Molecular Forms of Hypothalamic Ghrelin and Its Regulation by Fasting and 2-Deoxy-D-Glucose Administration
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内分泌学杂志 2005年第6期
Molecular Genetics, Institute of Life Sciences, Kurume University, Fukuoka 839-0864, Japan
Address all correspondence and requests for reprints to: Masayasu Kojima, Ph.D., Molecular Genetics, Institute of Life Sciences, Kurume University, Fukuoka 839-0864, Japan. E-mail: mkojima@lsi.kurume-u.ac.jp.
Abstract
Ghrelin, an endogenous ligand for the GH secretagogue receptor, is a hormone expressed in stomach and other tissues, such as hypothalamus, testis, and placenta. This hormone acts at a central level to stimulate GH secretion and food intake. Little is known, however, about the molecular forms and physiological roles of ghrelin within the hypothalamus. In this report, we detail the molecular forms, mRNA expression patterns, and peptide contents of ghrelin within the rat hypothalamus. Using the combination of reverse-phase HPLC and ghrelin-specific RIA, we determined that the rat hypothalamus contains both n-octanoyl-modified and des-acyl ghrelins. Fasting for 24 and 48 h significantly decreased ghrelin mRNA expression in the hypothalamus to 24% and 28% of control values, respectively. Both n-octanoyl-modified and des-acyl ghrelin content in the hypothalamus decreased after 24 and 48 h of fasting. These results contrast the changes in gastric ghrelin after fasting, which decreased in content despite increased mRNA expression. Two hours after injection of 2-deoxy-D-glucose (2-DG), a selective blocker of carbohydrate metabolism, ghrelin peptide levels also decreased. Thus, induction of glucoprivic states, such as fasting and 2-DG treatment, decreased ghrelin gene expression and peptide content within the hypothalamus.
Introduction
GHRELIN, ORIGINALLY purified from rat stomach, is an endogenous ligand of the GH secretagogue receptor (GHS-R) (1). Ghrelin is a 28-amino acid peptide existing in two major forms: n-octanoyl-modified ghrelin, which possesses an n-octanoyl modification on serine-3, and des-acyl ghrelin (2). Lipid modification of ghrelin is essential for ghrelin-induced GH release from the pituitary (3) and appetite stimulation (4, 5, 6, 7). Thus, the posttranscriptional regulation of octanoyl modification is an important step controlling the biological activity of ghrelin.
In addition to the stomach, ghrelin is localized to the hypothalamic arcuate nucleus (ARC) of rats and mice (1, 8, 9). The hypothalamus, especially the ARC, plays a central role in the integration of different metabolic signals and in appetite regulation. In the hypothalamus, ghrelin neurons contact the cell bodies and dendrites of neuropeptide Y (NPY)/agouti-related protein (AgRP) and proopiomelanocortin neurons, which produce orexigenic peptides and anorexigenic peptide, respectively (9). Intracerebroventricular injection of ghrelin potently promotes food intake in rats and mice (5, 10, 11), suggesting that hypothalamic ghrelin acts as an orexigen.
The molecular composition of hypothalamic ghrelin and its functions in that site, however, remains unclear. In this study, we investigated the molecular forms of hypothalamic ghrelin and the changes in hypothalamic ghrelin mRNA and peptide levels after fasting and 2-DG treatment.
Materials and Methods
Animals
Sprague Dawley male rats (CREA, Tokyo, Japan) were individually housed in a room maintained at a constant ambient temperature of 25 C with a 12-h light, 12-h dark cycle of 12 h (lights on at 0700 h, lights off at 1900 h). Animals were fed standard rodent chow pellets with water ad libitum. Body weight was approximately 310 g at the beginning of the experiments. All animal procedures were performed in accordance with the National Institutes of Health guidelines for the human care of laboratory animals and approved by the Institutional Animal Care Committee.
Fasting experiment
Rats were divided into three groups (12 rats per group) and subjected to experimental fasting for 0, 24, and 48 h. Samples from half of the animals in each group were used for ELISAs, whereas the other half were used for mRNA analysis.
2-DG injection
The rats were divided into two groups (16 rats per group). Rats were injected with either 2-DG (600 mg/kg, ip; Sigma-Aldrich, Tokyo, Japan) or saline (0.9% NaCl, ip). We collected the hypothalami from rats 2 h after injection. Half of the animals in each group were used for ELISAs, whereas the other half were used for mRNA analysis.
Quantification and molecular forms of immunoreactive ghrelin in rat hypothalamus
Fresh rat hypothalami (18 g) were diced and boiled for 5 min in 10 vol of water to inactivate intrinsic proteases. The solution was then adjusted to a final concentration of 1 M acetic acid (AcOH) and 20 mM HCl. The boiled hypothalami were homogenized with a Polytron mixer. The supernatants of the homogenized samples, obtained after centrifugation at 15,000 rpm (18,000 x g) for 40 min, was concentrated to approximately 20 ml in an evaporator. The residual concentrate was subjected to acetone precipitation in a concentration of 66% acetone. After removal of the precipitates, the acetone supernatant was evaporated. The sample was loaded onto a 10-g Sep-Pak C18 cartridge (Waters, Milford, MA) and washed with 10% CH3CN/0.1% trifluoroacetic acid (TFA). After elution with 60% CH3CN/0.1% TFA, samples were evaporated and lyophilized. The residual materials were redissolved in 1 M AcOH and adsorbed on an SP (sulphopropyl)-Sephadex C-25 (H+ form) column that had been preequilibrated in 1 M AcOH. Successive elution with 1 M AcOH, 2 M pyridine, and 2 M pyridine-AcOH (pH 5.0) provided three fractions, designated SP-I, SP-II, and SP-III. The lyophilized SP-III fraction was separated by reverse-phase HPLC (RP-HPLC) using a μBondasphere C18 (3.9 x 150 mm; Waters) column. A linear gradient of CH3CN from 10–60% in 0.1% TFA for 40 min served as the solvent system using a flow rate of 1 ml/min. Each fraction (0.5 ml) was lyophilized and subjected to RIAs specific for ghrelin.
RIAs for rat ghrelin
To characterize the molecular forms of immunoreactive ghrelin, we employed two polyclonal antibodies [no. 6-6 for amino-terminal RIA (N-RIA), and no. 1-7 for carboxyl-terminal RIA (C-RIA)] raised against the C-RIA (Gln13-Arg28) and amino- (Gly1-Lys11 with O-n-octanoylation at Ser3) terminal fragments of rat ghrelin (2). Each RIA incubation mixture contained 100 μl of either a ghrelin standard or an unknown sample and 200 μl of antiserum diluted in RIA buffer [50 mM PBS (pH 7.4), 0.5% BSA, 0.5% Triton X-100, 80 mM NaCl, 25 mM EDTA-2Na, and 0.05% NaN3] containing 0.5% normal rabbit serum. Two antisera were added at final dilutions of 1:12,000 (C-RIA) and 1:2,500,000 (N-RIA) and incubated for 12 h. Samples were then incubated with 100 μl of 125I-labeled ghrelin (20,000 cpm/tube) for 36 h. Next, 100 μl of antirabbit IgG goat serum was incubated with the samples for 24 h. Free and bound tracers were separated by centrifugation at 3,000 rpm for 30 min. Radioactivity in the pellet was quantitated with a counter (ARC-1000M, Aloka, Tokyo, Japan).
Immunohistochemistry
Porcine hypothalamus was immersed in 4% PFA solution, then immersed in a series of 10%, 20%, and 30% sucrose solutions with 10% alabia gum. Tissues were then embedded in OCT compound (Tissue-Tek Miles, Elkhart, IN). Sections were cut at a thickness of 20 μm using a cryostat (CM 3050S; Leica Microscopy and Scientific Instruments Group, Heerbrugg, Switzerland) and mounted on Matsunami adhesive slide-coated slides (Matsunami, Osaka, Japan). Ghrelin immunohistochemical staining was performed by the avidin-biotinylated-peroxidase complex (ABC) system using a VECTASTAIN ABC-PO kit (Vector Laboratories Inc., Burlingame, CA) as previously described (12). Briefly, sections were dried at 37 C for 30 min, washed in 10 mM PBS (pH 7.4), and pretreated with 3% hydrogen peroxide in methanol for 10 min to block endogenous peroxidase activity. Sections were treated with 0.01% saponin in PBS for 20 min. After rinsing with PBS, sections were treated with 3% normal goat serum for 1 h, then incubated in polyclonal rabbit antighrelin antibody (no. 6-6; diluted 1:80,000) for 16 h in 4 C. Sections were rinsed with PBS, then incubated with biotinylated antirabbit IgG for 40 min. After washing in PBS, sections were incubated with VECTASTAIN ABC Reagent for 1 h. Samples were visualized in 3,3'-diaminobenzidine using a Dako liquid diethylaminobenzidine substrate-chromogen system (Dako, Kyoto, Japan). The specificity of the antibodies was demonstrated by immunoabsorption and by the total loss of staining when primary antibodies were omitted. For an immunoabsorption test, we examined porcine ghrelin antigen at concentrations of 0, 0.01, 0.1, 1, and 10 (μg/ml).
Preparation of tissue and plasma samples
To prepare hypothalamus and stomach samples, rat tissues were quickly removed after the rats were killed. Each tissue was diced and boiled for 5 min in a 10-fold volume of water to inactivate intrinsic proteases. The solutions were adjusted to a final concentration of 1 M AcOH and 20 mM HCl after cooling. Tissues were then homogenized with a Polytron mixer and after centrifugation at 15,000 rpm for 10 min supernatants were obtained as tissue samples. To prepare plasma samples, whole blood was mixed with EDTA-2Na (2 mg/ml) and aprotinin (500 kIU/ml). Plasma was collected by centrifugation at 4 C. Tissues and plasma samples were loaded onto Sep-Pak C18 cartridges (Waters). The cartridges were washed in 0.9% NaCl and 10% CH3CN/0.1% TFA. Bound protein was eluted with 60% CH3CN/0.1% TFA. The eluate was lyophilized and subjected to ghrelin-specific ELISA.
ELISA
To quantify hypothalamus, stomach, and plasma ghrelin levels, we used an Active Ghrelin ELISA Kit (Mitsubishi Kagaku Iatron, Inc., Tokyo, Japan) to assess n-octanoyl modified ghrelin and a Desacyl-Ghrelin ELISA Kit (Mitsubishi Kagaku Iatron, Inc.) to measure des-acyl ghrelin according to the manufacturer’s instructions.
Real-time PCR of rat ghrelin, NPY, AgRP, and melanin-concentrating hormone (MCH)
Total RNA was extracted from frozen rat stomach and hypothalamus samples using TRIzol (Invitrogen, Tokyo, Japan). Poly(A)+ RNA was purified from 75 μg total hypothalamic RNA using Oligotex-dT30 (Roche, Tokyo, Japan), according to the manufacturer’s instructions. To synthesize cDNA, poly(A)+ RNA (0.4 μg/animal) derived from the hypothalamus and total RNA (1 μg/animal) from the stomach were used. Reaction mixtures were incubated at 37 C for 60 min. Reactions were stopped by incubation at 70 C for 15 min.
Real-time PCR was performed using a PE Applied Biosystems PRISM 7000 Sequence Detection System (PE Applied Biosystems, Foster City, CA). We measured the expression levels of the ghrelin, NPY, AgRP, and MCH genes in hypothalamus and the ghrelin gene in the stomach of rats. cDNA amplification was performed using SYBR Green PCR Core Reagents (PE Applied Biosystems) and uracil-N-glycosylase (Invitrogen) to prevent contamination with carried-over PCR products, as suggested by the manufacturer. All samples were amplified in a single MicroAmp Optical 96-well reaction plate (PE Applied Biosystems). Results reflect duplicate runs of at least two independent experiments. Primer pairs for each gene were designed using Primer3 software (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3_www.cgi). The gene names, forward and reverse primer sequences, and amplicon sizes are listed in Table 1. PCR cycling conditions were initiated by a 2-min incubation at 50 C to eliminate any deoxyuridine triphosphate-containing PCR products resulting from carryover contamination. After a 15-min period at 95 C to activate HotStarTaq DNA polymerase, PCR fragments were amplified by 40 cycles of 95 C for 30 sec, 60 C for 30 sec, and 1 min at 72 C. Each standard well contained the pGEM-T Easy vector, containing the standard cDNA fragment. The concentration of the standards covered at least 6 orders of magnitude. We also included no template controls on each plate. Experimental samples with a threshold cycle value within 2 SD of the mean threshold cycle value for the no template controls were considered to be below the limits of detection. The relative levels of mRNA were standardized to a housekeeping gene, glyceraldehyde-3-phosphate dehydrogenase, to correct for any bias among the samples caused by RNA isolation, RNA degradation, or the efficiencies of the RT. After amplification, PCR products were analyzed by melting curve to confirm amplification specificity. Amplicon size and reaction specificity were confirmed by agarose gel electrophoresis.
TABLE 1. Primer sequences used for real-time PCR analysis
Statistical analysis
Results are presented as mean ± SEM for each group. Data from the fasting experiment were analyzed with a Kluskal-Wallis nonparametric ANOVA followed by post hoc Fisher tests; and data from the experiment of 2-DG injection were analyzed with Student’s t test. P < 0.05 was accepted as statistically significance.
Results
Identification and molecular forms of ghrelin in hypothalamus
Ghrelin has been found in the hypothalamic ARC (1, 8, 9), an important region in the control of appetite. A recent study reported the presence of ghrelin in previously uncharacterized hypothalamic neurons adjacent to the third ventricle between the dorsal, ventral, paraventricular, and arcuate hypothalamic nuclei (9). It remains unclear, however, whether hypothalamic ghrelin is modified by an octanoic acid in a similar manner as gastric ghrelin.
To characterize the molecular forms of hypothalamic ghrelin, we analyzed peptide extracts from rat hypothalami by using RP-HPLC and ghrelin-specific RIAs. Two major ghrelin peaks were detected by ghrelin C-RIA (Fig. 1), which can recognize both n-octanoyl-modified and des-acyl ghrelins. One of the two major peaks eluted at a retention time of 21 min by HPLC (Fig. 1B, arrow d), the same position as that of n-octanoyl-modified ghrelin (Fig. 1A, arrow b). This major peak was detected by both ghrelin C-RIA and ghrelin N-RIA (Fig. 1C, arrow e), which specifically recognizes n-octanoyl-modified ghrelin. These results confirm the identity of the peak eluting at 21 min as n-octanoyl modified ghrelin, the major active form of ghrelin molecule.
FIG. 1. Representative RP-HPLC profiles of ghrelin immunoreactivity in the rat hypothalamus. A linear gradient of 10–60% CH3CN containing 0.1% TFA was run for 40 min at 1.0 ml/min. A, Chromatograph of rat hypothalamic extract. RP-HPLC of rat hypothalamus was monitored by C-RIA (B) and N-RIA (C) for ghrelin using fraction volumes of 0.5 ml. The arrows indicate the elution points of des-acyl rat ghrelin-(1–28) (arrow a) and n-octanoylated rat ghrelin-(1–28) (arrow b). The two major peaks observed were consistent with the elution points of des-acyl rat ghrelin-(1–28) (arrow c) and n-octanoylated rat ghrelin-(1–28) (arrows d and e).
An additional peak (Fig. 1B, arrow c), eluted at 12 min in HPLC, was only detected by ghrelin C-RIA. This elution position was identical with that of des-acyl ghrelin (Fig. 1A, arrow a). These results confirm identity of the peak as des-acyl ghrelin. Thus, in a manner similar to ghrelin in the stomach, the two major forms of ghrelin, n-octanoyl-modified and des-acyl ghrelins, also exist in the hypothalamus.
Immunohistochemical analysis of porcine ghrelin neurons
Although hypothalamic ghrelin was previously observed in the rat and mouse hypothalamus (1, 8, 9), there is little information concerning hypothalamic ghrelin in other mammals. We, therefore, confirmed the existence of ghrelin in the porcine hypothalamus by immunohistochemistry using an antibody that recognizes n-octanoyl-modified ghrelin. Our data indicated that ghrelin neurons were present in the porcine hypothalamus (Fig. 2), similar to the ghrelin neurons in the rat hypothalamus (1, 8, 9). As in rats and mice, porcine ghrelin-positive neurons were distributed in the ARC and periventricular areas; the fibers and terminals of these neurons projected onto other ghrelin-containing neurons in these areas. In the pig, multiple ghrelin-positive neurons were localized to the paraventricular nucleus (Fig. 2A). The cell shapes of porcine ghrelin-positive neurons were variable (Fig. 2B). The processes of a subset of these neurons projected onto both ghrelin-containing neurons (Fig. 2C) and ghrelin-negative neurons (Fig. 2D), suggesting contact regulation among ghrelin-positive neurons in the hypothalamus. Thus, we confirmed the presence of ghrelin-containing neurons in the porcine hypothalamus, indicating the general existence of n-octanoyl-modified ghrelin in the hypothalami of mammals.
FIG. 2. Localization of ghrelin-immunopositive neurons in the porcine hypothalamus. A, Ghrelin neurons distribution in the paraventricular nucleus. B, A ghrelin-producing neuron in paraventricular nucleus. A subset of ghrelin-positive neurons projected to cell bodies of either additional ghrelin-positive neurons (C, arrowheads) or ghrelin-negative neurons (D, arrowheads). 3V, Third ventricle. Bar, 200 μm (A), 20 μm (B–D).
mRNA expression of ghrelin in the hypothalamus and stomach under fasting condition
The most important influence on the regulation of ghrelin secretion is feeding (13, 14). Plasma ghrelin concentrations increase during fasting and decrease after food intake (13). To examine the role of hypothalamic ghrelin in feeding regulation, we investigated the changes in ghrelin gene expression and ghrelin content in the hypothalamus after fasting.
After 24 and 48 h of fasting, body weight decreased and blood glucose levels were lower than those in fed animals (Fig. 3A), indicating that the fasting experiment was performed properly.
FIG. 3. A, Body weights (left panel) and blood glucose concentrations (right panel) of rats after fasting. B, Ghrelin mRNA levels in the hypothalamus and stomach. C, NPY, AgRP, and MCH mRNA levels in the hypothalamus of rats fed ad libitum (control) or animals that fasted for 24 or 48 h. Note that the values of the longitudinal axes are different in each graph. Asterisks indicate the differences between each bar (P < 0.05).
Ghrelin mRNA expression in the hypothalamus was significantly decreased by 24% and 28% compared with those of control (ad libitum fed) when fasted for 24 and 48 h, respectively (Fig. 3B). As predicted by previous reports, ghrelin mRNA expression in the stomach increased by 75% for 48 h fasting (Fig. 3B). Thus, fasting for 24 and 48 h decreased ghrelin mRNA levels in the hypothalamus, but increased the levels present in the stomach. As the expression levels of hypothalamic appetite-regulating peptides, including NPY, AgRP, and MCH, increased upon fasting as expected (Fig. 3C) (15), the hypothalamic samples were processed and analyzed correctly. These results indicate that the regulatory mechanism(s) governing ghrelin secretion in the hypothalamus differ from that in the stomach.
Concentration of ghrelin in the hypothalamus and stomach under fasting condition
We next investigated the concentrations of ghrelin under fasting conditions. As seen at the mRNA level, ghrelin content in the hypothalamus also decreased after 48 h fasting (n-octanoyl-modified ghrelin: –64% and des-acyl ghrelin: –78%) (Fig. 4A). Whereas ghrelin concentrations in the stomach also decreased after 48 h of fasting, plasma ghrelin concentrations increased as previously reported (Fig. 4, B and C) (13). These results indicate that fasting stimulates the release of ghrelin from the stomach into the blood. In a similar fashion, hypothalamic ghrelin may be released from ghrelin-producing neurons in a fasting-dependent manner. The ratio of n-octanoyl-modified ghrelin to des-acyl ghrelin did not changed in either the stomach, plasma, or hypothalamus.
FIG. 4. Concentrations of n-octanoyl-modified and des-acyl ghrelin in the hypothalamus (A), stomach (B), and plasma (C) after fasting for 24 or 48 h. Note that the values of the longitudinal axes are different in each graph. Asterisks indicate the differences between each bar (P < 0.05).
mRNA expression in the hypothalamus and stomach after 2-DG treatment
Blood glucose levels are important factors for release of ghrelin; both oral and iv administration of glucose decreases plasma ghrelin concentrations (16). Because injection of 2-DG stimulates food intake by antagonizing glucose utilization (17), we investigated the effect of 2-DG on the ghrelin mRNA expression and concentration in the hypothalamus.
After administration of 2-DG, mean food intake for 2 h by rats was dramatically increased in comparison with food intake after saline administration (saline group: 0.4 ± 0.2 g, 2-DG group: 5.1 ± 1.1 g) (Fig. 5A), indicating the effectiveness of 2-DG administration. In rats, 2-DG treatment produced a significant decrease in ghrelin mRNA expression within the hypothalamus only (–50.2%); 2-DG did not alter ghrelin mRNA levels in the stomach (Fig. 5B). In contrast, additional orexigenic peptides produced by the rat hypothalamus, including NPY, AgRP, and MCH, increased after 2-DG treatment as reported (Fig. 5C) (17).
FIG. 5. A, Food intake in the 2 h after 2-DG injection. B, Ghrelin mRNA levels in the hypothalamus and stomach. C, NPY, AgRP, and MCH mRNA levels in the hypothalami of rats treated with 2-DG for 2 h. Asterisks indicate the differences between each bar (P < 0.05).
Concentration of ghrelin in the hypothalamus and stomach after 2-DG treatment
Ghrelin peptide levels in the hypothalamus were also decreased by 2-DG treatment (Fig. 6A). Both n-octanoyl-modified and des-acyl ghrelin decreased by 57% and 44% in comparison with control values (saline-treated group) values, respectively. There was no change in ghrelin peptide levels, however, in either the stomach or the plasma (Fig. 6, B and C). The ratio of n-octanoyl-modified ghrelin to des-acyl ghrelin in the hypothalamus was not changed by 2-DG treatment. Thus, antagonism of glucose utilization by 2-DG decreases ghrelin mRNA expression and protein levels in the hypothalamus only.
FIG. 6. n-Octanoyl and des-acyl ghrelin peptide content in the hypothalamus (A), stomach (B), and plasma (C) of rats treated with 2-DG for 2 h. Note that the values of the longitudinal axes differs in every graph. Asterisks indicate the differences between each bar (P < 0.05).
Discussion
The hypothalamus is one of the target tissues of ghrelin, a potent appetite-stimulating hormone. Previous studies have reported that ghrelin and its receptor are expressed within the hypothalamic ARC (1, 8, 9, 18) to function in appetite regulation (4, 5, 6, 7). The molecular forms of hypothalamic ghrelin have remained unclear; it was previously unknown whether hypothalamic ghrelin is also modified with an octanoic acid. Moreover, the regulation of hypothalamic ghrelin expression and concentrations remains to be elucidated.
In this study, we examined the molecular composition of hypothalamic ghrelin by HPLC and ghrelin RIAs. We identified two molecular forms of hypothalamic ghrelin, n-octanoyl-modified ghrelin and des-acyl ghrelin, as seen for ghrelin within the stomach (2). In the stomach, additional minor molecular forms of ghrelin, such as hexanoyl-, decenoyl-, and decanoyl-modified ghrelins, also exist at limited concentrations (19). Due to the low content of ghrelin in the hypothalamus, we could not detect these minor forms. Thus, the main active form of hypothalamic ghrelin is n-octanoyl-modified ghrelin, which is also the primary active form of ghrelin present in the stomach.
We next examined the changes in ghrelin mRNA expression levels and peptide concentrations in the hypothalamus after either fasting or treatment with 2-DG, an antagonist of glucose utilization. The results demonstrate that both ghrelin mRNA expression and peptide content in the rat hypothalamus decreased after fasting. These changes in the hypothalamus do not correlate with those previously seen in the stomach. Ghrelin mRNA expression in the stomach increases after fasting (Fig. 3), whereas peptide concentrations of ghrelin in the stomach are decreased. Because plasma ghrelin concentrations increase after fasting, fasting may induce the excessive secretion of ghrelin from the stomach into the blood, resulting in a decrease of ghrelin peptide content in the stomach. Thus, fasting likely also stimulates ghrelin release from the hypothalamus, resulting in a decrease in hypothalamic ghrelin concentrations. 2-DG treatment reduced hypothalamic ghrelin concentrations in the absence of any changes in the peptide content of either the stomach or plasma. Because 2-DG stimulates feeding by exerting central metabolic influences (20), hypothalamic ghrelin secretion should also be centrally regulated. Therefore, we think that glucoprivic states in hypothalamus, such as fasting or 2-DG treatment, promote hypothalamic ghrelin secretion.
In contrast to the increases in gastric ghrelin mRNA during fasting conditions, hypothalamic ghrelin mRNA decreased until 48 h of fasting. Although we cannot clearly explain this phenomenon as yet, one possibility is that ghrelin gene expression levels in the hypothalamus are suppressed after ghrelin release to prevent excessive ghrelin secretion. Because in case of long-term starvation, excessive food-exploratory behavior induced by orexigenic peptides may result in exhaustion and death. Thus, hypothalamic ghrelin might control effectual feeding behavior in response to a nutritional state.
Starvation in goldfish induces increases in hypothalamic ghrelin mRNA on d 7, whereas serum ghrelin levels increased at d 3 and 5 and returned on d 7 to the level of d 1 (21). These data for goldfish ghrelin are quite similar to our results. Thus, hypothalamic ghrelin gene expression may be regulated by multiple additional factors. Further studies will be necessary to understand the induction of hypothalamic ghrelin secretion by fasting and 2-DG.
In summary, we revealed that hypothalamic ghrelin exists as two major forms: the n-octanoyl-modified and des-acyl ghrelin peptides. The glucoprivic state of the hypothalamus, induced by fasting and 2-DG treatment, stimulates ghrelin secretion from ghrelin-producing neurons. Moreover, the contradictory expression patterns of ghrelin mRNA expression in the hypothalamus and the stomach after fasting imply that the mechanism of ghrelin synthesis differs between these two locations.
Acknowledgments
We thank Y. Yamashita (Kurume University, Fukuoka, Japan) for his helpful assistance.
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Address all correspondence and requests for reprints to: Masayasu Kojima, Ph.D., Molecular Genetics, Institute of Life Sciences, Kurume University, Fukuoka 839-0864, Japan. E-mail: mkojima@lsi.kurume-u.ac.jp.
Abstract
Ghrelin, an endogenous ligand for the GH secretagogue receptor, is a hormone expressed in stomach and other tissues, such as hypothalamus, testis, and placenta. This hormone acts at a central level to stimulate GH secretion and food intake. Little is known, however, about the molecular forms and physiological roles of ghrelin within the hypothalamus. In this report, we detail the molecular forms, mRNA expression patterns, and peptide contents of ghrelin within the rat hypothalamus. Using the combination of reverse-phase HPLC and ghrelin-specific RIA, we determined that the rat hypothalamus contains both n-octanoyl-modified and des-acyl ghrelins. Fasting for 24 and 48 h significantly decreased ghrelin mRNA expression in the hypothalamus to 24% and 28% of control values, respectively. Both n-octanoyl-modified and des-acyl ghrelin content in the hypothalamus decreased after 24 and 48 h of fasting. These results contrast the changes in gastric ghrelin after fasting, which decreased in content despite increased mRNA expression. Two hours after injection of 2-deoxy-D-glucose (2-DG), a selective blocker of carbohydrate metabolism, ghrelin peptide levels also decreased. Thus, induction of glucoprivic states, such as fasting and 2-DG treatment, decreased ghrelin gene expression and peptide content within the hypothalamus.
Introduction
GHRELIN, ORIGINALLY purified from rat stomach, is an endogenous ligand of the GH secretagogue receptor (GHS-R) (1). Ghrelin is a 28-amino acid peptide existing in two major forms: n-octanoyl-modified ghrelin, which possesses an n-octanoyl modification on serine-3, and des-acyl ghrelin (2). Lipid modification of ghrelin is essential for ghrelin-induced GH release from the pituitary (3) and appetite stimulation (4, 5, 6, 7). Thus, the posttranscriptional regulation of octanoyl modification is an important step controlling the biological activity of ghrelin.
In addition to the stomach, ghrelin is localized to the hypothalamic arcuate nucleus (ARC) of rats and mice (1, 8, 9). The hypothalamus, especially the ARC, plays a central role in the integration of different metabolic signals and in appetite regulation. In the hypothalamus, ghrelin neurons contact the cell bodies and dendrites of neuropeptide Y (NPY)/agouti-related protein (AgRP) and proopiomelanocortin neurons, which produce orexigenic peptides and anorexigenic peptide, respectively (9). Intracerebroventricular injection of ghrelin potently promotes food intake in rats and mice (5, 10, 11), suggesting that hypothalamic ghrelin acts as an orexigen.
The molecular composition of hypothalamic ghrelin and its functions in that site, however, remains unclear. In this study, we investigated the molecular forms of hypothalamic ghrelin and the changes in hypothalamic ghrelin mRNA and peptide levels after fasting and 2-DG treatment.
Materials and Methods
Animals
Sprague Dawley male rats (CREA, Tokyo, Japan) were individually housed in a room maintained at a constant ambient temperature of 25 C with a 12-h light, 12-h dark cycle of 12 h (lights on at 0700 h, lights off at 1900 h). Animals were fed standard rodent chow pellets with water ad libitum. Body weight was approximately 310 g at the beginning of the experiments. All animal procedures were performed in accordance with the National Institutes of Health guidelines for the human care of laboratory animals and approved by the Institutional Animal Care Committee.
Fasting experiment
Rats were divided into three groups (12 rats per group) and subjected to experimental fasting for 0, 24, and 48 h. Samples from half of the animals in each group were used for ELISAs, whereas the other half were used for mRNA analysis.
2-DG injection
The rats were divided into two groups (16 rats per group). Rats were injected with either 2-DG (600 mg/kg, ip; Sigma-Aldrich, Tokyo, Japan) or saline (0.9% NaCl, ip). We collected the hypothalami from rats 2 h after injection. Half of the animals in each group were used for ELISAs, whereas the other half were used for mRNA analysis.
Quantification and molecular forms of immunoreactive ghrelin in rat hypothalamus
Fresh rat hypothalami (18 g) were diced and boiled for 5 min in 10 vol of water to inactivate intrinsic proteases. The solution was then adjusted to a final concentration of 1 M acetic acid (AcOH) and 20 mM HCl. The boiled hypothalami were homogenized with a Polytron mixer. The supernatants of the homogenized samples, obtained after centrifugation at 15,000 rpm (18,000 x g) for 40 min, was concentrated to approximately 20 ml in an evaporator. The residual concentrate was subjected to acetone precipitation in a concentration of 66% acetone. After removal of the precipitates, the acetone supernatant was evaporated. The sample was loaded onto a 10-g Sep-Pak C18 cartridge (Waters, Milford, MA) and washed with 10% CH3CN/0.1% trifluoroacetic acid (TFA). After elution with 60% CH3CN/0.1% TFA, samples were evaporated and lyophilized. The residual materials were redissolved in 1 M AcOH and adsorbed on an SP (sulphopropyl)-Sephadex C-25 (H+ form) column that had been preequilibrated in 1 M AcOH. Successive elution with 1 M AcOH, 2 M pyridine, and 2 M pyridine-AcOH (pH 5.0) provided three fractions, designated SP-I, SP-II, and SP-III. The lyophilized SP-III fraction was separated by reverse-phase HPLC (RP-HPLC) using a μBondasphere C18 (3.9 x 150 mm; Waters) column. A linear gradient of CH3CN from 10–60% in 0.1% TFA for 40 min served as the solvent system using a flow rate of 1 ml/min. Each fraction (0.5 ml) was lyophilized and subjected to RIAs specific for ghrelin.
RIAs for rat ghrelin
To characterize the molecular forms of immunoreactive ghrelin, we employed two polyclonal antibodies [no. 6-6 for amino-terminal RIA (N-RIA), and no. 1-7 for carboxyl-terminal RIA (C-RIA)] raised against the C-RIA (Gln13-Arg28) and amino- (Gly1-Lys11 with O-n-octanoylation at Ser3) terminal fragments of rat ghrelin (2). Each RIA incubation mixture contained 100 μl of either a ghrelin standard or an unknown sample and 200 μl of antiserum diluted in RIA buffer [50 mM PBS (pH 7.4), 0.5% BSA, 0.5% Triton X-100, 80 mM NaCl, 25 mM EDTA-2Na, and 0.05% NaN3] containing 0.5% normal rabbit serum. Two antisera were added at final dilutions of 1:12,000 (C-RIA) and 1:2,500,000 (N-RIA) and incubated for 12 h. Samples were then incubated with 100 μl of 125I-labeled ghrelin (20,000 cpm/tube) for 36 h. Next, 100 μl of antirabbit IgG goat serum was incubated with the samples for 24 h. Free and bound tracers were separated by centrifugation at 3,000 rpm for 30 min. Radioactivity in the pellet was quantitated with a counter (ARC-1000M, Aloka, Tokyo, Japan).
Immunohistochemistry
Porcine hypothalamus was immersed in 4% PFA solution, then immersed in a series of 10%, 20%, and 30% sucrose solutions with 10% alabia gum. Tissues were then embedded in OCT compound (Tissue-Tek Miles, Elkhart, IN). Sections were cut at a thickness of 20 μm using a cryostat (CM 3050S; Leica Microscopy and Scientific Instruments Group, Heerbrugg, Switzerland) and mounted on Matsunami adhesive slide-coated slides (Matsunami, Osaka, Japan). Ghrelin immunohistochemical staining was performed by the avidin-biotinylated-peroxidase complex (ABC) system using a VECTASTAIN ABC-PO kit (Vector Laboratories Inc., Burlingame, CA) as previously described (12). Briefly, sections were dried at 37 C for 30 min, washed in 10 mM PBS (pH 7.4), and pretreated with 3% hydrogen peroxide in methanol for 10 min to block endogenous peroxidase activity. Sections were treated with 0.01% saponin in PBS for 20 min. After rinsing with PBS, sections were treated with 3% normal goat serum for 1 h, then incubated in polyclonal rabbit antighrelin antibody (no. 6-6; diluted 1:80,000) for 16 h in 4 C. Sections were rinsed with PBS, then incubated with biotinylated antirabbit IgG for 40 min. After washing in PBS, sections were incubated with VECTASTAIN ABC Reagent for 1 h. Samples were visualized in 3,3'-diaminobenzidine using a Dako liquid diethylaminobenzidine substrate-chromogen system (Dako, Kyoto, Japan). The specificity of the antibodies was demonstrated by immunoabsorption and by the total loss of staining when primary antibodies were omitted. For an immunoabsorption test, we examined porcine ghrelin antigen at concentrations of 0, 0.01, 0.1, 1, and 10 (μg/ml).
Preparation of tissue and plasma samples
To prepare hypothalamus and stomach samples, rat tissues were quickly removed after the rats were killed. Each tissue was diced and boiled for 5 min in a 10-fold volume of water to inactivate intrinsic proteases. The solutions were adjusted to a final concentration of 1 M AcOH and 20 mM HCl after cooling. Tissues were then homogenized with a Polytron mixer and after centrifugation at 15,000 rpm for 10 min supernatants were obtained as tissue samples. To prepare plasma samples, whole blood was mixed with EDTA-2Na (2 mg/ml) and aprotinin (500 kIU/ml). Plasma was collected by centrifugation at 4 C. Tissues and plasma samples were loaded onto Sep-Pak C18 cartridges (Waters). The cartridges were washed in 0.9% NaCl and 10% CH3CN/0.1% TFA. Bound protein was eluted with 60% CH3CN/0.1% TFA. The eluate was lyophilized and subjected to ghrelin-specific ELISA.
ELISA
To quantify hypothalamus, stomach, and plasma ghrelin levels, we used an Active Ghrelin ELISA Kit (Mitsubishi Kagaku Iatron, Inc., Tokyo, Japan) to assess n-octanoyl modified ghrelin and a Desacyl-Ghrelin ELISA Kit (Mitsubishi Kagaku Iatron, Inc.) to measure des-acyl ghrelin according to the manufacturer’s instructions.
Real-time PCR of rat ghrelin, NPY, AgRP, and melanin-concentrating hormone (MCH)
Total RNA was extracted from frozen rat stomach and hypothalamus samples using TRIzol (Invitrogen, Tokyo, Japan). Poly(A)+ RNA was purified from 75 μg total hypothalamic RNA using Oligotex-dT30
Real-time PCR was performed using a PE Applied Biosystems PRISM 7000 Sequence Detection System (PE Applied Biosystems, Foster City, CA). We measured the expression levels of the ghrelin, NPY, AgRP, and MCH genes in hypothalamus and the ghrelin gene in the stomach of rats. cDNA amplification was performed using SYBR Green PCR Core Reagents (PE Applied Biosystems) and uracil-N-glycosylase (Invitrogen) to prevent contamination with carried-over PCR products, as suggested by the manufacturer. All samples were amplified in a single MicroAmp Optical 96-well reaction plate (PE Applied Biosystems). Results reflect duplicate runs of at least two independent experiments. Primer pairs for each gene were designed using Primer3 software (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3_www.cgi). The gene names, forward and reverse primer sequences, and amplicon sizes are listed in Table 1. PCR cycling conditions were initiated by a 2-min incubation at 50 C to eliminate any deoxyuridine triphosphate-containing PCR products resulting from carryover contamination. After a 15-min period at 95 C to activate HotStarTaq DNA polymerase, PCR fragments were amplified by 40 cycles of 95 C for 30 sec, 60 C for 30 sec, and 1 min at 72 C. Each standard well contained the pGEM-T Easy vector, containing the standard cDNA fragment. The concentration of the standards covered at least 6 orders of magnitude. We also included no template controls on each plate. Experimental samples with a threshold cycle value within 2 SD of the mean threshold cycle value for the no template controls were considered to be below the limits of detection. The relative levels of mRNA were standardized to a housekeeping gene, glyceraldehyde-3-phosphate dehydrogenase, to correct for any bias among the samples caused by RNA isolation, RNA degradation, or the efficiencies of the RT. After amplification, PCR products were analyzed by melting curve to confirm amplification specificity. Amplicon size and reaction specificity were confirmed by agarose gel electrophoresis.
TABLE 1. Primer sequences used for real-time PCR analysis
Statistical analysis
Results are presented as mean ± SEM for each group. Data from the fasting experiment were analyzed with a Kluskal-Wallis nonparametric ANOVA followed by post hoc Fisher tests; and data from the experiment of 2-DG injection were analyzed with Student’s t test. P < 0.05 was accepted as statistically significance.
Results
Identification and molecular forms of ghrelin in hypothalamus
Ghrelin has been found in the hypothalamic ARC (1, 8, 9), an important region in the control of appetite. A recent study reported the presence of ghrelin in previously uncharacterized hypothalamic neurons adjacent to the third ventricle between the dorsal, ventral, paraventricular, and arcuate hypothalamic nuclei (9). It remains unclear, however, whether hypothalamic ghrelin is modified by an octanoic acid in a similar manner as gastric ghrelin.
To characterize the molecular forms of hypothalamic ghrelin, we analyzed peptide extracts from rat hypothalami by using RP-HPLC and ghrelin-specific RIAs. Two major ghrelin peaks were detected by ghrelin C-RIA (Fig. 1), which can recognize both n-octanoyl-modified and des-acyl ghrelins. One of the two major peaks eluted at a retention time of 21 min by HPLC (Fig. 1B, arrow d), the same position as that of n-octanoyl-modified ghrelin (Fig. 1A, arrow b). This major peak was detected by both ghrelin C-RIA and ghrelin N-RIA (Fig. 1C, arrow e), which specifically recognizes n-octanoyl-modified ghrelin. These results confirm the identity of the peak eluting at 21 min as n-octanoyl modified ghrelin, the major active form of ghrelin molecule.
FIG. 1. Representative RP-HPLC profiles of ghrelin immunoreactivity in the rat hypothalamus. A linear gradient of 10–60% CH3CN containing 0.1% TFA was run for 40 min at 1.0 ml/min. A, Chromatograph of rat hypothalamic extract. RP-HPLC of rat hypothalamus was monitored by C-RIA (B) and N-RIA (C) for ghrelin using fraction volumes of 0.5 ml. The arrows indicate the elution points of des-acyl rat ghrelin-(1–28) (arrow a) and n-octanoylated rat ghrelin-(1–28) (arrow b). The two major peaks observed were consistent with the elution points of des-acyl rat ghrelin-(1–28) (arrow c) and n-octanoylated rat ghrelin-(1–28) (arrows d and e).
An additional peak (Fig. 1B, arrow c), eluted at 12 min in HPLC, was only detected by ghrelin C-RIA. This elution position was identical with that of des-acyl ghrelin (Fig. 1A, arrow a). These results confirm identity of the peak as des-acyl ghrelin. Thus, in a manner similar to ghrelin in the stomach, the two major forms of ghrelin, n-octanoyl-modified and des-acyl ghrelins, also exist in the hypothalamus.
Immunohistochemical analysis of porcine ghrelin neurons
Although hypothalamic ghrelin was previously observed in the rat and mouse hypothalamus (1, 8, 9), there is little information concerning hypothalamic ghrelin in other mammals. We, therefore, confirmed the existence of ghrelin in the porcine hypothalamus by immunohistochemistry using an antibody that recognizes n-octanoyl-modified ghrelin. Our data indicated that ghrelin neurons were present in the porcine hypothalamus (Fig. 2), similar to the ghrelin neurons in the rat hypothalamus (1, 8, 9). As in rats and mice, porcine ghrelin-positive neurons were distributed in the ARC and periventricular areas; the fibers and terminals of these neurons projected onto other ghrelin-containing neurons in these areas. In the pig, multiple ghrelin-positive neurons were localized to the paraventricular nucleus (Fig. 2A). The cell shapes of porcine ghrelin-positive neurons were variable (Fig. 2B). The processes of a subset of these neurons projected onto both ghrelin-containing neurons (Fig. 2C) and ghrelin-negative neurons (Fig. 2D), suggesting contact regulation among ghrelin-positive neurons in the hypothalamus. Thus, we confirmed the presence of ghrelin-containing neurons in the porcine hypothalamus, indicating the general existence of n-octanoyl-modified ghrelin in the hypothalami of mammals.
FIG. 2. Localization of ghrelin-immunopositive neurons in the porcine hypothalamus. A, Ghrelin neurons distribution in the paraventricular nucleus. B, A ghrelin-producing neuron in paraventricular nucleus. A subset of ghrelin-positive neurons projected to cell bodies of either additional ghrelin-positive neurons (C, arrowheads) or ghrelin-negative neurons (D, arrowheads). 3V, Third ventricle. Bar, 200 μm (A), 20 μm (B–D).
mRNA expression of ghrelin in the hypothalamus and stomach under fasting condition
The most important influence on the regulation of ghrelin secretion is feeding (13, 14). Plasma ghrelin concentrations increase during fasting and decrease after food intake (13). To examine the role of hypothalamic ghrelin in feeding regulation, we investigated the changes in ghrelin gene expression and ghrelin content in the hypothalamus after fasting.
After 24 and 48 h of fasting, body weight decreased and blood glucose levels were lower than those in fed animals (Fig. 3A), indicating that the fasting experiment was performed properly.
FIG. 3. A, Body weights (left panel) and blood glucose concentrations (right panel) of rats after fasting. B, Ghrelin mRNA levels in the hypothalamus and stomach. C, NPY, AgRP, and MCH mRNA levels in the hypothalamus of rats fed ad libitum (control) or animals that fasted for 24 or 48 h. Note that the values of the longitudinal axes are different in each graph. Asterisks indicate the differences between each bar (P < 0.05).
Ghrelin mRNA expression in the hypothalamus was significantly decreased by 24% and 28% compared with those of control (ad libitum fed) when fasted for 24 and 48 h, respectively (Fig. 3B). As predicted by previous reports, ghrelin mRNA expression in the stomach increased by 75% for 48 h fasting (Fig. 3B). Thus, fasting for 24 and 48 h decreased ghrelin mRNA levels in the hypothalamus, but increased the levels present in the stomach. As the expression levels of hypothalamic appetite-regulating peptides, including NPY, AgRP, and MCH, increased upon fasting as expected (Fig. 3C) (15), the hypothalamic samples were processed and analyzed correctly. These results indicate that the regulatory mechanism(s) governing ghrelin secretion in the hypothalamus differ from that in the stomach.
Concentration of ghrelin in the hypothalamus and stomach under fasting condition
We next investigated the concentrations of ghrelin under fasting conditions. As seen at the mRNA level, ghrelin content in the hypothalamus also decreased after 48 h fasting (n-octanoyl-modified ghrelin: –64% and des-acyl ghrelin: –78%) (Fig. 4A). Whereas ghrelin concentrations in the stomach also decreased after 48 h of fasting, plasma ghrelin concentrations increased as previously reported (Fig. 4, B and C) (13). These results indicate that fasting stimulates the release of ghrelin from the stomach into the blood. In a similar fashion, hypothalamic ghrelin may be released from ghrelin-producing neurons in a fasting-dependent manner. The ratio of n-octanoyl-modified ghrelin to des-acyl ghrelin did not changed in either the stomach, plasma, or hypothalamus.
FIG. 4. Concentrations of n-octanoyl-modified and des-acyl ghrelin in the hypothalamus (A), stomach (B), and plasma (C) after fasting for 24 or 48 h. Note that the values of the longitudinal axes are different in each graph. Asterisks indicate the differences between each bar (P < 0.05).
mRNA expression in the hypothalamus and stomach after 2-DG treatment
Blood glucose levels are important factors for release of ghrelin; both oral and iv administration of glucose decreases plasma ghrelin concentrations (16). Because injection of 2-DG stimulates food intake by antagonizing glucose utilization (17), we investigated the effect of 2-DG on the ghrelin mRNA expression and concentration in the hypothalamus.
After administration of 2-DG, mean food intake for 2 h by rats was dramatically increased in comparison with food intake after saline administration (saline group: 0.4 ± 0.2 g, 2-DG group: 5.1 ± 1.1 g) (Fig. 5A), indicating the effectiveness of 2-DG administration. In rats, 2-DG treatment produced a significant decrease in ghrelin mRNA expression within the hypothalamus only (–50.2%); 2-DG did not alter ghrelin mRNA levels in the stomach (Fig. 5B). In contrast, additional orexigenic peptides produced by the rat hypothalamus, including NPY, AgRP, and MCH, increased after 2-DG treatment as reported (Fig. 5C) (17).
FIG. 5. A, Food intake in the 2 h after 2-DG injection. B, Ghrelin mRNA levels in the hypothalamus and stomach. C, NPY, AgRP, and MCH mRNA levels in the hypothalami of rats treated with 2-DG for 2 h. Asterisks indicate the differences between each bar (P < 0.05).
Concentration of ghrelin in the hypothalamus and stomach after 2-DG treatment
Ghrelin peptide levels in the hypothalamus were also decreased by 2-DG treatment (Fig. 6A). Both n-octanoyl-modified and des-acyl ghrelin decreased by 57% and 44% in comparison with control values (saline-treated group) values, respectively. There was no change in ghrelin peptide levels, however, in either the stomach or the plasma (Fig. 6, B and C). The ratio of n-octanoyl-modified ghrelin to des-acyl ghrelin in the hypothalamus was not changed by 2-DG treatment. Thus, antagonism of glucose utilization by 2-DG decreases ghrelin mRNA expression and protein levels in the hypothalamus only.
FIG. 6. n-Octanoyl and des-acyl ghrelin peptide content in the hypothalamus (A), stomach (B), and plasma (C) of rats treated with 2-DG for 2 h. Note that the values of the longitudinal axes differs in every graph. Asterisks indicate the differences between each bar (P < 0.05).
Discussion
The hypothalamus is one of the target tissues of ghrelin, a potent appetite-stimulating hormone. Previous studies have reported that ghrelin and its receptor are expressed within the hypothalamic ARC (1, 8, 9, 18) to function in appetite regulation (4, 5, 6, 7). The molecular forms of hypothalamic ghrelin have remained unclear; it was previously unknown whether hypothalamic ghrelin is also modified with an octanoic acid. Moreover, the regulation of hypothalamic ghrelin expression and concentrations remains to be elucidated.
In this study, we examined the molecular composition of hypothalamic ghrelin by HPLC and ghrelin RIAs. We identified two molecular forms of hypothalamic ghrelin, n-octanoyl-modified ghrelin and des-acyl ghrelin, as seen for ghrelin within the stomach (2). In the stomach, additional minor molecular forms of ghrelin, such as hexanoyl-, decenoyl-, and decanoyl-modified ghrelins, also exist at limited concentrations (19). Due to the low content of ghrelin in the hypothalamus, we could not detect these minor forms. Thus, the main active form of hypothalamic ghrelin is n-octanoyl-modified ghrelin, which is also the primary active form of ghrelin present in the stomach.
We next examined the changes in ghrelin mRNA expression levels and peptide concentrations in the hypothalamus after either fasting or treatment with 2-DG, an antagonist of glucose utilization. The results demonstrate that both ghrelin mRNA expression and peptide content in the rat hypothalamus decreased after fasting. These changes in the hypothalamus do not correlate with those previously seen in the stomach. Ghrelin mRNA expression in the stomach increases after fasting (Fig. 3), whereas peptide concentrations of ghrelin in the stomach are decreased. Because plasma ghrelin concentrations increase after fasting, fasting may induce the excessive secretion of ghrelin from the stomach into the blood, resulting in a decrease of ghrelin peptide content in the stomach. Thus, fasting likely also stimulates ghrelin release from the hypothalamus, resulting in a decrease in hypothalamic ghrelin concentrations. 2-DG treatment reduced hypothalamic ghrelin concentrations in the absence of any changes in the peptide content of either the stomach or plasma. Because 2-DG stimulates feeding by exerting central metabolic influences (20), hypothalamic ghrelin secretion should also be centrally regulated. Therefore, we think that glucoprivic states in hypothalamus, such as fasting or 2-DG treatment, promote hypothalamic ghrelin secretion.
In contrast to the increases in gastric ghrelin mRNA during fasting conditions, hypothalamic ghrelin mRNA decreased until 48 h of fasting. Although we cannot clearly explain this phenomenon as yet, one possibility is that ghrelin gene expression levels in the hypothalamus are suppressed after ghrelin release to prevent excessive ghrelin secretion. Because in case of long-term starvation, excessive food-exploratory behavior induced by orexigenic peptides may result in exhaustion and death. Thus, hypothalamic ghrelin might control effectual feeding behavior in response to a nutritional state.
Starvation in goldfish induces increases in hypothalamic ghrelin mRNA on d 7, whereas serum ghrelin levels increased at d 3 and 5 and returned on d 7 to the level of d 1 (21). These data for goldfish ghrelin are quite similar to our results. Thus, hypothalamic ghrelin gene expression may be regulated by multiple additional factors. Further studies will be necessary to understand the induction of hypothalamic ghrelin secretion by fasting and 2-DG.
In summary, we revealed that hypothalamic ghrelin exists as two major forms: the n-octanoyl-modified and des-acyl ghrelin peptides. The glucoprivic state of the hypothalamus, induced by fasting and 2-DG treatment, stimulates ghrelin secretion from ghrelin-producing neurons. Moreover, the contradictory expression patterns of ghrelin mRNA expression in the hypothalamus and the stomach after fasting imply that the mechanism of ghrelin synthesis differs between these two locations.
Acknowledgments
We thank Y. Yamashita (Kurume University, Fukuoka, Japan) for his helpful assistance.
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