The Effect of 2,3,7,8-Tetrachlorodibenzo-p-dioxin on Corticotrophin-Releasing Hormone, Arginine Vasopressin, and Pro-opiomelanocortin mRNA Levels in the Hypothalamus of the Cynomolgus Monkey

Surekha Shridhar*, Anne Farley{dagger}, Robert L. Reid{dagger}, Warren G. Foster{ddagger} and Dean A. Van Vugt*,{dagger},1

* Department of Physiology and {dagger} Department of Obstetrics and Gynecology, Queen's University, Kingston, Ontario, Canada; and {ddagger} Environmental and Occupational Toxicology Division, Health Protection Branch, Health Canada, Ottawa, Ontario, Canada

Received February 26, 2001; accepted July 13, 2001


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD) is a widespread environmental contaminant that has profound deleterious effects on development and reproduction. TCDD may act at one or more levels to alter the hypothalamic-pituitary-adrenal (HPA) and hypothalamic-pituitary-gonadal (HPG) axes. The objective of this study was to investigate whether TCDD modulates neuroendocrine systems by altering gene expression of arginine vasopressin (AVP), corticotrophin-releasing hormone (CRH), or pro-opiomelanocortin (POMC), which are important neuroregulators of the HPA and HPG axes. Four groups of female cynomolgus monkeys (Macaca fascicularis) were administered daily oral doses of gelatin capsule containing TCDD (0, 1, 5, or 25 ng/kg body weight) mixed with glucose 5 days a week for 1 year. At the end of the dosing period, animals were euthanized and brains were harvested. CRH, AVP, and POMC mRNA levels were semiquantified by in situ hybridization histochemistry on 30-µm coronal sections of the brain. Blood collected on the day of euthanasia was assayed for cortisol and progesterone. CRH mRNA levels in the paraventricular nucleus (PVN) were significantly increased by the 2 higher TCDD doses (5 and 25 ng/kg/day) compared to controls (p < 0.05). There was a trend towards increased AVP mRNA levels in both the supraoptic nucleus (SON) and PVN. No effect of TCDD on POMC was observed. Cortisol levels were significantly increased in TCDD-exposed animals. Progesterone concentrations and menstruation data indicated that TCDD did not interfere with ovulation. We conclude that TCDD stimulated the HPA axis by a central effect involving CRH, but had no effect on the HPG axis at the doses tested.

Key Words: 2,3,7,8-tetrachlorodibenzo-p-dioxin; cynomolgus monkeys; hypothalamic-pituitary-adrenal axis; corticotropin-releasing hormone; arginine vasopressin; pro-opiomelanocortin; mRNA; in situ hybridization histochemistry.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD) is a widespread environmental contaminant formed as a byproduct during the manufacture of phenoxy and chlorophenol herbicides (Lilienfeld and Gallo, 1989Go), bleaching of paper and pulp, and incineration of chlorine-containing waste (Rappe et al., 1979Go). Its slow biodegradation causes it to persist in the environment and food chain and thereby accumulate in adipose tissue. The mean background level of TCDD in adipose tissue in the American population is reported to be 5.4 ng/kg (Schecter et al., 1994Go), whereas the residue levels in individuals with occupational- or poisoning-type exposures can be much higher.

The effects of TCDD include wasting at higher doses in rats (Kociba et al., 1979Go) as well as in monkeys (McConnel et al., 1978), chloracne in humans (Taylor, 1974Go) and monkeys (McConnel et al., 1978), hepatotoxicity, as reported in rats (Kociba et al., 1979Go) and monkeys (Schulz et al., 1996Go), immunotoxicity (Neubert et al., 1991Go, 1993Go), and reproductive toxicity. Female reproductive toxicity includes altered estrous cycle, anovulation, and decreased fertility in rats (Gray and Ostby, 1995; Kociba et al., 1979Go). Similar adverse effects on reproduction were observed in monkeys (Allen et al., 1979Go; McNulty, 1984). The profound effects of TCDD on the hypothalamic-pituitary-gonadal (HPG) and hypothalamic-pituitary-adrenal (HPA) axes are believed to be responsible for reproductive toxicity. Alteration of blood steroid hormone and gonadotropin levels, abnormal estrous/menstrual cyclicity, and disruption of ovulation are indicative of the adverse effects of TCDD on the HPG axis, as reported in rats (Chaffin et al., 1996Go; Li et al., 1995Go, 1997Go) and monkeys (Barsotti et al., 1979Go). Decreased food intake and progressive weight loss in rats (Gasiewicz et al., 1980Go; Harris et al., 1973Go) and in monkeys (McConnel et al., 1978), altered blood levels of adrenocorticotropic hormone in rats (Bestervelt et al., 1993Go, 1998Go) or corticosteroids in rats (Balk and Piper, 1984Go; Gorski et al., 1988Go; Neal et al., 1979Go), and change in blood and tissue ß-endorphin levels in rats (Bestervelt et al., 1991Go; Pohjanvirta et al., 1993Go) suggest disruption of the HPA axis. However, the mechanisms responsible for eliciting these changes are not well documented. The brain contains aryl hydrocarbon receptors (AhRs; Carlstedt-Duke, 1979Go; Dolwick et al., 1993Go; Li et al., 1994Go), and TCDD accumulates in the brain of rodents following peripheral administration (Pohjanvirta et al., 1990Go; Russel et al., 1988). It is therefore possible that TCDD acts at the hypothalamic level of these axes.

The hypothalamus is the site for the synthesis and action of numerous neuropeptides that are involved in the regulation of both HPG and HPA axes. The HPA axis is regulated by the combined action of corticotrophin-releasing hormone (CRH) in the paraventricular nucleus (PVN) and arginine vasopressin (AVP) in the PVN and supraoptic nucleus (SON). These 2 peptides stimulate ACTH release through a synergistic action (Rivier and Vale, 1983Go). Furthermore, CRH, AVP, and pro-opiomelanocortin (POMC) in the arcuate nucleus (ARC) are important inhibitory neuromodulators of the HPG axis (Heisler et al., 1994Go; Olster and Ferin, 1987Go; Rasmussen et al., 1983Go; Rivier and Vale, 1983Go; Yen et al., 1985Go). Therefore, we hypothesized that TCDD-induced changes in the reproductive and adrenal axes are mediated, in part, by changes in the hypothalamic expression of CRH, AVP, and POMC.

The objective of this study was to investigate the effects of chronic TCDD exposure in female cynomolgus monkeys on gene expression of these neuropeptides by measuring mRNA levels of CRH in the PVN, AVP in the SON and PVN, and POMC in the ARC by in situ hybridization. This study was part of a larger study conducted at Health Canada, Ottawa (Yang et al., 2000Go) that examined the effect of TCDD on endometriosis.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Animals and dosing.
This study was conducted in 23 ovary-intact cynomolgus monkeys (Macaca fascicularis). All monkeys were housed in individual cages in light- and temperature-controlled rooms. Lights were on from 0600 to 1800 hours, and temperature was maintained between 20 and 22°C. Diet consisted of Purina Monkey Chow (Ralston-Purina, St. Louis, MO) and water ad libitum, with supplements of fruits and vegetables. All animal husbandry practices and experimental procedures were approved by the Institutional Animal Care and Use Committee of the Health Protection Branch and followed the guidelines of the Canadian Council of Animal Care

Monkeys were randomly assigned to 1 of 4 groups consisting of control (n = 5), 1, 5, or 25 ng TCDD/kg/day (n = 6 per group) for one year. Details of dose preparation and dosing have been described elsewhere (Yang, 2000). The doses of TCDD used in this study were selected so as to be relevant to human exposure in industrialized regions and comparable to those of a previous study in rhesus monkeys (Rier et al., 1993Go). The animals of the control group received gelatin capsules containing glucose only. Because TCDD was administered only on weekdays, the calculated daily doses were 0.71, 3.57, and 17.86 ng/kg/day.

At the end of one year of TCDD exposure, each animal was injected, im, with ketamine and removed from its cage. A 3-ml blood sample was collected by venipuncture prior to inducing rapid anesthesia with saffan (Pittman-Moore, Middlesex, UK; 1.2 mg/kg) and ketamine (10 mg/kg; 1:1 v/v). Both the right and left carotid arteries were cannulated and the brain was perfused with 100 ml PBS at 4°C, followed by 4% paraformaldehyde with 0.1 M borax (pH 9.5) at a rate of 10 ml/kg/min for 20 min. The brain was removed from the skull and postfixed in 300 ml of 4% paraformaldehyde and 0.1-M borax buffer at 4°C for 2 days. Brains were then transferred to 4% paraformaldehyde-borax buffer containing 10% sucrose at 4°C for 48 h prior to sectioning. 30-µm coronal sections were cut from the olfactory bulb to the caudal medulla. The slices were placed in cryoprotectant (0.05 M sodium phosphate buffer, pH 7.3, 30% ethylene glycol and 20% glycerol) and stored at –20°C until further processing.

In situ hybridization histochemistry.
Hybridization histochemical localization of each transcript was performed in one out of every 18 sections (approximately 0.5-mm intervals through the nuclei of interest; 8 to 10 sections/nucleus) in the SON, PVN, and ARC. Protocols for riboprobe synthesis and hybridization of mRNA were adapted from Simmons et al. (1989). All solutions were treated with diethyl pyrocarbonate (DEPC) and autoclaved to eliminate RNase activity. All glassware was autoclaved to minimize RNase contamination. Tissue sections were mounted onto gelatin and poly-L-lysine-coated slides and dried under vacuum, fixed in 4% paraformaldehyde for 20 min, incubated in potassium phosphate-buffered saline (KPBS) for 10 min, and then digested by proteinase K (5 µg/ml dissolved in 100 mM Tris-HCl, pH 8.0 and 50 mM EDTA, pH 8.0) at 37°C for 25 min. Sections were then rinsed with DEPC-treated water followed by 0.1 M triethanolamine (TEA, pH 8.0) and acetylated in 0.25% acetic anhydride in 0.1 M TEA. Thereafter the sections were rinsed in standard saline citrate (2X SSC) and dehydrated through graded concentrations of alcohol (50, 70, 95, and 100%). After vacuum drying the sections for at least 2 h, 130 µl of hybridization mixture containing 35S-labelled cRNA probes (107 cpm/ml) was spotted onto each section, cover-slipped, and incubated at 60°C in a humidified chamber for 12–36 h, depending on the probe. Cover slips were removed and the sections were rinsed in 4X SSC twice for 10 min. This was followed by digestion with RNase A (20 µg/ml, 37°C, 30 min.) in RNase buffer (0.5 M NaCl, 10 mM Tris-HCl, 0.5 mM EDTA). Tissue sections were then washed through 4 high-stringency washes of decreased salinity at room temperature (2X, 1X, 0.5X) followed by a high-temperature wash with 0.1X SSC at 60°C for 30 min and dehydrated in ascending concentrations of ethanol (50, 70, 95, 100%). After vacuum drying for at least 2h, slides were apposed to Biomax MR X-ray films (Eastman Kodak, Rochester, NY) at 4°C for 12–48 h, depending on the probe. Following film development, slides were defatted in xylene, dipped in NTB2 nuclear emulsion (Kodak) diluted 1:1 with distilled water, and exposed for 14–32 days at 4°C. Slides were developed in D19 developer (Kodak) for 3.5 min at 14–16°C and fixed in Kodafix fixer (Kodak) for 5 min. Slides were then rinsed in running distilled water for 1 h, counterstained with thionine (0.25%), dehydrated through graded concentrations of ethanol, cleared in xylene, and cover-slipped with DPX.

cRNA probe synthesis and preparation.
The CRH antisense cRNA probe was generated from an EcoRI fragment of CRH cDNA (1.2 Kb; from Dr. K. Mayo, Northwestern University), subcloned into a pGEM4 plasmid, and linearized with Hind III. The AVP antisense cRNA probe was generated from a SMA I-PST I fragment of AVP cDNA (230 bp; from Dr. D. Richter, Universitat Hamburg, Germany), subcloned into a pSP65 plasmid, and linearized with Hind III. POMC cRNA probe was generated from a Nepp 3a1 fragment of POMC cDNA (1040 bp; from Dr. S. Watson, University of Michigan, Ann Arbor), subcloned into a pGEM3 vector, and linearized with Hind III. The c-fos antisense cRNA probe was generated from an EcoRI fragment of c-fos cDNA (2 kb; from Dr. I. Verma, Salk Institute, La Jolla, CA), subcloned into a Bluescript SK plasmid, and linearized with SmaI.

Radioactive cRNA probes were synthesized by in vitro transcription by incubating 250 ng of linearized cDNA in 6 mM MgCl2, 40 mM Tris, pH 7.9, 2 mM spermidine, 10 mM NaCl, 100 mM DTT, 0.2 mM ATP/GTP/CTP, 200 µCi of {alpha}-35S-UTP (NENTM Life Science Products, Inc., Boston, MA), 40 U RNasin (Promega, Madison, WI), and 20 U RNA polymerase for 60 min at 37°C. After the transcription reaction, the DNA template was removed by adding 100 µl DNase solution (1 µl DNase, 2.5 µl tRNA 10 mg/ml, 94 µl of 10 mM Tris/10 mM MgCl2, and 2.5 µl DEPC H2O) and incubated at room temperature for 10 min. The unincorporated 35S-UTP nucleotides were removed by extraction with 100 µl of 1:1 vol/vol phenol-chloroform and precipitation with 80 µl ammonium acetate (5 M, pH 5.5) and 500 µl of 100% ethanol for 30 min at –70°C. After centrifugation, the pellet was dried and resuspended in 100 µl of 10 mM Tris/1 mM EDTA (pH 8.0), counted to determine the success of the probe synthesis, and stored at –70°C. On the day of hybridization, the probe was diluted to 107 cpm/ml of hybridization solution (1 ml hybridization solution = 500 µl formamide, 50 µl NaCl, 10 µl 1 M Tris, pH 8.0, 2 µl 0.5 µ EDTA, pH 8.0, 50 µl 20X Denhart's solution, 200 µl 50% dextran sulfate, 50 µl 10 mg/ml tRNA, 10 µl 1 M DTT, 118 µl DEPC-treated water minus the volume of probe used). The solution was mixed well by vortexing and heated at 65°C for 5 min before spotting 130 µl of this solution onto each section.

The hybridization signal-intensity of CRH mRNA in the PVN, AVP and c-fos mRNA in the SON and PVN were semiquantified for each animal using the X-ray films. POMC mRNA in the ARC was quantified using emulsion-dipped slides under dark field because of high background observed in the X-ray films. The X-ray images were captured (digitized) under a Northern Light Desktop Illuminator (Imaging Research System, Inc.; St. Catherines, ON, Canada) using a Sony Camera Video System attached to a Micro-Nikkor 55-mm lens (Nikon, Tokyo, Japan) and Vivitar extension tube set for a Nikon lens and coupled to an IBM computer running software from Imaging Research, Inc. (MCID-M5 Version 4.00, Brock University, St. Catherines, ON). The illuminator was set at an intensity of 1025 and a constant shading error was maintained throughout digitization. To establish that optical density measurements were linear, gray level/optical density calibration was done using a density step tablet (Eastman Kodak) of optical density ranging from 0.05 to 1.95. The transmittance values (or optical density) of the hybridization signals on the captured images were measured using Scion Image software, which is a Windows version of NIH Image written at NIH. Scion Image software is available for downloading at http://www.scioncorp.com. The optical density (OD) of the hybridization signal for SON, PVN, and ARC were measured bilaterally in all the sections of all the animals. The OD was corrected for the average background signal by subtracting the OD of areas without positive signal located immediately outside the digitized area. The integrated optical density (IOD) was calculated by multiplying the OD by the area. The average IOD of all sections for a specific nucleus was averaged for each animal prior to calculating a group average. The mRNA levels were expressed as IOD ± SEM. POMC mRNA levels were measured by capturing a region in the ARC with maximum signal intensity under dark-field illumination at a magnification of 10 x, using a microscope (Leitz, Germany) connected to the same camera system and computer as described above. The OD for the entire field was measured using Scion Image software as described before.

In order to determine if observed differences in optical density were due to increases in the number of cells expressing a particular transcript, the total number of cells positive for the hybridization signal were counted. The average number of positive cells in 2 sections through the nucleus of interest was determined for each animal before an average was calculated for each group. A grain density greater than 3 x the background was the criteria used for designating positive cells.

Blood collection and radioimmunoassays.
Blood samples were collected at the time of perfusion. Samples were left to clot overnight at 4°C before centrifugation for 15 min. at 1500 x g. Serum was separated and stored at –20°C until assayed. Progesterone and cortisol levels were measured with radioimmunoassay kits (Diagnostic Products Corporation, Los Angeles, CA). Assay sensitivities were 0.02 ng/ml and 0.22 µg/dl for progesterone and cortisol respectively. All serum samples were processed in a single assay for each hormone.

Statistical analyses.
Data were expressed as mean ± SEM and were analyzed by one-way analysis of variance (ANOVA) followed by Dunnett's multiple comparison tests in case of significance (SPSS analysis, Chicago, IL). The level of statistical significance was set at p < 0.05.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
No effect of TCDD on either body weight or menstrual cyclicity was observed. The mean body weights and menstrual cycle length of each group at the beginning, middle, and end of the study are shown in Table 1Go. TCDD did not alter either the menstrual cycle length or the menstrual bleeding duration. TCDD had no effect on ovulation as determined by progesterone levels on the day of euthanasia (see Table 1Go). Cortisol levels were significantly increased by TCDD; 41.8 ± 1.2 (control) versus 52.93 ± 2.2 (1 ng) and 51.61 ± 3.0 µg/dl (5 ng); p = 0.01. Cortisol measurements in the 25-ng dose group were restricted to only 2 animals (68.09 and 31.51 µg/dl) because of insufficient serum.


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TABLE 1 Effect of Dietary TCDD on Body Weight, Menstrual Cycle, and Progesterone and Cortisol Levels
 
The effect of TCDD on CRH mRNA in the PVN is shown in Figures 1–3GoGoGo. Figure 1Go shows representative examples of CRH hybridization in the PVN of control and TCDD-exposed animals. The optical density measurements of autoradiograms are shown in Figure 2Go. There was a 2- to 3-fold increase in integrated optical density representing CRH mRNA in the PVN of TCDD treated monkeys compared to controls (p = 0.027); 13.2 ± 2.24 (control) vs. 23.9 ± 3.8 (1 ng), 35.7 ± 4.85 (5 ng), and 33.1 ± 6.2 (25 ng). This increase was statistically significant at the 5- and 25-ng doses (p = 0.02 and 0.029 respectively). The increase at the 1-ng dose was not statistically significant (p = 0.34). The effect of TCDD on the number of cells in the PVN expressing CRH mRNA is shown in Figure 3Go. TCDD at doses 5 and 25 ng increased the average number of CRH mRNA-positive cells from 13.6 ± 2.90 (control) to 35.4 ± 5.65 (5 ng; p = 0.013) and 30.3 ± 4.13 (25 ng; p = 0.037). The increase in cell number observed in the 1-ng dose was not statistically significant (23.3 ± 3.86; p = 0.29).



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FIG. 1. Effect of TCDD on CRH mRNA in the PVN. Contrasted are bright-field photomicrographs depicting CRH-mRNA-positive cells in matched sections from a control (top panel) and a TCDD (5 ng/kg)-treated monkey. Cell nuclei are stained with thionine. Clusters of silver grains with a density greater than 3 times background are considered CRH-mRNA-positive. The arrows denote CRH-mRNA-positive cells (magnification x40).

 


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FIG. 2. Effects of TCDD on CRH mRNA in the PVN of nonhuman primates. Shown are mean integrated optical density (IOD) measurements for CRH mRNA in the PVN following TCDD treatment at doses of 0, 1, 5, and 25 ng/kg bw/day. Values are expressed as mean ± SEM. Asterisks denote statistical significance (p < 0.05); n = 4 in each group.

 


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FIG. 3. Effect of TCDD on CRH cell number in the PVN. Mean number of cells positive for CRH-mRNA hybridization signal in the PVN are shown. Cells were considered positive when the silver grain density over the nucleus was 3 times greater than background. Values are expressed as mean ± SEM. Asterisk denotes statistical significance (p < 0.05); n = 4, 5, 4, and 5 for TCDD doses of 0, 1, 5, and 25 ng/kg/day, respectively.

 
AVP mRNA in the SON of TCDD-exposed animals was increased compared to controls (31.2 ± 1.94 [control] vs. 29.4 ± 6.17 [1 ng], 39.0 ± 7.32 [5 ng], and 44.6 ± 4.67 [25 ng]; top panel of Fig. 4Go). However, analysis of variance determined this increase to be not statistically significant (p = 0.19). Similarly, AVP mRNA in the PVN was increased in TCDD-treated animals (bottom panel of Fig. 4Go), but this increase was not significant (34.2 ± 4.27 [control] vs. 39.3 ± 7.78 [1 ng], 54.0 ± 6.26 [5 ng], and 41.6 ± 2.32 [25 ng] ; p = 0.16).



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FIG. 4. Effect of TCDD on AVP mRNA levels in the SON and PVN. Mean integrated optical density (IOD) measurements for AVP mRNA in the SON (top) and PVN (bottom) are shown; n = 4, 5, 4, and 5 for TCDD doses of 0, 1, 5, and 25 ng/kg/day, respectively.

 
TCDD increased c-fos mRNA in the SON and PVN (Fig. 5Go). In the SON, this effect of TCDD narrowly failed to reach statistical significance ([p = 0.085]; 6.6 ± 1.88 [control] vs. 20.4 ± 4.96 [1 ng], 31.4 ± 4.84 [5 ng], and 16.3 ± 8.14 [25 ng]). TCDD had a similar effect on c-fos mRNA in the PVN; 4.4 ± 0.95 [control] vs. 17.6 ± 7.39 [1 ng], 21.9 ± 4.72 [5 ng], and 16.2 ± 5.66 [25 ng]). While differences were large, they were not statistically significance (p = 0.32). TCDD also increased the number of c-fos mRNA-positive cells in both the SON and PVN. This increase was statistically significant for the PVN; 14.7 ± 9.18 (control) vs. 56.8 ± 27.72 (1 ng), 178.1 ± 28.85 (5 ng), and 99.5 ± 46.99 (25 ng); p = 0.033. There was a positive correlation between c-fos mRNA and CRH mRNA levels in the PVN (correlation coefficient ± 0.690; p = 0.002). No significant correlation was observed between c-fos mRNA and AVP mRNA in either the SON or PVN.



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FIG. 5. Effect of TCDD on c-fos mRNA levels in the SON and PVN. Mean integrated optical density (IOD) measurements for c-fos mRNA in the SON (top) and PVN (bottom) and c-fos cell number (PVN only) at TCDD doses of 0, 1, 5, and 25 ng/kg/day are shown. The values are expressed as mean ± SEM; n = 3, 5, 4, and 4 for TCDD doses of 0, 1, 5, and 25 ng/kg/day, respectively.

 
No differences in ARC-POMC-mRNA levels were observed in TCDD-treated monkeys compared to control monkeys; 39.3 ± 3.11 (control), 33.6 ± 3.08 (1 ng), 33.9 ± 0.86 (5 ng), and 36.8 ± 2.2 (25 ng); p = 0.43. In contrast to CRH, AVP and c-fos, there were no trends for either increased or decreased POMC expression with increasing TCDD exposure.


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
To our knowledge, this is the first study that has examined the effects of TCDD on gene expression of CRH, AVP, c-fos, or POMC in the hypothalamus of nonhuman primates. While it is speculated that the effects of TCDD on reproduction and development may be mediated by its actions on the HPA and HPG axes, the mechanisms whereby TCDD affects the HPA and HPG axes are not well documented. This study attempted to shed new light on the mechanisms of TCDD actions by looking at the gene expression of neuropeptides that modulate the HPA and HPG axes.

TCDD, at doses relevant to accidental or occupational exposure levels in humans, increased CRH mRNA levels and the number of c-fos mRNA expressing cells in the PVN. In addition, cortisol concentrations were significantly increased with TCDD exposure. These data provide the first evidence that subchronic exposure to low, environmentally relevant TCDD doses can affect the HPA axis by stimulating CRH gene expression. The significant correlation between CRH and c-fos mRNA in the PVN favors the conclusion that the increase in CRH transcript levels was indeed the result of increased CRH transcription. Although the highest dose of TCDD did not produce the highest response in the parameters examined, a dose response was observed for most parameters, since CRH mRNA and cell number were increased over 3 doses (0, 1, and 5 ng/kg) as was AVP mRNA in the PVN, c-fos mRNA in the SON and PVN, as well as c-fos cell number in the PVN. The only exception was AVP in the SON. We believe that these differences produced by TCDD exposure are real and that they would achieve statistical significance if the number of observations/group were increased. The magnitude of the CRH response to TCDD is quite large. Compared to acute stimuli of CRH gene expression, the 2–3-fold increase in CRH observed in the present study is very large (Van Vugt et al., 1997Go). This is all the more remarkable when one considers that cortisol negative feedback is increased due to increased cortisol concentrations in these animals. It is possible that no further increase in CRH gene expression was observed at the 25-ng/kg TCDD dose because gene expression was maximally stimulated by the 5-ng/kg dose.

Several studies suggest that the hypothalamus may be a site of action of TCDD, even though the mechanisms are not fully understood (Bestervelt et al., 1991Go; Chaffin et al., 1996Go; Pohjanvirta et al., 1990Go; Russell et al., 1988Go). TCDD may act directly on the PVN to modulate CRH gene expression. This action may be mediated through AhRs that are present in the brain (Carlstedt-Duke, 1979Go; Dolwick et al., 1993Go; Li et al., 1994Go). AhR is a cytosolic receptor that, upon binding TCDD, translocates to the cell nucleus and interacts with a dioxin-response element (DRE) of a target gene to either increase or decrease expression of the target gene (Whitlock, 1990Go). While DRE is present in an array of genes that are induced by TCDD (Lai et al., 1996Go), the presence of DRE in the promoter region of the CRH gene is yet to be demonstrated. Nevertheless, the direct action of TCDD-AhR complex on CRH is a plausible mechanism for the increased CRH gene expression that we observed.

CRH neurons contain glucocorticoid receptors (GR) that can exert negative feedback inhibition of CRH neurons. The increase in CRH mRNA that we observed is unlikely the result of reduced feedback inhibition since TCDD increased cortisol secretion. Both stimulation and inhibition of corticosterone has been observed in response to TCDD in rats (Balk and Piper, 1984Go; Gorski et al., 1988Go; Neal et al., 1979Go). Alternatively, an increase in CRH mRNA could result from an effect of TCDD on GR. TCDD has been shown to decrease the binding capacities of the hepatic cytosolic GR (Sunahara et al., 1989Go). TCDD may similarly decrease the cortisol binding capacity of GR on CRH neurons, thereby reducing feedback inhibition of CRH.

Vasopressin mRNA in the SON and PVN were increased in TCDD-treated monkeys. However, this effect of TCDD did not reach statistical significance. In the absence of any similar reports in the literature on potential effects of TCDD on AVP, we cannot rule out the possibility that TCDD exerts similar effects on CRH and AVP gene expression.

We did not detect any effect of TCDD on POMC mRNA levels in the ARC. The POMC gene has a DRE on its promoter region (Drouin et al., 1985Go), and 2 reports demonstrated an effect of TCDD on ß-endorphin concentration, which is derived from its precursor POMC. A single oral dose of 50 µg/kg TCDD was reported to produce a biphasic effect, initially increasing then decreasing hypothalamic ß-endorphin concentrations in the rat hypothalamus (Bestervelt et al., 1991Go). The same dose of TCDD (50 µg/kg) was reported to decrease plasma ß-endorphin in female rats (Pohjanvirta et al., 1993Go). The relevance of these studies to the current study is arguable, given the high dose of TCDD, and since neither study measured POMC mRNA in the hypothalamus.

TCDD did not appear to negatively impact the menstrual cycle at the doses used in this study. Animals menstruated at regular intervals and progesterone concentrations indicative of ovulation were present at the expected frequency in the TCDD-treated animals. Only one monkey in the 5-ng group was deemed to be anovulatory during TCDD exposure. Similar results were reported in several studies that exposed female rats and monkeys to similarly low levels of TCDD. Kociba et al. (1978) reported no effects on the reproductive system in rats in a 2-year study using doses of 1 to 10 ng/kg/day. Rhesus monkeys fed a diet containing 500 ppt TCDD for 9 months, which translated to approximately 10 ng/kg/day, exhibited menstrual cycles of normal length, intensity, and duration (Barsotti et al., 1979Go). Allen et al. (1979) reported normal menstrual cycles in rhesus monkeys fed a diet containing 50 ppt TCDD for 6 months, while Bowman et al. (1989) reported no adverse effects on reproduction in female rhesus monkeys exposed to 5 or 25 ppt TCDD for 7 months. In contrast, inhibition of reproductive function has been clearly demonstrated in studies that employed higher TCDD doses. Inhibition of estrous cyclicity, ovulation, and decreased fertility in rats have been reported (Gao et al., 2001Go; Gray and Ostby, 1995Go; Kociba et al., 1979Go). Similar adverse effects on reproduction were observed in monkeys (Allen et al., 1979Go; McNulty, 1985Go).

The doses of TCDD used in this study had no effect on body weight. A loss in body weight has been shown to be the hallmark sign of TCDD toxicity when animals are exposed to lethal doses of TCDD (Peterson et al., 1984Go). Kociba et al. (1979) reported that body weights of male and female rats given low TCDD doses (1 ng TCDD/kg/day) were unaffected, whereas a higher dose (100 ng TCDD/kg/day) reduced body weights. Body weights of rhesus monkeys fed a diet containing 500 ppt TCDD for 9 months declined, although their food intake was unaltered (Allen et al., 1977Go).

A significant reduction in interleukin 6 was observed in the group of monkeys exposed to the highest TCDD dose, as reported by Yang et al. (2000) in their companion study. Chronic activation of the HPA axis is associated with compromised immune function (Bateman et al., 1989Go). Our observation of increased CRH mRNA combined with increased cortisol secretion is indicative of increased HPA axis activity and may offer an explanation of compromised immune function in these animals.


    ACKNOWLEDGMENTS
 
This work was supported by the Medical Research Council of Canada.


    NOTES
 
1 To whom correspondence should be addressed at the Department of Obstetrics and Gynecology, 3022 Etherington Hall, Queen's University, Kingston, Ontario, Canada K7L 3N6. Fax: (613) 533-6779. E-mail: vanvugtd{at}post.queensu.ca. Back


    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
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