Non‐Telomeric Role of RAP1 in Facilitating NF‐ κB Activation and Driving Hepatocellular Carcinoma Progression

ABSTRACT

RAP1 (TERF2IP) is a component of the shelterin complex that protects telomeric DNA and preserves chromosome stability. In addition to its telomeric function, accumulating evidence indicates that mammalian RAP1 also exerts multiple extra‐telomeric functions. Notably, RAP1 has been reported to regulate the NF‐κB signaling pathway and to function as a transcriptional regulator, suggesting potential roles in tumorigenesis. In this study, we found that RAP1 expression was significantly upregulated in hepatocellular carcinoma (HCC) cells. Functional analyses demonstrated that RAP1 promoted malignant phenotypes in HCC through a non‐telomeric mechanism. In cellular models, RAP1 overexpression enhanced cell proliferation while suppressing senescence and apoptosis, whereas RAP1 knockdown produced the opposite effects. Mechanistically, RAP1 functioned as an upstream activator of the NF‐κB signaling cascade, resulting in increased phosphorylation of the p65 subunit and upregulation of downstream targets, including IL‐1β and BCL‐2. Importantly, the oncogenic activity of RAP1 was shown to be dependent on NF‐κB signaling in vivo, as pharmacological inhibition of NF‐κB significantly suppressed RAP1‐driven tumor growth in a xenograft model. Collectively, these findings reveal a previously unrecognized role for RAP1 in promoting HCC progression through activation of NF‐κB signaling and identify the RAP1/NF‐κB axis as a potential therapeutic target for HCC.

1. Introduction

Hepatocellular carcinoma (HCC), the most common form of primary liver malignancy, represents a major global health burden and poses a significant challenge to healthcare systems worldwide [1]. Despite advances in diagnostic strategies and the development of novel therapeutic options, HCC remains the third leading cause of cancer‐related mortality worldwide, with a 5‐year survival rate of only 18% [2, 3]. This poor prognosis is largely attributable to the unique pathogenic context of HCC, in which more than 90% of cases develop in a background of chronic liver inflammation, such as chronic hepatitis infection or liver cirrhosis, forming a vicious cycle of inflammation, tissue damage, compensatory regeneration, and ultimately carcinogenesis. Furthermore, HCC cells possess strong intrinsic capabilities for rapid proliferation, evasion of both senescence and apoptosis, and progressive acquisition of resistance to chemotherapy, radiotherapy, and targeted therapies. These biological features collectively constitute the fundamental drivers of malignant progression [4, 5]. Consequently, identifying the critical molecular mechanisms that connect chronic inflammation, dysregulated cell fate decisions, and therapeutic resistance is essential for overcoming the current therapeutic limitations in HCC management.

The nuclear factor kappa‐light‐chain‐enhancer of activated B cells (NF‐κB) signaling pathway represents a central molecular node that links chronic inflammation to malignant transformation. As a transcription factor that is constitutively activated in many human solid tumors, including HCC, NF‐κB regulates the expression of more than 400 genes involved in cell survival, proliferation, inflammation, and immune regulation [6, 7, 8]. Under basal conditions, NF‐κB remains inactive in the cytoplasm through its association with inhibitory IκB proteins. Upon stimulation by various signals, including inflammatory cytokines, oxidative stress, and DNA damage, IκB proteins are phosphorylated and degraded, allowing NF‐κB to translocate into the nucleus. Once activated, NF‐κB initiates transcriptional programs that promote cell proliferation through targets such as cyclin D1 and c‐Myc, inhibit apoptosis through genes including BCL‐2, BCL‐XL, and XIAP, and enhance inflammatory responses through mediators such as IL‐6, TNF‐α, and COX‐2 [9, 10]. In addition to these canonical functions, NF‐κB has been shown to regulate telomerase activity and modulate the senescence‐associated secretory phenotype (SASP), thereby functioning as a key integrator of cellular senescence, survival signaling, and tumor microenvironment remodeling [11, 12, 13]. In particular, NF‐κB activation can stimulate transcription of telomerase reverse transcriptase (TERT), the catalytic subunit of telomerase, thereby enhancing telomerase activity. Through maintenance of telomere length and prevention of telomere attrition in malignant cells, the NF‐κB/TERT axis provides an additional mechanism through which cancer cells acquire limitless replicative potential and escape normal cellular senescence or apoptosis [14, 15, 16]. Consequently, through its combined regulation of telomerase‐dependent replicative immortality and SASP‐mediated paracrine signaling, NF‐κB acts as a critical regulator of the balance between senescence and survival in cancer cells.

Given the central importance of NF‐κB signaling in the malignant progression of HCC, identifying upstream regulators of this pathway has become an important research focus. RAP1, also known as TERF2IP, is one of the most evolutionarily conserved components of the shelterin complex and plays an essential role in telomere protection. In recent years, however, RAP1 has been increasingly recognized for its diverse extra‐telomeric functions. In addition to maintaining telomere integrity, RAP1 participates in the regulation of DNA damage responses, cellular metabolism, and inflammatory signaling pathways, including the NF‐κB pathway [17, 18, 19]. Mechanistically, RAP1 functions as an adaptor protein that facilitates IKK‐mediated phosphorylation of the NF‐κB p65 subunit at Ser536, thereby enhancing NF‐κB transcriptional activity and promoting the expression of anti‐apoptotic genes such as BCL‐2. This RAP1/NF‐κB signaling axis has been implicated in promoting tumor cell survival, invasion, and chemoresistance in several malignancies, including breast cancer and non‐small cell lung cancer [6].

Interestingly, the role of RAP1 in cancer appears to be highly context dependent. Although RAP1 is frequently overexpressed and exhibits oncogenic properties in several solid tumors [20, 21, 22, 23], reduced RAP1 expression has been reported in familial papillary thyroid cancer [24], and its role in hematological malignancies remains controversial [25, 26]. These findings suggest that the biological functions of RAP1 may vary substantially across different tumor types and cellular contexts. In the case of HCC, however, the expression pattern and functional role of RAP1 remain incompletely understood. In particular, the mechanistic relationship between RAP1 and NF‐κB signaling in HCC has not been fully elucidated. Moreover, it remains unclear whether RAP1, potentially through modulation of NF‐κB activity, regulates the critical balance between proliferation, senescence, and apoptosis that determines tumor progression and therapeutic responsiveness in HCC.

In the present study, we demonstrate that RAP1 expression is markedly elevated in HCC cells and promotes tumor progression by enhancing cellular proliferation while suppressing senescence and apoptosis. Mechanistic investigations further reveal that these effects are mediated through activation of the NF‐κB signaling pathway. Importantly, the tumor‐promoting activity of RAP1 was shown to be dependent on NF‐κB signaling in vivo, as pharmacological inhibition of NF‐κB significantly attenuated RAP1‐driven tumor growth. These findings identify the RAP1/NF‐κB axis as a key regulatory pathway in HCC progression and highlight its potential as a therapeutic target for HCC treatment.

2. Materials and Methods

2.1. HCC Cell Lines

The immortalized human benign hepatocyte cell line HL‐7702 and four human HCC cell lines, Hep3B, Huh‐7, HepG2, and QGY‐7703, were obtained from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China), and the identities of all cell lines were authenticated by the supplier. HL‐7702, HepG2, and Huh‐7 cells were cultured in DMEM (Solarbio, Beijing, China); Hep3B cells were maintained in MEM (Solarbio); and QGY‐7703 and SMMC‐7721 cells were maintained in RPMI‐1640 medium (Solarbio). All culture media were supplemented with 10% fetal bovine serum (FBS; Gibco, Grand Island, NY, USA). Cells were maintained at 37°C in a humidified incubator with 5% CO₂.

2.2. RNA Isolation and Quantitative Reverse Transcription PCR (qRT‐PCR)

Total RNA was extracted from HCC cell lines using TRIzol reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer's instructions. Extracted RNA was reverse transcribed into cDNA using the PrimeScript RT reagent kit (TaKaRa, Dalian, China). Quantitative real‐time PCR (qRT‐PCR) was performed using TB Green Premix Ex Taq II (TaKaRa) on a LightCycler 96 real‐time PCR system (Roche, Basel, Switzerland). Relative mRNA expression levels were calculated using the 2^−ΔΔCt method with an internal control gene. The primer sequences used for human genes in this study are provided in Table S1.

2.3. Artificial Alteration of RAP1 Expression in HCC Cell Lines

RAP1 overexpression was achieved using the eukaryotic expression vector pcDNA3.1(+) (Thermo Fisher Scientific, Waltham, MA, USA). The coding sequence of human RAP1 was synthesized and cloned into the multiple cloning site downstream of the CMV promoter. Small interfering RNA (siRNA) targeting human RAP1 and a negative control siRNA (siR‐NC) were purchased from GenePharma (Suzhou, China). The sequences of all siRNA oligonucleotides used in this study are listed in Table S2.

Transfection of plasmids or siRNAs was performed using Lipofectamine 2000 (Thermo Fisher Scientific) according to the manufacturer's protocol. Total RNA was extracted 48 h after transfection and subjected to qRT‐PCR analysis to verify gene expression changes.

2.4. Cell Counting Kit‐8 (CCK‐8) Cell Viability Assay

HCC cells were initially seeded in 24‐well plates at densities of 8 × 10⁴ cells per well for HepG2 cells or 5 × 10⁴ cells per well for Hep3B cells, followed by transfection the next day. After 24 h of transfection, cells were dissociated and reseeded into 96‐well plates at densities of 1 × 10⁴ cells per well for HepG2 cells or 5 × 10³ cells per well for Hep3B cells, with three replicates per group. Cell viability was assessed using the CCK‐8 reagent (Dojindo, Tokyo, Japan) according to the manufacturer's instructions. Absorbance at 450 nm (A₄₅₀) was measured using an EnSpire Multimode Plate Reader (PerkinElmer, Waltham, MA, USA). All experiments were performed independently at least three times.

2.5. 5‐Ethynyl‐2′‐Deoxyuridine (EdU) Cell Proliferation Assay

EdU is a thymidine analog that is incorporated into genomic DNA during the S phase of the cell cycle, allowing measurement of DNA synthesis and cellular proliferation. EdU staining was performed using the Cell‐Light EdU Apollo488 In Vitro Kit (RiboBio, Guangzhou, China). HCC cells were seeded and treated under the same conditions as those used in the CCK‐8 assay. Cells grown in 24‐well plates were stained according to the manufacturer's instructions and visualized using an IX71 fluorescence microscope (Olympus). Fluorescence intensity and EdU‐positive cells were quantified using ImageJ software.

2.6. β‐Galactosidase Staining

Cellular senescence was evaluated using a Senescence β‐Galactosidase Staining Kit (Beyotime, Shanghai, China) according to the manufacturer's protocol. Briefly, cells were washed once with PBS and fixed in 3% formaldehyde for 15 min at room temperature, followed by three washes with PBS. Cells were then incubated overnight at 37°C in a dry incubator without CO₂. Senescent cells were identified by the development of blue staining under light microscopy. At least 300 cells were counted from randomly selected microscopic fields for each experimental condition.

2.7. TUNEL Assay

Cells cultured on glass coverslips were fixed with 4% paraformaldehyde and permeabilized using 1% Triton X‐100. DNA strand breaks were detected using a TUNEL assay kit (KeyGEN BioTECH, KGA7051) according to the manufacturer's instructions. Cell nuclei were counterstained with DAPI. Fluorescent images were obtained using a fluorescence microscope (Nikon Eclipse Ni‐U, Japan). For each experimental group, apoptotic cells and total cells were counted in ten randomly selected microscopic fields at a magnification of ×20.

2.8. Western Blot

For Western blot analysis, two types of cell lysates were prepared: whole‐cell lysates and subcellular fractionation (nuclear/cytoplasmic) lysates. Whole‐cell lysates were generated using RIPA lysis buffer (Thermo Fisher Scientific) supplemented with protease and phosphatase inhibitors (Invitrogen); fractionated cell lysates were prepared using NE‐PER Nuclear and Cytoplasmic Extraction Reagents (Thermo Fisher Scientific, #78833), following the manufacturer's protocol. Briefly, cells were harvested, washed with PBS, and resuspended in ice‐cold CER I (200 μL) containing 1× protease and phosphatase inhibitors. After vortexing and ice incubation for 10 min, ice‐cold CER II (11 μL) was added, vortexed, and incubated on ice for 1 min. The lysate was centrifuged at 16,000 × g for 5 min at 4°C, and the supernatant was kept as the cytoplasmic fraction. The nuclear pellet was washed with PBS, then resuspended in ice‐cold NER (100 μL) containing 1× protease and phosphatase inhibitors, vortexed, and incubated on ice for 40 min with intermittent mixing. After centrifugation at 16,000 × g for 10 min at 4°C, the supernatant was collected as the nuclear fraction. Both fractions were stored at −80°C.

Protein concentrations were determined using a BCA Protein Assay Kit (Solarbio). Equal amounts of protein (60 μg) were separated on 12% SDS‐PAGE gels and transferred onto nitrocellulose membranes (Boster, Wuhan, China). Membranes were blocked with 5% skim milk for 60 min at room temperature and subsequently incubated overnight at 4°C with primary antibodies against cleaved caspase‐3 (C‐Cas3) (Huaan, Hangzhou, China, #ET‐1602‐47), NF‐κB p65 (p65) (Huaan, #HA721307), phospho‐NF‐κB p65 (p‐p65) (Cell Signaling Technology, Danvers, MA, USA, #3033), RAP1 (Abcam, Cambridge, MA, USA, #ab14404), p16 (Cell Signaling Technology, #92803), p21 (Cell Signaling Technology, #2947), BCL‐2 (Cell Signaling Technology, #3498), Histone H3 (Cell Signaling Technology, #4499) and GAPDH (Signalway Antibody, College Park, MD, USA, #21612). After washing three times with TBST (15 min each), the membranes were incubated for 1 h at room temperature with HRP‐conjugated secondary antibody (either goat anti‐rabbit IgG, Abbkine, Wuhan, China, #A21010, or goat anti‐mouse IgG, Abbkine, #A21020) as appropriate for the primary antibody host species. Immunoreactive bands were visualized using ECL and quantified with ImageJ software.

2.9. Animals and Tumor Model Establishment

Specific pathogen‐free BALB/c nude mice aged 6–8 weeks and weighing 15–20 g were purchased from Beijing Vital River Laboratories (China). All animal experiments were conducted in strict accordance with the International Ethics Guidelines and the National Institutes of Health Guidelines for the Care and Use of Laboratory Animals. Mice were housed in an air‐conditioned facility maintained at 23°C ± 2°C under a 12 h light/dark cycle, with free access to food and water. Animals were acclimated to these conditions for at least 7 days before the initiation of experiments.

HepG2 cells stably transfected with either an empty vector (pcDNA3) or a RAP1‐overexpressing plasmid (pcDNA3‐RAP1) were collected during the logarithmic growth phase and resuspended in PBS at a concentration of 5 × 10⁶ cells/mL. A volume of 0.1 mL of the cell suspension was injected subcutaneously into the right flank of each mouse. When tumor volumes reached approximately 50 mm³, mice were randomly assigned to four groups (n = 6): empty vector (pcDNA3), RAP1 overexpression (pcDNA3‐RAP1), RAP1 overexpression plus vehicle (pcDNA3‐RAP1 + Veh.), and RAP1 overexpression plus BAY 11–7082 (pcDNA3‐RAP1 + BAY). The NF‐κB inhibitor BAY 11–7082 was administered intraperitoneally at a dose of 3 mg/kg every other day for a total duration of 30 days, whereas control animals received the corresponding solvent. The number of injected tumor cells was identical for all groups.

Tumor dimensions were measured every 3 days using vernier calipers, and tumor volume was calculated using the formula volume = length × width² × 0.5. Body weights were recorded every 3 days throughout the experiment. At the end of the study, mice were euthanized and tumor tissues were harvested for further analysis.

2.10. Statistical Analysis

Data processing and figure generation were performed using GraphPad Prism version 9.0 (GraphPad Software, La Jolla, CA, USA). All experimental results are presented as mean ± SD. Statistical comparisons were performed using Student's t‐test or one‐way ANOVA, and p < 0.05 was considered statistically significant.

3. Results

3.1. RAP1 Is Required for the Growth of HCC Cell Lines

To investigate RAP1 (TERF2IP) expression in HCC, we first conducted a pan‐cancer analysis using The Cancer Genome Atlas (TCGA) database. The analysis revealed that RAP1 was upregulated across multiple tumor types, including liver hepatocellular carcinoma (LIHC), in which its expression was significantly higher than that in normal liver tissues (Figure S1A). Consistent with these findings, we detected an overall higher expression of RAP1 at both the mRNA and protein levels in HCC cell lines (Huh7, QGY‐7703, HepG2, and Hep3B) compared with the immortalized normal hepatocyte line HL‐7702, with the difference being particularly pronounced in the cytoplasmic fraction (Figure S1B,C). This led us to hypothesize that RAP1 facilitates malignancy by working in the cytoplasmic compartment. Among these cell lines, HepG2 (p53 wild‐type) and Hep3B (p53‐null) exhibited the highest RAP1 expression levels and were therefore selected for subsequent mechanistic investigations.

To examine the functional role of RAP1 in HCC cells, we manipulated its expression using pcDNA3 eukaryotic expression vectors and small interfering RNAs (siRNAs) to achieve gain‐ and loss‐of‐function in HepG2 and Hep3B cells (Figure S1D,E). CCK‐8 assays demonstrated that RAP1 knockdown significantly reduced cell viability, whereas RAP1 overexpression markedly increased it (Figure 1A), providing initial evidence that RAP1 promotes HCC cell proliferation.

FIGURE 1.

RAP1 promotes cell growth in HepG2 and Hep3B cells. RAP1 levels were experimentally manipulated in HCC cells. (A) Cell viability affected by RAP1 was measured using the CCK‐8 assay. (B, C) Expression levels of the proliferation marker genes MKI67 and PCNA in RAP1‐altered HCC cells were measured by qRT‐PCR. (D) DNA replication activity was assessed by EdU incorporation assay. EdU‐positive cells are labeled green. Scale bar: 50 μm. (E) Quantification of EdU‐positive cells. (n = 3). *p < 0.05, **p < 0.01.

To further substantiate this finding, we assessed the expression of key proliferation markers at the transcriptional level. qRT‐PCR analysis showed that RAP1 overexpression significantly increased the mRNA levels of the cell‐cycle‐related genes MKI67, which is primarily expressed from G1 to M phase, and PCNA, which is associated with DNA replication during the S phase and with DNA repair processes. In contrast, RAP1 knockdown resulted in decreased mRNA expression of MKI67 and PCNA (Figure 1B,C). Because genomic DNA replication is a fundamental process underlying cell proliferation, we next performed EdU incorporation assays to evaluate DNA synthesis activity. The results indicated that RAP1 expression was positively associated with the rate of DNA replication (Figure 1D,E). Collectively, these data demonstrate that RAP1 expression positively correlates with proliferative capacity in HCC cells.

3.2. RAP1 Inhibits Senescence and Apoptosis in HCC

Cellular senescence and apoptosis are critical biological processes that restrict uncontrolled cell proliferation. Having established that RAP1 promotes HCC cell proliferation, we next investigated whether this effect is mediated through the suppression of senescence and apoptosis. Senescence‐associated β‐galactosidase (SA‐β‐gal) staining was performed to identify senescent cells (Figure 2A). RAP1 knockdown markedly increased the proportion of SA‐β‐gal‐positive cells, indicating enhanced cellular senescence, whereas RAP1 overexpression significantly reduced the number of senescent cells (Figure 2B). We further examined the protein levels of p16 and p21, two key senescence regulators, by Western blot (Figure 2C). RAP1 knockdown significantly increased p16 and p21 expression, whereas RAP1 overexpression dramatically reduced their levels (Figure 2D,E). Notably, RAP1 overexpression reduced p21 more prominently in HepG2 cells, while it suppressed p16 preferentially in Hep3B cells. These results indicate that RAP1 suppresses cellular senescence in HCC cells, at least in part, through downregulation of the p16/p21 pathway, with cell‐type specificity that likely reflects the different p53 status of the two lines.

FIGURE 2.

RAP1 inhibits senescence in HepG2 and Hep3B cells. RAP1 expression was manipulated in HCC cells for in vitro assays. (A) Representative images of senescent cells detected by SA‐β‐gal staining. SA‐β‐gal‐positive cells are stained blue. Scale bar: 50 μm. (B) Quantification of SA‐β‐gal‐positive cells. (C) Western blot analysis of p16 and p21 protein levels. (D, E) Quantification of p16 (D) and p21 (E) protein levels normalized to GAPDH. (n = 3). *p < 0.05, **p < 0.01, ***p < 0.001.

We then examined whether RAP1 also influences apoptotic cell death. Apoptosis was assessed using the TUNEL assay (Figure 3A). RAP1 knockdown significantly increased the proportion of TUNEL‐positive cells in both HepG2 and Hep3B cell lines, whereas RAP1 overexpression produced the opposite effect and reduced apoptotic cell numbers (Figure 3B). To explore the molecular mechanisms underlying RAP1‐mediated inhibition of apoptosis, we examined key apoptotic regulators. qRT‐PCR analysis demonstrated that RAP1 overexpression upregulated the transcription of BCL‐2, a major anti‐apoptotic gene and downstream target of NF‐κB signaling, whereas RAP1 knockdown reduced BCL‐2 expression (Figure 3C). Western blot analysis showed that RAP1 overexpression significantly increased BCL‐2 protein levels while decreasing the protein level of cleaved caspase‐3 (C‐Cas3), a critical executioner of apoptosis; conversely, RAP1 knockdown produced opposite effects (Figure 3D,E). Taken together, these findings indicate that RAP1 suppresses apoptosis by upregulating BCL‐2 expression while concurrently inhibiting caspase‐3 activation.

FIGURE 3.

RAP1 inhibits apoptosis in HepG2 and Hep3B cells. (A) Representative images of apoptotic cells detected by TUNEL assay. TUNEL‐positive cells are shown in green. Scale bar: 50 μm. (B) Quantification of TUNEL‐positive cells. (C) qRT‐PCR analysis of BCL‐2 mRNA expression. (D) Western blot analysis of cleaved caspase‐3 (C‐Cas3) and BCL‐2 protein levels. (E, F) Quantification of C‐Cas3 (E) and BCL‐2 (F) protein levels normalized to GAPDH. (n = 3). *p < 0.05, **p < 0.01.

3.3. Non‐Telomeric Role of RAP1 Drives NF‐ κB Activation

Because RAP1 has a relatively limited role in telomere protection and genomic stability in this context, we hypothesized that its tumor‐promoting effects in HCC may arise from non‐telomeric functions. Based on previous reports suggesting that RAP1 can regulate inflammatory signaling pathways, we focused on its potential role in modulating the NF‐κB signaling pathway [27].

Our experimental results identify RAP1 as an upstream regulator of NF‐κB signaling in HCC cells. In HepG2 and Hep3B cells with RAP1 knockdown, the phosphorylation level of p65 (p‐p65), a key NF‐κB subunit, was significantly reduced (Figure 4A,C), while total p65 levels remained unchanged (Figure 4A,B). In contrast, RAP1 overexpression markedly enhanced p65 phosphorylation. Quantitative analysis of the p‐p65/p65 ratio further confirmed these changes (Figure 4D), indicating activation of the NF‐κB pathway (Figure 4A–D). These findings demonstrate that RAP1 positively regulates NF‐κB signaling activity.

FIGURE 4.

RAP1 promotes hepatocellular carcinoma cell proliferation by activating the NF‐κB signaling pathway. (A) Western blot analysis of NF‐κB signaling components, including total p65 and phospho‐p65 (p‐p65). (B–D) Quantification of total p65 (B), phospho‐p65 (C), and the p‐p65/p65 ratio (D) normalized to GAPDH. (E) qRT‐PCR analysis of IL‐1β mRNA expression, a downstream target of NF‐κB signaling. (n = 3). *p < 0.05, **p < 0.01; ns, not significant.

To determine whether RAP1‐mediated NF‐κB activation leads to downstream transcriptional responses, we examined the expression of classical NF‐κB target genes. qRT‐PCR analysis revealed that RAP1 overexpression significantly increased the mRNA level of the pro‐inflammatory cytokine IL‐1β, whereas RAP1 knockdown produced the opposite inhibitory effect (Figure 4E). These results are consistent with the observed changes in p65 phosphorylation and confirm that RAP1 activates the NF‐κB pathway, thereby promoting the transcription of downstream pro‐tumorigenic genes.

3.4. RAP1 Drives In Vivo Tumor Growth in an NF‐ κB‐Dependent Manner

To further validate the tumor‐promoting role of the RAP1/NF‐κB axis in vivo, we evaluated its function using a xenograft mouse model. Tumor weight analysis showed that overexpression of RAP1 via pcDNA3‐RAP1 significantly promoted tumor growth (Figure 5A,B), with an average increase of approximately 60%. Compared with the pcDNA3‐RAP1 + Veh. group, treatment with BAY 11–7082 (pcDNA3‐RAP1 + BAY group) significantly suppressed tumor growth (Figure 5B,C), resulting in an approximately 68% reduction in tumor weight. Importantly, no significant differences in body weight were observed among the experimental groups (Figure 5D). These results demonstrate that NF‐κB signaling is required for RAP1‐driven tumor growth in vivo. Pharmacological inhibition of NF‐κB effectively reversed the tumor‐promoting effect of RAP1, thereby confirming that activation of the NF‐κB pathway is a critical downstream mechanism through which RAP1 promotes HCC progression.

FIGURE 5.

NF‐κB activity is required for RAP1‐driven tumor growth in vivo. (A) Representative images of xenograft tumors. (B) Tumor weights and (C) tumor volumes. Compared with their respective controls, RAP1 overexpression increased tumor weight and volume, whereas treatment with BAY 11–7082 markedly suppressed RAP1‐driven tumor growth. (D) Body weights of mice were measured every 3 days. (n = 6). *p < 0.05, **p < 0.01, ***p < 0.001.

4. Discussion

Although RAP1 has previously been implicated in the regulation of NF‐κB signaling in other biological contexts, its mechanistic role in HCC progression remains insufficiently defined. The NF‐κB pathway is a well‐established driver of HCC development and progression; however, the upstream regulators responsible for sustaining its aberrant activation in the tumor microenvironment remain incompletely characterized. This knowledge gap highlights the importance of identifying molecular factors that initiate or maintain NF‐κB signaling in HCC. In the present study, we identify RAP1 as a critical non‐telomeric activator of NF‐κB signaling in HCC.

Our findings provide several important insights. First, RAP1 is markedly upregulated in HCC cells and promotes malignant phenotypes by enhancing cell proliferation while suppressing both cellular senescence and apoptosis. Second, mechanistic investigations demonstrate that RAP1 functions upstream of the NF‐κB signaling cascade, promoting phosphorylation of the p65 subunit and activating downstream transcriptional programs. This activation produces multiple pro‐tumorigenic effects. Specifically, it increases the expression of anti‐apoptotic genes such as BCL‐2 while suppressing activation of the executioner caspase‐3 (C‐Cas3), thereby inhibiting apoptotic cell death. At the same time, activation of the RAP1/NF‐κB axis likely contributes to the suppression of cellular senescence observed in HCC cells. Previous studies have shown that aberrant NF‐κB activation can facilitate escape from senescence by upregulating anti‐apoptotic genes such as BCL‐2 or by repressing key senescence regulators, including p16, thereby promoting tumor progression [28, 29]. These observations provide a mechanistic explanation for our finding that RAP1 overexpression delays cellular senescence, as reflected by reduced SA‐β‐gal positivity. In addition, activation of this signaling axis induces inflammatory mediators such as IL‐1β, which may contribute to the formation of a tumor‐promoting microenvironment. Third, and most importantly, our in vivo experiments demonstrate that the oncogenic activity of RAP1 is strongly dependent on NF‐κB signaling. Pharmacological inhibition of NF‐κB effectively abolished RAP1‐driven tumor growth in the xenograft model. Together, these results establish the RAP1/NF‐κB axis as a previously underappreciated driver of HCC progression (Figure 6) and suggest that it represents a promising target for therapeutic intervention.

FIGURE 6.

Schematic depiction of the oncogenic role of RAP1 in HCC by regulating the NF‐κB signaling pathway.

Manipulation of RAP1 expression revealed that RAP1 plays a central role in determining the fate of HCC cells by coordinately regulating both senescence and apoptosis. In particular, RAP1 exhibited a consistent suppressive effect on both processes. It is well established that apoptosis and senescence represent two major cellular stress responses capable of restricting cell proliferation [30, 31]. Despite their shared role in limiting cell growth, these pathways represent distinct biological programs and generally occur independently rather than simultaneously [32, 33]. Apoptosis is a form of programmed cell death that is typically triggered by severe or irreparable cellular damage, including extensive DNA damage [34, 35]. In contrast, cellular senescence represents a stable cytostatic program that arises in response to persistent proliferative or oncogenic stress and is frequently observed in aging tissues [36, 37]. In the present study, RAP1 knockdown suppressed cell proliferation while simultaneously inducing both senescence and apoptosis. This observation raises an important mechanistic question regarding which of these pathways plays the dominant role in mediating proliferation arrest following RAP1 inhibition. Although our data clearly demonstrate activation of both responses, further studies are required to determine their relative contributions and potential mechanistic interplay.

The NF‐κB pathway also serves as a central transcriptional regulator of the senescence‐associated secretory phenotype (SASP) [38]. SASP consists of a complex array of cytokines, chemokines, and proteases secreted by senescent cells that can dynamically reshape the surrounding tissue microenvironment [39]. During HCC progression, persistent NF‐κB activation may therefore exert dual effects. In addition to facilitating escape from senescence‐mediated tumor suppression, it may promote the formation of an immunosuppressive and pro‐angiogenic microenvironment that further supports tumor progression [40].

A limitation of the present study is that we did not comprehensively characterize the SASP profile of HCC cells following RAP1 modulation, nor did we directly investigate how the RAP1/NF‐κB axis influences immune cell populations such as macrophages or T cells within the tumor microenvironment. Moreover, our assessment of cellular senescence relied primarily on SA‐β‐gal staining as a phenotypic marker. Future studies should therefore examine additional molecular indicators of senescence, including the expression and activity of key regulators such as p16 and p21, to more precisely define how the RAP1/NF‐κB axis disrupts senescence pathways in HCC.

Finally, the findings of this study provide a conceptual basis for the development of more selective therapeutic strategies targeting HCC. Our in vivo rescue experiments demonstrated that RAP1‐driven tumor growth could be significantly reversed by treatment with the NF‐κB inhibitor BAY 11–7082, confirming that the tumor‐promoting effect of RAP1 depends on NF‐κB pathway activity. These observations suggest that targeting RAP1, an upstream regulator of the pathway, may represent a more selective therapeutic strategy than directly inhibiting core NF‐κB signaling components. By disrupting the specific source of aberrant NF‐κB activation in tumor cells, such an approach could potentially suppress tumor growth while preserving the physiological functions of NF‐κB signaling in normal tissues. This concept provides a new theoretical framework for overcoming some of the limitations associated with conventional NF‐κB‐targeted therapies.

Funding

This work was supported by Tianjin Health Research Project (TJWJ2025QN042), Joint Funds of the Natural Science Foundation of Tianjin (25JCLMJC00600), Youth Innovation Talents Training Project of Tianjin First Central Hospital.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Table S1: Primers used in this study.

Table S2: siRNA oligonucleotides used in this study.

Figure S1: RAP1 (TERF2IP) is upregulated in hepatocellular carcinoma tissues and cell lines. (A) Pan‐cancer analysis based on the TCGA database reveals that the expression of RAP1 (TERF2IP) is significantly higher in hepatocellular carcinoma (LIHC) tissues compared to adjacent normal tissues (labeled in the red rectangle). (B) In order to know RAP1 level in HCC cell lines, the relative RAP1 level in an immortalized benign hepatocyte cell line HL‐7702 and four HCC cell lines were detected by qRT‐PCR. The lncRNA level in HL‐7702 cells was set to 1. (C) qRT‐PCR analysis verified the efficiency of artificial manipulation of RAP1 levels in HepG2 and Hep3B cells. *p < 0.05, **p < 0.01, NC, negative control.

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.