The authors state that they have no conflicts of interest.
Published online on June 18, 2007
The AHR mediates many of the toxicological effects of aromatic hydrocarbons. We show that AHR expression in osteoblasts parallels the induction of early bone-specific genes involved in maturation. The AHR may not only mediate the effects of toxicants, but with an as yet unidentified ligand, be involved in the differentiation pathways of osteoblasts.
Introduction: Metabolic bone diseases arise as a result of an imbalance in bone cell activities. Recent evidence suggests that environmental toxicants may be contributing factors altering these activities. One candidate molecule implicated in mediating the toxic effects of exogenous compounds is the aryl hydrocarbon receptor (AHR).
Materials and Methods: Osteoblasts isolated from neonatal rat calvaria were analyzed for AHR expression by quantitative PCR, Western blot, and immunohistochemistry. In addition, AHR activation was evaluated by electromobility gel shift assay and fluorescence microscopy.
Results: Our findings showed AHR expression in mature osteoblasts in vivo. The pattern of AHR expression peaks after alkaline phosphatase and before induction of osteocalcin. We first show that AHR functions as a transactivating receptor in osteoblasts, as evidenced by its ligand-dependent migration to the nucleus and its association with known dioxin response elements. AHR activation by 2,3,7,8-tetrachlorodibenzo -p -dioxin (TCDD) mediated the induction of cytochrome p450 1A1 and cycloxygenase-2 protein levels. This effect could be inhibited by the potent AHR antagonist, 3′4 methoxynitroflavone. Furthermore, lead treatment of osteoblasts upregulates the expression of AHR mRNA and protein levels, supporting a novel mechanism whereby lead in the skeleton may increase the sensitivity of bone cells to toxicant exposure.
Conclusions: These data imply that the AHR mediates the effects of aromatic toxicants on bone and that AHR expression is regulated during osteoblast differentiation.
Osteoblast proliferation, differentiation, matrix synthesis, and mineralization comprise, an important series of events throughout the development and functioning of the skeleton. Osteoblasts are derived from mesenchymal stem cells and are responsible for synthesizing matrix proteins that subsequently become mineralized during the process of bone formation. Normal bone remodeling is a dynamic process that is characterized by a balance of bone resorption by osteoclasts and bone formation by osteoblasts.(1) Local cellular factors and hormones control the rate of remodeling, and an alteration in gene expression during osteoblast development may result in an imbalance in these processes.
Emerging evidence indicates that exposure to environmental toxicants influences osteoblast-specific functions. For example, 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) decreases bone nodule formation by osteoblasts treated in vitro.(2) However, little is known about the mechanism by which TCDD inhibits osteoblast differentiation. These findings prompted us to study a possible role of the aryl hydrocarbon receptor (AHR) in osteoblast function. This transcription factor mediates the toxicity of persistent environmental contaminants such as polycyclic-aromatic hydrocarbons and congeners of dioxin/furan compounds. AHR ligands elicit toxic responses such as endocrine disruption, tumor promotion, immunosuppression, and interference with fatty acid metabolism and cell differentiation.(3–6) Notably, AHR ligands are also prevalent in tobacco smoke.(7–9) It is not known whether this may be related to the findings that smoking is a risk factor for bone loss and osteoporosis.
AHR transformation by a ligand is followed by loss of chaperone heat shock protein 90, nuclear translocation, and heterodimerization with the AHR nuclear translocator (ARNT; also known as HIF-β).(10) The heterodimer complex binds DNA responsive elements and regulates expression of a variety of genes including those involved in phase I and phase II drug metabolism.(11,12) Ligand binding seems to be a prerequisite step for AHR activation, and no endogenous ligand has yet been conclusively identified.
The AHR is part of the basic helix-loop-helix Per-Arnt-Sim family of proteins involved in organogenesis and cellular development, and this evolutionarily conserved family of transcription factors displays a unique pattern of developmental expression emerging as early as mouse embryonic day 9.5.(13) As such, some as yet unknown, normal functional role of the AHR has been hypothesized. Inactivating the AHR in mice by gene knockout has shown that this protein is necessary for normal development of several tissues including liver, immune system, heart, and vascular tissues, as well as for normal reproduction.(14–19) Here we present data indicating that the AHR is upregulated during osteoblast differentiation and further hypothesize that altered activation of the AHR signaling pathway by TCDD may mediate adverse effects on osteoblasts by modulating differentiation.
Another environmental toxicant, lead, also strongly influences the bone microenvironment. The diverse effects of lead on skeletal tissue include impaired collagen synthesis, reduced circulating osteocalcin levels, and altered endocrine system function.(20–22) Lead interferes with intracellular calcium signaling in a way that impairs the production of osteocalcin.(23,24) The potential for lead to increase the sensitivity of bone tissue to further toxicological insults has not been previously studied. However, recent evidence showed that lead can increase the expression of AHR-regulated genes in mouse hepatoma cells.(25) Therefore, we studied the ability of lead to influence AHR expression in osteoblasts. Herein we also report that AHR expression is upregulated in osteoblasts in response to treatment with lead.
MATERIALS AND METHODS
TCDD (Cambridge Isotopes, Cambridge, MA, USA) was dissolved in DMSO (10 μM) and diluted to working concentrations in culture media. 3′-methoxy-4′-nitroflavone (MNF; 5 mM; synthesized as previously described(26)) stock solutions were diluted to working concentrations in culture media. MNF was used at concentrations previously reported to antagonize AHR activation by TCDD.(27) One molar stock concentrations of lead acetate (Sigma, St Louis, MO, USA) were diluted to working concentrations in culture media.
Rat osteoblast isolation and culture conditions
Osteoblasts were isolated from neonatal rat calvaria as previously described(28) and were plated at a density of 2 × 105 cells/ml in low-glucose α-MEM medium (Invitrogen, Carlsbad, CA, USA) with 5% bovine serum, 10% Pen-Strep, and 50 μg/ml ascorbate. Osteoblasts were cultured in 6-well plates (volume of 2 ml) for 2 days to confluence and used for experimentation. Fresh media were added to the cells every 2 days. β-glycerol phosphate (BGP; 10 mM) was added to the media 2 days after plating to promote osteoblast differentiation and mineralization.(29) No toxicity was seen to osteoblasts when cultured with 5 and 10 nM TCDD or 0.5 and 1 μM MNF (data not shown). Cellular toxicity was assessed by the MTT assay and trypan blue exclusion.
Neonatal rat calvaria and calvaria from 4-wk-old wildtype and AHR knockout mice were harvested, fixed in 10% formalin, decalcified, and paraffin embedded. B6.129-Ahr tml Bra/J (AHR-KO) mice were obtained from The Jackson Laboratories (Bar Harbor, ME, USA) to initiate a breeding colony at our facilities. These animals have been back-crossed into the C57Bl/6 background for at least 20 generations. Sections (4 μm) were cut and stained with a polyclonal goat anti-mouse antibody specific for AHR (Santa Cruz Biotechnology, Santa Cruz, CA, USA) or matched isotype control antibody. Visualization of the primary antibody was with a horseradish peroxidase (HRP)-conjugated rabbit anti-goat antibody (Jackson ImmunoResearch Laboratories, West Grove, PA, USA). Multiple calvaria sections were analyzed along with sections of mouse liver highly expressing AHR protein as a positive control. All sections were counterstained with eosin.
In vitro osteogenesis
Bone marrow cells were obtained from the femurs of 12- to 18-wk-old male wildtype and AHR knockout mice. Both femora were removed, and the soft tissues were detached aseptically. Bone marrow cells were collected by flushing the diaphysis with culture medium. A suspension of bone marrow cells was obtained by repeated aspiration of the cell preparation through a 22-gauge needle, and nucleated cells were counted with a hemocytometer. Bone marrow cells were incubated overnight and ascorbate supplemented medium was changed thereafter every 2 days for 1 wk. During the second week of culture, 10 mM BGP and 50 μg/ml ascorbate-supplemented medium was added to adherent cells to promote cell expansion and osteoblast differentiation. Nodule formation was assessed after 14 and 21 days in culture by alizarin red staining. Briefly, cells were washed with 2 ml of PBS and fixed with 10% formaldehyde at room temperature for 15 min. Cells were rinsed gently three times (5–10 min each) with an excess of distilled water. Cells were incubated with 1 ml of alizarin red solution (1% alizarin red/100% ethanol; Sigma) at room temperature for 20 min. Excess dye was removed, and cells were washed four times with deionized water. Differentiated cells containing mineral deposits were visualized on an Olympus BX51 microscope (Mellville, NY, USA).
Total cellular RNA was harvested from cultured osteoblasts on days 1, 3, 5, 10, 15, and 20 after treatment with BGP. RNA was extracted using the Qiagen RNAeasy mini kit (Valencia, CA, USA) following the supplier's protocol. One microgram of RNA was used in a two-step approach: RT-PCR for first-strand cDNA synthesis using Clontech RT kit primed by an oligo(dt)primer (Mountain View, CA, USA). RNA (10 ng) was used in each cDNA amplification reaction. The following primers were used for real-time PCR amplification: GAPDH, 5′-accacagtcccatgccatcac-3′, tccaccaccctgttgctgta (400 bp); AHR, 5′-caaagggcagcagcttattattctggg-3′, aagcgtgcattggactggac (197 bp); alkaline phosphatase, 5′-tgatcactcccacgttttca-3′, ctgggcctggtagttgttgt (202 bp); osteocalcin, 5′-ctcctgcttggacatgaagg-3′, tagcagacaccatgaggacc (419 bp). No change in GAPDH mRNA levels was detected with culture conditions. GAPDH was used to normalize real-time RT-PCR reactions for purposes of quantitation. Quantitative PCR using CYBERGREEN master mix as the fluorescent DNA intercalating agent was analyzed using Rotorgene Software (Corbett Research, Sydney, Australia). Real-time PCR melt curve analysis showed the formation of single products that were electrophoresed on a 1.2% agarose gel. Identification of PCR products was confirmed by ABI sequencing.
SDS-PAGE and Western blot
Total cellular protein was collected from cultured osteoblasts on days 1, 3, 5, 10, 15, and 20 after treatment with BGP. Osteoblasts were lysed in protein isolation buffer (1% IGEPAL, 150 mM sodium chloride, 50 mM TRIS, 10% protease inhibitor cocktail) and quantified by a BCA protein assay kit (Pierce, Rockford, IL, USA). Ten to twenty micrograms of protein was fractionated by 7.5% SDS-PAGE and electrophoretically transferred onto nitrocellulose membranes. Membranes were blocked in 10% nonfat powdered milk in 1× PBS/0.1% Tween 20 and incubated with either a primary goat anti-mouse cyclooxygenase-2 (Cox-2; Cayman Chemical, Ann Arbor, MI, USA), a monoclonal anti-rat cytochrome p450 1A1 (CYP1A1; Xenotech, Kansas City, KS, USA), a polyclonal rabbit anti-mouse AHR (BioMol, Plymouth Meeting, PA, USA), an anti-mouse aryl hydrocarbon nuclear transporter (ARNT), or a mouse anti-actin (Oncogene Research Products, San Diego, CA, USA) antibody for 2 h at 20°C. After washing, membranes were incubated with secondary HRP-conjugated antibodies (Jackson Immunoresearch) for 1 h at 20°C. Bands were visualized with enhanced chemiluminescence (Amersham Biosciences, Piscataway, NJ, USA) and autoradiography film according to the manufacturer's protocol. The molecular weights for each protein detected were as follows: AHR, 99 kDa; ARNT, 87 kDa; actin, 45 kDa; Cox-2, 68–72 kDa; Cyp1A1, 55 kDa.
Osteoblasts were cultured in the presence and absence of 10 nM TCDD for 60 min in 8-well chamber slides and analyzed for AHR nuclear localization by immunocytochemistry as previously described.(30) Cellular expression of AHR was detected in osteoblasts using rabbit polyclonal anti-AhR antibody (BioMol International) and visualized using Alexa-Fluor secondary antibody (Molecular Probes, Carlsbad, CA, USA). Incubation with secondary antibody alone was used as a control to assess nonspecific antibody binding. Nucleus was visualized by 4′,6′-diamidino-2-phenylindole (DAPI; Molecular Probes) staining. Cells were washed, lightly fixed in 2% paraformaldehyde (PFA) (5 min) and immediately visualized by fluorescence microscopy on an Axiovert 40 confocal microscope using an Axio camera.
Osteoblasts cultured with BGP were treated with DMSO or TCDD for 15, 30, or 60 min. Nuclear extracts were collected at each time-point. Cells were lysed with buffer containing 10 mM HEPES-KOH, 1.5 mM MgCl2, and 10 mM KCl to retain a nuclear pellet. Nuclear pellets were dissolved in a buffer containing 20 mM HEPES-KOH, 25% glycerol, 420 mM NaCl, 1.5 mM MgCl2, and 0.5 M EDTA. Five micrograms of nuclear protein was incubated in binding buffer [20 mM DTT, 25 mM HEPES, 10% glycerol, 1.5 mM EDTA, and 420 mM NaCl, and 0.05 mg/ml poly(dI:dC)] with a γ-32P end-labeled dioxin response element (DRE) oligonucleotide for 20 min. The sequence for DRE oligo was 5′-gatccggctcttctcacgcaactccgagctca-3′.(31) EMSA was performed at 60 V for 15 min and 140 V for 2.5–3 h. TCDD-treated liver lysate was used for a positive control for DRE binding, and untreated osteoblasts were a negative control.(10)
Alkaline phosphatase enzyme activity
Osteoblasts cultured for 5 days were treated with or without TCDD in the presence and absence of MNF for 24 h. Osteoblasts were washed three times in 0.9% NaCl and lysed with M-Per (Pierce). Twenty microliters of total lysate was added to 1 ml substrate p-nitrophenol phosphate dissolved in AMP buffer (BioRad, Hercules, CA, USA) at pH 10.5. After incubation at 37°C for 5 min, 0.5 ml stop solution (0.3 M Na3P04) was added, and alkaline phosphatase activity was measured at 410 nM on a Benchmark ELISA Plate reader (BioRad). Absorbance values were normalized to total protein levels that were quantified using a BCA kit (Pierce).
AHR expression by osteoblasts in vivo
Paraffin-embedded sections of neonatal rat calvarium were analyzed by immunohistochemistry to determine whether osteoblasts express AHR protein in vivo. Figure 1A shows AHR expression in osteoblasts from rat calvaria. Osteocytes do not express AHR protein in vivo. Figure 1B shows that AHR is expressed in adult mouse calvarial osteoblasts and that it is absent in AHR-deficient mice (Fig. 1C).
AHR upregulation during osteoblast differentiation
We next determined whether AHR expression was regulated in osteoblasts harvested from neonatal rat calvaria. Osteoblasts were induced to differentiate and form mineralized bone nodules in vitro with BGP and ascorbate.(29) To evaluate the AHR gene expression profile during this process, total RNA was harvested at different stages of osteoblast development and analyzed by RT-PCR. The data in Fig. 2A show that steady-state AHR mRNA levels were upregulated on days 3, 5, and 10 during osteoblast differentiation. The induction of steady-state alkaline phosphatase mRNA during differentiation and osteocalcin mRNA before mineralization are positive controls for the respective developmental stages of the tissue culture.(32) Further mRNA analysis using real-time RT-PCR showed a significant 4-fold induction of steady-state AHR mRNA levels on day 5 after BGP treatment (Fig. 2B). The peak 10-fold induction of AHR mRNA levels was detected on day 15 after BGP. The induction of AHR mRNA levels follows the increased alkaline phosphatase expression and precedes the induction of osteocalcin.
We next studied whether there was a concomitant upregulation of AHR protein during osteoblast differentiation as detected by Western blot. AHR protein was constitutively expressed in cultured osteoblasts and was elevated on days 5 and 10 after BGP treatment (Fig. 2C). These findings show that AHR protein expression was induced during differentiation and reduced to control levels during matrix maturation. An in vitro transcribed and translated AHR protein was used as a positive control and β-actin was used as a protein loading control. These findings of increased AHR mRNA and protein expression during critical stages of bone development may suggest a specific action for AHR in regulating skeletal development. Furthermore, these data also indicate that differentiating osteoblasts may be an important cellular target for the toxic AHR ligands.
Reduced in vitro osteogenesis by AHR-deficient compared with wildtype mice
The in vitro differentiation of bone marrow cells to mature osteoblasts with characteristics such as matrix maturation and eventual formation of multilayered nodules with a mineralized extracellular matrix was next evaluated in wildtype and AHR knockout mice. AHR knockout mice bone marrow cells showed a 70% reduction in bone nodule formation after 14 and 21 days of culture compared with wildtype (Fig. 3). Significant reductions in alkaline phosphatase activity were also measured in AHR knockout osteoblasts compared with wildtype. These findings showed that AHR is needed for optimal alkaline phosphatase activity in differentiating osteoblasts.
Osteoblast AHR activation by TCDD
These findings indicating regulated AHR expression during osteoblast differentiation prompted us to evaluate whether AHR could migrate to the nucleus and bind DNA in response to TCDD, a potent AHR ligand. Osteoblasts were treated with vehicle (DMSO) or TCDD for 15, 30, and 60 min, and nuclear localization was determined by fluorescence microscopy. Figure 4A shows that, in untreated osteoblasts, AHR is present both in the cytosol and nucleus, but with TCDD exposure for 30 min, the receptor translocates entirely into the nucleus. These data confirmed ligand-dependent activation and nuclear localization of the AHR in osteoblasts. The ability of nuclear translocated AHR to bind to the DNA consensus sequence for the DRE was next determined by EMSA. Figure 4B shows increased DNA binding of the AHR after 30 and 60 min of TCDD treatment compared with vehicle-treated control cells. Liver cell lysate treated with TCDD was used as a positive control for DNA binding, and untreated osteoblasts represent the negative control. These data are consistent with the AHR being a functional transcription factor and a direct molecular target for TCDD in osteoblasts.
TCDD-activated AHR induced CYP1A1 and Cox-2 protein expression
Given that the ligand-activated AHR in osteoblasts binds DNA, we investigated whether AHR could upregulate CYP1A1 protein levels, a conventional marker of AHR transcriptional activity.(33) As shown in Fig. 5A, exposure to 5 and 10 nM TCDD for 6 h increased CYP1A1 protein expression compared with vehicle-treated cells as detected by Western blot. A DRE was recently reported in the Cox-2 promoter,(34) and we therefore also examined Cox-2 expression after TCDD treatment in osteoblasts. Interestingly, a dramatic increase in Cox-2 expression by osteoblasts in response to AHR ligand compared with vehicle-treated osteoblasts was detected. The levels of AHR and ARNT remained relatively unchanged.
To determine whether the induction of CYP1A1 and Cox-2 protein by TCDD was mediated by AHR activation, we incubated osteoblasts with the AHR antagonist, MNF, in the presence and absence of TCDD for 6 h. As shown in Fig. 5B, the CYP1A1 and Cox-2 induction by 10 nM TCDD was substantially reduced by treatment with 0.5 or 1 μM MNF. These findings indicate that AHR activation by TCDD was responsible for increased CYP1A1 and Cox-2 protein levels.
TCDD reduced osteoblast alkaline phosphatase activity through an AHR-dependent mechanism
Increased alkaline phosphatase activity is a marker of osteoblast differentiation.(32) Alkaline phosphatase activity was measured in these studies to determine whether TCDD altered an osteoblast-specific function. Osteoblasts cultured with BGP were treated for 24 h with DMSO or increasing concentrations of TCDD. TCDD (2.5 and 5 nM) significantly reduced alkaline phosphatase enzymatic activity after 24 h. A statistically significant dose-dependent decrease in enzymatic activity was seen after 48 h with 1 and 2.5 nM TCDD (Fig. 6A). We determined that TCDD did not affect cell viability and was not directly inhibiting alkaline phosphatase enzyme activity (data not shown), and therefore, we next investigated whether this reduction occurred by an AHR-dependent mechanism. Osteoblasts were pretreated with the AHR antagonist, MNF, for 20 min and incubated in the presence and absence of TCDD for 24 h. The data in Fig. 6B show that the dose-dependent reduction in alkaline phosphatase activity by TCDD was completely reversed in osteoblasts treated with 1 μM MNF. These data support that TCDD reduced alkaline phosphatase activity in an AHR-dependent manner during osteoblast differentiation.
Lead induced upregulation of osteoblast AHR expression
Because bone is a storage depot for lead in the body, recent evidence confirmed that there are chronic adverse effects of lead on skeletal metabolism.(35–37) We studied whether lead modulates AHR expression. We first examined changes in steady-state AHR mRNA levels in response to increasing doses of lead as detected by real-time RT-PCR. Figure 7A shows that lead-treated osteoblasts increased steady-state AHR mRNA levels compared with untreated osteoblasts after 24 h. There was a significant 4-fold induction of AHR mRNA levels with 0.1 μM lead and an 8-fold induction in osteoblasts treated with 0.5 μM lead compared with untreated cells. The effect seemed to be biphasic in nature, because higher concentrations of lead did not evoke the same level of induction. We next determined whether the elevated AHR mRNA levels resulted in increased AHR protein expression. Consistent with the lead-induced AHR mRNA levels, a significant increase in AHR protein expression was detected in the 0.5 μM lead-treated osteoblasts compared with vehicle control (Fig. 7B). These data reveal a novel mechanism by which lead can influence bone tissue and implicate this heavy metal as an enhancer of the effects of toxic AHR ligands.
The data in this report showed a functional AHR signaling pathway in osteoblasts that may mediate both toxic and physiological responses during osteoblast development. AHR protein expression was clearly shown in neonatal rat calvarium in vivo (Fig. 1). Steady-state AHR mRNA levels and protein expression were regulated in osteoblasts that were induced to differentiate (Fig. 2). The pattern of AHR expression peaks after alkaline phosphatase and before induction of osteocalcin. Furthermore, AHR-deficient mouse bone marrow cells showed reduced differentiation to mature nodule forming osteoblasts compared with wildtype mice (Fig. 3). Together these data, along with ligand-activated AHR nuclear localization and binding to the DRE, support a possible novel function for this transcription factor in osteoblasts (Fig. 4). Our data showed that stimulation of AHR by TCDD decreases osteoblast alkaline phosphatase activity and suggests that this family of transcription factors modulates expression and/or activity of differentiation molecules in bone tissue. Moreover, the reduction in alkaline phosphatase activity by differentiating AHR-deficient osteoblasts indicates that a delicate balance of AHR activity is needed for nodule formation. Recent evidence supports a role for the AHR in osteoclastogenesis, because another AHR ligand, 3-methylcholanthrene, inhibited RANKL expression by osteoblastic cells.(38) Future studies that address whether the ligand-activated AHR alters other aspects of osteoblast differentiation (e.g., cell cycle progression, cytokine production) will enhance our understanding of the mechanisms by which AHR ligands influence the bone microenvironment. However, at present, it is not clear how possible endogenous AHR ligands influence the cellular growth and differentiation of osteoblasts compared with the toxic ligands.
Given that AHR expression peaks during osteoblast differentiation, this critical stage of bone development should be considered a susceptible target for toxic ligands to disrupt normal bone formation processes. AHR ligands may alter the normal processes of bone formation by inducing changes in gene expression. AHR transactivation by TCDD increased CYP1A1 and Cox-2 protein expression (Fig. 5). Although endogenous regulation of CYP1A1 and Cox-2 augment bone formation,(39) inappropriate upregulation of these enzymes by toxicants may exert adverse effects on the bone microenvironment.(40,41) Seidel et al.(42) reported activation of the AHR signaling pathway by prostaglandins in mouse hepatoma cells, suggesting a role for the AHR in mediating the effects of prostaglandins. However, our data revealed that AHR ligands may promote prostaglandin production at an inappropriate time in bone development and thus influence bone formation in ways that are not yet fully appreciated. Similarly, altered regulation of CYP1A1 drug-metabolizing activity could contribute to the overproduction of toxic metabolites and reactive oxygen species, thus interfering with tightly regulated bone formation processes. Moreover, we found that 1 μM MNF treatment alone increased Cox-2 levels (Fig. 5B), contrary to the ability of this AHR antagonist to completely reduce the TCDD-mediated upregulation of Cox-2. These findings support previous reports suggesting that MNF has low AHR agonist activity.(43)
Inappropriate activation of the AHR by TCDD during osteoblast differentiation not only influenced gene expression but also mediated significant reductions in alkaline phosphatase enzymatic activity (Fig. 6). We show that the reduction in alkaline phosphatase activity by TCDD was mediated through the AHR, because pretreatment with the AHR antagonist, MNF, dose-dependently restored osteoblast alkaline phosphatase activity. Future studies evaluating the potent AHR antagonist in osteoblasts and particularly the influence on their differentiation will provide more insight into the effects of TCDD that are AHR dependent and may guide the discovery of the role of AHR in bone physiology. These findings reported herein suggest that other bone-specific activities may be altered by TCDD exposure and warrant further study of the effects of AHR ligands on bone tissue in vivo.
In contrast to reports of AHR degradation after activation within isolated cells,(44) Franc et al.(45) reported in vivo upregulation of the AHR in rat liver in response to dioxin. The in vivo phenomenon suggests the possibility for continuous sensitization of the cells to the effects of persistent environmental chemicals. We report that lead upregulated AHR mRNA and protein expression in cultured osteoblasts, suggesting a similar mechanism by which osteoblasts may be more susceptible to insults by other toxic AHR ligands. The ability of lead to actively alter bone processes immediately after it becomes sequestered in the skeleton and throughout the life of an organism is appreciated, but still not yet well understood. A recent study showed differential effects of heavy metals on the constitutive and inducible expression of AHR-regulated genes in cultured hepatoma cells.(25) We postulate that lead-induced AHR expression influences endogenous AHR transactivation during osteoblast differentiation, and that exposure to exogenous ligands may only exacerbate the adverse effects of lead. Furthermore, the likelihood of successive and continuous exposure to organic and inorganic compounds in human populations supports the need for future studies to address the effects of mixtures, particularly because bone tissue is a reservoir for lead. Our finding of increased AHR expression by lead may have important implications for further increased sensitivity of bone to a variety of persistent environmental chemicals (e.g., dioxin, cigarette smoke, polychlorinated biphenyls) and warrants the active study of mixtures in vivo.
The authors thank Barbara Stroyer for assistance with the histology and immunohistochemistry. The work in this manuscript was supported by NIH Grants T32 ES07026, P30 ES01247, R01 ES09430, R25 CA102618, and P01 ES 11845.