Profiling of Environmental Mixtures Containing Metals for Their Toxicity Pathways and Mechanism of Action

Abstract

Superfund sites are where soil, air, and water are polluted with hazardous materials. Individuals residing and working in these areas are often exposed to metals and other hazardous materials, leading to many adverse health outcomes, including cancer. While individuals are often exposed to multiple chemicals simultaneously, the combined effect of such exposures remains largely unexplored. Here, we investigated the toxicity of metal mixtures in five categories of in vitro assays measuring cytotoxicity, oxidative stress, genotoxicity, cytokine release, and angiogenesis. After testing these mixtures in primary cells and cell lines, we discovered that the nickel/arsenic/cadmium and beryllium/arsenic/cadmium combinations exhibited higher cytotoxicity than their individual compounds, suggesting that the mixtures amplified the cytotoxic effect. To investigate the mechanism underlying their toxicity, we evaluated metal-induced oxidative stress, as oxidative stress is a common factor in most metal-related toxicities. Our results showed that cadmium-induced oxidative stress was increased in mixtures. Some mixtures that induced oxidative stress further increased DNA damage, inhibited DNA synthesis, and activated p53. In addition, some mixtures significantly increased interleukin-8 secretion and angiogenesis more than their component compounds. Our findings offer important insights into metal-related toxicity at Superfund sites.

Introduction

Metals are often used for industrial, agricultural, and medical products. The soil, water, and air around manufacturing facilities, processing plants, landfills, and mining sites are frequently contaminated with metals and metalloids (hereafter referred to as “metals”)¹. In response to public concerns about the health risks posed by metals, the U.S. Congress enacted the Comprehensive Environmental Response, Compensation, and Liability Act, commonly known as Superfund². However, metal exposure is not confined to the United States; similar issues related to heavy metal exposure have been observed in low- and lower-middle-income countries¹. Globally, one of the most pressing issues associated with these exposures is the increased risk of cancer among individuals living near these contaminated sites³. Compounding this issue is the bioaccumulative nature of metals, which makes chronic exposure particularly severe in specific regions and among occupational workers⁴. Major exposure pathways include the gastrointestinal tract, dermal exposure, and inhalation⁵. These exposure routes are closely associated with cancers in the lung, skin, bladder, liver, prostate, ovarian, kidney, and breast⁶.

As documented in the Report on Carcinogens⁷, contaminations of many metals pose serious concerns for human health. In particular, arsenic (As), cadmium (Cd), mercury (Hg), lead (Pb), and chromium (Cr) are considered priority metals due to their high toxicity and potential health risks⁸. These metals exert harmful effects through various mechanisms, such as disrupting cellular organelles, interacting with DNA, and inhibiting enzymes. Notably, their carcinogenicity is strongly linked to oxidative stress, particularly the production of reactive oxygen species (ROS)⁹.

Several studies have shown that each metal exhibits different toxicological effects. For example, cadmium mimics zinc, interfering with DNA binding to the zinc finger proteins¹⁰. Cadmium also disrupts homeostasis and impairs reproduction, which increases the risk of ovarian cancer and breast cancer⁶. Arsenic, on the other hand, can bind to thiol or sulfhydryl groups on proteins, resulting in enzyme dysfunction^{11, 12}. Since enzyme activity is essential for maintaining a healthy physiological balance, exposure to metals is closely linked with a wide range of diseases¹³.

Since multiple metals are often mined and processed at each site, individuals are frequently exposed to several metals simultaneously¹⁴. We investigated the metal mixtures commonly found at the U.S. Superfund sites and compiled a list of the most frequently observed mixtures at the Superfund sites¹⁵. We then refined the list based on the reported carcinogenicity of the compounds excluding volatile and nonsoluble compounds in dimethyl sulfoxide (DMSO). In this study, the mixtures, nickel (Ni)/Cd/As, beryllium (Be)/Cd/As, Ni/Be/As, Ni/Be/Cd, Cd/dimethylvinyl (DMV)/As, and Pb/Cd/As, were selected for testing their toxicity. Most current toxicity tests focus on the effects of the individual compounds. However, as suggested by Anyanwu et al., these metal compounds can exhibit more severe toxicity when humans are exposed to them simultaneously¹⁶. As noted by Martin et al, despite numerous studies on the toxicity of compound mixtures, the carcinogenic effect of these mixtures has not been well studied in mammalian cell models¹⁷. Consequently, there is a significant need to evaluate the carcinogenicity of metal mixtures and single compounds.

In this study, we used a panel of in vitro assays to assess the toxicity of metal compound mixtures frequently observed at Superfund sites. Based on solubility and stability in DMSO, 15 compounds, including 14 metal-containing compounds, were selected, and 24 mixtures were prepared for this study (Supporting Table 1). We first evaluated the cytotoxic effect of both the individual compounds and their mixtures in human primary cells, including lung, bladder, renal, liver epithelial cells, and peripheral blood mononuclear cells (BMC). Next, we conducted several assays to measure the oxidative stress induced by these mixtures. Finally, we examined the impact of these mixtures on several cancer-related biomarkers including p53, DNA damage, cytokine secretion, and angiogenesis. Our study offers valuable insights into the toxicity of metal containing mixtures by comparing their effects to those of the components across multiple aspects related to the hallmarks of cancer development.

Materials and methods

Compounds and their mixtures

2,2-Dimethylvinyl chloride, arsenic oxide (As₂O₃), beryllium sulfate tetrahydrate (BeSO₄ 4H2O), cadmium chloride, nickel(II) chloride (NiCl₂), nickel(II) sulfate hexahydrate (NiSO₄ 6H2O), dimethylarsinic acid, and lead(II) acetate trihydrate were purchased from Sigma-Aldrich (St. Louis, MO). Cadmium nitrate tetrahydrate (Cd(NO₃)₂ 4H₂O) was purchased from Acros Organics (Geel, Belgium). Lead(II) chloride and phenylarsine oxide (PhAsO) were purchased from Alfa Aesar (Haverhill, MA). Arsenobetaine was purchased from Cayman Chemical Co. (Ann Arbor, MI). Cadmium acetate dihydrate (Cd(CH₃COO)₂ 2H₂O) was purchased from ICN Biomedicals, Inc. (Costa Mesa, CA). Nickel dibutyldithiocarbamate was purchased from TCI America (Portland, OR). Nitarsone was purchased from Enamine Ltd. (Kyiv, Ukraine). All single-component compounds were dissolved in 100% DMSO prepared as 20 mM solutions, except arsenic oxide (0.75 mM), nickel(II) chloride (10 mM), and nickel(II) sulfate hexahydrate (10 mM). Two representative mixtures of Ni/Cd/As, Be/Cd/As, Ni/Be/As, Ni/Be/Cd, and Cd/DMV/As and one Pb/Cd/As mixture were tested for solubility. Solutions were prepared in DMSO at 60 mM or the highest concentration achievable below 60 mM, then mixed in equal parts to prepare 3-component mixtures. Mixtures had a stock concentration of 20 mM, except arsenic oxide (0.25 mM) and nickel(II) chloride (10 mM). The solutions were monitored for solubility for several hours through visual observation using bright light passing through the mixtures. While most mixtures remained soluble during the observation period, the Pb/Cd/As mixture developed cloudiness after ~3 h. This cloudiness was attributed to the formation of lead oxide due to the reaction of lead ions with oxygen in the DMSO. To prevent precipitation, 1 N hydrochloric acid and acetic acid were added to the DMSO solutions of lead(II) chloride and lead(II) acetate trihydrate. The final DMSO concentration in the assay wells was maintained at 0.45% for assays conducted at the 5 μL scale. The components in each mixture are detailed in Supporting Table 1. EC₅₀ and IC₅₀ values were calculated based on the highest concentration of the single-component compounds for each mixture to compare sensitivity across different cell types and assay end points. For example, the highest concentration of Mixture 1 was determined to be 92 μM in 5 μL of assay medium after transferring 23 nL of the compound using a Pintool station (Wako Automation, San Diego, CA). The mixture solution underwent a 1:3 serial dilution from the highest concentration using 100% DMSO to generate 11 concentration points ranging from a few nano molars to micro molars. The compound concentrations tested included 0.058 nM to 3.4 μM for arsenic oxide, 0.78 to 45.8 μM for nickel(II) chloride and nickel(II) sulfate hexahydrate, and 1.55 to 92 μM for the other compounds.

Cells and cell culture

Primary bronchial/tracheal epithelial cells (ATCC, Madison, WI, Cat No. PCS-300–010), primary bladder epithelial cells (ATCC, Cat No. PCS-420–010), primary renal mixed epithelial cells (ATCC, Cat No. PCS-400–012), primary liver epithelial cells (THLE-2, ATCC, Cat No. CRL-2706), primary peripheral blood mononuclear cells (ATCC, Cat No. PCS-800–011), HepG2 cell, HCT116 cells, and THP-1 cells and Angio-Ready Angiogenesis Assay Kit were obtained from ATCC. The p53RE-bla HCT-116 cell lines were obtained from Thermo Fisher Scientific (Waltham, MA).

Primary cells were cultured using the medium provided by ATCC. HCT116 wild-type cells were cultured in McCoy’s 5A medium supplemented with 10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin. p53RE-bla HCT-116 cells were cultured in McCoy’s 5A medium supplemented with 10% dialyzed FBS, 100 U/mL penicillin, 100 μg/mL streptomycin, and 5 μg/mL blasticidin. HepG2 cells were cultured in EMEM supplemented with 10% FBS, 100 U/mL penicillin, and 100 μg/mL streptomycin. THP-1 cells were cultured in RPMI-1640 supplemented with 10% FBS, 100 U/mL penicillin, 100 μg/mL streptomycin, and 50 μM 2-mercaptoethanol. The hTERT-immortalized mesenchymal stem cells and aortic ECs were cultured using the medium provided in the kit, supplemented with 25 U/mL penicillin and 25 μg/mL streptomycin.

Cytotoxicity assay

Cells were plated into a white solid-bottom 1536-well plate at a density of 2000 (kidney, liver, and bladder cells) or 2500 (lung cells) cells/well in 5 μL of culturing medium. After dispensing the cells, the assay plates were incubated at 37 °C for 5 h to allow cell attachment. Metal compounds, positive control (tetraoctylammonium bromide), and DMSO (23 nL each) were transferred to the wells by using a Pintool station (Wako Automation, San Diego, CA). The cells were then incubated at 37 °C for 72 h, except for blood mononuclear cells that were treated for 5 h. Following this, 5 μL of CellTiter-Glo (Promega) was added to each well, and the plates were incubated at RT for 30 min. The luminescence signal was read by using a ViewLux plate reader (PerkinElmer, Waltham, MA).

Caspase 3/7 assay

Cells were suspended in a culture medium and dispensed at a density of 1500 cells/5 μL/well in 1536-well white solid-bottom plates using a Multidrop Combi (Thermo Fisher Scientific) dispenser. The assay plates were incubated at 37 °C/5% CO2 for 5 h to allow for cell attachment. Then, 23 nL of metal compounds, positive control (staurosporine), and DMSO were transferred to the assay plates using a Pintool station. The plates were then further incubated at 37 °C/5% CO2 for 18 h, 5 μL/well of Caspase-Glo 3/7 reagent (Promega, Madison, WI) was added using a Flying Reagent Dispenser (Aurora Discovery, San Diego, CA). After an additional incubation period of 30 min at RT, luminescence intensity was measured with a ViewLux plate reader.

Mitochondrial membrane potential (MMP) assay

The cells were plated into a black-wall/clear-bottom 1536-well plate at a density of 2000 (kidney, liver, and bladder cells) or 2500 (lung cells)/well in 5 μL of culture medium. After dispensing cells to the 1536-well plates, the assay plates were incubated at 37 °C for 24 h to allow cell attachment. Next, 23 nL of metal compounds, positive control (p-trifluoromethoxyphenylhydrazone), and DMSO were transferred using a Pintool station. The cells were then incubated at 37 °C for 5 h. Following this, 5 μL of MPI reagent (Codex BioSolutions, Gaithersburg, MD) was added to each well, and the plates were incubated at 37 °C for 30 min. The fluorescence signal was subsequently read by using an Envision plate reader (PerkinElmer).

Reactive oxygen species (ROS) assay

HepG2 cells were plated into a white solid-bottom 1536-well plate at a density of 1000 cells/well in 5 μL of culture medium. After dispensing the cells, the assay plates were incubated at 37 °C for 5 h to allow cell attachment. Then, 23 nL of metal compounds and positive control (menadione) were transferred by using a Pintool station. The cells were then incubated at 37 °C for 16 h. Following this, 1 μL of H2O2 substrate was added to each well, and the plates were incubated for 6 h at 37 °C. Subsequently, 4 μL of ROS-Glo detection reagent (Promega) was added to each well. Immediately after the detection reagent was added, the luminescence signal was read using a ViewLux plate reader.

Glutathione (GSH) assay

HepG2 cells were plated into a white solid-bottom 1536-well plate at a density of 1000 cells/well in 5 μL of culture medium and incubated overnight at 37 °C. Next, 23 nL of metal compounds and positive control (tetraoctylammonium bromide) were transferred using a Pintool station. Following an incubation period of 1 h at 37 °C, the medium was discarded using a Blue Washer (Blue Cat Bio, Lebanon, NH), and the plates were washed with PBS. 4 μL of GSH reaction mixture (Promega) was then added to each well and incubated for 15 min at RT. Next, 4 μL of GSH-Glo detection reagent was added to each well, and the plates were incubated for 30 min at RT. The luminescence signal was read using a ViewLux plate reader.

p53-reporter gene assay

p53RE-bla HCT116 cells were resuspended in assay medium (Opti-MEM supplemented with 0.5% dialyzed FBS) and dispensed into 1536-well black-wall/clear-bottom plates at 4000 cells/5 μL/well. After incubation at 37 °C for 5 h, 23 nL of metal containing compounds and positive control (mitomycin C) were transferred using a Pintool station. The assay plates were then incubated for an additional 16 h. On the next day, 1 μL of LiveBLAzer (Life Technologies, Madison, WI) detection mixture was added to each well, and the plates were incubated at room temperature in the dark for 2 h. Next, an Envision reader measured the fluorescence intensity at emission at 460 and 530 nM and excitation at 405 nM excitation. The data was expressed as the ratio of the emission wavelengths (460/530). Following this, 3 μL of CellTiter-Glo reagent was added to each well, and plates were incubated in the dark at RT for 30 min. Luminescence readings were then obtained by using a ViewLux plate reader.

DNA synthesis assay

Cells were plated into a black-wall/clear-bottom 1536-well plate at a density of 1000 cells/well in 5 μL of culture medium. Following the plating, the assay plates were incubated at 37 °C for 5 h to allow cell attachment. Metal compounds and positive control (aphidicolin) were transferred in 23 nL volumes by using a Pintool station. The plates were then incubated at 37 °C for 24 h. Next, 1 μL of EdU reagent (Thermo Fisher) was added to each well, and the assay plates were then incubated for 1 h at 37 °C. The medium was removed using the blue washer, and 3 μL of fixation solution was added to each well, followed by a 30 min incubation period at RT. The wells were washed with wash buffer, before adding 2 μL of the 1× Click-iT EdU reaction cocktail to each well. The plates were then incubated at RT for 30 min. After removal of the Click-iT EdU reaction cocktail, 3 μL of 1.5% BSA Blocking Solution was added to each well and incubated for 5 min at RT, protected from light. The plates were then washed 3 times with 4 μL/well of wash buffer. Subsequently, 3 μL of the Amplex UltraRed reaction mixture was added to each well and incubated for 15 min at RT, protected from light. The reaction was stopped by adding 1 μL of 3 times diluted Amplex UltraRed stop solution to each well. Finally, the fluorescence signal was measured using a PHERASTR plate reader (BMG Labtech, Ortenberg, Germany)

γH2AX immunostaining assay

HCT116 cells were plated at a density of 1000 cells per well in 5 μL of culture medium into 1536-well clear-bottom/black-wall plates (Aurora Biosciences, San Diego, CA). The plates were incubated overnight to allow for cell attachment. The cells were treated with 23 nL of metal compounds, positive control (etoposide), or DMSO. After a 24 h compound treatment, the cells were fixed with 4% paraformaldehyde for 20 min and then were incubated with a permeabilization/blocking/nuclear-staining solution (PBS containing 5% BSA, 0.5% Tween-20, 1 μg/mL Hoechst). Next, the cells were treated with the anti-γH2AX antibody for 1 h and washed with PBS twice. Then, 2 μL of anti-mouse Alexa488 antibody was added to the plates and then the plates were incubated for 1 h at RT. After the incubation, the plate was washed 3 times with PBS. The assay plates’ fluorescence intensities (490 nm excitation and 550 nm emission) were measured on the Operetta CLS High-Content Analysis System (PerkinElmer) with a 20× confocal objective. Images from each well of the assay were obtained and analyzed using Harmony High-Content Imaging and Analysis (PerkinElmer). The number of foci was counted in each well and was normalized by a cell number to obtain the number of foci per cell.

Cytokine secretion assays

THP-1 cells were plated in a 1536-well white solid-bottom plate at a density of 2000 cells/well in 5 μL of culture medium. The cells were treated with 23 nL of metal compounds, positive control (lipopolysaccharides), and DMSO by using the Pintool station. After 24 h, 1 μL of IL-1, IL-8, or TNFα antibody mix (Cisbio, Saclay, France) was added to each well. The plates were incubated for 24 h at RT. The fluorescence signal was read using the PHERASTR plate reader.

Angiogenesis assay

The GFP-expressing tubular structure was used to assess the angiogenesis formation. Immortalized mesenchymal stem cells (hTERT-MSCs) and aortic ECs (TeloHAECs) were plated at a density of 5000 cells/7.5 μL/well into 1536-well black-wall/clear-bottom assay plates. The assay plates were incubated at 37 °C for 5 h to allow for cell attachment. Compounds were then added at a volume of 23 nL per well by using a Wako Pintool station. After a 48 h incubation at 37 °C, the plates were analyzed on the Operetta CLS High-Content Analysis System. Images from each well were acquired and analyzed using Harmony High-Content Imaging and Analysis Software.

Data analysis

Analysis of concentration–response data was performed as previously described^{18, 19}. Briefly, raw plate readings for each titration point were normalized against the positive control compound (100%) and DMSO-only wells (0%) using the following formula: % Activity = (V c_ompound – V _DMSO)/(V _pos-V _DMSO) × 100, where V _compound indicates the compound well values, V _pos indicates the median value of the positive control wells, and V _DMSO indicates the median values of the DMSO-only wells. The half-maximal inhibitory or effective concentration values (IC₅₀/EC₅₀) and efficacy values for each compound were determined by fitting the concentration–response curves of each compound to a four-parameter Hill equation²⁰. To identify the enhanced effect in mixtures, the Toxicity Unit (TU) approach was applied²¹. The TU_i was calculated as: TU_i = C_i/IC_50i, where C_i indicates the concentration of i^th compound in the mixture at its IC₅₀, and IC_50i represents the IC₅₀ of the i^th compound when tested individually. If ΣTU_i is below 0.8, the combined effect is defined as synergistic effects. Mixture 1–4 demonstrated ΣTU_i values below 0.8 in all the assays presented in the figures and Supporting Figures.

Results

Effect of mixtures on cell viability and caspase 3/7 activation

Human normal primary cells were treated with mixtures and their individual components to assess their cytotoxicity. Mixture 1 (NiSO₄/As₂O₃/Cd(NO₃)₂) and Mixture 2 (BeSO₄/As₂O₃/Cd(CH3COO)₂) exhibited higher potencies in viability assays in BMC, while Mixture 3 (BeSO₄/Cd(CH₃COO)/PhAsO) showed higher potencies in liver, renal, bladder, and lung cells compared to the individual component compounds (Figure 1 a, b, Supporting Figure 1a–d). Induction of caspase 3/7 activity, an indicator of apoptosis, was observed exclusively in blood cells against Mixtures 1, 2, and 4 (NiCl₂/As₂O₃/Cd(CH3COO)₂) (Figures 1c,d, and S1e). The other tested mixtures listed in Supporting Table 1 did not show an enhanced effect in these assays (Supporting Table 2).

Figure 1.

Cytotoxicity and activation of caspase 3/7 by metal mixtures

Cell viability was measured after 72 h treatment of Mixture 1 (a) and Mixture 2 (b) in blood mononuclear cells. Caspase 3/7 activity in blood mononuclear cells was measured after 18 h treatment with Mixture 1 (a) and Mixture 2 (b). Each mixture contains three compounds indicated in each plot. The error bars indicate the standard deviation from three independent experiments. NiSO₄·6H₂O: Nickel (II) sulfate hexahydrate, As₂O₃: Arsenic oxide, Cd(NO₃)₂·4H2O: Cadmium nitrate tetrahydrate, BeSO₄·4H₂O: Beryllium sulfate tetrahydrate, Cd(CH₃COO)₂·2H₂O: Cadmium acetate dihydrate

Effects of mixtures on cellular stress

Given that carcinogenic effects of metal compounds are often associated with oxidative stress, various assays related to oxidative stress, such as ROS and MMP assays, were conducted. The levels of ROS in HepG2 cells were induced by 3 cadmium compounds, nitarsone, and lead(II) chloride, with EC₅₀ values ranging from 2.65 to 6.84 μM (Supporting Table 3). However, no enhanced effects were observed in any mixtures. While phenylarsine oxide independently disrupted MMP, this effect was amplified in Mixture 3 in lung cells (Supporting Figure 2). These findings suggest that Mixture 3 is more toxic than phenylarsine oxide alone, consistent with the results of the caspase 3/7 assay results. Upon exposure to ROS, cellular GSH acts to counter oxidative stress, leading to a reduction in GSH levels²². Mixtures 1 and 2 caused a greater reduction in GSH compared to their component compounds in HepG2 cells, indicating that Mixtures 1 and 2 induced more oxidative stress (Figure 2a,b). In contrast, phenylarsine oxide-induced oxidative stress was not enhanced in Mixture 3 (Supporting Figure 3a) despite Mixture 3 exhibiting a higher cytotoxicity than phenylarsine oxide alone (Supporting Figure 1).

Figure 2.

Reduction in GSH and electrophilicity of metal mixtures

Cellular GSH level, one of the key defense mechanisms against oxidative stress, was measured in HepG2 cells after 1 h treatment with Mixture 1 (a) and Mixture 2 (b). Electrophilicity of the metals and their mixtures was measured using a MSTI assay against Mixture 1 (a) and Mixture 2 (b). Each mixture contains three compounds indicated in each plot. The error bars indicate the standard deviation from three independent experiments.

NiSO₄·6H₂O: Nickel (II) sulfate hexahydrate, As₂O₃: Arsenic oxide, Cd(NO₃)₂·4H2O: Cadmium nitrate tetrahydrate, BeSO₄·4H₂O: Beryllium sulfate tetrahydrate, Cd(CH₃COO)₂·2H₂O: Cadmium acetate dihydrate

Given that electrophilicity may also contribute to the toxicity of metal compounds, an MSTI assay was conducted to test the electrophilicity of the compounds and mixtures. Mixtures 1, 2, and 3 demonstrated higher electrophilicity and redox reactivity in the MSTI assay compared to their individual components (Figures 2c,d and S3b). However, the other tested mixtures did not show an enhanced effect in these assays (Supporting Table 3).

Effect of mixtures on genotoxicity

We next evaluated the genotoxicity of these compounds and mixtures by measuring p53 activation, DNA synthesis disruption, and DNA damage induction. p53 activation was measured using HCT116 cells integrated with a p53 reporter gene²³. The results showed that beryllium sulfate tetrahydrate and phenylarsine oxide activated p53, with this effect further enhanced by cadmium nitrate dihydrate (Supporting Figure 4a).

To assess the impact on DNA synthesis, an EdU incorporation assay was performed in liver, lung, and renal cells. Mixtures 1 and 2 had greater inhibitory effect on DNA synthesis across the cell types compared to their component compounds (Figures 3a,b and S3b–e). Additionally, DNA damage induction was evaluated using an immunostaining assay measuring γH2AX foci²⁴, a DNA damage biomarker (Supporting Figure 4f). In HCT116 cells, Mixtures 1 and 2 significantly increased γH2AX foci, despite the weak γH2AX-inducing capability of cadmium compounds and the absence of induction by the other two compounds (Figure 3c,d and S4g,h). Other tested mixtures did not exhibit an enhanced effect in these assays (Supporting Table 4).

Figure 3.

Effect of metals and their mixtures on genotoxicity

Activity of DNA synthesis in liver endothelial cells was measured using an EdU incorporation assay after 24 h treatment with Mixture 1 (a) and Mixture 2 (b). Induction of DNA damage was measured after the 24 h treatment with Mixture 1 (c) and Mixture 2 (d). Each mixture contains three compounds indicated in each plot. The error bars indicate the standard deviation from three independent experiments. NiSO₄·6H₂O: Nickel (II) sulfate hexahydrate, As₂O₃: Arsenic oxide, Cd(NO₃)₂·4H2O: Cadmium nitrate tetrahydrate, BeSO₄·4H₂O: Beryllium sulfate tetrahydrate, Cd(CH₃COO)₂·2H₂O: Cadmium acetate dihydrate

Effects of metal compounds on cytokine release and angiogenesis

The levels of IL-1, IL-8, and TNFα were measured in the THP-1 cells. Mixtures 1 and 2 were found to increase IL-8 secretion, while no significant changes were observed for IL-1 and TNFα (Figure 4, Supporting Table 5). Although other mixtures and single compounds also induced IL-8 secretion, no enhanced effects were detected in these cases.

Figure 4.

Effect of heavy metals on IL-8 secretion

The IL-8 secretion was measured using HTRF IL-8 assay in THP-1 cells after 24 h treatment with Mixture 1 (a) and 2 (b). Each mixture contains three compounds indicated in each plot. The error bars indicate the standard deviation from three independent experiments. NiSO₄·6H₂O: Nickel (II) sulfate hexahydrate, As₂O₃: Arsenic oxide, Cd(NO₃)₂·4H2O: Cadmium nitrate tetrahydrate, BeSO₄·4H₂O: Beryllium sulfate tetrahydrate, Cd(CH₃COO)₂·2H₂O: Cadmium acetate dihydrate

Tumor growth often relies on angiogenesis, the formation of new blood vessels, to overcome nutrient deprivation and hypoxic conditions. To study the impact of these mixtures on angiogenesis, a co-culture system of GFP-labeled aortic epithelial cells and immortalized mesenchymal stem cells was used²⁵. Representative images in Figure 5a show the concentration-dependent angiogenesis induction by Mixture 4. While several single metal compounds, such as cadmium acetate dihydrate, cadmium chloride, and cadmium nitrate tetrahydrate, independently induce angiogenesis (Supporting Table 5), Mixture 4 exhibited a stronger angiogenesis-inducing capability than its component compounds (Figure 5b). No enhanced effects were observed for the other tested mixtures in these assays (Supporting Table 5).

Figure 5.

Effect of heavy metals on angiogenesis

The induction of angiogenesis was measured using GFP-labeled endothelial cells and hTERT-immortalized mesenchymal stem cells. After cells were treated with metal compounds for 48 h, angiogenesis from each assay well was measured in high-content imaging analysis using an Operetta plate reader. The angiogenesis induction was quantified by the area of the GFP-labeled aortic epithelial cells. The representative images indicate concentrations of Mixture 4 (a) and induction of angiogenesis by Mixture 4 (b). Mixture 4 contains three compounds indicated in the plot. The error bars indicate the standard deviation from three independent experiments. NiCl₂: Nickel (II) chloride, As₂O₃: Arsenic oxide, Cd(CH₃COO)₂·2H₂O: Cadmium acetate dihydrate

Discussion

In this study, we evaluated the toxicity of several environmental chemical mixtures that contain metals commonly found at Superfund sites. We tested 24 mixtures in 5 assay categories, including cytotoxicity, oxidative stress, genotoxicity, cytokine release, and angiogenesis formation. Mixture 1 (NiSO₄/As₂O₃/Cd(NO₃)₂) and Mixture 2 (BeSO₄/As₂O₃/Cd(CH₃COO)₂) showed higher toxicity in cytotoxicity, oxidative stress, genotoxicity, and cytokine release assay. Mixture 3 (BeSO₄/Cd(CH₃COO)₂/PhAsO) showed enhanced activity in cytotoxicity, oxidative stress and genotoxicity assay. Mixture 4 (NiCl₂/As₂O₃/Cd(CH3COO)₂) showed only an enhanced effect on angiogenesis assay.

Several compounds, such as phenylarsine oxide, cadmium acetate dihydrate, cadmium chloride, cadmium nitrate tetrahydrate, lead(II) acetate trihydrate, and lead(II) chloride, were cytotoxic by themselves (Supporting Table 2). These six compounds were identified as active compounds in either the ROS or GSH assays. Their cytotoxicity was further enhanced when mixed with other compounds. For example, Mixture 1, comprising nickel(II) sulfate hexahydrate, cadmium nitrate tetrahydrate, and arsenic oxide, showed significant toxicity. Among these components, only cadmium nitrate tetrahydrate exhibited oxidative stress-inducing capability (Supporting Table 3). Mixtures 9, 12, 16, 17, and 23 shared two of the three component compounds with Mixture 1, but did not display enhanced effects in any assays compared to their component compounds. These findings indicated that the specific combination of compounds in Mixture 1 is crucial for amplifying the toxicity.

Four mixtures showed enhanced activities compared with their component compounds. As previously mentioned, potentiation was observed only in specific combinations. While ROS induction was not detected with arsenic oxide, nickel sulfate, or beryllium sulfonate treatment in our assay condition, oxidative stress induced by cadmium compounds was amplified when co-treated with arsenic oxide, nickel sulfate, and beryllium sulfonate (Figure 2). Arsenic is known to inhibit the antioxidant pathways and DNA repair proteins, which can amplify the genotoxic effect of DNA-damaging reagents²⁶. Similarly, the presence of arsenic oxide likely impaired the antioxidant defense against cadmium-induced oxidative stress, leading to increased DNA damage. However, this does not fully explain the potentiation mechanism, as several mixtures containing Ni/As/Cd or Be/As/Cd did not show enhanced effects..

The key difference between mixtures that exhibited potentiation and those that did not, despite containing the same metals, was the salt form. When salt is introduced into the assay medium, part of it dissociates into free metal cations and acid anions. Many mixtures that did not show potentiation were composed of chloride salts. Chloride ions may reduce the bioreactivity and bioavailability of metal compounds and influence the expression of metallothioneins (MTs), which play an important role in metal homeostasis and protection against metal toxicity^{27, 28}. In our assay conditions, chloride ions likely reduce the toxic effect of mixtures containing Ni/As/Cd or Be/As/Cd by limiting metal uptake, thereby masking the potentiation effects.

While only BMC showed caspase activation, epithelial cells showed a similar trend in the other assays, suggesting a difference in response to metal toxicity between epithelial cells and BMC (Supporting Table 6). The IC₅₀ values for GSH reduction of Mixtures 1 and 2 were 0.58 and 0.84 μM, respectively, which are close to the IC₅₀ values observed in the DNA synthesis assay (ranging from 0.23 to 0.34 μM). Given that the GSH assay was conducted for 1 h and the DNA synthesis assay for 24 h, it appears that ROS were induced first, followed by the inhibition of DNA synthesis. The EC₅₀ values for γH2AX induction were approximately 1 μM with a 24 h treatment and 2.5 μM with a 4 h treatment, slightly higher than those observed in the DNA synthesis assay. This suggests that the collapse of DNA replication caused by oxidative stress further induced a detectable amount of DNA damage.

Cell viability results indicated that BMCs were the most sensitive to Mixtures 1 and 2 among the tested primary cells. However, epithelial cells, particularly renal cells, were more sensitive to Mixture 3 than BMCs. Since our data suggests that metal-induced cytotoxicity is partially driven by DNA synthesis inhibition, cells with shorter doubling times are expected to be more sensitive to DNA damage²⁹. Therefore, the sensitivity differences among the epithelial cells may be attributed to variations in their doubling times.

In addition to the genotoxic potential, we found that cadmium compounds, nitarsone, and lead(II) chloride induced angiogenesis (Supporting Table 5). Consistent with the previous studies linking oxidative stress to angiogenesis³⁰, all mixtures and compounds that increased ROS in this study also induced angiogenesis. Cadmium, for instance, promotes angiogenesis by upregulating the vascular endothelial growth factor³¹. Since cytokine secretion can trigger tumor growth by activating proliferation signaling, we also tested these mixtures for their effects on IL-1, IL-8, and TNFα secretion^{32, 33}. IL-1 and TNFα are key mediators of the innate immune response, the first line of defense against pathogens³⁴, while IL-8 promotes monocyte-macrophage growth and differentiation, as well as endothelial cell survival, proliferation, and angiogenesis³⁵.

While none of the tested compounds induced IL-1 or TNFα, several mixtures and compounds did increase IL-8 secretion (Supporting Table 5). Mixtures 1 and 2, in particular, induced higher levels of IL-8 secretion compared to their individual compounds. Given that these mixtures also induce high oxidative stress (Figure 2), it is likely that the oxidative stress triggered the inflammatory response, resulting in higher secretion of IL-8³⁶.

Most current toxicity testing focuses on evaluating the toxicity of single compounds³⁷. However, this study demonstrates that some compounds can have more toxic effects when co-treated with other substances. Given the large number of compounds produced and released into the environment, testing all possible combinations would create an overwhelming amount of conditions³⁸. In this study, we prepared mixtures that simulate real-life exposure scenarios around Superfund sites². This approach is efficient for selecting relevant mixture components. For instance, combining pesticides and metals is of particular interest, as one of the main exposure pathways for metals is through consuming crops harvested near Superfund sites³⁹.

The concentration ranges tested in this study were 0.058 nM to 3.4 μM for arsenic oxide, 0.78 to 45.8 μM for nickel(II) chloride and nickel(II) sulfate hexahydrate, and 1.55 to 91.6 μM for other compounds. According to National Health and Nutrition Examination Survey conducted by the US Center for Disease Control, the average human serum concentration of metals in the general US population was 7.53 μg/L (36.3 nM) for lead, 0.241 μg/L for cadmium (2.15 nM), and 6.32 μg/L (84 nM) for arsenic⁴⁰. Blood levels in humans living near Superfund sites have been reported as 0.01–4.0 μg/L (1.3–53 nM) for arsenic, 0.01–3.5 μg/L (89 pM – 31 nM) for cadmium, and 0–33 μg/dL (0–1.50 μM) for lead⁴¹. The urine levels in occupational workers exposed to these metals ranged from 3.08 to 22.61 μg/L (27–200 nM) for cadmium, 20.26 to 89.6 μg/L (98–432 nM) for lead, and 4.45 to 79.5 μg/L (75 nM – 1.4 μM) for nickel⁴².

Most mixtures or compounds in this study had EC₅₀/IC₅₀ values around a few micromolar. Mixtures 1 and 2, phenylarsine oxide, and phenylarsine oxide-containing mixtures exhibited EC₅₀/IC₅₀ values below 1 μM in several assays (Supporting Tables 2–5). Considering that metals accumulate in the body, these metals and mixtures, along with other industrial compounds, may have toxic effects at real-life exposure, as several epidemiological studies have shown increased cancer incidences around Superfund sites³. Although our study found that several mixtures exhibited toxicity higher than that of their individual compounds, real-life exposure involves simultaneous exposure to multiple compounds. Moreover, our study indicates that cadmium-induced oxidative stress is enhanced in the presence of arsenic compounds, but the exact mechanisms remain unclear. Future studies utilizing proteomic and transcriptomic analyses could help identify the genes and proteins involved when cells are treated with mixtures compared to single compounds.

In this study, we measured several biomarkers that are considered hallmarks of carcinogenicity⁴³. We initially used primary cells to test whether the toxicity of metal compounds could be potentiated when co-treated with other compounds. However, some primary cells were not suitable for certain assays. For example, primary bladder cells could not be used for DNA synthesis assays because DNA replication was not very active in these cells, leading to weak signals (data not shown). Therefore, we turned to cancer cells to study the toxicity mechanisms. This approach, using a panel of in vitro assays, enabled us to identify potential carcinogens and investigate their mechanisms of toxicity. Since the mechanism of action varies among different compounds, testing the cytotoxicity of compounds could serve as a valuable first step in understanding their toxic effects. Follow-up studies, such as DNA damage quantification and IL-8 secretion assays, could further elucidate the mechanisms involved.

In summary, we evaluated the toxicity of mixtures containing metals commonly found in Superfund sites. We identified the two most toxic mixtures as: (1) NiSO₄/As₂O₃/Cd(NO₃)₂ and (2) BeSO₄/As₂O₃/Cd(CH3COO)₂. Although there are several toxicological studies on compound mixtures, their carcinogenic effects have not been studied well. We identified compound mixtures that show higher carcinogenicity through a series of biological assays using human cells. The toxicity of cadmium compounds was enhanced when co-treated with NiSO₄/As₂O₃or BeSO₄/As₂O₃. This enhanced toxicity was primarily related to oxidative stress, which induced DNA damage in the cells. Our findings suggest that NiSO₄/As₂O₃ or BeSO₄/As₂O₃ inhibits the cellular antioxidant pathway, thereby amplifying cadmium-induced oxidative damage. Given that soil, water, and air are contaminated by various metals, people living in these regions or occupational workers are often exposed to multiple metal compounds simultaneously. While many toxicological studies focus on the effects of single compounds, this study demonstrates that compound mixtures can exhibit greater toxicity. Therefore, the combined effects of the mixture must be considered when assessing the risks faced by regional populations or occupational workers.

Supplementary Material

Results for cytotoxicity related assays (Figure S1), Result for MMP assay (Figure S2), Results for oxidative stress related assay (Figure S3), Results for DNA damage related assays (Figure S4), List of metal mixtures used in this study (Table S1), EC₅₀/IC₅₀ values and efficacy for cytotoxicity related assays (Table S2), EC₅₀/IC₅₀ values and efficacy for oxidative stress related assays (Table S3), EC₅₀/IC₅₀ values and efficacy for DNA damage related assays (Table S4), EC₅₀/IC₅₀ values and efficacy for angiogenesis and cytokine related assays (Table S5), EC₅₀/IC₅₀ values and efficacy of representative mixtures (Table S6).

Highlights.

• Several metal mixtures showed higher cytotoxicity than the single metals alone

• Oxidative stress was enhanced by metal mixtures

• DNA damage was increased in metal mixtures

• Metal mixtures induced angiogenesis

Synopsis.

People are often exposed to several metal compounds simultaneously. We found that several metal compound mixtures have higher toxicity than their single component compounds