Immunotoxic effects of sodium tungstate dihydrate on female B₆C₃F₁/N mice when administered in drinking water

Abstract

Tungsten is a naturally occurring, high-tensile strength element that has been used in a number of consumer products. Tungsten has been detected in soil, waterways, groundwater, and human tissue and body fluids. Elevated levels of tungsten in urine were reported for populations exposed to tungstate in drinking water in areas where natural tungsten formations were prevalent. Published reports indicated that sodium tungstate may modulate hematopoiesis, immune cell populations, and immune responses in rodent models. The objective of this study was to assess potential immunotoxicity of sodium tungstate dihydrate (STD), a drinking water contaminant. Female B₆C₃F₁/N mice received 0–2000 mg STD/L in their drinking water for 28 d, and were evaluated for effects on immune cell populations in spleen and bone marrow, and humoral-mediated, cell-mediated, and innate immunity. Three different parameters of cell-mediated immunity were similarly affected at 1000 mg STD/L. T-cell proliferative responses against allogeneic leukocytes and anti-CD3 were decreased 32%, and 21%, respectively. Cytotoxic T-lymphocyte activity was decreased at all effector:target cell ratios examined. At 2000 mg STD/L, the absolute numbers of CD3⁺ T-cell progenitor cells in bone marrow were increased 86%, but the alterations in B-lymphocyte and other progenitor cells were not significant. There were no effects on bone marrow DNA synthesis or colony forming capabilities. STD-induced effects on humoral-mediated immunity, innate immunity, and splenocyte sub-populations were limited. Enhanced histopathology did not detect treatment-related lesions in any of the immune tissues. These data suggest exposure to STD in drinking water may adversely affect cell-mediated immunity.

Keywords:

• Cell-mediated immunity
• cytotoxic T-lymphocyte
• sodium tungstate dihydrate

Introduction

Exposure to tungsten is of widespread concern since it is a naturally occurring element in more than 20 minerals found in the Earth’s crust. In pure form, tungsten has the highest melting point and tensile strength of all metals, resists corrosion, and conducts electricity (Hammond 1987). Occupationally, it is used in welding, metal cutting, drilling, aerospace applications, and the manufacture of many consumer products. Tungsten alloys have been used in some types of ammunition as replacements for lead and depleted uranium.

Environmentally, geothermal activity, mining, manufacturing processes, and spent munitions all contribute to the deposition of soluble and insoluble forms of tungsten into soil, waterways, and groundwater. In both terrestrial and aquatic environments, tungsten oxidizes to the anion form and accumulates as monomeric (typically alkaline conditions) or polymeric (typically acidic conditions) species (Petkewich 2009; USEPA 2014). Environmental studies demonstrated trophic transfer of tungsten between plants and invertebrates (Kennedy et al. 2012), and bioaccumulation of tungsten in plants, animals, and humans (Koutsospyros et al. 2006; Rubin et al. 2007; Kennedy et al. 2012). Soluble tungsten has been detected in groundwater in areas that are near natural deposits of the mineral. The Agency for Toxic Substances and Disease Registry (ATSDR) reported that water-soluble tungsten compounds in the environment may pose a greater threat to human health than water-insoluble forms (ATSDR 2005). The Center for Disease Control (CDC) detected increased levels of tungsten in the drinking water and urine of residents in Churchill County, Nevada, where a high incidence of childhood acute lymphoblastic leukemia was also identified (Seiler et al. 2005). Although no causal relationship was established between tungsten levels in biological samples and disease (Rubin et al. 2007), this area is a prototypical example of populations exposed to tungstate in surface water, groundwater, and well water (ATSDR 2005). The U.S. Geological Survey (USGS) determined that the tungsten compounds present in groundwater were derived from natural sources, including upwelling of geothermal waters, erosion of tungsten-bearing mineral deposits, and reductive dissolution of metal oxides (Seiler et al. 2005). In alkaline waters, such as that found in the Carson Desert area, tungsten desorbs from metal oxides and other mineral surfaces to form soluble tungstate monomers (WO₄²⁻) (Johannesson et al. 2013).

Although evidence is limited, rodent studies suggest that tungsten exposure may affect the immune system, as indicated by localization of tungsten in bone and spleen (McDonald et al. 2007; Guandalini et al. 2011; Kelly et al. 2013); attenuation of the delayed-type hypersensitivity response; alteration of T-cell populations (Osterburg et al. 2014); transcriptomic changes associated with regulation of some immune responses and hematological/immunological disease (Fastje et al. 2008, 2009); and, induction of splenomegaly and neutrophilia following co-exposure to sodium tungstate and respiratory syncytial virus (RSV) (Fastje et al. 2012). Published reports indicated that sodium tungstate dysregulated cytokine production, the cell cycle, and apoptosis in human peripheral blood lymphocytes (Osterburg et al. 2010). However, there does not appear to be enough information in the literature to set a federal drinking water standard for tungsten (USEPA 2014). In addition, adequate information regarding the health effects of tungsten is not available (ATSDR 2005).

The objective of this study was to comprehensively evaluate potential adverse effects of sodium tungstate dihydrate (STD) in drinking water to the immune system in female B₆C₃F₁/N mice. Due to the reported deposition of tungsten in bone tissue, an evaluation of bone marrow cell populations and function was included in the standard testing battery for cell-mediated, humoral-mediated, and innate immunity (Luster et al. 1988). Drinking water was selected as the most likely route of exposure for the general population.

Materials and methods

Test substance

Sodium tungstate dihydrate (STD, Na₂WO₄·2H₂O, CAS #10213-10-2, Lot #12330JO) was obtained through an NTP analytical chemistry contract at Battelle (Columbus, OH). Stability data provided by Battelle indicated that formulations of STD in tap water were stable for at least 42 d both at 5 °C and at room temperature. Stock drinking water solutions were freshly prepared every 2 weeks in tap water and were stored at 4 °C in sealed Nalgene high-density polyethylene bottles with polypropylene caps. Samples of dose formulations were evaluated by scanning electron microscopy (SEM) with energy dispersive spectrometry (EDS) detection, X-ray diffraction (XRD) and ion chromatography to ensure that animal room samples were within 10% of the targeted concentration.

Animals and animal exposure

Female pathogen-free B₆C₃F₁/N mice were obtained from Taconic Farms Inc. (Germantown, NY) at 4–8 weeks of age and maintained on a 12-h light/dark cycle at 18–26 °C in an Association for Assessment and Accreditation of Laboratory Animal Care-accredited facility. All experiments were conducted at Virginia Commonwealth University under an approved Animal Care and Use Committee protocol. The animals received NTP2000 diet (Ziegler Brothers, Inc., Gardners, PA) and tap water or tap water containing the test substance ad libitum. At 8–9 weeks of age (17–26 g), the mice were weighed, randomized via an Apple computer-generated randomization procedure, identified by tattoo, and placed on treatment (N = 8/group). Due to the requirements for the evaluation of immunological and toxicological endpoints, 10 separate cohorts of mice were required to complete the study.

The mice were exposed to STD via the drinking water at 125, 250, 500, 1000, or 2000 mg/L for 28 d and through the morning of Day 29 until the time of study termination. The doses, vehicle, and route of exposure were selected for consistency with the 13-week NTP mouse toxicology studies, and were based on mechanistic immunotoxicology studies conducted by the Naval Health Research Center Detachment Environmental Health Effects Laboratory (NHRC Det) (Osterburg et al. 2014), and published long-term exposure studies (Barbera et al. 2001). Although these doses were higher than what is typically encountered in the natural environment, they represented the maximum concentration found in tap water (337 mg/L) (Koutsospyros et al. 2006) measured in Nevada, soil pore waters (400 mg/L) at military firing ranges (Clausen & Korte 2009), and doses used in a clinical study (200 mg/d) (Hanzu et al. 2010). Doses of >500 ppm represented geothermal fluids (1000 mg/L) (Seiler et al. 2005) in Nevada, surface/sub-surface soils at military firing ranges (>2000 mg/kg) (Clausen & Korte 2009), and doses used in animal research on the potential for sodium tungstate as a treatment for diabetes and obesity (2000 mg/L) (Barbera et al. 2001; Nakhaee et al. 2010).

Positive controls used in these studies included rabbit anti-asialo GM1 antibody (AAGM1, Wako BioProducts, Richmond, VA), maleic vinyl ether (MVE; Hercules Incorporated, Wilmington, DE) and cyclophosphamide (CPS; Sigma, St. Louis, MO). AAGM1, 0.2 ml of a 10% (v/v) solution in sterile physiological saline administered by intraperitoneal (IP) injection 24 h prior to necropsy, was the positive control for natural killer (NK) cell activity. MVE, 50 mg/kg administered in a single intravenous (IV) injection 24 h prior to necropsy, was the positive control for mononuclear phagocytic system (MPS) activity. CPS, given at a dose of 50 mg/kg by IP injection once daily during the last 4 d of the exposure period (last 5 d for the keyhole limpet hemocyanin [KLH] study), was the positive control for all other assays.

Toxicological studies

Drinking water consumption

Water bottles were changed twice a week; a control drip bottle, in an empty cage, was included for each treatment group to measure water loss due to movement of the cage or rack. Water consumption was determined as described previously (Auttachoat et al. 2009) and was expressed on an individual animal basis in terms of g/d. Water consumption data from all 10 cohorts were pooled for statistical analysis.

Body and organ weights

Mice were weighed prior to initiation of the study, and on Days 1, 8, 15, 22 and 29. Body weight data from all 10 cohorts were pooled for statistical analysis. On Day 29, animals were euthanized by CO₂ inhalation followed by cervical dislocation. Organ weights were obtained for the liver, spleen, lungs, thymus, kidneys and adrenal glands.

Hematology parameters

A panel of hematological parameters was analyzed in blood using a Hemavet 1500FS (Drew Scientific, Waterbury, CT). Hematology parameters evaluated included: erythrocyte and leukocyte numbers, leukocyte differentials, hemoglobin, hematocrit, mean corpuscular volume (MCV), mean corpuscular hemoglobin (MCH), mean corpuscular hemoglobin concentration (MCHC), and platelet number. Reticulocytes were evaluated using ReticCount™ stain and FACScan flow cytometer (BD Biosciences Immunocytometry Systems, San Jose, CA), according to the directions of the manufacturer.

Histopathology

At necropsy, liver, spleen, lungs, thymus, kidneys, adrenals, bone marrow (femur), gastrointestinal (GI) tract with Peyer’s patches, and mesenteric lymph nodes (LN), submandibular LN, and popliteal LN were collected and fixed in 10% neutral buffered formalin for histopathological evaluation. The lymphoid organs were evaluated using enhanced histopathology (EH) guidelines (Elmore 2006a–e); non-lymphoid organs were evaluated by traditional histopathology methods. All histopathology evaluations were conducted in accordance with the NTP Immunotoxicity Study Pathology Specifications (http://ntp.niehs.nih.gov/ntp/htdocs/levels/finalntpitoxspecs_508.pdf).

Immunological studies

Spleen cell immunophenotyping

Single-cell suspensions of splenocytes were analyzed by flow cytometry to quantify various cell populations, as previously described (Auttachoat et al. 2009). The following cell populations were evaluated: B-lymphocytes (B-cells, Ig⁺), total T-lymphocytes (T-cells, CD3⁺), T-cell subsets (CD4⁺CD8⁻, CD4⁻CD8⁺ and CD4⁺CD8⁺), NK cells (NK1.1⁺CD3⁻) and macrophages (Mac-3⁺). Isotype-matched irrelevant antibodies were used as controls. Cells were counted on a BD FACScan Flow Cytometer. Non-viable cells and red blood cells (RBC) were eliminated using a gate setting that excluded propidium iodide (PI) fluorescence and a forward scatter threshold above RBC size. For each sample, 5000 PI-negative events were counted. Data were analyzed with CellQuest software v. 3.2.1 (BD Biosciences).

T-dependent antibody responses to SRBC and KLH

The spleen IgM antibody-forming cell (AFC) response to sheep red blood cells (SRBC) was enumerated using a modified hemolytic plaque assay (Jerne et al. 1963; White et al. 2010) as previously described. The mice were immunized with SRBC on Day 25. The data were expressed in terms of specific activity (AFC/10⁶ splenocytes) and total spleen activity (AFC/spleen). Serum IgM antibody titers to SRBC were evaluated in the same animals using an enzyme-linked immunosorbant assay (ELISA) as previously described (Temple et al. 1993; Auttachoat et al. 2009). Serum anti-KLH IgM antibody levels were evaluated in mice immunized with KLH by IV injection on Day 24 of the study using ELISA, as previously described (Smith et al. 2013). Each sample was analyzed using 10 serial two-fold dilutions, multipoint analysis, and SoftMax (Molecular Devices Corp., Sunnyvale, CA) software.

MLR to DBA/2 mouse spleen cells

The one-way mixed leukocyte response (MLR) was conducted as previously described (Guo et al. 2000), with incorporation of [³H]-thymidine (Perkin Elmer, Inc., Waltham, MA) into proliferating cells as the assay endpoint. Samples were counted using a PerkinElmer 1450 Microbeta Trilux Liquid Scintillation and Luminescence Counter. Data were expressed as counts per minute (CPM)/10⁵ cells.

Anti-CD3 mediated spleen cell proliferation

T-cell proliferation following stimulation with anti-CD3 antibody was evaluated as described in Smith et al. (2010), using Biocoat T-cell activation plates (BD Biosciences). Samples were counted using the PerkinElmer 1450 Counter (Perkin Elmer). [³H]-thymidine incorporation into proliferating cells was used as the assay endpoint. Data were expressed as CPM/2 × 10⁵ cells.

CTL activity

The cytotoxic activity of T_CTL cells (CTL activity) against P815 mastocytoma cells was conducted in two phases as described previously (Bradley et al. 1994; Smith et al. 2010). After a 5-d induction phase, the cells were harvested and incubated for 4 h in 96-well plates in the presence of P815 cells labeled with [⁵¹Cr] (Na[⁵¹Cr]O₄, Perkin Elmer, Inc., Waltham, MA) (effector phase; 25:1 to 0.75:1 effector:target ratio). Supernatant from each well (100 μl) was then counted on a Wallac 1480 Wizard 3 in. γ-counter (Perkin Elmer, Shelton, CT). Release of [⁵¹Cr] into the supernatant was used as the endpoint of the assay, and results were expressed in terms of percent cytotoxicity, which was calculated as (experimental release CPM – spontaneous release CPM)/(maximum release CPM – spontaneous release CPM). Spontaneous and maximum release levels of [⁵¹Cr] into the media were determined by adding 0.1 ml of complete RPMI media or 0.1% Triton X-100, respectively, to each of 12 replicate cultures containing target cells.

Delayed-type hypersensitivity (DTH) response to Candida albicans

The DTH response to C. albicans was conducted as described by Smith and White (2010). Mice were immunized with formalin-fixed C. albicans (AlerChek Inc., Portland, ME) by subcutaneous (SC) injection on Day 21 and challenged on Day 29 with the C. albicans antigen chitosan in the right footpad. Footpad swelling was measured prior to challenge and 24 [±2] h post-challenge. Footpad swelling was calculated as [(post-measurement – pre-measurement) × 100] and reported in terms of mm × 100. A challenge only group, which received the chitosan injection on Day 29 without prior immunization with C. albicans, was included to control for non-specific footpad swelling.

NK-cell activity

NK-cell activity was assessed as earlier described (Wilson et al. 2001; Auttachoat et al. 2009) using [⁵¹Cr]-labeled YAC-1 cells as the target for NK-mediated cytotoxicity. Effector:target (E:T) ratios of 200:1, 100:1, 50:1, 25:1, 12.5:1, and 6.25:1 were used. Following a 4-h incubation, 100 μl of supernatant from each well was counted in the Wallac γ-counter (Perkin Elmer, Shelton, CT). Results were expressed as percent cytotoxicity, which was calculated in the same manner as for the CTL assay.

Mononuclear phagocyte system (MPS) activity

The functional activity of the MPS was evaluated by measuring the vascular clearance and uptake of [⁵¹Cr]-labeled SRBC ([⁵¹Cr]-SRBC) by fixed-tissue macrophages of the liver, spleen, lung, thymus and kidney, as previously described (White et al. 1985). In brief, mice were injected IV with [⁵¹Cr]-SRBC on Day 29, and blood was collected from the tail vein at multiple timepoints over a 30-min period (60 min for positive control animals) and counted in the Wallac γ-counter (Perkin Elmer, Shelton, CT) to determine the [⁵¹Cr]-SRBC vascular half-life. Sixty minutes after the [⁵¹Cr]-SRBC injection, these mice were euthanized, exsanguinated, and the liver, spleen, lungs, thymus, and kidneys were removed, weighed, and counted in the γ-counter to determine the organ uptake of the radiolabeled SRBC. Results were expressed in terms of percent uptake and in terms of CPM/mg of tissue (“specific activity”).

Bone marrow DNA synthesis, colony formation, and differentials

Bone marrow was obtained by flushing the medullary cavities of the left femur with RPMI containing 2% fetal bovine serum (FBS, ThermoFischer Scientific, Grand Island, NY). Single nucleated cell suspensions were prepared and adjusted to the desired concentration for each assay. All bone marrow assays were conducted as described previously (Smith et al. 2013). The following endpoints were evaluated: DNA synthesis, colony-forming units (CFU) and burst-forming units (BFU), and immunophenotyping. DNA synthesis was assessed through the incorporation of [³H]-thymidine into proliferating cells. Samples were counted using the PerkinElmer 1450 counter (Perkin Elmer, Shelton, CT), and the data were expressed as CPM/6 × 10⁵ cells. Commercially available MethoCult™ kits (STEMCELL Technologies, Vancouver, British Columbia, Canada) were used according to the kit instructions to assess colony formation

The CFU and BFU evaluated were CFU-E and BFU-E following culture for 2–5 d at 37 °C with 3 U erythropoietin/ml (STEMCELL Technologies, Vancouver, British Columbia, Canada), CFU-GM following culture for 12–13 d at 37 °C with 20 ng granulocyte-macrophage colony-stimulating factor (CSF-GM)/ml (R&D Systems, Minneapolis, MN), and CFU-M following culture for 12–13 d at 37 °C with 40 ng macrophage CSF (CSF-M)/ml (R&D Systems, Minneapolis, MN). Single cell suspensions of bone marrow cells were analyzed by flow cytometry to quantify various cell populations, as described in Smith et al. (2013), and in the Spleen cell immunophenotyping section above. The bone marrow cell populations evaluated were B-cell lineage (CD45R/B220⁺), T-cell lineage (CD3⁺), erythroid lineage (TER-119⁺), granulocytes/monocytes (CD11b⁺) and neutrophils (Gr-1⁺). High expression of Gr-1 indicated a predominantly neutrophil population, whereas intermediate expression was more indicative of a mixed population containing neutrophils, myelocytes, and other immature progenitors (Hestdal et al. 1991).

Statistical analysis

Results are presented as mean ± SE for 7–8 mice/group. Statistical analyses were conducted using Data Entry, Modification, and Statistics Programs Version 6.0 (Apple, Inc., Cupertino, CA) and SAS 9.3 (SAS Institute, Cary, NC). Data were evaluated for homogeneity of variances using Bartlett’s test or Levene’s test. Homogeneous data were evaluated using analysis of variance (ANOVA) and Dunnett’s test. Non-homogeneous data were evaluated using Welch ANOVA and, for pairwise comparisons with the control group, Wilcoxon Rank Sum Tests or Dunn’s test (bone marrow data). Student’s t-test was used to compare the vehicle and positive control groups, and for the DTH challenge only group. Jonckheere’s test (1954) was used to test for dose-related trends. Data that were different from control at two-sided p ≤ 0.05 and trends with one-sided p ≤ 0.05 were considered statistically significant.

Results

Analysis of test article

The 125, 1000, and 2000 mg STD/L dose formulations were analyzed using ion chromatography. The concentrations of all dose formulations tested were within 10% of the target, the NTP acceptance limit. The average (N = 5) determined concentration for the 125 mg STD/L formulation was 123.4 mg STD/L, the average for the 1000 mg STD/L formulation was 977.8 mg STD/L, and the average for the 2000 mg STD/L formulation was 1870 mg STD/L. At 1000 and 2000 mg/L, STD formed a white precipitate following the initial dissolution in tap water. The precipitate was analyzed via SEM with EDS detection and XRD. Calcium tungstate was the major component of the precipitate; sodium tungstate was the minor component. The EDS elemental analysis indicated that the precipitate was composed of calcium (11.6%), sodium (1.0%), tungsten (59%), and oxygen (28.4%). No precipitate formed when STD solutions were prepared in deionized water. These data indicated that despite the formation of the precipitate, the concentration of STD remained within acceptable limits, and that the precipitate may have been the result of a reaction between STD and a calcium ion that was present in tap water, but was not present in deionized water. Since the dose formulations were within NTP specifications, the tap water formulations were used throughout the study.

Drinking water consumption, body weight, and organ weights

The average drinking water consumption was evaluated as described in the Materials and methods section. With the exception of a 9% decrease (p ≤ 0.01) for animals in the 2000 mg/L exposure group at the 1-week timepoint, there were no significant differences in water consumption between control- and STD-exposed mice (data not shown). There were no significant differences in body weights throughout the study, or in overall body weight gain (Day 29–Day 1), between treated and control animals (data not shown). Weights of the liver, spleen, thymus, lungs, and kidneys were measured as a component of three studies, the MPS study, the hematology study, and the collection of tissues for histopathology. No effects were observed on the absolute or relative weights of these organs in any of the studies (data not shown), with two exceptions. In the hematology study, an increase (11%, p ≤ 0.01) in relative liver weight at 2000 mg STD/L was observed. In the MPS study, an increase (9%, p ≤ 0.05) in spleen weight at 250 mg STD/L was noted. These effects were not observed in the remaining studies and thus, were not considered to be biologically significant.

Clinical pathology

The absolute numbers of specific blood leukocyte populations were unaffected by STD exposure (Table 1). However, when the leukocyte differentials were evaluated on a percent basis, neutrophils were decreased 17–33% in all STD groups, and lymphocytes were increased 14% at 250–1000 mg STD/L, relative to the vehicle control. In addition, monocytes were decreased (15% and 21%) with 500 and 1000 mg STD/L. No effects were observed on blood hematology parameters, with the exception of MCH and MCHC. MCH was increased (4%) at 125 mg STD/L, and decreased (4%) at 250 and 500 mg STD/L; MCHC was decreased 5% with 500 mg STD/L. The effects on hematology are minimal and not sufficiently significant to contribute to the assessment of direct immunotoxicity (Descotes 2004).

Table 1. Hematology and blood leukocyte differentials in female B₆C₃F₁/N mice exposed to STD in the drinking water for 28 d.

		Sodium tungstate dihydrate (mg/L)					CPS 50 mg/kg	Trend analysis
Parameter	Vehicle	125	250	500	1000	2000
Hematology
Erythrocytes (10^6/mm^3)	9.00 ± 0.15	9.30 ± 0.22	8.84 ± 0.19	8.70 ± 0.18	9.19 ± 0.60	8.72 ± 0.14	7.88 ± 0.35*	p ≤ 0.05
Reticulocytes (%)	4.3 ± 0.2	4.6 ± 0.2	4.6 ± 0.2	4.1 ± 0.2	4.6 ± 0.3	4.8 ± 0.2	1.6 ± 0.1**	NS
Hemoglobin (g/dl)	12.8 ± 0.2	13.7 ± 0.2	12.1 ± 0.3	11.8 ± 0.3	12.4 ± 0.5	12.6 ± 0.2	9.9 ± 0.8**	p ≤ 0.05
Hematocrit (%)	47.9 ± 0.7	50.0 ± 1.2	46.9 ± 1.2	46.7 ± 0.9	49.4 ± 3.5	46.4 ± 0.9	41.8 ± 2.0*	p ≤ 0.05
MCV (fl)	53.3 ± 0.3	53.8 ± 0.2	53.1 ± 0.3	53.7 ± 0.2	53.6 ± 0.3	53.3 ± 0.3	53.1 ± 0.2	NS
MCH (pg)	14.2 ± 0.1	14.7 ± 0.2*	13.7 ± 0.3*	13.6 ± 0.2**	13.9 ± 1.1	14.4 ± 0.1	12.4 ± 0.7*	NS
MCHC (g/dl)	26.7 ± 0.2	27.4 ± 0.4	25.9 ± 0.5	25.3 ± 0.3*	26.0 ± 2.1	27.1 ± 0.2	23.3 ± 1.3*	NS
Platelets (10^3/μl)	684 ± 73	500 ± 68	782 ± 86	909 ± 101	684 ± 75	617 ± 65	917 ± 176	NS
Leukocytes (10^3/mm^3)	5.96 ± 0.60	5.00 ± 0.61	5.71 ± 0.41	5.56 ± 0.37	5.85 ± 0.41	4.68 ± 0.42	1.94 ± 0.21**	NS
Absolute leukocyte differentials (10^3/mm^3)
Lymphocytes	3.62 ± 0.29	3.28 ± 0.31	4.01 ± 0.30	3.92 ± 0.24	4.10 ± 0.29	3.13 ± 0.25	1.47 ± 0.17**	NS
Neutrophils	1.67 ± 0.24	1.18 ± 0.19	1.05 ± 0.10	1.03 ± 0.10	1.16 ± 0.09	1.07 ± 0.16	0.28 ± 0.04**	NS
Eosinophils	0.17 ± 0.05	0.14 ± 0.05	0.21 ± 0.04	0.19 ± 0.04	0.18 ± 0.03	0.10 ± 0.03	0.05 ± 0.01*	NS
Basophils	0.07 ± 0.03	0.05 ± 0.02	0.10 ± 0.02	0.07 ± 0.02	0.07 ± 0.01	0.05 ± 0.02	0.02 ± 0.01	NS
Monocytes	0.44 ± 0.06	0.35 ± 0.05	0.35 ± 0.03	0.35 ± 0.03	0.34 ± 0.03	0.33 ± 0.04	0.12 ± 0.02**	NS
Percent leukocyte differentials
% Lymphocytes	61.91 ± 2.54	67.11 ± 1.94	70.28 ± 2.20*	70.86 ± 1.82*	70.28 ± 1.44*	67.43 ± 2.25	75.16 ± 2.33**	p ≤ 0.05
% Neutrophils	27.31 ± 1.53	22.75 ± 1.03*	18.35 ± 1.2**	18.37 ± 0.92**	19.88 ± 0.8**	22.24 ± 1.33*	14.74 ± 1.51**	p ≤ 0.05
% Eosinophils	2.54 ± 0.72	2.37 ± 0.56	3.57 ± 0.66	3.33 ± 0.71	2.96 ± 0.43	2.23 ± 0.71	2.97 ± 0.60	NS
% Basophils	0.98 ± 0.38	0.84 ± 0.23	1.65 ± 0.40	1.27 ± 0.27	1.16 ± 0.19	1.04 ± 0.35	0.97 ± 0.36	NS
% Monocytes	7.25 ± 0.42	6.93 ± 0.66	6.15 ± 0.29	6.17 ± 0.24*	5.72 ± 0.32*	7.06 ± 0.65	6.18 ± 0.41	NS

MCV: mean corpuscular volume; MCH: mean corpuscular hemoglobin; MCHC: mean corpuscular hemoglobin concentration; CPS: cyclophosphamide; NS: not significant. Values represent the mean (± SE) from 7 to 8 animals/group.

Bold values indicates statistically significant STD-treatment related data.

*p ≤ 0.05.

**p ≤ 0.01.

Histology

No treatment-related lesions were detected in the thymus, spleen, mesenteric lymph node, popliteal lymph node, mucosa-associated lymphoid tissues, or bone marrow.

Spleen cell immunophenotyping

No effects on the total spleen cell numbers or on the absolute or percent values of splenic B-cells, T-cells, T-cell subsets, NK cells, or macrophages were observed following 28 d of exposure to STD (data not shown).

Humoral-mediated immunity

No STD exposure-related effects were observed on the AFC response to SRBC in the plaque assays (data not shown) or on serum IgM antibody levels to SRBC or KLH (data not shown).

Cell-mediated immunity

Treatment with 1000 mg STD/L resulted in decreased ex vivo activity of splenic T_CTL cells against P815 mastocytoma cells, relative to activity resulting from host exposure to the vehicle control, at all effector:target (E:T) ratios examined. The decreases in percent cytotoxicity at E:T ratios of 25:1, 12.5:1, 6:1, 3:1, 1.5:1, and 0.75:1 were 27, 34, 48, 57, 67, and 80%, respectively (Figure 1A). At 500 mg STD/L, T_CTL activity was increased at E:T ratios of 3:1 (50%), 1.5:1 (77%), and 0.75:1 (135%). No effects were observed with 125, 250, or 2000 mg STD/L. Similarly, as compared with the vehicle control mice cells, a decrease (32%) in ex vivo leukocyte proliferation in response to stimulation by allogeneic leukocytes was noted following host exposure to 1000 mg STD/L, with an increase in activity (32%) at 250 mg STD/L (Figure 1B). The response of the cells from animals in the high-dose group (2000 mg STD/L) was not different from that observed in the vehicle control mice cells. When stimulated with anti-CD3 antibody, splenocyte cell proliferation was decreased 21% with 1000 mg STD/L (Figure 2A), although this point was not statistically significant. The basal proliferative response in those cultures was decreased 11% at 125 mg STD/L, and increased 32% at 500 mg STD/L. No effects were observed on the delayed-type hypersensitivity response to C. albicans (Figure 2B).

Figure 1. Cytotoxic T-lymphocyte and mixed-leukocyte responses in female B₆C₃F₁/N mice exposed to STD in drinking water for 28 d. Panel A: cytotoxic T-lymphocyte (CTL) response. Splenocytes were cultured for 5 d with P815 mastocytoma cells (induction phase), harvested, and incubated in the presence of [⁵¹Cr]-labeled P815 cells for 4 h (effector phase). Release of [⁵¹Cr] into the supernatant was used as the endpoint of the assay. Spontaneous release over the 4-h incubation period was 11.1% of maximum release. Panel B: mixed-leukocyte response. Splenocytes were cultured for 5 d in the presence of mitomycin C-treated DBA/2 allogenic stimulator cells. R = responder (B₆C₃F₁/N) cells only, R + S = responder and stimulator (DBA/2) cells. All MLR cultures were labeled with [³H]-thymidine 18–24 h prior to harvest. [³H]-thymidine incorporation into proliferating cells was used as the endpoint of the assay. Results are expressed as percent cytotoxicity for the CTL, CPM/10⁵ splenocytes for the MLR. Asterisks indicate statistically significant differences from VH control; *p ≤ 0.05; N = 7–8 mice/group for VH, STD and CPS (MLR only); N = 4 for the CPS (CTL) group. Due to the reduction in spleen weight and cell number, it was necessary to pool splenocytes from two positive control CPS-treated animals to achieve the necessary cell concentrations for the CTL assay.

Figure 2. Anti-CD3-mediated splenic T-cell proliferation and delayed-type hypersensitivity responses in female B₆C₃F₁/N mice exposed to STD in drinking water for 28 d. Panel A: anti-CD3 mediated proliferation. Splenocytes were cultured in the presence (stimulated) or absence (unstimulated) of anti-CD3 antibody in 96-well plates for 3 d. All anti-CD3 cultures were labeled with [³H]-thymidine 18–24 h prior to harvest. Incorporation of [³H]-thymidine into proliferating cells was used as the endpoint of the assay. Panel B: delayed-type hypersensitivity response. Mice were sensitized on Day 21 with formalin-fixed C. albicans, and challenged in the right footpad on Day 29 with the C. albicans antigen, chitosan. Footpad swelling was determined 24 h post-challenge. Results are expressed as CPM/2 × 10⁵ splenocytes for the anti-CD3 assay, and as mm × 100 for the DTH assay. N = 8 mice/group. Asterisks indicate statistically significant differences from VH control; *p ≤ 0.05

Innate immunity

There was no alteration in the ability of fixed-tissue macrophages to phagocytize SRBC in liver, spleen, lung, or thymus (Table 2). Significant decreases were observed in the uptake of [⁵¹Cr]-SRBC in the kidney, which has been shown to quickly and efficiently eliminate the majority of tungsten (McDonald et al. 2007), with the 125 (percent) and 2000 (percent and specific activity) mg STD/L regimens. Natural killer cell activity was measured at E:T ratios of 6.25:1–200:1 for all doses (N = 8); no significant effect on NK cell activity was noted in any STD treatment group. NK cell activity of cells from spleens of mice treated with the AAGMI positive control was <1% at all E:T ratios, and was statistically significant at E:T ratios of 25:1 and above (data not shown)

Table 2. Functional activity of the mononuclear phagocytic system in mice exposed to STD in the drinking water for 28 d.

		Sodium tungstate dihydrate (mg/L)					MVE 50 mg/kg	Trend analysis
Parameter	Vehicle	125	250	500	1000	2000
Vascular half-life (min)	9.0 ± 0.6	7.9 ± 0.9	8.9 ± 0.8	9.3 ± 1.0	8.3 ± 0.7	9.4 ± 0.7	145.4 ± 34.7**	NS
Body weight (g)	24.1 ± 0.5	23.8 ± 0.4	24.6 ± 0.4	23.0 ± 0.7	22.8 ± 0.5	22.6 ± 0.4	28.1 ± 0.7**	p ≤ 0.01
Liver weight (mg)	947 ± 28	984 ± 25	981 ± 37	867 ± 32	896 ± 23	927 ± 26	1234 ± 38**	p ≤ 0.05
% Uptake	40.7 ± 2.8	41.7 ± 2.6	44.2 ± 1.7	40.6 ± 3.0	42.1 ± 2.1	39.4 ± 2.7	4.7 ± 0.4**	NS
cpm/mg	109 ± 9	106 ± 7	117 ± 4	113 ± 7	113 ± 6	100 ± 5	11 ± 1**	NS
Spleen weight (mg)	75 ± 4	78 ± 3	82 ± 1*	73 ± 4	75 ± 3	83 ± 13	96 ± 5**	NS
% Uptake	16.3 ± 1.1	14.3 ± 1.9	15.4 ± 1.4	12.9 ± 1.1	14.8 ± 0.9	14.0 ± 0.9	14.4 ± 2.1	NS
cpm/mg	558 ± 51	460 ± 62	484 ± 42	431 ± 33	480 ± 33	434 ± 44	436 ± 56	NS
Lung weight (mg)	187 ± 10	205 ± 8	219 ± 17	201 ± 13	217 ± 13	179 ± 6	210 ± 9	NS
% Uptake	0.8 ± 0.1	0.5 ± 0.1	0.8 ± 0.1	0.6 ± 0.1	0.7 ± 0.1	0.7 ± 0.2	4.2 ± 0.4**	NS
cpm/mg	11 ± 2	6 ± 1	10 ± 1	7 ± 1	8 ± 1	10 ± 2	60 ± 7**	NS
Thymus weight (mg)	58 ± 2	55 ± 3	53 ± 3	58 ± 2	55 ± 5	55 ± 2	72 ± 6	NS
% Uptake	0.04 ± 0.01	0.03 ± 0.004	0.02 ± 0.01	0.02 ± 0.01	0.03 ± 0.01	0.03 ± 0.01	0.35 ± 0.09**	NS
cpm/mg	2 ± 1	1 ± 1	1 ± 1	1 ± 1	1 ± 1	1 ± 1	13 ± 3**	NS
Kidney weight (mg)	309 ± 7	320 ± 14	336 ± 9	312 ± 7	310 ± 6	312 ± 8	374 ± 9**	NS
% Uptake	1.9 ± 0.1	1.5 ± 0.1*	1.8 ± 0.1	2.1 ± 0.2	1.8 ± 0.2	1.5 ± 0.1**	1.3 ± 0.2*	p ≤ 0.05
cpm/mg	15 ± 1	12 ± 1	14 ± 1	16 ± 2	14 ± 2	11 ± 1*	10 ± 2*	p ≤ 0.01

MVE: maleic vinyl ether positive control; NS: not significant. Values represent mean (± SE) from 7 to 8 animals/group.

Bold values indicates statistically significant STD-treatment related data.

*p ≤ 0.05.

**p ≤ 0.01.

Bone marrow

Total cell numbers in the femur were enumerated as a component of two studies, the immunophenotyping study and the DNA synthesis/CFU study. Exposure to STD in the drinking water induced a 41% increase in total bone marrow cell numbers in the immunophenotyping study, but not in the CFU study (Table 3). In the immunophenotyping study, the absolute values of CD3⁺ (T-cell lineage, 86%), B220⁺ (B-cell lineage, 53%), CD11b⁺ (granulocyte/macrophage lineage, 36%), TER-119⁺ (erythroid lineage, 50%), and intermediate (22%), high (35%), and total (31%) Gr1⁺ (neutrophils, myelocytes, and immature progenitors) cells were also increased, relative to the control, after host exposure to 2000 mg STD/L (Table 3). However, only the CD3⁺ data were statistically significant. There was no effect on DNA synthesis of bone marrow cells, or colony-forming units following in vitro stimulation of isolated bone marrow cells with CSF-M, CSF-GM, or CSF-E (data not shown).

Table 3. Bone marrow cell immunophenotyping in female B₆C₃F₁/N mice exposed to STD in the drinking water for 28 d.

Treatment	Cells/Femur	B220^+	CD3^+	Gr-1^+ (Int)	Gr-1^+ (High)	Gr-1^+ (Total)	CD11b^+	TER-119^+
Absolute values (×10^6)
Vehicle	7.47 ± 0.61	1.5 ± 0.1	0.1 ± 0.1	0.9 ± 0.1	2.6 ± 0.2	3.5 ± 0.3	3.3 ± 0.2	2.2 ± 0.2
STD
125 mg/L	8.82 ± 0.52	1.7 ± 0.2	0.1 ± 0.1	1.0 ± 0.1	3.2 ± 0.2	4.1 ± 0.2	3.8 ± 0.2	2.5 ± 0.2
250 mg/L	9.44 ± 0.68	2.0 ± 0.2	0.1 ± 0.1	1.1 ± 0.1	3.2 ± 0.2	4.2 ± 0.2	3.9 ± 0.2	2.8 ± 0.2
500 mg/L	8.60 ± 0.64	1.8 ± 0.1	0.1 ± 0.1	1.0 ± 0.1	2.7 ± 0.3	3.7 ± 0.3	3.5 ± 0.3	2.7 ± 0.2
1000 mg/L	7.25 ± 0.64	1.7 ± 0.2	0.1 ± 0.1	0.8 ± 0.1	2.4 ± 0.2	3.2 ± 0.2	3.1 ± 0.3	2.1 ± 0.2
2000 mg/L	10.57 ± 1.01	2.3 ± 0.3	0.2 ± 0.1*	1.1 ± 0.1	3.5 ± 0.3	4.6 ± 0.4	4.5 ± 0.4	3.3 ± 0.3
CPS (50 mg/kg)	4.07 ± 0.44**	0.1 ± 0.1**	0.1 ± 0.1**	0.1 ± 0.1**	0.1 ± 0.1**	0.2 ± 0.1**	0.2 ± 0.1**	3.6 ± 0.4**
Trend analysis	NS	p ≤ 0.05	p ≤ 0.05	NS	NS	NS	NS	NS
Percent values
Vehicle		20.1 ± 0.8	1.3 ± 0.1	11.4 ± 0.3	34.8 ± 1.0	46.1 ± 1.2	43.9 ± 1.0	29.3 ± 0.8
STD
125 mg/L		18.9 ± 1.8	1.5 ± 0.2	11.4 ± 0.3	35.9 ± 1.3	47.2 ± 1.5	43.4 ± 1.6	28.6 ± 1.9
250 mg/L		20.6 ± 1.1	1.4 ± 0.1	11.3 ± 0.4	34.3 ± 1.3	45.5 ± 1.5	41.2 ± 1.2	29.5 ± 1.0
500 mg/L		21.5 ± 1.0	1.5 ± 0.1	11.5 ± 0.2	31.2 ± 1.0	42.6 ± 1.0	40.5 ± 0.9	31.7 ± 0.6
1000 mg/L		23.7 ± 0.9	1.5 ± 0.1	11.4 ± 0.2	32.9 ± 1.3	44.2 ± 1.4	42.5 ± 0.9	29.1 ± 1.0
2000 mg/L		21.2 ± 1.2	1.5 ± 0.1	10.7 ± 0.3	32.9 ± 1.2	43.6 ± 1.3	42.7 ± 0.8	31.7 ± 1.1
CPS (50 mg/kg)		1.4 ± 0.1**	1.3 ± 0.1	2.6 ± 0.2**	2.7 ± 0.3**	5.3 ± 0.4**	4.5 ± 0.4**	88.1 ± 0.5**
Trend analysis		p ≤ 0.05	NS	NS	p ≤ 0.05	p ≤ 0.05	NS	NS

B220⁺: B-lymphocyte lineage; CD3⁺: T-lymphocyte lineage; Gr-1⁺ (int): Neutrophils, myelocytes and other immature progenitors; Gr-1⁺ (high): neutrophils; Gr-1 + (total): total Gr-1⁺ cells; CD11b⁺: granulocytes and monocytes; TER-119⁺: Erythroid lineage; STD: sodium tungstate dihydrate; CPS: cyclophosphamide (positive control); NS: not significant. Values represent the mean (± SE) from 8 animals/group.

Bold values indicates statistically significant STD-treatment related data.

* p ≤ 0.05.

** p ≤ 0.01.

Discussion

Exposure to tungsten has been associated with contaminated surface water, groundwater and well water in areas where the element is naturally occurring (ATSDR 2005), with inflammation and pneumonia in people following workplace exposure (Moriyama et al. 2007), and with persistence of the element in the body after clinical trials of medical therapies involving tungsten (ATSDR, 2005; Hanzu et al. 2010; Bolt et al. 2015). Published reports indicated that sodium tungstate may modulate hematopoiesis, immune cell populations, and immune responses in rodent models (Osterburg et al. 2010, 2014; Fastje et al. 2012).

The data presented herein suggest that exposure to STD in drinking water may adversely effect cell-mediated immunity following sensitization with an immune stimulating agent. The cytolytic function of antigen-specific T_CTL lymphocytes against P815 mastocytoma cells was diminished following exposure to 1000 mg STD/L. Splenocyte proliferation in response to allogeneic leukocytes or the anti-CD3 antibody, two other antigen-driven reactions, was also diminished with 1000 mg STD/L. There were no effects on the cell number of any T-lymphocyte subsets in animals that had not been sensitized. Both the C_TCL and MLR responses exhibited an increase in cell proliferation at lower doses, followed by a decrease with 1000 mg STD/L. Similar biphasic responses have been reported for insoluble tungsten (Peao et al. 1993) and for other trace elements (Llabjani et al. 2014). Llabjani et al. proposed that many elements induce a protective cellular response at low concentrations and a toxic effect at higher concentrations, possibly in conjunction with modulation of reactive oxygen species. Surprisingly, no effects were observed on cell-mediated immune responses at 2000 mg STD/L. Although an insoluble calcium tungstate precipitate formed at the two highest doses, the determined concentrations were within NTP specifications, and there was no apparent explanation for the lack of response.

Published immunophenotyping studies supported tungsten-induced modulation of T_CTL cell populations (CD8⁺) as seen in the current studies, and possibly acquired immunity and inflammation. However, the type of response varied depending on the form of tungsten and the route of exposure. Osterburg et al. (2014) demonstrated adverse effects on T-cell activation by soluble tungsten, and hypothesized that the suppression of acquired immunity could result in reduced host defense to immune challenges. Exposure to sodium tungstate in drinking water (200 mg/kg ≈1140 mg/L) led to a reduction in the percentage of activated (CD71⁺, transferrin receptor-positive) T_CTL cells and T-helper (T_H) cells in the spleens of C57BL/6 mice that had been challenged with T-cell activating bacterial antigen staphylococcal enterotoxin (SEB). Conversely, Moriyama et al. (2007) reported an increase in the numbers of CD8⁺ lymphocytes and CD163⁺ monocyte-macrophages in lungs (lesions, alveolar walls, and peribronchiole areas) of 17 human patients with occupationally induced hard metal lung disease, interstitial pneumonia, and centrilobular fibrotic lesions containing tungsten particles.

Both the current study and published reports indicated that adverse effects of tungsten exposure were observed primarily in models in which the animals or cells were co-exposed to an immune-stimulating agent, rather than through direct action. In the present study, the functional activity of T_CTL cells decreased at 1000 mg STD/L when splenic T_CTL lymphocytes from control and STD-treated mice were sensitized to P815 mastocytoma cells ex vivo, prior to functional assessment in the effector phase. Similarly, other studies have shown that tungstate-treated C57BL/6 mice developed significant splenomegaly when inoculated with the T-cell activating RSV (Fastje et al. 2012), and demonstrated suppression of activated T_H and T_CTL cells, when challenged with SEB (Osterburg et al. 2014). In the absence of RSV or SEB challenge, tungstate had no effect on spleen size or T-cell populations (Fastje et al. 2012; Osterburg et al. 2014). Exposure to tungsten in drinking water was also associated with increased tumor burden in BALB/c mice treated with 66Cl4 tumor cells (Bolt et al. 2015). Although the exact mechanisms remain unknown, Osterburg et al. (2014) and Kelly et al. (2013) proposed that tungstate exposure might induce suppression of adaptive immunity and/or leukemogenesis when in the presence of a stressor but that tungstate had little direct effect.

Modulation of reduction/oxidation (redox) balance and of apoptosis have been proposed as mechanisms for the immunotoxicity of tungstate. Disruption of redox processes leads to altered expression of FasL and Bcl2 in T-cells (Yang et al. 2013), and inhibition of CD8⁺ T-cell proliferation and function (Sklavos et al. 2008). Tungstate exposure has been associated with increased apoptosis in vitro in human and murine leukocytes (Osterburg et al. 2010; Guilbert et al. 2011); with DNA damage in vivo in bone marrow cells in C57BL/6J mice (Guilbert et al. 2011); with activation of antioxidant enzymes (Nakhaee et al. 2010; Donmez et al. 2014); and with up-regulation of pro-apoptotic genes (Lombaert et al. 2008; Harris et al. 2015) and proteins (Zhao et al. 2013) in rats and human and rodent cultured cells. However, in the present study, no increases in apoptosis, tingible body macrophages, or cell loss were evident in groups administered STD compared with control groups when hosts were evaluated using enhanced histopathology or spleen cell immunophenotyping.

Published absorption, distribution, metabolism and excretion (ADME) studies and biokinetic modeling indicated that tungstate was both absorbed and cleared rapidly from most tissues. The majority of the body burden of tungsten in rodents was retained in the bone, with some retention in the spleen and kidney (McDonald et al. 2007; Guandalini et al. 2011; McInturf et al. 2011; Kelly et al. 2013). In the current studies, T-lymphocyte lineage cells in bone marrow were increased at 2000 mg STD/L, but the alterations in B-lymphocyte and other progenitor cells were not significant. Damage to hematopoietic cells may require more extended exposure than the 28 d of the current study. The increase in late pro/large pre B-cells, and DNA damage to bone marrow cells, noted by Kelly et al. (2013) were observed following 16 weeks of exposure. The present studies did not reveal histopathological lesions in any tissue, although it is noteworthy that the exposure period was limited to 28 d and the mice evaluated for pathology were not co-exposed to an immune-stimulating agent. This is consistent with published reports showing no evidence of tungstate-induced histopathological injury to the immune organs in Sprague–Dawley rats at 5–125 mg/kg in a 70-d, two-generation, gavage study (McInturf et al. 2011).

Conclusions

In summary, exposure to STD in drinking water for 28 d at doses of 125–2000 mg/L had limited effect on humoral and innate immunity, on developing hematopoietic cells in the bone marrow, and on unstimulated splenocyte phenotypes in B₆C₃F₁/N mice. However, such exposure may have decreased the functional activity of T-lymphocytes at 1000 mg STD/L, as evidenced by the reduction of T_CTL activity, and the proliferative response to allogeneic leukocytes and the anti-CD3 antibody, while increasing the activity at lower doses. These data indicated that, under conditions of co-exposure to an immune-stimulating agent, such as tumor cells or genetically dissimilar leukocytes, STD may modulate the normal cell-mediated immune response.

Funding information

This research was supported by the NIH, National Institute of Environmental Health Sciences, NTP Contract N01-ES-55538, and the NTP Chemistry Support Services Contract N01-ES-05456.

Abbreviations:
AAGM1	=	rabbit anti-asialo GM1 antibody
ABC	=	antibody forming cell
ADME	=	absorption, distribution, metabolism, excretion
ANOVA	=	analysis of variance
ATSDR	=	Agency for Toxic Substances and Disease Registry
B-cells	=	B-lymphocytes
BFU	=	burst-forming units
CD	=	cluster designation
CDC	=	Center for Disease Control
CFU	=	colony-forming units
CPM	=	counts per minute
CPS	=	cyclophosphamide
CSF	=	colony-stimulating factor
CTL	=	cytotoxic T-lymphocyte
Cr	=	chromium
DTH	=	delayed-type hypersensitivity;
EDS	=	energy dispersive spectrometry
EH	=	enhanced histopathology
ELISA	=	enzyme-linked immunosorbant assay
E:T	=	effector:target
GI	=	gastrointestinal
GM	=	granulocyte macrophage
IgM	=	immunoglobulin M
IP	=	intraperitoneal
IV	=	intravenous
KLH	=	keyhole limpet hemocyanin
LN	=	lymph nodes
MAC	=	macrophage
MCH	=	mean corpuscular hemoglobin
MCHC	=	mean corpuscular hemoglobin concentration
MCV	=	mean corpuscular volume
MLR	=	mixed leukocyte response
MPS	=	mononuclear phagocytic system
MVE	=	maleic vinyl ether
NK	=	natural killer cell
NHRC Det	=	Naval Health Research Center Detachment Environmental Health Effects Laboratory
PI	=	propidium iodide
RBC	=	red blood cells
Redox	=	reduction/oxidation
RSV	=	respiratory syncytial virus
SC	=	subcutaneous
SE	=	standard error
SEB	=	bacterial antigen staphylococcal enterotoxin
SEM	=	scanning electron microscopy
SRBC	=	sheep red blood cells
STD	=	sodium tungstate dihydrate
T-cells	=	T-lymphocytes
USEPA	=	United States Environmental Protection Agency
USGS	=	United States Geological Survey
XRD	=	X-ray diffraction

Acknowledgements

Special thanks to Ronnetta Brown, Jackson Mitchell, Anthony Rapisardi, Dr. Wimolnut Auttachoat, and Dr. Tai Guo for outstanding technical assistance in conducting this research, and to Drs. Kristen Hobbie, and Warren Liewallen for pathology support. The authors also acknowledge the outstanding technical assistance of Deborah Musgrove, who was responsible for the conducting the technically challenging bone marrow colony assays. The authors also thank Drs. Matthew Stout and Erik Tokar for their critical review of the manuscript.

Disclosure statement

Dr. Kimber L. White Jr was the owner of a company, ImmunoTox^® Inc., that conducted immunotoxicological assessments under Good Laboratory Practices; however, none of the research presented here involved his company. Dr. Matthew J. Smith was an employee of ImmunoTox^® Inc., however, these studies were completed prior to his employment.