2026 Volume 51 Issue 5 Pages 315-320
Methylmercury (MeHg) is a potent environmental toxicant that frequently coexists with other heavy metals, raising concerns about combined toxic effects. Increasing evidence indicates that co-exposure to multiple metals can lead to synergistic or greater-than-additive effects; however, the molecular mechanisms underlying such interactions remain poorly understood. Among the tested metals, only co-exposure with Cd markedly enhanced MeHg cytotoxicity. Here, our objective was to evaluate the impact of MeHg co-exposure on cytotoxicity of various heavy metals in HeLa cells. We used cell viability assays, western blot analysis, and reverse-transcription-quantitative polymerase chain reactions to determine toxicity. Co-treatment with MeHg significantly reduced cell viability compared with that of Cd alone. Mechanistically, MeHg suppressed nuclear factor erythroid 2-related factor 2 (NRF2) expression more strongly at earlier time points than Cd alone, thereby impairing antioxidant and detoxification responses. This suppression was accompanied by increased intracellular mercury (Hg) retention, leading to enhanced cytotoxicity. Our results provide a mechanistic basis for metal–metal interactions and highlight the importance of considering co-exposure scenarios in environmental risk assessment.
Heavy metals such as cadmium (Cd), arsenic (As), and chromium (Cr), and methylmercury (MeHg) are major environmental hazards. While the toxic mechanisms of individual metals have been extensively studied, increasing evidence indicates that co-exposure to multiple metals can lead to synergistic or greater-than-additive toxic effects (Guo et al., 2022; van Strijp et al., 2023). However, the molecular mechanisms underlying such interactions remain poorly understood.
The transcription factor nuclear factor erythroid 2–related factor 2 (NRF2) acts as a master regulator of cellular defense against electrophiles and oxidative stress (Suzuki et al., 2023). Previous studies have reported conflicting effects of heavy metals on NRF2 regulation depending on the exposure context (Chen and Shaikh, 2009; Liu et al., 2019; Fan et al., 2021), and the impact of co-exposure remains unclear. Here, our objective was to determine the interactive toxicity of MeHg and Cd in HeLa cells, focusing on NRF2 regulation and intracellular Hg handling.
HeLa cells were cultured in DMEM (Sigma-Aldrich, St. Louis, MO, USA) supplemented with 10% fetal calf serum (FBS; Tissue Culture Biologicals, Seal Beach, CA, USA), 100 U/mL penicillin, 100 mg/L streptomycin, and 292 mg/L L-glutamine (Thermo Fisher Scientific, Rockford, IL, USA) following a previously established protocol (Takanezawa et al., 2023a). Cells were maintained at 37°C in a humidified atmosphere containing 5% CO2. Cells were seeded at a density of 3 × 105 cells/well in 6-well plates and allowed to attach overnight. The following day, cells were treated with (methylmercury chloride (MeHg) (Tokyo Kasei, Tokyo, Japan) and/or various concentrations of cadmium chloride (Cd) (Wako, Osaka, Japan), sodium arsenite (As) (Merck Millipore, Darmstadt, Germany), or hexavalent chromium (Cr) (Nacalai Tesque, Kyoto, Japan). Cd and MeHg were dissolved in water and dimethyl sulfoxide (DMSO), respectively, and control cells were treated with the same amount of water or DMSO.
Cell viability assayCell viability was evaluated by CCK-8 assay (Dojindo, Kumamoto, Japan). Cells were seeded in 96-well plates at a density of 1 × 104 cells/well and incubated for 24 hr with Cd and/or MeHg. Subsequently, 10 µL of CCK-8 solution was added to each well, followed by incubation at 37°C for 1 hr. Absorbance at 450 nm was measured using an iMark microplate reader (Bio-Rad, Hercules, CA, USA). Cell viability was presented as a percentage relative to the control group.
Western blot analysisCell lysates were prepared using RIPA buffer (20 mM Tris pH 7.4, 0.1% sodium dodecyl sulfate, 1% sodium deoxycholate, 1% Nonidet P40) containing protease inhibitors (Cell Signaling Technology, Danvers, MA, USA). Protein concentrations were determined using the BCA assay (Thermo Fisher Scientific). Proteins (10 μg) were resolved by SDS-PAGE and subsequently transferred onto polyvinylidene difluoride membranes. Membranes were blocked with 5% non-fat milk (Nacalai Tesque) in TTBS and incubated overnight at 4°C with primary antibodies against Nrf2 (Medical and Biological Laboratory, Nagoya, Japan, 1:1000), HO-1 (Cell Signaling Technology, 1:1000) and GAPDH (FUJIFILM Wako Pure Chemical Corporation, Osaka, Japan, 1:2000). Blots were washed and incubated with horseradish peroxidase-conjugated secondary antibodies (GE Healthcare, Buckinghamshire, United Kingdom 1:6000), and signals were detected using an ECL detection kit (Nacalai Tesque).
Reverse transcription-quantitative polymerase chain reaction (RT-qPCR)Total RNA was extracted and reverse transcription was performed following a previously described method (Takanezawa et al., 2023b). RT-PCR was performed in duplicate using PowerUp SYBR Green Master Mix (Thermo Fisher Scientific) with a CFX-96 thermal cycler system (Bio-Rad). The primers used for PCR were: Human NRF2: forward, 5′-CCAGGTTGCCCACATTCCCA-3′ and reverse, 5′-CGTAGCCGAAGAAACCTCATTGT-3′; Human GAPDH: forward, 5′- GCCAAGGTCATCCATGACAACT-3′ and reverse, 5′-GAGGGGCCATCCACAGTCTT-3′. GAPDH mRNA levels were used for normalization. Quantification cycle (Cq) values were recorded, and relative gene expression was calculated using the 2-∆∆Ct method.
Measurement of intracellular Hg and CdFollowing treatment, cells were washed with phosphate-buffered saline (PBS) and collected by scraping. The cells were divided into two tubes and centrifuged for 3 min at 220 ×g for 3 min to pellet the cells. Mercury levels were measured by solubilizing the cell pellets in RIPA buffer and analyzing them using a Mercury Analyzer MA3 solo (Nippon Instruments Co., Tokyo, Japan). Cadmium (Cd) levels were determined by digestion in nitric acid, Cd concentration was then analyzed by inductively coupled plasma-optical emission spectroscopy (ICP-OES; iCAP 7400 Duo, Thermo Fisher Scientific) as previously described (Nakamura et al., 2022). Mercury and cadmium concentrations were normalized to the total protein content, which was determined using the BCA assay.
Statistical analysisQuantitative data are expressed as means ± standard deviation. To determine the statistical significance (p < 0.05) of differences among groups, data were analyzed using a one-way analysis of variance (ANOVA) followed by Tukey’s HSD test. All statistical analyses were performed using R software (ver. 4.0.5) and Microsoft Excel (ver. 16.48; Microsoft, Redmond, WA, USA).
To investigate whether MeHg potentiates the toxicity of other heavy metals, we first examined the effects of co-treatment. Among the tested metals, only co-exposure with Cd markedly enhanced cytotoxicity, prompting further detailed experiments. When HeLa cells were treated with 8 µM MeHg in combination with different concentrations of Cd, a significant, dose-dependent decrease in viability was observed (Fig. 1A). In contrast, co-treatment with As or Cr did not significantly alter their cytotoxicity, suggesting that MeHg selectively exacerbates overall cytotoxicity during co-exposure with Cd. After 24 hr of exposure, Cd treatment alone induced modest cell rounding, whereas MeHg alone caused minimal changes. However, co-exposure to Cd and MeHg markedly increased the number of rounded cells (Fig. 1B).
MeHg enhances Cd-induced cell death. (A) HeLa cells were treated with various concentrations of Cd, As, and Cr in the absence (open bars) or presence of 8 µM MeHg (closed bars), respectively. Cell viability was determined 24 hr after treatment using CCK-8. Percent viability was expressed as a percentage relative to 0 µM Cd-, As-, or Cr-treatment. (B) HeLa cells were treated with 20 µM Cd in the absence or presence of 8 µM MeHg for 24 hr. The cells were photographed using a phase-contrast microscope to estimate overall morphology. Scale bars = 100 µm.
At the mRNA level, both Cd and MeHg decreased NRF2 expression at 24 hr, with no significant change at 8 hr (Fig. 2A). This temporal delay suggests that the heavy metal-induced transcriptional repression of NRF2 requires prolonged exposure and may reflect a secondary consequence of sustained cellular toxicity, contrasting with rapid post-translational regulation. Interestingly, despite the reduction in NRF2 mRNA at 24 hr, Cd treatment alone strongly induced the expression of the NRF2 target gene HO-1 (Fig. 2B), indicating activation of the NRF2 pathway at the protein level. In contrast, MeHg co-treatment tended to suppress this Cd-induced HO-1 upregulation.
Cd exposure suppresses NRF2 expression, and co-exposure to MeHg enhances the suppressive effect. (A) HeLa cells were treated with 5, 10, 20, 40 µM Cd or 8 µM MeHg for 8 or 24 hr, and NRF2 mRNA expression was quantified using real-time PCR. (B) Cells were treated with 20 µM Cd in the presence or absence of 8 µM MeHg for 24 hr, NRF2 and HO-1 mRNA expression were quantified using real-time PCR. mRNA levels were normalized to GAPDH and expressed as a relative of control. The values are the mean ± standard deviation of three experiments. (C) Cells were treated with 20 or 40 µM Cd in the presence or absence of 8 µM MeHg for indicated periods, and immunoblotted with anti-NRF2, anti-HO-1, and anti-GAPDH antibodies. (D) Line graphs showing the relative protein levels of NRF2 (left) and HO-1 (right) quantified from the immunoblots shown in (C). Band intensities were normalized to GAPDH. (E) Cells were treated with 5, 10, 20, 40 µM Cd in the presence or absence of 8 µM MeHg for 8 or 24 hr, and immunoblotted with anti-NRF2 and anti-GAPDH antibodies. (F) Line graphs showing the relative protein levels of NRF2 quantified from the immunoblots shown in (E). Band intensities were normalized to GAPDH. The line graphs in (D) and (F) represent the relative band intensities quantified from a single comprehensive experiment (n = 1) using whole-cell lysates.
Western blot analysis using whole-cell lysates revealed that NRF2 protein expression was increased as early as 2 hr after Cd treatment, indicating rapid post-translational activation (Fig. 2C). In contrast, co-treatment with MeHg markedly suppressed this Cd-induced NRF2 accumulation at all examined time points. A dose–response experiment (Fig. 2D) revealed that at 8 hr, Cd alone increased NRF2 protein levels dose-dependently, while a slight reduction was already apparent with MeHg co-treatment. By 24 hr, NRF2 protein levels declined markedly in cells treated with Cd at ≥ 10 µM, and this decrease was further enhanced by MeHg. These observations indicate that MeHg not only suppresses the early accumulation of NRF2 but also accelerates its late-phase degradation.
Strikingly, intracellular Hg accumulation was significantly increased by Cd co-treatment compared to MeHg alone, whereas intracellular Cd levels were not altered by MeHg (Fig. 3, Supplementary Table 1). These data strongly suggest that the enhanced cell death during co-exposure is primarily driven by an increased risk of MeHg toxicity–due to cellular retention–rather than solely an enhancement of Cd toxicity. Interestingly, while As and Cr showed minimal synergistic effects with MeHg in our current experimental conditions, Cd specifically exacerbated the cell death. Both MeHg and Cd are typical “soft” environmental electrophiles that highly target nucleophilic thiol groups, including reactive cysteines on Keap1. Consistent with reports that electrophiles potentiate cytotoxicity through disrupted redox signaling (Akiyama et al., 2020; Abiko et al., 2021; Akiyama et al., 2022), we hypothesize that the simultaneous presence of Cd and MeHg competitively exhausts intracellular thiol pools or critically disrupts Keap1–NRF2 regulation. This dual inhibition impairs antioxidant defenses and Hg efflux, leading to intracellular Hg retention and synergistic cytotoxicity.
Cd increases intracellular Hg concentration. HeLa cells were treated with 8 µM MeHg in the presence or absence of Cd (20 or 40 µM) for the indicated periods. Cells were harvested with RIPA buffer and protein concentrations were measured using a BCA assay kit. Intracellular Hg (top) and Cd (bottom) concentrations were measured using a mercury analyzer (MA3 Solo) and ICP-OES (iCAP7400-Duo), respectively. Values are presented as the mean ± standard deviation (n = 3). *p < 0.05, ***p < 0.001, N.S: Not significant.
We thank K. Sakai for technical assistance.
FundingThis work was supported in part by Grants-in-Aid for Scientific Research C (Grant Number 22K12392).
Conflict of interestThe authors declare that there is no conflict of interest.
Data availabilityThe data in this study are included in the article/supplementary materials. Contact the corresponding author(s) directly to request the underlying data.
Author contributionsY.T. designed the experiments. N.S., N.O., and Y.T. performed the experiments and analyzed the data. Y.T. wrote the manuscript. Y.O., R.N., S.U., and M.K. contributed to the development of the manuscript.
Ethical approval and consent to participateNot applicable.
Patient consent for publicationNot applicable.
