ToxSci Advance Access originally published online on June 8, 2007
Toxicological Sciences 2007 99(1):79-89; doi:10.1093/toxsci/kfm149
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Tumor Necrosis Factor-
Modulates Effects of Aryl Hydrocarbon Receptor Ligands on Cell Proliferation and Expression of Cytochrome P450 Enzymes in Rat Liver "Stem-Like" Cells
ina Zatloukalová*
má
,
,¶ek*,
* Laboratory of Cytokinetics, Institute of Biophysics, 62165 Brno, Czech Republic
Department of Chemistry and Toxicology, Veterinary Research Institute, 62100 Brno, Czech Republic
Graduate Center for Toxicology, University of Kentucky, Lexington, Kentucky 40536-0305
Molecular and Cell Nutrition Laboratory, College of Agriculture
¶ Graduate Center for Nutritional Sciences, University of Kentucky, Lexington, Kentucky 40536-0200
|| Department of Animal Physiology and Immunology, Institute of Experimental Biology, Faculty of Science, Masaryk University, 62100 Brno, Czech Republic
1 To whom correspondence should be addressed at Institute of Biophysics, Královopolská 135, 61265 Brno, Czech Republic. Fax: +42-05-41-21-12-93. E-mail: kozubik{at}ibp.cz.
Received March 2, 2007; accepted May 31, 2007
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUPPLEMENTARY DATAFUNDINGREFERENCES |
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Various liver diseases lead to an extensive inflammatory response and release of a number of proinflammatory cytokines, such as tumor necrosis factor-
(TNF-). This cytokine is known to play a major role in liver regeneration as well as in carcinogenesis. We investigated possible interactions of TNF- with ligands of the aryl hydrocarbon receptor (AhR) and known liver carcinogens, such as 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) and coplanar 3,3',4,4',5-pentachlorobiphenyl (PCB 126). These compounds have been previously found to disrupt cell cycle control in contact-inhibited rat liver WB-F344 cells, an in vitro model of adult liver progenitor cells. TNF- itself had no significant effect on the proliferation/apoptosis ratio in the WB-F344 cell line. However, it significantly potentiated proliferative effects of low picomolar range doses of both TCDD and PCB 126, leading to an increase in cell numbers, as well as an increased percentage of cells entering the S-phase of the cell cycle. The combination of TNF- with low concentrations of AhR ligands increased both messenger RNA (mRNA) and protein levels of cyclin A, a principle cyclin involved in disruption of contact inhibition. TNF- temporarily inhibited AhR-dependent induction of cytochrome P450 1A1 (CYP1A1). In contrast, TNF- significantly enhanced induction of CYP1B1 at both mRNA and protein levels, by a mechanism, which was independent of nuclear factor-
B activation. These results suggest that TNF- can significantly amplify effects of AhR ligands on deregulation of cell proliferation control, as well as on expression of CYP1B1, which is involved in metabolic activation of a number of mutagenic compounds.
Key Words: cell proliferation; tumor necrosis factor-; aryl hydrocarbon receptor; polychlorinated biphenyl; dioxin; xenobiotic metabolizing enzymes.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUPPLEMENTARY DATAFUNDINGREFERENCES |
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The liver cells are a target of various toxic compounds and infections that can lead to inflammatory responses within the liver associated with the production of a large array of inflammatory cytokines, such as interleukin-6 or tumor necrosis factor-
(TNF-) (Budhu and Wang, 2006
). These cytokines are important mediators of liver regeneration, but they can also play a role in the development of hepatocellular carcinoma (Budhu and Wang, 2006; Knight et al., 2005a; Lowes et al., 2003). Both infectious liver disease and lifestyle factors may significantly affect levels of TNF- within the liver (Pessayre et al., 2002). Among other effects, TNF- has been shown to promote proliferation of liver progenitor cells, also known as oval cells (Kirillova et al., 1999). These cells are activated during liver injury, in the event that the regenerative capacity of hepatocytes is compromised. A significant correlation exists between the number of parenchyma-infiltrated oval cells and the degree of lobular inflammation (Libbrecht et al., 2000). The proliferation of oval cell is often observed during the early stages of various liver disease states that may ultimately lead to cancer (Bennoun et al., 1993; Hsia et al., 1992). The oval cells are now considered to be both an important source of liver renewal and a possible target of factors involved in the etiology of hepatocellular carcinoma (Knight et al., 2005b; Roskams, 2006).
Environmental pollutants have been shown to contribute to the development of inflammatory reaction by multiple mechanisms. The aryl hydrocarbon receptor (AhR) agonists, such as coplanar polychlorinated biphenyls (PCBs) and 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD), activate nuclear factor-B (NF-B) in endothelial cells (Hennig et al., 2002; Toborek et al., 1995). AhR ligands have also been reported to increase levels of proinflammatory cytokines both in plasma and in various target organs, including liver (Fan et al., 1997; Wu et al., 2004; Zeytun et al., 2002). Importantly, a recent study of Pande et al. suggested that interleukin 1–like cytokines including TNF- might mediate hepatotoxic effects of dioxin, such as induction of hepatocyte apoptosis (Pande et al., 2005). Therefore, it seems important to describe potential interactions between inflammatory response and AhR activation. AhR is a key mediator of toxic effects induced by a number of organic pollutants (Gu et al., 2000; Van den Berg et al., 1998), which controls transcription of a number of xenobiotic metabolizing enzymes (Nebert and Dalton, 2006; Nebert et al., 2004). Nevertheless, the AhR also seems to be involved in regulation of cell growth, development, and differentiation (Bock and Köhle, 2006; Marlowe and Puga, 2005). Various AhR ligands, including polychlorinated dibenzo-p-dioxins, PCBs, or polycyclic aromatic hydrocarbons, have been shown to induce cell proliferation in contact-inhibited rat liver WB-F344 cells, a frequently used in vitro model of liver oval cells isolated from the liver of an adult male Fischer 344 rat (Chramostová et al., 2004; Dietrich et al., 2002; Tsao et al., 1984; Vondráek et al., 2005). Loss of contact inhibition is associated with abnormal growth and formation of multilayered foci in cell culture, which occurs commonly upon malignant transformation, and has been observed upon exposure to tumor promoters (Dietrich et al., 1997, 2002; Fagotto and Gumbiner, 1996; Oesch et al., 1988). Transactivation of AhR may thus lead to deregulation of cell proliferation in oval cells and contribute to tumor promotion in the liver (Andrysík et al., 2007). However, nothing is currently known about potential interactions of proinflammatory cytokines and AhR in deregulation of contact inhibition.
There is a substantial evidence for a mutual cross-talk between inflammatory mediators and AhR ligands. Expression of CYP1 family of enzymes is modulated by a variety of factors such as diet, hormone levels, or inflammation, which cause significant changes in the expression levels of CYPs in the liver (Morgan, 1997, 2001). In some reports, inflammation has been shown to suppress cellular responses to AhR activation, whereas other studies have suggested that inflammatory mediators may actually potentiate expression of some AhR target genes. For example proinflammatory cytokines, such as TNF-, or lipopolysaccharides suppress the expression of CYP1A1 (Barker et al., 1992; Muntane-Relat et al., 1995), which may be due to NF-B activation. A mutual inhibitory interaction between the AhR and NF-B signaling pathway has been indeed demonstrated (Ke et al., 2001; Tian et al., 1999). In a marked contrast, TNF- has been shown to increase expression of CYP1B1 in hepatic stellate cells (Piscaglia et al., 1999). This enzyme is involved in metabolic activation of a number of strong mutagens, such as benzo[a]pyrene or aflatoxin B1, the latter being an important factor in hepatocellular carcinoma etiology (Crespi et al., 1997; Kondraganti et al., 2003). Therefore, TNF- may both suppress and potentiate AhR response, based on the nature of its gene target.
Taken together, the available evidence suggests that the effects of AhR ligands could be significantly modulated under inflammatory conditions, which have been shown to contribute to liver carcinogenesis. This might be related both to the modulation of expression of CYP1 enzymes (involved in tumor initiation stage) and deregulation of cell proliferation (contributing to tumor promotion). However, nothing is currently known about possible interactions between proinflammatory cytokines and AhR ligands in liver progenitor cells. In the present study, we used rat liver epithelial cell line WB-F344 to study combined effects of model AhR ligands and proinflammatory cytokine (TNF-) on regulation of cell proliferation and expression of CYP1A1/1B1 enzymes. Our data also provide evidence that AhR ligand–induced deregulation of cell proliferation is markedly amplified in the presence of TNF-. This may have implications in understanding mechanisms of carcinogenesis induced by exposure to environmental pollutants.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUPPLEMENTARY DATAFUNDINGREFERENCES |
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Chemicals.
TCDD was purchased from Cambridge Isotope Laboratories (Andover, MA). 3,3'4,4',5-Pentachlorobiphenyl (PCB 126) was purchased from Promochem (Wesel, Germany). Stock solution was prepared in dimethyl sulfoxide (DMSO) (Merck, Darmstadt, Germany) and stored in the dark. Recombinant rat TNF-
(Sigma-Aldrich, Prague, Czech Republic) was dissolved in phosphate-buffered saline (PBS). Goat polyclonal antibody against CYP1A1 and rabbit polyclonal antibody against CYP1B1 were obtained from BD Biosciences (San Jose, CA). Rabbit polyclonal antibody to cyclin A and mouse monoclonal antibody to p65/RelA were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Mouse monoclonal antibody to ß-actin was obtained from Sigma-Aldrich. All other chemicals used were purchased from Sigma-Aldrich.
Cells.
WB-F344 rat liver epithelial cells (Tsao et al., 1984) were cultured in with Dulbecco's modified Eagle's (DMEM)/F12 Medium (Invitrogen, Carlsbad, CA), supplemented with 15mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid, L-Glutamine, penicillin/streptomycin (100,000 units/l and 100 mg/l, respectively), and 5% heat-inactivated fetal bovine serum. Only the cells at passage levels 15–22 were used for the study. Cells were incubated in a humidified atmosphere of 5% CO2 at 37°C. All tissue culture reagents were obtained from Sigma-Aldrich.
Assessment of cell proliferation and cell cycle distribution.
Cell proliferation and cell cycle assessment were performed as described previously (Chramostová et al., 2004). Briefly, cells were seeded at concentration of 30,000 cells/cm2 in 35-mm-diameter cell culture dishes (TPP, Trasadingen, Switzerland), and grown for 72 h to reach confluency. Afterward, cells were incubated with TNF- (20 ng/ml) or with TNF- in combination with TCDD or PCB 126. Following the exposure, medium was removed, cells were harvested by trypsinization and counted with a Coulter Counter (Model ZM, Coulter Electronics, Luton, UK). Cells were then washed with PBS and fixed in 70% ethanol at 4°C overnight. For cell cycle analysis, the fixed cells were washed with PBS and resuspended in 0.5 ml Vindelov solution (1M Tris–HCl, pH 8.0; 0.1% Triton-X100, vol/vol; 10mM NaCl; propidium iodide [PI], 50 µg/ml RNAse A 50 Kunitz units/ml) (Vindelov, 1977) and incubated in 37°C for 30 min. Cells were then analyzed by FACSCalibur, using a 488-nm (15 mW) air-cooled argon-ion laser for PI excitation, and CELLQuest software for data acquisition (Becton Dickinson, San Jose, CA). A minimum of 15,000 events were collected per sample. Data were analyzed using ModFit LT version 2.0 software (Verity Software House, Topsham, ME).
Detection of apoptosis.
The apoptosis was detected on the basis of morphological criteria (fragmentation of nuclei) as described in (Chramostová et al., 2004). The fixed cells were stained with DAPI (1 µg/ml MetOH final concentration) for 30 min at room temperature. After the incubation, the cells were centrifuged and mounted with MOWIOL solution (10% Mowiol 4-88 was prepared in 25% glycerol, 100mM Tris–HCl, pH 8.5). The slides were evaluated with a fluorescence microscope Olympus IX70 (Tokyo, Japan). A minimum of 200 cells were counted per sample.
Western blot analyses.
Cells were grown on 35-mm-diameter cell culture dishes and incubated with vehicle control (DMSO, 0.1%), TNF- and selected concentrations of TCDD and PCB 126 for 6 and 24 h. Cells were washed with PBS and lysed using 1% sodium dodecyl sulfate (SDS) lysis buffer (10% glycerol, 100mM Tris, pH 7.4) and protein concentration was estimated using Bio-Rad DC Protein Assay (Bio-Rad Laboratories, Inc., Hercules, CA). Equal amounts of protein were separated on 10% SDS-polyacrylamide gel and transferred onto a polyvinyldifluoride membrane. After incubation with primary and secondary antibodies, detection was performed using ECLPlus Western blotting detection system (GE Healthcare, Little Chalfont, UK). Densitometry was performed using AIDA Image Analyzer software (raytest Isotopenmeßgeräte, Starubenhardt, Germany).
Preparation of nuclear extracts and electrophoretic mobility shift assay.
Cells were grown on 60-mm-diameter cell culture dishes and treated with DMSO (0.1%), TCDD (5nM), or TNF- (20 ng/ml) 30 min or 3 h prior to harvesting. Nuclear extracts containing active proteins were prepared from cells according to the method of Dignam et al. (1983). Briefly, the cells were suspended in 1 ml of lysis buffer (10mM Tris–HCl, pH 8.0, 60mM KCl, 1mM ethylenediaminetetraacetic acid [EDTA], 1mM dithiothreitol, 100µM phenylmethylsulfonyl fluoride, 0.1% NP-40), lysed for 10 min on ice, and centrifuged at 600 x g for 4 min at 4°C to collect nuclei. Then, the nuclear pellets were rinsed with 1 ml of lysis buffer without NP-40, lysed 25 min in 50 µl of nuclear extract buffer (20mM Tris–HCl, pH 8.0, 420mM NaCl, 1.5mM MgCl2, 0.2mM EDTA, 25% glycerol) for 10 min on ice, and centrifuged at 12,000 x g for 15 min at 4°C. Nuclear extracts were collected and stored frozen in aliquots at 80°C. The amount of proteins in nuclear extracts was quantified using the Bradford assay. Binding reactions were performed as described in Eum et al. (2006), with minor modifications. DNA probes containing AhR (Generi Biotech, Hradec Králové, Czech Republic) and NF-B consensus oligonucleotides (Promega, Madison, WI) were labeled with 32P-
-adenosine triphosphate by T4-polynucleotide kinase (Promega) and purified by Mini QuickSpin columns (Roche Diagnostics, Mannheim, Germany), according to the manufacturer's instruction. Dioxin response element (DRE) sequences were published previously (Reiners et al., 1998). Binding reactions were performed in a 20-µl volume containing 4 µg of nuclear protein extracts, 10mM Tris–HCl, pH 7.5, 50mM NaCl, 1mM EDTA, 0.1mM dithiothreitol, 1M glycerol, and 0.5 µg of poly[dI-dC] (nonspecific competitor), and incubated at room temperature for 10 min. A 32P-labeled specific oligonucleotide probe (40,000 cpm) was added to the reaction and incubated for 20 min at room temperature. Following binding, the protein–DNA complexes were resolved by electrophoresis in a 6.5% (wt:vol) nondenaturing polyacrylamide gel and visualized by autoradiography.
Real-time reverse transcription-PCR.
The levels of CYP1A1, CYP1B1, and cyclin A messenger RNAs (mRNAs) were determined by real-time reverse transcription-PCR. The primers were designed to flank the exon junctions of the transcripts for amplification of complementary DNA only. The sequences of primers and probes for rat CYP1A1, CYP1B1, cyclin A2, and the porphobilinogen deaminase have been published previously (Vondráek et al., 2006; Zatloukalová et al., 2007). All probes were labeled with the fluorescent reporter dye 6-carboxyfluorescein on the 5'-end, and with the Black Hole 1 fluorescent quencher dye on the 3'-end. Total RNA was isolated from cells using the NucleoSpin RNA II purification kit (BD Biosciences). The reverse transcription and amplifications of the samples were carried out using QuantiTect Probe RT-PCR Kit (Qiagen, Valencia, CA) with final volume of 20 µl in reaction mixture containing 10 µl of QuantiTect Probe RT-PCR Master Mix, 0.2 µl of QuantiTect RT Mix (Qiaqen), 2 µl of solution of primers and probes, 5.8 µl of RNAse-free water, and 2 µl of template RNA. The amplifications were run on the Rotor-Gene 6000 (Corbett Research, Sydney, Australia) using the following program: reverse transcription at 50°C for 30 min and initial activation step at 95°C for 15 min, followed by 35 cycles of 95°C for 10 s and 60°C for 40 s.
Cell transfections with short interfering RNA.
Cells were plated at a density of 20,000/cm2 in 24-well plates in medium without antibiotics. After 24 h cultivation, transfections were performed, using the previously described short interfering RNA (siRNA) duplex directed against rat p65 mRNA sequence (sense, 5'-CUCAGAGUUUCAGCAGCUUdTdT-3'; antisense, 5'-AAGCUGCUGAAACUCUGAGdTdT-3') (Tao et al., 2006) or control siRNA directed against mRNA encoding the red fluorescence protein DsRed (Andrysík et al., 2007). Both specific and nonspecific siRNAs were provided by Ambion (Foster City, CA). The transfections were carried out in a total volume of 600 µl containing with 200 pmol siRNA and 1 µl of Lipofectamine 2000 (Invitrogene, Carlsbad, CA), according to the manufacturer's instructions. Transfection mix was removed 24 h later and cells were cultivated for another 24 h in DMEM/F12 medium with antibiotics, followed by exposure to TNF- (20 ng/ml) for 48 h for detection of apoptosis. Alternatively, transfected cells were exposed to TNF- (20 ng/ml) and/or 5nM TCDD for 6 h for detection of CYP1B1 expression.
Statistical analysis.
Data were expressed as means ± SD for at least three independent replications. Comparisons between treatments were made by one-way analysis of variance (ANOVA) with post hoc comparisons of the means made by Tukey range test. If the variances were nonhomogeneous, nonparametric Mann–Whitney U-test or Kruskal–Wallis ANOVA were used. A p value of less than 0.05 was considered significant.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUPPLEMENTARY DATAFUNDINGREFERENCES |
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Effects of TNF-
on Cell Proliferation/Apoptosis and Activation of NF-B in WB-F344 CellsSome studies (Yao et al., 2004
) suggest that high concentrations of TNF- can induce apoptosis in WB-F344 cells when combined with transcription inhibition, but on the other hand, TNF- has been shown to promote proliferation of liver progenitor cells (Kirillova et al., 1999; Nagy et al., 1998). Therefore, we first investigated the impact of TNF- on cell proliferation and apoptosis in WB-F344 cells. TNF- had no effect on either cell proliferation or apoptosis of confluent WB-F344 cells within the concentration range 1–50 ng/ml (data are presented as a supplementary material—Fig. S1). Similar to these results, we observed no effects in exponentially growing cells treated with the same concentrations of TNF- (data not shown). Since the lack of apoptotic effects of TNF- might be explained by activation of prosurvival transcription factor NF-B through TNF receptors, we determined the binding of NF-B to consensus oligonucleotide containing NF-B–binding site by electrophoretic mobility shift assay (EMSA). TNF- (20 ng/ml) induced both rapid (30 min) and sustained (3 h) activation of NF-B, while the model AhR ligand TCDD had no effect on its activation (Fig. 1A). Application of TNF- had no effect on AhR-DRE binding either at 30 min (Fig. 1B) or after 3 h (data not shown). Since the above data suggested that the lack of apoptosis induction upon TNF- treatment might be due to high level of NF-B activation, we next investigated effects of downregulation of p65/RelA subunit of this transcription factor on induction of apoptosis by TNF-, using RNA interference. As outlined in Figure 2, downregulation of p65 significantly increased percentage of apoptotic cells with fragmented nuclei upon TNF- treatment. These results confirmed that the lack of apoptotic effects of TNF- in WB-F344 cells is probably due to NF-B activation.
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TNF-
Potentiates Proliferation Induced by TCDD and PCB 126In our previous study, we found that AhR-activating PCBs, as well as their hydroxylated metabolites, induce cell proliferation in WB-F344 in a manner similar to the prototypical AhR ligand TCDD (Dietrich et al., 2002
; Vondráek et al., 2005). Therefore, we next investigated the impact of TNF- (20 ng/ml) on effects of low (picomolar range) concentrations of both TCDD and PCB126 on cell proliferation in contact-inhibited WB-F344 cells. The results were compared with effects of a high dose of TCDD that induces a maximal proliferative effect in WB-F344 cells (Chramostová et al., 2004). TNF- significantly increased cell numbers after 72 h of cotreatment with TCDD (10 pM) and PCB126 (100 pM) (Fig. 3). At higher doses, both compounds already induced significant proliferation themselves, without addition of TNF-. In order to confirm these data, we determined cell cycle distribution of cells treated with both AhR agonists alone or in combination with TNF-. TNF- significantly increased the percentage of S-phase cells after 24 h of incubation with TCDD (10 and 100 pM) or PCB 126 (10 pM) (Fig. 4). These data suggested that TNF- can significantly potentiate proliferative effects of both TCDD and PCB 126 in rat liver "stem-like" cells.
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Previous studies have suggested that cyclin A is downregulated during contact inhibition (Dietrich et al., 2002
; Yoshizumi et al., 1995) and that both TCDD and dioxin-like PCBs can increase cyclin A protein levels leading to enhanced cyclin A/cdk 2 activity and cell proliferation (Dietrich et al., 2002; Vondráek et al., 2005). Therefore, we next investigated, whether TNF- affects levels of cyclin A in WB-F344 cells treated with low concentrations of TCDD and PCB 126. As outlined in Figure 5, TNF- itself had no effect on either mRNA (Fig. 5A) or protein (Fig. 5B) levels of cyclin A. In contrast, it increased levels of cyclin A in combination with TCDD (10pM) or PCB 126 (100pM) to the levels comparable with maximum effective concentrations of either compound alone (Fig. 5B). The maximum effective concentrations were based on the results of previous studies (Chramostová et al., 2004; Vondráek et al., 2005).
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TNF-
Transiently Represses Induction of CYP1A1 and Potentiates Induction of CYP1B1 in WB-F344 CellsExpression of CYP1A1 is downregulated by a number of inflammatory mediators including TNF-
, and cross-talk between AhR and NF-B is thought to play a significant role in this effect (Tian et al., 2002). In order to confirm that this mechanism is effective also in WB-F344 cells, the impact of TNF- on CYP1A1 induction by TCDD (at a maximum effective concentration of 5nM) was investigated. It was observed that after 6 h of treatment, TNF- partially inhibited induction of both mRNA (Fig. 6A) and protein (Fig. 6B) levels of CYP1A1. This effect was less pronounced after 24 h treatment (Figs. 6A and 6B) and it disappeared after 48 h, probably due to TNF- degradation (data not shown). In contrast, TNF- significantly potentiated the TCDD-induced expression of CYP1B1 in WB-F344 cells at both mRNA (Fig. 7A) and protein (Fig. 7B) levels already after 6 h of treatment, and this induction was sustained for up to 24 h (Fig. 7). Because these effects were observed using a relatively high dose of TCDD, we next investigated effects of TNF- on CYP1A1 and 1B1 protein expression in combination with low concentrations of TCDD (10pM) and PCB 126 (100pM). TNF- markedly potentiated expression of CYP1B1, while the low doses of TCDD and PCB 126 used for cell proliferation experiments were not sufficient to induce detectable CYP1A1 protein expression (Fig. 8). In order to find out, whether the cross-talk between AhR agonists and TNF- leading to enhanced expression of CYP1B1 occurs at the level of NF-B activation, we used siRNA targeted against p65 subunit of NF-B in a similar manner as in experiments depicted in Figure 2. As outlined in Figure 9, siRNA targeted against p65 efficiently downregulated levels of p65 protein. However, no effects on synergistic effects of TNF- and TCDD on CYP1B1 expression were observed. TNF- had similar effects on CYP1B1 expression in cells transfected with siRNA against p65 as in control cells or in cells transfected with nonspecific siRNA. These results suggested that the cross-talk between AhR agonists and TNF- that lead to synergistic effects on CYP1B1 expression probably did not occur at the level of NF-B activation.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUPPLEMENTARY DATAFUNDINGREFERENCES |
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TNF-
is one of the principal inflammatory mediators, which have been implicated both in the liver injury and its regeneration (Budhu and Wang, 2006; Knight et al., 2005a; Lowes et al., 2003). It has been shown to promote proliferation of liver progenitor cells, known as oval cells (Kirillova et al., 1999), which are now considered to be a possible target of carcinogens (Knight et al., 2005b; Roskams, 2006). We have previously successfully used the WB-F344 cell line as an in vitro model of liver "stem-like" cells to study effects of PCBs and other environmental pollutants on intercellular communication and disruption of cell cycle control. One of the interesting findings was that unlike in mature liver cells, AhR ligands exert proliferative effects in this cell line, which might result in tumor promotion (Chramostová et al., 2004; Dietrich et al., 2002; Vondráek et al., 2005). The effects of AhR ligands could be significantly modulated under inflammatory conditions, which have been shown to contribute to liver carcinogenesis. However, currently there is no information on possible interactions between proinflammatory cytokines and AhR ligands in liver progenitor cells. Therefore, we investigated combined effects of model AhR ligands and TNF- on regulation of cell proliferation and expression of CYP1A1/1B1 enzymes in WB-F344 cell line.
In our experimental design, TNF- itself did not affect the proliferation/apoptosis ratio in WB-F344 cells. TNF- exerts a variety of effects that are mediated mainly by TNF-R1, resulting in induction of cell death. Simultaneously, TNF-/TNF-R signaling can lead to activation of the NF-B pathway that can inhibit the TNF-–induced cell death process (Ding and Yin, 2004) through control of the expression of antiapoptotic genes, such as the A20 zinc finger protein or manganese superoxide dismutase (Baichwal and Baeuerle, 1997). Activation of the NF-B pathway seems to be also important in proliferative effects (Hinz et al., 1999; Yao et al., 2004). The lack of proliferative effects of TNF- in WB-F344 cells might be due to the fact that additional cytokines and growth factors are necessary for activation and proliferation of oval cells observed in vivo (Knight et al., 2005b; Lowes et al., 2003). In the present study, TNF- induced high NF-B binding to its DNA response element, suggesting that NF-B activation might suppress apoptotic effects of TNF- in WB-F344 cells. The antiapoptotic effects of the NF-B pathway activation on TNF-–induced cell death were confirmed also in the present study, using the siRNA targeted against p65 subunit of NF-B. On the other hand, activation of NF-B in WB-F344 cells did not lead to cell proliferation, which has been reported previously in rat liver progenitor cells growth-arrested by serum deprivation (Kirillova et al., 1999).
PCBs are environmental pollutants that act as complete carcinogens in the liver (Safe, 1994; Silberhorn et al., 1990). The biological effects induced by PCBs depend on substitution pattern. The non-ortho–substituted coplanar PCBs evoke similar biological responses as TCDD and their principle mode of action is associated with the activation of AhR (Van den Berg et al., 1998). It has been shown that both TCDD and PCB 126 induce a release of WB-F344 cells from contact inhibition (Dietrich et al., 2002; Vondráek et al., 2005). TCDD was found to increase both liver expression (Fan et al., 1997) and systemic levels of TNF- (Wu et al., 2004). Importantly, interleukin 1–like cytokines such as TNF-, might mediate hepatotoxic effects of dioxin, such as induction of hepatocyte apoptosis (Pande et al., 2005). However, effects of TNF- might be significantly different in liver progenitor cells. We therefore studied the effect of TNF- on modification of proliferation induced by TCDD and PCB 126. TNF- significantly increased both cell numbers and the percentage of cells entering S-phase of the cell cycle in samples treated with low picomolar concentrations of AhR ligands. Under the same experimental conditions, TNF- also significantly potentiated expression of cyclin A, a principle regulator of G1-S phase transition and S-phase progression in contact-inhibited WB-F344 cells (Andrysík et al., 2007; Dietrich et al., 2002; Vondráek et al., 2005). These results demonstrated that inflammatory cytokines (e.g., TNF-) could augment cell cycle deregulation induced by AhR ligands.
The biological effects of TCDD and coplanar PCBs include induction of cytochrome P450 (CYP1A1/1B1) and other AhR-regulated genes. The cytochromes P450 1A1 and 1B1 are members of cytochromes P450 monooxygenase superfamily and are important during xenobiotic metabolism as well activation of precarcinogens (Nebert and Dalton, 2006; Nebert et al., 2004). It has been shown that proinflammatory cytokines suppress the expression of CYP1A1 in primary hepatocytes (Barker et al., 1992; Muntane-Relat et al., 1995). In the present study, we found that in model rat liver progenitor cells isolated from adult male rat, both AhR agonists–induced CYP1A1 mRNA and CYP1A1 protein levels are partially suppressed by TNF-. One possible explanation for this phenomenon is mutual functional repression between AhR and NF-B (Tian et al., 1999), which leads to inhibition of AhR-induced histone H4 acetylation at the promoter region of cyp1a1 (Ke et al., 2001). This corresponds with rapid activation of NF-B by TNF- observed in WB-F344 cells and indicates that a similar mechanism might be responsible for downregulation of CYP1A1 expression both in mature liver cells and in their progenitors.
In a marked contrast, the expression of CYP1B1 of WB-F344 cells treated with both high and low doses of TCDD or PCB 126 was significantly potentiated by TNF-. It has been established that there are at least two types of regulation for rat CYP1B1: hormonal regulation in steroidogenic tissues (Otto et al., 1991, 1992), where the constitutive expression predominates (Bhattacharyya et al., 1995); and AhR-dependent induction in liver, lung, and kidney (Bhattacharyya et al., 1995), where it can be upregulated by environmental pollutants. It has been demonstrated previously that TNF- potentiates CYP1B1 expression induced by 7,12-dimethylbenz[a]anthracene (DMBA), a model carcinogen, in hepatic stellate cells but not in mature hepatocytes (Piscaglia et al., 1999). This would suggest that mechanisms responsible for enhanced CYP1B1 expression are cell-type specific. The reason of the increased expression of CYP1B1 levels after TNF- cotreatment in WB-F344 cells, incubated with TCDD or PCB 126, could be the existence of other signaling pathways responsible for activation of CYP1B1 in liver epithelial "stem-like" cells, and analogous to those described in steroidogenic tissues (Otto et al., 1991, 1992; Zheng et al., 2003). Our data suggest that synergistic effects of TNF- cotreatment on CYP1B1 regulation might be independent of the NF-B activation (Fig. 9).
Importantly, CYP1B1 has been shown to participate in metabolic activation of some liver toxins, e.g., aflatoxin B1, which has been implicated in the etiology of hepatocelluar carcinoma (Crespi et al., 1997). Although CYP1A1 and CYP1B1 have partially overlapping substrate specificities (Shimada and Fujii-Kuriyama, 2004), there is a unique function for CYP1B1 emerging in carcinogenesis. In fact, CYP1B1–/– mice are protected from DMBA (Buters et al., 1999) and dibenzo[a,l]pyrene-induced carcinogenesis. One role CYP1B1 plays in tumor development is illustrated by the fact that increased expression was observed in a number of malignant tumors compared to corresponding normal tissues (Murray et al., 1997). In fact, CYP1B1 is considered as a target of some novel anticancer therapies (McFadyen and Murray, 2005). Thus, in addition to proliferative effects associated with tumor promotion, TNF- might also contribute to cancer development through potentiation of induction of CYP1B1 expression.
In conclusion, we studied the effects of model AhR ligands in combination with a principal inflammatory cytokine involved in liver regeneration and hepatocellular carcinoma development, using an in vitro model of liver progenitor cells. Our data suggest that TNF- significantly potentiates the AhR-dependent release of WB-F344 cells from contact inhibition, as well as expression of CYP1B1, a principal enzyme involved in metabolic activation of many promutagens. Proinflammatory cytokines may thus significantly influenced the effects of rodent liver carcinogens by modulation of the expression of the CYP1B1 enzyme (involved in tumor initiation) and deregulation of cell proliferation (contributing to tumor promotion). These data are significant in understanding mechanisms of carcinogenesis. Chronic and low level inflammation is associated with the etiology of numerous diseases, including cancer. Thus, apparently noneffective low doses of environmental toxins may be markedly procarcinogenic during an inflammatory response. This also suggests that the use of in vitro models may in fact underestimate toxicity of some xenobiotics. Thus, harmful effects of xenobiotics could be exacerbated under in vivo conditions, especially during coexposure to environmental factors producing inflammation. Further studies are warranted to understand exact mechanisms responsible for these important interactive biological events.
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SUPPLEMENTARY DATA |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONSUPPLEMENTARY DATAFUNDINGREFERENCES |
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Supplementary data are available online at http://toxsci.oxfordjournals.org/.
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FUNDING |
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Czech Science Foundation (grant no. 524/05/0595); National Institutes of Health/National Institute of Environmental Health Sciences (P42 ES007380); Academy of Sciences of the Czech Republic (Research Plans AV0Z50040507 and AV0Z50040702); and the Czech Ministry of Agriculture (MZE0002716201).
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ACKNOWLEDGMENTS |
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The authors thank Beth Oesterling for the comments on this manuscript.
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REFERENCES |
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