneously into the right flank of each nude mouse. The length (a) and width (b) of the tumors were measured daily to calculate their volume using the formula: V=(a×b2)/2.27 After the tumors grew to approximately 100 mm3, the mice were randomly divided into three groups: the PBS-treated group (100 µL/day), the SVA-treated group (6×106 TCID50/day), and the cisplatin (DDP)-treated group (20 mg/kg per day). Each group consisted of 12 mice. The DDP-treated group served as a positive control to assess the antitumor effect. Each mouse received a total of three injections on alternate days directly into the tumor. After twenty-eight days following the initial treatment, the mice were euthanized, and samples of their liver, heart, spleen, lungs, kidneys, blood, and tumors were collected.
RNA isolation and RT-qPCR
Total RNA was isolated from the mouse viscera, tumor tissues, and blood using TRIzol (Invitrogen, USA), as previously described.28 The synthesis of complementary DNAs was performed using a complementary DNA synthesis kit from Toyobo, Japan, following the manufacturer’s protocol. RT-qPCR analyses were conducted using NovoStart® SYBR qPCR SuperMix Plus (Novoprotein, China), following the manufacturer’s protocol. The SVA primers used were: Forward primer sequence: 5′-ATTGGTGTGGTCTGCGAGTT-3′, Reverse primer sequence: 5′-GTGCGAGGGCTAAGTCTTGT-3′. The GAPDH primers used were: Forward primer sequence: 5′-GCCTCCTGCACCACCAACTG-3′, Reverse primer sequence: 5′-ACGCCTGCTTCACCACCTTC-3′. The Ki67 primers used were: Forward primer sequence: 5′-ACGCCTGGTTACTATCAAAAGG-3′, Reverse primer sequence: 5′-CAGACCCATTTACTTGTGTTGGA-3′. The expression of Ki67 mRNA was standardized to the housekeeping gene GAPDH.
H&E staining
H&E staining was performed as previously described.29 The visceral tissues of mice were immersed in a 4% solution of paraformaldehyde for 24 h to preserve them. After that, the tissue sections underwent dewaxing using xylene, followed by hydration with a gradient of ethanol. Subsequently, hematoxylin staining (Biosharp, China) was applied for 5 m, followed by differentiation with 1% hydrochloric acid. After counterstaining with eosin (Biosharp, China) for 3 m, the sections were dehydrated and permeabilized. After sealing with neutral resin, they were observed under an optical microscope (Leica, Germany) at 400× magnification.
Immunohistochemistry
Immunohistochemistry was performed as described previously.30 The tumor tissues underwent fixation, dehydration, embedding, and sectioning, following the protocol outlined in the H&E staining assay mentioned earlier. The sections embedded in paraffin were dewaxed in a sequential manner. The Tris-EDTA antigen retrieval buffer from Servicebio, China, was utilized for antigen retrieval. The endogenous peroxidase activity was inhibited using a 3% solution of hydrogen peroxide obtained from the National Medicine Group in China for 25 m at room temperature in the dark. Non-specific binding was inhibited by treating with 5% normal goat serum (Solarbio, China) for 30 m at room temperature. Afterward, the sections were treated with rabbit polyclonal to Ki67 primary antibody (Abcam, UK) at 4°C overnight, followed by incubation with goat anti-rabbit secondary antibody (Servicebio, China) for 50 m at room temperature. The sections were stained with 3,3′-diaminobenzidine (Biosharp, China). Hematoxylin (Servicebio, China) was used to counterstain the cell nuclei for 30 s, and a neutral mounting medium was used for sealing. Microscopic images were acquired, and data analysis was conducted using ImageJ software.
TUNEL assay in tumor tissues
The tumor tissues underwent fixation, dehydration, embedding, and sectioning, in accordance with the protocol outlined in the H&E staining assay previously described. The detection of apoptosis was performed using an FITC TUNEL cell apoptosis detection kit (Servicebio, China), following the previously described method.31 In conclusion, the paraffin sections were subjected to a process of paraffin removal and proteinase K (Servicebio, China) treatment for 20 m at room temperature, which renders the sections permeable. Subsequently, the sections were incubated with TdT incubation buffer (recombinant TdT enzyme, FITC-12-dUTP Labeling Mix: Equilibration Buff=1:5:50) for 1 h at 37°C. Following this, the sections were stained with DAPI (Biosharp, China) for 10 m at room temperature in the dark. Cells that exhibited TUNEL positivity, as indicated by the presence of green nuclei, were visualized using an inverted fluorescence microscope. The data were analyzed using the ImageJ software.
Statistical analysis
The statistical analyses were conducted using GraphPad Prism version 8 software, developed by GraphPad Software in the United States. The results are presented as the mean value plus or minus the standard error of the mean. To evaluate the statistical significance of differences between the two groups, an unpaired two-tailed Student’s t-test was employed. The statistical significance of the results was determined using one-way ANOVA for three or more groups. A significance level of p<0.05 was employed to determine statistical significance.
Results
SVA suppresses the growth of HepG2 cells in vitro
To assess the influence of SVA on cell viability, we conducted a CCK-8 assay on HepG2 and MIHA cells, both with and without SVA infection. HepG2 and MIHA cells were exposed to SVA at three different MOIs – 0.1, 0.5, and 2.5 – for a duration of 48 h. The viability of HepG2 cells was found to have decreased significantly when compared to the control group treated with PBS. The viability of the HepG2 cells decreased significantly from 100.1±2.3% to 54.9±2.7% at MOI=0.1, 33±0.98% at MOI=0.5, and 14.3±1.45% at MOI=2.5 (Fig. 1A). Furthermore, HepG2 and MIHA cells were exposed to SVA at an MOI of 0.5, and cell viability was measured at 24 h, 48 h, and 72 h, respectively. A significant decrease in cell viability was observed over time in HepG2 cells. The viability of the HepG2 cells decreased from 100±0.6% to 73.5±2.6% at 24 h, 32.8±1.7% at 48 h, and 17.4±0.5% at 72 h. In contrast, there were no notable alterations in the survival of MIHA cells treated with SVA compared to the control group treated with PBS (Fig. 1B).
To validate the inhibitory impact of SVA on HCC cells, we conducted a colony formation assay. The number of clones formed by HepG2 cells significantly decreased from 152±2 to 46±6 and 27±6 after SVA infection at MOI of 0.1 and 0.5, respectively, in comparison to the control group treated with PBS. However, there were no notable alterations in the number of colonies produced by MIHA cells, regardless of whether they were infected with SVA or not (Fig. 1C, D). The results suggest that SVA specifically inhibits the proliferation of HCC cells.
SVA-induced cell cycle arrest at the S phase in HepG2 cells
Subsequently, we employed flow cytometry to examine the influence of SVA infection on the cell cycle advancement of HepG2 and MIHA cells that were infected with SVA. The percentage of HepG2 cells in the G0/G1 phase exhibited a significant decline from 68.3±0.5% to 62±0.6% and 53±0.4% following SVA infection at MOIs of 0.1 and 0.5, respectively, compared to the control group treated with PBS. Conversely, the percentage of HepG2 cells in the S phase increased from 24.1±0.2% to 32.8±1.1% (MOI=0.1) and 44.9±2.8% (MOI=0.5) (Fig. 2A, B). Nevertheless, the cell cycle distribution of MIHA cells did not exhibit any notable alterations following SVA infection (Fig. 2A, C). The results demonstrate that SVA induced the HepG2 cells to cease their cell cycle at the S phase.
SVA induced apoptosis in HepG2 cells
In order to examine the influence of SVA infection on apoptosis in HCC cells, we utilized Hoechst 33258 staining to label the nuclei of HepG2 and MIHA cells following SVA infection. Subsequently, the stained cells were observed using an inverted fluorescence microscope. HepG2 cells infected with SVA exhibited typical apoptotic nuclear morphology, characterized by nuclear condensation, fragmentation, formation of apoptotic bodies, and increased fluorescence intensity, in comparison to the control group treated with PBS. The fluorescence intensity of the nuclei in MIHA cells was not significantly altered in the absence or presence of SVA infection (Fig. 3A). The findings demonstrated that SVA stimulated apoptosis in HepG2 cells.
Flow cytometry was employed to quantitatively analyze apoptosis in HepG2 and MIHA cells following SVA infection. The apoptosis rates of HepG2 cells were significantly elevated in the SVA-infected group compared to the control group treated with PBS. The rates of early and late apoptosis in HepG2 cells were 6.6±0.5% and 7.7±0.5%, respectively, when infected with SVA at a MOI of 0.1. At a MOI of 0.5, the rates of early and late apoptosis were 13.3±0.8% and 9.3±0.2%, respectively (Fig. 3B, C). Nevertheless, no significant differences were observed in the rates of apoptosis between the MIHA cells and the control group treated with PBS (Fig. 3B, D). The results indicate that SVA induces apoptosis in HepG2 cells in a manner that is dependent on dosage. However, no similar effect was observed in MIHA cells.
JC-1 is a fluorescent dye that exhibits both lipophilic and cationic properties. JC-1 exists in two distinct forms: as individual molecules, referred to as monomers, or as larger structures, known as aggregates. In the absence of pathological conditions, JC-1 molecules aggregate within the mitochondria of healthy cells, resulting in the emission of red fluorescence that can be observed using the PI channel of a flow cytometer. During apoptosis, the reduction of MMP results in heightened permeability and subsequent liberation of JC-1 from the mitochondria, where it undergoes a transition to its monomeric state. The monomeric form of JC-1 can be identified by its green fluorescence when using the FITC channel of a flow cytometer. The ratio of green to red fluorescence in cells can be used to evaluate changes in MMP, which is regarded as one of the initial biological changes in cell apoptosis. The study revealed a notable rise in the ratio of green and red fluorescence in HepG2 cells, increasing from 5.6±1% to 19.4±3% and 22.9±2.7% following SVA infection at MOIs of 0.1 and 0.5, respectively (Fig. 4A, B). The presence of SVA during incubation did not result in any noticeable variation in MMP levels in MIHA cells (Fig. 4A, C). The results indicate that SVA induced apoptosis in HepG2 cells in a manner that depended on the dosage, while it had no impact on MIHA cells.
SVA inhibited HCC cells growth in vivo
In order to examine the potential suppressive impact of SVA on the growth of HepG2 cells in living organisms, a model of subcutaneous tumor xenografts in BALB/c nude mice was created (Fig. 5A). The PBS treatment served as the negative control, while the DDP treatment served as the positive control. One mouse in the group treated with PBS died following the second injection. The tumor growth was significantly suppressed in both the SVA- and DDP-treated groups compared to the PBS-treated group (Fig. 5B). On day 28 following the initial injection, the tumor volumes of the groups that received PBS, SVA, and DDP were 1,330±80.6 mm3, 808.9±102.2 mm3, and 672.6±88.7 mm3, respectively (Fig. 5C). The weights of eight tumors from each group were measured and recorded as follows: 0.9±0.07 g, 0.52±0.05 g, and 0.42±0.05 g (Fig. 5D). The results demonstrated that SVA effectively suppressed the proliferation of HepG2 cells in an in vivo setting.
Furthermore, the TUNEL assay was employed to evaluate apoptosis in tumor tissues from both the SVA-treated and PBS-treated groups. The SVA-treated group exhibited approximately double the number of TUNEL-positive cells in the tumor tissue compared to the PBS-treated group, suggesting an increase in apoptosis (Fig. 5E). Moreover, the SVA-treated group exhibited a reduction of approximately 50% in the expression of Ki67, a prominent indicator of tumor proliferation, at both the mRNA and protein levels (Fig. 5F, G, respectively). These findings indicate that SVA suppresses the growth of HepG2 cells in living organisms, most likely by enhancing programmed cell death.
Toxicity analysis of SVA in mice
In order to assess the toxicity of SVA on mice, we initially quantified the levels of SVA RNA in various tissues, including the liver, heart, spleen, lung, kidney, blood, and tumor samples obtained from the mice at the end of the experiment. The presence of SVA RNA was exclusively observed in the liver, tumor tissue, and blood of the SVA-treated mice. Conversely, no SVA RNA was detected in the heart, spleen, lung, and kidney (Fig. 6A). The level of SVA RNA in tumor tissue was significantly higher than that observed in the mouse liver. The histological examination of mouse heart, liver, spleen, lung, and kidney tissues revealed no notable pathological alterations when compared to the PBS-treated group, as indicated by H&E staining (Fig. 6B). These findings indicate that direct delivery of SVA into the tumor is a secure and efficacious approach for the treatment of HCC.
Discussion
SVV-001, an oncolytic picornavirus that occurs naturally, was initially discovered in 2002 in a cell culture that was likely contaminated with porcine trypsin or bovine serum containing SVV-001.32 SVV-001 has been proven to exhibit specific cytotoxicity towards tumor cells that exhibit neuroendocrine characteristics.32 In this study, we sought to examine the impact of SVA on HCC cells. Notably, this study represented the first demonstration of SVA’s strong oncolytic activity against HepG2 cells in both laboratory settings and live organisms. It is of note that SVA does not cause any harm to normal human liver cells or mouse organs.
The cell cycle serves as the foundation for cell proliferation, with a primary emphasis on two key events: the duplication of genomic DNA during the S phase and its subsequent distribution into two daughter cells. In the event of a disruption to the process of DNA replication, a series of DNA repair signaling events will be initiated, resulting in the extension of the S phase.33 A significant proportion of viruses employ the strategy of subverting the replication cycle of the host cell in order to create a cellular environment that is conducive to viral replication.34 The nucleocapsid protein of the porcine epidemic diarrhea virus coronavirus interacts with p53, resulting in cell cycle arrest specifically in the S phase.35 This interaction ultimately facilitates viral replication.35 Our study revealed that SVA infection in HepG2 cells leads to a decrease in the proportion of cells in the G0/G1 phase and an increase in the proportion of cells in the S phase. This indicated that the cell cycle is arrested and blocked, resulting in the accumulation of cells in the S phase and inhibition of cell proliferation. The inhibitory effect of SVA infection was confirmed by the decrease in HepG2 cell viability and clone formation, as well as the reduced expression of Ki67 protein in SVA-treated tumor tissues (Figs. 1 and 5F, G). The Ki67 protein is a widely recognized indicator used to evaluate cell proliferation.36,37 It is typically found only in cells that are actively dividing. It has been documented that Ki67 plays a crucial role in the normal progression of the S phase and the arrest of the cell cycle.38,39 Consequently, our findings indicated that SVA induced cell cycle arrest at the S phase in HepG2 cells, thereby inhibiting tumor cell proliferation. Nevertheless, it should be noted that our study is subject to certain limitations. Further experimentation is required to elucidate the precise signaling mechanisms involved in cell cycle arrest following SVA infection.
Furthermore, the inhibition of tumor cell proliferation was significantly correlated with the stimulation of apoptosis. Xu et al. provided evidence that the A-to-I edited form of miRNA-379-5p suppressed the growth of cancer cells by promoting apoptosis through CD97 activation.40 A separate study demonstrated that methanol derived from D. linearis effectively suppressed the growth of human breast cancer cells by causing cell cycle arrest in the S-phase and promoting apoptosis.41 Research has demonstrated that in the majority of cancer cases, the apoptosis process is suppressed to promote the growth of tumor cells.42 Consequently, a significant proportion of drugs exert their anti-tumor effect by stimulating apoptosis in tumor cells.43 Furthermore, it has been documented that SVV has the ability to trigger apoptosis in medulloblastomas.16 SVV proteins 2C and 3Cpro were found to trigger apoptosis in tumor cells through the intrinsic pathway, which is mediated by mitochondria.44 Moreover, Newcastle disease virus, reovirus, and vesicular stomatitis virus were found to induce apoptosis in tumor cells.45–47 In our study, we observed that SVA had a significant inhibitory effect on the proliferation of HepG2 cells. This was evidenced by the reduced cell viability observed in the CCK-8 assay and the decreased number of cell clones in the cell clone formation assay. Furthermore, our findings demonstrated that SVA induced apoptosis, as evidenced by nuclear condensation, fragmentation, and the formation of apoptotic bodies. Additionally, SVA was found to result in a reduction in MMP. In conclusion, our findings indicate that SVA infection in HepG2 cells results in the cessation of the cell cycle and the initiation of apoptosis, thereby impeding the proliferation of tumor cells.
Ensuring the safety is a paramount consideration in the field of oncolytic virus therapy. Prior studies have indicated that in A/J mice, a breed that is susceptible to viral replication, no toxic effects that would restrict the dosage were observed following a single systemic injection of SVV-001 at doses up to 1×1014 vp/kg.48 Moreover, a Phase I clinical trial of SVV-001 therapy for advanced solid tumors with neuroendocrine characteristics was conducted, administering doses ranging from 107 to 1011 vp/kg.49 The trial demonstrated that SVV-001 treatment was safe and well-tolerated. In this study, the presence of SVA RNA was observed in the mouse liver tissues 28 days after intratumorally injections of SVA. Nevertheless, the amount of viral RNA detected was considerably lower compared to the levels found in tumors. Additionally, it was observed that in a specific mouse utilized in the experiment, the tumor completely vanished. Notably, the presence of SVA RNA in the liver tissue was no longer detectable at two weeks after the tumor disappeared (data not shown). Therefore, we hypothesized that after undergoing significant replication within the tumor, SVA may infiltrate the bloodstream and subsequently migrate to the liver. However, it is unable to replicate within the liver tissues of healthy mice. Furthermore, it is crucial to acknowledge that the mice employed in this study exhibited a weakened immune system, which might have contributed to their inability to effectively eliminate SVA. This could also account for the observable presence of SVA in the bloodstream and liver of the mice, even 28 days after the initial injection into the tumor. Consequently, further investigation is necessary to obtain additional information regarding the infection and replication of SVA in the liver of mice with a fully functioning immune system. In addition, histological examination of liver, heart, spleen, lung, and kidney tissues from mice treated with SVA revealed no discernible pathological alterations when compared to the control group treated with PBS. These results are consistent with those of previous studies.48,49 These findings indicate that intratumorally administration of SVA treatment for HCC is a safe approach.
This study is subject to a number of limitations. Firstly, while the evidence presented in this study indicates that SVA inhibits the growth of HCC by causing cell cycle arrest and apoptosis, the specific mechanisms behind this need to be determined. Although we have confirmed that SVA causes cell apoptosis through various means, including the presence of apoptosis-related chromatin condensation, fragmentation, formation of apoptotic bodies, and increased fluorescence intensity observed using Hoechst 33258 dye, an increased proportion of apoptotic cells was detected by the Annexin V-FITC/PI apoptosis kit (FCM). Furthermore, disrupted mitochondrial membrane potential was detected by a specific detection kit (FCM) in cells, and DNA degradation in tumor tissues was observed using the TUNEL assay, it is necessary to validate additional apoptosis-related biomarkers such as caspase enzymes, cytosolic proteins, and DNA nucleosomes in the future. This will facilitate a more comprehensive comprehension of the specific mechanism by which SVA induces apoptosis. Moreover, a model of subcutaneous tumor xenograft in mice lacking immune function was created using a hepatocellular carcinoma cell line. Nevertheless, it remains unclear whether immunocompetent mice with patient-derived HCC cells will exhibit anti-tumor effects. Furthermore, the administration of SVA involved intratumoral multipoint injections. Nevertheless, it is worth exploring whether a more convenient method, such as intravenous tail injection, can be used. Although SVA injection did not result in significant pathological alterations in non-tumor tissues of mice, further research is required to determine the safety of SVA on non-tumor tissues.