Methylmercury induces ferroptosis by suppressing GPX4 protein levels in C17.2 mouse neural stem cells

Abstract

The causative agent of Minamata disease, which is characterized primarily by severe central nervous system dysfunction, is methylmercury; however, the mechanisms underlying methylmercury toxicity remain unclear. Ferroptosis is a type of programmed cell death that is mediated by iron-dependent lipid peroxidation and methylmercury has long been suggested to cause neuronal damage via lipid peroxidation, although the detailed mechanism of this action remains unknown. In this study we therefore investigated the involvement of ferroptosis in methylmercury-induced neuronal cell death using C17.2 mouse neural stem cells. First, we examined the effects of various ferroptosis inhibitors (ferrostatin-1, liproxstatin-1, and deferoxamine) on methylmercury-induced cell death. All the inhibitors tested attenuated methylmercury-induced cell death. We then examined the levels of intracellular reactive oxygen species and lipid peroxides, and found that these levels were increased prior to methylmercury-induced cell death. We also examined the levels of a cystine transporter (xCT/SLC7A11) and glutathione peroxidase 4 (GPX4), major factors involved in the suppression of ferroptosis. We found that both mRNA and protein levels of xCT were increased prior to methylmercury-induced cell death, whereas GPX4 mRNA levels were largely unaffected by methylmercury and its protein levels were decreased. C17.2 cells overexpressing FLAG-GPX4 exhibited greater resistance to methylmercury than control cells. These results indicate that methylmercury induces ferroptosis in C17.2 cells by suppressing GPX4 protein levels.

INTRODUCTION

Mercury is a toxic heavy metal that is widely distributed in the environment through natural processes and human activities. Its toxicity varies greatly depending on its chemical form. Minamata disease, characterized by severe central nervous system dysfunction, first arose in 1956 in Minamata, Japan in individuals who had consumed fish and shellfish heavily contaminated with methylmercury from industrial wastewater (Eto, 1997; Sakamoto et al., 2025). Methylmercury in seawater bioaccumulates in the food chain, concentrating in large fish, which can then be consumed by humans (Yoshimoto et al., 2016). Some of the methylmercury absorbed into the body binds to free cysteine in the blood, crosses the blood–brain barrier via neutral amino acid transporters, and accumulates in the brain (Simmons-Willis et al., 2002). Recently, reports have shown that children born to women who consumed seafood containing relatively high amounts of methylmercury during pregnancy exhibit motor and mental developmental disorders (Jacobson et al., 2015; Tatsuta et al., 2017, 2018), raising concerns about the adverse effects of methylmercury exposure on fetal neurodevelopment. Several research groups, including ours, have investigated the mechanisms involved in methylmercury toxicity and defense against it. In particular, tumor necrosis factor-α (Iwai-Shimada et al., 2016; Nakano et al., 2024), tumor necrosis factor receptor 3 (Toyama et al., 2023), oxidative stress-induced growth inhibitor 1 (Yamashita et al., 2024), and thioredoxin-interacting protein (Fukushima et al., 2025) are involved in enhancing methylmercury toxicity. In contrast, ornithine decarboxylase (Sato et al., 2018), transcription factor 3 (Toyama et al., 2021), sulfiredoxin-1 (Yamashita et al., 2025), and p62/SQSTM1 (Takanezawa et al., 2023) are involved in reducing methylmercury toxicity. However, the mechanisms underlying methylmercury toxicity still remain unclear, and further research is needed.

Ferroptosis is a recently described, novel type of programmed cell death (Li et al., 2020). Increased levels of intracellular free iron ions (Fe²⁺) generate reactive oxygen species (ROS) via the Fenton reaction. This increase in ROS promotes lipid peroxidation, thereby accelerating the formation of lipid radicals. Glutathione peroxidase 4 (GPX4) reduces lipid peroxide levels and exerts a protective effect on cells. However, intracellular glutathione depletion caused by reduced GPX4 activity or inhibition of the cystine transporter (xCT/SLC7A11) leads to the accumulation of ROS and lipid peroxidation, resulting in the induction of ferroptosis. xCT and GPX4 therefore play a central role in suppressing ferroptosis, and their dysfunction leads to increased susceptibility to ferroptosis (Dixon et al., 2012; Yang et al., 2014). Decreased levels of xCT and/or GPX4 and increased lipid peroxidation have been reported in models of neurological diseases, such as Parkinson’s disease (Liu et al., 2025), Alzheimer’s disease (Greenough et al., 2022), and ischemic brain injury (Lin et al., 2022), indicating the involvement of ferroptosis in neurodegenerative diseases.

Methylmercury induces not only necrosis but also apoptosis, a type of programmed cell death (Fujimura et al., 2009; Iijima et al., 2024; Watanabe et al., 2013). Furthermore, ROS production and lipid peroxidation are increased in the brains of mice treated with methylmercury (Andersen and Andersen, 1993; Sarafian and Verity, 1991). In addition, vitamin E, which is a peroxyl radical-scavenging antioxidant and inhibitor of lipid peroxidation, suppresses neuronal cell death caused by methylmercury (Kasuya, 1975). These findings indicate that ferroptosis, as well as apoptosis and necrosis, may be involved in methylmercury-induced neuronal cell death. Recently, methylmercury was shown to induce ferroptosis in rat brain astrocytes cells (Xu et al., 2023) and human embryonic kidney-derived cell lines (Chen et al., 2023). However, the involvement of ferroptosis in methylmercury-induced damage to central nervous system neurons has not been investigated. In this study, we investigated the involvement of ferroptosis in methylmercury-induced neuronal cell death and examined the underlying mechanism of this involvement using C17.2 mouse neural stem cells.

MATERIALS AND METHODS

Culture conditions

The mouse neural progenitor stem cell line, C17.2, was purchased from the European Collection of Cell Cultures. Cells were cultured in Dulbecco’s modified Eagle’s medium (Shimadzu Diagnostics Corporation, Tokyo, Japan) supplemented with 10% fetal bovine serum and 2 mM L-glutamine (Nacalai Tesque, Kyoto, Japan) in a humidified atmosphere of 5% CO₂ at 37°C.

Measurement of cell viability

Cell viability was evaluated using an alamarBlue assay (Invitrogen, Camarillo, CA, USA), according to the manufacturer’s protocol. Fluorescence was measured using a SpectraMax iD5 (Molecular Devices, Sunnyvale, CA, USA) at excitation and emission wavelengths of 540 nm and 590 nm, respectively.

Detection of intracellular ROS and lipid peroxidation

C17.2 cells were seeded onto bovine gelatin (Sigma-Aldrich, St. Louis, MO, USA) -coated PhenoPlate 96-well microplates (PerkinElmer, Waltham, MA, USA) at 1 × 10⁴ cells/well. Twenty-four hr after seeding, cells were exposed to methylmercury chloride (MeHgCl) (4 μM) for 2, 4, or 6 hr. Following removal of the culture medium, cells were incubated with 2 μM CellROX™ Green Reagent (Invitrogen, Carlsbad, CA, USA) or 2 μM BODIPY 581/591 C11 (Invitrogen) and Hoechst 33342 (Nacalai Tesque) for 30 min at 37°C in the dark. The wells were then gently washed twice with PBS, and fluorescence was measured using an Operetta CLS high-content imaging system (PerkinElmer). Each fluorescence intensity value was normalized to Hoechst 33342.

Quantitative polymerase chain reaction (qPCR)

Total RNA was isolated using ISOGEN II (Nippon Gene, Tokyo, Japan) and cDNA was synthesized from total RNA using a PrimeScript RT reagent kit (Takara, Shiga, Japan). qPCR was performed using SYBR Premix Ex Taq (Takara) and a LightCycler 96 System (Roche Diagnostics, Mannheim, Germany) with the following primers: mouse GPX4, F: 5′-GCTGGGAAATGCCATCAAATGGA-3′, R: 5′-ATGGGACCATAGCGCTTCACCA-3′; mouse xCT (SLC7A11), F: 5′-CCTGGGCAGGAGAAGGTAGT-3′, R: AGTATGCCCTTGGGGGAGAT-3′; mouse glyceraldehyde-3-phosphate dehydrogenase (GAPDH), F: 5′-AACTTTGGCATTGTGGAAGG-3′, R: 5′-ACACATTGGGGGTAGGAACA-3′. Each mRNA level was normalized against that of GAPDH using the relative standard curve method.

Western blotting

Cells were lysed in 2% sodium dodecyl sulfate (SDS) buffer and then incubated at 95°C for 5 min. Protein concentrations were measured using a DC Protein Assay Kit (Bio-Rad, Hercules, CA, USA). Cell lysates were separated by SDS–polyacrylamide gel electrophoresis and transferred to polyvinylidene fluoride membranes. The membranes were blocked for 1 hr in 5% skim milk (Nacalai Tesque). Western blotting was performed with anti-xCT/SLC7A11 (98051; Cell Signaling Technology, Danvers, MA, USA), anti-GPX4 (67763-1-1g; Proteintech, Chicago, IL, USA), and anti-GAPDH (015-25473; Fujifilm WAKO) primary antibodies, and horseradish peroxidase-conjugated secondary antibodies (Dako A/S, Glostrup, Denmark). Chemiluminescent images were obtained using a ChemiDoc Touch Imaging System and analyzed using Image Lab Software (Bio-Rad).

Plasmid construction and transfection

The coding sequence of mouse GPX4 (NM_001367995.1) with 3′ UTR containing the selenocysteine insertion sequence was amplified from C17.2 cell cDNA using KOD-plus neo (Toyobo, Osaka, Japan) and the primers, 5′-ATCATGTGTGCATCCCGCGATGATTG-3′ and 5′-TGAAGCTCGAGCCCAGGGCCACAG-3′ and inserted between the EcoRV (New England Biolabs, Beverly, MA, USA) and XhoI (New England Biolabs) sites of pCMV-3Tag-6 (Agilent Technologies, Santa Clara, CA, USA). C17.2 cells were transfected with control vector or FLAG-GPX4-expressing plasmid using PEI Max Reagent. Forty-eight hr after transfection, cells were exposed to methylmercury at the indicated concentrations. Cell viability was measured using an alamarBlue assay.

Statistical analysis

Data are expressed as mean ± SD. Multiple group comparisons were conducted using one-way analysis of variance followed by a post hoc Dunnett’s test. For two group comparisons, an unpaired Student’s t-test was used. Statistical analyses were performed using KaleidaGraph software (v4.1.1; Synergy Software, Eden Prairie, MN, USA), with statistical significance set at p < 0.05.

RESULTS

To clarify whether ferroptosis is involved in methylmercury-induced cell death, we examined the sensitivity of C17.2 cells to methylmercury after pretreatment with various ferroptosis inhibitors (deferoxamine, ferrostatin-1, and liproxstatin-1). All ferroptosis inhibitors attenuated methylmercury-induced cell death in a concentration-dependent manner (Fig. 1A–C).

Fig. 1

Effects of ferroptosis inhibitors on methylmercury-induced cell death. Cells were seeded into 24-well plates at 5 × 10⁴ cells/well. After incubation for 22 hr, the cells were pretreated with (A) deferoxamine (n=5), (B) ferrostatin-1 (n=3), and (C) liproxstatin-1 (n=3) for 2 hr and then exposed to the indicated concentrations of methylmercury chloride (MeHgCl) for 24 hr in the presence of each inhibitor. Cell viability was measured using an alamarBlue assay. Statistically significant compared with each control, as determined by Dunnett’s multiple comparisons test (*p < 0.05; **p < 0.01). Data are presented as the mean ± SD.

We next investigated the roles of ROS and lipid peroxidation, which are involved in ferroptosis, in methylmercury-induced cell death. Treatment of C17.2 cells with 4 µM methylmercury induced cell death in a time-dependent manner with the degree of cell death significant after 6 hr (Fig. 2A). Therefore, in subsequent experiments, the duration of methylmercury treatment was limited to 6 hr. The Fenton reaction is triggered by intracellular iron and increases intracellular ROS levels. Therefore, we examined the effect of methylmercury on ROS production using CellROX Green Reagent. We observed an increase in ROS levels after 4 hr of methylmercury treatment, indicating that ROS production is enhanced prior to the induction of cell death (Fig. 2B). ROS production promotes lipid oxidation and the excessive accumulation of lipid peroxides, thereby inducing ferroptosis. Therefore, we evaluated the effect of methylmercury on lipid peroxidation using BODIPY 581/591 C11 reagent. The amount of oxidized C11-BODIPY (indicating lipid peroxide levels) in C17.2 cells increased with methylmercury treatment in a time-dependent manner (Fig. 2C). These results indicate that methylmercury induces C17.2 cell death by triggering ferroptosis.

Fig. 2

Effects of methylmercury on cell death, intracellular reactive oxygen species, and lipid peroxidation. (A) Cells were seeded into 24-well plates at 5 × 10⁴ cells/well. After incubation for 24 hr, the cells were exposed to 4 µM methylmercury chloride (MeHgCl) for the indicated times. Cell viability was measured using an alamarBlue assay (n=5). (B, C) Cells were seeded on gelatin-coated PhenoPlate 96-well microplates at 1×10⁴ cells/well. After 24 hr, cells were exposed to 4 µM MeHgCl for 2, 4, and 6 hr. Following removal of the culture medium, cells were incubated with (B) CellROX Green Reagent (n=4) or (C) BODIPY 581/591 C11 (n=3) and Hoechst 33342 for 30 min at 37°C in the dark. The wells were then washed twice with PBS, and fluorescence was measured using an Operetta CLS high-content imaging system. Statistically significant compared with control, as determined by Dunnett’s multiple comparisons test (*p < 0.05; **p < 0.01). Data are presented as the mean ± SD.

As mentioned in the introduction, the cystine transporter, xCT, and GPX4, which inhibits lipid peroxidation in a glutathione-dependent manner, are well-known inhibitors of ferroptosis. Therefore, we examined the effects of methylmercury on the levels of these factors. Both mRNA and protein levels of xCT were increased prior to methylmercury-induced cell death (Fig. 3A, B). In contrast, GPX4 mRNA levels were slightly decreased by methylmercury, and the decrease in GPX4 protein levels was more pronounced than the decrease in its mRNA levels (Fig. 3C, D). These findings indicate that a decrease in GPX4 protein levels may be involved in the induction of ferroptosis by methylmercury.

Fig. 3

Effects of methylmercury on levels of xCT and GPX4 mRNA and protein. Cells were seeded into 24-well plates at 5 × 10⁴ cells/well. After incubation for 24 hr, the cells were exposed to 4 µM methylmercury chloride (MeHgCl) for 2, 4, and 6 hr (n=3). (A, C) xCT and GPX4 mRNA levels were determined by qPCR and the relative values normalized to GAPDH. (B, D) xCT and GPX4 protein levels were determined by western blotting (n=3). Statistically significant compared with control, as determined by Dunnett’s multiple comparisons test (*p < 0.05; **p < 0.01). Data are presented as the mean ± SD.

Finally, to clarify whether the decrease in GPX4 actually contributes to methylmercury-induced ferroptosis, we generated C17.2 cells overexpressing FLAG-tagged GPX4 (FLAG-GPX4) and examined their sensitivity to methylmercury. Overexpression of FLAG-GPX4 attenuated the sensitivity of C17.2 cells to methylmercury (Fig. 4A). We confirmed the overexpression of FLAG-GPX4 in C17.2 cells by western blotting using an anti-GPX4 antibody (Fig. 4B). These results strongly indicate that methylmercury induces ferroptosis in C17.2 cells by suppressing the levels of GPX4 protein.

Fig. 4

Effects of GPX4 overexpression on methylmercury-induced cell death. Cells were seeded into 24-well plates at 2 × 10⁴ cells/well. After incubation for 24 hr, the cells were transfected with an empty vector (Control) or FLAG-GPX4-expressing plasmid. Forty-eight hr later, the cells were exposed to the indicated concentrations of methylmercury chloride (MeHgCl) for 24 hr. (A) Cell viability was measured using an alamarBlue assay (n=6). Statistically significant compared with control, as determined by unpaired Student’s t-test (*p < 0.05; **p < 0.01). Data are presented as the mean ± SD. (B) Protein levels were detected by western blotting.

DISCUSSION

The findings of this study indicate that methylmercury induces ferroptosis in neuronal cells by suppressing GPX4 protein levels. Previous studies have reported that methylmercury induces ferroptosis in rat brain astrocytes cells, and that this may be mediated by the suppression of xCT and GPX4 (Xu et al., 2023). However, as shown in Fig. 3A, both mRNA and protein levels of xCT were significantly increased in C17.2 cells prior to methylmercury-induced cell death. This indicates that the effect of methylmercury on xCT levels differs depending on the cell type. At least in C17.2 cells, exposure to methylmercury may increase xCT levels and promote the synthesis of glutathione, a factor that alleviates methylmercury toxicity. Furthermore, the transcription factor, NRF2, is directly involved in inducing expression of the gene encoding xCT (Sasaki et al., 2002), and methylmercury activates NRF2 (Toyama et al., 2007); therefore, NRF2 may be involved in the induction of xCT gene expression by methylmercury. However, despite a marked increase in xCT levels prior to methylmercury-induced cell death, the protein levels of its downstream factor, GPX4, were decreased. Additionally, overexpression of FLAG-GPX4 suppressed methylmercury-induced cell death, indicating that suppression of GPX4 protein levels by methylmercury contributes to ferroptosis induction in C17.2 cells. Furthermore, the data in Fig. 3B indicate that methylmercury may suppress GPX4 levels through a post-translational mechanism rather than transcription. Autophagy and the ubiquitin-proteasome pathway are involved in the degradation of GPX4 protein (Ding et al., 2021; Xie et al., 2023; Xue et al., 2023) and conjugated fatty acids promote chaperone-mediated autophagy-dependent GPX4 degradation through mitochondrial ROS production (Hirata et al., 2024). We previously reported that methylmercury induces mitochondrial dysfunction and ROS generation in C17.2 cells (Sato et al., 2020). We have also observed that methylmercury can cause iron accumulation in the mitochondria of C17.2 cells (data not shown). Therefore, we suggested that mitochondrial dysfunction-mediated enhancement of GPX4 degradation may be involved in methylmercury-induced ferroptosis. Previous studies have shown that sodium selenite (Na₂SeO₃) pretreatment attenuated methylmercury-induced GPX4 downregulation in HEK293T cells (Chen et al., 2023). However, in C17.2 cells, although GPX4 protein levels were increased by Na₂SeO₃ pretreatment, the levels were still decreased by methylmercury under these conditions (data not shown). This result suggests that a mechanism other than intracellular selenium levels may be involved in the methylmercury-induced GPX4 downregulation in C17.2 cells.

Further investigation into the details of GPX4 suppression by methylmercury will contribute to determining the mechanism underlying methylmercury-induced ferroptosis.

As shown in Fig. 1, various ferroptosis inhibitors suppressed methylmercury-induced cell death by approximately 30%. In contrast, overexpression of FLAG-GPX4 suppressed cell death by approximately 10% (see Fig. 4), a weaker effect than that of the ferroptosis inhibitors. This is because methylmercury suppressed the expression of both plasmid-derived FLAG-GPX4 and endogenous GPX4 (data not shown), indicating that the suppression of ferroptosis by FLAG-GPX4 may have been limited. GPX4 has three isoforms: cytosolic isoform (cGPX4), mitochondrial isoform (mGPX4), and nucleolar isoform (nGPX4). Among these isoforms, cGPX4 is mainly involved in embryo development and ferroptosis suppression (Liang et al., 2009. Zheng and Conrad, 2020). Therefore, this study used a plasmid expressing cGPX4. However, mGPX4 has also been reported to suppress mitochondrial lipid peroxidation and ferroptosis (Gan, 2021). We plan to investigate the role of mGPX4 in methylmercury-induced ferroptosis in the future. In addition, besides xCT and GPX4, there are other regulators of ferroptosis, including factors that affect intracellular iron levels, such as iron storage factors (Fang et al., 2021) and iron chaperones (Jiang et al., 2024), as well as mechanosensitive cation channels (Hirata et al., 2023). Therefore, to fully elucidate the mechanism underlying methylmercury-induced ferroptosis, it is necessary to investigate the effects of methylmercury on ferroptosis regulators other than GPX4.

This study demonstrates the involvement of ferroptosis in methylmercury-induced neuronal cell death, and that methylmercury induces ferroptosis by suppressing GPX4 levels. Investigation using a mouse model of methylmercury-induced motor dysfunction will help clarify the relationship between methylmercury toxicity and ferroptosis.

ACKNOWLEDGMENTS

The authors would like to thank Fei Wu for technical assistance with the experiments and Jeremy Allen, PhD, from Edanz (https://jp.edanz.com/ac) for editing a draft of this manuscript.

Funding

This work was partially supported by JSPS KAKENHI Grant Number JP22H03752 for scientific research from the Ministry of Education, Culture, Sports, Science and Technology of Japan and the Study of the Health Effects of Heavy Metals organized by the Ministry of the Environment of Japan.

Conflict of interest

The authors declare that there is no conflict of interest.

Data availability

The data in this study are included in the article/supplementary materials. Contact the corresponding author directly to request the underlying data.

Author contributions

Conceptualization: Naoya Yamashita, Haruka Nozuki, Gi-Wook Hwang

Funding acquisition: Gi-Wook Hwang

Investigation: Naoya Yamashita, Haruka Nozuki, Tomoshi Yamashita, Kyotaro Tsubaki, Ryoko Fukushima, Ryota Yamagata, Gi-Wook Hwang

Supervision: Naoya Yamashita, Ryoko Fukushima, Ryota Yamagata, Gi-Wook Hwang

Visualization: Naoya Yamashita, Haruka Nozuki, Tomoshi Yamashita, Kyotaro Tsubaki, Ryoko Fukushima

Writing – original draft: Naoya Yamashita, Haruka Nozuki, Tomoshi Yamashita, Ryota Yamagata, Gi-Wook Hwang

Writing – review & editing: Naoya Yamashita, Ryota Yamagata, Gi-Wook Hwang

Ethical approval and consent to participate

Not applicable.

Patient consent for publication

Not applicable.

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