Sodium selenite promotes apoptosis and augments autophagic flux in cervical cancer cells by activating the YAP-Hippo signaling pathway

Introduction

Cervical cancer remains the fourth most frequently diagnosed malignancy and the fourth leading cause of cancer-related death among women worldwide, with an estimated more than 650,000 new cases and nearly 350,000 deaths each year (1,2). Although screening and human papillomavirus (HPV) vaccination have cut incidence in wealthy nations, cervical cancer is still rising in low- and middle-income regions, and standard therapy including surgery, radiotherapy, plus platinum chemoradiation, often leads to relapse, metastasis, or serious toxicity. These observations underscore the urgency of pinpointing novel therapeutic targets and developing innovative treatment approaches to treat cervical cancer (3).

Apoptosis and autophagy are two evolutionarily conserved catabolic processes central to cellular homeostasis. Programmed cell death eliminates genetically compromised or superfluous cells, whereas autophagy recycles damaged organelles and misfolded proteins to sustain metabolic balance under stress (4,5). Disturbances in either pathway can facilitate oncogenic transformation, promote tumor progression, and underlie resistance to chemotherapy or radiotherapy (6,7). Hence, pharmacological agents capable of reinstating apoptosis or modulating autophagic flux may restore therapeutic sensitivity and curb tumor growth.

The Hippo pathway serves as a master regulator of tissue size by integrating mechanical cues, metabolic status, and extracellular signals, thereby playing a key role in processes, such as cell proliferation, apoptosis, and stem cell self-renewal (8,9). Aberrant Yes-associated protein (YAP) has been documented in hepatocellular, colorectal, breast, and cervical cancers, among others, and often correlates with advanced stage, metastasis, and poor prognosis (10,11). Intriguingly, mounting evidence points to intricate crosstalk between Hippo signaling and the autophagy machinery, with YAP both regulating and being regulated by autophagic flux.

Selenium, an indispensable trace element incorporated into selenoproteins via selenocysteine, exerts diverse biological actions spanning antioxidant defense, immune modulation, and redox homeostasis (12). Sodium selenite (SS) is a readily bioavailable inorganic selenium compound widely used in nutritional supplementation and investigated for anticancer potential. Pre-clinical studies have shown that SS can generate reactive oxygen species, trigger DNA damage, and modulate signal-transduction pathways, culminating in growth arrest and apoptosis in several tumor types (13,14). Nonetheless, its impact on the apoptotic and autophagic programs of cervical cancer cells, and, importantly, the possible involvement of the YAP-Hippo axis, remains poorly defined.

Against this backdrop, the present study explores whether SS suppresses cervical-cancer cell growth by orchestrating apoptosis and autophagy through modulation of the YAP-Hippo pathway. Clarifying this relationship may not only broaden our mechanistic understanding of selenium’s anticancer activity but also unveil new molecular targets and combinatorial strategies to treat cervical cancer. A protocol was prepared before the study without registration. We present this article in accordance with the MDAR and ARRIVE reporting checklists (available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-1737/rc).

Methods

Cell lines and reagents

Human cervical cancer HeLa cells were obtained from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China). Analytical-grade SS (purity ≥98%) was obtained from Sigma-Aldrich (St. Louis, MO, USA). Biochemical reagents, including radioimmunoprecipitation assay (RIPA) buffer, bicinchoninic acid (BCA) quantification assay kit, Annexin V-FITC/propidium iodide (PI), apoptosis detection kit, and Cell Counting Kit-8 (CCK-8) were all purchased from Beyotime Biotechnology Co., Ltd. (Shanghai, China). Transwell chambers were supplied by Corning (Corning, NY, USA), and horseradish peroxidase (HRP)-conjugated goat anti-rabbit immunoglobulin G (IgG) secondary antibody was obtained from Cell Signaling Technology (Danvers, MA, USA).

Cell culture

HeLa cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM; Gibco, Grand Island, NY, USA) enriched with 10% (v/v) fetal bovine serum (FBS; Gibco), 100 U/mL penicillin and 100 µg/mL streptomycin (Gibco), incubated at 37 ℃ with 5% CO₂ in a humidified atmosphere (Thermo Fisher Scientific, Waltham, MA, USA). Growth medium was refreshed every 2–3 days, and sub-culturing was performed once cell confluency reached approximately 80–90%.

Experimental grouping and treatment

Exponentially growing HeLa cells were seeded into 6-well plates, 96-well plates, or Transwell chambers (Corning) as required for downstream assays. After overnight cell attachment, cultures were divided into four conditions: (I) vehicle control [0 µM SS; ≤0.1% dimethyl sulfoxide (DMSO; Sigma-Aldrich)], and (II) 5, (III) 10, or (IV) 15 µM SS. Treatments were applied for 24, 48, or 72 hours before further assays.

CCK-8 assay for cell proliferation

For cell viability analysis, 5×10³ cells were seeded in each well of a 96-well plate (five technical replicates per condition). The following day, cells were exposed to SS at 0, 2.5, 5, 7.5, 10, 15, 20, or 40 µM for 6, 12, or 24 hours. Subsequently, 10 µL of CCK-8 reagent (Beyotime Biotechnology) was added to each well. After 2 hours of incubation with CCK-8, absorbance at 450 nm was recorded with a microplate spectrophotometer (BioTek, Winooski, VT, USA). Relative proliferation was expressed as optical density (OD₄₅₀), and half-maximal inhibitory concentrations (IC₅₀) were calculated using GraphPad Prism 8.0.2 (GraphPad Software, San Diego, CA, USA).

Scratch (wound healing) cell migration assay

HeLa monolayers were established in 6-well plates. Once confluent, a sterile 200 µL pipette tip (Axygen, Union City, CA, USA) was used to disrupt (scratch) the monolayer in a single linear swipe. Detached cells were rinsed away twice with phosphate-buffered saline (PBS) (Gibco). Cultures were then exposed to SS at the previously determined IC₅₀ concentration. After 24 hours, five randomly chosen fields along the wound were imaged with an inverted microscope (Olympus, Tokyo, Japan). Wound closure was quantified with ImageJ (NIH, Bethesda, MD, USA), and migration was expressed as: Migration rate (%) = (initial scratch width − scratch width at 24 hours) / initial scratch width ×100.

Transwell assay

Matrigel-coated Transwell chambers were prepared by allowing the Matrigel matrix to polymerize at 37 ℃. HeLa cells (1×10⁵ per insert) were seeded in the upper compartment in a serum-free medium, while the lower wells contained DMEM supplemented with 20% FBS to serve as a chemoattractant. Following 24 hours of treatment, inserts were washed with PBS, fixed in methanol (15 min), and stained with 0.1% crystal violet (20 min). Non-migrated cells on the upper surface were gently removed. Invaded cells adhering to the underside of the membrane were counted in five random microscopic fields to quantify invasive capacity. For migration experiments, we do not lay substrate glue in the upper chamber.

Apoptosis analysis by flow cytometry

HeLa cell cultures treated for 24 hours with SS at the IC₅₀ dose were harvested, washed twice with ice-cold PBS, and re-suspended in 500 µL of binding buffer. Cells were incubated with 5 µL Annexin V-FITC and 10 µL PI for 15 min at ambient temperature in the dark. Stained samples were analyzed immediately on a flow cytometer, and percentages of early- and late-apoptotic cells were calculated with FlowJo software; total apoptosis represents the sum of these two fractions.

Immunofluorescence

Sterile glass coverslips were placed in 6-well plates and seeded with 1×10⁶ HeLa cells per well. After a 12-hour attachment period, cultures were treated with SS at the IC₅₀ dose for 24 hours. Cells were then fixed in 4% paraformaldehyde (PFA) (30 min), permeabilized with 0.5% Triton X-100 (10 min), and blocked in 5% bovine serum albumin (30 min). Cells were then incubated with primary antibodies at 4 ℃ overnight. The next day, cells were washed with PBS, and incubated for 1 hour with CoraLite-488- or CoraLite-594-labelled goat anti-rabbit IgG. Cell were then counter-stained with 4',6-diamidino-2-phenylindole (DAPI; Beyotime Biotechnology) (10 min), and mounted. The Zeiss LSM 880 (Carl Zeiss, Oberkochen, Germany) confocal microscope was used to acquire the images. Use ImageJ software to detect fluorescence intensity. Five randomly selected fields of view of each specimen were photographed at high magnification.

Western blotting

Cells were lysed on ice using RIPA buffer (30 min). Cell lysate was then centrifuged (12,000 ×g, 4 ℃, 15 min) and supernatant was collected as protein samples. Protein concentrations were determined with BCA kit, and equal amounts of protein were denatured (5 min, 100 ℃) in loading buffer. These protein samples were then resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), and transferred to polyvinylidene fluoride (PVDF) membranes. Later, membranes were blocked with 5% skim milk (1 hour, room temperature), and were incubated overnight at 4 ℃ with the indicated primary antibodies: LC3B (microtubule-associated protein 1 light chain 3) (1:2,000), sequestosome 1 (SQSTM1/p62) antibody (1:1,000), total YAP (1:1,000), phospho-YAP (1:1,000), total LATS1 (1:1,000), and phospho-LATS1 (Ser909, 1:1,000). Later, membranes were washed thrice in Tris-buffered saline with Tween-20 (TBST) and incubated for 1 hour with HRP-conjugated goat anti-rabbit IgG (1:5,000). Membranes were again washed thrice in TBST, and bands were visualized by enhanced chemiluminescence on a ChemiDoc imaging system. ImageJ software was used to detect protein grayscale values.

Animal experiments

All animal experiments were performed under a project license (No. 202402) granted by the Ethics Committee of Henan University of Chinese Medicine Animal Care and Use Committee, in compliance with the Guide for the Care and Use of Laboratory Animals. Five-week-old female BALB/c nude mice (BALB/c-nu/nu, Beijing Vital River Laboratory Animal Technology Co., Ltd., Beijing, China) were housed under specific-pathogen-free conditions for one week of acclimation. Log-phase HeLa cells (1×10⁶) suspended in 100 µL PBS were injected subcutaneously into the right forelimb. Tumor dimensions were recorded every other day with calipers (Mitutoyo Corp., Kawasaki, Japan), and volume was estimated as: V (mm³) = (L × W²)/2, where L and W denote length and width, respectively. When tumors reached 40–100 mm³, mice were randomized to receive either vehicle (saline; China National Pharmaceutical Group, Beijing, China) or SS (6 mg/kg, intraperitoneally) on alternate days. After 14 days of treatment, animals were anaesthetized with isoflurane (RWD Life Science Co., Ltd., Shenzhen, China) and euthanized; tumors were excised for subsequent analyses.

Histopathology

Tumors were fixed in 4% PFA (Servicebio, Wuhan, China) for 48 h, embedded in paraffin, and sectioned at 4 µm. For immunohistochemistry, re-hydrated sections were incubated overnight at 4 ℃ with anti-YAP antibody (1:1,000, Cell Signaling Technology, #8418S; Cell Signaling Technology, Danvers, MA, USA). After washing, a species-matched secondary antibody (ZSGB-Bio, Beijing, China) was applied for 1 hour at 37 ℃, and signal development followed the manufacturer’s protocol. Inflammatory infiltration and general tissue architecture were assessed on adjacent sections stained with hematoxylin and eosin (H&E; Beyotime Biotechnology).

Bioinformatics analyses

Microarray datasets GSE63678 and GSE89657 were retrieved from the Gene Expression Omnibus (GEO, https://www.ncbi.nlm.nih.gov/geo/). Raw CEL files were background-corrected and normalized with the Robust Multi-array Averaging (RMA) algorithm. Probe sets were collapsed to gene-level expression, and inter-study batch effects were mitigated using the ComBat procedure. Differentially expressed genes (DEGs) were identified with the limma package (version 3.58.1), applying thresholds of |log₂ fold-change| ≥1 (equivalent to ≥2-fold) and adjusted P<0.05 (Benjamini-Hochberg correction). Functional enrichment of DEGs was explored by Gene Ontology (GO) analysis, and Gene Set Enrichment Analysis (GSEA) was performed on a log₂ fold-change-ranked gene list. Visualizations were generated with the enrichplot package (version 1.24.0).

Statistical analyses

Group comparisons were carried out in SPSS 22.0 software (IBM Corp., Armonk, NY, USA). One-way analysis of variance (ANOVA) was used for multi-group tests; where significant, least significant difference (LSD) t-test was employed for post-hoc pairwise comparisons. Comparison between the two groups was conducted with an independent two-tailed Student’s t-test. A two-sided P<0.05 was considered statistically significant. Data are expressed as mean ± standard deviation (SD).

Results

SS inhibits the proliferative and metastatic potential of cervical-cancer cells

To establish whether SS directly restrains key malignant behaviors of cervical cancer cells, we examined its impact on HeLa cell proliferation, migration, and invasion. The CCK-8 proliferation assay demonstrated that SS exerts a concentration- and time-dependent inhibitory effect on HeLa cells. As the concentration of SS increased and the treatment duration extended, cell viability gradually decreased. The IC₅₀ values of SS intervention on HeLa cells at 6, 12, and 24 hours were 14.59, 6.84, and 4.92 µM, respectively (Figure 1A). Further investigations into the impacts of SS on the migration and invasion of HeLa cells revealed that treatment with 4.92 µM SS for 24 hours significantly reduced the migration and invasion of HeLa cells (Figure 1B-1G). These findings indicate that SS not only inhibits proliferation in cervical cancer cells but also markedly curtails their metastatic potential.

Figure 1 SS inhibits the proliferative and metastatic potential of cervical-cancer cells. (A) CCK-8 assay showing that SS reduces HeLa cell viability in a dose- and time-dependent manner after 6, 12, and 24 hours of treatment; calculated IC₅₀ values were 14.59, 6.84, and 4.92 µM, respectively. (B) Wound healing assay demonstrating that 4.92 µM SS markedly delays wound closure compared with the vehicle control (scale bar: 100 µm). (C) Quantification of wound healing migration rates confirming significant inhibition of migratory capacity. (D) Transwell invasion assay showing fewer SS-treated cells penetrating the Matrigel membrane (0.1% crystal violet, 200×). (E) Histogram of invasive cell numbers indicating reduced invasiveness upon SS exposure. (F) Transwell migration assay (without Matrigel) visualizing decreased numbers of migrated cells after SS treatment (0.1% crystal violet, 200×). (G) Histogram of cell migration demonstrating consistent attenuation of motility. *, P<0.05 vs. control. CCK-8, Cell Counting Kit-8; IC₅₀, half-maximal inhibitory concentration; SS, sodium selenite.

SS induces apoptosis in cervical cancer cells

As reduced viability can stem from either growth arrest or cell death (15), we next examined whether SS provokes apoptosis in HeLa cells. Flow cytometry analysis showed that treatment of HeLa cells with 4.92 µM SS for 24 hours markedly increased the proportion of annexin V-positive/PI-positive cells relative to the vehicle control (Figure 2A,2B). TUNEL staining revealed a significant increase in the percentage of apoptotic cells after SS treatment compared to that with the vehicle control (Figure 2C,2D). This finding confirms that SS diminishes HeLa-cell viability, at least in part, by activating apoptotic pathways.

Figure 2 SS induces apoptosis in cervical cancer cells. (A) HeLa cells were exposed to 4.92 µM SS for 24 hours or to vehicle (control), stained with annexin V-FITC and propidium iodide, and analyzed by flow cytometry, then shows an increased proportion of early and late apoptotic cells compared with the control group. (B) Quantitative analysis confirms a significant rise in total apoptosis percentage following SS treatment. (C) TUNEL staining showing enhanced nuclear fragmentation and apoptotic signal intensity in SS-treated cells. 400×. (D) Quantification of TUNEL-positive cells demonstrates consistent apoptotic elevation. *, P<0.05 vs. control. Annexin V-FITC, annexin V-fluorescein isothiocyanate; SS, sodium selenite.

SS promotes autophagic flux by enhancing autophagosome degradation

As autophagy often influences cancer-cell fate (16), we asked whether SS elicits a functional autophagic response in HeLa cells. To test this, we inhibited lysosomal enzyme activity by adding chloroquine (CQ, a lysosomotropic compound that blocks the final stage of autophagy) (17). Western blotting (Figure 3A,3B) and immunofluorescence assay (Figure 3C,3D) results showed a significant increase in the expression of LC3-II in the CQ treatment group in comparison to the control group. Furthermore, the combination treatment of CQ + SS further enhanced the expression of LC3-II compared to the CQ alone, indicating that SS further increases autophagosome formation (Figure 3A,3B). Next, to investigate whether the observed increase in LC3 II accumulation after SS treatment was in response to blocked or activated autophagic flux, we examined the expression levels of p62 in HeLa cells after 24 hours of SS treatment. Since p62 binds to polyubiquitinated proteins that are integrated into autophagosomes and degraded in autolysosomes, the association between p62 and LC3 can monitor autophagic flux (18). In this line, we observed a decrease in p62 expression after SS treatment, indicating that SS may enhance autophagic flux (Figure 3A-3D). Taken together, these findings show that SS amplifies autophagic flux in HeLa cells by both driving autophagosome biogenesis and promoting their lysosomal degradation, rather than merely causing a buildup of undegraded vesicles.

Figure 3 SS promotes autophagic flux by enhancing autophagosome degradation. (A) Western blot analysis of p62 and LC3 expression in HeLa cells treated with vehicle control, 4.92 µM SS for 24 hours, 20 µM CQ, or the combination SS + CQ. β-actin was used as the loading control. (B) Densitometric quantification showing that SS treatment increases LC3-II levels while decreasing p62 accumulation; combined SS + CQ further elevates LC3-II, confirming enhanced autophagosome formation and flux. (C) Immunofluorescence staining for LC3 (green), p62 (red) and nuclei (DAPI, blue) under the same treatment conditions. Merged images illustrate increased LC3 puncta and reduced p62 fluorescence in SS-treated cells compared with controls, whereas CQ blocks degradation and causes signal accumulation. 500×. (D) Quantitative analysis of immunofluorescence intensity for LC3 and p62 demonstrating that SS activates autophagic flux by simultaneously promoting autophagosome biogenesis and lysosomal degradation. Data are presented as mean ± SD. *, P<0.05; **, P<0.01; ***, P<0.001. CQ, chloroquine; SD, standard deviation; SS, sodium selenite.

SS regulates autophagy through YAP-Hippo pathway

As Hippo signaling can couple growth control to autophagy (19), we asked whether SS-induced autophagy depends on YAP-Hippo activity. In HeLa cells, SS markedly increased the phosphorylation of YAP (p-YAP) while reducing total YAP protein; phosphorylation of the upstream kinase LATS1 (p-LATS1) rose in parallel, indicating activation of the YAP-Hippo cascade (Figure 4A,4B). Recent work suggests that YAP turnover partly relies on the autophagy-lysosome system. Consistently, immunofluorescence showed strong co-localization of YAP with p62, confirming that YAP is routed to autolysosomes for degradation (Figure 4C,4D). To test functional relevance, we knocked down YAP (sh-YAP) and compared three groups: control, SS, and SS + sh-YAP. Western blotting revealed that SS alone lowered p62 and raised LC3-II, signatures of enhanced autophagic flux; these changes were largely reversed by YAP knockdown, with p62 increasing and LC3-II decreasing in the SS + sh-YAP group (Figure 4E,4F). Collectively, these results indicate that SS stimulates autophagy in cervical-cancer cells by activating and subsequently promoting lysosomal degradation of YAP, placing the Hippo pathway upstream of the observed autophagic flux.

Figure 4 SS regulates autophagy through YAP-Hippo pathway. (A) Western blot analysis showing that treatment of HeLa cells with 4.92 µM SS for 24 h markedly increases p-LATS1 and p-YAP while reducing total YAP protein levels, indicating activation of the Hippo cascade. β-actin was used as the loading control. (B) Densitometric quantification confirming upregulation of p-LATS1 and p-YAP and downregulation of total YAP protein following SS treatment. (C) Immunofluorescence staining of p62 (red), YAP (green), and nuclei (DAPI, blue) under the same treatment conditions as (A), showing increased cytoplasmic puncta and strong co-localization of YAP with p62 after SS exposure, suggesting YAP is targeted to autolysosomes for degradation. Magnification 400×. (D) Quantitative fluorescence intensity analysis demonstrating increased p62, YAP colocalization in SS-treated cells compared with controls. (E) Western blot analysis of p62 and LC3 in HeLa cells that were transfected with sh-YAP or scrambled control and then exposed to SS. β-actin was used as the loading control. SS alone decreased p62 and increased LC3-II, while YAP knockdown reversed these effects. (F) Densitometric quantification of p62 and LC3-II/LC3-I ratios confirming that SS-induced autophagic flux is abolished when YAP is silenced. *, P<0.05; **, P<0.01; ***, P<0.001. p-LATS1, phosphorylation of LATS1; p-YAP, phosphorylation of YAP; SS, sodium selenite.

Integrated RNA-seq meta-analysis and molecular docking identify YAP1 as a direct SS target driving hippo-pathway activation in cervical cancer

To complement the cellular assays and search for a direct molecular link between SS and Hippo signaling, we carried out transcriptome-level bioinformatics and in-silico docking studies. We selected the cervical-cancer datasets GSE63678 and GSE89657 for cross-dataset differential-expression analysis. After background correction, normalisation, and batch-effect removal, 104 intersecting DEGs were identified (|fold change| ≥2, P<0.05; Figure 5A-5C). GO enrichment showed that these DEGs participate in oocyte meiosis, regulation of hippocampal signaling, cellular senescence, and other processes (Figure 5D). Gene-set enrichment analysis (GSEA) further revealed significant up-regulation of Hippo-pathway signatures in cervical-cancer samples versus normal tissue (Figure 5E). Focusing on the Hippo subset, we built a protein-protein interaction (PPI) map with STRING and ranked hub genes with the CytoHubba MCC algorithm, in which YAP1 emerged as the core node (Figure 5F,5G). Molecular docking predicted a stable binding mode between SS and YAP1, with key contacts at Tyr407, Asn354, and Ser381 through a mix of hydrogen bonds and hydrophobic interactions (Figure 5H). These in-silico results place YAP1 at the intersection of Hippo-pathway dysregulation in cervical cancer and suggest that SS can directly engage YAP1, offering a mechanistic explanation for the pathway activation observed in our wet-lab experiments.

Figure 5 Integrated RNA-seq meta-analysis and molecular docking identify YAP1 as a direct SS target driving hippo-pathway activation in cervical cancer. (A,B) Volcano plots of DEGs from cervical cancer datasets GSE63678 and GSE89657 after normalization and batch-effect correction. Significantly dysregulated genes are highlighted (|log₂FC| ≥1, adjusted P<0.05). (C) Venn diagram displaying the 104 intersecting DEGs common to both datasets, representing shared transcriptional changes between studies. (D) Gene Ontology enrichment of the intersecting DEGs showing predominant involvement in oocyte meiosis, regulation of hippocampal signaling, and cellular senescence; Bar length reflects the number of genes per term, and color intensity denotes −log₁₀ (P value). (E) Gene Set Enrichment Analysis showing significant upregulation of Hippo-signaling pathway gene sets in tumor vs. normal samples, confirming pathway activation at the transcriptome level. (F) Protein-protein interaction network of Hippo-related DEGs generated with STRING, visualizing the interconnections among pathway components. (G) Hub-gene ranking by CytoHubba (MCC algorithm) in Cytoscape (version 3.9.1), identifying YAP1 as the top-scoring central node within the Hippo network. (H) Molecular-docking model showing the predicted binding conformation between SS and YAP1. The complex exhibits a stable interaction pattern involving Tyr407, Asn354, and Ser381 residues through a combination of hydrogen bonds and hydrophobic contacts, suggesting direct SS-YAP1 engagement. DEG, differentially expressed gene; FC, fold change; SS, sodium selenite.

SS inhibits cervical-cancer tumor growth in vivo

To determine whether the anti-proliferative effects of SS observed in vitro translate to a living system, we evaluated tumor growth in a nude-mouse xenograft model. Mice bearing established HeLa xenografts received intraperitoneal injections of SS (6 mg/kg) or saline. Tumors in SS-treated animals were visibly and significantly smaller than those in controls throughout the experiment (Figure 6A). Final tumor weights mirrored the volume data (Figure 6B). Hematoxylin-eosin staining showed markedly reduced inflammatory-cell infiltration in SS tumors compared with controls (Figure 6C). Immunohistochemistry revealed lower percentages of YAP-positive and Ki-67-positive cells in the SS group, indicating diminished Hippo-pathway activity and cellular proliferation (Figure 6D,6E). Together, these results confirm that SS significantly suppresses cervical-cancer tumor growth in vivo while attenuating inflammation, YAP signaling, and proliferative activity.

Figure 6 SS inhibits cervical cancer tumor growth in vivo. (A) Representative photographs of HeLa cell xenografts from BALB/c-nude mice treated intraperitoneally with SS (6 mg/kg, every other day for 14 days) or vehicle (control). Tumors in the SS-treated group were visibly smaller than those in controls throughout the treatment period. (B) Tumor-volume curves showing a significant reduction in tumor volume in SS-treated mice compared with controls; volumes were measured every 2 days with digital calipers and calculated as V = (L × W²)/2. Final tumor weights mirrored volume differences, confirming SS-mediated growth inhibition. (C) Hematoxylin and eosin staining of tumor sections demonstrating reduced inflammatory-cell infiltration and more compact tumor architecture in the SS-treated group relative to controls. Magnification 100×. (D) Immunohistochemistry for YAP and Ki-67 staining for the same treatment conditions, showing decreased nuclear YAP expression and a lower proportion of Ki-67-positive proliferating cells in SS-treated tumors. 200×. (E) Immunohistochemical histograms of YAP- and Ki-67-positive cell percentages confirming significant attenuation of Hippo-pathway activation and cell proliferation following SS treatment. *, P<0.05. SS, sodium selenite.

Discussion

Treating cervical cancer remains challenging, and new, effective therapies are urgently required. This work examined how SS influences apoptosis and autophagy in cervical-cancer cells and how these effects relate to Hippo-pathway signaling, yielding several noteworthy observations. Recent evidence shows that excessive activation of the oncogene YAP compromises innate immunity, rendering host cells more susceptible to HPV infection. In murine models, YAP overexpression alone can provoke cervical squamous-cell carcinoma within six to eight months (20,21). Co-expression of high-risk HPV E6/E7 with YAP shortens this latency to four months, indicating a cooperative acceleration of malignant progression. Genomic surveys of The Cancer Genome Atlas also reveal frequent 11q22 copy-number gains, encompassing YAP and its targets BACR2/3 (22). Elevated YAP protein is characteristic of cervical tumors displaying epithelial-mesenchymal transition (EMT) traits and predicts poor outcome (23). Taken together, these findings identify the Hippo/YAP axis as a promising therapeutic target. Autophagy and Hippo signaling interact bi-directionally to coordinate autophagic flux in response to environmental cues that govern tissue homeostasis and tumorigenesis (24,25). Only a few agents that modulate Hippo/YAP activity have been identified, and whether autophagy should be stimulated or inhibited for cancer therapy remains controversial. Exploring the Hippo-YAP-autophagy axis could therefore provide fresh therapeutic insight.

We first showed that SS suppresses HeLa-cell viability in a concentration- and time-dependent fashion (Figure 1). These results are in line with earlier reports on selenium compounds (26), suggesting that SS may hinder the normal growth and proliferation of cervical cancer cells. Flow-cytometric analysis showed a parallel, concentration-dependent increase in apoptosis, suggesting that SS limits cell growth partly by activating programmed cell death (27). Our findings suggest that SS may inhibit the growth of cervical cancer cells by triggering the apoptosis process (Figure 2). Additionally, we found that SS can induce autophagy in cervical cancer cells (Figure 3). Autophagy is an intracellular degradation process that plays a dual role in tumorigenesis and tumor progression (28). While moderate autophagy clears damaged organelles and supports survival, excessive autophagy can drive cell death; in this context, the SS-induced response is likely cytotoxic. In this study, the accumulation of autophagy-related protein LC3-II increased, and P62 levels decreased after SS treatment (Figure 4), indicating that SS induces autophagy in cervical cancer cells. This may be a cellular response to SS stress and could play a crucial role in inhibiting tumor cell growth.

When Hippo signaling is activated, phosphorylated YAP is sequestered in the cytoplasm and degraded, thereby suppressing proliferation and promoting apoptosis (29). We found that after SS treatment, the phosphorylation levels of YAP and LATS1 significantly increased, indicating the activation of the YAP-Hippo pathway by SS. This result provides new clues for explaining the SS-induced apoptosis and autophagy. Our data place Hippo activation upstream of both SS-induced apoptosis and autophagy. Consistent with cell-culture findings, SS markedly inhibited tumor growth in a mouse xenograft model and reduced intra-tumoral YAP expression, underscoring the pathway’s relevance in vivo (Figure 6).

In summary, this study shows for the first time that SS regulates both apoptosis and autophagy in cervical-cancer cells by modulating the YAP-Hippo pathway. These findings deepen our understanding of cervical-cancer biology and point to YAP-directed selenium therapy as a potential strategy. Future work should explore SS’s anti-tumor mechanisms in additional in vivo settings and assess combination regimens that may improve clinical efficacy.

Conclusions

Our findings demonstrate that SS exerts potent anti-tumor effects against cervical cancer by simultaneously inducing apoptosis and enhancing autophagic flux. Mechanistically, SS activates the Hippo pathway, increases LATS1/YAP phosphorylation, and promotes YAP degradation, thereby positioning YAP as a central mediator linking apoptotic and autophagic responses. These in vitro results were validated in vivo, where SS significantly suppressed xenograft tumor growth with minimal toxicity. Collectively, this work identifies SS as a promising Hippo-pathway-modulating agent and highlights YAP-centered signaling as a viable therapeutic target for cervical cancer. Further studies examining SS-based combination strategies and its translational potential in clinical settings are warranted.

Acknowledgments

The authors would like to thank all individuals who provided assistance and support for this study.

Footnote

Reporting Checklist: The authors have completed the MDAR and ARRIVE reporting checklists. Available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-1737/rc

Data Sharing Statement: Available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-1737/dss

Peer Review File: Available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-1737/prf

Funding: None.

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-1737/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. All animal experiments were performed under a project license (No. 202402) granted by the Ethics Committee of Henan University of Chinese Medicine Animal Care and Use Committee, in compliance with the Guide for the Care and Use of Laboratory Animals.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

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