Skip to main content
NCBI home page
Toxicological Sciences logoLink to Toxicological Sciences
. 2017 Apr 7;158(1):63–75. doi: 10.1093/toxsci/kfx067

From the Cover: Tributyltin Alters the Bone Marrow Microenvironment and Suppresses B Cell Development

Amelia H Baker *, Ting Hua Wu †, Alicia M Bolt †, Louis C Gerstenfeld ‡, Koren K Mann †, Jennifer J Schlezinger §,1
PMCID: PMC6075590  PMID: 28398592

Abstract

Organotins are industrial chemicals and agricultural pesticides, and they contaminate both outdoor and indoor environments. Organotins are detectable in human sera at biologically active concentrations and are immuno-and neuro-toxicants. Triphenyltin, tributyltin (TBT) and dibutyltin activate peroxisome proliferator-activated receptor γ in bone marrow multipotent mesenchymal stromal cells and promote adipogenesis. TBT also has been shown to suppress osteogenesis; osteoblasts not only support bone homeostasis but also support B lymphopoiesis. In addition, developing B cells are highly sensitive to exogenous insults. Thus, we hypothesized that bone marrow B cells may be negatively affected by TBT exposure both directly, through activation of apoptosis, and indirectly, through alterations of the bone marrow microenvironment. TBT activated apoptosis in developing B cells at environmentally relevant concentrations (as low as 80 nM) in vitro, via a mechanism that is distinct from that induced by high dose (μM) TBT and that requires p53. TBT suppressed the proliferation of hematopoietic cells in an ex vivo bone marrow model. Concurrent treatment of stromal cells and B cells or pretreatment of stromal cells with TBT induced adipogenesis in the stromal cells and reduced the progression of B cells from the early pro B (Hardy fraction B) to the pre B stage (Hardy fraction D). In vivo, TBT induced adipogenesis in bone marrow, reduced “aging-sensitive” AA4+CD19+ B cells in bone marrow, and reduced splenic B cell numbers. Immunosenescence and osteoporosis are adverse health effects of aging, we postulate that TBT exposure may mimic, and possibly intensify, these pathologies.

Keywords: B cell, organotin, apoptosis, adipocyte, bone marrow microenvironment

Tributyltin (TBT) has been a contaminant of concern in the marine environment due to its use as an antifouling agent; however, the use of organotins in agricultural pesticides, wood preservatives and the manufacturing of plastics has resulted in significant land-based sources of this environmental contaminant (Cornelissen et al., 2008). Indeed, organotins are even measurable in house dust (Fromme et al., 2005; Kannan et al., 2010). Significant human exposure is indicated by the presence of organotins in liver and blood (0.05–450 nM) (as reviewed in Antizar-Ladislao, 2008). Fortunately, human body burdens appear to be decreasing (Levine et al., 2015).

TBT is a well-known immunotoxicant. High dose, acute TBT exposure causes thymic atrophy and suppresses responses to infection in rodent models (Bressa et al., 1991; Vos et al., 1984). One explanation for these findings is the induction of thymocyte apoptosis (Baken et al., 2007; Raffray and Cohen, 1993; Stridh et al., 2001; Tomiyama et al., 2009). TBT exposure, beginning at 6 days gestation through postnatal day 60, results in a reduction in spleen weight (Cooke et al., 2004), suggesting that B lymphocytes also may be targets of TBT toxicity. At micromolar concentrations, TBT activates rapid calcium/calmodulin protein kinase II/mitogen-activated protein kinase-driven apoptosis and necrosis in B cells (Bissonnette et al., 2010). On the other hand, TBT has been shown to reduce survival, proliferation and differentiation of mature human B cells (tonsillar B lymphocytes) at 100 nM in vitro (De Santiago and Aguilar-Santelises, 1999), suggesting that B lymphocytes may be more sensitive to TBT-induced insults than T lymphocytes. Indeed, in human long-term bone marrow cultures, TBT (1 nM) reduces B cell, but not T cell, numbers (Carfi et al., 2010). These results suggest that B cells are targets of TBT toxicity at human relevant concentrations, but little is known about the mechanisms of action.

Two aspects of B cell development are likely to confer susceptibility to environmental toxicants: (1) apoptotic deletion of dysfunctional lymphocytes and (2) dependence of lymphopoiesis upon stromal cell support/interactions, ie, the bone marrow microenvironment. The minimal expression of BCL-2 that allows for negative selection in IL-7-dependent precursor B cells confers a general susceptibility to apoptosis induced by a variety of therapeutics (radiation, corticosteroid, etoposide, cisplatin) (Griffiths et al., 1994). Developing B cells also are highly sensitive to polycyclic aromatic hydrocarbon-induced apoptosis (Teague et al., 2010).

Lymphopoiesis is supported by bone marrow stromal elements. The bone marrow stromal compartment consists of multipotent mesenchymal stromal cells (MSCs) and their progeny, including adipocytes and osteoblasts (Bianco, 2011). Alterations in the balance of stromal elements may negatively impact hematopoietic stem cell populations and B cell development, both of which are supported by the cells of the osteoblast lineage and actively suppressed by adipocytes (Bilwani and Knight, 2012; Calvi et al., 2003; Naveiras et al., 2009; Visnjic et al., 2004; Wu et al., 2009; Zhu et al., 2007). TBT potently activates both peroxisome proliferator-activated receptor γ (PPARγ), the master regulator of adipocyte differentiation, and its DNA-binding partner retinoid X receptor (RXR) (Grun et al., 2006; le Maire et al., 2009). TBT induces adipocyte differentiation in the mouse bone marrow-MSC line, BMS2 (EC50 10 nM) and in primary mouse and human bone marrow cultures (Carfi et al., 2008; Yanik et al., 2011). TBT has been shown to suppress osteogenesis in rat calvarial osteoblasts, mouse bone marrow-derived MSC, and mouse adipose-derived MSC (Baker et al., 2015; Kirchner et al., 2010; Koskela et al., 2012; Tsukamoto et al., 2004; Watt and Schlezinger, 2015). Thus, TBT may compromise the supportive microenvironment created by osteoblast precursors.

Our previous studies demonstrating susceptibility of B lymphocytes to PPARγ agonist toxicity (Bissonnette et al., 2008; Schlezinger et al., 2004) and the ability of PPARγ agonists to alter BM-MSC physiology prompted testing the hypothesis that TBT may suppress B lymphopoiesis by 2 mechanisms: directly inducing apoptosis in early B cells and, indirectly, by altering the bone marrow microenvironment that supports B lymphopoiesis. Here, we demonstrate that a low concentration of TBT (100 nM) induces an apoptotic pathway that is distinct from the calcium-dependent necrosis/apoptosis induced by a high concentration of TBT (1 μM). BM-MSC cultures support hematopoietic cell differentiation without the addition of exogenous growth factors, including the differentiation of B cells, in an ex vivo model, and ligands which alter the stromal phenotype (TBT, rosiglitazone, and the RXR agonist bexarotene) suppress this phenomenon. Chronic, relatively low-dose TBT exposure in vivo reduced splenic B cells in C57BL/6 mice, which may be related to a reduction of “aging-sensitive” B cells in bone marrow.

MATERIALS AND METHODS

Materials

Rosiglitazone was from Cayman Chemical (Ann Arbor, Michigan). Bexarotene was from LC Laboratories (Woburn, Massachussetts). Human insulin, TBT chloride, and Protease Inhibitor Cocktail for Mammalian Cells were from Sigma-Aldrich (St Louis, Missouri). Plasmocin was from Invivogen (San Diego, California). Fluo-4-AM was from Molecular Probes (Eugene, Oregon). Murine rIL-7 was from Research Diagnostics (Flanders, New Jersey). Antibodies for immunoblotting were purchased from the following: β-actin (Sigma-Aldrich), cleaved caspase-3 (Cell Signaling Technology, Beverley, Massachusetts), cytochrome c (BD Biosciences, Franklin Lakes, New Jersey). Details of antibodies for fluorescence activated cell sorting (FACS) are in Supplementary Tables 1 and 2. All other reagents were from Thermo Fisher Scientific (Suwanee, Georgia) unless noted.

In vivo exposure

All animal studies were reviewed and approved by the Institutional Animal Care and Use Committee at Boston University or The Lady Davis Institute for Medical Research, McGill University. In vivo exposures were conducted using male, C57BL/6J mice (12 weeks of age) (Jackson Laboratories, Bar Harbor, ME). Animals were gavaged 3 times per week for 10 weeks with no substance, sesame oil (10 μl/g) or TBT (10 mg/kg). Mice were weighed prior to each dosing and at euthanasia. At euthanasia, the thymus and spleen were collected, weighed, and strained through a 70 μm cell strainer. Total live cells were determined by counting an aliquot by Trypan Blue exclusion and then were phenotyped by FACS analysis. The right tibia was fixed in 4% paraformaldehyde. The left tibia and femurs were collected, bone marrow was flushed, strained, pelleted, resuspended in freezing medium (FBS with 10% DMSO) and then stored in liquid nitrogen.

Cell culture

All cultures were maintained at 37°C in a humidified, 5% CO2 atmosphere. WEHI-231 cells (CRL-1702, ATCC, Manassas, Virginia) are an immature B lymphoma cell line isolated from (BALB/c × NZB) F1 mice. Stocks of WEHI-231 cells were maintained in DMEM with 5% fetal bovine serum (FBS), plasmocin, L-glutamine and 2-mercaptoethanol. BU-11 cells are a nontransformed, stromal cell-dependent B cell line isolated from C57BL/6J mice that express both CD43 and cytoplasmic Ig heavy chains (ie, a pro/pre B cell model) (Yamaguchi et al., 1997). BU-11 cells were co-cultured on cloned bone marrow-derived stromal cells (BMS2 cells) (Pietrangeli et al., 1988) (kindly provided by Dr P. Kincade, Oklahoma Medical Research Foundation). Stocks of BU-11 cells were maintained on BMS2 cell monolayers in an equal mixture of DMEM and RPMI 1640 medium with 5% FBS, plasmocin, L-glutamine, and 2-mercaptoethanol. For experiments, BU-11 cells were cultured (0.5–1 × 106 cells/ml medium) overnight without BMS2 cells in RPMI with 5% FBS and IL-7 (16 ng/ml). WEHI-231 cells were cultured (0.5 × 106 cells/ml medium) overnight in RPMI with 5% FBS. Cells were treated with Vh (DMSO, 0.1% final concentration) or TBT (1 nM–10 μM) for 15 min–48 h. In the low-dose TBT experiments, etoposide (0.1 μg/ml) was used as a positive control.

Primary B cell cultures were prepared from 4- to 8-week old, male C57BL/6J or B6.129S2-Trp53tm1Tyj/J mice (RRID:IMSR_JAX:002101) (Jackson Laboratories). Bone marrow was flushed from the femurs, and red blood cells were lysed. The remaining cells were cultured for 5–7 d in primary B cell medium (RPMI 1640 containing 10% FBS, penicillin/streptomycin, L-glutamine, 2- ME, and 16 ng/ml murine rIL-7). This procedure results in a B cell culture in which at least 95% of the cells express CD43 and B220. B cells were cultured (4 × 105 cells/ml medium) overnight in RPMI/DMEM with 7.5% FBS and IL-7 (16 ng/ml) and treated with Vh (DMSO, 0.1% final concentration) or TBT (40 nM–10 µM) for 24–48 h.

Primary bone marrow cultures were prepared from 8- to 10-week-old, male C57BL/6J mice (Jackson Laboratories). Bone marrow was prepared as above. Prior to dosing, the medium was changed to include osteogenic additives: ascorbate (12.5 μg/ml), β-glycerol phosphate (8 μM), dexamethasone (10 nM), and insulin (500 ng/ml), except for Naïve wells. Cultures received no treatment (Naïve) or were treated with Vh (DMSO, 0.1% final concentration), rosiglitazone, TBT or bexarotene (10–100 nM). Following treatment, the cells were cultured for 14 days. Medium was aspirated, new medium was added, and the cultures were redosed 5 times over the 14-day period. Given that DMSO is denser than water, the TBT was largely delivered to the cells, which were at the bottoms of the wells; therefore, cumulative doses were as high as 60–600 nM.

To evaluate B cell differentiation in vitro, pro B cells were maintained on a supportive stromal cell feeding layer. OP9 (gift from Dr Tarik Maroy, originally supplied by Dr J.C. Zuniga-Pflucker) stromal cells were maintained in AMEM media (Wisent, St Bruno, Qubec, Canada) supplemented with 20% FBS (HyClone, GE Healthcare Life Sciences, Mississauga, Ontario, Canada), sodium bicarbonate, β-mercaptoethanol and penicillin/streptomycin (Wisent). OP9 cells (4 × 103 cells/well) were seeded in 24-well plates and cultured overnight. Before adding the B cells, the culture medium was replaced with B cell culture medium: Optimem media (Life technologies) supplemented with 10% FBS (HyClone), β-mercaptoethanol and penicillin/streptomycin (Wisent).

Early pro B cells (Hardy fraction B (Hardy et al., 1991), see Supplementary Figure 1) were isolated from total bone marrow harvested from 5-week-old, male C57BL/6 mice as described previously in Kelly et al. (2013). Before cell sorting, CD19+ cells were enriched using magnetic-based column (LS Columns, Miltenyi Biotec, San Diego, California) and magnetic antibody-microbeads (CD19 MicroBeads, mouse, Miltenyl Biotec) following the standard protocol. CD19+ enriched bone marrow cells were blocked with rat antimouse CD16/CD32 Fc receptor block (BD Biosciences), and stained with fluorochrome-conjugated primary antibodies in 1x PBS, supplemented with 5% FBS. Information on the antibodies is provided in Supplementary Table 1. Fraction B cells were sorted directly into the 24-well plate (4 × 103 cells/well) with B cell culture medium on top of a feeding layer of OP9 cells using the FACSAria Fusion Cell Sorter (BD Biosciences) at the Lady Davis Flow Cytometry Core Facility. Two experimental treatment designs were used. First, OP9/B cell cultures were divided into 5 treatment groups: Vh (DMSO), 20, 40, 80, and 100 nM TBT. Second, OP9 stromal cells only were exposed to increasing concentrations of TBT (Vh (DMSO), 20, 40, 80, and 100 nM) for 7 days. The OP9 cultures were trypsinized, washed and replated prior to the addition of the B cells. After TBT exposure, sorted B cells were seeded on top of pretreated OP9 stromal cells. After 5 days of exposure or culture, suspension cells were collected and resuspended in FACS buffer prior to phenotyping.

MTT assay

3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT) labeling was used to quantitate cell death in multiple cell types over a wide range of TBT concentrations. After 42 h of treatment, MTT reagent (10 µl of 10 mg MTT in PBS/ml per well) was added, and the incubation was continued for 6 h. Solublization buffer (20% SDS, 50% DMF, pH 7.4; 100 ul per well) was added, and the cells incubated overnight at 37 °C. Absorbance was determined at 562 nm using a spectrophotometric plate reader.

Apoptosis assays

For analysis of cellular DNA content, cells were harvested into cold PBS containing 5% FBS and 10 µM azide. Cells were resuspended in hypotonic buffer containing 50 μg/ml propidium iodide (PI), 1% sodium citrate, and 0.1% Triton X-100 and analyzed for red fluorescence with FL-2 in the log mode on a FACSCalibur flow cytometer (Becton/Dickinson, Bedford, Massachusetts). The percentage of cells undergoing apoptosis was determined to be those having a weaker PI fluorescence than cells in the G0/G1 phase of the cell cycle. Caspase-3 activity was determined using the Caspase-Glo 3/7 Assay system (Promega, Madison, Wiscosin). Luminescence values in experimental wells were normalized by that measured in naïve wells to determine “Fold Change from Naïve” values.

Immunoblotting

For analysis of cleaved caspase-3 expression, cytoplasmic extracts were prepared as described previously in Schlezinger et al. (2006). For analysis of cytochrome c release, cytoplasmic fractions from digitonin-permeabilized cells were prepared as described previously in Schlezinger et al. (2007) and Waterhouse et al. (2004). Protein concentrations were determined by the Bradford method. Proteins (5–60 μg) were resolved on 15% gels, transferred to a 0.2 μm nitrocellulose membrane, and incubated with primary antibody, anticleaved caspase-3 (9661) or anticytochrome c (S2050). The secondary antibody was horseradish peroxidase-linked goat antirabbit or goat antimouse IgG (Biorad, Hercules, California). Immunoreactive bands were visualized with enhanced chemiluminescence. To control for equal protein loading, blots were reprobed with a mouse monoclonal antibody to β-actin (A5441) and analyzed as earlier. To quantify changes in protein expression, band densities were determined using the UVP Bioimaging System and the Labworks 4 program (UVP, Inc., Upland, California). The band density of the specific protein was divided by the band density of β-actin. β-actin-normalized data are reported as “Relative Expression”.

Free intracellular calcium

Cells were loaded with Fluo-4-AM (1 μM) for 30 min prior to treatment with Vh or TBT for 15 min–4 h. B cells were transferred to FACS tubes without washing and analyzed immediately for green fluorescence (FL-1) on a FACSCalibur flow cytometer.

Phenotyping by flow cytometry

In vitro: B cells from 24-well plates were removed from stromal cell layers by gentle pipetting and washed once with PBS. Cells were then stained with LIVE/DEAD Fixable Dead Cell Stain according to the manufacturer’s instructions, and the pellets were resuspended in FACS buffer with rat antimouse CD16/CD32 Fc receptor block (BD Biosciences). A master mix of stains or isotype controls was made in FACS buffer and added to each well. Information on the antibodies is provided in Supplementary Table 2. Cells were kept on ice and analyzed immediately on an LSR Fortessa Analyser (BD Biosciences) at the Lady Davis Flow Cytometry Core Facility.

In vivo: Cells collected from spleen and bone marrow were resuspended in FACS buffer. Cells were then stained with LIVE/DEAD Fixable Dead Cell Stain according to the manufacturer’s instructions, and the pellets were resuspended in FACS buffer with Mouse BD Fc Block™. A master mix of stains or isotype controls was made in FACS buffer and added to each well. Information on the antibodies is provided in Supplementary Table 3. Cells were kept on ice and analyzed immediately on a BD LSRII Flow Cytometer at the BU Flow Cytometry Core Facility.

Data were compensated and analyzed using FlowJo version 7 (Tree Star, Inc., Ashland, Oregon). Analyses of lineage staining and B cell development are depicted in Supplementary Figures 1 and 2. B cell development was analyzed using the Hardy/Hayakawa (Hardy et al., 1991) and the Miller/Allman methodologies (Miller and Allman, 2003).

Histology

Tibia were decalcified, embedded in paraffin and sectioned for histological analysis. Sections were stained with hematoxylin and eosin. For immunohistochemical analyses, rehydrated sections were incubated with Rodent Decloaker Reagent (Biocare Medical) for antigen retrieval. Sections were incubated with monoclonal rabbit-anti perilipin (9349), followed by incubation with Rabbit on Rodent HRP Polymer (Biocare Medical). To visualize the immune reactions, slides were incubated with 3,3-diaminobenzidine. Hematoxylin was used as a counterstain. Microscopy was carried out on an Olympus BX51 light microscope, and digital photography was carried out with a CoolSnap Pro Cf digital camera.

mRNA expression

Total RNA was extracted from whole humerus, and genomic DNA removed by double TRIzol extractions (Life Technologies, Grand Island, New York). RNA integrity was confirmed by gel electrophoresis. cDNA was prepared from total RNA using the GoScript Reverse Transcription System (Promega), with a 1:1 mixture of random and Oligo (dT)15 primers. All qPCR reactions were performed using the GoTaq qPCR Master Mix System (Promega). Validated primers were purchased from Qiagen (Rn18s: QT02448075, Fabp4: QT00091532, Plin1: QT00150360, Cfd: QT01051890). qPCR reactions (in duplicate) were performed using a 7300 Fast Real-Time PCR System (Applied Biosystems, Carlsbad, California): Hot-Start activation at 95 °C for 2 min, 40 cycles of denaturation (95 °C for 15 s) and annealing/extension (55 °C for 60 s). Relative gene expression was determined using the Pfaffl method (Pfaffl, 2001). The Cq value for 18s ribosomal RNA (Rn18s) was used for normalization. The average Cq value for all control mice was used as the reference point, and the data are reported as “Relative Expression”.

Statistics

Statistical analyses were performed with Prism 6 (GraphPad SoftwareInc., La Jolla, California). For in vivo experiments, individual data are presented with the mean indicated by a line. The Student’s t test was used to determine significant differences between control and TBT-treated animals. For in vitro experiments, data are presented as means ± SE. Student’s t tests, 1-factor ANOVAs, in conjunction with the Dunnett’s or Tukey-Kramer multiple comparisons tests, or 2-factor ANOVAs, in conjunction with the Bonferroni multiple comparisons test, were used to analyze the data and determine significant differences. All analyses were performed at alpha = 0.05.

RESULTS

Low Concentration TBT Induces Apoptosis in Early B Cells

First, we tested whether pro/pre B cells are sensitive to TBT-induced cell death at nM concentrations. BU-11 cells, primary pro B cells, and WEHI-231 cells from were treated with vehicle (DMSO, final concentration 0.1%) or TBT (0.001–10 μM) for 48 h. TBT -induced cell death, as indicated by reduced MTT labeling, in all 3 B cells types (Figure 1A). The EC50 was lowest for BU-11 cells (7*10−8 M), followed by primary pro B cells (2*10−7 M) and WEHI-231 cells (5*10−7 M). Apoptosis was readily detected after exposure of BU-11 cells to a concentration as low as 40 nM TBT, as indicated by the appearance of a population of cells with a subG0/G1 DNA content (Figure 1B). Significant cleavage of caspase-3 to the active, 17 kDa form was evident within 4 h of treatment of BU-11 cells (Figure 1C) and was accompanied by a significant increase in caspase-3 catalytic activity (Figure 1D). The increase in caspase-3 activation at 4 h was evident at a dose of TBT as low as 80 nM (data not shown).

FIG. 1.

FIG. 1

Open in a new tab

Low concentration TBT induces B cell apoptosis. A, BU-11, WEHI-231, and primary pro B cells were treated with Vh (DMSO, 0.1%), or TBT (1 nM–10 μM) for 48 h. Viability was determined by MTT labeling. Data are presented as means ± SE from 3 to 4 independent experiments. ANOVA (Dunnett’s test) was used to determine significance. B–D, BU-11 cells were treated with Vh (DMSO, 0.1%), etoposide (0.1 mg/ml, positive control) or TBT (20–120 nM) for 0.25–8 h. (B) Cells were analyzed for apoptosis by PI staining and FACS. (C) Caspase-3 activity was assessed using the Caspase-Glo 3/7 Assay. (D) Formation of active caspase-3 was analyzed by immunoblotting of cytoplasmic extracts. Data are presented as means ± SE from 3 to 5 independent experiments. Statistically different from Vh (*P < .05, **P < .01, ANOVA, Dunnett’s).

Given the indolent timing of nM TBT-induced apoptosis when compared with that of μM TBT (Bissonnette et al., 2010), we hypothesized that the mechanisms were distinct. In order to differentiate these death pathways, changes in cytoplasmic calcium were investigated. At 1 μM, TBT caused a catastrophic calcium release by 0.25 h (Figure 2A); however, at 100 nM, TBT did not cause a significant increase in cytoplasmic calcium. The slight, nonsignificant increase in cytoplasmic calcium continued to dissipate when followed to 4 h (data not shown). As there was no reason to believe that TBT would bind a death receptor and activate a cell-extrinsic apoptotic pathway, we examined BU-11 cell lysates for evidence of permeabilization of the mitochondria. Indeed, TBT treatment led to significant increase in cytoplasmic cytochrome c (Figure 2B).

FIG. 2.

FIG. 2

Open in a new tab

Low concentration TBT induces a mitochondrial-dependent apoptotic pathway that is distinct from that of high concentration exposure. A and B, BU-11 cells were treated with Vh (DMSO, 0.1%), etoposide (0.1 mg/ml, positive control) or TBT (100 nM–1 µM) for 0.25–8 h. (A) Cytoplasmic Ca2+ was detected by loading BU-11 cells with Fluo4-AM prior to treatment with Vh or TBT for 15 min, followed by flow cytometry. Statistically different from 100 nM (**P < .01, 2-factor ANOVA, Bonferroni). (B) Cytochrome c release was analyzed by immunoblotting of cytoplasmic extracts from digitonin-permeablized cells. Representative immunoblots from 3 independent experiments are presented. C, Primary pro B cells isolated from either Wt or p53 mutant mice were treated Vh (DMSO, 0.1%) or TBT (150 nM) for 24 h, and cells were analyzed for apoptosis by PI staining. The percentage of death measured in naive cell populations was subtracted prior to analysis. Data are presented as means ± SE from 3-4 independent experiments. Statistically different from Wt (**P < .01, 2-factor ANOVA, Bonferroni).

p53 is intimately tied to the process of mitochondrial outer membrane permeabilization. p53 controls the expression of proapoptotic BCL-2 family proteins and acts as a “super BH3-only protein” (Chipuk and Green, 2006). p53 can directly activate pore formation, as well as encumber the antiapoptotic BCL-2 proteins (Wolff et al., 2008). Therefore, we investigated the role of p53 in low-dose TBT-induced B cell death comparing primary pro B cells from wild-type and p53 mutant mice. In contrast to wild-type B cells, TBT did not induce apoptosis in p53 mutant B cells (Figure 2C). Taken together, the data indicate that at concentrations of as low as 40nM, TBT activates a mitochondrial-dependent apoptotic pathway that is mechanistically distinct from high-dose TBT.

TBT Suppresses B Lymphopoiesis in Bone Marrow Co-Cultures

Although the BU-11 studies demonstrated a direct effect of TBT on developing B cells, we moved to ex vivo models of the bone marrow microenvironment in order to investigate the hypothesis that TBT-induced alterations of BM-MSC differentiation negatively impact B lymphopoiesis. Osteoblast-lineage cells have been shown to create at least part of the niche that supports HSCs and hematopoiesis (as reviewed in Wu et al., 2009). Indeed, ex vivo BM-MSC cultures support a population of renewable hematopoietic cells.

Primary bone marrow cultures were estabilished from 8- to 10-week-old male C57BL/6J mice and induced to undergo osteogenic differentiation by culture in MSC medium supplemented with ascorbate, β-glycerol phosphate, and dexamethasone. After 14 days in culture, 4 times the number of suspension cells were present relative to undifferentiated cultures, and this population included cells that expressed CD11b, Gr-1, or B220, suggesting the presence of monocytes, granulocytes, and B cells (Figure 3A).

FIG. 3.

FIG. 3

Open in a new tab

BM-MSCs support hematopoietic cells in vitro and TBT suppresses this population. A and B, Primary bone marrow cultures. Bone marrow was isolated from 8- to 10-week-old, male C57BL/6 mice, plated, and allowed to adhere for 7 days. The medium was changed to include dexamethasone, ascorbate, and β-glycerol phosphate, except for Naïve wells. Cultures were treated as indicated. The medium was replaced and the cultures redosed every 2–3 days for a total of 6 dosings. After 14 days, the medium was collected, and suspension cells were analyzed. Data are presented as means ± SE from 3 to 5 independent experiments. (A) Flow cytometry analysis of B220+ cells. Statistically different from Vh-treated (*P < .05, Student t test). (B) Cell counts. Statistically different from Vh-treated (*P < .05, **P < .01, ANOVA, Dunnett’s). C and D, OP9/Developing B cell cultures. Bone marrow was isolated from 5-week-old C57BL/6 mice. CD19+ B cells were enriched from primary bone marrow by magnetic bead separation. Fraction B (IgM−, cKit+, B220+) B cells were sorted from enriched CD19+ B cells, seeded onto a OP9 stromal cell layer and cultured in vitro. The percentage and number of cells in each B cell fraction were calculated from OP9/Developing B-cell cultures that were either co-treated with Vh (DMSO, 0.1%) or TBT (20–100 nM) for 5 days (C) or the OP9 stromal cells were pretreated with Vh or TBT (20 - 100 nM) for 7 days then B cells were cultured on the OP9 feeder layer for an additional 5 days in the absence of TBT (D) was determined by flow cytometry using the Hardy B-cell antibody staining schematic (see Supplementary Figure 1). Data are presented as means ± SE from 3 independent experiments. Statistically different from Vh-treated (*P < .05, **P < .01, ***P < .001, ANOVA, Dunnett’s).

Next, we tested the effect of treatment with TBT. At the initiation of osteogenic differentiation, cultures were treated with Vh (DMSO, 0.1% final concentration) or TBT. Because TBT is a dual PPARγ/RXRαβ ligand, we compared TBT’s effects to rosiglitazone (PPARγ ligand) and bexarotene (RXR ligand). We have shown previously that rosiglitazone and bexarotene induce adipocyte differentiation in BM-MSCs similar to TBT (Baker et al., 2015; Watt and Schlezinger, 2015; Yanik et al., 2011). In addition, retinoids are known to influence B lymphocyte differentiation directly (Chen et al., 2008). Treatment with all 3 ligands significantly reduced the suspension cell population (Figure 3B). Treatment with bexarotene (100 nM, dosed 6 times) or TBT (10, 50 nM, dosed 6 times) reduced the suspension population to such a degree that phenotyping by flow cytometry was not possible. The reduction seen in the rosiglitazone-treated cultures was at least in part due to a reduction in B cells as demonstrated by the loss of the B220+ population in the rosiglitazone-treated wells (50 nM, dosed 6 times) (Figure 3A). These data suggest that TBT suppressed B lymphopoiesis in this ex vivo model.

To investigate the effect of TBT on B cell development specifically, we moved to an OP9/primary B cell co-culture model in which primary pro B cells (IgM−, cKit+, B220+, from 5-week-old, male C57BL/6 mice) were co-cultured with an OP9 stromal cell feeder layer that supports lymphopoiesis. This model allows for the monitoring of B cell differentiation (Hardy Fractions B-E) in vitro. Hardy Fraction B, early pro B cells were sorted onto an OP9 stromal layer and allowed to differentiate in vitro in the presence of increasing concentrations of TBT (20–100 nM) for 5 days. When the OP9 cells and B cells were co-treated with TBT, there was a dose-dependent decrease in the percentage of cells in the late pro B and pre B cell fractions (Hardy Fraction C/C’; 60 ± 2 vs 31 ± 4 and Fraction D; 19 ± 3 vs 9 ± 1% for Vh and 100 nM TBT, respectively) accompanied by an increase in the percentage of cells in the early pro B fraction (Hardy Fraction B; 18 ± 2 vs 58 ± 6% for Vh and 100 nM TBT, respectively) (Figure 3C). However, there was also a dose dependent decrease in the total number of B cells present in the cultures with increasing concentrations of TBT (significantly decreased by 56.0 ± 13.2% (40 nM), 78.2 ± 5.0% (80 nM) and 89.2 ± 0.1% (100 nM), ANOVA, Dunnett’s), which is reflected in all of the B cell fractions (Figure 3).

To test the effect of TBT on the B cell supportive microenvironment, OP9 cells were treated with TBT for 7 days, washed, trypsinized, and replated to insure that TBT was not transferred to the B cells. When B cells were seeded on OP9 cells that had been pretreated with Vh or TBT for 7 days, there was a significant increase in early pro B cells (Hardy Fraction B) in cultures with OP9 cells treated with 100 nM TBT (Figure 3D) and also a trend toward a decreased percentage of cells in the late pro B and pre B cell fractions (Hardy Fraction C/C’; 34 ± 4 vs 25 ± 5 and Fraction D; 25 ± 5 vs 17 ± 3% for Vh and 100 nM TBT, respectively). In addition, TBT pretreatment of the OP9 cells did not reduce overall B cell numbers (Figure 3D). These results suggest that the microenvironment that supports lymphopoiesis created by the OP9 cells is impaired by treatment with TBT. Indeed, this loss of support of lymphopoiesis was accompanied by the differentiation of the OP9 cells into adipocytes (Supplementary Figure 3). However, it cannot be ruled out that TBT also has direct effects on the B cells that are affecting lymphopoiesis.

Low-Dose TBT Exposure Reduces Peripheral B Cells

Previous studies of TBT-induced immunotoxicity used short-term high dose exposure schemes that resulted in systemic toxicity and thymic atrophy (Bressa et al., 1991; Vos et al., 1984). Here, we tested the hypothesis that long term, low dose exposure to TBT would result in significant suppression of lymphopoiesis in the absence of overt toxicity. Twelve-week-old, male, C57BL/6J mice were gavaged 3 times/week for 10 weeks with no substance, sesame oil (Vehicle, Vh) or TBT in sesame oil (10 mg/kg). Body weight loss and thymic atrophy were assessed as signs of toxicity. There was no difference in weight gain or liver/body weight ratios between treatment groups in male mice (Supplementary Figure 4). Thymic atrophy, as assessed by relative thymus weight and total thymocyte counts, also was not detected (Supplementary Figure 5).

In order to determine the effect of TBT on peripheral white blood cells, we examined myeloid and lymphoid populations in the spleen. Spleen/body weight and total splenocyte number were impacted minimally by treatment with TBT (Figs. 4A and B). Myeloid cell populations in the spleen, including granulocytes (Gr1+), macrophages (CD11b+), and regulatory myeloid cells (CD11b+Gr1+) were unaffected by TBT treatment (Figure 4C). Treatment with TBT also did not alter splenic T cell populations (total T cells, CD4+, CD8+, and the CD4/CD8 ratio) (Figure 4D). On the other hand, splenic B cells, as defined by B220 staining, were significantly reduced by TBT treatment (Figure 5). These data suggest that in vivo, long-term low dose exposure to TBT results in specific loss of peripheral B cells.

FIG. 4.

FIG. 4

Open in a new tab

In vivo TBT exposures does not alter total splenocyte, myeloid cell or T cell numbers. Twelve-week-old, male, C57BL/6 mice were gavaged 3 times/week for 10 weeks with no substance or sesame oil (Vh), rosiglitazone (25 mg/kg), or TBT (10 mg/kg). Spleens were dissected for analysis. A, Spleen/body weights. B, Total cell counts. Phenotyping by FACS analysis for (C) myeloid cells and (D) T cells. Individual data are presented with the average indicated by a line (n = 8–11). TBT was not statistically different from Vh (Student’s t Test).

FIG. 5.

FIG. 5

Open in a new tab

In vivo TBT suppresses peripheral B cell numbers. Twelve-week-old, male, C57BL/6 mice were treated as described in Figure 4. Spleens were removed, total cells counted, and then phenotyped by FACS analysis for the B cell marker B220. Individual data are presented with the average indicated by a line (n = 8–11). *Statistically different from Vh-treated (P < .05, Student’s t Test).

Early Bone Marrow B Cells Are Compromised by Low-Dose TBT Exposure InVivo

Given that peripheral B cells were suppressed in this low-dose, long-term exposure to TBT, we sought to determine if this may have been due to a reduced supply of maturing B cells from the bone marrow. Since TBT treatment can alter the stromal elements of the bone marrow microenvironment, we hypothesized that stromal-dependent stages of B cell development would be most evidently impacted.

In order to assess the bone marrow adipocyte content, histological sections of tibia were stained with hematoxylin and eosin (Figure 6A). TBT-treated mice had a greater number of adipocytes in bone marrow than Vh-treated mice, which was confirmed by immunohistological staining for perilipin (Figure 6B). Further, gene expression analysis of whole humerus revealed a significant increase in mRNA expression of the PPARγ target genes Fabp4, Plin1, and Cfd (Figure 6C), similar to TBT-induced effects on BM-MSC gene expression in vitro (Baker et al., 2015; Watt and Schlezinger, 2015).

FIG. 6.

FIG. 6

Open in a new tab

TBT increases adipocyte number and PPARγ target gene expression in bone marrow. Twelve-week-old, male, C57BL/6 mice were treated as described in Figure 4. A, Histological analyses are a panoramic view from stitched (200×) micrographic images for representative histological sections depicting epiphyseal through mid diaphyseal regions of the tibia. B, Immunohistochemical analysis of perilipin expression. Two immunoreactive adipocytes per section are indicated by the arrows. TB, trabecular bone; BV, blood vessel. C, Expression of adipocyte-related genes was assessed in whole humerus bone gene expression using RT-qPCR. Data from individual mice are presented with the mean indicated by a line (n = 8–11). Statistically different from Vh-treated (*P < .05, **P < .01 Student’s t test).

B cell subpopulations in bone marrow from tibia and femurs were analyzed by flow cytometry. Total bone marrow B cells were not significantly altered by treatment with TBT (Figure 7A). As bone marrow B cells mature, expression of B220 increases, resulting in B220L° and B220Hi subpopulations (Hardy et al., 1991). Although there was no significant change in the B220Hi subpopulation with treatment, the B220L° subpopulation decreased with TBT treatment (Figure 7B). TBT did not alter the number of pre-pro B cells (Hardy Fraction A; Figure 7C). Pro B cells (Hardy Fractions B and C) and pre B cells (Hardy Fraction D) showed trends towards suppression with treatment (Figs. 7C–E). Immature/mature B cells (Hardy Fraction E/F) were unchanged (Figure 7F). Although the bone marrow data are all presented as percentages of live cells, we confirmed that the same relationship in the B cell compartment is maintained when converted to approximate cell counts (using the % of live gate x total live cells from flushing left tibia, data not shown). Given that early B cells (B220L°) were significantly reduced (Figure 7B), a second staining method was used to delineate the subpopulation of B cells lost following TBT treatment. The “aging sensitive” pro B cell (B220+, CD43+, CD19+, AA4+) population defined by Miller and Allman (2003) was significantly reduced with TBT treatment (Figure 7G). Even though subpopulations of early B cells showed only trends towards suppression by treatment with TBT, as a whole, low-dose TBT treatment significantly suppressed early bone marrow B cells in vivo.

FIG. 7.

FIG. 7

Open in a new tab

In vivo TBT suppresses early B cells in the bone marrow. Twelve-week-old, male, C57BL/6 mice were treated as described in Figure 4. Bone marrow from femur and tibia was flushed and FACS analysis of B cells was carried out according to Supplementary Figures 1 and 2. Individual data are presented with the average indicated by a line (n = 8–11). *Statistically different from Vh-treated (P < .05, Student’s t test).

DISCUSSION

TBT is a well-known immunotoxicant because of its ability to cause thymic atrophy at high, acute exposures (Bressa et al., 1991; Vos et al., 1984). It is becoming increasingly clear that B cells are also a target of TBT’s toxic action, perhaps at exposure levels significantly below those required to compromise T cells (Carfi et al., 2010; De Santiago and Aguilar-Santelises, 1999). Here, we demonstrate that TBT directly induces apoptosis in a developing B cell line at environmentally relevant concentrations in vitro. In an ex vivo model, we demonstrate that TBT suppresses developing B cells likely through both direct mechanisms and by altering the microenvironment. Finally, we show, for the first time, that subtoxic, long-term exposure to TBT suppresses B cells in vivo.

At concentrations likely above what are relevant to human environmental exposures (Antizar-Ladislao, 2008), TBT (0.5–10 μM) is known to cause apoptosis at least in part by altering intracellular calcium dynamics in multiple cell types including a T lymphocyte cell line, thymocytes, and in B cells (Bissonnette et al., 2010; Gennari et al., 2000; Grundler et al., 2001; Stridh et al., 2001). At 1 μM, TBT caused necroapoptosis that followed a catastrophic release of calcium into the cytoplasm (Bissonnette et al., 2010). The apoptosis was dependent on a CaMKII-initiated phosphorylation cascade leading to release of cytochrome c from the mitochondria and caspase-3 activation. This mechanism of cell death is unlikely to be relevant to human toxicity, however.

Given that developing B cells have been shown to be particularly sensitive to exogenous chemical insults (Griffiths et al., 1994; Holladay and Smith, 1995; Schlezinger et al., 2004, 2007; Teague et al., 2010), we hypothesized that even environmentally relevant concentrations of TBT may likewise have a deleterious effect. Indeed, concentrations of TBT as low as 40 nM induced early B cell apoptosis. The data corroborate the observation that human developing B cells and human tonsillar B cells are highly sensitive to nM concentrations of TBT (Carfi et al., 2010; De Santiago and Aguilar-Santelises, 1999). Additionally, in a neuronal cell line, TBT induces apoptosis via distinct mechanisms depending on the concentration of exposure (500 nM vs 2 μM) (Nakatsu et al., 2007); thus, we also hypothesized that the apoptosis induced by 100 nM TBT may be distinct from that of 1 μM. Low-dose TBT (nM) activated calcium-independent, mitochondrially driven apoptosis that took an indolent rather than rapid course. The direct mechanism of activation of apoptosis is yet to be investigated; however, preliminary data suggest that p53 is an essential element in the apoptotic pathway.

Cells of the osteoblast lineage facilitate B lineage commitment and differentiation (Wu et al., 2009; Zhu et al., 2007). Osteoblast expression of Wnt proteins, osteoblast sclerostin signaling, and osteoprogenitor parathyroid hormone signaling all have been implicated in maintaining lymphopoiesis (Cain et al., 2012; Cao et al., 2015; Panaroni et al., 2015; Zhu et al., 2007). In contrast to the supportive nature of osteolineage cells, adipocytes release soluble factors that inhibit lymphopoiesis (Bilwani and Knight, 2012), which may include adiponectin and IL1 that favor myelopoiesis at the expense of lymphopoiesis (Kennedy and Knight, 2015; Yokota et al., 2003). Multipotent progenitors which differentiate into B cells are reduced 2- to 3-fold in adipocyte-rich skeletal regions when compared with adipocyte-scarce regions (Naveiras et al., 2009). Active negative regulation was indicated by the fact that the progenitors were rescued when adipocytes were either genetically or chemically suppressed (Naveiras et al., 2009).

When primary BM-MSC cultures are treated with rosiglitazone, bexarotene and TBT (nM) osteogenesis is suppressed and adipogenesis is promoted (Baker et al., 2015; Watt and Schlezinger, 2015). Here, we show that rosiglitazone (300 nM total applied dose, PPARγ agonist), bexarotene (600 nM total applied dose, RXR agonist) and TBT (60 nM total applied dose, dual PPARγ/RXR agonist) all significantly reduced hematopoeisis in bone marrow cultures. Further, we show that TBT suppressed OP9-supported lymphopoiesis. This likely occurred through induction of apoptosis when the B cells were co-cultured with the OP9 cells during treatment with TBT. The reduction of hematopoietic cells following repeated dosing of bone marrow cultures with 10 nM TBT was consistent with the reduction of total B cell numbers with a single dose of 80 nM TBT in the B cell/OP9 cultures. In co-cultures in which only the OP9 cells were treated with TBT, early pro B cells (Hardy Fraction B) accumulated suggesting that TBT interfered with the supportive microenvironment generated by the OP9 cells, in the absence of toxicity. Indeed, TBT induced the differentation of the OP9 cells into adipocytes. Interestingly, exposure of human long-term bone marrow cultures to TBT also results in formation of adipocytes and a concurrent reduction of leptin and IL-7 expression (Carfi et al., 2008).

No previous studies have examined the effect of TBT on bone marrow physiology in vivo; therefore this study is the first attempt at developing a model to study lymphopoiesis during long-term, low-dose exposure to TBT. It is not, as of yet, possible to define the exposure model presented here as “environmentally relevant” as we did not assess serum levels of TBT, nor has any other study attempted to do so in such a long-term exposure. A study of rats exposed to TBT showed that exposure to a dose of 2.5 mg/kg/d for 3 weeks resulted in a liver concentration in the range measured in humans from a fish eating population (100 ng/g liver) (Cooke et al., 2008; Takahashi et al., 1999). The exposure presented here equates to approximately 4 mg/kg/d. Therefore, using an exposure scenario that is of similar magnitude to exposure in a highly exposed human population, we show that lymphopoiesis is a target.

In this study, we demonstrate that peripheral B cells are suppressed by TBT. Given that TBT exposure also increased marrow adipogenesis and activated PPARγ in bone, we hypothesized that the peripheral reduction of B cells resulted from suppressed lymphopoiesis, and potentially alteration of the HSC niche as well. Although the present exposure scheme did not alter the proportion of HSCs (data not shown), early bone marrow B cells were significantly reduced, as we observed in vitro. Alternatively, the reduction in splenic B cells could result from a failure to recruit transitional B cells to the spleen due to changes in the splenic environment or direct effect on the transitional B cells in the process of migrating to the spleen (Wols et al., 2010).

Interestingly, in the bone marrow the strongest reduction occurred in the B220+, CD43+,AA4.1+,CD19+ pro B cells, the population most affected during aging (Miller and Allman, 2003). This early stage of B cell development is dependent up IL7 signaling, and defects in IL7 production by stromal elements and in IL7 responsiveness by pro B cells contribute to reduced B lymphopoiesis with age (Stephan et al., 1997, 1998). In fact, it is the AA4.1+ fraction of the B220+, CD43+ cells that expand robustly in the presence of IL7 (Miller and Allman, 2003). The early bone marrow B cell population also was a significant target in vitro, with TBT causing an apparent stall of differentiation at this stage. Because B lymphopoiesis thrives in an osteoblastic lineage-rich, adipocyte-poor microenvironment, it is interesting, but not surprising that immunosenescence occurs during aging (as reviewed in Dunn-Walters, 2016) and parallels an increase in adipogenesis in bone marrow (Rosen and Bouxsein, 2006) and osteoporosis (Chan and Duque, 2002). The combination of increased marrow adiposity and a potential suppression of aging-sensitive early B cells, suggests that in vivo exposure to TBT causes deleterious changes reminiscent of aging.

In summary, we show that TBT causes multifaceted effects on B cell differentiation. These effects may be due, in part, to induction of adipogenesis in BM-MSCs and subsequent alteration of the bone marrow microenvironment. Bone marrow adipogenesis also is a feature of human aging. Aging is associated with a decrease in both cellular and humoral immune responses, leading to an increased risk of more frequent and severe infectious diseases in the elderly, which may in part result from changes in the bone marrow microenvironment with age (as reviewed in Dunn-Walters, 2016; Pritz et al., 2014). By 2035, 1 in 5 Americans will be aged 65 or older, making immunosenescence an increasingly prevalent public health issue (Howden and Meyer, 2011). Environmental exposure to TBT has the potential to exacerbate these changes both by altering bone marrow physiology and by directly inducing apoptosis in developing B cells. Thus, exposure to this environmental contaminant may intensify age-related problems of the immune systems.

SUPPLEMENTARY DATA

Supplementary data are available at Toxicological Sciences online.

FUNDING

This work was supported by the National Institute of Environmental Health Sciences Superfund Research Program (P42ES007381), the Boston University Flow Cytomery Core, the Natural Sciences and Engineering Research Council of Canada (RGPIN-2015-04919), and the Lady Davis Institute for Medical Research Flow Cytometry Core.

Supplementary Material

Supplementary Data
Click here for additional data file. (645.2KB, pdf)

REFERENCES

  1. Antizar-Ladislao B. (2008). Environmental levels, toxicity and human exposure to tributyltin (TBT)-contaminated marine environment. A review. Environ. Int. 34, 292–308. [DOI] [PubMed] [Google Scholar]
  2. Baken K. A., Arkusz J., Pennings J. L., Vandebriel R. J., van Loveren H. (2007). In vitro immunotoxicity of bis(tri-n-butyltin)oxide (TBTO) studied by toxicogenomics. Toxicology 237, 35–48. [DOI] [PubMed] [Google Scholar]
  3. Baker A. H., Watt J., Huang C. K., Gerstenfeld L. C., Schlezinger J. J. (2015). Tributyltin engages multiple nuclear receptor pathways and suppresses osteogenesis in bone marrow multipotent stromal cells. Chem. Res. Toxicol. 28, 1156–1166. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Bianco P. (2011). Bone and the hematopoietic niche: A tale of two stem cells. Blood 117, 5281–5288. [DOI] [PubMed] [Google Scholar]
  5. Bilwani F. A., Knight K. L. (2012). Adipocyte-derived soluble factor(s) inhibits early stages of B lymphopoiesis. J. Immunol. 189, 4379–4386. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Bissonnette S. L., Haas A., Mann K. K., Schlezinger J. J. (2010). The role of CaMKII in calcium-activated death pathways in bone marrow B cells. Toxicol. Sci. 118, 108–118. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Bissonnette S. L., Teague J. E., Sherr D. H., Schlezinger J. J. (2008). An endogenous prostaglandin enhances environmental phthalate-induced apoptosis in bone marrow B cells: Activation of distinct but overlapping pathways. J. Immunol. 181, 1728–1736. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Bressa G., Hinton R. H., Price S. C., Isbir M., Ahmed R. S., Grasso P. (1991). Immunotoxicity of tri-n-butyltin oxide (TBTO) and tri-n-butyltin chloride (TBTC) in the rat. J. Appl. Toxicol. 11, 397–402. [DOI] [PubMed] [Google Scholar]
  9. Cain C. J., Rueda R., McLelland B., Collette N. M., Loots G. G., Manilay J. O. (2012). Absence of sclerostin adversely affects B-cell survival. J. Bone Miner. Res. 27, 1451–1461. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Calvi L. M., Adams G. B., Weibrecht K. W., Weber J. M., Olson D. P., Knight M. C., Martin R. P., Schipani E., Divieti P., Bringhurst F. R., et al. (2003). Osteoblastic cells regulate the haematopoietic stem cell niche. Nature 425, 841–846. [DOI] [PubMed] [Google Scholar]
  11. Cao J., Zhang L., Wan Y., Li H., Zhou R., Ding H., Liu Y., Yao Z., Guo X. (2015). Ablation of Wntless in endosteal niches impairs lymphopoiesis rather than HSCs maintenance. Eur. J. Immunol. 45, 2650–2660. [DOI] [PubMed] [Google Scholar]
  12. Carfi M., Bowe G., Pieters R., Gribaldo L. (2010). Selective inhibition of B lymphocytes in TBTC-treated human bone marrow long-term culture. Toxicology 276, 33–40. [DOI] [PubMed] [Google Scholar]
  13. Carfi M., Croera C., Ferrario D., Campi V., Bowe G., Pieters R., Gribaldo L. (2008). TBTC induces adipocyte differentiation in human bone marrow long term culture. Toxicology 249, 11–18. [DOI] [PubMed] [Google Scholar]
  14. Chan G. K., Duque G. (2002). Age-related bone loss: Old bone, new facts. Gerontology 48, 62–71. [DOI] [PubMed] [Google Scholar]
  15. Chen X., Esplin B. L., Garrett K. P., Welner R. S., Webb C. F., Kincade P. W. (2008). Retinoids accelerate B lineage lymphoid differentiation. J. Immunol. 180, 138–145. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Chipuk J. E., Green D. R. (2006). Dissecting p53-dependent apoptosis. Cell Death Differ. 13, 994–1002. [DOI] [PubMed] [Google Scholar]
  17. Cooke G. M., Forsyth D. S., Bondy G. S., Tachon R., Tague B., Coady L. (2008). Organotin speciation and tissue distribution in rat dams, fetuses, and neonates following oral administration of tributyltin chloride. J. Toxicol. Environ. Health 71, 384–395. [DOI] [PubMed] [Google Scholar]
  18. Cooke G. M., Tryphonas H., Pulido O., Caldwell D., Bondy G. S., Forsyth D. (2004). Oral (gavage), in utero and postnatal exposure of Sprague-Dawley rats to low doses of tributyltin chloride. Part 1: Toxicology, histopathology and clinical chemistry. Food Chem. Toxicol. 42, 211–220. [DOI] [PubMed] [Google Scholar]
  19. Cornelissen G., Pettersen A., Nesse E., Eek E., Helland A., Breedveld G. D. (2008). The contribution of urban runoff to organic contaminant levels in harbour sediments near two Norwegian cities. Mar. Pollut. Bull. 56, 565–573. [DOI] [PubMed] [Google Scholar]
  20. De Santiago A., Aguilar-Santelises M. (1999). Organotin compounds decrease in vitro survival, proliferation and differentiation of normal human B lymphocytes. Hum. Exp. Toxicol. 18, 619–624. [DOI] [PubMed] [Google Scholar]
  21. Dunn-Walters D. K. (2016). The ageing human B cell repertoire: A failure of selection?. Clin. Exp. Immunol. 183, 50–56. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Fromme H., Mattulat A., Lahrz T., Ruden H. (2005). Occurrence of organotin compounds in house dust in Berlin (Germany). Chemosphere 58, 1377–1383. [DOI] [PubMed] [Google Scholar]
  23. Gennari A., Viviani B., Galli C. L., Marinovich M., Pieters R., Corsini E. (2000). Organotins induce apoptosis by disturbance of [Ca(2+)](i) and mitochondrial activity, causing oxidative stress and activation of caspases in rat thymocytes. Toxicol. Appl. Pharmacol. 169, 185–190. [DOI] [PubMed] [Google Scholar]
  24. Griffiths S. D., Goodhead D. T., Marsden S. J., Wright E. G., Krajewski S., Reed J. C., Korsmeyer S. J., Greaves M. (1994). Interleukin 7-dependent B lymphocyte precursor cells are ultrasensitive to apoptosis. J. Exp. Med. 179, 1789–1797. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Grun F., Watanabe H., Zamanian Z., Maeda L., Arima K., Cubacha R., Gardiner D. M., Kanno J., Iguchi T., Blumberg B. (2006). Endocrine-disrupting organotin compounds are potent inducers of adipogenesis in vertebrates. Mol. Endocrinol. 20, 2141–2155. [DOI] [PubMed] [Google Scholar]
  26. Grundler W., Dirscherl P., Beisker W., Marx K., Stampfl A., Maier K., Zimmermann I., Nusse M. (2001). Early functional apoptotic responses of thymocytes induced by Tri-n-butyltin. Cytometry 44, 45–56. [PubMed] [Google Scholar]
  27. Hardy R. R., Carmack C. E., Shinton S. A., Kemp J. D., Hayakaya K. (1991). Resolution and characterization of pro-B and pre-pro-B cell stages in normal mouse bone marrow. J. Exp. Med. 173, 1213–1225. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Holladay S. D., Smith B. J. (1995). Benzo[a]pyrene-induced alterations in total immune cell number and cell-surface antigen expression in the thymus, spleen and bone marrow of B6C3F1 mice. Vet. Hum. Toxicol. 37, 99–104. [PubMed] [Google Scholar]
  29. Howden L. M., Meyer J. A. (2011). Age and Sex Composition: 2010. In (Bureau U. S. C., Ed.). U.S. Census Bureau, Washington, DC. [Google Scholar]
  30. Kannan K., Takahashi S., Fujiwara N., Mizukawa H., Tanabe S. (2010). Organotin compounds, including butyltins and octyltins, in house dust from Albany, New York, USA. Arch. Environ. Contam. Toxicol. 58, 901–907. [DOI] [PubMed] [Google Scholar]
  31. Kelly A. D., Lemaire M., Young Y. K., Eustache J. H., Guilbert C., Molina M. F., Mann K. K. (2013). In vivo tungsten exposure alters B-cell development and increases DNA damage in murine bone marrow. Toxicol. Sci. 131, 434–446. [DOI] [PubMed] [Google Scholar]
  32. Kennedy D. E., Knight K. L. (2015). Inhibition of B lymphopoiesis by adipocytes and IL-1-producing myeloid-derived suppressor cells. J. Immunol. 195, 2666–2674. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Kirchner S., Kieu T., Chow C., Casey S., Blumberg B. (2010). Prenatal exposure to the environmental obesogen tributyltin predisposes multipotent stem cells to become adipocytes. Mol. Endocrinol. 24, 526–539. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Koskela A., Viluksela M., Keinanen M., Tuukkanen J., Korkalainen M. (2012). Synergistic effects of tributyltin and 2,3,7,8-tetrachlorodibenzo-p-dioxin on differentiating osteoblasts and osteoclasts. Toxicol. Appl. Pharmacol. 263, 210–217. [DOI] [PubMed] [Google Scholar]
  35. le Maire A., Grimaldi M., Roecklin D., Dagnino S., Vivat-Hannah V., Balaguer P., Bourguet W. (2009). Activation of RXR-PPAR heterodimers by organotin environmental endocrine disruptors. EMBO Rep. 10, 367–373. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Levine K. E., Young D. J., Afton S. E., Harrington J. M., Essader A. S., Weber F. X., Fernando R. A., Thayer K., Hatch E. E., Robinson V. G., et al. (2015). Development, validation, and application of an ultra-performance liquid chromatography-sector field inductively coupled plasma mass spectrometry method for simultaneous determination of six organotin compounds in human serum. Talanta 140, 115–121. [DOI] [PubMed] [Google Scholar]
  37. Miller J. P., Allman D. (2003). The decline in B lymphopoiesis in aged mice reflects loss of very early B-lineage precursors. J. Immunol. 171, 2326–2330. [DOI] [PubMed] [Google Scholar]
  38. Nakatsu Y., Kotake Y., Ohta S. (2007). Concentration dependence of the mechanisms of tributyltin-induced apoptosis. Toxicol. Sci. 97, 438–447. [DOI] [PubMed] [Google Scholar]
  39. Naveiras O., Nardi V., Wenzel P. L., Hauschka P. V., Fahey F., Daley G. Q. (2009). Bone-marrow adipocytes as negative regulators of the haematopoietic microenvironment. Nature 460, 259–263. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Panaroni C., Fulzele K., Saini V., Chubb R., Pajevic P. D., Wu J. Y. (2015). PTH Signaling in Osteoprogenitors Is Essential for B-Lymphocyte Differentiation and Mobilization. J. Bone Miner. Res. 30, 2273–2286. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Pfaffl M. W. (2001). A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res. 29, e45. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Pietrangeli C., Hayashi S.-I., Kincade P. (1988). Stromal cell lines which support lymphocyte growth: Characterization, sensitivity to radiation and responsiveness to growth factors. Eur. J. Immunol. 18, 863–872. [DOI] [PubMed] [Google Scholar]
  43. Pritz T., Weinberger B., Grubeck-Loebenstein B. (2014). The aging bone marrow and its impact on immune responses in old age. Immunol. Lett. 162(1 Pt B), 310–315. [DOI] [PubMed] [Google Scholar]
  44. Raffray M., Cohen G. M. (1993). Thymocyte apoptosis as a mechanism for tributyltin-induced thymic atrophy in vivo. Arch. Toxicol. 67, 231–236. [DOI] [PubMed] [Google Scholar]
  45. Rosen C. J., Bouxsein M. L. (2006). Mechanisms of disease: Is osteoporosis the obesity of bone?. Nat. Clin. Pract. Rheumatol. 2, 35–43. [DOI] [PubMed] [Google Scholar]
  46. Schlezinger J. J., Emberley J. K., Bissonnette S. L., Sherr D. H. (2007). An L-tyrosine derivative and PPARg agonist, GW7845, activates a multifaceted caspase cascade in bone marrow B cells. Toxicol. Sci. 98, 125–136. [DOI] [PubMed] [Google Scholar]
  47. Schlezinger J. J., Emberley J. K., Sherr D. H. (2006). Activation of multiple mitogen-activated protein kinases in pro/pre-B cells by GW7845, a peroxisome proliferator-activated receptor gamma agonist, and their contribution to GW7845-induced apoptosis. Toxicol. Sci. 92, 433–444. [DOI] [PubMed] [Google Scholar]
  48. Schlezinger J. J., Howard G. J., Hurst C. H., Emberley J. K., Waxman D. J., Webster T., Sherr D. H. (2004). Environmental and endogenous peroxisome proliferator-activated receptor gamma agonists induce bone marrow B cell growth arrest and apoptosis: Interactions between mono(2-ethylhexyl)phthalate, 9-cis-retinoic acid, and 15-deoxy-Delta12,14-prostaglandin J2. J. Immunol. 173, 3165–3177. [DOI] [PubMed] [Google Scholar]
  49. Stephan R. P., Lill-Elghanian D. A., Witte P. L. (1997). Development of B cells in aged mice: Decline in the ability of pro-B cells to respond to IL-7 but not to other growth factors. J. Immunol. 158, 1598–1609. [PubMed] [Google Scholar]
  50. Stephan R. P., Reilly C. R., Witte P. L. (1998). Impaired ability of bone marrow stromal cells to support B-lymphopoiesis with age. Blood 91, 75–88. [PubMed] [Google Scholar]
  51. Stridh H., Cotgreave I., Muller M., Orrenius S., Gigliotti D. (2001). Organotin-induced caspase activation and apoptosis in human peripheral blood lymphocytes. Chem. Res. Toxicol. 14, 791–798. [DOI] [PubMed] [Google Scholar]
  52. Takahashi S., Mukai H., Tanabe S., Sakayama K., Miyazaki T., Masuno H. (1999). Butyltin residues in livers of humans and wild terrestrial mammals and in plastic products. Environ. Pollut. 106, 213–218. [DOI] [PubMed] [Google Scholar]
  53. Teague J. E., Ryu H. Y., Kirber M., Sherr D. H., Schlezinger J. J. (2010). Proximal events in 7,12-dimethylbenz[a]anthracene-induced, stromal cell-dependent bone marrow B cell apoptosis: Stromal cell-B cell communication and apoptosis signaling. J. Immunol. 185, 3369–3378. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Tomiyama K., Yamaguchi A., Kuriyama T., Arakawa Y. (2009). Analysis of mechanisms of cell death of T-lymphocytes induced by organotin agents. J. Immunotoxicol. 6, 184–193. [DOI] [PubMed] [Google Scholar]
  55. Tsukamoto Y., Ishihara Y., Miyagawa-Tomita S., Hagiwara H. (2004). Inhibition of ossification in vivo and differentiation of osteoblasts in vitro by tributyltin. Biochem. Pharmacol. 68, 739–746. [DOI] [PubMed] [Google Scholar]
  56. Visnjic D., Kalajzic Z., Rowe D. W., Katavic V., Lorenzo J., Aguila H. L. (2004). Hematopoiesis is severely altered in mice with an induced osteoblast deficiency. Blood 103, 3258–3264. [DOI] [PubMed] [Google Scholar]
  57. Vos J. G., de Klerk A., Krajnc E. I., Kruizinga W., van Ommen B., Rozing J. (1984). Toxicity of bis(tri-n-butyltin)oxide in the rat. II. Suppression of thymus-dependent immune responses and of parameters of nonspecific resistance after short-term exposure. Toxicol. Appl. Pharmacol. 75, 387–408. [DOI] [PubMed] [Google Scholar]
  58. Waterhouse N. J., Steel R., Kluck R., Trapani J. A. (2004). Assaying cytochrome C translocation during apoptosis. Methods Mol. Biol. 284, 307–313. [DOI] [PubMed] [Google Scholar]
  59. Watt J., Schlezinger J. J. (2015). Structurally-diverse, PPARgamma-activating environmental toxicants induce adipogenesis and suppress osteogenesis in bone marrow mesenchymal stromal cells. Toxicology 331, 66–77. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Wolff S., Erster S., Palacios G., Moll U. M. (2008). p53’s mitochondrial translocation and MOMP action is independent of Puma and Bax and severely disrupts mitochondrial membrane integrity. Cell Res. 18, 733–744. [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Wols H. A., Johnson K. M., Ippolito J. A., Birjandi S. Z., Su Y., Le P. T., Witte P. L. (2010). Migration of immature and mature B cells in the aged microenvironment. Immunology 129, 278–290. [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Wu J. Y., Scadden D. T., Kronenberg H. M. (2009). Role of the osteoblast lineage in the bone marrow hematopoietic niches. J. Bone Miner. Res. 24, 759–764. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Yamaguchi K., Matulka R. A., Schneider A. M., Toselli P., Trombino A. F., Yang S., Hafer L. J., Mann K. K., Tao X.-J., Tilly J. L., et al. (1997). Induction of preB cell apoptosis by 7,12-dimethylbenz[a]anthracene in long-term primary murine bone marrow cultures. Toxicol. Appl. Pharmacol. 147, 190–203. [DOI] [PubMed] [Google Scholar]
  64. Yanik S. C., Baker A. H., Mann K. K., Schlezinger J. J. (2011). Organotins are potent activators of PPAR{gamma} and adipocyte differentiation in bone marrow multipotent mesenchymal stromal cells. Toxicol. Sci. 122, 476–488. [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Yokota T., Meka C. S., Kouro T., Medina K. L., Igarashi H., Takahashi M., Oritani K., Funahashi T., Tomiyama Y., Matsuzawa Y., et al. (2003). Adiponectin, a fat cell product, influences the earliest lymphocyte precursors in bone marrow cultures by activation of the cyclooxygenase-prostaglandin pathway in stromal cells. J. Immunol. 171, 5091–5099. [DOI] [PubMed] [Google Scholar]
  66. Zhu J., Garrett R., Jung Y., Zhang Y., Kim N., Wang J., Joe G. J., Hexner E., Choi Y., Taichman R. S., et al. (2007). Osteoblasts support B-lymphocyte commitment and differentiation from hematopoietic stem cells. Blood 109, 3706–3712. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary Data
Click here for additional data file. (645.2KB, pdf)

Articles from Toxicological Sciences are provided here courtesy of Oxford University Press

RESOURCES