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Endocrinology, doi:10.1210/en.2005-1223
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Endocrinology Vol. 147, No. 8 3943-3951
Copyright © 2006 by The Endocrine Society

Molecular Cloning, Tissue Distribution, and Ontogenic Thyroidal Expression of the Chicken Thyrotropin Receptor

Sylvia V. H. Grommen, Shusuke Taniuchi, Tom Janssen, Liliane Schoofs, Sumio Takahashi, Sakae Takeuchi, Veerle M. Darras and Bert De Groef

Laboratories of Comparative Endocrinology (S.V.H.G., V.M.D., B.D.G.) and Developmental Physiology, Genomics, and Proteomics (T.J., L.S.), Katholieke Universiteit Leuven, B3000 Leuven, Belgium; and Department of Biology (S.Tan., S.Taka., S.Take.) Faculty of Science, Okayama University, Okayama 700-8530, Japan

Address all correspondence and requests for reprints to: Sylvia Grommen, Laboratory of Comparative Endocrinology, Naamsestraat 61, B-3000 Leuven, Belgium. E-mail: sylvia.grommen{at}bio.kuleuven.be.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
TSH and the interaction with its receptor (TSHR) in the thyroid gland play a crucial role in the pituitary-thyroid axis of all vertebrates. Released upon stimulation by TSH, thyroid hormones influence numerous processes in the body and are extremely important during the last week of chicken embryonic development. In this study, we have cloned and functionally characterized the chicken TSHR (cTSHR), which was found to be a G protein-coupled receptor consisting of 10 exons. Besides the full-length cDNA, we detected two splice variants lacking either exon 3, or exons 2 and 3, both part of the extracellular domain of the receptor. Bovine TSH increased intracellular cAMP levels in HEK-239 cells transiently expressing the full-length cTSHR (EC50 = 1.43 nM). In situ hybridization showed the expression of cTSHR mRNA in the thyroidal follicular cells. cTSHR mRNA expression, as determined by real-time PCR, was also found in several other tissues such as brain, pituitary, pineal gland, and retina, suggesting that the TSH-TSHR interaction is not only important in regulating thyroid function. TSHR mRNA expression in the thyroid gland did not change significantly during the last week of embryonic development, which suggests that an increased thyroidal sensitivity is not part of the cause of the concomitant increasing T4 levels.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
IT IS WELL ESTABLISHED that thyroid hormones (TH) are essential for normal growth, development, and function of several tissues including brain (1). Their release into the blood circulation is stimulated by the hypophyseal hormone TSH upon binding to its receptor (TSHR), a G protein-coupled glycoprotein. Several studies in rodents have demonstrated that TSH, by binding to its receptor, transcriptionally regulates the expression of genes involved in iodide metabolism such as thyroid peroxidase and the sodium-iodide symporter (2, 3, 4, 5). Besides the synthesis and release of TH, TSH seems to play an important role in terminal thyroid gland maturation and growth. Recent experiments in rat suggest that TSH, acting through its receptor, is not involved in early thyroid organogenesis or migration but plays an important role in terminal thyroid maturation and therefore in the development of a fully functional thyroid gland (6, 7). For the reasons mentioned above, the TSH/TSHR interaction can be regarded as the major regulator of thyroid function.

In chicken, TH concentrations are relatively low during the first half of embryonic life and gradually increase around 10 d of incubation, reaching maximal levels at 19–20 d (8, 9). A sharp increase in T4 is observed between d 15.5 and 19.5. Thyroid function in embryonic chickens is said to be autonomous from hypothalamic-pituitary control during the first half of embryonic development, whereas thyroid activity increases during the second half and then appears to be dependent on pituitary control (10, 11).

This paper describes the molecular cloning of the chicken TSHR (cTSHR). We have functionally characterized the full-length cTSHR and confirmed its presence in thyroid cells using in situ hybridization. To widen our knowledge about the central and peripheral target tissues of TSH, we have also examined cTSHR mRNA distribution in 24 different chicken tissues. Finally, we have investigated whether the strong rise in plasma T4 concentration in the last week before hatching could be partially caused by an increased sensitivity of the thyroid gland for TSH resulting from a higher TSHR expression as suggested by Scanes et al. (12). We have therefore measured cTSHR mRNA expression using real-time PCR in the thyroid gland starting from d 14 of embryonic development (E14) up until 1-d-old chicks (C1).


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
All experiments were approved by the ethical committee for animal experiments of the Katholieke Universiteit Leuven and the Okayama University.

Cloning of the cTSHR
Animals.
Seven-day-old chicks of a commercial broiler strain (Okayama chicken) were kindly provided by the Okayama Prefecture General Livestock Center (Okayama, Japan). Fertilized Rock Cornish eggs were purchased from Fukuda Poultry Breeding Farm (Okayama, Japan). One-day-old Ross broiler chickens (C1) were purchased from Vervaeke hatchery (Tielt, Belgium).

Sequence analysis of the full-length cTSHR cDNAs.
Since March 1, 2004, the first draft of the chicken genome sequence has been available at the following URL: http://www.ensembl.org/Gallus_gallus/. A predicted sequence of the cTSHR cDNA, XM_426455, was reported by the Ensembl pipeline analysis system. Primers used to clone the cTSHR were therefore based on XM_426455 and are indicated in Table 1Go.


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  		TABLE 1. Primers used to amplify the cTSHR with RT-PCR

 
Total RNA was prepared from pituitaries of 7-d-old Okayama chicks and from thyroid glands of 19-d-old Rock Cornish embryos and 1-d-old Ross chickens using Trizol reagent (Invitrogen, San Diego, CA) or the RNAgents Total RNA Isolation System (Promega, Madison, WI). The RNA was subsequently treated with DNase I (Invitrogen) or the DNA-free kit (Ambion, Austin, TX) according to the manufacturer’s directions.

For primers used to clone TSHR from pituitary gland (described in Table 1Go), 1 µg total RNA was reverse transcribed in a total volume of 20 µl using 15 U Thermoscript reverse transcriptase (Invitrogen) according to the manufacturer’s guidelines. A 0.5-µl aliquot of the reaction product was used in each PCR. PCR was carried out in a final volume of 25 µl using 0.625 U Takara Taq DNA polymerase (Takara, Otsu, Japan) in the Gene Amp PCR System 9600 (Applied Biosystems, Warrington, UK). The PCR cycle program was as follows: 1 min at 94 C, 35 cycles of 30 sec at 94 C and 30 sec at 66 C (FP5-RP4) or 60 C (FP4-RP2 and FP3-RP1), followed by an additional incubation of 10 min at 72 C. A 10-µl aliquot of each reaction was analyzed on a 2.0% agarose gel, stained with ethidium bromide, and photographed under UV illumination. The PCR-amplified cDNA fragments from pituitary total RNA were excised from the agarose gel and extracted and purified using the Concert Rapid Gel Extraction System (Invitrogen). Each of the cDNA fragments was amplified by a PCR with the same primers as the first PCR or with nested primers, and the resulting products were subsequently purified using the Concert Rapid PCR Purification System (Invitrogen). Ten to 15 ng PCR product was used for direct sequencing according to the protocol of the ABI PRISM Big Dye Terminator Cycle Sequencing Ready Reaction kit using the automatic sequencer PE ABI Prism 373A (Applied Biosystems). Sequence analyses were carried out using the GENETYX software.

For the primers used to clone TSHR from thyroid gland (Table 1Go), approximately 1 µg total RNA from the thyroid glands of C1 chicks was heated for 5 min at 70 C with oligo(dT) primer and then reverse transcribed to cDNA in a volume of 20 µl containing 2.5 U avian myeloblastosis virus (AMV) reverse transcriptase and reaction buffer (Roche Diagnostics, Basel, Switzerland), 0.5 mM of each dNTP, 10 U RNasin ribonuclease inhibitor (Promega), and 0.01 M dithiothreitol. Five microliters of the RT product were used for amplification in a 20-µl PCR containing 10x PCR buffer (100 mM Tris-HCl, 15 mM MgCl2, 500 mM KCl, pH 8.3), 1 mM of each dNTP, 1 µM forward primer and reverse primer, and 2.5 U Taq polymerase (Roche Diagnostics). PCR was performed in the GeneAmp PCR System 9700 (PerkinElmer, Wellesley, MA) with the following thermal cycle parameters: 1 min at 94 C, 32 cycles of 30 sec at 94 C and 30 sec at 59 C, followed by 10 min at 72 C. PCR products were analyzed on a 1.5% agarose gel and visualized with ethidium bromide. The PCR product was excised from the agarose gel, extracted, and purified using the QIAEX II Gel Extraction kit (QIAGEN, Hilden, Germany). The gel extraction products were subsequently subcloned with the TOPO TA Cloning kit (Invitrogen). Plasmid isolation was performed using the High Pure Plasmid Isolation kit (Roche Diagnostics). Sequencing of the products was done using the ABI PRISM Big Dye Terminator Cycle Sequencing Ready Reaction kit and the automatic sequencer ABI PRISM 310 Genetic Analyzer (Applied Biosystems).

The obtained cTSHR sequence was analyzed and protein domains were predicted using Prosite on the ExPASy Proteomics Server (http://us.expasy.org/). The seven potential transmembrane domains were determined using the TMHMM program (TMHMM Server version 2.0). The location of the putative signal peptide was predicted using the SignalP 3.0 Server (http://www.cbs.dtu.dk/services/SignalP/) (13). Multiple sequence alignments and a dendrogram were made using CLUSTAL W.

Functional characterization of the full-length cTSHR
Construction of a cTSHR expression vector.
To confirm that the cloned full-length cTSHR is encoding a functional receptor, total RNA was isolated from thyroid glands of 4-d-old Ross chickens and reverse transcribed using the AMV reverse transcriptase (Invitrogen) according to the manufacturer’s guidelines. The entire coding region was amplified using the Platinum Pfx DNA polymerase (Invitrogen) with reverse primer 8 (Table 1Go) and the forward primer 5'-CACCATGCTGTGGCTGCCTGTCGCC-3', which contains a Kozak sequence and bp 35–55 (Fig. 1Go). The PCR product corresponding to the full-length cTSHR was subsequently subcloned into the pcDNA3.1/V5-His TOPO expression vector (Invitrogen) and sequenced.


Figure 1
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  		FIG. 1. Nucleotide and deduced amino acid sequence of the cTSHR cDNA. Amino acids are annotated by the single-letter code. Numbers on the left refer to the position of the nucleotides (top) and the amino acids (bottom). The positions of the 10 exons are shown with vertical dotted lines. The predicted signal peptide is indicated in italics. The four potential glycosylation sites are indicated (bold and underlined) as well as a possible protein kinase C phosphorylation site (boxed). The sequence has been submitted to GenBank and is available under accession number AB234613.

 
Cell culture and transfections.
HEK-293 cells were maintained in DMEM supplemented with 10% fetal bovine serum and 100 IU/ml of a penicillin/streptomycin solution. They were split every 3 d (1:5) and grown at 37 C in a humidified atmosphere of 5% CO2 in air. Cells grown to 70% confluence were transiently transfected with Fugene 6 (Roche Diagnostics) using a 5:1 ratio of pcDNA3.1/cTSHR and a multimerized CRE-luciferase reporter construct (pCRE(6X)-Luc), kindly provided by Dr. Paul H. Taghert (Washington University, St. Louis, MO).

CRE-luciferase assay.
Twenty-four hours after transfection, cells were exposed to bovine TSH (bTSH) (Sigma-Aldrich, Bornem, Belgium) for 4 h in serum-free medium supplemented with 200 µM 3-isobutyl-1-methylxanthine. The luciferase activity was quantified using a LucLite Kit (PerkinElmer Life Sciences) and the luminescence measured on a Microlumat Plus LB96V microplate luminometer (EG&G Berthold, Bad Wildbad, Germany).

In situ hybridization
Cellular localization studies were performed on 18-d-old embryos from the Ross broiler strain. Animals were killed by decapitation, and the thyroid glands were removed and kept in 4% paraformaldehyde in PBS (pH 7.4) at 4 C. After 24 h, the tissues were cryoprotected overnight at 4 C in the same solution containing 30% sucrose. They were subsequently stored at –80 C until sectioning, and 7-µm sections were cut using a cryostat. Sections were kept at –20 C until usage.

Riboprobes were transcribed from 1 µg linearized pCR II-TOPO plasmid containing an insert of the cloned cTSHR (bp 35–391, Fig. 1Go) in the presence of digoxigenin (DIG) RNA labeling mix and 40 U RNA polymerase (SP6 for antisense and T7 for sense probes) (Roche Diagnostics). A spotting test was performed to test the efficiency of the labeling. The in situ hybridization was performed as described previously (14) using 400 ng/ml DIG-labeled riboprobe.

Tissue distribution of cTSHR mRNA
Tissue sampling.
Studies were performed on 4-d-old chicks (C4) of the Ross broiler strain. Animals were killed by decapitation, and the following tissues were collected, immediately frozen in liquid nitrogen, and stored at –80 C: thyroid glands (pooled per five pairs), pituitary glands (pooled per five), pineal glands (pooled per five), adrenal glands (pooled per three pairs), testes (pooled per three pairs), ovaries (pooled per three), retinal tissue, telencephalon, diencephalon, optic lobes, cerebellum, brain stem, proventriculus, gizzard, duodenum, intestine, liver, pancreas, kidneys, bursa, heart, spleen, lungs, and skeletal muscle.

RNA isolation and real-time PCR.
Total RNA was isolated using the RNAgents Total RNA Isolation System. The samples were subsequently treated with DNase I using the DNA-free kit to make sure no genomic DNA was present in the sample during PCR. For all tissues, except the ones that were pooled, 50 mg was used for isolation. Three individual samples or three pools were used to study the cTSHR distribution. Approximately 1 µg RNA was reverse transcribed using AMV reverse transcriptase (Invitrogen). The RT product was diluted 5-fold in water, and 5 µl of each diluted cDNA sample, corresponding to approximately 50 ng total RNA, was used for real-time PCR. Primers for cTSHR and the active reference ß-actin (accession no. NM_205518) were chosen using the Primer Express software version 2.0 (Applied Biosystems) and are listed in Table 2Go. Real-time PCR was performed in an ABI PRISM 7000 Sequence Detection System thermal cycler in a total volume of 25 µl containing 1x SYBR Green PCR Master Mix (Applied Biosystems), 300 nM forward primer, and 300 nM reverse primer. The following thermal cycle parameters were used: 2 min at 50 C, 10 min at 95 C, and 40 cycles of 15 sec at 95 C and 1 min at 60 C. A dissociation protocol followed the amplification program to detect nonspecific amplification. Each sample was analyzed in triplicate. For each gene, a nontemplate control and a sample that did not undergo RT were added as negative controls. Values were calculated using the {Delta}{Delta}Ct method after performing a validation experiment as described in Applied Biosystems User Bulletin No. 2 (P/N 4303859). Diencephalon was used as the calibrator.


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  		TABLE 2. Primers used to amplify the cTSHR, ß-actin, and GAPDH with real-time PCR

 
Thyroidal expression of cTSHR during the last week of embryonic development
Tissue sampling.
Fertilized eggs from the Ross broiler strain were purchased from Avibel (Halle-Zoersel, Belgium) and incubated in a laboratory incubator for 21 d as described previously (15). The start of incubation was called d 1 (E1). At d 20 of embryonic development (E20), we made a distinction between animals that had or had not perforated the membrane of the air chamber, which we indicated as the internal pipping (E20IP) and nonpipping stage (E20NP), respectively. Blood and tissue samples were taken every day starting from E14 until C1. After collecting blood samples by heart puncture (embryos) or by decapitation (posthatch chicks), thyroid glands were pooled per four pairs, frozen on dry ice, and stored at –80 C.

RNA isolation and real-time PCR.
Total RNA was isolated and treated with DNase I as described for the tissue distribution. RT was performed with the Taqman reverse transcription reagents (Applied Biosystems). The RT product was diluted 5-fold in water, and 5 µl of each diluted cDNA sample, corresponding to approximately 20 ng total RNA, was used for real-time PCR. Primers for cTSHR and the active reference glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (accession no. NM_204305) were chosen using the Primer Express software version 2.0 and are listed in Table 2Go. Real-time PCR was performed in an ABI PRISM 7000 Sequence Detection System thermal cycler as mentioned above. A 1:5 dilution series of thyroid gland cDNA was included as a standard on every 96-well optical reaction plate. Each sample was analyzed in triplicate, and standards were measured in duplicate.

Statistical analysis
Statistical analysis was performed using the SAS program (SAS Institute, Cary, NC). The values represented are the mean ± SEM. For the relative expression of the cTSHR in the thyroid gland, the Wilcoxon (rank sum) test was used to evaluate statistical differences between E14 and every other developmental stage.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
cTSHR sequence and structure
To clone the cTSHR, we used the predicted TSHR cDNA and genomic sequence with accession numbers XM_426455 and NW_060388, respectively. Preliminary experiments (not shown), using forward primers located at the 5' end and in exon 2 of XM_426455, indicated that neither predicted exon 1 nor exon 2 were present in the actual cTSHR cDNA sequence. The amino acid sequence deduced from NW_060388 and corresponding to the 3' part of intron 2 of the XM_426455 showed high levels of similarity with the N-terminal region of the reported mammalian TSHR sequences. We therefore chose an additional forward primer (FP5) corresponding to the region upstream from the predicted start codon. The actual full-length chicken cDNA sequence is shown in Fig. 1Go together with the deduced amino acid sequence. The 10 different exons are indicated. The nucleotide sequence is 2286 bp long and contains an open reading frame encoding a protein of 761 amino acids. The first 18 amino acids of the receptor protein constitute a putative signal peptide. The cTSHR has an extracellular domain containing four potential N-linked glycosylation sites, followed by a region with seven membrane-spanning domains, and a cytoplasmic region containing one potential protein kinase C phosphorylation site. In Fig. 2Go, the cTSHR amino acid sequence is aligned with that of human, rat, and African catfish, and the seven potential transmembrane domains as well as 11 conserved cysteine residues in the extracellular regions are indicated. To investigate the relationship of the cTSHR with some of the other known vertebrate TSHRs, a phylogenetic dendrogram was made using CLUSTAL W. The dendrogram (Fig. 3Go) shows that the cTSHR is more closely related to the mammalian TSHRs than to the fish TSHRs.


Figure 2
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  		FIG. 2. Deduced amino acid sequence of the cTSHR (Gallus gallus, Gg) compared with the sequence of Rattus norvegicus (Rn), Homo sapiens (Hs), and Clarias gariepinus (Cg). The conserved cysteine residues in the extracellular regions are marked with arrows. Identical amino acids have a black background, and similar amino acids have a gray background.

 

Figure 3
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  		FIG. 3. Phylogenetic dendrogram showing the evolutionary relationship of the cTSHR to some of the known vertebrate TSHRs.

 
Besides the full-length cTSHR cDNA, two additional splice variants were detected in the pituitary as well as in the thyroid gland, using primer pair FP5-RP4 (Figs. 4Go and 5Go). The sequence of the two different pituitary cTSHR variants was determined. The first splice variant, indicated as cTSHRb, seemed to lack exon 3 of the extracellular domain, whereas the second splice variant, designated as cTSHRc, lacked both exons 2 and 3. The cTSHR cDNA sequences obtained in the present study are available from DDBJ, EMBL, and GenBank data libraries under accession numbers AB234613, AB234614, and AB234615.


Figure 4
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  		FIG. 4. Representation of the cTSHR and the two splice variants, TSHRb lacking exon 3 and TSHRc lacking exons 2 and 3. The boxes represent the 10 different exons. The black box corresponds to the 34 bp preceding the translation start codon. The primers used to clone the cTSHR from pituitary and thyroid gland are indicated with arrows. Also the number of base pairs within each exon is shown. FP, Forward primer; RP, reverse primer.

 

Figure 5
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  		FIG. 5. cDNA fragments obtained from chicken pituitary and thyroid gland after RT-PCR with primer couples FP5-RP4, FP4-RP2, and FP3-RP1. The number of cycles used for PCR is indicated above each product. In the RT negative control (RT–), reverse transcriptase was omitted. To verify the integrity of the cDNA product, a PCR with GAPDH was performed, the primers of which are listed in Table 1Go. The different receptor variants are indicated with arrows. A 100-bp ladder was used with the 500-bp mark indicated on the left.

 
Functional characterization of the full-length cTSHR
Cultured HEK-293 cells transiently cotransfected with the pcDNA3.1/cTSHR construct and the pCRE(6X)-Luc reporter plasmid were challenged with various concentrations of bTSH in quadruplicate, followed by measurement of intracellular cAMP levels (Fig. 6Go). Bovine TSH was able to activate the cTSHR in a dose-dependent way with an EC50 value of 1.43 ± 0.04 nM.


Figure 6
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  		FIG. 6. Effect of bTSH on the intracellular cAMP production in HEK-293 cells transiently expressing the full-length cTSHR and CRE-luciferase. Data are presented as means ± SEM of four replicates from a single representative experiment indicated as percent activation. The value in parentheses represents the EC50 value of bTSH.

 
Cellular localization of the cTSHR in the thyroid gland
cTSHR mRNA transcripts were detected in the thyroid gland of a C4 chick using in situ hybridization, with a DIG-labeled riboprobe. The probe was directed against the first 356 bp of the full-length cTSHR cDNA, which include the second and third exon. The antisense probe gave a clear signal in the follicular cells (Fig. 7AGo). Hybridization with a sense probe produced no signal (Fig. 7BGo).


Figure 7
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  		FIG. 7. A, Localization of the cTSHR in the thyroid follicular cells of a 4-d-old chick using an antisense riboprobe; B, result of hybridization with the sense probe as a negative control.

 
Tissue distribution of cTSHR mRNA
To study the distribution of cTSHR mRNA in C4 chicks, a real-time PCR assay was performed using primers located in a region of the extracellular domain of the receptor that is common for the three receptor variants. The cTSHR expression was clearly most abundant in thyroid gland, with a 100-fold higher expression level compared with diencephalon. Also in brain (especially diencephalon, brain stem, and optic lobes), pineal gland, retina, and pituitary, cTSHR mRNA could be detected. From all the peripheral tissues studied, skeletal muscle, heart, and pancreas showed the highest expression (Fig. 8Go). In the other tissues, low or no cTSHR expression was seen.


Figure 8
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  		FIG. 8. Distribution of the cTSHR in different tissues of a representative 4-d-old chick, measured with real-time PCR. Results are calculated with the {Delta}{Delta}Ct method and expressed as relative values ± SEM, using diencephalon as the calibrator (n = 3 tissues or 3 pools of tissues).

 
Thyroidal expression of cTSHR during the last week of embryonic development
To investigate the expression of the cTSHR in the thyroid gland during the last week of development of the chicken embryo, a real-time PCR assay was performed with the same primer sets that were used for studying the tissue distribution. The highest cTSHR expression during the last week of development was observed at E14, whereas most of the mRNA levels at the other ages (except for E20NP and C1) were significantly lower in comparison with E14 (Fig. 9Go).


Figure 9
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  		FIG. 9. TSHR expression measured with real-time PCR and expressed as relative values ± SEM (n = 3 pools of four pairs of thyroid glands). The days of incubation are indicated. At E20, a distinction is made between the nonpipping stage (E20NP) and the internal pipping stage (E20IP). Newly hatched chicks are indicated as C0 and 1-d-old chicks as C1. *, Significantly different results compared with E14 (P < 0.05).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In the present study, we report the cloning of the cTSHR from thyroid gland and pituitary. Although cDNA sequences of several mammalian species, such as human (16, 17, 18, 19), rat (20), dog (21, 22), and some fish species (23, 24) have already been determined, this is the first sauropsid TSHR to be cloned. The cTSHR gene is located on chromosome 5 and contains 10 exons. The coding sequence of the cTSHR consists of a 2283-bp open reading frame, which encodes a 761-amino-acid seven-transmembrane G protein-coupled glycoprotein. The extracellular domain is encoded by the first nine exons and part of the last exon, whereas the transmembrane and intracellular domains are encoded entirely by the last exon, which is the case for most vertebrate glycoprotein hormone receptors. The overall cDNA homology with TSHR sequences of Rattus norvegicus (NM_012888), Homo sapiens (NM_000369), and Clarias gariepinus (AY129556) is 67, 71, and 59%, respectively (Fig. 2Go), indicating a higher identity to the mammalian sequences. Comparison of their deduced amino acid sequences gives similar results with homologies of 68, 69, and 54%, respectively. Consequently, the cTSHR is grouped in the same cluster as the mammalian TSHRs in the phylogenetic dendrogram (Fig. 3Go). The homology between the cTSHR and the FSH and LH/choriogonadotropin receptor is lower: 56 and 53% (cDNA sequences) and 50% (amino acid sequences). The highest sequence resemblance can be found in the transmembrane and intracellular components. Besides the full-length cTSHR, two additional splice variants were detected in the pituitary as well as the thyroid gland, cTSHRb lacking exon 3 and cTSHRc lacking both exons 2 and 3. Although a variant dog TSHR cDNA with a similar deletion in exon 3 has already been described (M90047) (21, 22), there are, to our knowledge, no reports of splice variants lacking both exon 2 and exon 3. Several smaller transcripts, however, have been observed in human thyroid cells (16). For example, a thyroidal mRNA encoding both the signal peptide and ligand-binding region of the human TSHR (exon 1–8) but not the seven transmembrane helices has been described (19, 25). Also, a truncated bTSHR lacking half of the fifth segment of the transmembrane domain up to the C-terminal domain of the full-length TSHR has been identified (26).

A functional analysis was carried out to determine whether the cloned full-length cTSHR encoded a functional receptor. This was done by challenging cTSHR-expressing HEK-293 cells with bTSH and measuring intracellular cAMP release. Because cTSH is not available as such, and previous studies have shown that bTSH is highly effective in elevating TH levels in birds (27), bTSH was used to stimulate the cTSHR. The functional assays indicated that bTSH was able to increase intracellular cAMP levels significantly in a dose-dependent manner and with a very high efficacy (EC50 = 1.43 nM). Nevertheless, a thorough characterization of the binding characteristics of the cTSHR awaits the availability of the homologous cTSH. Also, the functionality of the two cTSHR splice variants, cTHSRb and cTSHRc, remains to be determined.

The presence of cTSHR mRNA in thyroid tissue was confirmed with in situ hybridization, where a clear signal was observed in the follicular cells. However, the TSHR is not merely important in regulating function, growth, and differentiation of the thyroid gland, as suggested by the detection of TSHR mRNA in several mammalian and teleost tissues such as lymphocytes (28, 29), pituitary (30, 31), kidney (32), and testis (24, 33, 34). Using real-time PCR, we measured cTSHR mRNA expression in 24 different chicken tissues. Besides the thyroid gland, we found clear expression of cTSHR mRNA in the pituitary gland, pineal gland, retina, and in most brain parts. TSHR expression has been demonstrated in mammalian brain in both neuronal cells and astrocytes. In young rats, the TSHR was predominantly found in neuron-rich areas such as the hippocampus and hypothalamic nuclei. As for the expression in astrocytes, TSHR mRNA was mostly detected in the ependymal cell layer and the subependymal zone (35). Also in ovine hypothalamus, cerebellum, and cortex, TSHR mRNA was found (36). Although in situ hybridization experiments are required to locate the specific cell types that express cTSHR mRNA, our real-time PCR results already indicate that its distribution in the chicken brain is widespread. The possible presence of this receptor in the brain may have various physiological roles. Some authors suggest that TSHR expression in the hypothalamus is consistent with a neuroendocrine feedback of TSH upon TRH release or expression (37). As in humans, we also found TSHR mRNA in the pituitary gland. In humans, pituitary TSHR is expressed in a set of nonendocrine cells, the folliculo-stellate cells, which are known to influence neighboring cells in a paracrine way (30). Also in chicken heart, pancreas, and skeletal muscle, cTSHR mRNA could be detected. Although mRNA levels do not always reflect protein expression, our results suggest that TSH, acting through its receptor, regulates diverse processes in the body and that its function is not limited to the stimulation of the thyroid gland.

It has been suggested that an elevated thyroidal and pituitary sensitivity to secretagogues coupled with an increased gland size may be responsible for the rise in plasma T4 concentration toward hatching in birds (12). Our results show no rise in cTSHR mRNA expression during the last week of embryonic development, which suggests that the thyroid cells do not become more sensitive for pituitary TSH during this period. Furthermore, a rise in sensitivity of the pituitary gland for hypothalamic stimuli has been discarded recently (38), and experimental evidence confirms the presence of a functional negative feedback regulation on TSH gene expression and secretion by TH during this critical period (39). Most likely, an increased TSH secretion resulting from a higher release of hypothalamic releasing factors and an increasing number of thyrotropes is the main cause of the change in plasma T4 levels during this crucial last week of embryonic development. A rise in pituitary TSHß mRNA expression has indeed been observed during this period (40, 38), but circulating levels of TSH have not been accurately determined yet, because of the lack of a homologous RIA. Also, the increasing thyroid size (10, 41) and changes in the expression and activity of components mediating TH action such as iodothyronine deiodinases, TH transporters, TH receptors, and cofactors, should be taken into account.

Previous studies have shown that the numerical densities of TSH-binding cells in chicken thyroid are highest between d 11.5 and 12.5 (42). Afterward, these numbers decrease and remain stable from d 14.5–17.5. The peak values in the density of TSH-binding cells correspond to an increased pituitary-thyroid activity at d 11.5–12.5. TSH is already found in chicken pituitary gland by d 6.5 (43, 44) when vascularization of the adenohypophysis occurs, and by d 12.5, the hypothalamo-adenohypophyseal portal vascular system has been fully formed (45). At E14, we found higher cTSHR mRNA expression levels than at any of the other ages examined. The cTSHR mRNA expression was 2-fold lower at C0. The lower expression levels starting from d 15 could reflect a down-regulation of the cTSHR by the increasing plasma T4 levels during that period.

In summary, we have cloned the cTSHR and two splice variants. Whereas the full-length cTSHR is a functional receptor, the activity of the splice variants remains to be determined. The presence of cTSHR mRNA in tissues other than the thyroid gland suggests that TSH exerts other roles than merely the control of thyroid function. The relative expression of thyroidal cTSHR mRNA did not change during the last week of embryonic development, indicating that thyroidal sensitivity to TSH is not an important factor contributing to the increasing T4 concentrations toward hatching.


   Acknowledgments
 
We thank L. Noterdaeme, W. Van Ham, and F. Voets for their technical assistance and V. Beck for her help with the tissue sampling.


   Footnotes
 
B.D.G. and T.J. are financially supported by the Research Foundation–Flanders (FWO–Vlaanderen). This work was partially funded by FWO project 0582.06, grants-in-aid from the Ministry of Education, Science, and Culture, Japan, to S.Take., and Excellent Research Project Grant (ERPG) of Okayama University to S.Taka.

Disclosure summary: S.V.H.G., S.Tan., T.J., L.S., S.Taka., S.Take., V.M.D., and B.D.G. have nothing to declare.

First Published Online May 18, 2006

Abbreviations: AMV, Avian myeloblastosis virus; bTSH, bovine TSH; cTSHR, chicken TSH receptor; DIG, digoxigenin; E14, d 14 of embryonic development; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; TH, thyroid hormones; TSHR, TSH receptor.

Received September 23, 2005.

Accepted for publication May 9, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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