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cDNA cloning of ghrelin and ontogeny of ghrelin mRNA expression in the gastrointestinal tract of African ostrich chicks
Regulatory Peptides 167 (2011) 50–55
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Regulatory Peptides j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / r e g p e p
cDNA cloning of ghrelin and ontogeny of ghrelin mRNA expression in the gastrointestinal tract of African ostrich chicks J.X. Wang a,b, P. Li b, K.M. Peng a,⁎, S.H.Z. Jin b a b
College of Animal Science and Veterinary Medicine, Huazhong Agricultural University, Wuhan 430070, PR China College of Animal Science, Yangtze University, Jingzhou 434103, PR China
a r t i c l e
i n f o
Article history: Received 16 July 2010 Received in revised form 18 November 2010 Accepted 25 November 2010 Available online 3 December 2010 Keywords: African ostrich Ghrelin Gastrointestinal tract Gene expression
a b s t r a c t Ghrelin is the endogenous ligand for the growth hormone secretagogue receptor (GHS-R). The sequence of ghrelin has been determined in many species ranging from fish to mammals. The ostrich is the largest herbivorous bird in the world. Although the distribution, morphological characteristics, and developmental changes of ghrelin-producing cells in the gastrointestinal tract of African ostrich chicks have recently been determined, the sequence and structure of ghrelin and its expression in the gastrointestinal tract of African ostrich chicks have not been studied. In the present study, the sequence and structure of ghrelin and its expression in the gastrointestinal tract of African ostrich chicks were investigated by reverse-transcriptase polymerase chain reaction (RT-PCR). Results of cDNA cloning revealed that African ostrich ghrelin is composed of 28 amino acid residues and the sequence of the 7 amino acids of the N-terminal region of African ostrich ghrelin was identical with that of other birds. Ninety-day-old female African ostriches were used to investigate the expression of ghrelin in the gastrointestinal tract. The results showed that ghrelin mRNA existed in the proventriculus, gizzard, duodenum, ileum, cecum, and rectum; there was no expression in the jejunum and colon. We observed developmental changes in the ghrelin mRNA expression in the stomach and small intestine of African ostriches. The results of the present study showed that ghrelin mRNA existed on day 1 in the proventriculus, but there was no expression in other tissues. On day 45, ghrelin mRNA existed in the proventriculus, gizzard, and ileum; however, there was no expression in the duodenum and jejunum. On day 90 and 334, we detected ghrelin mRNA in the proventriculus, gizzard, duodenum, and ileum, but there was no expression in the jejunum. The results of the present study clearly demonstrate that ghrelin mRNA exists and the distribution of ghrelin mRNA in the gastrointestinal tract of African ostriches changes with age (from postnatal day 1 to day 334). © 2010 Published by Elsevier B.V.
1. Introduction Ghrelin is a brain-gut peptide that has been isolated as an endogenous ligand for the growth hormone secretagogue receptor (GHS-R) from the rat stomach. This peptide consists of 28 amino acids, of which the third serine residue (Ser3) is n-octanoylated. This side chain is essential for its biological activity [1]. Ghrelin is mainly produced in the stomach in all animals investigated so far [2]. Expression of ghrelin has also been found in many other organs, including the gastrointestinal tract, lung, thyroid, heart, breast, fat, placenta, lymph nodes, liver, kidney, adrenal gland, pancreas, testis, and central nervous system in mammals [3]. Sakata et al. reported the level of weak ghrelin mRNA expression was low in the postnatal period but then increased in a dimorphic pattern, i.e., transient ⁎ Corresponding author. College of Animal Science and Veterinary Medicine, Huazhong Agricultural University, South Lake, Wuhan, PR China. Tel.: +86 27 8728 6970; fax: +86 27 87280408. E-mail address: [email protected] (K.M. Peng). 0167-0115/$ – see front matter © 2010 Published by Elsevier B.V. doi:10.1016/j.regpep.2010.11.009
stagnation at 4 weeks in the male rats and at 5 weeks in the female rats [4]. In mammals, ghrelin stimulates growth hormone (GH) release, regulates food intake, energy balance, body weight, gastric motility, acid secretion, endocrine pancreas functions, and glucose metabolism [1,5]. In addition to mammals, ghrelin has also been identified in nonmammalian species. In the chicken, ghrelin consists of 26 amino acids. The Ser3 is conserved between the chicken and mammalian species, as is its acylation by either n-octanoic or n-decanoic acid [8]. Expression of ghrelin has also been found in the gastrointestinal tract, lung, heart, liver, kidney, pancreas, testis, and brain in chicks [6–9]. Wada et al. reported that ghrelin mRNA expression was only detected in the proventriculus of the gastrointestinal tract in hatching chicken. On the other hand, a high level of ghrelin mRNA expression was detected in the proventriculus and small amounts of reaction products were found in the pylorus and duodenum of adult chickens [10]. Ghrelin also stimulates GH release in chickens [6,11]; however, in contrast to its action in mammals, ghrelin inhibits feeding when injected intracerebroventricularly, particularly in neonatal chicks [12–14].
J.X. Wang et al. / Regulatory Peptides 167 (2011) 50–55
Ghrelins have been identified in other avian species, such as duck, goose, emu, and turkey [15]. Wang et al. [16] reported the distribution, morphological characteristics, and developmental changes of ghrelinimmunopositive (ghrelin-ip) cells in the gastrointestinal tract of African ostrich chicks. Their results indicated that African ostrich ghrelin-ip cells were found to be localized in the mucous membrane of the entire gastrointestinal tract, and the number of ghrelin-ip cells in the gastrointestinal tract of African ostriches increased with age (from postnatal day 1 to day 90) [16]. However, there have been no studies on the sequence and structure of ghrelin and its expression in the gastrointestinal tract of African ostrich chicks. Therefore, in this study, the sequence and structure of ghrelin and its expression in the gastrointestinal tract were studied in detail using reverse-transcriptase polymerase chain reaction (RT-PCR), in order to provide the theoretic basis for further studies on the effects of ghrelin in the development of the African ostrich and associated regulatory mechanisms. 2. Materials and methods 2.1. Animals Female African ostriches (age: neonatal day 1, 45, 90, and 334) were used in the present study. African ostrich chicks (24 females) were obtained from a standard ostrich farm in the Guangdong Province, China, on postnatal day 1 (newly hatched chicks) and were transported within 10 h to a battery house, where feed and water were made available ad libitum. The 24 birds were divided into 4 groups (6 ostriches per group) on the basis of their body weight (BW) in order to adjust for the BW and the variance among the groups. All birds were maintained in a heated room with slatted plastic flooring and were fed a starter diet for postnatal days 1–334, which was formulated according to the specifications of the Elsenburg Ostrich Feed Database [17]. All procedures were approved by the Animal Care and Welfare Committee of our institute.
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2.4. Primer design According to the published sequences of the broiler chicken ghrelin mRNA (GenBank accession no. AB075215), primer sets were designed using the Primer premier 5.0 software. The primer sequences generating a fragment of 191 bp were as follows: sense primer: 5′ CTCTGGCTGGCTCTAGTTTTTTAAG 3′; antisense primer: 5′ CGCTTCTGTTATCTTGACACCAATTTC3′. 2.5. Polymerase chain reaction (PCR) RT products (3.0 μL) were amplified in a 25-μL PCR reaction containing 3.0 μL cDNA, 1.0 μL each for the primer (10 μM), 1.5 μL each for dNTP (10 mM), 0.25 μL Taq DNA polymerase (5 U/μL) (Takara), 1.00 μL MgCl2 (25 mM), and 2.5 μL of 10×PCR buffer. The thermal cycling parameters were 1 cycle at 94 °C for 5 min, followed by 30 cycles at 94 °C for 1 min, 60 °C for 1 min, 72 °C for 2 min, with a final extension step at 72 °C for 10 min. Under these conditions, reactions were conducted within the linear phase of amplification for each PCR product. The resulting PCR products were analyzed by electrophoresis on a 1% agarose gel, which was subsequently analyzed with a computer flatbed scanner. 2.6. Cloning and sequence analysis of the amplified fragments The PCR products of the proventriculus on day 45 were excised after the correct size had been confirmed by electrophoresis on a 1% agarose gel, purified using the OMEGA DNA Purification Kit (V-gene Biotechnology Ltd., Hangzhou, China) according to the manufacturer's manual, and then cloned into the pMD18-T simple vector. Subsequently, the ligation products were transformed into competent DH5α cells. Successful insertion was then confirmed by PCR using the above primers. Positive clones based on colony PCR selection were picked out for sequencing. Bacteria-liquids containing the fragment of the right size were sequenced by the Jin SHIRUI Biotechnology Co., Ltd. (Nanjing, China).
2.2. Tissue preparation 2.7. Ghrelin tissue distribution and mRNA expression On postnatal days 1, 45, 90, and 334 (control group), the birds were weighed, deeply anesthetized with 10% urethane (Caoyang Secondary Chemical Plant, Shanghai, China) at a dose of 1 g/kg BW, and initially perfused with 1000 mL of 0.85% normal saline (containing 0.075% sodium citrate). Then, the abdomen was cut open. Segments of their gastrointestinal tracts, approximately 1 cm in length, were quickly removed and opened along their longitudinal axes. The specimens were obtained from the following portions of the gastrointestinal tracts: the proventriculus, the gizzard, the duodenum, the jejunum, the ileum, the cecum, the colon, and the rectum. Tissues were removed, immediately snap frozen in liquid nitrogen, and then stored at −80 °C until use. 2.3. RNA extraction, cDNA synthesis, and reverse transcription The weight of the frozen tissues was determined and 1 mL of Trizol (Invitrogen, USA) was added for every 100 mg of tissue. Total RNA was extracted from the gastrointestinal tract tissues of developing African ostrich chicks using the Trizol reagent (Invitrogen) according to the manufacturer's instructions. RNA pellets were suspended in 20 μL of DEPC-treated water, and the RNA concentration was determined by spectrophotometry. Total RNA (2 μg) was reverse transcribed in a final volume of 25 μL containing 1.0 μL AMV reverse transcriptase (200 U) (Takara, Japan), 1.5 μL dNTPs (10 mM) (Takara), 0.5 μL RNasin ribonuclease inhibitor (25 U) (Takara), and 0.5 μg of Oligo dT primer (Takara) in DEPC-treated water and buffer supplied by the manufacturer. The reaction was incubated at 42 °C for 60 min. The cDNA was then heated to 70 °C for 5 min to inactivate the reverse transcriptase. All cDNA samples were stored at − 20 °C until use.
RT-PCR analyses of ghrelin expression in different tissues were performed as follows. Total RNA was isolated using standard Trizol methods (Invitrogen, Inc., Carlsbad, CA). The RT was performed using the same methods with Section 2.3. The PCR were performed using the same methods with Section 2.5. The resulting PCR products were loaded onto a 1% agarose gel and visualized by ethidium bromide staining. Each sample was repeated three times and the average mRNA expression was obtained for further analysis. 2.8. Statistics methods The homology of ghrelin among chicken, turkey, emu, goose, Japanese quail and duck was studied. The sequences of the amplified fragments were compared with other birds using the Nucleotide Blast programs at NCBI (http://www.ncbi.nlm.nih.gov/BLAST/) and identity percentages were obtained from amino acid BLAST with the DNASTAR software (http://www.dnastar.com/) (accession no. for emu is AY338467, turkey AY333783, chicken AB075215, duck AY338466, Japanese quail AB244056 and goose AY338465) The obtained sequences of ghrelin were aligned with the Mega 3.1 software (http://www.megasoftware.net/). 3. Results 3.1. RT-PCR of ghrelin genes Total RNA from the proventriculus of a 45-day old African ostrich was used as an initial sample to amplify ghrelin genes by RT-PCR, which yielded a 191-bp cDNA fragment (Fig. 1).
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J.X. Wang et al. / Regulatory Peptides 167 (2011) 50–55 Table 1 The identity comparison among ghrelins and other birds.
Fig. 1. PCR result of ghrelin gene of African ostrich. M. DL 2000; 1. negative control; 2. PCR product (ghrelin).
3.2. Structure of African ostrich ghrelin genes The amplified ghrelin cDNA fragments were then cloned into the pMD18-T simple vector. After ligation, the products were identified by PCR using the above primers. Those bacteria-liquids containing the fragment of the right size were then sequenced. PCR products of 191 bp length were obtained (Fig. 2), and the sequences of the amplified fragments were compared with the corresponding reported sequences of other birds. The results showed that there were 84, 84, 95, 86, 82, and 84% sequence identity with the chicken, duck, emu, goose, Japanese quail and turkey cDNA, respectively (Table 1). 3.3. Structure of the African ostrich ghrelin protein A search for the translation of the nucleotide sequences into the amino acids was performed using the DNA MAN software. The result indicated that the amino acid sequence of the mature ghrelin protein is a peptide of 28 amino acids, in which the Ser3 could be n-octanoylated although we have not confirmed it in this study (Fig. 2). Comparisons of the mature ghrelin protein from different avian species showed a high degree of amino acid sequence similarity with complete conservation of the first 7 N-terminal amino acids (GSSFLSP) of the mature ghrelin peptide that contains the site (serine 3) of fatty acid acylation (Fig. 3). In addition, the sequence of the amino acid sequence of the mature ghrelin protein was compared with the corresponding reported sequences of the ghrelin protein from 6 species. The results showed that there were 53.5, 51.8, 62.3, 53.5, 54.8, and 53.5% sequence identity with chicken, duck, emu, goose, Japanese quail and turkey ghrelin, respectively (Table 2). To further eliminate the noise caused by saturation of synonymous nucleotide substitution, protein sequences were used to construct the phylogenetic tree using a homology tree of the DNASTAR software. The results showed that the coefficient of relationship between the African ostrich mature ghrelin and emu mature ghrelin was neighbor (Fig. 4). These results indicated that the amplified cDNA fragments were African ostrich ghrelin.
Accession
Description
Length (bp)
Identity (%)
AB075215 AY338466 AY338467 AY338465 AB244056 AY333783
Chicken Duck Emu Goose Japanese quail Turkey
210 208 304 226 196 210
84% 84% 95% 86% 82% 84%
3.4. Distribution of ghrelin mRNA expression in the gastrointestinal tract of the African ostrich On postnatal days 1, 45, 90, and 334, ghrelin mRNA expression was found in different tissues in female African ostriches; the distribution of ghrelin mRNA in the gastrointestinal tract changed with age from postnatal day 1 to day 90. Ninety-day-old female African ostriches were used to investigate the distribution of ghrelin mRNA expression in various tissues by RT-PCR. It was found that ghrelin mRNA existed in the proventriculus, gizzard, duodenum, ileum, cecum, and rectum. There was no expression in the jejunum and colon (Fig. 5). 3.5. Developmental changes in ghrelin mRNA expression in the stomach and small intestine of the African ostrich The result of the RT-PCR revealed that the ghrelin mRNA expression in stomach and small intestine of African ostriches changed with age: on day 1, the ghrelin mRNA was detected in the proventriculus, but there was no expression in other tissues (Fig. 6A). On day 45, ghrelin mRNA was detected in the proventriculus, gizzard, and ileum, but there was no expression in the duodenum and jejunum (Fig. 6B). On day 90 and 334, ghrelin mRNA was found in the proventriculus, gizzard, duodenum, and ileum, but there was no expression in the jejunum (Figs. 5 and 6C). 4. Discussion 4.1. Structure of African ostrich ghrelin genes This is the first report on the gene sequence of African ostrich ghrelin. The cDNA has a length of 191 bp and showed high homologies with those of other species. Kaiya et al. [6] reported that the fulllength chicken ghrelin cDNA was 836-bp long, containing 147 bp in the 5′-untranslated region (5′-UTR), 351 bp of coding region, and 338 bp in the 3′-untranslated region (3′-UTR) using 5′- and 3′-rapid amplification of cDNA ends (RACE) PCR methods. Richards et al. [9] and Yuan et al. [18] also reported that the full-length ghrelin cDNA of turkey, goose, duck, and emu was 869 bp, 886 bp, 745 bp, and 682 bp, respectively. The size of the coding region was also 351 bp. The deduced protein of the turkey, goose, duck, and emu preproghrelin contains 116 amino acids, consisting of 3 parts: the signal peptide,
Fig. 2. The gene and translated product sequence of ghrelin. Graphic representation and nucleotide and amino acid (AA) sequence for African ostrich. The nucleotide sequence of the CDS is underlined. Translated AA sequence of the mature ghrelin hormone is highlighted by the shaded box.
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Fig. 3. Sequence comparison of avian ghrelins. Amino acid comparisons of preproghrelin proteins (116 AA) for different avian species. The locations of mature ghrelin peptide (28 AA) is indicated. Also indicated is the site within the mature ghrelin peptide (serine 3) for acylation by n-octanoic acid binding and signaling. Amino acid sequences shown for chicken, duck, emu goose, turkey and Japanese quail were obtained from GenBank accession nos. AB075215, AY338466, AY338467, AY338465, AY333783 and AB244056, respectively.
Table 2 The sequence of the amino acid sequence of the mature ghrelin protein and a comparison of birds and African ostrich ghrelins. Homology matrix Ostrich Chicken Duck Distance matrix Ostrich Chicken Duck Emu Goose Japanese quail Turkey
100.% 46.5% 48.2% 37.7% 46.5% 45.2% 46.5%
53.5% 100.% 23.1% 21.8% 17.7% 3.1% 6.8%
Emu
Goose
Japanese Turkey quail
51.8% 62.3% 53.5% 54.8% 76.9% 78.2% 82.3% 96.9% 100.% 78.9% 91.2% 75.0% 21.1% 100.% 82.3% 83.3% 8.8% 17.7% 100.% 82.3% 25.0% 16.7% 17.7% 100.% 25.2% .218% 18.4% 4.2%
53.5% 93.2% 74.8% 78.2% 81.6% 95.8% 100.%
mature ghrelin, and C-terminal peptide [9,18]. In this study, the primers were designed according to the chicken cDNA sequence of mature ghrelin (accession no. AB075215), and it could amplify a 191-bp fragment, which only contains the sequence of the mature form of ghrelin. The sequence of African ostrich ghrelin has high homologies with those of other species as shown by the highly conserved parts in the sequence of mature ghrelin. However, the sequence of the full-length ostrich ghrelin gene remains to be further studied. 4.2. Structure of the African ostrich ghrelin protein The present study showed that the mature ghrelin is a peptide consisting of 28 amino acids, in which the Ser3 could be n-octanoylated
although we have not confirmed it in this study. Comparisons of the sequences of mature ghrelin from different avian species showed a high degree of amino acid sequence similarity with complete conservation of the first 7 N-terminal amino acids (GSSFLSP), which contain the site (serine 3) of fatty acid acylation. These results further prove that ghrelin is unambiguously identified as an [n-octanoyl-Ser 3]-peptide and suggest that the ghrelins are highly conserved. The acylated peptide and n-octanoylation at serine 3 is essential for ghrelin activity [1,19]. In birds, ghrelin stimulates GH release in vivo and in vitro [6,11,20], decreases food intake in neonatal chicks [7,14], inhibits water intake [21], controls gonadal functions, such as proliferation, apoptosis, and hormone secretion, in chicken [8], and stimulates the contraction of the crop and proventriculus [22]. However, the physiological functions of the African ostrich ghrelin remains to be further studied. In the chicken, the mature ghrelin peptide consists of 26 amino acids and is processed from the precursor at the dibasic sequence ArgArg (RR) located at the C-terminal end of the peptide [6]. In the turkey this sequence is Pro-Arg (PR), and the change may indicate a potential difference in the size of the mature turkey ghrelin peptide (28 vs. 26 amino acids) as compared to the chicken due to different C-terminal end proteolytic processing [9]. In the African ostrich, this sequence is Cys-Arg(CR), and the change may also indicate a potential difference in the size of the mature ostrich ghrelin peptide (28 vs. 26 amino acids) as compared to the chicken due to the generation of a 28-amino acid peptide by enzymatic cleavage in the C-terminal. All other avian species reported to date, except for turkey, contain the Arg-Arg (RR)
Fig. 4. Cladogram analysis of birds and African ostrich ghrelins. The coefficient of relationship between the African ostrich mature ghrelin and emu mature ghrelin was near.
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the ghrelin gene expression in the gastrointestinal tract of African ostrich chicks. However, the reason remains to be further studied. 4.4. Developmental changes of ghrelin mRNA expression in the stomach and small intestine of the African ostrich Fig. 5. RT-PCR analysis of ghrelin mRNA from gastrointestinal tract tissues of African ostrich on postnatal days 90. Agarose gel (1%) electrophoresis of the RT-PCR products for ghrelin showed 191 bp fragments. M, DNA molecular weight marker; 1. negative control; 2. jejunum; 3. proventriculus; 4. gizzard; 5. duodenum; 6. ileum; 7. cecum; 8. rectum; 9. colon.
sequence at the C-terminal end of the ghrelin peptide which makes the C-terminal sequence of mature ghrelin unique in ostrich, like in the case of turkey. 4.3. Distribution of ghrelin mRNA expression in the gastrointestinal tract of the African ostrich Using RT-PCR, ghrelin mRNA expression has been found in the proventriculus, duodenum, and ileum of chickens, broiler chickens, and layer chicks [6,9,23]. In agreement with these results, ghrelin mRNA expression was also found in the gastrointestinal tract of African ostriches, including the proventriculus, duodenum, and ileum. In the present study, RT-PCR revealed that ghrelin was further expressed in the gizzard, cecum, and rectum of African ostriches. There was no expression in the jejunum and colon. Comparison of the present data with chickens revealed that this expression pattern was not identical because ghrelin was not expressed in the gizzard of chickens. Shao et al. [24] reported that ghrelin is expressed in the proventriculus, proventriculus–gizzard junction, gizzard, and duodenum, but not in the ileum and large intestine in Peking duck [24]. It is possible that the difference of ghrelin expression in the gastrointestinal tract of African ostriches is an individual difference among species. These results indicate that there is a species-specific difference in ghrelin expression in birds. We reported the distribution of ghrelin-ip cells in the gastrointestinal tract of African ostrich chicks and indicated that African ostrich ghrelinip cells were found to be localized in the mucous membrane of the entire gastrointestinal tract [16]. The results of the present study indicated that ghrelin is not expressed in the jejunum and colon of African ostriches. Thus, there are differences between distribution of ghrelin-ip cells and
Fig. 6. RT-PCR analysis of ghrelin mRNA from gastrointestinal tract tissues of African ostrich at three stages of development. Agarose gel (1%) electrophoresis of the RT-PCR products for ghrelin showed 191 bp fragments. Expression of ghrelin mRNA on posthatching-day 1 (A, P1), 45 (B, P45) and 334 (C, P334). M, DNA molecular weight marker; 1. negative control; 2. ileum; 3. duodenum; 4. jejunum; 5. gizzard; 6. proventriculus.
The most interesting result obtained in the present study was with regard to the ontogenic pattern of the ghrelin mRNA expression. Comparison of the present data with the results obtained from birds showed some similarities and differences. For example, in chickens, ghrelin mRNA expression was only detected in the proventriculus of newly hatched chicken, whereas mRNA expression was also detected in the duodenum of adult chickens [10]; Chen et al. [25] reported that the level of chicken ghrelin mRNA in the proventriculus was low from embryonic day 15 (E 15) to E 19, but dramatically increased at posthatching day 2 (P 2), then remained constant until P 30 and followed by a significant decrease at P 44 when there was a diet transition at P 31 and thereafter. The decreased level was reversed at P 58. In Peking duck, ghrelin mRNA expression was detectable at very low levels on E 14 in the proventriculus, and the expression increased by E 21. The expression level remained unchanged on P 1 and P 60 [24]. Our results show that the ghrelin mRNA expression in the gastrointestinal tract of African ostriches changes with age (from postnatal day 1 to day 90). These results were consistent with the number of ghrelin-ip cells that increased with age in the gastrointestinal tract of African ostriches (from postnatal day 1 to day 90) [16]. These results suggest that the part of ghrelin mRNA expression in birds is tissue- and ontogenic stage-specific. Acknowledgements We would like to thank Dr. Liu Huazhen of the Department of Anatomy, Histology and Embryology, College of Animal Science and Veterinary Medicine, Huazhong Agricultural University for her valuable comments on the experiments. This study was supported by the National Natural Science Foundation Project of China, No. 30471249 and No. 39970547. References [1] Kojima M, Hosoda H, Date Y, Nakazato M, Matsuo H, Kangawa K. Ghrelin is a growth hormone-releasing acylated peptide from stomach. Nature 1999;402: 656–60. [2] Ariyasu H, Takaya K, Tagami T, Ogawa Y, Hosoda K, Akamizu T, et al. Stomach is a major source of circulating ghrelin, and feeding state determines plasma ghrelinlike immunoreactivity levels in humans. J Clin Endocrinol Metab 2001;86:4753–8. [3] Gnanapavan S, Kola B, Bustin SA, Morris DG, McGee P, Fairclough P, et al. The tissue distribution of the mRNA of ghrelin and subtypes of its receptor, GHS-R, in humans. J Clin Endocrinol Metab 2002;87:2988–91. [4] Sakata I, Tanaka T, Matsubara M, Yamazaki M, Tani S, Hayashi Y, et al. Postnatal changes in ghrelin mRNA expression and in ghrelin-producing cells in the rat stomach. J Endocrinol 2002;174:463–71. [5] Ueno H, Yamaguchi H, Kangawa K, Nakazato M. Ghrelin: a gastric peptide that regulates food intake and energy homeostasis. Regul Pept 2005;126:11–9. [6] Kaiya H, Van der Geyten S, Kojima M, Hosoda H, Kitajima Y, Matsumoto M, et al. Chicken ghrelin: purification, cDNA cloning, and biological activity. Endocrinology 2002;143:3454–63. [7] Saito ES, Kaiya H, Tachibanat T, Tomonaga S, Denbow DM, Kangawa K, et al. Inhibitory effect of ghrelin on food intake is mediated by the corticotropinreleasing factor system in neonatal chicks. Regul Pept 2005;125:201–8. [8] Sirotkin AV, Grossmann R, Maria-Peon MT, Roa J, Tena-Sempere M, Klein S. Novel expression and functional role of ghrelin in chicken ovary. Mol Cell Endocrinol 2006;257–258:15–25. [9] Richards MP, Poch SM, McMurtry JP. Characterization of turkey and chicken ghrelin genes, and regulation of ghrelin and ghrelin receptor mRNA levels in broiler chickens. Gen Comp Endocrinol 2006;145(3):298–310. [10] Wada R, Sakata I, Kaiya H, Nakamura K, Hayashi Y, Kangawa K, et al. Existence of ghrelin-immunopositive and expressing cells in the proventriculus of the hatching and adult chicken. Regul Pept 2003;111:123–8. [11] Baudet ML, Harvey S. Ghrelin-induced GH secretion in domestic fowl in vivo and in vitro. J Endocrinol 2003;179:97–105. [12] Furuse M, Tachibana T, Ohgushi A, Ando R, Yoshimatsu T, Denbow DM. Intracerebroventricular injection of ghrelin and growth hormone releasing factor inhibits food intake in neonatal chicks. Neurosci Lett 2001;301:123–6.
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Contents lists available at ScienceDirect
Regulatory Peptides j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / r e g p e p
cDNA cloning of ghrelin and ontogeny of ghrelin mRNA expression in the gastrointestinal tract of African ostrich chicks J.X. Wang a,b, P. Li b, K.M. Peng a,⁎, S.H.Z. Jin b a b
College of Animal Science and Veterinary Medicine, Huazhong Agricultural University, Wuhan 430070, PR China College of Animal Science, Yangtze University, Jingzhou 434103, PR China
a r t i c l e
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Article history: Received 16 July 2010 Received in revised form 18 November 2010 Accepted 25 November 2010 Available online 3 December 2010 Keywords: African ostrich Ghrelin Gastrointestinal tract Gene expression
a b s t r a c t Ghrelin is the endogenous ligand for the growth hormone secretagogue receptor (GHS-R). The sequence of ghrelin has been determined in many species ranging from fish to mammals. The ostrich is the largest herbivorous bird in the world. Although the distribution, morphological characteristics, and developmental changes of ghrelin-producing cells in the gastrointestinal tract of African ostrich chicks have recently been determined, the sequence and structure of ghrelin and its expression in the gastrointestinal tract of African ostrich chicks have not been studied. In the present study, the sequence and structure of ghrelin and its expression in the gastrointestinal tract of African ostrich chicks were investigated by reverse-transcriptase polymerase chain reaction (RT-PCR). Results of cDNA cloning revealed that African ostrich ghrelin is composed of 28 amino acid residues and the sequence of the 7 amino acids of the N-terminal region of African ostrich ghrelin was identical with that of other birds. Ninety-day-old female African ostriches were used to investigate the expression of ghrelin in the gastrointestinal tract. The results showed that ghrelin mRNA existed in the proventriculus, gizzard, duodenum, ileum, cecum, and rectum; there was no expression in the jejunum and colon. We observed developmental changes in the ghrelin mRNA expression in the stomach and small intestine of African ostriches. The results of the present study showed that ghrelin mRNA existed on day 1 in the proventriculus, but there was no expression in other tissues. On day 45, ghrelin mRNA existed in the proventriculus, gizzard, and ileum; however, there was no expression in the duodenum and jejunum. On day 90 and 334, we detected ghrelin mRNA in the proventriculus, gizzard, duodenum, and ileum, but there was no expression in the jejunum. The results of the present study clearly demonstrate that ghrelin mRNA exists and the distribution of ghrelin mRNA in the gastrointestinal tract of African ostriches changes with age (from postnatal day 1 to day 334). © 2010 Published by Elsevier B.V.
1. Introduction Ghrelin is a brain-gut peptide that has been isolated as an endogenous ligand for the growth hormone secretagogue receptor (GHS-R) from the rat stomach. This peptide consists of 28 amino acids, of which the third serine residue (Ser3) is n-octanoylated. This side chain is essential for its biological activity [1]. Ghrelin is mainly produced in the stomach in all animals investigated so far [2]. Expression of ghrelin has also been found in many other organs, including the gastrointestinal tract, lung, thyroid, heart, breast, fat, placenta, lymph nodes, liver, kidney, adrenal gland, pancreas, testis, and central nervous system in mammals [3]. Sakata et al. reported the level of weak ghrelin mRNA expression was low in the postnatal period but then increased in a dimorphic pattern, i.e., transient ⁎ Corresponding author. College of Animal Science and Veterinary Medicine, Huazhong Agricultural University, South Lake, Wuhan, PR China. Tel.: +86 27 8728 6970; fax: +86 27 87280408. E-mail address: [email protected] (K.M. Peng). 0167-0115/$ – see front matter © 2010 Published by Elsevier B.V. doi:10.1016/j.regpep.2010.11.009
stagnation at 4 weeks in the male rats and at 5 weeks in the female rats [4]. In mammals, ghrelin stimulates growth hormone (GH) release, regulates food intake, energy balance, body weight, gastric motility, acid secretion, endocrine pancreas functions, and glucose metabolism [1,5]. In addition to mammals, ghrelin has also been identified in nonmammalian species. In the chicken, ghrelin consists of 26 amino acids. The Ser3 is conserved between the chicken and mammalian species, as is its acylation by either n-octanoic or n-decanoic acid [8]. Expression of ghrelin has also been found in the gastrointestinal tract, lung, heart, liver, kidney, pancreas, testis, and brain in chicks [6–9]. Wada et al. reported that ghrelin mRNA expression was only detected in the proventriculus of the gastrointestinal tract in hatching chicken. On the other hand, a high level of ghrelin mRNA expression was detected in the proventriculus and small amounts of reaction products were found in the pylorus and duodenum of adult chickens [10]. Ghrelin also stimulates GH release in chickens [6,11]; however, in contrast to its action in mammals, ghrelin inhibits feeding when injected intracerebroventricularly, particularly in neonatal chicks [12–14].
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Ghrelins have been identified in other avian species, such as duck, goose, emu, and turkey [15]. Wang et al. [16] reported the distribution, morphological characteristics, and developmental changes of ghrelinimmunopositive (ghrelin-ip) cells in the gastrointestinal tract of African ostrich chicks. Their results indicated that African ostrich ghrelin-ip cells were found to be localized in the mucous membrane of the entire gastrointestinal tract, and the number of ghrelin-ip cells in the gastrointestinal tract of African ostriches increased with age (from postnatal day 1 to day 90) [16]. However, there have been no studies on the sequence and structure of ghrelin and its expression in the gastrointestinal tract of African ostrich chicks. Therefore, in this study, the sequence and structure of ghrelin and its expression in the gastrointestinal tract were studied in detail using reverse-transcriptase polymerase chain reaction (RT-PCR), in order to provide the theoretic basis for further studies on the effects of ghrelin in the development of the African ostrich and associated regulatory mechanisms. 2. Materials and methods 2.1. Animals Female African ostriches (age: neonatal day 1, 45, 90, and 334) were used in the present study. African ostrich chicks (24 females) were obtained from a standard ostrich farm in the Guangdong Province, China, on postnatal day 1 (newly hatched chicks) and were transported within 10 h to a battery house, where feed and water were made available ad libitum. The 24 birds were divided into 4 groups (6 ostriches per group) on the basis of their body weight (BW) in order to adjust for the BW and the variance among the groups. All birds were maintained in a heated room with slatted plastic flooring and were fed a starter diet for postnatal days 1–334, which was formulated according to the specifications of the Elsenburg Ostrich Feed Database [17]. All procedures were approved by the Animal Care and Welfare Committee of our institute.
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2.4. Primer design According to the published sequences of the broiler chicken ghrelin mRNA (GenBank accession no. AB075215), primer sets were designed using the Primer premier 5.0 software. The primer sequences generating a fragment of 191 bp were as follows: sense primer: 5′ CTCTGGCTGGCTCTAGTTTTTTAAG 3′; antisense primer: 5′ CGCTTCTGTTATCTTGACACCAATTTC3′. 2.5. Polymerase chain reaction (PCR) RT products (3.0 μL) were amplified in a 25-μL PCR reaction containing 3.0 μL cDNA, 1.0 μL each for the primer (10 μM), 1.5 μL each for dNTP (10 mM), 0.25 μL Taq DNA polymerase (5 U/μL) (Takara), 1.00 μL MgCl2 (25 mM), and 2.5 μL of 10×PCR buffer. The thermal cycling parameters were 1 cycle at 94 °C for 5 min, followed by 30 cycles at 94 °C for 1 min, 60 °C for 1 min, 72 °C for 2 min, with a final extension step at 72 °C for 10 min. Under these conditions, reactions were conducted within the linear phase of amplification for each PCR product. The resulting PCR products were analyzed by electrophoresis on a 1% agarose gel, which was subsequently analyzed with a computer flatbed scanner. 2.6. Cloning and sequence analysis of the amplified fragments The PCR products of the proventriculus on day 45 were excised after the correct size had been confirmed by electrophoresis on a 1% agarose gel, purified using the OMEGA DNA Purification Kit (V-gene Biotechnology Ltd., Hangzhou, China) according to the manufacturer's manual, and then cloned into the pMD18-T simple vector. Subsequently, the ligation products were transformed into competent DH5α cells. Successful insertion was then confirmed by PCR using the above primers. Positive clones based on colony PCR selection were picked out for sequencing. Bacteria-liquids containing the fragment of the right size were sequenced by the Jin SHIRUI Biotechnology Co., Ltd. (Nanjing, China).
2.2. Tissue preparation 2.7. Ghrelin tissue distribution and mRNA expression On postnatal days 1, 45, 90, and 334 (control group), the birds were weighed, deeply anesthetized with 10% urethane (Caoyang Secondary Chemical Plant, Shanghai, China) at a dose of 1 g/kg BW, and initially perfused with 1000 mL of 0.85% normal saline (containing 0.075% sodium citrate). Then, the abdomen was cut open. Segments of their gastrointestinal tracts, approximately 1 cm in length, were quickly removed and opened along their longitudinal axes. The specimens were obtained from the following portions of the gastrointestinal tracts: the proventriculus, the gizzard, the duodenum, the jejunum, the ileum, the cecum, the colon, and the rectum. Tissues were removed, immediately snap frozen in liquid nitrogen, and then stored at −80 °C until use. 2.3. RNA extraction, cDNA synthesis, and reverse transcription The weight of the frozen tissues was determined and 1 mL of Trizol (Invitrogen, USA) was added for every 100 mg of tissue. Total RNA was extracted from the gastrointestinal tract tissues of developing African ostrich chicks using the Trizol reagent (Invitrogen) according to the manufacturer's instructions. RNA pellets were suspended in 20 μL of DEPC-treated water, and the RNA concentration was determined by spectrophotometry. Total RNA (2 μg) was reverse transcribed in a final volume of 25 μL containing 1.0 μL AMV reverse transcriptase (200 U) (Takara, Japan), 1.5 μL dNTPs (10 mM) (Takara), 0.5 μL RNasin ribonuclease inhibitor (25 U) (Takara), and 0.5 μg of Oligo dT primer (Takara) in DEPC-treated water and buffer supplied by the manufacturer. The reaction was incubated at 42 °C for 60 min. The cDNA was then heated to 70 °C for 5 min to inactivate the reverse transcriptase. All cDNA samples were stored at − 20 °C until use.
RT-PCR analyses of ghrelin expression in different tissues were performed as follows. Total RNA was isolated using standard Trizol methods (Invitrogen, Inc., Carlsbad, CA). The RT was performed using the same methods with Section 2.3. The PCR were performed using the same methods with Section 2.5. The resulting PCR products were loaded onto a 1% agarose gel and visualized by ethidium bromide staining. Each sample was repeated three times and the average mRNA expression was obtained for further analysis. 2.8. Statistics methods The homology of ghrelin among chicken, turkey, emu, goose, Japanese quail and duck was studied. The sequences of the amplified fragments were compared with other birds using the Nucleotide Blast programs at NCBI (http://www.ncbi.nlm.nih.gov/BLAST/) and identity percentages were obtained from amino acid BLAST with the DNASTAR software (http://www.dnastar.com/) (accession no. for emu is AY338467, turkey AY333783, chicken AB075215, duck AY338466, Japanese quail AB244056 and goose AY338465) The obtained sequences of ghrelin were aligned with the Mega 3.1 software (http://www.megasoftware.net/). 3. Results 3.1. RT-PCR of ghrelin genes Total RNA from the proventriculus of a 45-day old African ostrich was used as an initial sample to amplify ghrelin genes by RT-PCR, which yielded a 191-bp cDNA fragment (Fig. 1).
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J.X. Wang et al. / Regulatory Peptides 167 (2011) 50–55 Table 1 The identity comparison among ghrelins and other birds.
Fig. 1. PCR result of ghrelin gene of African ostrich. M. DL 2000; 1. negative control; 2. PCR product (ghrelin).
3.2. Structure of African ostrich ghrelin genes The amplified ghrelin cDNA fragments were then cloned into the pMD18-T simple vector. After ligation, the products were identified by PCR using the above primers. Those bacteria-liquids containing the fragment of the right size were then sequenced. PCR products of 191 bp length were obtained (Fig. 2), and the sequences of the amplified fragments were compared with the corresponding reported sequences of other birds. The results showed that there were 84, 84, 95, 86, 82, and 84% sequence identity with the chicken, duck, emu, goose, Japanese quail and turkey cDNA, respectively (Table 1). 3.3. Structure of the African ostrich ghrelin protein A search for the translation of the nucleotide sequences into the amino acids was performed using the DNA MAN software. The result indicated that the amino acid sequence of the mature ghrelin protein is a peptide of 28 amino acids, in which the Ser3 could be n-octanoylated although we have not confirmed it in this study (Fig. 2). Comparisons of the mature ghrelin protein from different avian species showed a high degree of amino acid sequence similarity with complete conservation of the first 7 N-terminal amino acids (GSSFLSP) of the mature ghrelin peptide that contains the site (serine 3) of fatty acid acylation (Fig. 3). In addition, the sequence of the amino acid sequence of the mature ghrelin protein was compared with the corresponding reported sequences of the ghrelin protein from 6 species. The results showed that there were 53.5, 51.8, 62.3, 53.5, 54.8, and 53.5% sequence identity with chicken, duck, emu, goose, Japanese quail and turkey ghrelin, respectively (Table 2). To further eliminate the noise caused by saturation of synonymous nucleotide substitution, protein sequences were used to construct the phylogenetic tree using a homology tree of the DNASTAR software. The results showed that the coefficient of relationship between the African ostrich mature ghrelin and emu mature ghrelin was neighbor (Fig. 4). These results indicated that the amplified cDNA fragments were African ostrich ghrelin.
Accession
Description
Length (bp)
Identity (%)
AB075215 AY338466 AY338467 AY338465 AB244056 AY333783
Chicken Duck Emu Goose Japanese quail Turkey
210 208 304 226 196 210
84% 84% 95% 86% 82% 84%
3.4. Distribution of ghrelin mRNA expression in the gastrointestinal tract of the African ostrich On postnatal days 1, 45, 90, and 334, ghrelin mRNA expression was found in different tissues in female African ostriches; the distribution of ghrelin mRNA in the gastrointestinal tract changed with age from postnatal day 1 to day 90. Ninety-day-old female African ostriches were used to investigate the distribution of ghrelin mRNA expression in various tissues by RT-PCR. It was found that ghrelin mRNA existed in the proventriculus, gizzard, duodenum, ileum, cecum, and rectum. There was no expression in the jejunum and colon (Fig. 5). 3.5. Developmental changes in ghrelin mRNA expression in the stomach and small intestine of the African ostrich The result of the RT-PCR revealed that the ghrelin mRNA expression in stomach and small intestine of African ostriches changed with age: on day 1, the ghrelin mRNA was detected in the proventriculus, but there was no expression in other tissues (Fig. 6A). On day 45, ghrelin mRNA was detected in the proventriculus, gizzard, and ileum, but there was no expression in the duodenum and jejunum (Fig. 6B). On day 90 and 334, ghrelin mRNA was found in the proventriculus, gizzard, duodenum, and ileum, but there was no expression in the jejunum (Figs. 5 and 6C). 4. Discussion 4.1. Structure of African ostrich ghrelin genes This is the first report on the gene sequence of African ostrich ghrelin. The cDNA has a length of 191 bp and showed high homologies with those of other species. Kaiya et al. [6] reported that the fulllength chicken ghrelin cDNA was 836-bp long, containing 147 bp in the 5′-untranslated region (5′-UTR), 351 bp of coding region, and 338 bp in the 3′-untranslated region (3′-UTR) using 5′- and 3′-rapid amplification of cDNA ends (RACE) PCR methods. Richards et al. [9] and Yuan et al. [18] also reported that the full-length ghrelin cDNA of turkey, goose, duck, and emu was 869 bp, 886 bp, 745 bp, and 682 bp, respectively. The size of the coding region was also 351 bp. The deduced protein of the turkey, goose, duck, and emu preproghrelin contains 116 amino acids, consisting of 3 parts: the signal peptide,
Fig. 2. The gene and translated product sequence of ghrelin. Graphic representation and nucleotide and amino acid (AA) sequence for African ostrich. The nucleotide sequence of the CDS is underlined. Translated AA sequence of the mature ghrelin hormone is highlighted by the shaded box.
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Fig. 3. Sequence comparison of avian ghrelins. Amino acid comparisons of preproghrelin proteins (116 AA) for different avian species. The locations of mature ghrelin peptide (28 AA) is indicated. Also indicated is the site within the mature ghrelin peptide (serine 3) for acylation by n-octanoic acid binding and signaling. Amino acid sequences shown for chicken, duck, emu goose, turkey and Japanese quail were obtained from GenBank accession nos. AB075215, AY338466, AY338467, AY338465, AY333783 and AB244056, respectively.
Table 2 The sequence of the amino acid sequence of the mature ghrelin protein and a comparison of birds and African ostrich ghrelins. Homology matrix Ostrich Chicken Duck Distance matrix Ostrich Chicken Duck Emu Goose Japanese quail Turkey
100.% 46.5% 48.2% 37.7% 46.5% 45.2% 46.5%
53.5% 100.% 23.1% 21.8% 17.7% 3.1% 6.8%
Emu
Goose
Japanese Turkey quail
51.8% 62.3% 53.5% 54.8% 76.9% 78.2% 82.3% 96.9% 100.% 78.9% 91.2% 75.0% 21.1% 100.% 82.3% 83.3% 8.8% 17.7% 100.% 82.3% 25.0% 16.7% 17.7% 100.% 25.2% .218% 18.4% 4.2%
53.5% 93.2% 74.8% 78.2% 81.6% 95.8% 100.%
mature ghrelin, and C-terminal peptide [9,18]. In this study, the primers were designed according to the chicken cDNA sequence of mature ghrelin (accession no. AB075215), and it could amplify a 191-bp fragment, which only contains the sequence of the mature form of ghrelin. The sequence of African ostrich ghrelin has high homologies with those of other species as shown by the highly conserved parts in the sequence of mature ghrelin. However, the sequence of the full-length ostrich ghrelin gene remains to be further studied. 4.2. Structure of the African ostrich ghrelin protein The present study showed that the mature ghrelin is a peptide consisting of 28 amino acids, in which the Ser3 could be n-octanoylated
although we have not confirmed it in this study. Comparisons of the sequences of mature ghrelin from different avian species showed a high degree of amino acid sequence similarity with complete conservation of the first 7 N-terminal amino acids (GSSFLSP), which contain the site (serine 3) of fatty acid acylation. These results further prove that ghrelin is unambiguously identified as an [n-octanoyl-Ser 3]-peptide and suggest that the ghrelins are highly conserved. The acylated peptide and n-octanoylation at serine 3 is essential for ghrelin activity [1,19]. In birds, ghrelin stimulates GH release in vivo and in vitro [6,11,20], decreases food intake in neonatal chicks [7,14], inhibits water intake [21], controls gonadal functions, such as proliferation, apoptosis, and hormone secretion, in chicken [8], and stimulates the contraction of the crop and proventriculus [22]. However, the physiological functions of the African ostrich ghrelin remains to be further studied. In the chicken, the mature ghrelin peptide consists of 26 amino acids and is processed from the precursor at the dibasic sequence ArgArg (RR) located at the C-terminal end of the peptide [6]. In the turkey this sequence is Pro-Arg (PR), and the change may indicate a potential difference in the size of the mature turkey ghrelin peptide (28 vs. 26 amino acids) as compared to the chicken due to different C-terminal end proteolytic processing [9]. In the African ostrich, this sequence is Cys-Arg(CR), and the change may also indicate a potential difference in the size of the mature ostrich ghrelin peptide (28 vs. 26 amino acids) as compared to the chicken due to the generation of a 28-amino acid peptide by enzymatic cleavage in the C-terminal. All other avian species reported to date, except for turkey, contain the Arg-Arg (RR)
Fig. 4. Cladogram analysis of birds and African ostrich ghrelins. The coefficient of relationship between the African ostrich mature ghrelin and emu mature ghrelin was near.
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the ghrelin gene expression in the gastrointestinal tract of African ostrich chicks. However, the reason remains to be further studied. 4.4. Developmental changes of ghrelin mRNA expression in the stomach and small intestine of the African ostrich Fig. 5. RT-PCR analysis of ghrelin mRNA from gastrointestinal tract tissues of African ostrich on postnatal days 90. Agarose gel (1%) electrophoresis of the RT-PCR products for ghrelin showed 191 bp fragments. M, DNA molecular weight marker; 1. negative control; 2. jejunum; 3. proventriculus; 4. gizzard; 5. duodenum; 6. ileum; 7. cecum; 8. rectum; 9. colon.
sequence at the C-terminal end of the ghrelin peptide which makes the C-terminal sequence of mature ghrelin unique in ostrich, like in the case of turkey. 4.3. Distribution of ghrelin mRNA expression in the gastrointestinal tract of the African ostrich Using RT-PCR, ghrelin mRNA expression has been found in the proventriculus, duodenum, and ileum of chickens, broiler chickens, and layer chicks [6,9,23]. In agreement with these results, ghrelin mRNA expression was also found in the gastrointestinal tract of African ostriches, including the proventriculus, duodenum, and ileum. In the present study, RT-PCR revealed that ghrelin was further expressed in the gizzard, cecum, and rectum of African ostriches. There was no expression in the jejunum and colon. Comparison of the present data with chickens revealed that this expression pattern was not identical because ghrelin was not expressed in the gizzard of chickens. Shao et al. [24] reported that ghrelin is expressed in the proventriculus, proventriculus–gizzard junction, gizzard, and duodenum, but not in the ileum and large intestine in Peking duck [24]. It is possible that the difference of ghrelin expression in the gastrointestinal tract of African ostriches is an individual difference among species. These results indicate that there is a species-specific difference in ghrelin expression in birds. We reported the distribution of ghrelin-ip cells in the gastrointestinal tract of African ostrich chicks and indicated that African ostrich ghrelinip cells were found to be localized in the mucous membrane of the entire gastrointestinal tract [16]. The results of the present study indicated that ghrelin is not expressed in the jejunum and colon of African ostriches. Thus, there are differences between distribution of ghrelin-ip cells and
Fig. 6. RT-PCR analysis of ghrelin mRNA from gastrointestinal tract tissues of African ostrich at three stages of development. Agarose gel (1%) electrophoresis of the RT-PCR products for ghrelin showed 191 bp fragments. Expression of ghrelin mRNA on posthatching-day 1 (A, P1), 45 (B, P45) and 334 (C, P334). M, DNA molecular weight marker; 1. negative control; 2. ileum; 3. duodenum; 4. jejunum; 5. gizzard; 6. proventriculus.
The most interesting result obtained in the present study was with regard to the ontogenic pattern of the ghrelin mRNA expression. Comparison of the present data with the results obtained from birds showed some similarities and differences. For example, in chickens, ghrelin mRNA expression was only detected in the proventriculus of newly hatched chicken, whereas mRNA expression was also detected in the duodenum of adult chickens [10]; Chen et al. [25] reported that the level of chicken ghrelin mRNA in the proventriculus was low from embryonic day 15 (E 15) to E 19, but dramatically increased at posthatching day 2 (P 2), then remained constant until P 30 and followed by a significant decrease at P 44 when there was a diet transition at P 31 and thereafter. The decreased level was reversed at P 58. In Peking duck, ghrelin mRNA expression was detectable at very low levels on E 14 in the proventriculus, and the expression increased by E 21. The expression level remained unchanged on P 1 and P 60 [24]. Our results show that the ghrelin mRNA expression in the gastrointestinal tract of African ostriches changes with age (from postnatal day 1 to day 90). These results were consistent with the number of ghrelin-ip cells that increased with age in the gastrointestinal tract of African ostriches (from postnatal day 1 to day 90) [16]. These results suggest that the part of ghrelin mRNA expression in birds is tissue- and ontogenic stage-specific. Acknowledgements We would like to thank Dr. Liu Huazhen of the Department of Anatomy, Histology and Embryology, College of Animal Science and Veterinary Medicine, Huazhong Agricultural University for her valuable comments on the experiments. This study was supported by the National Natural Science Foundation Project of China, No. 30471249 and No. 39970547. References [1] Kojima M, Hosoda H, Date Y, Nakazato M, Matsuo H, Kangawa K. Ghrelin is a growth hormone-releasing acylated peptide from stomach. Nature 1999;402: 656–60. [2] Ariyasu H, Takaya K, Tagami T, Ogawa Y, Hosoda K, Akamizu T, et al. Stomach is a major source of circulating ghrelin, and feeding state determines plasma ghrelinlike immunoreactivity levels in humans. J Clin Endocrinol Metab 2001;86:4753–8. [3] Gnanapavan S, Kola B, Bustin SA, Morris DG, McGee P, Fairclough P, et al. The tissue distribution of the mRNA of ghrelin and subtypes of its receptor, GHS-R, in humans. J Clin Endocrinol Metab 2002;87:2988–91. [4] Sakata I, Tanaka T, Matsubara M, Yamazaki M, Tani S, Hayashi Y, et al. Postnatal changes in ghrelin mRNA expression and in ghrelin-producing cells in the rat stomach. J Endocrinol 2002;174:463–71. [5] Ueno H, Yamaguchi H, Kangawa K, Nakazato M. Ghrelin: a gastric peptide that regulates food intake and energy homeostasis. Regul Pept 2005;126:11–9. [6] Kaiya H, Van der Geyten S, Kojima M, Hosoda H, Kitajima Y, Matsumoto M, et al. Chicken ghrelin: purification, cDNA cloning, and biological activity. Endocrinology 2002;143:3454–63. [7] Saito ES, Kaiya H, Tachibanat T, Tomonaga S, Denbow DM, Kangawa K, et al. Inhibitory effect of ghrelin on food intake is mediated by the corticotropinreleasing factor system in neonatal chicks. Regul Pept 2005;125:201–8. [8] Sirotkin AV, Grossmann R, Maria-Peon MT, Roa J, Tena-Sempere M, Klein S. Novel expression and functional role of ghrelin in chicken ovary. Mol Cell Endocrinol 2006;257–258:15–25. [9] Richards MP, Poch SM, McMurtry JP. Characterization of turkey and chicken ghrelin genes, and regulation of ghrelin and ghrelin receptor mRNA levels in broiler chickens. Gen Comp Endocrinol 2006;145(3):298–310. [10] Wada R, Sakata I, Kaiya H, Nakamura K, Hayashi Y, Kangawa K, et al. Existence of ghrelin-immunopositive and expressing cells in the proventriculus of the hatching and adult chicken. Regul Pept 2003;111:123–8. [11] Baudet ML, Harvey S. Ghrelin-induced GH secretion in domestic fowl in vivo and in vitro. J Endocrinol 2003;179:97–105. [12] Furuse M, Tachibana T, Ohgushi A, Ando R, Yoshimatsu T, Denbow DM. Intracerebroventricular injection of ghrelin and growth hormone releasing factor inhibits food intake in neonatal chicks. Neurosci Lett 2001;301:123–6.
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