Neuropharmacology and analgesia

Effects of cannabidiol on the function of α7-nicotinicacetylcholine receptors

Mohamed Mahgoub a, Susan Yang Keun-Hang b, Vadym Sydorenko a,c, Abrar Ashoor a,Nadine Kabbani d, Lina Al Kury a, Bassem Sadek a, Christopher F. Howarth e, Dmytro Isaev a,c,Sehamuddin Galadari f, Murat Oz a,n

a Laboratory of Functional Lipidomics, Department of Pharmacology, College of Medicine and Health Sciences, UAE University, Abu Dhabi, Al Ain,United Arab Emiratesb Department of Biological Sciences, Schmid College of Science and Technology, Chapman University, One University Drive, Orange, CA 92866, USAc Department of General Physiology of Nervous System, Bogomoletz Institute of Physiology, Bogomoletz Str, Kiev 01024, Ukrained Neuroscience Program, George Mason University, 4400 University Drive, Fairfax, VA 22030, USAe Laboratory of Functional Lipidomics, Department of Physiology, College of Medicine and Health Sciences, UAE University, Al Ain, United Arab Emiratesf Laboratory of Functional Lipidomics, Department of Biochemistry, College of Medicine and Health Sciences, UAE University, Al Ain, United Arab Emirates

a r t i c l e i n f o

Article history:Received 15 June 2013Received in revised form16 September 2013Accepted 7 October 2013Available online 18 October 2013

Keywords:Nicotinic receptorsCannabidiolCannabinoidsXenopus oocyte

a b s t r a c t

The effects of cannabidiol (CBD), a non-psychoactive ingredient of cannabis plant, on the function of thecloned α7 subunit of the human nicotinic acetylcholine (α7 nACh) receptor expressed in Xenopus oocyteswere tested using the two-electrode voltage-clamp technique. CBD reversibly inhibited ACh (100 μM)-induced currents with an IC50 value of 11.3 mM. Other phytocannabinoids such as cannabinol andΔ9-tetrahydrocannabinol did not affect ACh-induced currents. CBD inhibition was not altered bypertussis toxin treatment. In addition, CBD did not change GTP-γ-S binding to the membranes of oocytesinjected with α7 nACh receptor cRNA. The effect of CBD was not dependent on the membrane potential.CBD (10 mM) did not affect the activity of endogenous Ca2þ-dependent Cl� channels, since the extent ofinhibition by CBD was unaltered by intracellular injection of the Ca2þ chelator BAPTA and perfusion withCa2þ-free bathing solution containing 2 mM Ba2þ . Inhibition by CBD was not reversed by increasing AChconcentrations. Furthermore, specific binding of [125I] α-bungarotoxin was not inhibited by CBD (10 mM)in oocytes membranes. Using whole cell patch clamp technique in CA1 stratum radiatum interneurons ofrat hippocampal slices, currents induced by choline, a selective-agonist of α7-receptor induced currentswere also recoded. Bath application of CBD (10 mM) for 10 min caused a significant inhibition of cholineinduced currents. Finally, in hippocampal slices, [3H] norepinephrine release evoked by nicotine (30 mM)was also inhibited by 10 mM CBD. Our results indicate that CBD inhibits the function of the α7-nAChreceptor.

& 2013 Elsevier B.V. All rights reserved.

1. Introduction

Cannabidiol (CBD) is one of the most abundant cannabinoidsfound in Cannabis sativa constituting up to 40% of cannabisextracts (for a review, Mechoulam and Hanus, 2002). It wasreported to have antioxidant, anti-inflammatory and antiemeticeffects (Izzo et al., 2009). Recent in vitro and in vivo studiesindicate that CBD can be used in the treatment of various

pathological conditions such as epilepsy, glaucoma, anxiety andcancer (Izzo et al., 2009; Pertwee, 2010; Hill et al., 2012).Importantly, CBD is a devoid of psychoactive properties due to alow affinity for the CB1 and CB2 receptors (Mechoulam et al.,2007; Izzo et al., 2009; Pertwee, 2009). Thus, pharmaceuticalinterest in this compound has risen significantly in recent years(for reviews, Izzo et al., 2009; Pertwee, 2009; Scuderi et al., 2009).

Nicotinic acetylcholine (nACh) receptors are important mem-bers of the ligand-gated ion channel family and are divided intotwo groups: muscle receptors, which are found at the skeletalneuromuscular junction where they mediate neuromusculartransmission and neuronal receptors, which are found throughoutthe peripheral and central nervous systems where they areinvolved in fast synaptic transmission and in the modulation oftransmitter release (for reviews, Hogg et al., 2003; Albuquerque

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European Journal of Pharmacology

0014-2999/$ - see front matter & 2013 Elsevier B.V. All rights reserved.http://dx.doi.org/10.1016/j.ejphar.2013.10.011

Abbreviations: ACh, acetylcholine; ANOVA, analysis of variance; BAPTA, 1,2-bis(o-aminophenoxy) ethane-N, N,N′,N′-tetraacetic acid; DMSO, dimethyl sulfoxide;HEPES, 4-(2-hydroxyethyl) piperazineethane sulfonic acid; MBS, modified Barth'ssolution; PTX, pertussis toxin.

n Corresponding author. Tel.: þ971 3 713 7523; fax: þ971 3 767 2033.E-mail address: Murat_Oz@uaeu.ac.ae (M. Oz).

European Journal of Pharmacology 720 (2013) 310–319

et al., 2009). α7-nACh receptors are widely distributed throughoutthe cortical, thalamic and hippocampal region in the centranervous system (Albuquerque et al., 2009). Importantly, α7-nAChreceptors which have considerably high permeability to Ca2þ havebeen shown to be located on both glutamatergic and GABAergicnerve terminals suggesting that both excitatory and inhibitorysynaptic transmissions can be modulated by the activity of thesereceptors (Hogg et al., 2003; Albuquerque et al., 2009).

The effects of endocannabinoids such as N-arachidonoy-lethanolamine and 2-arachidonoylglycerol (Oz et al., 2003,2004c; Baranowska et al., 2008) and synthetic cannabinoid recep-tor agonists such as WIN55, 212-2, and CP55, 940 (Oz et al., 2004c)on the functional properties of α7-nACh receptors have beenshown in earlier in vitro and in vivo studies. However, the effectof non-psychotropic cannabinoids such as CBD on the α7-nACh-receptor function remains unknown. We investigated the effects ofCBD on human α7-nACh receptors expressed in Xenopus oocytesand on α7-nACh receptor-mediated responses in rat hippocampalneurons. In this study, we provide novel evidence on a functionalinteraction between CBD and α7-nACh receptor.

2. Materials and methods

2.1. Recordings from oocytes

Mature female Xenopus laevis frogs were purchased fromXenopus Express (Haute-Loire, France) and housed in dechlori-nated tap water at 19–21 1C with a 12/12-h light/dark cycle and fedwith food pellets supplied by Xenopus Express. The proceduresfollowed in this study were in accordance with the Guide for theCare and Use of Laboratory Animals of the National Institutes ofHealth (Bethesda, MD) and approved by the Institutional AnimalCare and Use Committee at the College of Medicine and HealthSciences, UAEU. Clusters of oocytes were removed surgically underbenzocaine (Sigma, St. Louis, MO) local anesthesia (0.03%, w/v) andindividual oocytes were dissected away manually in a solutioncontaining (in mM): NaCl, 88; KCl, 1; NaHCO3, 2.4; MgSO4, 0.8;HEPES, 10 (pH 7.5). Dissected oocytes were then stored for 2–7days in modified Barth's solution (MBS) containing (in mM): NaCl,88; KCl, 1; NaHCO3, 2.4; CaCl2, 2; MgSO4, 0.8; HEPES, 10 (pH 7.5),supplemented with 2 mM, sodium pyruvate, 10,000 IU/L penicillin,10 mg/L streptomycin, 50 mg/L gentamicin, and 0.5 mM theophyl-line. Briefly, oocytes were placed in a 0.2 mL recording chamberand superfused at a rate of 2–3 mL/min. Under these conditions,solution exchange time was less than 100 ms. The bathing solutioncontained (in mM): NaCl, 96; KCl, 2; CaCl2, 1.8; MgCl2, 1; HEPES, 5(pH 7.5). The cells were impaled with two glass microelectrodesfilled with a 3 M KCl (0.5–2 MΩ). The oocytes were routinelyvoltage clamped at a holding potential of �70 mV using aGeneClamp-500 amplifier (Axon Instruments Inc., Burlingame,CA) and current responses were recorded and stored digitally forfurther analysis. Oocyte capacitance was measured by a paired-ramp method described earlier (Oz et al., 2004a). Briefly, voltage-ramps were employed to elicit constant capacitive current (Icap)and the charge associated with this current was calculated by theintegration of Icap. Ramps had slopes of 2 V/s and durations of20 ms and started at a holding potential of �90 mV. A series of 10paired-ramps was delivered at 1 s intervals and averaged traceswere used for charge calculations. In each oocyte, the averages of5–6 measurements were used to obtain values for membranecapacitance (Cm) Currents for Icap recordings were filtered at 20 kHzand sampled at 50 kHz.

Drugs were applied by gravity flow via a micropipette posi-tioned about 2 mm from the oocyte. Some of the compounds wereapplied externally by the addition to the superfusate (flow rate of

3–4 mL/min). CBD, CBN and THC were generously provided byNIDA Drug Supply/NIH, Rockville, MD, USA. Acetylcholine, PNU-282987, methyllycaconitine, α-bungarotoxin and all chemicalsused were obtained from Sigma (St. Louis, MO). Procedures forthe injections of BAPTA (50–100 nl, 100 mM) were performed asdescribed previously (Oz et al., 1998). BAPTA was prepared inCs4-BAPTA and injections were performed 1 h prior to recordingsusing an oil-driven ultra microsyringe pump (Micro4, WPI, Inc.Sarasota, FL). Stock solutions of CBD used in this study wereprepared in ethanol at a concentration of 30 mM. At the highestfinal concentrations used (0.1%, v/v), ethanol did not have asignificant effect on ACh (100 mM)-induced currents (n¼8).

The cDNA clone of human α7-nACh receptor was kindlyprovided by Dr. J. Lindstrom (University of Pennsylvania, PA).Capped cRNA transcripts were synthesized in vitro using amMESSAGE mMACHINE kit from Ambion (Austin, TX) and ana-lyzed on 1.2% formaldehyde agarose gel to check the size and thequality of the transcripts. Approximately 3–5 ng of cRNA wasinjected into each oocyte.

2.2. Radioligand binding studies

Oocytes were injected with 5 ng human α7-nACh receptorcRNA, and the functional expression of the receptors was testedby electrophysiology on day three. Isolation of oocyte membraneswas carried out by modification of a method described earlier (Ozet al., 2004b). Briefly, oocytes (200–300 oocytes per assay) weresuspended (approximately 20 μl/oocyte) in a homogenizationbuffer containing (in mM) HEPES 10, EDTA 1, PMSF 0.1, and0.02% NaN3, 50 μg/mL bacitracin (pH 7.4) at 4 1C on ice andhomogenized using a motorized Teflon homogenizer (six strokes,15 s each at high speed). The homogenate was centrifuged for10 min at 800 � g. The supernatant was collected and the pelletwas resuspended in homogenization buffer and recentrifuged at800 � g for 10 min. Supernatants were then combined and cen-trifuged for 1 h at 36,000 � g. The membrane pellet was resus-pended in homogenization buffer and used for the binding studies.

Binding assays were performed in 500 μL of 10 mM HEPES(pH 7.4) containing 50 μL of oocyte preparation and 0.1–5 nM [125I]α-bungarotoxin (2200 Ci/mmol; Perkin-Elmer, Inc. Waltham, MA).Non-specific binding was determined using 10 μM α-bungarotoxin.Oocyte membranes were incubated with [125I] α-bungarotoxin inthe absence and presence of drugs, for 1 h at room temperature(22–24 1C). The radioligand was separated by rapid filtration ontoGF/C filters presoaked in 0.2% polyethyleneimine. Filters were thenwashed with two 5 mL washes of ice-cold HEPES buffer, and theradioactivity was determined by counting samples in a BeckmanGamma-300 γ-counter.

2.3. [125I] α-bungarotoxin binding assays in intact oocytes

Two to three days after injection, [125I] α-bungarotoxin bindingassays were performed subsequent to the voltage-clamp measure-ment on the same intact oocytes and only those with a currentresponse (to 100 mM acetylcholine) of more than 2000 nA wereselected for the binding assay. Binding assays in single intactoocytes were carried out by modification of a method describedearlier (Fenster et al., 1999). Briefly, oocytes were incubated in20 nM [125I] α-bungarotoxin, 5 mg/mL BSA, MBS at room tempera-ture for 2 h. Non-injected oocytes were incubated under the sameconditions to measure non-specific binding. The excess of toxinwas removed by washing each oocyte with 25 mL of MBS. Radio-activity was measured using a Beckman Gamma-300 γ-counter.Counts per minute (cpm) values were calculated from the meansof four separate experiments. In each experiment, 5–8 oocyteswere used for each group.

M. Mahgoub et al. / European Journal of Pharmacology 720 (2013) 310–319 311

2.4. [35S]GTPγS binding

Oocyte membranes for [35S]GTPγS binding experiments wereprepared as described earlier (Lipinsky and Oron, 1996). Onehundred and fifty oocytes were gently homogenized at 4 1C bypassing through a 27 G needle in 4 mL of homogenization mediumcontaining (in mM): NaCl, 5; Na-HEPES, 5; PMSF, 0.6; leupeptin,4 pM, pH 7.5. The homogenate was centrifuged for 5 s in a conicalcentrifuge tube at 8000 � g. The supernatant was decanted fromthe precipitated cellular organelles and the procedure repeated.Following the aspiration of the lipid layer at the surface of thesupernatant, it was centrifuged for 20 min at 8000 � g. The pelletwas resuspended by re-homogenization in 1.8 mL of bindingmedium containing (in mM): KC1 80, Na-HEPES 20, MgC1 1, PMSF1.7, and leupeptin 0.013, pH 7.5. Each tube contained (in 25 μlvolume) the following: membrane preparation equivalent to1 oocyte and the following additions (in mM): KCl 45, Na-HEPES11, MgCl 0.5, PMSF 1.0, leupeptin 7.3 pM, pH¼7.5 and [35S]GTPγS0.5 nM–10 pM. [35S]GTPγS (1000–1500 Ci/mmol) was from NewEngland Nuclear (Boston, MA). Non-specific binding was assayedby adding 0.5 mM non-radioactive GTPγS and 50 μM ATP. Themixture was incubated at room temperature (22 1C) for 30 minand filtered rapidly through Whatman (Clifton, NJ) GFIC 25 mmfilters and washed with 5 mL of ice-cold wash solution (in mM:KC1 80, Na-HEPES 20, MgC1 1, pH 7.5). The filters, once washed,were subjected to scintillation counting.

2.5. Nicotine-induced norepinephrine release

The release of norepinephrine (NE) was assessed by modifica-tions of a method described earlier (Ashoor et al., 2011). Briefly,male Sprague–Dawley rats (10–30 days old) were decapitated, thebrains were rapidly removed and hippocampi were dissected.Coronal hippocampal slices (400 μm, 3–5 mg) were incubated inKrebs' buffer containing (in mM) 118 NaCl, 4.7 KCl, 1.2 MgCl2,2.5 CaCl2, 1.0 NaH2PO4, 11.1 glucose, 25 NaHCO3, 0.11 L-ascorbicacid and 4.0 disodium ethylenediamine tetraacetate; pH 7.4, andsaturated with 95% O2/5% CO2) at 34 1C for 30 min. Subsequently,slices were incubated in fresh buffer (5–6 slices/2 mL) containing[3H]NE (0.1 μM, final concentration) for an additional 30 min. Afterrinsing in fresh buffer, each slice was transferred to 1 of 8 Plexiglaschambers maintained at 34 1C and superfused (1 mL/min) withoxygenated Krebs' buffer containing the monoamine oxidaseinhibitor pargyline (10 μM), to prevent metabolism of NE. Sliceswere superfused initially for a 45-min period and then three5-min samples (5 mL/sample) were collected to determine basal[3H]NE outflow. In a separate series of experiments, inhibitionproduced by cannabinoids on nicotine-evoked [3H]NE overflowwas determined. After collection of the third basal sample, eachhippocampal slice was superfused in the absence (control) orpresence of CBD, and superfusate samples were collected every5 min for 30 min. Subsequently, nicotine (30 μM) was added to thebuffer, and slices were superfused for 30 min and samples col-lected every 5 min. A control slice in each experiment was super-fused for 30 min with buffer in the absence of inhibitor, followedby superfusion with nicotine (30 μM) to determine [3H]NE over-flow. At the end of each experiment, slices were solubilized withTS-2 tissue solubilizer (RPI Corp, Mt. Prospect, IL), and the [3H]-content of the tissue and samples was determined using a liquidscintillation counter.

Fractional release for each superfusate sample was calculatedby dividing the amount of tritium in each 5-min sample by thetotal tissue-[3H] at the time of sample collection. Basal [3H]outflow was calculated as the average fractional release in thethree samples just before addition of the inhibitor to the super-fusion buffer. Total [3H] overflow was calculated by summing the

increases in fractional release above basal [3H] outflow resultingfrom exposure to nicotine, either in the absence or presence ofinhibitor and subtracting [3H] outflow for an equivalent period ofinhibitor exposure.

2.6. Whole-cell patch clamp recordings in rat hippocampal slices

Male Sprague–Dawley rats (10–30 days old) were killed by CO2

narcosis followed by decapitation. Their brains were removed in icecold artificial cerebrospinal fluid (ACSF), which was composed of(in mM): NaCl, 125; NaHCO3, 25; KCl, 2.5; NaH2PO4, 1.25; CaCl2, 2,MgCl2, 1; glucose, 25. Slices of 350–400 mm thickness were cut usinga Vibratome and stored at room temperature in an immersionchamber containing ACSF bubbled with 95% O2 and 5% CO2. Whole-cell patch-clamp experiments were performed on visually identifiedCA1 stratum raditum (SR) interneurons of rat hippocampal slices(Isaev et al., 2007; Singhal et al., 2007). Whole-cell currents wererecorded from the soma of SR interneurons according to thestandard patch-clamp technique using an Axopatch 200B amplifier(Axon Instruments, Foster City, CA). The slices were superfused withACSF at 2 mL/min in the presence of the tetrodotoxin (1 mM), themuscarinic antagonist atropine (0.5 mM), the 6-cyano-7-nitroqui-noxaline-2,3-dione (CNQX, 10 mM), the 2-amino-5-phosphonovalericacid (APV, 10 mM), and the bicuculline (10 mM). Choline (1 mM) wasapplied every 2 min to interneuron cell bodies via a quartz tube (i.d.,300 mm) positioned about 150–200 mm from the cell; drug deliverywas controlled by a computer-driven valve system. Other drugswere applied via bath superfusion. Signals were filtered at 5 kHz andrecorded by a microcomputer using the pCLAMP8.1 program (AxonInstruments, Foster City, CA). Patch pipettes were pulled fromborosilicate glass capillary (1.2-mm outer diameter), and when filledwith internal solution had resistance between 3 and 5 MΩ. At�70 mV holding potential, the leak current was generally between100 and 250 pA, and when it exceeded 250 pA, the data were notincluded in the analysis. The internal pipette solution contained (inmM): ethylene-glycol bis(b-amino-ethyl ether)-N-N0-tetraaceticacid, 10; HEPES, 10; Cs-methane sulfonate, 130; CsCl, 10; MgCl2, 2;lidocaine N-ethyl bromide (QX-314), 5 (pH adjusted to 7.2 withCsOH; 330 mΩ). All recordings were done at room temperature (20–22 1C). Only a single neuron was studied in a given slice, therefore,the number of neurons represents the number of hippocampal slicesanalyzed.

2.7. Data analysis

Average values were calculated as the mean7standard errormeans (S.E.M.). Statistical significance was analyzed using Stu-dent's t test or ANOVA as indicated. Two-way ANOVA was used toanalyze the effect of the inhibitors on fractional [3H]NE release.Concentration–response curves were obtained by fitting the datato the logistic equation,

y¼ Emax=ð1þ½x=EC50��nÞ;where x and y are concentration and response, respectively, Emax isthe maximal response, EC50 is the half-maximal concentration,and n is the slope factor (apparent Hill coefficient).

3. Results

At the highest concentration used in this study, bath applica-tion of neither ACh (1 mM) nor CBD (100 mM) produced detectablecurrents in oocytes injected with distilled water (30 nl per oocyte,n¼8). Application of 100 μM ACh for 3–4 s activated fast inwardcurrents that desensitized rapidly in oocytes injected with cRNAtranscribed from cDNA encoding the α7-subunit of human nACh

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receptor. Moreover, ACh-induced inward currents were abolishedcompletely with 100 nM α-bungarotoxin (n¼7, data not shown),further indicating that these responses are mediated by the α7-nACh receptors.

In the first set of experiments, the effect of cannabidiol (CBD)was tested on ACh (100 mM)-induced ion currents. An effectof 10 min incubation with CBD (10 μM) on α7-nACh receptormediated currents are shown in Fig. 1A. Time-courses of effectsof CBD and the vehicle applications on the amplitudes of ACh-induced currents are presented in Fig. 1B. CBD (10 μM) caused asignificant inhibition of the current which was partially reversedduring 10 to 15 min washout period. In the absence of CBD, vehicle(0.1% ethanol) alone did not alter the amplitude of the ACh-induced current, suggesting that CBD selectively acts on nAChreceptors in these cells. (Fig. 1B, controls versus CBD treatmentgroup at 10 min of exposure, ANOVA, n¼5–7; Po0.05).

The inhibitory effect of CBD was significantly dependent on theapplication mode. For example, without CBD preincubation, coap-plication of CBD (10 μM) and ACh (100 μM) did not alter theamplitudes of maximal currents (Fig. 1C). However, when oocyteswere preincubated with CBD, the drug was found to inhibitmaximal ACh-induced currents in a time dependent mannerreaching a maximal level within 10–15 min (with a half-time (τ1/2) of 2.170.3 min; Fig. 1C). Since the magnitude of the CBD effectwas time-dependent, 10–15 min CBD application time was usedroutinely to ensure equilibrium conditions. CBD was found toinhibit the function of α7-nACh receptor in a concentration-dependent manner with IC50 and slope value of 11.371.8 mMand 1.270.1, respectively (Fig. 1D).

The participation of Gi/o-proteins in the signaling of nAChreceptors and activation via cannabinoids and certain CBD analogs

has been reported (for reviews Sharir and Abood, 2010; Nordmanand Kabbani, 2012). Furthermore the relatively slow time course ofCBD action shown in Fig. 1B and C also suggests an involvement ofG proteins and/or G protein coupled receptors. We tested theeffect of CBD in control (distilled-water injected) and pertussistoxin (PTX)-injected oocytes expressing α7-nACh receptors. Therewas no significant difference in CBD inhibition of ACh responsesbetween controls and PTX-injected cells (Fig. 2A; P¼0.644;F¼0.223; ANOVA, n¼7–8) suggesting a G alpha i/o independentmechanism of channel regulation in these cells.

In addition, we investigated the effect of 30 μM CBD on thespecific binding of [35S]GTPγS in oocyte membranes. Equilibriumcurves for the binding of [35S]GTPγS, in the presence and absenceof the CBD are shown in Fig. 2B (n¼6–8). Cannabidiol, at aconcentration of 30 μM, did not significantly change the specificbinding of [35S]GTPγS. Maximum binding activities (Bmax) of [35S]GTPγS were 4.7570.37 and 4.5470.51 pmol/oocyte (means7S.E.M.) for controls and CBD, respectively. The apparent affinity(KD) of the receptor for [35S]GTPγS was 1.0870.15 and0.9670.17 μM for controls and CBD, respectively. There were nostatistically significant differences between control and CBD trea-ted groups with respect to both KD (ANOVA, F¼0.11; P¼0.74,n¼7–9) and the Bmax values (ANOVA, F¼0.31; P¼0.59, n¼7–9).

Activation of α7-nACh receptors allows sufficient Ca2þ entry toactivate endogenous Ca2þ-dependent Cl� channels in Xenopusoocytes (Sands et al., 1993; Uteshev, 2012). Therefore, it wasimportant to determine whether the effect of CBD was exertedon nACh receptor mediated currents or on Cl� currents induced byCa2þ entry. For this reason, extracellular Ca2þ was replaced withBa2þ since Ba2þ can pass through α7-nACh receptors but causeslittle, if any, activation of Ca2þ-dependent Cl� channels (Sands

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Fig. 1. Effects of cannabidiol (CBD) on α7-nicotinic acetylcholine receptor-mediated ion currents. (A) Records of currents activated by acetylcholine (ACh, 100 μM) in controlconditions (left), during coapplication of 10 μM CBD and ACh after 10 min pretreatment with 10 μM CBD (middle), and 15 min following CBD washout (right). (B) Time-course of the effect of vehicle (0.3% ethanol; filled circles) and CBD (10 μM; open circles) on the peaks of the ACh-induced currents. Each data point represents the normalizedmean7S.E.M. of 5–7 experiments. Duration of drug application is indicated by the horizontal bar in the figure. (C) Effect of pre-application time on the CBD inhibition ofα7-nACh receptor. Inhibition of the α7-nACh receptor increases with the prolongation of CBD pre-application time. Each data point represents the mean7S.E.M. of5–6 oocytes. (D) Cannabidiol inhibits α7-nACh receptor function in a concentration-dependent manner. Each data point represents the mean7S.E.M. of 5–6 oocytes. Thecurve is the best fit of the data to the logistic equation described in Section 2.

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et al., 1993). Since CBD has been shown to increase intracellularCa2þ levels in neurons and glia (Drysdale et al., 2006; Ryan et al.,2006) we have also injected the Ca2þ chelator BAPTA into oocytes.Under these conditions, we tested the effect of CBD in a solutioncontaining 2 mM Ba2þ in BAPTA-injected oocytes. CBD (10 mM)produced the same level of inhibition (4375 in controls versus4174 in BAPTA-injected oocytes; ANOVA, F¼ 0.21, P¼0.66; n¼6–7)

on ACh-induced currents when BAPTA-injected oocytes wererecorded in Ca2þ free solutions containing 2 mM Ba2þ (Fig. 2C). Atthis point, it is important to mention that in the oocyte expressionsystem, CBD-induced changes in intracellular Ca2þ levels can bedetected by Ca2þ-activated Cl� channels and concomitant alterationsin the holding currents and membrane input resistance. However, incontrol experiments, CBD (30 mM for 15 min) did not alter themagnitudes of holding-currents in oocytes voltage-clamped at�70mV (n¼12), indicating that intracellular Ca2þ levels were notaltered by these drugs. Similarly, in controls and in the presence ofCBD (30 mM), there were no statistically significant differencesbetween the means of membrane resistance (Rm) membrane capaci-tance (Cm) and resting membrane potentials (Vm) values were1.470.3 and 1.270.3 MΩ, 254717 and 241719 nF, and �3874and �3673 mV, respectively (n¼7–9, ANOVA, P40.05). Thus, ourdata indicate that CBD does not cause a significant effect on passivemembrane properties of oocytes.

CBD has been shown to modulate functional properties of T-type Ca2þ channels in a voltage-dependent manner (Ross et al.,2008). We examined the effect of membrane potential on CBD-inhibition of α7-nACh receptors. Each tested membrane potentialwas held for 20–30 s and then returned to holding potential of�70 mV. As indicated in Fig. 3A, the inhibition of ACh (100 mM)-induced currents by CBD (10 mM) does not appear to be voltage-dependent. The extent of CBD inhibition was similar at all testedmembrane potentials from �120 to �20 mV. Evaluation of thecurrent–voltage relationship (Fig. 3B) indicates that the extent ofinhibition by CBD does not change significantly at differentholding potentials (P40.05, n¼6, ANOVA).

An open-channel blockade, by definition, requires the openingof the channel by the binding of the agonist to the receptor. Thus,in the absence of an agonist, the degree of blockade should berelated to the frequency of channel opening. Therefore, the extentof CBD inhibition of the nACh receptor was compared in oocytesexposed to ACh at 5-min intervals with those exposed at 10- and20-min intervals (Fig. 3C). Application of CBD (10 μM for 20 min)was equally effective in inhibiting the currents activated at 5-,10- and 20-min intervals (Fig. 3D on the left; between 5, 10, and20 min interval groups, P¼0.814, F¼0.207, n¼5–6 for eachgroup, ANOVA), indicating that the frequency of the channelopening does not alter the extent of CBD inhibition and that thechannel does not need to be opened by the agonist for CBD to beeffective. Similarly, recoveries from an open channel blockerwould be facilitated by increases in opening frequency. We findthat recovery from CBD effect was not altered by ACh stimulationintervals, suggesting that CBD is not trapped in the channel whenthe channel closes, as can occur with open channel blockers(Fig. 3D; between 5, 10, and 20 min interval groups P¼0.892,F¼0.115, n¼5–6 for each group, ANOVA).

CBD may decrease the binding of the agonist to the receptor byacting as a competitive antagonist. For this reason, we determinedthe effect of CBD at different ACh concentrations. Concentration–response curves for ACh in the absence and presence of 30 mM CBDare presented in Fig. 4A. In the presence of CBD, the maximalresponse induced by ACh decreased significantly (n¼6–8). How-ever, the potency of the ACh (EC50) remained unaltered. In theabsence and presence of ACh, the EC50 values were 79.1722.6 μMand 83.4720.8 μM (Po0.05, ANOVA, n¼5–6) and slope valueswere 2.170.2 and 2.370.3, respectively, suggesting that CBDinhibits the ACh responses in a non-competitive manner.

[125I] α-bungarotoxin competes with ACh, an endogenousactivator of α7-nACh receptors by binding to ACh binding site onthe receptor (Albuquerque et al., 2009). For this reason, the effectof 10 μM CBD was investigated on the specific binding of [125I]α-bungarotoxin. Equilibrium curves for the binding of [125I]α-bungarotoxin, in the presence and absence (controls) of CBD

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Fig. 2. Effects of cannabidiol on nACh receptors of either pertussis toxin treated orBAPTA injected oocytes and on [35S]GTPγS binding of oocytes membranes. (A) Barpresentation of the effects of 10 mM CBD application (15 min) on the maximalamplitudes of ACh induced currents in oocytes injected with 50 nl distilled-water,controls (n¼6) or 50 nl of PTX (50 mg/mL, n¼7). Bars represent the means7S.E.M.(B) The effect of 30 μM CBD on [35S]GTPγS binding to oocyte membrane prepara-tion. Membranes were incubated with different concentrations of [35S]GTPγS for20 min at room temperature, and the results analyzed as described in Section 2.Data points for controls and CBD are indicated by filled and open circles,respectively (n¼6–8). (C) Bar presentation of the effects of 10 mM CBD application(20 min) on the maximal amplitudes of ACh induced currents in oocytes injectedwith 50 nl distilled-water, controls (n¼5) or BAPTA (50 nl, 200 mM, n¼7). Barsrepresent the means7S.E.M. of 6–8 experiments. The number of experiment ispresented on top of each bar. There was no statistically significant difference incannabidiol (10 μM) inhibition in the presence or in the absence of BAPTAinjections (P40.05, n¼6–7, ANOVA).

M. Mahgoub et al. / European Journal of Pharmacology 720 (2013) 310–319314

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Fig. 3. The effects of change in membrane potential and acetylcholine application interval on cannabidiol inhibition of nicotinic acetylcholine receptors. (A) Current–voltagerelationships of acetylcholine-activated currents in the absence and presence of cannabidiol (CBD, 10 μM). Normalized currents activated by 100 μM ACh before (control, filledcircles) and after 15 min treatment with 10 μM CBD (open circles). Each data point presents the normalized means and S.E.M. of 5–6 experiments. (B) Quantitative evaluation ofthe effect of cannabidiol as percent inhibition at different voltages. (C) Time-course of the effect of cannabidiol on the maximal amplitudes of the currents induced by 100 mMAChat 5 (open circles), 10 (filled circles), and 20 (filled triangles) min intervals. Data points represent means7S.E.M of 6–8 cells. (D) The percentage of CBD inhibition of the ACh-induced currents recorded at the end of a 20-min application period was not different among oocytes stimulated with ACh application at 5, 10, and 20 min intervals (P40.05,ANOVA). The percentage recovery from CBD inhibition of the ACh receptor-mediated currents recorded at the end of a 20-min recovery period was not different between oocytesstimulated with ACh every 5 min (P40.05, ANOVA) and non-stimulated controls. Duration of CBD application is indicated by the horizontal bar.

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Fig. 4. Concentration–response curves for acetylcholine-induced currents and binding of [125I] α-bungarotoxin in control and in the presence cannabidiol. (A) Effect of cannabidiol(CBD) on the acetylcholine (ACh) concentration–response relationship. Oocytes were voltage-clamped at �70mV and currents were activated by applying acetylcholine (1 μM to3mM). Oocytes were exposed to 10 mM CBD for 15 min and ACh was reapplied. Paired concentration–response curves were constructed and responses normalized to maximalresponse under control conditions. EC50 and slope values were determined by fitting the curves from 6 to 8 oocytes to the standard logistic equation as described in the methodssection. Data points obtained before (control) and after 15 min treatment with cannabidiol (10 μM) were indicated by filled and open circles, respectively. Each data point presents thenormalized means and S.E.M. of 6–8 experiments. (B) The effects of cannabidiol on the specific binding of [125I] α-bungarotoxin to oocyte membrane preparations. In the presence andabsence of cannabidiol, specific binding as a function of the concentration of [125I] α-bungarotoxin is presented. Data points for controls and cannabidiol (10 μM) are indicated by filledand open circles, respectively. Data points are the means of three independent experiments carried out in triplicate. (C) Effects of increasing concentration of cannabidiol on the specificbinding of [125I] α-bungarotoxin. Experiments were conducted in the presence of 1 nM of [125I] α-bungarotoxin. The results present data from 8 to 11 measurements. Data pointsindicate mean7S.E.M. (D) Effect of cannabidiol on the specific binding of [125I] α-bungarotoxin in single, intact oocytes expressing α7-nACh receptors.

M. Mahgoub et al. / European Journal of Pharmacology 720 (2013) 310–319 315

are presented in Fig. 4B. At a concentration of 30 μM, CBD didnot cause a significant inhibition of the specific binding of [125I]α-bungarotoxin. Maximum binding activities (Bmax) of [125I]α-bungarotoxin were 2.4870.19 and 2.6370.32 pM/mg (means7S.E.M.) for controls and CBD-treated preparations, respectively(Fig. 4B). The apparent affinity (KD) of the receptor for [125I]α-bungarotoxin was 0.5570.14 and 0.6770.22 pM for controlsand CBD, respectively. There was no statistically significant differ-ence between controls and CBD-treated groups with respect to KD

and Bmax values (P40.05, ANOVA, n¼5–6). In another set ofexperiments, the effect of increasing CBD concentrations on thespecific binding of [125I] α-bungarotoxin (1 nM) was tested (Fig. 4C).In this concentration range (1–100 mM), CBD did not cause asignificant alteration in [125I] α-bungarotoxin binding (P40.05,ANOVA, n¼8–11) suggesting that CBD does not compete witha-bungarotoxin at the same binding site.

The radioligand binding assays from oocyte membranes involvea disruption in cell integrity. It is possible that these subcellularfractions contain intracellular membranes, and therefore, themeasured binding results are confounded by inconsistent phar-macological effects. We have performed radioligand bindingassays in intact oocytes. CBD (30 μM) did not cause a significantinhibition of the specific binding of [125I] α-bungarotoxin (20 nM)in oocytes injected with the α7-nACh receptor cRNA. Specificbinding of [125I] α-bungarotoxin was 21937321 cpm and23477339 cpm (means7S.E.M.) for controls and CBD (30 μM)-treated, respectively. There was no statistically significant differ-ence detected between controls and CBD-treated groups (Fig. 4D;Po0.05, ANOVA, n¼17–19) using this assay.

Cannabidiol, within the concentration range used in this study, hasbeen reported to inhibit 5-HT3 receptors another member of cyc-loopfamily of ion channels with structural similarities to α7-nACh receptor(Yang et al., 2010; Xiong et al., 2011). In these studies, the extent ofCBD inhibition of 5-HT3 receptors has been shown to inverselycorrelate to the density of 5-HT-induced currents in oocytes (Yanget al., 2010; Xiong et al., 2011). We examined if the extent of CBDactions on the α7-nACh receptor shows a correlation with the densityof ACh-induced currents in the same cell. In Fig. 5A, the extent of CBDinhibition is plotted as a function of maximal currents induced by100 mM ACh. The results indicate that there is no correlation betweenmaximal current amplitudes and CBD inhibition (n¼36; r¼0.32).

We also compared the effects of CBD with other phytocannabi-noids such as cannabinol (CBN) and Δ9-tetrahydrocannabinol (THC) onthe function of α7-nACh receptors. As shown in Fig. 5B, neither CBNnor THC (10 mM) appeared to affect the maximal amplitude of ACh(100 mM)-induced currents in oocytes expressing α7-nACh receptors.These findings confirm a selective function of CBD on α7-nAChreceptor.

We have also tested the effect of CBD on α7-nACh receptors inthe CA1 region of stratum radiatum interneurons in rat hippo-campal brain slices. In whole cell patch clamp mode, focalapplication of 10 mM choline, a selective agonist for α7-nAChreceptor (Albuquerque et al., 2009) for a short duration (0.5–1 s)caused a rapidly activating and fast desensitizing inward currentsthat were completely inhibited by the bath application of 1 mMmethyllycaconitine, a selective antagonist for α7-nACh receptor(data not shown, n¼3). Choline-induced currents were signifi-cantly inhibited by 10 min bath application of 10 mM CBD (Fig. 6A).Time-courses of effects of CBD and the vehicle applications on theamplitudes of choline-induced currents are presented in Fig. 6B.CBD (10 μM) caused a significant inhibition of the current whichwas partially reversed during 10 min washout period. In theabsence of CBD, vehicle (0.3% ethanol) alone did not alter theamplitude of the ACh-induced current (Fig. 6B, controls). We alsocompared the effects of CBD with CBN and THC on the choline-induced currents in hippocampal interneurons. As shown in

Fig. 6C, neither CBN nor THC (10 mM) appeared to affect themaximal amplitude of choline (10 mM)-induced currents.

Although it is not specific for α7-nACh receptors, in the rathippocampus, nicotine-induced [3H] norepinephrine (NE) releasefrom synaptic membranes has been used to assess indirectly thefunction of α7-nACh receptors (Smith et al., 2009). Therefore, wehave also investigated the effects of CBD on [3H]NE releaseinduced by nicotine (30 mM), in rat hippocampal slices. A 10 minapplication of nicotine increased [3H]NE release at a maximumwithin 5 min. In contrast, following a 10 min preincubation withCBD (10 μM), agonist-evoked [3H]NE release was decreased sig-nificantly (Fig. 7A). The area above the basal [3H]NE release duringthe agonist-application was used to determine the effects of CBD,CBN and THC (Fig. 7B). The results indicated that although CBD(10 mM) inhibited nicotine-evoked [3H]NE release, CBN and THCdid not cause a significant alteration of [3H]NE release.

4. Discussion

In the present study, using electrophysiological and biochem-ical methods, we provide evidence that CBD causes a significantinhibition of human α7-nACh receptors expressed in Xenopus

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Fig. 5. Lack of correlation between current amplitudes and the effect of cannabidioland a comparison of the effects of cannabidiol (CBD), cannabinol (CBN), and Δ9-tetrahydrocannabinol (THC) on human α7-nicotinic receptors. (A) Cannabidiolinhibition of nACh receptors is not correlated with the mean current density(MCD) ACh (100 mM)-induced currents. Lack of correlation between the percentCBD inhibition of mean current density is shown in the plot (linear regression,r¼0.32). (B) Comparison of the effect of 10 mM of CBD, CBN, and THC on ACh(100 mM)-induced currents in oocytes expressing human α7-nACh receptors. Barsrepresent the means7S.E.M. of 4–7 experiments. The number of experiments ispresented on top of each bar.

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oocytes in a time and concentration dependent manner. In addi-tion, the results indicate that the inhibitory effect of CBD on α7-nACh receptors is not a common pharmacological property of allphytocannabinoids, since THC and CBN do not appear to alterfunctional characteristics of this receptor.

Cannabidiol has been found to function as an antagonist of bothCB1 and CB2 receptors (for a review, Pertwee, 2008). Since CBreceptors are not expressed in Xenopus oocytes (Hejazi et al., 2006;Oz et al., 2007) it is unlikely that PTX sensitive receptors mediateinteraction between CBD and nACh receptor. Furthermore, bindingof GTPγS to oocyte membranes was not altered by CBD, suggestingthat G-protein coupled receptors are not involved in the observedactions of CBD.

Cannabidiol, in the concentration range used in this study, hasbeen shown to elevate intracellular Ca2þ levels in cultured hippo-campal neurons (Drysdale et al., 2006; Ryan et al., 2006) glia (Matoet al., 2010) and immune system cells (Rao and Kaminski, 2006). InXenopus oocytes, activation of α7-nACh receptors, due to their highCa2þ permeability, allows sufficient Ca2þ entry to activate endo-genous Ca2þ-dependent Cl� channels (Sands et al., 1993; Hartzellet al., 2005). Therefore, the direct actions of CBD on Ca2þ-dependentCl� channels may contribute to the observed effects of CBD on ACh-activated currents in this expression system. In oocytes injectedwith BAPTA and recorded in a solution containing 2 mM Ba2þ , CBDcontinued to inhibit α7-nACh receptor-mediated ion currents,

suggesting that Ca2þ-dependent Cl� channels were not involvedin CBD inhibition of nicotinic responses. In addition, the reversalpotential in solutions containing Ba2þ was not altered in thepresence of CBD, suggesting that the inhibitory effect of CBD is notdue to alterations in the Ca2þ permeability of the α7-nACh receptor–channel complex. Furthermore, since Ca2þ-activated Cl� channelsare highly sensitive to intracellular Ca2þ levels (KD of Ca2þ-activatedCl� channels for Ca2þ is less than 1 mM, for a review (Hartzell et al.,2005), alterations in intracellular Ca2þ levels would be reflected bychanges in the holding current under voltage-clamp conditions.However, during our experiments, application of CBD (10 μM), didnot cause alterations in baseline or holding currents, suggesting thatCBD does not affect intracellular Ca2þ concentrations.

In this study, CBD was applied in the concentration range of1–30 mM and was found to successfully inhibit the function ofα7-nACh receptors in a concentration-dependent manner (IC50value of 12.7 mM). The concentration of CBD in plasma and itsability to pass the blood brain barrier following intraperitoneal,oral and intravenous administration has been studied previously(Lodzki et al., 2003; Varvel et al., 2006; Hill et al., 2012).Commonly used doses of CBD (3–10 mg/kg) are found to promoteCBD brain levels of 200 nM to 3 μM, respectively (Varvel et al.,2006). Since CBD is a highly lipophilic compound with LogP(octanol–water partition coefficient) value ranges between 6 and8 (Lodzki et al., 2003) its membrane concentration is expected to

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Fig. 6. The effect of cannabidiol on choline-induced ion currents recorded in CA1 area stratum radiatum interneurons of rat hippocampal slices. (A) Recordings of choline-induced currents before (control, left panel), during (10 min of CBD) and after (10 min of recovery) the bath application of 10 mM CBD in hippocampal interneurons. Cholineapplication time was indicated by a solid bar on top of the current traces. Dashed line indicates continuing bath application of CBD. (B) Time-course of the effect of vehicle(0.3% ethanol; filled circles) and CBD (10 μM; open circles) on the peaks of the ACh-induced currents. Each data point represents the normalized mean7S.E.M. of 4–5experiments. Duration of drug application is indicated by the horizontal bar in the figure. (C) Comparison of the effect of 10 mM of CBD, CBN, and THC on Choline (10 mM)-induced currents in oocytes expressing human α7-nACh receptors. Bars represent the means7S.E.M. of 4–5 experiments. The number of experiments is presented on top ofeach bar.

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be considerably higher than blood levels. Therefore, the functionalmodulation of nACh receptors demonstrated in this study can bepharmacologically relevant.

In earlier studies, the actions of CBD on several integral mem-brane proteins including glycine receptors (Ahrens et al., 2009; EC50vaules are 12.3 mM and 18.1 mM for α1 and α1β1 subunits), mus-carinic receptors (Cluny et al., 2011; 3–30 mM);5-HT receptors (Russo et al., 2005; 30 mM displaces 50% of ketan-serin binding), opioid receptors (Kathmann et al., 2006, IC50¼8–12 mM), transient receptor potential channels (Bisogno et al., 2001,TRPV1 activation EC50¼3.5 mM; De Petrocellis et al., 2008, TRPA1and TRPM8 activation EC50¼01–1 mM; Qin et al., 2008, TRPV2activation EC50¼3.7 mM), and T-type Ca2þ channels (Ross et al.,2008; IC50¼1–3 mM) have also been described (for recent reviews,Izzo et al., 2009; Hill et al., 2012). It is plausible that CBD acts as an

allosteric modulator for various receptors and ion channels at themembrane accounting for some of its analgesic, antiepileptic andanti-inflammatory actions (for a review, Izzo et al., 2009).

Open-channel blockade is a widely used model to describe theblock of ligand-gated ion channels (Hille, 2001). However, this modelcannot account for the results of the present study. Firstly, for openchannel blockers, the presence of the agonist is required to allow theblocker to enter the channel after an agonist-induced conformationalchange. In contrast to open channel blockers, preincubation wasfound to augment CBD's effect on α7-nACh receptor activity (Fig. 1),suggesting that CBD interacts with the closed state of the receptor.Secondly, inhibition by CBD is not voltage sensitive, suggesting thatthe CBD-binding site is not affected by the transmembrane electricfield. Similarly, there was an absence of use-dependent blockade(Fig. 3) and CBD had little effect when co-administered with AChwithout CBD preincubation (only coapplication data in Fig. 1C).Thirdly, recovery from CBD inhibition occurred independent ofagonist application intervals (Fig. 3C), indicating that CBD is nottrapped in the channel when the channel closes, as can occur withopen channel blocking drugs.

Allosteric modulators alter the functional properties of ligand-gated-ion channels by interacting with site(s) that are topogra-phically distinct from the ligand binding sites (for a review;(Onaran and Costa, 2009). In electrophysiological studies, althoughthe potency of ACh, a natural ligand (agonist) for the receptor, wasnot altered, its efficacy was significantly inhibited by CBD, indicat-ing that CBD did not compete with the ACh binding site on thereceptor. In agreement with these findings, radioligand bindingstudies indicated that the specific binding characteristics of [125I]α-bungarotoxin were not affected by CBD, further suggesting thatCBD does not interact with the ACh binding site on the receptor.Collectively, these findings indicate that CBD acts as an allo-steric inhibitor of the α7-nACh receptor i.e., CBD binds to site(s) topographically distinct from the ACh binding sites on thereceptor-ion channel complex. The non-competitive property ofthe allosteric CBD inhibition puts it in an advantageous position,since increases in the concentration of the endogenous agonist(ACh) in the synaptic cleft will not alter the efficacy of CBD.

It is likely that CBD, a highly lipophilic agent, first dissolves intothe lipid membrane and then diffuses into a non-annular lipidspace to inhibit the ion channel-receptor complex. Consistent withthis idea, the effect of CBD on α7-nACh receptor reached a maximallevel within 10–15 min of application. Similarly, actions of severalhydrophobic allosteric modulators such as cannabinoids (Oz et al.,2004c; Spivak et al., 2007), and general anesthetics (Zhang et al.,1997) on ligand-gated ion channels require 5–20 min to reachtheir maxima (for a review, Oz, 2006), suggesting that the bindingsite(s) for these allosteric modifiers is located inside the lipidmembrane and require a relatively slow (in minutes) time courseto modulate the function of the receptor. It is likely that thesehydrophobic agents (for a review, Oz, 2006) affect the energyrequirements for gating-related conformational changes in ligand-gated ion channels (Spivak et al., 2007).

In rat hippocampal interneurons, currents activated by cholinea selective agonist for α7-nACh receptors were significantly inhib-ited after 10 min application of CBD, indicating that CBD canregulate the functions of native α7-nACh receptors expressed inintact neurons. Nicotine-induced [3H]NE release from rat hippo-campal slices has also been used for an indirect functionalassessment of α7-nACh receptors (Smith et al., 2009). The inhibi-tion of choline-induced currents and nicotine-induced [3H]NErelease by CBD suggest that the actions of CBD observed in theheterologous expression system (oocytes) also occur in neuronsand may therefore, contribute to neuronal circuitry and function.In conclusion, our results indicate that CBD inhibits the function ofthe α7-nACh receptor.

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Fig. 7. The effects of cannabidiol and other phytocannabinoids on agonist-evoked[3H]NE release in rat hippocampal slices. (A) Fractional [3H]NE release from rathippocampal slices evoked by 30 μM nicotine alone (control) or in the presence ofcannabidiol (CBD; 10 μM) after 10 min CBD preincubation. Data are expressed as %of basal release (BL¼mean release of 3 preceding fractions). The solid bar showsthe duration of nicotine application. Cannabinoids were applied in perfusionstream for 10 min before nicotine and continued in the solution after coapplicationwith nicotine. (B) Effects of CBD, CBN, and THC on nicotine-evoked [3H]NE releasefrom rat hippocampal slices. The areas under the release curves (AUC) shown inpanel A were calculated for controls (nicotine only) and in the presence of nicotineplus cannabinoids. Basal release levels for each condition were calculated from thelinear fit of beginning fractions (dashed line in Fig. 7A) and subtracted fromnicotine-induced release. The means7S.E.M. of subtracted values (Δ–AUC) wereexpressed for each condition in the bar graph. The numbers of measurements from3 to 4 independent experiments are presented at the top of each column.

M. Mahgoub et al. / European Journal of Pharmacology 720 (2013) 310–319318

Acknowledgments

The research in this study was supported by the grants fromCMHS, UAE University. Research in our laboratory is also sup-ported by LABCO partner of Sigma–Aldrich. The authors gratefullyacknowledge Dr. Jon Lindstrom for providing cDNA clones of thehuman α7-nACh receptor subunit. We thank Dr. Syed Nurulain ofDepartment of Pharmacology, CMHS, for his expertise and assis-tance in some of the experiments. We also thank Mr. Nadeem U.Rahman for his invaluable support in establishing the data-acquisition system in our laboratory.

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