Published online Jun 15, 2026. doi: 10.4251/wjgo.v18.i6.118121
Revised: January 18, 2026
Accepted: February 25, 2026
Published online: June 15, 2026
Processing time: 167 Days and 18.5 Hours
Gastric cancer (GC) is a globally prevalent, lethal malignancy with high recur
To investigate the specific biological functions of CDH6 in the transforming grow
We performed in vitro experiments using HGC-27/AGS cells with stable overexpression of CDH6, as well as in vivo xenograft experiments in BALB/c-nu mice. Bioinformatics analysis and clinical validation with GC samples were also condu
CDH6 overexpression significantly enhanced migration, invasion, and prolife
CDH6 promotes progression and oxaliplatin resistance of GC by activating the TGF-β/Smad pathway and pro
Core Tip: Gastric cancer (GC) is among the most prevalent and lethal malignancies globally and the molecular mechanisms of cadherin 6 (CDH6) in GC remain unclear. We verified that CDH6 is a key regulator in GC progression and oxaliplatin resistance. Mechanistically, CDH6 exerts its oncogenic effects by activating the transforming growth factor-β pathway and promoting epithelial-mesenchymal transition. Targeting the CDH6/transforming growth factor-β/Smad axis may serve as a promising therapeutic strategy to overcome oxaliplatin resistance and improve clinical outcomes in GC patients.
- Citation: Zhao ZX, Liu QL, Fan JX, Zhu S, Wang FS, Hu ZJ. Cadherin 6 drives epithelial-mesenchymal transition and oxaliplatin resistance in gastric cancer via transforming growth factor-β/Smad signaling axis. World J Gastrointest Oncol 2026; 18(6): 118121
- URL: https://www.wjgnet.com/1948-5204/full/v18/i6/118121.htm
- DOI: https://dx.doi.org/10.4251/wjgo.v18.i6.118121
Gastric cancer (GC) is among the most prevalent and lethal malignancies globally, ranking as the fifth most commonly diagnosed cancer and the fourth leading cause of cancer-related mortality worldwide according to epidemiological surveillance data from GLOBOCAN 2020[1]. The disease burden is particularly substantial in East Asia; China alone accounts for approximately 44% of the global incidence of GC and approximately 50% of GC-related mortality, which can be largely attributed to endemic Helicobacter pylori infection and delayed diagnosis[2-4].
Contemporary management strategies for GC predominantly rely on surgery-based multimodal approaches in which curative gastrectomy is integrated with a combination of adjuvant chemotherapy, targeted therapies such as trastuzumab for human epidermal growth factor receptor 2-positive tumors, and immune checkpoint inhibitor therapy[5-8]. However, despite the widespread use of such regimens, 50% of patients who undergo radical resection followed by standardized adjuvant chemotherapy experience recurrence and distant metastasis within 3 years[9,10]. These poor outcomes are due in part to the occurrence of resistance to chemotherapy; moreover, the key molecules and genes and the mechanisms underlying the pathogenesis of GC remain unclear, limiting the development of novel therapeutic agents[11].
Cadherin 6 (CDH6), a transmembrane calcium-dependent adhesion protein, has emerged as a key coordinator of gastric oncogenesis. In previous work, we found that CDH6 overexpression was correlated with aggressive tumor behavior and poor patient prognosis[12]. A bioinformatics analysis showed that CDH6 could function as a master regu
The transforming growth factor (TGF)-β signaling pathway has stage-dependent dual functions in GC. During early tumorigenesis, it acts as a tumor suppressor by inducing apoptosis, blocking EMT, and restraining proliferation. However, in the advanced stages of the disease, it undergoes a functional switch to drive tumor progression through EMT-mediated metastasis, immune evasion, angiogenesis, and cancer stem cell maintenance. Oxaliplatin is a key chemotherapeutic agent that is frequently used in GC treatment; however, it is common for oxaliplatin resistance to occur, resulting in tumor recurrence[11]. Emerging evidence suggests that this phenomenon is driven by the TGF-β pathway via multiple mechanisms, including EMT-mediated drug exclusion, tumor microenvironment remodeling, and dysregulation of the DNA damage response[15,16]. However, the specific mechanisms underlying the development of oxaliplatin resistance in GC remain unclear. Thus, there is an urgent need for further elucidation of the pathways involved in this process. In the present study, we conducted an in-depth investigation of the mechanisms by which CDH6 promotes EMT via the TGF-β signaling pathway in GC. We also explored the molecular mechanisms through which CDH6 may participate in the development of oxaliplatin resistance.
Gene expression profiles and associated clinicopathological data of patients with gastric adenocarcinoma were downloaded from the Genomic Data Commons Data Portal of The Cancer Genome Atlas (https://portal.gdc.cancer.gov/repository) on January 10, 2025 for further analysis. Twenty-five GC patients who had received oxaliplatin-based neoadjuvant chemotherapy at Fuyang People’s Hospital from January 2023 to June 2024 were enrolled. Tumor tissue samples were collected from these patients, together with clinicopathological data. All cases were pathologically confirmed, and participants completed three courses of neoadjuvant chemotherapy as scheduled. Enhanced computed tomography re-examination data were available after treatment. We adopted the response evaluation criteria in solid tumors version 1.1 to define clinical oxaliplatin resistance. Patients who achieved complete response, partial response, or stable disease with a tumor size reduction ≥ 10% after neoadjuvant chemotherapy were defined as the sensitive group; and patients who experienced progressive disease, including an increase in tumor size ≥ 20% or the appearance of new lesions, or stable disease with a tumor size reduction < 10% were defined as the resistant group.
GC cell line HCG-27, AGS was purchased from Procell Life Science and Technology Company (Wuhan, Hubei Province, China). Cells were cultured in high-glucose Dulbecco’s modified Eagle’s medium (DMEM; RG-CE-2, Ketu Biotech, Hefei, Anhui Province, China) containing 10% fetal bovine serum (FBS; CG-SR-01, Ketu Biotech, Hefei, Anhui Province, China) and 1% penicillin-streptomycin (RG-CE-8, Ketu Biotech, Hefei, Anhui Province, China) in a 5% CO2 atmosphere at 37 °C. The culture medium was replaced every 48 hours. Cells were screened periodically for mycoplasma contamination using a One-step Quickcolor Mycoplasma Detection Kit (Shanghai, China).
HGC27/AGS cells were seeded in 24-well plates at a density of 30%-50%. The next day, they were transfected with a CDH6-overexpression lentivirus (multiplicity of infection: 10) obtained from Ketu Biotech in the presence of 8 μg/mL polybrene. After 24 hours, the medium was replaced with fresh culture medium, and green fluorescent protein expression efficiency was assessed via fluorescence microscopy 48 hours post-transfection. Selection was performed with puromycin (2 μg/mL) for 7 days until all nontransfected control cells had been eliminated, allowing transfected cells to survive and propagate stably. Monoclonal colonies were isolated using the dilution method, and individual colonies were expanded to yield stable CDH6-overexpressing (OE-CDH6) cell lines. CDH6 expression levels in these cells were evaluated by quantitative real-time polymerase chain reaction (qRT-PCR) and western blotting to confirm transfection efficiency.
Transfected cells were seeded into six-well plates (5 × 105 cells/well) and cultured in medium until they reached 90% confluence. A 200-μL pipette tip was used to make a scratch on each plate. At 0, 24, and 48 hours post-scratch, wound closure was measured. ImageJ was used to evaluate the scratch area or width at each time point and calculate the scratch closure rate.
Cells were digested, and their concentration was adjusted to 5 × 104 cells/mL in FBS-free DMEM. Then, 24-well Transwell chambers were coated overnight at 4 °C with 50 μL Matrigel matrix (1:8) or left uncoated; after which, 200 μL cell suspension was added to the upper compartment and 800 μL DMEM containing 10% FBS to the lower compartment of each chamber. The chambers were incubated at 37 °C in a 5% CO2 atmosphere for 24 hours. Finally, the cells were fixed with 4% paraformaldehyde for 20 minutes and stained with 0.1% crystal violet for observation and counting.
Cells were digested with trypsin (without EDTA) and centrifuged at 1000 rpm for 5 minutes. The cell pellets were collected, washed twice with precooled phosphate-buffered saline (PBS) and resuspended in 1 × binding buffer at a concentration of 106 cells/mL. Then, 100 μL aliquots of cell suspension were transferred to 5 mL flow cytometry tubes, followed by 5 μL Annexin V-fluorescein isothiocyanate and 5 μL propidium iodide (PI) staining solution. The mixture was gently vortexed and incubated in the dark at room temperature for 15 minutes. After incubation, 400 μL 1 × binding buffer was added to each tube, and apoptosis was analyzed immediately using a flow cytometer. Percentages of apoptotic cells (where Annexin V+/PI- cells were considered to be in early apoptosis and Annexin V+/PI+ cells in late apoptosis) were calculated using FlowJo (version 10).
Control and transfected OE-CDH6 cells were seeded in six-well plates (800 cells/well) and incubated at 37 °C for 10 days to allow single-cell colonies to form. For the oxaliplatin intervention group, the medium was replaced and supplemented with 2 μM oxaliplatin-containing medium 24 hours after inoculation. In the control and OE-CDH6 groups, equal volumes of drug-free medium were added. The medium was changed every 2-3 days throughout the culture period. Subsequently, the cells were fixed in 4% paraformaldehyde, and stained with 0.1% crystal violet for observation and counting.
The cytotoxicity of oxaliplatin at varying concentrations (0, 0.5, 1, 2, 4, 6, 8, and 10 μM) toward the cells was evaluated via cell counting kit-8 (CCK-8) assay. Cells were digested, counted, and seeded at 4000 cells/well in 96-well plates (three replicates per concentration). After 12 hours incubation at 37 °C with 5% CO2 to allow cell adhesion to occur, the medium was replaced with fresh medium containing oxaliplatin or a drug-free control. Following 48 hours treatment, CCK-8 reagent was added, and optical density values at 450 nm were measured after 1 hour incubation. A concentration-inhibition curve was plotted to calculate half-maximal inhibitory concentration (IC50) values. A 1.25-fold increase in IC50 value was defined as chemoresistance.
Cells were digested and counted, and adjusted to a concentration of 4000 cells/100 μL. Then, 100 μL of cell suspension was seeded into 96-well plates, with three replicate wells for each group. After incubation at 37 °C in 5% CO2 for 12 hours to allow cell adhesion, fresh medium was added as follows. For the control group and OE-CDH6 group, cells were supplied with fresh complete medium, whereas in the oxaliplatin group and OE-CDH6 + oxaliplatin group, cells were treated with 2 μM oxaliplatin. Following 48 hours incubation, 10 μL CCK-8 reagent was added to each well; after which, the plates were gently shaken and incubated for a further 1 hour. The optical density at 450 nm was measured using a microplate reader.
Total RNA was extracted from cells using an RNAprep Pure Tissue Kit (MG-TQ-01; Ketu Biotech, Hefei, Anhui Province, China). Complementary DNA was reverse transcribed using KT First-strand complementary DNA Synthesis Mix for qPCR (with dsDNase) (MG-QZ-01; Ketu Biotech, Hefei, Anhui Province, China), and qRT-PCR was performed using KT AntiQ qPCR SYBR Green Fast Mix (MG-QP-01; Ketu Biotech, Hefei, Anhui Province, China). mRNA expression levels were calculated using the 2-ΔΔCt method. The primer sequences are shown in Supplementary Table 1. All gene primers were obtained from General Biol (Anhui Province, China). The thermocycling conditions were as follows: Initial denaturation at 95 °C for 30 seconds, followed by 40 cycles of 95 °C for 10 seconds, 55 °C for 10 seconds, and 72 °C for 30 seconds.
RIPA lysis buffer (BL504A, Biosharp, Ketu Biotech, Hefei, Anhui Province, China) was used to extract proteins from cells, and protein concentrations were determined using a bicinchoninic acid kit (P0012; Beyotime, China). Protein extracts were placed in centrifuge tubes together with 5 × loading buffer and incubated in a water bath at 95 °C for 5 minutes. The proteins were separated by sodium-dodecyl sulfate gel electrophoresis at 80 V for 20 minutes and 120 V for 60 minutes and transferred to membranes. The membranes were blocked with 5% skimmed milk powder at room temperature for 1 hour, followed by incubation with the respective primary antibodies [CDH6: 27703-1-AP; TβRI: 84453-1-RR; Smad2: 12570-1-AP; phosphorylated Smad (p-Smad): 80427-2-RR; E-cadherin: 20874-1-AP; zonula occludens-1 (ZO-1): 21773-1-AP; vimentin: 10366-1-AP; a-SMA: 14395-1-AP; Proteintech, China] overnight at 4 °C and with secondary antibody (GB23301; Servicebio, China) at room temperature for 1 hour. Bands were developed using luminescent liquid.
For immunohistochemical staining, paraffin-embedded tissue sections were baked at 60 °C for 1 hour; dewaxed with xylene twice; and rinsed with absolute ethanol twice, followed by 95%, 85%, and 75% ethanol, sequentially. Sections were immersed in antigen retrieval solution, heated in a microwave, and washed with PBS three times (for 5 minutes each). They were next incubated with 3% H2O2 for 15 minutes and washed with PBS three times, and nonspecific antigens were blocked with 5% bovine serum albumin. Diluted (1:200) primary antibodies [CDH6: Ab197845, Abcam, China; proliferating cell nuclear antigen (PCNA): 10205-2-AP; Ki67: 27309-1-AP; p53: 10442-1-AP, Proteintech, China] were added, followed by incubation at 4 °C overnight in a humid chamber. The next day, the tissues were washed with PBS three times (5 minutes each); then, secondary antibody (GB23301, Servicebio, China) was added, followed by incubation for 30 minutes and washing with PBS. Finally, the sections were stained with 3,3’-diaminobenzidine, counterstained with hematoxylin, dehydrated, and mounted.
Nude mouse xenograft models (BALB/c-nu) were established for in vivo experiments by subcutaneous injection of HGC27 or OE-CDH6 cells (1 × 107 cells/mL in 100 μL PBS/Matrigel mixture) into nude mice. Tumor-bearing mice (tumor volume 100-200 mm3) were randomized into groups (n = 3), and oxaliplatin (5 mg/kg) or saline was intraperitoneally injected twice weekly for 3-4 weeks, with monitoring of body weight. Tumors were excised after death, and volumes were calculated based on the long and short diameters.
All experiments were performed in triplicate (n = 3), and data were analyzed and visualized using GraphPad Prism 8 and R 4.2.1. Categorical variables are expressed as n (%) and continuous variables as mean ± SD. The survival analysis was performed using the online tool Kaplan-Meier plotter (https://kmplot.com/). Significant differences for categorical variables were analyzed using the χ2 test, whereas continuous variables were analyzed using Student’s t-test or one-way analysis of variance. The Shapiro-Wilk test was used to verify the normality of continuous data distribution, while Bonferroni post hoc tests were applied to correct for potential multiple testing biases. Correlations were analyzed using the Spearman method, and drug sensitivity was predicted with the oncoPredict R package. A two-tailed P < 0.05 was considered to indicate statistical significance.
Our experimental data demonstrated that overexpression of CDH6 in GC was closely associated with enhanced cell migration, invasion, proliferation, and poor prognosis (Figure 1). In the wound-healing assay, initial scratch distances (at 0 hour) in the control and OE-CDH6 groups were similar. However, at 24 hours and 48 hours, the OE-CDH6 group showed significantly greater closure of the scratch compared to the control group (Figure 1A). In the Transwell migration and invasion assays, the OE-CDH6 group had significantly higher numbers of migrated and invading cells compared to the control group (Figure 1B). The CCK-8 assay showed that overexpression of CDH6 promoted cell viability (Figure 1C). These results suggested that CDH6 overexpression accelerated processes related to cell proliferation and migration. Annexin V/PI double-staining flow cytometry analysis revealed a significantly lower proportion of apoptotic cells in the OE-CDH6 group compared to the control group, suggesting that CDH6 overexpression suppressed apoptosis (Figure 1D). The plate colony formation assay showed that CDH6 overexpression promoted colony formation. Kaplan-Meier survival curves for multiple datasets consistently demonstrated that patients with high CDH6 expression had poorer survival outcomes than those with low expression (Figure 1E-G). We used another GC cell line (AGS) for validation, and the results were consistent with above observations (Supplementary Figure 1).
The bioinformatics analysis indicated that there was a positive correlation between CDH6 expression and the probability of oxaliplatin resistance (Figure 2A), with the CDH6 high expression group showing significantly higher oxaliplatin resistance scores compared to the low-expression group (Figure 2B). Similarly, qRT-PCR results confirmed that CDH6 expression levels in the oxaliplatin-resistant group were significantly higher than those in the oxaliplatin-sensitive group (Figure 2C). Dose-response curves found that the IC50 of oxaliplatin in the OE-CDH6 group (3.38 ± 0.15 μM) was significantly higher than that in the control group (2.61 ± 0.14 μM) (Figure 2D and E). In the colony formation assay, the number of colonies formed in the OE-CDH6 group was significantly higher than that in the control group following oxaliplatin treatment (Figure 2F).
In vivo animal experiments, the body weights of mice in all groups showed a stable increasing trend, indicating that the experimental intervention (oxaliplatin administration) did not induce severe toxic side effects leading to weight loss (Figure 3A and B). Tumor volumes were significantly greater in the OE-CDH6 group than in the control group (Figure 3C and D). By contrast, oxaliplatin monotherapy effectively inhibited tumor growth, with tumor volumes significantly lower in the oxaliplatin group compared to the control group (P < 0.001). However, tumors were notably significantly larger in the oxaliplatin + OE-CDH6 group compared to the oxaliplatin-only group (P < 0.05). These results demonstrate that CDH6 overexpression promotes proliferation of GC and mediates its resistance to oxaliplatin.
We observed upregulated transcriptional levels of key TGF-β pathway genes (TβR, Smad2/3) in the OE-CDH6 group (Figure 4A). Western blotting confirmed that there were also significant increases in proteins encoded by these genes (Figure 4B). Both western blotting and qRT-PCR demonstrated that CDH6 overexpression led to decreased levels of epithelial markers (E-cadherin, ZO-1) and increased levels of mesenchymal markers (vimentin and α-SMA). These findings indicate that CDH6 may promote EMT via the TGF-β pathway in GC.
Immunohistochemical analysis showed significant upregulation of TGF-β and Smad2 protein expression following CDH6 overexpression, indicating activation of the TGF-β signaling pathway (Figure 5). CDH6 promoted EMT, given the decreased expression of epithelial marker E-cadherin and increased expression of mesenchymal marker N-cadherin in CDH6 overexpression tissues. CDH6 overexpression promoted expression of proliferation-associated PCNA and Ki67, while suppressing protein levels of tumor suppressor p53. These findings confirm that CDH6 overexpression synergistically promoted GC cell proliferation via activation of the TGF-β pathway and drives EMT progression.
To confirm that CDH6 exerted its effects through the TGF-β pathway, we performed rescue experiments by treating CDH6 overexpression cells with a TGF-β inhibitor at a concentration of 10 μM (OE-CDH6 + TGF-β inhibitor group) and comparing phenotypic characteristics (migration, invasion, and expression of EMT markers) and TGF-β/Smad signaling pathway activity between the OE-CDH6 and OE-CDH6 + TGF-β inhibitor groups. In the wound healing (Figure 6A) and Transwell (Figure 6B) assays, treatment of cells with the TGF-β inhibitor significantly rescued (reversed) the migration and invasion conferred by CDH6 overexpression. Colony formation assays (Figure 6C) showed that TGF-β inhibitor treatment suppressed colony formation in OE-CDH6 cells, suggesting that CDH6-mediated proliferation enhancement is dependent on the TGF-β pathway. Comparison of oxaliplatin IC50 between the OE-CDH6 group and OE-CDH6 + TGFβ inhibitor group showed that TGF-β inhibition reversed the oxaliplatin resistance induced by CDH6 overexpression. qRT-PCR (Figure 6D) and western blotting (Figure 6E and F) revealed that TGF-β inhibitor treatment reduced TβRI and p-Smad2 expression (P < 0.05), indicating suppression of pathway activation, and restored levels of epithelial markers (E-cadherin and ZO-1) while downregulating those of mesenchymal markers (N-cadherin, vimentin, and α-SMA; P < 0.05), thereby reversing the EMT phenotype induced by CDH6 overexpression. Collectively, these rescue experiments demonstrate that inhibition of the TGF-β pathway reverses multiple CDH6-induced phenotypic changes: Enhanced migration, invasion, proliferation, resistance to oxaliplatin, and EMT.
GC remains a global healthcare challenge with high mortality rates, which are largely attributed to its malignant biological properties and chemoresistance[17]. The biological mechanisms governing the occurrence and progression of GC remain unclear, and important therapeutic targets are still under investigation. CDH6, as a member of the cadherin family, holds potential as a therapeutic target for GC, yet its specific molecular mechanisms of action in GC remain incompletely elucidated. In this study, we systematically explored the role of CDH6 in GC progression and oxaliplatin resistance and found that CDH6 overexpression drives EMT and chemoresistance via activation of the TGF-β/Smad signaling axis. Specifically, through experiments using an inhibitor of the TGF-β/Smad signaling pathway, we confirmed that inhibiting this pathway reverses the aforementioned phenotypes induced by CDH6 overexpression. This finding suggests that the regulation of EMT and drug resistance by CDH6 is dependent on the activation of the TGF-β/Smad signaling axis. In detail, our results demonstrate that CDH6 overexpression enhances GC cell migration, invasion, and proliferation while suppressing apoptosis, consistent with our previous observation that high CDH6 expression is correlated with poor prognosis in GC patients[12]. Mechanistically, we found that CDH6 exerts these effects by activating the TGF-β signaling pathway, as demonstrated by upregulation of TβRI, Smad2, and p-Smad2, and concurrent induction of EMT (downregulation of epithelial markers E-cadherin and ZO-1 and upregulation of mesenchymal markers vimentin and α-SMA) in OE-CDH6 cells.
EMT is a major driver of tumor metastasis, as it enables cancer cells to acquire invasive properties via loss of epithelial polarity[16,18]. Our results are consistent with those of a bioinformatics study suggesting that CDH6 is a master regulator of EMT in GC[13]. However, we added to previous findings by experimentally validating the TGF-β/Smad pathway as a critical mediator. CDH6-mediated EMT was accompanied by increased expression of proliferation markers (PCNA and Ki67) and decreased expression of p53. The CCK-8 cell proliferation assay demonstrated that CDH6 overexpression can promote the proliferation of GC cells. Therefore, we hypothesize that CDH6 can regulate uncontrolled cell proliferation and promote cancer cell metastasis in GC, forming a dual phenotype, thereby accelerating tumor progression[19].
Oxaliplatin is a first-line chemotherapeutic agent that is widely used in GC treatment; however, the development of oxaliplatin resistance can severely limit its effectiveness, resulting in poor patient prognosis[20]. In the present study, we identified CDH6 as a novel biomarker for predicting oxaliplatin resistance, with CDH6 overexpression significantly increasing the IC50 of oxaliplatin and reducing drug-induced apoptosis, and in vivo xenograft experiments showing that OE-CDH6 tumors were less responsive to oxaliplatin. Mechanistically, this resistance was linked to sustained TGF-β pathway activation and EMT induced by CDH6 overexpression, consistent with emerging evidence that TGF-β-driven EMT promotes drug resistance by enhancing cancer stem-cell-like properties and drug efflux[16,21].
The TGF-β/Smad signaling pathway often interacts with and synergistically promotes multiple tumor-related signaling pathways, regulating tumor progression and chemoresistance[22,23]. After activation of TGF-β receptors, the phosphoinositide 3-kinases (PI3K)/protein kinase B (AKT) pathway can be directly activated via the PI3K regulatory subunit p85[24]. Smad3 can cross-promote the phosphorylation of AKT, and these two mechanisms synergistically amplify EMT and chemoresistance. At the level of the mitogen-activated protein kinase signaling pathway, Smad2/3 can form a complex with phosphorylated extracellular signal-regulated kinase in this pathway, significantly enhancing the binding efficiency to the promoters of EMT-related genes and thereby synergistically promoting gene transcription. Additionally, TGF-β/Smad signaling stabilizes β-catenin by inhibiting glycogen synthase kinase 3β, which promotes the nuclear translocation of β-catenin and activates the transcription of EMT and Wnt pathway target genes[25]. The aforementioned signaling crosstalk collectively forms a complex regulatory network, which is a key mechanism underlying oxaliplatin resistance mediated by molecules such as CDH6. This field urgently requires more in-depth research in the future.
An important strength of our study was the use of rescue experiments to confirm that inhibition of the TGF-β pathway reversed CDH6-induced phenotypes, including migration, invasion, proliferation, and EMT. Specifically, TGF-β inhibitor treatment reduced TβRI/p-Smad2 activation, restored epithelial marker expression, and suppressed colony formation in CDH6 overexpression cells. These results establish a causal relationship in which CDH6 does not act through off-target or non-specific mechanisms but functions via the TGF-β/Smad axis to drive GC malignancy. The identification of this mechanism supports CDH6 as a potential therapeutic target. CDH6-targeted therapies (e.g., HKT288) have shown efficacy in renal and ovarian cancers[14]. Our data suggest that such strategies could be repurposed for use in GC treatment, particularly in combination with TGF-β inhibition to block both CDH6 and its downstream pathway, as a means of overcoming resistance to oxaliplatin-based chemotherapy[26]. For future studies, CDH6-targeted therapies can be used to systematically assess how CDH6 inhibition affects GC cell proliferation, invasion, and oxaliplatin resistance, and whether such intervention reverses or alleviates chemoresistance.
Our findings have two key clinical implications. First, CDH6 expression could serve as a prognostic biomarker for GC patients and guide personalized treatment decisions, as high CDH6 levels are correlated with poor survival rates and may predict oxaliplatin resistance. Second, targeting the CDH6/TGF-β axis could improve therapeutic outcomes. In particular, our preclinical data suggest that the combination of oxaliplatin and TGF-β inhibitor may reverse EMT and restore oxaliplatin sensitivity for GC patients with CDH6 high expression.
The roles of cadherin family members in GC are complex and not yet fully elucidated[27]. Among them, CDH1 is a calcium-dependent transmembrane adhesion molecule and a classic tumor suppressor gene. Studies have shown that CDH1 maintains the polarity and structural stability of gastric mucosal epithelium by mediating tight junctions between epithelial cells, preventing cell detachment or abnormal spread, and thus serves as a key protein inhibiting EMT[28]. Dysfunction of CDH1 (such as mutation and methylation) impairs the tumor-suppressive effect of E-cadherin, promotes the progression of sporadic GC, and is also a core pathogenic factor in hereditary diffuse GC[29]. CDH11 can relieve inhibition of β-catenin, allowing it to enter the nucleus and activate downstream oncogenes (c-myc and cyclin D1), thereby promoting the abnormal proliferation and cell cycle progression of GC[30]. High expression of CDH11 can downregulate the expression of epithelial markers (such as E-cadherin) and simultaneously upregulate the levels of mesenchymal markers (such as vimentin and α-SMA), inducing EMT, which is similar to the function of CDH6[31]. CDH17 mainly exerts an oncogenic role in GC, which inhibits the apoptosis of GC cells, promotes cell cycle progression, and is associated with chemotherapeutic drug resistance by activating the PI3K/AKT signaling pathway[32]. Unlike CDH6, the resistance mechanism of CDH17 primarily involves upregulating the expression of multidrug resistance-related proteins (such as P-glycoprotein), enhancing the efflux capacity of GC cells for chemotherapeutic drugs (such as 5-fluorouracil and cisplatin), reducing the intracellular accumulation concentration of drugs, and ultimately leading to chemotherapeutic resistance[33].
However, our study had several limitations. Our in vitro experiments primarily used the HGC-27 and AGS cell lines. Validation in additional GC cell lines (e.g., MKN-45 and NCI-N87) and organoids would improve the generalizability of the results. For the in vivo experiments, we used nude mouse xenografts, which lack a functional immune system. This limited the insights that could be obtained regarding the role of CDH6 in the immunosuppressive microenvironment; a known mediator of chemoresistance[34,35]. Future studies should incorporate syngeneic models or patient-derived xenografts to better mimic clinical conditions. There was a lack of relevant evidence (co-immunoprecipitation or reporter assays) to clarify whether CDH6 interacts directly with components of the TGF-β pathway. Although our data implicate CDH6 in oxaliplatin resistance, its potential roles in resistance to other chemotherapeutics (e.g., 5-fluorouracil or paclitaxel) remain unexplored. Our oxaliplatin-resistant cohort was small, potentially causing overfitting and weakening result robustness. Thus, validation in expanded clinical cohorts is essential to confirm CDH6 as a reliable predictive biomarker for GC oxaliplatin resistance. Longitudinal clinical studies are needed to confirm the predictive value of CDH6 with respect to oxaliplatin response and to evaluate the safety and efficacy of CDH6/TGF-β-targeted combination therapies. Future studies could also use single-cell RNA sequencing to investigate the heterogeneity of CDH6 expression within tumors, potentially identifying subpopulations that are particularly dependent on the CDH6/TGF-β pathway for survival and resistance. Whether the expression level of CDH6 can serve as a biomarker for TGF-β-targeted therapy also holds high research value and potential.
Our results identify CDH6 as a critical driver of GC progression and oxaliplatin resistance through activation of the TGF-β/Smad/EMT axis. These findings not only expand our understanding of GC pathogenesis but also highlight CDH6 as a promising biomarker and therapeutic target that could be used to reverse chemoresistance. Targeting the CDH6/TGF-β pathway may represent a novel strategy that could improve outcomes for GC patients with advanced disease.
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