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Molecular characterization of chicken growth hormone secretagogue receptor gene
GENERAL AND COMPARATIVE
ENDOCRINOLOGY General and Comparative Endocrinology 134 (2003) 198–202 www.elsevier.com/locate/ygcen
Short Communication
Molecular characterization of chicken growth hormone secretagogue receptor gene Minoru Tanaka,a,* Takashi Miyazaki,b Ichiro Yamamoto,c Naoya Nakai,d Yoshiyuki Ohta,a Nobumichi Tsushima,a Masaaki Wakita,b and Kiyoshi Shimadac a
Department of Animal Science, Faculty of Applied Life Science, Nippon Veterinary and Animal Science University, Musashino, Tokyo 180-8602, Japan b Department of Animal Science, Faculty of Bioresources, Mie University, Tsu, Mie 514-8507, Japan c Laboratory of Animal Physiology, Bioagricultural Sciences Graduate School of Nagoya University, Chikusa, Nagoya, Japan d Department of Biochemistry, Faculty of Medicine, Mie University, Tsu, Mie 514-8507, Japan Accepted 3 July 2003
Abstract Synthetic growth hormone secretagogues stimulate growth hormone secretion by binding to a specific receptor, growth hormone secretagogue receptor (GHS-R). In this study, we investigated the cDNA and the genomic structure of chicken GHS-R. Chicken GHS-R gene is composed of two exons separated by an intron. Two GHS-R mRNA species, cGHS-R1a and cGHS-R1a-variant (cGHS-R1aV) are generated by alternative splicing of a primary transcript. cGHS-R1a protein is predicted to have seven transmembrane domains by a high degree of amino acid sequence identity with mammalian and teleost homologs. cGHS-R1aV lacks the transmembrane-6 domain due to a 48 bp deletion. RT-PCR analysis showed widespread tissue distributions of cGHS-R1a and cGHS-R1aV mRNAs with much higher amounts of cGHS-R1a in all the tissues. Ó 2003 Elsevier Inc. All rights reserved. Keywords: Growth hormone secretagogue receptor; Ghrelin receptor; Chicken; Gene; cDNA
1. Introduction Growth hormone (GH) plays an essential role in post-natal growth of vertebrate. The secretion of GH from the anterior pituitary gland is known to be positively and negatively regulated by two hypothalamic hormones, GH-releasing hormone (GHRH) and somatostatin, respectively (Bertherat et al., 1995; Frohman and Jansson, 1986). In addition, synthetic molecules termed GH-secretagogues (GHS) have been shown to stimulate the pulsatile GH release by acting on a receptor distinct from the GHRH receptor (Howard et al., 1996; Patchett et al., 1995; Smith et al., 1997, 2001). Recently, ghrelin, a peptide of 28 amino acid, has been discovered from mammalian stomach as an endogenous ligand for the GHS receptor (GHS-R) (Kojima et al., 1999, 2001), and more recently, ghrelin * Corresponding author. Fax: +81-422-39-0722. E-mail address: [email protected] (M. Tanaka).
0016-6480/03/$ - see front matter Ó 2003 Elsevier Inc. All rights reserved. doi:10.1016/S0016-6480(03)00247-8
has been identified in nonmammalian vertebrates such as bullfrog, chicken, and goldfish (Kaiya et al., 2001, 2002; Unniappan et al., 2002). In addition to the GHrelease, ghrelin has been shown to be involved in other functions such as gastric acid secretion, adiposity, and food intake (Kamegai et al., 2000; Masuda et al., 2000; Nakazato et al., 2001; Shintani et al., 2001; Tschop et al., 2000; Wren et al., 2000). GHS-R belongs to the family of G-protein-coupled receptors containing seven transmebrane (TM) domains. Mammalian GHS-R gene is composed of two exons, and two types of GHS-R mRNAs, GHS-R1a and 1b, are generated by alternative transcription process of the gene (Howard et al., 1996). GHS-R1a is produced by the splicing of exon 1 and exon 2 in the primary transcript of the gene. On the other hand, GHS-R1b mRNA is generated by the termination of transcription at a position in intron 1, and therefore, GHS-R 1b protein is not functional as GHS-R due to lack of TM 6 and TM 7 domains (Howard et al., 1996). Recently, three GHS-R
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cDNA clones referred to 78B7, 75E7, and 1H9 have been cloned in a teleost species, the Pufferfish, (Palyha et al., 2000). In avian species, a peptidyl and a nonpeptidyl GHSs stimulated GH secretion (Bowers et al., 1984; Geris et al., 1998) and GHS-R-related genomic fragments have been detected in the chicken by Southern blot analysis with human and Pufferfish cDNA probes (Palyha et al., 2000). However, the molecular structure of avian GHS-R gene is not yet known. In the present study, we report the structure of chicken GHS-R gene and the generation of two GHS-R transcripts from the gene.
digested with NcoI restriction enzyme of which recognition site was found in the 30 -region of GHS-R gene. After self-ligation, the circularized genomic fragments were subjected to PCR with primers 3 (50 -AGCTTG ACCTTCCTCTTGGTGATG-30 ) and 4 (50 -ACAGATC CATGCCTGGTCACTGAGCA-30 ) derived from the sequence of the 30 -region of GHS-R gene. Finally, the 50 region of GHS-R cDNA was cloned by RT-PCR with primers 5 (50 -AAGAACGCACGCCATCGCCCTTT30 ) and 6.
2. Materials and methods
Five micrograms each of the total RNA from various tissues of chicken was transcribed with oligo-dT primer by SuperScript II RNase H reverse transcriptase (Life Technologies, Tokyo, Japan) according to the manufacturerÕs instruction. The resulting cDNA samples were subjected to PCR of 25 cycles with primers 6 and 7 (50 CTGCTCACCATCATGGTGTGGATC-30 ) to detect cGHS-R1a and cGHS-R1aV mRNAs simultaneously. PCR amplification was performed in a total volume of 25 ll with 1.25 U of ExTaq polymerase. Each PCR cycles consisted of 1 min of denaturing at 95 °C, 1 min of annealing at 60 °C, and 1 min of extension at 72 °C. The PCR products were separated by electrophoresis on a 2.0% agarose gel and transferred to a nylon membrane. The membrane was hybridized with radiolabeled oligodeoxynucleotide probe (positions 676–699 at cGHS-R genomic sequence in Fig. 1).
2.1. Cloning of chicken GHS-R cDNA Total RNA was isolated from the pituitary of chicken (male white leghorn at 8 weeks of age) by acid guanidium/phenol/chloroform method and poly(A)þ RNA was prepared with Oligotex-dT30 (Takara, Tokyo, Japan). A pituitary cDNA library was prepared from the poly(A)þ RNA by Marathon cDNA amplification kit (Clontech, Tokyo, Japan). First, a cDNA fragment (635 bp) for chicken GHS-R was amplified from the cDNA library by 30 cycles of polymerase chain reaction (PCR) using sense (50 -TGGCAGTACCGGCCCTGGA ACTT-30 : positions 310–332) and antisense (50 -CTGAG GTAGAAGAGGACAAAGGA-30 : positions 922–944) primers designed from human GHS-R cDNA (Howard et al., 1996). PCR was performed in 25 ll reaction mixtures containing 1.25 U of ExTaq polymerase (Takara, Osaka, Japan), 5 ll of 5 buffer supplied by the manufacturer and 200 lM each of dNTP. Each PCR cycle consisted of 1 min of denaturing at 95 °C, 2 min of annealing at 60 °C, and 3 min of extension at 72 °C. The amplified fragments were cloned into pGEM-T Easy plasmid vector (Promega, Tokyo, Japan), and sequenced. The 50 - and 30 -regions of GHS-R cDNA were cloned by the method of rapid amplification of cDNA ends (RACE) (Frohman, 1989) with primer 1 (50 -TTGG CAGTACCGGCCCTGGAACTT-30 ) and primer 6 (50 GGCTGATCACTGCTATCTCCAAGG-30 ), respectively, derived from the sequence of the chicken GHS-R cDNA fragment. 2.2. Cloning of chicken GHS-R gene Genomic DNA was prepared from chicken kidney by the SDS–Proteinase K method. The 30 -region of GHS-R gene was cloned by PCR with primers 1 and 2 (50 -ACGTGACATCTCCCAGCAAATCC-30 ) designed from the sequence of cGHS-R1a cDNA. The 50 -region of GHS-R gene was cloned by the Inverse PCR method (Ochman et al., 1990). Briefly, the genomic DNA was
2.3. RT-PCR analysis of GHS-R mRNA expression in chicken tissues
3. Results and discussion 3.1. Structure of chicken GHS-R cDNA and gene A GHS-R cDNA (cGHS-R1a) and its variant form cDNA (cGHS-R1aV) containing a 48 bp deletion were cloned from the chicken pituitary cDNA library, and genomic fragments for cGHS-R1a were cloned by PCR technology. As shown in Fig. 1, the genomic sequence contained a noncontiguous sequence for cGHS-R1a, indicating that chicken GHS-R gene is composed of two exons separated by an intron. The position of the intron is comparable to that of the intron in mammalian GHSR gene (McKee et al., 1997; Petersenn et al., 2001). The 48 bp deletion in cGHS-R1aV is observed at the 50 -end of exon 2, and followed by ag which is the common sequence at the splicing acceptor site. These observations indicate that cGHS-R1aV mRNA is produced by the splicing of exon 1 to the position of 48 bp downstream from the 50 -end of exon 2. Although the transcription start site is not yet defined, no canonical TATA box is observed in the 525 bp upstream region of the translation start site, suggesting that chicken GHS-R
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Fig. 1. Genomic sequence of chicken GHS-R. Exons are shown by capitals. Intron and 50 - and 30 -untranslated regions are indicated by small letters. The deleted region in cGHS-R1aV is boxed. Double underlines indicate the splicing acceptor sites for cGHS-R1a and cGHS-R1aV transcripts. An arrow head indicates the cleavage site by NcoI used in Inverse PCR. Right- and left-oriented arrows indicate the sense and antisense primers, respectively, used for the PCR. A potential polyadenylation signal is underlined. The nucleotide sequences will appear in the DDBJ nucleotide detabase with the Accession Nos. AB095994 for cGHS-R gene, AB095995 for cGHS-R1a, and AB095996 for cGHS-R1aV.
gene is a TATA-less gene, the same as human GHS-R gene (Petersenn et al., 2001). A potential polyadenylation signal is found at 30 bp upstream from the 30 -end of the genomic sequence. cGHS-R1a shows a high degree of amino acid sequence identity with mammalian and teleost homologs
(68% with human GHS-R1a and 56% with Pufferfish 78B7). As shown in Fig. 2, the seven TM domains in human GHS-R1a are highly conserved in chicken GHSR1a as well as in the Pufferfish GHS-R, suggesting that cGHS-R1a also consists of seven TM domains. In cGHS-R1aV, TM-6 domain is lost as a result of the
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Fig. 2. Multiple aligment of the amino acid sequences of chicken (cGHS-R1a), human (hGHS-R 1a and hGHS-R1b), and Pufferfish (78B7) GHS-Rs. Residues identical to those of cGHS-R1a are shown by asterisks. The predicted seven TM regions are boxed, and the deleted sequence in cGHSR1aV is underlined.
48 bp deletion, and therefore, the C-terminal region is predicted to be located in the extracellular side, while the corresponding region of cGHS-R1a is located in the intracellular side. Although the ligand-binding ability of cGHS-R1aV has not yet been investigated, if cGHSR1aV binds ghrelin, it could modulate ghrelin clearance. 3.2. Expression of GHS-R mRNAs in chicken tissues In mammals, the expression of GHS-R mRNA was detected in the pituitary and various brain areas including hypothalamas and hippocampus by in situ hybridization analysis and RNase protection assay (Guan et al., 1997; Howard et al., 1996). Recently, human GHS-R1a and 1b mRNAs have been detected in a wide range of tissues by RT-PCR analysis (Gnanapavan et al., 2002). As shown in Fig. 3, cGHS-R1a mRNA were detected in the pituitary, brain, thymus, spleen, pancreas, liver, lung, heart, intestine, kidney, stomach, gizzard, and muscle by RT-PCR. The expression levels were the highest in the pituitary and brain, and moderately high in the liver, intestine, and spleen. The preferential expression of cGHS-R1a mRNA in the pituitary and brain is consistent with recent findings that chicken ghrelin is involved in the regulation of GH secretion and the feeding behavior of the chick (Kaiya et al., 2002; Saito et al., 2002). It has been shown that
Fig. 3. Tissue distribution of cGHS-R1a and cGHS-R1aV mRNAs. Positions of PCR primers and the hybridization probe are shown by arrowheads and a square in the diagrams under the autoradiogram. A black box in the diagram indicates the deleted region in cGHS-R1aV.
ghrelin is expressed in various peripheral tissues such as the gastro-intestinal tract, spleen, and lung where GHSR1a is also expressed, suggesting autocrine or paracrine effects as well as endocrine effects of ghrelin in these tissues. By the RT-PCR analysis, cGHS-R1aV mRNA was detected in the pituitary, brain, thymus, spleen, liver, lung, intestine, kidney, and stomach. The amount detected was much smaller than that of cGHS-R1a mRNA (Fig. 3). The physiological function of cGHSR1aV remains to be elucidated.
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References Bertherat, J., Bluet-Pajot, M.T., Epelbaum, J., 1995. Neuroendocrine regulation of growth hormone. Eur. J. Endocrinol. 132, 12–24. Bowers, C.Y., Momany, F.A., Reynolds, G.A., Hong, A., 1984. On the in vitro and in vivo activity of a new synthetic hexapeptide that acts on the pituitary to specifically release growth hormone. Endocrinology 114, 1537–1545. Frohman, M.A., 1989. RACE: Rapid amplification of cDNA ends. In: Innes, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocols. Academic Press, San Diego, CA, pp. 28–38. Frohman, L.A., Jansson, J.Q., 1986. Growth hormone-releasing hormone. Endocr. Rev. 7, 223–253. Geris, K.L., Hickey, G.J., Berghman, L.R., Visser, T.J., Kuhn, E.R., Darras, V.M., 1998. Pituitary and extrapituitary action sites of the novel nonpeptidyl growth hormone (GH) secretagogue L-692,429 in the chicken. Gen. Comp. Endocrinol. 111, 186–196. Gnanapavan, S., Kola, B., Bustin, S.A., Morris, D.G., McGee, P., Fairclough, P., Bhattacharya, S., Carpenter, R., Grossman, A.B., Korbonits, M., 2002. The tissue distribution of the mRNA of ghrelin and subtypes of its receptor, GHS-R, in humans. J. Clin. Endocrinol. Metab. 87, 2988–2991. Guan, X.-M., Yu, H., Palyha, O.C., McKee, K.K., Feighner, S.D., Sirinathsinghji, D.J., Smith, R.G., Van der Ploeg, L.H., Howard, A.D., 1997. Distribution of mRNA encoding the growth hormone secretagogue receptor in brain and peripheral tissues. Brain Res. Mol. Brain Res. 48, 23–29. Howard, A.D., Feighner, S.D., Cully, D.F., Arena, J.P., Liberator, P.A., Rosenblum, C.L., Hamelin, M., Hreniuk, D.L., Palyha, O.C., Anderson, J., Paress, P.S., Diaz, C., Chou, M., Liu, K.K., McKee, K.K., Pong, S.S., Chaung, L.Y., Elbrecht, A., Dashkevicz, M., Heavens, R., Rigby, M., Sirinathsinghji, D.J., Dean, D.C., Melillo, D.G., Patchett, A.A., Nargund, R., Griffin, P.R., DeMartino, J.A., Gupta, S.K., Schaeffer, J.M., Smith, R.G., Van der Ploeg, L.H.T., 1996. A receptor in pituitary and hypothalamus that functions in growth hormone release. Science 273, 974–977. Kaiya, H., Kojima, M., Hosoda, H., Koda, A., Yamamoto, K., Kitajima, Y., Matsumoto, M., Minamitake, Y., Kikuyama, S., Kangawa, K., 2001. Bullfrog ghrelin is modified by n-octanoic acid at its third threonine residue. J. Biol. Chem. 276, 40441–40448. Kaiya, H., Van Der Geyten, S., Kojima, M., Hosoda, H., Kitajima, Y., Matsumoto, M., Geelissen, S., Darras, V.M., Kangawa, K., 2002. Chicken ghrelin. Purification, cDNA cloning, and biological activity. Endocrinology 143, 3454–3463. Kamegai, J., Tamura, H., Shimizu, T., Ishii, S., Sugihara, H., Wakabayashi, I., 2000. Central effect of ghrelin, an endogenous growth hormone secretagogue, on hypothalamic peptide gene expression. Endocrinology 141, 4797–4800. Kojima, M., Hosoda, H., Date, Y., Nakazato, M., Matsuo, H., Kangawa, K., 1999. Ghrelin is a growth-hormone-releasing acylated peptide from stomach. Nature 402, 656–660. Kojima, M., Hosoda, H., Matsuo, H., Kangawa, K., 2001. Ghrelin: discovery of the natural endogenous ligand for the growth hormone secretagogue receptor. Trends Endocrinol. Metab. 12, 118–122. Masuda, Y., Tanaka, T., Inomata, N., Ohnuma, N., Tanaka, S., Itoh, Z., Hosoda, H., Kojima, M., Kangawa, K., 2000. Ghrelin stimulates gastric acid secretion and motility in rats. Biochem. Biophys. Res. Commun. 276, 905–908.
McKee, K.K., Palyha, O.C., Feighner, S.D., Hreniuk, D.L., Tan, C.P., Phillips, M.S., Smith, R.G., Van der Ploeg, L.H., Howard, A.D., 1997. Molecular analysis of rat pituitary and hypothalamic growth hormone secretagogue receptors. Mol. Endocrinol. 11, 415– 423. Nakazato, M., Murakami, N., Date, Y., Kojima, M., Matsuo, H., Kangawa, K., Matsukura, S., 2001. A role for ghrelin in the central regulation of feeding. Nature 409, 194–198. Ochman, H., Mendhora, M.M., Garza, D., Hartl, D.L., 1990. Amplification of flanking sequences by inverse PCR. In: Innes, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocols. Academic Press, San Diego, CA, pp. 219–227. Palyha, O.C., Feighner, S.D., Tan, C.P., McKee, K.K., Hreniuk, D.L., Gao, Y.D., Schleim, K.D., Yang, L., Morriello, G.J., Nargund, R., Patchett, A.A., Howard, A.D., Smith, R.G., 2000. Ligand activation domain of human orphan growth hormone (GH) secretagogue receptor (GHS-R) conserved from Pufferfish to humans. Mol. Endocrinol. 14, 160–169. Patchett, A.A., Nargund, R.P., Tata, J.R., Chen, M.H., Barakat, K.J, Johnston, D.B., Cheng, K., Chan, W.W., Butler, B., Hickey, G., Jacks, T., Schleim, K., Pong, S.S., Chaugen, L.Y.P., Chen, H.Y., Frazier, E., Leugen, K.H., Chiu, S.H.L., Smith, R.G., 1995. Design and biological activities of L-163,191 (MK-0677): a potent, orally active growth hormone secretagogue. Proc. Natl. Acad. Sci. USA 92, 7001–7005. Petersenn, S., Rasch, A.C., Penshorn, M., Beil, F.U., Schulte, H.M., 2001. Genomic structure and transcriptional regulation of the human growth hormone secretagogue receptor. Endocrinology 142, 2649–2659. Saito, E., Kaiya, H., Takagi, T., Yamasaki, I., Denbow, D.M., Kangawa, K., Furuse, M., 2002. Chicken ghrelin and growth hormone-releasing peptide-2 inhibit food intake of neonatal chicks. Eur. J. Pharmacol., 453–475. Shintani, M., Ogawa, Y., Ebihara, K., Aizawa-Abe, M., Miyanaga, F., Takaya, K., Hayashi, T., Inoue, G., Hosoda, K., Kojima, M., Kangawa, K., Nakao, K., 2001. Ghrelin, an endogenous growth hormone secretagogue, is a novel orexigenic peptide that antagonizes leptin action through the activation of hypothalamic neuropeptide Y/Y1 receptor pathway. Diabetes 50, 227–232. Smith, R.G., Van der Ploeg, L.H., Howard, A.D., Feighner, S.D., Cheng, K., Hickey, G.J., Wyvratt, M.J., Fisher Jr., M.H., Nargund, R.P., Patchett, A., 1997. A peptidomimetic regulation of growth hormone secretion. Endocr. Rev. 18, 621–645. Smith, R.G., Leonard, R., Bailey, A.R., Palyha, O., Feighner, S., Tan, C., Mckee, K.K., Pong, S.S., Griffin, P., Howard, A., 2001. Growth hormone secretagogue receptor family members and ligands. Endocrine 14, 9–14. Tschop, M., Smiley, D.L., Heiman, M.L., 2000. Ghrelin induces adiposity in rodents. Nature 407, 908–913. Unniappan, S., Lin, X., Cervini, L., Rivier, J., Kaiya, H., Kangawa, K., Peter, R.E., 2002. Goldfish ghrelin: molecular characterization of the complementary deoxyribonucleic acid, partial gene structure and evidence for its stimulatory role in food intake. Endocrinology 143, 4143–4146. Wren, A.M., Small, C.J., Ward, H.L., Murphy, K.G., Dakin, C.L., Taheri, S., Kennedy, A.R., Roberts, G.H., Morgan, D.G., Ghatei, M.A., Bloom, S.R., 2000. The novel hypothalamic peptide ghrelin stimulates food intake and growth hormone secretion. Endocrinology 141, 4325–4328.
ENDOCRINOLOGY General and Comparative Endocrinology 134 (2003) 198–202 www.elsevier.com/locate/ygcen
Short Communication
Molecular characterization of chicken growth hormone secretagogue receptor gene Minoru Tanaka,a,* Takashi Miyazaki,b Ichiro Yamamoto,c Naoya Nakai,d Yoshiyuki Ohta,a Nobumichi Tsushima,a Masaaki Wakita,b and Kiyoshi Shimadac a
Department of Animal Science, Faculty of Applied Life Science, Nippon Veterinary and Animal Science University, Musashino, Tokyo 180-8602, Japan b Department of Animal Science, Faculty of Bioresources, Mie University, Tsu, Mie 514-8507, Japan c Laboratory of Animal Physiology, Bioagricultural Sciences Graduate School of Nagoya University, Chikusa, Nagoya, Japan d Department of Biochemistry, Faculty of Medicine, Mie University, Tsu, Mie 514-8507, Japan Accepted 3 July 2003
Abstract Synthetic growth hormone secretagogues stimulate growth hormone secretion by binding to a specific receptor, growth hormone secretagogue receptor (GHS-R). In this study, we investigated the cDNA and the genomic structure of chicken GHS-R. Chicken GHS-R gene is composed of two exons separated by an intron. Two GHS-R mRNA species, cGHS-R1a and cGHS-R1a-variant (cGHS-R1aV) are generated by alternative splicing of a primary transcript. cGHS-R1a protein is predicted to have seven transmembrane domains by a high degree of amino acid sequence identity with mammalian and teleost homologs. cGHS-R1aV lacks the transmembrane-6 domain due to a 48 bp deletion. RT-PCR analysis showed widespread tissue distributions of cGHS-R1a and cGHS-R1aV mRNAs with much higher amounts of cGHS-R1a in all the tissues. Ó 2003 Elsevier Inc. All rights reserved. Keywords: Growth hormone secretagogue receptor; Ghrelin receptor; Chicken; Gene; cDNA
1. Introduction Growth hormone (GH) plays an essential role in post-natal growth of vertebrate. The secretion of GH from the anterior pituitary gland is known to be positively and negatively regulated by two hypothalamic hormones, GH-releasing hormone (GHRH) and somatostatin, respectively (Bertherat et al., 1995; Frohman and Jansson, 1986). In addition, synthetic molecules termed GH-secretagogues (GHS) have been shown to stimulate the pulsatile GH release by acting on a receptor distinct from the GHRH receptor (Howard et al., 1996; Patchett et al., 1995; Smith et al., 1997, 2001). Recently, ghrelin, a peptide of 28 amino acid, has been discovered from mammalian stomach as an endogenous ligand for the GHS receptor (GHS-R) (Kojima et al., 1999, 2001), and more recently, ghrelin * Corresponding author. Fax: +81-422-39-0722. E-mail address: [email protected] (M. Tanaka).
0016-6480/03/$ - see front matter Ó 2003 Elsevier Inc. All rights reserved. doi:10.1016/S0016-6480(03)00247-8
has been identified in nonmammalian vertebrates such as bullfrog, chicken, and goldfish (Kaiya et al., 2001, 2002; Unniappan et al., 2002). In addition to the GHrelease, ghrelin has been shown to be involved in other functions such as gastric acid secretion, adiposity, and food intake (Kamegai et al., 2000; Masuda et al., 2000; Nakazato et al., 2001; Shintani et al., 2001; Tschop et al., 2000; Wren et al., 2000). GHS-R belongs to the family of G-protein-coupled receptors containing seven transmebrane (TM) domains. Mammalian GHS-R gene is composed of two exons, and two types of GHS-R mRNAs, GHS-R1a and 1b, are generated by alternative transcription process of the gene (Howard et al., 1996). GHS-R1a is produced by the splicing of exon 1 and exon 2 in the primary transcript of the gene. On the other hand, GHS-R1b mRNA is generated by the termination of transcription at a position in intron 1, and therefore, GHS-R 1b protein is not functional as GHS-R due to lack of TM 6 and TM 7 domains (Howard et al., 1996). Recently, three GHS-R
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cDNA clones referred to 78B7, 75E7, and 1H9 have been cloned in a teleost species, the Pufferfish, (Palyha et al., 2000). In avian species, a peptidyl and a nonpeptidyl GHSs stimulated GH secretion (Bowers et al., 1984; Geris et al., 1998) and GHS-R-related genomic fragments have been detected in the chicken by Southern blot analysis with human and Pufferfish cDNA probes (Palyha et al., 2000). However, the molecular structure of avian GHS-R gene is not yet known. In the present study, we report the structure of chicken GHS-R gene and the generation of two GHS-R transcripts from the gene.
digested with NcoI restriction enzyme of which recognition site was found in the 30 -region of GHS-R gene. After self-ligation, the circularized genomic fragments were subjected to PCR with primers 3 (50 -AGCTTG ACCTTCCTCTTGGTGATG-30 ) and 4 (50 -ACAGATC CATGCCTGGTCACTGAGCA-30 ) derived from the sequence of the 30 -region of GHS-R gene. Finally, the 50 region of GHS-R cDNA was cloned by RT-PCR with primers 5 (50 -AAGAACGCACGCCATCGCCCTTT30 ) and 6.
2. Materials and methods
Five micrograms each of the total RNA from various tissues of chicken was transcribed with oligo-dT primer by SuperScript II RNase H reverse transcriptase (Life Technologies, Tokyo, Japan) according to the manufacturerÕs instruction. The resulting cDNA samples were subjected to PCR of 25 cycles with primers 6 and 7 (50 CTGCTCACCATCATGGTGTGGATC-30 ) to detect cGHS-R1a and cGHS-R1aV mRNAs simultaneously. PCR amplification was performed in a total volume of 25 ll with 1.25 U of ExTaq polymerase. Each PCR cycles consisted of 1 min of denaturing at 95 °C, 1 min of annealing at 60 °C, and 1 min of extension at 72 °C. The PCR products were separated by electrophoresis on a 2.0% agarose gel and transferred to a nylon membrane. The membrane was hybridized with radiolabeled oligodeoxynucleotide probe (positions 676–699 at cGHS-R genomic sequence in Fig. 1).
2.1. Cloning of chicken GHS-R cDNA Total RNA was isolated from the pituitary of chicken (male white leghorn at 8 weeks of age) by acid guanidium/phenol/chloroform method and poly(A)þ RNA was prepared with Oligotex-dT30 (Takara, Tokyo, Japan). A pituitary cDNA library was prepared from the poly(A)þ RNA by Marathon cDNA amplification kit (Clontech, Tokyo, Japan). First, a cDNA fragment (635 bp) for chicken GHS-R was amplified from the cDNA library by 30 cycles of polymerase chain reaction (PCR) using sense (50 -TGGCAGTACCGGCCCTGGA ACTT-30 : positions 310–332) and antisense (50 -CTGAG GTAGAAGAGGACAAAGGA-30 : positions 922–944) primers designed from human GHS-R cDNA (Howard et al., 1996). PCR was performed in 25 ll reaction mixtures containing 1.25 U of ExTaq polymerase (Takara, Osaka, Japan), 5 ll of 5 buffer supplied by the manufacturer and 200 lM each of dNTP. Each PCR cycle consisted of 1 min of denaturing at 95 °C, 2 min of annealing at 60 °C, and 3 min of extension at 72 °C. The amplified fragments were cloned into pGEM-T Easy plasmid vector (Promega, Tokyo, Japan), and sequenced. The 50 - and 30 -regions of GHS-R cDNA were cloned by the method of rapid amplification of cDNA ends (RACE) (Frohman, 1989) with primer 1 (50 -TTGG CAGTACCGGCCCTGGAACTT-30 ) and primer 6 (50 GGCTGATCACTGCTATCTCCAAGG-30 ), respectively, derived from the sequence of the chicken GHS-R cDNA fragment. 2.2. Cloning of chicken GHS-R gene Genomic DNA was prepared from chicken kidney by the SDS–Proteinase K method. The 30 -region of GHS-R gene was cloned by PCR with primers 1 and 2 (50 -ACGTGACATCTCCCAGCAAATCC-30 ) designed from the sequence of cGHS-R1a cDNA. The 50 -region of GHS-R gene was cloned by the Inverse PCR method (Ochman et al., 1990). Briefly, the genomic DNA was
2.3. RT-PCR analysis of GHS-R mRNA expression in chicken tissues
3. Results and discussion 3.1. Structure of chicken GHS-R cDNA and gene A GHS-R cDNA (cGHS-R1a) and its variant form cDNA (cGHS-R1aV) containing a 48 bp deletion were cloned from the chicken pituitary cDNA library, and genomic fragments for cGHS-R1a were cloned by PCR technology. As shown in Fig. 1, the genomic sequence contained a noncontiguous sequence for cGHS-R1a, indicating that chicken GHS-R gene is composed of two exons separated by an intron. The position of the intron is comparable to that of the intron in mammalian GHSR gene (McKee et al., 1997; Petersenn et al., 2001). The 48 bp deletion in cGHS-R1aV is observed at the 50 -end of exon 2, and followed by ag which is the common sequence at the splicing acceptor site. These observations indicate that cGHS-R1aV mRNA is produced by the splicing of exon 1 to the position of 48 bp downstream from the 50 -end of exon 2. Although the transcription start site is not yet defined, no canonical TATA box is observed in the 525 bp upstream region of the translation start site, suggesting that chicken GHS-R
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Fig. 1. Genomic sequence of chicken GHS-R. Exons are shown by capitals. Intron and 50 - and 30 -untranslated regions are indicated by small letters. The deleted region in cGHS-R1aV is boxed. Double underlines indicate the splicing acceptor sites for cGHS-R1a and cGHS-R1aV transcripts. An arrow head indicates the cleavage site by NcoI used in Inverse PCR. Right- and left-oriented arrows indicate the sense and antisense primers, respectively, used for the PCR. A potential polyadenylation signal is underlined. The nucleotide sequences will appear in the DDBJ nucleotide detabase with the Accession Nos. AB095994 for cGHS-R gene, AB095995 for cGHS-R1a, and AB095996 for cGHS-R1aV.
gene is a TATA-less gene, the same as human GHS-R gene (Petersenn et al., 2001). A potential polyadenylation signal is found at 30 bp upstream from the 30 -end of the genomic sequence. cGHS-R1a shows a high degree of amino acid sequence identity with mammalian and teleost homologs
(68% with human GHS-R1a and 56% with Pufferfish 78B7). As shown in Fig. 2, the seven TM domains in human GHS-R1a are highly conserved in chicken GHSR1a as well as in the Pufferfish GHS-R, suggesting that cGHS-R1a also consists of seven TM domains. In cGHS-R1aV, TM-6 domain is lost as a result of the
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Fig. 2. Multiple aligment of the amino acid sequences of chicken (cGHS-R1a), human (hGHS-R 1a and hGHS-R1b), and Pufferfish (78B7) GHS-Rs. Residues identical to those of cGHS-R1a are shown by asterisks. The predicted seven TM regions are boxed, and the deleted sequence in cGHSR1aV is underlined.
48 bp deletion, and therefore, the C-terminal region is predicted to be located in the extracellular side, while the corresponding region of cGHS-R1a is located in the intracellular side. Although the ligand-binding ability of cGHS-R1aV has not yet been investigated, if cGHSR1aV binds ghrelin, it could modulate ghrelin clearance. 3.2. Expression of GHS-R mRNAs in chicken tissues In mammals, the expression of GHS-R mRNA was detected in the pituitary and various brain areas including hypothalamas and hippocampus by in situ hybridization analysis and RNase protection assay (Guan et al., 1997; Howard et al., 1996). Recently, human GHS-R1a and 1b mRNAs have been detected in a wide range of tissues by RT-PCR analysis (Gnanapavan et al., 2002). As shown in Fig. 3, cGHS-R1a mRNA were detected in the pituitary, brain, thymus, spleen, pancreas, liver, lung, heart, intestine, kidney, stomach, gizzard, and muscle by RT-PCR. The expression levels were the highest in the pituitary and brain, and moderately high in the liver, intestine, and spleen. The preferential expression of cGHS-R1a mRNA in the pituitary and brain is consistent with recent findings that chicken ghrelin is involved in the regulation of GH secretion and the feeding behavior of the chick (Kaiya et al., 2002; Saito et al., 2002). It has been shown that
Fig. 3. Tissue distribution of cGHS-R1a and cGHS-R1aV mRNAs. Positions of PCR primers and the hybridization probe are shown by arrowheads and a square in the diagrams under the autoradiogram. A black box in the diagram indicates the deleted region in cGHS-R1aV.
ghrelin is expressed in various peripheral tissues such as the gastro-intestinal tract, spleen, and lung where GHSR1a is also expressed, suggesting autocrine or paracrine effects as well as endocrine effects of ghrelin in these tissues. By the RT-PCR analysis, cGHS-R1aV mRNA was detected in the pituitary, brain, thymus, spleen, liver, lung, intestine, kidney, and stomach. The amount detected was much smaller than that of cGHS-R1a mRNA (Fig. 3). The physiological function of cGHSR1aV remains to be elucidated.
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References Bertherat, J., Bluet-Pajot, M.T., Epelbaum, J., 1995. Neuroendocrine regulation of growth hormone. Eur. J. Endocrinol. 132, 12–24. Bowers, C.Y., Momany, F.A., Reynolds, G.A., Hong, A., 1984. On the in vitro and in vivo activity of a new synthetic hexapeptide that acts on the pituitary to specifically release growth hormone. Endocrinology 114, 1537–1545. Frohman, M.A., 1989. RACE: Rapid amplification of cDNA ends. In: Innes, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocols. Academic Press, San Diego, CA, pp. 28–38. Frohman, L.A., Jansson, J.Q., 1986. Growth hormone-releasing hormone. Endocr. Rev. 7, 223–253. Geris, K.L., Hickey, G.J., Berghman, L.R., Visser, T.J., Kuhn, E.R., Darras, V.M., 1998. Pituitary and extrapituitary action sites of the novel nonpeptidyl growth hormone (GH) secretagogue L-692,429 in the chicken. Gen. Comp. Endocrinol. 111, 186–196. Gnanapavan, S., Kola, B., Bustin, S.A., Morris, D.G., McGee, P., Fairclough, P., Bhattacharya, S., Carpenter, R., Grossman, A.B., Korbonits, M., 2002. The tissue distribution of the mRNA of ghrelin and subtypes of its receptor, GHS-R, in humans. J. Clin. Endocrinol. Metab. 87, 2988–2991. Guan, X.-M., Yu, H., Palyha, O.C., McKee, K.K., Feighner, S.D., Sirinathsinghji, D.J., Smith, R.G., Van der Ploeg, L.H., Howard, A.D., 1997. Distribution of mRNA encoding the growth hormone secretagogue receptor in brain and peripheral tissues. Brain Res. Mol. Brain Res. 48, 23–29. Howard, A.D., Feighner, S.D., Cully, D.F., Arena, J.P., Liberator, P.A., Rosenblum, C.L., Hamelin, M., Hreniuk, D.L., Palyha, O.C., Anderson, J., Paress, P.S., Diaz, C., Chou, M., Liu, K.K., McKee, K.K., Pong, S.S., Chaung, L.Y., Elbrecht, A., Dashkevicz, M., Heavens, R., Rigby, M., Sirinathsinghji, D.J., Dean, D.C., Melillo, D.G., Patchett, A.A., Nargund, R., Griffin, P.R., DeMartino, J.A., Gupta, S.K., Schaeffer, J.M., Smith, R.G., Van der Ploeg, L.H.T., 1996. A receptor in pituitary and hypothalamus that functions in growth hormone release. Science 273, 974–977. Kaiya, H., Kojima, M., Hosoda, H., Koda, A., Yamamoto, K., Kitajima, Y., Matsumoto, M., Minamitake, Y., Kikuyama, S., Kangawa, K., 2001. Bullfrog ghrelin is modified by n-octanoic acid at its third threonine residue. J. Biol. Chem. 276, 40441–40448. Kaiya, H., Van Der Geyten, S., Kojima, M., Hosoda, H., Kitajima, Y., Matsumoto, M., Geelissen, S., Darras, V.M., Kangawa, K., 2002. Chicken ghrelin. Purification, cDNA cloning, and biological activity. Endocrinology 143, 3454–3463. Kamegai, J., Tamura, H., Shimizu, T., Ishii, S., Sugihara, H., Wakabayashi, I., 2000. Central effect of ghrelin, an endogenous growth hormone secretagogue, on hypothalamic peptide gene expression. Endocrinology 141, 4797–4800. Kojima, M., Hosoda, H., Date, Y., Nakazato, M., Matsuo, H., Kangawa, K., 1999. Ghrelin is a growth-hormone-releasing acylated peptide from stomach. Nature 402, 656–660. Kojima, M., Hosoda, H., Matsuo, H., Kangawa, K., 2001. Ghrelin: discovery of the natural endogenous ligand for the growth hormone secretagogue receptor. Trends Endocrinol. Metab. 12, 118–122. Masuda, Y., Tanaka, T., Inomata, N., Ohnuma, N., Tanaka, S., Itoh, Z., Hosoda, H., Kojima, M., Kangawa, K., 2000. Ghrelin stimulates gastric acid secretion and motility in rats. Biochem. Biophys. Res. Commun. 276, 905–908.
McKee, K.K., Palyha, O.C., Feighner, S.D., Hreniuk, D.L., Tan, C.P., Phillips, M.S., Smith, R.G., Van der Ploeg, L.H., Howard, A.D., 1997. Molecular analysis of rat pituitary and hypothalamic growth hormone secretagogue receptors. Mol. Endocrinol. 11, 415– 423. Nakazato, M., Murakami, N., Date, Y., Kojima, M., Matsuo, H., Kangawa, K., Matsukura, S., 2001. A role for ghrelin in the central regulation of feeding. Nature 409, 194–198. Ochman, H., Mendhora, M.M., Garza, D., Hartl, D.L., 1990. Amplification of flanking sequences by inverse PCR. In: Innes, M.A., Gelfand, D.H., Sninsky, J.J., White, T.J. (Eds.), PCR Protocols. Academic Press, San Diego, CA, pp. 219–227. Palyha, O.C., Feighner, S.D., Tan, C.P., McKee, K.K., Hreniuk, D.L., Gao, Y.D., Schleim, K.D., Yang, L., Morriello, G.J., Nargund, R., Patchett, A.A., Howard, A.D., Smith, R.G., 2000. Ligand activation domain of human orphan growth hormone (GH) secretagogue receptor (GHS-R) conserved from Pufferfish to humans. Mol. Endocrinol. 14, 160–169. Patchett, A.A., Nargund, R.P., Tata, J.R., Chen, M.H., Barakat, K.J, Johnston, D.B., Cheng, K., Chan, W.W., Butler, B., Hickey, G., Jacks, T., Schleim, K., Pong, S.S., Chaugen, L.Y.P., Chen, H.Y., Frazier, E., Leugen, K.H., Chiu, S.H.L., Smith, R.G., 1995. Design and biological activities of L-163,191 (MK-0677): a potent, orally active growth hormone secretagogue. Proc. Natl. Acad. Sci. USA 92, 7001–7005. Petersenn, S., Rasch, A.C., Penshorn, M., Beil, F.U., Schulte, H.M., 2001. Genomic structure and transcriptional regulation of the human growth hormone secretagogue receptor. Endocrinology 142, 2649–2659. Saito, E., Kaiya, H., Takagi, T., Yamasaki, I., Denbow, D.M., Kangawa, K., Furuse, M., 2002. Chicken ghrelin and growth hormone-releasing peptide-2 inhibit food intake of neonatal chicks. Eur. J. Pharmacol., 453–475. Shintani, M., Ogawa, Y., Ebihara, K., Aizawa-Abe, M., Miyanaga, F., Takaya, K., Hayashi, T., Inoue, G., Hosoda, K., Kojima, M., Kangawa, K., Nakao, K., 2001. Ghrelin, an endogenous growth hormone secretagogue, is a novel orexigenic peptide that antagonizes leptin action through the activation of hypothalamic neuropeptide Y/Y1 receptor pathway. Diabetes 50, 227–232. Smith, R.G., Van der Ploeg, L.H., Howard, A.D., Feighner, S.D., Cheng, K., Hickey, G.J., Wyvratt, M.J., Fisher Jr., M.H., Nargund, R.P., Patchett, A., 1997. A peptidomimetic regulation of growth hormone secretion. Endocr. Rev. 18, 621–645. Smith, R.G., Leonard, R., Bailey, A.R., Palyha, O., Feighner, S., Tan, C., Mckee, K.K., Pong, S.S., Griffin, P., Howard, A., 2001. Growth hormone secretagogue receptor family members and ligands. Endocrine 14, 9–14. Tschop, M., Smiley, D.L., Heiman, M.L., 2000. Ghrelin induces adiposity in rodents. Nature 407, 908–913. Unniappan, S., Lin, X., Cervini, L., Rivier, J., Kaiya, H., Kangawa, K., Peter, R.E., 2002. Goldfish ghrelin: molecular characterization of the complementary deoxyribonucleic acid, partial gene structure and evidence for its stimulatory role in food intake. Endocrinology 143, 4143–4146. Wren, A.M., Small, C.J., Ward, H.L., Murphy, K.G., Dakin, C.L., Taheri, S., Kennedy, A.R., Roberts, G.H., Morgan, D.G., Ghatei, M.A., Bloom, S.R., 2000. The novel hypothalamic peptide ghrelin stimulates food intake and growth hormone secretion. Endocrinology 141, 4325–4328.