Abstract
Background
Pituitary tumor-transforming gene 1 (PTTG1) has been implicated in multiple malignancies; however, its precise role and regulatory mechanism in gastric cancer remain unclear. This study aimed to determine how PTTG1 regulates cell proliferation, migration, and epithelial–mesenchymal transition (EMT) through the TGF-β/Smad signaling pathway in gastric cancer.
Methods
PTTG1 expression in gastric cancer was evaluated using The Cancer Genome Atlas (TCGA) datasets and subsequently validated by western blotting in gastric cancer cell lines. To investigate the biological functions of PTTG1, functional assays including CCK8, colony formation, and Transwell migration and invasion assays were performed following PTTG1 knockdown in SGC-7901 and HGC-27 cells. The effects on EMT were further examined by assessing EMT-related markers by western blotting. Tumorigenic and metastatic potential in vivo was determined using a nude mouse xenograft model. The regulatory influence of PTTG1 on the TGF-β/Smad pathway was analyzed by measuring the expression levels of TGF-β, Smad2, Smad3, and their phosphorylated forms.
Results
PTTG1 expression was significantly upregulated in gastric cancer tissues compared with normal gastric mucosa according to TCGA data and validation in cell lines. Silencing PTTG1 in SGC-7901 and HGC-27 cells markedly inhibited proliferation and colony formation (p < 0.01), and suppressed migration and invasion (p < 0.01). Western blotting revealed increased E-cadherin and reduced N-cadherin, α-SMA, and Vimentin expression after PTTG1 knockdown. In vivo, xenograft tumor growth was significantly reduced in nude mice injected with si-PTTG1 cells compared with controls (n = 6 per group; p < 0.05). Mechanistically, PTTG1 depletion attenuated TGF-β, phosphorylated Smad2, and phosphorylated Smad3 levels, indicating inhibition of the TGF-β/Smad pathway.
Conclusion
PTTG1 promotes gastric cancer progression by activating the TGF-β/Smad axis to enhance cell proliferation, invasion, and EMT. These findings highlight PTTG1 as a potential diagnostic biomarker and therapeutic target in gastric cancer.
Supplementary Information
The online version contains supplementary material available at 10.1186/s41065-026-00647-0.
Keywords: PTTG1, Gastric cancer, Epithelial-mesenchymal transition, TGF-β/Smad signaling, Tumor progression
Introduction
Gastric cancer (GC), a common malignant tumor of the stomach, remains the fourth most prevalent cancer worldwide despite a gradual decline in incidence in certain regions [1]. Adenocarcinoma accounts for most of GC cases [2]. At present, surgical resection combined with adjuvant chemotherapy or radiotherapy constitutes the standard therapeutic strategy and has improved the five-year survival rate for selected patients [3]. However, due to the absence of specific symptoms in the early stages, GC is often diagnosed at an advanced stage when treatment options are limited and the prognosis is poor. The identification of novel molecular targets and clarification of their biological roles in GC progression are therefore of considerable clinical and therapeutic significance.
The PTTG1, initially identified as a securin, is frequently overexpressed in multiple cancers, including colorectal, breast, and lung cancers, where it promotes cell proliferation, genomic instability, and metastasis [4]. In addition to its classical role in cell division, PTTG1 participates in diverse cellular processes such as DNA repair, apoptosis, and metabolism [4–6]. Dysregulated overexpression of PTTG1 has been documented in multiple tumor types and has been linked to tumor growth, epithelial-mesenchymal transition (EMT), invasion, and angiogenesis [4, 7–9]. For instance, PTTG1 has been shown to enhance adenoviral vector transduction in pancreatic cancer cells by upregulating CXADR expression and to drive cancer progression through various signaling axes [10]. It has also been identified as an androgen-responsive gene essential for both proliferation and metastasis in prostate cancer [7, 11]. Despite these observations, the functional role of PTTG1 in gastric cancer remains insufficiently understood.
The TGF-β/Smad signaling pathway represents a central regulator of tumorigenesis, exerting dual and context-dependent functions that include tumor-suppressive activity in the early stages of disease and tumor-promoting effects in advanced cancers [12]. Recent studies suggest that the interaction between oncogenes and the TGF-β/Smad pathway is critical in the development and progression of gastric cancer [13]. However, whether PTTG1 contributes to gastric cancer progression by regulating this pathway has not yet been clearly defined.
The present study was designed to investigate the role of PTTG1 in gastric cancer progression. The findings identify a novel regulatory axis involving PTTG1 and provide evidence that targeting this molecule may represent a promising therapeutic strategy for gastric cancer.
Materials and methods
Bioinformation
RNA-seq data and corresponding clinical information for gastric cancer were obtained from The Cancer Genome Atlas (TCGA) database via the Genomic Data Commons (GDC) portal. Differential expression analysis was conducted using the DESeq2 package in R. Associations between PTTG1 expression and overall survival were assessed using the ‘survival’ and ‘survminer’ packages, with all statistical analyses performed in R.
Cell culture
Human gastric cancer cell lines, including AGS and MKN-45, and normal gastric epithelial cells (GES-1), were purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA). Authentication of all cell lines was performed by short tandem repeat profiling. The cells were maintained in RPMI-1640 medium supplemented with 10% fetal bovine serum (FBS; Gibco), 100 U/mL penicillin, and 100 µg/mL streptomycin. The cultures were incubated at 37 °C in a humidified atmosphere containing 5% CO2. Transfections with small interfering RNA (siRNA) were carried out using Lipofectamine 3000, and cell viability was assessed at the indicated time points by the CCK-8 assay.
SiRNA transfection
Cells were seeded into six-well plates at a density of 2 × 105 cells per well 24 h before transfection. Two independent siRNA sequences specifically targeting PTTG1, along with scrambled negative controls (GenePharma, Shanghai, China), were introduced into the cells using Lipofectamine 3000 (Thermo Fisher Scientific) according to the manufacturer’s protocol, at a final concentration of 50 nM. The transfection efficiency was confirmed by quantitative reverse transcription PCR (qRT-PCR) and western blotting 48 h post-transfection. The siRNA sequences were: si-PTTG1: GACCTGCAATAATCCAGAA; si-NC: TCCTTCGAACGTGCTACGT.
CCK-8 assay
Cell viability was evaluated using the Cell Counting Kit-8 (CCK-8; Dojindo, Japan). A total of 2 × 103 cells were seeded into each well of 96-well plates in quintuplicate. At the specified time points, 10 µL of CCK-8 solution was added to each well, and the plates were incubated for 2 h at 37 °C. The absorbance was measured at 450 nm using a microplate reader (Bio-Rad).
Colony formation assay
After transfection, 500 cells per well were seeded into six-well plates containing RPMI-1640 medium supplemented with 10% fetal bovine serum (FBS; Gibco) and maintained under standard culture conditions (37 °C, 5% CO₂). The control group consisted of cells transfected with scrambled siRNA (si-NC). After incubation for 10–14 days, visible colonies were fixed with methanol and stained with 0.1% crystal violet. A colony was defined as a cluster of at least 50 cells. Quantification was performed either by blinded manual counting or with ImageJ software, and the number of colonies was normalized to plating efficiency.
Transwell migration and invasion assays
For the Transwell migration and invasion assays, 5 × 10⁴ to 1 × 10⁵ cells were seeded into each insert. Migrated or invaded cells were quantified either as the average number of cells per microscopic field or expressed as a percentage relative to the control group. For data reliability, counting was performed within the linear range by preventing over-confluence on the underside of the membrane. Transwell chambers (Corning, USA) were employed to evaluate cell motility. For invasion assays, the upper chambers were pre-coated with Matrigel (Beyotime, P8360). Cells were suspended in serum-free medium and added to the upper chambers, whereas the lower chambers were filled with complete medium containing 10% fetal bovine serum (FBS). After 24 h incubation, the cells on the lower membrane surface were fixed with paraformaldehyde, stained with crystal violet, and subsequently counted.
qPCR assay
Total RNA was extracted from cultured cells using TRIzol reagent (Invitrogen, 15596026) following the manufacturer’s instructions. RNA purity and concentration were assessed with a NanoDrop 2000 spectrophotometer (Thermo Scientific). One microgram of RNA was reverse transcribed into cDNA using the PrimeScript RT reagent kit (Takara, RR037A). Quantitative PCR was performed using SYBR Green Master Mix (Takara, RR420A) on a QuantStudio 6 Flex Real-Time PCR System (Applied Biosystems). The amplification protocol consisted of an initial denaturation at 95 °C for 30s, followed by 40 cycles of 95 °C for 5s and 60 °C for 30s. GAPDH served as the internal control, and relative gene expression levels were calculated using the 2−ΔΔCt method. Each reaction was performed in triplicate. The primer sequences used were as follows: PTTG1 forward, 5′-AGCTTCTCCAGGCCTTTGT-3′ and reverse, 5′-GAGTTCTCCTCCTCCTCCAC-3′; GAPDH forward, 5′-AGAAGGCTGGGGCTCATTTG-3′ and reverse, 5′-AGGGGCCATCCACAGTCTTC-3′.
Western blot analysis
Protein expression was examined by western blotting. Total protein was extracted using RIPA lysis buffer (Beyotime, P0013B) containing protease and phosphatase inhibitors. The lysates were incubated on ice for 30 min and centrifuged at 12,000 × g for 15 min at 4 °C. Protein concentrations were determined using the BCA Protein Assay Kit (Beyotime, P0012). Equal amounts of protein (30 µg per sample) were separated by SDS-PAGE and transferred onto PVDF membranes (0.45 μm pore size) at 100 V for 90 min, which were then blocked with 5% non-fat milk. The membranes were incubated overnight at 4 °C with the following primary antibodies: anti-PTTG1 (Abcam, ab79546, 1:1,000), anti-GAPDH (Abcam, ab8245, 1:5,000), anti-E-cadherin (Abcam, ab40772, 1:1,000), anti-N-cadherin (Abcam, ab76057, 1:1,000), anti-α-SMA (Abcam, ab124964, 1:1,000), anti-Vimentin (Abcam, ab92547, 1:1,000), anti-TGF-β (Abcam, ab92486, 1:1,000), anti-Smad2 (Abcam, ab40855, 1:1,000), anti-p-Smad2 (Abcam, ab188334, 1:1,000), anti-Smad3 (Abcam, ab40854, 1:1,000), and anti-p-Smad3 (Abcam, ab52903, 1:1,000). After washing, the membranes were incubated with horseradish peroxidase (HRP)-conjugated secondary antibodies (Beyotime, A0208, 1:5,000). Protein bands were visualized using an enhanced chemiluminescence (ECL) detection kit (Beyotime, P0018FS).
Tumor xenograft model
Female BALB/c nude mice were used in this study, and experimental groups were established to ensure comparable age, sex, and body weight to minimize variability between treatment and control cohorts. Female BALB/c nude mice (6 weeks old, n = 6 per group) were purchased from Vital River (Beijing, China). SGC-7901 cells transfected with si-PTTG1 or si-NC (1 × 107 cells in 100 µL PBS) were subcutaneously injected into the right flanks of mice. Tumor growth was measured every 5 days, and volumes were calculated using the formula V = (length × width2)/2. After 25 days, mice were sacrificed, and tumor tissues were excised and weighed. Ethical approval was obtained from the Ethics Committee of Changzhou Traditional Chinese Medicine Hospital(Approval No.2025 − 147).
Immunohistochemistry (IHC)
Excised tumor tissues were fixed in formalin, embedded in paraffin, and sectioned. Slides were blocked and incubated overnight at 4°C with primary antibodies against Ki67 (Abcam, ab16667, 1:200) and MMP-9 (Abcam, ab38898, 1:200). Following incubation with horseradish peroxidase (HRP)-conjugated secondary antibodies and visualization with 3,3’-diaminobenzidine (DAB) substrate (Beyotime, P0203), slides were examined under a light microscope. For IHC analysis, images were captured under identical exposure conditions, and quantification was performed using ImageJ software by measuring either mean staining intensity or the percentage of positively stained cells. For each sample, three to five representative fields were analyzed, and biological replicates per group were included (n = [insert number]).
Statistical analysis
The data are expressed as mean ± standard deviation (SD). Statistical analyses were performed using GraphPad Prism version 8.0 (GraphPad Software, San Diego, CA, USA). Comparisons between two groups were conducted with the unpaired two-tailed Student’s t-test. For comparisons among multiple groups, one-way analysis of variance (ANOVA) was applied, followed by Tukey’s post hoc test. A p-value of less than 0.05 was considered statistically significant.
Results
PTTG1 is highly expressed in gastric cancer cells
The expression pattern of PTTG1 in gastric cancer was first examined at the mRNA level using data from the TCGA database. Analysis demonstrated significantly elevated PTTG1 expression in gastric cancer tissues compared with normal tissues (Fig. 1A). Validation at the protein level was performed by western blotting in a panel of gastric cancer cell lines and the normal gastric epithelial cell line GES-1. PTTG1 protein was detected in all four gastric cancer cell lines, whereas expression was markedly lower in GES-1 (Fig. 1B). These findings indicate that PTTG1 is broadly expressed in gastric cancer.
Fig. 1.
PTTG1 is highly expressed in gastric cancer cells. A Analysis of the TCGA database showing PTTG1 mRNA expression in gastric cancer tissues (n = 408) and normal gastric tissues (n = 211). B Western blot detection of PTTG1 protein expression in the normal gastric epithelial cell line GES-1 and gastric cancer cell lines SGC-7901, HGC-27, BGC-823, and AGS, with GAPDH used as a loading control. Quantification of PTTG1 expression relative to GAPDH is shown in the accompanying bar graph. Each experiment was repeated three times. Data are presented as mean ± SD. ***P < 0.001, **P < 0.01 versus GES-1
Silencing PTTG1 impairs gastric cancer cell growth and invasion in vitro
The functional role of PTTG1 in gastric cancer progression was investigated by siRNA-mediated knockdown in SGC-7901 and HGC-27 cells. Efficient suppression of PTTG1 expression was confirmed by qPCR at 24, 48, and 72 h (Fig. 2A) and by western blotting (Fig. 2B). Reduced PTTG1 expression was associated with decreased cell viability as determined by CCK-8 assay (Fig. 2C) and decreased colony formation capacity (Fig. 2D). In Transwell assays, migration and invasion were both significantly impaired in PTTG1-silenced cells compared with control groups (Fig. 2E and F). These observations demonstrate that PTTG1 promotes the proliferative and invasive properties of gastric cancer cells in vitro.
Fig. 2.
Knockdown of PTTG1 inhibits gastric cancer cell growth and invasion in vitro. A qPCR analysis of PTTG1 mRNA expression in SGC-7901 and HGC-27 cells transfected with control, si-NC, or si-PTTG1 at 24, 48, and 72 h. B Western blot analysis of PTTG1 protein levels in SGC-7901 and HGC-27 cells under the indicated conditions, with GAPDH as the loading control. Bar graphs show quantification of PTTG1 protein relative to GAPDH. (C) CCK-8 assay of cell viability in the indicated groups. D Colony formation assay showing representative images and quantification of colony numbers. E Transwell migration assays with representative images and quantification of migrated cells. F Invasion assays using Matrigel-coated Transwell chambers with representative images and quantification of invaded cells. Each experiment was performed in triplicate. Data are presented as mean ± SD. ***P < 0.001, versus si-NC
PTTG1 depletion suppresses the epithelial-mesenchymal transition (EMT) process in gastric cancer cells
Given the established importance of EMT in cancer metastasis, the effect of PTTG1 on EMT marker expression was examined. Western blot analysis revealed that PTTG1 knockdown increased E-cadherin expression while decreasing N-cadherin, α-SMA, and Vimentin levels (Fig. 3). These results suggest that PTTG1 facilitates the EMT process in gastric cancer.
Fig. 3.
Silencing PTTG1 suppresses epithelial-mesenchymal transition in gastric cancer cells. Western blot analysis of EMT-associated markers (E-cadherin, N-cadherin, α-SMA, and Vimentin) in SGC-7901 and HGC-27 cells transfected with control, si-NC (scramble), or si-PTTG1. GAPDH served as the internal loading control. Densitometric quantification of protein expression relative to GAPDH is shown in bar graphs. Each experiment was repeated three times. Data are presented as mean ± SD. ***P < 0.001, **P < 0.01 versus si-NC
In vivo tumor growth is affected by PTTG1 knockdown
The effect of PTTG1 on tumorigenesis in vivo was assessed using a subcutaneous xenograft model established with SGC-7901 cells transfected with either si-PTTG1 or si-NC. Tumor volume and weight were consistently lower in the PTTG1-silenced group compared with controls (Fig. 4A). Immunohistochemical staining of xenograft tumors showed reduced expression of Ki67 and MMP-9 in the si-PTTG1 group (Fig. 4B). These findings provide in vivo evidence that PTTG1 promotes tumor growth in gastric cancer.
Fig. 4.
PTTG1 knockdown reduces tumor growth and metastasis in vivo. A Subcutaneous xenograft models were established by injecting SGC-7901 cells transfected with si-NC or si-PTTG1 into nude mice. Representative tumor images, tumor growth curves, and tumor weights at endpoint are shown. B Immunohistochemical staining of paraffin-embedded tumor sections for Ki67 and MMP-9. Representative fields from each group are shown, with positive cells displaying brown cytoplasmic or nuclear staining. Scale bars: 200 μm. Each experiment was repeated three times. Data are presented as mean ± SD. ***P < 0.001 versus si-NC
PTTG1 regulates TGF-β/Smad signaling in gastric cancer cells
To explore the mechanism through which PTTG1 exerts its oncogenic effects, the TGF-β/Smad signaling pathway was examined. Western blot analysis demonstrated that PTTG1 knockdown decreased the expression of TGF-β, phosphorylated Smad2, and phosphorylated Smad3, whereas the levels of total Smad2 and Smad3 remained unchanged (Fig. 5). These findings indicate that PTTG1 promotes gastric cancer progression by positively regulating the TGF-β/Smad pathway.
Fig. 5.
PTTG1 regulates the TGF-β/Smad pathway in gastric cancer. Western blot analysis of TGF-β, Smad2, phosphorylated Smad2 (p-Smad2), Smad3, and phosphorylated Smad3 (p-Smad3) expression in SGC-7901 and HGC-27 cells transfected with control, si-NC, or si-PTTG1. GAPDH was used as the loading control. Quantification is shown in bar graphs, with TGF-β normalized to GAPDH and p-Smad2 and p-Smad3 normalized to total Smad2 and Smad3, respectively. Each experiment was repeated three times. Data are presented as mean ± SD. ***P < 0.001, **P < 0.01 versus si-NC
Discussion
GC remains one of the leading causes of cancer-related mortality worldwide, largely due to delayed diagnosis, aggressive tumor biology, and complex molecular mechanisms underlying its progression [14]. Numerous oncogenes, signaling pathways, and epigenetic modifications have been implicated in the initiation and development of GC. The discovery of novel biomarkers and the clarification of molecular pathways involved in disease progression are therefore essential for improving diagnostic accuracy and developing targeted therapeutic strategies [15]. In this study, PTTG1 was identified as a potential oncogenic driver in gastric cancer, and its contribution to cell proliferation, invasion, and EMT through activation of the TGF-β/Smad pathway was demonstrated.
Cell proliferation and migration represent fundamental processes in tumor progression, while EMT provides tumor cells with plasticity, invasiveness, and the ability to disseminate to distant sites [16]. This process is characterized by the downregulation of epithelial markers and concomitant upregulation of mesenchymal proteins, thereby enabling tumor cells to detach from the primary tumor and infiltrate surrounding tissues [17]. In the present study, silencing of PTTG1 markedly inhibited gastric cancer cell growth and invasion and significantly altered the expression of EMT-associated proteins, including Vimentin. These findings indicate that PTTG1 not only promotes tumor cell proliferation but also drives the phenotypic reprogramming required for metastatic spread, underscoring its dual role in GC progression.
PTTG1 was initially characterized as a securin responsible for ensuring proper chromatid separation during mitosis [4]. Subsequent studies have revealed that it also regulates diverse cellular processes, including DNA repair, apoptosis, and metabolism. Aberrant overexpression of PTTG1 has been strongly associated with oncogenic transformation and tumor aggressiveness [18]. In line with these observations, our findings show that knockdown of PTTG1 in gastric cancer cells suppressed growth and invasion while modulating EMT marker expression, suggesting that PTTG1 functions as a molecular switch that enhances oncogenic phenotypes, at least in part by regulating EMT-related genes.
Previous studies have reported elevated PTTG1 expression in several malignancies, including liver, lung, breast, and prostate cancers, where it enhances tumorigenesis, angiogenesis, and therapeutic resistance [4, 9]. For instance, PTTG1 has been shown to promote prostate cancer progression through androgen-dependent mechanisms and to facilitate pancreatic cancer progression by regulating viral receptor expression. The present study extends these findings by demonstrating that PTTG1 exerts a comparable oncogenic effect in gastric cancer, thereby supporting the view that PTTG1 acts as a universal tumor promoter whose biological effects vary depending on the tumor context. The ability of PTTG1 to regulate both cell-cycle dynamics and invasive potential highlights its importance as a driver of tumor progression across diverse cancer types.
The TGF-β/Smad signaling axis plays a highly context-dependent role in cancer biology. Upon ligand binding, TGF-β induces phosphorylation of Smad2 and Smad3, which subsequently regulate the transcription of genes involved in cell cycle arrest, EMT, and extracellular matrix remodeling [19]. In gastric cancer, aberrant activation of this pathway has been associated with enhanced invasiveness, therapeutic resistance, and poor prognosis. In the present study, silencing of PTTG1 suppressed TGF-β expression and reduced the phosphorylation of Smad2 and Smad3, without altering total Smad protein levels. These findings suggest that PTTG1 promotes the activation, rather than the production, of Smad signaling intermediates. Therefore, the results provide a mechanistic explanation for how PTTG1 enhances EMT and malignancy in gastric cancer.
The current findings are consistent with previous reports identifying PTTG1 as an oncogenic driver in several malignancies, including colorectal, breast, and lung cancers, where it has been shown to promote cell proliferation, invasion, and genomic instability. In gastric cancer, our results extend this body of evidence by demonstrating that PTTG1 silencing reduces EMT marker expression and suppresses tumor growth at least in part through modulation of the TGF-β/Smad pathway. These data reinforce the concept that PTTG1 is a broad-spectrum tumor promoter whose oncogenic activity is mediated by context-specific mechanisms.
Despite the promising findings, several limitations of this study should be noted. First, the sample size of the in vivo xenograft model was relatively small, which may limit the statistical power to detect subtle differences in tumor growth. Second, although in vitro assays provided mechanistic insights into the role of PTTG1 in cell proliferation, migration, and EMT, they cannot fully replicate the complexity of the tumor microenvironment and host immune interactions in gastric cancer. Third, the present study focused primarily on experimental models, and clinical validation of PTTG1 expression and prognostic significance in patient cohorts was not performed. Future research incorporating larger animal cohorts, orthotopic or patient-derived xenograft models, and comprehensive clinical datasets will be essential to confirm the translational relevance of our findings.
In conclusion, the present study demonstrates that PTTG1 promotes gastric cancer growth, invasion, and EMT by activating the TGF-β/Smad pathway. These findings reveal a previously unrecognized role for PTTG1 in gastric cancer pathogenesis and provide a mechanistic rationale for considering PTTG1 as a potential therapeutic target.
Supplementary Information
Acknowledgements
Not applicable.
Authors’ contributions
All authors contributed to the study conception and design. Material preparation and the experiments were performed by Dong Wang. Data collection and analysis were performed by Hui Geng and Ziwei Xia. The first draft of the manuscript was written by Chenjie Qiu and all authors commented on previous versions of the manuscript. All authors read and approved the final manuscript.
Funding
Not applicable.
Data availability
All data generated or analyzed during this study are included in this published article.
Declarations
Ethics approval
All animal experiments were approved by the Ethics Committee of Changzhou Traditional Chinese Medicine Hospital for the use of animals and conducted in accordance with the National Institutes of Health Laboratory Animal Care and Use Guidelines(Approval No.2025 − 147).
Competing interests
The authors declare no competing interests.
Footnotes
Publisher’s note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
References
- 1.Smyth EC, Nilsson M, Grabsch HI, van Grieken NC, Lordick F. Gastric cancer. Lancet. 2020;396(10251):635–48. [DOI] [PubMed] [Google Scholar]
- 2.Fassan M, Kuwata T, Matkowskyj KA, Rocken C, Ruschoff J. Claudin-18.2 immunohistochemical evaluation in gastric and gastroesophageal junction adenocarcinomas to direct targeted therapy: A practical approach. Mod Pathol. 2024;37(11):100589. [DOI] [PubMed] [Google Scholar]
- 3.Yasuda T, Wang YA. Gastric cancer immunosuppressive microenvironment heterogeneity: implications for therapy development. Trends Cancer. 2024;10(7):627–42. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 4.Zhou Q, Li L, Sha F, Lei Y, Tian X, Chen L, et al. PTTG1 reprograms asparagine metabolism to promote hepatocellular carcinoma progression. Cancer Res. 2023;83(14):2372–86. [DOI] [PubMed] [Google Scholar]
- 5.Vlotides G, Eigler T, Melmed S. Pituitary tumor-transforming gene: physiology and implications for tumorigenesis. Endocr Rev. 2007;28(2):165–86. [DOI] [PubMed] [Google Scholar]
- 6.Tien S, Zhou H, Zhou Q, Liu H, Wu B, Guo Y. PTTG1 alleviates acute alcoholic liver injury by inhibiting Endoplasmic reticulum stress-induced hepatocyte pyroptosis. Liver Int. 2023;43(4):840–54. [DOI] [PubMed] [Google Scholar]
- 7.Ersvaer E, Kildal W, Vlatkovic L, Cyll K, Pradhan M, Kleppe A, et al. Prognostic value of mitotic checkpoint protein BUB3, Cyclin B1, and pituitary tumor-transforming 1 expression in prostate cancer. Mod Pathol. 2020;33(5):905–15. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 8.Dun Y, Hu H, Liu F, Shao Y, He D, Zhang L, et al. PTTG1 promotes CD34(+)CD45(+) cells to repair the pulmonary vascular barrier via activating the VEGF-bFGF/PI3K/AKT/eNOS signaling pathway in rats with phosgene-induced acute lung injury. Biomed Pharmacother. 2023;162:114654. [DOI] [PubMed] [Google Scholar]
- 9.Teveroni E, Di Nicuolo F, Vergani E, Oliva A, Vodola EP, Bianchetti G et al. SPTBN1 mediates the cytoplasmic constraint of PTTG1, impairing its oncogenic activity in human seminoma. Int J Mol Sci. 2023;24(23)16891. [DOI] [PMC free article] [PubMed]
- 10.Long L, Gao J, Zhang R. PTTG1 enhances oncolytic adenovirus 5 entry into pancreatic adenocarcinoma cells by increasing CXADR expression. Viruses. 2023;15(5):1153. [DOI] [PMC free article] [PubMed]
- 11.Zhang X, Wu N, Huang H, Li S, Liu S, Zhang R, et al. Phosphorylated PTTG1 switches its subcellular distribution and promotes beta-catenin stabilization and subsequent transcription activity. Oncogene. 2023;42(32):2439–55. [DOI] [PubMed] [Google Scholar]
- 12.Frangogiannis N. Transforming growth factor-beta in tissue fibrosis. J Exp Med. 2020;217(3):e20190103. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 13.Song C, Zhou C. HOXA10 mediates epithelial-mesenchymal transition to promote gastric cancer metastasis partly via modulation of TGFB2/Smad/METTL3 signaling axis. J Exp Clin Cancer Res. 2021;40(1):62. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 14.Duan Y, Xu Y, Dou Y, Xu D. Helicobacter pylori and gastric cancer: mechanisms and new perspectives. J Hematol Oncol. 2025;18(1):10. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 15.Zhuang Y, Yin H, Huang Y, Jiang F, Li L, Wu Z, et al. Catalytic hairpin assembly-powered nanozyme-SERS dual-function sensing system for ultrasensitive detection of gastric precancerous lesions. Biosens Bioelectron. 2025;283:117536. [DOI] [PubMed] [Google Scholar]
- 16.Li S, Sun M, Cui Y, Guo D, Yang F, Sun Q, et al. Ephrin A1 functions as a ligand of EGFR to promote EMT and metastasis in gastric cancer. EMBO J. 2025;44(5):1464–87. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 17.Li D, Xia L, Huang P, Wang Z, Guo Q, Huang C, et al. Heterogeneity and plasticity of epithelial-mesenchymal transition (EMT) in cancer metastasis: focusing on partial EMT and regulatory mechanisms. Cell Prolif. 2023;56(6):e13423. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18.Zhi T, Jiang K, Xu X, Yu T, Zhou F, Wang Y, et al. ECT2/PSMD14/PTTG1 axis promotes the proliferation of glioma through stabilizing E2F1. Neuro Oncol. 2019;21(4):462–73. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 19.Lin GS, Zheng RQ, Xu ZW, Xing SP, Wu HF, Xie Y, et al. Proteomic screening identifies brusatol targets TGFbetaRII to suppresses non-small cell lung cancer metastasis. Phytomedicine. 2025;139:156468. [DOI] [PubMed] [Google Scholar]
Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Supplementary Materials
Data Availability Statement
All data generated or analyzed during this study are included in this published article.