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1 Department of Animal and Avian Sciences, University of Maryland, College Park, Maryland 20742, USA
2 Molecular and Cell Biology Program, University of Maryland, College Park, Maryland 20742, USA
(Requests for offprints should be addressed to T E Porter; Email: TEPorter{at}umd.edu)
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-subunit. Thyrotropin-releasing hormone (TRH) stimulates secretion of TSH-like bioactivity from pituitaries cultured in vitro (Scanes 1974), and increases plasma levels of thyroxine (T4) in vivo (Kuhn et al. 1988). In addition to TRH, corticotropin-releasing hormone (CRH) has been shown to induce TSH release in chickens, and somatostatin can diminish the CRH- and TRH-induced TSH response through the type 2 and type 5 somatostatin receptors (Geris et al. 1996, 2003a,Geris et al. b, DeGroef et al. 2003). Thyroid-stimulating activity has been localized to the cephalic lobe of the anterior pituitary (Brasch & Betz 1971, Radke & Chiasson 1974, 1977), and cells containing immunoreactive TSH have been localized to the cephalic lobe in histological sections stained with antisera against mammalian TSH (Sharp et al. 1979, Thommes et al. 1983, Mikami 1986, Murphy & Harvey 2001, 2002, Sasaki et al. 2003) or with an antiserum raised against a peptide based on the chicken TSHß sequence (Iwasawa et al. 2002, Nakamura et al. 2004). Similarly, pituitary cells that stained with monoclonal antibodies against the chicken -subunit and that did not also stain with anti-chicken luteinizing hormone (LH)ß and follicle-stimulating hormone (FSH) ß monoclonal antibodies were localized to the cephalic lobe (Berghman et al. 1993). Thommes et al.(1983) reported that the thyrotrophs were first detectable on embryonic day (e) 6.5 and dramatically increased on e11.5, the time in embryonic development when the HPT axis is thought to become functionally mature in chickens (Thommes et al. 1983). However, Murphy & Harvey (2001, 2002) found no immunoreactive TSH in the pituitary on e7, and Nakamura et al.(2004) first detected TSH-containing cells on e10. The -subunit mRNA is expressed as early as e3.5 (Kameda et al. 2000), while TSHß mRNA levels are present from e9 and increase at the end of embryonic development (Gregory et al. 1998, Nakamura et al. 2004). Depression of plasma thyroid hormone levels by goitrogen administration has been shown to increase the size of purported thyrotrophs and the thyroid-stimulating bioactivity of the pituitary, suggesting that thyroid hormones exert negative feedback on TSH production in mature birds (Sharp et al. 1979), and treatment of e19 pituitary cells in vitro with triiodothy-mRNA ronine (T3) significantly decreased levels of TSHß (Gregory et al. 1998). However, nothing is known about the regulation of TSH protein production by endogenous thyroid hormones in the developing chick embryo, even though production of thyroid hormones is essential for hatching and progression to endothermic life. Pituitary TSHß mRNA levels increase prior to hatching, coincident with an increase in plasma T4 and T3, raising the possibility that pituitary TSH production is free from the negative effects of thyroid hormones at this critical juncture of development. In the present study we validated an antiserum raised against purified rat TSHß subunit to detect chicken TSHß peptide. This antiserum was used in immunocytochemistry, along with whole-mount in situ hybridization, to characterize the distribution of thyrotrophs within the cephalic lobe of the anterior pituitary and to quantify the abundance of thyrotrophs during embryonic development. Finally, TSHß gene and peptide expression were examined following in vivo goitrogen treatment of developing embryos, to assess involvement of thyroid hormone negative feedback in controlling TSH production prior to hatching. We report that TSHß gene expression and pituitary content are regulated by negative feedback from endogenous thyroid hormones by the end of chick embryonic development, even though levels of thyroid hormones, pituitary TSH gene expression, and pituitary thyrotrophs all increase prior to hatching. These findings suggest that hypothalamic feed-forward stimulation, rather than a lack of thyroid hormone negative feedback, is the predominant cause of the increased HPT axis activity at the end of embryonic development necessary for hatching and endothermic existence. |
Materials and Methods |
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All animals were Avian x Avian broiler strain chicken embryos obtained from Allen’s Hatchery (Seaford, DE, USA). Eggs were set in a humidified incubator at 37.5 °C, and that day was designated as e0. Ethanol-swabbed eggs were injected with a sterile 1 ml syringe by inserting the needle (25-gauge) into the albumen at the junction of the air cell, which was visualized by candling and marked. Chicken eggs received injections (in 100 µl sterile water) of 125 µg of the thyroid hormone synthesis inhibitor methimazole (MMI) on e13, 250 µg on e15, 500 µg on e17, and 750 µg on e19. MMI doses were chosen based on preliminary studies that defined doses effective at suppressing development and hatching but which were non-lethal. Control eggs were injected with sterile water. Water was used as the vehicle because the injections were made into the water-containing albumen. The injection hole was sealed with cellophane tape. Animals were used with approval from our Institutional Animal Care and Use Committee.
Western blotting
Chicken pituitaries were isolated, pooled, and homogenized in either 0.1 M HCl, or a protein extraction buffer (10% glycerol, 15 mM HEPES pH 7.9, 300 mM NaCl, 50 mM KCl, 0.5% deoxycholate, 0.5% CHAPS, 0.5% Triton-X 100, 1 x proteinase inhibitor cocktail (Roche Molecular Biochemicals, Mannheim, Germany) and 2 mM EDTA pH 8.0) in a 1 ml Dounce homogenizer (Wheaton, Millville, NJ, USA). Protein concentrations in pituitary extracts were quantified by the bicinchoninic acid assay (Pierce; Rockford, IL, USA) according to manufacturer’s directions and extracts were separated on a 12.5% SDS-polyacrylamide gel using a Tris–Tricine-SDS running buffer under a current of 60 mA. Proteins were transferred overnight to a 0.45µm supported-nitrocellulose membrane (Bio-Rad; Hercules, CA, USA) using a 0.2 M phosphate buffer at 22 V. The membrane was washed twice for 5 min in PBS (pH 7.4) and then incubated for 30 min in 0.3% H2O2 in PBS. The membrane was washed again for 5 min in PBS and then blocked with 1% normal goat serum (NGS) for 1 h. After blocking, the membrane was incubated with the rabbit anti-rat TSHß antiserum (NIDDK-anti-rBetaTSH-IC-1) diluted 1:10 000 in 0.05% Tween 20-PBS (T-PBS) with 1% NGS for 3 h. The membrane was washed five times for 5 min with T-PBS and then incubated with a goat anti-rabbit biotinylated IgG secondary antibody (Vector Laboratories, Burlingame, CA, USA) in T-PBS with 1% NGS for 2 h. Again the membrane was washed 5 times for 5 min with T-PBS. After washing, the membrane was incubated with Vectastain ABC reagent (Vector Laboratories), an avidin–biotin signal amplification complex, for 30 min. The membrane was washed again with PBS and developed using the Vectastain VIP horseradish peroxidase substrate kit (Vector Laboratories). When statistical tests were applied, bands were scanned and quantified using ImageQuant software (Molecular Dynamics; Sunnyvale, CA, USA). The NIDDK-anti-rBetaTSH-IC-1 and purified rat TSH were obtained through NHPP, NIDDK and AF Parlow (Harbor UCLA Medical Center, Torrance, CA, USA). L Berghman (Texas A&M University, College Station, TX, USA) and J Proudman (Germplasm and Gamete Physiology Laboratory, USDA, Beltsville, MD, USA) kindly provided the chicken LH and FSH, respectively, used in the validation of the rat TSHß antiserum for chicken TSHß.
Immunocytochemistry (ICC)
Cell culture reagents were from Invitrogen Gibco (Grand Island, NY, USA) unless otherwise noted. Pituitaries were removed from embryos using a dissecting microscope and dissociated by trypsin digestion (1 mg/ml) in SMEM (supplemented with 0.1% BSA, 100 U/ml penicillin and 100 µg/ml streptomycin) and mechanical agitation at 37.5 °C under 5% CO2/95% O2 for 45 min, triturating and gassing every 15 min as previously described (Porter et al. 1995). ICC was performed as previously described (Liu et al. 2003, Bossis et al. 2004), with the following changes. Rat TSHß antiserum (NIDDK-anti-rBetaTSH-IC-1) was used as the primary antiserum at 1:10 000. Cells were washed and then incubated with Vectastain ABC reagent. Finally, the plate was washed again with PBS and developed using the Vectastain VIP horseradish peroxidase substrate kit. Cells were counted using an inverted light microscope and positively stained cells expressed as a percentage of all cells counted. Counting of pituitary cells began in the center of each well, and a minimum of 300 cells was evaluated for each duplicate well for each age in each replicate experiment.
Whole-mount in situ hybridization
The whole-mount in situ hybridization procedure, with modifications noted below, was performed as previously described (Bossis & Porter 2000). Briefly, pituitaries were isolated from e19 embryos and fixed. Pituitaries were then dehydrated in an ethanol series, stored overnight, rehydrated the next day, permeabilizied with Proteinase K, and then post-fixed. Pituitaries were hybridized with digoxigenin-labelled riboprobes overnight, washed with 2 x and 0.2 x SSC, incubated with alkaline phosphatase-conjugated sheep anti-digoxigenin antibody (1:2000), and developed with 4-nitro blue tetrazolium chloride and 160 µg/ml 5-bromo-4-chloro-3-indolylphosphate (Roche Applied Science, Indianapolis, IN, USA). The sense and antisense digoxigenin-labelled riboprobes (Boehringer Mannhein, Mannheim, Germany) were generated from a plasmid previously described (Gregory et al. 1998).
Whole-mount immunocytochemistry
Pituitaries from e19 embryos were isolated and fixed in 3.7% formaldehyde at 4 °C for 30 min. Pituitaries were washed in PBS and dehydrated in an increasing methanol series (50%, 75%, 95%, and 100%), quenched with 0.3% in methanol for 5 min, and stored overnight in H2O2 methanol at –20 °C. Pituitaries were rehydrated the next day, permeabilized with 0.02% trypsin in 0.1% TritonX-100 in PBS (PBT) for approximately 4 min, and then the digestion was stopped by quenching in 0.1% soybean trypsin inhibitor in PBT with 1 mM CaCl2 (Sigma, St Louis, MO, USA) for 2 min. Pituitaries were washed in PBT three times for 5 min. The pituitaries were blocked in 2% NGS for 1 h and incubated overnight at 4 °C with the rabbit anti-rat TSHß antiserum (1:5000) in PBS with 1% NGS. Pituitaries were then washed six times for 5 min in PBT and then incubated with biotinylated anti-rabbit IgG (Vector Laboratories) for 2 h. Pituitaries were washed in PBT three times for 5 min, then three times for 10 min, and incubated in Vectastain ABC reagent for 1 h. Pituitaries were washed in PBT three times for 5 min, three times for 20 min, and then developed with Vectastain VIP horseradish peroxidase substrate kit.
Plasma collection and radioimmunoassays (RIAs)
Plasma was collected by removing the shell apical to the air sac junction, visualizing the blood vessels by moistening the inner shell membrane, nicking a vessel with a 25-gauge needle, and drawing the blood into a 1 ml syringe wetted with 0.5 M EDTA. Blood samples were discarded if egg albumen became mixed with blood. The hematocrit was checked by visual inspection. Collected blood samples were transferred to 1.5 ml microcentrifuge tubes on ice containing 2.5 µl of 0.5 M EDTA. Blood samples were centrifuged, and the plasma was collected and frozen at –20 °C until assayed. RIA was performed using the Coat-A-Count Free T4 RIA kit from Diagnostic Products Corporation (Los Angeles, CA, USA) according to the manufacturer’s protocol. Two different assays were used in attempts to assess plasma levels of TSH, the rat TSH RIA from American Laboratory Products Company (Windham, NH, USA) and the human TSH immuno-radiometric assay from Diagnostic Products Corporation. Neither TSH assay proved effective at measuring chicken TSH.
RNA extraction and Northern blotting
Pituitaries were isolated from embryos and snap frozen in liquid nitrogen and stored at –80 °C until RNA extraction. Pituitaries were homogenized in a 1 ml Dounce homogenizer and RNA was isolated using TRIzol according to the manufacturer’s protocol (Invitrogen Gibco). RNA was suspended in nuclease-free water, quantified by absorbance at 260 nm, and then stored at –80 °C until use. Approximately 5 µg of total RNA/sample in RNA Loading Buffer with Ethidium Bromide (Sigma) were separated on 1% agarose-6% formaldehyde gels with MOPS (Invitrogen Gibco) running buffer at 200 V and transferred overnight to a Zeta-Probe nylon membrane (Bio-Rad, Hercules, CA, USA) by capillary action. The RNA was cross-linked to the membrane in a UV cross-linker and pre-hybridized at 65 °C in PerfectHyb Plus hybridization solution (Sigma) for 3 h in a rotating hybridization oven. 32-P dCTP (Amersham Biosciences, Piscataway, N)-labelled TSHß cDNA probes were generated by PCR using Taq polymerase (MBI Fermentas; Hanover, MD, USA). The TSHß template: GTGGAG AAGCGGGAGTGTGCCTACTGCCTGGCCACAAC ACCACCATCTGCGCTGGATTTTGCATGACACG GGACAGCAACGGCAAGAAGCTGCTACTCAAA AGTGCTCT and primer: AGAGCACTTTTGAGTAG CAG were synthesized by Sigma-Genosys. PCR products were separated from unincorporated nucleotides by centrifugation through a Spin Column-30 (Sigma) and counted in a beta counter to determine radioactivity. Approximately 1x106 counts per minute (c.p.m.) of probe was added per ml of hybridization solution and allowed to hybridize overnight at 65 °C. The membrane was then washed twice at low stringency with 2 x SSC (300 mM NaCl, 30 mM sodium citrate) –0.1% SDS at room temperature. The membrane was washed with 0.2 x SSC–0.1% SDS twice at 65 °C, and then twice with 0.1x SSC–0.1% SDS at 65 °C for high stringency washes. Then, the membrane was placed between two sheets of transparency acetate inside a film cassette, allowed to expose X-ray film (X-OMAT, Eastman Kodak Co, Rochester, NY, USA) at –80 °C for 2 days with intensification screens and the film was developed. Bands were scanned and quantified using ImageQuant software (Amersham Biosciences).
Statistical analyses
Data are reported as the least squares means±S.E.M. Analysis of variance (ANOVA) was used to compare means in the ontogeny experiment and the treatment effects of MMI injections (SAS version 8.02, PROC GLM or PROC MIXED procedures, SAS Institute, Cary, NC, USA). Differences were considered significant at P<0.05.
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Results |
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Initial experiments evaluated whether the anti-rat TSHß antiserum could detect chicken TSHß without cross-reacting with the other pituitary glycoprotein hormones, namely LH and FSH, since they all share the same -subunit. Chicken pituitaries were dissected from three d1, d6, and e18 embryos, and five e20 embryos, pooled and extracted. Protein from heart, liver, spleen, and muscle tissues of 3 week-old chicks was also extracted. 10 ng of purified rat TSH, 1 µg each of chicken FSH and chicken LH, and 5 µg of total protein from pituitary, heart, liver, spleen, and muscle were separated on a polyacrylamide gel and immunoblotted with the anti-rat TSHß antiserum. Two bands of approximately 30 kMr and 17 kMr were detected in the chicken pituitary extract (Fig. 1
). The rat TSH lane also contained 2 bands of nearly identical molecular weight. The lower band at 17 kMr is presumably the ß-subunit, and the higher band at 30 kMr likely represents the /ß heterodimer. The antiserum only detected bands in the pituitary extracts and not in other tissues. More importantly, no cross-reaction with either chicken LH or FSH was seen, even though 100-fold more protein was tested for each than for TSH. This indicates that the antiserum recognizes chicken TSHß and does not recognize LH and FSH or the common -subunit.
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-subunit. Taken together, these two experiments indicate that the antiserum is specific for chicken TSHß and can be used for both Western blotting and ICC. Ontogeny of thyrotrophs
Although levels of TSHß mRNA have been characterized during development (Gregory et al. 1998, Nakamura et al. 2004), changes in the abundance of anterior pituitary thyrotrophs have not been quantified. Pituitaries were isolated from embryos on e11, e13, e15, e17, e19, and d1 and dispersed, plated, and subjected to ICC using the anti-rat TSHß antiserum. Results are shown in Figure 2. TSHß-containing cells comprised 0.7±0.1%, 4.0±0.4%, 5.3±0.4%, 7.0±0.8%, 6.7±0.7%, and 3.9±0.4% of all pituitary cells on e11, e13, e15, e17, e19, and d1, respectively. Thus, thyrotrophs were rare on e11, and their abundance was maximal on e17 (P<0.01, n = 4 separate trials).
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Early reports indicate that thyroid-stimulating activity and immunoreactive TSH are found in the cephalic lobe of the anterior pituitary gland in birds (Brasch & Betz 1971, Radke & Chiasson 1974, 1977, Sharp et al. 1979, Thommes et al. 1983, Murphy & Harvey 2001). More recently, cells containing the -subunit but not the ß-subunits for LH and FSH were found in the cephalic lobe in sagittal histological sections (Berghman et al. 1993). To evaluate three-dimensional distribution of thyrotrophs within the cephalic lobe prior to hatching, whole-mount in situ hybridization for TSHß mRNA and whole-mount ICC for TSHß peptide were performed. Pituitaries were isolated on e19, when thyrotrophs were abundant and TSHß mRNA levels were reported to be maximal (Gregory et al. 1998). Individual pituitaries were probed with either a TSHß antisense riboprobe or the anti-rat TSHß antiserum. Representative results are presented in Figure 3. The presence of individual thyrotrophs is evident from the specific punctate staining. Thyrotrophs were not distributed uniformly among pituitary glands. TSHß mRNA-containing cells were localized within the cephalic lobe of the e19 pituitary in three general patterns: a focal spot in the ventral medial region of the cephalic lobe (Fig. 3A); a "saddle" pattern extending from the ventral medial to the dorsal lateral regions of the cephalic lobe along the cephalic–caudal boundary (Fig. 3B); and an even distribution within the entire cephalic lobe (Fig. 3C). Hybridization with a sense riboprobe was negative (Fig. 3D). Whole-mount ICC produced similar results. TSHß-containing cells were found in a focal spot in the ventral region of the cephalic lobe (Fig. 3E), in a "saddle" pattern (Fig. 3F), and distributed throughout the cephalic lobe (Fig. 3 G). No staining was seen when the anti-rat TSHß antiserum was omitted (Fig. 3H). Thus, results from both whole-mount in situ hybridization and whole-mount ICC indicate heterogeneous distribution of thyrotrophs within the cephalic lobe. To confirm cephalic lobe localization of thyrotrophs, the cephalic and caudal distal ends (approximately one third) of anterior pituitaries were isolated in situ within the sella turcica on e19, as previously described (Lopez et al. 1995). By using only the caudal or cephalic third of each individual pituitary, separation of the lobes was ensured. Pools were then made of caudal and cephalic lobes from at least three embryos for each replicate experiment. Growth hormone (GH)- and prolactin (PRL)-containing cells were identified by ICC, as described previously (Lopez et al. 1995, Liu et al. 2003, Bossis et al. 2004). The lobes were then dispersed, plated, and subjected to ICC for TSHß, GH (a caudal lobe marker), and PRL (a cephalic lobe marker). TSHß- and PRL-containing cells were found among cells isolated from the cephalic lobe, while GH-positive cells were predominantly found in the caudal lobe (Fig. 4). Taken together, these findings from single cell ICC, whole-mount ICC, and whole-mount in situ hybridization confirm that thyrotrophs are localized to the cephalic lobe of the chicken anterior pituitary gland. Furthermore, results from whole-mount ICC and in situ hybridization indicate that thyrotroph distribution is heterogeneous within the cephalic lobe among individual embryos, with a "saddle-shaped" pattern being the most common distribution pattern seen on e19.
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We reported previously that TSHß mRNA levels and plasma thyroid hormone levels increase prior to hatching (Gregory et al. 1998). We also found thatT3 could suppress TSHß mRNA levels in cultures of e19 pituitary cells, indicating the potential for feedback regulation. However, the contribution of negative feedback from endogenous thyroid hormones on TSHß mRNA levels and pituitary TSH content during this critical period of embryonic development has not been reported. Endogenous thyroid hormone production in the chick embryo was suppressed with MMI injections on e13, e15, e17, and e19. MMI-injected embryos were less developed than the controls and did not hatch. This is consistent with reported effects of MMI on chicken embryos by a continuous flow delivery system (Iqbal et al. 1987). To confirm MMI suppression of thyroid hormones, a RIA for free T4 was performed. In control embryos, T4 increased more than 6-fold, from 0.025±0.004 ng/dl on e17, to 0.072 ± 0.03 ng/dl on e19, to 0.162±0.044 ng/dl on d1 (Fig. 5). T4 was undetectable in the plasma of MMI-treated embryos. Next, identical experiments were performed, and pituitaries were collected and pooled from five to seven embryos for each treatment on e17, e19, and d1. Total RNA was isolated from pituitaries of MMI-injected and control embryos on e17, e19, and d1, and analyzed by northern blotting. As seen in Fig. 6, TSHß mRNA levels were increased by MMI treatment (n = 6 replicate experiments; P<0.01; across all ages), with significant differences between MMI and control groups found on e19 and d1 (P<0.05). Levels of TSHß mRNA in control groups were: e17, 1.61±0.40 R.D.U. (relative densitometric units; ratio of TSHß/ß-actin mRNA in arbitrary densitometric units); e19, 1.21±0.42 R.D.U.; d1, 1.24±0.48 R.D.U. Levels of TSHß mRNA in MMI-treated groups were: e17, 2.00±0.40 R.D.U.; e19, 2.07±0.40 R.D.U.; d1, 2.58±0.42 R.D.U. These results indicate that TSHß gene expression during perinatal development of the chick is under negative feedback regulation from endogenous thyroid hormones. Next, experiments were performed to determine the effect of MMI on TSHß protein levels. Protein was extracted from pituitaries pooled from five to seven MMI-injected and control embryos on e17, e19, and d1, and analyzed by Western blotting (Fig. 7). Control pituitaries contained more TSHß protein than the MMI treated groups (n = 3 replicate experiments; P<0.01; across all ages), with significant differences between MMI and control groups found on e17 and e19 (P<0.05). TSHß protein levels in R.D.Us (intensity of the 17 kMr band normalized to the 17 kMr band of rat TSHß standard in arbitrary densitometric units) for the control groups were: e17, 0.336±0.08 R.D.U.; e19, 0.350±0.08 R.D.U.; d1, 0.246±0.08 R.D.U. Levels of TSHß protein in MMI-treated pituitaries were: e17, 0.108±0.08 R.D.U.; e19, 0.090±0.08 R.D.U.; d1, 0.140±0.08 R.D.U. Thus, inhibition of endogenous thyroid hormone synthesis resulted in decreased pituitary TSHß protein content.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Thyrotrophs have been localized in the cephalic lobe of the anterior pituitary by ICC on histology sections and by assessment of biological activity in tissue extracts (Brasch & Betz 1971, Radke & Chiasson 1974, 1977, Sharp et al. 1979, Thommes et al. 1983, Berghman et al. 1993, Murphy & Harvey 2001, 2002, Iwasawa et al. 2002, Nakamura et al. 2004). However, the three-dimensional distribution of thyrotrophs within the cephalic lobe of the anterior pituitary has not been determined. To accomplish this, in situ hybridization and ICC were performed in the present study on whole-mounted pituitary glands. As expected, the cTSHß riboprobe detected TSHß mRNA in the cephalic lobe of the anterior pituitary. Interestingly, different patterns of expression were noted. In some pituitaries, the thyrotrophs were localized to a small region in the medial ventral area of the cephalic lobe; in other pituitaries, cells throughout the entire cephalic lobe were stained for TSHß mRNA. There was also a "saddle-shaped" staining pattern observed, extending from the medial ventral region of the cephalic lobe to the lateral dorsal regions along the cephalic-caudal boundary of the anterior pituitary. Similar patterns were seen when ICC for TSHß protein was performed on whole-mounted pituitaries. The saddle-shaped pattern was most commonly observed. These two sets of experiments, probing for both TSHß mRNA and TSHß peptide, prove conclusively that the thyrotrophs are localized to the cephalic lobe of the chick anterior pituitary. This is consistent with the reports of immunopositive cells in cephalic sections and homogenates using anti-peptide and heterologous antisera (Sharp et al. 1979, Thommes et al. 1983, Mikami 1986, Murphy & Harvey 2001, 2002, Iwasawa et al. 2002, Sasaki et al. 2003, Nakamura et al. 2004). These findings also indicate that the distribution of thyrotrophs is heterogeneous among individual embryos at this stage of development. Interestingly, in mice a population of thyrotrophs exists transiently during development at the rostral tip of the pituitary in addition to the caudomedial, permanent thyrotrophs (Lin et al. 1994). It is unknown whether this is also true in chickens, as only pituitaries on e19 were examined. However, the different patterns of thyrotroph localization noted in the present study using both ICC and in situ hybridization make it tempting to speculate that thyrotroph differentiation during chick development initiates in the ventral medial region of the cephalic lobe and proceeds in a dorsal and then rostral fashion so that thyrotrophs are eventually dispersed throughout the cephalic lobe of the anterior pituitary gland.
Classically, thyroid hormones are thought to feed back to suppress the release of TSH from the pituitary. However, this has not been determined during chick embryonic development. T4 and T3 secretion increases prior to hatching (Gregory et al. 1998). Our finding of increased numbers of thyrotrophs and previous findings of elevated TSHß mRNA levels in the pituitary at the end of embryonic development (Gregory et al. 1998) raises the possibility that TSH production is not regulated by negative feedback from endogenous thyroid hormones during this period of development. To explore this possibility, endogenous thyroid hormone synthesis was suppressed by injecting eggs with MMI. Effectiveness of MMI injections was confirmed by measuring plasma levels of T4. T4 was below the limit of detection for all three ages of MMI-treated embryos assayed. The control embryos showed increasing levels of T4 prior to hatching, as expected. MMI treatment increased levels of TSHß mRNA in the pituitary. This increase in TSHß mRNA suggests that TSHß gene expression is normally under negative feedback by thyroid hormones during this period of development. Somewhat surprisingly, control pituitaries contained more TSHß protein than the MMI-treated pituitaries. Thus, MMI-treated pituitaries contain more TSHß mRNA but less TSHß protein. This finding is likely due to increased secretion of TSH protein by the thyrotrophs into the bloodstream in MMI-treated embryos. The pituitary, therefore, is depleted of TSH as compared with control embryos. Alternatively, this may indicate that TSHß protein synthesis was decreased in response to MMI treatment. However, given that TSHß mRNA levels were increased by MMI, this possibility would seem unlikely. Similar findings of decreased staining intensities in purported thyrotrophs were noted when older, post-hatch birds were fed MMI (Sharp et al. 1979, Berghman et al. 1993). Taken together, the increase in TSHß mRNA levels in combination with diminished pituitary TSHß protein content in MMI-treated embryos clearly indicate that pituitary thyrotrophs are normally under tonic inhibition by endogenous thyroid hormones prior to hatching. Furthermore, these findings indicate that the increase in thyroid hormone levels, in pituitary thyrotrophs, and in pituitary TSHß mRNA and protein levels prior to hatching are not due to a lack of negative feedback inhibition within the HPT axis. Rather, our findings suggest that the increase in thyroid hormone levels critical to hatching and the transition to endothermic life is driven either by an autonomous increase in pituitary thyrotrophs and TSH secretion or by an increase in hypothalamic stimulation of pituitary thyrotrophs. Levels of hypothalamic TRH are known to increase prior to hatching (Geris et al. 1998, 1999), supporting the notion that ultimately the hypothalamus controls the metabolic jump to endothermic life.
In conclusion, we report that the number of thyrotrophs increases dramatically between e11 and hatching, that their distribution within the cephalic lobe is heterogeneous on e19 but most commonly in a saddle-shaped pattern, and that TSHß gene expression and protein secretion are regulated by negative feedback from endogenous thyroid hormones during the perinatal period of chick embryonic development, when the transition to endothermic life occurs. These findings indicate that the increased activity of the HPT axis necessary for hatching and endothermic life is not due to the absence of thyroid hormone negative feedback on pituitary TSH production.
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Acknowledgements |
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References |
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Received 28 September 2004
Accepted 11 November 2004
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