Tumor tissue samples were obtained from Ruijin Hospital, Shanghai Jiao Tong University. Patients underwent curative GC resection between 2018 and 2019, and samples from these patients were used for immunohistochemistry (IHC). The Ethics Committee of Ruijin Hospital, Shanghai Jiao Tong University approved this project. All samples were anonymously coded in accordance with local ethical guidelines (as stipulated by the Declaration of Helsinki), and written informed consent was obtained.

Normal gastric cell (GES-1) and GC cell lines (AGS, SGC-7901, MGC-803, MKN-45, HGC-27) were stored at Shanghai Institute of Digestive Surgery, Ruijin Hospital. Cell lines were cultured in RPMI-1640 medium or Dulbecco’s modified Eagle’s medium (DMEM) with 10% fetal bovine serum (Gibco, United States) and 5 μg/ml penicillin-streptomycin in a humidified incubator at 37°C with 5% CO₂.

Total cellular RNA was isolated from GC cells according to the EZB RNA reagent kit protocol, and 200 ng of total RNA was reverse transcribed to cDNA using a PrimeScript RT Master Mix Kit (Takara Bio). mRNA levels were measured by quantitative real-time PCR using SYBR Premix Ex Taq, and human GAPDH was used as an internal control.

Western blotting was performed as described previously (Jiang et al., 2018). Antibodies against FOXC1, DKK1, c-MYC, and β-catenin were purchased from Abcam, and the secondary antibody (anti-rabbit IgG or anti-mouse IgG) was purchased from Proteintech. An anti-GAPDH antibody was used as an internal control. Detail information could be obtained in the Supplementary Material 2.

Plasmid transfection process was performed using Lipofectamine 2000 Reagent (Life Technology, Thermo Fisher Scientific, DE, United States). Full-length FOXC1 cDNAs were cloned into the pLVXyu-2Flag-3C-2 vector for FOXC1 expression. The c-MYC plasmid and DKK plasmid were gifts from Professor Guohong Hu at The Key Laboratory of Stem Cell Biology, Institute of Health Sciences, Shanghai Institutes for Biological Sciences. Stably transfected GC cells were established using puromycin selection after transfection with the expression vector or control plasmid. CRISPR/Cas9 vector plasmid was obtained from the Molecular and Cell Biology Laboratory, Fudan University. The sgRNA target sequence for FOXC1 was 5′-GGGTGCGAGTACACGCTCAT-3′, and CRISPR/Cas9 vector plasmid was used as a control. Stably transfected GC cells for 24 h were established using puromycin selection for 2 weeks after transfection. Knockout effiency were validated by Western blot with FOXC1 antibody.

The pGL3-Basic Luciferase Reporter vector was a gift from Fudan University Shanghai Cancer Center. The 2.1-kb DKK1 and c-MYC promoter was cloned into the pGL3-Basic Luciferase Reporter vector. Activity of the DKK1 and c-MYC promoters was normalized by co-transfection with the Renilla luciferase reporter plasmid, which was a gift from Fudan University. Firefly and Renilla luciferase activities were measured at 48 h after transfection using a Dual-Luciferase Reporter Assay System (Promega).

A TOP-flash/FOP-flash-dependent luciferase reporter assay was used to evaluate Wnt/LEF/TCF- activity. TOP-flash/FOP flash and Renilla plasmids to demonstrate transfection efficiency were incubated for 24 h. Relative luciferase activity was determined using a dual-luciferase assay (Promega). Transfection efficiency was normalized according to Renilla luciferase activity. The cells were lysed at 16 h after transfection, and luciferase activities were determined.

GC cells (2^∗10^⁷) were used for chromatin immunoprecipitation assays, and an anti-FOXC1 antibody was used to pull down DKK1 and c-MYC promoter-protein complexes. This assay was performed using CST Chromatin Immunoprecipitation Kit (CST9002) according to the manufacturer’s instructions. The DNA samples were analyzed by PCR for potential binding sites. The PCR products were separated and visualized on a 2.5% agarose gel. The primers used for PCR are listed in Table 1.

AGS and MKN-45 cells were seeded onto coverslips before they reached 80% confluence. The cells were fixed with 4% paraformaldehyde for 15 min, and the samples were kept in phosphate-buffered saline (PBS) containing 0.1% Triton X-100 (PBS-T), quenched with 50 mM NH4Cl in PBS-T, and blocked with 1% BSA in PBS-T. Immunostaining was performed using the appropriate primary and secondary antibodies, and images were acquired using a confocal microscope (Zeiss, LSM510).

Apoptosis was evaluated by flow cytometry. Briefly, cells were stained with Annexin V-FITC conjugate and propidium iodide solution. A FACSCalibur system (BD Biosciences, United States) was used to analyze apoptosis.

The cells were fixed in 75% ethanol and stained with PI/RNase Staining Buffer (BD Biosciences). The cell cycle was analyzed by flow cytometry using the FACSCalibur system (BD Biosciences, United States).

GC cell proliferation was evaluated by the CCK-8 assay. For the colony formation assay, cells were seeded into six-well culture plates at a density of 1,000–2,000 cells/well and grown for 10–14 days. The cells were fixed with methanol and stained with 0.1% crystal violet.

Ten nude mice were sorted into two groups, and GC cells (5 × 10⁶ per mouse) were subcutaneously injected. The mice were sacrificed 5 weeks later. The tumors were removed and weighed, processed and embedded in paraffin for further study.

All experiments that were presented in this work were repeated more than three times. Data was presented as mean ± standard error of the mean. For statistical analysis, two-tailed independent Student’s t-test was applied to a demonstration of homogeneity of variance with the F test or one-way or two-way ANOVA for more than two groups. Statistical significance was set as P < 0.05.

To investigate the role of FOXC1 in human gastric cancer, the mRNA level data of FOXC1 in GC and adjacent normal tissues were obtained from GEPIA and analyzed. And the result indicated that the expression level of FOXC1 was increased in GC tissues compared with normal tissues (Figure 1A). To confirm the expression level of FOXC1 in GC, IHC staining was applied to detect the protein level of FOXC1. As shown in Figure 1B, compared to normal tissues, the protein level of FOXC1 was higher in 85.7% (18/21) of the GC tissues. qPCR and western blot were applied to evaluate mRNA and protein levels of FOXC1 in non-malignant (GES-1) and gastric cancer cell lines (AGS, SGC-7901, MGC-803, MKN-45, HGC-27). The results demonstrated that both mRNA and protein levels were significantly upregulated in GC cell lines (Figures 1C,D). The data was obtained from Kaplan-Meier Plotter and affy ID of FOXC1(213260_at) dataset used for the analysis¹. A total of 422 patients were concluded in our analysis and cut-off value for the high and low expression is 141. And number of patients in high and low expression groups are 300 and 122, respectively. The median survival time for high and low expression groups are 89.43 and 34.3 months respectively (p = 0.0038). In general, high expression of FOXC1 indicated a poor prognosis in GC (Figure 1E). These results indicate that FOXC1 might be an important oncogene in GC.

To investigate the role of FOXC1 in the tumorigenesis and progression of GC, we applied CRISPR–Cas9-mediated gene-editing system to knock out FOXC1 expression in AGS and MKN-45 cells. Meanwhile, FOXC1 sequence was cloned into pCDNA plasmids overexpressed FOXC1 in AGS and MKN-45 cells. The efficiencies were confirmed by western blot and qPCR assays (Figure 2A). We observed that FOXC1 knockout (KO) significantly suppressed colony formation and cell proliferation, while FOXC1 overexpression dramatically enhanced cell viability and proliferation (p < 0.05) (Figures 2B,C). To study the oncogenic functions of FOXC1 in GC, we checked the effect of FOXC1 knockout on tumor growth and found that FOXC1 KO significantly decreased the tumor growth of MKN-45 cells (p < 0.05) (Figures 2D–G). FOXC1 expression level in xenografts was determined by IHC (Figure 2H).

Apoptosis and cell cycle are two main factors that affect tumor growth. To investigate the role of FOXC1, we applied flow cytometry for verification. The results demonstrated that FOXC1 KO in GC cells didn’t affect apoptosis (Figure 3A), but decreased cell cycle progression (p < 0.05) (Figure 3B), indicating FOXC1 regulates the proliferation of GC cells through cell cycle. To search for downstream effectors of FOXC1-induced GC proliferation, we examined the expression levels of several signaling molecules (e.g., Wnt/β-catenin, Notch, and Hippo) implicated in the regulation of cell proliferation during gastric carcinogenesis. qPCR and western blotting analyses revealed unchanged expression of the core components of those signaling pathways with FOXC1 KO (Figures 3C,D). To better understand the role of FOXC1 in the cell cycle of GC, we further measured mRNA levels of other target genes, such as baculoviral IAP repeat-containing protein 5 (BIRC5), cyclin D1 (CCND1), c-MYC, secreted phosphoprotein 1 (SPP1), and glypican 3 (GPC3) BAX and PUMA and found that CCND1 and c-MYC gene expression was downregulated in FOXC1 KO AGS cells (Figure 3C), demonstrating that FOXC1 promotes cell cycle via regulation of cyclin D1 and c-MYC.

Due to c-MYC is a critical transcription factor in regulating cell proliferation and development, we focused on c-Myc in regulating GC proliferation. To investigate the role of c-MYC in FOXC1-mediated GC proliferation, we next analyzed whether c-MYC expression is changed in AGS and MKN-45 cells with KO and overexpression of FOXC1, respectively. The results of qPCR and western blotting indicated that FOXC1 KO in AGS and MKN-45 cells decreased c-MYC expression, and consistently, FOXC1 overexpression dramatically enhanced c-MYC expression (Figures 4A,B). Furthermore, direct overexpression of c-MYC restore the FOXC1-deficient phenotype on cell proliferation in AGS and MKN-45 cells (Figures 4C,D). The results of luciferase activity assays indicated that FOXC1 could regulate c-MYC expression at the transcriptional level (Figures 4E,F). Given that FOXC1 is an important transcription factor, we next examined whether FOXC1 acts as a transcription factor to activate c-Myc expression in AGS and MKN-45 cells. We employed ChIP assay to confirmed whether there was a potential binding site in the c-MYC promoter. However, the results showed no direct binding of FOXC1 to the c-MYC promoter (Figure 4G).

Considering c-MYC is an important target gene of the Wnt pathway, we hypothesized that FOXC1 may activate Wnt signaling to enhance c-MYC expression. Given that AGS and MKN-45 cells expressed higher levels of FOXC1 than SGC-7901, MGC-803, HGC-27 cells. Further, Wnt activity was aberrant higher in AGS and MKN-45 cells than in SGC-7901, MGC-803, HGC-27 cells, as determined by TOP-Flash reporter assay (Figure 1A), indicating that Wnt activity is regulated and correlated with the FOXC1 levels in the cells (Figure 5A). β-Catenin aberrant cumulation in the cytoplasm is an important part for Wnt activation. We conducted qPCR and WB assay to explore the potential correlation of FOXC1 and β-Catenin, and the results indicated that FOXC1 did not have a significant influence on β-catenin expression at transcription or translation level (Figures 5B,C). At the same time, we learned that the activity of the wnt signaling pathway is related to β-Catenin in the nucleus (Zhang et al., 2011). Thus, we further explore the distribution of β-catenin in cytoplasm and nucleus. Interestingly, the results showed that β-catenin distribution in the nucleus increased with upregulation of FOXC1, while decreased with FOXC1 knockout, supporting the role of FOXC1 in activation of classic Wnt pathway (Figures 5D,E). In addition, β-Catenin will be labeled with E3 ubiquitin ligase after β-Catenin phosphorylation at Ser33/37/Thr41 in the cytoplasm (Reya and Clevers, 2005; Clevers and Nusse, 2012; Zhan et al., 2017). We also found β-Catenin phosphorylation at Ser33/37/Thr41 increased with FOXC1 KO and decreased with FOXC1 overexpression (Figure 5F), supporting the conclusion that WNT signaling pathway is activated under the condition of high FOXC1 expression. Previous studies indicated FOXM1 could directly bound to β-catenin in vivo and in vitro and Promoted β-catenin nuclear localization and controls Wnt target-gene expression (Zhang et al., 2011). Considering FOXC1 and FOXM1 belong to forkhead box family, thus we further explore the potential role of FOXC1in nuclear localization of β-catenin. Our results showed that FOXC1bound to β-catenin to form a complex in AGS and MKN-45 cells (Figures 5G,H). However, we also found a contradiction, that is, in Figure 4G, the ChIP experiment showed that there was no potential FOXC1 binding site in c-MYC promoter. Thus, we conducted COIP assays in the cytoplasm and nucleus, and the results showed FOXC1 bound to β-catenin in cytoplasm, but not in nucleus (Figures 5I–K). These results were consistent with previous result demonstrating that FOXC1 does not directly bind to the c-MYC promoter.

To find out the mechanism by which FOXC1 activates the Wnt signaling pathway, we first applied the TopFlash/FopFlash assay to determine the activity of Wnt pathway in FOXC1 KO GC cells. The results indicated that the activity of the Wnt signaling pathway was decreased in FOXC1 KO and increased in FOXC1 overexpression AGS and MKN-45 cells (Figure 6A). Considering the role of DKK1 in classic Wnt signaling, We next examined the level of DKK1 expression, which is the antagonist of Wnt signaling, and found that the DKK1 mRNA level increased in FOXC1 KO GC cells and decreased in FOXC1-overexpressing GC cells (Figure 6B; Zhu et al., 2021). At the same time, we applied western blot and EILSA assays were to confirm the correlation of DKK1 protein and FOXC1, and results indicated that FOXC1 negatively regulated DKK1 expression at translational level (Figures 6C,D). In addition, siDKK1 restored the cell proliferation repressed by FOXC1 KO in AGS and MKN-45 cells (Figures 6E,F). Those results indicated FOXC1 regulated GC cells proliferation through DKK1.

A previous study indicated that AAAYA-rich sequences are the main DNA-binding sequence of FOX transcription factors (Berry et al., 2002), and we found 12 potential binding sites (AAAYAs) in the DKK1 promoter (Figure 7A). To evaluate possible interaction between DKK1 and FOXC1, the sequence from + 100bp to −2000bp of the human DKK1 promoter was cloned into a luciferase reporter plasmid (pGL3-DKK1-Luc), and pGL3-DKK1-Luc plasmids with FOXC1 plasmids or control vectors were transiently transfected into AGS and MKN-45 cells. Luciferase assays indicated that FOXC1 significantly decreased the luciferase activity of DKK1 in both cell lines (Figure 7B). According to ChIP analysis, FOXC1 directly binds to the DKK1 promoter (sites #2 and #5) (Figure 7C). To further validate whether DKK1 is regulated directly by FOXC1, we used the wildtype (WT) or mutant (MT) promoter of DKK1 in the luciferase assay (Figure 7D). We found that FOXC1 reduced the luciferase activity of the WT DKK1 promoter to approximately 75% of the corresponding control but did not affect the activity of MT1 or MT123, suggesting that FOXC1 binds to the site #1 (–1514–1518 bp) in the DKK1 promoter (Figure 7E).