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. 2010 May 12;151(7):3237–3246. doi: 10.1210/en.2010-0219

Hindbrain Catecholamine Neurons Modulate the Growth Hormone But Not the Feeding Response to Ghrelin

Alan J Emanuel 1, Sue Ritter 1
PMCID: PMC2903929  PMID: 20463049

Abstract

The gastrointestinal peptide, ghrelin, elicits feeding and secretion when administered systemically or centrally. Previous studies have suggested that hypothalamic projections of hindbrain catecholamine neurons are involved in both of these actions of ghrelin. The purpose of this study was to further assess the role of hindbrain catecholamine neurons in ghrelin-induced feeding and GH secretion and to determine the anatomical distribution of the catecholamine neurons involved. We lesioned noradrenergic and adrenergic neurons that innervate the medial hypothalamus by microinjecting the retrogradely transported immunotoxin, saporin (SAP) conjugated to antidopamine-β-hydroxylase (DSAP) into the paraventricular nucleus of the hypothalamus. Controls were injected with unconjugated SAP. We found that the DSAP lesion did not impair the feeding response to central or peripheral ghrelin administration, indicating that these neurons are not required for ghrelin’s orexigenic effect. However, the GH response to ghrelin was prolonged significantly in DSAP-lesioned rats. We also found that expression of Fos, an indicator of neuronal activation, was significantly enhanced over baseline levels in A1, A1/C1, C1, and A5 cell groups after ghrelin treatment and in A1, A1/C1, and A5 cell groups after GH treatment. The similar pattern of Fos expression in catecholamine cell groups after GH and ghrelin and the prolonged GH secretion in response to ghrelin in DSAP rats together suggest that activation of hindbrain catecholamine neurons by ghrelin or GH could be a component of a negative feedback response controlling GH levels.

Hindbrain catecholamine neurons are activated after ghrelin treatment but are not required for the feeding response to ghrelin; instead, these neurons likely modulate GH secretion.

Ghrelin is a peptide secreted primarily by endocrine cells in the oxyntic gastric mucosa (1,2). Ghrelin was originally discovered as an endogenous ligand for the secretogogue receptor (GHSR) type 1a (1,3,4,5). It has been suggested that ghrelin takes part in the energy state-dependent regulation of GH (6), which is released in a pulsatile pattern and is regulated through the GHRH and somatostatin axis. Ghrelin stimulates GH release both by acting directly on GHSRs in the anterior pituitary and by increasing release of GHRH from hypothalamic parvocellular neurons (7,8). Dysregulation of GH release results in abnormal growth characteristics as evidenced by acromegaly patients with excess GH and growth failure in patients with GH deficiency (9).

A second major physiological effect of ghrelin is stimulation of food intake (10,11,12,13,14,15,16). During fasting and glucoprivation, ghrelin peptide content is decreased in the hypothalamus and stomach and Ghrelin expression is increased in stomach. This indicates that food deprivation is a stimulus for ghrelin production and secretion (17,18,19). Ghrelin-induced stimulation of feeding appears to be mediated largely by the arcuate nucleus of the hypothalamus (Arc) (11,20). Ghrelin activates orexigenic neurons in the Arc that express neuropeptide Y (NPY) and Agoutigene-related peptide (AGRP) and inhibits anorexigenic neurons that express proopiomelanocortin (20,21,22). Lesions of the Arc that destroy NPY/AGRP and proopiomelanocortin neurons impair or abolish ghrelin-induced stimulation of feeding (23,24).

Although there is much evidence to support the view that ghrelin-induced GH secretion and feeding are dependent on ghrelin’s actions within the pituitary and/or Arc, ghrelin and ghrelin receptors are also expressed in other brain sites (25) and on the vagus nerve (26). In addition, ghrelin injections into hypothalamic sites outside the Arc and into nonhypothalamic sites stimulate feeding (19,27,28,29), and both ghrelin-induced feeding and GH secretion are reported to be blocked by vagotomy (26). Therefore, ghrelin’s actions at these various extrahypothalamic sites are potentially important for its major effects. Indeed, Date et al. (30) recently reported that noradrenergic (NE) neurons in the A2 cell group within the nucleus of the solitary tract (NTS) are a required substrate for control of food intake by circulating ghrelin. Specifically, Date et al. (30) proposed that circulating ghrelin acts initially on vagal sensory neurons, which then activate A2 neurons that innervate the Arc. In support of their hypothesis, these investigators showed that ghrelin increases dopamine-β-hydroxylase (Dbh) expression in A2 cell bodies and NE release in the Arc, and conversely, that ghrelin-induced feeding is attenuated by intracerebroventricular (icv) α1 and β2 NE receptor antagonists or lesion of the catecholamine projection to the Arc by midbrain transection or by Arc injection of the retrogradely transported catecholamine immunotoxin, anti-DBH-saporin (SAP) [SAP conjugated to a monoclonal antibody against DBH (DSAP)]. They also reported that both the feeding and GH responses to circulating ghrelin were eliminated by vagotomy, suggesting that ghrelin does not act directly on the A2 neurons (26).

Although the ability of ghrelin to stimulate feeding and GH secretion by action within the hypothalamus itself is compelling and widely accepted, known functions of hindbrain catecholamine neurons are also consistent with this proposed role in mediating ghrelin’s actions. Some hindbrain catecholamine neurons, like ghrelin, exert potent stimulatory effects on food intake (31,32,33), influence GH secretion (34,35,36), and their activation increases expression of hypothalamic Npy and Agrp (37). However, although implicating A2 neurons in ghrelin-induced feeding, previous work (30) did not examine the potential contribution of hypothalamically projecting catecholamine neurons other than those in A2 to ghrelin-induced feeding. NE and epinephrine (E) neurons from other hindbrain cell groups not only contribute to the innervation of the hypothalamus (38,39,40), but a subpopulation of these non-A2 neurons is activated by glucose deficit (41,42,43), and the activation of these neurons stimulates food intake (40,44,45). Localized gene silencing or retrograde lesion of the hypothalamic NE/E projection with DSAP impairs this feeding response (40,44,45). Similarly, activation of hindbrain catecholamine neurons inhibits or stimulates GH secretion via hypothalamic α1 and α2 adrenergic receptors, respectively (34,35,36,46), but the specific catecholamine cell population involved and whether ghrelin activates this population are not known.

Therefore, the goal of the present study was to determine which hypothalamically projecting catecholamine cell groups are involved in these actions of ghrelin. We used double label immunohistochemistry to examine the distribution of hindbrain catecholamine neurons expressing Fos, an indicator of neuronal activation (47), in response to ghrelin. Our results indicate that hindbrain catecholamine neurons activated by ghrelin are concentrated in the ventrolateral medulla (A1, A1/C1, C1, and A5) but are not present in the A2 neurons suggested by Date et al. (30) to be critical for ghrelin-induced feeding. We also examined GH secretion and food intake after targeted retrograde lesion of hypothalamically projecting catecholamine neurons with DSAP. In contrast to Date et al. (30), we found that DSAP lesions that destroyed hypothalamically projecting catecholamine neurons did not alter the feeding response to either central or systemic ghrelin administration. However, the DSAP lesion caused a significant prolongation of the GH secretory response to central ghrelin administration, suggesting a role for NE in feedback inhibition of ghrelin-induced GH secretion. Finally, we measured GH-induced Fos expression in the hindbrain to determine whether activation of the catecholamine neurons observed after ghrelin administration could be mediated by GH itself. We found that the pattern of Fos expression was similar in ghrelin and GH-treated animals, supporting this possibility.

Materials and Methods

Animals

Adult male Sprague Dawley rats from Simonsen Laboratories (Gilroy, CA) were housed in suspended wire cages within a temperature controlled room illuminated between 0700 and 1900 h and used for all experiments. Rats had ad libitum access to water and standard pelleted rat chow (Harlan Teklad F6 Rodent Diet W; Harlan, Madison, WI). The Washington State University Institutional Animal Care and Use Committee, which conforms to National Institute of Health rules and regulations, approved all experimental animal protocol.

Experiment 1: Effect of DSAP lesions on ghrelin-induced feeding and GH secretion

In this experiment, DSAP or SAP was injected into the paraventricular nucleus of the hypothalamus (PVH), as described previously (40,45), to retrogradely lesion catecholamine neurons innervating medial hypothalamic nuclei. DSAP and SAP rats were subsequently tested for feeding and GH secretion responses to systemic and fourth ventricle (4V) administration of ghrelin. Feeding in response to 2-deoxy-d-glucose (2DG)-induced glucoprivation was used as a behavioral screening test to determine lesion effectiveness, because this response requires the catecholamine neurons targeted by the DSAP lesion (40,45). At the end of the experiment, the DSAP lesion was evaluated using immunohistochemical approaches.

DSAP and SAP lesions

Rats were anesthetized with chloropent anesthesia (3 ml/kg ip), made by combining 21.25 g of chloral hydrate, 10.6 g of magnesium sulfate, 4.43 g pentobarbital sodium, 75.26 ml ethyl alcohol, and 169.00 ml propylene glycol, brought to 500 ml with sterile double distilled H2O, and filtered. DSAP (42 ng/200 nal 0.1 m phosphate buffer; Advanced Targeting Systems, San Diego, CA) was stereotaxically microinjected over 8 min into the PVH (1.8 mm caudal to bregma, 0.4 mm lateral to the midline on both sides, 7.35 mm ventral to the dura mater) with a glass micropipette (30 μm tip diameter) attached to a Picospritzer with polyethylene tubing to lesion DBH-expressing neurons projecting to the PVH. Unconjugated SAP (42 ng/200 nal 0.1 m phosphate buffer; Advanced Targeting Systems) was administered in the same manner to control animals. The progress of the injection solution through the micropipette was monitored microscopically. The animals were allowed to recover from surgery for at least 2 wk before further experimentation.

4V cannula implantation

Cannulas were implanted into the 4V of SAP and DSAP-lesioned rats for administration of ghrelin and artificial cerebrospinal fluid (aCSF) control solution. Rats were anesthetized for cannula implantation with 1 ml/kg ketamine/xylazine/acepromazine cocktail [5 ml ketamine HCl, 100 mg/ml (Fort Dodge Animal Health, Fort Dodge, IA); 2.5 ml xylazine, 20 mg/ml (Ben Venue Laboratories, Bedford, OH); 1 ml acepromazine, 10 mg/ml (Vedco, Saint Joseph, MO); and 1.5 ml 0.9% saline solution]. Twenty-six gauge cannulas, occluded with stainless steel obturators, were implanted stereotaxically at the midline, 2.0 mm rostral to the occipital suture and 6.5 mm ventral to the dura mater. The animals recovered to their presurgical body weight before further experimentation.

2DG and ghrelin-induced feeding tests

Systemic feeding tests were conducted during the light phase, beginning at 1000 h in DSAP-treated (n = 14) and SAP-treated (n = 13) rats. There was at least 1 d between each feeding test. Glucoprivic feeding was measured in a 4-h test immediately after injection of 2DG (200 mg/kg, sc; Sigma, St. Louis, MO) or saline (0.9%, 1 ml/kg) to assess the effectiveness of the DSAP lesion. Subsequently, feeding in response to systemic ghrelin (15 μg/kg, ip; Peptide Institute, Osaka, Japan) or saline control injection (0.9%, 1 ml/kg, ip or sc) was measured in a 2-h test in DSAP- and SAP-treated rats. Feeding responses were also measured in different groups of DSAP (n = 8) and SAP (n = 6) rats at 1000 h in response to 4V ghrelin (150 pmol/1 μl or 30 pmol/200 nl) and aCSF (1 μl or 200 nl) to ensure that the systemic feeding response was not an artifact of ghrelin access to GHSRs at forebrain circumventricular organs. Solutions were delivered into the 4V using 33 gauge injectors connected with polyethylene tubing to a Hamilton glass syringe.

GH response to 4V ghrelin injection

Whole blood was collected for determination of GH concentrations in DSAP (n = 7) and SAP (n = 8) rats in response to 4V ghrelin (2 μg/6 μl) or aCSF (6 μl) injection. Samples (150 μl) were collected from the saphenous vein using glass capillary tubes at four time points: immediately before and 20, 40, and 60 min after ventricular injection. Samples were transferred to microcentrifuge tubes containing 4 μl 15% EDTA and promptly centrifuged for 10 min at 8000 rpm. Plasma was transferred to new microcentrifuge tubes and stored at −20 C until analyzed. Rat GH concentration was measured in plasma using an ELISA kit (Millipore Corp., Billerica, MA). The immunoassay has a sensitivity of 0.07 ng/ml and a 2.6% coefficient of variation for GH concentrations at baseline levels. Plates were read using an uQuant microplate (Biotek, Winooski, VT) reader with Gen5 software (Biotek).

Immunohistochemical analysis of DSAP lesions

After the completion of behavioral testing, rats were euthanized by deep anesthesia induced by inhalation of isoflurane and transcardially perfused with a 0.1 m PBS followed by 4% formaldehyde in PBS. The brains were immediately extracted and postfixed in 4% formaldehyde in PBS overnight and then submerged into 25% sucrose in PBS overnight. The hindbrain and hypothalamus were sectioned (30 μm) using a cryostat. Sections were collected sequentially into four vials with 0.1 m Tris-sodium phosphate buffer (TPBS) so that each vial contained a set of anatomically equivalent sections. The immunohistochemistry procedure was started immediately after sectioning.

Free-floating hindbrain and hypothalamic sections were processed for immunohistochemical detection of DBH. The sections were rinsed three times for 5 min in TPBS and blocked overnight in 10% normal horse serum (NHS) made in TPBS. After removing the blocking solution, the sections were incubated in mouse anti-DBH (1:50,000 for hindbrain tissue or 1:20,000 for hypothalamic tissue; Millipore Corp.) antibody diluted in 10% NHS for 48 h. This solution was removed, and the tissue was washed in TPBS (3× 5 min). Hypothalamic tissue was then incubated in biotinylated donkey antimouse IgG (1:500; Jackson ImmunoResearch, West Grove, PA) diluted in 1% NHS overnight, washed, and incubated in ExtrAvidin-Peroxidase (1:1500; Sigma) for 4 h. Hypothalamic tissue was washed and reacted for visualization of nickel-intensified diaminobenzidine to reveal black reaction product. Hindbrain tissue was incubated in Alexa 488 donkey antimouse secondary antibody for 3 h (Invitrogen, Carlsbad, CA) and was washed again, mounted on slides (VWR, West Chester, PA), and cover-slipped using Prolong Gold (Invitrogen).

DBH-immunoreactive (ir) terminals in the medial hypothalamus were evaluated in all DSAP and SAP animals with direct light microscopy. To quantify the effectiveness of the DSAP lesions, all DBH-ir cell bodies in one set of tissue were counted in the A1, A2, and C1 catecholaminergic cell groups between the calamus scriptorius to the caudal border of the facial nucleus in a random subset of rats (SAP, n = 6; DSAP, n = 5). Data from animals (n = 3) that did not have visibly decreased terminals innervating the hypothalamus and did not lose their responsiveness to 2DG (consumed ≥2 g over baseline) were not quantified and were removed from additional experimental analysis.

Experiment 2: Effect of ghrelin and GH on Fos expression in hindbrain catecholamine cell groups

The purpose of this experiment was to determine whether ghrelin and GH activate hindbrain catecholaminergic neurons in rats without immunotoxin injections and, if so, to examine the distribution of the activated neurons. We used double-label immunofluorescence to evaluate Fos expression in DBH-ir cells after ghrelin and GH treatment.

Lateral ventricle (LV) cannula implantation

Cannulas were stereotaxically implanted into the LV, as described above for 4V cannula implantation, for administration of ghrelin or aCSF. Cannulas were implanted 1.0 mm caudal to bregma, 1.5 mm lateral to the midline, and 3.9 mm ventral to the dura mater. The animals recovered to their presurgical body weight before further experimentation.

Ghrelin injection and tissue collection and preparation

Ghrelin (2 μg/6 μl; n = 5) or aCSF (6 μl; n = 2) was injected into the LV through cannulas prepared as described in experiment 1 and human recombinant GH (75 μg/100 g body weight; n = 4; A. F. Parlow, National Hormone and Peptide Program, Torrance, CA) or 0.9% saline (1 ml/kg; n = 2) was injected sc for induction of Fos signaling 90 min before killing. Rats were killed by deep anesthesia induced by inhalation of isoflurane and transcardially perfused with a 0.1 m PBS followed by 4% formaldehyde in PBS. The brains were immediately extracted and postfixed in 4% formaldehyde in PBS overnight and then submerged into 25% sucrose in PBS overnight. The hindbrain and hypothalamus were sectioned (30 μm) using a cryostat and collected sequentially into four vials containing TPBS so that each vial contained a set of anatomically equivalent sections. The immunofluorescence procedure was started immediately after sectioning was completed.

Immunohistochemical detection of Fos protein and DBH

Free-floating hindbrain sections were processed for immunohistochemical detection of DBH and Fos protein, using the protocol described for experiment 1. Mouse anti-DBH (1:50,000; Millipore Corp.) and rabbit anti-Fos (1:10,000, Ab5; EMD Biosciences, San Diego, CA) antibodies diluted in 10% NHS for 48 h were used as the primary antibodies. According to the manufacturer, the Fos antibody recognizes 55-kDa c-Fos and 62-kDa v-Fos proteins but does not cross-react with the 39-kDa Jun proteins. Secondary antibodies were Alexa 555 donkey antirabbit and Alexa 488 donkey antimouse (Invitrogen).

Quantification of Fos expression in DBH-ir neurons

All counting of the tissue was done immediately after staining to eliminate reduction of fluorescent signal. All DBH positive neurons in cell groups A1, A2, C1, C2, C3, A5, and the area postrema were counted in every section of one of the four sets of tissue. Immediately thereafter, neurons expressing both DBH and Fos in the same cell groups were counted using direct fluorescence microscopy by switching between red and green filters. We confirmed the number of coexpressing neurons in each cell group of each animal by comparing the quantified results obtained using direct microscopy with the number of neurons expressing both markers in photomicrographs. Cell group boundaries were defined according to Paxinos and Watson (48).

Statistical analysis

Data for experiments 1 and 2 are presented as mean ± sem and were analyzed using three-way or two-way ANOVA for repeated measures or Student’s t test as appropriate. Differences with P ≤ 0.05 were considered to be significant. When a significant F value was revealed by ANOVA, the Holm-Sidak post hoc test was used to isolate significant differences.

Results

Experiment 1

Lesion verification

Rats with DSAP injections exhibited reduced total numbers of DBH-ir neurons in A1 (t9 = 7.245, P < 0.001) and A1/C1 (t9 = 10.471, P < 0.001) cell groups compared with SAP controls (Fig. 1). Neurons in the A2 cell group form two morphologically distinct subtypes. The large multipolar cells predominately localized in the medial and commissural NTS were reduced by the DSAP lesion (t9 = 6.643, P < 0.001), as reported previously (40,49). Conversely, the small, round DBH-ir cells localized in the dorsal NTS were not reduced by the PVH DSAP injections (t9 = 0.968, P = 0.358) (49). DBH-ir terminals in medial hypothalamic nuclei (including ARC and PVH) of animals injected with DSAP were visibly reduced in comparison with those injected with SAP. An example of this reduction can be seen in Fig. 1.

Figure 1.

Figure 1

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Immunohistochemical and quantitative confirmation of DSAP lesion. The micrographs show typical DBH-ir in anatomically matched sections of a SAP control animal (A and C) and DSAP-lesioned animal (B and D). There was a dramatic reduction of DBH immunoreactivity in both Arc and PVH with DSAP treatment at both rostrocaudal levels depicted. Total hindbrain DBH-ir neurons were counted in A1, A2, and C1 cell groups for confirmation of reduced neuronal counts after DSAP lesion (E). DSAP rats (n = 5) have significantly fewer A1 and A1/C1 neurons than the SAP controls (n = 6). The reduction in A2 neurons was cell-type specific; the larger neurons in the medial NTS (A2-large) were reduced, whereas the smaller neurons in the dorsal strip and dorsal NTS (A2-small) were not significantly reduced. 3V, Third ventricle. *, P < 0.05 in comparison with SAP control. Scale bar, 500 μm.

Catecholamine neurons are essential for glucoprivic feeding but not ghrelin-induced feeding

Rats were tested for an intact glucoprivic feeding response after DSAP lesion (n = 14) or SAP (n = 13) control injection. Analysis of the results with two-way repeated measures ANOVA indicated that there were significant main effects for 2DG compared with saline control (F1,25 = 79.00, P < 0.001), as well as the lesion (F1,25 = 41.290, P < 0.001). There was an interaction between the two factors (F1,25 = 41.555, P < 0.001), indicating that the response to the 2DG treatment depends on whether or not rats were subjected to the DSAP lesion. Post hoc analysis revealed that 2DG significantly elevated feeding over baseline in SAP rats but not in DSAP rats (Fig. 2A).

Figure 2.

Figure 2

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Feeding in response to 2DG and ghrelin in 2- and 4-h tests. SAP rats increased their food intake significantly in response to 200 mg/kg 2DG than after 1 ml/kg saline control injection, whereas DSAP rats did not increase their intake significantly in response to 2DG (A). Both SAP and DSAP rats responded to ghrelin with elevated 2-h feeding whether ghrelin was injected systemically (15 μg/kg ghrelin or 1 ml/kg saline) (B) or into the 4V in different rats (SAP, n = 6; DSAP, n = 8) (C). There were no significant differences in ghrelin treatment groups in the SAP and DSAP animals. *, P < 0.05 vs. vehicle control.

DSAP and SAP rats were tested for ghrelin-induced feeding, and a two-way repeated measures ANOVA was used to analyze the resulting data. There was a significant main effect of ip ghrelin or saline treatment (F1,25 = 29.725, P < 0.001) but no main effect due to the DSAP lesion (F1,25 = 0.0000234, P = 0.996). Post hoc analysis revealed that both SAP and DSAP rats had significantly increased food intake 2 h after ip ghrelin treatment in comparison with the 2-h ip saline baseline (Fig. 2B). There was a significant main effect of the three levels of icv ghrelin treatment vs. the saline control (F2,22 = 37.558, P < 0.001), but there were no effects due to lesion status and no interaction between ghrelin treatment and lesion status. Increased food intake was also observed after 4V injection of ghrelin when compared with aCSF control (Fig. 2C). There was a slight, but not significant, trend toward enhanced icv ghrelin-induced feeding in DSAP rats over ghrelin-induced feeding in SAP rats.

GH secretion induced by 4V ghrelin administration was increased in DSAP-lesioned rats

Figure 3 shows plasma GH concentration in DSAP and SAP rats immediately before (0) and 20, 40, and 60 min after aCSF or ghrelin treatment. A significant interaction among the three factors (2 × 2 × 4), ghrelin treatment, DSAP lesion, and time was found with a three-way repeated measures ANOVA (F3,104 = 2.975, P = 0.035). The data at each time point was analyzed with two-way repeated measures ANOVA. There were no main effects due to treatment or lesion status at the 0- or 60-min time points. At 20 min after injection, there was a main effect for ghrelin treatment of the animal (F1,13 = 37.80, P < 0.001). At 40 min after treatment, there were significant main effects for both lesion status (F1,13 = 5.531, P = 0.035) and ghrelin treatment (F1,13 = 5.332, P = 0.038). Post hoc analysis indicated that ghrelin treatment resulted in elevated plasma GH levels over GH levels after saline treatment in DSAP rats at 20 and 40 min after treatment and at 20 min after treatment in SAP rats. Treatment of DSAP and SAP rats with aCSF did not result in any significant differences between sampling times or between groups.

Figure 3.

Figure 3

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Concentration (Conc) of rat plasma GH after icv ghrelin or control aCSF stimuli in SAP and DSAP rats. GH concentration in both SAP and DSAP rats peaked 20 min after treatment. The plasma GH concentration in SAP animals returned to baseline levels at 40 min, whereas that in DSAP animals remained elevated at near peak concentration at that time point. *, P < 0.05 DSAP ghrelin in comparison with DSAP aCSF; †, P < 0.05 for SAP ghrelin vs. SAP aCSF.

Experiment 2

Ghrelin and GH increase Fos expression in hindbrain catecholamine neurons

Typical immunofluorescent labeling can be seen in the A1/C1 and A2 cell groups in Fig. 4. Physiological saline and aCSF-treated animals were pooled into a vehicle group because they were not different. Higher levels of Fos expression in DBH-ir cells were observed in rats treated with ghrelin or GH than those treated with vehicle (Fig. 5). One-way ANOVA revealed significant differences between treatment groups in the percentage of cells coexpressing Fos and DBH in A1 (F2,10 = 12.95, P = 0.002), A1/C1 (F2,10 = 54.79, P < 0.001), C1 (F2,10 = 8.22, P = 0.008), and A5 (F2,10 = 53.95, P < 0.001) cell groups. Holm-Sidak post hoc analysis revealed that ghrelin significantly elevated Fos expression in DBH-positive neurons in A1, A1/C1, C1, and A5 cell groups and that sc GH significantly increased Fos and DBH coexpression in A1, A1/C1, and A5 cell groups, compared with vehicle control levels. There were no significant differences in Fos expression in A2, C2, C3, or area postrema catecholamine neurons in ghrelin-treated animals or in A2, C1, C2, C3, or area postrema catecholamine neurons in GH-treated animals.

Figure 4.

Figure 4

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Representative coronal brain sections showing Fos and DBH immunofluorescence in A1/C1 and A2 cell groups. Both GH and ghrelin increased the number of DBH positive neurons (green) that coexpress Fos-ir nuclei (red) in the A1/C1 region. Although there was a large increase in Fos expression in the NTS of ghrelin-treated animals, the majority of these neurons did not coexpress DBH immunofluorescence. Scale bar, 50 μm.

Figure 5.

Figure 5

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Coexpression of Fos and DBH after icv ghrelin (n = 5), sc GH (n = 4), or vehicle (n = 4) injections, shown as the percentage of DBH cell bodies in each cell group that were positive for Fos expression. *, P < 0.05 vs. vehicle control; †, P < 0.05 vs. ghrelin.

Discussion

Immunohistochemical results presented here reveal that ghrelin administration increased Fos expression in DBH-ir neurons in cell groups A1, in the area of A1/C1 overlap, to a limited extent in C1, and in A5. The largest percentage of activated cells was in cell group A1. Thus, NE neurons were the predominant catecholamine phenotype activated by ghrelin. We also found that ghrelin increased Fos expression in the NTS, as others have found (50), but Fos was not expressed in A2 neurons. The absence of ghrelin-induced Fos expression in A2 neurons is consistent with other results (51,52). Failure of A2 neurons to express Fos in response to ghrelin does not eliminate the possibility that ghrelin’s effects are mediated by inhibition of these neurons. However, absence of Fos expression in A2 neurons was surprising, because Date et al. (30) reported that ghrelin increased DBH mRNA in A2 neurons and increased hypothalamic catecholamine release, suggesting that they are activated, not inhibited, by ghrelin.

Cell groups A1 and caudal C1, where DBH- and Fos-ir were coexpressed after ghrelin administration, provide dense innervation of the medial hypothalamus, including the PVH and Arc (38,39,40,45). Stimulation of feeding in response to glucose deficit is an important function of neurons in these cell groups (40,44,45). Injections of DSAP into either the Arc or PVH virtually eliminate DBH terminals throughout the medial hypothalamus, reduce cell numbers in catecholamine cell groups (mainly A1, A2, and caudal C1), and significantly reduce or eliminate glucoprivic feeding (37,40,45). In the present study, we used the glucoprivic feeding response as an independent behavioral test of the DSAP-induced lesion of the hypothalamic NE/E projection. We found that DSAP lesions eliminated the glucoprivic feeding response in animals in which the immunohistochemical analysis of the hypothalamus showed the typical pattern of DSAP-induced loss of DBH terminals in Arc, PVH, and other medial hypothalamic sites, as well as reduction in the number of hindbrain cell bodies in the cell groups known to contribute to the hypothalamic innervation. Judged by these two criteria, the DSAP lesion was effective in 14 of 17 rats. However, this lesion did not impair the feeding response to either central or systemic ghrelin administration. Together, these findings fail to support the hypothesis, proposed previously by Date et al. (30), that ghrelin-induced feeding requires NE neurons that innervate the Arc. It should be noted that even hypothalamically projecting NE and E neurons that might be inhibited by ghrelin would have been lesioned by the DSAP injection, and therefore, an essential role for such neurons in ghrelin-induced feeding can also be ruled out.

The discrepancy between our DSAP data and that reported by Date et al. (30) might be attributable to the difference in the DSAP injection sites in the two experiments. In Date et al.’s experiment, DSAP was injected into the Arc, whereas in ours, DSAP was injected into the PVH. Differences in the pattern of DBH-terminal loss are not likely to have caused the differences in experimental outcomes, because injection of DSAP into either site causes a loss of DBH terminals in the Arc and medial hypothalamus. However, with injections into the Arc, nonspecific damage to Arc ghrelin-sensitive neurons, rather than the loss of NE cells and terminals, could conceivably have accounted for the failure of the DSAP-injected rats to respond to ghrelin. Published findings using a variety of approaches have shown that Arc lesions abolish the feeding responses to central and peripheral ghrelin administration (23,24). Evidence helping to rule out this possibility was not provided by Date et al. (30).

Date et al. (30) also supported their hypothesis that hindbrain NE neurons mediate ghrelin-induced feeding by demonstrating that this response was reduced after midbrain transection of the ascending NE fiber bundle. These data are also inconclusive, however, because midbrain transection eliminates both afferent and efferent fiber pathways between the forebrain and hindbrain, such that the reduced feeding could be due to the elimination of efferent fibers essential for appetitive behavior and overall motor function. Without data showing that the midbrain transected animals were capable of maintaining their body weight and of responding to other appetitive stimuli, it is not possible to interpret the loss of the feeding response to ghrelin as being a specific consequence of NE fiber damage.

It has been suggested previously (51) that ghrelin enhances feeding by antagonizing the effects of peripheral satiety peptides, such as cholecystokinin, on vagal sensory neurons. The signal generated by this interaction is then proposed to be transmitted to the NTS, where vagal sensory fibers terminate, and then to the hypothalamus by A2 neurons. However, our data show that DSAP lesions, which eliminate A2 neurons that project to the hypothalamus, neither change the magnitude of ghrelin-induced feeding (this experiment) nor impair the response to cholecystokinin (40). Thus, if ghrelin stimulates feeding by antagonizing vagally mediated satiety signals or by activating A2 neurons that do not express Fos, central transmission of this effect does not require NE neurons that innervate the hypothalamus. However, A2 neurons projecting to other sites, such as the small-A2 neurons described in this study, could still be involved. It is important to note, however, that the importance of the vagal sensory neurons in ghrelin-induced feeding and GH secretion has been challenged by conflicting bodies of evidence both in rats and in humans (26,53,54,55).

Ghrelin is a GH secretagogue capable of acting directly on GH-secreting cells in the pituitary and on parvocellular GHRH cells in the Arc. Release of NE at the level of the hypothalamus is capable of producing either excitatory or inhibitory effects on GH secretion, depending on the receptor type activated, but the predominant effect of NE appears to be the inhibitory effect mediated by α-1 adrenoreceptor activation (36). In our study, we found that GH secretion after ghrelin administration was prolonged in DSAP-treated animals. This indicates that hindbrain catecholamine neurons suppress GH release. Our finding that GH induces Fos expression in many of the same catecholaminergic cell groups as ghrelin indicates that the NE neurons in the ventrolateral medulla likely react to high blood GH concentration as a regulatory component of GH negative feedback. The inability to respond appropriately to high GH levels could result in a chronic overexposure of GH, which would result in increased anabolism and growth. In fact, DSAP animals tend to gain more weight than their SAP counterparts (40), an effect that could potentially be due to the dysregulation of GH.

Glucocorticoids and stress have been shown to inhibit GH secretion (56). Therefore, if our rats were stressed by the blood sampling protocol, then nonspecific damage to PVH neurons expressing corticotrophin-releasing factor (CRF) could account for the prolonged ghrelin-induced GH secretion that we observed in DSAP rats. However, in previously published work, we extensively investigated the possibility of nonspecific damage to these neurons by PVH DSAP (57). Using the same injection protocol described here, combined with in situ hybridization and immunohistochemistry, we found that the DSAP injection did not reduce CRF gene expression or protein in the PVH. In addition, PVH DSAP injections did not impair either the circadian rhythm of corticosterone secretion or the response to swim stress. These data demonstrate that the PVH DSAP lesion does not produce significant nonspecific damage to the CRF neurons. Therefore, we believe that it is unlikely that nonspecific damage in combination with any stress produced by the blood sampling protocol, in which a total of only 600 μl was collected, is accountable for our finding.

In summary, the present data indicate that a population of catecholamine neurons with projections to the medial hypothalamus serves as a negative feedback mechanism to modulate ghrelin-induced GH release, and possibly to control GH secretion induced by other stimuli as well. However, our data do not indicate a role for hypothalamically projecting catecholamine neurons in ghrelin-induced feeding and certainly do not support the proposal that they are required for this response. Because injection of ghrelin directly into the Arc stimulates feeding (11), ghrelin receptors are present in the Arc and accessible to circulating ghrelin (4) and Arc lesions impair feeding induced by both systemic administration of ghrelin agonist (24) and icv ghrelin (23), it seems most likely from present data that circulating ghrelin, or arguably ghrelin produced in the hypothalamus (25,58), induces feeding primarily through its direct actions in the Arc.

Footnotes

This work was supported by National Institutes of Health Grants DK 40498 and DK00345072 (to S.R.).

Disclosure Summary: The authors have nothing to disclose.

First Published Online May 12, 2010

Abbreviations: aCSF, Artificial cerebrospinal fluid; AGRP, Agoutigene-related peptide; Arc, arcuate nucleus of the hypothalamus; CRF, corticotrophin-releasing factor; DBH, dopamine-β-hydroxylase; 2DG, 2-deoxy-d-glucose; DSAP, SAP conjugated to a monoclonal antibody against DBH; E, epinephrine; GHSR, GH secretogogue receptor; icv, intracerebroventricular; ir, immunoreactive; LV, lateral ventricle; NE, noradrenergic NHS, normal horse serum; NPY, neuropeptide Y; NTS, nucleus of the solitary tract; PVH, paraventricular nucleus of the hypothalamus; SAP, saporin; TPBS, Tris-sodium phosphate buffer; 4V, fourth ventricle.

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