Diet-induced obesity suppresses ghrelin in rat gastrointestinaltract and serum

Ibrahim Sahin • Suleyman Aydin • Yusuf Ozkan •

Adile Ferda Dagli • Kadir Okhan Akin • Saadet Pilten Guzel •

Zekiye Catak • Mehmet Resat Ozercan

Received: 28 February 2011 / Accepted: 28 April 2011 / Published online: 10 May 2011

� Springer Science+Business Media, LLC. 2011

Abstract The aims of the present study were to examine

ghrelin expression in serum and gastrointestinal tract (GIT)

tissues, and to measure tissue ghrelin levels and obesity-

related alterations in some serum biochemical variables in

rats with diet-induced obesity (DIO). The study included 12

male rats, 60 days old. The rats were randomly allocated to

two groups (n = 6). Rats in the DIO group were fed a caf-

eteria-style diet to induce obesity, while those in the control

group were fed on standard rat pellets. After a 12 week diet

program including an adaptation period all rats were

decapitated, tissues were individually fixed, ghrelin expres-

sion was examined by immunohistochemistry , and tissue

and serum ghrelin levels were measured by radioimmuno-

assay. Serum biochemical variables were measured using an

autoanalyzer. When the baseline and week 12 body mass

index and GIT ghrelin expression were compared between

DIO and control rats, BMI had increased and ghrelin

expression decreased due to obesity. The RIA results were

consistent with these findings. Serum glucose, LDL choles-

terol, and total cholesterol levels were elevated and HDL

cholesterol significantly decreased in the DIO group. A

comparison of GIT tissues between the control and obese

groups demonstrated that ghrelin was decreased in all tissues

of the latter. This decrease was brought about a decline in the

circulating ghrelin pool. This suggests that rather than being

associated with a change in a single tissue, obesity is a

pathological condition in which ghrelin expression is chan-

ged in all tissues.

Keywords Diet-induced obesity �Immunohistochemistry � Gastrointestinal tract �Acylated ghrelin � Desacylated ghrelin

Introduction

Obesity results from impairment of the balance between

the body weight and body height in favor of the former.

This impairment is brought about by genetic factors,

dietary habits, physical activity, and orexigenic and

anorexigenic hormone levels. Ghrelin, a hormone of lipo-

peptide-structure, was discovered by Kojima et al. in 1999 in

the X/A cells (ghrelin cells) of the fundus and pyloric parts of

the rat stomach [1]. It is composed of 28 amino acids and

contains a fatty acyl (octanoyl) group on the third amino acid

(serine) from the N-terminal. This bioactive peptide is found

in two major forms, acylated (with octanoyl-bioactive) and

desacylated (without octanoyl-inactive). These two forms

are present in both the plasma and tissues [2–6].

Levels of ghrelin are lower in obese humans than in thin

humans [7, 8]. Research on experimental animals has

I. Sahin

Department of Nutrition and Dietetics, Erzincan University,

Erzincan, Turkey

I. Sahin � S. Aydin (&) � S. P. Guzel � Z. Catak

Department of Medical Biochemistry and Clinical Biochemistry

(Firat Hormones Research Group), Firat University Hospital,

Medical Faculty of Firat University, 23100 Elazig, Turkey

e-mail: saydin1@hotmail.com

Y. Ozkan

Department of Internal Medicine, School of Medicine, Firat

University Hospital, Elazig, Turkey

A. F. Dagli � M. R. Ozercan

Department of Pathology, School of Medicine, Firat University

Hospital, Elazig, Turkey

K. O. Akin

Department of Medical Biochemistry and Clinical Biochemistry,

Kecioren Education and Research Hospital, Ankara, Turkey

123

Mol Cell Biochem (2011) 355:299–308

DOI 10.1007/s11010-011-0867-5

supported this finding [2, 9, 10]. Weight loss brings about an

increase in circulating ghrelin levels [11]. The fall in ghrelin

levels after food intake is less marked in obese individuals

than in healthy controls [12]. Although, endogenous ghrelin

and PYY(3-36) in diet-induced obese rats are not altered in

the fasting state, peripheral administration of ghrelin and

PYY (3-36) exhibit blunted responses in food intake [13].

As stated above, the stomach is the main site for ghrelin

production (reviewed by Schubert) [14], but the hormone is

synthesized in various other parts of the gastrointestinal tract

(GIT) and elsewhere in the body: hypothalamus, hypophysis,

salivary glands, thyroid gland, small intestines, kidneys,

heart, pancreas, central nervous system, lungs, placenta,

gonads, immune system, breasts, and teeth (reviewed by

Aydin) [2]. The amount of ghrelin released from these organs

accounts for about 30% of the blood ghrelin pool [3]. In the

GIT, ghrelin has been identified in the esophagus, duodenum,

jejunum, ileum, and colon, decreasing in intensity from the

duodenum toward the colon [2]. Ghrelin mRNA is present in

almost all tissues, as demonstrated by Gnanapavan et al. [15].

Quantities of mRNA were reported in the following, in

descending order: stomach fundus, jejunum, duodenum,

antrum, lung, pancreas, vein, gall bladder, lymph node,

esophagus, left colon, cheek, somatotropinoma, hypophysis,

breast, kidney, ovarium, prostate, right colon, ileum, liver,

spleen, fallopian tubes, lymphocyte, testis, adipose tissue,

placenta, adrenal gland, muscle, urinary bladder, atrium,

thyroid, myocardium, and skin [15]. Diet-induced models of

obesity in rats aptly represent obesity in humans. There is a

significant correlation between the development of obesity

and circulating lipid parameters. Furthermore, acylated and

desacylated ghrelin are transported in the circulation by

binding to very high density lipoprotein (VHDL) and HDL

[16]. Similarly, a significant relationship has been reported

between ghrelin and weight gain in several studies [17–19].

On the basis of the available data, the aims of the present

study were to create diet-induced obesity in rats to determine

and compare: (I) ghrelin expression in GIT tissues by

immunohistochemistry, (II) ghrelin expression in GIT tis-

sues by RIA, (III) serum ghrelin levels by RIA, and (IV)

serum glucose, alanine aminotransferase (ALT), aspartate

aminotransferase (AST), gamma glutamyltransferase

(GGT), total cholesterol, triglycerides, HDL cholesterol,

LDL cholesterol, total protein, albumin, calcium, and iron

levels using an autoanalzyer.

Materials and methods

Experimental animals and diet protocol

The study was carried out at the Experimental Research

Unit of Firat University (FUDAM), after the approval by

the Ethics Committee of Firat University Medical School

with the decision dated May 25, 2006 and number 20. The

study included 12 two-month-old male rats of the Wistar

albino type. After random allocation to two groups, the rats

were put into four cages, each sheltering three rats. Fol-

lowing a 1 week period of adaptation, the heights and

weights of the rats were measured to determine their BMIs.

In order to induce diabetes through diet, rats in the DIO

group were fed on a cafeteria-style diet consisting of 33%

rat pellet, 33% whole-fat sugary milk (Nestle), 7% sucrose,

and 27% water, while the rats in the control group were fed

ad libitum on standard rat pellet containing 51% carbohy-

drate, 4% fat, 21% protein, and 24% water [20]. All rats

were given water ad libitum. In order to determine the

development of DIO, BMI values were recorded at three-

week intervals.

Body mass index (BMI) and obesity criteria in rats

BMI is commonly determined according to the following

formula: BMI = body weight (kg)/height2 (m2). The

height of each rat was measured as the distance between

the tip of the nose and the anus. In order to determine their

weight, the rats were weighed in a capped plastic container

without tare. BMI value in healthy rats is considered to be

0.72 kg/m2 [21]. Despite the extensive literature search

undertaken, no BMI value indicating obesity in rats could

be found. Human individuals with BMI between 18.5 and

24.9 are accepted as having normal weight, those with BMI

between 25 and 29.9 are accepted as overweight, and those

with BMI at and above 30 are accepted as obese [22]. In

order to determine a BMI for obese rats, the ratios between

normal and obese BMI in humans were used. Accordingly,

the mean BMI of normal people was calculated

(18.5 ? 24.9 = 43.4/2 = 21.7). This value was subtracted

from the obesity borderline value in humans

(BMI C 30 kg/m2) (30-21.7 = 8.3) to determine what

percentage of BMI in normal humans should be added to

the mean to obtain an obese value (38.248%). This coef-

ficient (38.248%) was added to the BMI values of the

healthy rats to determine the probable BMI value of obese

rats (38.248% of 0.72 is 0.275; 0.72 ? 0.275 = 0.995 g/

cm2). As BMI is expressed in g/cm2 in rats, the obese BMI

was established to be C1.00 g/cm2 for rats.

Preparation and storage of samples

At the end of the diet program, the rats were sacrificed

using sterile equipment. Half of the blood obtained was put

into regular biochemistry tubes containing aprotinin and

the other half into regular empty biochemistry tubes. The

blood samples were centrifuged for 5 min at 4,000 rpm at

room temperature. Sera containing aprotinin were added to

300 Mol Cell Biochem (2011) 355:299–308

123

1/10 N HCl and transferred to Eppendorf tubes to be stored

in the deep freeze at -20�C pending analyses [23]. Bio-

chemical variables were measured immediately in the

serum samples without aprotinin. The tissue samples were

also divided into two for immunohistochemical (IHC)

staining and determination of ghrelin levels. Samples for

the IHC staining were immediately put into 10% formal-

dehyde. Samples for quantifying ghrelin were put into a

boiling water bath immediately after dissection and boiled

for 5 min, preventing proteolysis of the ghrelin. After

boiling, the tissue samples (100 mg) were transferred into

regular biochemistry tubes containing 10 ml 5% w/v

phosphate buffered saline (PBS), carefully ground with a

glass rod, and centrifuged at 4,000 rpm for 10 min at

?4�C. To adjust its temperature, the centrifuge was run

several times before loading. The resulting tissue super-

natants were added to 50 ll/ml (500 KIU) aprotinin and

stored in a deep freeze at -20�C pending analysis of the

hormone.

Hormone analysis

Levels of ghrelin in sera and tissues were measured by

RIA. Serum samples and tissue homogenates prepared in

advance and stored at -20�C were used. RIA was per-

formed in accordance with the procedures described in the

catalogues. As the measurement limits of the kits are set

(7.8 pg/ml for acylated ghrelin and 93 pg/ml for total

ghrelin), ghrelin levels below these amounts were mea-

sured as follows: the tubes were supplemented with pure

standards of 7.8 pg/ml for acylated ghrelin and 93 pg/ml

for total ghrelin. At the end of the experiment, these

quantities were subtracted from the values obtained to

determine the original ghrelin levels in the samples. The

quantities of desacylated ghrelin were established by sub-

tracting the amount of acylated ghrelin from the total

ghrelin. RIA analysis of ghrelin in serum and tissues was

carried out in three stages: preparation, analysis, and cal-

culation. Both IHC and RIA analyses recognize the octa-

noylated and des-octanylated porsion of ghrelin.

Antibodies rose against positions 1–11(N-terminal) and

13–28 (C-terminal) of ghrelin protein.

Immunohistochemistry

The tissues were stained immunohistochemically using

Avidin–Biotin-Peroxidase Complex (ABC) as suggested

by Hsu et al. [24]. IHC analyses consisted of two stages:

preparation and staining. The solutions to be used were

freshly prepared before staining. The chromogen solution

(3-amino-9-ethyl carbazole, AEC) was used within 15 s

after it was prepared.

Preparation: 1. Sections of 4 lm from the blocks of

tissue samples in paraffin were transferred on to polylysine

microscope slides. 2. They were kept in the drying oven

(?80�C) for 20 min. 3. They were passed through a pure

xylol series four times, 5 min each (4 9 50). 4. They were

passed through an ethanol series (99.5 ? 96 ? 90 ?80 ? 70%) (5 9 30). 5. They were kept in distilled water

for 10 min. 6. They were kept in 3% H2O2 in methanol

(2 9 50).Staining: 1. Samples were incubated in 10% citrate

buffer, pH 6.0 for 15 min in a microwave heater (750 mw),

and left to cool for 20 min at room temperature. 2. They

were kept in PBS (0.01 M, pH 7.4) for 5 min. 3. They were

incubated with horse radish peroxidase blocking for

10 min. 4. They were incubated with ghrelin tissue anti-

body for 30 min in a water bath (?38�C). 5. They were

kept in PBS for 5 min. 6. They were incubated with bio-

tinylated goat anti-mouse for 10 min in a water bath

(?38�C). 7. They were kept in PBS for 5 min. 8. They

were incubated with streptavidin peroxidase for 10 min in

a water bath (?38�C). 9. They were kept in PBS for 5 min.

10. They were incubated with AEC for 10 min in a water

bath (?38�C). 11. They were washed with distilled water

for 1–2 min. 12. They were rinsed (not thoroughly dried)

and subjected to contrast staining with Mayer hematoxylin

for 1–2 min. 13. They were washed in distilled water for

1–2 min and dried. 14. They were covered with lamellae to

obtain permanent preparations. 15. The preparations were

examined under a light microscope and photographed.

Salivary gland, which is known to express ghrelin, was

used as a positive control [17]. When preparing negative

tissue samples, PBS was used instead of primary antibody

(ghrelin tissue antibody). Absence of staining in tissue

preparations was evaluated as negative (-) and different

staining intensities were evaluated as ?1, ?2 and ?3 in the

IHC evaluation.

Statistical analyses

Statistical analyses were conducted using SPSS 10.00. The

data are presented as mean ± standard deviation. Levels of

acylated ghrelin, desacylated ghrelin, and total ghrelin

obtained from the statistical analyses were rounded to two

digits after the decimal point and the percentage differ-

ences in these values between the DIO and control groups

were rounded to one digit after the decimal point. A Mann–

Whitney U test was used to compare data between groups,

and a Wilcoxon rank test to determine the statistical sig-

nificance of repeated measurements. Possible correlations

between the data were evaluated using a Pearson correla-

tion test. The level of statistical significance was set at

P B 0.05.

Mol Cell Biochem (2011) 355:299–308 301

123

Results

There was no incidence of death in either group throughout

the diet program. BMI was regularly recorded at three-

week intervals throughout the diet program. Baseline, week

3, week 6, week 9, and week 12 BMI measurements are

presented in Table 1. The baseline BMI values were

0.55 ± 0.02 g/cm2 in the DIO group and 0.53 ± 0.03 g/

cm2 in the control group, with no statistical difference

between the two (P [ 0.05). The BMI values at the end of

the 12 week diet program were 1.02 ± 0.05 g/cm2 in the

DIO group and 0.73 ± 0.04 g/cm2 in the control group,

with a statistically significant difference (P \ 0.01). The

weight gain changes values were from the beginning

156.92 ± 13.37 g to the ending 290.30 ± 29.02 g for

controls, while the weight gain changes values were from

the beginning 166.63 ± 13.91 g to the ending

451.85 ± 33.62 g for DIO group

Biochemical measurements in the DIO and control rats

are presented in Table 2. Serum glucose, total cholesterol,

and LDL cholesterol levels in the DIO group were signif-

icantly greater than those in the control group (P \ 0.01).

HDL cholesterol, however, was significantly lower in the

DIO group than the control group (P \ 0.01). No statisti-

cally significant difference was established between the

DIO and control groups in terms of ALT, AST, GGT, tri-

glyceride, total protein, albumin, and calcium and iron

values.

Serum levels of ghrelin in the DIO and control group

rats are presented in Table 3. In comparison with the

control group, the DIO group had lower levels of acylated

ghrelin, desacylated ghrelin, and total ghrelin. The

decreases in desacylated and total ghrelin in the DIO group

were statistically significant (P \ 0.01), while the decrease

in acylated ghrelin was not (P [ 0.05).

Tissue ghrelin expression was analyzed in the GIT tis-

sues by RIA and IHC. IHC showed that the nuclei of all

cells were stained blue with Mayer hematoxylin. Ghrelin

was expressed in the cytoplasm, and the specific parts of

the cells were stained with a variety of colors ranging from

red to brown, depending on the intensity of staining.

In the GIT, the parotid gland, esophagus, stomach fun-

dus, stomach pylorus, duodenum, jejunum, ileum, cecum,

colon, rectum, liver, and pancreas tissues were examined.

Acylated ghrelin levels measured by RIA are shown in

Table 3. The decrease in acylated ghrelin in the DIO group

relative to the control group was 36.6% in the parotid

gland, 16.7% in the esophagus, 11.0% in the stomach

fundus, 10.5% in the stomach pylorus, 31.3% in the duo-

denum, 22.4% in the jejunum, 15.8% in the ileum, 23.8%

in the cecum, 22.7% in the colon, 36.0% in the rectum,

13.3% in the liver, and 22.1% in the pancreas.

Levels of desacylated ghrelin measured in the GIT by

RIA are presented in Table 3. The relative decreases in

desacylated ghrelin in the DIO group were 37.0% in the

parotid gland, 16.8% in the esophagus, 2.5% in the stom-

ach fundus, 14.5% in the stomach pylorus, 27.0% in the

duodenum, 26.3% in the jejunum, 8.9% in the ileum,

18.2% in the cecum, 12.7% in the colon, 21.3% in the

rectum, 7.9% in the liver, and 42.6% in the pancreas. An

Table 1 Changes in BMI of rats from the beginning to the end of the experiment

Parameter BMI (g/cm2)

Baseline 3rd week 6th week 9th week 12th week

Control (n:6) 0.53 ± 0.03a 0.57 ± 0.05 0.62 ± 0.05 0.67 ± 0.04 0.73 ± 0.04c

DIO (n:6) 0.55 ± 0.02b 0.59 ± 0.02 0.67 ± 0.02 0.83 ± 0.04 1.02 ± 0.05d

BMI Body mass index

P [ 0.05, a Baseline control group versus b baseline diet-induced obese group

P \ 0.05, a Baseline control group versus c 12th week control group

P \ 0.05, b Baseline diet-induced obese group versus d 12th week diet-induced obese group

P \ 0.01, c 12th week control group versus d 12th week diet-induced obese group

Table 2 Serum levels of biochemical variables

Parameter Control (n:6) DIO (n:6) P-value

Glucose (mg/dl) 90.83 ± 5.85 182.17 ± 6.85 <0.01*

ALT (U/l) 53.83 ± 13.39 62.33 ± 14.99 [0.05

AST (U/l) 217.83 ± 92.09 209.33 ± 47.13 [0.05

GGT (U/l) 4.50 ± 0.84 5.00 ± 1.87 [0.05

Total cholesterol (mg/dl) 149.33 ± 8.71 185.00 ± 6.93 <0.01*

Triglyceride (mg/dl) 128.67 ± 51.58 143.83 ± 33.35 [0.05

HDL cholesterol (mg/dl) 43.00 ± 6.78 30.00 ± 5.14 <0.01*

LDL cholesterol (mg/dl) 84.33 ± 8.55 126.23 ± 6.22 <0.01*

Total protein (mg/dl) 7.25 ± 0.81 6.53 ± 0.55 [0.05

Albumin (mg/dl) 3.43 ± 0.41 3.18 ± 0.17 [0.05

Calcium (mg/dl) 11.33 ± 0.81 10.73 ± 0.48 [0.05

Iron (lg/dl) 267.67 ± 71.03 238.33 ± 40.00 [0.05

ALT Alanine aminotransferase, AST Aspartate aminotransferase, DIODiet-induced obese, GGT Gamma glutamyltransferase HDL High density

lipoprotein LDL Low density lipoprotein. All data points are the average

of six results

302 Mol Cell Biochem (2011) 355:299–308

123

evaluation of ghrelin in the GIT by IHC in terms of

immunoreactivity is presented in Table 4, and IHC pho-

tographs showing ghrelin expression by the tissues are

presented in Figs. 1 and 2.

Discussion

It is estimated that there are currently 400 million obese

adults in the world and this figure will rise to more than 700

million by 2015. Obesity, which has become a serious

health problem in developed and developing countries

alike, places a heavy burden on the health systems of

countries where it is common. In order to eliminate the

health problems associated with obesity such as high blood

pressure, osteoarthritis, sleep apnea, dyslipidemia, type II

diabetes, coronary heart disease, and some types of cancer

(breast, colon and endometrial), researchers endeavoring to

identify and prevent the causes of obesity are allocated a

generous share of state budgets [25].

At the end of the 12 week diet program, the BMI values

of the DIO group rats rose above 1.02 g/cm2. Healthy rats

whose baseline body weights were 0.55 ± 0.02 g/cm2 and

were fed ad libitum on a cafeteria-style diet for 12 weeks

Table 3 Comparison of acylated and desacylated ghrelin levels in serum and GIT tissues of DIO and control rats

Parameter Acylated ghrelin (pg/mg tissue) Desacylated ghrelin (pg/mg tissue)

Control (n:6) DIO (n:6) Control (n:6) DIO (n:6)

Parotid gland 17.75 ± 7.89 11.25 ± 4.57 169.75 ± 24.39 107.00 ± 13.14a

Esophagus 24.00 ± 7.53 20.00 ± 8.04 218.25 ± 20.61 181.75 ± 14.80a

Stomach fundus 27.17 ± 7.73 24.17 ± 7.81 242.17 ± 39.25 236.17 ± 43.83

Stomach pylorus 23.83 ± 5.04 21.33 ± 7.34 222.00 ± 28.12 189.83 ± 29.10

Duodenum 16.75 ± 5.74 11.50 ± 3.87 151.75 ± 10.72 110.75 ± 10.47a

Jejunum 12.25 ± 3.59 9.50 ± 5.45 114.25 ± 17.80 84.25 ± 7.14a

Ileum 9.50 ± 4.95 8.00 ± 7.07 95.00 ± 11.31 86.50 ± 24.75

Cecum 10.50 ± 7.78 8.00 ± 2.83 104.50 ± 23.33 85.50 ± 19.09

Colon 11.00 ± 4.24 8.50 ± 4.95 98.50 ± 24.75 86.00 ± 31.11

Rectum 12.50 ± 4.95 8.00 ± 5.66 110.00 ± 31.11 86.50 ± 43.13

Liver 26.25 ± 7.37 22.75 ± 5.44 233.25 ± 28.43 214.75 ± 20.45

Pancreas 19.25 ± 7.54 15.00 ± 8.37 129.25 ± 10.84 74.25 ± 8.22a

Serum (pg/ml) 46.20 ± 13.95 34.00 ± 5.96 1240.20 ± 88.17 948.20 ± 34.22b

All data points are the average of six results and values are given as mean ± SD

DIO Diet-induced obesea Diet-induced obese versus control group P \ 0.05b Diet-induced obese versus control group P \ 0.01

Table 4 Ghrelin expression by

GIT tissues

DIO Diet-induced obese, IHCEImmunohistochemical (IHC)

evaluation

Tissue Ghrelin location IHCE

Control DIO

Parotid gland Interlobular ducts ?2 ?1

Esophagus Stratum corneum ?3 ?3

Stomach fundus Stratum corneum and stratum granulosum ?3 ?3

Stomach pylorus Gland epithelium ?3 ?3

Duodenum Gland epithelium ?2 ?1

Jejunum Gland epithelium ?1 –

Ileum Gland epithelium and mucosa epithelium ?1 –

Cecum Gland epithelium ?1 –

Colon Gland epithelium ?1 –

Rectum Gland epithelium ?1 –

Liver Hepatocytes ?3 ?3

Pancreas Serous acinus ?3 ?2

Mol Cell Biochem (2011) 355:299–308 303

123

were seen to develop obesity. The units of BMI values used

for rats vary. Some studies use kg/m2 [26, 27] and others

use g/cm2 [28, 29]. Novelli et al. [30] suggested the use of

g/cm2 in their study. We also employed this unit (g/cm2)

for BMI values. Although this kind of diet can cause

obesity in rats over the period we used, we are of the

opinion that BMI values rather than dietary period should

be employed to determine the development of DIO in rats.

When the serum glucose levels of the obese rats were

compared with those of the control group, the former was

seen to be statistically significantly elevated. Many studies

have shown that the amount of adipose tissue increases

with obesity [30, 31]. Adipose tissue that is metabolically

active continuously supplies non-esterified or free fatty

acids (FFA) to the liver. While providing the energy

required for glucogenesis, high FFA levels lead to hepatic

insulin resistance owing to the increase in hepatic glucose

output. Thus, this mechanism impairs glucose tolerance by

reducing cellular glucose intake and increasing the blood

glucose level.

The effects of obesity, which leads to liver dysfunction,

and its relationship to other parameters, have been topics of

Fig. 1 Negative control with no primary antibody added to GIT

tissues (a–f). Positive immunoreactivity: ghrelin distribution is

indicated by dark brown color (shown with arrow) in the control

rat’s parotid gland, esophagus, stomach fundus, stomach pylorus,

duodenum, and jejunum (a1–f1). Positive immunoreactivity: ghrelin

distribution is indicated by dark brown color (shown with arrow) in

the diet-induced obese rat’s parotid gland, esophagus, stomach

fundus, stomach pylorus, duodenum, and jejunum (a2–f2). Magnifi-

cation: 2009

Fig. 2 Negative control with no primer primary antibody added to

GIT tissues (a–f). Positive immunoreactivity: ghrelin distribution is

indicated by dark brown color (shown with arrow) in the control rat’s

ileum, cecum, colon, rectum, liver, and pancreas (a1–f1). Positive

immunoreactivity: ghrelin distribution is indicated by dark browncolor (shown with arrow) in the diet-induced obese rat’s ileum,

cecum, colon, rectum, liver, and pancreas (a2–f2). Magnification:

2009

304 Mol Cell Biochem (2011) 355:299–308

123

many recent studies. Therefore, some liver function tests

(ALT, AST, GGT, and albumin) were conducted in the

present study to investigate changes in serum levels. The

results revealed that ALT levels were higher while AST

levels were lower in obese rats than in controls. In two

independent studies, Brucket et al. [32] and Clark et al.

[33] established that ALT levels in obese individuals were

higher than normal. Studies of experimental animals have

also revealed that ALT increases with obesity [34].

Increased ALT and AST indicate hepatocellular damage. It

appears that ALT (EC. 2.6.1.2) is more markedly affected

by obesity than AST (EC. 2.6.1.1). A possible explanation

may be that although ALT is synthesized only in the liver,

AST is synthesized and stored in the pancreas, liver, lung,

heart, spleen, muscle, erythrocytes, and brain as well as the

liver.

It was also found in this study that the serum GGT level

was almost 10% higher in the DIO group than the controls.

Elevated GGT activity is mainly seen in diseases of the

liver, biliary tract, and pancreas. The increase associated

with DIO indicates hepatocellular damage. It is understood

that kidney, heart, brain, pancreas, and spleen tissues are

afflicted with inflammation in obesity. GGT (EC. 2.3.2.2),

which is a cell surface glycoprotein, is found in liver,

pancreas, kidney, seminal glands, heart, spleen, and brain,

and these organs may have increased in capacity by 10%

because of inflammation. Recent studies have also shown a

strong correlation between elevated GGT levels and the

metabolic syndrome [35].

It was established in the present study that total cho-

lesterol, triglyceride, and LDL cholesterol levels were

higher, and HDL cholesterol was significantly lower, in the

obese rats than the controls. A possible reason why tri-

glycerides and cholesterol were higher in the obese rats is

that the cafeteria-style diet contains more fat than normal

rat pellet, and more of this fat is absorbed by the intestine.

As stated previously, insulin resistance is among the major

components of obesity. The development of insulin resis-

tance brings about an increase in the plasma concentration

of free fatty acids, which stimulate the accumulation of

triglyceride in the liver. Also, insulin resistance causes a

decrease in plasma lipoprotein lipase activity, which leads

to dyslipidemia and hypertriglyceridemia (elevated plasma

triglycerides) on the one hand, and destruction of HDL

cholesterol on the other [36]. In addition, increased hepatic

glucogenesis creates a predisposition to glucose intolerance

in the liver and muscles [37].

Metabolic and endocrine changes take place during the

preliminary stages of obesity. Ghrelin is a lipopeptide-

structure hormone that plays a pivotal part in the etiopa-

thology of obesity. Although this hormone is mainly

synthesized in the stomach, both its mRNA and the peptide

itself have been demonstrated in many tissues [15]. It is

found in two major forms in the circulation, acylated and

desacylated. This study compared serum ghrelin levels

between DIO rats and controls. Both the forms were

observed to decrease in obesity. Clinical studies have

demonstrated that ghrelin levels decline in the obese [8].

Similarly, the present study showed decreased ghrelin in

obese rats. Obesity is closely associated with inflammation

and oxidative stress. Ghrelin not only fulfills a hormonal

function, but also is an endogenous antioxidant [38].

Therefore, ghrelin may have been consumed in eliminating

the inflammation and oxidative stress caused by obesity,

and this may have caused the decline in levels. In addition,

increases in the glucose level are accompanied by decreases

in ghrelin, and so the increased glucose level observed in

this study may also have brought about a decrease in

ghrelin; the decreased ghrelin levels may be associated with

energy regulation. All these factors may jointly or severally

account for the decrease in ghrelin levels. Furthermore, the

loss of ghrelin expression in tissues caused by obesity may

have contributed. The following part of the discussion will

deal with ghrelin expression in the GIT tissues and their

interactions with obesity.

The present study compared ghrelin expression in the

GIT between DIO rats and controls. Ghrelin expression in

the parotid gland, a part of the GIT, showed positive

staining with ghrelin antibody in its striated, intralobular,

sinus, and asinus parts. The intensity of this staining was

lower in the obese group. These results are consistent with

the tissue ghrelin levels analyzed by RIA. They also show

that the salivary gland secretes ghrelin. Our literature

review revealed no study indicating whether the salivary

glands express ghrelin in control or DIO group rats.

However, Gnanapavan et al. [15] reported ghrelin mRNA

in the salivary gland using RT-PCR. Since ghrelin is

known to be present in human saliva [5, 17], we think that

ghrelin synthesized by the salivary gland may be trans-

ferred from saliva to blood or from blood to saliva after it

reaches a certain level. The presence of ghrelin in the

human salivary gland is consistent with the ghrelin posi-

tivity of the salivary gland in rats [5, 17, 39]. Furthermore,

several researchers have noted that various salivary pro-

teins may be transported [16, 40, 41].

Ghrelin expression in the esophagus, which is located in

the upper GIT, was demonstrated by both IHC and RIA.

The presence of ghrelin mRNA was demonstrated previ-

ously [15]. The present study established that ghrelin

expression in the esophagus declined with obesity.

Although ghrelin expression by the esophagus has been

demonstrated by IHC in humans, no published study

indicates that the esophagus synthesizes ghrelin in rats

[42].

The present study also explored how obesity changed

ghrelin expression in the stomach, the organ in which

Mol Cell Biochem (2011) 355:299–308 305

123

ghrelin was first discovered [1]. It has already been

reported that the stomach synthesizes ghrelin in X/A cells;

this study confirmed those results and showed that ghrelin

expression declined with obesity. The amount of ghrelin

synthesized in the fundus and pyloric parts of the stomach

was parallel to the ghrelin synthesized in the liver.

Although it has been reported that the stomach is respon-

sible for the predominantly produced circulating ghrelin

[43], the present study demonstrated that the liver synthe-

sizes almost as much ghrelin as the stomach, or even more.

The pancreas has also been shown to synthesize ghrelin,

reportedly in the beta or epsilon cells or another unknown

cell group [44]. The present study not only confirmed

ghrelin synthesis by the pancreas, but also showed a

decrease with the development of obesity.

Examination of ghrelin expression by the duodenum,

jejunum, ileum, cecum, colon, and rectum, which are other

organs of the GIT, showed a decrease from the duodenum

toward the ileum and a partial increase from the ileum to

the rectum in the control group. This pattern of change in

the GIT was found to be parallel in the control and DIO

groups, and ghrelin levels in the obese group were lower

than those in the control group. These data, obtained from

IHC, were confirmed by RIA. When the levels of ghrelin

expression were compared in our study, more ghrelin was

synthesized by the pyloric part of the stomach than the

duodenum. Although, this part of our study is not consis-

tent with the IHC studies by Date et al. [45], there is a

consistency with respect to the ghrelin levels in the stom-

ach fundus, jejunum, ileum, and colon. Besides, ghrelin

synthesis per mg tissue as measured by RIA was parallel to

the IHC results. Moreover, tissue ghrelin expression by the

GIT was consistent with the IHC evaluation data.

Further and more detailed studies are needed to identify

the functions of ghrelin, which is synthesized by a variety

of tissues. However, the available data suggest that it may

undertake several functions such as anti-apoptosis, anti-

angiogenesis, cell proliferation, growth hormone secretion,

cell division, reducing organ injury associated with burns,

reducing inflammation, and energy regulation [46, 47].

Obesity not only leads to damage and fibrosis in many

organs, most notably the liver (hepatosteatosis) but also the

parotid gland, stomach fundus, lung, artery, heart, spleen,

thymus, muscle, brain, and adipose tissue. Our results

revealed elevated ALT and GGT, which are indicators of

liver damage. Furthermore, there are studies showing

increased alkaline phosphatase (ALP) and bilirubin in

obese rats, and the development of obesity leads to various

degrees of liver damage [48]. The most crucial correlation

among these is the negative relationship between BMI and

ghrelin. These results confirm the findings of Jeyakumar

et al. [49].

Conclusion

It was established in the present study that serum and tissue

expression levels of both acylated and desacylated ghrelin

levels decreased in obesity. It is believed that the major

cause for this decrease in the blood pool is that synthesis of

ghrelin by tissues is inhibited to restore energy regulation.

As a matter of fact, one study has demonstrated that blood

ghrelin levels decrease in obese individuals. However, no

previous study has investigated tissue ghrelin levels in all

organs. Therefore, the present study provides empirical

evidence for the reduction of ghrelin expression in obesity.

Although, this is an experimental animal study, it may offer

a new perspective on the etiopathology of obesity in

humans. Furthermore, it indicates that the amount of

ghrelin is important in energy regulation for losing weight.

The best way to eliminate obesity, which has currently

become a pandemic, is to eradicate the agents that stimu-

late appetite, and the most significant of these is ghrelin; it

creates a sensation of hunger and leads people to food [6].

Elimination of the effect of acylated ghrelin by pharma-

cological means will retard weight gain. The decrease in

ghrelin due to obesity is a significant metabolic event for

maintaining energy homeostasis.

Acknowledgments The authors would like to acknowledge TUBI-

TAK (project no: 106S350) and FUBAP (project no: 1445) for their

financial support.

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