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Abstract
We tested the hypothesis that cannabinoids modulate feeding in male guinea pigs, and correlated cannabinoid-induced changes in feeding behavior with alterations in glutamatergic synaptic currents impinging upon proopiomelanocortin (POMC) neurons of the hypothalamic arcuate nucleus. Feeding experiments were performed as follows: after a three-day acclimation period, animals were weighed and injected with either the CB1 receptor agonist WIN 55,212-2 (1 mg/kg, s.c.), antagonist AM251 (3 mg/kg, s.c.) or their cremophore/ethanol/saline vehicle (1
18; 1 ml/kg, s.c.) each day for seven days. WIN 55,212-2 increased, whereas AM251 decreased, the rate of cumulative food intake. The agonist effect was manifest primarily by increases in meal frequency and the amount of food eaten per meal. By contrast, the antagonist effect was associated with decreases in meal frequency, duration and weight loss. For the electrophysiological experiments, we performed whole-cell patch clamp recordings from POMC neurons in hypothalamic slices. WIN 55,212-2 decreased the amplitude of evoked, glutamatergic excitatory postsynaptic currents (eEPSCs) and increased the S2:S1 ratio. Conversely, AM251 increased eEPSC amplitude per se, and blocked the inhibitory effects of the agonist. WIN 55,212-2 also decreased miniature EPSC (mEPSC) frequency; whereas AM251 increased mEPSC frequency per se, and again blocked the inhibitory effect of the agonist. A subpopulation of cells exhibited an agonist-induced outward current, which was blocked by AM251, associated with increased conductance and reversed polarity near the Nernst equilibrium potential for K+. These data demonstrate that cannabinoids regulate appetite in the guinea pig in part through both presynaptic and postsynaptic actions on anorexigenic POMC neurons.
Introduction
There is a growing appreciation of the role cannabinoids play in regulating homeostatic behaviors governed by the hypothalamus including, but not limited to, reproduction and feeding. For example, cannabinoids increase appetite in humans and various rodent animal models [1—5], and it is thought that the hyperphagia induced by cannabinomimetics can be of clinical utility in combating the body wasting observed with cancer chemotherapy and AIDS [2,6,7]. On the other hand, cannabinoid CB1 receptor antagonists are being used as a pharmacological intervention in the treatment of obesity [4,8,9] and the metabolic syndrome [10].
CB1 receptors are located in the myenteric plexus of the enteric nervous system [11], where they play an important role in regulating gastric emptying and intestinal motility [12]. CB1 receptors also are expressed in vagal afferents emanating from the gastrointestinal (GI) tract [13], and activation of these receptors inhibits glutamatergic synaptic currents in the dorsal motor nucleus of the vagus [14]. While it is clear that cannabinoids can influence appetite at the level of both the GI tract and brainstem, there is also considerable evidence to suggest that cannabinoids stimulate feeding by also acting within the hypothalamus. The hypothalamic feeding circuitry comprises orexigenic components with orexin- and melanin-concentrating hormone-containing somata in the lateral hypothalamic area [15—18], and neuropeptide Y (NPY)- and ghrelin-containing neurons in the arcuate nucleus [19—23]. The neuroanatomical substrates of the anorexigenic component include the ventromedial nucleus [24], proopiomelanocortin (POMC) neurons in the arcuate nucleus [25,26], cocaine-amphetamine-regulated transcript (CART) neurons in the arcuate, dorsomedial, lateral hypothalamic and paraventricular nuclei [27], and corticotropin-releasing hormone (CRH) neurons in the paraventricular nucleus [28]. This latter population appears to be a point of convergence for both orexigenic and anorexigenic inputs that integrates incoming information into an efferent response [18,22,28—30]. CB1 receptors are expressed in several nuclei of the hypothalamic feeding circuitry, where they are found on nerve terminals impinging upon MCH, CRH and POMC neurons [31]. In addition, hypothalamic levels of endogenous cannabinoids such as 2-arachidonyl glycerol (2-AG) are decreased by the adipostat leptin [32], which also depolarizes and increases the firing rate of POMC neurons [25,33]. Furthermore, food intake is decreased in CB1 receptor knockout mice [27,32], whereas transgenic mice deficient in the byproduct of posttranslational POMC processing, β-endorphin, exhibit rampant hyperphagia [34]. Thus, when considering the possible substrates upon which cannabinoids act to increase appetite, one must also include POMC neurons of the hypothalamic arcuate nucleus.
The purpose of the present study was test the hypothesis that cannabinoids stimulate feeding behavior in part by decreasing the excitability of POMC neurons. To this end, we employed a guinea pig animal model used in a number of different feeding, gastroenterological and metabolic studies [35—37]. We undertook a multidisciplinary approach to evaluate cannabinoid-induced changes in feeding behavior, as well as alterations in both glutamatergic synaptic input and postsynaptic K+ conductances occurring within POMC neurons. We utilized several indices of feeding behavior, including the rate of food intake, meal frequency and duration, and the amount of food eaten per meal. Intracellular patch-clamp recordings from hypothalamic arcuate neurons were performed using an in vitro hypothalamic slice preparation with biocytin-filled electrodes that enabled subsequent phenotypic identification of the cells. The results reveal that the CB1 receptor agonist (R)-(+)-[2,3-dihydro-5-methyl-3-(4-morpholinylmethyl)pyrrolo[1,2,3-de]-1,4-benzoxazin-6-yl]-1-naphthalenylmethanone mesylate (WIN 55,212-2) and antagonist N-(piperidin-1-yl)-5-(4-iodophenyl)-4-methyl-1H-pyrazole-3-carboxamide (AM251) increase and decrease feeding respectively. These behavioral changes correlate in a compelling fashion with changes in glutamatergic neurotransmission and postsynaptic K+ conductances in POMC neurons, suggesting that these neurons are indeed a critical cellular link to the orexigenic response elicited by cannabinoids.
Materials and Methods
Animals and Treatments
All animal procedures described in this study are in accordance with institutional guidelines based on NIH standards. Male Topeka guinea pigs (250—410 g) were obtained from Elm Hill Breeding Labs (Chelmsford, MA, USA), kept under controlled temperature (69—73°F) and light (12 h on: 12 h off), and provided with food and water ad libitum.
Drugs
Unless otherwise specified, all drugs were purchased from Tocris Cookson, Inc. (Ellisville, MO, USA.) For the behavioral experiments, both WIN 55,212-2 and AM251 were dissolved in a mixture of cremephor/ethanol/0.9% saline (118; v/v/v). For the electrophysiology experiments, tetrodotoxin (TTX) with citrate (Alomone Labs, Jerusalem, Israel) was dissolved in UltraPure H2O to a stock concentration of 1 mM. Arachidonyl-2′-chloroethylamide (ACEA) was dissolved in ethanol to a stock concentration of 1 mM. cis-4-[Phosphomethyl]-2-piperidinecarboxylic acid (CGS 19755; 10 mM) was dissolved in 0.1N NaOH and then diluted to the final volume with UltraPure H20. WIN-55,212-2, AM251 and 2,3-dioxo-6-nitro-1,2,3,4-tetrahydrobenzo[f]quinoxaline-7-sulfonamide (NBQX) were dissolved in dimethyl sulfoxide to stock concentrations of 10 mM. Stock solutions of 6-imino-3-(4-methoxyphenyl)-1(6H)-pyridazinebutanoic acid hydrobromide (SR 95531; 10 mM) and tertiapin (1 μM; Alomone Labs) were prepared with UltraPure H2O. Baclofen (40 mM) was dissolved in 0.1 N HCl.
Feeding behavior paradigm
The behavioral experiments were conducted using a four-cage, Comprehensive Lab Animal Monitoring System (Columbus Instruments, Columbus, OH, USA) from which we monitored four parameters of feeding: food intake, meal frequency, meal duration and the amount of food eaten per meal. An electronic balance measured the amount of food (certified guinea pig diet 5026, powdered formulation; Newco Distributors, Inc., Rancho Cucamonga, CA, USA), contained within the food dish and separate spillage container that sit atop it, with the difference being the actual amount of food consumed. Two photoelectric sensors are arrayed above the food dish, which detect each time the animal moves in for a meal. The signals generated are transmitted to an A/D converter, and digitally stored on computer. Food intake is defined as the amount of food consumed (in g) normalized to the total amount eaten over the designated six-hour observation period ranging from 08:30 — 14:30. Meal frequency equals the number of meals consumed per unit time, with a meal defined as the amount of food (in g) consumed in one sitting. Meal duration refers to the amount of time necessary to eat a meal consisting of at least 10 mg of food. The amount of food eaten per meal is defined as the amount of food eaten in a given hour relative to the total amount consumed over the six-hour evaluation period divided by the number of meals in that same hour. The animals were allowed to acclimate in the feeding chambers for six hours per day over a three-day period. Each morning they were weighed, handled, placed in their respective chambers for six hours, and then returned to their home cages. After the three-day acclimation session, we initiated the seven-day monitoring phase during which the animals were weighed, injected each morning with either WIN 55,212-2 (1 mg/kg; s.c.), AM251 (3 mg/kg; s.c.) or its cremephor/ethanol/0.9% saline vehicle (1 ml/kg; s.c.), immediately placed in their feeding chambers and then subject to continuous monitoring of each of the four above-described feeding parameters over the six-hour time span.
Tissue preparation
On the day of experimentation the animal was decapitated, its brain removed from the skull and the hypothalamus dissected. We then mounted the resultant hypothalamic block on a cutting platform that was then secured in a vibratome well filled with ice-cold, oxygenated (95% O2, 5% CO2) artificial cerebrospinal fluid (aCSF, in mM: NaCl, 124; NaHCO3 26; dextrose, 10; HEPES, 10; KCl, 5; NaH2PO4, 2.6; MgSO4, 2; CaCl2, 1). Five coronal slices (350 μm) through the arcuate nucleus were then cut. The slices were transferred to an auxiliary chamber containing oxygenated aCSF, and kept there until electrophysiological recording.
Electrophysiology
Whole-cell patch recordings in hypothalamic slices were performed as previously described [38]. Briefly, during the recording session, slices were maintained in a chamber perfused with a warmed (35°C), oxygenated aCSF that was identical to that described above, the exception being an increase in the CaCl2 concentration to 2 mM. Artificial CSF and all drugs (diluted with aCSF) were perfused via a peristaltic pump at a rate of 1.5 ml/min. Patch electrodes were assembled from borosilicate glass (World Precision Instruments; Sarasota, FL, USA; 1.5 mm O.D.) pulled on a P-97 Flaming Brown puller (Sutter Instrument Co., Novato, CA, USA), and filled with the following (in mM): potassium gluconate, 128; NaCl, 10; MgCl2, 1; EGTA, 11; HEPES, 10; ATP, 1.2; GTP, 0.4; 0.35 — 0.4% biocytin; adjusted to a pH of 7.3 — 7.4 with 1.0 N KOH; 297 — 321 mOsm. Electrode resistances varied from 3 — 8 M Ω A Multiclamp 700A preamplifier (Axon Instruments, Foster City, CA, USA) amplified potentials and passed current through the electrode. The patch electrode was advanced into the arcuate nucleus with a positive pressure applied to the back of the electrode using a 5-ml syringe connected by polyethylene tubing to the electrode holder. Once the electrode came up against a cell membrane, as indicated by a small decrease in the monitored current deflection (20—50% of the initial amplitude), a slight negative pressure was then applied via suction to form a stable gigaohm seal. We obtained whole cell recordings by applying additional suction to rupture the membrane under the electrode. Membrane currents were recorded in voltage clamp and underwent analog-digital conversion via a Digidata 1322A interface coupled to pClamp 8.2 software (Axon Instruments). The access resistance, as well as the resting membrane potential (RMP) and the input resistance (Rin), were monitored throughout the course of the recording. If the access resistance deviated greater than 10% of its original value, the recording was ended. Low-pass filtering of the currents was conducted at a frequency of two KHz. The liquid junction potential was calculated to be −10 mV, and is corrected for during data analysis using pClamp software.
In the studies designed to evaluate eEPSCs in arcuate neurons, concentric bipolar tungsten stimulating electrodes (World Precision Instruments) were placed 0.5 — 1.0 mm lateral to the recording electrode to deliver square-wave pulses 500 μsec in duration (0.1 Hz; 3—18 V) generated by a model S88K stimulator (Astro-Med, Inc., West Warwick, RI, USA) and converted to constant voltage by a model SIU5 stimulus isolation unit (Astro-Med). We recorded eEPSCs from a holding potential of −75 mV in the presence of SR 95531 (10 μM) to block γ-aminobutyric acid (GABA)A receptor-mediated synaptic currents. Peak eEPSC amplitudes were measured by averaging those obtained from 20 consecutive stimulations. Rare failures of the stimulus to evoke PSCs were excluded from the analyses. The time-to-peak reflects the time necessary for the inward current to descend to its maximum amplitude. The rate of eEPSC decay was calculated as the time necessary for the current to decline to one-third of its peak value. Cells were perfused with WIN 55,212-2 (1 - 3 μM) for four min and the stimulation paradigm is repeated in the presence of the agonist. To help confirm a presynaptic site of cannabinoid action, we employed the paired-pulse paradigm in which we presented successive stimuli that were separated by 75 msec from one another. CB1 receptor specificity was ascertained by evaluating CB1 receptor activation in slices pretreated with AM 251 (1 μM). We verified that the eEPSCs were ionotropic glutamate receptor-mediated by repeating the stimulation in the presence of CGS 19755 (10 μM) and NBQX (3 μM).
We recorded mEPSCs in the presence of both SR 95531 (10 μM) and TTX (500 nM) from a holding potential of −75 mV. After collecting a 3—4 min segment of baseline data, we perfused either WIN 55,212-2 (100 nM — 10 μM) or ACEA (1 μM) for four min, and then recorded mEPSCs in the presence of agonist. The threshold for mEPSC detection was set at least three pA below the baseline holding current as assessed from the headstage output, and continuously monitored throughout each 3—4 min recording period. Information on interval and amplitude was obtained from at least 100 contiguous mEPSCs, which we used to evaluate cannabinoid-induced alterations in mEPSC frequency and amplitude as assessed from cumulative probability plots. CB1 receptor specificity was again ascertained by evaluating CB1 receptor activation in slices pretreated with AM 251 (1 μM), and the mEPSCs were pharmacologically identified using CGS 19755 (10 μM) and NBQX (3 μM).
To characterize the CB1 receptor-mediated activation of GIRK, either WIN 55,212-2 or ACEA were perfused until a new steady-state holding current was established (3—5 min). Current-voltage relationships were generated before and immediately following agonist application over a range centering on the equilibrium potential for K+. This is accomplished with step command potentials ranging from −50 to −130 mV (1 sec duration; 10 mV increments). Blockade of the agonist-induced outward current by AM 251 served as the indication that the effect was CB1 receptor-mediated. For comparison, the ability of the GABAB receptor agonist baclofen (100 μM) to elicit its characteristic outward current was ascertained prior to, and in the presence of, the GIRK channel blocker tertiapin (10 nM).
Immunohistofluorescence
After recording, some slices were processed for phenotypic identification as described previously [39]. Briefly, slices were fixed with either 4% paraformaldehyde (PFM) in Sorensen's phosphate buffer (pH 7.4) for 90—180 min. They then were immersed overnight in 20% sucrose dissolved in Sorensen's buffer, and frozen in Tissue-Tek embedding medium (Miles, Inc., Elkhart, IN, USA) the next day. Coronal sections (20 μm) were cut on a cryostat and mounted on charged slides. These sections were washed with 0.1 M sodium phosphate buffer (pH 7.4). All sections were then processed with streptavidin-cy2 (Amersham Biosciences, Piscataway, NJ, USA) at a 1:300 dilution. After localizing the biocytin-filled neuron, the slides containing the appropriate sections were reacted with either a β-endorphin antiserum [40] or an antibody directed against α-melanocyte-stimulating hormone (α-MSH; Immunostar, Inc., Hudson, WI, USA) at a 1:1000 or 1:200 dilution, respectively, using fluorescence immunohistochemistry [39].
Statistics
Variance homogeneity was evaluated using Levene's test. Comparisons between groups were performed using either the Student's t-test or the Mann-Whitney U-test. Comparisons between more than two groups were accomplished via the analysis of variance (ANOVA) followed by either the Least Significant Difference (LSD) test or median-notched box-and-whisker analysis, or the Kruskal-Wallis test followed by either the Mann-Whitney U-test or median-notched box-and-whisker analysis. Differences were considered statistically significant if the probability of error was less than 5%.
Results
Figure 1 depicts food intake as a function of time on four representative days (days 2—5) over the course of the seven-day monitoring period. Multifactorial ANOVA revealed that, compared to vehicle-treated controls, systemic administration of either the CB1 receptor agonist WIN 55,212-2 (1 mg/kg; s.c.) or antagonist AM251 (3 mg/kg; s.c.) resulted in a significant main effect (day 2: F-ratio = 81.07, p<0.0001; day 3: F-ratio = 49.54, p<0.0001; day 4: F-ratio = 61.43; p<0.0001; day 5: F-ratio = 152.2; p<0.0001) on the rate of food consumption. WIN 55,212-2 increased the rate of food intake, whereas AM251 decreased it. These effects became apparent 1—2 hour following their administration just prior to beginning the evaluation period, and lasted up to 4—6 hours into the monitoring. The agonist-induced increase in the cumulative rate of food intake shown for day 5 (like all other days) was completely abolished by co-administration of antagonist. There was no change in the total amount of food eaten over the course of the six-hour evaluation period (not shown). We also examined cannabinoid effects on meal frequency, meal duration and the amount of food eaten per meal. Figure 2 shows the effect of WIN 55,212-2 and AM251 on meal frequency ascertained every hour during the evaluation period. Multifactorial ANOVA again revealed significant main drug effect (F-ratio = 13.41, p<0.0001), as well as a significant interaction (F-ratio = 4.77, p<0.0001). WIN 55,212-2 increased meal frequency during the second hour, followed by a pronounced decrease during hours 3—6. AM251 completely blocked the agonist-induced increase meal frequency, and attenuated the prolonged decrease in meal frequency observed during the mid- to latter stages of the monitoring period. On the other hand, AM251 produced a decrease in meal frequency during hour 1, which was reversed by co-administration of the agonist. The increase in the rate of food consumption and meal frequency produced by WIN 55,212-2 was associated with a modest increase in meal duration (Figure 3) during hour 1 and a dramatic increase in the amount of food eaten per meal (Figure 4). These effects were eliminated by co-administration of the antagonist. WIN 55,212-2 was without effect on the rate of weight gain (Figure 5). Conversely, AM251 markedly decreased meal duration (Figure 3) and elicited a significant reduction in weight gain (Figure 5), and these effects were at least partially reversed by co-administration of the agonist. AM251 did not affect the amount of food eaten per meal, per se (Figure 4). Collectively, this indicates that CB1 receptor activation by both exogenous and endogenous cannabinoid agonists increases feeding behavior, whereas CB1 receptor antagonism suppresses it.
We then wanted to determine if the increased appetite produced by activation of CB1 receptors was associated with a decrease in glutamatergic synaptic input onto anorexigenic POMC neurons. Accordingly, we performed whole-cell patch-clamp recordings from arcuate neurons in hypothalamic slices. Figure 6 shows traces of excitatory postsynaptic currents (EPSCs) evoked by square-wave pulses delivered by a bipolar tungsten stimulating electrodes positioned 0.5—1.0 μm lateral to the recording electrode. Bath application of WIN 55,212-2 (1 μM) reduced the amplitude of evoked EPSCs (eEPSCs) that were subsequently abolished in the presence of the ionotropic glutamate receptor antagonists NBQX (3 μM) and CGS 19755 (10 μM; Figures 6A and 6C). On the other hand, AM251 (1 μM) per se increased eEPSC amplitude, and when co-administered with WIN 55,212-2 it blocked the inhibitory effect of the agonist (Figures 6B and 6C). Neither WIN 55,212-2 nor AM251 had any effect on the rate of rise or the decay of the eEPSCs (not shown). Further analysis using the paired-pulse paradigm revealed that the agonist-induced reduction in eEPSC amplitude was associated with an increase in the S2:S1 ratio (Figures 6D—6F). To verify that the agonist-induced reduction could be attributed to a presynaptic mechanism of action, we next performed an analysis of miniature EPSCs (mEPSCs) recorded in the presence of 500 nM TTX. Figures 7 and and88 show representative traces of mEPSCs recorded from arcuate neurons. Varying doses of WIN 55,212-2 (100 nM & 10 μM) produced graded increases in the interval between contiguous mEPSCs, which effectively translates into a decrease in mEPSC frequency, with no discernable effect on mEPSC amplitude (Figure 7A). These observations are substantiated further by the cumulative probability plots shown in Figure 7B. WIN 55,212-2 (100 nM) similarly, yet slightly less effectively, reduced the frequency of miniature, inhibitory postsynaptic currents (mIPSCs) recorded in the presence of TTX, NBQX and CGS 19755 from a holding potential of −30 mV (mIPSCs: 68.3 ± 7.6% of control (n = 3) vs. mEPSCs: 52.6 ± 6.5% of control (n = 5)). In addition, the selective effect of WIN 55,212-2 on mEPSC frequency was mimicked by the anandamide analog ACEA (not shown). On the other hand, AM251 both increased mEPSC frequency per se and blocked the diminution caused by WIN 55,212-2 (Figure 8A). The composite effects of WIN 55,212-2 and AM251 on mEPSC frequency and amplitude are summarized in Figure 8B. These data suggest that CB1 receptor activation by both exogenous and endogenous cannabinoid agonists presynaptically inhibits glutamatergic synaptic input onto arcuate neurons.
We noticed during the mEPSC experiments that approximately one-third of arcuate neurons (10/31) responded to both WIN 55,212-2 and ACEA with a change in the holding current, and thus explored this observation in greater detail. Figures 9A1 and 9B1 show representative examples of the outward current elicited by bath application of 1 μM ACEA. Current-voltage relationships generated prior to, and in the presence of, the agonist reveal that the current reverses polarity very near the Nernst equilibrium potential for K+, and exhibits an increase in slope conductance that grows more pronounced as the membrane potential is hyperpolarized below the reversal potential (Figure 9A2). This agonist-induced outward current was blocked by AM251 (Figure 9B2), and was found to be both qualitatively (Figure 9C) and quantitatively (Figure 9D) very similar to the G-protein-gated, inwardly-rectifying K+ current (GIRK) elicited by activation of metabotropic GABAB receptors. The activation of this postsynaptic K+ current, as well as the presynaptic inhibition of glutamatergic synaptic input described above, was observed in POMC neurons (n = 6) subsequently identified through immunohistofluorescent visualization (Figure 10). Taken together, these data demonstrate that cannabinoids increase feeding behavior, which is associated with a presynaptic inhibition of glutamatergic neurotransmission and a postsynaptic activation of an inwardly-rectifying K+ current within POMC neurons. This indicates that the cannabinoid modulation of appetite is mediated, at least in part, through interactions with the hypothalamic feeding circuitry.
Discussion
The results of the present study demonstrate unequivocally that cannabinoids stimulate appetite, which correlates with a decrease in the excitability of POMC neurons. These conclusions are based on the following observations: 1) CB1 receptor activation and blockade respectively increases and decreases feeding behavior as determined using four different indices: the rate of food intake, meal frequency, meal duration, the amount of food eaten per meal and change in body weight, 2) CB1 receptor activation and blockade respectively decreases and increases glutamatergic neurotransmission via a presynaptic site of action, 3) cannabinoids affect a CB1 receptor-mediated activation of an inwardly rectifying K+ current and 4) the latter two cellular effects of cannabinoids occur in arcuate neurons subsequently identified as POMC neurons via immunohistofluorescence. The current findings also suggest that glutamatergic synaptic input onto POMC neurons is tonically inhibited by endogenous cannabinoids, which provides further evidence that these neurons are a critical target in the cannabinoid modulation of appetite.
While WIN 55,212-2 has affinity for both CB1 and CB2 receptor subtypes [41], and CB2 receptors are found in the brainstem [42], the fact that the CB1 receptor antagonist AM251 largely abolished the agonist-induced hyperphagia, decrease in eEPSC amplitude, mEPSC frequency and the activation of the inwardly-rectifying K+ conductance clearly indicates that the effects observed in the present study are CB1 receptor-mediated. In the present study, CB1 receptor activation upon the exogenous administration of WIN 55,212-2 increased the rate of food intake, meal frequency, meal duration and the amount of food eaten per meal. Thus, exogenously administered cannabinoids increased eating efficiency. On the other hand, CB1 receptor antagonism decreased the rate of food intake, meal frequency and meal duration, indicating that endogenous cannabinoids drive appetite and food seeking behavior. The cannabinoid-induced hyperphagia that we observed in our guinea pig animal model is consistent with that reported in humans and other rodent models. Despite a limited number of controlled scientific studies, the ability of cannabinoids, in particular Δ 9-tetrahydrocannabinol (Δ 9-THC), to cause the sensation of hunger and increase food intake in humans has been recognized for centuries [2—5]. In rats and mice, exogenously administered cannabinoid agonists stimulate appetite under ad libitum and food-deprived conditions [1,2,43], whereas CB1 receptor antagonists suppress it [8,9]. In addition, Δ 9-THC promotes feeding in rats when focally injected into the ventromedial or lateral hypothalamus [2,4], and the satiety hormone leptin decreases hypothalamic levels of endogenous 2-AG [32]. Moreover, both ob/ob mice (which do not produce leptin) and fa/fa rats (which do not express leptin receptors) exhibit rampant hyperphagia and weight gain [32,44,45], as well as elevated hypothalamic levels of endogenous cannabinoids [32], with respect to their wildtype counterparts. Thus, the cannabinoid regulation of appetite is clearly conserved among mammalian species, and involves, at least in part, interactions with various neuroanatomical substrates within the hypothalamus.
The cannabinoid-induced presynaptic inhibition of glutamatergic synaptic input onto guinea pig arcuate POMC neurons is also consistent with that reported in several other regions of the central nervous system, including the prefrontal cortex [46], hippocampus [47,48], PAG [49] and the dorsal motor nucleus of the vagus [14]. As mentioned above, we also found that CB1 receptor antagonism unmasked a tonic inhibition of glutamate release onto POMC neurons, suggesting that endogenous cannabinoids are a principal neuromodulator influencing synaptic input onto these cells. In the hippocampus [50], cerebellum [51], ventral tegmental area [52] and other hypothalamic nuclei [53], endogenous cannabinoids released from postsynaptic neurons in response to depolarizing stimuli retrogradely inhibit glutamate release from nerve terminals impinging upon them. Indeed, the retrograde inhibition of glutamatergic synaptic input released by endogenous cannabinoids released from parvocellular neurons in the paraventricular nucleus of the hypothalamus is instrumental in mediating the rapid feedbacks of corticosteroids on the stress axis [53]. In transgenic mice whose POMC neurons are labeled with enhanced green fluorescent protein, however, endogenous cannabinoids retrogradely inhibit GABAergic but not glutamatergic synaptic input onto these cells [54]. This suggests that amino acid neurotransmission within POMC neurons is differentially regulated in the two species. Future studies will determine if endogenous cannabinoids synthesized de novo in guinea pig POMC neurons serve to tonically inhibit glutamate neurotransmission in a retrograde fashion. Our finding of a CB1 receptor-mediated activation of an inwardly-rectifying K+ current in POMC neurons is also in agreement with that observed in AtT20 cells transfected with the CB1 receptor [55], and in oocytes co-expressing the CB1 receptor and GIRK1 channel subtype [56,57]. This current cannot be attributed to other voltage-dependent K+ currents such as the A-type K+ current [38], and is virtually identical to that elicited by the activation of GABAB receptors, which have long been known to couple to GIRK in the arcuate nucleus, preoptic area, supraoptic nucleus and many other regions of the central nervous system [58—61].
Along with the ventromedial nucleus of the hypothalamus, the POMC neurons in the arcuate nucleus are the principal anorexigenic component of the hypothalamic feeding circuitry. These neurons release posttranslational byproducts of POMC such as α-melanocyte-stimulating hormone (α-MSH) β-endorphin, as well as CART that is co-expressed in these cells [34,45]. Indeed, β-endorphin and melanocortin (MC)4 receptor knockout mice display extensive hyperphagia [34] and are resistant to disease-induced cachexia [26], respectively. In addition, the anorexigenic effect of the CB1 receptor antagonist rimonabant is eliminated in CART-deficient transgenic mice [62]. POMC neurons are a key target of the satiety hormone leptin, which not only depolarizes and increases the firing rate of these cells, but also decreases hypothalamic levels of endogenous cannabinoids [25,32,33]. Glutamate increases β-endorphin secretion and POMC gene expression, and glutamatergic synaptic input is observed in POMC neurons in the arcuate nucleus and the nucleus of the tractus solitarius of transgenic mice [54,63,64]. While our findings clearly indicate that POMC neurons are a major hypothalamic locus for the hyperphagic actions of cannabinoids, there are in reality a multiplicity of hypothalamic sites and mechanisms involved in the induction of feeding behavior. For example, cannabinoids increase orexigenic NPY release from hypothalamic explants [65]. On the other hand, NPY neurons do not express CB1 receptors, and so the cannabinoid-induced increase in NPY release is due presumably to a decrease in GABA input onto these cells [31], much like the retrograde inhibition of GABA release onto orexigenic MCH neurons in the lateral hypothalamus [66]. Moreover, cannabinoid interactions with the brain stem, gut, pancreas and adipose tissue also play a role in the regulation of appetite and energy homeostasis [31]. Nevertheless, the presynaptic inhibition of glutamatergic neurotransmission within guinea pig POMC neurons must account, at least in part, for the cannabinoid-induced hyperphagia observed in the present study.
In conclusion, these results reveal that both endogenous and exogenous cannabinoids stimulate feeding behavior via a mechanism that involves, at least in part, a presynaptic inhibition of glutamate neurotransmission in anorexigenic POMC neurons. Our findings convey important insight into how cannabinoids increase appetite through their interactions with the hypothalamic feeding circuitry.
Source, Graphs and Figures: CANNABINOID-INDUCED HYPERPHAGIA: CORRELATION WITH INHIBITION OF PROOPIOMELANOCORTIN NEURONS?
We tested the hypothesis that cannabinoids modulate feeding in male guinea pigs, and correlated cannabinoid-induced changes in feeding behavior with alterations in glutamatergic synaptic currents impinging upon proopiomelanocortin (POMC) neurons of the hypothalamic arcuate nucleus. Feeding experiments were performed as follows: after a three-day acclimation period, animals were weighed and injected with either the CB1 receptor agonist WIN 55,212-2 (1 mg/kg, s.c.), antagonist AM251 (3 mg/kg, s.c.) or their cremophore/ethanol/saline vehicle (1
18; 1 ml/kg, s.c.) each day for seven days. WIN 55,212-2 increased, whereas AM251 decreased, the rate of cumulative food intake. The agonist effect was manifest primarily by increases in meal frequency and the amount of food eaten per meal. By contrast, the antagonist effect was associated with decreases in meal frequency, duration and weight loss. For the electrophysiological experiments, we performed whole-cell patch clamp recordings from POMC neurons in hypothalamic slices. WIN 55,212-2 decreased the amplitude of evoked, glutamatergic excitatory postsynaptic currents (eEPSCs) and increased the S2:S1 ratio. Conversely, AM251 increased eEPSC amplitude per se, and blocked the inhibitory effects of the agonist. WIN 55,212-2 also decreased miniature EPSC (mEPSC) frequency; whereas AM251 increased mEPSC frequency per se, and again blocked the inhibitory effect of the agonist. A subpopulation of cells exhibited an agonist-induced outward current, which was blocked by AM251, associated with increased conductance and reversed polarity near the Nernst equilibrium potential for K+. These data demonstrate that cannabinoids regulate appetite in the guinea pig in part through both presynaptic and postsynaptic actions on anorexigenic POMC neurons.Introduction
There is a growing appreciation of the role cannabinoids play in regulating homeostatic behaviors governed by the hypothalamus including, but not limited to, reproduction and feeding. For example, cannabinoids increase appetite in humans and various rodent animal models [1—5], and it is thought that the hyperphagia induced by cannabinomimetics can be of clinical utility in combating the body wasting observed with cancer chemotherapy and AIDS [2,6,7]. On the other hand, cannabinoid CB1 receptor antagonists are being used as a pharmacological intervention in the treatment of obesity [4,8,9] and the metabolic syndrome [10].
CB1 receptors are located in the myenteric plexus of the enteric nervous system [11], where they play an important role in regulating gastric emptying and intestinal motility [12]. CB1 receptors also are expressed in vagal afferents emanating from the gastrointestinal (GI) tract [13], and activation of these receptors inhibits glutamatergic synaptic currents in the dorsal motor nucleus of the vagus [14]. While it is clear that cannabinoids can influence appetite at the level of both the GI tract and brainstem, there is also considerable evidence to suggest that cannabinoids stimulate feeding by also acting within the hypothalamus. The hypothalamic feeding circuitry comprises orexigenic components with orexin- and melanin-concentrating hormone-containing somata in the lateral hypothalamic area [15—18], and neuropeptide Y (NPY)- and ghrelin-containing neurons in the arcuate nucleus [19—23]. The neuroanatomical substrates of the anorexigenic component include the ventromedial nucleus [24], proopiomelanocortin (POMC) neurons in the arcuate nucleus [25,26], cocaine-amphetamine-regulated transcript (CART) neurons in the arcuate, dorsomedial, lateral hypothalamic and paraventricular nuclei [27], and corticotropin-releasing hormone (CRH) neurons in the paraventricular nucleus [28]. This latter population appears to be a point of convergence for both orexigenic and anorexigenic inputs that integrates incoming information into an efferent response [18,22,28—30]. CB1 receptors are expressed in several nuclei of the hypothalamic feeding circuitry, where they are found on nerve terminals impinging upon MCH, CRH and POMC neurons [31]. In addition, hypothalamic levels of endogenous cannabinoids such as 2-arachidonyl glycerol (2-AG) are decreased by the adipostat leptin [32], which also depolarizes and increases the firing rate of POMC neurons [25,33]. Furthermore, food intake is decreased in CB1 receptor knockout mice [27,32], whereas transgenic mice deficient in the byproduct of posttranslational POMC processing, β-endorphin, exhibit rampant hyperphagia [34]. Thus, when considering the possible substrates upon which cannabinoids act to increase appetite, one must also include POMC neurons of the hypothalamic arcuate nucleus.
The purpose of the present study was test the hypothesis that cannabinoids stimulate feeding behavior in part by decreasing the excitability of POMC neurons. To this end, we employed a guinea pig animal model used in a number of different feeding, gastroenterological and metabolic studies [35—37]. We undertook a multidisciplinary approach to evaluate cannabinoid-induced changes in feeding behavior, as well as alterations in both glutamatergic synaptic input and postsynaptic K+ conductances occurring within POMC neurons. We utilized several indices of feeding behavior, including the rate of food intake, meal frequency and duration, and the amount of food eaten per meal. Intracellular patch-clamp recordings from hypothalamic arcuate neurons were performed using an in vitro hypothalamic slice preparation with biocytin-filled electrodes that enabled subsequent phenotypic identification of the cells. The results reveal that the CB1 receptor agonist (R)-(+)-[2,3-dihydro-5-methyl-3-(4-morpholinylmethyl)pyrrolo[1,2,3-de]-1,4-benzoxazin-6-yl]-1-naphthalenylmethanone mesylate (WIN 55,212-2) and antagonist N-(piperidin-1-yl)-5-(4-iodophenyl)-4-methyl-1H-pyrazole-3-carboxamide (AM251) increase and decrease feeding respectively. These behavioral changes correlate in a compelling fashion with changes in glutamatergic neurotransmission and postsynaptic K+ conductances in POMC neurons, suggesting that these neurons are indeed a critical cellular link to the orexigenic response elicited by cannabinoids.
Materials and Methods
Animals and Treatments
All animal procedures described in this study are in accordance with institutional guidelines based on NIH standards. Male Topeka guinea pigs (250—410 g) were obtained from Elm Hill Breeding Labs (Chelmsford, MA, USA), kept under controlled temperature (69—73°F) and light (12 h on: 12 h off), and provided with food and water ad libitum.
Drugs
Unless otherwise specified, all drugs were purchased from Tocris Cookson, Inc. (Ellisville, MO, USA.) For the behavioral experiments, both WIN 55,212-2 and AM251 were dissolved in a mixture of cremephor/ethanol/0.9% saline (1
18; v/v/v). For the electrophysiology experiments, tetrodotoxin (TTX) with citrate (Alomone Labs, Jerusalem, Israel) was dissolved in UltraPure H2O to a stock concentration of 1 mM. Arachidonyl-2′-chloroethylamide (ACEA) was dissolved in ethanol to a stock concentration of 1 mM. cis-4-[Phosphomethyl]-2-piperidinecarboxylic acid (CGS 19755; 10 mM) was dissolved in 0.1N NaOH and then diluted to the final volume with UltraPure H20. WIN-55,212-2, AM251 and 2,3-dioxo-6-nitro-1,2,3,4-tetrahydrobenzo[f]quinoxaline-7-sulfonamide (NBQX) were dissolved in dimethyl sulfoxide to stock concentrations of 10 mM. Stock solutions of 6-imino-3-(4-methoxyphenyl)-1(6H)-pyridazinebutanoic acid hydrobromide (SR 95531; 10 mM) and tertiapin (1 μM; Alomone Labs) were prepared with UltraPure H2O. Baclofen (40 mM) was dissolved in 0.1 N HCl.Feeding behavior paradigm
The behavioral experiments were conducted using a four-cage, Comprehensive Lab Animal Monitoring System (Columbus Instruments, Columbus, OH, USA) from which we monitored four parameters of feeding: food intake, meal frequency, meal duration and the amount of food eaten per meal. An electronic balance measured the amount of food (certified guinea pig diet 5026, powdered formulation; Newco Distributors, Inc., Rancho Cucamonga, CA, USA), contained within the food dish and separate spillage container that sit atop it, with the difference being the actual amount of food consumed. Two photoelectric sensors are arrayed above the food dish, which detect each time the animal moves in for a meal. The signals generated are transmitted to an A/D converter, and digitally stored on computer. Food intake is defined as the amount of food consumed (in g) normalized to the total amount eaten over the designated six-hour observation period ranging from 08:30 — 14:30. Meal frequency equals the number of meals consumed per unit time, with a meal defined as the amount of food (in g) consumed in one sitting. Meal duration refers to the amount of time necessary to eat a meal consisting of at least 10 mg of food. The amount of food eaten per meal is defined as the amount of food eaten in a given hour relative to the total amount consumed over the six-hour evaluation period divided by the number of meals in that same hour. The animals were allowed to acclimate in the feeding chambers for six hours per day over a three-day period. Each morning they were weighed, handled, placed in their respective chambers for six hours, and then returned to their home cages. After the three-day acclimation session, we initiated the seven-day monitoring phase during which the animals were weighed, injected each morning with either WIN 55,212-2 (1 mg/kg; s.c.), AM251 (3 mg/kg; s.c.) or its cremephor/ethanol/0.9% saline vehicle (1 ml/kg; s.c.), immediately placed in their feeding chambers and then subject to continuous monitoring of each of the four above-described feeding parameters over the six-hour time span.
Tissue preparation
On the day of experimentation the animal was decapitated, its brain removed from the skull and the hypothalamus dissected. We then mounted the resultant hypothalamic block on a cutting platform that was then secured in a vibratome well filled with ice-cold, oxygenated (95% O2, 5% CO2) artificial cerebrospinal fluid (aCSF, in mM: NaCl, 124; NaHCO3 26; dextrose, 10; HEPES, 10; KCl, 5; NaH2PO4, 2.6; MgSO4, 2; CaCl2, 1). Five coronal slices (350 μm) through the arcuate nucleus were then cut. The slices were transferred to an auxiliary chamber containing oxygenated aCSF, and kept there until electrophysiological recording.
Electrophysiology
Whole-cell patch recordings in hypothalamic slices were performed as previously described [38]. Briefly, during the recording session, slices were maintained in a chamber perfused with a warmed (35°C), oxygenated aCSF that was identical to that described above, the exception being an increase in the CaCl2 concentration to 2 mM. Artificial CSF and all drugs (diluted with aCSF) were perfused via a peristaltic pump at a rate of 1.5 ml/min. Patch electrodes were assembled from borosilicate glass (World Precision Instruments; Sarasota, FL, USA; 1.5 mm O.D.) pulled on a P-97 Flaming Brown puller (Sutter Instrument Co., Novato, CA, USA), and filled with the following (in mM): potassium gluconate, 128; NaCl, 10; MgCl2, 1; EGTA, 11; HEPES, 10; ATP, 1.2; GTP, 0.4; 0.35 — 0.4% biocytin; adjusted to a pH of 7.3 — 7.4 with 1.0 N KOH; 297 — 321 mOsm. Electrode resistances varied from 3 — 8 M Ω A Multiclamp 700A preamplifier (Axon Instruments, Foster City, CA, USA) amplified potentials and passed current through the electrode. The patch electrode was advanced into the arcuate nucleus with a positive pressure applied to the back of the electrode using a 5-ml syringe connected by polyethylene tubing to the electrode holder. Once the electrode came up against a cell membrane, as indicated by a small decrease in the monitored current deflection (20—50% of the initial amplitude), a slight negative pressure was then applied via suction to form a stable gigaohm seal. We obtained whole cell recordings by applying additional suction to rupture the membrane under the electrode. Membrane currents were recorded in voltage clamp and underwent analog-digital conversion via a Digidata 1322A interface coupled to pClamp 8.2 software (Axon Instruments). The access resistance, as well as the resting membrane potential (RMP) and the input resistance (Rin), were monitored throughout the course of the recording. If the access resistance deviated greater than 10% of its original value, the recording was ended. Low-pass filtering of the currents was conducted at a frequency of two KHz. The liquid junction potential was calculated to be −10 mV, and is corrected for during data analysis using pClamp software.
In the studies designed to evaluate eEPSCs in arcuate neurons, concentric bipolar tungsten stimulating electrodes (World Precision Instruments) were placed 0.5 — 1.0 mm lateral to the recording electrode to deliver square-wave pulses 500 μsec in duration (0.1 Hz; 3—18 V) generated by a model S88K stimulator (Astro-Med, Inc., West Warwick, RI, USA) and converted to constant voltage by a model SIU5 stimulus isolation unit (Astro-Med). We recorded eEPSCs from a holding potential of −75 mV in the presence of SR 95531 (10 μM) to block γ-aminobutyric acid (GABA)A receptor-mediated synaptic currents. Peak eEPSC amplitudes were measured by averaging those obtained from 20 consecutive stimulations. Rare failures of the stimulus to evoke PSCs were excluded from the analyses. The time-to-peak reflects the time necessary for the inward current to descend to its maximum amplitude. The rate of eEPSC decay was calculated as the time necessary for the current to decline to one-third of its peak value. Cells were perfused with WIN 55,212-2 (1 - 3 μM) for four min and the stimulation paradigm is repeated in the presence of the agonist. To help confirm a presynaptic site of cannabinoid action, we employed the paired-pulse paradigm in which we presented successive stimuli that were separated by 75 msec from one another. CB1 receptor specificity was ascertained by evaluating CB1 receptor activation in slices pretreated with AM 251 (1 μM). We verified that the eEPSCs were ionotropic glutamate receptor-mediated by repeating the stimulation in the presence of CGS 19755 (10 μM) and NBQX (3 μM).
We recorded mEPSCs in the presence of both SR 95531 (10 μM) and TTX (500 nM) from a holding potential of −75 mV. After collecting a 3—4 min segment of baseline data, we perfused either WIN 55,212-2 (100 nM — 10 μM) or ACEA (1 μM) for four min, and then recorded mEPSCs in the presence of agonist. The threshold for mEPSC detection was set at least three pA below the baseline holding current as assessed from the headstage output, and continuously monitored throughout each 3—4 min recording period. Information on interval and amplitude was obtained from at least 100 contiguous mEPSCs, which we used to evaluate cannabinoid-induced alterations in mEPSC frequency and amplitude as assessed from cumulative probability plots. CB1 receptor specificity was again ascertained by evaluating CB1 receptor activation in slices pretreated with AM 251 (1 μM), and the mEPSCs were pharmacologically identified using CGS 19755 (10 μM) and NBQX (3 μM).
To characterize the CB1 receptor-mediated activation of GIRK, either WIN 55,212-2 or ACEA were perfused until a new steady-state holding current was established (3—5 min). Current-voltage relationships were generated before and immediately following agonist application over a range centering on the equilibrium potential for K+. This is accomplished with step command potentials ranging from −50 to −130 mV (1 sec duration; 10 mV increments). Blockade of the agonist-induced outward current by AM 251 served as the indication that the effect was CB1 receptor-mediated. For comparison, the ability of the GABAB receptor agonist baclofen (100 μM) to elicit its characteristic outward current was ascertained prior to, and in the presence of, the GIRK channel blocker tertiapin (10 nM).
Immunohistofluorescence
After recording, some slices were processed for phenotypic identification as described previously [39]. Briefly, slices were fixed with either 4% paraformaldehyde (PFM) in Sorensen's phosphate buffer (pH 7.4) for 90—180 min. They then were immersed overnight in 20% sucrose dissolved in Sorensen's buffer, and frozen in Tissue-Tek embedding medium (Miles, Inc., Elkhart, IN, USA) the next day. Coronal sections (20 μm) were cut on a cryostat and mounted on charged slides. These sections were washed with 0.1 M sodium phosphate buffer (pH 7.4). All sections were then processed with streptavidin-cy2 (Amersham Biosciences, Piscataway, NJ, USA) at a 1:300 dilution. After localizing the biocytin-filled neuron, the slides containing the appropriate sections were reacted with either a β-endorphin antiserum [40] or an antibody directed against α-melanocyte-stimulating hormone (α-MSH; Immunostar, Inc., Hudson, WI, USA) at a 1:1000 or 1:200 dilution, respectively, using fluorescence immunohistochemistry [39].
Statistics
Variance homogeneity was evaluated using Levene's test. Comparisons between groups were performed using either the Student's t-test or the Mann-Whitney U-test. Comparisons between more than two groups were accomplished via the analysis of variance (ANOVA) followed by either the Least Significant Difference (LSD) test or median-notched box-and-whisker analysis, or the Kruskal-Wallis test followed by either the Mann-Whitney U-test or median-notched box-and-whisker analysis. Differences were considered statistically significant if the probability of error was less than 5%.
Results
Figure 1 depicts food intake as a function of time on four representative days (days 2—5) over the course of the seven-day monitoring period. Multifactorial ANOVA revealed that, compared to vehicle-treated controls, systemic administration of either the CB1 receptor agonist WIN 55,212-2 (1 mg/kg; s.c.) or antagonist AM251 (3 mg/kg; s.c.) resulted in a significant main effect (day 2: F-ratio = 81.07, p<0.0001; day 3: F-ratio = 49.54, p<0.0001; day 4: F-ratio = 61.43; p<0.0001; day 5: F-ratio = 152.2; p<0.0001) on the rate of food consumption. WIN 55,212-2 increased the rate of food intake, whereas AM251 decreased it. These effects became apparent 1—2 hour following their administration just prior to beginning the evaluation period, and lasted up to 4—6 hours into the monitoring. The agonist-induced increase in the cumulative rate of food intake shown for day 5 (like all other days) was completely abolished by co-administration of antagonist. There was no change in the total amount of food eaten over the course of the six-hour evaluation period (not shown). We also examined cannabinoid effects on meal frequency, meal duration and the amount of food eaten per meal. Figure 2 shows the effect of WIN 55,212-2 and AM251 on meal frequency ascertained every hour during the evaluation period. Multifactorial ANOVA again revealed significant main drug effect (F-ratio = 13.41, p<0.0001), as well as a significant interaction (F-ratio = 4.77, p<0.0001). WIN 55,212-2 increased meal frequency during the second hour, followed by a pronounced decrease during hours 3—6. AM251 completely blocked the agonist-induced increase meal frequency, and attenuated the prolonged decrease in meal frequency observed during the mid- to latter stages of the monitoring period. On the other hand, AM251 produced a decrease in meal frequency during hour 1, which was reversed by co-administration of the agonist. The increase in the rate of food consumption and meal frequency produced by WIN 55,212-2 was associated with a modest increase in meal duration (Figure 3) during hour 1 and a dramatic increase in the amount of food eaten per meal (Figure 4). These effects were eliminated by co-administration of the antagonist. WIN 55,212-2 was without effect on the rate of weight gain (Figure 5). Conversely, AM251 markedly decreased meal duration (Figure 3) and elicited a significant reduction in weight gain (Figure 5), and these effects were at least partially reversed by co-administration of the agonist. AM251 did not affect the amount of food eaten per meal, per se (Figure 4). Collectively, this indicates that CB1 receptor activation by both exogenous and endogenous cannabinoid agonists increases feeding behavior, whereas CB1 receptor antagonism suppresses it.
We then wanted to determine if the increased appetite produced by activation of CB1 receptors was associated with a decrease in glutamatergic synaptic input onto anorexigenic POMC neurons. Accordingly, we performed whole-cell patch-clamp recordings from arcuate neurons in hypothalamic slices. Figure 6 shows traces of excitatory postsynaptic currents (EPSCs) evoked by square-wave pulses delivered by a bipolar tungsten stimulating electrodes positioned 0.5—1.0 μm lateral to the recording electrode. Bath application of WIN 55,212-2 (1 μM) reduced the amplitude of evoked EPSCs (eEPSCs) that were subsequently abolished in the presence of the ionotropic glutamate receptor antagonists NBQX (3 μM) and CGS 19755 (10 μM; Figures 6A and 6C). On the other hand, AM251 (1 μM) per se increased eEPSC amplitude, and when co-administered with WIN 55,212-2 it blocked the inhibitory effect of the agonist (Figures 6B and 6C). Neither WIN 55,212-2 nor AM251 had any effect on the rate of rise or the decay of the eEPSCs (not shown). Further analysis using the paired-pulse paradigm revealed that the agonist-induced reduction in eEPSC amplitude was associated with an increase in the S2:S1 ratio (Figures 6D—6F). To verify that the agonist-induced reduction could be attributed to a presynaptic mechanism of action, we next performed an analysis of miniature EPSCs (mEPSCs) recorded in the presence of 500 nM TTX. Figures 7 and and88 show representative traces of mEPSCs recorded from arcuate neurons. Varying doses of WIN 55,212-2 (100 nM & 10 μM) produced graded increases in the interval between contiguous mEPSCs, which effectively translates into a decrease in mEPSC frequency, with no discernable effect on mEPSC amplitude (Figure 7A). These observations are substantiated further by the cumulative probability plots shown in Figure 7B. WIN 55,212-2 (100 nM) similarly, yet slightly less effectively, reduced the frequency of miniature, inhibitory postsynaptic currents (mIPSCs) recorded in the presence of TTX, NBQX and CGS 19755 from a holding potential of −30 mV (mIPSCs: 68.3 ± 7.6% of control (n = 3) vs. mEPSCs: 52.6 ± 6.5% of control (n = 5)). In addition, the selective effect of WIN 55,212-2 on mEPSC frequency was mimicked by the anandamide analog ACEA (not shown). On the other hand, AM251 both increased mEPSC frequency per se and blocked the diminution caused by WIN 55,212-2 (Figure 8A). The composite effects of WIN 55,212-2 and AM251 on mEPSC frequency and amplitude are summarized in Figure 8B. These data suggest that CB1 receptor activation by both exogenous and endogenous cannabinoid agonists presynaptically inhibits glutamatergic synaptic input onto arcuate neurons.
We noticed during the mEPSC experiments that approximately one-third of arcuate neurons (10/31) responded to both WIN 55,212-2 and ACEA with a change in the holding current, and thus explored this observation in greater detail. Figures 9A1 and 9B1 show representative examples of the outward current elicited by bath application of 1 μM ACEA. Current-voltage relationships generated prior to, and in the presence of, the agonist reveal that the current reverses polarity very near the Nernst equilibrium potential for K+, and exhibits an increase in slope conductance that grows more pronounced as the membrane potential is hyperpolarized below the reversal potential (Figure 9A2). This agonist-induced outward current was blocked by AM251 (Figure 9B2), and was found to be both qualitatively (Figure 9C) and quantitatively (Figure 9D) very similar to the G-protein-gated, inwardly-rectifying K+ current (GIRK) elicited by activation of metabotropic GABAB receptors. The activation of this postsynaptic K+ current, as well as the presynaptic inhibition of glutamatergic synaptic input described above, was observed in POMC neurons (n = 6) subsequently identified through immunohistofluorescent visualization (Figure 10). Taken together, these data demonstrate that cannabinoids increase feeding behavior, which is associated with a presynaptic inhibition of glutamatergic neurotransmission and a postsynaptic activation of an inwardly-rectifying K+ current within POMC neurons. This indicates that the cannabinoid modulation of appetite is mediated, at least in part, through interactions with the hypothalamic feeding circuitry.
Discussion
The results of the present study demonstrate unequivocally that cannabinoids stimulate appetite, which correlates with a decrease in the excitability of POMC neurons. These conclusions are based on the following observations: 1) CB1 receptor activation and blockade respectively increases and decreases feeding behavior as determined using four different indices: the rate of food intake, meal frequency, meal duration, the amount of food eaten per meal and change in body weight, 2) CB1 receptor activation and blockade respectively decreases and increases glutamatergic neurotransmission via a presynaptic site of action, 3) cannabinoids affect a CB1 receptor-mediated activation of an inwardly rectifying K+ current and 4) the latter two cellular effects of cannabinoids occur in arcuate neurons subsequently identified as POMC neurons via immunohistofluorescence. The current findings also suggest that glutamatergic synaptic input onto POMC neurons is tonically inhibited by endogenous cannabinoids, which provides further evidence that these neurons are a critical target in the cannabinoid modulation of appetite.
While WIN 55,212-2 has affinity for both CB1 and CB2 receptor subtypes [41], and CB2 receptors are found in the brainstem [42], the fact that the CB1 receptor antagonist AM251 largely abolished the agonist-induced hyperphagia, decrease in eEPSC amplitude, mEPSC frequency and the activation of the inwardly-rectifying K+ conductance clearly indicates that the effects observed in the present study are CB1 receptor-mediated. In the present study, CB1 receptor activation upon the exogenous administration of WIN 55,212-2 increased the rate of food intake, meal frequency, meal duration and the amount of food eaten per meal. Thus, exogenously administered cannabinoids increased eating efficiency. On the other hand, CB1 receptor antagonism decreased the rate of food intake, meal frequency and meal duration, indicating that endogenous cannabinoids drive appetite and food seeking behavior. The cannabinoid-induced hyperphagia that we observed in our guinea pig animal model is consistent with that reported in humans and other rodent models. Despite a limited number of controlled scientific studies, the ability of cannabinoids, in particular Δ 9-tetrahydrocannabinol (Δ 9-THC), to cause the sensation of hunger and increase food intake in humans has been recognized for centuries [2—5]. In rats and mice, exogenously administered cannabinoid agonists stimulate appetite under ad libitum and food-deprived conditions [1,2,43], whereas CB1 receptor antagonists suppress it [8,9]. In addition, Δ 9-THC promotes feeding in rats when focally injected into the ventromedial or lateral hypothalamus [2,4], and the satiety hormone leptin decreases hypothalamic levels of endogenous 2-AG [32]. Moreover, both ob/ob mice (which do not produce leptin) and fa/fa rats (which do not express leptin receptors) exhibit rampant hyperphagia and weight gain [32,44,45], as well as elevated hypothalamic levels of endogenous cannabinoids [32], with respect to their wildtype counterparts. Thus, the cannabinoid regulation of appetite is clearly conserved among mammalian species, and involves, at least in part, interactions with various neuroanatomical substrates within the hypothalamus.
The cannabinoid-induced presynaptic inhibition of glutamatergic synaptic input onto guinea pig arcuate POMC neurons is also consistent with that reported in several other regions of the central nervous system, including the prefrontal cortex [46], hippocampus [47,48], PAG [49] and the dorsal motor nucleus of the vagus [14]. As mentioned above, we also found that CB1 receptor antagonism unmasked a tonic inhibition of glutamate release onto POMC neurons, suggesting that endogenous cannabinoids are a principal neuromodulator influencing synaptic input onto these cells. In the hippocampus [50], cerebellum [51], ventral tegmental area [52] and other hypothalamic nuclei [53], endogenous cannabinoids released from postsynaptic neurons in response to depolarizing stimuli retrogradely inhibit glutamate release from nerve terminals impinging upon them. Indeed, the retrograde inhibition of glutamatergic synaptic input released by endogenous cannabinoids released from parvocellular neurons in the paraventricular nucleus of the hypothalamus is instrumental in mediating the rapid feedbacks of corticosteroids on the stress axis [53]. In transgenic mice whose POMC neurons are labeled with enhanced green fluorescent protein, however, endogenous cannabinoids retrogradely inhibit GABAergic but not glutamatergic synaptic input onto these cells [54]. This suggests that amino acid neurotransmission within POMC neurons is differentially regulated in the two species. Future studies will determine if endogenous cannabinoids synthesized de novo in guinea pig POMC neurons serve to tonically inhibit glutamate neurotransmission in a retrograde fashion. Our finding of a CB1 receptor-mediated activation of an inwardly-rectifying K+ current in POMC neurons is also in agreement with that observed in AtT20 cells transfected with the CB1 receptor [55], and in oocytes co-expressing the CB1 receptor and GIRK1 channel subtype [56,57]. This current cannot be attributed to other voltage-dependent K+ currents such as the A-type K+ current [38], and is virtually identical to that elicited by the activation of GABAB receptors, which have long been known to couple to GIRK in the arcuate nucleus, preoptic area, supraoptic nucleus and many other regions of the central nervous system [58—61].
Along with the ventromedial nucleus of the hypothalamus, the POMC neurons in the arcuate nucleus are the principal anorexigenic component of the hypothalamic feeding circuitry. These neurons release posttranslational byproducts of POMC such as α-melanocyte-stimulating hormone (α-MSH) β-endorphin, as well as CART that is co-expressed in these cells [34,45]. Indeed, β-endorphin and melanocortin (MC)4 receptor knockout mice display extensive hyperphagia [34] and are resistant to disease-induced cachexia [26], respectively. In addition, the anorexigenic effect of the CB1 receptor antagonist rimonabant is eliminated in CART-deficient transgenic mice [62]. POMC neurons are a key target of the satiety hormone leptin, which not only depolarizes and increases the firing rate of these cells, but also decreases hypothalamic levels of endogenous cannabinoids [25,32,33]. Glutamate increases β-endorphin secretion and POMC gene expression, and glutamatergic synaptic input is observed in POMC neurons in the arcuate nucleus and the nucleus of the tractus solitarius of transgenic mice [54,63,64]. While our findings clearly indicate that POMC neurons are a major hypothalamic locus for the hyperphagic actions of cannabinoids, there are in reality a multiplicity of hypothalamic sites and mechanisms involved in the induction of feeding behavior. For example, cannabinoids increase orexigenic NPY release from hypothalamic explants [65]. On the other hand, NPY neurons do not express CB1 receptors, and so the cannabinoid-induced increase in NPY release is due presumably to a decrease in GABA input onto these cells [31], much like the retrograde inhibition of GABA release onto orexigenic MCH neurons in the lateral hypothalamus [66]. Moreover, cannabinoid interactions with the brain stem, gut, pancreas and adipose tissue also play a role in the regulation of appetite and energy homeostasis [31]. Nevertheless, the presynaptic inhibition of glutamatergic neurotransmission within guinea pig POMC neurons must account, at least in part, for the cannabinoid-induced hyperphagia observed in the present study.
In conclusion, these results reveal that both endogenous and exogenous cannabinoids stimulate feeding behavior via a mechanism that involves, at least in part, a presynaptic inhibition of glutamate neurotransmission in anorexigenic POMC neurons. Our findings convey important insight into how cannabinoids increase appetite through their interactions with the hypothalamic feeding circuitry.
Source, Graphs and Figures: CANNABINOID-INDUCED HYPERPHAGIA: CORRELATION WITH INHIBITION OF PROOPIOMELANOCORTIN NEURONS?
