The Important Role of GPX1 and NF-ĸB Signaling Pathway in Human Gastric Cancer: Implications for Cell Proliferation and Invasion

Abstract

Background/Aim

Glutathione peroxidases (GPXs) are crucial antioxidant enzymes, counteracting reactive oxygen species (ROS). GPX overexpression promotes proliferation and invasion in cancer cells. Glutathione peroxidase-1 (GPX1), the most abundant isoform, contributes to invasion, migration, cisplatin resistance, and proliferation in various cancers. Nuclear factor-kappa B (NF-ĸB) participates in cell proliferation, apoptosis, and tumor progression. The inhibition of NF-ĸB expression reduces the malignancy of esophageal squamous cell carcinoma. This study aimed to explore the GPX1 and NF-ĸB signaling pathways and their correlation with gastric cancer cell proliferation and invasion.

Materials and Methods

Materials and Methods: Cell culture, complementary DNA microarray analysis, western blotting, reverse transcription-polymerase chain reaction, zymography, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay, GPX1 knock-down with short hairpin RNA (shRNA), standard two-chamber invasion assay, chromatin immunoprecipitation assay.

Results

Hepatocyte growth factor (HGF) up-regulated GPX1 expression in gastric cancer cells. The NF-ĸB inhibitor, pyrrolidine dithiocarbamate down-regulated HGF-induced GPX1 protein levels. Furthermore, NF-ĸB and urokinase-type plasminogen activators were down-regulated in GPX1-shRNA-treated cells. Treatment with an Akt pathway inhibitor (LY294002) led to the down-regulation of GPX1 and NF-ĸB gastric cancer cells. GPX1 knockdown resulted in decreased HGF-mediated in vitro cell proliferation and invasion. The study identified the putative binding site of the GPX1 promoter containing the NF-ĸB binding site, confirmed through chromatin immunoprecipitation.

Conclusion

HGF induced GPX1 expression through the NF-ĸB and Akt pathways, suggesting a central role in gastric cell proliferation and invasion. Hence, GPX1 emerges as a potential therapeutic target for gastric cancer.

Gastric cancer presents formidable treatment challenges due to its aggressive nature, rapid progression, and tendency for hematogenous and lymphatic spread (1-3). The multi-step progression includes involvement of growth factors, such as hepatocyte growth factor (HGF), vascular endothelial growth factor (VEGF), epidermal growth factor (EGF), fibroblast growth factor (FGF), and transforming growth factor (TGF)-α (4). Notably, HGF has been linked to gastric cancer (5).

HGF is produced by mesenchymal cells, a type of connective tissue cell (5). Its primary impact is on growth, differentiation, and survival of hepatocytes (5). Furthermore, HGF is expressed in various tissues and cell types and is involved in various physiological functions, such as angiogenesis and inflammation regulation (6). HGF promotes cell proliferation, survival, and invasion (6). HGF activates signaling pathways crucial for tumor growth and metastasis, including the phosphoinositide 3-kinase (PI3K)/protein kinase B (Akt) pathway (5,7-9). In gastric cancer cells, HGF induces vascularization through the PI3K/Akt pathway (10).

Glutathione peroxidases (GPXs) are a family of eight antioxidant enzymes, all of which are located on different chromosomes (11). The primary role of GPXs is to efficiently remove reactive oxygen species (ROS) from the body, thereby protecting against oxidative stress and DNA damage (12). Abnormal expression of GPXs has been widely observed in studies investigating cancer development when it plays a role in promoting cell proliferation, invasion, and metastasis (11,13). Among the eight GPX groups, glutathione peroxidase-1 (GPX1) is the most abundant isoform and is highly responsive to oxidative damage (11). GPX1 overexpression is associated with cancer development and cisplatin resistance in diverse cancer types, including colon, breast, lung, bladder, prostate, ovarian, and oral adenoid cystic carcinomas (13-16). Additionally, nuclear factor-kappa B (NF-ĸB) can act as an upstream regulator of GPX1 expression (14,17).

NF-ĸB is a ubiquitous nuclear transcription factor that plays a crucial role in diverse cellular processes, including cell apoptosis, proliferation, and malignant progression in specific cancer cells (13,18). In specific cancers, such as esophageal squamous cell carcinoma, prostate cancer, and melanoma, anti-inflammatory and antioxidant agents, including vitamin D, have demonstrated the ability to suppress GPX1 gene expression through the NF-ĸB pathway, leading to decreased malignancy (13). GPX1 inhibition reduced the invasion and migration of cancer cells, often associated with a decline in the expression or activity of urokinase-type plasminogen activator (uPA) (13,14,19).

However, there are reports showing that GPX1 and GPX3 are down-regulated in gastric cancer, but little is known on the signaling pathway or relationship between GPXs and gastric cancer (20). The current study identified the signaling pathway triggered by HGF in gastric cancer cell lines, such as NUGC3 and MKN28, mediated by NF-ĸB and GPX1. The impact of these signaling pathways on cancer progression was elucidated in vitro, presenting a novel target in gastric cancer.

Materials and Methods

Cell culture. Two human gastric cancer cell lines, poorly differentiated adenocarcinoma (NUGC3) and moderately differentiated tubular adenocarcinoma (MKN28) were obtained from the Korea Cell Line Bank in Seoul, Republic of Korea. Cells were cultured in RPMI 1640 medium (Life Technologies Inc., Gaithersburg, MD, USA) supplemented with 10% fetal bovine serum (FBS). The cells were maintained in a CO₂ incubator at 37˚C under a humidified atmosphere consisting of 5% CO₂ and 95% air.

Semi-quantitative reverse transcription-polymerase chain reaction. Complementary DNA (cDNA) was synthesized using MMLV reverse transcriptase (Promega Corp., Madison, WI, USA) with the oligo (dT) priming method in a 10 μl reaction mixture. Polymerase chain reaction (PCR) was performed in a 10 μl reaction volume containing 10 mM Tris-HCl (pH 8.5), 50 mM KCl, 1 μl cDNA, 200 μM dNTPs, 1 mM MgSO₄, 1 U Platinum Pfx Taq polymerase, and 2 μM primers. The PCR protocol included an initial denaturation at 95˚C for 4 min; 27 cycles at 94˚C for 15 s, 60˚C for 15 s, and 72˚C for 30 s; and a final extension at 72˚C for 10 min. PCR products were separated on a 1.5% agarose gel containing SYBR™ Green I Nucleic Acid Gel Stain (Thermo Fisher Scientific, Waltham, MA, USA) and visualized using a UV Vilber (Vilber, San Sebastiàn, Spain).

Complementary DNA microarray analysis. The cDNA microarray with 17,448 sequence-verified human cDNA clones was supplied by GenomicTree, Inc. (Daejeon, Republic of Korea). Experimental procedures on cDNA microarray followed the protocol outlined by Yang et al. (19). In brief, 100 μg of total RNA was reverse-transcribed in the presence of Cy3-dUTP or Cy5-dUTP (25 mM stock; NEN Life Science Products, Boston, MA, USA) at 42˚C for 2 h. The labeled cDNA was hybridized to the cDNA microarray at 65˚C for 16 h. Following hybridization, the slides were washed, scanned using an Axon GenePix 4000B microarray scanner (Axon Instruments, Union city, CA, USA), and analyzed using GenePix Pro 4.0 (Molecular Devices Corp.). Raw data were normalized and analyzed utilizing GeneSpring Software (Silicon Genetics version 6.0, Redwood City, CA, USA). Genes were filtered based on their control channel intensities. Genes with control channel values exceeding 80 in all samples were considered unreliable. Intensity-dependent normalization (LOWESS) was implemented, wherein the ratio was reduced to the residual of the LOWESS fit of the intensity-versus-ratio curve. The average normalized ratios were calculated by dividing the average normalized signal channel intensity by the average normalized control channel intensity. Welch’s ANOVA test was performed with p≤0.01 and 0.05 to pinpoint genes in differentially expressed samples. Correlation analysis involved Pearson correlation coefficients ranging from –1 to 1. Spots demonstrating changes greater than 2-fold were deemed significant.

Western blot analysis. The cells (5×10⁵) were harvested and treated with a lysis buffer containing 50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 10% glycerol, 1 mM PMSF, 1 mM sodium vanadate, and 5 mM NaF along with protease inhibitors. The mixture was then centrifuged at 15,000 rpm at 4˚C for 10 min. Proteins (50 μg) were separated on 10% sodium dodecyl sulfate-polyacrylamide gels and transferred to nitrocellulose membranes. These membranes were incubated with 5% non-fat dried milk in TTBS (composed of 10 mM Tris-HCl, pH 7.5, 150 mM NaCl, and 0.05% Tween-20) for 30 min and left overnight with a primary antibody at 4˚C. Following six washes with TTBS for 5 min each, the membranes were exposed to a horseradish peroxidase-conjugated secondary antibody for 1 h and 30 min at 4˚C. Subsequently, the membranes were washed three times with TTBS for 30 min each, and the antigen-antibody complexes were detected using an enhanced chemiluminescence detection system. The analyses were performed in a minimum of three independent experiments.

Zymography. The culture supernatants were denatured with a reducing agent subjected to electrophoresis on a 10% polyacrylamide gel containing 0.1% (w/v) plasminogen for urokinase-type plasminogen activator analysis. The gel was incubated at room temperature for 2 h in the presence of 2.5% Triton X-100 and subsequently at 37˚C overnight in a buffer containing 10 mM CaCl₂, 0.15 mM NaCl, and 50 mM Tris (pH 7.5). Following incubation, the gel was stained for proteins using a 0.25% Coomassie brilliant blue solution in methanol, acetic acid, and water (4:1:5) and then de-stained in the same solution without the dye. Enzyme activity was visualized in negatively stained regions. Zymographic analyses were performed in a minimum of three independent experiments.

Assessment of cell metabolic activity. The cells and GPX1-shRNA (1×10⁴) cells per well) were seeded in 96-well plates using RPMI 1640 medium supplemented with 5% FBS and incubated for 24 h. Subsequently, the cells were serum-starved for an additional 24 h and treated for 72 h with or without HGF (10 ng/ml). Following this incubation period, 50 μl of a 2 mg/ml solution of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) was added, and the cells were allowed to incubate for 3 h at 37˚C. After removing the supernatant via aspiration, the converted dye was dissolved in 100 μl of dimethyl sulfoxide. The plates were placed on a microplate shaker for 5 min, and the absorbance was measured at 570 nm using a Bio-Rad Multiscan plate reader (Hercules, CA, USA). Each result was the mean of triplicate assays.

GPX1 knockdown with short hairpin RNA. A human GPX1-specific shRNA expression vector (GPX1-shRNA), incorporating a GPX1-targeted shRNA sequence, was procured from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Cells were transfected with GPX1-shRNA using Lipofectamine (Life Technologies Inc., Gaithersburg, MD, USA). The clonal selection was performed by culturing with puromycin (10 μg/ml), and cells were serially diluted. Stable transfectant clones displaying low target gene expression were identified using western blotting.

Standard two-chamber invasion assay. Control cells and transfected cells (1×10³) were seeded in the upper chamber of a Matrigel migration chamber with 0.8-micron pores (Fisher Scientific, Houston, TX, USA) in media containing 5% FBS, with or without HGF (10 ng/ml). Following a 48-h incubation, the cells were fixed and stained using a HEMA 3 stain set (Curtis Matheson Scientific, Houston, TX, USA) according to the manufacturer's instructions. The stained filter membrane was cut and mounted on a glass slide. Migrated cells were counted under a light microscope (10 fields at 200× magnification). The assays were performed in a minimum of three independent experiments.

Chromatin immunoprecipitation assay. Following the manufacturer's instructions, a chromatin immunoprecipitation (ChIP) assay kit was used using an Upstate Biotechnology (Waltham, MA, USA) ChIP assay. Briefly, cells (5×10⁵) were fixed with 1% formaldehyde at 37˚C for 10 min. After two washes with ice-cold PBS containing protease inhibitors (1 mM phenylmethylsulphonyl fluoride, 1 mg/ml aprotinin, and 1 mg/ml pepstatin A), cells were scraped and pelleted via centrifugation at 4˚C. The cell pellet was resuspended in lysis buffer (1% SDS, 10 mM EDTA, and 50 mM Tris-HCl, pH 8.1), incubated for 10 min on ice, and sonicated to shear the DNA. Following sonication, the lysate was centrifuged for 10 min at 13,000 rpm at 4˚C. The resulting supernatant was diluted in ChIP dilution buffer (0.01% SDS, 1% Triton X-100, 2 mM EDTA, 16.7 mM Tris-HCl (pH 8.1, 167 mM NaCl, and protease inhibitors).

Primary antibodies were added, and the mixture was incubated with rotation overnight at 4˚C. The immunocomplex was then collected using protein A/G agarose beads and washed successively with low salt washing buffer (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 200 mM Tris-HCl, pH 8.1, and 150 mM NaCl), high salt buffer (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 200 mM Tris-HCl, pH 8.1, and 500 mM NaCl), LiCl washing buffer (0.25 M LiCl, 1% NP40, 1% deoxycholate, 1 mM EDTA, and 10 mM Tris-HCl, pH 8.1), and finally 1* TE buffer (10 mM Tris-HCl, and 1 mM EDTA, pH 8.0). Subsequently, the immunocomplex was eluted using elution buffer (1% SDS, 0.1 M NaHCO3, and 200 mM NaCl), and the cross-links were reversed by heating at 65˚C for 4 h. Following the reaction, the samples were adjusted to 10 mM EDTA, 20 mM Tris-HCl, pH 6.5, and 40 mg/ml proteinase K, and incubated at 4˚C for 1 h. DNA was then recovered and subjected to PCR amplification of the GPX1 promoter region using the primers 5’-gcctagatccctctggctgt-3’ (forward) and 5’-aggcggcagcggtccaggtt-3’ (reverse).

Reagents and antibodies. Horseradish peroxidase-conjugated anti-mouse and anti-rabbit antibodies were purchased from Bio-Rad (Hercules, CA, USA). Recombinant human HGF was obtained from R&D Systems, Inc. (Minneapolis, MN, USA) and purchased from Sigma-Aldrich (Saint Louis, MO, USA). Antibodies against phospho-AKT and AKT were sourced from Cell Signaling Technology (Beverly, MA, USA), while NF-ĸB antibodies were purchased from Santa Cruz (Santa Cruz). GPX1 antibodies were acquired from Abcam (Cambridge Biomedical Campus, Cambridge, UK).

Results

GPX1 is up-regulated by HGF in gastric cancer cells. We detected GPX1 expression in gastric cancer cells, such as NUGC3 and MKN28. We conducted reverse transcription-polymerase chain reaction (RT-PCR) to confirm GPX1 RNA expression levels, and western blot analysis to confirm GPX1 protein levels in gastric cancer cells with or without HGF. As expected, GPX1 mRNA levels increased 1 and 6 h after HGF treatment (Figure 1A). Furthermore, GPX1 protein levels increased by 1 and 6 h after HGF treatment (Figure 1B). To confirm the effect of HGF dosage, the degree of GPX1 mRNA and protein expressions were confirmed by varying the dosage of HGF (0, 10, and 40 ng/ml) in gastric cancer cells. As the HGF dose increased, PT-PCR revealed that GPX1 mRNA expression increased (Figure 2A). In addition, western blot analysis showed that GPX1 protein expression increased with increasing HGF dosage (Figure 2B). These results confirmed that HGF up-regulates GPX1 mRNA and protein expression in gastric cancer cells.

Figure 1. The impact of HGF on GPX1 expression in NGUC3 and MKN28 cells was assessed. Cells (5×10⁵) were serum-starved for 24 h, treated with or without HGF (10 ng/ml) for the specified durations, and subsequently harvested. GPX1 RNA expression levels were validated via reverse transcription-polymerase chain reaction (A). GPX1 protein expression levels were confirmed using western blot analysis (B). The presented data represent typical results obtained from three independent experiments. HGF, Hepatocyte growth factor; NGUC3, poorly differentiated adenocarcinoma; MKN28, moderately differentiated tubular adenocarcinoma; GPX1, glutathione peroxidase-1.

Figure 2. HGF up-regulated GPX1 expression in NGUC-3 and MKN-28 cells. Cells (5×10⁵) were serum-starved for 24 h and treated with varying concentrations of HGF (0, 10, and 40 ng/ml) for 1 h before harvesting. Expression levels of GPX1 RNA were validated using RT-PCR (A). Concurrently, the expression levels of GPX1 protein were confirmed using western blot analysis (B). Presented data are representative results from three independent experiments. HGF, Hepatocyte growth factor; NGUC3, poorly differentiated adenocarcinoma; MKN28, moderately differentiated tubular adenocarcinoma; GPX1, glutathione peroxidase-1; RT-PCR, reverse transcriptase polymerase chain reaction.

NF-ĸB is located upstream in the signaling pathway and up-regulates GPX1 expression. Pyrrolidine dithiocarbamate (PDTC) was used to examine the correlation between NF-ĸB and GPX1 expression in gastric cancer cells. PDTC is known as an inhibitor of NF-ĸB (21). We treated gastric cancer cells with HGF (0, 10 ng/ml) and PDTC according to the dosage (0, 50, 100, and 200 μM). NF-ĸB expression was down-regulated, as confirmed by western blot analysis (Figure 3). To confirm the correlation between GPX1 and NF-ĸB, GPX1 was knocked down using GPX1 shRNA, following which NF-ĸB protein expression level was confirmed. GPX1 expression was reduced by GPX1 shRNA in gastric cancer cells, but NF-ĸB expression was not considerably affected (Figure 4). These results indicate that NF-ĸB is located upstream on the pathway and up-regulates GPX1.

Figure 3. Effect of PDTC, an NF-ĸB inhibitor, on HGF-induced GPX1. Serum-starved cells were pre-treated with varying concentrations of PDTC (50, 100, and 200 μM) and then exposed to 10 ng/ml of HGF. The expression levels of GPX1 and NF-ĸB were assessed using western blot analysis. Presented data are representative results from three independent experiments. PDTC, Pyrrolidine dithiocarbamate; HGF, hepatocyte growth factor; GPX1, glutathione peroxidase-1; NF-ĸB, nuclear factor-kappa B; NGUC3, poorly differentiated adenocarcinoma; MKN28, moderately differentiated tubular adenocarcinoma.

Figure 4. The levels of NF-ĸB and GPX1 proteins in response to HGF were reduced in cells expressing GPX1-shRNA. Control cells and stable GPX1-shRNA cells (5×10⁵/well) were plated overnight in a complete medium, starved for 24 h, then treated with or without HGF (10 ng/ml) for 1 h before harvesting. Expression levels of NF-ĸB and GPX1 were analyzed using western blot analysis. Presented data are representative results from three independent experiments. NF-ĸB, Nuclear factor-kappa B; GPX1, glutathione peroxidase-1; shRNA, short hairpin RNA; NGUC3, poorly differentiated adenocarcinoma; MKN28, moderately differentiated tubular adenocarcinoma.

GPX1 regulated uPA expression and activity. HGF expression also up-regulated uPA (22). To confirm the relationship between GPX1 and uPA, we conducted western blot analysis and zymography following GPX1 knockdown by shRNA with HGF treatment (0, 10 ng/ml) to examine uPA expression and activity. In the absence of GPX1 shRNA treatment, uPA expression remained normal; however, GPX1 knockdown resulted in reduced uPA expression (Figure 5A). This was also confirmed by the zymography results (Figure 5B). Therefore, GPX1 was confirmed to regulate uPA expression and activity.

Figure 5. Expression levels and enzyme activity of the uPA protein in response to HGF were down-regulated in cells expressing GPX1-shRNA. Control cells and stable GPX1-shRNA cells (5×10⁵) were treated with or without HGF (10 ng/ml) and subsequently harvested. The levels of uPA proteins in the culture media were analyzed using western blot analysis (A). Furthermore, the enzyme activity levels of uPA proteins in the culture media were assessed by zymography (B). Presented data are representative results from three independent experiments. uPA, Urokinase-type plasminogen activator; HGF, hepatocyte growth factor; GPX1, glutathione peroxidase-1; shRNA, short hairpin RNA; NGUC3, poorly differentiated adenocarcinoma; MKN28, moderately differentiated tubular adenocarcinoma.

GPX1 and NF-ĸB are regulated by pAkt. HGF regulates signaling pathways by mediating the activation of the PI3K/Akt pathway, also known as pAkt (23). HGF binds to its receptor c-Met and triggers downstream signaling cascades, including activation (24,25). To investigate the correlation between pAkt, GPX1, and NF-ĸB, gastric cancer cells were treated with LY294002, a pAkt inhibitor, and analyzed using western blotting. Inhibition of pAkt expression by LY294002 treatment (1, 5, and 10 μg/ml) at various concentrations, confirmed the decrease in NF-ĸB and GPX1 expression decrease in both gastric cancer cell lines (NUGC3 and MKN28) (Figure 6A). Thus, the down-regulation of pAkt leads to reduced NF-ĸB and GPX1 expression.

Figure 6. Down-regulation of GPX1, NF-ĸB, and pAkt expression was observed, and was influenced by LY294002, acting as a pAkt inhibitor. Cells (5×10⁵) were pre-treated with different concentrations of LY294002 (1, 5, and 10 μM) and then treated with 10 ng/ml of HGF. The expression levels of GPX1, NF-ĸB, and pAkt proteins were evaluated by western blotting. Presented data are representative results from three independent experiments. GPX1, Glutathione peroxidase-1; NF-ĸB, nuclear factor-kappa B, pAkt; phosphor-protein kinase; NGUC3, poorly differentiated adenocarcinoma; MKN28, moderately differentiated tubular adenocarcinoma.

NF-ĸB is a transcription factor for binding the GPX1 promoter. To validate the interaction of NF-ĸB with the GPX1 promoter, a ChIP assay was conducted using an anti-NF-ĸB antibody following amplification of the GPX1 promoter (Figure 7). To confirm NF-ĸB binding activity on the GPX1 promoter, short-hairpin RNA of GPX1 was used to compare GPX1 knockdown and control cells. NF-ĸB binding was not observed in GPX1 knockdown cells but was observed in control cells. Moreover, we verified that the binding activity of NF-ĸB to the GPX1 promoter was augmented by HGF (10 ng/ml). Consequently, HGF enhanced the GPX1 promoter-binding activity of NF-ĸB; however, NF-ĸB binding was not observed in GPX1 knockdown cells, irrespective of the presence or absence of HGF. In conclusion, NF-κB can be speculated to bind to the GPX1 promoter and act as a transcription factor.

Figure 7. The effect of NF-ĸB knockdown on HGF regulation of GPX1 was investigated. ChIP assay results demonstrate the amplification of a fragment from the proximal GPX1 promoter, which contains the NF-ĸB binding site (A). Immunoprecipitation was conducted using an anti-NF-ĸB antibody (B). Presented data are representative results from three independent experiments. NF-ĸB, Nuclear factor-kappa B; HGF, hepatocyte growth factor; GPX1, glutathione peroxidase-1; CHIP; chromatin immunoprecipitation assay; NGUC3, poorly differentiated adenocarcinoma; MKN28, moderately differentiated tubular adenocarcinoma.

GPX1 is up-regulated by cell proliferation and invasion into gastric cells. To confirm the effect of GPX1 on cell proliferation, an MTT assay was performed with or without HGF treatment (10 ng/ml) for 72 h in GPX1 shRNA cells and control cells. Compared to control cells, HGF-induced cell proliferation was diminished in GPX1 knockdown cells (Figure 8). Furthermore, we conducted an in vitro invasion assay using Matrigel-coated migration chambers to assess the impact of GPX1 on cell invasion. The analysis of cell proliferation both in the absence or presence of HGF (10 ng/ml) for 48 h, in GPX1 knockdown and control cells revealed a decrease in HGF-mediated cell invasion in GPX1 knockdown cells compared to control cells (Figure 9). Consequently, GPX1 emerges as a crucial factor in regulating cancer cell proliferation, invasion, and migration.

Figure 8. The effect of GPX1 on cell proliferation in the presence of HGF was investigated. Control cells (1×103/well) and stable GPX1-shRNA cells were seeded in 96-well plates with DMEM supplemented with 5% FBS and incubated for 24 h. After serum starvation for an additional 24 h, cells were treated or untreated with HGF (10 ng/ml) for 72 h. Cell proliferation was measured using 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assays and expressed as a percentage of HGF-untreated control cells. The values represent means±standard deviation of three independent experiments. GPX1, Glutathione peroxidase-1; HGF, hepatocyte growth factor; shRNA, short hairpin RNA; DMEM, Dulbecco's Modified Eagle Medium, FBS; fetal bovine serum; NGUC3, poorly differentiated adenocarcinoma; MKN28, moderately differentiated tubular adenocarcinoma.

Figure 9. Impact of GPX1 on HGF-mediated cell invasion. Stable GPX1-shRNA cells and control cells (1×10³) were treated with or without HGF (10 ng/ml) for 48 h. Cell invasion capacity was measured using the standard two-chamber invasion assay with Matrigel migration chambers. The presented values represent means±standard deviation of three independent experiments. GPX1, Glutathione peroxidase-1; HGF, hepatocyte growth factor; shRNA, short hairpin RNA; NGUC3, poorly differentiated adenocarcinoma; MKN28, moderately differentiated tubular adenocarcinoma.

Discussion

Gastric cancer is one of the most common cancers in South Korea and worldwide (26). It has a relatively high prevalence; as of 2020, it is the fourth most common cancer in Korea (26,27). Gastric cancer is radioresistant, metastasizes to other organs, and has rapid progression (1). Clinical symptoms appear late, and survival rates are poor (28). Gastric cancer, notably, is frequently associated with peritoneal carcinomatosis, leading to a generally poor prognosis (29). Despite the development and utilization of various anticancer drugs, including targeted agents (VEGF and HER-2), and immune-oncologic agents known as checkpoint inhibitors, the treatment of gastric cancer continues to pose significant challenges, marked by a high mortality rate (30). The lack of effective targeted therapies against specific genetic mutations in gastric cancer is one of the reasons for the difficulty of its treatment. A recent and credible study has identified genes linked to peritoneal carcinomatosis, presenting them as potential targets for further investigation and intervention strategies (29). Therefore, novel targets are required to prevent or suppress rapid proliferation, metastasis, and carcinogenesis resulting from DNA damage.

Carcinogenesis can be attributed, in part, to DNA damage induced by ROS (12). The development of human cancer often involves mutations arising from the destabilization of genomic integrity due to ROS (12,18,31). Therefore, antioxidant mechanisms, including the activity of enzymes, for example, GPXs, are crucial for protecting cells from ROS-induced DNA damage and maintaining cellular health. The relationship between intracellular antioxidants, such as GPX, glutathione, and cancer is complex and can have varying effects depending on the context and stage of cancer (11,12,31-33). Intracellular antioxidant expression increases, ROS decreases, and carcinogenesis is enhanced (12,31).

The roles and functions of GPX1 in cancer have been reported in several types of carcinomas, such as prostate, esophageal, ovarian, and hepatocellular carcinomas (13,15,16,34). GPX1 contributes to cancer cell invasion and proliferation via uPA overexpression (35-38). Furthermore, the HGF-induced PI3K/Akt pathway is known to enhance the vascularization, migration, and invasion of gastric cancer cells (5,10). GPX1 expression is a known prognostic factor in gastric cancer (20,39). However, the mechanism by which the PI3K/Akt pathway, mediated by HGF and GPX1, activates the gastric cancer cell remains unclear. The present study experimentally demonstrated an in vitro correlation between PI3K/Akt pathway, mediated by HGF and GPX1 expression. Moreover, the presence or absence of GPX1 expression affected the proliferation and invasion of gastric cancer cells in vitro.

NF-ĸB plays an important role in proliferation, aromorphosis, apoptosis, and metastasis in carcinoma and gastrointestinal cancers (18,40). Many studies have confirmed changes in NF-ĸB expression and activity in carcinoma cells compared with those in normal tissues. Changes in the expression of downstream proteins have also been observed (41,42). GPX1 expression has already been shown to be regulated via the NF-κB pathway (41,43). However, this relationship remains unclear in gastric cancer cells. Therefore, this study confirms the regulation of GPX1 expression via the NF-ĸB pathway in gastric cancer cells in vitro.

The present study confirmed that HGF up-regulated GPX1 expression in NUGC3 and MKN28 cells (Figure 1 and Figure 2). Therefore, NF-ĸB was involved as a transcription factor in HGF-mediated GPX1 expression (Figure 3, Figure 4, and Figure 7). Additionally, GPX1 expression was affected by the AKT pathway and uPA expression and activity (Figure 5 and Figure 6). Finally, GPX1 knockdown reduced the proliferation and invasion of NUGC3 and MKN28 cells (Figure 8 and Figure 9). We presented the experimental process and results of this article in a study flow-chart (Figure 10) and schematic diagram (Figure 11 and Figure 12).

Figure 10. Study flow-chart. After cell culture and stimulation with HGF, the expression of GPX1 was analyzed at both mRNA and protein levels. Following inhibition of the NF-ĸB and Akt pathways, the levels of GPX1 were analyzed. Subsequently, to identify downstream genes, GPX1 knockdown was performed and analyzed. Then, using MTT assay and Two-chamber invasion assay, differences in cell proliferation and invasion were confirmed based on GPX1 knockdown. GPX1, Glutathione peroxidase-1; HGF, hepatocyte growth factor; shRNA, short hairpin RNA; NF-ĸB, nuclear factor-kappa B.

Figure 11. Schematic diagram of HGF-mediated GPX1 up-regulation, HGF mediated GPX1 up-regulation is regulated through the PI3K/Akt pathway and NF-ĸB, which was confirmed by pretreatment with LY294002 and PDTC, HGF-mediated GPX1 up-regulation increased expression uPA. Increased uPA mediates cell proliferation and invasion. GPX1, Glutathione peroxidase-1; HGF, hepatocyte growth factor; shRNA, short hairpin RNA; NF-ĸB, nuclear factor-kappa B; pAkt, phosphorprotein kinase; uPA, urokinase-type plasminogen activator; PDTC, pyrrolidine dithiocarbamate.

Figure 12. Schematic diagram of HGF-mediated GPX1 up-regulation, HGF mediated GPX1 up-regulation is regulated through the PI3K/Akt pathway and NF-ĸB, which was confirmed by pretreatment with LY294002 and PDTC, HGF-mediated GPX1 up-regulation increased expression uPA. Increased uPA mediates cell proliferation and invasion. GPX1, Glutathione peroxidase-1; HGF, hepatocyte growth factor; shRNA, short hairpin RNA; NF-ĸB, nuclear factor-kappa B; pAkt, phosphor-protein kinase; uPA, urokinase-type plasminogen activator; PDTC, pyrrolidine dithiocarbamate.

Our results suggest that GPX1 is a crucial factor in HGF-mediated cancer cell proliferation, invasion, and metastasis in gastric cancer. GPX1 up-regulation by HGF and its regulation by NF-ĸB highlight its importance in the progression of gastric cancer. These results have notable implications for identifying biomarkers and developing targeted therapeutic strategies for gastric cancer. GPX1 has emerged as a potential biomarker for assessing treatment efficacy and prognosis in patients with gastric cancer.

Monitoring GPX1 levels may provide valuable insights into the dynamic behavior of gastric cancer cells, and thus GPX1 could serve as a prognostic biomarker. This, in turn, could facilitate a more nuanced evaluation of treatment response and disease prognosis. This could guide clinicians in tailoring therapeutic interventions based on the individual molecular profiles of patients.

Targeting GPX1 as a therapeutic strategy represents a novel approach to combating gastric cancer. The involvement of GPX1 in critical cellular processes associated with cancer progression suggests that modulating its activity could potentially impede the malignant phenotype. However, the translational potential of this strategy necessitates a comprehensive understanding of the underlying molecular mechanisms and the development of targeted interventions that are both effective and safe clinically.

This study highlights the potential of GPX1 as both a biomarker and a therapeutic target for managing gastric cancer. Further research, including detailed molecular investigations and clinical trials, is essential to validate these observations and develop personalized and effective therapeutic approaches for patients with gastric cancer.

Conflicts of Interest

The Authors have no conflicting interests.

Authors’ Contributions

Kyung Hee Lee conceived and supervised the study. Byeong Il Jang, Sung Ae Koh, and Ji Yoon Jung designed the experiments. Byeong Il Jang performed the experiments. Byeong Il Jang, Sung Ae Koh, Ji Yoon Jung, and Kyung Hee Lee analyzed the data. Byeong Il Jang and Ji Yoon Jung wrote the manuscript. Sung Ae Koh and Kyung Hee Lee confirm the authenticity of all the raw data. All Authors read and approved the final version of the manuscript.