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TOXICOLOGY
Institut National de la Santé et de la Recherche Médicale U620, Faculté de Pharmacie, Rennes, France (J.v.G., S.L., O.F.); Etablissement Français du Sang, Rennes, France (C.L.B.); and Service d'Hématologie Clinique (M.B.) and Laboratoire d'Hématologie-Immunologie (T.F., O.F.), Hôpital Pontchaillou–Centre Hospitalier Universitaire, Rennes, France
Received February 9, 2005; accepted April 26, 2005.
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-naphthoflavone blocked both BP adduct formation and BP toxicity; and 3) benzo(a)pyrene-trans-7,8-dihydrodiol-9,10-epoxide, a highly reactive BP metabolite, exerted a marked toxicity toward CD34+ cell cultures. Overall, these data indicate that human hematopoietic CD34+ cells can bioactivate chemical carcinogens such as PAHs and, in this way, constitute targets for such carcinogenic environmental contaminants.
; Phillips, 1999). Some of them, including BP, are well recognized as major chemical carcinogens for rodents and humans (Zedeck, 1980). This has been linked to mutagenic properties of PAH metabolites, such as benzo(a)pyrene-trans-7,8-dihydrodiol-9,10-epoxide (BPDE), which form covalent adducts with DNA (Melendez-Colon et al., 1999). The generation of these toxic metabolites is mediated by cytochromes P450 (P450s), especially CYP1A1 and 1B1, whose expression can be markedly up-regulated by PAHs (Whitlock, 1999). This induction of xenobiotic metabolizing enzymes is caused by PAH-mediated activation of the aryl hydrocarbon receptor (AhR), a ligand-activated basic-helix-loop-helix transcription factor. Indeed, PAHs bind to cytoplasmic AhR, thereby triggering its translocation into the nucleus, association with the AhR nuclear translocator (ARNT), and interaction with xenobiotic responsive elements found in the 5'-flanking regions of PAH-regulated genes, including P450s (Whitlock, 1999).
In addition to carcinogenic properties, PAHs exert a wide range of deleterious effects toward tissues and cells. PAHs are thus potent immunotoxic agents that impair functional activation of lymphocytes (Burchiel et al., 1990; Davila et al., 1996) and inhibit macrophagic differentiation (van Grevenynghe et al., 2003). They reduce fertility, which may reflect altered survival of ovocytes (Matikainen et al., 2001), favor the development of cardiovascular diseases (Ambrose and Barua, 2004), and trigger apoptosis in various cell lines (Salas and Burchiel, 1998; Solhaug et al., 2004).
Bone marrow is also affected by PAHs, at least in rodents. Indeed, acute treatment of mice by 7-12 dimethylbenzanthracene (DMBA) results in a profound hypocellularity in the marrow (Galvan et al., 2003; Page et al., 2004). In addition, B-cell development, which takes place in the bone marrow, is markedly altered by PAHs (Mann et al., 1999). Moreover, PAHs are well known chemicals used for inducing experimental leukemias in rodents (Heidel et al., 2000). Together, these data suggest that hematopoietic stem cells may constitute important targets for PAHs. To investigate this hypothesis and to gain insights about the cellular and mechanisms involved, the present study was designed to analyze the effects of PAHs toward normal isolated human CD34+ progenitor cells. Such progenitor cells, characterized by the expression of the immaturity marker CD34, can differentiate into different hematological cell lineages and constitute a suitable in vitro model for analyzing the effects of xenobiotics on hematopoiesis, especially in humans (Bartolovic et al., 2004). Our data indicate that BP, used here as a prototypical PAH, markedly impairs proliferation and differentiation of these hematopoietic stem cells and induces their apoptosis. Moreover, such toxic effects are demonstrated to be related, at least in part, to P450-dependent BP metabolite formation. These data thus provide evidence that human hematopoietic CD34+ cells are capable of bioactivating chemical carcinogens such as PAHs and, in this way, constitute targets for these major environmental contaminants.
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Materials and Methods |
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-naphthoflavone (NF) were provided by Sigma-Aldrich (St. Louis, MO), whereas 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) was from Cambridge Isotope Laboratories, Inc. (Andover, MA). The pure AhR antagonist 3'-methoxy-4'-nitroflavone (3'M4'NF) was a generous gift from Professor Thomas A. Gasiewicz (University of Rochester, Rochester, NY), and BPDE was obtained from Promochem (Molsheim, France). All chemicals were used as stock solutions in dimethyl sulfoxide, except BPDE, which was dissolved in anhydrous tetrahydrofuran (Sigma-Aldrich). Final concentrations of solvent in culture medium did not exceed 0.2% (v/v), and control cultures received the same dose of solvent. Recombinant human stem cell factor, Flt3-ligand, and interleukin-3 were provided by Tebu-bio (Le Perray-en-Yvelines, France) and StemCell Technologies (Meylan, France).
Cellular Culture. Peripheral human CD34+ cells mobilized by granulocyte colony-stimulating factor were purified from samples of blood cytapheresis products using anti-CD34 Ab-coated magnetic beads (Miltenyi Biotec, Paris, France). Purity of isolated CD34+ cellular fractions was routinely greater than 98%, as determined by immunophenotyping analysis. CD34+ cells were then cultured for 3 to 10 days in conditions triggering their proliferation in the absence or presence of PAHs. Such mitogenic culture conditions consisted of Iscove's modified Dulbecco's medium (Invitrogen, Cergy-Pontoise, France) containing 10% fetal bovine serum, 2 mM L-glutamine, antibiotics, and the following cytokines: 20 ng/ml stem cell factor, 50 ng/ml Flt3-ligand, and 50 ng/ml interleukin-3 (Kohler et al., 1999). Initial cellular plating concentration was 5.104 CD34+ cells/ml.
Quantitation of Clonogenic Progenitors in Semisolid Assays. The ability of CD34+ cells to differentiate along the erythroid, granulomacrophagic, and megakaryocytic cell lineages was analyzed using methylcellulose- or collagen-based progenitor assays for colony-forming units (CFU). Briefly, for erythroid and granulomacrophagic lineages, 1000 CD34+ cells/dish were cultured in STEM.IA semisolid medium (Tebu-bio) containing erythropoietin and granulocyte-macrophage colony stimulating factor for 2 weeks at 37°C in a humidified atmosphere with 5% CO2 in the absence or presence of BP. Burst-forming units-erythroid (BFU-E), CFU-granulocyte-macrophage (CFU-GM), and CFU-macrophage (CFU-M) were then scored on an inverted microscope. For megacaryocyte lineage, 5000 CD34+ cells/dish were cultured for 2 weeks in collagen-Megacult-C complete medium (StemCell Technologies) containing 50 ng/ml thrombopoietin. After 14 days of culture, CFU-megacaryocytes (CFU-MK) were immunolabeled with mouse mAb anti-CD41a (BD Biosciences, San Jose, CA) and thereafter visualized using an alkaline phosphatase system (Dako, Trappes, France).
Flow Cytometric Immunolabeling Assays. Cells were labeled with FITC-conjugated mAbs purchased by Immunotech (Marseille, France) and directed against CD34, CD11b, or CD15. Isotypic control labeling was performed in parallel. Cells were then analyzed analysis using a FACSCalibur flow cytometer (BD Biosciences).
Cytotoxicity and Apoptosis Detection. Cellular viability in CD34+ cell cultures was evaluated using the trypan blue dye exclusion test (Eurobio, Les Ullis, France). Assessment of apoptosis was performed using the FITC-conjugated annexin V Kit (Beckman Coulter, Marseille, France) for detection of externalized phosphatidylserine membrane residues of apoptotic cells. Briefly, after washing in phosphate-buffered saline at 4°C, cells were incubated with 2 µl of FITC-annexin V and 5 µl of propidium iodide in Ca2+ binding buffer for 10 min at 4°C and were finally analyzed by flow cytometry.
Caspase 3 Activity Measurement. Caspase 3 activity was determined using the caspase 3 fluorogenic substrate Ac-DEVD-AMC as previously described (Gorman et al., 1999). Briefly, protein cellular lysates (10 µg) were incubated with 50 µM Ac-DEVD-AMC in buffer assay [20 mM PIPES (pH 7.2), 100 mM NaCl, 1 mM EDTA, 0.1% CHAPS, 10% sucrose, and 10 mM dithiothreitol]. Caspase-mediated release of amino-4-methylcoumarin was then monitored continuously over a 30-min period at 37°C using a spectrofluorimeter SpectraMax Gemini (Molecular Devices, Sunnyvale, CA); excitation and emission wavelengths were 380 and 460 nm, respectively. Data were expressed as fluorescence arbitrary units per 30 min.
Measurement of Mitochondrial Membrane Potential (
m). Mitochondrial energization was evaluated through the analysis of mitochondrial retention of the cationic fluorescent dye 3-3'-dihexyl-oxocarbocyanine [DiOC6(3)], which depends on m (Kalbacova et al., 2003). Briefly, cells were first washed with phosphate-buffered saline before incubation with 50 nM DiOC6(3) at 37°C for 30 min. Cells were then washed twice with phosphate-buffered saline, and DiOC6(3)-related fluorescence was analyzed by flow cytometry using a FACSCalibur cytometer. Positive control of mitochondrial potential depolarization was performed in parallel using cyanide p-(trifluoromenthoxy) phenyl hydrazone (Kalbacova et al., 2003).
RNA Isolation and RT-PCR Assay. Total RNA was isolated from cells using the TRIzol method (Invitrogen). Total RNA (1 µg) was reverse-transcribed using the Superscript II reverse transcriptase (Invitrogen), and aliquots of cDNA were subsequently amplified with the PCR Master Mix from Promega (Madison, WI). The gene-specific primers used were as follows: CYP1B1 sense, 5'-AAAGAGGTACAACATCACCT-3'; CYP1B1 antisense, 5'-GTATATTGTTGAAGAGACAG-3'; AhR sense, 5'-CTCATACAACACAGCTTCTCC-3'; AhR antisense, 5'-TACTGAAGCAGAGCTGTGCA-3'; ARNT sense, 5'-ACAGAAAGCCATCTGCTGCC-3'; ARNT antisense, 5'-CGGAACA AGATGACAGCCTAC-3'; GAPDH sense, 5'-TTCACCACCATGGAGAAGGC-3'; and GAPDH antisense, 5'-GGCATGGACTGTGGTCATGA-3'. The primers used for CYP1A1 detection were exactly those used by van Grevenynghe et al. (2003). The analysis of GAPDH mRNA levels, not affected by PAHs, was routinely performed as a control. The numbers of PCR cycles were 32 (for AhR, ARNT, CYP1A1, and CYP1B1 detection) and 26 (for GAPDH detection). PCR products were resolved by electrophoresis on 1% agarose gels and visualized by ethidium bromide.
Western Blotting. Total cell lysates were obtained by incubating cells in a lysis buffer containing 50 mM HEPES, 150 mM NaCl, 1 mM EGTA, 0.1% Tween 20, 10% glycerol, 100 µM phenylmethylsulfonyl fluoride, 10 mM dithiothreitol, 2 µg/ml leupeptin, and 1 µg/ml pepstatin. Protein lysates were then subjected to 12% SDS-polyacrylamide gel electrophoresis under reducing conditions and transferred onto nitrocellulose membranes. After blocking with 5% bovine serum albumin in 50 mM Tris-HCl (pH 7.6), 150 mM NaCl, and 0.1% Tween 20, membranes were incubated overnight at 4°C with the following primary Abs provided by Santa Cruz Biotechnology, Inc. (Santa Cruz, CA): rabbit anti-caspase 3, anti-caspase 9, or mouse anti-HSC70 Abs. Anti-rabbit or anti-mouse peroxidase-conjugated Abs were then used as secondary Abs, and the immunocomplexes formed were detected by autoradiography using the chemiluminescence ECL detection kit (Amersham Biosciences Inc., Orsay, France). Analysis of HSC70 expression, not affected by PAHs, was performed as a control.
Measurement of BP Metabolite Adducts. CD34+ cells, previously exposed to 10 nM TCDD for 24 h to induce CYP1A1/1B1 expression, were treated by 0.1 µg/ml [3H]BP (specific activity, 50 Ci/mmol) (Isobio, Fleurus, Belgium) in the absence or presence of 1 µM NF for 4 h. After two phosphate-buffered saline washes, total proteins and nucleic acids were extracted using a trichloroacetic acid precipitation method. Amounts of tritiated BP metabolites covalently bound to cellular nucleophile macromolecules were then determined by scintillation counting as previously described (Solhaug et al., 2004). Background levels of BP-related radioactivity found in cell-free incubation experiments were subtracted, and data were normalized to amounts of total proteins quantified by the Bradford method using the principle of protein-dye binding (Bradford, 1976).
Statistical Analysis. Data were analyzed with the nonparametric Wilcoxon test or with Student's t test. The level of significance was p < 0.05.
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), CD34+ cells actively proliferated in such culture conditions, yielding approximately an expansion factor of 18-fold after 10 days of culture. This cellular proliferation was associated with partial myeloid differentiation, i.e., increased expression of the myeloid markers CD11b and CD15 and concomitant down-regulation of the relative percentage of CD34+ cells (Fig. 1B). In the presence of 10 µM BP, cell expansion in CD34+ cell cultures was altered; such a BP-mediated effect, although observed for a 3-day treatment, was more pronounced after a 6- or 10-day treatment (Fig. 2A). This inhibitory role of BP toward CD34+ cell expansion was similarly observed for the doses of 1, 5, and 10 µM, whereas a lower BP dose (0.1 µM) had no effect (Fig. 2B). In addition to BP, other PAHs such as DMBA and 3-methylcholanthrene were found to decrease the number of viable cells in CD34+ cell cultures, whereas, by contrast, benzo(e)pyrene and pyrene had no significant effect (Fig. 2C). We then analyzed whether a continuous exposure to BP was required for altering expansion in CD34+ cell culture. For this, isolated CD34+ cells were treated with 10 µM BP for 1, 2, or 3 days, washed, and cultured in BP-free medium until day 10; the number of viable cells was then compared with those found in untreated cultures and in cultures permanently exposed to BP for 10 days. As shown in Fig. 2D, amounts of viable cells were similarly down-regulated regardless of the time of BP treatment (i.e., 1, 2, or 3 days) compared with untreated cultures; they were similar to those found in CD34+ cell cultures continuously exposed to BP for 10 days, suggesting that an initial short 1-, 2-, or 3-day exposure to BP was sufficient to markedly impair CD34+ cell expansion.
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m, as shown by decreased DiOC6(3) labeling (Fig. 4D) and an increased expression of cleaved forms of caspase 9 (Fig. 4C); such events have been reported to represent important steps in numerous mitochondrion-related apoptotic processes (Delhalle et al., 2003).
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; Matikainen et al., 2001), and its cofactor ARNT were found to be expressed in CD34+ cells, as assessed by RT-PCR assays (Fig. 5A). Their levels of mRNA expression were lower, however, than those detected in human hepatocytes, used here as positive controls. BP and TCDD, a very potent agonist of the AhR (Landers and Bunce, 1991), were found to induce the expression of prototypical AhR-regulated genes such as CYP1A1 and CYP1B1 in cultured CD34+ cells (Fig. 5B), indicating that AhR was fully functional in these cells. CYP1B1 mRNAs, however, were constitutively present at low levels in untreated CD34+ cell cultures (Fig. 5B). To determine whether AhR may play a role in PAH toxicity toward CD34+ cells, we then analyzed the effects of cotreatment by 3'M4'NF, used at a 0.1 µM dose, for which this compound acts as a pure AhR antagonist (Lu et al., 1995). As indicated in Fig. 5C, 3'M4'NF was able to partially suppress BP toxicity toward CD34+ cells, as assessed by counting viable cells in CD34+ cell cultures, indicating an involvement of AhR. We then determined whether AhR activation was sufficient to trigger alteration of CD34+ cell proliferation. For this, cells were treated by 10 nM TCDD, a potent AhR ligand fully active in CD34+ cell cultures, as revealed by its inducing effects on CYP1A1/1B1 expression (Fig. 5B). As shown in Fig. 5C, TCDD failed to alter expansion of CD34+ cells, suggesting that activation of AhR is not sufficient by itself for exerting toxicity in CD34+ cell cultures.
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Involvement of P450-Dependent BP Metabolites and Adducts in BP Toxicity toward CD34+ Cell Cultures. A major difference between TCDD and PAHs lies in the fact that the former is poorly metabolized, if at all, whereas the latter can be converted in toxic-reactive metabolites through the action of P450, especially CYP1A1 and CYP1B1, and, in this way, form covalent adducts to cellular macromolecules (Zedeck, 1980). Interestingly, BP-exposed CD34+ cells exhibited up-regulation of CYP1A1 and CYP1B1 expression (Fig. 5B), suggesting that PAHs can be metabolized and form adducts in these cells. To examine this point, the formation of reactive BP adducts was analyzed through measuring covalent binding of radiolabeled metabolites of BP to macromolecules in the presence or absence of the CYP1A1/1B1 inhibitor NF (Chang et al., 1994). As shown in Fig. 6A, BP adducts were detected in CD34+ cells not coexposed to NF. Such formation of BP adducts, however, was greatly reduced in the presence of NF (Fig. 6A), most likely indicating that PAHs such as BP can generate reactive metabolites and adducts in CD34+ cell cultures through their P450-dependent metabolism.
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To further investigate the potential role of P450-related metabolism of PAHs in the toxicity of these environmental contaminants in CD34+ cell cultures, we analyzed the effects of NF on BP-mediated alteration of CD34+ cell culture expansion. NF was able to fully abolish the toxic effects of BP toward cell expansion in 6-day-old CD34+ cell cultures (Fig. 6B). To confirm the involvement of BP metabolites, we then directly added BPDE, one of the major toxic BP metabolites (Zedeck, 1980), to CD34+ cell cultures for 6 days and studied the effects of this reactive compound on cell proliferation. BPDE was found to markedly inhibit cell growth; this effect was dose-dependent, and even very low doses (10–50 nM) exerted significant toxicity (Fig. 6C). In addition, BPDE induced the apoptotic process, as assessed by annexin V labeling (data not shown). Moreover, the antioxidant NAC, already described as capable of preventing the toxicity of BP metabolites, i.e., the formation of BPDE adducts to liver and lung DNA (De Flora et al., 1991; Izzotti et al., 1995), was demonstrated to significantly counteract BP- and BPDE-mediated alteration of cell proliferation in CD34+ cell cultures (Fig. 6D).
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). Moreover, BP triggered apoptosis in CD34+ cell cultures, as shown by annexin V labeling, measurement of caspase 3 activity, detection of cleaved forms of caspase 3 and 9, and the loss of mitochondrial potential. Interestingly, a short exposure to BP (24 or 48 h), starting at the beginning of CD34+ cell cultures, was sufficient to markedly impair proliferation after 10 days of culture, although BP was withdrawn after the initial incubation. Such results therefore provide evidence that CD34+ cells, which represent more than 90% of cultured cells during the first 2 days of culture (data not shown), were targets by themselves for PAHs. An additional toxicity of PAHs toward more differentiated myeloid CD34- cells, which appeared during cell expansion with time in culture, can, however, not be excluded. Indeed, several studies have indicated that differentiated myeloid cells such as monocytes/macrophages (van Grevenynghe et al., 2004) or dendritic cells (Laupeze et al., 2002) constitute targets for PAHs.
AhR seemed to be expressed and fully functional in progenitor cells, as notably demonstrated by the up-regulation of CYP1A1/1B1 expression in BP-treated CD34+ cells. Its activation was required for the deleterious effects of PAHs since 1) the pure AhR antagonist 3'M4'NF partially counteracted BP toxicity and 2) PAHs such as benzo(e)pyrene or pyrene, known to not or only poorly interact with the AhR (Zedeck, 1980), failed to exert toxicity toward CD34+ cell cultures. The lack of deleterious effects of TCDD, however, a very potent ligand of the AhR, clearly demonstrated that activation of the AhR is not sufficient by itself and, therefore, that additional events are required. In this context, we have focused on PAH metabolites since 1) their formation requires prior AhR-dependent up-regulation of CYP1A1/1B1 and BP-exposed CD34+ cells effectively expressing CYP1A1/1B1; 2) they are well known to play a major role in many PAH deleterious effects, especially via formation of adducts to macromolecules (Melendez-Colon et al., 1999); and 3) careful analysis of the effects of the different PAHs used in the present study reveals that only PAHs known to generate toxic-reactive intermediates, i.e., BP, DMBA, and 3-methylcholanthrene (Zedeck, 1980), affected CD34+ cell expansion. Our results fully supported an involvement of PAH metabolites capable of forming cellular DNA and protein adducts. Indeed, BP-related adducts were observed in CD34+ cell cultures, and their formation required P450-dependent metabolism of BP, as illustrated by the inhibitory effects of NF. Importantly, NF was capable of significantly preventing toxic effects of BP toward CD34+ cell expansion, linking formation of adducts and BP toxicity. The fact that 1) BPDE, a major toxic metabolite of BP, was found to directly exert toxicity in CD34+ cell cultures and 2) that NAC, known to counteract PAH metabolite effects by preventing DNA adduct formation (Izzotti et al., 1995), partially inhibited toxicity of both BP and BPDE toward CD34+ cell expansion, also fully favors the idea of a critical role of PAH metabolites and adducts. It is noteworthy that, in addition to CD34+ cells, lymphocytes (Salas and Burchiel, 1998; Mann et al., 1999) and macrophages (van Grevenynghe et al., 2004) have been shown to be targeted by PAH metabolites. This emphasizes the fact that PAH metabolism and adduct formation with prior AhR-dependent CYP1A1/1B1 up-regulation probably represent key steps in PAH toxicity toward these different hematological cells. Interestingly, CD34+ hematopoietic progenitor cells have been previously shown to exhibit slow DNA repair and high sensitivity to DNA-damaging drugs, notably when compared with hematological CD34- cells (Buschfort-Papewalis et al., 2002). Our results fully support this conclusion since CD34+ cells were found to be very sensitive to low doses of the mutagenic agent BPDE.
Apoptosis due to BP exposure in CD34+ cell culture involved executioner caspase 3. It also incriminated mitochondrial events; indeed, m was strongly reduced in response to BP, and caspase 9 was activated. Such a participation of both caspases and mitochondria to PAH-related apoptotic processes has already been described in several cell types undergoing apoptosis in response to PAHs, such as lymphoma, hepatoma, and macrophagic cells (Salas and Burchiel, 1998; van Grevenynghe et al., 2004; Solhaug et al., 2004). In such cells, PAH-induced apoptosis has been linked to altered expression of both pro- and antiapoptotic factors such as bax, p53, c-FLIP, and Bcl-XL (Salas and Burchiel, 1998; Matikainen et al., 2001; Solhaug et al., 2004). Whether these proteins may also be affected by PAH exposure in CD34+ cell cultures will deserve further studies. Another factor linked to apoptosis is the overproduction of reactive oxygen species (ROS) (Delhalle et al., 2003). Using the ROS-sensitive probes 2',7'-dichlorodihydrofluorescein diacetate and dihydroethidium, we failed, however, to demonstrate an increased formation of ROS in BP-treated cultures of CD34+ cells (data not shown). This likely indicates that the antioxidant NAC protected CD34+ cells from BP toxicity not by counteracting ROS overproduction, but rather by down-modulating BP-related adduct formation (Izzotti et al., 1995) as already described above.
The potential contribution of PAH-related deleterious effects toward hematopoietic CD34+ cells to the global toxicity of these ubiquitous environmental contaminants remains to be determined, notably after in vivo exposure. It is noteworthy, however, that previous experimental data in rodents have thoroughly established the bone marrow toxicity of PAHs. Indeed, DMBA treatment of mice resulted in a profound hypocellularity of the bone marrow, with a depletion of various cell lineages, i.e., granulocytes, erythroid precursors, and lymphocytes (Galvan et al., 2003; Page et al., 2004). In rats, BP exposure also triggered bone marrow cytotoxicity, micronuclei formation in erythroid precursors (Shimada et al., 1990), and the development of aplastic anemia (Nebert et al., 1977). These data suggest that hematopoiesis is markedly altered in response to PAHs. Our data, which indicates that CD34+ progenitor cells constitute targets for PAHs, fully support this hypothesis and provide information about the cellular bases of PAH hematotoxicity, i.e., a direct deleterious action on human hematopoietic CD34+ cells. In addition, PAHs are well recognized as immunotoxic agents that have been linked to alteration of the functions of lymphocytes (Davila et al., 1996) and to perturbation of lymphopoiesis (Mann et al., 1999); moreover, differentiation of antigen-presenting cells such as dendritic cells (Laupeze et al., 2002) and macrophages (van Grevenynghe et al., 2003) may be compromised. Toxicity of PAHs toward CD34+ hematopoietic cells may also directly contribute to their immunosuppressive properties by down-regulating the production of cell lineages playing a major role in the immune response, such as lymphocytes, macrophages, and neutrophils.
Our study, which used human cells, suggests that PAH hematoxicity, already known for rodents, may concern humans. The fact that human lymphocytes (Davila et al., 1996), dendritic cells (Laupeze et al., 2002), and macrophages (van Grevenynghe et al., 2003) constitute targets for PAHs favors this hypothesis. Interestingly, BP was still active on human CD34+ cells when used at 0.5 µM (125 ng/g), a relatively low concentration close to PAH amounts found in some foods (up to 50 ng/g) (Lijinsky, 1991; Phillips, 1999) or in tobacco smoke (20–40 ng/cigarette) (Izzotti et al., 1991). This suggests that humans may be exposed to PAH concentrations putatively affecting hematopoietic cells.
A major effect of PAHs on health is to favor the development of cancers, including malignant hemopathies (Mills et al., 1990). Indeed, DMBA elicits leukemia and lymphomas in rodents (Ball, 1970; Heidel et al., 2000), whereas oral exposure to BP in mice results in leukemias (Zhu et al., 1995). In this context, the fact that normal hematopoietic stem cells express AhR-related-inducible metabolizing enzymes such as CYP1A1/1B1 and therefore can generate reactive PAH metabolites when exposed to these environmental chemicals is probably important to consider. Indeed, mutagenic damage to CD34+ bone marrow cells is thought to constitute an important initial event in leukemogenesis (Irons and Stillman, 1996), and that PAHs can cause adducts in CD34+ cells after their P450-dependent metabolism may thus provide a molecular basis for the potential leukemogenic effects of these chemicals. It is noteworthy that CYP1B1, constitutively present in bone marrow cells, may play a major role in PAH bioactivation as recently suggested (Galvan et al., 2003). The fact that low doses of NF (0.1 and 0.2 µM), thought to block CYP1B1 activity unlike that of CYP1A1 (Shimada et al., 1998), remain capable of decreasing BP toxicity toward CD34+ cell proliferation (data not shown) fully supports this conclusion. In addition to CYP1A1/1B1, other AhR-related genes may also be incriminated in PAH-induced leukemogenesis, owing to the fact that some genes controlling cell division and apoptosis are known to be AhR-responsive (Nebert et al., 2000).
In conclusion, we have demonstrated that PAHs, potent carcinogenic and immunotoxic agents, can markedly alter cell expansion and differentiation in human progenitor CD34+ cell cultures. This is associated with induction of apoptosis and most likely depends on AhR-dependent PAH metabolite formation. These data therefore provide evidence that human hematopoietic CD34+ cells are capable of bioactivating chemical carcinogens such as PAHs and, in this way, constitute potent targets for such major environmental contaminants.
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: PAH, polycyclic aromatic hydrocarbon; BP, benzo(a)pyrene; BPDE, benzo(a)pyrene-trans-7,8-dihydrodiol-9,10-epoxide; P450, cytochrome P450; AhR, aryl hydrocarbon receptor; ARNT, AhR nuclear translocator; DMBA, 7-12 dimethylbenzanthracene; NAC, N-acetylcysteine; NF, naphthoflavone; TCDD, 2,3,7,8-tetrachlorodibenzo-p-dioxin; Ab, antibody; mAb, monoclonal antibody; 3'M4'NF, 3'-methoxy-4'-nitroflavone; CFU, colony-forming unit(s); BFU-E, burst-forming units-erythroid; CFU-GM, CFU-granulocyte-macrophage; CFU-M, CFU-macrophage; FITC, fluorescein isothiocyanate; Ac-DEVD-AMC, N-acetyl-Asp-Glu-Val-Asp-amino-4-methylcoumarin; PIPES, 1,4-piperazinediethanesulfonic acid; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid; DiOC6(3), 3-3'-dihexyl-oxocarbocyanine; RT-PCR, reverse transcriptase-polymerase chain reaction; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; ROS, reactive oxygen species.
Address correspondence to: Dr. Olivier Fardel, INSERM U620, Faculté de Pharmacie, 2 avenue du Pr. Léon Bernard, 35043 Rennes, France. E-mail: olivier.fardel{at}univ-rennes1.fr
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