Excitability of prefrontal cortical pyramidal neurons is modulated by activation of intracellular type-2 cannabinoid receptors

Abstract

The endocannabinoid (eCB) system is widely expressed throughout the central nervous system (CNS) and the functionality of type-1 cannabinoid receptors in neurons is well documented. In contrast, there is little knowledge about type-2 cannabinoid receptors (CB₂Rs) in the CNS. Here, we show that CB₂Rs are located intracellularly in layer II/III pyramidal cells of the rodent medial prefrontal cortex (mPFC) and that their activation results in IP₃R-dependent opening of Ca²⁺-activated Cl⁻ channels. To investigate the functional role of CB₂R activation, we induced neuronal firing and observed a CB₂R-mediated reduction in firing frequency. The description of this unique CB₂R-mediated signaling pathway, controlling neuronal excitability, broadens our knowledge of the influence of the eCB system on brain function.

The endocannabinoid (eCB) system is involved in many functions of the CNS, including executive functions associated with the prefrontal cortex, such as decision-making and working memory (1). The eCB system consists of at least two G protein-coupled receptors (GPCRs), type-1 cannabinoid receptor (CB₁R) and type-2 cannabinoid receptor (CB₂R), lipid endogenous ligands (e.g., anandamide and 2-arachidonoylglycerol), and various enzymes responsible for the synthesis and degradation of the endogenous ligands (2–6). CB₁Rs are among the most abundantly expressed GPCRs in the rat brain and their role, predominantly as presynaptic receptors, in modulating neurotransmission is clearly established (5, 7, 8). In contrast with CB₁R, the presence and function of CB₂R in the brain has long been a matter of debate (9). CB₂Rs are found primarily in the immune system and were initially regarded as the “peripheral” cannabinoid receptor (10, 11). This generally accepted idea is challenged by the description of CNS CB₂R gene expression in rats and wild-type mice (12–14) and the identification of functional CB₂Rs on glial cells and neurons (15–18). In addition to the current view that supports the expression of functional CB₂Rs in neurons upon brain stress or damage (19), it has been reported that CB₂Rs could play a role in general CNS physiology (20–22). These developments emphasize the importance of understanding how CB₂R activation affects neuronal functioning. To demonstrate the presence of functional CB₂Rs in the rodent medial prefrontal cortex (mPFC) and to elucidate their functional role, we used Western blotting, a radioactive binding assay, and electrophysiological techniques (whole-cell current and voltage clamp) on layer II/III pyramidal neurons.

Results

Functional CB₂Rs in the mPFC.

The presence of CB₂Rs in the rat mPFC was demonstrated by a Western blot performed on homogenated mPFC samples (Fig. 1A). A band of the expected molecular weight for CB₂R was detected, which was absent when the primary antibody was incubated with immunizing peptide (Fig. 1A). Immunoblots on tissues from spleen and brainstem (positive controls) and skeletal muscle tissue (negative controls) confirmed the specificity of the CB₂R immunodetection (Fig. 1A).

Fig. 1.

CB₂Rs are present in the rodent mPFC, and their activation evokes a delayed depolarization. (A) Western blot detection of CB₂Rs in the rat mPFC (Upper). Primary antibody preincubated with immunogen peptide showed no CB₂R bands (Lower). Positive controls: spleen (Spl) and brainstem (Brs); negative control: skeletal muscle (SkM). (B) Bath-applied JWH-133 (1 μM, arrow indicates the start of the application) induced a delayed (10 min) depolarization of a layer II/III pyramidal neuron. (C) Bath application of 1 and 5 μM JWH-133 induced similar depolarizations (ΔV = 29 ± 3 mV and 25 ± 2 mV, both P < 0.001). After preincubation with 5 μM Sch.356036, application of 1 μM JWH-133 resulted in a significantly reduced depolarization (**P < 0.01, Mann–Whitney U test). JWH-133 (1 μM) depolarized neurons of Wt mice, but not of CB₂R KO mice (ΔV = 15 ± 4 mV and 0 ± 0.3 mV; **P < 0.01). Local application of JWH-133 (5 μM) with a Picospritzer device and introduction of JWH-133 (5 μM) into neurons via the patch pipette also depolarized neurons (ΔV = 38 ± 4 mV and 22 ± 6 mV; P < 0.0001 and P < 0.05). Sch.356036 (1 μM), introduced into neurons via the patch pipette, largely prevented the depolarization by JWH-133 (1 μM, ΔV = 4 ± 2 mV; ***P < 0.001). (D) The delays of onset of depolarization after bath application of JWH-133 (1 and 5 μM) and local application of JWH-133 (5 μM) were 855 ± 177 s, 490 ± 76 s, and 529 ± 125 s, respectively. Introduction of JWH-133 (5 μM) via the patch pipette reduced the delay of onset of the effect compared with the other application methods with 5 μM JWH-133 (103 ± 50 s; both **P < 0.01, Mann–Whitney U tests). Numbers of observations are indicated in bars (C and D); error bars represent SEM.

To investigate whether CB₂R activation can modulate ion conductances, we performed whole-cell current clamp recordings in layer II/III pyramidal neurons of the mPFC at an experimental membrane potential (Vm = −80 mV) that differed from all experimental ion equilibrium potentials. We observed that bath applications of the selective CB₂R agonist JWH-133 (1 μM and 5 μM) resulted in a transient depolarization in most (86%) layer II/III pyramidal neurons of the mPFC, after a delay that lasted several minutes (Fig. 1 B and C). The depolarization induced by bath application of 1 μM JWH-133 was significantly smaller after preincubation of the slice with a selective CB₂R antagonist, Sch.356036 (5 μM) (Fig. 1C). We confirmed the pharmacological evidence that the depolarization is mediated by CB₂R activation by performing similar experiments in CB₂R knockout (KO) and wild-type (Wt) mice. We did not observe a delayed depolarizing response in mPFC neurons from KO mice, whereas in Wt mice such responses could be evoked (Fig. 1C).

To investigate the origin of the depolarization delay, we repeated the experiments but replaced the bath application by fast local pressure ejection of 5 μM JWH-133 onto the soma of patched layer II/III pyramidal neurons. Because the depolarization delay was hardly affected (Fig. 1 C and D), we concluded that it did not originate from diffusion limitations in the bath application method. We hypothesized that CB₂Rs are located intracellularly in these neurons and that the depolarization delay most likely originates from the time required for the lipophilic ligands to pass the plasma membrane and to reach the intracellularly located target. To test this hypothesis, we introduced 5 μM JWH-133 into the cell via the patch pipette and we still observed a depolarization (Fig. 1C), but with a much reduced delay (≈1.5 min) after going whole-cell (Fig. 1D). Introduction of the antagonist Sch.356036 (1 μM) into the cell via the patch pipette largely prevented the membrane depolarization induced by bath application of 1 μM JWH-133 (Fig. 1C). When the antagonist Sch.356036 was bath applied, 5 μM was needed to antagonize the JWH-133 (1 μM) effect (Fig. 1C). Taken together, the combined electrophysiological and pharmacological experiments suggest an intracellular localization of the CB₂R.

CB₂Rs Are Intracellularly Localized in the mPFC.

To obtain further evidence for an intracellular localization for CB₂Rs, we performed additional Western blot experiments. Whole mPFC samples were fractionated into a plasma membrane and an intracellular fraction. The identity of the fractions was established by using antibodies against the plasma membrane marker Na⁺/K⁺-ATPase and the intracellular marker nucleoporin. The Western blot on fractionated samples showed that CB₂Rs are abundantly present in the intracellular fraction, whereas they are hardly detectable in the plasma membrane fraction (Fig. 2A). Experiments using a radioactive binding assay on similarly fractionated mPFC samples corroborated the intracellular localization of CB₂Rs (Fig. 2B). Intracellular binding of [³H]CP55.940 (a mixed CB₁R/CB₂R agonist) was reduced by ≈50% in the presence of the selective CB₂R antagonist SR2 (SR144528), but not when the selective CB₁R antagonist SR1 (SR141716) was present. Incubating the plasma membrane fraction with SR1 or SR2 reduced the binding of [³H]CP55.940. However, this reduction in binding was much larger in the presence of SR1 (≈70% reduction) than in the presence of SR2 (≈20% reduction). These results confirm that CB₂R binding sites are predominantly located intracellularly in the rat mPFC, whereas CB₁R binding sites appear to be mainly present in the plasma membrane. In additional fluorescence imaging experiments on a neuronal cell line (human SH-SY5Y neuroblastoma cells transiently transfected with GFP-tagged CB₂Rs), we demonstrated that CB₂Rs are almost exclusively localized in intracellular membranous structures and that they are not present on the plasma membrane (Fig. S1 and Table S1).

Fig. 2.

CB₂Rs are localized intracellularly in the rat mPFC. (A) Western blot on subcellular mPFC fractions, consisting of an intracellular fraction (Intr) and a plasma membrane fraction (PM), demonstrated a predominantly intracellular localization of CB₂R. Membranes were probed with antibodies against CB₂R, the intracellular marker nucleoporin, and the plasma membrane marker Na⁺/K⁺-ATPase. (B) Radioactive binding experiments show that intracellular binding of the mixed CB₁R/CB₂R agonist [³H]CP55.940 (CP; 46 ± 3 fmol/mg) was reduced in the presence of SR2 (a CB₂R antagonist; 24 ± 1 fmol/mg; **P < 0.01) and not in the presence of SR1 (a CB₁R antagonist; 44 ± 3 fmol/mg). Plasma membrane binding of [³H]CP55.940 (126 ± 6 fmol/mg) was reduced after incubation with SR1 (36 ± 1 fmol/mg; ***P < 0.001) or SR2 (102 ± 4 fmol/mg; *P < 0.05). The reduction of plasma membrane binding was significantly larger in the presence of SR1 compared with SR2 (71 ± 1% and 19 ± 4%, respectively; P < 0.001). Numbers of observations are indicated in bars; error bars represent SEM.

CB₂R Activation Opens Ca²⁺-Activated Cl⁻ Channels via IP₃R.

The signaling cascade after CB₂R activation can lead, through phospolipase C production, to Ca²⁺ release via IP₃R (23) and, thus, to the potential activation of Ca²⁺-activated conductances (24). Introduction of the fast Ca²⁺ chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid (BAPTA) at a high concentration (10 mM) into the cell via the patch pipette, strongly reduced the amplitude of the delayed depolarization (Fig. 3A). Bath application of the IP₃R blocker 2-aminoethyl diphenylborinate (2-APB, 0.1 mM) also reduced the depolarization evoked by JWH-133 (Fig. 3A). These results indicate that a rise in [Ca²⁺]_i after IP₃R activation is necessary for the CB₂R-mediated depolarization. A series of voltage clamp experiments was performed to determine which conductance underlies the CB₂R-mediated depolarization. Application of JWH-133 (1 μM) decreased the input resistance (R_input; Fig. 3B), implying that the signal transduction pathway after CB₂R activation opens ion channels in the plasma membrane. Using a series of step potentials before and during the JWH-133 application allowed the construction of a current-voltage (I/V) relationship that reversed near the calculated reversal potential (E_rev) for Cl⁻ (E_Cl-) (Fig. 3C). Blocking K⁺, Ca²⁺, and Na⁺ channels with 4-aminopyridine (4-AP), tetraethylammonium-Cl (TEACl), Cd²⁺, and tetrodotoxin (TTX) hardly had an effect on the magnitude and the reversal potential of the currents evoked by JWH-133 (Fig. 3D). In addition, decreasing [Cl⁻]_i to 4.95 mM shifted E_rev toward the newly established E_Cl- (Fig. 3D). We observed similar currents when we used a different selective CB₂R agonist, HU-308 (1 μM) (Fig. 3D, Inset). Finally, the JWH-133–evoked currents were markedly reduced in the presence of the Cl⁻ channel blocker 4,4′-diisothiocyanatostilbene-2,2′-disulfonic acid (DIDS) (Fig. 3E). In summary, these results show that CB₂R activation leads to the opening of Ca²⁺-activated Cl⁻ channels (CaCCs) that may contribute to the control of the membrane potential of layer II/III pyramidal neurons of the rat mPFC.

Fig. 3.

CB₂R activation evokes an IP₃R-dependent opening of Ca²⁺-activated Cl⁻ channels. (A) In the presence of either 10 mM BAPTA in the patch pipette or 0.1 mM 2-APB in the bath, 5 μM or 1 μM JWH-133 evoked significantly reduced (both ***P < 0.001) depolarizations (ΔV = 4 ± 1 mV and 8 ± 2 mV). The average response evoked with 5 μM JWH-133 (in the absence of BAPTA) is the same as is depicted in Fig. 1C. (B) The input resistance (R_input) was reduced by JWH-133 (182 ± 20 MΩ and 128 ± 25 MΩ; *P < 0.05). (C) I/V relationship of CB₂R-mediated currents (evoked with 1 μM JWH-133) that reverse at −40 ± 2mV, close to E_Cl- (−38.3 mV); Inset shows representative current traces. (D) In the presence of 4-AP, TEACl, Cd²⁺, and TTX (to block K⁺, Ca²⁺, and Na⁺ channels) and recorded with [Cl⁻]_i = 30 or 4.95 mM, E_rev of CB₂R-mediated currents (−26 ± 2 mV and −76 ± 4 mV) followed E_Cl- (−32.6 mV and -80 mV). (D Inset) Another selective CB₂R agonist, HU-308 (1 μM), evoked similar currents (E_rev: −31 ± 2 mV, E_Cl-: −32.6 mV). (E) The Cl⁻ channel blocker DIDS (0.2 mM) reduced the amplitude of CB₂R-mediated currents. Numbers of observations are indicated in bars/brackets; error bars represent SEM.

CB₂R Activation Reduces Neuronal Excitability.

The hypothesis that the CB₂R-mediated signaling pathway contributes to neuronal excitability was further investigated in a series of current clamp experiments under physiological Cl⁻ conditions (E_Cl- = −70 mV). A “slow” automatic feedback system ensured that recordings always started at a membrane potential of −70 mV. Neuronal firing was evoked by Gaussian current input into the soma, via the patch pipette, that evoked fluctuations around resting membrane potential. For each neuron, the variance of the input signal was adjusted to cause a stable spiking rate (≈0.85 Hz). Application of 1 μM JWH-133 reduced the firing rate by 45%, and this reduction could be prevented by preincubation (of at least 10 min) with the CB₂R antagonist, Sch.356036 (5 μM) (Fig. 4). In a separate experiment, we tested the effect of Sch.356036 by itself on the firing rate. Application of 5 μM Sch.356036 increased the firing rate (mean baseline firing frequency of 0.86 ± 0.09 Hz was normalized over slices to 100 ± 8% and increased to 121 ± 8% in the presence of Sch.356036, P < 0.05, n = 12). These results indicate an endogenous tonus of eCBs and/or constitutive activity of the receptor and show that, when firing is evoked with an input that could resemble spontaneous background synaptic activity, further CB₂R activation modulates the firing rate of mPFC neurons.

Fig. 4.

CB₂R activation decreases firing activity of rat mPFC layer II/III pyramidal neurons. (A) Application of 1 μM JWH-133 led to a reduction of neuronal firing rate. Firing was induced by Gaussian current input into the soma. Mean baseline firing frequency of 0.88 ± 0.10 Hz was normalized over slices to 100 ± 3% and reduced to 55 ± 6% in the presence of 1 μM JWH-133 (***P < 0.001). After preincubation (of at least 10 min before going whole-cell) with and continuous presence of 5 μM Sch.356036, baseline firing frequency 0.83 ± 0.15 Hz, normalized to 100 ± 3%, JWH-133 (1 μM) could not induce a response (92 ± 6%, JWH-133 plus Sch.356036). (B) Representative traces of current clamp recordings showing action potential firing of layer II/III neurons in the absence (baseline) and presence of 1 μM JWH-133 and in the presence of 5 μM Sch.356036 (Sch.356036 baseline) and 1 μM JWH-133 plus 5 μM Sch.356036.

Discussion

Our results provide evidence for functional neuronal CB₂Rs that are located intracellularly, in the rodent mPFC. Their activation results, via IP₃R activation, in the opening of CaCCs. Furthermore, this opening of CaCCs is most likely responsible for the observed reduction in neuronal excitability upon CB₂R activation. The presence of CB₂R mRNA in the brain and CB₂Rs in neurons of the brainstem, the cerebellum, and the hippocampus has been reported (12, 13, 18, 25–28), but their presence in the cortex remained to be further characterized. The current view on the presence of functional CB₂Rs in the CNS supports the expression of CB₂Rs in neurons essentially upon brain stress and damage (19). Commonplace problems with the visualization of CB₂Rs by immunohistochemical stainings have been described. These problems may arise from the difficulty of producing reliable antibodies against CB₂Rs, slight differences in diaminobenzidine (DAB) staining protocols, species-specific isoform expression patterns and complications with negative controls such as CB₂R-KO (9, 14, 28, 29). Therefore, we used a combination of biochemical (Western blotting and radioactive binding assay) and functional (in vitro electrophysiology and pharmacology) techniques to provide evidence that functional CB₂Rs are expressed intracellularly in cortical neurons of the healthy brain. The results of the radioactive binding assay did not depend on the quality of antibodies and provided additional support for the intracellular presence of CB₂Rs in the mPFC where this technique confirmed that CB₁Rs are primarily located in the plasma membrane. In addition, by means of fluorescence imaging, we could demonstrate that in a neuronal cell line (human neuroblastoma cells; SI Materials and Methods) CB₂Rs are predominantly located in intracellular membranous structures and not in the plasma membrane. In accordance with our results, a few publications support the idea that, like other functional GPCRs (30, 31), functional CB₂Rs may have an intracellular localization. A report by Currie et al. (32) shows the intracellular localization of functional CB₂Rs in guinea pig heart cells. Others report that CB₂Rs are associated with the rough endoplasmic reticulum and Golgi apparatus in hippocampal pyramidal neurons, but in these studies, the functional role of intracellular CB₂Rs was not investigated (25, 26, 28).

The CB₂R signaling pathway shows great diversity and complexity (33) and has not been fully elucidated in neurons. The main downstream signaling pathway of CB₂R activation involves Gi/o and the subsequent modulation of adenylate cyclase and MAP kinase activity (34), but coupling to the modulation of [Ca²⁺]_i has also been reported (33). In calf pulmonary endothelial cells, activation of CB₂Rs by anandamide resulted in an increase in [Ca²⁺]_i, mediated by PLC activation and IP₃ production (23). In cardiac cells (32) and adult dorsal root ganglion neurons (35), CB₂R activation was reported to be negatively coupled to IP₃R-mediated Ca²⁺ release. In addition to the known CB₂R-mediated pathways, we demonstrate that in rodent layer II/III cortical pyramidal neurons, the CB₂R signaling cascade involves IP₃R activation and results in the opening of CaCCs. CaCCs are known to control excitability in various types of peripheral and central neurons (24, 36), but have not been described in pyramidal neurons of the mPFC.

Although some publications report a functional role for CB₂Rs in the CNS (20–22), the precise mode of action is not known. In this study, we demonstrate that CB₂R activation—under physiological conditions—leads to a decrease in neuronal firing rate, probably via the opening of CaCCs. If E_Cl- is close to the resting membrane potential of cortical neurons, the activation of CB₂Rs will stabilize or even clamp the membrane potential around that level. This reduction of neuronal excitability upon CB₂R activation is reminiscent of shunting inhibition, described for GABA_A receptors (37, 38), that also mediate a Cl⁻ conductance. Furthermore, we observed that the application of a CB₂R antagonist slightly increased the firing rate, indicative of a certain eCB tonus and/or a constitutive activity of the receptor. These results are in line with both the emerging idea that the eCB system consists of a basal and an “on demand” pool of endocannabinoids and with the possibility of constitutively active CB₂Rs (39–41).

The mode of action we report here could be the underlying mechanism involved in the reduction of firing activity by JWH-133 in wide-range dorsal horn neurons in a rat model of acute, inflammatory, and neuropathic pain (20) and in thalamic neurons in a rat model of neuropathic pain (42).

The unique aspects of eCB signaling we describe in this study, uncover a modulatory role for the eCB system in mPFC function, through the regulation of neuronal excitability by CB₂R activation. Because high frequency stimulation leads to eCB synthesis (43), activation of CB₂Rs after increased neuronal activity may prevent excessive neuronal firing via an intracellularly organized feedback system. Through this mechanism, CB₂Rs could play a protective role in the brain (44). More generally, the differential (sub)cellular localization of CB₁Rs and CB₂Rs and their downstream pathways diversify the response repertoire of the neuronal eCB system beyond the generally accepted modulation of neurotransmission processes.

Materials and Methods

Western Blotting.

Experiments were approved by the animal welfare committee of the University of Amsterdam. mPFC, brainstem, spleen, and skeletal muscle of 14-d-old male Wistar rats (Harlan) were rapidly dissected out in ice-cold homogenization buffer (0.05 mM PBS) with 320 mM sucrose and protease inhibitor mixture (Complete, pH 7.4; Roche) and homogenized with a glass douncer. For subcellular fractionation of the mPFC, the same buffer was used. The homogenate was centrifuged at 800 × g for 10 min to discard undisrupted tissue. The supernatant was centrifuged at 100,000 × g for 1 h to separate plasma membrane fractions (pellet) from intracellular fractions (supernatant). Equal amounts of protein, 10 or 20 μg, were loaded and separated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) on 10% Tris-glycine gel and transferred onto nitrocellulose membrane. The membranes were washed in Tris-buffered saline (TBS) and blocked in TBS with 1% (vol/vol) Tween (TBST) containing 4% (wt/vol) nonfat milk and incubated overnight at 4 °C with antibodies against CB₂R (Abcam; 1:800) alone or in combination with antibodies against the plasma membrane marker Na⁺/K⁺-ATPase α3 (Santa Cruz Biotechnology; 1:500) and against the nuclear envelope marker nucleoporin p62 (Beckton Dickinson; 1:800). In some cases, the CB₂R antibody was preincubated with the immunogen peptide (Abcam; 1:40). Membranes were washed with TBST and incubated for 1 h with HRP-conjugated goat anti-rabbit antibody (Bio-Rad; 1:3,000), goat anti-mouse antibody (Bio-Rad, 1:3,000), and donkey anti-goat antibody (Santa Cruz Biotechnology; 1:5,000) and processed for immunoreactivities using enhanced chemiluminesence (ECL) Plus Western Blotting detection reagents (Amersham). Bands were visualized with Hyperfilm ECl (Amersham) or the Odyssey Infrared Imaging System (Licor).

Radioactive Binding Assay.

CB₁R and CB₂R binding was assessed by rapid filtration assays, using 400 pM [³H]CP55.940 as reported (45). mPFC tissue of 14-d-old male Wistar rats (Harlan) was used in rapid filtration assays, after subcellular fractionation as described for the Western blot samples in 2 mM Tris⋅EDTA, 320 mM sucrose, and 5 mM MgCl₂ at pH 7.4. Unspecific binding was determined in the presence of “cold” agonist (1 μM CP55.940) and was further corroborated by selective antagonists [0.1 μM SR141716 (SR1) for CB₁R or 0.1 μM SR144528 (SR2) for CB₂R]. Binding data were expressed as femtomole ligand bound per milligram of protein.

Electrophysiology.

Coronal slices (300 μm) of the mPFC were obtained from male Wistar rats (Harlan) aged 14–19 d postnatal and male C57BL/6 Wt mice or male C57BL/6 CB₂R KO mice (The Jackson Laboratory) aged 14–19 d postnatal. Animals were killed by decapitation, and their brains rapidly were removed and placed in oxygenated (95% O₂–5% CO₂) ice cold (4 °C) adapted artificial cerebrospinal fluid (aaCSF: 120 mM choline chloride, 3.5 mM KCl, 0.5 mM CaCl₂, 6 mM MgSO₄, 1.25 mM NaH₂PO₄, 25 mM d-glucose, and 25 mM NaHCO₃). Slices were cut in aaCSF on a vibratome (VT1200S; Leica) and placed for 30 min in aCSF (120 mM NaCl, 3.5 mM KCl, 25 mM NaHCO₃, 25 mM d-glucose, 2.5 mM CaCl₂, 1.3 mM MgSO₄, and 1.25 mM NaH₂PO₄; [Cl⁻]_out = 128.5 mM) at 32 °C. Slices were kept at room temperature for at least 1 h before recording. Glass recording pipettes were pulled from borosilicate glass (Science Products) and had a resistance of 2–3 MΩ when filled with pipette solution (110 mM KGluconate, 30 mM KCl, 0.5 mM EGTA, 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (Hepes), 4 mM Mg-ATP, and 0.5 mM Na-GTP). For local application of JWH-133, a Picospritzer II device (General Valve) was used to deliver the compound to the recorded neurons with brief pressure pulses (500 ms). Modified aCSF with K⁺-, Ca²⁺-, and Na⁺-channel blockers contained 70 mM NaCl, 3.5 mM KCl, 25 mM NaHCO₃, 25 mM d-glucose, 2.5 mM CaCl₂, 1.3 mM MgSO₄, 1.25 mM NaH₂PO₄, 25 mM TEACl, 5 mM 4-AP, 0.2 mM CdCl₂, and 0.0005 mM TTX; [Cl⁻]_out = 103.7 mM. Modified pipette solution with 30 mM [Cl⁻]_i contained 110 mM CH₃O₃SCs, 30 mM CsCl, 0.5 mM EGTA, 10 mM Hepes, 4 mM Mg-ATP, and 0.5 mM Na-GTP). Modified pipette solution with 4.95 mM [Cl⁻]_i contained: 135.05 mM CH₃O₃SCs, 4.95 mM CsCl, 0.5 mM EGTA, 10 mM Hepes, 4 mM Mg-ATP, and 0.5 mM Na-GTP. Introduction of CB₂R ligands into neurons was achieved by backfilling the recording pipettes. Whole-cell current and voltage clamp recordings were made at 32 °C from the soma of layer II/III pyramidal neurons. In the whole-cell current clamp configuration, we used a slow feedback system that guaranteed that current clamp recordings started at a membrane voltage of −80 mV (Figs. 1 and 3) or −70 mV (Fig. 4). In the whole-cell voltage clamp configuration with [Cl⁻]_i = 30 mM, currents were evoked, every 30 s until JWH-133–mediated currents were recorded, by a series of rectangular voltage steps (200-ms duration) from a holding potential of −80 mV to voltage potentials ranging from −90 to −20 mV in 10-mV increments. For experiments with [Cl⁻]_i = 4.95 mM, the currents were evoked from a holding potential of −50 mV. Input resistance was calculated from current responses to hyperpolarizing steps of −5 mV. Frozen filtered Gaussian noise (time constant = 10 ms) was injected via the patch pipette with a variance adjusted for each neuron to result in a mean spike frequency (≈0.85 Hz). The firing frequency was calculated by using 1-min bins. Before drug application (or at least 10 min after preincubation with Sch.356036), 5-min recordings were used as control. Drug effects were determined 3–15 min after application. Recordings were made by using an EPC9 patch-clamp amplifier controlled by PULSE software (HEKA Electronic) and in-house software running under Matlab (MathWorks). Signals were filtered at 2.9 kHz and sampled at 10 kHz. Series resistance ranged from 5 to 15 MΩ and was compensated to ≈65%. Signals were corrected for liquid junction potential. Current densities were calculated by using cell capacitance and expressed in pA/pF.

Data Analysis.

Data were statistically tested with paired and unpaired Student's t tests unless otherwise stated. In the figures, the significance is indicated with asterisks (*P < 0.05, **P < 0.01, and ***P < 0.001).

Drugs.

For the radioactive binding assay, the synthetic cannabinoid CP55.940 {5-(1,10-dimethyheptyl)-2-[1R,5R-hydroxy-2R-(3-hydroxypropyl)-cyclohexyl] phenol} was purchased from Sigma Chemical. [³H]CP55.940 (126 Ci/mmol) was from PerkinElmer Life Sciences. SR141716 [N-piperidino-5-(4-chlorophenyl)-1-(2,4-dichlorophenyl)-4-methyl-3-pyrazole-carboxamide], and SR144528 {N-[(1S)-endo-1,3,3-trimethy-1-bicyclo[2.2.1]-heptan-2-yl]5-(4-choro-3-methyl-phenyl)-1-(4-methyl-benzyl)-pyrazole-3-carboxamide} were kind gifts from Sanofi-Aventis Recherche (Paris, France). For the electrophysiological experiments, JWH-133, HU-308, and Sch.356036 were generous gifts from Abbott Laboratories (Weesp, the Netherlands). Cannabinoid receptor ligands were dissolved in DMSO to 50 mM and diluted in aCSF that never contained a final concentration of DMSO higher than 0.1%. BAPTA, 2-APB, and DIDS were all purchased from Sigma-Aldrich. TTX (0.5 μM, Latoxan) was present during all recordings, with the exception of the experiments shown in Fig. 4.