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Institute for Marine Biosciences, 1411 Oxford Street, Halifax, Nova Scotia, B3H 3Z1 Canada
(Correspondence should be addressed to A J Manning; Email: amanning{at}munalum.ca)
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Ghrelin undergoes post-translational modifications, often appearing in acylated and unacylated forms in the blood. The acylation occurs at the third residue of the ‘active core’ consisting of a Gly-Ser-Ser-Phe motif at the N-terminus of the mature peptide. An octanoylated form, often seen in mammals, is responsible for the GHS effects (Kojima et al. 1999). Unacylated ghrelin has no effect on GH secretion and thus had been considered biologically inactive. However, recent studies have shown otherwise, with unacylated peptide having actions that mitigate acylated ghrelin effects on insulin secretion (Broglio et al. 2004, Gualillo et al. 2006). A further complication to the understanding of ghrelin regulation is the discovery that another peptide with potentially opposing effects on appetite, obestatin, is encoded in the preproghrelin mRNA transcript (Zhang et al. 2005). This peptide slows gastric emptying activity, antagonizing the stimulatory action of ghrelin (Zhang et al. 2005). However, obestatin secretion does not respond to fasting, nor does it induce GH secretion (Zhang et al. 2005). An obestatin encoding sequence has been recently identified in black sea bream, Acanthopagrus schlegelii (Yeung et al. 2006).
The wide spectrum of ghrelin actions has prompted research into its physiological functions in different animal models including fishes. Ghrelin has thus far been identified in nine different teleosts, from anguilliforms to perciforms: Japanese eel, Anguilla japonica (Kaiya et al. 2003a), goldfish, Carassius auratus (Unniappan et al. 2002), zebrafish, Danio rerio (GenBank Acc. AM055940), channel catfish, Ictalurus punctatus (Kaiya et al. 2005), rainbow trout, Oncorhynchus mykiss (Kaiya et al. 2003b), two tilapia species, Oreochromis mossambicus (Kaiya et al. 2003c) and Oreochromis niloticus (Parhar et al. 2003), sea bass, Dicentrarchus labrax (GenBank Acc. DQ665912), and black sea bream (Yeung et al. 2006). The size of ghrelin peptides is variable among teleosts, and more than one preproghrelin transcript may occur within a species (e.g., rainbow trout, Kaiya et al. 2003b). Post-translational modifications of teleostean ghrelin include acylation as in mammals, C-terminal amidation unique to teleosts, and glycine C-terminus extension as found in channel catfish (Kaiya et al. 2005, Unniappan & Peter 2005). Depending on the species, ghrelin is expressed either exclusively in the stomach or is distributed in other tissues as well, namely, brain, gill, heart, intestine, kidney, or spleen (Unniappan & Peter 2005; above studies).
Physiological roles common to both teleostean and mammalian ghrelins include: orexigenic actions (Unniappan et al. 2004, Riley et al. 2005); stimulation of GH transcription and release (Kaiya et al. 2003a,b,c, 2005, Parhar et al. 2003, Ran et al. 2004); and stimulation of prolactin secretion (Kaiya et al. 2003a,c). Increased ghrelin transcription and peptide secretion have been reported in goldfish under short- and long-term fasting conditions (Unniappan et al. 2004). However, a response to fasting was not seen in sexually mature Nile tilapia (Parhar et al. 2003). Regulation of ghrelin secretion beyond meal frequency is not clear for teleosts, but recent evidence has shown that GH can stimulate ghrelin expression in sea bream (Yeung et al. 2006). Positive feedback may occur between GH and ghrelin under certain physiological conditions in fish.
Here, we examine the expression of ghrelin in a large pleuronectid flatfish, the Atlantic halibut, Hippoglossus hippoglossus. Halibut larvae hatch in an altricial state and spend their early development in a yolk-sac larval stage during which time growth is dependent on endogenous reserves supplied during maternal oogenesis. Many organs are in a rudimentary state and develop functionality during the larval period. For instance, the mouth is not functional and a differentiated stomach is absent at hatching (Pittman et al. 1990, Murray et al. 2006). Major developmental stages during larval growth include the onset of exogenous feeding (first feeding) and metamorphosis from the larval to juvenile stage. Metamorphosis is a complex and highly energy-dependent process during which physiological relationships mature, and key events such as ossification, blood pigmentation, and skin pigmentation take place. Flatfish metamorphosis is particularly complex due to eye migration and changes in body orientation. Larvae of the same natal age can exhibit considerable size variation, possibly due to differential appetite, food intake, or GH secretion. In this study, we present the full cDNA sequence of preproghrelin and, for the first time in fishes, the ontogenetic pattern of ghrelin expression through larval development. In addition, tissue and cellular sites of expression are investigated, and associations between larval size/developmental stage and ghrelin expression level are examined. Ghrelin's role as a feeding inducer may be pertinent for the survival of altricial halibut larvae, particularly at first feeding, since energy reserves are limited following yolk-sac absorption.
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Materials and Methods |
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Adult halibut stomach tissue was homogenized using a Polytron (Kinematica AG, Luzern, Switzerland) and total RNA was extracted according to the protocol specified by the RNeasy Midi-tissue kit (Qiagen). Following DNase treatment (DNA-free kit, Ambion, Austin, TX, USA), 2 µg total RNA were used for first-strand cDNA (fscDNA) synthesis using Retroscript (Ambion) and a 3' universal primer (RT UP; Table 1). The resulting cDNA was extracted with phenol:chloroform extraction (1:1), precipitated with ethanol, and resuspended in 50 µl H2O.
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The 5' end of the cDNA was obtained by RACE using 1 µg halibut stomach total RNA with primers Race 1R and Race 2NR (Table 1) and the SMART RACE cDNA Amplification Kit (BD Biosciences Clontech). RACE reactions were performed according to the manufacturer's recommendations using the following cycling conditions: 94 °C for 5 s; 5 cycles of 94 °C for 5 s, 65 °C for 10 s; 72 °C for 1 min; 25 cycles of 94 °C for 5 s, 62 °C for 10 s, 72 °C for 1 min; 72 °C for 10 min. A nested PCR was performed using a 1/150 dilution of the primary reaction with primer Race 2NR and the same cycling conditions as the first reaction except that annealing temperatures of 62 °C and 60 °C were used. The 350 bp PCR product was purified using a Montage kit (Millipore, Bedford, MA, USA) and sequenced directly. The full-length cDNA was amplified from fscDNA using primers Full 1F and Full 2R (Table 1). PCR conditions were 94 °C for 1 min; 30 cycles of 94 °C for 30 s, 53 °C for 30 s, 72 °C for 90 s; final extension at 72 °C for 3 min. The amplicon was purified, cloned, and sequenced as described above.
Sampling of larval halibut
Atlantic halibut gametes were obtained from captive broodstock and the larvae reared at Scotian Halibut Limited (Clark's Harbour, NS, Canada; 43° 26.3' N; 65° 37.8' W) according to standard industry protocols. Samples of larvae were collected at five developmental periods: hatching (S1), mouth opening (S2), post-notochord flexion/pro-metamorphosis (S6), active metamorphosis (S7–S9), and late metamorphosis/juvenile (S9+) (Table 2). The size and morphology of the larvae in these collections were compared with published staging descriptions for Atlantic halibut (Pittman et al. 1990, Sæle et al. 2004). Additional samples of larvae from pro-metamorphic to early climax metamorphosis (S7–S9) were taken and graded according to size and developmental stage (Table 3). For this latter sample, visceral material was dissected from five individuals from each stage for investigation of ghrelin gene expression.
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Sampling of tissues from 3-year-old immature halibut
Halibut aged 3 years old (43–51 cm total length), originally reared at Scotian Halibut Limited, were sampled from juvenile populations held at the National Research Council Marine Research Station (Sandy Cove, NS, Canada; 44° 30.4' N; 63° 36.6' W). Three males and three females were given an overdose of TMS and then killed by post-cranial section of the spinal cord. Pieces of different tissues were dissected, placed directly in RNAlater (Ambion) for 16 h at 4 °C, removed from RNAlater, and stored at –80 °C until RNA isolation. Tissues examined include: gill, heart, liver, head kidney, body kidney, muscle, spleen, stomach, pyloric caecae/pancreas, intestine, gonad, as well as the pituitary and brain. Brain tissue was further dissected into five areas: olfactory bulb and telencephalon, optic lobe, thalamus and hypothalamus, cerebellum, and medulla oblongata.
RNA extraction of larval and tissue samples from 3-year-old halibut
Total RNA preparations used to investigate ghrelin expression through larval development were based upon extractions of pools of whole larvae. For early stages, 15 or 20 larvae were collected, whereas for later stages only five larvae were collected (Table 2). RNA was prepared from three replicate pools per stage. In an additional study, 15 individual larvae sampled during active metamorphosis (S7–S9) were divided among three size/developmental groups (n=5 larvae per group; Table 3). The viscera were dissected from each individual and total RNA extracted for each larva. Further investigation of visceral ghrelin expression was analyzed based on these separate extracts.
Whole larvae pools or individual viscera samples were weighed, placed in the recommended proportions of Trizol reagent (Invitrogen), and homogenized using a Polytron (Kinematica AG). Total RNA was extracted using Trizol in combination with Eppendorf Phase Lock Gel Heavy tubes (Brinkman, Mississauga, ON, Canada) as specified by the manufacturers' protocols. Following precipitation, RNA pellets were washed twice with 75% (v/v) ethanol prior to resuspension in DEPC-treated water. RNA was quantified using a NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies Inc., Wilmington, DE, USA) and 10 µg aliquots were treated with DNAse I using DNA-free (Ambion). Samples were requantified before storage at –80 °C.
An alternate extraction method was employed for tissues sampled from the six 3-year-old halibut. RNeasy Mini- or Midi-kits (Qiagen) were used following hand homogenization of tissue in RNeasy kit RLT buffer. Homogenates were passed through 18- then 21-gauge needles to disrupt tissue thoroughly. Eluted RNA isolates were treated with DNA-free (Ambion). Each individual was represented by its own tissue-based series of total RNA extracts that were processed separately in subsequent analyses.
Localization of ghrelin expression in larvae by in situ hybridization (ISH)
Probes for halibut ghrelin were prepared by in vitro transcription of the linearized full-length cDNA clone using the DIG RNA labeling kit (Roche Diagnostics) with modifications previously described in Murray et al. (2003).
Halibut sampled from larval pro-metamorphosis to post-metamorphic juvenile stages were killed with an overdose of TMS, fixed overnight at 4 °C by immersion in 4% (v/v) paraformaldehyde in Tris–HCl (pH 7.8), and processed for paraffin embedding (Murray et al. 2006). For larger fish, the visceral region and the head were removed and processed into paraffin separately.
Sections (7 µm) were cut on a rotary microtome, placed on 3-aminopropyl-triethoxysilane-coated slides (Sigma) and baked overnight at 60 °C. After deparaffinizing in xylene and rehydrating through an ethanol series to fish saline (Murray et al. 2003), sections were equilibrated in 0.1 M triethanolamine (TEA, pH 7.5) for 5 min at room temperature with agitation and then acetylated in two further washes of fresh TEA containing 0.025% (v/v) acetic anhydride for an additional 5 min with vigorous agitation. The reaction was stopped in 2x SSC for 2 min and sections were subsequently dehydrated through ethanol series, air-dried, and stored in an RNase-free container.
The ISH protocol, including prehybridization, riboprobe hybridization, and signal detection, was carried out using a Freedom EVO liquid handling robot (Tecan, San Jose, CA, USA). Slide assembly into Teflow chambers (Tecan) and instrument setup were as described in Yaylaoglu et al. (2005). The instrument was directed by a program script written using Gemini software (Tecan) and paralleled the manual ISH protocol as described in Murray et al. (2003). All liquid transfers and incubations took place on the automated platform. Briefly, sections were equilibrated in methanol, washed in fish saline and treated with proteinase K (2.5 µl/ml) for 30 min at 37 °C. Sections were prehybridized in hybridization buffer for 30 min at 25 °C and then hybridized at 47 °C with two consecutive applications of probe (antisense 400 ng/ml, and sense 200 ng/ml) 
2.5 h apart. Total hybridization time was 5 h. After hybridization, sections were incubated in 50% (v/v) formamide with 2x SSC and then washed 30 min in each of 2x SSC, 0.5x SSC, and 0.1x SSC at 47 °C. Following a blocking step in antibody buffer (0.1 M Tris–HCl, pH 7.5, 150 mM NaCl) with 1% (w/v) BSA and 10% (v/v) goat serum, the hybridization signal was detected by incubating the sections for 30 min at 2 °C with a sheep anti-digoxigenin-alkaline phosphatase conjugate (Roche). The reaction was visualized by incubating for 15 min in a chromogenic buffer (0.1 M Tris–HCl pH 9.5, 0.1 M NaCl, 0.05 M MgCl2, 0.1% (v/v) Tween 20, and 1 mM levamisol) followed by 20 min incubation in fresh buffer containing 3.5 µl/ml 5-bromo-4-chloro-3-indolyl-phosphate (Roche) and 4.5 µl/ml nitro blue tetrazolium (Roche). The reaction was stopped by washing in system water from the instrument and the reaction product stabilized by fixing in 4% (v/v) paraformaldehyde in fish saline for 10 min followed by a final system water wash. Hybridization chambers were disassembled in clean water, the slides air-dried at room temperature, mounted in an aqueous mounting media (Hydro-Matrix, Micro-Tech-Lab, Graz, Austria), and cover slipped for observation using a Leica DMRE microscope and image capture with an Orca camera system (Hamamatosu Inc., Bridgewater, NJ, USA) and Simple PCI software (Compix Inc. Imaging Systems, Cranberry Township, PA, USA). Image plates were created using Adobe Photoshop 6.0.1. (Adobe System Incorporated).
Quantitative real-time PCR assays for gene expression in ontogenetic studies
First-strand cDNA synthesis for each RNA extract was conducted using 2 µg DNase I-treated total RNA and an oligo-dT primer with the RetroScript kit (Ambion). Amplification of cDNA was performed using an iCycler (Bio-Rad Laboratories). Data obtained for ghrelin were compared with data obtained from amplification of two reference genes, elongation factor 1-
(EF1) and 40S ribosomal protein S4 (40S). Samples of cDNA were diluted 50-fold for use in the reference gene assays, while those for ghrelin were diluted either 10-fold or were used undiluted (pooled larval extracts). PCR (20 µl) contained 2 µl cDNA solution, 0.8 µl (10 µM) forward and reverse primers (Table 4), 6.4 µl nuclease-free water, and 10 µl iQ SYBR Green Supermix (Bio-Rad). PCR product sizes for ghrelin, EF1, and 40S were 305 bp, 141 bp, and 138 bp respectively. All reactions were performed in duplicate within a PCR assay and under the same cycling conditions: 95 °C for 10 min; 42 cycles of 95 °C for 30 s; 57 °C for 30 s; and 72 °C for 1 min. Fluorescent data for quantification were collected at 82 °C. Melt curve analysis was performed over a range of 55–95 °C in order to verify single product generation at the end of the assay.
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RT-PCR and quantitative real-time PCR assays of tissue-specific ghrelin expression in 3-year-old halibut
First-strand cDNA synthesis was performed on 5 µg DNase-treated total RNA using oligo-dT with SuperScript II RNase H- reverse transcriptase (Invitrogen) according to the protocol detailed in Matsuoka et al. (2006). The RT-PCR amplification of ghrelin cDNA was carried out as follows: denaturation at 95 °C for 3 min; 31 cycles of denaturation at 95 °C for 30 s, annealing at 57 °C for 30 s, and extension at 72 °C for 30 s; and a final extension step at 72 °C for 5 min. Primers used for ghrelin amplification were the same as those used for quantitative PCR (Table 4). Amplification of glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was performed on the cDNA samples using the same protocol but with an annealing temperature of 53 °C (GAPDH forward primer, 5'-GTG TCA GTG GTT GAC CTG A-3'; GAPDH reverse primer, 5'-AGC TTG ACA AAG TGG TCA TT-3'). PCR amplicons were visualized on a 2% (w/v) agarose gel with SYBR Safe DNA Gel Stain (Invitrogen).
Quantitative PCR was additionally performed on the tissue cDNA samples using the same protocol as outlined for the larval ontogenetic studies, with the exception that EF1 and another reference gene, ribosomal protein L13A (RPL13A), were used as reference genes for normalization of tissue expression.
Statistical analysis of expression data
Normalized expression values obtained for different samples were averaged and the mean values were used for graphical representation along with their associated S.D. Kruskal–Wallis tests were performed to determine differences in normalized preproghrelin expression in developmental stage comparisons. The Wilcoxon two-sample test was used for post hoc comparisons. Statistical analysis was done using SYSTAT Version 11 (Systat Software Inc., San José, CA, USA).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The initial PCR to isolate preproghrelin gene using degenerate primer P1 amplified three products, a prominent band of 800 bp and two weaker bands of 640 and 200 bp. The 800 bp band was excised and identified as preproghrelin. The smaller weaker bands were therefore not studied further. Following 5' RACE and further cloning and sequencing, the full 899 bp cDNA sequence was obtained for halibut. The sequence includes a 5'-untranslated region of 51 bp, an open reading frame of 315 bp that encodes an 105 amino acid preproghrelin, and a 3'-untranslated region of 533 bp (Fig. 1). For the latter, two potential polyadenylation signals (aataaa) were seen, one at position 882–887 near the end of the cloned sequence and another well upstream of the poly-A tail at position 619–624.
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Ontogenetic ghrelin expression in larval halibut
Quantitative PCR analysis on three replicate sets of pooled larvae collected at five different developmental stages demonstrated that the ghrelin gene is expressed as early as hatching (Fig. 3). Low levels of ghrelin expression were seen over the yolk-sac stage (S1 and S2) with little change in normalized expression occurring even by prometamorphosis (S6 post-notochord flexion). Increased expression was associated with metamorphosis (S7 to S9). The highest expression level was seen for fish staged as either juveniles or advanced climax metamorphic stage 9 larvae. Changes in normalized ghrelin expression over larval development were statistically significant over time (P<0.05, Fig. 3)
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Cellular localization of ghrelin by ISH
In terms of methodology, the present study is the first to employ an automated liquid handling platform to perform ISH on fish tissues. In contrast to the method described in Yaylaoglu et al. (2005), the process here does not utilize an amplification step. The procedure is directly adapted from a previously published manual protocol (Murray et al. 2003) and seems to be effective in detecting moderately expressed genes such as halibut ghrelin. The expression signal was very specific to cell type and showed no significant background.
Halibut of varying size and developmental stage (larval stage 7 to post-metamorphic juvenile) were examined for the presence and distribution of ghrelin-expressing cells. The greatest number of cells was observed in stage 9 to post-metamorphic fish; no difference in cell number or distribution was noted across the size range of fish sampled among these stages. Although all tissues of the fish were examined, ghrelin-expressing cells were only observed in the stomach mucosa (Fig. 4A). Some of these cells were triangular or pyramidal in shape and showed a distinct luminal orientation with a basal nucleus, but they were never observed to reach the luminal surface (Fig. 4B). Other ghrelin-expressing cells were more rounded in shape and did not appear to have a luminal orientation. A strong signal was evident throughout the cytoplasm of ghrelin-expressing cells. No signal was observed for tissues hybridized with a sense probe (Fig. 4C).
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Assessment of ghrelin expression by RT-PCR showed that among the tissues sampled from six halibut (three males and three females), the strongest expression was seen for stomach tissue (Fig. 5). Expression was also evident for most individuals in the following tissues after 31 cycles of amplification: pyloric caecae (n=6 fishes), intestine (n=5), and both the ovaries (n=2–3) and testes (n=3). Examining RT-PCR data obtained after an additional two cycles of amplification suggested that ghrelin was either weakly expressed or absent in other tissues. Tissues with low-level expression include the brain (n=3 fishes), gills (n=3), heart (n=1), and kidney (n=2). Individuals with expression in the brain demonstrated general weak- to low-level expression among the different regions of the brain. RT-PCR did not show prominent expression in any one region that was consistent among individuals.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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500 bp (<535 bp). The detection of two putative poly-adenylation signals within the cloned sequence for halibut suggests the presence of an additional smaller transcript. Multiple transcripts of varying stability can be expressed, and more than one ghrelin transcript has been found in channel catfish, rainbow trout, mouse, and human (Kaiya et al. 2003b, 2005, Gualillo et al. 2006). Although the cloned sequence was the most prominent band found in initial isolation PCR, the weaker 640 bp band detected could represent an alternate transcript associated with the second poly-adenylation signal. Halibut preproghrelin had the highest amino acid identity with preproghrelin sequences from other acanthopterygian teleosts represented by the four perciform species (i.e., tilapias, sea bass, and black sea bream). The deduced 105 amino acid preproghrelin sequence is within the range seen in other fishes (103–111 aa). The active core of the mature ghrelin peptide, represented by the first four amino acids (GSSF), is conserved in all vertebrates studied thus far with the exception of goldfish (Unniappan et al. 2002), zebrafish, and the bullfrog Rana catesbeiana (Kaiya et al. 2001). Generally, the first seven amino acids of the mature peptide are highly conserved among the vertebrates, but if one examines the teleosts alone the conserved area increases to the first 12 amino acids.
A conserved region that aligns with human obestatin was seen within halibut preproghrelin, similar to the previously reported sea bream sequence (Yeung et al. 2006). Mammalian obestatin has been reported to be anorexigenic (Zhang et al. 2005, Lagaud et al. 2007). However, effects on appetite suppression in normal or ghrelin-stimulated animals remain controversial (Seoane et al. 2006, Gourcerol & Taché 2007). Whether obestatin is translated in fishes and forms a functional peptide is unknown. Convertase cleavage sites (represented by the basic amino acids Arg or Lys) that flank mammalian obestatin are not seen at the amino termini of the teleost sequences, but a number of species may have sites at the carboxy termini of their sequences. In addition, the carboxy terminus of the mammalian obestatin sequence is flanked by a glycine residue indicating amidation, which is important for obestatin bioactivity (Zhang et al. 2005). This is not observed in the putative obestatin sequences of teleosts, but is seen among their functional ghrelin peptides. The potential role of obestatin in fishes requires further investigation.
The examination of ghrelin expression through halibut larval development revealed that ghrelin transcripts were detectable from hatching. This is the first report of ghrelin gene expression during teleostean larval development. The biological significance of ghrelin expression during the yolk-sac stage is unclear as growth and nutrition at this time are based on the endogenous energy supplies of the yolk. A role in embryonic development, rather than appetite, may be more relevant for ghrelin at hatching. Studies in mammals that support this idea include reports of ghrelin expression in early embryo development (morula cells, Kawamura et al. 2003) and ghrelin-mediated induction of fetal neurogenesis (Sato et al. 2006).
Normalized ghrelin expression changed little between hatching, mouth-opening, and prometamorphosis (stage 6). Notable increases in expression were seen in larvae undergoing metamorphosis, particularly the later stages of climax metamorphosis and entry into the post-metamorphic juvenile stage (S9+). Metamorphosis is a period when anatomical relationships and physiological systems mature. This is evident in the endocrine systems as well. For instance Einarsdóttir et al. (2006) reported that the pituitary completes development by stage 7 and an increase in pituitary endocrine cells is seen with climax metamorphosis. In terms of ghrelin, increases in expression during advanced metamorphosis are likely tied to the concurrent changes in stomach development. The glandular area of the stomach develops from first feeding in halibut larvae (66 dph; Murray et al. 2006). Initial gastric gland formation was confirmed histologically for stage 6 larvae of the present study (data not shown). Organized glandular areas were detected in metamorphic stages 7–9, but separate mucosal and glandular layers of the stomach were not detected until late climax metamorphosis and post-metamorphosis. These results are consistent with patterns detailed in Murray et al. (2006), which indicate that stomach development, glandular/mucosal layer differentiation, and gastric functionality, shown by pepsinogen expression, are closely associated with metamorphosis. The differentiation of the stomach layers may influence the increase in ghrelin expression since ghrelin-expressing cells were detected in both glandular and mucosal layers of the stomach in halibut larvae.
Two morphological types of ghrelin-expressing cells were evident in the halibut stomach. These were similar to the open- and closed-type cells described in the gastric region of rainbow trout (Sakata et al. 2004). The triangular-shaped cells seen in the surface mucosal layer of the halibut stomach may represent open-type cells as they had a definite luminal orientation. In contrast, other ghrelin-positive cells noted in the glandular region of the stomach, but lacking any discernable orientation, appear to be closed-type cells. These observations are supported by Murray et al. (1994) in an ultrastructural study that described two enteroendocrine cells from the halibut stomach, one of which was seen to have a distinct luminal orientation. Mammalian ghrelin-positive cells (enteroendocrine X/A-like cells, Date et al. 2000) are only seen as the closed-type variety in human and rat stomachs, but are found as both open- and closed-type cells in the lower gastrointestinal tract of rat (Sakata et al. 2002).
Although ghrelin expression in 3-year-old halibut was strongest in stomach, significant expression was also detected in other digestive tissues and in the gonads using sensitive PCR assays. In contrast, ISH results for stage 9 larvae and post-metamorphic juveniles revealed discernable transcripts only in the stomach, despite numerous attempts to detect expression with serial sections of cranial and abdominal regions. This discrepancy suggests that the sensitivity of the ISH method was insufficient for detection of ghrelin expressed in other post-metamorphic tissues than stomach. Alternatively, ghrelin-producing cells may become more numerous in non-gastric tissue with further maturation of the digestive system and can only be detected by ISH in older fish.
Ghrelin expression in fishes is variously reported to be confined to the stomach (Nile tilapia, Parhar et al. 2003; sea bream, Yeung et al. 2006) or to be more widely distributed. In mammals, ghrelin-producing cells, although concentrated in the stomach, are distributed along the digestive tract and have also been detected in human insulin-secreting β-cells of the pancreas (Volante et al. 2002). Goldfish, which lack a distinct stomach, express ghrelin mainly in the intestine (Unniappan et al. 2002). Intestinal expression was also identified by RT-PCR in rainbow trout and Japanese eel (Kaiya et al. 2003a,b). Low-level ghrelin expression in pyloric caecae has been reported in rainbow trout (Kaiya et al. 2003b). For halibut, expression in the pyloric caecae may originate from the diffuse pancreas associated with these digestive structures (Murray et al. 2006). Pancreatic expression has been noted in channel catfish (Kaiya et al. 2005).
Only weak ghrelin expression was observed in the halibut brain, which agrees with the general pattern reported for other teleosts (Unniappan & Peter 2005). Ghrelin expression in the brain has been observed in Japanese eel (Kaiya et al. 2003a), rainbow trout (Kaiya et al. 2003b), Mozambique tilapia (Kaiya et al. 2003c), and goldfish (Unniappan et al. 2002). Studies for rainbow trout and goldfish have demonstrated ghrelin expression in the telencephalon and hypothalamus, with hypothalamic expression having been originally observed in the rat brain (Kojima et al. 1999). Despite the evidence of hypothalamic ghrelin expression in two other teleosts, our analyses using quantitative PCR could only confirm ghrelin expression in the medulla oblongata of male 3-year-old halibut. Expression levels in other areas of the brain, including the thalamus and hypothalamus, were only at the threshold of detection.
Halibut is the first species for which gonadal ghrelin expression has been found among teleosts. Only two other studies have examined fish gonads for ghrelin expression, but no expression was detected (Unniappan et al. 2002, Parhar et al. 2003). Mammalian studies suggest autocrine/paracrine roles for ghrelin in both the ovary and the testis. Ghrelin expression has been shown in steroidogenic and other somatic cells of the mammalian gonads, while expression of its receptor (GHS-R1a) has been noted in both the somatic and germ cells (Barreiro & Tena-Sempere 2004). Thus, the detection of gonadal ghrelin production in halibut prompts the question as to whether this peptide might have a regulatory role in fish reproduction.
In conclusion, we have cloned preproghrelin from Atlantic halibut, the first pleuronectiform teleost to be examined in terms of ghrelin expression. Similarities in mRNA transcript length and amino acid sequence reflect the evolutionary links between halibut and other acanthopterygian teleosts. This study is the first to examine ghrelin expression during fish larval development. The data indicate low levels of ghrelin expression during the prolonged yolk-sac larval stage, prior to exogenous feeding and the differentiation of the stomach. This suggests that ghrelin has activities beyond appetite stimulation and GH secretion in early stages. It may also indicate expression in non-gastric tissues, or a role in development, as has been shown in mammals. Increased expression during larval development was clearly associated with climax metamorphosis and the differentiation of the stomach. ISH showed that dominant expression occurs in stomach tissue. Expression is localized within two cell types associated with the luminal and gastric areas of the developing stomach. The examination of 3-year-old halibut demonstrated that ghrelin is expressed in other tissues including intestine, pyloric caecae, and the immature gonads. Ghrelin effects on appetite and GH release in vertebrates suggest that this hormone could be physiologically important both during and after the larval stage of fishes.
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References |
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Received in final form 8 October 2007
Accepted 17 October 2007
Made available online as an Accepted Preprint 17 October 2007
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