Human gastric epithelial cell lines, HFE-145 (immortalized, non-cancerous) and AGS (human diffuse type of gastric cancer) were cultured in DMEM and DMEM/F12, containing 10% FBS (Gibco), 100 U/ml penicillin, and 100 μg/ml streptomycin at 37°C in an atmosphere of 5% CO2.

CagA + VacA+ H. pylori strain PMSS1 (pre-mouse Sydney strain 1) and its isogenic cagA mutant were established in our previous research. CagA + VacA + H. pylori strains 7.13 and an isogenic cagA were also included in this study, which were kindly provided by Dr. Richard Peek Jr. from the Vanderbilt Digestive Disease Research Center. All H. pylori strains were cultured on Campylobacter agar plates containing 10% sheep serum at 37°C under microaerophilic conditions. The measure methods of H. pylori bacterial density were as same as the previous study (Hu et al., 2019). Gastric epithelial cells were cocultured with H. pylori strains at MOI of 100.

A total of 40 human chronic non-atrophic gastritis (n = 20) and gastric carcinoma (n = 20) tissues were acquired from The First Affiliated Hospital of Nanchang University. Each group were divided into two subgroups: H. pylori-positive (n = 10) and H. pylori-negative (n = 10) individuals. The study protocol and exemption of informed consent were approved by the Ethics Committee of The First Affiliated Hospital of Nanchang University. Status of H. pylori infection for these clinical specimens was determined with a rapid urease test or Giemsa staining. Immunohistochemical staining was performed to examine expression profiles of TAZ on these samples, which were evaluated and scored for intensity (scaled 0–3) and frequency (scaled 0–4) by two pathologists blinded to sample identity. For statistical analysis, expression levels of TAZ proteins were illustrated by an expression score in range of 0 to 12 using the formula intensity × frequency (Li et al., 2018).

Animal care and experimental protocols were in accordance with guidelines established by the Institutional Animal Care and Use Committee of Nanchang University. The INS-GAS transgenic mice overexpressed pancreatic gastrin were purchased from Jackson Laboratory (Bar Harbor, ME). INS-GAS mice were orogastrically challenged with Brucella Broth (n = 6) or with 2 × 10⁹ CFU/ml H. pylori strain PMSS1 (n = 8) once every other day for a total of 5 times (Arnold et al., 2011; Li et al., 2020). Mice were euthanized at 4 months post infection, and gastric tissues were collected.

The recombinant plasmids such as TAZ, β-catenin cDNAs were designed and synthesized by hitrobio (Beijing, China). The recombinant CagA plasmid was a generous gift from Prof. Shiming Yang, Third Military Medical University of China. Small interfering (si) RNA duplexes were obtained from GenePharma (Shanghai, China). Cells were transfected with appropriate plasmid or siRNA using Lipofectamine 3,000 (Thermo Scientific, Waltham, MA, United States) according to manufacturer’s instructions.

Immunohistochemistry analysis was performed for gastric tissues from human and INS-GAS mice as previously described (22). These specimens were incubated with rabbit polyclonal anti-TAZ (Proteintech, Wuhan, China) at dilution of 1:400, followed by incubation with secondary antibody (PV6000, Zhongshan Golden-bridge, Beijing, China). Immunostained tissue slides were imaged on an upright confocal microscope (Nikon ECLIPSE Ni). Tissue sections were imaged at 200× and 400× magnification.

Western blotting analysis was conducted as described previously (Xie et al., 2020). Primary anti-TAZ (#83669), anti-β-catenin (#37447), anti-β-tubulin (#2128), anti-Histone 3 (#4499), anti-GAPDH (#2118), anti-HA-Tag (#3724), anti-Myc (#9402), anti-Cyclin D1(#2978), anti-CYR61(#14479), anti-CTGF (#86641) antibodies were purchased from Cell Signal Technology (Beverly, MA, United States); anti-CagA (sc-28,368) were from Santa Cruz (Dallas, TX, United States), anti-β-actin (#20536-1-AP) from proteintech (Wuhan, China); anti-Flag (F1804) was purchased from Sigma-Aldrich. All primary antibodies were used at a dilution of 1:1000, except for the internal control antibodies including GAPDH and β-actin which were diluted at 1:2000. Briefly, the proteins were extracted after adding lysis buffer supplemented with protease inhibitors (Invitrogen, GA, United States). Equal amounts of the sample proteins were separated on SDS-PAGE gels and transferred to nitrocellulose membranes. After blocking with 5% milk, the membranes were incubated with the primary antibodies overnight at 4°C, and then incubated with HRP-conjugated secondary antibodies (Invitrogen, GA, United States) for 1 h at room temperature. The protein bands were visualized using Super Signal West Pico stable peroxide solution (Thermo Scientific) and collected using iBright imaging system (Thermo Scientific). β-actin and GAPDH were used as internal control to normalize protein expression in cells sample and animal tissues sample, respectively.

The qRT-PCR analysis was performed as described in our previous studies (Xie et al., 2020). Briefly, total RNA was extracted using TRIzol reagent (Invitrogen, GA, United States). The primer sequences are presented in Supplementary Table S1. The qPCR assays were performed with a QuantStudio 5 Real-time PCR system (Life Technologies) according to the manufacturer’s protocol. The GAPDH gene was used as an internal control.

The TCF/LEF reported plasmid for β-catenin activity (M50 Super 8 × TOPFlash; plasmid#12456; Addgene, Cambridge, MA, United States) was a gift from Randall Moon (Veeman et al., 2003). For dual luciferase assay, cells were seeded into 12-well plates. The cells were transfected with 8 × TOPFlash plasmid and indicated siRNA. After transfection for 48 h, cells were co-cultured with H. pylori strain. The cell lysates were collected and subjected to a dual luciferase assay system (Yeasen Biotech, Shanghai, China) according to the manufacturer’s protocol.

For immunofluorescence staining, cells were washed with ice-cold PBS and fixed with 4% formaldehyde in PBS for 15 min. Then, the cells were permeabilized with 0.5% Triton X-100 for 10 min at room temperature. After blocking with 3% BSA for 30 min, the cells were incubated with primary antibodies against TAZ (dilution 1:100) or β-catenin (dilution 1:200) overnight at 4°C, and then incubated with Secondary Antibody (Alexa Fluor Plus 488 and Alexa Fluor Plus 555; Thermo Scientific). Cell nuclei were counters with DAPI. All images were acquired using a confocal fluorescence microscope (Leica Stellaris).

For immunoprecipitation assay, cells were transfected with the indicated plasmids, and then collected with lysis buffer. In brief, the cell lysates were incubated overnight with the mixture of 1 ug of antibodies and beads at 4°C. Then the beads were washed three times with 1 ml of lysis buffer and then boiled in loading buffer. The samples were subjected to western blot analysis as described above.

Cells were transfected with indicated plasmid vector or siRNA, and then seeded into 96-well plates at a density of 2 × 10³ cells/100 μl per well. The cells were infected with H. pylori strain at an MOI of 50 for the indicated time. The CCK-8 assay (TransGen Biotech, Beijing, China) was performed according to the manufacturer’s instructions. The optical density values were measured at a wavelength of 450 nm using a Molecular Devices SpectraMax M2e. For EDU assay, the relative viability of cells was determined by Cell-Light EDU Apollo 488 in Vitro imaging kit (RiboBio) following the kit protocol. All images were acquired and quantified using the high-content screening platform In-Cell Analyzer 2,200 (GE Healthcare).

Boyden chamber assay is useful tool to study cell migration and invasion. For cell invasion detection, about 2 × 10⁵ cells were plated into the upper Boyden chamber (8 μm pore size, Corning, NY, United States) with Matrigel-precoated inserts (BD Science, United States). For cell migration detection, Matrigel was not required. DMEM/F12 medium supplemented with 20% FBS was added in the lower chamber. After transfection and H. pylori infection, Cells adhering to the lower surface were fixed with methanol for 30 min, stained with 1% crystal violet for 15 min. The cells on the upper surface of the filters were gently wiped and counted under the inverted microscope (Nikon, ECLIPSE Ti).

All the statistical analysis was performed using SPSS 21.0 software. Data are presented as mean ± SD of three independent experiments. Studies for continuous variables were statistically analyzed using Student’s t-test or One-way ANOVA. The immunohistochemical data from human clinical specimens was statistically analyzed using Mann–Whitney test. The results were considered statistically significant at p < 0.05 (***, p < 0.001; **, p < 0.01; *, p < 0.05).

We previously reported augmented YAP expression in preneoplastic lesions of human gastric tissues (Li et al., 2018). In this study, we first examined the expression patterns of the YAP homologue TAZ in clinical specimens. As shown in Figures 1A,B, the expression of gastric TAZ was strongly increased in human gastric cancer tissues compared to that observed in chronic nonatrophic gastritis tissues. H. pylori infection is linked to the initiation of chronic active gastritis and significantly increases the risk of gastric adenocarcinoma (Crowe, 2019). Therefore, the TAZ expression levels were further compared between H. pylori-positive and H. pylori-negative subjects. Interestingly, H. pylori-positive gastritis tissues tended to have higher levels of TAZ than H. pylori-negative gastritis tissues. However, no significant difference in TAZ expression was noted between H. pylori-infected and -uninfected gastric cancer individuals (Figures 1A,C). Next, we utilized INS-GAS mice, which overexpress human pancreatic gastrin and have been widely used for studying the pathogenesis of H. pylori infection (14), to characterize the effects of H. pylori on TAZ. Western blot analysis showed that gastric TAZ expression was significantly increased in H. pylori-infected mice compared with sham controls (Figures 1D,E). In addition, a significant increase in gastric TAZ expression was noted in H. pylori-infected mice compared to uninfected animals based on immunohistochemistry analysis (Figure 1F) and quantitative image analysis (Figure 1G). These results indicated that H. pylori infection triggers TAZ activation, which may be implicated in the initiation of gastric neoplastic lesions.

To recapitulate the effects of H. pylori infection on gastric TAZ expression in human gastric tissues and INS-GAS mice, human gastric epithelial AGS cells were cocultured with the CagA+ H. pylori strain NCTC11637 or 7.13. H. pylori infection significantly increased TAZ protein expression in both an MOI-dependent (50, 100, 200 and 400 MOI) (Figure 2A) and a time-dependent manner (2 h, 4 h, 6 h, and 8 h) (Figure 2B). Similarly, an increase was noted in total TAZ expression in normal human gastric epithelial HFE145 cells infected with H. pylori (Figure 2C). In addition, the intracellular localization of TAZ was increased after treatment with H. pylori strains, as shown by immunofluorescence staining. Importantly, the increase of H. pylori MOIs was correlated with the promotion of TAZ translocation from the cytoplasm to the nucleus (Figure 2D). These in vitro data clearly indicated that H. pylori infection leads to TAZ upregulation and activation.

CagA, which is delivered into gastric epithelial cells via the type IV secretion system (T4SS), has been considered an oncoprotein that induces gastric carcinogenesis (30). To determine the role of CagA in H. pylori-induced activation of TAZ, AGS cells were cultured alone or cocultured with H. pylori PMSS1 wild-type or its CagA-deficient isogenic mutant strain. We found that CagA knockout significantly inhibited the induction of TAZ in response to H. pylori infection (Figure 2E). Furthermore, transient overexpression of CagA via plasmid transfection into AGS cells strongly increased TAZ expression (Figure 2F). Furthermore, the recombinant CagA overexpression caused an marked increase in the nuclear localization of TAZ. These findings suggest that bacterial CagA is integral for H. pylori-induced TAZ activation.

Given that TAZ activation promotes cell survival, proliferation, and metastasis by stimulating its downstream transcription factors (Pocaterra et al., 2020), we next dissected the role of TAZ in the H. pylori infection-induced malignant transformation of gastric epithelial cells. Knockdown of endogenous TAZ by siRNA inhibited cell proliferation (Figures 3A,B, EDU assay) and cell viability (Figure 3C, CCK8 assay) induced by CagA+ H. pylori strain NCTC11637 or 7.13. compared with H. pylori infection alone in AGS cells. Similarly, the number of AGS cell clones was diminished after treatment with TAZ siRNA in combination with the H. pylori strain compared to H. pylori infection alone (Figure 3D). To explore how TAZ mediates cell invasion and migration induced by H. pylori infection, AGS cells were transfected with TAZ siRNA for 48 h and then cocultured with the H. pylori strain for a transwell assay. The siRNA-mediated knockdown of YAP in combination with CagA+ H. pylori NCTC11637 infection significantly decreased invasion and migration capacities (Figures 3E–G). These findings suggest that impaired TAZ activation abolishes H. pylori infection-induced malignant transformation including cell survival, proliferation, invasion and migration.

YAP/TAZ are closely linked to the Wnt/β-catenin pathway because they share some target genes and biological processes (Li et al., 2019). Given that our data indicated that H. pylori infection led TAZ overexpression and nuclear accumulation, we next investigated whether TAZ induction promotes β-catenin pathway activation in response to H. pylori infection. TAZ knockdown significantly downregulated the β-catenin protein levels induced by H. pylori (Figure 4A). In addition, qRT-PCR data showed that H. pylori infection upregulated the transcription of β-catenin-targeted genes including Lgr5 and Myc which were then downregulated after treatment with TAZ siRNA (Figure 4B). Given that β-catenin activation is correlated with increased nuclear β-catenin abundance, we examined the effect of TAZ knockdown on the subcellular expression of β-catenin following H. pylori infection in AGS cells by separating the nuclear and cytoplasmic proteins. As shown in Figure 4C, H. pylori infection promoted nuclear TAZ and β-catenin compartmentalization, which was significantly suppressed by TAZ knockdown. Consistent with these findings, immunofluorescence assay showed that H. pylori infection-induced nuclear translocation was significantly suppressed by inhibition of TAZ activation (Figure 4D). Moreover, we performed a TOP/FOP flash luciferase reporter assay to detect β-catenin transcriptional activity. As shown in Figure 4E, H. pylori infection increased TCF/β-catenin reporter activity, and this effect was significantly inhibited by siRNA-mediated TAZ knockdown. To further confirm the effect of TAZ on the β-catenin pathway, AGS cells were transiently transfected with a TAZ overexpression plasmid. Exogenous TAZ overexpression increased the protein levels of Myc and Cyclin D1, the downstream targets of the Wnt/β-catenin pathway (Figure 4F). Consistent with these results, TAZ overexpression led to significant transcriptional induction of the downstream genes of Wnt signaling, such as Lgr5, Cyclin D1, and Myc, which was remarkably blocked following β-catenin knockdown by siRNA (Figure 4G). Collectively, these findings suggest that H. pylori infection leads to β-catenin activation and target gene expression through TAZ upregulation.

We next explored the molecular mechanism of TAZ activation in response to H. pylori infection. Given the close relationship between the Hippo and Wnt signaling pathways, we hypothesized that H. pylori infection activates the TAZ pathway through the regulation of β-catenin. To test this hypothesis, gastric epithelial AGS cells were transfected with β-catenin siRNA and then cocultured with the H. pylori NCTC11637 strain. Knockdown of β-catenin significantly suppressed TAZ expression induced by H. pylori (Figure 5A). Moreover, cytoplasmic and nuclear proteins were effectively separated for the detection of the intracellular TAZ expression. Notably, H. pylori infection promoted the nuclear accumulation of TAZ and β-catenin, whereas β-catenin knockdown by siRNA remarkably inhibited this induction (Figure 5B). Immunofluorescence staining for TAZ and β-catenin confirmed that H. pylori-induced nuclear translocation of TAZ was inhibited by knockdown of β-catenin (Figure 5C). Furthermore, transient overexpression of β-catenin upregulated the protein levels of CTGF and CYR61 downstream of the Hippo signaling pathway (Figure 5D). Taken together, these findings suggest that a positive feedback loop exists between TAZ and the Wnt/β-catenin pathway in H. pylori-induced gastric tumorigenesis.

It has been reported that TAZ is associated with the destruction complex in the Wnt/β-catenin pathway, including Axin1, β-TrCP, β-catenin and GSK3β (Azzolin et al., 2014; Lee et al., 2020). Therefore, we delineated the possible interaction between TAZ and β-catenin in H. pylori-associated gastric carcinogenesis. The immunofluorescence assay was first performed to determine the expression and cellular localization of TAZ and β-Catenin in human gastric tissues. β-catenin was normally expressed in the membrane of epithelial cells of gastric mucosa with chronic gastritis, and TAZ was present in the plasma membrane and cytoplasm. In contrast, increased overall expression of gastric TAZ was observed, and TAZ colocalization with β-catenin was noted in human gastric cancer tissues but not gastritis tissues (Figure 6A). The interaction of endogenous TAZ with β-catenin was further demonstrated using the co-immunoprecipitation assay (Figure 6B). These data suggest that TAZ may be translocated from the plasma membrane and cytoplasm to the cellular nucleus, and directly interact with β-catenin in gastric tumorigenesis. Moreover, HA-tagged β-catenin was transiently coexpressed with Flag-tagged TAZ in gastric AGS cells, and the cell lysates were subjected to immunoprecipitation with an anti-HA antibody, demonstrating the strong exogenous interaction between HA-β-catenin and Flag-TAZ (Figure 6C). This interaction was significantly enhanced by coinfection with CagA+ H. pylori strain NCTC11637 or 7.13 (Figure 6D). Our data and recent studies have indicated that CagA is required for H. pylori-induced TAZ and β-catenin (Yong et al., 2016); w therefore, we investigated whether CagA is involved in the interaction between TAZ and β-catenin in response to H. pylori infection. Coimmunoprecipitation of TAZ with β-catenin in wild-type H. pylori-infected cells was stronger than that in CagA-deficient mutant strain-infected cells (Figure 6E). These data collectively indicated that H. pylori CagA plays an important role in the H. pylori-enhanced interaction between TAZ and β-catenin.

As our data indicated that H. pylori infection promotes the Wnt/β-catenin signaling pathway via the activation of TAZ, we next investigated whether β-catenin plays a role in regulating TAZ-mediated phenotypic alterations of gastric cancer cells. TAZ overexpression induced gastric cell proliferation, which was blocked by β-catenin siRNA (Figure 7A). Furthermore, β-catenin knockdown alleviated TAZ-induced gastric cancer cell migration and invasion (Figures 7B,C). Taken together, knockdown of β-catenin suppressed TAZ-mediated cell proliferation, migration and invasion.