* Department of Biology, University of Waterloo, Ontario, Canada N2L 3G1, and Department of Biomedical Sciences, Ontario Veterinary College, University of Guelph, Guelph, Ontario, Canada N1G 2W1
1 To whom correspondence should be addressed at Department of Biology, University of Waterloo, Waterloo, Ontario, N2L 3G1 Canada. Fax: (519) 746-0614. E-mail: mvijayan{at}uwaterloo.ca.
Received November 9, 2004; accepted December 15, 2004
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ABSTRACT |
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Key Words: arylhydrocarbon receptor; -naphthoflavone; ß-naphthoflavone; fish; salmonid; CYP1A1; cortisol; stress.
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INTRODUCTION |
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The main proteins involved in the corticosteroidogenic pathway comprise steroidogenic acute regulatory (StAR) protein, cytochrome P450 cholesterol side chain cleavage (P450scc/ CYP11A), 17-hydroxylase, 3ß-hydroxysteroid dehydrogenase, and 21- and 11ß-hydroxylases (Payne and Hales, 2004
). The rate-limiting steps in steroidogenesis are the StAR protein, translocating cholesterol from outer to inner mitochondrial membrane, and P450scc enzyme, catalyzing the conversion of cholesterol to pregnenolone (Payne and Hales, 2004
; Stocco and Clark, 1996
). These two key steps in steroidogenesis are impacted by xenobiotics (Sugawara et al., 2001
; Walsh and Stocco, 1998
, 2000
; Walsh et al., 2000a
,b
,c
), including AhR agonists (Dasmahapatra et al., 2000
; Fukuzawa et al., 2004
; Moore et al., 1991
). Other enzyme pathways, downstream of P450scc, appear less sensitive to toxicant insult (Walsh et al., 2000b
), perhaps because these proteins are abundant with longer half-life and retain normal steroidogenic capacity even in the absence of trophic hormone stimulation (Payne and O'Shaughnessy, 1996
).
Most studies on xenobiotic impact on steroidogenesis have focused on reproductive steroids, but very little is known about the mechanism of action of toxicants on corticosteroidogenesis. As corticosteroids play an important role in the homeostatic process, including growth, metabolism, ion and water balance, reproduction, and immune function, any impact on this endocrine axis could potentially affect animal performance (Sapolsky et al., 2000). Unlike the situation in mammals, in teleost fishes the steroidogenic cells of the adrenal cortex are distributed around the posterior cardinal veins of the head kidney region (the interrenal tissue) (Wendelaar Bonga, 1997
). However, as in mammals, the steroid secretion by the interrenal tissue is under the control of the hypothalamus-pituitary axis (Mommsen et al., 1999
; Wendelaar Bonga, 1997
). Cortisol is the main corticosteroid produced by these steroidogenic cells in response to pituitary secretions, of which adrenocorticotropic hormone (ACTH) is the major secretagogue (Wendelaar Bonga, 1997
). ACTH secretion in turn is under the control of corticosteroid releasing factor (CRF) from the hypothalamus (Huising et al., 2004
; Pepels et al., 2004
), and this axis is tightly regulated by a negative feedback loop, including glucocorticoid receptor (GR) signaling in the brain (Aluru et al., 2004
; Bernier and Peter, 2001
). Indeed the hypothalamus-pituitary-interrenal axis is impacted by xenobiotics, including PCBs and other AhR agonists, but the mechanism of action remains unknown (Aluru and Vijayan, 2004
; Aluru et al., 2004
; Hontela, 1998
, 2005
; Vijayan et al., 2005
; Wilson et al., 1998
).
In this study we specifically asked the question whether the AhR-mediated suppression of cortisol production seen before in trout (Aluru and Vijayan, 2004; Wilson et al., 1998
) involves changes in the transcription levels of StAR, P450scc, and 11ß-hydroxylase. To this end, we fed fish AhR agonist, ß-naphthoflavone (BNF), and antagonist,
-naphthoflavone (ANF), either singly or in combination for 5 days to tease out the mechanisms involved in AhR-mediated suppression of cortisol biosynthesis. In addition, the steroidogenic capacity of the interrenal tissue was assessed in vitro using radiolabelled steroid substrate ([7-3H (N)]-pregnenolone) and using high performance liquid chromatography (HPLC) to separate the products formed. We also measured plasma cortisol and glucose concentration and liver glucocorticoid receptor (GR) protein expression in the brain and liver as indicators of target tissue responsiveness.
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MATERIALS AND METHODS |
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Experimental conditions.
Juvenile rainbow trout (Oncorhynchus mykiss, average body mass 200 g) were obtained from Humber springs trout hatchery, Mono Mills, ON, Canada. Fish were maintained in 200-l tanks with continuous running water at 13°C and 12-h L:12-h D photoperiod and were acclimated for 2 months before the start of the experiment. The fish were fed to satiety (3 point sinking food, Martin Mills Inc., Elmira, ON, Canada) once daily 5 days a week during the acclimation period.
Experimental design.
Groups of 67 fish each were randomly assigned to four aquaria (100 l). Fish were fed with feed to which was added ethanol alone (control) or -naphthoflavone (ANF), ß-naphthoflavone (BNF) or
/ß-naphthoflavone dissolved in ethanol (ABNF) to levels that would provide 10 mg/kg body mass/day. The fish were fed the diets daily for 5 days. Briefly, the feed was evenly submerged in 95% ethanol alone (control) or ethanol containing ANF, BNF, or ABNF to ensure adequate coating of the pellets. The ethanol was allowed to evaporate by air-drying, and the feed was stored in a cool and dry place. This method provides an easy means of administering the drugs without stressing the fish as is the case with intraperitoneal injections or implants (Gamperl et al., 1994
). At the end of the experimental period, fish were quickly netted from the aquaria and killed with an overdose of 2-phenoxyethanol (1:1000).
Each fish was bled by caudal puncture, and the blood was collected into heparinized tubes. Plasma was collected after centrifugation (6000 x g for 10 min) and stored frozen for plasma cortisol and glucose determination at a later date. Pieces of liver and head kidney (containing the interrenals) were frozen immediately on dry ice for protein and mRNA determinations (see below). The head kidney tissues, for in vitro incubations, were placed in M199 media and processed as outlined below.
In vitro head kidney incubations.
Head kidney tissue from treatment groups was excised immediately and sliced in ice-cold M199 medium (supplemented with 0.1% BSA, 0.01% L-glutamine and 0.1% ß-D-glucose) before incubating with porcine ACTH (139) and 3H-P5 to measure cortisol production rate and metabolism of radioactive pregnenolone, respectively. Cortisol production rate was measured as described previously (Wilson et al., 1998). Briefly, the incubation consisted of distributing the tissues from each fish equally into two wells (i.e., one each for control and ACTH) in a multiwell (24 wells) Falcon tissue culture plate. Tissue pieces were allowed to incubate with gentle shaking for 2 h at 13°C (equilibration period), after which the supernatant was replaced with fresh medium and incubated for 1 h, and the media was collected and stored for determination of cortisol concentration (basal cortisol production rate). The media was replaced with either fresh media alone (control) or fresh media containing ACTH (0.5 IU/ml) and incubated for an additional 3 h, after which the supernatant was frozen for later determination of cortisol concentration (stimulated cortisol production rate). Preliminary time-course and dose-response studies were conducted to establish the concentration of ACTH required for eliciting a maximal response in trout head kidney tissues. The wet weight of the tissue in each well was recorded, and cortisol production rate was expressed as ng/h/mg wet weight. The head kidney tissue was frozen for later determination of steroidogenic enzymes mRNA abundance.
For 3H-P5 incubations, head kidney tissue slices were equilibrated for 2 h at 13°C in a 24-well plate (one well per fish per treatment). The media was exchanged, and the tissue slices were incubated in 3H-P5 at a final concentration of 0.75 µCi/ml for 3 h. At the end of the incubation period media was collected and stored at 20°C for later separation of steroids using high performance liquid chromatography (HPLC) analysis. The concentration of the 3H-P5 used was based on previous studies on rainbow trout ovarian follicles and embryo liver incubations (Petkam et al., 2003). The duration of the incubation was established based on the preliminary time-course studies. The tissue slices were weighed and frozen for later determination of the protein content using BioRad protein assay kit and bovine serum albumin (BSA) as the standards.
HPLC Analysis
Sample extraction.
Steroids in the samples were separated using solid phase extraction (SPE), followed by HPLC as per the previously established protocols (Khan et al., 1997). Briefly, Sep-Pak C18 cartridges (Waters Corp, Milford, MA) were used for the SPE. The cartridges were primed with methanol and then double-distilled water. The incubation medium was percolated with the cartridge followed by 5 ml of hexane. The free and conjugated steroid fraction was eluted with 5 ml diethyl ether and 5 ml of methanol, respectively. The recovery of the radiolabelled steroid was measured by mixing 100 µl of the eluate with 4 ml of scintillation cocktail (Ecolite, ICN Biochemicals) and counted in a Searle Delta 300 ß-counter. Subsequently, they were dried under nitrogen at 35°C. The dried ether residues were dissolved in 200 µl of acetonitrile:water (1:1) containing authentic standard reference steroids, and 25 µl was injected into either one of the following chromatographic systems.
HPLC methods.
This system was equipped with a Waters model 2690 alliance separation module and Waters 996 photodiode array detector. Millennium 32 software was used to create a binary gradient of acetonitrile and water or a trinary acetonitrile-methanol-water on a Nova-Pak C18 column. The acetonitrile-water gradient consisted of mobile phase with 27% acetonitrile in water from 0 to 5 min, increasing to 36% at 9.5 min, remaining at 36% from 9.5 to 21 min, increasing to 60% at 27 min, and then running from 85 to 100% between 27 and 35 min. The flow rate was 0.6 ml/min. The ternary gradient of acetonitrile-methanol-water gradient consisted of mobile phase that started with a methanol:acetonitrile:water mix of 46.8:4.2:49 that was changed to 70:15:15 mixture over a 30-min period using Waters gradient curve # 7 at a flow rate of 0.5 ml/min. Two gradients were used to confirm the authenticity of the separated steroid products. Detection of the steroids was done by generating chromatograms at 254 and 280 nm from the photodiode array detector, and the radioactive labeled metabolites were detected using a mix of scintillant and eluant (3:1 ratio) by Canberra-Packard 500TR flow scintillation analyzer. The radiolabelled steroid metabolites were identified based on their co-elution with authentic reference steroid standards from the two columns (Lowartz et al., 2003). The two solvent gradients and the shifts in retention times for a given steroid provided a means for testing the authenticity of any particular peak.
Cortisol and glucose analyses.
Plasma cortisol concentrations were measured using a commercially available ImmuChemTM 125I RIA kit (ICN Biomedicals, Costa Mesa, CA) according to established protocols (Aluru and Vijayan, 2004). Plasma glucose levels were determined colorimetrically using a commercially available kit (Trinder method; Sigma).
SDSPAGE and Western blotting.
The protein concentrations of the tissues were determined using the bichinchoninic acid (BCA) method with BSA as the standard. The procedure for SDSPAGE and Western blotting were according to established protocols (Boone et al., 2002). Briefly, samples (40 µg protein/sample) were separated on 8% polyacrylamide gels using the discontinuous buffer system of Laemmli (1970)
. The proteins were transferred onto nitrocellulose membrane (20 V for 30 min) with a semidry transfer unit (BioRad) using transfer buffer (25 mM Tris pH 8.3, 192 mM glycine, and 20% (v/v) methanol) and were blocked with 5% skimmed milk in TBS-t (20 mM Tris pH 7.5, 300 mM NaCl and 0.1% (v/v) Tween 20 with 0.02% sodium azide) for 60 min. Primary and secondary antibodies were diluted in the blocking solution to appropriate concentrations as indicated. CYP1A was detected using a mouse monoclonal anti-cod antibody at 1:3000 dilution, and for AhR, a rabbit polyclonal antibody against trout AhR was used at 1:1000 dilution. The glucocorticoid receptor (GR) was detected using a polyclonal trout GR antibody at 1:1000 dilution. The secondary antibodies were alkaline phosphatase-conjugated goat anti-mouse for CYP1A (1:1000 dilution), goat anti-rabbit for GR (1:3000 dilution), and goat anti-rabbit conjugated with horseradish peroxidase for AhR (1:3000 dilution). The membranes were incubated in primary antibody for 60 min at room temperature, washed with TBS-t (2 x 5 min), incubated with secondary antibody for 60 min, and finally washed with TBS-t (1 x 15 min). Visualization of bands was carried out with either NBT (0.033% w/v) and BCIP (0.017% w/v) or enhanced chemiluminescense (ECL) plus for AhR (Amersham Biosciences). The molecular weight was estimated by comparison with prestained low-range molecular weight markers (phosphorylase B 112 kDa, bovine serum albumin 81 kDa, ovalbumin 49.9 kDa, carbonic anhydrase 36.2 kDa, soybean trypsin inhibitor 29.9 kDa, lysozyme 21.3 kDa). The intensities of the protein bands were determined by scanning the blots with either Chemi imagerTM (for CYP1A1 and GR) or Typhoon 9400 (for AhR) (Amersham Biosciences) and quantifying band intensity using the AlphaEase software (Alpha Innotech, CA).
Quantitative Real-Time (RT)-PCR (qPCR)
RNA isolation and first strand cDNA synthesis.
Total RNA was isolated from head kidney using RNeasy mini-kit (Qiagen, ON) following the manufacturer's protocol and quantified spectrophotometrically at 260 nm. During RNA isolation, DNase treatment was done to avoid any genomic contamination. The first strand cDNA was synthesized from 1 µg of total RNA using First Strand cDNA synthesis kit (MBI Fermentas). Briefly, total RNA was heat denatured (70°C) and cooled on ice. The sample was used in a 20 µl reverse transcriptase reaction using 0.5 µg oligo d(T) primers and 1 mM each dNTP, 20 U ribonuclease inhibitors, and 40 U MMuLV reverse transcriptase. The reaction was incubated at 37°C for 1 h and stopped by heating at 70°C for 10 min. A cDNA stock for constructing a relative standard curve and cDNA from samples were synthesized using this method.
Relative standard curve.
Primers were designed using rainbow trout CYP1A, StAR, P450scc, 11ß-hydroxylase, and ß-actin cDNA sequences to amplify either a 100 bp for CYP1A or
500 bp product for other genes used in qPCR. The primer sequences and their accession numbers are given in Table 1. A relative standard curve was constructed for target genes (CYP1A, StAR, P450scc, and 11ß-hydroxylase) and housekeeping gene (ß-actin), using either cDNA stock or plasmid vectors with inserted target sequences according to established protocols (Sathiyaa and Vijayan, 2003
). Briefly, the concentration of the cDNA or plasmid vector stock was assumed to be 500 pg/µl, and reactions were set-up with different concentrations ranging from 10 to 3000 pg per reaction for the standard curve. Platinum® Quantitative PCR SuperMix-UDG (Invitrogen, CA) used was 2x concentrated, and every 25-µl reaction had 1.5 U Platinum Taq DNA polymerase, 20 mM TrisHCl (pH 8.4), 50 mM KCl, 3 mM MgCl2, 200 µM dGTP, 200 µM dATP, 200 µM dCTP, 400 µM dUTP, and 1 U UDG; the reaction also contained 0.2 µM forward and reverse primers and 1:100,000 SYBR green I nucleic acid gel stain (Roche). Master mixes, to reduce pipetting errors, were prepared at every stage for triplicate reactions (3 x 25 µl) for each standard.
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Statistical analysis.
All statistical analyses were performed with SPSS version 11.5 (SPSS Inc., Chicago, IL), and data were expressed as mean ± standard error of the mean (SEM). The data were transformed (logarithmic), wherever necessary, for homogeneity of variance (confirmed using Levene's test), but nontransformed values are shown in the figures and tables. One-way ANOVA was used to compare treatment effects on tissue CYP1A1 and AhR protein expression, plasma cortisol and glucose levels, and GR protein expression, in vitro cortisol production rate and StAR, P450scc, and 11ß-hydroxylase gene expression, as well as radiolabelled steroids/metabolite accumulation. Student's t-test for independent samples was used to test for significant differences between the means of unstimulated control and ACTH-mediated changes in cortisol production rate and StAR, P450scc, and 11ß-hydroxylase mRNA abundance. A post hoc (Tukey's) test was used for pair-wise comparison wherever significant differences were observed. A probability level of p 0.05 was considered significant.
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RESULTS |
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StAR, P450scc (CYP11A), and 11ß-Hydroxylase Gene Expression
Incubation of head kidney pieces with ACTH for 3 h significantly increased mRNA abundance of StAR and P450scc, but not 11ß-hydroxylase compared to the control fish (Fig. 3A). This ACTH-induced StAR mRNA abundance was significantly attenuated by ANF, BNF, and ABNF compared to control group (Fig. 3B). ACTH-stimulated P450scc mRNA abundance was significantly lower in BNF and ABNF groups compared to the control group (Fig. 3C). None of the treatments had any significant effect on ACTH-mediated 11ß-hydroxylase mRNA levels (Fig. 3D).
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DISCUSSION |
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StAR protein, which translocates cholesterol from the outer to the inner mitochondrial membrane, is considered to be a rate-limiting step in steroidogenesis (Stocco, 2000; Stocco and Clark, 1996
). Also, the conversion of cholesterol to pregnenolone, catalyzed by P450scc, is the primary rate-limiting step in the steroidogenic pathway (Payne and Hales, 2004
). Both these proteins are acutely regulated in the steroidogenic tissues, as is evident from the transient increase in their mRNA and protein levels in response to trophic hormone stimulation, including LH and ACTH (Lehoux et al., 1998
; Le Roy et al., 2000
; Stocco, 2000
). This was also true in fish interrenal tissue, where exogenous ACTH administration and/or stress manipulations, which increase endogenous ACTH and cortisol levels, enhanced StAR and P450scc mRNA abundance (Geslin and Auperin, 2004
; Kasakabe et al., 2002; Li et al., 2003
; present study). The production of both StAR and P450scc, which have key roles in steroid biosynthesis, appear to be sensitive to xenobiotic-mediated impairment (Dasmahapatra et al., 2000
; Fukuzawa et al., 2004
; Moore et al., 1991
; Walsh et al., 2000a
,b
,c
). Most of those studies, however, utilized mammalian cell line models to understand the mechanism(s) of action, and consequently, very little is known about the physiological implications.
In the present study we show that in vivo exposure of trout to BNF also impacts StAR and P450scc mRNA content in the interrenal tissues, suggesting a mechanistic link for the attenuated cortisol production in response to xenobiotics activating AhR in fish (Aluru and Vijayan, 2004; Hontela, 2005
; Wilson et al., 1998
). In this context, TCDD has been shown to depress P450scc in mammalian cell systems (Dasmahapatra et al., 2000
; Moore et al., 1991
), but to our knowledge no study had examined the impact of AhR activation on StAR protein regulation. Recently, however, an in vitro reporter assay using StAR promoter showed that BNF modulated the transactivational capacity (Sugawara et al., 2001
). Although these studies point to a direct impact of AhR activation on genes involved in steroidogenesis, the precise mechanism(s) of action still remains unclear. Apart from the direct interaction of AhR with XREs, this receptor heterocomplex may also indirectly affect target gene transcription. For instance, steroidogenic factor (SF-1), a member of nuclear receptor super family which regulates gene expression of StAR and many steroidogenic enzymes (P450scc, CYP11B1, CYP11B2, CYP19), competes with AhR for binding to steroid receptor coactivator-1 (SRC-1), cAMP response binding protein (CBP), C/EBP, and Sp1, thereby modulating the expression of steroidogenic genes (Suguwara et al., 2001
; Val et al., 2003
). As steroid production is of utmost importance for all aspects of animal performance, including reproduction, growth and metabolism, and ion homeostasis, the impairment of these key proteins in the steroidogenic pathway may be a mechanism(s) involved in the PCB's impact on animal function (Aluru et al., 2004
; Hontela et al., 1992
; Jorgensen et al., 2002
; Quabias et al., 1997
, 2000
).
In addition to StAR and P450scc, there are several other enzymes involved in steroidogenesis, including other P450s, dehydrogenases, and hydroxylases (Payne and Hales, 2004; Sewer and Waterman, 2003
). However, unlike StAR and P450scc, the enzymes downstream of pregnenolone are not thought to be acutely regulated by trophic hormones (Payne and O'Shaughnessy, 1996
). This seems to be the case in our study, as 11ß-hydroxylase, the final step in the cortisol biosynthesis, showed no alteration in its mRNA abundance upon ACTH stimulation (Fig. 3D). This may be due to the high basal levels of this enzyme, including high abundance of 11ß-hydroxylase mRNA transcripts, required for producing large amounts of cortisol within minutes of exposure to stressful stimuli (ACTH stimulation) (Jiang et al., 1998
). Further confirmation that the enzyme pathways downstream of P450scc were not unduly affected by AhR activation arises from radiolabelled studies using P5 as the substrate (Table 2). There was a significant treatment effect on radioactive P4 accumulation, but not on other steroid metabolites. While this may suggest a potential impact of AhR activation on 3ß-hydroxysteroid dehydrogenase (3ß-HSD) activity, overall there was very little impact on flux through the steroidogenic pathway (Table 2). Considered together, StAR and P450scc appear to be the key targets for AhR-mediated impairment of interrenal steroidogenesis in rainbow trout.
One of our objectives of this study was to determine whether AhR signaling was directly involved in BNF-mediated impairment of cortisol synthesis. Consequently, we used ANF, a well-established AhR antagonist (Lee and Dasmahapatra, 1994; Miranda et al., 1998
; Navas et al., 2003
; Navas and Segner, 2000
; Santostefano et al., 1993
), to block the BNF-induced changes. However, unlike other studies, we were not only unable to block AhR signaling, but also for the first time we are showing that ANF behaves like an AhR agonist in fish. Specifically, the induction of CYP1A1 gene and protein expression in rainbow trout with ANF (Figs. 1B and 2A) was not seen in previous studies (Aluru and Vijayan, 2004
; Navas et al., 2003
; Navas and Segner, 2000
). This CYP1A1 induction by ANF or BNF did not involve AhR protein regulation (Fig. 1A), supporting post-receptor signaling modifications (Sadar et al., 1996
; Santostefano et al., 1993
). The significantly higher CYP1A1 in the BNF groups relative to ANF group is not surprising, given the lower affinity of the AhR for ANF compared to BNF (Santostefano et al., 1993
). However, the two ligands together seem to have a permissive effect on AhR protein content, but this was not reflected in a significantly higher CYP1A1 expression relative to the drugs alone, further supporting the notion that post-receptor signaling modulates Ah-responsive gene induction (Delescluse et al., 2000
).
In our earlier study ANF failed to block BNF-induced CYP1A1 mRNA and protein expression (Aluru and Vijayan, 2004), clearly arguing against ANF as an AhR antagonist in rainbow trout. The induction of CYP1A1 expression in this study, but not in the earlier study may be related to the dosage and route of administration. For instance, we used a slow-release intraperitoneal implant for drug administration previously (Aluru and Vijayan, 2004
), whereas in this study the fish were fed a relatively large dose of ANF (compared to the implant) on a daily basis for 5 days. Consequently, the dosage difference between the two studies could account for the activation of AhR. This notion was confirmed by dose-dependent studies with trout hepatocytes that clearly showed significant CYP1A1 gene and protein expression only at a high ANF dose (110 µM; unpublished data). Considered together, ANF behaves as an AhR agonist in rainbow trout. The similarity in ANF- and BNF-mediated steroidogenic responses, coupled with the lack of any inhibition of this pathway in the ABNF groups (Figs. 1B, 2, and 3), strengthens the above argument.
Interestingly, the impairment of corticosteroidogenesis by these drugs was not reflected in the plasma cortisol concentration, which was generally low (unstressed levels reported in trout) in all groups (Table 3). This is not surprising given the fact that xenobiotics impact acute regulation (in response to trophic hormone stimulation) of the steroidogenic enzymes and proteins. Also, previously we did see a lack of cortisol effect in unstressed trout treated with BNF, but there was clear attenuation of the stress-induced cortisol response in these animals (Wilson et al., 1998). However, other studies did show that plasma cortisol levels were higher with BNF or TCBP treatment (Aluru and Vijayan, 2004
; Vijayan et al., 1997
). The reason for the differing plasma cortisol levels with BNF treatment is not clear; however, the lack of a consistent glucose response to cortisol stimulation in those other studies as well as in this study (Table 3) clearly attests to a lack of tissue responsiveness (Aluru and Vijayan, 2004
). The reasons are unclear, especially since the higher GR content with BNF in the liver and brain would suggest enhanced tissue responsiveness. The higher liver GR content with BNF was also evident in our previous study, despite higher plasma cortisol level, suggesting that BNF-mediated GR changes are independent of plasma cortisol levels in rainbow trout. The recent observation that BNF also increases GR protein expression in trout hepatocytes in primary culture in vitro (S. Wiseman and M. M. Vijayan, unpublished) supports the above contention that tissue GR protein responses to BNF treatment is independent of changes in plasma cortisol levels (Aluru and Vijayan, 2004
). This, coupled with the fact that ANF abolished the BNF-induced GR response (Figs. 5A and 5B), suggests an antagonistic effect of ANF. This is intriguing, because the mode of action of ANF appeared similar to BNF with respect to CYP1A1 expression and steroidogenesis, but the dissimilar GR response leads us to propose that the antagonistic effect of ANF may be gene specific, suggesting post-receptor modifications. However, as trout have multiple AhR isoforms with different sensitivities to ligand (Abnet et al., 1999
), one cannot discount the possibility that these receptors may be distinct with respect to ANF activation and/or gene transactivation. Studies focusing on these receptor isoforms and their signaling pathways are warranted to shed light on the functional significance of multiple AhR isoforms on endocrine disruption in fish.
In conclusion, this study suggests that AhR activation impacts corticosteroidogenesis by affecting StAR and P450scc gene expression in rainbow trout. AhR activation did not affect the steroidogenic enzyme activities downstream of P450scc, as determined by incubation of head kidney tissues with radiolabelled pregnenolone. Overall, the endocrine disruption of xenobiotics acting via AhR may involve impaired cortisol production and abnormal target tissue receptor response in rainbow trout. Finally, our study indicates that ANF does not act as an AhR antagonist in rainbow trout.
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ACKNOWLEDGMENTS |
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