Enriched protein diet-modified ghrelin expression and secretion in rats

Regulatory Peptides 121 (2004) 113 – 119 www.elsevier.com/locate/regpep

Enriched protein diet-modified ghrelin expression and secretion in rats M.T. Vallejo-Cremades, L. Go´mez-Garcı´a, M. Chacatas-Cortesao, C. Moreno, M. Sa´nchez, E. De Miguel *, I.A. Go´mez de Segura Research Unit, Servicio de Cirugı´a Experimental, Hospital Universitario La Paz, Paseo de la Castellana 261, 28046 Madrid, Spain Received 26 November 2003; received in revised form 22 April 2004; accepted 29 April 2004 Available online 11 June 2004

Abstract Gastrointestinal (GI) integrity and function are regulated by nutrition and growth factors. The discovery of ghrelin, a natural growth hormone (GH) secretagogue produced by the gastrointestinal (GI) tract, is a potential link between diet and growth signals. The aim of this study was to evaluate macronutrient effect on ghrelin expression and secretion in addition to some possible function in intestinal trophic status. Wistar rats were fed a high-carbohydrate, high-protein (HP), high-fat or standard (St) diet. Animals received the same daily food volume and caloric intake. After 7 days, animals were fasted for 24 h and blood and tissue samples were obtained just before feeding or at 2 or 6 h after feeding. Fasting high-protein-fed rats had higher ghrelin plasma levels than with rats fed the high-carbohydrate, high-fat or standard diets. Two-hours after refeeding, ghrelin plasma levels had decreased in all groups with a slight recovery at 6 h after refeeding, except in the high-protein group. Ghrelin plasma levels in rats fed with the high-protein diet correlated negatively with their GH and insulin-like growth factor 1 (IGF-1) plasma concentrations which were also the lowest among the study groups. In conclusion, ghrelin secretion was nutritionally manipulated because a protein-enriched diet increased its levels. D 2004 Elsevier B.V. All rights reserved. Keywords: Ghrelin; GH; IGF-1; Intestinal morphometry; Protein; Fat; Carbohydrate; Diet

1. Introduction Ghrelin is a natural ligand of the growth hormone (GH) secretagogue receptor and a gastrointestinal (GI) hormone involved in the control of appetite and energy balance that may link nutritional status with neuroendocrine function [1]. This peptide is mainly produced and secreted in the gastric fundus [2,3] while hypothalamic ghrelin production seems to be more closely related with orexigenic pathways [4]. Serum ghrelin levels increase with fasting and decrease after refeeding in normal conditions [5,6]. However, the regulatory mechanism that governs the biosynthesis and secretion of ghrelin in the digestive tract has not yet been clarified. Secretory and circulating ghrelin levels also seem to be influenced by dietary nutrient content; * Corresponding author. Tel.: +34-917-277-154; fax: +34-917-277050. E-mail address: [email protected] (E. De Miguel). 0167-0115/$ - see front matter D 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.regpep.2004.04.016

a fat diet stimulates ghrelin secretion (75% fat in total caloric content [7]) while glucose decreases ghrelin serum levels after fasting [8]. A recent study in humans has shown higher ghrelin secretion after enriched protein intake [9]. Ghrelin is a potent GH secretagogue that acts on specific pituitary and hypothalamic receptors [10], physiologically regulating the pulsatile secretion of GH, as do GH-releasing hormone (GHRH) and somatostatin [11]. The dual action of ghrelin on GH secretion and food intake, as well as the dual localization of this peptide and its receptors in the hypothalamus and gastrointestinal tract, strongly suggests interdependency between these actions. Nutritional status is a critical element in the regulation of GH expression [12,13] and can act independently of ghrelin in certain conditions [14]. In addition, ghrelin can stimulate distinct populations of hypothalamic neurons that have opposing effects on GH secretion [15]. Various peptide growth factors have been found to regulate epithelial cell function within the mucosal epithe-

114

M.T. Vallejo-Cremades et al. / Regulatory Peptides 121 (2004) 113–119

lium of the gastrointestinal tract [12,16 – 19]. Growth hormone [20], whether administered exogenously or produced endogenously [21], stimulates small intestine growth. Growth hormone and high-protein (HP) diets have been used to improve intestinal function after injury [22,23]. Although GH treatment is controversial, many studies are attempting to regulate endogenous GH secretion with GH secretagogues [24]. In summary, because nutrient composition may greatly affect gastrointestinal structure and function, and, specifically, high-protein diets can affect the gastrointestinal tract in pathological conditions [25], we hypothesized that the effect of a macronutrient could also modify ghrelin response as well as gastrointestinal proliferative status and the insulin-like growth factor 1 (IGF-1) – GH axis. Our experimental design included a 24-h fast prior to refeeding and sacrifice in order to intensify the regulatory peptide plasma levels; determining the level of IGF-1 in plasma corroborated GH plasma levels indirectly [24] because IGF-1 expression in liver and other tissues is stimulated by GH. We have found that ghrelin secretion and synthesis can be manipulated with different macronutrients.

rats were fed at the same time for 1 week to condition them to the feeding schedule and avoid variations in their hormonal levels. All the rats completely consumed their food within that period. On the seventh day, they were fasted (24 h) and on the eighth day, the animals were sacrificed to collect samples. Animals in each dietary group were sacrificed either immediately before refeeding or at either 2 or 6 h after refeeding. Body weight was measured on the first and last day of the experiment. 2.2. Extraction of tissue and plasma samples Animals were anesthetized with 2% – 2.5% Isoflurane (ForaneR, Abbott), for laparotomy and blood (4 ml) was collected from the abdominal aorta into chilled polypropylene tubes containing disodium EDTA (1-mg/ml blood) and aprotinin (500-units/ml blood, Bayer) and was then centrifuged for 15 min at 3000  g. Sampling was done 3 –5 min after inducing anesthesia with isoflurane and the first procedure was to extract the blood sample to minimize the potential impact of anesthesia on hormonal levels. Plasma was stored at 80 jC. Tissue samples were also collected from the gastric fundus, duodenum and jejunum, were rinsed with saline and then stored at 80 jC.

2. Material and methods 2.3. Intestinal morphometry 2.1. Animals and dietary manipulations Adult Wistar rats (n = 96; Centre d’Elevage Janvier; le Genest, Saint-Isle, France) aged 12 weeks and weighing 237 F 12 g were used in this study. The animals were allowed 1 week of acclimatization followed by another week in individual wire floor cages in a standard environment at 22 jC, 50% humidity and 12-h light/dark cycle (European Directive 609/86). Rats were randomly distributed into four groups (n = 24 per group). Groups received an equal volume (70 ml) and calorie count (1 kcal/ml) of one of four diets: a high-carbohydrate diet (75% carbohydrate, 15% fat, 10% protein); a high-protein diet (45% carbohydrate, 25% fat, 30% protein); a high-fat diet (30% carbohydrate, 60% fat, 10% protein); and finally, a standard diet (St; 55% carbohydrate, 30% fat, 15% protein). Diets were obtained by mixing the appropriate proportions of three commercial food presentations: carbohydrates (starch hydrolysate; SHS Maxijul Super Soluble, SHS International; Liverpool, UK); protein (composed by a 98.5% lactoalbumin hydrolisate and 1.5% L-cystine; Oligopeptides, Clinical Nutrition S.A, Mataro; Barcelona, Spain); and lipids (Supracal SHS International; Liverpool, UK) in order to provide a 70-kcal/rat/day balanced diet. All diets were supplemented daily with 5 Al per rat of a vitamin complex (CernevitR, Clintec Parenteral SA, Maurepas; France) and rats had free access to water. Experiment schedule. After the first week of acclimatization, the four diet groups of rats had access to the corresponding diet from 0700 until 1100 for 6 days. All

A 1-cm section of the intestine was fixed in 10% buffered formaldehyde solution for 24 h. The samples were prepared in paraffin for 5-Am serial cuts; one section was obtained from each block and stained with Hematoxilin – Eosin. All samples were sectioned and reoriented by successive slices in the search for the sample with the longest villi. This sample was used because it yielded much more homogenous results than standard techniques based merely on the measurement of the 10 longest villi for a single sample. The length of the 10 longest villi in each sample and their corresponding crypts were determined. The samples were studied in an image analysis system (Videoplan, Kontron). 2.4. Proliferation (ki-67 labelling) To assess cell growth in the tissue samples, a proliferation antigen was studied with a specific antibody (NCLKi67-MM1 Novo Castra Laboratories; Newcastle. UK). Counts were carried out for 20 full-length crypts per preparation from 5-Am sections from each sample. Proliferating nuclei are immunohistochemically stained dark brown by the Ki-67 antigen whereas nonproliferating nuclei are stained blue. The wavelengths of both colours are recognised by means of a computerized system (CAS-200 Image Analyzer) that determines the area covered by a full-length section of crypt in order to obtain the ratio of the area of Ki67-positively stained tissue (nuclear area) to the total area (positively + negatively stained nuclear area). Morphometric

M.T. Vallejo-Cremades et al. / Regulatory Peptides 121 (2004) 113–119

and proliferative analyses were performed under a blind protocol. Finally, a relationship between proliferation and crypt length was established.

115

products were separated on a 2% agarose gel, the bands were visualized with ethidium bromide staining and then quantified with a densitometer (1D Manager, TDI; Madrid, Spain).

2.5. Hormone determinations 2.8. Statistical analysis Serum ghrelin, GH and IGF-1 and ghrelin tissue levels were measured by radioimmunoassay (RIA). Aliquots of plasma and tissue samples (2.5 Ag) were analysed using the rat ghrelin RIA kits (Phoenix, Pharmaceuticals; Mountain View, CA, USA) and the rat GH [I125] Biotrakk Assay System with magnetic separation (RPA551, Amersham Biosciences UK; Buckinghamshire, UK), according to the manufacturer’s instructions. The tracer was radioiodinated (I125) rat hormone and the minimum detectable amount for ghrelin was 10 pg/tube, with 5% inter- and 13% intra-assay CV. GH RIA kit sensitivity was 0.16 ng/ tube and the intraassay CV was 3.0% with an interassay CV of 10%. Because the design would not reflect pulsatile GH secretion, we also determined IGF-1 to screen response to GH to the high-protein and standard diets. Total serum IGF-1 was measured in the high-protein and standard diet groups by RIA after acid ethanol extraction with a commercially available kit (Diagnostic Systems Laboratories; Webster, TX, USA). The interassay CV was 5.5% and the intraassay CV was 3%. The sensitivity of this assay is 0.8 Ag/ml. 2.6. RNA and protein preparation Total RNA and protein from the gastric fundus were obtained by the Tri ReagentR method (Molecular Research Center; Ohio, USA) according to the manufacturer’s instructions. The concentration of RNA and the degree of protein contamination were assessed by spectrophotometric analysis. The protein concentration was measured with bicinchoninic acid protein assay (BCA Protein Assay Kit, Ref. 23227, Pierce; Rockford, IL, USA) using bovine serum albumin as the reference standard. 2.7. RT-PCR for ghrelin mRNA expression in gastric fundus Ghrelin mRNA expression was detected using a RT-PCR kit (Retrotools cDNA/DNA polymerase, Biotools). Reverse transcription (RT) was carried out with 1-Ag total gastric fundus RNA for 30 min at 42 jC in a 20-Al reaction volume. Polymerase chain reaction (PCR) was performed in a 50Al reaction volume using primers identical to those used by Kojima et al. [26] forward primer, 5VGAGCCCAGAGCCCAGAGCACCAGAAA3V; reverse primer, 5VAGTTGCAGAGGAGGCAGAAGCT3V. 18S rRNA (Quantum, Ambion) was used as an internal control because it remains stable in different treatments and physiological conditions (data not shown). Amplification involved 32 cycles of PCR, an initial period of denaturation (94 jC, 5 s), annealing (65 jC, 10 s) and an extension (72 jC, 1 min). The PCR

Statistical analysis of data was performed by a commercial computer program (Statview 4.5 Abacus Concepts). The Kolmogorov – Smirnov test was applied to the data to determine the normality of the distribution of the data to be used in the parametric test. One-way analysis of variance (ANOVA Factorial) and the Fishers test were used to compare the diet groups and a p value < 0.05 was considered statistically significant. Nonparametric tests (Mann –Whitney U-test and the Wilcoxon test) were also employed and confirmed the results. The regression lines were calculated when the F value was significant and a probability of < 5% (two-tailed) was considered significant.

3. Results 3.1. Dietary effect on rat weight During the 7 days of the study, no significant differences were found in body weight between the different dietary groups, neither in the fasting nor refeeding groups of animals. 3.2. Dietary effect on jejunum and duodenum morphometry and proliferation The duodenum of high-protein-fed rats showed a significant increase in crypt and villous length ( p < 0.05). Maximum differences in length were found in the crypts of rats fed the high-protein diet; their crypts were 20%, 27% and 28% longer, respectively, than those in rats fed with highcarbohydrate, high-fat and standard diets, respectively (Table 1). The villous length of high-protein-fed rats was 16%, 14% and 9% longer respectively, than those of rats on the high-carbohydrate, high-fat and standard diets, respectively (Table 1). Diet did not have the same effect in the jejunum as in the duodenum. Jejunal crypt length in high-fat rats was significantly longer ( p < 0.05) than in rats on the high-carbohydrate or standard diets (7% and 8%, respectively; Table 1). However, nonsignificant differences in villous length were found between the four diet groups (Table 1). Knowledge of enterocyte proliferation is vital in elucidating how nutrition and growth factors regulate intestinal cell turnover. A possible cause of longer villi could be a higher crypt cell proliferation rate. We studied proliferation with Ki-67 labelling and found that dietary content did not modify the duodenum or jejunum crypt labelling indices (Table 1).

116

M.T. Vallejo-Cremades et al. / Regulatory Peptides 121 (2004) 113–119

Table 1 Dietary effect on jejunum and duodenum morphometry and proliferation Diet

Villous length (Am)

Crypt length (Am)

Proliferation index

Duodenum

Jejunum

Duodenum

Jejunum

Duodenum

Jejunum

Standard High-carbohydrate High-fat High-protein

286 F 12 266 F 16 271 F 6 315 F 12*

157 F 6 154 F 1 162 F 9 161 F 2

65 F 1 67 F 6 66 F 2 77 F 3*

49 F 1 50 F 1 54 F 4* 51 F 5

930 F 43 870 F 75 910 F 23 900 F 32

880 F 21 870 F 83 860 F 22 900 F 71

Villous and crypt lengths (Am) were measured in each group of rats fed with one of the four study diets. Significant differences in crypt, villous length or proliferation index are indicated by *, p < 0.05 vs. length of animals fed the standard diet. Proliferation index ( 1000) was calculated as indicated in the Material and Method section from crypts of rats fed with each diet. No significant differences were found between groups.

3.3. Dietary manipulation affects plasma hormone concentration The ghrelin plasma concentration was increased by fasting in all the studied diets (Table 2). Rats on the highprotein diet had the highest plasma ghrelin levels in the fasting measurement. Ghrelin immunoreactivity in blood 2 h after refeeding was 60%, 53%, 43% and 56% (highcarbohydrate, high-protein, high-fat and standard, respectively) lower than in the fasting period (Table 2). In animals receiving a high-carbohydrate, high-fat or a standard diet, there was a slight recovery in plasma ghrelin levels that did not reach fasting levels at 6 h after refeeding. This recovery was not observed in the high-protein diet-fed animals at 6 h after refeeding. Comparable to systemic ghrelin concentrations, GH plasma levels were also regulated by food intake. The four diets had a similar effect on the GH secretion pattern during fasting and after feeding (2 and 6 h). GH plasma levels were the lowest during fasting while the concentration after feeding increased by 1.5– 2-fold (Table 2). The concentration of GH in plasma in the animals on the high-protein diet was significantly lower than in the animals fed with the standard diet. High-carbohydrate and high-protein diet-fed rats showed a 45 – 50% lower GH concentration than rats on the standard and high-fat diets at 6 h after feeding ( p < 0.05). Plasma GH concentrations were negatively correlated with plasma ghrelin levels (r = 0.33 p < 0.05). Because the GH plasma levels in the rats on the highprotein diet were much lower than we had expected, based

on the literature, we used IGF-1 as a screening tool to indirectly measure the GH plasma levels in the rats fed the high-protein diet, with the objective of establishing a relationship between the morphometric and proliferative results and the hormonal levels in this dietary group. IGF1 has a longer half-life than GH and gives information about integrated GH secretion because there are spontaneous pulsatile fluctuations in the GH level. Fasting IGF-1 plasma levels were lower than after refeeding in the group on the high-protein diet than in the standard diet group (Table 2). The IGF-1 concentration was the highest 6 h after refeeding in the standard-fed rats ( p < 0.05). Two hours after refeeding, plasma IGF-1 levels were similar in rats fed with a high protein and those on a standard diet. Plasma IGF-1 levels in rats fed with a high-protein diet correlated negatively with their ghrelin plasma levels (r=-0.49; p < 0.05) and positively with their GH plasma levels (r = 0.49; p < 0.05). 3.4. mRNA and protein tissue expression of ghrelin The effects of the high-protein diet were further analyzed by determining stomach ghrelin mRNA expression and peptide levels compared to those in rats fed the standard diet. Ghrelin tissue expression was reverse that found in plasma where the highest ghrelin levels were observed at 6 h after refeeding (Fig. 1A) while the fasting the tissue concentration was lowest. As observed in the plasma levels, ghrelin tissue levels were significantly higher in rats fed the protein-enriched diet than in rats receiving a standard diet (Fig. 1A). Ghrelin mRNA expression followed the same

Table 2 Dietary effect on hormone plasma level Diets

Hormones Ghrelin (pg/ml) Fasting

Standard High-carbohydrate High-fat High-protein

1526 F 187 1531 F 166 1610 F 286 2079 F 224*

GH (ng/ml) +2 h

+6 h #

678 F 145 610 F 162# 919 F 173*,# 970 F 198*,#

Fasting #

984 F 197 817 F 172# 1108 F 156# 827 F 100#

19.4 F 5.4 17.3 F 3.2 15.4 F 8.7 10.0 F 3.5*

IGF-1 (ng/ml) +2 h 15.6 F 6.5 21.3 F 5.4 15.7 F 5.8 20.7 F 13.3

+6 h

Fasting #

43.6 F 7.5 28.4 F 8.9# 39.9 F 7.3# 20.6 F 5.1*,#

513 F 3 n.d. n.d. 373 F 4*

+2 h

+6 h #

487 F 2 n.d. n.d. 448 F 1#

630 F 7# n.d. n.d. 512 F 3#

Ghrelin and GH plasma concentrations were measured during fasting and at 2 and 6 h after refeeding in all diets studied. IGF-I plasma concentrations were measured just in rats fed with high-protein and standard diet as mentioned in the Results section. Mean F S.E. and significant differences between ghrelin plasma levels are indicated as: *, p < 0.05 groups vs. standard diet; #, p < 0.05 groups vs. fasting. n.d., Not determined.

M.T. Vallejo-Cremades et al. / Regulatory Peptides 121 (2004) 113–119

Fig. 1. Gastric fundus ghrelin protein levels and mRNA expression. (A) Ghrelin concentration in gastric fundus was measured by RIA in 2.5 Ag of tissue. *p < 0.05 vs. HP diet; #p < 0.05 vs. fasting. (B) Ghrelin mRNA in fundus was measured by RT-PCR with 1 Ag of total mRNA in eight assays. MW, Molecular weight standards; HP, high-protein diet; St, standard diet; and C, control.

pattern as the plasma ghrelin levels in the high-protein dietfed rats. During the fasting period, ghrelin mRNA expression in rats fed the protein-enriched diet was higher than that observed in rats on the standard diet (Fig. 1B). Two hours after feeding, mRNA decreased in animals on the highprotein diet and was even lower in rats receiving the standard diet (Fig. 1B).

4. Discussion Ghrelin secretion and expression were nutritionally manipulated. Fasting is known to increase plasmatic ghrelin in rodents [27] and, according to our results, we found that dietary content affects these increases. This finding indicates that ghrelin synthesis and secretion respond to dietary nutrients as well as to fasting [7]. Since after a fast, being fed returns ghrelin levels to the prefeeding values at 2– 6 h after feeding [28 –31] we believe that the ghrelin levels at these times were comparable to the levels existing before the fast. In addition, fasting would have magnified the amplitude of ghrelin secretion and thereby made it possible to detect possible variations in the plasma ghrelin concentrations due to diet more easily.

117

Interestingly, ghrelin was especially sensitive to the highprotein diet because the rats receiving this diet had the highest fasting ghrelin levels of all the groups and, together with the high-fat diet rats, the highest postfasting ghrelin levels. Ghrelin levels decrease after feeding before recovering and reaching a new peak in plasma [8,27,30,32]. In our study, ghrelin levels were decreased (40 – 60%) at 2 h after feeding and had begun recovering at 6 h, except in the rats on the high-protein diet. The increase in ghrelin levels 6 h after feeding was greater than 20% in all the diet groups except for animals on the high-protein diet; this indicated that the ghrelin levels in the rats after feeding tend to reach the levels that existed before feeding in all the dietary groups except for animals on the high-protein diet. The contradictory data on the effect of diet on ghrelin secretion are very probably the result of the diversity of experimental models used to study these phenomena [7,33 – 35]. Our experimental model has demonstrated that ghrelin secretion and synthesis were regulated by the high-protein diet with a controlled daily food caloric content (70 kcal/day) and volume (70 ml). Although endogenous ghrelin levels did vary with the different diets, there was not significant variation over the short term and no relation could be established between diet and ghrelin level. Nevertheless, several studies have suggested that ghrelin may be important in long-term control of food intake and body weight [36]. Although most studies have employed long feeding periods (4 – 14 weeks), studies on hormonal secretion regulation have used short adaptation to the diet (1 – 2 weeks) [5,37 – 39]. In addition, evidence of short-term intestinal adaptation to changes or injures have been reported [23,40]. The differences resulting from calorie content [7], the supply of other macronutrients [33], length of the diets administration [6] or the influence of other hormones, such as insulin [41], can affect the results for plasma ghrelin levels. It has been shown that a low-protein, high-carbohydrate diet has been shown to increase plasma ghrelin levels [33], probably due to the effect of carbohydrates on ghrelin secretion in conditions of negative energy balance [41]. Measurement of mRNA and protein expression in the gastric fundus proved that nutritional manipulation mainly regulated ghrelin biosynthesis and secretion, and this regulation was not affected by the metabolic balance or feeding conditions. The increase in ghrelin synthesis under the highprotein diet was corroborated by the high mRNA levels in the fundus of rats on this diet while intracellular protein decreased because fasting increases stomach ghrelin secretion while feeding decreases the secretion [27,33]. De novo ghrelin mRNA synthesis and plasma ghrelin levels decrease at 2 h after refeeding in rats on the high-protein diet; however, their ghrelin concentration in fundus at 2 and 6 h after refeeding is higher than in fasting conditions or in rats fed the standard diet. This ghrelin may have been stored until feeding triggers secretion. There is evidence indicating

118

M.T. Vallejo-Cremades et al. / Regulatory Peptides 121 (2004) 113–119

a positive relationship between trophic peptides and diet [23]. Previous studies have shown that the high-protein diet has a trophic effect on the intestine [25]. Our results further confirm this effect and the greatest length increase for villi and duodenal crypts was obtained in animals on the highprotein diet although no relation was found between diet and the proliferation index. Nonproliferative effects may be related to the low plasma levels of GH and IGF-1 in the rats receiving the high-protein diet. More exhaustive studies are needed to prove whether the trophic effect observed in rats on the high-protein diet is related with the high ghrelin levels induced by the diet or with the existence of some other direct effect on the intestine [42]. Plasma GH decreased with fasting as was expected because fasting abolishes the pulsatile pattern of GH secretion in the male rat [43], suggesting an underlying mechanism that would help conserve energy stores when adverse metabolic conditions prevail. In our study, pulsatile GH secretion was not measured because of the sampling time schedule but it was indirectly assessed through IGF-1. Fasting IGF-1 plasma levels were lower than those observed during the refeeding period; in addition, this hormone had the lowest plasma levels at any study time in rats fed with high-protein diet. Plasma IGF-1 increase some 10% at 2 – 3 days after being fed following a 48– 72 h fast [44], while our results showed an increase of 33% and 36% after a 24 h fast, suggesting that a shorter fast results in a faster response to feeding that a prolonged fast. In our model, the high-protein diet induced elevated ghrelin expression, but, contrary to what would be expected, the GH levels were lower than in the other dietary groups. Although Ghrelin induces GH secretion in vivo and in vitro [45,46], recent studies have shown that chronic ghrelin administration decreases GH secretion [45,47], probably due to resistance mechanisms similar to those affecting leptin secretion after long-term high-fat diets [48]. Therefore, a negative regulation by GH itself [49], ghrelin-related secretagogues, such as motilin [50], or by other neuroendocrine hormones, such as insulin [41], somatostatin (possibly regulated via leptin; [51,52]) or NPY [53] which might counteract the effect of ghrelin on GH expression, cannot be ruled out. In conclusion, our study demonstrates that diet macronutrient content can regulate ghrelin release in rats. The highest ghrelin plasma levels are found in fasting rats fed a protein-enriched diet, and this observation was further confirmed by ghrelin mRNA expression in their gastric fundus. On the other hand, the high-protein diet increases villi and crypt length, as previously reported [23]. Although small differences in GH were noted with the different diets, the biological relevance of these differences in intestinal trophic phenomena remains to be determined. Thus, the increased ghrelin secretion induced by fasting and elevated dietary protein may cause a signalling feedback among macronutrient intake, the GH – IGF-I axis and the intestinal trophic state. The mechanisms underlying these relations require further study.

References [1] Cummings DE, Shannon MH. Roles for ghrelin in the regulation of appetite and body weight. Arch Surg 2003;138:389 – 96. [2] Ariyasu H, Takaya K, Tagami T, Ogawa Y, Hosoda K, Akamizu T, et al. Stomach is a major source of circulating ghrelin, and feeding state determines plasma ghrelin-like immunoreactivity levels in humans. J Clin Endocrinol Metab 2001;86:4753 – 8. [3] 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 – 61. [4] Toshinai K, Date Y, Murakami N, Shimada M, Mondal MS, Shimbara T, et al. Ghrelin-induced food intake is mediated via the orexin pathway. Endocrinology 2003;144:1506 – 12. [5] Cummings DE, Purnell JQ, Frayo RS, Schmidova K, Wisse BE, Weigle DS. A preprandial rise in plasma ghrelin levels suggests a role in meal initiation in humans. Diabetes 2001;50:1714 – 9. [6] Tschop M, Weyer C, Tataranni PA, Devanarayan V, Ravussin E, Heiman ML. Circulating ghrelin levels are decreased in human obesity. Diabetes 2001;50:707 – 9. [7] Beck B, Musse N, Stricker-Krongrad A. Ghrelin, macronutrient intake and dietary preferences in long-evans rats. Biochem Biophys Res Commun 2002;292:1031 – 5. [8] Tschop M, Wawarta R, Riepl RL, Friedrich S, Bidlingmaier M, Landgraf R, et al. Post-prandial decrease of circulating human ghrelin levels. J Endocrinol Invest 2001;24:RC19 – 21. [9] Erdmann J, Lippl F, Schusdziarra V. Differential effect of protein and fat on plasma ghrelin levels in man. Regul Pept 2003;116:101 – 7. [10] Wren AM, Small CJ, Ward HL, Murphy KG, Dakin CL, Taheri S, et al. The novel hypothalamic peptide ghrelin stimulates food intake and growth hormone secretion. Endocrinology 2000;141:4325 – 8. [11] Tannenbaum GS, Epelbaum J, Bowers CY. Interrelationship between the novel peptide ghrelin and somatostatin/growth hormone-releasing hormone in regulation of pulsatile growth hormone secretion. Endocrinology 2003;144:967 – 74. [12] Thissen JP, Ketelslegers JM, Underwood LE. Nutritional regulation of the insulin-like growth factors. Endocrinol Rev 1994;15:80 – 101. [13] Thissen JP, Underwood LE, Ketelslegers JM. Regulation of insulin-like growth factor-I in starvation and injury. Nutr Rev 1999;57:167 – 76. [14] Kletke ML, Cummings DE, Scott Frayo D, et al. Evidence against an obligatory role for circulating ghrelin in physiological growth hormone regulation. Proceedings of the Endocrine Society’s 85th annual meeting; 19-22 June 2003; Philadelphia, USA. Chevy Chase, MD, USA: The Endocrine Society Press; 2003. p. 500. [15] Bluet-Pajot MT, Tolle V, Zizzari P, Robert C, Hammond C, Mitchell V, et al. Growth hormone secretagogues and hypothalamic networks. Endocrine 2001;14:1 – 8. [16] Wheeler EE, Challacombe DN. The trophic action of growth hormone, insulin-like growth factor-I, and insulin on human duodenal mucosa cultured in vitro. Gut 1997;40:57 – 60. [17] Beck PL, Rosenberg IM, Xavier RJ, Koh T, Wong JF, Podolsky DK. Transforming growth factor-beta mediates intestinal healing and susceptibility to injury in vitro and in vivo through epithelial cells. Am J Pathol 2003;162:597 – 608. [18] Dignass AU, Lynch-Devaney K, Podolsky DK. Hepatocyte growth factor/scatter factor modulates intestinal epithelial cell proliferation and migration. Biochem Biophys Res Commun 1994;202:701 – 9. [19] Hoffmann P, Dignass AU, Zeeh JM, Mazurkiewicz J, Holtmann G, Gerken G. Epithelial cell-derived components induce transforming growth factor-alpha mRNA expression and epithelial cell proliferation in vitro. Eur J Gastroenterol Hepatol 2001;13:1333 – 40. [20] Leblond C, Carriere R. The effect of growth hormone and thyroxine on the mitotic rate of the intestinal mucosa of the rat. Endocrinology 1955;56:261 – 6. [21] Ohneda K, Ulshen MH, Fuller CR, D’Ercole AJ, Lund PK. Enhanced

M.T. Vallejo-Cremades et al. / Regulatory Peptides 121 (2004) 113–119

[22]

[23]

[24]

[25] [26]

[27]

[28]

[29]

[30]

[31] [32]

[33]

[34]

[35]

[36]

[37]

growth of small bowel in transgenic mice expressing human insulinlike growth factor I. Gastroenterology 1997;112:444 – 54. Chen K, Nezu R, Inoue M, Wasa M, Iiboshi Y, Fukuzawa M, et al. Beneficial effects of growth hormone combined with parenteral nutrition in the management of inflammatory bowel disease: an experimental study. Surgery 1997;121:212 – 8. Vazquez I, Gomez-de-Segura IA, Grande AG, Escribano A, Gonzalez-Gancedo P, Gomez A, et al. Protective effect of enriched diet plus growth hormone administration on radiation-induced intestinal injury and on its evolutionary pattern in the rat. Dig Dis Sci 1999; 44:2350 – 8. Petersenn S. Growth hormone secretagogues and ghrelin: an update on physiology and clinical relevance. Horm Res 2002;58(Suppl. 3): 56 – 61. Miller AL. Therapeutic considerations of L-glutamine: a review of the literature. Altern Med Rev 1999;4:239 – 48. Kojima M, Hosoda H, Date Y, Nakazato M, Matsuo H, Kangawa K. Ghrelin is a growth-hormone-releasing acylated peptide from stomach. Nature 1999;402:656 – 60. Toshinai K, Mondal MS, Nakazato M, Date Y, Murakami N, Kojima M, et al. Upregulation of ghrelin expression in the stomach upon fasting, insulin-induced hypoglycemia, and leptin administration. Biochem Biophys Res Commun 2001;281:1220 – 5. Ariyasu H, Takaya K, Hosoda H, Iwakura H, Ebihara K, Mori K, et al. Delayed short-term secretory regulation of ghrelin in obese animals: evidenced by a specific RIA for the active form of ghrelin. Endocrinology 2002;143:3341 – 50. Bagnasco M, Dube MG, Kalra PS, Kalra SP. Evidence for the existence of distinct central appetite, energy expenditure, and ghrelin stimulation pathways as revealed by hypothalamic site-specific leptin gene therapy. Endocrinology 2002;143:4409 – 21. Tolle V, Bassant MH, Zizzari P, Poindessous-Jazat F, Tomasetto C, Epelbaum J, et al. Ultradian rhythmicity of ghrelin secretion in relation with GH, feeding behavior, and sleep – wake patterns in rats. Endocrinology 2002;143:1353 – 61. Tschop M, Smiley DL, Heiman ML. Ghrelin induces adiposity in rodents. Nature 2000;407:908 – 13. Murakami N, Hayashida T, Kuroiwa T, Nakahara K, Ida T, Mondal MS, et al. Role for central ghrelin in food intake and secretion profile of stomach ghrelin in rats. J Endocrinol 2002;174:283 – 8. Lee HM, Wang G, Englander EW, Kojima M, Greeley Jr GH. Ghrelin, a new gastrointestinal endocrine peptide that stimulates insulin secretion: enteric distribution, ontogeny, influence of endocrine, and dietary manipulations. Endocrinology 2002;143:185 – 90. Levin BE, Dunn-Meynell AA. Reduced central leptin sensitivity in rats with diet-induced obesity. Am J Physiol Regul Integr Comp Physiol 2002;283:R941 – 8. Schaffhauser AO, Madiehe AM, Braymer HD, Bray GA, York DA. Effects of a high-fat diet and strain on hypothalamic gene expression in rats. Obes Res 2002;10:1188 – 96. Wren AM, Small CJ, Abbott CR, Dhillo WS, Seal LJ, Cohen MA, et al. Ghrelin causes hyperphagia and obesity in rats. Diabetes 2001; 50:2540 – 7. Chin E, Bondy CA. Dietary protein-induced renal growth: correlation between renal IGF-I synthesis and hyperplasia. Am J Physiol 1994; 266:C1037 – 45.

119

[38] Dardevet D, Manin M, Balage M, Sornet C, Grizard J. Influence of low- and high-protein diets on insulin and insulin-like growth factor-1 binding to skeletal muscle and liver in the growing rat. Br J Nutr 1991;65:47 – 60. [39] Smith SR, de Jonge L, Zachwieja JJ, Roy H, Nguyen T, Rood JC. Fat and carbohydrate balances during adaptation to a high-fat. Am J Clin Nutr 2000;71:450 – 7. [40] Gomez dS I, Castell T, Vazquez I, Mata A, De Miguel E. Somatostatin antagonism prevents elemental diet-induced intestinal atrophy in the rat. Dig Dis Sci 2001;46:1985 – 92. [41] Flanagan DE, Evans ML, Monsod TP, Rife F, Heptulla RA, Tamborlane WV, et al. The influence of insulin on circulating ghrelin. Am J Physiol Endocrinol Metab 2003;284:E313 – 6. [42] King IS, Paterson JY, Peacock MA, Smith MW, Syme G. Effect of diet upon enterocyte differentiation in the rat jejunum. J Physiol 1983; 344:465 – 81. [43] Harel Z, Tannenbaum GS. Long-term alterations in growth hormone and insulin secretion after temporary dietary protein restriction in early life in the rat. Pediatr Res 1995;38:747 – 53. [44] Frystyk J, Delhanty PJ, Skjaerbaek C, Baxter RC. Changes in the circulating IGF system during short-term fasting and refeeding in rats. Am J Physiol 1999;277:E245 – 52. [45] Date Y, Murakami N, Kojima M, Kuroiwa T, Matsukura S, Kangawa K, et al. Central effects of a novel acylated peptide, ghrelin, on growth hormone release in rats. Biochem Biophys Res Commun 2000;275: 477 – 80. [46] Tolle V, Zizzari P, Tomasetto C, Rio MC, Epelbaum J, Bluet P. In vivo and in vitro effects of ghrelin/motilin-related peptide on growth hormone secretion in the rat. Neuroendocrinology 2001;73:54 – 61. [47] Yamazaki M, Nakamura K, Kobayashi H, Matsubara M, Hayashi Y, Kangawa K, et al. Regulational effect of ghrelin on growth hormone secretion from perifused rat anterior pituitary cells. J Neuroendocrinol 2002;14:156 – 62. [48] Frederich RC, Hamann A, Anderson S, Lollmann B, Lowell BB, Flier JS. Leptin levels reflect body lipid content in mice: evidence for dietinduced resistance to leptin action. Nat Med 1995;1:1311 – 4. [49] Minami S, Kamegai J, Sugihara H, Suzuki N, Wakabayashi I. Growth hormone inhibits its own secretion by acting on the hypothalamus through its receptors on neuropeptide Y neurons in the arcuate nucleus and somatostatin neurons in the periventricular nucleus. Endocrinol J 1998;45 Suppl.:S19 – 26. [50] Folwaczny C, Chang JK, Tschop M. Ghrelin and motilin: two sides of one coin? Eur J Endocrinol 2001;144:R1 – 3. [51] Arosio M, Ronchi CL, Gebbia C, Cappiello V, Beck-Peccoz P, Peracchi M. Stimulatory effects of ghrelin on circulating somatostatin and pancreatic polypeptide levels. J Clin Endocrinol Metab 2003;88: 701 – 4. [52] Scanlon MF, Issa BG, Dieguez C. Regulation of growth hormone secretion. Horm Res 1996;46:149 – 54. [53] Seoane LM, Lopez M, Tovar S, Casanueva FF, Senaris R, Dieguez C. Agouti-related peptide, neuropeptide Y, and somatostatin-producing neurons are targets for ghrelin actions in the rat hypothalamus. Endocrinology 2003;144:544 – 51.