HCC cell lines, including HepG2 and Huh-7, were purchased from Beyotime Biotechnology Company (Shanghai, China). HepG2 cells are widely used in tumor biology research due to their stable proliferation but are non-permissive to hepatitis B virus (HBV) and hepatitis C virus (HCV) infection. Huh-7 cells are well-differentiated hepatocellular carcinoma cells and are permissive to HCV infection. Cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM) containing 10% fetal bovine serum (FuHeng, Shanghai, China). Cells were maintained in a humidified incubator under 5% CO₂ at 37°C.

HCC cells were seeded into 96-well plates at a final density of 3.0 × 10³ cells per well. Cells were incubated for 0, 24, 48, 72, or 96 hours. At each time point, 10 μl of CCK-8 reagent (Dojindo, Japan) was added to each well, followed by a 2-hour incubation at 37°C in the dark. A microplate reader (TECAN, Switzerland) was used to measure the OD value at 450 nm (32).

Cell proliferation was evaluated using the BeyoClick™ EdU-594 Cell Proliferation Kit (Beyotime Biotechnology, China). HCC cells were seeded into 24-well plates at a density of 5.0 × 10⁴ cells per well. After 24 hours, cells were incubated with EdU solution for 4 hours, then fixed with 4% paraformaldehyde and permeabilized using the supplied permeabilization buffer. Cells were subsequently stained with Alexa Fluor 594, and nuclei were counterstained with Hoechst 33342. Fluorescence images were acquired using a fluorescence microscope, and the percentage of EdU-positive cells was quantified using ImageJ software (NIH, Rockville, USA).

Cell invasion was assessed using a Transwell chamber assay. Briefly, Transwell inserts (Corning, USA) were pre-soaked at 37°C for 30 minutes and coated with Matrigel matrix diluted 1:8 in DMEM. For the invasion assay, 5 × 10⁴ cells suspended in 200 μL of serum-free DMEM were seeded into the upper chamber, while 500 μL of DMEM containing 20% fetal bovine serum was added to the lower chamber as a chemoattractant. After 24 hours of incubation at 37°C, non-invading cells on the upper surface were removed, and the invaded cells on the lower membrane were fixed and stained with 0.1% crystal violet. Images were acquired using an inverted microscope. For the migration assay, Transwell inserts were used without Matrigel coating. Migratory cells were stained with Calcein-AM (Beyotime, China) and visualized under a fluorescence microscope.

HCC cells were seeded in 6-well plates and allowed to grow to 90 – 100% confluence. A linear scratch was created using a sterile 200 μL pipette tip. The wells were washed gently with sterile PBS to remove debris and non-adherent cells. Wound closure was monitored by capturing images at 0 and 20 hours using an optical microscope. The migration rate was quantified by measuring the gap distance using ImageJ software.

Total protein was extracted using RIPA lysis buffer and quantified by a BCA assay. Equal amounts of protein were separated via SDS-PAGE and transferred onto nitrocellulose membranes. After blocking with 5% non-fat milk for 1 hour at room temperature, membranes were incubated overnight at 4°C with primary antibodies, including anti-FBXO2 (14590-1-AP, Proteintech), anti-USP49 (18066-1-AP, Proteintech), anti-Myc (16286-1-AP, Proteintech), anti-GAPDH (GB15004-100, Servicebio), anti-Flag (F1804, Sigma), and anti-ubiquitin (sc-166553, Santa Cruz Biotechnology). The membranes were then washed and incubated with appropriate HRP-conjugated secondary antibodies. Signals were detected using Omni-ECL™ chemiluminescent substrate (YEASEN, Shanghai, China) (33).

Cell lysates were prepared from HepG2 or Huh-7 cells using NP-40 RIPA buffer. Co-immunoprecipitation was performed by incubating 5 μg of the indicated antibody with cell lysates overnight at 4°C, followed by the addition of protein A/G magnetic beads (B23202, Selleck). After incubation, the beads were washed thoroughly, and bound proteins were eluted by boiling in loading buffer. The eluates were then subjected to SDS-PAGE and Western blot analysis (34).

For in vivo ubiquitination assays, cells with or without plasmid transfection were treated with 20 μM MG-132 (a proteasome inhibitor) for 6 hours. Cells were then lysed, and ubiquitinated proteins were detected by immunoprecipitation followed by Western blotting using the appropriate antibodies. For CHX chase experiments, HepG2 and Huh-7 cells were treated with 100 μg/mL cycloheximide (CHX) to inhibit protein synthesis. Cells were harvested at 0, 2, 4, 6, and 8 hours post-treatment. Protein lysates were prepared using RIPA buffer and analyzed by SDS-PAGE followed by immunoblotting to evaluate protein stability over time.

HepG2-shNC and HepG2-shFBXO2 cells (5 × 10⁶ in 100 μL PBS) were subcutaneously injected into BALB/c nude mice (n = 10). Once tumor volumes reached approximately 100 mm³, mice were randomly assigned into two groups and treated with either 20 mg/kg sorafenib or vehicle control via daily intraperitoneal injection for 20 days. Tumor volumes were measured every four days and calculated using the formula: V = (L × W²) × 0.52, where L is the longest diameter and W is the shortest diameter of the tumor. At the end of the treatment period, tumors were excised and weighed for further analysis.

All statistical analyses were performed using GraphPad Prism 8.0 or SPSS Statistics software. Data are presented as the mean ± standard deviation (SD). Comparisons between two groups were made using Student’s t-test, while one-way ANOVA was used for comparisons among multiple groups. A p-value < 0.05 was considered statistically significant.

To investigate the expression pattern of FBXO2 in HCC, we analyzed publicly available transcriptomic data from The Cancer Genome Atlas (TCGA). The results revealed that FBXO2 expression was significantly upregulated in tumor tissues compared to adjacent normal liver tissues. To further explore the clinical relevance of FBXO2, we utilized the UALCAN database to assess its association with various clinicopathological features of HCC. Elevated FBXO2 expression was found to be significantly correlated with several clinical parameters, including TP53 mutation status, patient gender, race, and tumor stage ( Figure 1A ). In addition, immunohistochemistry (IHC) analysis confirmed that FBXO2 protein levels were markedly elevated in HCC tissues relative to adjacent normal liver tissues ( Figures 1B–C ). Furthermore, FBXO2 expression is associated with patient survival in HCC, as analyzed using the KMplot database (https://kmplot.com/analysis). These findings suggest that FBXO2 overexpression may play a role in HCC progression and could serve as a potential prognostic biomarker.

To investigate the biological function of FBXO2 in HCC, we silenced its expression by transfecting HepG2 and Huh-7 cells with lentivirus encoding FBXO2-specific shRNA ( Figure 2A ). Knockdown of FBXO2 significantly reduced the proliferative capacity of HCC cells, as demonstrated by both CCK-8 and EdU assays ( Figures 2B–D ). To assess the impact on cell motility, we performed Transwell and wound healing assays. FBXO2 depletion markedly inhibited the invasion and migration of HepG2 and Huh-7 cells compared to control groups ( Figures 2E–G ). These results collectively indicate that FBXO2 knockdown significantly suppresses HCC cell proliferation and motility.

To further validate the role of FBXO2, we generated FBXO2-overexpressing cell models by transfecting HepG2 and Huh-7 cells with a Flag-FBXO2 lentiviral construct ( Figure 3A ). CCK-8 and EdU assays revealed that FBXO2 overexpression significantly enhanced the proliferation of both cell lines ( Figures 3B–D ). In Transwell invasion assays, FBXO2-overexpressing HCC cells exhibited a significantly greater invasive ability compared to wild-type controls ( Figures 3E–F ). Furthermore, both Transwell and wound healing assays confirmed that FBXO2 overexpression significantly increased the migratory capacity of HCC cells ( Figures 3G–I ). Collectively, these results demonstrate that FBXO2 functions as a positive regulator of HCC cell proliferation, invasion, and migration.

To identify potential substrates of Fbxo2, we performed liquid chromatography-tandem mass spectrometry (LC-MS/MS). Following transient transfection of Flag-tagged Fbxo2 into HEK293T cells, Fbxo2-associated protein complexes were immunoprecipitated and subjected to LC-MS/MS analysis, which revealed USP49-derived peptides as candidate substrates. To determine whether USP49 is a substrate of FBXO2, we performed co-immunoprecipitation (co-IP) assays in HepG2 and Huh-7 cells. The results confirmed both endogenous and exogenous interactions between FBXO2 and USP49 ( Figures 4A–B ). Given this interaction, we next examined how FBXO2 expression affects USP49 protein levels. Western blot analysis revealed that FBXO2 knockdown significantly increased USP49 protein levels, while FBXO2 overexpression led to a marked decrease ( Figures 4C, D ). Furthermore, the reduction in USP49 levels caused by FBXO2 overexpression was reversed by treatment with MG132, a proteasome inhibitor, indicating that FBXO2 mediates USP49 degradation through the ubiquitin-proteasome pathway ( Figure 4E ). To assess the effect of FBXO2 on USP49 protein stability, we performed a CHX chase assay. The results showed that FBXO2 depletion markedly delayed the degradation of USP49 ( Figures 4F–G ), further supporting the role of FBXO2 in promoting USP49 proteasomal degradation.

To further investigate the mechanism by which FBXO2 regulates USP49, we performed in vivo ubiquitination assays in HepG2 and Huh-7 cells. Overexpression of FBXO2 significantly enhanced the ubiquitination of USP49 ( Figure 5A ). To determine which domain of FBXO2 is responsible for USP49 recognition, we conducted co-IP experiments with FBXO2 deletion mutants. The results indicated that the F-box-associated (FBA) domain of FBXO2 is required for its interaction with USP49 ( Figures 5B–C ). Consistently, deletion of the FBA domain abolished FBXO2-mediated ubiquitination of USP49, as confirmed by in vivo ubiquitination assays ( Figure 5D ). Since ubiquitination typically occurs on lysine residues, we used the GPS-Uber prediction tool to identify potential ubiquitination sites on USP49. Six lysine-to-arginine mutants were constructed (K93R, K106R, K161R, K241R, K348R, and K353R). Among these, the K353R mutant exhibited markedly reduced ubiquitination compared to the wild-type and other mutants, suggesting that lysine 353 is a critical site for FBXO2-mediated ubiquitination ( Figure 5E ). Collectively, these results demonstrate that FBXO2 binds to USP49 via its FBA domain and promotes its degradation by enhancing ubiquitination at lysine 353 in HCC cells.

To determine whether FBXO2 regulates HCC cell proliferation and motility through USP49, we performed co-transfection experiments in HepG2 and Huh-7 cells using shFBXO2 and shUSP49 ( Figure 6A ). CCK-8, EdU, and Transwell assays revealed that FBXO2 knockdown significantly suppressed cell proliferation, migration, and invasion. Notably, these inhibitory effects were reversed upon USP49 silencing, indicating that USP49 mediates the tumor-suppressive effects of FBXO2 depletion ( Figures 6B–H ). Together, these results demonstrate that FBXO2 promotes HCC cell proliferation and motility, at least in part, by modulating USP49 stability and activity.

Beyond its role in tumor progression, we investigated whether FBXO2 contributes to sorafenib resistance in HCC. Given that sorafenib is a frontline multikinase inhibitor for advanced HCC, we examined whether FBXO2 knockdown affects sorafenib sensitivity in HCC cells. FBXO2 was silenced in HepG2 and Huh-7 cells, and their response to sorafenib was evaluated ( Figure 7A ). CCK-8 assay results showed that FBXO2-depleted cells exhibited significantly increased sensitivity to sorafenib compared to control cells. Consistent with in vitro findings, in vivo experiments using HepG2 xenograft models in nude mice demonstrated that FBXO2 knockdown significantly enhanced the antitumor efficacy of sorafenib. Tumor volumes and weights were markedly reduced in the FBXO2-depleted group treated with sorafenib compared to controls ( Figures 7B–E ). Collectively, these findings suggest that targeting FBXO2 may sensitize HCC cells to sorafenib and improve therapeutic outcomes in both in vitro and in vivo settings.