Tropomyosin 3 regulates tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma to promote malignant progression in gastric cancer

Abstract

BACKGROUND

Tropomyosin 3 (TPM3) has been implicated in the progression of several cancers; however, its specific role and underlying molecular mechanisms in gastric cancer (GC) remain unclear. The current research aimed to investigate the role of TPM3 in the onset and advancement of GC, along with the related molecular pathways.

AIM

To investigate TPM3’s role in enhancing GC malignancy and elucidates the underlying molecular mechanisms.

METHODS

TPM3 expression in GC tissues was evaluated using bioinformatics analysis. Protein expression levels were determined using western blotting, and mRNA levels were measured through quantitative real-time PCR. Cellular functional assays, including cell counting kit-8 assay, 5-ethynyl-2-deoxyuridine incorporation, colony formation, wound-healing, and transwell migration/invasion, were conducted to assess the effects of TPM3 and tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma (YWHAG) on GC cell proliferation, migration, invasion, and metastatic potential. Cell cycle progression and apoptosis were analyzed using flow cytometry. A subcutaneous xenograft model in nude mice was employed to assess the tumorigenic potential of GC cells, while a tail vein injection model was used to evaluate their invasive and metastatic potential in vivo.

RESULTS

TPM3 was significantly upregulated in GC tissues and cells, and its high expression correlated with poor patient prognosis. Silencing TPM3 markedly inhibited GC cell proliferation, invasion, and metastasis while promoting apoptosis. TPM3 interacted with YWHAG, and TPM3 knockdown reduced YWHAG expression. Notably, the inhibitory effects of TPM3 silencing on GC progression were partially reversed by overexpressing YWHAG, confirming YWHAG as a key downstream effector. Furthermore, TPM3 knockdown suppressed activation of the mitogen-activated protein kinase pathway in a YWHAG-dependent manner. In vivo experiments demonstrated that increased TPM3 expression significantly promoted GC tumor growth and metastasis, whereas silencing YWHAG effectively attenuated this effect.

CONCLUSION

TPM3 promotes GC progression by regulating YWHAG and activating the mitogen-activated protein kinase signaling pathway. These findings identify the TPM3/YWHAG axis as a potential therapeutic target for GC intervention.

Key Words: Tropomyosin 3; Tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma; Mitogen-activated protein kinase signaling pathway; Gastric cancer; Molecular mechanism

Core Tip: In this study, we investigated the role of tropomyosin 3 (TPM3) in gastric cancer (GC). We found that TPM3 is highly expressed in GC tissues and is associated with poor prognosis. Mechanistically, TPM3 promotes GC cell proliferation, invasion, and metastasis through interaction with tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma and subsequent activation of the mitogen-activated protein kinase signaling pathway. Both in vitro and in vivo experiments demonstrated that the TPM3/tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma axis plays a critical role in GC progression.

INTRODUCTION

Gastric cancer (GC) is a common malignant tumor of the digestive system and ranks third in incidence and fourth in mortality worldwide. The global survival rate over five years for patients with GC is estimated to be 20%[1]. Recently, with rapid economic development, China has become a high-incidence region for GC. Statistical analyses of cancer mortality in China demonstrate that GC ranks second among female cancer patients and third among male cancer patients[2]. Early-stage GC often presents without obvious clinical symptoms, making timely and effective screening challenging[3]. As a result, many patients are diagnosed at advanced stages, leading to an unfavorable prognosis[4]. Consequently, investigating GC pathogenesis is of considerable importance. Although there is a relatively comprehensive understanding of the etiology of GC, the molecular mechanisms underlying its initiation and progression remain unclear. A deeper understanding of these molecular mechanisms may facilitate the discovery of new therapeutic targets and advance precision medicine.

Tropomyosin (TPM) is a family of actin-binding proteins located in the thin filaments of skeletal muscle, smooth muscle, and certain non-muscle tissues[5]. These proteins play an important role in regulating muscle contraction, and are prevalent in different eukaryotic cells[6]. Aberrant expression of TPM family genes initially induces alterations in cellular cytoskeletal morphology and ultimately contributes to fibrosis and carcinogenesis. The major TPM isoforms in mammals include TPM1, TPM2, TPM3, and TPM4, each of which exists as multiple splice variants[7]. TPM1 has been widely acknowledged as a tumor-inhibiting factor, with notably reduced levels reported in several cancers, including gastric, prostate, and non-small cell lung carcinomas. Loss of TPM1 expression is consistently associated with poor clinical outcomes and tumor progression[8-10]. Additionally, TPM2 plays a tumor-suppressive role in prostate cancer by inhibiting PD-LIM7-mediated nuclear translocation of YAP1, thereby suppressing malignant progression[11]. Conversely, TPM4 exhibits oncogenic characteristics across multiple malignancies, frequently promoting tumor progression[12].

Located on human chromosome 1q21.3, TPM3 is a crucial member of the TPM family that helps stabilize cytoskeletal microfilaments. This gene is composed of a single exon and encodes a 284-amino-acid protein[13]. Initial research indicates that in non-muscle eukaryotic tissues, actin microfilaments contribute to various cellular functions by assembling into unique actin structures, promoting cell migration, intracellular vesicle trafficking, and other essential processes[7]. Recent studies have demonstrated that TPM3 is involved in tumor initiation and progression in non-muscle tissue. Structural alterations and dysregulated TPM3 expression are associated with the development of several malignancies[6,14]. Although the role of TPM3 in tumorigenesis is context-dependent, it is predominantly oncogenic. TPM3 is frequently overexpressed in cancers such as gliomas, liver cancer, and colorectal cancer, where it facilitates tumor development through mechanisms including gene fusion events and the activation of epithelial-mesenchymal transition (EMT)[15-17]. Moreover, TPM3 regulates key proliferative and survival signaling pathways, particularly the phosphatidylinositol 3-kinase/protein kinase B pathway, and its elevated expression has been significantly associated with poor prognosis in multiple tumor types[18,19]. A previous proteomic analysis demonstrated that TPM3 is significantly upregulated in diffuse and intestinal subtypes of GC[20]. However, studies investigating the molecular functions and regulatory mechanisms of TPM3 in GC cells remain limited, emphasizing the need for further investigations.

The 14-3-3 proteins constitute a group of regulatory factors that bind to phosphoserine and phosphothreonine. In mammals, it comprises seven isoforms (β, γ, ε, δ, ζ, τ, and η), and is widely expressed in human tissues[21]. These proteins regulate biological processes by binding to phosphorylated proteins, altering their activity, stability, or protein-protein interactions[22,23]. Studies have found that each isoform of 14-3-3 proteins exhibits distinct expression patterns and possesses unique interaction networks[24]. Tumor progression and treatment responses have been associated with the unusual expression of specific 14-3-3 isoforms. For instance, elevated levels of 14-3-3ζ have been associated with chemotherapy resistance in patients with non-small cell carcinoma and diffuse large B-cell lymphoma[25]. Conversely, 14-3-3δ expression is significantly reduced in primary breast cancer[26]. Located on human chromosome 7q11.23, tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma (YWHAG), also known as 14-3-3γ, comprises seven exons and encodes a 245-amino-acid protein[27]. Increasing evidence suggests that YWHAG plays an important role in tumor metastasis by controlling gene networks associated with EMT and protecting cancer cells against oxidative stress through autophagy[22]. Furthermore, YWHAG modulates the mitogen-activated protein kinase (MAPK) signaling pathway in bladder cancer[28]. However, studies investigating the role of YWHAG in GC are limited, and its functional significance and underlying molecular mechanisms in GC require further investigation.

The MAPK signaling pathway was significantly enriched in the transcriptomic analyses of TPM3-correlated genes in GC. This bioinformatic evidence, together with the previously reported regulatory role of YWHAG in the MAPK pathway in bladder cancer, prompted us to hypothesize that TPM3 promotes GC progression through a previously unrecognized TPM3/YWHAG/MAPK signaling axis. In this study, we aimed to systematically evaluate TPM3 expression in GC and investigate its impact on tumor proliferation, migration, and clinical prognosis using transcriptomic data from The Cancer Genome Atlas (TCGA). Our findings demonstrate that TPM3 modulates YWHAG expression and is essential in GC development. Moreover, it was identified that the TPM3/YWHAG axis is involved in regulating the MAPK signaling pathway. Collectively, these findings provide new insights into TPM3’s role in GC and suggest it as a potential therapeutic target for GC treatment.

MATERIALS AND METHODS

Bioinformatics analysis

The pan-cancer expression profile of TPM3 was visualized using Tumor Immune Estimation Resource (version 2.0) web server. Publicly available transcriptomic data and corresponding clinical information for stomach adenocarcinoma were retrieved from the TCGA database. The analysis of TPM3 differential expression between tumor and nearby normal tissues was conducted using the limma R package, with statistical significance criteria of |log₂(fold change)| > 1 and an adjusted P value < 0.05. Overall survival (OS) was analyzed through Kaplan-Meier survival curves, and the log-rank test was applied to evaluate the association between TPM3 expression (dichotomized by median) and OS. To investigate signaling cascades and associated biological functions, enrichment analysis of gene sets was performed using the R software package clusterProfiler (version 4.0). Gene sets obtained from Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) databases were employed for the analysis. A GO or KEGG term was considered significantly enriched when the false discovery rate-adjusted P value was less than 0.05.

Clinical tissue samples

All GC tissues and corresponding adjacent non-cancerous tissues were collected from patients undergoing surgical resection at the Second Affiliated Hospital of Nanchang University. Before surgery, none of the patients received radiotherapy or chemotherapy. Right after surgical removal, newly obtained tissue samples were promptly preserved in 4 g/L paraformaldehyde (PFA) (P1110; Solarbio, Beijing, China). Tumor purity was evaluated by experienced pathologists through histopathological assessment of hematoxylin and eosin (HE)-stained slides. The study was approved by Institutional Review Board of the Second Affiliated Hospital of Nanchang University, and all patients provided written informed consent. Complete research processes that included human subjects were conducted in full compliance with the guidelines specified by the Helsinki Declaration.

Cell culture and transfection

In this study, all human GC cell lines were obtained from the Shanghai Cell Institute of the Chinese Academy of Sciences. Human GC cell lines AGS, HGC27, MKN45, and MKN74 were cultured in roswell park memorial institute 1640 (RPMI-1640) medium (31800; Solarbio, Beijing, China) supplemented with 10 mL/L fetal bovine serum (FSP500; ExCell Bio, Shanghai, China). All cell lines were maintained at 37 °C in a humidified incubator containing 50 mL/L CO₂. All three MAPK pathway inhibitors used in this study were obtained from Ambeed, Inc. (Chicago, IL, United States).

Lentiviral short hairpin RNAs (shRNAs) targeting TPM3 (sh-TPM3#1, sh-TPM3#2, and sh-TPM3#3), control shRNA (sh-NC), overexpression plasmid (OE-TPM3), and corresponding control vector (vector) were obtained from HanHeng Biotechnology. Similarly, lentiviral shRNA targeting YWHAG (sh-YWHAG), sh-NC, overexpression plasmid (OE-YWHAG), and the corresponding control vector (vector) were obtained from HanHeng Biotechnology. The ‘sh-NC’ group served as a non-targeting scramble control, and the ‘vector’ group served as an empty vector control in all the transfection experiments. The specific shRNA sequences used in this study are listed in Supplementary Table 1. Lentiviral transfection was performed according to the manufacturer’s instructions, and puromycin (ST551; Beyotime, Shanghai, China) was used to select stable transfected cells.

Quantitative real-time PCR and western blotting

Total RNA was extracted using TriQuick Reagent (R1100, Solarbio, Beijing, China), and cDNA was synthesized using a cDNA synthesis kit (AE301-02, TransGen, Beijing, China) under the following conditions: (1) Genomic DNA digestion at 42 °C for 2 minutes; (2) Followed by reverse transcription at 25 °C for 5 minutes; (3) 55 °C for 15 minutes; and (4) 85 °C for 5 minutes. Quantitative real-time PCR (qRT-PCR) was performed using SYBR Green Master Mix (RR820A; Takara, Kyoto, Japan). The qRT-PCR protocol was as follows: (1) Initial denaturation at 95 °C for 5 minutes; (2) Followed by 35 cycles of 95 °C for 30 seconds; (3) 58 °C for 30 seconds; and (4) 72 °C for 30 seconds. The qRT-PCR results for cell lines were analyzed using the 2^-ΔΔCT method, while the 2^-ΔΔCT method was applied for tissue samples. The PCR primers used in this study were synthesized by Servicebio Biotech (Wuhan, Hubei Province, China), and their sequences are provided in Supplementary Table 2.

Total cell proteins were extracted on ice for 30 minutes using radio immunoprecipitation assay (RIPA) lysis buffer (R0010, Solarbio, Beijing, China). The proteins were denatured by boiling at 100 °C for 10 minutes in 1× sodium dodecyl sulfate (SDS) protein loading buffer (IR9158; Solarbio, Beijing, China). Following standard SDS-polyacrylamide gel electrophoresis electrophoresis and membrane transfer, the membrane was blocked with 50 mL/L skimmed milk (D8340; Solarbio, Beijing, China) for 1 hour and incubated overnight at 4 °C with the primary antibody. After 2-3 washes with Tris-buffered saline containing 0.1 mL/L Tween 20 (T8220; Solarbio, Beijing, China), the membrane was incubated with the corresponding secondary antibody at room temperature for 1 hour. Detailed information on all key reagents, including antibodies and chemical inhibitors (supplier, catalog number, and working dilution or concentration), is provided in Supplementary Table 3 to ensure transparency and reproducibility. After three washes with Tris-buffered saline containing 0.1 mL/L Tween 20, the signal was detected using a chemiluminescent detection kit (S6009M; UElandy, Suzhou, Jiangsu Province, China), and the immunoblot was visualized and analyzed using the ImageJ software. For proteins displaying multiple bands, the total intensity of all specific bands was quantified to represent the overall protein expression.

Co-immunoprecipitation

Pre-chilled RIPA lysis solution (1.5 mL) containing PMSF (P0100; Solarbio, Beijing, China) was added to a 10 cm culture dish (TCD010100; Jet BioFil, Guangzhou, Guangdong Province, China), and proteins were extracted as described for western blotting (WB). After incubation with 3 μg of the primary antibody at 4 °C for 2 hours, 30 μL of protein A/G magnetic beads (P2080S; Beyotime, Shanghai, China) were added, and the mixture was incubated overnight with gentle rotation. The following day, the samples were washed thrice with RIPA lysis buffer; after which 30 μL of 2× SDS-polyacrylamide gel electrophoresis loading buffer (P0015, Beyotime, Shanghai, China) was added, and the samples were incubated and boiled at 100 °C for 10 minutes for further WB.

Immunohistochemistry

Fresh tumor tissues were embedded in paraffin and sectioned into 4 μm thick slices. The tissue sections were incubated with primary antibodies, followed by the corresponding secondary antibodies. 3,3′-diaminobenzidine (DA1010; Solarbio, Beijing, China) was used for signal visualization, and the immunohistochemical staining results of patient or mouse tissues were quantified using a positive cell counting method. The percentage of positively stained cells was classified into four grades, each given a score: (1) 0 points (1%-25% positive cells); (2) 1 point (26%-50% positive cells); (3) 2 points (51%-75% positive cells); and (4) 3 points (76%-100% positive cells).

Immunofluorescence

GC cells were seeded at a density of 1 × 10⁵ cells /well in a 24-well plate (TCP010024; Jet BioFil, Guangzhou, Guangdong Province, China) and incubated at 37 °C for 24-36 hours. The cells were then washed thrice with phosphate buffered saline (PBS) (G4202-500ML; Servicebio, Wuhan, Hubei Province, China) and fixed with 4 g/L PFA for 30 minutes. The cells were then permeabilized with 0.5 mL/L Triton X-100 (GC204003-100ML; Servicebio, Wuhan, Hubei Province, China) at room temperature for 20 minutes. The cells were then incubated with 50 mL/L goat serum (G1208-5ML; Servicebio, Wuhan, Hubei Province, China) for 30 minutes to block non-specific binding and prevent background fluorescence interference.

Cell cycle and apoptosis

Cell cycle analysis: (1) The cells were trypsinized, centrifuged; and (2) Washed thrice with PBS. Cells were then fixed in 750 mL/L ethanol (80176961; Sinopharm Chemical Reagent Co., Ltd., Shanghai, China) and then incubated with propidium iodide staining solution (C6031 L; UElandy, Suzhou, Jiangsu Province, China) for 30 minutes duration at room temperature in the dark. Cell cycle distribution was then analyzed using flow cytometry.

Apoptosis analysis: (1) Cells were trypsinized centrifuged; (2) Washed with PBS; (3) Stained with annexin V fluorescein isothiocyanate and propidium iodide (F6012 L; UElandy, Suzhou, Jiangsu Province, China) according to the manufacturer’s instructions; and (4) Followed by flow cytometry analysis.

Cell proliferation assay

To evaluate cell proliferation, multiple methods were employed, including the cell counting kit-8 (CCK-8) assay, 5-ethynyl-2-deoxyuridine (EdU) assay, and colony formation experiments. For the CCK-8 test, transfected cell samples were plated at a concentration of 5 × 10³ cells per well in a 96-well plate (TCP010096; Jet BioFil, Guangzhou, Guangdong Province, China). The CCK-8 solution (C6005M; UElandy, Suzhou, Jiangsu Province, China) was administered at 6 hours, 24 hours, 48 hours, and 72 hours after plating. Following a 2-hour incubation period at 37 °C, the optical density at 450 nm was determined to assess cell growth. For the EdU experiment, detection of DNA replication was conducted using an EdU detection kit (C6043M; UElandy, Suzhou, Jiangsu Province, China) following the manufacturer’s guidelines, and fluorescent microscopy was used to observe cells that were EdU-positive. For the colony formation experiment, cells were plated in a six-well culture plate (TCP011006; Jet BioFil, Guangzhou, Guangdong Province, China) at a seeding density of 1000 cells per well and then incubated for a two-week period. These cultured cells were subjected to fixation with 4 g/L PFA and then stained using 1 g/L crystal violet (G1062; Solarbio, Beijing, China). To assess cellular growth activity, the cell clusters were imaged, and the quantity of round-shaped clusters was enumerated.

Cell migration and invasion assay

To assess cell capabilities in invasion and migration, transwell-based tests for migration and invasion were employed, respectively. The upper chambers were pretreated through coating with Matrigel (356234; Solarbio, Beijing, China), which was diluted in a 1:8 ratio. For the invasion experiments and migration tests, cells were plated at a concentration of 2 × 10⁴ cells per well in the top chamber (TCS018024; Jet BioFil, Guangzhou, Guangdong Province, China) using medium without serum, whereas the bottom chamber was supplied with 20 mL/L fetal bovine serum. Following a 48-hour incubation period, the cells were immobilized using a 4 g/L PFA solution and dyed with 1 g/L crystal violet dye. Subsequently, the treated cells were observed and enumerated under a microscope.

Cell migratory capacity was evaluated via a scratch wound assay. For this purpose, cell cultures were plated in six-well plates at a seeding density of 60000 cells per well. After the cells developed into a completely confluent sheet, a scratch was generated with the tip of a sterile 200 μL pipette (T-200 L; Servicebio, Wuhan, Hubei Province, China). Afterwards, the cell culture was continued in medium without serum, and the process of wound healing was examined using a microscope at the 0 hour and 24 hours time points.

Subcutaneous tumorigenicity model in nude mice

Hangzhou Resource Company supplied 18 four-week-old female BALB/c nude mice, each with a body weight ranging from 18 g to 21 g. All these animals were maintained in a specific pathogen-free environment. These mice were randomly allocated into three experimental groups: (1) Empty vector group (vector); (2) OE-TPM3 group; and (3) OE-TPM3 + sh-YWHAG group. For each group, stably transfected cell lines in the exponential growth stage were subjected to trypsin treatment and then re-suspended in a medium without serum. After disinfecting the skin of nude mice using alcohol, a 1 mL sterile syringe was used to aspirate 150 μL of this cell suspension, which was then subcutaneously administered into the right inguinal area of each mouse. Once tumors were formed, tumor diameter and body weight were measured every two days. The tumor volume was calculated using the following formula: V = 0.52 × length × width². About 40 days following the injection procedure, the animals were humanely sacrificed via CO₂ exposure. Subsequently, the neoplasms were dissected, imaged, weighed, and then preserved in a tissue fixative to facilitate immunohistochemical and HE staining procedures.

Tail vein injection lung metastasis model in nude mice

The nude mice used in this experiment were sourced and housed under the same conditions as described previously. The mice were randomly divided into three groups: (1) Empty vector (vector); (2) OE-TPM3; and (3) OE-TPM3 + sh-YWHAG. Stable transfected cells in the logarithmic growth phase were trypsinized and resuspended in PBS to a final concentration of 1 × 10⁷ cells/mL. The nude mice were restrained on an operating table, and their tail veins were disinfected with alcohol. Using a sterile 1 mL syringe, 150 μL of the cell suspension was carefully injected into the tail vein, with the tail held in position. Subsequently, the needle was gently penetrated along the surface of the skin to locate the vein. The mice were monitored for general conditions, including activity, feeding, and behavior, and body weight was recorded every other day. Four weeks post-injection, the mice were euthanized by CO₂ inhalation, and the lung tissues were excised, photographed, and fixed for HE staining. All animal experiments were approved by the Institutional Animal Care and Use Committee of the Nanchang Royo Biotechnology Co., Ltd. (No. RYE2025021202).

HE staining

Lung specimens freshly obtained from nude mice underwent paraffin embedding and were cut into sections with a thickness of 4 μm. These sections were subsequently placed in an oven for a 1-hour baking process. Following this baking step, the sections underwent sequential deparaffinization using xylene I and II (10023418; Sinopharm Chemical Reagent Co., Ltd., Shanghai, China). These sections were rehydrated via a series of ethanol solutions of decreasing concentration, and subjected to hematoxylin staining (G1004-100ML; Servicebio, Wuhan, Hubei Province, China). For the differentiation process, hydrochloric acid ethanol (80070560; Sinopharm Chemical Reagent Co., Ltd., Shanghai, China) was utilized, followed by eosin staining (G1002-100ML; Servicebio, Wuhan, Hubei Province, China). Next, the slides were subjected to dehydration via a sequence of ethanol solutions with increasing concentration, and then clearing was performed using xylene I and II. Subsequently, neutral resin (WG10004160; Servicebio, Wuhan, Hubei Province, China) was applied to mount the slides, which were then examined under a microscope.

Statistical analysis

The study’s data analysis utilized GraphPad Prism (version 8.0.1) and R Studio (version 4.0.3) software. Student’s t-test evaluated the significance of differences between the two groups. All experiments were independently repeated thrice (n = 3). A P < 0.05 was considered statistically significant.

RESULTS

TPM3 is upregulated in GC and correlates with poor prognosis

We examined TPM3’s involvement in GC by assessing its mRNA and protein expression using bioinformatics and experimental data. Additionally, we assessed the relationship between TPM3 expression and patient outcomes by analyzing clinical data. The results of the bioinformatics analysis are summarized below.

First, we analyzed TPM3 expression across multiple human tumors using the Tumor Immune Estimation Resource database. The results revealed that TPM3 was expressed at higher levels in most tumors, including GC, compared to normal tissues (Figure 1A). We further integrated data from TCGA and GTEx databases using the GEPIA platform (version 2.0) to assess differential TPM3 mRNA expression. This analysis confirmed that TPM3 was significantly upregulated in GC samples (Figure 1B). Analysis of GSE54129 and GSE66229 datasets from the Gene Expression Omnibus database also demonstrated that TPM3 expression was significantly higher in GC specimens compared to adjacent normal tissues (Figure 1C). Patients with high TPM3 expression demonstrated significantly poorer OS in Kaplan-Meier Plotter analysis compared to those with low TPM3 expression (P < 0.0001; Figure 1D), suggesting that increased TPM3 levels correlate with a poor prognosis in GC. Moreover, an examination conducted via the UALCAN database revealed that TPM3 expression was positively associated with patients’ age, gender, tumor stage, histological type, lymph node involvement status, as well as TP53 mutation status in GC (Supplementary Figure 1).

Figure 1 Tropomyosin 3 is upregulated in gastric cancer sample and associated with poor prognosis. A: Differential expression analysis of tropomyosin 3 (TPM3) in multiple tumors from Tumor Immune Estimation Resource database; B: Comparative analysis of the expression differences of TPM3 in normal tissues and gastric cancer (GC) tissues using the GEPIA2.0 database; C: Gene Expression Omnibus databases (GSE54129 and GSE66229) analysis of TPM3 mRNA expression in normal tissues and GC tissues; D: Analyze the difference in overall survival between the high-expression group and the low-expression group of TPM3 on the Kaplan-Meier plotter website; E: Quantitative real-time PCR analysis of TPM3 mRNA expression in 62 pairs of tumorous and para-carcinoma tissue from the Second Affiliated Hospital of Nanchang University; F: Western blotting showed the protein expression of TPM3 in tumorous and para-carcinoma tissue (it shows that TPM3 protein expression in GC tissues presents as dual bands, representing different subtypes); G: Immunohistochemistry showed the protein expression of TPM3 in tumorous and para-carcinoma tissue; H: Overall survival of GC patients with low or high levels of TPM3; I: Protein and mRNA expression of TPM3 in GC cell lines. ^aP < 0.05, ^bP < 0.01, ^cP < 0.001, ^dP < 0.0001. HR: Hazard ratio; STAD: Stomach adenocarcinoma; TPM3: Tropomyosin 3.

Second, we collected 62 pairs of GC samples from the Department of Gastrointestinal Surgery at the Second Affiliated Hospital of Nanchang University for analysis. The qRT-PCR analysis revealed that TPM3 mRNA levels were notably elevated in GC tissues compared to the surrounding normal tissues (Figure 1E). WB and immunohistochemistry analyses further demonstrated that TPM3 protein levels were notably elevated in GC tissues compared to adjacent normal tissues (Figure 1F and G). To investigate the association between TPM3 expression and patient outcomes, individuals were divided into high and low expression cohorts according to the median expression value of TPM3. An investigation into how TPM3 expression levels relate to clinicopathological traits was conducted, and the outcomes are summarized in Table 1. Specifically, clinical observations indicated that elevated TPM3 expression levels were significantly correlated with tumor T classification, tumor node metastasis classification, and pathological differentiation grade (Table 1). Kaplan-Meier survival analysis of patient follow-up data further demonstrated that those with elevated TPM3 expression exhibited notably poorer OS (Figure 1H). Collectively, the results from the analyses suggest that TPM3 is notably overexpressed in GC and strongly linked to prognostic outcomes in patients.

Table 1 The association between tropomyosin 3 expression and clinicopathological, n (%).

Characteristics	Expression of TPM3		P value
	Low expression of TPM3 (n = 31)	High expression of TPM3 (n = 31)
Age			0.607
≤ 60	14 (22.58)	12 (19.35)
> 60	17 (27.42)	19 (30.65)
Gender			0.799
Male	16 (25.81)	17 (27.42)
Female	15 (24.19)	14 (22.58)
T stage			0.039
Tis-2	17 (27.42)	9 (14.52)
T3-4	14 (22.58)	22 (35.48)
N stage			0.103
N0	13 (20.97)	7 (11.29)
N1-3	18 (29.03)	24 (38.71)
Distant metastasis			0.783
Negative	22 (35.48)	21 (33.87)
Positive	9 (14.52)	10 (16.13)
Tumor node metastasis stage			0.037
I and II	16 (25.81)	8 (12.90)
III and IV	15 (24.19)	23 (37.10)
Tumor differentiation			0.039
Moderate-well	17 (27.42)	9 (14.52)
Poor	14 (22.58)	22 (35.48)

TPM3: Tropomyosin 3.

TPM3 promotes the proliferation, migration, and invasion of GC cells in vitro

Previous studies have demonstrated that TPM3 is expressed at high levels in GC cells and has a strong correlation with patient prognosis. The prognosis for individuals with cancer is frequently linked to changes in the proliferation, invasion, and migration of tumor cells[29]. Consequently, we assessed the effects of TPM3 on GC cell proliferation, invasion, and migration in vitro. To compare TPM3 expression among different GC cell lines, we assessed protein and mRNA levels in MKN74, HGC27, MKN45, and AGS cells using WB and qRT-PCR. The results indicated that TPM3 expression was highest in AGS cells and lowest in HGC27 cells, at the protein and mRNA levels (Figure 1I). Following these results, AGS and HGC27 cell lines were chosen for further experiments. Short interfering RNA encoded by a lentivirus was used to silence TPM3 expression. After transfection, viral transfection efficiency was evaluated using WB and qRT-PCR. In AGS cells, the knockdown efficiency of sh-TPM3#1 and sh-TPM3#2 was higher than that of sh-TPM3#3. Conversely, in HGC27 cells, OE-TPM3 overexpression was more effective than empty vector control (Figure 2A). Consequently, subsequent experiments were conducted using sh-TPM3#1, sh-TPM3#2, and OE-TPM3.

Figure 2 Tropomyosin 3 promotes proliferation, migration and invasion of gastric cancer cells. A: Validation of tropomyosin 3 (TPM3) knockdown and overexpression efficiency; B-D: Cell counting kit-8 assay, colony formation assay, and 5-ethynyl-2-deoxyuridine assay were conducted to evaluate the effect of TPM3 knockdown or overexpression on the proliferative capacity of gastric cancer cells; E and F: Transwell assay and wound healing assay were performed to assess the impact of TPM3 knockdown or overexpression on the migration and invasion ability of gastric cancer cells. ^bP < 0.01, ^cP < 0.001, ^dP < 0.0001. EdU: 5-ethynyl-2-deoxyuridine; sh-NC: Control shRNA; TPM3: Tropomyosin 3; Vector: Empty vector control.

We assessed the effect of TPM3 on GC cell proliferation using CCK-8, colony formation, and EdU assays. The results demonstrated that inhibiting TPM3 significantly hindered AGS cell growth, whereas enhancing TPM3 expression promoted HGC27 cell proliferation (Figure 2B-D). The impact of TPM3 on the migration of GC cells was evaluated through transwell migration and wound-healing assays. The results revealed that a decrease in TPM3 resulted in a significant reduction in AGS cell migration, whereas TPM3 overexpression enhanced the migration of HGC27 cells (Figure 2E and F). Finally, transwell invasion assays were used to evaluate how TPM3 affects GC cell invasiveness. The results indicated that TPM3 silencing decreased the number of AGS cells invading the Matrigel-coated chambers (Figure 2E). Collectively, these results indicate that TPM3 enhances proliferation, migration, and invasion of GC cells.

TPM3 regulates cell cycle and suppresses apoptosis in GC cells

Previous findings have indicated that TPM3 promotes the proliferation, motility, and invasive capabilities of GC cells. Prior research has identified connections among cancer cell growth, cell cycle irregularities, and apoptosis, whereby apoptosis acts to suppress growth[30]. Consequently, an investigation was conducted into the impact of TPM3 expression on cell cycle advancement and apoptosis in GC cells.

First, cell cycle distribution and apoptosis were examined using flow cytometry. Compared to the control group, AGS cells displayed significant changes with TPM3 knockdown (sh-TPM3#1 and sh-TPM3#2), as more cells were found in the S phase and fewer in the G1 phase. Conversely, OE-TPM3 was significantly reduced in the S phase population and an increase in the G1 phase cell proportion in HGC27 cells compared to the vector group (Figure 3A and B). Additionally, TPM3 silencing significantly increased the proportion of apoptotic AGS cells, whereas TPM3 overexpression reduced the apoptosis of HGC27 cells (Figure 3C and D). WB revealed that the expression of cell cycle and apoptosis-related proteins CDK4, CDK6, Cyclin D1, and Bax decreased in sh-TPM3#1 and sh-TPM3#2 groups, whereas BCL2 expression was upregulated (Figure 3E and Supplementary Figure 2A). Collectively, these results indicate that TPM3 silencing inhibits GC progression by blocking the G1/S phase transition and promoting apoptosis, whereas TPM3 overexpression exerts the opposite effect.

Figure 3 Tropomyosin 3 facilitates cell cycle progression and suppresses apoptosis in gastric cancer cells. A: Flow cytometry analysis of cell cycle distribution in AGS and HGC27 cells following tropomyosin 3 knockdown or overexpression; B: Quantification of cell cycle phase distribution (G1, S, and G2) in AGS and HGC27 cells; C: Flow cytometry analysis of apoptosis using annexin V fluorescein isothiocyanate/propidium iodide staining in AGS and HGC27 cells following tropomyosin 3 knockdown or overexpression; D: Quantification of apoptosis rate in AGS and HGC27 cells; E: Western blotting analysis of proteins associated with cell cycle and apoptosis; F: Western blotting analysis was performed to examine the expression levels of epithelial-mesenchymal transition-related proteins and matrix metalloproteinase family proteins. ^bP < 0.01, ^cP < 0.001, ^dP < 0.0001. NS: Not significant; FITC: Fluorescein isothiocyanate; OE: Overexpression plasmid; sh-NC: Control shRNA; TPM3: Tropomyosin 3; Vector: Empty vector control.

TPM3 promotes EMT in GC cells

The progression of tumors relies on the MAPK signaling pathway, particularly in advancing EMT. Consequently, we investigated whether TPM3 affects EMT in GC cells. WB demonstrated that in AGS cells, TPM3 knockdown (sh-TPM3#1 and sh-TPM3#2) significantly decreased the protein levels of N-cadherin and vimentin, while increasing E-cadherin expression. Conversely, in HGC27 cells, TPM3 overexpression (OE-TPM3) significantly reduced E-cadherin levels and increased N-cadherin and vimentin expression (Figure 3F and Supplementary Figure 2B). These findings indicate that TPM3 effectively promotes EMT in GC cells.

TPM3 regulates the MAPK signaling pathway to promote GC progression

To examine the biological function of TPM3 in GC progression, transcriptome sequencing was conducted on AGS cells from the sh-TPM3#1, sh-TPM3#2, and control groups, with subsequent GO and KEGG enrichment studies. GO enrichment analysis suggested that focal adhesion may represent a potential mechanism through which TPM3 affects GC progression (Figure 4A). Additionally, KEGG analysis revealed significant enrichment of the MAPK signaling pathway (Figure 4B). Previous research has reported that CPNE8 drives GC metastatic processes via the cell adhesion-related pathway, with several core genes in this pathway being closely associated with enhanced metastatic capacity and unfavorable clinical outcomes in GC[31,32]. Based on these observations regarding signaling pathways in GC, it was hypothesized that TPM3 modulates GC development through the MAPK signaling pathway. To test this hypothesis, WB analysis was performed. These analyses revealed that knockdown of TPM3 significantly decreased the expression levels of p-extracellular signal-regulated kinase (ERK), p-P38, and p-c-Jun N-terminal kinase (JNK) proteins (Figure 4C and Supplementary Figure 2C).

Figure 4 Tropomyosin 3 enhances the mitogen-activated protein kinase signaling pathway in gastric cancer cells. A and B: Gene Ontology and Kyoto Encyclopedia of Genes and Genomes enrichment analyses showed that tropomyosin 3 was involved in gastric cancer progression via the mitogen-activated protein kinase pathway; C: Western blotting analysis of tropomyosin 3 knockdown or overexpression effects on the mitogen-activated protein kinase pathway. Akt: Protein kinase B; BP: Biological process; CC: Cellular component; ERK: Extracellular signal-regulated kinase; GO: Gene Ontology; JAK: Janus kinase; JNK: C-Jun N-terminal kinase; KEGG: Kyoto Encyclopedia of Genes and Genomes; MAPK: Mitogen-activated protein kinase; MF: Molecular function; PI3K: Phosphoinositide 3-kinase; sh-NC: Control shRNA; STAT: Signal transducer and activator of transcription; TNF: Tumor necrosis factor; TPM3: Tropomyosin 3; Vector: Empty vector control.

TPM3 can interact with YWHAG

Our study aimed to identify the intermediary molecules through which TPM3 regulates GC cell growth. Using immunoprecipitation combined with mass spectrometry analysis, we identified 1,158 proteins that potentially interact with TPM3. Among these, YWHAG emerged as a protein of particular interest. Using the GEPIA database (version 2.0), correlation analysis exhibited a strong positive link between TPM3 and YWHAG expression (Figure 5A). Furthermore, analysis of GSE54129 and GSE66229 datasets from the Gene Expression Omnibus database demonstrated that YWHAG expression was notably higher in GC tissues compared to adjacent normal tissues (Figure 5B). Consistently, GEPIA analysis (version 2.0) confirmed that YWHAG was significantly overexpressed in GC tissues (Figure 5C). Moreover, WB and qRT-PCR analyses indicate that the expression levels of YWHAG protein and mRNA were significantly increased in GC tissues compared to adjacent normal tissues (Figure 5D and E).

Figure 5 Tropomyosin 3 interacts with tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma in gastric cancer cells and positively regulates the expression of tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma. A: Tropomyosin 3 (TPM3) was significantly positively correlated with tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma (YWHAG) in The Cancer Genome Atlas cohort from GEPIA 2.0 database; B: Gene Expression Omnibus databases (GSE54129 and GSE66229) analysis of YWHAG mRNA expression in normal tissues and gastric cancer (GC) tissues; C: Comparative analysis of the expression differences of YWHAG in normal tissues and GC tissues using the GEPIA2.0 database; D: Quantitative real-time PCR analysis of YWHAG mRNA expression in 62 pairs of tumorous and para-carcinoma tissue from the Second Affiliated Hospital of Nanchang University; E: Western blotting showed the protein expression of YWHAG in tumorous and para-carcinoma tissue; F: Co-immunoprecipitation indicated the presence of direct or indirect binding between TPM3 and YWHAG; G: The colocalization between TPM3 and YWHAG was visualized by a confocal laser scanning microscope in different GC cell lines; H: Western blotting analysis was performed to observe the expression changes of YWHAG following TPM3 knockdown or overexpression; I: Quantitative real-time PCR analysis was performed to examine the expression changes of YWHAG following TPM3 knockdown or overexpression. ^aP < 0.05, ^bP < 0.01, ^cP < 0.001, ^dP < 0.0001. NS: Not significant; Co-IP: Co-immunoprecipitation; IgG: Immunoglobulin G; sh-NC: Control shRNA; STAD: Stomach adenocarcinoma; TPM3: Tropomyosin 3; YWHAG: Tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma; Vector: Empty vector control.

Co-immunoprecipitation assays revealed a direct protein–protein interaction between TPM3 and YWHAG in GC cells (Figure 5F). Furthermore, WB and qRT-PCR analyses of the four GC cell lines demonstrated that YWHAG expression was highest in HGC27 cells and lowest in AGS cells at the protein level (Supplementary Figure 3A). These results demonstrated a consistent pattern at the mRNA level, indicating concordance between transcriptional and protein expression. Immunofluorescence experiments in GC cells confirmed that TPM3 and YWHAG were localized in the cytoplasm (Figure 5G). WB demonstrated that in AGS cells, TPM3 knockdown decreased YWHAG protein expression, whereas in HGC27 cells, TPM3 overexpression increased YWHAG protein levels (Figure 5H and Supplementary Figure 3B). Notably, qRT-PCR analysis revealed no significant changes in YWHAG mRNA expression between the groups (Figure 5I). Based on YWHAG expression profiles across the four GC cell lines, HGC27 and AGS cells were selected for subsequent experiments. WB demonstrated that YWHAG knockdown or overexpression non-significantly affects TPM3 protein levels. Moreover, qRT-PCR analysis indicated no significant differences in TPM3 mRNA expression levels (Supplementary Figure 4A).

TPM3 affects proliferation, migration, and invasion of GC in vitro through YWHAG modulation

From prior experimental data, it was hypothesized that YWHAG modulates TPM3's regulatory role in cancer progression. To confirm that YWHAG acts downstream of TPM3, rescue experiments were conducted by overexpressing YWHAG in TPM3-knockdown cells. Cellular functional analyses, such as CCK-8, colony formation, and EdU assays, demonstrated that overexpressing YWHAG restored cell proliferation to levels comparable to the negative control. These results indicate that the inhibitory effect of TPM3 silencing on cell proliferation relies on YWHAG. Conversely, in stable cell lines with TPM3 overexpression, silencing YWHAG counteracted the proliferative impact caused by TPM3 overexpression in GC cells (Figure 6A-C). Transwell invasion and migration experiments further revealed that elevated YWHAG levels restored the compromised migratory and invasive capacities of GC cells resulting from TPM3 silencing. Conversely, YWHAG suppression notably reduced the strengthened migratory and invasive characteristics induced by TPM3 overexpression (Figure 6D). Consistent results were observed in wound-healing experiments. This evidence indicated that the migration impairment caused by TPM3 silencing could be partially alleviated through YWHAG upregulation, while the migratory promotion resulting from TPM3 overexpression was partially counteracted via YWHAG depletion (Figure 6E). Such findings suggest that TPM3 demonstrates a positive regulatory effect on YWHAG, thereby enhancing the proliferation, migration, and invasion of GC cells.

Figure 6 Tropomyosin 3 influences proliferation, migration, and invasion of gastric cancer cells by regulating tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma. A-C: Cell counting kit-8 assay, colony formation assay, and 5-ethynyl-2-deoxyuridine assay were used to compare the changes in cell proliferation ability across different gastric cancer cell lines; D and E: Transwell assay and wound healing assay were used to compare the changes in cell migration and invasion abilities across different gastric cancer cell lines. ^aP < 0.05, ^bP < 0.01, ^cP < 0.001, ^dP < 0.0001. NS: Not significant; EdU: 5-ethynyl-2-deoxyuridine; sh-NC: Control shRNA; TPM3: Tropomyosin 3; YWHAG: Tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma; Vector: Empty vector control.

TPM3 regulates cell cycle, apoptosis, and EMT in GC cells through modulation of YWHAG and also controls the MAPK signaling pathway

To further explore whether TPM3 impacts the cell cycle, apoptosis, and EMT in GC cells via YWHAG, a WB analysis was performed. This analysis revealed that heightened YWHAG expression promoted tumor molecular profile. Notably, YWHAG expression increased levels of cell cycle-related proteins (CDK4, CDK6, and CyclinD1), the apoptosis-promoting protein Bax, and EMT-associated markers (N-cadherin and vimentin), while reducing expression of the apoptosis-inhibiting protein BCL2 and the epithelial marker E-cadherin. Notably, the molecular alterations triggered by YWHAG were significantly counteracted to levels comparable with those of the control cohort when TPM3 was silenced. Conversely, overexpressing TPM3 restored the protein expression alterations triggered by YWHAG depletion (Figure 7A and Supplementary Figure 4B). In a previous investigation, Tian and team found that YWHAG boosts TMOD3 expression, sets off the MAPK pathway, and strengthens the invasive and metastatic potential of bladder cancer[27]. Based on these results, it was put forward that YWHAG controls the MAPK signaling pathway, and this control is affected by TPM3. WB analysis indicated that silencing YWHAG in HGC27 cells significantly decreased the expression levels of p-ERK, p-P38, and p-JNK, while co-overexpressing TPM3 counteracted this reduction. Similarly, in AGS cells, overexpressing YWHAG diminished the levels of proteins in the MAPK signaling pathway that resulted from TPM3 depletion (Figure 7B and Supplementary Figure 5A). Furthermore, when HGC27 cells were treated with three separate MAPK pathway blockers, the impacts induced by TPM3 overexpression were counteracted (Figure 7C and Supplementary Figure 5B). These findings indicate that TPM3 promotes cell cycle and EMT while suppressing apoptotic processes in GC cells via YWHAG. Moreover, the signaling pathway consisting of TPM3, YWHAG, and MAPK is vital for driving the cancerous progression of GC cells.

Figure 7 Tropomyosin 3 influences cell cycle, apoptosis, matrix metalloproteinase family proteins, and epithelial-mesenchymal transition in gastric cancer cells through tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma, and activates the mitogen-activated protein kinase signaling pathway. A and B: Western blotting analysis confirmed that tropomyosin 3 influences the cell cycle, apoptosis, matrix metalloproteinase family proteins, and epithelial-mesenchymal transition in gastric cancer cells through tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma, and activates the mitogen-activated protein kinase signaling pathway; C: Western blotting analysis of HGC27 cells overexpressing tropomyosin 3, followed by tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma knockdown or treatment with mitogen-activated protein kinase pathway inhibitors, including SCH772984 (extracellular signal-regulated kinase inhibitor, 20 μM), SB203580 (p38 inhibitor, 20 μM), and SP600125 (c-Jun N-terminal kinase inhibitor, 20 μM). ERK: Extracellular signal-regulated kinase; JNK: C-Jun N-terminal kinase; OE: Overexpression plasmid; MMP: Matrix metalloproteinase; TPM3: Tropomyosin 3; YWHAG: Tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma.

YWHAG knockdown in vivo alleviates the malignant progression of GC cells induced by TPM3 overexpression

For a deeper exploration of the in vivo roles of TPM3 and YWHAG, researchers employed subcutaneous tumor models and tail vein lung metastasis models in nude mice. For the subcutaneous tumor model, HGC27 cells were engineered via transfection with TPM3 upregulation vectors (OE-TPM3), OE-TPM3 paired with YWHAG suppression constructs (sh-YWHAG), or a control plasmid, and then inoculated into nude mice. Compared to the reference cohort, the TPM3-overexpressing group demonstrated increased tumor mass and the speed of volume expansion, while concurrent suppression of YWHAG significantly reduced such outcomes (Figure 8A and B). Based on immunohistochemistry assessment of subcutaneous tumor specimens, Ki-67 protein levels were notably higher in the OE-TPM3 group compared to the control group. Conversely, YWHAG suppression significantly decreased the Ki-67 expression (Figure 8C). Histopathological analysis using HE staining revealed that tumors in the OE-TPM3 group exhibited the greatest size, while YWHAG silencing reduced tumor development (Figure 8D).

Figure 8 Tropomyosin 3 regulates gastric cancer cell progression through tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma in vivo. A and B: Tumor volumes and weights measured at the endpoint; C: Immunohistochemical analysis showed the expression of Ki-67 in xenograft tumor tissues from different groups; D: Hematoxylin and eosin staining analysis revealed the tumor tissues in different groups; E and F: Fresh lung tissues were harvested from mice intravenously injected with HGC27 cells through the tail vein, including the control group, tropomyosin 3 (TPM3) overexpression group, and TPM3 overexpression + tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma knockdown group. The number of lung metastatic nodules was then counted and hematoxylin and eosin staining was used to observe the lung metastatic nodules in different groups. Primary tumor samples were collected from mice injected with HGC27 cells, including the control group, TPM3 overexpression group, and TPM3 overexpression + tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma knockdown group. ^aP < 0.05, ^cP < 0.001, ^dP < 0.0001. NS: Not significant; TPM3: Tropomyosin 3; YWHAG: Tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein gamma; Vector: Empty vector control.

For the pulmonary metastasis model established through tail vein delivery, identically transfected HGC27 cell lines were introduced into the tail veins of nude mice. After 30 days, widespread metastatic growth was observed in the pulmonary tissues of the OE-TPM3 group, while metastatic lesions were significantly diminished in the OE-TPM3 combined with sh-YWHAG group (Figure 8E). Similarly, in the pulmonary metastasis model, the volume of metastatic lesions in the OE-TPM3 cohort was significantly larger when contrasted with the control group, and this augmentation was attenuated through YWHAG silencing (Figure 8F). Collectively, these findings demonstrate that reducing YWHAG expression within living organisms lessens the cancerous development of GC cells brought about by TPM3 overexpression.

DISCUSSION

GC is a common malignancy of the digestive system. Identifying effective therapeutic targets is crucial for improving GC prevention and treatment. Moreover, demonstrating the molecular mechanisms underlying GC is essential for the discovery of prognostic biomarkers. These biomarkers help monitor tumor recurrence, assess prognosis, and identify potential therapeutic targets, thereby supporting the development of personalized treatment approaches for patients with GC.

Our research explored the roles of TPM3 and YWHAG in GC, both of which are widely studied in various diseases, particularly cancer. While TPM1 functions as a tumor suppressor by inhibiting malignant progression, TPM3 frequently acts as an oncogene in non-muscle tissue tumors[33]. Accumulating evidence indicates that TPM3 promotes tumorigenesis and cancer progression in multiple malignancies. For instance, in cervical cancer, high TPM3 expression significantly accelerates malignant tumor progression, and patients with elevated TPM3 levels exhibit a poor prognosis. Furthermore, TPM3 expression can effectively differentiate between different stages of cervical cancer[34]. Similarly, in liver cancer cell lines, TPM3 amplification induces EMT and downregulates E-cadherin, promoting invasion and metastasis, correlated with worse clinical outcomes[35]. Additionally, patients with glioma and high TPM3 expression exhibit poorer prognosis[15]. Consistently, in our study, TPM3 was highly expressed in GC tissues, and elevated TPM3 levels were significantly associated with poor OS. This strong correlation suggests that TPM3 may function as a prognostic biomarker and a potential therapeutic target in GC. Additionally, functional experiments demonstrated that TPM3 significantly promoted GC cell proliferation, while TPM3 knockdown increased cell apoptosis. Moreover, TPM3 inhibition impairs cell proliferation by arresting the cell cycle at the G1-to-S phase transition. WB revealed that TPM3 knockdown decreased the expression of key cell cycle regulators, including CDK4, CDK6, and Cyclin D1. These results indicated that TPM3 may act as a crucial effector directly controlling GC cell cycle progression. Moreover, TPM3 doublets were observed exclusively in tissue samples, whereas GC cell lines consistently exhibited a single band. Database analysis from national center for biotechnology information confirmed the presence of 17 distinct TPM3 gene isoforms in human tissues. We propose that this variability reflects the greater complexity of clinical specimens, where TPM3 may exist in multiple isoforms or undergo distinct post-translational modifications, the specific roles of which remain to be explored. Research has also confirmed that TPM3 can form fusion genes with other partners, resulting in cytoskeletal alterations that promote cell invasion and metastasis, thereby driving tumorigenesis and cancer progression[36]. As an actin-associated protein, TPM3 directly regulates cell morphology and motility, providing a fundamental driving force for cell migration and invasion[5,37]. Similarly, transwell experiments revealed that silencing TPM3 significantly reduced the migration and invasion abilities of GC cells, consistent with its well-established role in cytoskeletal regulation. Based on these findings, we propose a potential synergistic mechanism in which TPM3-mediated cytoskeletal remodeling establishes the cellular state that facilitates GC cell migration and invasion. Subsequently, the downstream YWHAG/MAPK signaling axis amplifies and sustains this malignant phenotype.

In such a scenario, the family of matrix metalloproteinases (MMPs) plays a critical role in cancer development. MMP2 can hydrolyze components of the intercellular matrix and type IV collagen in the basement membrane, whereas MMP9 can degrade the extracellular matrix or remodel its dynamic balance[38]. Chen et al[14] demonstrated that TPM3 promotes the migration and invasion of esophageal squamous carcinoma cells by regulating MMP2 and MMP9, thereby inducing EMT. Consistently, our GC study demonstrated that TPM3 similarly enhances EMT, migration, and proliferation of GC cells, accompanied by upregulation of MMP2 and MMP9. Collectively, these studies underscore the diverse functions of TPM3 in regulating tumor progression across the cellular and extracellular components of the tumor microenvironment.

The 14-3-3 protein family consists of dimeric phosphoserine-binding proteins that interact with numerous signaling proteins through their phosphoserine-binding domains[39]. For instance, in the Hippo signaling pathway, 14-3-3 proteins bind to the tumor suppressors YAP/TAZ, preventing their nuclear translocation and inhibiting their transcriptional activity, thereby contributing to sustained tumor cell growth[40]. YWHAG, a key member of the 14-3-3 family, has attracted increasing attention due to its crucial role in tumor pathogenesis. Multiple studies have reported that YWHAG is consistently upregulated in various tumor cell lines and metastatic tissues. For instance, in pancreatic cancer, CERS6-AS1 functions as a competitive endogenous RNA for miR-217, leading to YWHAG upregulation and activation of the threonine kinase-1 (RAF1)/ERK signaling pathway[22]. In colorectal carcinoma, YWHAG modulates the CTTN-Wnt/β-catenin signaling cascade, which facilitates cell growth, invasion, and metastasis[27]. In our study, mass spectrometry analysis and co-immunoprecipitation experiments identified a direct interaction between TPM3 and YWHAG, and we confirmed that TPM3 positively regulates YWHAG at the protein level. Immunofluorescence further demonstrated that TPM3 and YWHAG co-localized in the cytoplasm of GC cells, suggesting a potential functional coupling that may contribute to the rewiring of oncogenic pathways in GC. To validate YWHAG as a critical downstream mediator of TPM3-driven GC progression, we conducted a reciprocal genetic rescue experiment. These experiments confirmed that the TPM3/YWHAG axis positively regulates GC cell proliferation and invasion. Consistent with previous findings in other cancers, such as endometrial cancer, MMP2 and MMP9 appear to act downstream facilitate tumor cell invasion and migration[41]. Our previous data demonstrated that TPM3 knockdown reduces the expression of the invasion-related proteases MMP2 and MMP9. Notably, this reduction was rescued by YWHAG overexpression, confirming that YWHAG acts downstream of TPM3. Collectively, these complementary genetic manipulations support the conclusion that the TPM3/YWHAG axis promotes GC cell invasion, at least in part, by upregulating MMP2 and MMP9. Notably, our preliminary experiments revealed that altering TPM3 expression led to corresponding changes in YWHAG protein levels, whereas YWHAG mRNA levels remained largely unchanged. These observations suggest that TPM3 may regulate YWHAG via post-translational mechanisms, the precise nature of which requires further investigation.

The onset and malignant progression of tumors are commonly driven by the aberrant activation of two major signaling cascades: (1) Phosphatidylinositol 3-kinase/protein kinase B/mammalian target of rapamycin; and (2) The Ras-RAF-MEK-ERK pathways, also known as the MAPK pathway[42]. The MAPK family comprises three major subgroups: (1) ERKs (ERK MAPKs); (2) The c-Jun N-terminal kinases, also known as stress-activated protein kinases (JNK or SAPK); and (3) The p38 MAPKs[43]. Additionally, ERK1 and ERK2 are referred to as MAPK3 and MAPK1, respectively, whereas the stress-activated MAPKs p38α and p38β correspond to MAPK14 and MAPK11, respectively[42]. Functionally, MAPKs are traditionally classified into mitogen-activated and stress-activated kinases, with ERK primarily mediating mitogenic responses and JNK and p38 mediating stress responses[44]. These MAPK signaling pathways regulate various cellular activities, such as proliferation, differentiation, apoptosis, autophagy and stress responses[45-48]. To demonstrate the downstream pathways regulated by TPM3 in GC cells, we performed mRNA transcriptomic sequencing followed by GO and KEGG enrichment analyses. The results revealed significant enrichment of the MAPK signaling pathway. Based on these findings, further experiments indicated that TPM3 knockdown decreased the levels of phosphorylated MAPK proteins, including p-ERK, p-P38, and p-JNK, whereas TPM3 overexpression led to their upregulation. These results indicate that TPM3 promotes GC development and progression, at least in part, by activating the MAPK signaling pathway.

Early studies have established that activation of the MAPK signaling pathway can induce EMT, a key process in tumor metastasis that enhances the invasive and migratory potential of tumor cells[49]. EMT is the process where epithelial cells change into mesenchymal cells, marked by a reduction in epithelial markers such as E-cadherin and cytokeratins, and an increase in mesenchymal markers such as vimentin and metalloproteinases[47]. Our results demonstrate that TPM3 regulates EMT in GC cells, indicated by increased N-cadherin and vimentin levels and decreased E-cadherin expression. Notably, the ability of TPM3 to drive EMT and MAPK pathway activation was functionally dependent on YWHAG, as genetic manipulation of YWHAG could either rescue or suppress these phenotypes. This establishes YWHAG as a critical downstream effector through which TPM3 coordinates oncogenic signaling and regulates cellular plasticity. Moreover, in our WB experiments, we observed variability in the number of vimentin bands across different treatment groups. Vimentin can appear as either a single band or a doublet, depending on the cell line and experimental conditions, a phenomenon widely reported in the literature. This variability likely reflects state-dependent post-translational modifications and differential susceptibility to proteolysis. For quantitative analysis, the dominant and consistently observed bands were used across all samples.

Based on previous studies, we propose that TPM3 enhances the activation of multiple MAPK pathways by upregulating YWHAG, a central regulatory protein. Prior research has partially demonstrated how the other 14-3-3 family members regulate downstream effectors and target genes of the MAPK pathway. For instance, 14-3-3ζ has been reported to potentiate the MAPK/c-Jun signaling pathway, thereby promoting breast cancer cell proliferation. Conversely, Lam et al[50] demonstrated that in dermal fibroblasts, 14-3-3δ rapidly induces increases in c-jun and c-fos mRNA levels via MAPK signaling[51]. Given the high functional conservation within the 14-3-3 family, YWHAG may function similarly to a 14-3-3 dimer by binding to phosphorylated sites such as pS365 and pS729 on RAF to activate RAF[52]. Moreover, YWHAG may facilitate JNK/p38 activation by binding to and stabilizing upstream kinases, including ASK1 or MEKs[53,54]. Some studies have also reported that certain 14-3-3 proteins can shuttle into the nucleus and directly regulate gene transcription through interactions with transcription factors or epigenetic regulators[55-57]. Accordingly, we speculate that nuclear YWHAG may exert similar regulatory functions in GC, a possibility that warrants further investigation in future studies. Additionally, we propose an intriguing hypothesis that TPM3, as a core cytoskeletal component, may sense intracellular tension changes during processes such as cell adhesion and migration. In response to these mechanical cues, TPM3 may undergo conformational alterations to function as a “signaling protein”. Through interaction with YWHAG, TPM3 may amplify and transduce these mechanical cues into biochemical signals that activate the MAPK pathway. Our findings further revealed that altering YWHAG expression non- significantly affects TPM3 levels, suggesting a unidirectional regulatory relationship that ensures a committed, non-reversible signal flow from the cytoskeletal regulator (TPM3) to the signaling hub/scaffold protein (YWHAG). This prevents signal attenuation or interference from feedback loops and enables more efficient translation of cytoskeletal remodeling into robust pro-proliferative and promigratory signaling outputs. Moreover, TPM3 upregulation may be an early oncogenic event in GC. By inducing the expression of the multifunctional adaptor YWHAG, TPM3 can coordinately activate multiple downstream pathways, including the MAPK cascade, thereby converting a single molecular alteration into an integrated malignant cellular phenotype.

Building on in vitro findings, we extended the functional validation of the TPM3/YWHAG axis to in vivo models using xenograft and metastasis assays. TPM3 overexpression notably promoted tumor growth and lung colonization, effects that were critically dependent on YWHAG, as evidenced by their significant attenuation following YWHAG knockdown. Consistently, immunohistochemical analysis of proliferation (Ki-67) and histopathological assessment of the metastatic burden directly linked the pro-tumorigenic functions of TPM3 to YWHAG-mediated signaling in vivo. Collectively, these results demonstrate that the TPM3/YWHAG axis is necessary and sufficient to drive GC progression in physiologically relevant contexts, solidifying its role as a key mechanistic driver of tumorigenesis and metastasis.

Overall, these data support the conclusion that TPM3 exerts its oncogenic effects in GC primarily by regulating the YWHAG/MAPK axis. Despite these findings, our study has several limitations. First, functional conclusions are mainly based on established cell lines and xenograft models, which may not fully capture the complexity, heterogeneity, and microenvironmental context of human GC. Second, although we have provided evidence that TPM3 regulates YWHAG expression, precise molecular mechanisms, including potential post-translational modifications, remain unclear, leaving the possibility of alternative regulatory pathways. Third, detecting multiple TPM3 isoforms in GC tissues suggests functional diversity that our study, which focused on overall TPM3 activity, could not fully dissect. Finally, reported clinical associations are retrospective; prospective validation in independent patient cohorts is imperative to establish TPM3’s translational potential as a biomarker or therapeutic target. These limitations emphasize directions for future research and provide an essential context for interpreting the scope of our current findings.

CONCLUSION

Overall, the current research revealed that TPM3 acts as a tumor promoter in GC. The research initially confirmed that TPM3 is significantly overexpressed in GC tissues, and this overexpression is also associated with unfavorable clinical outcomes. Through integrated bioinformatics and functional analyses, we identified YWHAG as a critical downstream effector of TPM3. Mechanistically, TPM3 interacts with YWHAG, triggering MAPK signaling, which promotes GC cell growth, invasion, and spread in both laboratory and in vivo environments. Collectively, our findings define a novel TPM3/YWHAG/MAPK signaling axis that is essential for GC progression and identify TPM3 as a potential therapeutic target. Future research should prioritize the development of small-molecule inhibitors that disrupt the TPM3-YWHAG interaction and validate the combined expression of TPM3 and YWHAG as a robust prognostic biomarker in prospective clinical cohorts.