Benzo(a)pyrene and 7,12-dimethylbenz(a)anthrecene differentially affect bone marrow cells of the lymphoid and myeloid lineages

Toxicology and Applied Pharmacology 213 (2006) 105 – 116 www.elsevier.com/locate/ytaap

Benzo(a)pyrene and 7,12-dimethylbenz(a)anthrecene differentially affect bone marrow cells of the lymphoid and myeloid lineages Noe´ Galva´n a, Todd J. Page c, Charles J. Czuprynski a,c, Colin R. Jefcoate a,b,* a

Molecular and Environmental Toxicology, University of Wisconsin, 1300 University Avenue, Madison, WI 53706, USA b Department of Pharmacology, University of Wisconsin, 1300 University Avenue, Madison, WI 53706, USA c Department of Pathological Sciences, University of Wisconsin, 2015 Linden Drive, Madison, WI 53706, USA Received 1 March 2005; revised 14 September 2005; accepted 14 September 2005 Available online 22 November 2005

Abstract Polycyclic aromatic hydrocarbons (PAHs) are common environmental contaminants that are carcinogenic and immunosuppressive. Benzo(a)pyrene (BP) and 7,12-dimethylbenz(a)anthracene (DMBA) are two prototypic PAHs known to impair the cell-mediated and humoral immune responses. We have previously shown that, in C57BL/6J mice, total bone marrow (BM) cellularity decreased two-fold following intraperitoneal DMBA treatment but not BP treatment. Here, we have used flow cytometry to demonstrate that BP and DMBA differentially alter the lymphoid and myeloid lineages. Following DMBA treatment, the pro/pre B-lymphocytes (B220lo/IgM ) and the immature B-lymphocytes (B220lo/IgM+) significantly decreased, while the mature B-lymphocytes (B220hi/IgM+) remained unaffected. In contrast, BP treatment decreased the pro/pre B-lymphocytes, and did not affect the immature B-lymphocytes or mature B-lymphocytes. The Gr-1+ cells of the myeloid lineage were depleted 50% following DMBA treatment and only minimally depleted following BP treatment. Interestingly, the monocytes (7/4+1A8lo) and neutrophils (7/4+1A8hi) within this Gr-1+ population were differentially affected by these PAHs. Monocytes and neutrophils were depleted following DMBA treatment whereas neutrophils decreased and monocytes increased following BP treatment. Although TNFa and CYP1B1 are implicated as essential mediators of hypocellularity, the similar induction of TNFa mRNA and CYP1B1 mRNA in the BM by BP and DMBA suggests that they are not limiting factors in mediating the different effects of these PAHs. Given that similar amounts of BP and DMBA reach the BM when administered intraperitoneally, their differential effects on the lymphoid and myeloid lineages probably stem from differences in reactive metabolites such as PAH quinones and PAH-dihydrodiol-epoxides. D 2005 Published by Elsevier Inc. Keywords: Polycyclic aromatic hydrocarbons; Benzo(a)pyrene; 7,12-dimethylbenz(a)anthracene; Bone marrow; Hematopoiesis; B-lymphocytes; Monocytes; Neutrophils

Introduction Polycyclic aromatic hydrocarbons (PAHs) are ubiquitous environmental contaminants that are produced during incomplete combustion of organic materials. The in vivo effects of PAH treatment of mice include a decrease in bone marrow cellularity (Heidel et al., 2000), which could contribute to Abbreviations: AHR, aryl hydrocarbon receptor; BP, benzo(a)pyrene; CLPs, common lymphoid progenitors; CMPs, common myeloid progenitors; DMBA, 7,12-dimethylbenz(a)anthracene; HSCs, hematopoietic stem cells; PAHs, polycyclic aromatic hydrocarbons; PAHDEs, PAH-dihydrodiol-epoxides; TCDD, 2,3,7,8-tetrachlorodibenzo-p-dioxin. * Corresponding author. Department of Pharmacology, University of Wisconsin, 1300 University Avenue, Madison, WI 53706, USA. Fax: +1 608 265 1257. E-mail address: [email protected] (C.R. Jefcoate). 0041-008X/$ - see front matter D 2005 Published by Elsevier Inc. doi:10.1016/j.taap.2005.09.018

impairment of humoral and cell-mediated immunity (Ward et al., 1984; Dean et al., 1985; Ladics et al., 1991; White et al., 1994). PAHs typically require activation by cytochrome P450 (CYP) to exert their biological effects. The induction of CYP transcription by PAHs is tissue-specific and dependent on activation of the aryl hydrocarbon receptor (AHR) (Schmidt and Bradfield, 1996; Rowlands and Gustafsson, 1997). The major responses are provided by members of the CYP1 family (CYP1A1, CYP1A2, and CYP1B1) which each metabolize PAHs but with different selectivities (Pottenger and Jefcoate, 1990; Savas et al., 1993, 1997; Kleiner et al., 2002). CYP1A2 is primarily expressed in the liver and is expressed at appreciable constitutive levels whereas CYP1B1 is almost exclusively expressed in extrahepatic tissues (Gonzalez and Lee, 1996; Murray et al., 2001; Shimada et al., 2003). CYP1A1

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is expressed in liver and many cell types, but it is only expressed at functional levels after induction through the AHR (Hankinson, 1995; Gonzalez and Lee, 1996; Whitlock, 1999). This paper addresses bone marrow toxicity following intraperitoneal administration of two PAHs, benzo(a)pyrene (BP) and 7,12-dimethylbenz(a)anthracene (DMBA), which are both carcinogenic (Pelkonen and Nebert, 1982) and immunosuppressive (White et al., 1994). These PAHs differ substantially in their activation of the AHR (BP>>DMBA). The toxicity of chemicals in peripheral tissues like the bone marrow is dependent on the balance of metabolic processes in the liver and in the peripheral tissue. The liver is able to remove PAHs but can also deliver toxic metabolites to peripheral tissues. The liver contribution is highly dependent on the route of administration. Recent work shows that when BP is administered intragastrically, CYP1A1 in the liver removes most of the BP within 5 h (Uno et al., 2004). However, the liver contribution is much less for intraperitoneal delivery where the levels of BP are sustained for over 24 h and equal levels of BP and DMBA reach the bone marrow (Galvan et al., 2005). Furthermore, metabolism of PAHs in the bone marrow can provide reactive metabolites directly at the site of toxicity. CYP1B1, which is expressed in the bone marrow but not the liver, is essential for the toxicity of BP and DMBA (Galvan et al., 2003). Although not the case, the induction of CYP1B1 in the bone marrow via AHR activation was expected to be greater for BP than for DMBA. Phase II enzymes (NAD(P)H:quinone oxidoreductase (NQO1), UDP-glucuronosyltransferases (UGTs)), which remove reactive metabolites are also responsive to AHR activation and are likely to respond selectively to BP and DMBA metabolites generated in both the liver and the bone marrow (Ritter, 2000; Noda et al., 2003; Nioi and Hayes, 2004). The effect of PAHs on hematopoietic stem cells (HSCs) is probably mediated by bone marrow stromal cells, which express CYP1B1 largely to the exclusion of other CYPs (Heidel et al., 1999; Allan et al., 2003). Stromal cells also generate reactive metabolites and secrete cytokines and other proteins in response to chemical stress that affect hematopoiesis (Jensen et al., 2003). In vitro, there is very little difference between BP and DMBA with respect to toxicity generated from bone marrow stromal fibroblasts. BP and DMBA are examples of PAHs that, when administered intraperitoneally, are capable of inducing CYP1B1 within the bone marrow, liver, and lung (Galvan et al., 2003, 2005). However, only DMBA depletes bone marrow cellularity in wild-type C57BL/6J mice (Ahr b ). We have recently demonstrated that congenic mice (Ahr d ) with an AHR that is unresponsive to PAHs are equally susceptible to bone marrow depletion by BP and DMBA (Galvan et al., 2003). This finding implicates AHR activation in providing protection against the depletion of bone marrow by BP. The PAH activation process involves two CYP-dependent steps, which lead to the production of PAH-dihydrodiolepoxides (PAHDEs) (Christou et al., 1994; Miyata et al., 1999). These PAHDEs are primarily responsible for the initiation of carcinogenesis through covalent binding with DNA (Luch et al., 1998; Dipple et al., 1999; Buters et al., 2003). They also contribute to the immunosuppressive effects

of PAHs (Kawabata and White, 1987; Ladics et al., 1991; Nagasawa et al., 1996; Burchiel and Luster, 2001). However, alternative metabolism pathways that form quinones and phenols may also contribute to PAH toxicity (Kodama and Nagata, 1977; Moorthy et al., 2003; Cavalieri and Rogan, 2004). We recently demonstrated that the formation of PAHDE-DNA adducts is associated with bone marrow hypocellularity (Galvan et al., 2003, 2005). Similarly, others have shown that splenic leukocytes can metabolize PAHs to PAHDEs, which then form PAHDE-DNA adducts (Ginsberg et al., 1989). The presence of DNA adducts is taken as evidence for PAHDEs in these cells, and of possible modification of other macromolecules as well. These DNA adducts can evoke a well characterized DNAdamage response that can potentially arrest cell growth, initiate apoptosis, or alter the fate of progenitor cells. Hematopoiesis begins with hematopoietic stem cells (HSCs) that give rise to all blood cell lineages (Fig. 1) and is regulated by a tight network of cytokines and hormones, which are essential for the survival and differentiation of progenitor cells. This process can be altered by exogenous chemical stress (Kincade et al., 1994; Smithgall, 1998; Lotem and Sachs, 2002). For example, DMBA treatment causes a marked reduction in the number of bone marrow granulocytes and Blymphocytes in mice (Heidel et al., 2000; Galvan et al., 2003). In addition, we recently used Tnfr-null mice to demonstrate that TNFRSF1A (TNFR1), and to a lesser extent TNFRSF1B (TNFR2), were essential for DMBA induced bone marrow hypocellularity (Page et al., 2004). We also showed that the stress response mediators PKR and P53 are each essential for DMBA induced bone marrow hypocellularity (Page et al., 2003, 2004). NFnB signaling, which is linked to both of these proteins also plays a key modulatory role (Mann et al., 2001). These findings suggest that reactive metabolites generated by CYP1B1 metabolism of DMBA in the bone marrow might stimulate TNFa release that in turn contributes to the reduction of lymphoid and myeloid cell maturation or survival. We have recently shown that intraperitoneal administration of DMBA generates more stable DNA adducts than administration of BP (Galvan et al., 2005). In this study, we test the possibility that BP and DMBA activation leads to distinct induction of TNFa, which may explain the difference in their specific toxicity. Our previous emphasis on bone marrow hypocellularity as a measure of bone marrow toxicity might obscure differential effects of BP and DMBA on the various lineages of hematopoietic cells. In addition, these PAHs differ substantially in their generation of reactive metabolites like PAH quinones and PAHDEs, which may be selective in their effects on different bone marrow cell lineages (Twerdok et al., 1992; Savas et al., 1997). The intraperitoneal administration used here delivers near equal levels of BP and DMBA to the bone marrow, thus, providing a good test of such metabolism differences. This study resolves the effects of BP and DMBA on bone marrow hematopoietic stem cells (HSCs), and bone marrow cell population committed to the myeloid and lymphoid lineages. We have used antibodies against specific cell surface markers to distinguish the effects of PAH treatment on these populations. Our results indicate that DMBA sustains or

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Fig. 1. Simplified scheme of hematopoiesis from hematopoietic stem cells. Adapted from Akashi et al. (2000). Scheme includes surface markers used to identify Blymphocyte, myeloid, monocyte, and neutrophil populations. HSCs, hematopoietic stem cells; CLPs, common lymphoid progenitors; CMPs, common myeloid progenitors; MEPs, megakaryocyte/erythrocyte progenitors; GMPs, granulocyte/monocyte progenitors.

increases HSCs, while decreasing the number and proportion of cells committed to the myeloid and lymphoid lineages. Although BP does not cause a net loss of HSCs, it does cause shifts among bone marrow lineages, sometimes to a greater extent than does DMBA. Materials and methods Reagents and antibodies. Benzo[a]pyrene (BP) and 7,12-dimethylbenz[a]anthracene (DMBA) were purchased from Sigma Chemical (St. Louis, MO), and dissolved in olive oil at a concentration of 5 mg/ml for intraperitoneal injection. RPMI 1640 was purchased from Sigma Chemical and was supplemented with 5% FBS (v/v; Atlanta Biologicals), 50 IU penicillin/ml, and 50 Ag streptomycin/ml (w/v). The following monoclonal antibodies (mAbs) were purchased from BD Pharmingen: CD45/B220-phycoerythrin (PE), IgM-PE, Gr-1-fluorescein isothiocyanate (FITC), 1A8-FITC c-kit-cytochrome (Cy), and Sca-1-FITC. The following mAbs where purchased from Caltag: 7/4-PE. Animals and treatments. C57BL/6J (wild-type, Ahr b ) mice were purchased from The Jackson Laboratories (Bar Harbor, ME). Animals were housed at the AAALAC certified University of Wisconsin – Madison Medical School Animal Care Unit and used in accordance with the NIH Guide for the Care and Use of Laboratory Animals. Female and male mice (30 T 2 days old) were randomly selected and injected intraperitoneally with 50 mg/kg BP or DMBA in olive oil. This dose results in maximum bone marrow toxicity in wild-type mice for DMBA (Heidel et al., 2000). Untreated control animals were injected with an equivalent volume of olive oil. Bone marrow cell isolation. Mice were sacrificed 12, 24, and 48 h after intraperitoneal injection with oil vehicle, BP, or DMBA. The femurs and tibias were dissected free of muscle tissue, and the ends of the bones removed with a surgical blade. For the total bone marrow cell counts, cells from both femurs were flushed from the bones with 5 ml of culture medium using a syringe equipped with a 25-gauge needle. The bone marrow cells were dispersed into single cell suspensions by successive passage through 22- and 25-guage needles. Following centrifugation, red blood cells were lysed in ACK buffer (150 mM NH4Cl, 10 mM KHCO3, and 100 mM Na2EDTA pH 7.3). Viable cells were identified and enumerated in a hemocytometer by their exclusion of 0.05% Trypan Blue. Cells were then used for flow cytometry or isolation of total RNA for real-time RTPCR analysis. Flow cytometry staining and analysis. Following enumeration of bone marrow cells with the hemocytometer, freshly isolated cells were suspend at

1  106 cells per 100 Al media. Aliquots of cells (1  106) were maintained on ice for 10 min with FcgIII/II (0.5 Ag/106 cells, Caltag) to block Fc receptors. The cells were then incubated with 1 Ag of the primary antibody for 30 min on ice. Following incubation, the cells were washed once with 200 Al of media. Cells were resuspended in 500 Al media and propidium iodide (PI) was added (2 Ag/ml) to exclude dead cells from analysis. Aliquots containing positive and negative controls with isotype antibodies were processed simultaneously for each experiment. Fifty thousand cells were acquired for each sample with a FACSCalibur flow cytometer (BD Biosciences, San Jose, CA) and Blymphocytes, myeloid cells, monocytes, and neutrophils were analyzed using Flow Jo 4.5.8 software. Hematopoietic stem cell staining and analysis. Staining and analysis were performed as described previously (Goodell et al., 1996). Briefly, cells were resuspended at 1  106 per ml in pre-warmed medium (RPMI 1640 2% FBS, 50 IU penicillin/ml, 50 Ag streptomycin/ml (w/v), and 5 Ag per ml Hoechst 33342) and incubated for 90 min at 37 -C. The cells were resuspended with Hanks’ Balanced Salt Solution (HBSS) containing 2% FBS (HBSS+) at 1  108 cells per ml and incubated for 10 min with a cocktail consisting of the following monoclonal antibodies (mAb): CD4, CD8, CD5, B220, Mac-1, and Gr-1 (Ab 1/50 or 1/100) and then washed with HBSS+. Mouse lineage cells were resuspended in HBSS+ containing a PE conjugated anti-mouse Ab and incubated for 10 min on ice. Following incubation, cells were stained with 1Ag of mAb to Sca-1-FITC and c-kit-Cy, which are specific cell surface markers of HSCs (Morrison et al., 1997), washed, and filtered through 70 Am nylon filter. Cells were then resuspended in HBSS+ containing 2 Ag/ml PI, analyzed with a FACSVantage SE (BD Biosciences, San Jose, CA), and further analyses were performed using Flow Jo 4.5.8 software. Real-time RTPCR. Total RNA was extracted from bone marrow cells isolated from femurs and tibias using TRIzol Reagent (Life Technologies, Inc., Grand Island, NY), according to the manufacturer’s directions. RNA concentrations were measured via a spectrophotometer. Each reverse transcriptase reaction was performed using 1 Ag RNA and SuperScript II RNase H Reverse Transcriptase (Invitrogen, Carlsbad, CA) as per the manufacturer’s instructions. Oligonucleotide primers for TNFa (PubMed Accession # M13049), TNFRSF1A (TNFR1) (PubMed Accession # BC004599), AHR (PubMed Accession # AF405563), CYP1B1 (PubMed Accession # U03283), and Cyclophilin (PubMed Accession # X52803) were designed with Primers Express, version 2.0.0 (Applied Biosystems). The primer sequences are as follows: TNFa forward, 5V-tcatgcaccaccatcaagga-3V; TNFa reverse, 5V-ggctccagtgaattcggaaa-3V; TNFR1 forward, 5Vatgcagaccttgcgattctgt-5V; TNFR1 reverse, 3V-ccatgaaacgcatgaactcct-5V; AHR forward, 5V-agaatcccacatccgcatga-3V; AHR reverse, 5V-tgcaagaagccggaaaactg-3V; CYP1B1 forward, 5V-tggctgctcatcctctttac-3V; CYP1B1 reverse, 5V-aggttgggctggtcactcat-3V; Cyclophilin forward, 5V-agcgttttgggtccaggaat-3V; Cyclophilin

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reverse, 5V-aaatgcccgcaagtcaaaag-3V. Each treatment is presented as marrow isolated from three different mice and each PCR reaction carried out in duplicates. Real-time RTPCR was performed on an ABI 7000 Sequence Detection System. Each 20 Al reaction contained 2 Al cDNA (diluted 1:20), 1 SYBR Green PCR Master Mix (Applied Biosystems, Warrington, UK), and 100 nM of each forward and reverse primer. All reactions were run with the following parameters: 2 min at 50 -C then 10 min at 95 -C followed by 40 cycles of 95 -C for 15 s and 60 -C for 1 min. Standard curves for all genes presented were plotted separately using the cycle number at which the fluorescence signal exceeds background levels (Ct value) versus log cDNA dilution. For each PCR sample, the mean Ct for all genes was normalized against the mean Ct value for cyclophilin. Intracellular staining for TNFa. Bone marrow cells were harvested from the femurs of untreated C57BL/6J mice, and red blood cells removed by hypotonic lysis (ACK buffer), as described above. The cells were incubated for 5 h at 37 -C in 6 well tissue culture plates (2  106 cells per well) in the presence of 1 AM DMBA (or vehicle for the control cultures) and Brefeldin A (Golgi-Plug, BD Pharmingen), to prevent protein secretion. The cells were then permeabilized (Cytofix/Cytoperm, BD Pharmingen), stained with PE-labeled anti-TNFa mAb (BD Pharmingen), and fixed with 4% paraformaldehyde. The cells were then analyzed by flow cytometry as described above. Statistical analysis. Statistical analysis was performed by ANOVA followed by Tukey multiple comparison test for multiple groups. Student’s t test was

used when comparing two groups. Significance was set at P < 0.05 for all analysis.

Results Differential effects of BP and DMBA on bone marrow B-lymphocyte subpopulations We have previously demonstrated that wild-type C57BL/6J mice (Ahr b ) given a single intraperitoneal injection of DMBA (50 mg/kg) exhibit severe bone marrow cell hypocellularity of both the lymphoid and myeloid bone marrow cell lineages at 48 h post-treatment (Heidel et al., 2000). In contrast, this is not observed following equivalent treatment with BP (Heidel et al., 2000; Galvan et al., 2003). In the following experiments, we tested whether BP treatment has selective effects on bone marrow cell populations. Based on a previously reported flow cytometry analysis of B-lymphocyte subpopulations (Thurmond and Gasiewicz, 2000), a live gate parameter was established and maintained, which divided the bone marrow cells into region 1 (R1) and region 2 (R2) (Fig. 2A). Most B-lymphocytes are within R1

Fig. 2. Flow cytometric analysis of B-lymphocyte subpopulations. (A) Analysis of forward-scatter (FSC) versus propidium iodide (PI) for bone marrow cells from untreated control mice shows representative gating coordinates to identify viable cells. Based on staining for B220 and IgM, more than 90% of B-lymphocytes identified were found in region 1 (R1), whereas region 2 (R2) contains more than 90% of myeloid cells (Gr-1+). Representative dot plot for B220 versus IgM of lymphocyte subpopulation gate illustrates the three B-lymphocyte subpopulations for bone marrow cells from (B) control-, (C) DMBA-, and (D) BP-treated mice. Cells were assigned to these regions as previously described by Thurmond et al. (2000) (pro/pre-B cells, B220lo/IgM ; immature B-cells, B220lo/IgM+; and mature B-cells, B220hi/IgM+).

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and most myeloid derived cells are within R2. In the present study, we used the mAbs B220-FITC and IgM-PE to detect pro/pre B-lymphocytes (B220lo, IgM ), immature B-lymphocytes (B220lo, IgM+), and mature B-lymphocytes (B220hi, IgM+). Representative dot plots identifying B-lymphocyte subpopulations are shown for control-, DMBA-, and BP-treated mice (Figs. 2B – D). The proportion of lymphocytes in the bone marrow was not affected by either treatment even though DMBA halved the total number of cells (Galvan et al., 2003). The relative proportion of pre/pro-B-lymphocytes was reduced following DMBA treatment, while the proportion of mature Blymphocytes increased. The immature B-lymphocytes were not appreciably affected by DMBA treatment. BP treatment produced similar trends, albeit with smaller differences that were only significant for the loss of pre/pro-B-lymphocytes (Table 1). The total number of cells in each B-lymphocyte subpopulation was obtained by multiplying the percentage of cells in that population, Table 1, by the total number of bone marrow cells recovered from that mouse. This confirmed that the pro/pre and immature B-lymphocytes were highly susceptible to DMBA, and to a lesser extent to BP, while neither PAH affected the total number of mature B-lymphocytes (Fig. 3). Differential effects of BP and DMBA on bone marrow myeloid cells Myeloid cells were identified by their binding of the Gr-1 mAb which recognizes myeloid differentiation antigens (Ly6G and Ly-6C), and whose intensity of expression is directly correlated with myeloid cell differentiation and maturation into granulocytes (Fleming et al., 1993). DMBA treatment caused a 60% reduction in total myeloid cells (Gr-1+), whereas, BP had no significant effect (Fig. 5, Table 2). This is consistent with our previous results (Heidel et al., 2000; Galvan et al., 2003). We then used mAbs 7/4 and 1A8 (Ly-6G) to subdivide the Gr-1+ myeloid cells into predominantly neutrophil (7/4+, 1A8hi) and monocyte (7/4+, 1A8lo) populations (Henderson et al., 2003). The combination of 7/4 and 1A8 mAbs allows easier distinction of the two myeloid populations by flow cytometry than traditional means of using Gr-1 and Mac-1 mAbs to identify monocytes (Gr-1lo, Mac-1hi) and neutrophils (Gr-1hi, Mac-1hi). The flow cytometric results showed that both BP and DMBA significantly reduced the relative Table 1 Effect of BP and DMBA treatment on bone marrow B-lymphocytes and Blymphocyte subpopulations Treatment

Control DMBA BP

% B-lymphocyte (B220+)

% pro/pre B-lymphocyte (B220lo/lgM )

% immature B-lymphocyte (B220lo/lgMhi)

% immature B-lymphocyte (B220hi/lgMhi)

38.1 T 1.8 40.1 T 1.1a 34.0 T 1.0a

21.3 T 1.0 12.7 T 1.0a 16.1 T 0.7a

8.5 T 1.8 10.9 T 1.1a 9.5 T 1.0

8.3 T 0.7 16.5 T 1.4a 8.4 T 0.3

Note. For each treatment group, n  7; mean T SEM. Values represent the relative % of B-lymphocytes populations within the viable cells (see Fig. 2). a Significantly different from the control group (P < 0.05).

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Fig. 3. BP and DMBA effects on total bone marrow cells and B-lymphocyte subpopulations. Bone marrow cells of wild-type mice were eluted 48 h posttreatment from the femurs and red blood cells were removed. Flow cytometry data in Table 1 were multiplied by the number of total bone marrow cells for each mouse with the percentage of each population to obtain the total cells for each B-lymphocyte subpopulation. Viable cells were analyzed for pro/pre (B220lo/IgM ), immature (B220lo/IgM+), and mature (B220hi/IgM+) B-lymphocytes by flow cytometry. B-lymphocytes subpopulations are expressed as the mean T SEM as compared to the untreated control group (n  7). *P < 0.05.

proportion of neutrophils. In contrast, the relative proportion of monocytes was decreased by DMBA and increased by BP (Table 2). Fig. 4 shows representative dot plots of neutrophils and monocytes from control-, DMBA-, and BP-treated mice. DMBA treatment decreased the total number of bone marrow monocytes and neutrophils by more than two-fold (Fig. 5). However, BP treatment caused an increase in the total number of bone marrow monocytes and a decrease in total number of bone marrow neutrophils that was statistically significant (Fig. 5). Unlike their effects on the lymphoid lineage, BP and DMBA exerted different effects on the myeloid lineage. Fig. 4, bottom right, shows a substantial proportion of Gr-1+ cells that are negative for both 7/4 and 1A8 (Ly-6G) mAbs. These cells are likely to be committed myeloid progenitors (CMPs) and they roughly doubled following either BP or DMBA treatment. The numbers of Gr-1+ cells (7/4 , 1A8 ) not identified as neutrophils or monocytes in Table 2 were 5.3, 8.6, and 12.5% for control, BP, and DMBA treatments, respectively. Because the total bone marrow number of Gr-1+ cells did not decrease following BP treatment, this represents a net increase in the total number of the CMPs. The total number of cells in this population remains about the same after DMBA treatment (Fig. 5).

Table 2 Effect of BP and DMBA treatment on bone marrow myeloid lineage cells Treatment % myeloid % neutrophils % monocytes % myeloid cells cells (Gr-1+) (7/4+/1A8hi) (7/4+/1A8lo) (Gr-1+, 7/4 /1A8 ) Control DMBA BP

59.1 T 1.0 44.7 T 2.0a 58.6 T 2.0

43.9 T 2.2 30.3 T 1.2a 31.9 T 1.6a

9.9 T 0.6 5.8 T 0.3a 14.2 T 0.9a

5.3 8.6 12.5

Note. For each treatment group, n  4; mean T SEM. Values represent the % of cells within the viable cells (see Fig. 2). a Significantly different from the control group (P < 0.05).

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Fig. 4. Flow cytometric analysis of myeloid populations. Based on staining for 7/4 and 1A8, cells were identified as monocytes (7/4+1A8lo) and neutrophils (7/4+1A8hi). Representative dot plots are shown for monocytes and neutrophils of (A) control-, (B) DMBA-, and (C) BP-treated mice.

Hematopoietic stem cells are not depleted following DMBA treatment The PAHs could potentially affect the maturation of hematopoietic stem cells (HSCs) to the committed lymphoid progenitors (CLPs) and committed myeloid progenitors (CMPs), which give rise to the respective lineages (Fig. 1). It has been previously demonstrated that in vivo treatment with 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) results in a sustained increase in HSCs (Lin , Sca1+, and ckit+), within 2 days post-treatment (Murante and Gasiewicz, 2000). HSCs can be identified through their ability to efflux Hoechst 33342 through ATP-dependent transporters. Cells that share this characteristic are referred to as Hoechstlow cells or side population (SP) cells, and represent approximately 0.1% of the total bone marrow cells (Fig. 6A) (Goodell et al., 1996). The proportion of total bone marrow SP fraction was not affected following DMBA treatment (data not shown). The HSCs present in the SP fraction were further resolved using monoclonal antibodies (mAbs) that identify (Lin , Sca1+, and c-kit+) cells (Fig. 6A). About 12% of the SP fraction (0.1% of the total bone marrow cells) from the control mice were viable HSCs (SP, Lin , Sca-1+, and c-kit+), and the SP fraction doubled after 24 h of

Fig. 5. BP and DMBA effects on total bone marrow cells and myeloid populations. Bone marrow cells were collected 48 h post-treatment as described in Fig. 3. Myeloid cells (Gr-1+), monocytes (7/4+1A8lo), and neutrophils (7/4+1A8hi) were analyzed by flow cytometry. Flow cytometry data in Table 2 were multiplied by the number of total bone marrow cells for each mouse with the percentage of each population to obtain the total cells for monocyte and neutrophil populations. Results are expressed as the mean T SEM normalized to the untreated control group (n  4). *P < 0.05.

DMBA treatment (Fig. 6B). The small net increases in total HSCs (Fig. 6C), which parallel the effects reported previously with TCDD, could arise from their diminished differentiation to the CMPs and CLPs (Fig. 1). The increase in Gr-1+ (7/4 , 1A8 ) cells following DMBA treatment suggests that there is no impairment of the maturation of HSCs to CMPs. However, the substantial decrease in pre/ pro-B-lymphocytes suggests that maturation to CLPs may be impaired. CYP1B1 is induced prior to reduction in bone marrow cellularity We previously demonstrated that bone marrow cell depletion and formation of PAHDE-DNA-adducts are dependent on CYP1B1 (Heidel et al., 2000; Galvan et al., 2003). In addition, we previously used fluorescent in situ hybridization to show qualitative increases in CYP1B1 following BP and DMBA treatments (Galvan et al., 2003). These increases are diminished in the bone marrows of Ahr d mice (Galvan et al., 2005). In the present study, we found that BP and DMBA were equally effective in increasing the levels of CYP1B1. There was a four-fold increase in CYP1B1 mRNA at 12 h and a two-fold increase after 24 h (Figs. 7A and B). The similar response to BP and DMBA was consistent with our previously published fluorescence in situ hybridization analysis, but contrasted with liver and lung induction of CYP1A1, and lung induction of CYP1B1, where BP was a more effective inducer (Galvan et al., 2003). This increase in CYP1B1 precedes the onset of bone marrow hypocellularity, which first becomes apparent at about 24 h. Interestingly, CYP1B1 mRNA is expressed constitutively, and is induced by DMBA, in bone marrow fibroblastoid stromal cells, and was observed to be equally distributed among adherent and non-adherent bone marrow cells, indicating expression in multiple cell types (Heidel et al., 1998; Galvan et al., 2003). Expression of AHR mRNA was apparent in bone marrow cells (Fig. 7C), and has previously been identified in bone marrow stroma (Heidel et al., 1998; Lavin et al., 1998). The 30% decrease in AHR at 24 h after treatment with BP or DMBA (Fig. 7C) is unlikely to contribute to CYP1B1 regu-

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Fig. 6. Hematopoietic stem cells are increased following DMBA treatment. (A) The boxed region is the side population (SP) region (left panel), which represents approximately 0.1% of the total bone marrow cells of untreated control mice. HSCs (Lin , Sca-1+, and c-kit+) represent 12% of the viable cells that fall in the SP region. (B) The percentage of HSC (Lin , Sca-1+, and c-kit+) in the SP region increased following 24 h DMBA treatment. (C) Total number of HSC (SP, Lin , Sca1+, and c-kit+) was slightly increased following 24 h DMBA treatment (P > 0.05). HSCs were analyzed by flow cytometry as described in the Materials and methods and these data are expressed as the mean T SEM (n = 2). *P < 0.05.

lation, but may affect cytokines that have been linked to AHR regulation (Dohr and Abel, 1997). TNFa expression is equally stimulated by DMBA and BP We recently reported that DMBA-treated Tnfr-null mice did not exhibit bone marrow hypocellularity, implicating TNFa in the adverse response to DMBA (Page et al., 2004). To further investigate the role of TNFa in PAH toxicity, we measured the expression levels of TNFa and TNFRSF1A in bone marrow cells that were eluted from the bone marrow of

mice at 24 h after DMBA treatment (Fig. 8). This time point was chosen because it is prior to the point at which maximum bone marrow cell depletion is seen following DMBA treatment. We found that TNFa expression levels doubled following both DMBA and BP treatment (Fig. 8A). TNFRSF1A expression also increased modestly in response to DMBA, but not to BP (Fig. 8B). TNFa is expressed in fibroblastic cells, macrophages, and neutrophils (Aggarwal, 2000; Page et al., 2004). We suspected that the modest TNFa response to BP and DMBA might reflect large increase in a small number of responsive cells. We

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is no decrease in total bone marrow cellularity following BP treatment, there are alterations in specific bone marrow cell populations. Our data show that BP and DMBA differ appreciably in their effects on lymphoid and myeloid lineages, but show similar increase in bone marrow cells at an early stage of the response. Although CYP1B1 and TNFa increase following BP and DMBA treatment, it seems likely that they are not limiting factors for the large differences seen in the response of BP and DMBA. The substantial differences in the respective metabolic pathways subsequent to the essential metabolism by CYP1B1 provide a probable source in the observed differences. The larger amounts of DNA adducts formed from DMBA as compared to those from BP demonstrate this difference (Galvan et al., 2005). BP and DMBA exerted similar effects on B-lymphocyte subpopulations, although DMBA had a greater effect that was reflected in the decrease of total number of cells in this lineage. Both BP and DMBA decreased the number of pre/pro-Blymphocytes, without affecting mature B-lymphocytes (Fig. 3). The very small effects of DMBA on the immature Blymphocytes indicate that the progenitor cells are not lost through maturation and a sufficient progression to the immature B-lymphocytes cells is sustained despite the loss of pre/pro-B-lymphocytes. DMBA produced a greater affect on total myeloid cells (Gr1+) than on total lymphoid cells (60% and 40% decrease, respectively) while total cells in these lineages were again unaffected by BP. Furthermore, DMBA substantially decreased the number of neutrophils and monocytes, whereas BP

Fig. 7. BP and DMBA effects on bone marrow expression of CYP1B1 and AHR. Expression of CYP1B1 is greater at 12 h (A) than 24 h (B) posttreatment with either BP or DMBA. (C) BP and DMBA treatment decreases AHR expression in bone marrow. Values shown were normalized to Cyclophilin mRNA expression and presented as the mean T SEM (n = 3). *P < 0.05.

assessed bone marrow cell TNFa expression by treating freshly isolated bone marrow cells with 1 AM DMBA for 5 h in vitro in the presence of Brefeldin A to prevent TNFa secretion (Suresh et al., 2005). Flow cytometry revealed that a small proportion of cells expressed TNFa at 5 –10 times the levels in the bulk of the cells after DMBA treatment (Fig. 9A). We assessed the impact of DMBA by gating the TNFa response to near the maximum level from the untreated bone marrow cells (Fig. 9A). The increase in intracellular TNFa produced by DMBA corresponded to increases above this threshold in about 1.3% of the cells (Fig. 9B). Discussion In this study, we have used flow cytometry to compare the effects of BP and DMBA on bone marrow lymphoid and myeloid populations. We previously showed that DMBA produced an extensive hypocellularity that was not seen with BP (Galvan et al., 2003). Here, we show that even though there

Fig. 8. BP and DMBA effects on bone marrow expression of TNFa and TNFRSF1A. Bone marrow cells were eluted from the femurs and tibias of mice treated 24 h either with BP or DMBA. Expression of TNFa (A) and TNFRSF1A (B) was greater for cells obtained from DMBA than BP treated mice. Values shown were normalized to Cyclophilin mRNA expression and presented as the mean T SEM (n = 3). *P < 0.05.

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Fig. 9. Intracellular staining for TNFa in DMBA-treated bone marrow cells. Bone marrow cells were harvested from the femurs of untreated C57BL/6J mice and then incubated for 5 h with DMBA or vehicle in the presence of Brefeldin A, to prevent protein secretion. The cells were then permeabilized, stained with PE-labeled anti-TNFa mAb, and analyzed by flow cytometry. Panel A illustrates the representative pattern for staining by DMBA-treated cells, and panel B illustrates the mean number of TNFa positive cells for untreated control (n = 2) and DMBA-treated bone marrow cells (n = 5).

decreased the number of neutrophils while producing an increase in the number of monocytes (Fig. 5). Surprisingly, both BP and DMBA appreciably increased the proportion of Gr-1+ (7/4 , 1A8 ), which are likely to be CMPs. Overall, the changes in myeloid populations point to suppression by DMBA in the maturation of CMPs to monocytes and neutrophils, and a selective effect of BP that favors monocytes. BP and DMBA are metabolized by CYP1B1 to generate similar reactive metabolites, but differ in the amounts generated (Pottenger and Jefcoate, 1990; Savas et al., 1997; Shimada et al., 1999; Kleiner et al., 2002). Many of these metabolites also differ in their reactivity and secondary metabolism, which may account for the different effects of BP and DMBA on various bone marrow cell populations. CYP1B1 is found at high constitutive levels in bone marrow stromal cells, and in monocytes and neutrophils (Baron et al., 1998; Heidel et al., 1998; Spencer et al., 1999; Toide et al., 2003). Here, we found that BP and DMBA equally stimulate CYP1B1 mRNA levels at a time when changes in bone marrow cellularity are minimal (Fig. 7A). This finding suggests that CYP1B1 increases precede changes in bone marrow cellularity. The similar response to BP and DMBA is compatible with the CYP1B1 distribution seen in the bone marrow after 24 h treatment (Galvan et al., 2003, 2005). However, the similar response provides further evidence that the CYP1B1 increase is not a

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limiting factor for the response to BP and DMBA or to the differences in DNA adducts formed. We previously came to the conclusion that low levels of CYP1B1 were sufficient for the PAH induced hypocellularity. This was based on the low expression of CYP1B1 in the bone marrow of Ahr d mice, which show enhanced response to BP (Galvan et al., 2003). The similar induction observed in the bone marrow by BP and DMBA contrasts with the larger stimulation by BP of CYP1B1 in the lung and of CYP1A1 in the liver (Galvan et al., 2005). A more complex regulation in bone marrow cells is indicated by the appreciable expression of CYP1B1 in Ahr-null bone marrow fibroblasts (Heidel et al., 1998). Thus, the small decrease in AHR produced by both PAHs is unlikely to affect the induction of CYP1B1 (Fig. 7C). DMBA treatment produced a two-fold increase in the percentage of HSCs (SP, Lin , Sca-1+, and c-kit+) at 24 h (Fig. 6B), at a time when total bone marrow cells decreased by only about 25%. This reflected a small net increase in number of HSCs in response to DMBA (Fig. 6C), which contrast with the very large depletion of the cells in the myeloid and lymphoid lineage following DMBA treatment. As noted above, we have some evidence that common myeloid progenitors (CMPs) increase in response to DMBA in part due to diminished maturation to neutrophils and monocytes. One possible explanation for the modest increase in HSCs is that their commitment to the lymphoid lineage was blocked or slowed. Another possibility is that there is a switch of HSCs from a differentiation to a self-renewal program. Similar to our observations, it was reported previously that TCDD activation of AHR also elicits an increase in HSCs (Lin , Sca-1+, and ckit+) (Murante and Gasiewicz, 2000). In the present study, the effects of DMBA depend on CYP1B1 dependent metabolite formation, rather than on AHR activation per se. These experiments, however, raise the issue of whether HSCs are susceptible targets for chemical disruption of hematopoiesis. These changes in bone marrow cell populations differ from those previously reported for TCDD where AHR activation occurs without metabolism (Thurmond and Gasiewicz, 2000; Thurmond et al., 2000). In contrast to the effects of PAHs noted in the present study, TCDD treatment increased the number of mature B-lymphocytes at 24 h without significantly altering the number of pro/pre, immature, and progenitor B-lymphocytes at this time point. It was suggested that TCDD initially accelerated maturation of immature to mature B-lymphocytes, and subsequently suppressed conversion of B-lymphocyte progenitors to immature B-lymphocytes. The levels of TCDD that produced the decrease in mature B-lymphocytes were relatively high (30 Ag/kg). Because BP is a less potent agonist for the AHR, it is not likely that BP provides sufficient AHR activation to replicate these effects. The need for a high TCDD concentration could be explained by the low AHR levels in mouse B-lymphocytes (Yamaguchi et al., 1997). Hematopoiesis is regulated by a tight network of cytokines, produced by bone marrow stromal cells, that are responsible for the self-renewal, survival, and differentiation of HSCs (Smithgall, 1998). Studies of benzene toxicity have implicated changes in stromal cytokines that regulate hematopoiesis, as

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an important part of benzene’s hematoxicity (Irons and Stillman, 1996). TNFa, which is produced by stromal cells, neutrophils, and macrophages in response to a variety of stimuli, has been implicated in PAH induced bone marrow hypocellularity (Aggarwal, 2000; Page et al., 2004). The majority of the biological effects of TNFa are attributed to signaling through TNFRSF1A (TNFR1), while TNFRSF1B (TNFR2) provides a contributory role in some processes (Engelmann et al., 1990; Heller et al., 1992; Chan and Lenardo, 2000). Recently, our laboratory demonstrated that mice lacking TNFR1A, or both TNFRs, were fully protected against DMBA bone marrow depletion (Page et al., 2004). Here, we have shown that both BP and DMBA appreciably elevated TNFa mRNA. The similar effects of BP and DMBA indicate that TNFa alone does not determine the appreciable differences between the effects of BP and DMBA on bone marrow cellularity. By performing intracellular staining and flow cytometry, we found that TNFa was stimulated by DMBA exposure in only a small fraction of cells (Fig. 9). This finding suggests that TNFa induced by PAH exposure may work locally, and perhaps synergistically with reactive PAH metabolites, to alter normal hematopoiesis and deplete bone marrow cellularity. The similar response in TNFa also contrasts with the large differences between the PAHs with respect to dihydrodiol epoxide generation in bone marrow as indicated by DNA adduct formation (Galvan et al., 2005). BP, unlike DMBA, readily forms BP-quinones through direct and enzymatic oxidation, which have been implicated in the toxic effects on bone marrow cells (Legraverend et al., 1980; Trush et al., 1985). This divergence of responses suggests that TNFa does not arise from this type of chemical stress. The failure of BP to substantially deplete number of myeloid and lymphoid cells, despite alterations in the relative proportions of individual bone marrow cell populations, suggests that BP may exert a diminished effect on HSCs. The differing effects of BP and DMBA on bone marrow suggest that there are multiple targets for toxicity in the complex differentiation processes that regulate hematopoiesis. Furthermore, different bone marrow cell populations may display different susceptibilities to the various PAH metabolites and can work synergistically with TNFa to elicit a selective response. Acknowledgments This work was supported by NIH grant 2R01 CA081493. N.G. was supported by a Comprehensive Minority Biomedical Branch Research Supplement for Underrepresented Minorities and by the Molecular and Environmental Toxicology Training Grant (NIEHS) ES07015. References Aggarwal, B.B., 2000. Tumour necrosis factors receptor associated signalling molecules and their role in activation of apoptosis, JNK and NF-kappaB. Ann. Rheum. Dis. 59, 6 – 16.

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