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Endocrinology Vol. 145, No. 5 2197-2205
Copyright © 2004 by The Endocrine Society

Characterization of the Rhesus Monkey Ghrelin Gene and Factors Influencing Ghrelin Gene Expression and Fasting Plasma Levels

Stephen V. Angeloni, Nicole Glynn, Grazia Ambrosini, Michael J. Garant, J. Dee Higley, Stephen Suomi and Barbara C. Hansen

The Center for Vaccine Development (S.V.A.), The Obesity and Diabetes Research Center (G.A., B.C.H.), Division of Endocrinology, Diabetes, and Nutrition (N.G., M.J.G.), University of Maryland School of Medicine, Baltimore, Maryland 21201; and the National Institute on Alcohol Abuse and Alcoholism, Laboratory of Clinical Studies-Primate Section Unit (J.D.H.), National Institute of Child Health and Human Development, Laboratory of Comparative Ethology (S.S.), National Institutes of Health, Poolesville, Maryland 20837

Address all correspondence and requests for reprints to: Stephen V. Angeloni, Ph.D., Center for Vaccine Development, University of Maryland School of Medicine, 685 West Baltimore Street, Room 480, Baltimore, Maryland 21201-1509. E-mail: sangelon{at}medicine.umaryland.edu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Ghrelin stimulates release of GH from the pituitary, stimulates appetite, and may influence metabolic processes in other tissues expressing the GH secretagogue receptor. Ghrelin can thus influence behaviors and endocrine pathways contributing to weight gain. In this study we characterized the ghrelin gene from the rhesus monkey and analyzed the association of plasma ghrelin levels with metabolic and endocrine markers. Rhesus ghrelin is 97, 91, and 96% homologous to the human cDNA, gene, and peptide, respectively. Ghrelin expression was highest in the stomach with lower levels found in muscle and duodenum. In these tissues, ghrelin expression in calorie-restricted and obese animals was about 40–99% lower than in lean animals. In addition, ghrelin expression in muscle was fairly high and may allow this tissue to contribute significantly to plasma levels. Fasting plasma ghrelin concentrations were also inversely correlated with body mass index and exhibited a nonlinear association with age with increased levels in younger and older monkeys and lower levels in middle-aged monkeys. Although a significant inverse correlation between fasting plasma ghrelin and fasting insulin levels were found, iv glucose and insulin administration did not significantly alter ghrelin levels. These studies demonstrate that ghrelin levels are influenced by age-related factors and adiposity in the rhesus monkey. These similarities between the rhesus monkey and human ghrelin genes and plasma ghrelin responses suggest a unique opportunity to study the mechanisms regulating ghrelin secretion and gene expression in different tissues in normal and disease states using this model system.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GHRELIN AND GHRH are the two known natural ligands for the GH secretagogue receptor (GHS-R). Ghrelin was first characterized in the rat as a prepropeptide hormone of 117 amino acids, which is processed to an active 28-amino acid peptide with a 3-n-octanoylated serine at position 3 (1). Stomach is the major source of circulating ghrelin as shown by immunohistochemistry (2), in gene expression studies (1), and by the reduction to 35% of plasma ghrelin levels in totally gastrectomized patients (3). Ghrelin has also been reported to be expressed in low amounts in various tissues such as duodenum, kidney, placenta, and pituitary in humans and rodents (4) and has been shown to be expressed in many human tissues (5). GHS-R (6) and its subtypes are expressed not only in the hypothalamus and pituitary (7, 8, 9, 10) but also in tissues such as adipose, pancreas, liver, heart, and endothelial cells (5, 11). In addition to its GH-releasing properties, ghrelin administration exerts a potent orexigenic response, even in ghrelin knockout mice and can produce weight gain in rodents (6, 12, 13, 14). As in rodents, iv administration of ghrelin to healthy human volunteers substantially increased the intake of a subsequent meal (15). These effects in rodents and humans may be via the antagonizing of leptin action through activation of the hypothalamic neuropeptide Y/Y1 receptor pathway (12), although the exact orexigenic mechanism of ghrelin is not yet known.

Ghrelin’s role in regulating appetite is supported by studies that reported an increase in plasma concentration before a meal and rapid decrease after eating (3, 16), a response that is lost after gastric bypass surgery (17, 18). Whereas the mechanism regulating fluctuations in plasma ghrelin levels around meals is not well understood, some studies have indicated that glucose and/or insulin can lower ghrelin levels (19, 20, 21, 22). Caixas et al. (23) found that a meal or oral glucose consumption induced reductions in ghrelin but did not find an effect of parenteral glucose or insulin, and Schaller et al. (24) concluded that the meal-related suppression of ghrelin was not a direct effect of either glucose or insulin regulation. Recently ghrelin was reported to up-regulate gluconeogenesis and affect insulin signaling in hepatoma cells (25), suggesting peripheral effects of ghrelin on glucose homeostasis. Contrary to these findings, Sun et al. (13) did not see any significant change in feeding behavior or other metabolic-related changes in ghrelin-null mice.

In humans, plasma ghrelin levels were found to be inversely related to body fat content, with obese subjects having lower fasting plasma ghrelin concentrations than lean individuals (26, 27). The suppression of fasting ghrelin levels by increased body fat content can be reversed by weight loss as a result of gastric bypass surgery (17), participation in a weight loss program (28), or calorie restriction as seen in rodents (29). Conversely, plasma ghrelin levels are elevated in anorexia nervosa (3) and return to normal levels after partial weight recovery (30). The link of ghrelin to metabolic disorders is also supported by analysis of ghrelin mutations. Ukkola et al. (31) reported that the mutation at codon 72 outside the coding region of the human mature ghrelin peptide tended to be associated with lower age onset of obesity.

These data support the importance of ghrelin in regulating behavioral and possibly metabolic responses that have a significant impact on energy homeostasis and illustrate the complexity of the mechanisms regulating plasma ghrelin levels. In this study we examined the genetic and physiological characteristics of the rhesus monkey ghrelin gene and parameters associated with changes in ghrelin plasma levels. We hypothesized that the setting of ghrelin fasting plasma levels are mediated by factors influencing changes in ghrelin gene expression levels. To study this hypothesis, we used tissue and plasma samples from a colony of rhesus monkeys with well-characterized physiological, metabolic and anthropometric traits associated with aging, obesity, and the onset of type 2 diabetes (32, 33, 34, 35, 36, 37). We isolated and characterized the coding region of the rhesus monkey ghrelin mRNA and compared its predicted amino acid sequence with that of human, rat, and mouse. The monkey ghrelin gene was cloned and compared with the human homolog. Using RT-PCR methods, the tissue distribution of ghrelin expression was determined and expression levels compared between lean and obese animals. Total fasting plasma ghrelin levels were compared with anthropometric and metabolic markers such as age, body weight, body mass index (BMI), and insulin in four groups of monkeys: 1) ad libitum-fed lean, 2) ad libitum-fed obese, 3) long-term calorie-restricted (CR), and 4) spontaneous natural type 2 diabetic animals. In addition, we analyzed plasma ghrelin levels during euglycemic-hyperinsulinemic clamps and iv glucose tolerance tests to determine whether insulin and glucose influence plasma ghrelin levels.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
Eighty-three rhesus macaques (Macaca mulatta) from the Obesity and Diabetes Research Center at the University of Maryland School of Medicine (Baltimore) and the NIH primate colony (Poolesville, MD) were used for these studies. The ad libitum population, ranging in age from 3 to 33 yr, included 49 normal lean adult monkeys, 16 normal obese adult animals (body fat content over 22%), and 12 diabetic monkeys that spontaneously developed type 2 diabetes during adulthood. BMI was calculated as the weight in kilograms divided by the square of the crown-to-rump height in meters (kg/m2) and ranged from 4.4 and 37.2 kg/m2 for the animals used in this study. Six of the animals in this study have been calorie restricted for over 20 yr on a diet designed to maintain their body weight at the mature lean adult level of 10 kg with a body fat composition of 17–22% (a normal body weight clamp maintained by constant calorie adjustments) (34). Monkeys were provided water ad libitum, whether or not they were caloric restricted and maintained in individual stainless steel cages in accordance with the recommendations of the NIH Guide for the Care and Use of Laboratory Animals. All procedures for handling, treatment, and sample collection were approved by the University of Maryland School of Medicine’s Institutional Animal Care and Use Committee.

Blood plasma procedures
All fasting blood samples were collected under 10 mg/kg ketamine anesthesia after a 16-h fast. A glucose dose of 0.25 mg/kg was administered during iv glucose tolerance tests (IVGTT) (35), and euglycemic-hyperinsulinemic clamps were performed as previously described (38). Briefly, a continuous infusion of insulin was administered to produce maximally stimulated circulating insulin levels (> 3000 µU/ml), and a variable infusion of a 20% glucose solution was started at time 0 and periodically adjusted to clamp the plasma glucose concentration between 80 and 90 mg/dl. Plasma glucose levels were determined using the glucose oxidase assay measured on a Beckman glucose analyzer 2 (Beckman Instruments Inc., Fullerton CA). Analysis of total plasma ghrelin (RIA, pg/ml), primate leptin (ELISA, ng/ml), and human insulin (ELISA, µU/ml) was performed by Linco Diagnostics (St. Charles, MO). The inter- and intraassay coefficients of variation for ghrelin and insulin were 6.7 and 16.3% and 4.5 and 3.3%, respectively. Conversion of ghrelin values from picograms per milliliter to picomoles per liter was done by multiplying picograms per milliliter by 0.296 (17). The number of fasting plasma ghrelin levels determined for each monkey ranged from one to three samples. When more than one value was obtained, they were averaged for each monkey, resulting in a single data point.

Cloning and sequencing of the M. mulatta ghrelin gene
Isolation of RNA from monkey tissues was performed using standard Trizol methods (Invitrogen, Inc., Carlsbad, CA). RNA quality was checked with a cDNA integrity kit according to the manufacture’s directions (KPL, Gaithersburg, MD). Reverse transcriptase (RT) reactions were performed using Superscript II RT (Invitrogen) and PCRs were performed using primers derived from the human ghrelin nucleotide sequence (accession no. BC025791) as follows: forward primer 5'-CCATGCCCTCCCCAGGGACC-3' and reverse primer 5'-ATCACTTGTCGGCTGGGGCCTC-3'. All oligonucleotide primers were synthesized by MWG (High Point, NC). Hot-start PCR using ThermalAce DNA polymerase (Invitrogen) was performed with 30 cycles of 95 C for 30 sec, 60 C for 30 sec, and 72 C for 1 min, plus a final extension time of 2 min at 72 C and held at 4 C. The resulting 354-nucleotide product was cloned into pCR 4-TOPO sequencing vector (Invitrogen) and sequenced using the Big Dye Terminator sequencing kit, version 2.0 (ABI Systems, Applied Biosystems, Foster City, CA). Nucleotide sequence analysis and contiguous sequence construction was preformed using Sequencher (Gene Codes Corp., Ann Arbor, MI) and analysis of amino acid sequence homology was performed using the IBM Bioinformatics Group multiple alignment program (http://cbcsrv.watson.ibm.com/Tmsa.html).

Genomic segments of the ghrelin gene were obtained by PCR using genomic DNA. Genomic DNA was isolated from muscle using the procedure of Miller et al. (39) with added phenol:chloroform and chloroform/isoamyl alcohol extractions before the ethanol precipitation step. Primers for genomic PCR were designed from coding regions of the rhesus cDNA sequence. PCR products were cloned into pCR 4-TOPO and verified by sequence analysis as described above. Flanking regions of the ghrelin gene were cloned using the Universal Genome Walker kit (Clonetech, Palo Alto, CA) according to the manufacturer’s directions. This method involves using four separate restriction-digested genomic libraries ligated to the Universal Genome Walker adaptor. The adaptor forward primer, AP1F, was 5'-GTAATACGACTCACTATAGGGC-3' and the adaptor reverse primer, AP1R, was 5'-GCCCTATAGTGAGTCGTATTA-3'. Gene-specific primers were used along with the corresponding AP1 primer for PCR. The 5' upstream genomic region was obtained by PCR from these libraries with the appropriate primer pairs. PCR products were isolated, cloned, and verified by sequence analysis as described above.

Ghrelin tissue distribution and mRNA expression
Analysis of ghrelin expression in different tissues by RT-PCR was performed using the Titan One-Tube RT-PCR system (Roche Applied Science, Nutley, NJ), and monkey total RNA was isolated using RNA STAT-60 according to the manufacturer’s directions (Tel-Test Inc., Friendswood, TX). Primers for ghrelin gave products from exon 1 to exon 3. Primers for ß-actin were used as a control. The RT reaction contained 1 µg total RNA and the following primers: 5'-CTGGCTGGACTTGGCCATGGCAGGCTCCAGCTTCCTGAGCC-3' and 5'-GAAGAAACTTCCCCAGGGCCTGGCTGTGCTGCTGGTACTG-3' for ghrelin or 5'-GCTACGAGCTGCCTGACGGCCAGG-3' and 5'-GAAGCATTTGCGGTGGACGATGG-3' for ß-actin. Cycling conditions were as follows: for ghrelin, 94 C for 2 min and then 25 cycles at 94 C for 30 sec, 60 C for 30 sec and 68 C for 30 sec. For ß-actin, cycling conditions were 94 C for 2 min and then 25 cycles at 94 C for 30 sec, 50 C for 30 sec, and 68 C for 1 min. The resulting PCR products were loaded onto a 1.5% agarose gel and visualized by ethidium bromide staining.

Real-time RT-PCR analyses of differences in ghrelin expression were performed as follows. Total RNA was isolated using RNA STAT-60. The RT reactions were carried out in 96-well plates using QuantiTect RT-PCR kit (Qiagen, Valencia, CA). The PCR were performed in triplicate using iQ SYBR Green Supermix (Bio-Rad Laboratories, Hercules, CA). The samples were run in identical well positions for the ghrelin and ß-actin PCR. The ß-actin reactions were run with the primers listed above, whereas the ghrelin primers were: forward primer, 5'-GGGCGGAGGATGAACTGGAA-3', and reverse primer, 5'-CCTGGCTGTGCTGCTGGTA-3'. The PCR conditions for ghrelin and ß-actin were 95 C for 5 min for one cycle and then 94 C for 20 sec, 60 C for 30 sec, and 72 C for 30 sec for 40 cycles on an iCycler (Bio-Rad). Data were collected during the 60 C extension step. The amount of transcript was calculated from amplicon-specific standard curves of stomach total RNA. Ghrelin levels in each sample were normalized to the actin level in that sample, and the relative quantitation of ghrelin mRNA expression was calculated by comparing the ghrelin/ß-actin ratios as percent expression between lean, CR, and obese animals.

Statistical analysis
Results are shown as means ± SE. The relationship of ghrelin with age, BMI, glucose, and insulin were evaluated by regression analysis, allowing for first- and second-order effects (SAS Institute Inc., Cary, NC). Statistically significant differences among diabetic, normal, or CR monkeys were examined by an unpaired Student’s two-tailed t test.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Sequence analysis of rhesus ghrelin
The nucleotide sequences for the coding region of the monkey ghrelin mRNA and the predicted prepropeptide sequence were determined (Fig. 1AGo). The monkey prepropeptide cDNA sequence of ghrelin is 97 and 86% homologous to that of the human and rat, respectively (data not shown). Sequence analysis of the monkey ghrelin gene shows it to have the same exon and intron structure as the human gene (Fig. 1BGo) and is 91% homologous to the human gene including approximately 1.7 kb of sequence upstream of the ATG start codon.



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  		FIG. 1. Rhesus ghrelin sequence. The nucleotide sequences were determined for the coding region of the monkey ghrelin mRNA and for about 4000 bp of the monkey gene. A, The 354 nucleotide coding sequence of rhesus monkey preproghrelin (GenBank accession no. AY371699). The predicted amino acid sequence of the prepropeptide is shown with the mature 28-amino acid sequence of ghrelin indicated with bold italic letters. B, Gene structure of monkey ghrelin, not to scale (GenBank accession no. AY372274). Sizes of exon coding regions and noncoding regions (NCRs) such as introns and the gene sequence upstream of the ATG start site are indicated.

 
The predicted monkey preproghrelin amino acid sequence is 96, 83, and 84% homologous to human, rat, and mouse preproghrelin sequences, respectively (Fig. 2Go). The mature 28-amino acid protein is 96% homologous between monkey and all species studied, resulting from single amino acid differences. Compared with human ghrelin, monkey ghrelin has an A35V substitution and an N34K substitution compared with the rodent amino acid sequence. These are conservative amino acid differences, suggesting that the mature 28-amino acid peptide structure and function is maintained.



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  		FIG. 2. Alignment of predicted amino acid sequence. The predicted amino acid sequence for rhesus ghrelin was compared with human, rat, and mouse. Amino acid differences are noted in bold, lower-case lettering. The mature 28-amino acid peptide is underlined. Amino acid sequence alignments were performed using the IBM Bioinformatics Group multiple alignment program. Sequences are derived from the translation of human (accession no. NM_016362), rat (accession no. Q9QYH7), and mouse (accession no. BAB19046) GenBank sequences.

 
Tissue distribution and mRNA expression
Analysis of the tissue distribution of ghrelin expression was performed by RT-PCR from multiple tissues of a nondiabetic, nonobese monkey. Using this method, ghrelin mRNA was most abundant in stomach and was expressed at lower levels in muscle and duodenum (Fig. 3AGo). We were not able to detect ghrelin expression in brain, aorta, pancreas, ileum, and omental fat or sc fat by RT-PCR. This does not rule out the possibility that the level of expression of the transcript was below the detectable limits of the RT-PCR assay. Ghrelin mRNA expression was also examined by real-time PCR in stomach, muscle, and duodenum of CR, lean normal, obese normal, and diabetic monkeys. Tissues from as many as two monkeys of each group were analyzed. These data show that for each tissue, expression was lowest in obese animals and highest in lean animals (Fig. 3BGo). Surprisingly, CR animals had levels between that of obese and lean. Ghrelin expression was also highest in stomach, regardless of health or nutrition status (Table 1Go, percent in health-nutrition status). However, when ghrelin expression levels are compared with the highest expressing tissue, lean stomach, the tissues with the next highest levels of expression are CR stomach (63%) followed by lean muscle (44.73%) and obese stomach (44.6%) (Table 1Go, percent of lean stomach). These data suggest that muscle may contribute significantly to ghrelin plasma levels.



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  		FIG. 3. Analysis of ghrelin expression. RT-PCR analysis was performed to determine tissue distribution and expression patterns. A, RT-PCR analysis of tissue distribution. Samples in each lane are marker (1), brain (2), aorta (3), muscle (4), stomach (5), pancreas (6), duodenum (7), ilium (8), omental fat (9), and sc fat (10). B, Real-time PCR analysis of ghrelin expression levels in stomach, skeletal muscle, and duodenum of lean (open bar), obese (solid bar), and CR (gray bar) animals.

 

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  		TABLE 1. Real time RT-PCR analysis of ghrelin expression levels

 
Fasting plasma ghrelin concentrations
Fasting plasma ghrelin concentrations ranged from 573 to 3878 pg/ml, with a mean concentration of 1738 ± 84 pg/ml. In humans, plasma ghrelin concentrations have been reported to be inversely correlated with BMI (22, 27), whereas plasma leptin is positively correlated with BMI (40). To determine whether these relationships exist in the rhesus monkeys, fasting plasma ghrelin concentrations were plotted as a function of BMI. Consistent with previous findings in humans, plasma ghrelin was inversely correlated with BMI in the rhesus monkey (Fig. 4Go). Although leptin and ghrelin have been reported to be inversely regulated in humans (41, 42, 43, 44), we did not find a statistically significant correlation between the fasting plasma levels of these two hormones (data not shown).



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  		FIG. 4. Relationship between plasma ghrelin levels and BMI. Average fasting plasma ghrelin levels from a monkey population ranging from normal to diabetic and ranging in age from 3 to 33 yr, compared with BMI (kg/m2, n = 83, R2 = 0.04, P = 0.08).

 
Fasting plasma ghrelin concentrations were previously reported to change with age in humans as children mature to adulthood (16, 26, 45). To examine the relationship between age and ghrelin levels in monkeys, fasting plasma ghrelin concentrations were analyzed as a function of age over a range of 3–33 yr. This analysis revealed a nonlinear relationship between age and fasting ghrelin levels (Fig. 5AGo). To further statistically clarify this relationship, the data were stratified into two age groups, young (3–10 yr) and older adult (over 11 yr). For young monkeys, fasting plasma ghrelin levels were inversely related to age (Fig. 5BGo) as previously reported in human studies (26); however, a statistically significant gradual increase was found going from middle-aged to elderly monkeys (Fig. 5CGo).



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  		FIG. 5. Ghrelin plasma levels compared with age. Relationship between fasting plasma ghrelin levels and age from all study animals over different age ranges. A, Analysis in all animals from 3 to 33 yr (n = 83, R2 = 0.15, P = 0.04). B, Analysis of animals from 3 to 10 yr (n = 33, R2 = 0.16, P = 0.02). C, Analysis of animals from 11 to 33 yr (n = 50, R2 = 0.17, P = 0.003). Best-fitting model for A included age and age2.

 
Plasma ghrelin level regulation by glucose
Prior reports provide conflicting evidence concerning whether iv glucose administration could affect plasma ghrelin levels (19, 22, 24). To explore this issue, IVGTTs were performed on CR, normal nondiabetic, and diabetic ad libitum-fed monkeys. Administration of glucose to monkeys, with its concomitant increase in insulin secretion, induced no significant changes in plasma ghrelin levels within or between the groups studied (Fig. 6Go).



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  		FIG. 6. Glucose regulation of plasma ghrelin levels. Plasma samples from CR (n = 5), normal nonobese ad libitum (n = 9), and diabetic (n = 5) monkeys during IVGTTs. Fasting (solid bar), 30 (gray bar) and 60 (open bar) min post glucose administration values are shown. SEs are indicated.

 
Plasma ghrelin level regulation by insulin
Studies have shown that plasma ghrelin levels fluctuate on a diurnal basis with a secretion pattern reciprocal to that of insulin in humans (16, 46), and preprandial peaks in ghrelin levels occur with rapid decreases after a meal. To test whether ghrelin was regulated by insulin, fasting plasma levels of ghrelin and insulin were compared and showed a small but significant inverse correlation (Fig. 7Go) that did not differ between younger and older groups (data not shown). Ghrelin levels were also determined in CR and nondiabetic ad libitum-fed rhesus monkeys subjected to a euglycemic-hyperinsulinemic clamp. No significant changes in plasma ghrelin levels in response to maximal insulin stimulation during the euglycemic-hyperinsulinemic clamp were observed (Fig. 8Go). Whereas there were also no significant differences in ghrelin levels between CR and lean animals, obese animals had the lowest fasting and insulin stimulated levels of the three groups.



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  		FIG. 7. Fasting plasma ghrelin levels compared with insulin concentration. Fasting plasma ghrelin levels compared with the natural log (In) of fasting plasma insulin concentrations on a monkey population ranging from normal to diabetic (n = 63; R2 = 0.06; P < 0.05).

 


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  		FIG. 8. Insulin regulation of plasma ghrelin levels. Plasma samples were collected during euglycemic-hyperinsulinemic clamps from CR animals (n = 4), and ad libitum-fed nondiabetic animals that were lean (n = 11) or obese (n = 3). Fasting (solid bars) and insulin stimulated (open bars) ghrelin levels are shown and SEs indicated.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Numerous studies have reported that supraphysiological stimulation with ghrelin increases food intake in rodents and humans (14, 15, 47, 48). In ghrelin-null mice, ghrelin administration increased food intake, but the lack of functional ghrelin did not effect other facets of mouse physiology (13). Yet the presence of GHS-Rs in a wide range of tissues allows ghrelin to play a more direct role in other facets of energy metabolism by affecting adipogenesis (49) and insulin signaling (25). These characteristics define ghrelin as an important but not an essential factor in the regulation of energy homeostasis.

The importance of conserving ghrelin function across species is exemplified by the high degree of homology (96%) of the mature 28-amino acid peptide among monkeys, humans, and rodents, resulting from a single amino acid difference. Our data demonstrate that the rhesus ghrelin prepropeptide has 96, 83, and 84% homology to human, rat, and mouse, respectively. Although the mature ghrelin is well conserved, differences in the precursor amino acid sequence may affect the hormone’s posttranslational processing among species. Monkey ghrelin cDNA is 97% homologous to the human cDNA sequence. The intron/exon structure of the monkey ghrelin gene was identical with the human gene, with a 91% homology at the nucleotide level.

In this study we also elucidated the tissue distribution of ghrelin gene expression in rhesus monkeys. Monkey ghrelin mRNA was highly expressed in the stomach, whereas muscle and duodenum expressed low levels of the transcript. This pattern of expression is in agreement with studies done in human (3) and rat (1) tissues by Northern blot analysis. In contrast, Gnanapavan et al. (5) reported a broader tissue distribution for ghrelin in humans using a more sensitive real-time PCR method. In humans and rodents, feeding behavior (9, 14, 50) and adiposity (27) have been correlated with changes in ghrelin mRNA expression in the stomach (51). In the present study, ghrelin gene expression was examined in tissues from long-term CR and ad libitum-fed lean and obese monkeys that were either normal or diabetic. Independent of disease status, real time RT-PCR analysis showed that ghrelin gene expression in each tissue was highest in lean animals and lowest in obese animals, whereas CR animals had levels higher than obese but significantly lower than lean animals. These results indicate that whether normal or diabetic, increased body fat content corresponds to decreased expression of ghrelin in stomach, muscle, and duodenum. Comparison of ghrelin expression levels in various tissues to the highest expressing tissue (lean stomach) showed that expression was 36% lower in the CR stomach and about 45% lower in obese stomach and lean muscle. The relatively high expression in muscle is interesting because muscle is the body’s most abundant tissue and highest consumer of energy and may indicate that muscle plays a significant role in contributing to ghrelin plasma levels. These changes in ghrelin expression in muscle and duodenum may represent a physiological adaptation to the positive energy balance associated with obesity (27, 52). This is supported by monkey fasting plasma ghrelin levels that were significantly lower in animals with higher BMI, a result similar to those reported in humans and rodents (22, 26, 27, 44, 53).

Age also appears to have a significant influence on fasting plasma ghrelin levels (16, 26). Interestingly, our data indicate that fasting plasma ghrelin levels have a significant nonlinear correlation with age across the entire life span, with fasting plasma levels being higher in younger and older monkeys and lower in middle-aged monkeys, similar to observations made in previous human studies (16, 45). Regulation of age-dependent plasma ghrelin levels may be the result of changes in body fat content or hormonal changes that occur during the aging process. During the aging process, levels of a number of hormones change with age. For example several studies have shown that GH secretion decreases with age in both humans and monkeys (54, 55), and age-related decreases of GH in old female rhesus monkeys correlate with a decrease of circulating GHRH and an increase of somatostatin release from the hypothalamus (56). Ghrelin strongly stimulates GH release via the GHS-Rs, and this response is markedly higher in younger human subjects than elderly subjects (57). Although we did not measure GH levels in our study, we speculate that GH would decrease in older monkeys as reported in humans due to reduced GHS-R expression in the aged hypothalamus (58) and the age-related decrease in number and size of somatotroph cells in the pituitary (59). In some older individuals, these or other hormonal changes could result in alterations of ghrelin sensitivity or ghrelin expression and may contribute to the slight increase in fasting ghrelin levels seen in some of the elderly.

Several studies indicate that ghrelin and leptin exert opposite actions in the regulation of food intake (30, 42). Ghrelin promotes hunger and adipogenesis (48), whereas leptin elicits satiety and adipolysis (60, 61, 62, 63). Circulating levels of leptin and ghrelin have been previously shown to be inversely correlated in humans and rodents (27, 42). However, our observations in monkeys showed no relationship between fasting plasma ghrelin and plasma leptin concentrations. This result may be attributable to the heterogeneity of our study population in terms of adiposity, insulin resistance, and age.

In addition to the mechanisms responsible for establishing fasting plasma ghrelin levels, daily fluctuations of ghrelin levels have been observed in response to meals (16). Whereas these fluctuations are mediated by changes in secretion rather than changes in expression levels, the exact molecular mechanisms involved are not known. Some studies (20, 21, 22, 64, 65) have suggested that these variations may be controlled by changes in glucose and/or insulin levels, whereas others (23, 24) could not demonstrate an effect on ghrelin levels after a glucose or insulin infusion. Some of these differences in the reported literature result from differences in the BMI of the populations studied as well as differences in the amount and mode of glucose administration. Our data showed that glucose administration during an IVGTT to CR and ad libitum-fed monkeys (including lean and obese nondiabetic monkeys) did not induce a significant decrease in ghrelin levels at 30 or 60 min. Shiiya et al. (22) reported a rapid decrease of ghrelin within 15 min after iv glucose administration followed by an immediate return to basal levels. To see a response in this animal model, we may need to look for changes in ghrelin levels before 30 min combined with increasing the amount of glucose administered during an IVGTT.

Our data also show that fasting plasma ghrelin levels have a significant inverse correlation with fasting insulin concentrations. Nevertheless, high circulating insulin during a euglycemic-hyperinsulinemic clamp did not affect ghrelin levels in any group tested. During the revision of this manuscript, Murdolo et al. (20) showed that insulin could stimulate a more dramatic decrease in plasma ghrelin after a meal in type 1 diabetics within the first 30–60 min of insulin administration. The absence of a suppressive effect of insulin administration (as well as glucose administration) on ghrelin levels in our monkeys may suggest the need to monitor changes at earlier time points as noted above and that the presence of food in the stomach and intestine, or other factors, triggers the suppression of ghrelin release after a meal (23).

In summary, we have determined the genomic and coding sequences of ghrelin from the rhesus monkey and have shown them to be highly conserved between monkeys and humans. Analysis of fasting ghrelin levels resulted in a nonlinear correlation with age and an inverse correlation with BMI and insulin, similar to findings in humans. However, fasting ghrelin levels were not related to plasma leptin levels and were not affected by glucose during an IVGTT or by insulin during a euglycemic-hyperinsulinemic clamp. In addition, the finding that muscle expression of ghrelin is dramatically affected by body fat content may suggest that muscle plays an active role in energy homeostasis through ghrelin’s orexigenic actions. Further studies of the tissue-specific regulation of ghrelin gene expression may help elucidate the possible mechanisms involved in regulating ghrelin expression and the role of ghrelin and other tissues in the short- and long-term regulation of energy homeostasis. In addition, the genetic and physiological similarities between the human and monkey ghrelin sequences and factors influencing plasma levels should allow this model to serve as a useful tool to study the mechanisms regulating ghrelin gene expression, secretion, and impact on energy homeostasis under normal and disease states.


   Acknowledgments
 
We thank Dr. Rongze Yang and Dr. Da-Wei Gong for initial monkey RNA samples; Braxton D. Mitchell, Kathy Ryan, and Toni Pollin for statistical analysis; and Stephen G. Lindell, Tami L. Gura, Kimberly Wojteczko, Courtney Shannon, Kenneth Vaughan, and Pete Roma for their assistance with collecting samples from the animals in Poolesville.


   Footnotes
 
This work was supported by National Institutes of Aging Grant NIA NOI-AG-0-2100 (to B.C.H.).

Abbreviations: BMI, Body mass index; CR, calorie-restricted; GHS-R, GH secretagogue receptor; IVGTT, iv glucose tolerance test; RT, reverse transcriptase.

Received August 25, 2003.

Accepted for publication January 13, 2004.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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