1 Introduction
Hepatocellular carcinoma (HCC) ranks among the most prevalent malignancies of the digestive system worldwide. Characterized by insidious onset, high rates of recurrence and metastasis, HCC significantly impacts patient prognosis. Currently, it stands as the third leading cause of cancer-related mortality globally [
1]. Although significant advancements have been made in HCC treatment, a considerable proportion of patients are diagnosed at an advanced stage during their initial presentation due to the lack of prominent clinical symptoms. Furthermore, the propensity for early metastasis and rapid recurrence contributes to the dismal prognosis associated with this disease [
2,
3]. To date, the precise pathogenesis of HCC remains incompletely elucidated. Existing research indicates that factors such as alcohol consumption, non-alcoholic fatty liver disease (NAFLD), viral hepatitis, consumption of mold-contaminated food, and genetic predisposition can all contribute to HCC development [
4]. Therefore, elucidating the molecular mechanisms underlying HCC pathogenesis and identifying effective therapeutic targets are of paramount importance for improving patient outcomes.
P53 and DNA damage-regulated gene 1 (PDRG1), located on chromosome 20q13.12, encodes a small protein of 133 amino acids (13.6 kDa). Luo
et al. first identified in 2003 that PDRG1 expression is regulated by ultraviolet (UV) radiation and the tumor suppressor p53 [
5]. Furthermore, PDRG1 is implicated in various cellular physiologic processes, including cell proliferation, growth, apoptosis, cell cycle regulation, and DNA damage repair [
6]. Recent studies have revealed that PDRG1 expression is significantly elevated in multiple tumor types, including colorectal, ovarian, lung, gastric, breast, and uterine cancers, compared to adjacent normal tissues. Consequently, PDRG1 has been proposed as a novel tumor biomarker. Mechanistically, Xu
et al. demonstrated that PDRG1 participates in the malignant progression of colorectal cancer by modulating the p21-mediated signaling pathway [
7]. Zhang
et al. reported that PDRG1 silencing suppresses the growth and metastasis of gastric cancer cells via activation of the ataxia–telangiectasia mutated (ATM)/p53 pathway [
8]. Additionally, PDRG1 silencing has been shown to inhibit the Wnt signaling pathway in esophageal cancer cells [
9]. However, the expression profile of PDRG1 in HCC and its precise mechanistic role in regulating HCC cell behavior remain incompletely understood.
The Wnt/β-catenin pathway, a canonical Wnt signaling cascade, regulates diverse biological processes including proliferation, apoptosis, migration, and invasion [
10]. Recent research indicates that aberrant Wnt/β-catenin signaling is prevalent in numerous malignancies and exerts significant clinical impacts on tumor biology, patient prognosis, and therapeutic response [
11,
12]. In the absence of Wnt ligand activation, the destruction complex—comprising Axin, adenomatous polyposis coli (APC), and GSK3β proteins—binds and phosphorylates β-catenin, targeting it for ubiquitin-mediated degradation and preventing its cytoplasmic accumulation under physiologic conditions [
13]. Upon Wnt pathway activation, binding of Wnt ligands to frizzled receptors (FZDs) on the cell membrane activates dishevelled (Dvl) protein. This inhibits the destruction complex, allowing non-phosphorylated β-catenin to accumulate in the cytoplasm. Subsequently, non-phosphorylated β-catenin translocates to the nucleus, where it associates with transcription factors of the T cell factor/lymphoid enhancer factor (TCF/LEF) family, initiating transcription of Wnt-regulated target genes such as c-Myc [
14,
15].
Specificity protein 1 (SP1), a sequence-specific DNA binding protein, has been documented in early studies to exhibit dysregulated expression and activation in tumor tissues [
16]. As a prominent member of the transcription factor family, SP1 activates the transcription of numerous cellular genes harboring GC-rich binding sites within their promoters [
17]. During tumor progression across various cancers, SP1 can transactivate numerous oncogenes while simultaneously suppressing certain tumor suppressor genes [
18]. Multiple studies demonstrate the involvement of SP1 in regulating tumor cell proliferation, invasion, angiogenesis, and other biological functions. Furthermore, its expression level in tumor tissues correlates with patient prognosis and treatment outcomes [
19,
20].
In the present study, we demonstrate that PDRG1 is overexpressed in HCC tissues, correlating with poor prognosis in HCC patients. Inhibition of PDRG1 suppresses tumor formation, proliferation, and invasion by modulating the Wnt/β-catenin pathway. Furthermore, we identify PDRG1 as a direct downstream transcriptional target of SP1. Critically, knockdown of PDRG1 reverses the tumor-promoting functions induced by SP1 overexpression. Collectively, our findings establish that the SP1/PDRG1/Wnt/β-catenin axis regulates HCC tumorigenesis and progression. These results highlight PDRG1 as a potential therapeutic target for developing novel treatment strategies against HCC.
2 Materials and methods
2.1 Public data analysis
The expression levels of PDRG1 and SP1 in HCC versus normal liver tissues were analyzed using the UALCAN web portal and the Gene Expression Omnibus (GEO). UALCAN employs Level 3 standardized data from The Cancer Genome Atlas (TCGA) and Clinical Proteomic Tumor Analysis Consortium (CPTAC) databases. For CPTAC, protein expression Z-values represent standard deviations from the median across samples, based on log2 spectral count ratios normalized within and across samples. Correlations between PDRG1 expression and tumor stage/grade in TCGA were evaluated using Welch’s t-test. Survival analysis was performed using the GepLiver platform with the TCGA-LIHC cohort. Kaplan–Meier curves were generated and compared by the log-rank test, with patients stratified according to the median expression level of PDRG1. This analytical approach aligns with standard bioinformatic methodologies for cancer prognosis studies using TCGA data, with a significance threshold of P < 0.05. The GEPIA2 portal was used to analyze correlations between PDRG1 and selected genes in the TCGA-LIHC data set. Expression values were log2-transformed, and Pearson correlation analysis was applied, reporting the correlation coefficient (R) and P-value. A significance threshold of P < 0.05 was applied to all analyses. Detailed information on all data sets used in this study is provided in Table S1.
2.2 Gene set enrichment analysis (GSEA)
GSEA was performed using GSEA software (version 3.0). Samples were stratified into PDRG1 high-expression (≥ 50%) and low-expression (< 50%) groups based on PDRG1 expression levels. The h.all.v7.4.symbols.gmt gene set was acquired from the Molecular Signatures Database for pathway enrichment analysis. The analysis parameters were set as follows: minimum gene set size = 5, maximum gene set size = 5000, with 1000 permutations. A nominal P-value < 0.05 was considered statistically significant.
2.3 Protein–protein interaction (PPI) network analysis
The PPI network of PDRG1 was constructed using the STRING database (version 12.0). The search was performed with PDRG1 as the query protein in Homo sapiens, requiring an interaction score > 0.7 (high confidence). The resulting network included both known and predicted interactions, covering physical associations and functional linkages. The network graph was exported and further visualized using Cytoscape software (version 3.9.1).
2.4 Clinical specimens
Sixty-six pairs of tumor tissues and matched adjacent non-tumorous tissues (collected 2 cm from the tumor margin) were obtained from patients undergoing hepatectomy at the Department of Hepatobiliary and Pancreatic Surgery, The Second Affiliated Hospital of Harbin Medical University (Harbin, China) between May 2024 and June 2025. All diagnoses were pathologically confirmed postoperatively. Inclusion criteria were: (1) Child-Pugh class A or B; (2) No prior antitumor therapy; (3) R0 radical resection; (4) Postoperative pathological diagnosis of HCC; (5) Availability of microvascular invasion (MVI) description in the pathology report. Exclusion criteria included: (1) Recurrent HCC; (2) Extrahepatic metastasis; (3) Previous history of malignancy; (4) Presence of portal/hepatic vein tumor thrombus; (5) History of immunomodulatory drug use. All specimens were immediately snap-frozen in liquid nitrogen and stored at 80 °C. Informed consent was obtained from all participants, and the study protocol was approved by the Institutional Review Board (IRB) of The Second Affiliated Hospital of Harbin Medical University (Approval No. YSJKY2024-235).
2.5 Immunohistochemical (IHC) analysis
IHC staining for PDRG1 protein expression was performed on a subset of 30 HCC cases randomly selected from the 66 cases collected between May 2024 and June 2025 at The Second Affiliated Hospital of Harbin Medical University. IHC staining intensity was evaluated using a semiquantitative scoring system based on the percentage of strongly stained cells within the observed field. The association between PDRG1 expression levels in HCC tumors and adjacent tissues was determined using the paired samples t-test.
2.6 Cell culture
The HCC cell lines HCCLM3, Huh7, MHCC97-H, and Hep3B were obtained from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China). The PLC/PRF/5 cell line was purchased from the American Type Culture Collection (ATCC). Cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM) (Gibco) supplemented with 10% fetal bovine serum (FBS) (Abcells) and 1% Penicillin-Streptomycin Solution (Seven). All cells were maintained in a humidified incubator at 37 °C with 5% CO2. The Wnt/β-catenin pathway-specific inhibitor XAV939 was purchased from MedChemExpress (USA) and used to treat relevant HCC cells at a concentration of 3.5 µmol/L.
2.7 Plasmids, lentivirus, and siRNA transfection
Lentiviruses encoding short hairpin RNAs (shRNAs) targeting PDRG1 (shPDRG1-1) or a non-targeting control (shNT) sequence were purchased from Shanghai Nobio Biotechnology Co., Ltd. (China). Lentiviruses co-expressing luciferase (Luc) with either OE-PDRG1 or empty vector (Vector) were obtained from Wuhan GeneCreate Biological Engineering Co., Ltd. (China). Small interfering RNAs (siRNAs) targeting PDRG1 (si-PDRG1-1 and si-PDRG1-2), and plasmids containing the empty vector, full-length PDRG1 cDNA (OE-PDRG1), or full-length SP1 cDNA (OE-SP1) were purchased from Wuhan GeneCreate Biological Engineering Co., Ltd. (China). For lentiviral infection, HCC cells were incubated overnight with virus-containing supernatant supplemented with 8 μg/mL Polybrene®. Plasmid and siRNA transfection was performed using the Lipofectamine™ 3000 Transfection Reagent (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer’s instructions; complexes were formed by incubating the nucleic acid with the reagent for 20 min before addition to the cell culture medium. Cells were subjected to gain-of-function or loss-of-function experiments 72 h post-infection/transfection.
2.8 Cell counting kit-8 (CCK-8) assay
HCC cells seeded in 96-well plates were incubated with 10 μL of CCK-8 solution (Sevenbio, Beijing, China) per well for 2 h at 37 °C. The optical density (OD) at 450 nm was measured using a Synergy HTX Multi-Mode Microplate Reader (BioTek Instruments, Inc., Winooski, VT, USA).
2.9 Colony formation assay
HCC cells were plated in 6-well plates at a density of 800 cells per well and cultured for 14 days under standard conditions (37 °C, 5% CO2). Resulting colonies were fixed with 4% paraformaldehyde for 15 min, washed twice with phosphate-buffered saline (PBS), and stained with 1 mL of 0.1% crystal violet solution (Beyotime, Shanghai, China) for 20 min. After washing and air-drying, colonies (defined as clusters > 50 cells) were counted manually or using ImageJ software (National Institutes of Health, Bethesda, MD, USA).
2.10 Wound healing assay
HCC cells were seeded in 6-well plates and cultured to form a confluent monolayer. A straight scratch wound was created across the monolayer using a sterile 200-μL pipette tip, guided by a straightedge. The wells were gently washed three times with PBS to remove detached cells and debris, and serum-free DMEM was added. Initial images (0 h) of the wound were captured using phase-contrast microscopy. After 48 h of incubation, the wells were washed gently with PBS, and final images were captured. Wound closure percentage was quantified by comparing the remaining wound area at 48 h to the initial wound area at 0 h using ImageJ software.
2.11 Transwell migration assay
Cell migration was assessed using Transwell chambers (8.0-μm pore size, Corning Inc., Corning, NY, USA). Briefly, 5 × 104 HCC cells resuspended in 200 μL of serum-free DMEM were seeded into the upper chamber. The lower chamber was filled with 600 μL of DMEM containing 10% FBS as a chemoattractant. After incubation for 48 h at 37 °C with 5% CO2, non-migrated cells on the upper surface of the membrane were gently removed with a cotton swab. Cells that had migrated to the lower surface were fixed with 4% paraformaldehyde for 15–20 min, stained with 0.1% crystal violet solution for 20–40 min (with intermittent PBS washes), and air-dried. Migrated cells were photographed using an inverted microscope and quantified by counting cells in three random fields per chamber.
2.12 Transwell invasion assay
Transwell invasion assays were performed using chambers pre-coated with Matrigel® Matrix (Corning Inc., Corning, NY, USA). Briefly, Matrigel® was thawed at 4 °C, diluted 1:3 with ice-cold serum-free DMEM, and 100 µL of the mixture was added to the upper chamber of a Transwell insert (8.0-µm pore size) placed in a 24-well plate, ensuring even coverage of the membrane. The plate was incubated at 37 °C for 4 h to allow gel polymerization. Excess liquid was then carefully aspirated. 1 × 105 HCC cells suspended in 200 µL of serum-free DMEM were seeded onto the solidified Matrigel® layer in the upper chamber. The lower chamber was filled with 600 µL of DMEM supplemented with 10% FBS as a chemoattractant. After 48 h of incubation at 37 °C with 5% CO2, non-invading cells on the upper surface of the membrane were removed with a cotton swab. Cells that had invaded through the Matrigel® and membrane to the lower surface were fixed with 4% paraformaldehyde for 15–20 min, stained with 0.1% crystal violet solution for 40 min (with three washes in PBS during staining), and air-dried. Invaded cells were photographed using an inverted microscope and quantified by counting cells in three randomly selected fields per membrane.
2.13 Reverse transcription-quantitative polymerase chain reaction (RT-qPCR)
Total RNA was extracted from HCC cell lines and clinical tissue specimens (paired tumor and adjacent normal tissues) using the Sevenfast® Total RNA Extraction Kit (Sevenbio, Beijing, China). For tissue samples, approximately 30 mg of each specimen was homogenized prior to RNA extraction. RNA concentration and purity were verified spectrophotometrically. Then, 1000 ng of total RNA was reverse transcribed into cDNA using the All-in-one First Strand cDNA Synthesis Kit II (with dsDNase) (Sevenbio). Quantitative PCR was performed with 2× SYBR Green qPCR MasterMixII (Universal) (Sevenbio) on a QuantStudio™ Real-Time PCR System (Thermo Fisher Scientific). Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as the internal control, and relative expression levels were calculated by the 2−ΔΔCt method. All human tissues were obtained with informed consent, and the study was approved by the Institutional Review Board of The Second Affiliated Hospital of Harbin Medical University (Approval No. YSJKY2024-235). The primer sequences used were:
PDRG1: Forward 5′-ATTGTGGACCTGGACACTAAAAGG-3′, Reverse 5′-TGTCTCAGGGTGAGGCATCTTG-3′
SP1: Forward 5′-CACCAGAATAAGAAGGGAGG-3′, Reverse 5′-GGTGGTAATAAGGGCTGAA-3′
GAPDH: Forward 5′-GGTGTGAACCATGAGAAGTATGA-3′, Reverse 5′-GAGTCCTTCCACGATACCAAAG-3′
PDRG1 promoter E1 region: Forward 5′-CGCCTTTTACAGCCATGTGG-3′, Reverse 5′-AACAGCCAGCGTATGTGGAA-3′
PDRG1 promoter E2 region: Forward 5′-TAGTGTTAGAGCCTGCGCAC-3′, Reverse 5′-GATAGCATAGCGCCCACCAA-3′
PDRG1 promoter E3 region: Forward 5′-TGTGTTCACTGCTGCATCCT-3′, Reverse 5′-CTGCCTCTGACCCGTTACTC-3′
2.14 Western blot
Total protein was extracted from cells or tissues using RIPA lysis buffer supplemented with protease and phosphatase inhibitors (Beyotime, Shanghai, China). Protein concentration was determined using the BCA Protein Assay Kit (Beyotime). Equal amounts of protein (30 µg) were separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred onto PVDF membranes. Membranes were blocked with 5% non-fat milk in Tris-buffered saline containing 0.1% Tween-20 (TBST) for 1 h at room temperature and then incubated with primary antibodies diluted in blocking buffer overnight at 4 °C. After washing three times with TBST, membranes were incubated with appropriate horseradish peroxidase (HRP)-conjugated secondary antibodies for 50 min at room temperature. Following three additional TBST washes, protein bands were visualized using enhanced chemiluminescence (ECL) substrate (Beyotime) and imaged using a Bio-Rad ChemiDoc Touch Imaging System (Bio-Rad Laboratories, Hercules, CA, USA). Band intensity was quantified using ImageJ software (National Institutes of Health).
2.15 Co-immunoprecipitation (Co-IP) assay
PPI between RUVBL1 and PDRG1 was analyzed by co-immunoprecipitation using the Pierce™ Magnetic Direct Co-IP Kit (Thermo Fisher Scientific, Cat# 88828). Total proteins were extracted from Hep3B and PLC/PRF/5 cells using IP Lysis/Wash Buffer with protease inhibitors. Cell lysates were diluted to 2 mg/mL with the same buffer. For each reaction, 5 µg of antibody—anti-RUVBL1 (Proteintech, Cat# 10210-2-AP), anti-PDRG1 (Santa Cruz Biotechnology, Cat# sc-398815), or normal rabbit IgG (Proteintech, Cat# 30000-0-AP, as a negative control)—was cross-linked to magnetic beads. Then, 500 µL of diluted lysate was added to the antibody-conjugated beads and incubated overnight at 4 °C. After washing, bound proteins were eluted and analyzed by Western blot.
2.16 Nuclear protein extraction
Nuclear and cytoplasmic fractions were isolated using the Nuclear and Cytoplasmic Protein Extraction Kit (Beyotime, Shanghai, China) strictly following the manufacturer’s instructions. Histone H3 (H3) was used as a nuclear fraction loading control.
2.17 Chromatin immunoprecipitation (ChIP)
ChIP assays were performed using the SimpleChIP® Enzymatic Chromatin IP Kit (Cell Signaling Technology, Danvers, MA, USA; #9005S). Briefly, cells were cross-linked with 37% formaldehyde for 10 min at room temperature, and the reaction was quenched by adding 125 mmol/L glycine for 5 min. Cells were washed with ice-cold PBS, harvested, and lysed. Chromatin was digested with Micrococcal Nuclease to obtain DNA fragments predominantly between 150 bp and 900 bp. Chromatin was immunoprecipitated overnight at 4 °C with rotation using anti-SP1 antibody (CST #9389S) or normal rabbit IgG as a negative control. Protein G Magnetic Beads were added for 2 h to capture the antibody-chromatin complexes. Complexes were washed sequentially with low salt, high salt, LiCl, and TE buffers. Cross-links were reversed by incubation with 200 mmol/L NaCl at 65 °C for 30 min, followed by proteinase K digestion. Immunoprecipitated DNA was purified using spin columns. Enrichment of specific DNA fragments was analyzed by qPCR using primers targeting the predicted binding sites (E1, E2, E3) within the PDRG1 promoter region (sequences listed above).
2.18 Luciferase reporter assay
The pmirGLO vector containing the PDRG1 promoter region was purchased from Wuhan GeneCreate Biological Engineering Co., Ltd. (China). HCC cells were co-transfected with the PDRG1-promoter reporter construct and either OE-SP1 plasmid or empty vector control using Lipofectamine™ 3000. Renilla luciferase plasmid (pRL-TK) was co-transfected as an internal control for normalization. Forty-eight hours post-transfection, luciferase activity was measured using the Dual-Luciferase® Reporter Assay System (Vazyme Biotech Co., Ltd., Nanjing, China) according to the manufacturer's instructions. Firefly luciferase activity was normalized to Renilla luciferase activity for each sample.
2.19 In vivo tumorigenesis and metastasis assay
All procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of The Second Affiliated Hospital of Harbin Medical University (Approval No. YSJKY2024-235).
2.20 Subcutaneous tumor model
Six-week-old male BALB/c nude mice (SPF conditions) received subcutaneous injections of Hep3B cells stably transduced with lentivirus encoding shNT or shPDRG1-1. Cells (1 × 106) were resuspended in 100 µL of sterile PBS and injected into the left flank of each mouse (n = 6 per group). Tumor volume was measured every three days using the formula: Volume = 0.5 × length × width2. Mice were euthanized at four weeks, and tumors were collected for analysis.
2.21 Lung metastasis model
Hep3B cells stably transduced with lentivirus co-expressing luciferase (Luc) and OE-PDRG1 or empty vector (Vector) were injected via tail vein (1 × 106 cells in 100 µL PBS) into male BALB/c nude mice. Mice were divided into four groups (n = 6/group): Vector + PBS, Vector + XAV939, OE-PDRG1 + PBS, and OE-PDRG1 + XAV939. XAV939 was prepared from a 100 mmol/L DMSO stock solution and diluted to a final concentration of 10 mg/mL (DMSO ≤ 0.1%). Mice in the XAV939 groups received intraperitoneal injections (10 mg/kg) every other day for 6 weeks, while control groups received PBS. At six weeks post-injection, all mice underwent bioluminescence imaging using an IVIS Lumina XRMS system (PerkinElmer) and were subsequently euthanized for lung tissue collection and further analysis.
2.22 Statistical analysis
All statistical analyses and graphing were performed using GraphPad Prism software (version 9.5.1) and SPSS Statistics (version 22.0). Quantitative data are presented as the mean ± standard deviation (SD). Comparisons between two groups were performed using Student’s t-test. Comparisons among multiple groups were analyzed using one-way analysis of variance (ANOVA) followed by Tukey’s post-hoc test. The association between PDRG1 expression and clinicopathological parameters was assessed using the Chi-square test or Fisher’s exact test, as appropriate. All cell-based experiments were independently repeated at least three times. A P < 0.05 was considered statistically significant.
3 Results
3.1 Elevated PDRG1 predicts poor prognosis in HCC
Analysis of The Cancer Genome Atlas (TCGA) and Gene Expression Omnibus (GEO) data sets using the UALCAN platform revealed that PDRG1 mRNA expression was significantly higher in HCC tissues compared to normal liver tissues (P < 0.0001, Fig. 1A and 1B). Further analysis of the relationship between PDRG1 expression and specific etiological factors showed that among alcohol consumption, non-alcoholic fatty liver disease (NAFLD), hepatitis B, and hepatitis C, only NAFLD demonstrated a significant inverse association with PDRG1 expression (P < 0.05), while the other three factors showed no statistically significant association (Fig. S1A). Furthermore, data from the Clinical Proteomic Tumor Analysis Consortium (CPTAC) demonstrated upregulation of PDRG1 protein in HCC tissues versus normal liver controls (P < 0.0001, Fig. 1C). Validation using our clinical specimens (n = 66 pairs) via Western blot, immunohistochemistry (IHC), and RT-qPCR consistently confirmed that PDRG1 expression was significantly elevated in HCC tissues at both protein and mRNA levels compared to matched adjacent non-tumorous tissues (P < 0.0001, Fig. 1D–1F).
PDRG1 mRNA expression increased progressively with higher tumor grade, a trend that was statistically significant. However, such a consistent increase was not observed across pathological tumor stages (stage I–IV) (P < 0.05, Fig. 2A and 2B). Analysis of our cohort, in which patients were stratified into high- and low-expression groups based on the median PDRG1 protein level, further established significant associations between elevated PDRG1 expression and key clinicopathological parameters, including tumor number, capsular invasion, tumor diameter, and MVI (P < 0.05, Table 1). The relationship between PDRG1 expression levels and these statistically significant parameters was further visualized using box plots with overlaid scatter points. (P < 0.05, Fig. S1B). Critically, analysis of TCGA data via the GepLiver platform demonstrated that HCC patients with high PDRG1 mRNA expression had significantly worse survival outcomes. This included reduced overall survival (OS), disease-specific survival (DSS), disease-free survival (DFS), and progression-free survival (PFS) (P < 0.05 for all, Fig. 2C–2F). Collectively, these results indicate that PDRG1 is a potential biomarker predictive of poor prognosis in HCC.
3.2 PDRG1 facilitates HCC cell proliferation, migration, and invasion
Analysis of five HCC cell lines (Huh7, HCCLM3, Hep3B, MHCC97-H, and PLC/PRF/5) via Western blot and RT-qPCR revealed significantly higher levels of both PDRG1 protein and mRNA in Hep3B and PLC/PRF/5 cells, while HCCLM3 cells exhibited the lowest expression (Fig. 3A and 3B). Transfection of Hep3B and PLC/PRF/5 cells with two distinct PDRG1-targeting siRNAs (si-PDRG1-1 and si-PDRG1-2) significantly downregulated PDRG1 expression at both mRNA and protein levels (P < 0.0001, Fig. 3C–3F). Conversely, overexpression of PDRG1 (OE-PDRG1) in HCCLM3 cells markedly elevated PDRG1 mRNA and protein levels (P < 0.001, Fig. 3G and 3H).
Functional assays demonstrated that PDRG1 knockdown significantly inhibited HCC cell proliferation, as evidenced by CCK-8 and colony formation assays (P < 0.05, Fig. 3I–3M). Overexpression of PDRG1 significantly enhanced proliferation (Fig. 3I–3M). Furthermore, PDRG1 knockdown significantly attenuated HCC cell migration and invasion in wound healing and Transwell assays, while PDRG1 overexpression markedly promoted both processes (P < 0.01, Fig. 3N and 3O). Collectively, these data establish PDRG1 as an oncogene in HCC.
3.3 PDRG1 regulates the Wnt/β-Catenin pathway
Analysis of 374 HCC patients from the TCGA database, stratified by median PDRG1 expression into high- and low-expression groups, identified differentially expressed genes (DEGs) (Fig. 4A and 4B). KEGG pathway analysis showed significant enrichment of DEGs in Spliceosome, RNA transport, and Ubiquitin-mediated proteolysis (Fig. 4C). Gene Ontology (GO) enrichment analysis of DEGs revealed significant enrichment in: biological process (BP): RNA splicing and mRNA metabolic process (Fig. 4D); cellular component (CC): nucleoplasm and nuclear part (Fig. 4E); molecular function (MF): RNA binding, nucleic acid binding, and chromatin binding (Fig. 4F).
GSEA indicated that PDRG1 expression was significantly positively correlated with the Wnt/β-catenin and p53 signaling pathways, and negatively correlated with bile acid metabolism in HCC (Fig. 5A). To further substantiate these findings, we constructed a PPI network for PDRG1, which identified RUVBL1, PFDN6, URI1, and UXT as its direct interacting partners (Fig. 5B). Correlation analysis using the GEPIA2 platform in the TCGA-LIHC cohort revealed that among these candidates, RUVBL1 exhibited the strongest positive correlation with PDRG1 mRNA expression (Fig. 5C; Fig. S2A–S2C). Co-immunoprecipitation (Co-IP) assays in Hep3B and PLC/PRF/5 cells confirmed a physical interaction between PDRG1 and RUVBL1 proteins (Fig. 5D). Critically, RUVBL1 has been established as a key regulator that activates the Wnt/β-catenin pathway through chromatin remodeling [
21]. Recent evidence demonstrates that RUVBL1-containing complexes can drive the expression of β-catenin and facilitate its nuclear translocation [
22]. Therefore, the specific interaction between PDRG1 and RUVBL1 suggests a plausible mechanism whereby PDRG1 may modulate Wnt/β-catenin signaling by engaging this central chromatin regulator. Based on this integrated evidence from GSEA, PPI analysis, and the established role of its binding partner, we focused subsequent investigations on the Wnt/β-catenin pathway.
Western blot analysis confirmed that PDRG1 knockdown for 72 h in Hep3B and PLC/PRF/5 cells significantly decreased the levels of β-catenin, c-Myc, phosphorylated GSK-3β (p-GSK-3β) in the cytoplasm, and nuclear β-catenin (Fig. 5E and 5F). Conversely, PDRG1 overexpression in HCCLM3 cells significantly increased these key Wnt pathway components in both compartments (Fig. 5E and 5F). Collectively, these data, combined with the observed physical interaction between PDRG1 and RUVBL1 and the strong correlation between PDRG1 expression and Wnt pathway activity, establish PDRG1 as a functional regulator of the Wnt/β-catenin signaling pathway.
3.4 XAV939 reverses PDRG1-mediated HCC progression through Wnt/β-catenin pathway inhibition
To determine whether PDRG1 exerts its oncogenic effects through the Wnt/β-catenin pathway, we performed rescue experiments using the Wnt/β-catenin pathway-specific inhibitor XAV939 (3.5 µmol/L). Overexpression of PDRG1 significantly enhanced HCCLM3 cell proliferation; however, co-treatment with XAV939 completely reversed this pro-proliferative effect (Fig. 6A, 6B, and 6G). Similarly, the enhanced migration and invasion induced by PDRG1 overexpression were also effectively abolished by XAV939 treatment (Fig. 6C–6F).
Extending these observations to in vivo settings, we first confirmed in the subcutaneous tumor model that PDRG1 knockdown significantly suppressed Hep3B-derived tumor growth (Fig. 6H). Prior to the lung metastasis experiments, Hep3B cells stably overexpressing PDRG1 were established and validated by both Western blot and RT-qPCR, confirming significantly elevated PDRG1 levels at both protein and mRNA levels (P < 0.001) (Fig. 6I). Furthermore, in the lung metastasis model, tail vein injection of OE-PDRG1 Hep3B cells markedly enhanced metastatic burden, which was again substantially attenuated by XAV939 treatment (Fig. 6J).
Collectively, these consistent results from both in vitro and in vivo experiments demonstrate that PDRG1 promotes HCC proliferation and metastasis primarily via activation of the Wnt/β-catenin pathway.
3.5 SP1 transcriptionally activates PDRG1 by binding its promoter
To identify upstream transcriptional regulators of PDRG1, we intersected predictions from the GTRD, TFDB, PROMO, and KnockTF databases, yielding four candidate transcription factors: CEBPB, SP1, TP53, and MYB. Based on literature review and clinicopathological correlation, SP1 was prioritized for further investigation (Fig. 7A). Analysis of TCGA data via GEPIA2 confirmed a significant positive correlation between SP1 and PDRG1 mRNA expression in HCC tissues (R = 0.38, P = 2.9 × 10−14; Fig. 7B). Furthermore, data from TCGA and CPTAC demonstrated significant upregulation of both SP1 mRNA and SP1 protein in HCC tissues compared to adjacent non-tumorous controls (Fig. 7C).
JASPAR database analysis predicted three potential SP1 binding sites within the PDRG1 promoter region: E1 (−813 to −803), E2 (−1908 to −1898), and E3 (−1927 to −1917) (Fig. 7D). Chromatin Immunoprecipitation qPCR (ChIP-qPCR) in HCC cells confirmed that SP1 specifically bound only to the predicted E3 site (Fig. 7E), validated further by agarose gel electrophoresis (Fig. 7F). Dual-luciferase reporter assays corroborated that SP1 bound to the E3 site, significantly enhancing promoter activity (P < 0.0001, Fig. 7G and 7H). Critically, overexpression of SP1 (OE-SP1) in HCCLM3 cells significantly increased both PDRG1 mRNA and protein levels (P < 0.001, Fig. 7I and 7J). These findings establish SP1 as a direct transcriptional activator of PDRG1.
3.6 SP1/PDRG1/Wnt/β-catenin axis promotes HCC malignancy
To investigate whether SP1 regulates the Wnt/β-catenin pathway via PDRG1, we performed rescue experiments assessing Wnt pathway proteins. Overexpression of SP1 increased PDRG1 expression and activated Wnt/β-catenin signaling (elevated β-catenin, c-Myc, p-GSK-3β). Remarkably, this activation was reversed by PDRG1 knockdown (si-PDRG1-1) (Fig. 8A and 8B), demonstrating that SP1 modulates Wnt/β-catenin signaling through PDRG1. Importantly, SP1 overexpression significantly enhanced HCC cell proliferation, migration, and invasion. Co-treatment with si-PDRG1-1 effectively reversed these SP1-induced oncogenic phenotypes (Fig. 8C–8I).
Complementary rescue experiments were performed to further validate this regulatory hierarchy. SP1 knockdown suppressed both PDRG1 expression and Wnt/β-catenin signaling activation (Fig. 9A and 9B), whereas concurrent PDRG1 overexpression effectively restored pathway activity (Fig. 9C–9I). These results conclusively demonstrate that the SP1/PDRG1/Wnt/β-catenin signaling axis promotes the proliferation, migration, and invasion of HCC cells.
4 Discussion
PDRG1 has been implicated in the pathogenesis of diverse malignancies, including testicular, liver, cervical, lung, ovarian, and gastric cancers [
23]. Since its initial discovery in 2003 as a gene regulated by UV radiation and p53 [
5], subsequent research has focused on elucidating its molecular mechanisms and associated signaling pathways in both neoplastic and non-neoplastic tissues [
24–
26]. In this study,
in silico analyses revealed significantly elevated PDRG1 mRNA and protein expression in HCC tissues compared to adjacent non-tumorous liver. Critically, high PDRG1 mRNA expression correlated positively with advanced tumor stage and grade, and negatively with OS, DSS, DFS, and PFS in HCC patients. These findings were validated experimentally: Western blot and RT-qPCR analyses of 66 paired HCC and adjacent normal tissues confirmed PDRG1 overexpression in tumors. IHC analysis on a subset of 30 paired samples further corroborated this, and subgroup analysis established significant associations between elevated PDRG1 protein expression and adverse clinicopathological parameters, including increased tumor number, capsular invasion, larger tumor diameter, higher incidence of MVI, and elevated aspartate aminotransferase (AST) levels. Notably, PDRG1 expression showed no significant correlation with gender, age, portal vein tumor thrombus, or liver cirrhosis. Collectively, these results provide compelling evidence that PDRG1 is overexpressed in HCC and associated with poor prognosis. This prompted our investigation into the biological functions and molecular mechanisms of PDRG1 in HCC pathogenesis.
Recent studies support PDRG1’s oncogenic role: it is overexpressed in colorectal cancer and promotes malignant progression via the p21-mediated signaling pathway [
7]. Similarly, PDRG1 has been linked to Wnt signaling activation in esophageal cancer [
9]. While our study confirms PDRG1 overexpression in human HCC, its specific biological functions and mechanisms in this context remain incompletely defined. To address this, we established PDRG1 gain-of-function (OE-PDRG1) and loss-of-function (si-PDRG1 and si-PDRG1-2) HCC cell models. Our functional assays demonstrated that PDRG1 knockdown significantly suppressed HCC cell proliferation, migration, and invasion. To investigate the underlying mechanism, we hypothesized that PDRG1 might regulate the Wnt/β-catenin pathway. This hypothesis was based on two key lines of predictive evidence. First, our PPI analysis revealed that PDRG1 physically binds to RUVBL1, a known chromatin remodeling ATPase. Importantly, RUVBL1 has been established as a direct activator of the Wnt/β-catenin pathway in cancer. Critically, RUVBL1 is a key regulator that activates the Wnt/β-catenin pathway through chromatin remodeling [
21], and recent evidence demonstrates that RUVBL1-containing complexes can drive the expression of β-catenin and facilitate its nuclear translocation [
22]. Second, GSEA indicated a significant positive correlation between PDRG1 expression and Wnt/β-catenin signaling activity in HCC. To experimentally test this hypothesis, we examined the core components of the Wnt pathway. Mechanistically, and consistent with our prediction, PDRG1 depletion downregulated key Wnt/β-catenin pathway components, including p-GSK-3β and β-catenin. Conversely, PDRG1 overexpression enhanced these malignant phenotypes and activated Wnt signaling by upregulating the same components. Crucially, rescue experiments using the specific Wnt/β-catenin pathway inhibitor XAV939 reversed the pro-proliferative and pro-metastatic effects induced by PDRG1 overexpression in both
in vitro and
in vivo settings. These data provide robust evidence that PDRG1 acts as an oncogene in HCC, primarily mediated through Wnt/β-catenin signaling, and identify it as a potential therapeutic target.
While previous studies described PDRG1’s association with cancer and its downstream signaling [
27,
28], the upstream transcriptional regulators of PDRG1 remained unexplored. Our study is the first to report that the transcription factor SP1 binds to the PDRG1 promoter and directly activates its transcription. SP1 is a well-established transcription factor implicated in diverse cancers, regulating the transcription of numerous genes critical for tumorigenesis [
29–
31]. For instance, SP1 promotes lung cancer growth by transcriptionally activating MALAT1 [
32] and facilitates gastric cancer progression by enhancing PD-L1 transcription [
33]. Our preliminary findings indicated SP1-mediated upregulation of PDRG1 expression. Dual-luciferase reporter assays identified the specific binding site E3 (−1927 to −1917) within the PDRG1 promoter responsible for SP1 binding. Furthermore, rescue experiments demonstrated that the SP1/PDRG1/Wnt/β-catenin signaling axis regulates the proliferation, migration, and invasion of HCC cells. These novel discoveries highlight the SP1/PDRG1 axis as a promising therapeutic target for HCC.
Our study has several limitations. First, the detailed molecular interplay between SP1 and PDRG1 in HCC, particularly concerning potential epigenetic modifications (e.g., histone acetylation/methylation), warrants further investigation. Second, due to technical constraints, we were unable to perform double immunofluorescence staining to precisely co-localize SP1 and PDRG1 expression within tumor tissues. Finally, in vivo rescue experiments validating whether SP1 promotes HCC proliferation, metastasis, and invasion specifically through PDRG1 are necessary to fully establish the translational relevance of this axis.
In summary, our study identifies PDRG1 as an oncogenic driver in HCC, where its overexpression correlates with advanced tumor progression and poor prognosis. Mechanistically, PDRG1 activates Wnt/β-catenin signaling to promote HCC proliferation, migration, and invasion. We further demonstrate that SP1 transcriptionally upregulates PDRG1 by binding to its promoter, forming a novel SP1/PDRG1/Wnt/β-catenin axis that drives HCC malignancy. Rescue experiments confirm the therapeutic potential of targeting this pathway. These findings establish the SP1-PDRG1-Wnt/β-catenin axis as a promising therapeutic target for the treatment of HCC.
4.0.0.0.1 Acknowledgements
This work was supported by National Natural Science Foundation of China (Nos. 82472743, 82300921, and 82573164); Development of a Precision Management System for Biliary Tract Tumors (No. SC2024ZX12C0024); Heilongjiang Provincial Natural Science Foundation of China (No. PL2024H115); Guangxi Science and Technology Program (No. AD25069077); Natural Science Foundation of Heilongjiang Province (No. LH2023H043); Open Funds of State Key Laboratory of Oncology in South China (No. HN2025-02); Open Funds of Shanghai Key Laboratory of Cancer Systems Regulation and Clinical Translation (No. CSRCT-KF0002); Open Research Project from Key Laboratory of Clinical Laboratory Medicine of Guangxi Department of Education (No. GXGXLCJYZDX2025001); Guangxi Science and Technology Program (No. AD25069077); Open Research Fund of Key Laboratory of Gastrointestinal Cancer (Fujian Medical University), Ministry of Education (No. FMUGIC-202501); Open Funds of Key Laboratory of Clinical Laboratory Technology for Precision Medicine (No. FKLCLT-202504); Open Research Fund of Anhui Province Key Laboratory of Non-coding RNA Basic and Clinical Transformation (No. NcRNA202508); Thematic Research Support Scheme of State Key Laboratory of Liver Research, The University of Hong Kong (No. SKLLR/TRSS/2025/08).
4.0.0.0.2 Compliance with ethics guidelines
Conflicts of interest Xudong Zhao, Shihua Lv, Haikuan Wang, Zeyi Lu, Pengcheng Kang, Jinglin Li, Liang Yu, Shihui Ma, Changxing Hua, Junqi You, Ziqiang Ge, Yi Xu, and Yunfu Cui declare that they have no conflict of interest.
This study was approved by the Ethics Committee of the Second Affiliated Hospital of Harbin Medical University (Approval No. YSJKY2024-235) and was performed in accordance with the ethical standards as laid down in the 1964 Declaration of Helsinki and its later amendments or comparable ethical standards. Written informed consent was obtained from all participants, and all animal procedures followed the institutional and national guidelines for the care and use of laboratory animals.
The Author(s). This article is published with open access at link.springer.com and journal.hep.com.cn
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