cPDS Promotes Cell Apoptosis by Reducing the Translational Efficiency of BIRC3 mRNA in HCC

Abstract

Background

Hepatocellular carcinoma (HCC) is one of the primary liver tumors with high incidence and mortality. RNA G-quadruplexes (rG4) are nucleic acid structures involved in gene expression and genome duplication. rG4 exerts its function by interacting with rG4-binding proteins. The carboxypyridostatin (cPDS), a specific ligand of rG4, are widely studied in numerous tumors. However, the role of cPDS in HCC and its regulatory mechanisms are not yet fully understood. Our study aimed to discuss the regulatory mode of cPDS on Baculoviral IAP Repeat Containing 3 (BIRC3) expression and its impact on proliferation, apoptosis, and other biologic functions in HCC.

Methods

We conducted colony formation, CCK8, Edu incorporation, scratch healing, and cell spheroid formation assays to analyze the function of cPDS on cell proliferation and migration. Additionally, we explored the role of cPDS in regulating BIRC3 expression by Western blot and qRT-PCR. Furthermore, we evaluated the impact of BIRC3 on cell proliferation and subcutaneous tumor formation in nude mice. Finally, we analyzed the regulatory mechanisms of cPDS on cell apoptosis by Western blot, qRT-PCR, flow cytometry, and Annexin V-FITC staining.

Results

Our results demonstrated that cPDS inhibited HCC cells proliferation and migration. Moreover, cPDS elevated the mRNA level while inhibiting the protein expression of BIRC3 in HCC cells. Overexpression of BIRC3 significantly enhanced the proliferation of HCC cells. In the nude mice model, BIRC3 significantly increased the tumor volume and weight. Mechanistically, cPDS promoted cell apoptosis via inhibiting BIRC3-mediated anti-apoptotic effect.

Conclusion

Our findings revealed a critical role of rG4 ligand cPDS in HCC progression and indicate that cPDS may be used for HCC treatment considering its tumor inhibitory properties by regulating cell apoptosis.

Supplementary Information

The online version contains supplementary material available at 10.1007/s10620-025-08916-0.

Introduction

Hepatocellular carcinoma (HCC) is a highly malignant tumor, its prognosis is generally poor due to its complex biologic characteristics and a variety of genetic and epigenetic variations [1, 2]. The current standard treatments for HCC include surgical resection, liver transplantation, and locoregional therapies, such as transarterial chemoembolization (TACE), systemic treatments with tyrosine kinase inhibitors like sorafenib and lenvatinib, and immunotherapy, such as the combination of atezolizumab (Tecentriq) and bevacizumab (Avastin) for first-line treatment [3–7]. However, these treatments are faced with numerous challenges due to the difficulties in diagnosis at an early-stage and chemotherapy resistance. Thus, developing new therapies and improving existing therapeutic strategies are crucial for improving patients’ outcomes.

G-quadruplexes (G4) intramolecular structures are present in a multitude of different mRNAs or DNAs and affect their functionality [8]. RNA G-quadruplex (rG4) is a stable structure composed of multiple guanines (Gs) interacting to form multiple planar arrangements [7, 9]. This stable secondary mRNA structure is connected by Hoogsteen hydrogen bonds [9]. The rG4 structure is enriched and functional in many mRNAs [10–12]. Generally, rG4 located in the 5’ UTR or 3’ UTR plays a significant role in biologic processes, such as mRNA stability, RNA localization, RNA processing, and protein translation [13–20]. Given the ubiquity of rG4 structures, they may exist in certain oncogenes and affect their translation. However, external factors such as environmental temperature and pH can disrupt the rG4 structure, further affecting protein expression [21–23]. Therefore, the stability of the rG4 structure is important for its function.

Carboxypyridostatin (cPDS) is a selective small-molecule ligand of rG4 structure exhibiting stabilizing effects on the rG4 conformation. cPDS has been reported to disrupt cell proliferation and promote cell cycle escape in nervous system [24, 25]. In addition, cPDS participates in cancer development via regulating the expression of proteins involved in cell proliferation. Baculoviral IAP Repeat Containing 3 (BIRC3) is an apoptosis regulatory factor and a member of the inhibitor of apoptosis proteins (IAP) family [26]. BIRC3 has been found in many cancers and can be utilized for developing anti-cancer drugs. Currently, there is evidence suggesting that BIRC3 has an anti-apoptotic effect in cancer cells [27]. Additionally, it plays a significant role in the nuclear factor-κB (NF-κB) signaling pathway, which is influential for many cancer cells [28–30].

In this study, we found that the exon region of BIRC3 contains rG4 structures. The addition of the rG4 stabilizer cPDS ensures more rG4 structures maintain intact, leading to a decreasement of BIRC3 protein expression in HCC cells. Interestingly, the mRNA level of BIRC3 was not affected. In a word, cPDS regulates BIRC3 expression at translational levels, thereby influencing the biologic properties of HCC cells by modulating cell apoptosis, which ultimately exert anti-cancer benefits.

Results

cPDS Supressed the Proliferation and Migration of HCC Cells

Considering that the location of G4 structures varies across different mRNAs. The rG4 structures located at different positions may have distinct functions. Therefore, we attempted to introduce the specific rG4 stabilizer, cPDS, into HCC cell lines to observe its impact on cell malignant behaviors. From colony formation and CCK8 assays, we found that cPDS inhibited the proliferation of MHCC97H and HEPa1-6 cells (Fig. 1A). Furthermore, cPDS also prevented the migration of these two cell lines, as indicated by scratch healing experiments (Fig. 1B). However, cPDS had no effect on the proliferation and migration of the HEP1 cell line. To further verify this finding, we conducted spheroid formation assays on the agarose gel in MHCC97H cells and Hepa1-6 cells and observed that the spheroid formation of cells treated with cPDS was restricted compared to the control group (Fig. 1C). Similarly, the spheroid capacity of HEP1 cells was not affected by cPDS. Therefore, cPDS can impede the proliferation, migration, and aggregation of HCC cells to in vitro in a concentration-dependent manner.

Fig. 1.

cPDS supressed the proliferation and migration of HCC cells. A Plate cloning assays were conducted using the MHCC97H, HEPa1-6, and HEP1 cell lines, with experimental groups treated with 10 μM and 20 μM concentrations of cPDS. CCK8 cell proliferation assays were performed on the three cell lines at 12-h, 24-h, and 48-h time points to assess cell viability. ****P < 0.0001. B Scratch assays were conducted on the three cell lines, with experimental groups treated with 10 μM and 20 μM concentrations of cPDS, and photographs were taken at 24 h for quantification. ****P < 0.0001. C Sphere formation assays were carried out on the three cell lines, with photographs taken on the 5th day

cPDS Regulated the Translational Efficiency of BIRC3 by Affecting rG4

To further explore the downstream molecules of cPDS that mediate the biologic behaviors of HCC cells, we performed RNA sequencing on MHCC97H cells treated with different concentrations of cPDS (0 μM, 10 μM, and 20 μM). From the volcano plot, we observed that the majority of genes were upregulated by cPDS at a concentration of 20 μM (Fig. 2A). We put together the top 10 genes that are upregulated and downregulated (Supporting Table 1). We sorted and prioritized the pathway enrichment analysis of upregulated genes and selected 23 pathways with the lowest false discovery rate (FDR). Based on these results, we identified six genes regulated by the TNF signaling pathway. Among these, BIRC3 drew our attention as it was one of the most significantly upregulated genes (Supporting Table 2), so we performed GO enrichment analysis of these genes and found that the functions of these genes are primarily related to cellular hypoxia stress, migration, and angiogenesis (Fig. 2D). The pathway enrichment analysis revealed that the TNF signaling pathway was most significantly influenced (Fig. 2B, C, E). Based on this, we identified six genes regulated by the TNF signaling pathway, and BIRC3 attracted our attention, because it was one of the most markedly upregulated genes. Previous study reported that BIRC3 can interact with the TNF receptor complex through its BIR domain, thereby influencing cellular apoptosis and survival signals. Our qRT-PCR results exhibited that cPDS upregulated BIRC3 mRNA levels in MHCC97H and HEPa1-6 cells (Fig. 2F and G). Intriguingly, the protein expression of BIRC3 showed a trend of downregulation by cPDS in the above two cell lines (Fig. 2H, I). This phenomenon may attributed to the rG4 structure in BIRC3 mRNA. Thus, under the influence of the rG4 structure, the translation efficiency of BIRC3 is inhibited.

Fig. 2.

cPDS regulated the translational efficiency of BIRC3 by affecting rG4. A Volcano plot analysis of differentially expressed genes in the experimental group compared to the control group. D Enrichment analysis of the functions of differentially expressed genes, listing the top 20 functions. B Pathway enrichment analysis of upregulated differential genes. C Pathway enrichment analysis of downregulated differential genes. E Overall pathway enrichment analysis of differential genes. F, G qRT-PCR analysis of the MHCC97H and HEPa1-6 cell lines treated with 20 μM concentration of cPDS. **P < 0.01; ***P < 0.001; ****P < 0.0001. H, I Western blot analysis of the two cell lines and the quantification of the blots. ****P < 0.0001

BIRC3 Accelerated the Proliferation of HCC Cells

Next, we performed a predictive analysis of the rG4 structure in the BIRC3 mRNA sequence and found that it has a typical rG4-forming sequence in the 5’ UTR region (Fig. 3A). Then, we constructed a schematic model of the rG4 structure (Fig. 3B). To explore the impact of BIRC3 on the biologic behavior of HCC cells, we constructed a BIRC3 overexpression plasmid and transfected it into the MHCC97H and Hepa1-6 cell lines. Western blot and qRT-PCR analyses confirmed the successful overexpression of BIRC3 in these two cell lines (Fig. 3C–F). The plate cloning and CCK8 experiment both demonstrated that BIRC3 significantly enhanced the proliferative capacity of these cells (Fig. 3G, H). EdU incorporation experiments further verified that cell proliferation was significantly promoted after overexpression of BIRC3 (Fig. 3I).

Fig. 3.

BIRC3 accelerated the proliferation of HCC cells A Gene sequence of the 5’ UTR of BIRC3. B Schematic illustration of the rG4 structure. C Western blot analysis and quantification of regular MHCC97H cells and those overexpressing BIRC3. ****P < 0.0001. D qRT-PCR analysis of regular MHCC97H cells and those overexpressing BIRC3. *****P< 0.0001. E Western blot analysis and quantification of regular HEPa1-6 cells and those overexpressing BIRC3. ****P < 0.0001. F Rt-qPCR analysis of regular HEPa1-6 cells and those overexpressing BIRC3. ****P < 0.0001. G Plate cloning assays for normal and BIRC3-overexpressing cells. H CCK8 cell proliferation assays for normal and BIRC3-overexpressing cells. ****P < 0.0001. I EDU chromogenic assay of MHCC97H and HEPa1-6 cells

BIRC3 Promotes Tumor Proliferation In Vivo

Due to the promotive effects of BIRC3 to cell proliferation in vitro, we constructed orthotopic HCC models to further explore the function of BIRC3 in HCC development by injecting Hepa1-6 cells transfected with BIRC3 overexpressed plasmid or empty vector control plasmid into mice livers. We discovered that BIRC3 enhanced intrahepatic tumorigenesis compared to control mice, as evidenced by IVIS bioluminescence images and tumor nodules of liver photos (Fig. 4A–C). We injected BIRC3-overexpressing MHCC97H cells and control cells subcutaneously into nude mice. Results showed that the volume and weight of tumors in nude mice injected with BIRC3 overexpression plasmid were notably enhanced compared to the control mice (Fig. 4D, E). Next, we performed immunohistochemistry staining on the removed tumors and discovered that the positive staining of BIRC3 and tumor proliferation markers (KI67 and PCNA) were uniformly intensified in BIRC3 overexpression tumors (Fig. 4F). Combined with our previous results (Fig. 1), we concluded that BIRC3 could accelerate tumor growth.

Fig. 4.

BIRC3 promotes tumor proliferation in vivo. A Representative images of tumor formation in liver tissues. B, C Representative live IVIS images of mice bearing HEPa1-6-Luc-BIRC3 tumors and HEPa1-6-Luc-control tumors. D Tumors harvested from nude mice after subcutaneous tumor formation. E Quantitative analysis of tumor diameters and tumor weights in the control group and the BIRC3-overexpressing group. ***P < 0.001, ****P < 0.0001. F Immunohistochemistry of tumor sections and their quantification. Antibodies used were against BIRC3, KI67, and PCNA. **P < 0.01

cPDS Induced Cell Apoptosis by Modulating Caspase-3

Previous research reported that BIRC3 is involved in the regulation of apoptosis by inhibiting the activity of caspase-3. We then explored the effects and regulatory mechanisms of cPDS on cell apoptosis by stimulating MHCC97H and HEPa1-6 cells with caspase-E3-IN-1, a specific inhibitor of caspase-3 before 20-μM cPDS treatment. We found that the addition of IN-1 had no effect on BIRC3 expression. What’s more, cPDS-induced upregulation of Bax and c-caspase-3 and downregulation of Bcl-2 were reversed by IN-1 addition (Fig. 5A, C). To further validate the effect of IN-1on BIRC3, we performed qRT-PCR on these two cell lines and found that the mRNA level of BIRC3 did not change by IN-1 (Fig. 5B, D). Next, we performed fluorescence labeling on these two cells double stained with Annexin V-FITC and PI and found the promoting effect of cPDS on apoptosis and the restoration of apoptosis after the addition of IN-1 (Fig. 5E). Furthermore, we performed flow cytometry assays and observed that the addition of cPDS enhanced cells apoptotic abilities, while this influence was alleviated after the IN-1 treatment (Fig. 5F). However, cell cycle distribution was not affected (Fig. 5G). In conclusion, we found that cPDS greatly reduced the expression of BIRC3 protein, which is responsible for inhibiting cleaved caspase-3 activity in regulating cell apoptosis, thereby exerting pro-apoptotic effect. And these regulatory effects could be restored by caspase-3 inhibitor IN-1.

Fig. 5.

cPDS induced cell apoptosis by modulating caspase-3. A Western blot analysis of MHCC97H cells treated differently. Quantitative analysis of BIRC3, c-caspase-3, and the ratio of BAX to BCL2 protein expression. ****P < 0.0001. B qRT-PCR analysis of control and treated MHCC97H cells. C Western blot analysis of HEPa1-6 cells treated differently. Quantitative analysis of BIRC3, c-caspase-3, and the ratio of BAX to BCL2 protein expression. ****P < 0.0001. D qRT-PCR analysis of control and treated HEPa1-6 cells. E Fluorescence staining of control and treated MHCC97H and Hepa1-6 cells. Green fluorescence represents Annexin V-FITC, and red fluorescence represents PI. F Flow cytometric analysis of controlled and treated MHCC97H and HEPa1-6 cells to determine apoptosis and cell cycle

Discussion

In this article, we reported the rG4 structure within the BIRC3 mRNA and explored its regulatory role in BIRC3 translation and HCC progression [31]. We found that with the addition of cPDS, a specific ligand of rG4, the protein level of BIRC3 was significantly inhibited, while its mRNA level did not decrease and even increased. This made us speculate that the inhibitory effect of the rG4 structure on the translation efficiency of BIRC3. Interestingly, cPDS restrained HCC cells growth both in vivo and in vitro and its pro-apoptotic effects might be associated with the downregulation of BIRC3 protein levels.

rG4 structures consist of guanine (G)-rich quartets stacked by planar guanine tetramers and their formation is prevalent observed in telomeres and the promoter regions of human oncogene [32, 33]. The stability and function of rG4 can be affected by various factors, including ion concentration, specific structural features of the RNA sequence, and interactions with proteins or small molecules [20, 34, 35]. rG4 is related to various biologic processes in the cell, such as DNA replication, mRNA splicing, transcriptional, and post-translational regulation [36]. In liver cancer cells, increased expression of BIRC3 is associated with tumor aggressiveness and poor prognosis [27, 37, 38]. rG4 structures located in the 5’ UTR of mRNA generally regulate gene translation efficiency [39]. BIRC3, also known as c-IAP2, is a member of the IAPs family [40]. In rheumatoid arthritis (RA), high expression of BIRC3 is associated with abnormal proliferation and anti-apoptotic ability of fibroblast-like synovial cells (FLS), which aggravated chronic inflammation and joint damage [41, 42]. Moreover, BIRC3 can interact with caspase-3, a key executor of apoptosis through its BIR3 domain, thereby inhibiting the activity of caspase-3 and thus inhibiting apoptosis [31, 43, 44]. The relationship between BIRC3 and caspase-3 activity is of great significance in tumor biology. High expression of BIRC3 exhibited pro-survival and anti-apoptotic characteristics in various tumors, such as Oral squamous cell carcinoma (OSCC), Chronic lymphocytic leukemia (CLL), and Brest cancer [28, 45–49].

Considering the role of TNF signaling pathway in regulating cell survival and apoptosis, our enrichment analysis pointed to the TNF signaling pathway drew our attention [50]. Our data indicate that BIRC3 is an identified component of this pathway and may be a key mediator of the effects of cPDS on HCC cells. However, the exact mechanism by which BIRC3 and the TNF pathway intersect to determine cell fate under the background of rG4 stabilization by ligands requires further research. Although we have provided valuable insights into the participation of cPDS and BIRC3 in HCC, there are many aspects that remain deeply digging. For example, the mechanism by which cPDS interacts with rG4 structures to regulate the expression and function of BIRC3 needs further study. Besides, we have not been able to directly explore the rG4 structure in BIRC3, as well as the impact of the environment, pH value, and temperature on the stability of rG4. In addition, more HCC cell lines and the animal models should be included to expand the generalizability of our findings. Future research should focus on the role of rG4 structures and its ligand in a broader range of cancer types and genetic backgrounds and investigate the potential of targeting rG4 structures and BIRC3 as a therapeutic strategy for HCC.

Surprisingly, our results showed that the impact of cPDS is varied in different HCC cell lines; the proliferation and migration of MHCC97H and HEPa1-6 cells are inhibited in response to cPDS stimulation, while HEP1 cells are relatively unaffected. These results suggested that the impact of cPDS may be highly dependent on the environment, possibly due to changes in the genomic landscape, including the presence, distribution, and accessibility of rG4 motifs in different cell lines. The cell line-specific response to cPDS emphasizes the complexity of rG4 biology and highlights the need for a comprehensive understanding of its role in cancer. Moreover, we speculate that the regulation of apoptosis by cPDS depends on the activity of caspase-3 regulated by BIRC3. In future, we will focus not only on the biologic functions of cPDS but also on exploring the specific mechanism between cPDS- and BIRC3-mediated an-apoptotic effects. What’s more, the expression of BIRC3 in different HCC cell lines is inconsistently affected by the rG4 ligand cPDS, the mechanism need further excavation.

In summary, our study revealed a complex regulatory network involving cPDS, BIRC3, and apoptotic pathways in HCC. These findings emphasize the potential of targeting rG4 structures and BIRC3 as new avenues for HCC treatment. Further research is needed to fully understand the molecular basis of these observations and to translate these insights into clinical applications.

Methods

Cell Culture

MHCC97H, HEP1, and HEPa1-6 cells were grown in Dulbecco’s Modified Eagle’s Medium supplemented with 10% Fetal Bovine Serum and 100-U/ml penicillin–streptomycin solution. These cells were kept at 37 degrees Celsius in a CO2 incubator with a 5% CO2 atmosphere.

Plate Cloning Assay

First, cells were digested with trypsin and resuspended with 10% complete culture medium to obtain a single-cell suspension. Then, cells were seeded in a 6-well plate at 500 cells per well. After cultured in a 37 °C incubator for 4 days, the complete culture medium were replaced and treated with 10-μM and 20-μM cPDS. On the 7th and 10th days, culture medium and cPDS were refreshed. On the 14th day, the cells were washed three times with PBS, fixed with 4% paraformaldehyde for 10 min, washed three times with PBS, then 0.5% crystal violet were added to each well, and then stained for 30 min. Finally, after washing off the crystal violet and drying the 6-well plate, cells were photographed under a microscope.

CCK8 Cell Proliferation Assay

Cells were seeded onto 96-well plate 5 × 10^{^3} cells per well. After 24-h incubation, 10-μL CCK8 solution were added to each well containing 100-μL complete culture medium. 2 h after reaction, the absorbance at 450 nm was measured under a microplate reader. The average absorbance values for each group were calculated and normalized to the control group or the 0-h time point. Relative cell viability was expressed as a percentage of the control group.

Cell Scratch Assay

The cells were cultured in a 6-well plate for 48 h until reached a confluence of 80%. Then, the culture medium were aspirated. After creating a wound in the well by drawing a straight line with a sterile pipette tip, the detached cells were washed with PBS. The 5% complete culture medium were added to the well and the plate was placed in the incubator to allow cells migrating into the wound area over 0–48 h. The images of the wound area were captured at 0 h, 24 h, and 48 h using a microscope equipped with a camera. Finally, the wound closure was measured by comparing images taken at different time points.

Cell Spheroid Formation Assay

First, 15-mg agarose were added to 1-ml DMEM medium and then heated in an 80 °C water bath for 30 min. After placing the vial in a sterilizer, it is sterilized at 115 °C for 30 min, taken out from the vial, then placed on a clean bench, and then added to 96-well plate at a volume of 60 μL per well. The 96-well plate was kept for 30 min to allow the agarose to solidify. Then, a single-cell suspension containing 1 × 10^{^4} cells per well in a 96-well plate was cultivated in an incubator for 1 h and then photographed using a light microscope. On the 3rd and the 5th day, half of the complete culture medium were refreshed. On the 5th day, spheroid formation was monitored using a light microscope and captured for documentation and analysis.

Western Blot

Collected cells were lysed in RIPA buffer to extract total protein. The lysates were centrifuged at 12,000 rpm for 10 min to remove debris and collect the supernatant. 5X Loading buffer were then added to the supernatant, and the mixed proteins were heated to denature. Proteins were subjected onto an SDS-PAGE gel and underwent electrophoresis for separation. The separated proteins were transferred from the gel to a PVDF membrane using a wet transfer system. The membrane were blocked with 5% BSA in Tris-buffered saline with Tween-20 (TBST) for 2 h to prevent non-specific binding. The membrane were incubated with the primary antibody diluted in TBST (dilution ratio 1:500) overnight at 4 °C with gentle shaking. After washing the membrane three times with TBST for 10 min each to remove unbound primary antibody, the membrane is incubated with the secondary antibody diluted in TBST (dilution ratio 1:1000) for 1 h at room temperature with gentle shaking. The membrane were washed three times with TBST for 10 min each. Finally, the target protein’s relative abundance was visualized under a chemiluminescence imaging system. GAPDH were used as an internal reference. Triplicates were performed for each sample. Primary antibodies used in this study included BIRC3 (Proteintech, 24,304–1-AP), BAX (Proteintech, 50,599–2-Ig), BCL2 (Proteintech, 12,789–1-AP), c-caspase-3 (Proteintech, 68,773–1-Ig), and GAPDH (Proteintech, 60,004–1-Ig).

qRT-PCR

Total RNA were extracted using Trizol according to the manufacturer’s instructions. 2 µg of total RNA were reverse-transcribed using a reverse transcription kit. Design primers involving BIRC3 and GAPDH (Table 1). PCR were performed under SYBR Green conditions: 95 °C for 2 min, followed by 40 cycles of 95 °C for 15 s, the corresponding annealing temperature for 15 s, and 72 °C for 30 s. GAPDH were used as an internal reference. Each sample were performed triplicates.

Table 1.

Primer sequences used for qRT-PCR

BIRC3	Forward: 5′-AAGCTACCTCTCAGCCTACTTT-3′
	Reverse: 5′-CCACTGTTTTCTGTACCCGGA-3′
GAPDH	Forward: 5ʹ-ACCCAGAAGACTGTGGATGG-3ʹ
	Reverse: 5ʹ-TTCTAGACGGCAGGTCAGGT-3ʹ

Plasmid Transfection

Plasmid DNA were purified using a standard plasmid purification kit according to the manufacturer’s instructions. Cells were seeded in a culture dish and at a density of 60–70% confluence on the day of transfection. Plasmid DNA were mixed with transfection reagent according to the manufacturer’s protocol. The DNA reagent mixture were added to the cells and then incubated for 4–6 h. The transfection medium was replaced with fresh complete culture medium and continued incubation for 24–48 h to allow stable integration and expression of the BIRC3 gene. The selective antibiotic were added into the culture medium at the final concentration recommended by the manufacturer. The selection medium was changed every 2–3 days to construct stably transfected cells and store aliquots in liquid nitrogen for future use.

5-Ethynyl-20-deoxyuridine (EdU) Incorporation Assay

Cells were seeded on glass coverslips in a 24-well plate. Then, EdU labeling and DNA staining procedures were conducted using BeyoClick™ EdU Cell Proliferation Kit with TMB (Beyotime Biotechnology, Shanghai). The number of Edu-positive cells were visualized under a fluorescent microscope.

Subcutaneous Tumor Formation

Cell suspension was prepared in sterile PBS at a concentration of 1 × 10^{^7} cells/mL. The cell suspensions were mixed with an equal volume of Matrigel on ice and 100-μL cell Matrigel mixtures were injected subcutaneously into the flank of nude mice using a 1-mL syringe. Mice were monitored daily for signs of distress or tumor-related morbidity and provided with appropriate care. Tumor size was measured with calipers every 2–3 days. The nude mice were euthanized according to ethical guidelines when the tumor reaches a predetermined size in the second week and then tumors were harvested for further analysis.

Immunohistochemistry Staining

The tissue sections were deparaffinized by immersing the slides in xylene for 10 min and rehydrated through a graded series of ethanol (100%–95%–85%–75%) for 5 min each, followed by a 5-min rinse in PBS. The slides were microwaved in antigen retrieval solution and cooled for 2 h. Endogenous peroxidase activity was blocked by incubating the slides in 35% H₂O₂ for 30 min. The slides were rinsed in PBS for 5 min and blocked with 5% BSA solution for 30 min to block non-specific binding sites. The slides were incubated using diluted primary antibody (dilution ratio 1:50) overnight at 4 °C in a humidified chamber. The slides were rinsed three times in PBS for 5 min, followed by incubation with the diluted secondary antibody (dilution ratio 1:500) for 1 h at room temperature in the dark. After rinsing the slides three times in PBS for 5 min, applying DAB chromogen solution to stand for color development for 2 min. The slides were counterstained with hematoxylin, dehydrated through a graded series of ethanol (75%–85%–95%–100%) for 5 min, and then cleared using xylene for 10 min. The slides were mounted with a coverslip and captured using a microscope. Primary antibodies used in this study included BIRC3 (Proteintech, 24,304–1-AP), KI67 (Proteintech, 27,309–1-AP), and PCNA (Proteintech, 10,205–2-AP).

Establishment of Orthotopic HCC Models

Hepa1-6 HCC cell lines with or without BIRC3 overexpression were co-transfected with firefly luciferase. Before intrahepatic implantation, 2 × 106 Hepa1-6 cell suspensions were collected. Then, the C57BL/6 mice aged 6 weeks were anesthetized and their livers were exposed by a median abdominal incision. Prepared cell suspensions were orthotopically injected into the right liver lobe. After closing the abdominal cavity, their vital signs were closely observed. 2 weeks later, VivoGlo luciferin was intraperitoneal injected into mice and then the tumor growth was observed under an in vivo imaging system (IVIS). Finally, intrahepatic tumors were photographed, measured, and then removed for further analyses.

Flow Cytometry

The prepared cells were digested with trypsin and washed with staining buffer to remove any culture medium or serum that may interfere with staining. The cells were centrifuged for 5 min, and the precipitations were gently resuspended in 195-μL Annexin V-FITC binding buffer and 5 μL Annexin V-FITC and gently mix. 10-μL propidium iodide staining solution were added and gently mixed again. The cells were incubated at room temperature in the dark for 10–20 min and then placed on ice. A flow cytometer were employed to analyze the cell apoptosis and cell cycle using appropriate gates and parameters for acquiring the data.

In Situ Fluorescence Detection

Cells were seeded onto a 24-well plate and cultured for 24 h until reaching 80% confluence. The cells were washed with PBS and then treated with 195-μL Annexin V-FITC binding buffer combined with 5-μL Annexin V-FITC. After gently mixing, 10-μL propidium iodide staining solution were added and gently mixed to incubate the cells at room temperature in the dark for 10–20 min. Finally, apoptotic cells with positive staining were observed under a fluorescence microscope.

Supplementary Information

Below is the link to the electronic supplementary material.

Author Contributions

Yiheng Liu contributed to conceptualization, investigation, and methodology. Qingqing Liu contributed to conceptualization and methodology. Tianyi Huang contributed to conceptualization and writing—original draft. Shengjie Zhang contributed to conceptualization. Zicheng Zhou contributed to data curation. Chiyu Gu contributed to investigation. Cui-Hua Lu contributed to conceptualization, funding acquisition, resources, and supervision. All authors approved the final version of the manuscript.

Funding

We greatly appreciate the financial support for this study provided by Nantong University. This study was supported by the National Natural Science Foundation of China (NO. 82070624) and Jiangsu Commission of Health (NO. ZDB202006).

Data Availability

Data are available upon reasonable request.

Declarations

Conflict of interest

The authors declare no conflicts of interest.