Bertille Debris

Degree project for Master of Science (Two Year) in

Biology

Zoophysiology 60 hec Spring 2013 - Autumn 2013

Department of Biological and Environmental Sciences

University of Gothenburg

Examiner: Michael L. Axelsson

Department of Biological and Environmental Sciences

University of Gothenburg

Supervisor: Elisabeth Jönsson Bergman

Department of Biological and Environmental Sciences

University of Gothenburg

The effect of ghrelin on the gene expression of

appetite regulatory neuropeptides and leptin, and

on plasma levels of growth regulatory hormones

in rainbow trout (Oncorhynchus mykiss)

1

Table of contents

ABSTRACT ............................................................................................................................................ 1

1. INTRODUCTION ............................................................................................................................... 3

1.1. APPETITE REGULATION ............................................................................................................. 3

1.2. GHRELIN ........................................................................................................................................ 4

1.2.1 Physiological functions of ghrelin ............................................................................................. 5

1.2.2 Ghrelin role in appetite control in fish ...................................................................................... 7

1.3. AIM OF THE STUDY ..................................................................................................................... 7

2. MATERIALS AND METHOD .......................................................................................................... 8

2.1. EXPERIMENTAL ANIMALS ........................................................................................................ 8

2.2. GHRELIN IMPLANTS ................................................................................................................... 8

2.3. EXPERIMENTAL PROCEDURE ................................................................................................... 8

2.4. TISSUE SAMPLING ....................................................................................................................... 9

2.5. TISSUE ANALYSIS ........................................................................................................................ 9

2.5.1. Gene expression ....................................................................................................................... 9

2.5.2. Plasma analysis ...................................................................................................................... 13

2.6. STATISTICAL ANALYSIS .......................................................................................................... 15

3. RESULTS .......................................................................................................................................... 16

3.1. GHRELIN EFFECT ON GENE EXPRESSION ........................................................................... 16

3.2. GHRELIN EFFECT ON GH AND IGF-I PLASMA LEVELS ..................................................... 18

3.3. GHRELIN EFFECT ON GROWTH .............................................................................................. 19

4. DISCUSSION ................................................................................................................................... 19

5. REFERENCES .................................................................................................................................. 23

2

Abstract

Ghrelin, a peptide hormone produced in the stomach, is known for its orexigenic effects in

most species. But in rainbow trout (Oncorhynchus mykiss), ghrelin acts as an anorexigenic

factor. Ghrelin is also believed to be involved in other mechanisms such as growth through

the stimulation of growth hormone (GH) secretion, along with appetite and metabolism.

In the present work, the effect of ghrelin on central neuropeptides gene expression (NPY,

POMC, CART, CRH and CCK) was measured, as well as the interaction of ghrelin with other

hormones involved in appetite regulation (leptin mRNA expression in liver) and growth

(plasma GH and IGF-I levels). Juvenile rainbow trout were implanted with ghrelin or control

pellets for 2 or 10 days. Gene expression was analyzed with qPCR and plasma hormone

levels were measured with species specific RIAs. There was no significant effect of ghrelin

on gene expression but trends towards increased expression of the anorexigenic signals CCK

and CRH. Plasma GH and IGF-I level were not influenced by ghrelin treatment, nor was the

leptin expression. Overall, these findings support an anorexigenic role of ghrelin through the

CRH pathway in the hypothalamus in rainbow trout.

3

1. Introduction

In all organisms energy balance is essential for survival. Energy balance consists of the

equilibrium between energy input and energy output. Energy input comes from the intake and

uptake of food. Organisms use energy for many biological processes such as growth,

reproduction, locomotion or thermoregulation. In order to maintain energy balance, food

intake in animals is regulated by physiological mechanisms where hormones play important

roles.

1.1. Appetite regulation

The main appetite control center is situated in the brain, in the hypothalamus. The

hypothalamus is situated between the thalamus and the brain stem and is composed of several

nuclei which are responsible for its different functions. The hypothalamic regions involved in

appetite regulation are the primary nuclei including three different areas. These are the arcuate

nucleus (ARH) responsible for food intake regulation as well as stimulation of hormone

secretion from the pituitary, the dorsomedial hypothalamic nucleus (DMH) involved in the

stimulation of the gastrointestinal tract, and the ventromedial hypothalamic nucleus (VMH)

which controls the sensation of satiety (see Castañeda et al. 2010, see Kojima and Kangawa

2005). In addition to its function in appetite control, the hypothalamus stimulates or inhibits

hormone secretion from the pituitary gland and controls many mechanisms such as

reproduction, growth or the immune system.

The hypothalamus can receive and integrate different signals: from other neuronal cells in the

brain, from hormones secreted by peripheral glands (e.g. ghrelin, leptin, cholecystokinin

(CCK), insulin), and from nutrients such as glucose or fatty acids. The signals can be either

stimulatory (having an orexigenic effect) or inhibitory (anorexigenic effect) on food intake.

For the orexigenic pathway, ghrelin secreted by the stomach stimulates orexigenic neurons

situated in the ARH in most species. It induces the release and/or production of neuropeptide

Y (NPY) and Agouti-related peptide (AgRP) in the paraventricular nucleus which stimulates

secondary order neurons containing melanin concentrating hormone (MCH) and orexin,

causing an increase of food intake. The anorexigenic pathway acts through the stimulation of

anorexigenic neurons in the ARH by leptin and insulin and the inhibition of orexigenic

4

neuron by leptin. The anorexigenic cells contain pro-opiomelanocortin (POMC) and cocaine

and amphetamine-regulated transcript (CART) which will then be released in the

paraventricular nucleus and increase the secretion of corticotropin releasing hormone (CRH).

CRH has an inhibitory effect on food intake (Figure 1) (see Castañeda et al. 2010, see Kojima

and Kangawa 2005).

Figure 1: Hypothalamic appetite regulation: ghrelin from the stomach stimulates orexigenic neurons in the

ARH which produce NPY and AgRP. They induce the production of MCH and orexin in the paraventricular

nucleus which leads to a stimulation of food intake. Leptin inhibits NPY and AgRP release and, with insulin,

stimulates anorexigenic neurons to release POMC and CART to increase CRH production in the

paraventricular nucleus. CRH will then decrease food intake.

1.2. Ghrelin

Ghrelin is a 14-28 amino acid hormone which was first discovered in the rat stomach after

intensive research on its receptor, the growth hormone secretagogue receptor (GHSR)

(Kojima et al. 1999). GHSR is a G protein-coupled receptor identified in 1982 by Guillemin

et al. (1982). Growth hormone (GH) release from the pituitary was found to be stimulated by

synthetic peptides binding to this GHSR. Since this receptor was activated by synthetic

peptides, it was suggested that a natural ligand should exist for this receptor. The orphan

receptor strategy was used in order to identify the natural ligand (Kojima et al. 1999). This

strategy means that a synthetic ligand is used as a positive control to build an assay for the

5

identification of a natural ligand (Pong et al. 1996). At first, research focused on the brain

since GHSR is present in the ARH and pituitary, but this approach was not successful.

Finally, ghrelin was identified from stomach extract (see review from Kojima and Kangawa

2005). The mode of action of GHSR to stimulate GH release differs from that of the GH

releasing hormone-receptor (GHRH-R) by the fact that GHSR stimulates GH release through

an augmentation in intracellular Ca2+

concentration whereas GHRH-R stimulates GH release

by increase of cAMP intracellular concentration (Figure 2) (Kojima and Kangawa 2005).

Figure 2: Stimulation of GH secretion in the pituitary gland by growth hormone-releasing hormone (GHRH)

and ghrelin and growth hormone secretagogues (GHS). GHRH from the hypothalamus binds to GHRH-R and

provokes the increase of cAMP and the release of GH. Ghrelin or synthetic GHS bind to GHS-R and stimulates

intracellular Ca2+ which leads to GH release (reproduced from Kojima and Kangawa, 2005).

Ghrelin can be found in two major forms, a biologically active form which has an N-decanoic

or an N-octanoic acid modification on its third amino-acid; and an inactive form which does

not present any modification. They are called acyl-ghrelin and des-acyl-ghrelin, respectively

(Kojima et al. 1999). Des-acyl-ghrelin does not have any effect on appetite regulation, but it

has been shown that it can bind, as well as acyl-ghrelin, to receptors in cardiomyocytes where

GHSR is not present. Besides, it can also have an effect on cell proliferation (Cassoni et al.

2004).

1.2.1 Physiological functions of ghrelin

Ghrelin and its receptor are present in many tissues, suggesting that ghrelin has many

physiological functions. Ghrelin and its receptor has been found in all vertebrate groups such

as mammals (Kojima et al. 1999), birds (Kaiya et al. 2002, Wang et al. 2009), amphibians

6

(Kaiya et al. 2001), reptiles (Kaiya et al. 2004) and also in all fish groups, for example,

goldfish (Carassius auratus) (Miura et al. 2009), rainbow trout (Oncorhynchus mykiss)

(Kaiya et al. 2003a), Atlantic cod (Gadus morhua) (Xu and Volkoff 2009), Atlantic halibut

(Hippoglossus hippoglossus) (Manning et al. 2008) and sharks (Sphyrna lewini and

Carcharhinus melanopterus) (Kawakoshi et al. 2007). Ghrelin is mainly produced in the

stomach (Ariyasu et al. 2001) but also in other areas such as the hypothalamus, pituitary,

pancreas, testis, heart, lung and kidney (De Vriese and Delporte 2008). In stomachless species

such as goldfish for example, ghrelin is mainly produced in the intestine (Miura et al. 2009).

Ghrelin can act locally in the brain and peripherally through the vagus nerve or via the blood

(see Kojima and Kangawa 2005). It is possible that ghrelin passes the blood brain barrier in

fish as seen in mammals (Banks et al. 2002, see Castañeda et al. 2010), but this has not been

studied yet. In general, very little is known about the characteristics of the blood brain barrier

in fish, but data indicate that other peptide hormones e.g. GH can pass over from the blood to

the brain (Johansson 2004). As explained above, ghrelin act as a GH releasing hormone by its

action on the GHSR receptor. Ghrelin injected both centrally and peripherally resulted in an

increase in GH release in rat (Kojima et al. 1999), humans (Peino et al. 2000), birds (Kaiya

and al. 2002), fish (Kaiya et al. 2003a; Kaiya et al. 2003b; Unniappan et al. 2002) and

amphibian (Kaiya et al. 2001). Besides its effects on GH release, ghrelin acts on appetite

regulation. In mammals, ghrelin is an orexigenic hormone which stimulates food intake by

increasing the release of orexigenic neuropeptides like NPY (Cowley et al. 2003). Plasma

ghrelin levels increase before a meal and during fasting (Cummings et al. 2001) and decrease

postprandially due to lower production by the stomach (De Vriese and Delporte 2007).

Ghrelin also has gastrointestinal functions e.g. modulating acid gastric production and gastric

motility (Masuda et al. 2000), and cardiovascular functions such as lowering blood pressure

(Nagaya et al. 2001) and improving cardiac cachexia (Nagaya and Kangawa 2003). In fish,

not only does ghrelin alter food intake, but it also has metabolic functions which are

important for energy balance as well. Indeed, in tilapia (Oreochromis mossambicus), ghrelin

treatment showed an increase of the condition factor due to higher body weight, and of liver

lipid content (Riley et al. 2005). Similar results were observed in rainbow trout (Jönsson et al.

2007).

7

1.2.2 Ghrelin role in appetite control in fish

Ghrelin has been identified in many fish species. Its number of amino acid varies from one

species to another: 27 or 28 in eel (anguilla japonica), 20 in tilapia, 14, 17, 18 or 19 in

goldfish and 21 or 24 in rainbow trout (see Kaiya et al. 2011). In teleost fish, ghrelin

stimulates food intake in goldfish and tilapia. In goldfish, ghrelin injection, both centrally and

peripherally stimulated an increase in food intake and a higher production of orexigenic

neuropeptides such as NPY and orexin (Miura et al. 2007, Unniappan et al. 2002). Unniappan

et al. (2004) found that goldfish fasted for seven days had a higher ghrelin mRNA expression

in the brain. Moreover, as in mammals, goldfish present a preprandial increase and a

postprandial decrease of plasma ghrelin levels (Unniappan et al. 2004) supporting its

orexigenic effect. In tilapia, ghrelin stimulates food intake after long-term treatment (Riley et

al. 2005) but no change in ghrelin mRNA levels were detected after a seven day fast (Parhar

et al. 2003). In rainbow trout ghrelin treatment resulted in a decrease in food intake, so ghrelin

appears to be an anorexigenic hormone in this species (Jönsson et al. 2010). In goldfish, the

possible mechanisms behind ghrelin's orexigenic effect have been studied both by

investigating the gene expression of neuropeptides involved in appetite control (Miura et al.

2006) and by blocking the receptors for these neuropeptides (Miura et al. 2007) in the brain.

But in rainbow trout, the target of action of ghrelin on the CRH pathway in the hypothalamus

has only been shown after central injection of a CRH receptor antagonist in a short term

experiment (Jönsson et al. 2010). The more long term and peripheral influence of ghrelin on

appetite regulatory neuropeptides in the hypothalamus is still unknown in rainbow trout.

1.3. Aim of the study

The aim of this study was to better understand the actions of ghrelin on food intake in

rainbow trout by determining the long term, peripheral effect of ghrelin on the gene

expression of neuropeptides regulating appetite (NPY, POMC, CART, CRH and CCK). In

addition, the effect of ghrelin on metabolic and appetite regulatory hormones GH, leptin and

IGF-I was investigated.

8

2. Materials and method

2.1. Experimental animals

The experiment was conducted at the Department of Biological and Environmental Sciences,

University of Gothenburg. Juvenile rainbow trout were bought from a local hatchery

(Vänneåns fiskodling). The fish were kept in tanks at 10°C and with aerated and recirculating

freshwater. Fish were fed with commercial feed during the 2 weeks acclimation prior to the

start of the experiment. The experiment was carried out under the ethical permit 85-2012.

2.2. Ghrelin implants

Ghrelin and control pellets were prepared in order to be implanted into the fish. The control

pellets were composed of 95% cholesterol and 5% coconut fat. For the ghrelin pellets, 76 ng

of rainbow trout ghrelin (rtghrelin) per gram fish was added. The pellets weight was 5 mg.

The ghrelin used was the rainbow trout octanoylated 23-amino acid form, synthesized by

Peptide Institute Inc, Osaka, Japan.

2.3. Experimental procedure

The fish were anesthetized with 2-phenoxyethanol (ICN Biomedicals Inc., Germany) at a

concentration of 0.4 ml 2-phenoxyethanol per 1 liter water. The fish were weighed and the

fork length was measured. A small incision was made on the left side to the peritoneal cavity

and the pellet were implanted (control n=20 and ghrelin n=20). The fish were then placed in

individual tanks for 2 days (control n=8 and ghrelin n=8) or 10 days (control n=12 and ghrelin

n=12). The tanks were filled with oxygenated and recirculating freshwater at 10°C. They were

covered with opaque plastic on the sides to minimize stress. The fish were not fed during the

experimental time.

9

2.4. Tissue sampling

After 2 or 10 days, the fish were killed by overdose of anesthetics (2-phenoxyethanol at 0.5

ml per liter water). The weight and the fork length were measured and blood sample was

taken from the caudal vein with heparinized syringes and put in eppendorf tubes on ice. The

animal was decapitated to allow bleeding from the brain region and facilitate sampling. The

brain and the hypothalamus were quickly dissected and removed and directly placed in

microtubes and put into liquid nitrogen. The liver and muscle were sampled and placed on dry

ice. The blood samples were then centrifuged for 5 minutes at 5000 rpm and the plasma

removed and placed into eppendorf tubes. All samples were stored at -80°C until analysis.

2.5. Tissue analysis

2.5.1. Gene expression

Gene expression of neuropeptides was measured from the hypothalamus and the brain. The

expressions of the following genes were measured: NPY, POMCA1, CCK, CART and CRH.

The expression of the growth hormone receptor (GHR) and leptin was measured in the liver.

Leptin expression was also measured in the brain, but not in the hypothalamus because of a

too low yield of RNA.

o RNA extraction

Samples from hypothalamus, brain and liver were processed for RNA extraction using the

RNeasy®

Plus Mini Kit (QIAGEN). Pieces of brain and liver, weighing between 20 and 30

mg were cut and directly put into homogenizing tubes (Precellys Lysing Kit, Bertin

Technologies) containing 600 µl of Buffer RLT Plus mixed with β-mercaptoethanol (β-ME).

The hypothalami were small (weight < 10 mg) therefore the whole tissue was transferred into

homogenizing tubes filled with 350 µl Buffer RLT Plus mixed with β-ME. All tissues were

put into the tubes for homogenization when still frozen to prevent deterioration during

melting. The tubes were then placed into a tissue homogenizer (Precellys® 24, Bertin

Technologies) and samples were homogenized and then centrifuged at 14.000 g during 3

minutes. All centrifugations were performed at room temperature. The supernatant was

10

transferred into a gDNA-eliminator spin column placed in a 2 ml tube and was centrifuged

again for 30 seconds at 9.900 g in order to avoid the clogging of the gDNA-eliminator spin

column from suspended material. 600 µl (for brain and liver) or 350 µl (for hypothalamus) of

70% ethanol (for hypothalamus and brain) or 50% ethanol (for liver) was added for

purification of the RNA. The solution was put into an RNeasy spin column placed in a 2 ml

tube and centrifuged for 15 seconds at 9.900 g for RNA collection into the filter. The column

was then washed several times first with 700 µl RW1 followed by 15 seconds centrifugation

at 9.900 g, then with 500 µl Buffer RPE followed by 15 seconds centrifugation at 9.900 g and

finally with 500 µl Buffer RPE and centrifuged at 9.900 g for 2 minutes in order to dry the

column before the RNA elution and make sure that no ethanol or buffer solution was still

present. RNA elution was finally done using RNase free water. RNA concentrations were

measured with Nanodrop 2000c spectrophotometer (Thermo Scientific).

o cDNA synthesis

For the RNA from the hypothalamus, the RNA concentrations were low, so it was decided to

use the Invitrogen Superscript III First-Strand-Synthesis-System (Life technologies Ltd). To

each tube 1 µl of primers (50 µM oligo(dT)20), 1 µl of 10 mM dNTP mix as well as RNA

solution and RNase free water was added. The volume of RNase free water and RNA solution

used was calculated for each sample in order to get 300 ng of RNA into a final volume of 10

µl. The tubes were incubated at 65 °C for 5 minutes, and then put on ice for at least 1 minute.

Next, 10 µl of cDNA synthesis Mix was added. This synthesis kit is composed of 2 µl of

10×RT Buffer, 4 µl of 25 mM MgCl2, 2 µl of 0.1 M DTT, 1 µl of RNase OUT (40 U/µl) and

1 µl of SuperScript III RT (200 U/µl). The tubes were incubated at 50°C for 50 minutes in a

thermal cycler (MyCyclerTM

, BIO-RAD) followed by 5 minutes at 85°C to terminate the

reactions and then placed on ice. 1 µl of RNase H was added to each tube which was then

incubated for 20 minutes at 37°C. The cDNA solution was stored at -20°C until use for qPCR.

For the RNA from brain and liver, the iScriptTM

cDNA synthesis Kit (BIO RAD) was used for

cDNA synthesis. First, the RNA solutions from the liver were diluted 1:10 because of their

high RNA concentrations. To each tube, 4 µl of iScript reaction mix and 1 µl of iScript

reverse transcriptase were added. The volume of RNase free water and RNA added was

calculated for each sample in order to get 1000 ng (for liver) or 500 ng (for brain) of RNA

into each tube. The tube were then placed into the thermal cycler and incubated for a cycle of

11

5 minutes at 25°C, then 30 minutes at 42°C and finally 5 minutes at 85°C. The samples were

kept at -20°C until use for qPCR.

o qPCR

The primers used for qPCR were bought from eurofins MWG Synthesis GmbH. Their

sequences are presented in Table 1. Efficiency tests were performed for every primer pair and

all tissues in order to test the primers concentrations and to get the efficiency value (Table 1)

and the slope needed for the calculations and to detect which cDNA concentration was the

best to use for the qPCR. The reference gene used for the three tissues was GADPH. The

genes tested were, for the hypothalamus: NPY, POMC, CART, CCK and CRH; for the brain:

NPY, POMC, CART, CCK, CRH and leptin; for the liver: GHR and leptin. The primer

concentration used was 0.5 µM for all primers except for leptin in the liver where 0.3 µM was

better. cDNA (diluted to get 10 ng for the hypothalamus, 12.5 ng for the liver and 8 ng for the

brain) was put as duplicates for each sample in a 96-well plate. Then, primers (forward and

reverse), iQTM

SYBR® Green Supermix (BIO-RAD) and RNAse free water were added into

each well according to Table 2. The plate was placed into the qPCR machine (MyQTM

single

color Real-time PCR detection system, BIO-RAD or CFX ConnectTM

Real-Time system,

BIO-RAD). The thermal cycles for the qPCR reactions were as follow: 1 cycle at 95°C for 30

seconds; 40 cycles at 95°C for 20 seconds, 59°C for 20 seconds and 72°C for 30 seconds; 61

cycles at 65°C for 30 seconds and finally kept on hold at 16°C. The optimal temperature for

the primers was 59°C.

12

Table 1: Sequence and efficiency of the primers for the measure of gene expression

sequence (5' → 3') Efficiency (%)

GADPH Hypothalamus Brain Liver

gadph-f1 CCACTCCATCTCCGTATTCC 100,5 91,1 91,3

gadph-r1 ACTTGTCTTCGTTGACTCCC

NPY

NPY-f CTCGTCTGGACCTTTATATGC 105,1 97,6 -

NPY-r GTTCATCATATCTGGACTGTG

POMC

POMC-A1-f CTCGCTGTCAAGACCTCAACTCT 105,3 98,7 -

POMC-A1-r GAGTTGGGTTGGAGATGGACCTC

CART

CART-f GAACCATGGAGAGCTCCAGG 98,1 98,1 -

CART-r GCGCACTGCTCTCCAACGT

CCK

CCKL-f TCCCAGCCACAAGATAAAGG 97,1 97,1 -

CCKL-r GATGGATTTAGTGGTGGTGC

CRH

CRF1-f TCTGCTCATTGCTTTCTTACC 81,2 86,3 -

CRF1-r AGTCGGATGTAGTATTCCTCTC

Leptin

Lep-rt-f GGTGATTAGGATCAAAAAGCTGGA - 109,2 108,3

Lep-rt-r GACGAGCAGTAGGTCCTGGTAGAA

GHR

GHR2aFwOm TGGGAAGATGAGTGCCAGACT - - 105,0

GHR2aReOm CACAAGACTACTGTCCTCTGTTGG

Table 2: Volume (in µl) of primers, Supermix, mRNAse free water and cDNA added to each well. The MyQTM

single color Real-Time PCR detection system was used to measure gene expression of GADPH, POMC and NPY from the hypothalamus with a primers concentration of 0.5 µM. CFX Connect

TM Real-Time system was used to measured gene

expression of: CART, CRH and CCK from the hypothalamus; GADPH, POMC, NPY, CART, CRH, CCK and leptin from the brain; GHR and leptin from the liver; the primers concentration for all genes was of 0.5 µM except for leptin from the liver where the primers concentration was of 0.3 µM.

MyQTM single color Real-Time PCR detection system

CFX ConnectTM Real-Time system

Final primers concentrations 0,5 µM 0,5 µM 0,3 µM

Primer (forward) 1 0,5 0,3

Primer (reverse) 1 0,5 0,3

Supermix 10 5 5

mRNAse free water 3 1,5 1,9

cDNA 5 2,5 2,5

Final volume 20 10 10

13

2.5.2. Plasma analysis

Growth hormone and IGF-I plasma levels were measured with a radioimmunoassay (RIA). In

a RIA, a labeled antigen is competing with an endogenous antigen to bind to the antibody. In

this case, the antigen is GH or IGF-I. The assay has to be conducted under a specific pH and

with specific reagents. At the start of the assay, a standard curve is elaborated to test the

specificity of the antibody with the antigen. If the specificity is high, only one standard curve

will be produced. But if the specificity is lower, there can be binding with other substances

which produces other standard curves. The label was the complex: GH-I125

for GH RIA and

IGFI-I125

for IGF-I RIA. It was made from commercial I125

(PerkinElmerTM

Life Sciences).

o GH RIA

The RIA protocol used has been created for salmon GH (Björnsson et al. 1994). The GH RIA

is conducted during 3 days. The plate set up include vials for the measure of the total

radioactivity (T) containing only the label, the non specific binding (NSB), the maximum

binding (B0), the reference plasma (RP) and the plasma blank (PB). All of these vials are

present in triplicates. For the establishment of the standard curve, a dilution series of 8

concentrations is set up in duplicates. The measure of the samples requires duplicates as well.

During the first day, all vials are set up in the plate as described previously. The standard

dilution series is made with RIA buffer at the following concentrations: 0.1, 0.39, 0.78, 1.56,

3.13, 6.25, 12.5 and 50 ng/ml. Then, standards, sample plasma, reference plasma, primary

antibody (HU-85, diluted 1:10000) and other reagents are added according to Table 3. The

primary antibody is added at the end and the vials are incubated at +4°C overnight.

Table 3: Volume in µl of reagents added to each vial during the first day of GH RIA. T: total radioactivity, NSB: non specific binding, PB: plasma blank, B0: maximum binding, RP: reference plasma.

Vial RIA buffer 1% normal

rabbit serum Standard

sample plasma

reference plasma

1° Ab

T - - - - - -

NSB 50 50 - - - -

PB - 50 - 50 - -

B0 50 - - - - 50

RP - - - - 50 50

Standard - - 50 - - 50

Sample - - - 50 - 50

14

On day 2, the pure label is diluted in order to get 4000 CPM in 50 µl. 50 µl of the diluted

label is added to all vials, including the totals which are then closed with a lid. The vials are

incubated again at +4°C overnight.

On the last day, 100 µl of the secondary antibody (R0881 anti rabbit serum, diluted 1:10 in

2% PEG) is added to all vials (except totals) and the vials are then incubated at room

temperature for 2 hours. After incubation, 100 µl of RIA buffer is added (not for the totals)

and the vials are centrifuged at +4°C and 3000 rpm for 1 hour. The supernatant is aspirated

and the radioactivity of the pellets is counted in a gamma counter (PerkinElmerTM

Life

Sciences). The GH concentrations from the samples and the standard series are calculated

using the ASSAY-ZAP program.

o IGF-I RIA

The IGF-I RIA was realized following the protocol by Shimizu et al. (2000). The assay

requires 2 days. The IGF-I RIA requires an additional step for plasma extraction from the

samples before the start of the assay. This step is performed in RIA vials. 25 µl of plasma is

placed in the vials and 100 µl of acid/ethanol solution (87.5% ethanol, 12.5% 2M HCl) is

added. The vials are incubated at room temperature for 30 minutes and then centrifuged at

3000 rpm at +4°C for 30 minutes. The supernatant is transferred into new vials and 50 µl of

Tris is added. The samples are then ready for the assay. The vials are set up on the plate the

same way as for the GH RIA, but with a dilution series composed of 12 concentrations

(instead of 8 for the GH assay). On the first day, the vials are set up on the plate and the

dilution series is established for the standard. The standard plasma is diluted into RIA buffer

to get the highest concentration of 25 ng/ml, and then a dilution series of 1:2 is made to get

the other 11 concentrations. The standards, samples, label, reagents and primary antibody

(Gro Pep IGF-I Ab, diluted 1:4000 in RIA buffer) are added to the vials according to Table 4.

The label has been diluted so that in contains 5000 CPM in 50 µl. The vials are incubated at

+4°C overnight.

During the second day, 50 µl of the secondary antibody (R0881 anti rabbit serum, diluted

1:10 in RIA buffer) and 25 µl of gammaglobuline (diluted 1:200 in RIA buffer) are added,

followed by incubation at +4°C for 30 minutes. 1 mL of 6% PEG (diluted in ddH2O at a

concentration of 2.8 mg/ml) is added, the vials are then incubated at +4°C for 10 minutes. The

15

vials are then centrifuged at for 1 hour at +4°C and 3200 rpm . After centrifugation, the

supernatant is aspirated and the pellets are counted in a gamma counter (PerkinElmerTM

Life

Sciences). The calculations of the IGF-I concentrations has been realized using the program

ASSAY-ZAP.

Table 4: Volume in µl of reagents added to the vials for the IGF-I RIA. T: total radioactivity, NSB: non specific binding, B0: maximum binding

T NSB B0 Standard

Plasma blank

sample

Standard - - - 100 - -

Standard dilution solution

- 100 100 - - -

Sample - - - - 20 20

RIA buffer - - - - 80 80

Total volume - 100 100 100 100 100

RIA buffer - 150 100 100 150 100

1° Ab - - 50 50 - 50

Label 50 50 50 50 50 50

Final volume 50 300 300 300 300 300

2.6. Statistical analysis

For the calculation of gene expression, the mRNA expression (converted copy number) has

been calculated following the formula:

10^(-CT target/Etarget)/10^(-CT ref/Eref)

CT Target represents the CT value of the target genes (NPY, POMC, CART, CCK, CRH, GHR

and leptin), Etarget is the slope of the efficiency curve for the target gene, CT ref represents the

CT value of the reference gene (GADPH) and Eref is the slope of the efficiency curve for the

reference gene.

The converted copy number of all genes as well as the GH and IGF-I concentrations has been

analysed by a 2-way ANOVA test using SPSS (IBM SPSS Statistics 21) with time (2 and 10

days) and treatment (grelin and control) as fixed factors.

16

3. Results

3.1. Ghrelin effect on gene expression

In the hypothalamus, there was no significant difference between treatment groups for the

orexigenic neuropeptide, NPY (p > 0.4, Fig. 3). For the anorexigenic peptides: POMCa1,

CART, CCK and CRH, there were no significant differences in the gene expression between

control and ghrelin-treated fish (p > 0.05 for all tests) (Fig. 4). However, for CCK and CRH

there is a trend for an interaction between treatment and time (p = 0.059 for CCK and p =

0.079 for CRH), with higher expression level in the 10 days group in ghrelin treated fish than

in control fish, while the levels are similar between treatment groups after 2 days.

Figure 3: NPY mRNA expression in the hypothalamus for control (n = 5) and ghrelin treated fish (n = 7) after 2 days treatment and control (n = 9) and ghrelin treated fish (n = 8) after 10 days treatment. Data are showed as the mean value. Error bars represent ± standard error.

17

00,20,40,60,8

11,21,41,61,8

2

2 days 10 days

mR

NA

exp

ress

ion

Time

CCK expression

control

ghrelin

0

0,05

0,1

0,15

0,2

0,25

0,3

0,35

2 days 10 days

mR

NA

exp

ress

ion

Time

CART expression

control

ghrelin

0

0,05

0,1

0,15

0,2

0,25

0,3

0,35

0,4

0,45

2 days 10 days

mR

NA

exp

ress

ion

Time

CRH expression

control

ghrelin

00,0020,0040,0060,008

0,010,0120,0140,0160,018

2 days 10 days

mR

NA

exp

ress

ion

Time

POMCa1 expression

control

ghrelin

A B

C D

Figure 4: Gene expression in the hypothalamus for control (n = 5) and ghrelin treated fish (n = 7) after 2 days treatment and control (n = 9) and ghrelin treated fish (n = 8) after 10 days treatment A. POMCa1, B. CRH, C. CCK and D. CART mRNA expression. Data are showed as the mean value. Error bars represent ± standard error.

For gene expression in the brain, there was a significant effect of time on NPY expression

with a higher expression in the 10 days groups than for the 2 days groups in both control and

ghrelin (p = 0.028) (Fig. 5). There was no effect of treatment (p > 0.100), no time effect (p >

0.100) and no interaction time-treatment (p > 0.400) for any of the anorexigenic genes in the

brain (Table 5).

GHR expression in the liver showed no significant change with time (p = 0.802) and there

was no interaction time-treatment (p = 0.563) but a trend can be seen for a treatment effect (p

= 0.095) with lower expression for ghrelin-treated fish compared to control after 10 days.

There was no significant difference in leptin mRNA expression in liver between groups (p >

0.150 for all tests) (Table 6).

18

Figure 5: NPY expression in the brain for control (n = 7) and ghrelin treated fish (n = 7) after 2 days treatment and control (n = 11) and ghrelin treated fish (n = 11) after 10 days treatment. Data are showed as the mean value. Error bars represent ± standard error. Asterisk (*) shows significant time effect (p = 0.028) with higher expression after 10 days compared to 2 days.

Table 5: Gene expression (expressed as converted copy number) of the listed neuropeptides in the brain for control (n = 7) and ghrelin treated fish (n = 7) after 2 days treatment and control (n = 11) and ghrelin treated fish (n = 11) after 10 days treatment. Data are showed as the mean value with ± standard error.

2 days 10 days

Control Ghrelin Control Ghrelin

POMC 1,19E-05 ± 0,19E-05 6,16E-05 ± 3,97E-05 4,83E-05 ± 2,52E-05 8,87E-05 ± 3,69E-05

CART 2,53E-02 ± 0,34E-02 2,64E-02 ± 0,52E-02 2,86E-02 ± 0,34E-02 2,84E-02 ± 0,37E-02

CCK 1,60E-02 ± 0,24E-02 1,34E-02 ± 0,25E-02 1,99E-02 ± 0,43E-02 1,87E-02 ± 0,24E-02

CRH 1,32E-02 ± 0,23E-02 1,24E-02 ± 0,18E-02 1,01E-02 ± 0,21E-02 1,28E-02 ± 0,19E-02

Leptin 3,82E-06 ± 1,06E-06 3,88E-06 ± 1,13E-06 4,28E-06 ± 1,17E-06 3,58E-06 ± 0,73E-06

Table 6: Gene expression for GHR and leptin in liver for control (n = 8) and ghrelin treated fish (n = 7) after 2 days treatment and control (n = 11) and ghrelin treated fish (n = 11) after 10 days treatment. Data are showed as the mean value with ± standard error.

2 days 10 days

Control Ghrelin Control Ghrelin

GHR 1,19 ± 0,14 1,31 ± 0,13 2,47 ± 0,56 1,54 ± 0,18

Leptin 0,055 ± 0,010 0,108 ± 0,047 0,046 ± 0,0106 0,057 ± 0,0163

3.2. Ghrelin effect on GH and IGF-I plasma levels

GH and IGF-I plasma level are presented in Table 7. There were no significant effects of

time, treatment or a time-treatment interaction on GH (p > 0.250). There was no treatment

19

effect or interaction time-treatment effect on IGF-I levels (p > 0.500), but there is a time

effect (p = 0.014) with higher expression after 10 days compared to 2 days.

Table 7: GH and IGF-I plasma levels (ng/ml) for control (GH n = 8; IGF-I n = 7) and ghrelin treated fish (GH n = 8; IGF-I n = 8) after 2 days treatment and control (GH n = 11; IGF-I n = 9) and ghrelin treated fish (GH n = 11; IGF-I n = 9) after 10 days treatment. Data are showed as the mean value with ± standard error. Asterisk (*) indicates a significant time effect (p = 0.014).

2 days 10 days

Control Ghrelin Control Ghrelin

GH plasma level 2,46 ± 0,38 2,82 ± 0,55 3,87 ± 0,83 2,98 ± 0,72

IGF-I plasma level (*) 11,06 ± 1,27 10,56 ± 1,95 17,71 ± 1,38 16,01 ± 3,43

3.3. Ghrelin effect on growth

There was no significant difference between the treatment groups in weight, length, condition

factor, liver somatic index and SGRw at the start and the end of the experiment (p > 0.100 for

all tests) (Table 8).

Table 8: Initial and final body weight (g) (Wi, Wf), length (cm) (Li, Lf), condition factor ((weight/length3)*100) (CFi, CFf),

liver somatic index (% wet tissue weight) (LSI) and specific growth rate for the weight (% body weight gain/day) (SGRw) for control (n = 8) and ghrelin treated fish (n = 8) after 2 days treatment and control (n = 12) and ghrelin treated fish (n = 11) after 10 days treatment. Data are showed as the mean value with ± standard error.

2 days 10 days

Control Ghrelin Control Ghrelin

Wi 26,89 ± 2,34 27,64 ± 1,95 26,02 ± 2,13 27,38 ± 2,07

Wf 26,84 ± 2,16 29,04 ± 1,82 25,53 ± 2,17 26,95 ± 2,06

Li 14,11 ± 0,52 14,13 ± 0,25 13,56 ± 0,37 13,85 ± 0,29

Lf 14,13 ± 0,52 14,14 ± 0,25 13,68 ± 0,38 13,76 ± 0,34

LSI 9,59E-03 ± 0,42E-03 8,51E-03 ± 0,58E-03 9,27E-03 ± 0,58E-03 9,95E-03 ± 0,74

CFi 0,998 ± 0,098 0,971 ± 0,031 1,024 ± 0,031 1,010 ± 0,028

CFf 0,988 ± 0,045 1,020 ± 0,029 0,977 ± 0,020 1,017 ± 0,035

SGRw 0,419 ± 1,015 0,010 ± 0,337 -0,202 ± 0,099 -0,155 ± 0,213

4. Discussion

In the present study, the ghrelin treatment did not change the expression of the orexigenic

neuropeptide, NPY, in the hypothalamus or brain, suggesting that ghrelin does not influence

NPY gene expression in rainbow trout. In goldfish, brain NPY expression was significantly

higher after central injection of ghrelin, but, similar to our study, not after peripheral

treatment (Miura et al. 2006). In brown trout (Salmo trutta), peripheral ghrelin stimulates

food intake but not NPY expression in the hypothalamus (Tinoco et al. 2014). However, in

20

rainbow trout, ghrelin is anorexigenic, so it could be expected that NPY expression is lower in

ghrelin treated fish than control. The results observed here may be explained by the fact that

peripheral ghrelin might act posttranscriptionally on NPY, for example on its release rather

than on its expression, as was suggested for goldfish (Miura et al. 2006) and brown trout

(Tinoco et al. 2014) or that peripheral and central administration of ghrelin has different

effects on NPY expression. A time effect was present for the brain NPY mRNA expression,

with higher expression after 10 days. This may be explained by a lesser stress after 10 days

than 2 days, perhaps because the fish had more time to acclimatized to their individual aquaria

and/or recover from the treatment handling. Also, the fish were not fed during the experiment,

which could explain a higher expression of orexigenic neuropeptite due to appetite increase.

Although the ghrelin treatment did not result in any statistically significant effect on the

mRNA expression for the tested anorexigenic neuropeptides in the brain or the hypothalamus,

some interesting patterns were observed. For CCK and CRH expression in the hypothalamus,

a trend can be seen where ghrelin treated fish are indicated to have a higher expression after

10 days treatment compared to control. But due to a high variance, more individuals would

have been needed in this study to achieve statistical significance. However, even if this trend

is only partly supported by the statistics, we can speculate that this study confirms the role of

ghrelin as an anorexigenic hormone in rainbow trout, and hypothesize that it acts by

stimulating neuropeptides involved in anorexigenic pathways. This hypothesis lends support

from the findings of Jönsson et al. (2010), where central injection of homologous ghrelin

decreased food intake in juvenile rainbow trout by acting on CRH. Future studies are needed

to test if also peripheral ghrelin acts on the CRH system. The present findings are in line with

the fact that ghrelin has a similar effect in birds where central injection of rat or chicken

ghrelin inhibits food intake in neonatal chicks (Furuse et al. 2001, Saito et al. 2002).

Interactions between ghrelin and leptin have been shown in other species. In mammals, where

leptin and ghrelin have opposite effects on food intake, an increase of ghrelin expression in

the stomach was coupled with a decrease of gastric leptin expression in fasted rat (Zhao et al.

2008). Salmerón et al. (Submitted article) showed that rainbow trout adipocytes exposed to

ghrelin had an increase in leptin secretion. However, our results show that leptin mRNA

expression presents no significant change in the brain or the liver in response to the ghrelin

treatment. The interaction between the two hormones might be at another level than

transcriptional, for example, it is possible than ghrelin acts on leptin secretion.

21

Although ghrelin is considered a GH secretagogue (see Kaiya et al. 2011), the effect of

ghrelin on GH appears limited in our study. GH may influence growth, which was not

affected here, as well as appetite in rainbow trout (Björnsson 1997). In the Persian sturgeon

(Acipenser persicus), a peripheral injection of ghrelin led to increased GH plasma level

(Miandare et al. 2011). Ghrelin also increases GH release in rainbow trout (Kaiya et al.

2003a), goldfish (Unniappan and Peter 2004), tilapia (Kaiya et al. 2003b), as it does in

mammals (see Kojima and Kangawa 2005). However, these studies only examined the effect

of ghrelin on plasma GH levels in the short term (hours). Long term studies are limited; in

tilapia, a 21-day ghrelin treatment had no effect on plasma GH levels (Riley et al. 2005). The

present study agrees with these more long term findings, indicating that ghrelin's effect on

plasma GH levels may be acute. Negative feedback or other physiological mechanisms may

then stabilize the circulating levels of GH. Although it did not reach a statistical difference,

there was a trend towards a reduced liver GHR mRNA expression in ghrelin treated fish after

10 days, which may indicate that ghrelin suppresses GHR expression. Future studies with

more individuals should be conducted to confirm the results. However, it may be speculated

that if the change in GH mRNA expression is translated to functional GH receptors, this

would mean that ghrelin fish are less sensitive to the action of GH and potentially would have

higher levels of GH, which was not the case here. Hence a change in GH receptor dynamics is

not a likely mechanism that explains that the expected increase in GH in ghrelin treated fish

did not occur. GH stimulates the release of IGF-I mainly from the liver to the blood. Ghrelin

treatment had no effect on plasma levels of IGF-I. This is opposite to the previous study on

tilapia where ghrelin treatment induced a reduction in IGF-I (Riley et al. 2005). IGF-I plasma

levels were generally higher after 10 days. One can speculate that this can be explained by

lower stress levels after 10 days, that the fish had more time to acclimatize to their new

aquaria and/or recover from the handling. Finally, no significant change is seen for the

weight, length, liver somatic index, condition factor, or specific growth rates for weight. This

was probably due to the fact that the fish were fasted during the experiment and that the

experimental time was too short to see any potential effects.

In conclusion, this study confirms the action of ghrelin as an anorexigenic hormone in

rainbow trout by acting on the hypothalamic CRH system as seen in birds but different from

other fish species and from mammals. It seems likely that ghrelin acts on both gene

expression as well as the release of CRH in the hypothalamus. Further studies are needed with

more individuals to see if statistical significance can be reach. Also, it would be interesting to

22

look at the effect of ghrelin on NPY and leptin release and on GH production and secretion in

the pituitary in addition to plasma levels.

23

5. References

Ariyasu H, Takaya K, Tagami T, Ogawa Y, Hosoda K, Akamizu T, Suda M, Koh T, Natsui

K, Toyooka S, Shirakami G, Usui T, Shimatsu A, Doi K, Kojima M, Kangawa K, Nakao K.

2001. Stomach is a major source of circulating ghrelin, and feeding state determines plasma

ghrelin-like immunoreactivity levels in humans. The Journal of Clinical Endocrinology and

Metabolism, 86: 4753-4758.

Banks WA, Tschöp M, Robinson SM, Heiman ML. 2002. Extent and direction of ghrelin

transport across the blood-brain barrier is determined by its unique primary structure. The

Journal of Pharmacology and Experimental Therapeutics, 302: 822-827.

Björnsson BT. 1997. The biology of salmon growth hormone: from daylight to dominance.

Fish Physiology and Biochemistry, 17: 9-24.

Björnsson BT, Taranger GL, Hansen T, Stefansson SO, Haux C. 1994. The interrelation

between photoperiod, growth hormone and sexual maturation of adult Atlantic salmon (Salmo

salar). General and Comparative Endocrinology, 93: 70-81.

Cassoni P, Ghe C, Marrocco T, Tarabra E, Allia E, Catapano F, Deghenghi R, Ghigo E,

Papotti M, Muccioli G. 2004. Expression of ghrelin and biological activity of specific

receptors for ghrelin and des-acyl ghrelin in human prostate neoplasms ans related cell lines.

European Journal of Endocrinology, 150: 173-184.

Castañeda TR, Tong J, Datta R, Culler M, Tschöp MH. 2010. Ghrelin in the regulation of

body weight and metabolism. Frontiers in Neuroendocrinology, 31: 44-60.

Cowley MA, Smith RG, Diano S, Tschöp M, Pronchuk N, Grove KL, Strasburger CJ,

Bidlingmaier M, Esterman M, Heiman ML, Garcia-Segura LM, Nillni EA, Mendez P, Low

MJ, Sotonyi P, Friedman JM, Liu H, Pinto S, Colmers WF, Cone RD, Horvath TL. 2003. The

Distribution and Mechanism of Action of Ghrelin in the CNS Demonstrates a Novel

Hypothalamic Circuit Regulating Energy Homeostasis. Neuron, 37: 649-661.

Cummings DE, Purnell JQ, Frayo RS, Schmidova K, Wisse BE, Weigle DS. 2001. A

preprandial rise in plasma ghrelin levels suggests a role in meal initiation in humans.

Diabetes, 50: 1714-1719.

De Vriese C and Delporte C. 2008. Ghrelin: A new peptide regulating growth hormone

release and food intake. The International Journal of Biochemistry and cell physiology, 40:

1420-1424.

Furuse M, Tachibana T, Ohgushi A, Ando R, Yoshimatsu T,Denbow DM. 2001.

Intracerebroventricular injection of ghrelin and growth hormone releasing factor inhibits food

intake in neonatal chicks. Neuroscience Letters, 301: 123-126.

Guillemin R, Brazeau P, Bohlen P, Esch F, Ling N and Wehrenberg WB. 1982. Grpwth

hormone-releasing factor from a human pancreatic tumor that caused acromegaly. Science,

218: 585-587.

24

Johansson V. 2004. Behavioural effects and central nervous system actions of growth

hormone in salmonids fish. PhD-thesis. University of Gothenburg.

Jönsson E, Kaiya H, Björnsson BT. 2010. Ghrelin decreases food intake in juvenile rainbow

trout (Oncorhyncus mykiss) through the central anorexigenic corticotropin-releasing factor

system. General and comparative endocrinology. 166: 39-46.

Jönsson E, Forsmann A, Einarsdottir IE, Kaiya H, Ruohonen K, Björnsson BT. 2007. Plasma

ghrelin levels in response to fasting, feeding and food composition, and effects of ghrelin on

voluntary food intake. Comparative Biochemistry and Physiology, Part A, 147: 1116-1124.

Kaiya H, Miyazato M, Kangawa K. 2011. Recent advances in the phylogenetic study of

ghrelin. Peptides, 32: 2155-2174.

Kaiya H, Sakata I, Kojima M, Hosoda H, Sakai T, Kangawa K. 2004. Structural

determination and histochemical localization of ghrelin in the red-eared slider turtle,

Trachemys scripta elegans. General and Comparative Endocrinology, 38: 50-57.

Kaiya H, Kojima M, Hosoda H, Moriyama S, Takahashi A, Kawauchi H, Kangawa K. 2003a.

Peptide purification, complementary deosyribonucleic acid (DNA) and genomic DNA

cloning, and functional characterization of ghrelin in rainbow trout. Endocrinology, 144:

5215-5226.

Kaiya H, Kojima M, Hosoda H, Riley LG, Hirano T, Grau EG, Kangawa K. 2003b.

Identification of tilapia ghrelin and its effects on growth hormone and prolactin release in the

tilapia Oreochromis mossambicus. Comparative biochemistry and physiology part B:

biochemistry and molecular biology, 135: 421-429.

Kaiya H, Van Der Greyten S, Kojima H, Hosoda H, Kitajima Y, Matsumoto M, Geelissen S,

Darras VM, Kangawa K. 2002. Chicken ghrelin: purification, cDNA cloning and biological

activity. Endocrinology, 143: 3454-3463.

Kaiya H, Kojima H, Hosoda H, Koda A, Yamamoto K, Kitajima Y, Matsumoto M,

Minamitake Y, Kikuyama S, Kangawa K. 2001. Bullfrog ghrelin is modified by n-octanoic

acid at its thrid threonine residue. Journal of Biological Chemistry, 276: 40441-40448.

Kawakoshi A, Kaiya H, Riley LG, Hirano T, Grau EG, Miyazato M, Hosoda H, Kangawa K.

2007. Identification of a ghrelin-like peptide in two species of sharks, Sphyrna lewini and

Carcharhinus melanopterus. General and Comparative Endocrinology, 151: 259-268.

Kojima M and Kangawa K. 2005. Ghrelin: structure and function. Physiological reviews, 85:

495-522.

Kojima M, Hosoya H, Data Y, Nakazato M, Matsuo H, Kangawa K. 1999. Ghrelin is a

growth-hormone-releasing acylated peptide from stomach. Nature, 402: 656-660.

Manning AJ, Murray HM, Gallant JW, Matsuoka MP, Radford E, Douglas SE. 2008.

Ontogenetic and tissue-specific expression of preproghrelin in the Atlantic halibut,

Hippoglossus hippoglossus L. Journal of Endocrinology, 196: 181-192.

25

Masuda Y, Tanaka T, Inomata N, Ohnuma N, Tanaka S, Itoh Z, Hosoda H, Kojima M,

Kangawa K. 2000. Ghrelin stimulates gastric acid secretion and motility in rats. Biochemical

and Biophysical Research Communications, 276: 905-908.

Miandare HK, Farahmand H, Ramezanpour S, Nematollahi MA, Rafiee G, Amiri BM. 2011.

Effects of ghrelin on some plasma hormonal changes in juvenile sturgeon (Acipenser

persicus). Comparative Clinical Pathology, 21: 1099-1102.

Miura T, Maruyama K, Shimakura S, Kaiya H, Uchiyama M, Kangawa K, Shioda S, Matsuda

K. 2006. Neuropeptide Y mediates ghrelin-induced feeding in the goldfish, Carassius

auratus. Neurosciences Letters, 407: 279-283.

Miura T, Maruyama K, Shimakura S, Kaiya H, Uchiyama M, Kangawa K, Shioda S, Matsuda

K. 2007. Regulation of food intake in the goldfish by interaction between ghrelin and orexin.

Peptides, 28: 1207-1213.

Miura T, Maruyama K, Kaiya H Miyazati M, Kangawa K, Uchiyama M, Shioda S, Matsuda

K. 2009. Purification and properties of ghrelin from the intestine of the goldfish, Carassius

auratus. Peptides, 30: 758-765.

Nagaya N and Kangawa K. 2003. Ghrelin improves left ventricular dysfunction and cardiac

cachexia in heart failure. Current Opinion in Pharmacology, 3: 146-151.

Nagaya N, Kojima M, Uematsu M, Yamagishi M, Hosoda H, Oya H, Hayashi Y, Kangawa K.

2001. Hemodynamic and hormonal effects of human ghrelin in healthy volunteers. American

Journal of Physiology: Regulatory, Integrative and Comparative Physiology, 280: R1483-

R1487.

Nieminen P, Mustonen AM, Hyvarinen H. 2003. Fasting reduces plasma leptin-and ghrelin-

immunoreactive peptide concentrations of the burbot (Lota lota) at 2 degrees C but not at 10

degrees C.

Parhar IS, Sato H, Sakuma Y. 2003. Ghrelin gene in cichlid fish is regulated by sex and

development. Biochemical and biophysical research communications, 305: 169-175.

Peino R, Baldelli R, Rodriguez-Garcia J, Rodriguez Segade S, Kojima M, Kangawa K, Ghigo

E, Dieguez C, Casanueva FF. 2000. Ghrelin-induced growth hormone secretion in humans.

European Journal of Endocrinology, 143: R11-R14.

Pong SS, Chaung LY, Dean DC, Nargund RP, Patchett AA, Smith RG. 1996. Identification of

a new G-protein-linked receptor for growth hormone secretagogues. Molecular

endocrinology, 10: 57-61.

Riley LG, Fox BK, Kaiya H, Hirano T, Grau EG. 2005. Long-term treatment of ghrelin

stimulates feeding, fat deposition, and alters the GH/IGF-I axis in the tilapia, Oreochromis

mossambicus. General and Comparative Endocrinology, 142: 234-240.

Salmerón S, Johansson M, Angotzi AR, Rønnestad I, Jönsson E, Björnsson BT, Gutiérrez J,

Navarro I, Capilla E. Effects of nutrional status on plasma leptin levels and in vitro regulation

of adipocyte leptin expression and secretion in rainbow trout. Submitted article.

26

Saito E, Kaiya H, Takagi T, Yamasaki I, Denbow DM, Kangawa K, Furuse M. 2002. Chicken

ghrelin and growth hormone-releasing peptide-2 inhibit food intake of neonatal chicks.

European Journal of Pharmacology, 453: 75-79.

Shimizu M, Swanson P, Fukada H, Hara A, Dickhoff WW. 2000. Comparison of Extraction

Methods and Assay Validation for Salmon Insulin-like Growth Factor-I Using Commercially

Available Components. General and Comparative Endocrinology, 119: 26-36.

Tinoco AB, Näslund J, Delgado MJ, Pedro N, Johnsson JI, Jönsson E. 2014. Ghrelin

increases food intake, swimming activity and growth in juvenile rainbow trout (Salmo trutta).

Physiology and Behavior, 124: 15-22.

Unniappan S, Peter RE. 2004. In vitro effects of ghrelin on luteinizing hormone and growth

hormone release in goldfish. Regulatory, Integrative and Comparative Physiology, 286:

R1093-R1101.

Unniappan S, Canosa LF, Peter RE. 2004. Orexigenic actions of ghrelin in goldfish: feeding-

induced changes in brain and gut mRNA expression and serum levels, and responses to

central and peripheral injections. Neuroendocrinology, 79: 100-108.

Unniappan S, Lin X, Cervini L, Rivier J, Kaiya H Kangawa K, Peter RE. 2002. Goldfish

ghrelin: molecular characterization of the complementary deoxyribonucleic acid, partial gene

structure and evidence for its stimulatory role in food intake. Endocrinology, 143: 4143-4146.

Wang JX, Peng KM, Liu HZh, Song H, Chen X, Liu M. 2009. Distribution and

developmental changes in ghrelin-immunopositive cells in the gastrointestinal tract of African

ostrich chicks. Regulatory Peptides, 154: 97-101.

Xu M, Volkoff H. 2009. Molecular characterization of ghrelin and gastrin-releasing peptide in

Atlantic cod (Gadus morhua): cloning, localization, developmental profile and role in food

intake regulation. General and Comparative Endocrinology, 160: 250-258.

Zhao Z, Sakata I, Okubo Y, Koike K, Kangawa K, Sakai T. 2008. Gastric leptin, but not

estrogen and somatostatin, contributes to the elevation of ghrelin mRNA expression level in

fasted rats. Journal of Endocrinology, 196: 529-538.