Dopaminergic Modulation of Endocannabinoid-Mediated Plasticity at GABAergic Synapses in the Prefrontal Cortex

Abstract

Similar to dopamine (DA), cannabinoids strongly influence prefrontal cortical functions, such as working memory, emotional learning, and sensory perception. Although endogenous cannabinoid receptors (CB₁Rs) are abundantly expressed in the prefrontal cortex (PFC), very little is known about endocannabinoid (eCB) signaling in this brain region. Recent behavioral and electrophysiological evidence has suggested a functional interplay between the dopamine and cannabinoid receptor systems, although the cellular mechanisms underlying this interaction remain to be elucidated. We examined this issue by combining neuroanatomical and electrophysiological techniques in PFC of rats and mice (both genders). Using immunoelectron microscopy, we show that CB₁Rs and dopamine type 2 receptors (D₂Rs) colocalize at terminals of symmetrical, presumably GABAergic, synapses in the PFC. Indeed, activation of either receptor can suppress GABA release onto layer 5 pyramidal cells. Furthermore, coactivation of both receptors via repetitive afferent stimulation triggers eCB-mediated long-term depression of inhibitory transmission (I-LTD). This I-LTD is heterosynaptic in nature, requiring glutamate release to activate group I metabotropic glutamate receptors. D₂Rs most likely facilitate eCB signaling at the presynaptic site as disrupting postsynaptic D₂R signaling does not diminish I-LTD. Facilitation of eCB–LTD may be one mechanism by which DA modulates neuronal activity in the PFC and regulates PFC-mediated behavior in vivo.

Introduction

Dopaminergic innervation of the prefrontal cortex (PFC) strongly modulates high-level cognitive processes (Le Moal and Simon, 1991; Goldman-Rakic, 1998; Seamans and Yang, 2004). Unlike fast ionotropic neurotransmitters, dopamine (DA) does not directly mediate synaptic transmission but affects it by altering the cellular or synaptic properties of target neurons (Greengard, 2001; Seamans and Yang, 2004; Iversen and Iversen, 2007). The recent link between DA and endocannabinoids (eCBs) (van der Stelt and Di Marzo, 2003; Laviolette and Grace, 2006) is particularly interesting in the PFC given the well known hallucinogenic effects of exogenous cannabinoids and the potential eCB association with neuropsychiatric disorders that reflect PFC dysfunction (Ujike and Morita, 2004; Koethe et al., 2009; Sewell et al., 2009). However, it remains unclear exactly how eCBs regulate PFC function and whether DA plays a role in this regulation.

Short- and long-term synaptic plasticity mediated by eCBs has been reported in several brain regions (for recent reviews, see Chevaleyre et al., 2006; Lovinger, 2008; Heifets and Castillo, 2009; Kano et al., 2009). Recent evidence in the midbrain suggests that DA, via D₂-like receptors (D₂Rs), can facilitate eCB signaling and eCB-mediated plasticity (Melis et al., 2004; Kreitzer and Malenka, 2005; Yin and Lovinger, 2006; Kreitzer and Malenka, 2007; Shen et al., 2008). Consistent with the observation that D₂R activation increases striatal levels of the eCB anandamide in vivo (Giuffrida et al., 1999), it was suggested that DA acts via postsynaptic D₂Rs by promoting the production of eCBs. However, like CB₁Rs, D₂Rs are coupled to G_i/o-proteins and their activation leads to the inhibition of adenylyl cyclase (AC), resulting in a reduction of cAMP levels and protein kinase A (PKA) activity. The observation that D₂Rs may be present at presynaptic locations in the PFC (Fadda et al., 1984; Seamans et al., 2001) raises the possibility that CB₁Rs and D₂Rs may colocalize on the same axon terminals and perhaps be coactivated by physiological synaptic activity. It is unknown how coincident activation of presynaptic CB₁Rs and D₂Rs in the PFC may affect transmitter release.

Most studies on eCB-mediated plasticity have concentrated on excitatory synapses, and the PFC is no exception. Pharmacological activation of CB₁Rs suppresses excitatory inputs to layer 5 pyramidal cells in rodent PFC (Auclair et al., 2000; Lafourcade et al., 2007), and repetitive stimulation in PFC can elicit long-term depression of excitatory transmission (E-LTD) (Lafourcade et al., 2007). However, given that a balance of excitation and inhibition shapes neuronal excitability and network firing patterns, knowledge of how eCBs influence inhibitory transmission in the PFC will be necessary to predict the functional impact of eCB signaling in this structure. Here, we investigate eCB signaling at inhibitory synapses in the PFC and demonstrate facilitation of inhibitory LTD (I-LTD) by the dopaminergic system. Our findings suggest a potential synaptic link between eCB and DA function in the PFC. Dysregulation of this crosstalk may play a role in neuropsychiatric disorders, such as schizophrenia.

Materials and Methods

Double CB₁R/D₂R immunocytochemistry for electron microscopy.

Female mice (n = 3, 2 months old, C57BL/6 strain) were deeply anesthetized with chloral hydrate (400 mg/kg body weight). CB₁R^−/− knock-out mice (n = 2; kindly provided by Dr. Giovanni Marsicano, Institute François Magendie, Bordeaux, France) (Marsicano et al., 2002) were used as controls of CB₁R antibody specificity. All animals were transcardially perfused with phosphate-buffered solution (PB) (0.1 m, pH 7.4) and then fixed by 500 ml of 0.1% glutaraldehyde, 4% formaldehyde (freshly depolymerized from paraformaldehyde), and 0.2% picric acid in 0.1 m PB, pH 7.4, prepared at 4°C. Tissue blocks were extensively rinsed in 0.1 m PB, pH 7.4. Rostral vibrosections containing the PFC were cut at 50 μm and collected in 0.1 m PB, pH 7.4, at room temperature (RT). Sections were preincubated in a blocking solution of 10% bovine serum albumin (BSA), 0.1% sodium azide, and 0.02% saponin prepared in Tris-HCl-buffered saline (TBS), pH 7.4, for 30 min at RT.

A preembedding silver-intensified immunogold method and an immunoperoxidase method were used for the colocalization of CB₁R and D₂R in mouse PFC sections. Two primary polyclonal antibodies were used in this study: goat CB₁R raised against a 31 aa C-terminal sequence (GenBank accession number NM007726) of the mouse CB₁R (CB1-Go-Af450-1; Frontier Science) and rabbit D₂R raised against a 28 aa sequence (amino acids 284-311) within the third cytoplasmic loop of the human D₂R, which is common to both long and short forms of the receptor (AB5084P; Millipore Bioscience Research Reagents). The CB₁R (2 μg/ml) and D₂R (5 μg/ml) antibodies prepared in 10% BSA/TBS containing 0.1% sodium azide and 0.004% saponin were incubated simultaneously in PFC tissue on a shaker for 2 d at 4°C.

After several washes in 1% BSA/TBS, sections were incubated in a secondary 1.4 nm gold-labeled rabbit anti-goat IgG (Fab′ fragment, 1:100; Nanoprobes) for the detection of CB₁R and in a biotinylated donkey anti-rabbit IgG (1:200; Jackson ImmunoResearch) for the detection of D₂R, in both 1% BSA/TBS with 0.004% saponin on a shaker for 4 h at RT. PFC tissue was washed in 1% BSA/TBS and processed by a conventional avidin–biotin horseradish peroxidase complex method (ABC Elite; Vector Laboratories). The sections were washed in 1% BSA/TBS overnight at 4°C and postfixed in 1% glutaraldehyde in TBS for 10 min at RT. After washes in double-distilled water, gold particles were silver intensified with an HQ Silver kit (Nanoprobes) for ∼12 min in the dark and then washed in 0.1 m PB, pH 7.4. The tissue was preincubated subsequently with 0.05% DAB in 0.1 m PB for 5 min, incubated by adding 0.01% hydrogen peroxide to the same solution for 5 min, and washed in 0.1 m PB for 2 h at RT. Stained sections were osmicated (1% OsO₄ in 0.1 m PB, pH 7.4, 20 min), dehydrated in graded alcohols to propylene oxide, and plastic-embedded flat in Epon 812. Ultrathin sections (80 nm) were collected on mesh nickel grids, stained with lead citrate, and examined in a Philips EM2008S electron microscope. Tissue preparations were photographed by using a digital camera coupled to the electron microscope.

Specificity of the immunostainings was assessed as follows: CB₁R antibodies were incubated in CB₁R^−/− PFC tissue in the same conditions as above. In this case, CB₁R immunolabeling was not observed in cortical synapses of CB₁R deficient mice (data not shown). The D₂R antibodies gave a similar faint cortical pattern as described in the light microscopy in previous publications using the same antiserum (Wang and Pickel, 2002; Lei et al., 2004). In addition, no specific staining was detected in sections in which the primary D₂R antibodies were replaced by goat serum (data not shown). Figure compositions were scanned at 300 dots per inch. Labeling and minor adjustments in contrast and brightness were made using Adobe Photoshop (CS; Adobe Systems).

Quantitative analysis of electron microscopy images.

The quantitative analysis was made in layer 5/6 of PFC sections obtained from three mice processed for CB₁R/D₂R colocalization with preembedding immunocytochemistry. PFC sections, 50 μm thick, from each animal showing good and reproducible DAB immunoreaction (D₂R) and silver-intensified gold particles (CB₁R) were cut at 80 nm. Electron micrographs (18,000–28,000×) were taken from grids (132 μm side) containing DAB immunodeposits; all ultrathin sections showed a similar DAB labeling intensity, indicating that selected areas were at the same depth. Positive CB₁R labeling was determined by observation of at least one immunoparticle within ∼30 nm of the plasmalemma of axon terminals forming symmetrical-type synapses. Presynaptic boutons with D₂R immunodeposits making symmetrical synapses were also counted. Percentages of presynaptic inhibitory-like profiles positive for CB₁R or D₂R were analyzed and displayed using a statistical software package (GraphPad Prism 4; GraphPad Software). Finally, the percentage of CB₁R-positive terminals at symmetrical synapses that colocalize with D₂R as well as the percentage of immunoreactive D₂R axonal boutons with CB₁R immunolabeling that form symmetrical synapses were also analyzed. Data are presented as mean ± SEM. A total of 250 inhibitory-like presynaptic axon terminals in an area of 642.492 μm² were measured and analyzed by NIH ImageJ (version 1.36).

Electrophysiological recordings.

All animal experiments were performed in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Acute prefrontal cortical slices were prepared from postnatal day 16 (P16) to P25 Wistar rats and P24–P46 C57BL/6 mice from Charles River Laboratories or from our colony of Zimmer line CB₁R^−/− mice (Zimmer et al., 1999). The CB₁R^−/− mouse colony was maintained as heterozygotes and genotyped before use as described previously (Takahashi and Castillo, 2006). Male and female animals were anesthetized with isoflurane and decapitated, and their brains were removed to a chilled cutting solution consisting of the following (in mm): 215 sucrose, 2.5 KCl, 26 NaHCO₃, 1.6 NaH₂PO₄, 1 CaCl₂, 4 MgCl₂, 4 MgSO₄, and 20 glucose. The posterior end of the brain was truncated flat with a razor blade, and then the brain was positioned in the cutting chamber with the cut end on the bottom. Coronal slices (350 μm thick) were cut with a DTK-2000 (Dosaka) or Leica VT1200 S (Leica Microsystems) vibrating microslicer. Rostral sections containing the prelimbic–infralimbic region of the PFC were stored at room temperature in holding solution containing the following (in mm): 63 NaCl, 2.5 KCl, 22 NaHCO₃, 1.4 NaH₂PO₄, 1.1 CaCl₂, 3.2 MgCl₂, 2 MgSO₄, 15.5 glucose, and 107.5 sucrose. Thirty minutes after sectioning, slices were placed in extracellular recording solution containing the following (in mm): 126 NaCl, 2.5 KCl, 18 NaHCO₃, 1.2 NaH₂PO₄, 1.2 CaCl₂, 2.4 MgCl₂, and 11 glucose. All solutions were saturated with 95% O₂ and 5% CO₂, pH 7.4. Slices were incubated for at least 30 min in the recording solution before experiments.

All experiments were conducted in a submersion-type recording chamber perfused at ∼2 ml/min, and the recording temperature was maintained at 30 ± 1°C using a TC-344B dual-channel heater controller (Warner Instruments). Whole-cell patch-clamp experiments were performed in 10 μm 2,3-dioxo-6-nitro-1,2,3,4-tetrahydrobenzo[f]quinoxaline-7-sulfonamide (NBQX) or CNQX (AMPAR/kainate receptor antagonists) and 25 μm APV (NMDAR antagonist). In recordings of miniature IPSCs (mIPSCs), 1 μm tetrodotoxin was additionally applied in the bath. To record GABA_AR-mediated IPSCs, layer 5 pyramidal cells were voltage clamped at 0 mV using patch-type pipettes filled with intracellular solution containing the following (in mm): 131 Cs-gluconate, 8 NaCl, 1 CaCl₂, 10 EGTA, 10 glucose, 10 HEPES, 5 MgATP, and 0.4 Na₃GTP, pH 7.2 (285 mmol/kg). In experiments with postsynaptic calcium chelators, 20 mm Cs and 10 mm EGTA was replaced with 20 mm BAPTA. In some experiments, we assessed the electrophysiological properties of the patched pyramidal cells using a K-based internal solution containing the following (in mm): 130 K-gluconate, 10 NaCl, 0.2 EGTA, 10 HEPES, 5 MgATP, and 0.4 Na₃GTP, pH 7.2 (285 mmol/kg). Series resistance (typically 5–15 MΩ) was monitored throughout the experiment with a −4 mV, 80 ms voltage step, and recordings with a >10% change in series resistance were excluded from analysis.

Stimulating electrodes were filled with extracellular recording solution and placed in layer 2/3 to stimulate distal inhibitory inputs and layer 5 to stimulate proximal inhibitory inputs. Patched cells were typically no more than 50 μm lateral from the tips of stimulating electrodes. Each pathway was stimulated every 20 s, and synaptic responses typically ranged from 200 to 800 pA. I-LTD in the rat was triggered in layer 2/3 inputs by a 5 Hz train of synaptic stimulation lasting 10 min, at twice the test stimulus intensity. In the mouse PFC, a shorter 5 Hz train (5 min) was required to induce I-LTD. Stimulation was triggered by a Master-8 pulse generator (A.M.P.I.), and acquisition was controlled by custom-written software in Igor Pro 4.09A (Wavemetrics). Whole-cell patch-clamp recordings were performed using a Multiclamp 700B amplifier (Molecular Devices).

To assess the efficacy of loading protein kinase A inhibitor (PKI) 6–22 peptide in the patch pipette to inhibit postsynaptic PKA activity, experiments were performed in the CA1 region to monitor slow afterhyperpolarization potential currents (I_AHP), which is modulated by PKA activation (Pedarzani and Storm, 1993). Transverse hippocampal slices (400 μm) were prepared from P23–P25 Wistar rats. Slow I_AHP were elicited by a depolarizing step (between 60 and 70 mV) for 100 ms in CA1 pyramidal cells voltage clamped between −50 to −55 mV with intracellular solution containing the following (in mm): 120 K-gluconate, 20 KCl, 15 HEPES, 2 Mg-ATP, 0.2 GTP, 1 MgCl₂, and 0.1 EGTA. Sp-cAMPS at 50 μm was used to inhibit the slow I_AHP component, and the I_AHP amplitude was measured 15–20 min after Sp-cAMPS application under control conditions or in cells loaded with 2.5 or 100 μm PKI 6–22.

Quantitative analysis of electrophysiological data.

All values are provided as mean ± SEM. The paired-pulse ratio (PPR) was defined as the ratio of the amplitude of the second IPSC over the amplitude of the first IPSC. The magnitude of drug effects was calculated as the percentage change between baseline (averaged responses for the 10 min before drug application) and 20–30 min after start of WIN 55,212-2 (WIN) [R-(+)-[2,3-dihydro-5-methyl-3-(4-morpholinylmethyl)pyrrolo[1,2,3-de]-1,4-benzoxazin-6-yl]-1-naphthalenyl methanone mesylate] or H89 (N-[2-[[3-(4-bromophenyl)-2-propenyl]amino]ethyl]-5-isoquinolinesulfonamide dihydrochloride) application, 10–15 min after start of quinpirole application or 20–30 min after the start of bromocriptine application. The magnitude of I-LTD was calculated as the percentage change between baseline and 20–30 min after complete delivery of tetanus. Statistical analysis was performed using Student's independent t test at the p < 0.05 significance level in OriginPro 7.0 software (OriginLab Corp), unless otherwise stated.

Drugs.

WIN 55,212-2, (−)-quinpirole hydrochloride, bromocriptine mesylate, (S)-(−)-sulpiride, SCH23390 [R(+)-7-chloro-8-hydroxy-3-methyl-1-phenyl-2,3,4,5-tetrahydro-1H-3-benzazepine] hydrochloride, LY367385 [(S)-(+)-α-amino-4-carboxy-2-methylbenzeneacetic acid], OR 486 (3,5-dinitrocatechol), and picrotoxin were purchased from Tocris Bioscience. PKI 6–22 and PKI 14–22 peptides were obtained from either Tocris Bioscience or BIOMOL Research Laboratories. Sp-cAMPS was acquired from BIOMOL Research Laboratories. d-APV, NBQX, CNQX, 2-methyl-6-(phenylethynyl)-pyridine (MPEP) hydrochloride, and AM 251 [N-(piperidin-1-yl)-1-(2,4- dichlorophenyl)-5-(4-iodophenyl)-4-methyl-1H-pyrazole-3-carboxamide] were obtained from Ascent Scientific. H89 was procured from LC Labs. Total DMSO in the bath solution was maintained at 0.1% or below in all experiments.

Results

Anatomical evidence for CB₁R and D₂R presynaptic colocalization in the mouse PFC

We first assessed anatomically whether CB₁Rs and D₂Rs colocalize at inhibitory synaptic terminals in the PFC. Double immunoelectron microscopy was used to precisely determine the localization of CB₁Rs and D₂Rs in layers 2/3 (Fig. 1A,B) and 5/6 (Fig. 1C–F) of the mouse PFC. Immunoperoxidase labeling of D₂Rs (thick arrows) and immunogold labeling of CB₁Rs (thin arrows) was frequently observed within axonal profiles in both layers (Fig. 1A–F), which formed symmetrical synapses onto small postsynaptic dendrites and somatic membranes. CB₁R immunoparticles were consistently associated with portions of presynaptic membranes away from the active zones, as expected for its known perisynaptic localization. To assess the degree of colocalization between CB₁Rs and D₂Rs, quantitative measurements were performed in layer 5/6 of PFC sections (Fig. 1G). Notably, ∼50% of relatively small presynaptic boutons with restricted immunoreaction for the D₂R colocalized with CB₁R immunoparticles along their membranes. A similarly high proportion of inhibitory presynaptic boutons with CB₁R immunolabeling also contained D₂R immunoreactivity. Although this number is probably an underestimation, ∼30% of presynaptic boutons with pleomorphic synaptic vesicles and symmetrical synapses were immunonegative for both CB₁R and D₂R in layers 5/6. The high number of symmetrical synapses expressing either CB₁R or D₂R suggests that eCBs and DA play significant roles in modulating inhibitory transmission in the PFC. The superficial layer 2/3 also exhibited the same pattern of CB₁R/D₂R colocalization in presynaptic terminals impinging symmetrically on postsynaptic membranes of cell bodies and small-diameter dendrites (Fig. 1A,B). Thus, our electron microscopy (EM) images reveal a strong pattern of colocalization between CB₁Rs and D₂Rs at terminals of symmetrical, presumably inhibitory, synapses in both layers 2/3 and 5/6 of the mouse PFC.

Figure 1.

Colocalization of presynaptic CB₁R and D₂R at symmetrical synapses in the mouse PFC. Double labeling of receptors by combining a pre-embedding immunogold (CB₁R) and an immunoperoxidase (D₂R) method for electron microscopy (A–F). D₂R-immunoreactive (thick arrows) presynaptic axon terminals (ter) forming symmetrical synapses (arrowheads) with small dendrites (den) and cell bodies (soma) colocalize with CB₁R immunoparticles (thin arrows) at their perisynaptic and extrasynaptic membranes. Note in A CB₁R immunolabeling (thin arrows) in a D₂R-immunonegative synaptic bouton making a somatic symmetrical synapse (arrowheads). Scale bars, 0.5 μm. G, Left, percentages (mean ± SEM) of the total number of symmetrical synapses analyzed (n = 280) containing CB₁R alone, D₂R alone, both CB₁R and D₂R, and neither CB₁R nor D₂R in layers 5/6 of the mouse PFC. Right, Approximately 50% of CB₁R-immunopositive inhibitory-like synaptic terminals have D₂R immunoreactivity, and 50% of D₂R immunoreactive symmetrical boutons are also equipped with CB₁R immunometals.

Pharmacological activation of CB₁Rs or D₂Rs suppresses GABAergic transmission in mouse and rat PFC, most likely via a presynaptic mechanism

Because both CB₁Rs and D₂Rs are present at terminals of symmetrical presumably GABAergic synapses, we next tested whether activation of these receptors has any functional effect on GABAergic synaptic transmission. Whole-cell recordings of pharmacologically isolated IPSCs were obtained from layer 5 pyramidal cells in acute mouse PFC slices. The identity of pyramidal cells was confirmed by assessment of electrophysiological properties and pyramidal cell-like morphology under infrared-differential interference contrast (IR-DIC) microscopy (Fig. 2A). Exogenous bath application of the CB₁R agonist WIN 55,212-2 (5 μm) suppressed IPSCs evoked by focal stimulation via electrodes placed distally in layer 2/3 as well as proximally in layer 5 (Fig. 2B). IPSC amplitudes after WIN application were 49.4 ± 7.9 and 56.3 ± 9.9% of baseline in layer 2/3 and layer 5, respectively (n = 4). Consistent with a reduction in GABA release, PPR increased from 0.83 ± 0.03 to 0.94 ± 0.04 in layer 2/3 (p < 0.05) and from 0.84 ± 0.03 to 0.92 ± 0.04 in layer 5 (p < 0.05). Similarly, activation of D₂Rs by the agonist quinpirole (1 μm) also depressed IPSCs evoked in both layers (Fig. 2C). IPSC amplitudes after quinpirole application were 61.3 ± 8.6 and 59.3 ± 8.9% of baseline in layer 2/3 and layer 5, respectively (n = 6). This reduction was associated with a change in PPR in both layers (layer 2/3, from 0.84 ± 0.06 to 0.94 ± 0.05; layer 5, from 0.76 ± 0.03 to 0.93 ± 0.06, p < 0.05). Thus, activation of CB₁Rs or D₂Rs decreases GABAergic transmission in the mouse PFC, most likely via a reduction in GABA release.

Figure 2.

Pharmacological activation of CB₁Rs or D₂Rs suppresses GABAergic responses evoked by stimulation in layers (L) 2/3 and 5 in the mouse PFC. A, Left, Representative response of whole-cell patched layer 5 pyramidal cell in current clamp (above) while depolarizing or hyperpolarizing current injections were delivered (below) to show that targeting cells in layer 5 by their shape is a reliable way of patching pyramidal cells. Note the initial spike adaption, followed by a slow regular spiking behavior. Right, Photograph of image under IR-DIC microscopy of a patched cell in layer 5 of the PFC exhibiting a pyramidal cell-like morphology. B, Time course of WIN 55,212-2 suppression on both proximal (black circles) and distal (white circles) GABAergic inputs in the mouse PFC (n = 4 cells). Top, Representative average IPSC traces from a single experiment, obtained at the time points in the time course graph. Right, Bar graph of the average PPR before and after WIN application. Open symbols represent the PPR of individual experiments. C, Time course of quinpirole (Quin) suppression on both proximal (black circles) and distal (white circles) GABAergic inputs in the mouse PFC (n = 6 cells). Top, Representative average IPSC traces from an individual experiment, obtained at the time points indicated. Right, Bar graph of the average PPR before and after quinpirole application. Open symbols represent the PPR of individual experiments. *p < 0.05.

WIN and quinpirole inhibition of GABAergic transmission is not exclusive to the mouse PFC. In rat PFC slices, IPSCs evoked by stimulating in layer 2/3 and recording from layer 5 pyramidal cells were also suppressed by 5 μm WIN to 41.4 ± 4.5% of baseline (n = 5) after WIN application, and PPR increased from 0.77 ± 0.09 to 0.89 ± 0.09 (p < 0.01, paired t test) (Fig. 3A). The suppression was indeed mediated by CB₁Rs because preincubation with the antagonist AM 251 (4 μm) abolished the WIN effect (91.8 ± 3.9% of baseline, n = 5, p < 0.005). GABAergic inputs in the rat were also sensitive to quinpirole because bath application of this D₂R agonist reduced IPSCs to 66.9 ± 4.6% of baseline (n = 8), an effect that is accompanied by a change in PPR from 0.71 ± 0.07 to 0.81 ± 0.05 (p < 0.05) (Fig. 3B). Preincubation with the D₂R antagonist sulpiride (10 μm) blocked quinpirole-induced suppression of GABAergic transmission (99.0 ± 3.8% of baseline, n = 6, p < 0.005), supporting a role for D₂Rs in this quinpirole effect. Moreover, a different and selective D₂R agonist, bromocriptine (2 μm), also suppressed IPSCs, reducing the amplitude to 72.5 ± 4.2% of baseline (n = 6, p < 0.005) along with a change in PPR from 0.85 ± 0.04 to 0.99 ± 0.07 (p < 0.05, paired t test). Sulpiride also blocked the suppression by bromocriptine (102.5 ± 3.6% of baseline, n = 6, p < 0.005). In the striatum, D₂R activation may lead to eCB mobilization (Giuffrida et al., 1999; Yin and Lovinger, 2006), leading to a reduction in neurotransmitter release via CB₁R activation (Yin and Lovinger, 2006). To explore this possibility in the PFC, we examined the role of CB₁Rs in quinpirole suppression of IPSCs; however, AM 251 did not block suppression by quinpirole (60.2 ± 9.5% of baseline, n = 5, p < 0.05). Together with the EM data showing presynaptic localization of CB₁Rs and D₂Rs, the PPR changes after WIN or quinpirole application support a functional role for CB₁Rs and D₂Rs in modulating GABA release in both mouse and rat PFC.

Figure 3.

Activation of CB₁R or D₂R suppresses GABAergic transmission in the rat PFC. A, Time course of WIN suppression of IPSC amplitudes in the rat PFC (black circles), which is abolished in CB₁R blockade with 4 μm AM 251 (white circles). Top left, Representative average IPSC traces from a single control experiment, obtained at time points indicated. Top right, PPR plot before and after WIN application. PPR of layer (L) 2/3 IPSCs significantly increased. Open symbols represent the PPR of individual experiments. B, Time course of quinpirole (Quin) suppression of IPSC amplitudes in the rat PFC in control (black circles), under D₂R antagonism with 10 μm sulpiride (Sulp; white circles), and under CB₁R block with 4 μm AM 251 (gray circles). Top left, Representative average IPSC traces from a single control experiment. Top right, PPR plot before and after quinpirole application. PPR of layer 2/3 IPSCs significantly increased under control conditions and in AM 251. Open symbols represent the PPR of individual experiments. Bottom right, Summary plot of suppression mediated by D₂R agonists quinpirole (1 μm) and bromocriptine (Brcriptine; 2 μm). Effects of both drugs were blocked by sulpiride. However, AM 251 had no effect on quinpirole suppression. The number of experiments for each condition is indicated in parentheses. *p < 0.05, **p < 0.01, ***p < 0.005.

Endogenous PKA activity is critical for basal release of GABA and is most likely involved in the suppression of inhibitory transmission by WIN and quinpirole in the PFC

Both CB₁Rs and D₂Rs are coupled to G_i/o-proteins, and their activation leads to inhibition of the AC/cAMP/PKA pathway (Howlett et al., 2004; Neve et al., 2004), which may be responsible for the suppression of IPSCs in the PFC by WIN (Figs. 2B, 3A) and by quinpirole (Figs. 2C, 3B). If this were the case, we expect that directly blocking PKA activity with the inhibitor H89 should also reduce basal GABA release in the PFC. As in the hippocampus (Chevaleyre et al., 2007), bath application of 10 μm H89 resulted in a reduction of IPSCs to layer 5 pyramidal cells (65.0 ± 7.4% of baseline, n = 6), which was accompanied by a change in PPR (from 0.72 ± 0.03 to 0.84 ± 0.03, p < 0.05, paired t test) (Fig. 4A). It is known that inhibition of voltage-gated calcium channels (VGCCs) and PKA contribute to CB₁R-mediated depression of inhibitory transmission (Chevaleyre et al., 2006; Lovinger, 2008; Heifets and Castillo, 2009; Kano et al., 2009); thus, we examined the contribution of PKA in WIN suppression in the PFC by measuring spontaneous miniature IPSCs (mIPSCs) to reduce contamination by the VGCC-sensitive component (Fig. 4B). As reported previously (Vaughan et al., 1999; Takahashi and Linden, 2000; Trettel and Levine, 2002; Chevaleyre et al., 2007), WIN (5 μm) reduced mIPSC frequency (from 4.14 ± 0.99 to 2.00 ± 0.41 Hz, n = 4, p < 0.05, paired t test) but not amplitude (from 11.3 ± 1.8 to 10.5 ± 1.5 pA, p > 0.05, paired t test). Preincubating PFC slices in 10 μm H89 lowered mIPSC frequency (1.62 ± 0.29 Hz, n = 5, p < 0.05) and occluded additional reduction by subsequent WIN application (1.50 ± 0.19 Hz, n = 5, p > 0.05, paired t test). The mechanism by which D₂Rs suppress GABA release is less clear. One study in the ventral tegmental area (VTA) suggests that activation of D₂Rs at the presynaptic terminal may suppress inhibitory transmission solely via a decrease in PKA activity (Pan et al., 2008). Another study using two-photon calcium imaging of axonal varicosities of VTA medium spiny neurons also implicate calcium sources in D₂R-mediated inhibition (Mizuno et al., 2007). We tested whether quinpirole suppression is sensitive to PKA inhibition and found that, in slices preincubated in H89, evoked IPSC amplitudes were unchanged after quinpirole application (108.9 ± 9.4% of baseline, n = 4) (Fig. 4C). To confirm that the lack of quinpirole suppression in H89 indeed resulted from inhibition of PKA activity, we also used the potent cell-membrane-permeable PKA inhibitor peptide PKI 14–22 and found similar block of the quinpirole effect (97.4 ± 6.1% of baseline, n = 6, p < 0.005). Thus, our results strongly suggest that PKA activity is involved in basal GABA release, and, by inhibiting this activity, CB₁Rs or D₂Rs can suppress GABAergic transmission.

Figure 4.

Inhibiting PKA activity with H89 reduces GABA release and occludes WIN and quinpirole suppression of GABAergic transmission in the rat PFC. A, Time course of H89 suppression of IPSC amplitudes in the rat PFC (n = 6 cells). Top left, Representative average IPSC traces from a single experiment, obtained at the time points indicated. Top right, PPR plot before and after H89 application. PPR of layer (L) 2/3 IPSCs significantly increased. Open symbols represent the PPR of individual experiments. B, WIN suppression of mIPSC frequency in control is absent in slices preincubated in 10 μm H89. Top left, Three representative mIPSC traces from a single control experiment before and after WIN application. Top right, Three representative mIPSC traces from a single H89 experiment before and after WIN application. C, Quinpirole (Quin) suppression of IPSC amplitudes in control (black circles) is absent in slices preincubated in either 10 μm H89 (white circles) or 2.5 μm PKI 14–22 (gray circles). Top, Representative average IPSC traces from single experiments. Bottom right, Summary plot of quinpirole suppression in control and block under PKA inhibition by H89 and PKI 14–22. The number of experiments for each condition is indicated in parentheses. *p < 0.05, ***p < 0.005.

Pharmacological activation of D₂Rs facilitates induction of I-LTD

Our EM images show colocalization of CB₁Rs and D₂Rs at terminals of symmetrical synapses in the PFC, and we have provided functional evidence that independent activation of either receptor suppresses GABA release. Given that both CB₁Rs and D₂Rs can signal via PKA inhibition (Howlett et al., 2004; Neve et al., 2004), we next wondered whether simultaneous activation of both receptors will reveal an interaction between D₂R- and CB₁R-mediated effects. To this aim, we applied a low dose of quinpirole (500 nm) in the absence or presence of a submaximal concentration of WIN (50 nm) (Fig. 5A). Interestingly, although by itself 500 nm quinpirole had very little effect of basal IPSC amplitude (96.2 ± 6.2% of baseline, n = 10), it significantly decreased IPSCs when CB₁Rs were coactivated (78.4 ± 10.7% of baseline, n = 8, p < 0.05). We next tested whether D₂R activation could enhance eCB signaling in the PFC by delivering a 5 Hz train of synaptic stimulation (see Materials and Methods) in the presence of 500 nm quinpirole. Although on its own the 5 Hz stimulation was below threshold to trigger I-LTD, it became suprathreshold when D₂Rs were activated (Fig. 5B). IPSC amplitudes after train were 97.6 ± 11.0% of baseline (n = 5) and 60.6 ± 3.3% of baseline (n = 14) in the absence and presence of quinpirole, respectively (p < 0.005). Consistent with a role for D₂Rs, I-LTD was abolished in the presence of sulpiride (98.6 ± 2.2% of baseline, n = 4, p < 0.01) (Fig. 5C). Importantly, I-LTD was dependent on CB₁R activation because it was blocked in AM 251 (99.6 ± 11.9% of baseline, n = 5, p < 0.01) (Fig. 5D) and was occluded by preincubation with 5 μm WIN (99.1 ± 7.6% of baseline, n = 6, p < 0.01) (Fig. 5E).

Figure 5.

I-LTD can be triggered by a 5 Hz train of synaptic stimulation in the presence of low-dose quinpirole. A, Time course of 500 nm quinpirole effect on IPSC amplitude in the presence (black circles) or absence (white circles) of 50 nm WIN. Preincubation with a submaximal dose of WIN (+WIN) reveals the suppressive effect of an otherwise ineffective dose of quinpirole (control). Top, Representative average IPSC traces from single experiments obtained from the time points indicated. B, Time course plot of average IPSC amplitudes before and after 5 Hz stimulation train in the absence (white circles) or presence (black circles) of quinpirole. Responses only persistently depressed as a result of 5 Hz stimulation when D₂R agonist is in the bath (+Quin). Top left, Representative average IPSC traces from single experiments obtained from the time points indicated. Top right, Plot of PPR before and after train in the presence of D₂R agonist. Open symbols represent single experiments. C, Time course of control I-LTD (black circles) and the absence of I-LTD under D₂R antagonism with 10 μm sulpiride (white circles). Top, Representative average IPSC traces from a single control and sulpiride block experiment. D, Time course of control I-LTD (black circles) and the absence of I-LTD in the presence of 4 μm AM 251 (white circles). Top, Representative average IPSC traces from a single control and AM 251 block experiment. E, Time course of control I-LTD (black circles) and its absence in slices that were preincubated in 5 μm WIN (white circles). Top, Representative average IPSC traces from a single control and WIN occlusion experiment. F, Time course of control I-LTD (black circles) and the lack of I-LTD in the presence of group I mGluR antagonists (white circles). Top, Representative average IPSC traces from single control and block experiments. *p < 0.05. L, Layer; LY, LY367385.

In many brain regions, eCB mobilization in LTD is triggered by activation of group I metabotropic glutamate receptors (mGluRs) (Chevaleyre et al., 2006; Heifets and Castillo, 2009; Kano et al., 2009). Given evidence of the expression of these receptors in cell bodies and dendritic bundles of PFC pyramidal cells in layer 5 (Defagot et al., 2002; Lafourcade et al., 2007), we next examined the involvement of group I mGluRs in I-LTD. Indeed, I-LTD was abolished by the group I mGluR antagonists LY367385 and MPEP (98.9 ± 8.9% of baseline, n = 6, p < 0.005), whereas interleaved I-LTD controls were normal (55.7 ± 5.2% of baseline, n = 6) (Fig. 5F). Thus, despite the need for quinpirole, I-LTD in the PFC is similar to the more classical form of eCB–LTD in other brain regions in that its induction requires group I mGluR activation.

DA facilitation of I-LTD most likely occurs via synergism between presynaptic D₂R and CB₁R signaling

D₂Rs may act at several sites to facilitate I-LTD. We investigated the different possibilities in the following experiments. First, we tested whether D₂Rs postsynaptically enhance eCB production and release in the PFC as has been suggested in the striatum and VTA (Giuffrida et al., 1999; Melis et al., 2004; Yin and Lovinger, 2006). Given that the AC/cAMP/PKA pathway is a principal effector of D₂R-mediated effects (Neve et al., 2004), we blocked PKA in the entire slice by bath application of H89 (10 μm) or PKI 14–22 (2.5 μm) and compared this effect with inhibiting PKA specifically in the postsynaptic cell (Fig. 6A). Consistent with a presynaptic site of action by D₂Rs, bath application of H89 or PKI 14–22 abolished I-LTD (92.6 ± 4.9% of baseline, n = 6, p < 0.01; and 97.0 ± 14.3% of baseline, n = 6, p < 0.01, respectively), whereas loading the membrane-impermeable PKI 6–22 peptide (2.5 or 100 μm) into the patch pipette did not prevent I-LTD (57.9 ± 5.6% of baseline, n = 4; and 53.8 ± 5.1% of baseline, n = 5, respectively). We verified that this loading procedure could reliably inhibit postsynaptic PKA by assaying the ability of the PKI peptide to reduce the PKA-dependent inhibition of the slow AHP current (I_AHP) in CA1 pyramidal cells. Interestingly, loading PKI 6–22 at 2.5 or 100 μm were equally effective at decreasing the inhibition of slow I_AHP induced by bath application of the PKA activator Sp-cAMPS (Fig. 6B). Hence, although I-LTD in the PFC is dependent on PKA activity, it appears that postsynaptic PKA does not play a significant role.

Figure 6.

Neither postsynaptic PKA nor intracellular calcium rise is required for I-LTD. A, Time course of I-LTD in interleaved controls (black circles), bath-applied H89 (10 μm; white triangles), bath-applied PKI 14–22 peptide (2.5 μm; white squares), and cells loaded with 2.5 μm PKI 6–22 peptide (white circles). Inhibiting PKA activity in the slice blocks I-LTD, but inclusion of PKI (at either 2.5 or 100 μm) in the postsynaptic cell does not, as depicted in the summary bar plots on the top and bottom right. Top, Representative average IPSC traces from single H89 and PKI experiments obtained from the time points indicated. B, Verification that intracellular loading of the PKI 6–22 peptide effectively inhibits postsynaptic PKA activity. PKI loaded into CA1 pyramidal cells at either 2.5 or 100 μm significantly reduced the inhibition of slow AHP current (I_AHP) induced by bath application of a specific PKA activator (Sp-cAMPS). Top, Representative average AHP traces from single experiments at the time points indicated. Bottom right, Summary bar plot depicting the effect of loading PKI on slow I_AHP inhibition. C, Time course of I-LTD in control (black circles) and in cells loaded with 20 mm BAPTA (white circles). Chelating postsynaptic calcium has no effect on I-LTD. Top, Representative average IPSC traces from single experiments. The number of experiments for each condition is indicated in parentheses. **p < 0.01, ***p < 0.005. L, Layer; Quin, quinpirole.

Although inhibition of PKA activity is a main target of D₂R signaling, alternative molecular players such as intracellular calcium stores have been linked to D₂R-mediated effects (Nishi et al., 1997; Hernandez-Lopez et al., 2000). We tested whether intracellular calcium rise is necessary for eCB mobilization in I-LTD by loading the postsynaptic cell with the fast calcium chelator BAPTA (Fig. 6C). In the presence of quinpirole, the 5 Hz train of synaptic stimulation still led to a long-lasting depression of IPSCs in the PFC (59.6 ± 6.6% of baseline, n = 5). Thus, similar to the hippocampus (Chevaleyre and Castillo, 2003) and amygdala (Azad et al., 2004), I-LTD in the PFC does not require postsynaptic calcium increases, further bolstering the idea that postsynaptic D₂Rs are not involved. Our findings that D₂Rs also localize to presynaptic inhibitory-like terminals and that activation of these receptors may reduce GABA release via PKA inhibition make presynaptic D₂Rs the most likely mediators of I-LTD facilitation.

In the hippocampus, the level of spontaneous GABAergic activity can determine whether I-LTD occurs (Heifets et al., 2008). It was shown that I-LTD requires calcium increases at GABAergic terminals during the induction step, via the recruitment of calcium-dependent protein kinases and phosphatases. In the PFC, activation of D₂Rs may increase interneuronal excitability and thus spontaneous inhibitory activity (Retaux et al., 1991; Grobin and Deutch, 1998). Although this effect is not seen by all (Gorelova and Yang, 2000; Seamans et al., 2001; Tseng and O'Donnell, 2007), it is formally possible that quinpirole may indirectly facilitate I-LTD in the PFC by enhancing the activity of interneurons whose terminals express CB₁Rs. We examined whether sIPSC frequency increased in our hands after 500 nm quinpirole application but found no difference (20.8 ± 4.0 Hz before vs 17.7 ± 2.8 Hz after, p > 0.05, n = 9). Thus, an increase in spontaneous inhibitory activity cannot account for the facilitatory effect of quinpirole on eCB signaling.

Endogenous DA facilitates I-LTD when catechol-O-methyltransferase is inhibited

Thus far, we have activated D₂Rs by applying the specific receptor agonist quinpirole. We next sought to determine whether endogenous DA can also facilitate eCB signaling. It is possible that DA deafferentation that is inherent to the slicing procedure prevented the 5 Hz train from triggering I-LTD in the absence of quinpirole. To compensate for a reduction of DA levels that is highly expected in PFC slices, we pharmacologically blocked DA catabolism and tested whether synaptic stimulation will trigger I-LTD under this recording condition (Fig. 7). The major mammalian enzymes involved in the degradation of DA are catechol-O-methyltransferase (COMT) and monoamine oxidases, with relative contribution tilted toward COMT in the PFC (Karoum et al., 1994; Matsumoto et al., 2003; Yavich et al., 2007). Thus, to effectively increase DA levels in PFC slices, we pharmacologically blocked COMT activity with the inhibitor OR 486 (2 μm). The 5 Hz train failed to trigger I-LTD in the presence of OR 486 alone (100.4 ± 21.1% of baseline, n = 4) (supplemental Fig. S1, available at www.jneurosci.org as supplemental material). Because DA can activate both D₁Rs and D₂Rs, with potentially opposing effects (Cepeda et al., 1993; Nishi et al., 1997; Seamans et al., 2001; Trantham-Davidson et al., 2004; Tseng and O'Donnell, 2004; Xu et al., 2009), an effect by D₂Rs may potentially be masked by a conflicting D₁R effect, depending on which receptor was more strongly activated. Therefore, we next tested I-LTD by coapplication of OR 486 with the D₁R antagonist SCH23390 (10 μm). Indeed, in the presence of OR 486 and SCH23390, I-LTD could be triggered by the 5 Hz synaptic stimulation (IPSC amplitude, 49.6 ± 4.6% of baseline, n = 10) (Fig. 7A). Consistent with a reduction in probability of GABA release, PPR in I-LTD experiments increased from 0.82 ± 0.03 to 0.92 ± 0.04 (p < 0.05, paired t test). A role for D₂Rs was confirmed by the ability of sulpiride to reduce I-LTD in the presence of OR 486 (88.5 ± 8.4% of baseline, n = 8, p < 0.005) (Fig. 7A). As was observed for the quinpirole-facilitated form, I-LTD in OR 486 and SCH23390 was also sensitive to CB₁R blockade with AM 251 (93.0 ± 9.8% of baseline, n = 6, p < 0.01) (Fig. 7B) and was not present in the CB₁R knock-out mice (knock-out, 92.2 ± 4.8% of baseline, n = 7 vs WT, 70.7 ± 3.8% of baseline, n = 7, p < 0.005) (Fig. 7C). Consistent with a role for PKA in D₂R- and CB₁R-mediated effects on GABAergic transmission in the PFC, I-LTD in the presence of OR 486 and SCH23390 was reduced in H89 (91.0 ± 5.7% of baseline, n = 6, p < 0.005) (Fig. 7D). Thus, together, our results strongly suggest that endogenous DA can activate D₂Rs in the presynaptic GABAergic terminal to enhance eCB signaling and trigger I-LTD.

Figure 7.

Increasing endogenous DA levels by inhibiting COMT also enables I-LTD. A, The 5 Hz stimulation in the presence of the COMT inhibitor OR 486 and the D₁R antagonist SCH23390 elicited I-LTD (black circles) that was sensitive to D₂R antagonist sulpiride (white circles). Top left, Representative average traces from single experiments obtained at the time points indicated. Top right, Plot of PPR before and after delivery of 5 Hz train, showing a change in PPR. B, This I-LTD (black circles) was also sensitive to the CB₁R antagonist AM 251 (white circles). Top, Representative average traces from single experiments obtained at time points indicated. C, In the wild-type mouse (CB₁^+/+), a weaker 5 Hz train (for 5 min) triggers I-LTD in the presence of OR 486 and SCH23390. This I-LTD is not present in the CB₁ knock-out littermates (CB₁^−/−). Top, Representative average traces from single experiments obtained at the time points indicated. D, Time course of I-LTD in the presence of OR 486 and SCH23390 in control (black circles) and in slices bath applied with H89 (white circles). Top, Representative average IPSC traces from single control and H89 block experiments in the rat PFC obtained at the time points indicated. L, Layer.

Discussion

In this study, we combine anatomical and electrophysiological techniques to investigate the cellular mechanisms underlying the link between DA and eCB function in the PFC. Using EM, we first show immunolabeling of D₂Rs and CB₁Rs at terminals of symmetrical presumably GABAergic synapses in the PFC. Notably, quantitative analysis reveal a robust pattern of CB₁R and D₂R colocalization at these presynaptic sites. Consistent with this localization, activation of either receptor suppresses GABA release as indicated by an increase in PPR. Importantly, repetitive afferent stimulation in the presence of a D₂R agonist induces eCB-dependent I-LTD that is insensitive to interference of postsynaptic D₂R signaling. Together, our results strongly suggest a synergistic interaction between colocalized D₂Rs and CB₁Rs in regulating long-term depression of inhibitory transmission in the PFC.

Previous studies have shown that activation of CB₁Rs in neocortex (Trettel and Levine, 2002; Bodor et al., 2005; Hill et al., 2007; Galarreta et al., 2008) or D₂Rs in prefrontal cortex (Retaux et al., 1991; Seamans et al., 2001) can suppress inhibitory transmission, presumably by reducing GABA release. Our combined anatomical and physiological data support similar roles of CB₁Rs and D₂Rs on GABAergic transmission in the PFC. Furthermore, in keeping with a recent report in the nucleus accumbens (Pickel et al., 2006), we have found that D₂Rs and CB₁Rs can colocalize to sites presynaptic to symmetrical synapses in the PFC; this colocalization may provide an anatomical substrate for receptor interaction when coactivated. Coactivation of CB₁Rs and D₂Rs can have several potential effects on signal transduction of the respective individual receptors. In heterologous systems and striatal cultures, pharmacological coactivation of coexpressed CB₁Rs and D₂Rs has been shown to alter G-protein signaling, converting G_i/o-mediated pathways to G_s-coupled ones (Glass and Felder, 1997; Kearn et al., 2005), thereby enhancing rather than reducing transmitter release. There is also the possibility of synergistic or antagonistic crosstalk between the receptor systems in the presynapse (Kearn et al., 2005; Marcellino et al., 2008).

We have tested for a potential interaction between the eCB and DA systems in the PFC and found that nominal activation of D₂Rs with a low dose of quinpirole can facilitate eCB signaling at GABAergic synapses to trigger I-LTD. D₂Rs most likely act by enhancing a process downstream of eCB mobilization from the postsynaptic cell because blocking postsynaptic PKA activity has no effect on I-LTD, whereas global block of PKA in the slice does. D₂Rs may signal via pathways other than PKA. In the striatum, activation of D₂Rs has been shown to lead to stimulation of phospholipase Cβ isoforms and mobilization of intracellular calcium stores (Nishi et al., 1997; Hernandez-Lopez et al., 2000). However, two lines of evidence argue against a role for postsynaptic D₂Rs in facilitating I-LTD in the PFC. First, quinpirole suppression of IPSCs is completely insensitive to AM 251, indicating that D₂R activation is insufficient to trigger eCB production in the PFC. Second, intracellular calcium rise is not required for I-LTD because loading the postsynaptic cell with BAPTA has no effect on plasticity. Because both CB₁R and D₂R signaling can lead to inhibition of the PKA pathway, we postulate that coactivation of CB₁Rs and D₂Rs enables I-LTD in the PFC by cooperatively lowering PKA activity below a threshold level as supported by our observation of an enhancement of quinpirole-mediated suppression by a submaximal concentration of WIN (Fig. 5A). A similar mechanism has been suggested recently for I-LTD in the VTA (Pan et al., 2008). The precise molecular PKA target underlying I-LTD remains to be identified.

It is widely accepted that DA modulates PFC function. Here, we show that DA may influence synaptic transmission and plasticity in the PFC by facilitating eCB signaling via presynaptic D₂Rs. Interestingly, our results using the COMT inhibitor OR 486 alone, in the absence of the D₁R antagonist SCH23390 (supplemental Fig. S1, available at www.jneurosci.org as supplemental material), suggest that D₁R activation may antagonize D₂R-facilitated I-LTD. This raises the question of whether DA likely facilitates I-LTD in the intact brain. However, some evidence suggests that D₁Rs and D₂Rs may be selectively activated in vivo. D₂Rs show comparatively higher affinity to DA than D₁Rs (Creese et al., 1983), and thus D₂R activation may require lower concentrations of DA. It has been proposed, at least in the nucleus accumbens, that basal DA release continuously activate D₂Rs, whereas phasic DA release is needed to elevate DA levels enough to stimulate D₁Rs (Grace, 1991). Moreover, tonic versus phasic DA transmission can be differentially evoked by distinct sets of afferents to DA neurons (Goto and Grace, 2005). It is possible that D₂Rs may be preferentially activated during tonic DA release conditions in the PFC as well, making GABAergic synapses primed for eCB signaling and I-LTD.

It is unclear how activation of D₁Rs opposes PFC I-LTD in our experiments; D₁Rs may directly antagonize downstream D₂R/CB₁R signaling in the GABAergic terminal and/or mediate I-LTP at the same or different GABAergic synapse. We have shown with EM that a high proportion of presynaptic D₂Rs are colocalized with CB₁Rs (Fig. 1), but the extent of overlap between presynaptic D₁Rs and CB₁Rs remains unknown. Furthermore, given the high expression of postsynaptic D₁Rs in the PFC (Ariano and Sibley, 1994; Smiley et al., 1994; Lidow et al., 2003), it is possible that these receptors mask I-LTD by mediating a potentiation of GABAergic transmission via an increase in number or conductance of GABA_A receptors as has been shown for D₁R and AMPAR (Sun et al., 2005).

In the PFC, eCBs have been shown to depress excitatory transmission (Lafourcade et al., 2007), and we now demonstrate that inhibitory transmission is also under eCB regulation. What, then, is the overall effect of eCBs on the balance of excitation and inhibition in the PFC and how is PFC output modulated? The answers to these questions may lie in the differential induction of E-LTD and I-LTD. It is interesting to note that eCB-mediated E-LTD in the PFC can be triggered by a slightly stronger synaptic stimulation train (e.g., 10 vs 5 Hz) and does not require DA transmission. Furthermore, postsynaptic calcium rise is necessary for E-LTD but not I-LTD, suggesting that multiple mechanisms underlying eCB production may exist. Future studies will be needed to examine whether DA can modulate this form of E-LTD. Interestingly, a form of E-LTD, induced by brief high-frequency (50 Hz) stimulation, has been described previously in the PFC to depend on DA (Otani et al., 1998); the involvement of eCB signaling in this E-LTD is yet to be determined. Another provocative question is whether synaptic activity can simultaneously trigger both eCB-mediated E-LTD and I-LTD. If so, it will be important to assess changes in the firing properties of the output layer 5 pyramidal cells in PFC under conditions of intact excitation and inhibition.

In humans, cannabis consumption has dramatic effects on PFC-mediated function and can trigger schizophrenia-like states in normal individuals, exacerbate psychotic symptoms in schizophrenic patients, and increase the risk of developing schizophrenia in predisposed individuals (Ujike and Morita, 2004; Koethe et al., 2009; Sewell et al., 2009). In addition, studies using CB₁R-deficient mice have revealed an important role of eCBs in the extinction of learned behaviors, such as fear (Marsicano et al., 2002; Varvel and Lichtman, 2002), which are partly mediated by the PFC (Morgan et al., 1993; Morrow et al., 1999). It is currently unknown how eCBs and cannabinoid agonists can lead to these effects. The involvement of DA in the central actions of cannabinoids has been suggested (van der Stelt and Di Marzo, 2003; Laviolette and Grace, 2006). Cannabinergic signaling may lead to DA release (Cadogan et al., 1997; van der Stelt and Di Marzo, 2003) and can modulate D₂R agonist-induced behavior (Beltramo et al., 2000; Gorriti et al., 2005). Indeed, microdialysis measurements have revealed an increase of DA in the PFC after in vivo administration of the cannabinoid Δ(9)-tetrahydrocannabinol (Pistis et al., 2002). Notably, a decrease of GABA in the PFC was also observed in this study. Given that an imbalance of excitatory and inhibitory transmission in the PFC has been proposed to contribute to schizophrenia (Lewis et al., 2005; Gonzalez-Burgos and Lewis, 2008), these changes in neurotransmitter levels provide insight into how schizophrenic symptoms may emerge after cannabis consumption. We propose that a synergic interaction between DA and eCB function at the synaptic level in the PFC (i.e., to trigger I-LTD) may play a role in schizophrenia in predisposed cannabis users.