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Endocrinology, doi:10.1210/en.2007-1164
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Endocrinology Vol. 149, No. 5 2306-2312
Copyright © 2008 by The Endocrine Society

The Gastrointestinal Hormone Ghrelin Modulates Inhibitory Neurotransmission in Deep Laminae of Mouse Spinal Cord Dorsal Horn

Angela M. Vergnano, Francesco Ferrini, Chiara Salio, Laura Lossi, Mario Baratta and Adalberto Merighi

Department of Veterinary Morphophysiology (A.M.V., F.F., C.S., L.L., M.B., A.M.), University of Turin, 10095 Grugliasco (Torino) Italy; and Istituto Nazionale di Neuroscienze (L.L., A.M.), 10125 Torino, Italy

Address all correspondence and requests for reprints to: Adalberto Merighi, Department of Veterinary Morphophysiology, Via Leonardo da Vinci 44, 10095 Grugliasco (TO), Italy. E-mail: adalberto.merighi{at}unito.it.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Ghrelin is mainly described for its effects on feeding behavior and metabolism. However, central nervous system distribution of its receptor [type 1a GH secretagogue receptor (GHSR)] and modulation of neurotransmission in the hypothalamus suggest broader effects than originally predicted. Systemically administrated ghrelin inhibits inflammatory pain after behavioral observations. Therefore, we investigated the expression and function of type 1a GHSR in mouse spinal cord by molecular biology, biochemistry, histology, and electrophysiology. The mRNA and protein were detected in tissue extracts by RT-PCR and Western blotting. In situ, receptor mRNA and immunoreactivity were localized to cell bodies within the medial aspect of the deep dorsal horn. Patch clamp recordings on laminae IV–VI demonstrated that bath-applied ghrelin (100 nM) induced a strong increase of spontaneous {gamma}-aminobutyric acid/glycine-mediated current frequency (463 ± 93% of the control) and amplitude (150 ± 16% of the control) in about 60% of recorded neurons. Specificity of type 1a GHSR activation was confirmed by the lack of effect of the deacylated form of ghrelin (des-acyl-ghrelin) and after preincubation with the specific receptor antagonist [D-Lys3]GHRP-6. In the presence of tetrodotoxin, the effect of the peptide was strongly reduced, mainly indicating an action potential-dependent mechanism. The functional link between ghrelin and pain was confirmed by inhibition in vitro of the c-fos response to capsaicin activation of nociceptive fibers, after quantification of Fos-immunoreactive nuclei in laminae IV–VI. Our results are the first demonstration of the presence of functional type 1a GHSRs in the spinal cord and indicate that ghrelin may exert antinociceptive effects by directly increasing inhibitory neurotransmission in a subset of deep dorsal horn neurons.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
GHRELIN, AN ENDOGENOUS peptide ligand of the GH secretagogue receptor (GHSR), has gained increasing attention as a brain-gut hormone with GH-releasing and appetite-inducing functions (1). Ghrelin was originally synthesized in vitro (1, 2) and subsequently shown to be mainly produced by X/A-like cells of the oxyntic stomach mucosa (3, 4). The peptide is biologically active in acylated form (5). However, most circulating ghrelin (~80–90%) is deacylated (des-acyl-ghrelin), and still it remains unclear whether des-acyl-ghrelin represents a precursor or a degradation product of the acylated peptide (1, 6).

The GHSR is a typical G protein-coupled receptor. Two cDNAs encoding for two different GHSR isoforms were cloned, and named types 1a and 1b (7). Type 1a encodes for the full-length biologically active receptor, whereas type 1b encodes for a truncated isoform. Affinity studies have shown that GH secretagogues only bind GHSR type 1a. Nonetheless, in HEK293 cells coexpressing full-length and truncated receptors, it has been recently observed that type 1b may play a regulatory/inhibitory role on type 1a receptor activity (8).

Whereas ghrelin is present at low levels in the brain (9, 10), the GHSR type 1a protein is highly expressed in the arcuate and ventromedial nuclei of the hypothalamus (11), substantial levels having also been found in the hippocampus, dentate gyrus, and piriform cortex (12). In addition, Zigman et al. (13) have demonstrated a strong mRNA signal in several brainstem nuclei. These authors have also investigated the cervical spinal cord but failed to find any significant in situ hybridization signal.

Functionally, brain ghrelin was initially found to modulate synaptic transmission and regulate neuron excitability in the hypothalamic nuclei involved in the central control of feeding (10, 14, 15), but it was more recently observed that the peptide also promotes long-term potentiation in the hippocampus (16). Although it is still under debate whether or not circulating ghrelin may cross the blood-brain barrier, thereby reaching appropriate concentrations in the central nervous system (17), Diano et al. (16) have provided some experimental observations in support.

No studies have been performed so far concerning the involvement of ghrelin in central spinal cord sensory mechanisms. Nonetheless, ip, intraplantar, and intracerebroventricular administrations of ghrelin in rat reduce the hyperalgesia and edema provoked by intraplantar injections of carrageenan (18).

In an attempt to investigate the role of ghrelin in spinal cord neurotransmission, we analyzed the expression and activity of the peptide and type 1a GHSR in dorsal horn (DH) neurons.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals
All experimental procedures were approved by the Committee of Bioethics and Animal Welfare of the University of Torino, and were maintained according to the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Postnatal (P8-P14; n = 57) or adult (P > 30; n = 5) mice were deeply anesthetized with a lethal dose of pentobarbital (30 mg/kg). The cervical/lumbar spinal cord, hypothalamus, and stomach were quickly dissected out and further processed as follows: 1) deep frozen in liquid nitrogen and powdered in a mortar for tube RT-PCR or Western blotting; 2) placed in oxygenated artificial cerebrospinal fluid (ACSF) (in mM: 125 NaCl, 2.5 KCl, 25 NaHCO3, 1 NaHPO4, 25 glucose, 1 MgCl2, and 2 CaCl2) and sliced at 300 µm in a transversal plane for patch clamp recordings or in vitro capsaicin challenge and analysis of c-fos response (spinal cord only); and 3) fixed in 4% paraformaldehyde in 0.1 M phosphate buffer (pH 7.2), dehydrated, and embedded in paraffin wax for histological procedures.

RT-PCR
For the tube procedure, total RNA was extracted by RNAWiz (Ambion, Inc., Austin, TX) with a standard protocol. RT was performed by a Ready-to-Go First strand kit (Amersham Pharmacia Biotech, Uppsala, Sweden), amplification with Ready-to-Go PCR beads (Amersham Pharmacia Biotech). GHSR primers were: forward 5'-CTGCTCTGCAAACTCTTCCA, reverse 5'-GAGCACAGTGAGGCAGAAGA; for controls 18S PCR Alternate Primer Pair (5 µM; Ambion) and 18S PCR Competitimers (5 µM) (ratio 3:7) were used. Amplification consisted of 33 cycles (denaturation 94 C for 60 sec, annealing 64 C for 70 sec, extension 72 C for 90 sec). PCR products were visualized with ethidium bromide after electrophoresis on 2% agarose gel. The size of the amplified cDNAs for GHSR and 18S was 354 and 323 bp, respectively. The in situ procedure was performed on dewaxed sections after pretreatments with proteinase K and triethanolamine. RT-PCR was performed by the Reverse Transcription System with Oligo(dT)15 primer (Promega Corp., Madison, WI), amplification with PCR master mix (Promega), 11-digoxigenin-d-uridine 5'-triphosphate (1 mM), and the GHSR primers used previously. Amplification was the same as the tube procedure. To reveal the amplification products, slices were then incubated in antidigoxigenin goat antibodies conjugated with alkaline phosphatase (Roche Diagnostics, Mannheim, Germany), followed by the nitro blue tetrazolium/5-bromo-3-indolylphosphate-p-toluidine salt.

Immunocytochemistry
Localization of type 1a GHSR was performed on free-floating sections after incubation in a rabbit polyclonal antibody (1:500) directed against the C terminus of the rat type 1a GHSR (Alpha Diagnostics International Inc., San Antonio, TX). A rabbit anti-c-Fos antibody (1:100; Abcam plc, Cambridge, UK) was used to detect immunoreactivity in paraffin sections obtained from acute slices subjected to capsaicin challenge (see Results). In both instances standard avidin-biotin complex (ABC) procedures were used.

Western blotting
Proteins were extracted with standard procedures in general lysis buffer containing protease inhibitors (1 mM phenylmethylsulfonylfluoride, 1 mg/ml leupeptin, and 5 mg/ml aprotinin). Lysates were separated by SDS-PAGE in 10% acrylamide gels and blotted onto nitrocellulose paper. Blots were blocked with Blotto and probed with the antitype 1a GHSR antibody (1:250), followed by peroxidase-labeled antirabbit antibody (1:15,000; Promega). Protein bands were detected by an enhanced chemiluminescence Western blotting system (Amersham Biosciences Inc., Piscataway, NJ).

Specificity controls for type 1a GHSR were obtained by preincubating the primary antibody with its control peptide (Alpha Diagnostics International).

Electrophysiology
Patch clamp recordings were performed on DH neurons of postnatal mouse spinal cord, as previously described (19). Briefly, slices were placed in a recording chamber constantly perfused with oxygenated ACSF. Whole-cell recordings were obtained from DH neurons held at –63 mV. Spontaneous excitatory postsynaptic currents (sEPSCs) were recorded with a low chloride intracellular solution containing (in mM): 145 Csmethansulfonate, 5 EGTA, 2 MgCl2, 10 HEPES, 2 ATP.Na, 0.2 GTP.Na (pH 7.2) (with CsOH). Spontaneous inhibitory postsynaptic currents (sIPSCs) were recorded with a high chloride intracellular solution containing (in mM): 145 KCl or CsCl, 5 EGTA, 2 MgCl2, 10 HEPES, 2 ATP.Na, 0.2 GTP.Na (pH 7.2) (with KOH or CsOH), and 0.1% Lucifer yellow (LY) (Sigma-Aldrich, St. Louis, MO) to allow further morphological analysis. Neurons were classified as ghrelin responsive if the interevent interval differences were statistically significant (Kolmogorov-Smirnov test, P < 0.01) and if there were more than 20% increases in sIPSC frequency. The Wilcoxon matched-pairs test was used to identify statistical significance among groups. Tests were performed on rough values, unless otherwise indicated. Electrophysiological data are expressed as a percentage of the predrug control value ± SEM, with n indicating the number of neurons. Data were considered significantly different when P < 0.05.

All drugs were bath applied. Ghrelin (human), Des-acyl-ghrelin, [D-Lys3]GHRP-6, NBQX were from Tocris (Bristol, UK); tetrodotoxin, bicuculline methiodide, and strychnine methiodide were from Sigma-Aldrich. Ghrelin was used at 100 nM (10).

Fos response to slice functional stimulation
Spinal cord acute slices were maintained in oxygenated ACSF for 30 min at room temperature to recover from cutting, and then subjected to one of the following experimental treatments: 1) maintained in ACSF for a further 3 h (control); 2) incubated with capsaicin 1 µM for 10 min and then washed in ACSF for 3 h; and 3) preincubated with ghrelin 100 nM for 30 min, then incubated with capsaicin and washed as previously described in the constant presence of ghrelin. Slices were then fixed, dehydrated, embedded, and cut to be processed further for Fos immunoreactivity. Immunostained sections were photographed with a x20 lens, and the cell density (number of cells per mm2) within the superficial (laminae I–III) and deep (laminae IV–VI) DH was calculated with the ImageJ software (National Institutes of Health, Bethesda, MD). Statistics was performed with one-way ANOVA (Bonferroni post hoc, P < 0.05). Data are expressed as means ± SEM, with n indicating the number of slices.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Type 1a GHSR mRNA and protein are expressed in spinal cord
Type 1a GHSR mRNA was detected in the spinal cord after tube and in situ RT-PCR (Fig. 1AGo, lane b). The corresponding band was also detected in extracts of hypothalamus (Fig. 1AGo, lane a) and gastric mucosa (Fig. 1AGo, lane c), albeit at different levels of expression. Type 1a GHSR protein was detected in Western blots from spinal cord extracts (Fig. 1BGo, lane a), and staining was specifically abolished after preabsorption (Fig. 1BGo, lane b). After RT-PCR in situ, the mRNA was observed in neuronal cell bodies scattered across several laminae at the base and neck of the DH (Fig. 1CGo). These neurons were of large size and mainly located in the medial aspect of the DH, close to the dorsal funiculus. The reaction product was also observed at the level of the motor neurons in lamina IX (data not shown). There was no positive reaction in glia.


Figure 1
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  		FIG. 1. Type 1a GHSR expression in mouse spinal cord. A, RT-PCR amplified products from total mRNA extracts of the hypothalamus (a), spinal cord (b), and gastric mucosa (c). The corresponding amplified 18S products are shown in lanes d–f. B, Western blotting of spinal cord protein extracts after immunodetection with the same antitype 1a GHSR primary antibody used for histological procedures. A band of about 35 x 103 is shown in lane a; no signal is detected after preabsorption of the primary antibody with its control peptide (lane b). C, Localization of neurons expressing the type 1a GHSR mRNA after in situ RT-PCR. The nitro blue tetrazolium-5-bromo-3-indolylphosphate-p-toluidine salt reaction product (blue) is clearly seen in a group of neurons at the limits between laminae IV and V in the medial aspect of the DH. D and E, Immunocytochemical localization of type 1a GHSR protein in the spinal cord. D, Three immunopositive neurons are shown in laminae V–VI. The large immunoreactive cell body in lamina VI (circled) is shown at higher magnification in E to display better its major dendritic trunks (arrowheads). F, Immunoreactive neurons in the arcuate nucleus (arc). Two neuronal cell bodies (circled) are shown at higher magnifications in the insets. Roman numerals indicate the laminae of the DH. Bars, C and D = 100 µm; E = 30 µm; F = 50 µm; insets = 10 µm. bdh, Base of the DH; df, dorsal funiculus; me, median eminence.

 
After immunohistochemical labeling, positive neurons of corresponding sizes were observed in the same locations (Fig 1Go, D and E). Immunopositive cells tended to be multipolar with round or fusiform cell bodies and a few large dendritic trunks (Fig. 1EGo). There was no staining in glia. Antibody specificity was confirmed after staining of hypothalamic sections cut through the arcuate nucleus as a positive control (Fig. 1FGo).

Deep DH neurons respond to ghrelin with an increase of sIPSCs
Whereas ghrelin had no statistically significant effects on sEPSCs (n = 5; frequency: 103 ± 8% of control, P > 0.05; amplitude: 97 ± 2% of control, P > 0.05), the peptide strongly affected sIPSCs in the presence of 10 µM NBQX (n = 36; frequency: 328 ± 63% of control, P < 0.0001; amplitude: 128 ± 11% of control, P > 0.05; Fig. 2Go, A–D). Addition of bicuculline (10 µM) and strychnine (2 µM) completely suppressed all ongoing events (n = 5), thus confirming their inhibitory nature. In detail, frequency increase was clearly detected in 22 out of the 36 neurons recorded (Fig. 2Go, C and D, black circles). The effect was reversible upon washout (Fig. 2EGo, hatched bars): frequency and amplitude during washout were 121 ± 15% (n = 15; P > 0.05) and 116 ± 6% (P > 0.05) of control, respectively. In five responsive neurons, we also observed a small inward current of 6.6 ± 0.5 pA under ghrelin. There were 13 responsive neurons filled with LY. They were located in the medial aspect of laminae IV–VI, had round to fusiform cells bodies of medium-to-large size (20.3 ± 1.3 µm latero-lateral diameter and 16.0 ± 1.4 µm dorsoventral diameter), giving stem to a few major dendritic trunks (Fig. 2Go, F and G). The dendritic arborization had an overall spheroid distribution, with numerous branches directed toward the superficial aspect of the DH.


Figure 2
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  		FIG. 2. Effect of ghrelin (ghre) onto sIPSCs recorded from deep DH neurons. A, sIPSCs recorded from a representative neuron responding to bath-applied ghrelin (arrow). B, Time course of the recording (same neuron as in A). In this cell sIPSC increase took place after about 6-min ghrelin perfusion. C and D, sIPSC frequency (C) and amplitude (D) values from 22 ghrelin responsive (black circles) and 14 ghrelin unresponsive (open circles) neurons in control (CTR) and in the presence of ghrelin (ghre). E, Pooled data of frequency (top; 463 ± 93% of control, n = 22; P < 0.001) and amplitude (bottom; 150 ± 16% of control; P < 0.05) from responsive neurons (n = 22). Values are normalized with respect to the control. sIPSC increase is reversible upon washout (hatched bars). F, Morphology of a ghrelin-responsive neuron. G, Laminar distribution of LY injected responsive neurons. ***, P < 0.001. *, P < 0.05. cc, Central canal.

 
Although histological results were strongly indicative of the absence of type 1a GHSR mRNA and protein in the superficial DH, we also investigated the effects of ghrelin onto lamina II neurons, giving the pivotal role of these cells in the first processing of nociceptive information. The peptide did not significantly change the frequency (107 ± 12% of control) and amplitude (106 ± 6%; n = 8; P > 0.05) of sEPSCs. The lack of effects was confirmed also for sIPSCs (frequency: 106 ± 26% of control; amplitude: 106 ± 7% of control) upon blockade of glutamate neurotransmission in the presence of 10 µM NBQX (n = 6; P > 0.05) (data not shown).

To investigate further the specificity of the effect, we performed experiments using the competitive GHSR antagonist [D-Lys3]GHRP-6 (20) in seven neurons previously shown to be responsive to ghrelin (Fig. 3Go, A–C). [D-Lys3]GHRP-6 (10–4 M) reduced per se the frequency of sIPSCs to 67 ± 11% (P < 0.05), without changing their amplitude (102 ± 10%; P > 0.05) and elicited a slow outward current (30.4 ± 6.8 pA) in three neurons. Frequency and amplitude under ghrelin plus [D-Lys3]GHRP-6 were not significantly affected (Fig. 3Go, B and C, white bars), and the effect of ghrelin was reduced by 77 ± 11% after preincubation with the antagonist (P < 0.05).


Figure 3
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  		FIG. 3. Specificity of the ghrelin effect (A–E) and action potential involvement (F and G). A, Time course of sIPSCs under ghrelin, before and after superfusion with GHS receptor antagonist [D-Lys3]GHRP-6 (D-lys). B and C, Blockade of ghrelin effect in seven responsive neurons. Frequency (B) and amplitude (C) under ghrelin are 807 ± 209% (P < 0.05) and 170 ± 41% (P > 0.05) of control, respectively (black bars). Frequency and amplitude under ghrelin plus the antagonist are 283 ± 154% (P > 0.05) and 114 ± 8% (P > 0.05) of control, respectively (white bars). D, A representative trace showing a transient effect of des-acyl-ghrelin (Des-ghre) (upper trace). The same neuron strongly responded to a subsequent administration of ghrelin. E, Pooled data of frequency (top) and amplitude (bottom) from 10 neurons under des-acyl-ghrelin (white bars) and ghrelin (black bars). F and G, mIPSC frequency and amplitude values from 10 neurons from deep laminae of the DH in control (CTR) and in the presence of ghrelin (ghre). mIPSC increase has been detected in three neurons (black circles). **, P < 0.01. *, P < 0.05.

 
The effect of des-acyl-ghrelin was investigated on 10 neurons that were subsequently shown to be responsive to ghrelin (Fig. 3Go, D and E). In these experiments frequency and amplitude under des-acyl-ghrelin were not significantly affected [293 ± 157% (P > 0.1) and 105 ± 8% (P > 0.9) of control; Fig. 3EGo, white bars]. Frequency and amplitude under ghrelin were 628 ± 246% (P < 0.01) and 122 ± 11% (P > 0.05) of control (Fig. 3EGo, black bars). Moreover, frequency values under ghrelin and des-acyl-ghrelin normalized to their respective controls were found to be significantly different (n = 10; P < 0.01).

Some recordings were performed in the presence of tetrodotoxin (1 µM) to block action potential sodium-mediated neurotransmission, given the axonal distribution of GHSR reported in the hypothalamus (10). In these experiments ghrelin had no effects on miniature IPSCs (mIPSCs) (Fig. 3Go, F and G). Frequency and amplitude were 198 ± 57% (P > 0.05; n = 10) and 109 ± 8% (P > 0.05; n = 10) of control, respectively.

Ghrelin prevents the capsaicin-induced increase of Fos immunoreactivity in the deep DH
Capsaicin, the pungent vanilloid of hot chili pepper, specifically activates peptidergic nociceptive primary afferent terminals in the DH (21), and several in vivo studies have shown that peripheral injections of capsaicin induce the expression of the early gene c-fos in DH (e.g. see Ref. 21). To assess whether or not the increase of inhibitory neurotransmission observed after patch clamp recordings of deep DH neurons could be more directly related to a pain stimulus, we have challenged slices with capsaicin with or without the concurrent application of ghrelin (Fig. 4Go). Incubation with capsaicin induced a significant 2-fold increase of Fos immunoreactivity in deep DH neurons (58.6 ± 8.4 cells per mm2, n = 24) compared with controls (24.6 ± 3.6 cells per mm2, n = 24; one-way ANOVA, P < 0.001; Bonferroni post hoc test, P < 0.001). Notably, in the presence of ghrelin, the effect of capsaicin was prevented (27.1 ± 5.1 cells per mm2, n = 24; Bonferroni post hoc, P > 0.05).


Figure 4
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  		FIG. 4. Ghrelin inhibition of the capsaicin (CAP)-induced Fos response in the deep DH. A, Fos immunoreactivity in the spinal DH under control (CTR) condition. The insets on the right show Fos immunoreactivity in deep DH neurons observed in controls (CTR), after capsaicin administration (CAP), and after capsaicin administration in the presence of ghrelin 100 nM (CAP + GHRE). Microscopic fields were taken from the regions indicated by the dashed lines in A. Arrows indicate immunoreactive nuclei. B, Mean density of Fos immunoreactive nuclei observed in the deep DH under control condition (black bar) compared with that observed in capsaicin and capsaicin plus ghrelin experiments (white bars; one-way ANOVA, P < 0.001). A significant increase in cell density was observed after capsaicin, but not after capsaicin in the presence of ghrelin. ***, P < 0.001. Bars, A = 100 µm; insets = 50 µm. WM, White matter; CC, central canal.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Expression of type 1a GHSR in the spinal cord
The present study is the first demonstration that functional type 1a GHSRs are expressed in the mammalian spinal cord and increase inhibitory neurotransmission in deep laminae of the DH, as a consequence of an augmented {gamma}-aminobutyric acid/glycine release.

The mRNA and protein detected by RT-PCR and Western blotting showed the presence of type 1a GHSRs in spinal cord extracts from cervical and lumbar enlargements. Histologically, by in situ RT-PCR and immunocytochemistry, we localized the neurons containing the mRNA and protein in deep laminae of the DH. It was previously reported that several problems may be related to the use of RT-PCR for detection of GHSR isoforms (22). In addition, a recent in situ hybridization study failed to detect the GHSR mRNA at the cervical spinal cord level (13). However, biochemical and histological data are corroborated by patch clamp experiments, in which ghrelin induces a strong enhancement of sIPSC frequency in about 60% laminae IV–VI neurons, an effect that, in a subpopulation of cells, occurs together with a significant amplitude increase. The strong reduction of ghrelin response in the presence of the antagonist [D-Lys3]GHRP-6 and the lack of significant effects of des-acyl-ghrelin confirms the specific activation of type 1a GHSR. However, it should be mentioned that the antagonist was not capable to block completely the response to ghrelin in all neurons, likely as a consequence of weak receptor affinity (23), and that a subset of ghrelin-responsive neurons also showed an increase in sIPSC frequency under des-acylghrelin (Fig. 3DGo). Therefore, interactions with other receptor systems or with an unknown additional ghrelin receptor cannot be excluded.

The results on spontaneous and mIPSCs indicate that the effect of the ghrelin is mainly due to an action potential-dependent presynaptic release of inhibitory neurotransmitters. Although the present data are not conclusive about the neuronal circuits involved, it seems reasonable that release occurs directly from neurons expressing type 1a GHSRs, with or without subsequent activation of local circuit interneurons. Nonetheless, the slow inward currents observed in some neurons after peptide stimulation, and the slow outward currents under GHSR antagonist are also consistent with a postsynaptic effect. These considerations are strengthened by the significant reduction of sIPSC frequency in the presence of the antagonist alone. Thus, one is tempted to speculate that the GHSR type 1a may be tonically active in the spinal cord.

Functional implications
Sibilia et al. (18) have recently observed that systemically administrated ghrelin inhibits inflammatory pain in rats. Their behavioral observations were not informative about the site and mechanism of action of the peptide. Our present results offer a functional demonstration that: 1) ghrelin modulates inhibitory neurotransmission in laminae IV–VI of the DH; and 2) in these laminae the peptide inhibits in vitro the c-fos response to capsaicin, which specifically activates nociceptive circuitry in the spinal cord (20).

A direct correlation with painful states can be hardly demonstrated in a slice preparation, nonetheless, our results strongly implicate ghrelin as a central modulator of pain in deep DH neurons. Functionally, many neurons in this area of the spinal cord are wide dynamic range neurons, responding to both noxious and non-noxious stimuli (see Ref. 24). After LY filling, ghrelin-responsive neurons are relatively large multipolar cells with spherically shaped dendritic arbors. Some of them clearly fit into the category of the "antenna" cells representing the major output of lamina II neurons. These neurons are known to receive a direct nociceptive input at their dendrites in laminae I–II from somatic and visceral fine afferent fibers or an indirect feed-forward inhibitory/excitatory input from laminae I–III upon capsaicin-induced activation of polysynaptic circuits (24).

The cell body size of ghrelin-responsive neurons and of type 1a GHSR mRNA and protein positive cells, indicates them as projection neurons. Most projection neurons in lamina V are involved in the transmission of noxious stimuli to the thalamus, lateral cervical nucleus, dorsal column nuclei, reticular formation, and cerebellum. Among these, spinoreticular neurons are traditionally involved in arousal and autonomic activation after a noxious stimulus, and subjected to plastic changes in chronic painful states (24). Thus, it seems reasonable that ghrelin-responsive neurons are wide dynamic range and/or nociceptive-specific neurons. The inhibitory tone of these neurons may be increased not only when the peptide is bath applied on slices and, thus, has direct access to spinal cord type 1a GHSRs (as also confirmed by the Fos response that can be elicited in the DH), but, assuming that the hormone is indeed capable of crossing the blood-brain barrier (see below), also when it is given systemically.

Physiopathological relevance of type 1a GHSRs in the spinal cord
Ghrelin is only one of the several peptides that in the past years have been shown to participate to the control of appetite and food intake. Interestingly, many of these orexigenic peptides also have a more or less well-defined role in pain modulation (25). Experimental and clinical data indicate that ghrelin plays different roles in physiological and pathological status (1, 2, 17).

Under physiological conditions ghrelin, after being secreted from the stomach (5), binds to type 1a GHSRs in the arcuate nucleus of the hypothalamus, which is influenced by circulating hormones diffusing from the median eminence and area postrema, two brain areas characterized by a deficiency of the blood-brain barrier (26). Acting on arcuate neurons, ghrelin stimulates hunger and food intake. In parallel, the hormone promotes neuronal release of neuropeptide Y (10), which negatively modulates nociception (27).

Physiologically, a reduced pain perception in feeding has been interpreted as functional to the maintenance of an appropriate vigilance status, and can be understood as part of an adaptive constellation of changes in the physiology and behavior of animals that must find food by foraging in potentially dangerous environments (28).

Under pathological conditions, especially in chronic diseases leading to poor nutritional status, high levels of ghrelin are reached in the blood (29). Blood-brain barrier mechanisms regulating ghrelin accumulation by the brain appear to be influenced by pathophysiological events (30). In these circumstances ghrelin might be allowed to cross the blood-brain barrier and get access to central type 1a GHSRs in locations other than the arcuate nucleus. Brain type 1a GHSRs expressed in the hypothalamus, pons, and medulla oblongata (13), i.e. the centers for descending control of nociception (31), can be targeted and inhibit pain at supraspinal level, whereas activation of spinal cord type 1a GHSRs, as shown here, enhances inhibitory neurotransmission in the deep DH, where it blocks the response of peptidergic nociceptive afferents to painful stimuli.

Central inhibition of pain mechanisms by ghrelin, in parallel with a potent antiinflammatory action at the periphery (32), may represent an adaptive response to counteract abnormal pain signaling conditions related to nutritional status.


   Acknowledgments
 
We thank Professor E. Ghigo for his precious comments and suggestions.


   Footnotes
 
This work was supported by Progetto di Ricerca di Rilevante Interesse Nazionale (PRIN) 2006 Grant 2006070912, and grants from Compagnia di San Paolo e Fondazione Cassa di Risparmio di Torino (CRT), Torino, Italy.

Disclosure Statement: The authors have nothing to disclose.

First Published Online January 17, 2008

Abbreviations: ACSF, Artificial cerebrospinal fluid; DH, dorsal horn; GHSR, GH secretagogue receptor; LY, Lucifer yellow; mIPSC, miniature inhibitory postsynaptic current; sEPSC, spontaneous excitatory postsynaptic current; sIPSC, spontaneous inhibitory postsynaptic current.

Received August 22, 2007.

Accepted for publication January 9, 2008.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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L. S. Stone and D. C. Molliver
In Search of Analgesia: Emerging Poles of GPCRs in Pain
Mol. Interv., October 1, 2009; 9(5): 234 - 251.
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