Ghrelin is a gut-brain peptide hormone, which binds to the growth hormone secretagogue receptor (GHS-R) to regulate a wide variety of biological processes in fish. Despite these prominent physiological roles, no studies have reported the anatomical distribution of preproghrelin transcripts using in situ hybridization in a non-mammalian vertebrate, and its mapping within the different encephalic areas remains unknown. Similarly, no information is available on the possible 24-h variations in the expression of preproghrelin and its receptor in any vertebrate species. The first aim of this study was to investigate the anatomical distribution of ghrelin and GHS-R1a ghrelin receptor subtype in brain and gastrointestinal tract of goldfish (Carassius auratus) using immunohistochemistry and in situ hybridization. Our second aim was to characterize possible daily variations of preproghrelin and ghs-r1 mRNA expression in central and peripheral tissues using real-time reverse transcription-quantitative PCR. Results show ghrelin expression and immunoreactivity in the gastrointestinal tract, with the most abundant signal observed in the mucosal epithelium. These are in agreement with previous findings on mucosal cells as the primary synthesizing site of ghrelin in goldfish. Ghrelin receptor was observed mainly in the hypothalamus with low expression in telencephalon, pineal and cerebellum, and in the same gastrointestinal areas as ghrelin. Daily rhythms in mRNA expression were found for preproghrelin and ghs-r1 in hypothalamus and pituitary with the acrophase occurring at nighttime. Preproghrelin, but not ghs-r1a, displayed a similar daily expression rhythm in the gastrointestinal tract with an amplitude 3-fold higher than the rest of tissues. Together, these results described for the first time in fish the mapping of preproghrelin and ghrelin receptor ghs-r1a in brain and gastrointestinal tract of goldfish, and provide the first evidence for a daily regulation of both genes expression in such locations, suggesting a possible connection between the ghrelinergic and circadian systems in teleosts.
Citation: Sánchez-Bretaño A, Blanco AM, Unniappan S, Kah O, Gueguen M-M, Bertucci JI, et al. (2015) In Situ Localization and Rhythmic Expression of Ghrelin and ghs-r1 Ghrelin Receptor in the Brain and Gastrointestinal Tract of Goldfish (Carassius auratus). PLoS ONE 10(10): e0141043. https://doi.org/10.1371/journal.pone.0141043
Editor: Juan Fuentes, Centre of Marine Sciences & University of Algarve, PORTUGAL
Received: July 29, 2015; Accepted: October 2, 2015; Published: October 27, 2015
Copyright: © 2015 Sánchez-Bretaño et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Data Availability: All relevant data are within the paper and its Supporting Information files.
Funding: This collaborative research was supported by the Spanish Ministerio de Economía y Competitividad (AGL2013-46448-C3-2-R) to MJD, and partly by a Discovery Grant and Discovery Accelerator Supplement award from the Natural Sciences and Engineering Research Council of Canada, an Establishment grant from Saskatchewan Health Research Foundation and John R. Evans Leaders Fund from the Canada Foundation for Innovation to SU. SU is a recipient of the Canadian Institutes of Health Research New Investigator Award. ASB and AMB are predoctoral fellows from the Spanish Ministerio de Economía y Competitividad and Ministerio de Educación y Ciencia, respectively. JIB is a predoctoral fellow from the Argentinian Agencia Nacional de Promoción Científica y Tecnológica, and a recipient of the Emerging Leaders of the Americas Program funded by the Government of Canada. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Ghrelin, a peptide hormone mainly synthesized by the gut, was originally purified in 1999 by Kojima and colleagues . The main site for the synthesis of ghrelin in all the vertebrates so far studied is the stomach or its equivalent , although gene expression of ghrelin by PCR shows a widespread tissue distribution, with low expression levels in peripheral tissues (apart from stomach) and brain in both mammals [3,4] and fish [5–7]. Imaging techniques have reported the presence of ghrelin gene and peptide in the brain  and gastrointestinal tract [9–12] of mammals. Similarly, ghrelin peptide has been localized by immunohistochemistry in the hypothalamus of goldfish  and in the gastrointestinal tract of goldfish (Carassius auratus) , sea bass (Dicentrarchus labrax) , zebrafish (Danio rerio) , and rainbow trout (Oncorhynchus mykiss) . However, no studies to date have reported the anatomical distribution of the preproghrelin transcripts using in situ hybridization in a non-mammalian vertebrate, and its mapping within the different encephalic areas remains unknown.
Ghrelin is suggested to have a key role in energy balance regulation by promoting food intake, carbohydrate utilization and adiposity [17,18], and other physiological processes [19,20]. A unique aspect of this peptide is the presence of a posttranslational acyl modification catalysed by a recently discovered member of the membrane-bound O-acyltransferase family, named ghrelin O-acyl transferase [21,22]. This modification is essential for most of the bioactivity of the peptide, enabling the binding to its receptor, the G protein-coupled growth hormone secretagogue receptor (GHS-R). In contrast to mammals and other tetrapods with only one GHS-R gene, ancestral teleost underwent a genome duplication, and two paralog genes (GHS-R1 and GHS-R2) have been identified in otophysi teleosts . Particularly, goldfish has experienced a tetraploidization, and four subtypes of GHS-Rs have been described and characterized in this teleost, named GHS-R1a1, GHS-R1a2, GHS-R2a1, and GHS-R2a2 . In addition, each one of these receptor subtypes presents a type ‘b’ isoform obtained by alternative mRNA splicing. Among all of these different GHS-R, the GHS-R1a has been widely studied in vertebrates in terms of structure, tissue abundance, mechanism of action, dynamics and regulation, and seems to be involved in many of the physiological actions of ghrelin [3,24,25]. Nevertheless, little information is available about its anatomical location and distribution. In this sense, GHS-R1a has been localized in the mammalian brain [26–28], but no neuroanatomic mapping using imaging techniques has been performed in the brain of non-mammalian vertebrates. Among fishes, the distribution of ghrelin receptor in the gastrointestinal tract has been described only in zebrafish .
Most behaviour and physiology of living organisms follow daily rhythms due to the presence of endogenous clocks that synchronize biological processes to the 24-h light/dark cycle, enabling them to anticipate periodic changes in the environment. A growing interest in the relationships between energy balance and the circadian system has occurred in the last years. In fact, daily oscillations have been found in fish for many hormones involved in food intake regulation and metabolism, including neuropeptide Y (NPY)  and leptin . In relation with the ghrelinergic system, only one study carried out in humans described the 24-h secretion profile of ghrelin, showing a circadian oscillation of this peptide with higher levels of circulating ghrelin during the night . However, no information is available about the possible 24-h variations in the expression of this hormone and its receptor in vertebrates.
Goldfish is a member of the teleostean Cyprinidae family and has been widely used for studying the hormonal regulation of feeding in fish [32,33]. The biological activities of ghrelin have been previously examined in this teleost, [34–37] but, despite the growing interest and importance of the ghrelinergic system, no studies have reported a brain mapping of the elements composing this system in fish, and nothing is known about its possible rhythmic expression. Therefore, the aim of the present study was first, to investigate the anatomical distribution of ghrelin and the ghrelin receptor subtype GHS-R1a in the brain and gastrointestinal tract of goldfish by using immunohistochemistry and in situ hybridization techniques. We also characterized the daily profile of preproghrelin and ghs-r1 mRNA expression in both central and peripheral tissues using RT-qPCR.
Material and Methods
All procedures carried out in France were conducted in accordance with the guidelines of Ethical Committee at our institutions (University of Rennes 1, CNRS and INSERM) and in accordance with European Union regulations concerning the protection of experimental animals (Directive 86/609/EEC). The protocols were approved by the Ethical Committee CREEA (Comité Rennais d'Ethique en matière d'Expérimentation Animale) and performed under the supervision of authorized investigators (Permit number: EEA B-35-040). The study conducted in Canada strictly followed the regulations of the Canadian Council for Animal Care and were approved by the University of Saskatchewan Animal Research Ethics Board (Protocol # 2012–0082). The euthanasia was performed by deep anesthesia and all efforts were made to minimize suffering.
Animals, experimental designs and sampling
For the anatomical studies, goldfish (2.0 ± 0.5 g) obtained from a local supplier (Rennes, France) were maintained in 60 l aquaria with filtered and aerated fresh water (22 ± 1°C) under a 12 h light: 12 h darkness (12L:12D) photoperiod (lights on at 9 AM). Fish were daily fed at zeitgeber time 2 (ZT-2, or 11 AM, as ZT-0 corresponds to lights-on) with food pellets (1% body weight; Novo GranoMix, JBL, GmbH & Co., Neuhofen, Germany). Goldfish were fasted for 48 h, and at ZT-2 of the experimental day were anesthetized with phenoxyethanol 1 ml/l (ICN Biomedicals Inc., Irvine, CA, USA) and sacrificed. Then, fish were immersed overnight in 4% paraformaldehyde diluted in 0.1 M sodium phosphate buffer with saline (PBS, pH 7.4). The following day, the brain and the gastrointestinal tract (esophagus, intestinal bulb, j-loop and anterior intestine) were removed and post-fixed 3 h in the same solution. Then, the samples were cryoprotected with 30% sucrose (MP Biomedical, LLC, Illkirch, France) overnight, included in the frozen section medium Richard-Allan ScientificTM Neg-50 (Thermo Shandon Scientific, Cheshir, UK) and stored at -80°C.
For the study of daily changes in gene expression, goldfish (22 ± 8 g) obtained from a commercial supplier (Aquatic Imports, Calgary, Alberta, Canada) were maintained in 200 L aquaria with filtered and aerated fresh water (21 ± 2°C). Fish were maintained under a 12L:12D photoperiod (lights on at 7 AM) and daily fed at ZT-4 with a commercial pellet diet (1% body weight; Martin Profishent, Ontario, Canada). The day of the experiment, fish were sacrificed in 4 h intervals (6 fish/sampling time) throughout a complete 24-h cycle beginning at ZT-0. Fish were randomly collected from two different tanks (21 fish/tank). Food was offered as scheduled the day of the experiment. Once sacrificed, samples of forebrain (including telencephalon and diencephalon without hypothalamus), hypothalamus, hindbrain (including mesencephalon and rombencephalon), pituitary and gastrointestinal tract (including esophagus, intestinal bulb, j-loop and anterior intestine) were collected and immediately frozen in liquid nitrogen. Sampling during darkness was conducted under dim red lighting.
Molecular cloning of preproghrelin and ghs-r1a genes of goldfish and riboprobe synthesis
For the molecular cloning of preproghrelin and ghs-r1a genes, specific primers (Table 1) were designed based on the goldfish sequences (GenBank accession numbers AF454389.1 for preproghrelin, AB504275.1 for ghs-r1a1, and AB504276.1 for ghs-r1a2) and were purchased from Sigma (Sigma-Aldrich, Steinheim, Germany). To clone preproghrelin gene (366 bp), the reaction mixture contained cDNA from goldfish gastrointestinal tract, GoTaq® Green MasterMix (Promega, Madison, WI, USA), and forward and reverse primers (3 nM each). To clone ghs-r1a gene (979 bp), the reaction mixture contained cDNA from goldfish gastrointestinal tract, Taq DNA Polymerase with its buffer (0.00125 U; Invitrogen), dNTPs (10 mM each), MgCl2 (6 mM), and forward and reverse primers (3 nM each). These primers were able to distinguish between the ghs-r1a and ghs-r1b splicing variants, but unable to distinctly identify ghs-r1a1 and ghs-r1a2 subtypes. PCRs were performed in a total volume of 25 μl. PCR conditions were set at 95°C for 3 min followed by 40 cycles of 95°C for 10 sec, 60°C for 30 sec and 72°C for 1 min, and a final extension step of 72°C for 10 min. The amplified products were run on an agarose gel and purified using PCR clean-up gel extraction (MACHEREY-NAGEL GmbH & Co., Düren, Germany) for preproghrelin gene and GenElute gel extraction kit (Sigma, Steinheim, Germany) for ghs-r1a gene. Purified preproghrelin and ghs-r1a PCR products were ligated into pCR®II- TOPO® (Invitrogen, Carlsbad, CA, USA) or pCR™4-TOPO® (Invitrogen, Carlsbad, CA, USA) vectors, respectively, and employed to transform Escherichia coli One Shot TOP10 cells (Invitrogen, Carlsbad, CA, USA) or JM109 cells (Promega, Madison, WI, USA), respectively. Positive clones were collected and plasmid DNA extraction was performed by using a routine miniprep protocol. Plasmids with the insert were linearized with SpeI and NotI. Antisense and sense mRNA probes were obtained with DIG RNA labeling MIX (Roche Diagnostic, Mannheim, Germany) by in vitro transcription with T7 and SP6 RNA polymerases (Promega, Madison, WI, USA) for preproghrelin gene, and with T7 and T3 RNA polymerases (Promega, Madison, WI, USA) for ghs-r1a gene probes. The specificity of the probes was confirmed with parallel series of slides hybridized with the correspondent sense RNA probes.
Localization of preproghrelin and ghs-r1a by in situ hybridization (ISH)
The presence and anatomical distribution of preproghrelin and ghs-r1a transcripts in the brain and gastrointestinal tract of goldfish were studied by in situ hybridization. For this purpose, samples obtained as described earlier were embedded in TissueTek and sectioned at 8-μm thickness using a cryostat. Transverse sections were mounted onto superfrost slides (Thermo scientific, Braunschweig, Germany). The protocol for ISH was performed as previously described  with minor modifications. In brief, cryostat sections were washed in PBS two times during 10 min before post-fixation in Antigenfix (DiaPath, Martinengo, Italy) for 20 min. After washing in PBS, sections were treated for 5 min at 37°C with proteinase K (2 μg/ml, Sigma, Steinheim, Germany) diluted in PBS, and fixed in 4% paraformaldehyde for 15 min. Sections were rinsed twice in 2X standard saline citrate (SSC). Hybridization was performed at 65°C overnight in a humidified chamber using 100 μl hybridization buffer (50% deionized formamide; 2x SSC; 5x Denhardt's solution; 50 μg/ml of yeast tRNA; 4 mM EDTA; 2.5% dextran sulfate) containing the DIG-labeled probe (3 μg/ml). After hybridization, slides were washed in 2x SSC at 65°C (2x30 min), 2x SSC/50% formamide at 65°C (2x30 min), 0.2x SSC (1x15 min) and 0.1x SSC (1x15 min) at room temperature. Slides were next washed in 100 mM Tris-HCl (pH 7.5) containing 150 mM NaCl for 10 min, then washed in the same buffer containing 0.1% Triton and 0.5% of skimmed milk powder (2x30 min), and incubated overnight at room temperature with anti-digoxigenin alkaline phosphatase Fab fragments (1:2,000; Roche Pharma, Mannheim, Germany). The next day, slides were incubated for 4.5 h with an HNPP (2-hydroxy-3-naphtoic acid -2′-phenylanilide phosphate)/FastRED detection kit (Roche Pharma, Mannheim, Germany), according to the manufacturer's instructions. Finally, Vechashield mounting medium containing 4′,6-diamidino-2-phenylindole (DAPI; Vector Laboratories, Burlingame, CA, USA) was applied and coverslips were placed. Slides were observed with an epifluorescence microscope (Olympus Provis, equipped with a DP71 digital camera). Images were processed with either the Olympus Analysis or Zeiss Cell software. Micrographs were generated in the “TIFF” format and adjusted linearly for light and contrast before being assembled on plates using Photoshop CS6.
Localization of ghrelin by immunohistochemistry (IHC)
Immunohistochemical staining of goldfish brain and gastrointestinal tract samples was used to study the anatomical distribution of ghrelin. The immunohistochemistry study was carried out as previously described . Briefly, the above described cryostat sections were washed twice in 0.1M PBS and incubated twice in PBS containing 0.2% Triton and 0.5% of skimmed milk powder (45 min at room temperature). After overnight incubation with primary monoclonal antibody (mouse anti human ghrelin 1:200; ab57222, Abcam, Cambridge, MA, USA), previously used in goldfish by Kerbel and Unniappan , sections were washed three times in 0.2% Triton PBS and subsequently incubated with rabbit antimouse Alexa Fluor 488 (1:400; Invitrogen Molecular Probes, Eugene, OR, USA) for 2 hours at room temperature. A separate set of negative control slides were only treated with the secondary antibody (S1 Fig). After washing in PBS, slides were coverslipped with Vectashield containing DAPI and observed with an epifluorescence microscope (Olympus Provis, equipped with a DP71 digital camera). Imaging processing was conducted as described previously for ISH.
Analysis of daily preproghrelin and ghs-r1 mRNA expression by RT-qPCR
The possible 24-h rhythmic expression of preproghrelin and ghs-r1 in the brain and gastrointestinal tract of goldfish was studied using Real-time or Reverse Transcription-quantitative PCRs (RT-qPCR). Total RNA from forebrain, hypothalamus, hindbrain, pituitary and gastrointestinal tract was isolated using TRIzol RNA isolation reagent (Invitrogen, Carlsbad, CA, USA). RNA purity was validated by optical density absorption ratio (260/280 nm) using a NanoDrop 2000c (Thermo, Vantaa, Finland). Then, an aliquot of 1 μg of total RNA was reverse transcribed into cDNA in a 20 μl reaction volume using iScript cDNA synthesis kit (BioRad, Hercules, USA) according to the manufacturer’s instructions. The RT reactions condition consisted in 25°C for 5 min, an extension of 30 min at 42°C and a denaturalization step at 85°C for 5 min, and were carried out in a T100 Thermal Cycler (BioRad, Hercules, USA).
RT-qPCRs were performed using iQ SYBR Green Supermix (BioRad, Hercules, USA). The specific primer sequences used for target genes preproghrelin and ghs-r1, and reference genes β-actin (accession number AB039726.2), elongation factor 1α (EF1α; accession number AB056104) and 18s (accession number FJ710820.1) were ordered to IDT (Ontario, Canada) and are shown in Table 1. Primers used for quantifying ghrelin receptor were designed on the exon 1 (common for ghs-r1a and ghs-r1b splicing variants) and in a region conserved between the ghs-r1a1 and ghs-r1a2 sequences, so PCR products correspond to the sum of all ghs-r1 mRNA isoforms mentioned. Genes were amplified in duplicated qPCR runs using a 96-well plate loaded with 1 μL of cDNA and 500 nM of each forward and reverse primer in a final volume of 20 μL. Each PCR run included a standard curve for the corresponding gene made of two replicates of three serial dilution points and water controls to ensure that the reagents were not contaminated. RT-qPCR cycling conditions consisted of a ramp of 95°C for 5 min, 35 cycles of 95°C for 30 sec, 56.6°C/60°C (preproghrelin and ghs-r1, respectively) for 30 sec and 73°C for 30 sec, and a final step of 95°C for 10 min. A melting curve was systematically monitored (temperature gradient at 0.5°C/5 sec from 65 to 95°C) at the end of each run to confirm the specificity of the amplification reaction. In addition, PCR products were electrophoresed on a 1% agarose gel, and single bands for each gene were purified using GenElute™ Gel Extraction Kit (Sigma-Aldrich, Madrid, Spain) and sequenced (Secugen, Madrid, Spain). The efficiency of the amplification for all studied genes was around 100%. All runs were performed using a CFX Connect Real-time System (BioRad, Hercules, USA). The 2-ΔΔCt method  was used to determine the relative mRNA expression, assigning the relative value of ‘1’ to the sampling time with the lowest expression values.
Analysis of mRNA relative abundance among time points was conducted using one-way ANOVA followed by post-hoc Student-Newman-Keuls multiple comparison test. All analyses were carried out using SigmaStat 12.0 statistics package. In addition, to evaluate rhythmicity of gene expression, cosinor analysis was performed by fitting periodic sinusoidal functions to the expression values for the genes across the seven time points. The formula used was f(t) = M+Acos(tπ/12−Π), where f(t) is the gene expression level in a given time, the mesor (M) is the mean value, A is the sinusoidal amplitude of oscillation, t is time in hours and Π is the acrophase (time of peak expression). Non-linear regression allows the estimation of M, A, and Π, and their standard error (SE) . Significance of cosinor analysis was tested using the zero-amplitude test, which indicates if the sinusoidal amplitude differs from 0 with a given probability . The time series data were considered to display a significant 24-h rhythm when p<0.05 by ANOVA, and p<0.005 by the zero-amplitude test with cosinor analysis.
Brain and gastrointestinal distribution of preproghrelin, ghrelin and ghs-r1a in goldfish
The preproghrelin and ghs-r1a mRNAs were observed surrounding the nucleus of the cells (A, B, D, E in S1 Fig), while the sense riboprobes yielded no signal (C, F in S1 Fig), supporting the specificity of the obtained signal under the conditions employed. Only some blood cells show unspecific labeling in both sense and antisense riboprobes (C, F in S1 Fig). For the immunohistochemical analysis, specific signal of the peptide is observed at the cytoplasm level in the gastrointestinal tract cells (G in S1 Fig) and no staining was observed in the control sections stained only with the secondary antibody (H in S1 Fig).
Preproghrelin mRNA expression and ghrelin were found by ISH and IHC in the mucosal and submucosal layers of all the sections analyzed from the esophagus to the anterior intestine (Fig 1). Preproghrelin transcripts were widely expressed along the esophagus (Fig 1A), while in the intestinal bulb, j-loop and anterior intestine, signal of preproghrelin probes was mainly located in the most basal cells of the mucosal epithelium and in the submucosal layer, with lower expression in the muscular layer (Fig 1C, 1E and 1G). Ghrelin-like immunoreactivity (ir) was higher in cells of the submucosal layer in the esophagus, intestinal bulb and j-loop, while in the anterior intestine, it was predominant in cells of the mucosal layer although still detected in the submucosa (Fig 1B, 1D, 1F and 1H). Ghrelin-like ir was also observed in the muscular layer of the gastrointestinal tract (Fig 1D and 1H). Preproghrelin mRNA expression and ghrelin were undetected in goldfish brain neither by ISH nor by IHC.
(A, B) Esophagus. (C, D) Intestinal bulb. (E, F) j-loop. (G, H) Anterior intestine. M, mucose; Mus, muscular layer; SM, submucose. Scale bars are indicated in each image.
The goldfish brain showed widespread distribution of ghs-r1a mRNA (Figs 2–5). Expression of ghs-r1a gene within the telencephalon was found in almost all the different areas in the pallial and subpallial regions (Figs 2A, 2B, 2C and 3). In the hypothalamus, ghs-r1a expressing cells were detected in the nucleus of the posterior recess (Figs 2D and 4D), the preoptic region, specifically in the periventricular preoptic nucleus (Figs 2B and 4A), and in the nucleus of the lateral recess (Figs 2D, 4E and 4G), with the highest levels of signal observed in the anterior periventricular nucleus (homologous to the mammalian suprachiasmatic nucleus) and in the magnocellular area of the preoptic nucleus (Figs 2B, 2C, 4B and 4C). The ghs-r1a expression showed a specific pattern in the nucleus of the lateral recess: it was found only in the lateral area of the most anterior part (Figs 2D and 4E), but it extends to the medial area (Figs 2E and 4F) until ghs-r1a mRNA surrounds completely this nucleus in posterior sections (Figs 2F and 4G). There were ghs-r1a expressing cells in the pineal gland and in the habenular nuclei (Figs 2D, 5A and 5B). Ghs-r1a transcripts were also detected in the torus longitudinalis (Figs 2F and 5C) and the highest expression was observed in the valvula of the cerebellum of the metencephalon (Figs 2F and 5D).
The brain areas shown in the transversal sections are indicated with a schematic representation in the upper part of each image. Intensity of signal is represented by density of red dots. Dc, central portion of the dorsal telencephalon; Dd, dorsal portion of the dorsal telencephalon; Dl, lateral portion of the dorsal telencephalon; Dld, dorsal part of the lateral portion of the dorsal telencephalon; Dlv ventral part of the lateral portion of the dorsal telencephalon; Dm, medial portion of the dorsal telencephalon; NAPv, anterior periventricular nucleus; NH habenular nucleus; NPO, preoptic nucleus; NPP, periventricular preoptic nucleus; NRL, lateral recess nucleus; NRP, posterior recess nucleus; OT, optic tract; P, pineal; TL, torus longitudinalis; Vc, valvula of the cerebellum; Vd, dorsal portion of the ventral telencephalon; Vl, lateral portion of the ventral telencephalon; Vp, postcommissural portion of the ventral telencephalon; Vs, supracommisural portion of the ventral telencephalon; Vv, ventral portion of the ventral telencephalon.
(A, B) Overview of telencephalon. (C) Lateral portion of the dorsal telencephalon. (D) Ventral portion of the ventral telencephalon (arrowheads indicate riboprobe signaling). (E) Example of telencephalic nucleus surrounded by ghs-r1a mRNA riboprobe (arrowhead). Dc, central portion of the dorsal telencephalon; Dd, dorsal portion of the dorsal telencephalon; Dl, lateral portion of the dorsal telencephalon; Dm, medial portion of the dorsal telencephalon; Vd, dorsal portion of the ventral telencephalon; Vv, ventral portion of the ventral telencephalon. Scale bars are indicated in each image.
(A) Periventricular preoptic nucleus. (B) Preoptic recess. (C) Preoptic nucleus. (D) Posterior recess nucleus. E, F, G. Lateral recess nucleus. NAPv, anterior periventricular nucleus; NPO, preoptic nucleus; NPP, periventricular preoptic nucleus; NRL, lateral recess nucleus; NRP, posterior recess nucleus; OT, optic tract. Scale bars are indicated in each image.
NH habenular nucleus; P, pineal; TL, torus longitudinalis; Vc, valvula cerebelli. Scale bars are indicated in each image.
The ghs-r1a expression along the gastrointestinal tract of goldfish is shown in Fig 6. A wide distribution was observed in esophagus (Fig 6A), whereas in the intestinal bulb (Fig 6B), j-loop (Fig 6C) and anterior intestine (Fig 6D), ghs-r1a expression was observed only in most apical cells of the mucosal epithelium and in the submucosal layer.
Daily expression of preproghrelin and ghs-r1 ghrelin receptor in goldfish
The daily pattern of preproghrelin expression in goldfish forebrain, hypothalamus, hindbrain, pituitary and gastrointestinal tract during a 12L:12D photocycle is shown in Fig 7. Statistical analysis (ANOVA and cosinor) showed that preproghrelin transcripts displayed significant rhythmic oscillations as a function of the 24-h cycle in hypothalamus (Fig 7B), pituitary (Fig 7D) and gastrointestinal tract (Fig 7E), with acrophases at nighttime. The acrophases in hypothalamus and gastrointestinal tract were advanced around 3–4 h as compared to pituitary (Table 2). The amplitude of preproghrelin daily rhythm was 3-fold higher in gastrointestinal tract compared to the hypothalamus and pituitary. No significant differences throughout the 24-h cycle were detected in preproghrelin expression in the forebrain and hindbrain.
Relative mRNA amounts were quantified by RT-qPCR. Data are expressed as mean ± SEM (n = 6/time point). The grey area indicates the dark phase of the daily photocycle, and the arrow indicates the scheduled feeding time (ZT-4). Dashed lines represent the periodic sinusoidal functions determined by the cosinor analysis when a significant rhythm was detected. Different letters indicate significant differences by ANOVA and post-hoc SNK test (p<0.05).
The quantitative analysis of ghs-r1 daily expression demonstrates the existence of a 24-h rhythmic profile in the hypothalamus and pituitary, with higher abundance of transcripts during the dark phase of the daily photocycle (Fig 8). In both tissues, the acrophase and amplitude of ghs-r1 daily rhythms were similar to those found for preproghrelin (Table 2). Expression of ghs-r1 was not significantly modified throughout the 24-h cycle in goldfish forebrain, hindbrain and gastrointestinal tract.
Relative mRNA amounts were quantified by RT-qPCR. Data are expressed as mean ± SEM (n = 6/time point). The grey area indicates the dark phase of the daily photocycle, and the arrow indicates the scheduled feeding time (ZT-4). Dashed lines represent the periodic sinusoidal functions determined by the cosinor analysis when a significant rhythm was detected. Different letters indicate significant differences by ANOVA and post-hoc SNK test (p<0.05).
The ghrelinergic system is a key regulator of numerous physiological processes, particularly metabolism and reproduction in fish. The brain and gastrointestinal tract are sources of ghrelin, and are major contributors to the endocrine regulation of both, metabolism and reproduction. However, two critically missing information on ghrelin biology are cellular localization of ghrelin transcripts and ghrelin, and the circadian profile of ghrelin in fish tissues. The novel results of this research address both of these paucities in fish ghrelin literature. This study shows for the first time the detailed neuroanatomical distribution of ghrelin and ghs-r1 in a non-mammal, and the existence of a daily rhythmic expression of preproghrelin and ghs-r1 in the hypothalamus, pituitary, and gastrointestinal tract of goldfish.
Several studies using RT-PCR reported a wide expression of preproghrelin mRNA in the brain of fish [5–7] and mammals , but its anatomical distribution in discrete encephalic areas remains to be described. Present data support such a broad expression of preproghrelin in the goldfish brain by RT-qPCR analyses, but it was undetectable by in situ hybridization and immunohistochemical analysis. A possible explanation for these discrepancies might lie on the different age and body weight of fish used for both studies. Thus, it is possible that ghrelin transcript levels in small and immature fish are considerably lower than in big and mature fish, and so be undetected by less sensitive techniques such as ISH and IHC. Furthermore, this is in contrast to a recent study using imaging techniques in which preproghrelin was detected in goldfish hypothalamus  by IHC. Again, the reproductive stage and metabolic status of fish used in both experiments were significantly different (1.5–2.5 g vs 10–20 g) and so might be influencing in the different results observed in one study and the other. This may indicate that detecting ghrelin transcripts in goldfish brain using imaging techniques requires delicate refinement of the techniques used, including higher concentrations of probes/antibodies, longer incubation times, etc.
In the gastrointestinal tract, preproghrelin mRNA expression and ghrelin are found by in situ hybridization and immunohistochemistry, respectively, with a similar intense signal in all the studied sections. The strong signal detected for preproghrelin mRNA expression and ghrelin-like immunoreactivity in proximal sections of the gastrointestinal tract is in accordance with previous studies on the enteric location of this hormone in other vertebrates, where ghrelin was predominantly detected in the anterior part of the gastrointestinal tract. In this respect, ghrelin immunoreactivity decreased from the small to the large intestine was observed in rodents , and several studies have reported the stomach as the portion of the intestine with more ghrelin-immunoreactive cells in mammals [1,9,11,12]. Also, in chickens, ghrelin immunoreactive cells were found in the proventriculus and in the small intestine . Finally, in fish, ghrelin immunostaining was predominantly found in the stomach of rainbow trout  and sea bass , and in the proximal intestine of goldfish . Then, the location of this peptide in the gastrointestinal tract is highly conserved through phylogeny. Furthermore, results from our study show that ghrelin-like immunoreactivity within the gastrointestinal sections is most abundant in the mucosa, which is consistent with previous observations in both mammals [9–12] and fish [14–16]. This observation, together with the fact that preproghrelin mRNA was found in the same locations than the peptide, support the mucosal cells as the primary synthesizing site of ghrelin in goldfish.
A second integral component in the ghrelinergic system is its receptor. Our results show that ghs-r1a gene is widely expressed in the goldfish brain, from telencephalon to the cerebellum. This is consistent with a previous study describing the expression of the same gene in goldfish brain areas by RT-q PCR, in which expression of ghs-r1a1 and ghs-r1a2 was predominantly observed in telencephalon, diencephalon and vagal lobe . In addition, rich expression of ghs-r1a is detected in the present study in specific hypothalamic nucleus, such as the lateral recess nucleus, in agreement with the well known orexigenic role of the ghrelinergic system. An interesting observation derived from the present study is that encephalic areas where ghs-r1a expression is predominant are known to contain cells that also express other appetite-regulating hormones. For instance, all hypothalamic nuclei expressing ghs-r1a, and even some extrahypothalamic locations where ghs-r1 was found, including the valvula cerebelli, the habenula, the pineal gland and the torus longitudinalis, all have been previously related with the orexinergic system in zebrafish [45,46]. Similarly, we report here an important expression of ghs-r1a in the periventricular preoptic nucleus, the anterior part of the periventricular nucleus, the preoptic nucleus and the nucleus of the lateral recess, areas that take part of the NPY system in the European seabass  and goldfish . Indeed, a colocalization of the ghrelin receptor and NPY in the hypothalamic arcuate nucleus of the rat, a key nucleus involved in food intake regulation, has been previously reported . This co-expression of both ghrelin receptor and appetite-regulating neuropeptides in feeding-related specific brain areas, as well as the reported physiological interactions among the ghrelinergic system and other orexigenic agents [49–51], support the cross-talking between ghrelin and other orexigenic agents in the feeding regulation.
In the gastrointestinal tract, ghs-r1a gene was found mainly in the mucosal epithelium of all the areas studied, matching the expression of preproghrelin gene. However, interestingly, the expression of the receptor was highest in the most apical cells of mucosal folds, while highest levels of the prepropeptide were observed in the most basal cells of this epithelium. Ghrelin receptor immunoreactivity in the gastrointestinal tract has only been reported in zebrafish by Olsson and co-workers , who described the presence of the receptor in numerous endocrine cells of the mucosa and in the muscle layers along the entire intestine. This important presence of ghrelin receptors in the muscle layers of the zebrafish gut was suggested to be related with motility functions of ghrelin in this teleost . In the present study, ghs-r1a expressing cells in the muscle layer were only observed in the esophagus, but not in the intestinal bulb, j-loop and anterior intestine. In fact, it is to note that ghrelin was ineffective on gut motility in rainbow trout and goldfish , which is consistent with present results on the absence of ghrelin receptors in the muscle layers of goldfish gastrointestinal tract, but in contrast with results on zebrafish . Together, here we show the brain-gut mapping of ghrelin and its receptor, two main components of the ghrelinergic system, in goldfish. GOAT, the third peptide in this hormonal system, is yet to be identified in goldfish and future studies warrant its localization.
Previous research  shows ghrelin is a multifunctional peptide in fish, and our current results show extensive, yet cell specific localization of ghrelin and its receptor in goldfish tissues. Is there a daily pattern of expression for ghrelin and GHS-R in goldfish? In fact, recent studies in this teleost show that some food intake regulatory hormones, such as NPY  and leptin , display daily oscillations in response to the 24-h light/dark cycle, suggesting a relationship between the orexigenic system and circadian organization. Such a relationship between ghrelin and the circadian system has been pointed out in mammals, where the stomach ghrelin-secreting cells were described to contain the machinery that constitutes a food entrainable oscillator . However, no studies to date have analyzed daily changes in expression of preproghelin and its receptors in any vertebrate group. Our results demonstrate daily rhythms of preproghrelin and ghs-r1 expression in hypothalamus and pituitary of goldfish maintained under scheduled photoperiod and feeding regime. Considering the key role played by the hypothalamus in the functional organization of the circadian system , the existence of daily rhythms in the hypothalamic ghrelinergic system supports such interplay between ghrelin and the circadian system. Moreover, it is important to note that a high expression of ghs-r1a was detected in specific hypothalamic nucleus related with the circadian organization, such as the anterior periventricular nucleus, which is homologous to the mammalian suprachiasmatic nucleus, the master clock regulating circadian functions in these vertebrates . In the periphery, the daily expression pattern found for preproghrelin, but not for ghs-r1 in the gastrointestinal tract, might suggest that the daily regulation of ghrelin-related genes in peripheral organs is exerted only on bioactive peptide synthesis, without effect on its receptors. It is also interesting that the amplitude of the preproghrelin expression rhythm is nearly 3-fold higher in the gastrointestinal tract compared to hypothalamus and pituitary. This strength in rhythmicity reinforces the proposal that ghrelin might be an important output of the intestinal oscillator.
Present results revealed a major finding regarding the nocturnal acrophase of the preproghrelin and ghs-r1 expression rhythms in both central and peripheral locations. The biological significance of preproghrelin and ghs-r1 transcripts peaking during the night remains to be elucidated, although it could be related with any of the wide variety of physiological functions that ghrelin is known to exert apart from feeding regulation in non-mammalian vertebrates, i.e. growth, reproduction, immunity . Interestingly, two main observations are to note for a possible crosstalking between the ghrelinergic and the circadian systems. On one hand, the 24-h ghrelinergic expression profile overlaps the one described for melatonin, a key component of the circadian system in vertebrates . On the other, the rhythms are also in phase with the daily rhythms of the negative loop clock genes of the clock molecular machinery (gper1a, gper1b, gper3 and gcry3 in the hypothalamus [58,59] and gut [60,61] of goldfish), and are in antiphase with the genes of the positive loop (clock1a and bmal1 in rainbow trout  and senegale sole ). Aditionally, ghrelin was found to induce the expression of some per and cry genes, but not bmal1a, in goldfish central and peripheral tissues . The possible connection between the ghrelinergic system with negative elements of the circadian molecular machinery, such as per, is also supported by the similar anatomical distribution reported in present study for ghs-r1a gene in brain, and the one previously reported for gper1b gene in goldfish maintained under the same environmental conditions . All these parallelisms clearly suggest a connection between the ghrelinergic system and clock genes expression, although further studies are required to demonstrate the specific implication of ghrelin in this function.
In summary, present results demonstrate that preproghrelin, ghrelin and ghrelin receptor ghs-r1a are widely expressed in the goldfish brain and gastrointestinal tract, and show for the first time a rhythmical pattern of expression in hypothalamus, pituitary and gastrointestinal tract of goldfish. These rhythmic patterns indicate an important connection between the ghrelinergic system and the circadian system of teleosts, and suggest that ghrelin might be acting alternatively as an input/output of the food entrainable oscillator.
S1 Fig. Specificity of preproghrelin and ghs-r1a mRNA riboprobes and the antibody anti-human ghrelin.
A, B. Anterior intestine showing preproghrelin antisense riboprobes signaling (arrowheads) surrounding the nucleus. C. Anterior intestine showing preproghrelin sense riboprobes staining. D, E. Telencephalon showing ghs-r1a antisense riboprobes signaling (arrowheads) staining. F. Telencephalon showing ghs-r1a sense riboprobes staining. G. Antibody anti- human ghrelin cytoplasmic signal (arrowheads) in the anterior intestine. H. Control anterior intestine without the primary antibody, incubated only with the secondary one. #: Blood cells with unspecific staining.
Conceived and designed the experiments: ASB AMB SU OK ALA AIV EI MJD. Performed the experiments: ASB AMB SU OK MMG JIB. Analyzed the data: ASB AMB SU OK MMG JIB. Contributed reagents/materials/analysis tools: SU OK MJD. Wrote the paper: ASB AMB SU ALAG AIV EI MJD.
- 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: 656–660. pmid:10604470
- 2. Kojima M, Kangawa K. Ghrelin: structure and function. Physiol Rev. 2005; 85: 495–522. pmid:15788704
- 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: 2988–2991. pmid:12050285
- 4. Ghelardoni S, Carnicelli V, Frascarelli S, Ronca-Testoni S, Zucchi R. Ghrelin tissue distribution: comparison between gene and protein expression. J Endocrinol Invest. 2006; 29: 115–121. pmid:16610236
- 5. Unniappan S, Lin X, Cervini L, Rivier J, Kaiya H, Kangawa K, et al. Goldfish ghrelin: molecular characterization of the complementary deoxyribonucleic acid, partial gene structure and evidence for its stimulatory role in food intake. Endocrinology. 2002; 143: 4143–4146. pmid:12239128
- 6. Kaiya H, Kojima M, Hosoda H, Moriyama S, Takahashi A, Kawauchi H, et al. Peptide purification, complementary deoxyribonucleic acid (DNA) and genomic DNA cloning, and functional characterization of ghrelin in rainbow trout. Endocrinology. 2003; 144: 5215–5226. pmid:12970156
- 7. Feng K, Zhang G-R, Wei K-J, Xiong B-X. Molecular cloning, tissue distribution, and ontogenetic expression of ghrelin and regulation of expression by fasting and refeeding in the grass carp (Ctenopharyngodon idellus). J Exp Zool. 2013; 319: 202–212.
- 8. Cowley MA, Smith RG, Diano S, Tschöp M, Pronchuk N, Grove KL, et al. The distribution and mechanism of action of ghrelin in the CNS demonstrates a novel hypothalamic circuit regulating energy homeostasis. Neuron. 2003; 37: 649–661. pmid:12597862
- 9. Date Y, Kojima M, Hosoda H, Sawaguchi A, Mondal MS, Suganuma T, et al. Ghrelin, a novel growth hormone-releasing acylated peptide, is synthesized in a distinct endocrine cell type in the gastrointestinal tracts of rats and humans. Endocrinology. 2000; 141: 4255–4261. pmid:11089560
- 10. Rindi G, Necchi V, Savio A, Torsello A, Zoli M, Locatelli V, et al. Characterisation of gastric ghrelin cells in man and other mammals: studies in adult and fetal tissues. Histochem Cell Biol. 2002; 117: 511–519. pmid:12107501
- 11. Hayashida T, Murakami K, Mogi K, Nishihara M, Nakazato M, Mondal MS, et al. Ghrelin in domestic animals: distribution in stomach and its possible role. Domest Anim Endocrinol. 2001; 21: 17–24. pmid:11524171
- 12. Yabuki A, Ojima T, Kojima M, Nishi Y, Mifune H, Matsumoto M, et al. Characterization and species differences in gastric ghrelin cells from mice, rats and hamsters. J Anat. 2004; 205: 239–246. pmid:15379929
- 13. Kerbel B, Unniappan S. Nesfatin-1 suppresses energy intake, co-localises ghrelin in the brain and gut, and alters ghrelin, cholecystokinin and orexin mRNA expression in goldfish. J Neuroendocrinol. 2012; 24: 366–377. pmid:22023656
- 14. Arcamone N, Neglia S, Gargiulo G, Esposito V, Varricchio E, Battaglini P, et al. Distribution of ghrelin peptide in the gastrointestinal tract of stomachless and stomach-containing teleosts. Microsc Res Tech. 2009; 72: 525–533. pmid:19322896
- 15. Olsson C, Holbrook JD, Bompadre G, Jönsson E, Hoyle CHV, Sanger GJ, et al. Identification of genes for the ghrelin and motilin receptors and a novel related gene in fish, and stimulation of intestinal motility in zebrafish (Danio rerio) by ghrelin and motilin. Gen Comp Endocrinol. 2008; 155: 217–226. pmid:17582410
- 16. Sakata I, Mori T, Kaiya H, Yamazaki M, Kangawa K, Inoue K, et al. Localization of ghrelin-producing cells in the stomach of the rainbow trout (Oncorhynchus mykiss). Zool. Sci. 2004; 21: 757–762. pmid:15277719
- 17. Abizaid A, Horvath TL. Ghrelin and the central regulation of feeding and energy balance. Indian J Endocrinol Metab. 2012; 16: S617–626. pmid:23565498
- 18. Depoortere I. Targeting the ghrelin receptor to regulate food intake. Regul Pept. 2009; 156: 13–23. pmid:19362579
- 19. Sato T, Nakamura Y, Shiimura Y, Ohgusu H, Kangawa K, Kojima M. Structure, regulation and function of ghrelin. J Biochem. 2012; 151: 119–128. pmid:22041973
- 20. Delporte C. Structure and physiological actions of ghrelin. Scientifica. 2013; 2013: 1–25.
- 21. Gutiérrez JA, Solenberg PJ, Perkins DR, Willency JA, Knierman MD, Jin Z, et al. Ghrelin octanoylation mediated by an orphan lipid transferase. Proc Natl Acad Sci USA. 2008; 105: 6320–6325. pmid:18443287
- 22. Yang J, Brown MS, Liang G, Grishin NV, Goldstein JL. Identification of the acyltransferase that octanoylates ghrelin, an appetite-stimulating peptide hormone. Cell. 2008; 132: 387–396. pmid:18267071
- 23. Kaiya H, Miura T, Matsuda K, Miyazato M, Kangawa K. Two functional growth hormone secretagogue receptor (ghrelin receptor) type 1a and 2a in goldfish, Carassius auratus. Mol Cell Endocrinol. 2010; 327: 25–39. pmid:20558240
- 24. Kaiya H, Kangawa K, Miyazato M. Ghrelin receptors in non-mammalian vertebrates. Front Endocrinol. 2013; 4:1–16.
- 25. Yin Y, Li Y, Zhang W. The growth hormone secretagogue receptor: Its intracellular signaling and regulation. Int J Mol Sci. 2014; 15: 4837–4855. pmid:24651458
- 26. Zigman JM, Jones JE, Lee CE, Saper CB, Elmquist JK. Expression of ghrelin receptor mRNA in the rat and the mouse brain. J Comp Neurol. 2006; 494: 528–548. pmid:16320257
- 27. Tannenbaum GS, Lapointe M, Beaudet A, Howard AD. Expression of growth hormone secretagogue-receptors by growth hormone-releasing hormone neurons in the mediobasal hypothalamus. Endocrinology. 1998; 139: 4420–4423. pmid:9751527
- 28. Willesen MG, Kristensen P, Rømer J. Co-localization of growth hormone secretagogue receptor and NPY mRNA in the arcuate nucleus of the rat. Neuroendocrinology. 1999; 70: 306–316. pmid:10567856
- 29. Hoskins LJ, Volkoff H. Daily patterns of mRNA expression of two core circadian regulatory proteins, Clock2 and Per1, and two appetite-regulating peptides, OX and NPY, in goldfish (Carassius auratus). Comp Biochem Physiol A Mol Integr Physiol. 2012; 163: 127–136. pmid:22643337
- 30. Tinoco AB, Nisembaum LG, de Pedro N, Delgado MJ, Isorna E. Leptin expression is rhythmic in brain and liver of goldfish (Carassius auratus). Role of feeding time. Gen Comp Endocrinol. 2014; 204: 239–247. pmid:24932715
- 31. Yildiz BO, Suchard MA, Wong M-L, McCann SM, Licinio J. Alterations in the dynamics of circulating ghrelin, adiponectin, and leptin in human obesity. Proc Natl Acad Sci USA. 2004; 101: 10434–10439. pmid:15231997
- 32. Volkoff H, Canosa LF, Unniappan S, Cerdá-Reverter JM, Bernier NJ, Kelly SP, et al. Neuropeptides and the control of food intake in fish. Gen Comp Endocrinol. 2005; 142: 3–19. pmid:15862543
- 33. Lin X, Volkoff H, Narnaware Y, Bernier NJ, Peyon P, Peter RE. Brain regulation of feeding behavior and food intake in fish. Comp Biochem Physiol A Mol Integr Physiol. 2000; 126: 415–434. pmid:10989336
- 34. Unniappan S, Peter RE. In vitro and in vivo effects of ghrelin on luteinizing hormone and growth hormone release in goldfish. Amer J Physiol. Regul Integr Comp Physiol. 2004; 286: R1093–R1101.
- 35. Matsuda K, Miura T, Kaiya H, Maruyama K, Shimakura S- I, Uchiyama M, et al. Regulation of food intake by acyl and des-acyl ghrelins in the goldfish. Peptides. 2006; 27: 2321–2325. pmid:16687192
- 36. Matsuda K, Miura T, Kaiya H, Maruyama K, Uchiyama M, Kangawa K, et al. Stimulatory effect of n-octanoylated ghrelin on locomotor activity in the goldfish, Carassius auratus. Peptides. 2006; 27: 1335–1340. pmid:16297501
- 37. Nisembaum LG, de Pedro N, Delgado MJ, Isorna E. Crosstalking between the “gut-brain” hormone ghrelin and the circadian system in the goldfish. Effects on clock gene expression and food anticipatory activity. Gen Comp Endocrinol. 2014; 205: 287–295. pmid:24681192
- 38. Escobar S, Servili A, Espigares F, Gueguen M-M, Brocal I, Felip A, et al. Expression of kisspeptins and kiss receptors suggests a large range of functions for kisspeptin systems in the brain of the European sea bass. PLoS ONE. 2013; 8: e70177. pmid:23894610
- 39. Diotel N, Servili A, Gueguen M-M, Mironov S, Pellegrini E, Vaillant C, et al. Nuclear progesterone receptors are up-regulated by estrogens in neurons and radial glial progenitors in the brain of zebrafish. PLoS ONE. 2011; 6: e28375. pmid:22140581
- 40. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. 2001; 25: 402–408.
- 41. Delgado MJ, Alonso-Gómez AL, Gancedo B, de Pedro N, Valenciano AI, Alonso-Bedate M. Serotonin N-acetyltransferase (NAT) activity and melatonin levels in the frog retina are not correlated during the seasonal cycle. Gen Comp Endocrinol. 1993; 92: 143–150. pmid:8282167
- 42. Koukkari W, Sothern R. Chronobiometry: analyzing for rhythms. Introducing Biological Rhythms. New York: Springer Science; 2006. pp. 577–602.
- 43. Peter RE, Gill VE. A stereotaxic atlas and technique for forebrain nuclei of the goldfish, Carassius auratus. J Comp Neurol. 1975; 159: 69–101. pmid:1088950
- 44. Neglia S, Arcamone N, Esposito V, Gargiulo G, de Girolamo P. Presence and distribution of ghrelin-immunopositive cells in the chicken gastrointestinal tract. Acta Histochem. 2005; 107: 3–9. pmid:15866281
- 45. Kaslin J, Nystedt JM, Ostergård M, Peitsaro N, Panula P. The orexin/hypocretin system in zebrafish is connected to the aminergic and cholinergic systems. J Neurosci. 2004; 24: 2678–2689. pmid:15028760
- 46. Panula P. Hypocretin/orexin in fish physiology with emphasis on zebrafish. Acta Physiol 2010; 198: 381–386.
- 47. Cerdá-Reverter JM, Anglade I, Martínez-Rodríguez G, Mazurais D, Muñoz-Cueto JA, Carrillo M, et al. Characterization of neuropeptide Y expression in the brain of a perciform fish, the sea bass (Dicentrarchus labrax). J Chem Neuroanat. 2000; 19: 197–210. pmid:11036237
- 48. Kah O, Pontet A, Danger JM, Dubourg P, Pelletier G, Vaudry H, et al. Characterization, cerebral distribution and gonadotropin release activity of neuropeptide Y (NPY) in the goldfish. Fish Physiol Biochem. 1989; 7: 69–76. pmid:24221756
- 49. Miura T, Maruyama K, Shimakura S-I, Kaiya H, Uchiyama M, Kangawa K, et al. Regulation of food intake in the goldfish by interaction between ghrelin and orexin. Peptides. 2007; 28: 1207–1213. pmid:17481778
- 50. Guan J-L, Wang Q-P, Kageyama H, Takenoya F, Kita T, Matsuoka T, et al. Synaptic interactions between ghrelin- and neuropeptide Y-containing neurons in the rat arcuate nucleus. Peptides. 2003; 24: 1921–1928. pmid:15127943
- 51. Miura T, Maruyama K, Shimakura S- I, Kaiya H, Uchiyama M, Kangawa K, et al. Neuropeptide Y mediates ghrelin-induced feeding in the goldfish, Carassius auratus. Neurosci Lett. 2006; 407: 279–283. pmid:16979293
- 52. Kitazawa T, Itoh K, Yaosaka N, Maruyama K, Matsuda K, Teraoka H et al. Ghrelin does not affect gastrointestinal contractility in rainbow trout and goldfish in vitro. Gen Comp Endocrinol. 2012: 178, 539–545. pmid:22776445
- 53. Kaiya H, Miyazato M, Kangawa K, Peter RE, Unniappan S. Ghrelin: a multifunctional hormone in non-mammalian vertebrates. Comp Biochem Physiol A 2008; 149: 109–128.
- 54. LeSauter J, Hoque N, Weintraub M, Pfaff DW, Silver R. Stomach ghrelin-secreting cells as food-entrainable circadian clocks. Proc Natl Acad Sci USA. 2009; 106: 13582–13587. pmid:19633195
- 55. Dibner C, Schibler U, Albrecht U. The mammalian circadian timing system: organization and coordination of central and peripheral clocks. Annu Rev Physiol. 2010; 72: 517–549. pmid:20148687
- 56. Tsang AH, Barclay JL, Oster H. Interactions between endocrine and circadian systems. J Mol Endocrinol. 2013; 52: R1–R16. pmid:23997239
- 57. Falcón J, Migaud H, Muñoz-Cueto JA, Carrillo M. Current knowledge on the melatonin system in teleost fish. Gen Comp Endocrinol. 2010; 165: 469–482. pmid:19409900
- 58. Feliciano A, Vivas Y, de Pedro N, Delgado MJ, Velarde E, Isorna E. Feeding time synchronizes clock gene rhythmic expression in brain and liver of goldfish (Carassius auratus). J Biol Rhythms. 2011; 26: 24–33. pmid:21252363
- 59. Sánchez-Bretaño A, Gueguen M-M, Cano-Nicolau J, Kah O, Alonso-Gómez AL, Delgado MJ, et al. Anatomical distribution and daily profile of gper1b gene expression in brain and peripheral structures of goldfish (Carassius auratus). Chronobiol Int. 2015; 32: 889–902. pmid:26171989
- 60. Velarde E, Haque R, Iuvone PM, Azpeleta C, Alonso-Gómez AL, Delgado MJ. Circadian clock genes of goldfish, Carassius auratus: cDNA cloning and rhythmic expression of period and cryptochrome transcripts in retina, liver, and gut. J Biol Rhythms. 2009; 24: 104–113. pmid:19346448
- 61. Nisembaum LG, Velarde E, Tinoco AB, Azpeleta C, de Pedro N, Alonso-Gómez AL, et al. Light-dark cycle and feeding time differentially entrains the gut molecular clock of the goldfish (Carassius auratus). Chronobiol Int. 2012; 29: 665–673. pmid:22734567
- 62. López-Patiño MAL, Rodríguez-Illamola A, Conde-Sieira M, Soengas JL, Míguez JM. Daily rhythmic expression patterns of clock1a, bmal1, and per1 genes in retina and hypothalamus of the rainbow trout, Oncorhynchus mykiss. Chronobiol Int. 2011; 28: 381–389. pmid:21721853
- 63. Martín-Robles AJ, Whitmore D, Sánchez-Vázquez FJ, Pendón C, Muñoz-Cueto JA. Cloning, tissue expression pattern and daily rhythms of Period1, Period2, and Clock transcripts in the flatfish Senegalese sole, Solea senegalensis. J Comp Physiol B. 2012; 182: 673–685. pmid:22373774