Neurophysiological evidence for the presence of cannabinoid CB1 receptors in the laterodorsal tegmental nucleus.

European Journal of Neuroscience, Vol. 40, pp. 3635–3652, 2014

doi:10.1111/ejn.12730

MOLECULAR AND SYNAPTIC MECHANISMS

Neurophysiological evidence for the presence of cannabinoid CB1 receptors in the laterodorsal tegmental nucleus Neeraj Soni,1 Shankha Satpathy2 and Kristi A. Kohlmeier1 1

Department of Drug Design and Pharmacology, Faculty of Health and Medical Sciences, University of Copenhagen, Universitetsparken 2, DK-2100, Copenhagen, Denmark 2 The Novo Nordisk Foundation Center for Protein Research, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark Keywords: arousal, cholinergic, electrophysiology, endocannabinoids, synaptic transmission

Abstract Marijuana, which acts within the endocannabinoid (eCB) system as an agonist of the cannabinoid type 1 receptor (CB1R), exhibits addictive properties and has powerful actions on the state of arousal of an organism. The laterodorsal tegmental nucleus (LDT), as a component of the reticular activating system, is involved in cortical activation and is important in the development of drug addiction-associated behaviours. Therefore, eCBs might exert behavioural effects by actions on the LDT; however, it is unknown whether eCBs have actions on neurons in this nucleus. Accordingly, whole-cell voltage- and current-clamp recordings were conducted from mouse brain slices, and responses of LDT neurons to the CB1R agonist WIN-2 were monitored. Our results showed that WIN-2 decreased the frequency of spontaneous and miniature inhibitory postsynaptic currents (sIPSCs and mIPSCs). Ongoing activity of endogenous eCBs was confirmed as AM251, a potent CB1R antagonist, elicited sIPSCs. WIN-2 reduced the firing frequency of LDT neurons. In addition, our RT-PCR studies confirmed the presence of CB1R transcript in the LDT. Taken together, we conclude that CB1Rs are functionally active in the LDT, and their activation changes the firing frequency and synaptic activity of neurons in this nucleus. Therefore, endogenous eCB transmission could play a role in processes involving the LDT, such as cortical activation and motivated behaviours and, further, behavioural actions of marijuana are probably mediated, in part, via cellular actions within the LDT induced by this addictive and behavioural state-altering drug.

Introduction As part of the classic reticular activating system, the laterodorsal tegmental nucleus (LDT) is believed to participate in controlling cortical activity across sleep and wakefulness states (Hallanger et al., 1987; el Mansari et al., 1989). Recently, attention has been drawn to a role of the LDT in manifestation of drug addictionassociated behaviours (Forster & Blaha, 2000; Nelson et al., 2007; Lammel et al., 2012). The acetylcholine (ACh)-containing neurons of the LDT provide the major cholinergic input to dopamine (DA)containing ventral tegmental area (VTA) neurons (Cornwall et al., 1990; Oakman et al., 1995) and allow the burst firing pattern required for the large release of DA necessary for evaluation and assignment of reward to environmental stimuli such as drugs of abuse (Schultz, 1998; Lodge & Grace, 2006). The induction of behaviour associated with drug addiction by optogenetic stimulation selectively of neurons of the LDT-VTA pathway in the absence of drug cues has provided compelling evidence that the LDT can regulate motivated behaviour (Lammel et al., 2012).

Correspondence: Dr Kristi Anne Kohlmeier, as above. E-mail: [email protected] Received 27 May 2014, revised 13 August 2014, accepted 20 August 2014

Marijuana, which is the most commonly abused illicit drug, has actions on behavioural state and exhibits addictive potential (Mechoulam et al., 1970; Nutt et al., 2007, 2010). The psychoactive compound in marijuana is tetrahydrocannabinol (Gaoni & Mechoulam, 1964), which acts as an agonist at one of the two known cloned and pharmacologically-identified, cannabinoid receptors, the cannabinoid type-1 receptor (CB1R) (Matsuda et al., 1990). Endogenous activation of the CB1R occurs by activity-induced (e.g. depolarisation, metabotropic receptor mediated) production and secretion of natural endocannabinoids (eCBs) (Devane et al., 1992). Activation of presynaptic CB1Rs can result in reducing the release of GABA or glutamate onto the postsynaptic cell and can involve a retrograde process known as suppression of inhibition (Pitler & Alger, 1994; Wilson & Nicoll, 2001) or suppression of excitation (Melis et al., 2004), respectively, when activity in the postsynaptic cell activates eCBs which serve as a reverse neurotransmitter. Mechanisms underlying the inhibition of presynaptic transmission have been shown to include decreases in intracellular Ca2+ via inhibition of voltage-gated calcium channels (Caulfield & Brown, 1992; Mackie & Hille, 1992; Guo & Ikeda, 2004) and activation of inwardly rectifying potassium channels (Henry & Chavkin, 1995; Guo & Ikeda, 2004).

© 2014 Federation of European Neuroscience Societies and John Wiley & Sons Ltd

3636 N. Soni et al. Although the LDT is involved in cortical activity and drug addiction-associated cellular processes and behaviours, and marijuana is well known to have actions on arousal state and exhibit addictive properties, whether agonists of the CB1R have actions on LDT neurons is unexplored. Accordingly, we tested the hypothesis that functional CB1Rs are present in the LDT by examining the actions of WIN-2, a CB1R agonist, on membrane currents, postsynaptic currents and firing of LDT neurons using patch-clamp recordings from mouse LDT neurons in brain slices. In addition, by using a CB1R antagonist we examined whether endogenous activation of CB1Rs in the LDT was present; this could suggest production of eCBs in the brain slice. Our data indicate that exogenous application of CB1R agonists, such as marijuana, would have actions on LDT cells and, further, they suggest that endogenous stimulation of the eCB system could influence processes and behaviours with which the LDT is involved, such as cortical activation during different behavioural states and drug addiction-associated behaviours.

Materials and methods Ethical approval All animal studies complied with the European Communities Council Directive of 24 November 1986 (86/609/EEC) and with Danish laws regulating experiments on animals. Animal use studies were permitted by the Animal Welfare Committee, appointed by the Danish Ministry of Justice. Mice Brain slices were taken from 10- to 25-day-old NMRI wild-type mice of either sex (10–25 g; Harlan Mice laboratories, Denmark) that were housed in a temperature-controlled room (22–23 °C) with a 12:12 h light : dark cycle (lights on at 07.00 h). Data in this report are sourced from a total of 53 animals. Tap water and laboratory chow were available ad libitum. Brain slice preparation Preparation of brain slices containing the LDT was conducted as previously described (Kohlmeier et al., 2006). Briefly, before extraction of the brain, mice were anaesthetised with isofluorane. Sufficient depth of anesthesia was assayed by failure to display a reaction upon a strong pinch of the paw. Following decapitation, the brain was rapidly removed and placed into ice-cold (0–4 °C) artificial cerebrospinal fluid (ACSF) and sectioned into 250-lm-thick coronal slices which contained the LDT, using a vibratome which had been calibrated right before sectioning to reduce horizontal vibration thereby facilitating cell survival and health (Leica VT 1200S; Leica, Germany). It was possible to obtain one or two brain slices containing the LDT from each animal. Dissections and sectioning were performed in ACSF containing (in mM): NaCl, 124; KCl, 5; Na2HPO4, 1.2; CaCl2, 2.7; MgSO4, 1.2; glucose, 10; and NaHCO3, 2.6; bubbled with 95% O2 and 5% CO2, resulting in a pH of 7.4. Slices were incubated in ACSF at 37 °C for 15–20 min and thereafter maintained at room temperature for 1 h to allow cellular equilibration before placement within the recording chamber. All recordings were conducted at room temperature. Whole-cell voltage- and current-clamp electrophysiological recording Slices were placed in a recording chamber (volume ~ 1 mL) in continuously oxygenated (95% O2, 5% CO2) ACSF at room temperature

(16–20 °C) at a fixed flow rate of 2–3 mL/min. The recording chamber was fitted onto the fixed stage of an upright microscope (Olympus BX51WI, Germany) and neurons were viewed under Olympus optics with 409 water-immersion objective. Patch-clamp recordings were initiated using visual guidance of the electrode to the cell surface with an Imago CCD camera (PCO, Sensicam, TILL Photonics, Germany). A maximum of two neurons were utilised in each slice, with a minimum of a 30-min recovery period between drug applications. Electrodes were made from borosilicate glass capillary tubing (1.5 mm O.D., 1–3 GO), capacitance cancellation was minimised to the slowest component of capacitive current elicited by a 5-mV voltage step. Electrophysiological recordings were conducted using an Axopatch 200B amplifier (Molecular Devices: Axon Instruments, USA) operated in whole-cell voltage-clamp gap-free mode, and all recorded membrane currents were filtered at 2 KHz (low-pass Bessel filter) at the amplifier output and digitised at 20 KHz with subsequent analysis being performed using CLAMPEX 10.3 (Molecular Devices, USA). Recordings were discontinued if access resistance, which was calculated with an online tool by the exponential fitting of the uncompensated capacitance transient in response to a brief step of the voltage, was unstable, or varied by > 20%. Holding currents were monitored continuously during recordings and, if found to increase by > 50 pA, the recordings were terminated. For some voltage-clamp recordings and all current-clamp recordings, patch electrodes were filled with an internal patch solution containing (in mM): K-gluconate, 144; MgCl2, 3; HEPES, 10; NaGTP, 0.3; and Na2ATP, 4. The osmolarity of the pipette solution was adjusted to k295–300 mOsM and pH to 7.4. To record spontaneous and miniature inhibitory postsynaptic currents (sIPSCs and mIPSCs), a high-chloride recording solution was used to optimise detection of inhibitory events by increasing the drive for, and reversing the polarity of, chloride-mediated currents. This patch solution contained (in mM): KCl, 144; EGTA, 0.2; MgCl2, 3; HEPES, 10; NaGTP, 0.3; and Na2ATP, 4; osmolarity range between 295 and 300 mOsM). The intracellular electrode was also filled with Alexa-594 (25 lM; Sigma-Aldrich, St Louis, MO, USA), which was allowed to passively diffuse into the cell once the membrane of the cell had been ruptured by the pipette. This dye remained in the cell if the patch pipette was carefully removed at the end of the recording and could be visualised later with appropriate optics. This allowed detection of recorded cells following immunohistochemical procedures for identification of neurotransmitter content of recorded cells. Drugs (R)-(+)-[2,3-dihydro-5-methyl-3-(4-morpholinylmethyl)pyrrolo[1,2,3de]-1,4-benzoxazin-6-yl]-1-naphthalenylmethanone mesylate (WIN55, 212-2, abbreviated to WIN-2), which is a synthetic analogue of delta-9-tetrahydrocannabinol, the psychoactive component of marijuana, was used as a potent cannabinoid agonist for CB1R (10 lM; Sigma-Aldrich). 1-(2,4-dichlorophenyl)-5-(4-iodophenyl)-4-methyl-N(1-piperidyl) pyrazole-3-carboxamide (AM251; 10 lM; Tocris, Bristol, UK) was used as an antagonist for CB1Rs. WIN-2 and AM251 were dissolved in dimethyl sulfoxide (DMSO) and were stored at 20 °C. The final concentration of DMSO applied to cells was 10 min) of WIN-2, the amplitude of sIPSCs was found to decrease by 21.5 4.5% of control (amplitude after WIN-2: 22.8 2.2 pA, n = 22/25 from 16 slices, P = 0.0011, paired t-test; Fig. 1, C2). WIN2 application reduced the frequency of sIPSCs to 49.7 4.0% of control (Control, 0.4 0.05 Hz; WIN-2, 0.2 0.04 Hz; n = 22/25, P < 0.0001, paired t-test; Fig. 1, C1). As reported in other studies, actions of WIN-2 on IPSCs were slow to develop, exhibited a wide range in latency (Roberto et al., 2010) and full effects were not noted until WIN-2 had been continuously applied for 10–15 min (Hoffman & Lupica, 2000). In preliminary experiments, there were no observable differences across age or gender and therefore data have been pooled. WIN-2 decreased the frequency of mIPSCs but did not affect their amplitude within the LDT To determine whether the WIN-2-induced changes in frequency and amplitude of sIPSCs were reliant on action potentials generated within the slice, we performed recordings in the presence of TTX (500 nM), which blocks voltage-dependent sodium channels. During the blockade of TTX-dependent action potentials, we recorded mIPSCs and found that WIN-2 had a significant effect on reducing the frequency of mIPSCs, but did not have a significant effect on the amplitude. Application of WIN-2 resulted in a reduction in mIPSC frequency of 58.1 5.6% from control (Control, 0.25 0.05 Hz; WIN-2, 0.09 0.01 Hz; n = 15/19 from 12 slices, P = 0.0016, paired t-test; Fig. 2, C1) and a non-significant decrease in amplitude of 12 7% from control (Control, 21.9 1.8 pA; WIN-2, 19.8 2.9 pA; n = 15, P = 0.2971, paired t-test; Fig. 2, C2). The WIN-2-mediated decrease in the frequency of mIPSCs without any change in their amplitude indicates that the actions of WIN-2 on mIPSCs frequency are action potential-independent and, further, these data suggest that WIN-2 is acting at presynapticallylocated CB1Rs located on inhibitory terminals, rather than at more distally located receptors. Taken together, our data suggest that CB1R agonists in the LDT would significantly affect inhibitory tran-smission directed to postsynaptic LDT neurons.

WIN-2, a cannabinoid CB1R agonist, decreased sIPSCs within the LDT

WIN-2 did not induce any significant changes in frequency and amplitude of spontaneous EPSCs (sEPSCs) and miniature EPSCs (mEPSC) within the LDT

While postsynaptic localisation of CB1Rs has been reported, including in spinal cord neurons of the rat dorsal horn, trigeminal neurons, and

As a heavy glutamatergic input is directed to LDT cells (Wang & Morales, 2009) and this excitatory input is functionally active in

© 2014 Federation of European Neuroscience Societies and John Wiley & Sons Ltd European Journal of Neuroscience, 40, 3635–3652

Functional presence of CB1Rs in the LDT 3639 A

B

C

Fig. 1. WIN-2 reduced sIPSC frequency and amplitude. (A) WIN-2 altered sIPSC frequency and amplitude in LDT neurons, shown in representative voltageclamp recordings of a cell in control conditions (ACSF; A1, left trace) and during the application of WIN-2 (A2; left trace). Higher gain records are to the right of these traces where it is clearly apparent that WIN-2 induces a change in the frequency and amplitude of IPSCs. (B1) WIN-2 induced a decrease in the amplitude of sIPSCs, apparent when comparison is made of the amplitude of a composite sIPSC composed of the average of several sIPSCs recorded before WIN-2 (44.5 pA; n = 76 events, black) and an average composite of several inhibitory events recorded following WIN-2 exposure in the same cell (17.7 pA; n = 35 events, gray). (B2 and 3) Examination of cumulative distribution plots of sIPSC inter-event intervals and amplitudes before (black) and during (gray) WIN-2 application from an individual cell which was identified as bNOS-positive show that WIN-2 induced a significant decrease in sIPSC amplitudes (K-S test, P < 0.05) and induced a shift toward the right in the distribution of inter-event intervals which was significant (K-S test, P < 0.05), indicating a decrease in sIPSC frequency. A total of three cells that exhibited this response to WIN-2 were identified as bNOS-positive. (C) Histogram showing the significant effect of WIN-2 on reducing (C1) the frequency and (C2) the amplitude of sIPSCs from the population of LDT neurons examined. Data were analysed using Student’s t-test (two-tailed, paired); statistically significant differences were assumed at P < 0.05. Errors represent SEM. #P ≤ 0.001, ##P ≤ 0.0001.

brain slices (Kohlmeier et al., 2012), we examined whether WIN-2 had actions on EPSCs recorded in LDT cells. Recordings were conducted in the presence of the glycine receptor antagonist strychnine (2.5 lM) and the GABAA receptor antagonist gabazine (10 lM) to block inhibitory input. In the majority of cells, WIN-2 failed to induce a significant decrease in the frequency of sEPSCs (15.1 7.1% reduced from control: Control, 1.5 0.5 Hz; WIN-2, 1.4 0.5 Hz; n = 7/9 from six slices, three animals, P = 0.1506, paired t-test; Fig. 3, C1). WIN-2 also failed to result in a significant change in sEPSC amplitude in these cells (8.6 7.2% reduced from control: Control, 29.9 5.1 pA; WIN-2, 27.1 5.5 pA; n = 7/9, P = 0.1867, paired t-test; Fig. 3, C2). However, in two of these cells, WIN-2 did induce a significant decrease in frequency (Control, 0.5 0.2 Hz; WIN-2, 0.3 0.1 Hz; n = 2/9, P < 0.05, K-S test; data not shown), but failed in these cells to induce a

significant change in amplitude (Control, 22.0 6.3 pA; WIN-2, 21.0 4.7 pA; n = 2, P > 0.05, K-S test). In the presence of TTX, WIN-2 failed to induce any significant change in mEPSC frequency in the majority of cells. (8.6 3.0% reduced from control: Control, 0.60 0.03 Hz; WIN-2, 0.55 0.02 Hz; n = 5/6 from six slices, three animals, P = 0.0759, paired t-test), nor did WIN-2 induce a significant change in amplitude (3.8 2.1% reduced from control: Control, 39.0 5.6 pA; WIN-2, 38.0 5.2 pA; n = 5/6, P = 0.0836, paired t-test). In one cell recorded in TTX, WIN-2 did induce a significant decrease in frequency (Control, 0.37 Hz; WIN-2, 0.20 Hz; n = 1/6, K-S test, P = 0.0027, data not shown) but did not significantly alter the amplitude (Control, 26.0 pA; WIN-2, 27.6 pA; n = 1/6, K-S test, P = 0.7300). Taken together, these data suggest that in the majority of cases WIN-2 does not activate CB1Rs located on glutamatergic


3640 N. Soni et al. A

B

C

Fig. 2. WIN-2 reduced mIPSC frequency without affecting mIPSC amplitude. (A) Voltage-clamp traces displaying mIPSCs in control conditions (ACSF + TTX, A1, left) and during the applications of WIN-2 + TTX (A2, left), and higher gain records (right), show that WIN-2 decreased the frequency but not the amplitude of mIPSCs in LDT neurons. (B1) The amplitude of the composite mIPSC composed of the average of several mIPSCs before drug application (13.4 pA; n = 116 events; black) was not different from the amplitude measured from an average of several events following WIN-2 exposure (11.5 pA; n = 30 events, gray). (B2 and 3) Cumulative distribution plots of mIPSC inter-event intervals and amplitude before (black) and during (gray) WIN-2 application for the cell shown in A. The cumulative plots indicate that WIN-2 induced a significant decrease in the mIPSC frequency (K-S test, P < 0.05) but did not induce a significant change in amplitude (K-S test, P > 0.05). (C) Summary of the effect of WIN-2 on (C1) the frequency and (C2) the amplitude of mIPSCs across a population of cells, indicating that a significant change was noted only in a decrease in the frequency of mIPSCs (two-tailed paired Student’s t-test, with significance assumed at P < 0.05). Error bars are SEM. #P ≤ 0.001.

presynaptic inputs within the LDT but do suggest that, in a small group of LDT cells, CB1R activation can affect glutamatergic release. The cannabinoid CB1R antagonist AM251 increased the frequency of sIPSCs within the LDT In the next series of experiments, we wanted to examine whether endogenously present eCBs were exerting actions on LDT neurons within the brain slice. To this end, we applied a potent synthetic CB1R antagonist, AM251 (10 lM), and recorded changes in the activity of ongoing sIPSCs of LDT neurons. Application (> 5–8 min) of AM251 was found to significantly increase the frequency of sIPSCs to 71.2 38.7% of control values in the majority of the LDT neurons recorded (Control, 1.4 0.36 Hz; AM251, 1.9 0.38 Hz; n = 5/6 from four slices, three animals,

P = 0.0388, paired t-test; Fig. 4, C1); however, AM251 failed to significantly increase the amplitude of sIPSCs in these cells (1.9 9.3% of control: Control, 33.2 5.03 pA; AM251, 31.9 4.9 pA; n = 5/6, P = 0.7193, paired t-test; Fig. 4, C2). Our findings using the CB1R antagonist provide further evidence that CB1Rs are present within the LDT and can be functionally activated. Further, our data, with the caveat that studies were conducted in brain slices, suggest that production of eCBs sufficient to activate CB1Rs and influence LDT synaptic activity occurs endogenously. The cannabinoid CB1R antagonist AM251 suppressed WIN-2-induced inhibition of mIPSCs within the LDT To test whether the actions of WIN-2 were specific to agonism at the CB1R, we examined whether antagonism of the CB1R



B

C

Fig. 3. WIN-2 did not affect sEPSC frequency and amplitude in the majority of cells. (A) Example traces displaying sEPSCs in (A1, left) control conditions and (A2, left) in the presence of WIN-2. As is apparent in (A1) the low-gain voltage-clamp records and (A2) higher gain inserts to the right, WIN-2 did not induce a change in frequency or amplitude of recorded sEPSCs in LDT neurons in this cell, nor were significant changes induced in these parameters in the majority of LDT neurons recorded. (B1) The amplitude of a composite sEPSC composed of the average of several sEPSCs before drug application (12.8 pA; n = 213 events; black) was not different from the amplitude measured from an average of several events following WIN-2 exposure (11.5 pA; n = 203 events, gray). (B2 and 3) Cumulative distribution plots of sEPSC (B2) inter-event intervals and (B3) amplitudes before (black) and during (gray) WIN-2 application for the experiment shown in A. The cumulative plots show that WIN-2 did not induce any significant change in the sEPSC frequency (K-S test, P > 0.05) or amplitude (K-S test, P > 0.05). (C) Summary of the effects of WIN-2 on (C1) the frequency and (C2) the amplitude of sEPSCs in a population of LDT neurons, indicating that WIN-2 failed to have a significant action on sEPSCs in the majority of neurons examined (two-tailed paired Student’s t-test, with significance assumed at P < 0.05). Error bars are SEM.

prevented the effect of WIN-2 on mIPSCs. In the presence of AM251, WIN-2 failed to significantly decrease the frequency of mIPSCs. In a group of cells in which TTX and AM251 were present, WIN-2 reduced the frequency of mIPSCs by only 12.00 8.10% of control, which was not significant (AM251 + TTX, 0.70 0.21 Hz; AM251 + TTX+WIN-2, 0.66 0.21 Hz; n = 6/6 from five slices, four animals, P = 0.4065, paired t-test; Fig. 5, C1). While the mean amplitude was slightly changed, the reduction in AM251 in this parameter was not significant (6.0 5.2% changed from control: AM251 + TTX, 44.0 4.9 pA; AM251 + TTX+ WIN-2, 40.0 3.9 pA; n = 6/6, P = 0.2050, paired t-test; Fig. 5, C2). Taken together, our data suggest the effect of WIN-2 in reducing mIPSC frequency is specific to actions at the CB1R, and that CB1Rs are engaged at terminals presynaptic to LDT cells in controlling GABAergic input to LDT neurons.

WIN-2 reduced the firing frequency of LDT neurons As we found that CB1R activation reduced the frequency of mIPSCs directed to LDT neurons, we hypothesised that WIN-2 appli cation might be associated with an enhancement in the excitability of LDT neurons which might be reflected in alterations in action potential firing rates. To examine this issue, we recorded the effects of WIN-2 on the steady-state firing frequency of LDT neurons using current-clamp recordings. Firing frequency was recorded during blockade of excitatory transmission (AP5, CNQX). Contrary to our expectations, we found that WIN-2 application resulted in a decrease in firing frequency across the population of cells studied, and in no single recorded neuron did we detect an increase in firing following WIN-2 exposure. In a population of matched cells in which it was possible to obtain firing before and after WIN-2, in the presence of



B

C

Fig. 4. The cannabinoid CB1R antagonist AM251 increased the frequency but did not affect the amplitude of sIPSC in the LDT. (A1 and 2, left) Whole-cell recording in voltage-clamp mode showing the actions of the CB1R antagonist AM251 on inducing a change in sIPSCs. Higher gain recordings are shown to the right. (B1) Composite sIPSC of the average of several events before (black) (41.9 pA; n = 227 events; black) and after AM251 exposure (gray) (45.7 pA; n = 320 events; black), indicating that this receptor antagonist did not influence the amplitude of sIPSCs. (B2 and 3) Cumulative distribution plots of sIPSC (B2) inter-event intervals and (B3) amplitudes before (black) and during AM251 (gray) application for the cell shown in A indicate that AM251 significantly increased the frequency of sIPSCs (K-S test, P < 0.05), but did not significantly influence the amplitude (K-S test, P > 0.05). (C) Summary histograms indicate that AM251 significantly decreased (C1) the frequency but (C2) not the amplitude of sIPSCs across a population of LDT neurons (two-tailed paired Student’s t-test, with significance assumed at P < 0.05). Error bars are SEM. *P ≤ 0.05.

WIN-2 the mean steady-state firing frequency was 14.0 2.3 Hz (n = 6/6), which was significantly lower than the mean firing frequency in absence of WIN-2 (in ACSF, 22.9 2.9 Hz; n = 6/6 from four slices, two animals, P = 0.0007, paired t-test; Fig. 6, A3). Comparison of firing rates in a larger population of cells in which it was not possible to obtain the firing frequency before and after in the same matc-hed cells revealed that the firing rate in the presence of WIN-2 (10.4 2.2 Hz; n = 22, from 10 slices) was significantly lower than the mean firing frequency in the absence of WIN-2 (17.9 2.0 Hz; n = 30, P < 0.0001, unpaired t-test; data not shown). In a separate population of cells, we examined the ability of AM251 to alter the WIN-2-induced decrease in firing rate. In this group of cells, WIN-2 reduced the firing frequency to 10.0 1.0 Hz (n = 7/7) from a higher rate of firing in control conditions. During continued application of WIN-2, AM251 was then applied continuously (> 10 min) and we observed an increase in firing frequency by 34.0 6.0% from WIN-2 conditions (WIN-2, 10.0 0.8 Hz; AM251 + WIN-2, 14.0 1.4 Hz; n = 7/7 from four slices, three animals, P = 0.0021, paired t-test; Fig. 6, B3), suggesting a CB1R mediated inhibition of firing within the slice which, once induced, could be partially antagonised by inhibition of the CB1R. Application of AM251 alone did not have a significant

action on firing rate, suggesting that within the slice, endogenous activation of CB1Rs is not sufficient to influence steady-state firing rate and, accordingly, CB1R-mediated IPSCs were not likely to be involved in the WIN-2-induced decrease in steady-state firing rate. In many cases, induction of an action potential train sufficient to determine the steady-state firing frequency required that injection currents were much larger than those that had been necessary in preWIN-2 conditions. Current strengths needed were often 100–300 pA, which was higher than current strengths sufficient to elicit a train of spikes in control conditions (10–50 pA). Consistent with this observation, the rheobase, the current needed to induce a single action potential, was significantly higher (644 153%) in WIN-2-treated neurons when compared to that in control conditions (Control, 41.0 8.4 pA; WIN-2, 235.0 37.0 pA; n = 10 from five slices, P = 0.0001, paired t-test; Fig. 6D). As these experiments required quite long recording periods, to ensure that decreases seen in cell firing were not due to cell rundown we ran a separate study. We pre-incubated the brain slice in WIN-2 (10–15 min) and, following this exposure period, patchclamp recordings were initiated. Firing was monitored immediately and after a washout period in which slices were no longer exposed to WIN-2. Under these conditions, firing rates obtained soon after



B

C

Fig. 5. Cannabinoid CB1R antagonist AM251 suppressed WIN-2-induced inhibition of mIPSC within the LDT. (A) Lower (left) and higher (right) gain recordings from a representative cell displaying the effect of WIN-2 on mIPSCs recorded in presence of AM251 and TTX in which it can be seen that there was no change in mIPSCs when WIN-2 was applied in the presence of the receptor antagonist. (B1) The composite mIPSC before (black) (19.7 pA; n = 124 events; black) and after WIN-2 (gray) (18.6 pA; n = 108 events; black) suggest that there was no change in amplitude with WIN-2. (B2 and 3) Cumulative distribution plots of mIPSC (B2) inter-event intervals and (B3) amplitudes in the presence of AM251 + TTX (black) and during (gray) WIN-2 application for the single experiment show that WIN-2 in the presence of the receptor antagonist did not induce any significant changes in the cumulative distribution of inter-event intervals or amplitudes of mIPSCs (K-S test, P > 0.05). (C) As is apparent from these histograms presenting data from the population of neurons examined, in the presence of AM251 + TTX, WIN-2 (C1) did not induce a significant decrease in the frequency of mIPSCs, nor was there (C2) a significant change in the mean mIPSC amplitude, suggesting that the actions of WIN-2 were specific to involvement of the CB1R (two-tailed paired Student’s t-test, with significance assumed at P < 0.05). Error bars are SEM.

the patch was established were found to be reduced from those obtained following the washout period in the same cells in the majority of cases (preincubation in WIN-2, 15.0 3.1 Hz; after washout of WIN-2 with ACSF, 21.0 4.4 Hz; n = 7/7 from five slices, P = 0.0262, paired t-test; Fig. 6C). These data strongly suggest that the effects on firing rate associated with WIN-2 exposure were not due to exhaustion of cellular constituents, nor in eroding patch conditions during the long recordings, and were specific to actions mediated by CB1R activation. WIN-2 elicited a small shift in membrane holding current of the postsynaptic cell and an increase in a tetraethylammonium (TEA)-sensitive potassium conductance After determination that a CB1R agonist induced a significant decrease in cellular excitability within neurons of the LDT, we examined

the possibility that WIN-2 had actions leading to alterations in intrinsic membrane properties such as membrane potential and resistance. In voltage-clamp mode, we observed that WIN-2 induced a small outward current in the majority of cells when clamped at 60 mV, indicating that a conductance had been activated (5.8 0.55 pA; n = 25/35 from 20 slices; Fig. 7, A1). To elucidate the identity of the conductance underlying this small WIN-2-induced shift in holding current, in responding cells we evaluated drug-induced changes in input resistance by stepping the voltage of the cell from 60 to 70 mV and monitoring the current required to achieve this step. This examination showed that the WIN-2-induced outward current was associated with a decrease in membrane resistance (Control, 566 22 MO; WIN-2, 223 29 MO; n = 11/11 from six slices, P < 0.0001, paired t-test; Fig. 7, B1 and 2). The most parsimonious explanation for the mechanism underlying the WIN-2-induced outward current was that a potassium conductance had been activated.



B

C

D

Fig. 6. eCB CB1R agonist WIN-2 reduced the firing frequency of LDT neurons. (A) Representative traces recorded from an individual LDT neuron, showing the firing of action potentials evoked by a depolarising pulse (300 ms, 10 pA) in (A1) control conditions and in (A2) the presence of WIN-2. The WIN-2-induced decrease in steady-state firing frequency of action potentials is clearly evident in this cell which was identified as bNOS-positive, and this effect was apparent in all cells recorded, of which three were identified as bNOS-positive. (A3) Histogram summarising the firing frequency before and after WIN-2 application, from the population of neurons recorded from the LDT, indicating that WIN-2 significantly decreased the steady-state firing frequency. (B) The steady-state firing frequency partially recovered in the presence of AM251 as indicated by this current-clamp recording showing the firing of action potentials of a bNOS-positive LDT neuron during (B1) exposure to WIN-2, and (B2) subsequently during concurrent presence of AM251 in the same cell. Only one cell from this series was recovered for immunohistochemistry. (B3) Histogram showing action potential firing for a population of recorded LDT neurons in the presence of WIN-2 and the recovery of firing rate when AM251 was also included. (C) A significant increase in steady-state firing frequency was observed in a population of neurons within the LDT after washout of WIN-2, as apparent from this histogram from a population of cells which were pre-incubated in WIN-2 before whole-cell recordings were established. (D) In the presence of WIN-2, the rheobase, the amount of current required to induce a single action potential, was increased, indicating further that WIN-2 influences cellular excitability of LDT neurons (two-tailed paired Student’s t-test, with significance assumed at P < 0.05). Error bars are SEM. *P ≤ 0.05; **P ≤ 0.01; #P ≤ 0.001.

To determine whether a potassium conductance was involved in WIN-2-induced actions on these cells, we examined the reversal potential of the WIN-2-induced current to determine whether it was near the calculated equilibrium potential for potassium. We generated I–V input–output curves by stepping the voltage of the cell between 100 and 40 mV from the holding voltage of 60 mV in 10-mV increments and calculating the current necessary to achieve this voltage once a steady-state holding current had been reached (Fig. 7, C1 and 2). Subtraction of control and curves collected after WIN-2 application for the population of cells revealed that the WIN-2-induced current exhibited an average reversal potential of 77.0 1.6 mV (n = 10), which was near the reversal potential for potassium ions calculated under the recording conditions utilised (Ek = 80 mV: with [K+]o 5.0 mm and [K+]i 144.0 mm at RT ~ 16 °C using the Nernst equation). Indicating that the WIN-2-stimulated conductance was mediated by an increase in a potassium ion conductance, we found the WIN-2-activated current was sensitive to the non-selective potassium channel blocker TEA. In the presence of this potassium channel blocker,

there was no detected crossing of the control and TEA I–V curves, and subtraction of control and WIN-2 I–V curves failed to show the reversal of current noted in non-TEA conditions (n = 4; Fig. 7, D1 and 2). Our next step was to determine whether WIN-2 induced changes in steady-state firing during blockade of potassium conductances. In the presence of TEA, which itself altered firing rate by 13 3% from control (Control, 18.0 5.0 Hz; TEA, 16.0 4.1 Hz; n = 3 from two slices, two animals, P = 0.1565, paired t-test; Fig. 7, E1 and 2 and F1), WIN-2 failed to further influence the firing frequency (1 10% change from control; TEA+WIN-2, 17.0 5.5 Hz; n = 3/3, P = 0.5632, paired t-test; Fig. 7, E3 and F2). When compared across a population of cells, the change in steady-state firing induced by WIN-2 applied in the presence of TEA was not significant (4.9 15.0% reduction from control; control in ACSF, 14.0 2.1 Hz; TEA+WIN-2, 13.0 1.9 Hz; n = 12/12 from six slices, P = 0.7289, paired t-test; data not shown). Taken together, our studies using a non-specific channel blocker suggest that WIN-2 application leads to activation of a TEA-sensitive potassium



C

B

D

E

F

Fig. 7. WIN-2 elicited a small TEA-sensitive shift in membrane holding current of the postsynaptic cell and a decrease in resistance. (A1) As apparent from this voltage-clamp recording in a single cell, WIN-2 induced a small outward current in the majority of cells studied. (B1) Underlying the increase in outward current was a decrease in input resistance as shown in this example from another cell recorded in current-clamp mode of the reduced amplitude of the membrane response to injection of hyperpolarising current following application of WIN-2 when compared to the amplitude elicited in control conditions (B1). A significant change in input resistance was noted in a population of cells (B2) (two-tailed paired Student’s t-test, with significance assumed at P < 0.05; error bars are SEM. ##P ≤ 0.0001). (C1) I–V curve in control conditions (black) and following exposure to WIN-2 (gray). (C2) Subtraction of these currents revealed the WIN-2-induced current which exhibited a reversal potential consistent with mediation by a K+ ion conductance. (D) The nonselective K+ channel blocker TEA inhibited the WIN-2-induced current, as evident from these I-V curves showing the responses of a cell to a series of voltage steps recorded in the presence of TEA (black) and TEA+WIN-2 (gray), where no crossing of these curves is apparent. (D2) Subtraction of these currents revealed that, in the presence of TEA, WIN-2 failed to induce a current with a reversal consistent with a K+ conductance (D2). (E) WIN-2 did not induce a significant decrease in steady-state firing when applied in the presence of TEA, as shown in this example from a single cell (E1–3). (F1) While TEA induced a small decrease in firing, this effect was not significant in the group of cells studied. (F2) Across the matched population of cells studied, WIN-2 in the presence of TEA failed to induce a significant change in firing, suggesting the involvement of a WIN-2 activated K+ conductance in alterations of the steady-state firing rate of LDT cells.


3646 N. Soni et al. conductance that results in a small outward current of the cell which is sufficient to reduce cellular excitability.

A

B

C

D

E

F

Immunohistochemical characterisation of recorded LDT cells While considered a cholinergic nucleus, the LDT nucleus is in fact heterogeneous with distinct populations of GABAergic, glutamatergic and cholinergic neurons (Clements & Grant, 1990; Wang & Morales, 2009), as well as a small population of neurons co-expressing both ACh and GABA (Jia et al., 2003; Mieda et al., 2011, 2013). In our study, so as to optimise the numbers of recordings obtained from the principal cell type, cholinergic neurons, we recorded from brain slices containing the medial subdivision of this nucleus, which has been shown to contain the highest concentration (36 2%) of ACh-containing cells (Wang & Morales, 2009), and we targeted large cells with a phenotype characteristic of cholinergic LDT cells (Boucetta & Jones, 2009; Boucetta et al., 2014). To unambiguously identify the phenotype of cells which responded to WIN-2, we performed immunohistochemistry on recorded cells to identify their neurotransmitter content (Vincent et al., 1983; Vincent & Kimura, 1992). However, because the actions of WIN-2 took a long time to develop and we were interested in obtaining cellular and synaptic recoveries after drug effects, recordings in this study needed to be conducted for extended periods of time. Long recording times meant that recovery of cells suitable for immunohistochemistry was extremely low in this study when compared to that of our other studies where shorter durations of recordings were possible (Ishibashi et al., 2009; Christensen et al., 2014). Despite this complication, we successfully recovered nine cells from which data for this study were collected. Seven of these recovered cells were found to be positive for both bNOS and Alexa-594 (showing one of seven in Fig. 8A–D), whereas two of these were bNOS-negative (Fig. 8E and F, showing two of two). All of the seven cells identified as being cholinergic showed a significant decrease in IPSCs (n = 3) or a decrease in firing rate on WIN-2 application (n = 3). One bNOS-positive cell showed a partial recovery in steady-state firing upon AM251 application. While the two cells identified as non-cholinergic were found to be among those few that did not exhibit a change in synaptic currents following WIN-2 exposure, this sample size is too low to conclude that non-cholinergic neurons do not receive input from inhibitory terminals modulated by CB1Rs. In addition, we do not conclude that all cholinergic cells in the LDT exhibit responses noted to CB1R agonist as it is possible that the long recording times dialysed cells, and false negatives for bNOS stain could have been obtained. Regardless, we can infer from our data that activity of a population of cholinergic neurons is affected by stimulation of CB1Rs within the LDT. Presence of CB1R mRNA within the LDT nucleus Currently, the Allen Mice Brain Atlas is the only available reference documenting presence of CB1R mRNA in the LDT. Therefore, we decided to verify the presence of CB1R mRNA in this nucleus. Accordingly, we performed quantitative RT-PCR on total RNA from isolated mouse LDT. Several published reports have suggested a heavy presence of CB1R in rat cerebellum (Herkenham et al., 1991); therefore, total RNA isolated from cerebellum was used as a positive control. b-ACTIN, which is ubiquitous in neurons was used as a housekeeping control. To prevent amplification of contaminant gDNA, if any, we used primers that were designed to span an intron (Acuna-Goycolea et al., 2010). Presence of CB1R transcript in the cerebellum as well as in the LDT of the mouse brain was noted

G

H

Fig. 8. Identification of cholinergic cells using immunohistochemistry after patch-clamp recording within the LDT and RT-PCR within the LDT. (A) Representative LDT neuron from this study viewed under illuminated light optics in which the patch pipette filled with the dye, Alexa-594, is apparent. During voltage- or current-clamp recordings, the dye passively filled the cell from the pipette. (B) View of the same cell under fluorescent optics after the patch pipette had been carefully removed, indicating that the cell can be retained, and visualised in slices. (C and D) Following immunohistochemical staining, Alexa-594-filled recorded cells can (C) be identified under 580 nm optics, and (D) it can be determined whether they are bNOS-positive when viewed under 488 nm, as is apparent in this example. In this way, cells in this study could be identified as cholinergic or non-cholinergic. (E and F) Cells that were negative for bNOS, and hence non-cholinergic, were also recovered; however, in these two cells, actions of WIN-2 on synaptic currents were not detected. (G) Shown is the display of the expression of CB1R (~ 250 bp) PCR products from cDNA isolated from the cerebellum and the LDT displayed on a 2% agarose gel. b-ACTIN (~ 450 bp) was used as an equal loading control and its expression in both brain regions is also displayed. (H) Histogram representing the DCT values of CB1R expression relative to b-ACTIN in the cerebellum and the LDT.


Functional presence of CB1Rs in the LDT 3647 (Fig. 8G). Examination of the relative CB1R transcript levels in the cerebellum and those in the LDT suggested the possibility that there is an approximately two-fold lower expression of CB1R mRNA in the LDT than in cerebellar tissue (Fig. 8H; n = 2 independent samples); however, a more detailed examination of this point would have to be performed before firm conclusions can be drawn. Although the sample size is limited, our RT-PCR studies contribute to the available reference literature documenting the presence of mRNA for CB1R in the LDT. Our RT-PCR data when combined with our electrophysiological findings strongly suggest translation of mRNA into functional CB1R protein in the LDT.

Discussion We provide for the first time strong evidence that CB1R-mediated effects are present in the LDT by showing that a CB1R agonist has synaptic and membrane actions on cells in this nucleus. WIN-2 induced a decrease in the frequency of both sIPSCs and mIPSCs. As a significant effect of WIN-2 on amplitude was only noted for sIPSCs and not mIPSCs, we conclude this effect was mediated by activation of local presynaptic receptors, probably on GABAergic terminals directed to postsynaptic LDT cells. Specificity of CB1R actions was demonstrated by partial blockade of the WIN-2-mediated decrease in sIPSC frequency by a CB1R antagonist, AM251. WIN-2 also decreased the firing frequency of LDT cells, an effect involving CB1Rs as it was reduced by the presence of AM251. WIN-2 was found to induce a small outward current mediated by an increase in a TEA-sensitive potassium conductance which, via a concurrent membrane hyperpolarisation and shunting of input current, probably lead to reductions in cell excitability. In a limited number of recovered cells, we were able to determine that WIN-2 actions extended to cholinergic LDT cells. RT-PCR studies confirmed the presence of mRNA for CB1R within the LDT, and our electrophysiological findings clearly suggest that functional CB1Rs are present in the LDT. Taken together with our immunohistochemical identification of recor-ded cells, our findings suggest CB1R-mediated actions on LDT cells, including a population of cholinergic cells, and presynaptic GABAergic terminals in this nucleus. Further, our data using a CB1R antagonist indicate that CB1Rs are physiologically engaged in the LDT brain slice, which suggests that they are endogenously active in vivo and, therefore, probably involved in processes governed by this nucleus. Pharmacological agents: caveats While fatty-acid endogenous ligands of the CB1R are available, we utilised WIN-2 as most cellular electrophysiological studies have used this agent, and we chose a concentration used by others. WIN-2 is within a structurally unique class of compounds of aminoalkylindoles that bind to the same CB1R as endogenously-produced agonists, such as anandamide and Sn-2-arachidonoyl-glycerol (2-AG), and the plantderived agonist Δ9-tetrahydrocannabinol (Kuster et al., 1993). WIN2, when compared to THC, is a full agonist at the CB1R, displays high affinity (~ 1.9 nM) and binds reversibly (Compton et al., 1992; Kuster et al., 1993). Despite its high affinity, WIN-2, when compared to THC, actions require long periods of time to achieve their maximal effect, making it difficult to compare effects on the same cell in the presence and absence of antagonists in brain slices. We do not believe changes of activity of LDT neurons noted in the present study are a consequence of reported receptor-independent mechanisms (Oz, 2006a,b), as a CB1R antagonist attenuated WIN-2-induced cellular actions. AM251 was chosen as an antagonist for CB1Rs as it has been well documented to be a highly effective and selective receptor ligand

that, as a reverse agonist, readily prevents or reverses CB1R-mediated effects (Gatley et al., 1996). CB1R receptor activation on presynaptic terminals reduced the frequency of IPSCs Our data suggest that presynaptically located CB1Rs mediate a decrease in the release of inhibitory neurotransmitter, as occurs in several other brain regions (Katona et al., 1999; Hoffman & Lupica, 2000; Kreitzer & Regehr, 2001; Wilson & Nicoll, 2001; Diana et al., 2002). Our data suggest that exogenous exposure to CB1R agonists, which would occur upon use of tetrahydrocannabinol-containing products such as marijuana or exposure to eCBs via endogenous production of CB1R agonists, would suppress inhibitory synaptic activity directed to LDT neurons. Endogenous activation of presynaptic CB1Rs in the LDT could be involved in shaping the output of postsynaptic cells as inhibitory conductances, probably via shunting, have been well established to determine spike rate and timing, and their reduction would be expected to alter temporal firing patterns (Gauck & Jaeger, 2000). The localisation of presynaptic CB1Rs would also be expected to determine the effect of their activation on the output on the postsynaptic LDT cell, with perisomatic inhibitory inputs shown to play a role in thresholds for activation and with dendritic inputs positioned to modulate excitatory inputs in other cell types where this has been studied (Qian & Sejnowski, 1990; Miles et al., 1996). Accordingly, actions mediated by endogenous or exogenous activation of presynaptic CB1Rs in the LDT resulting in alterations in IPSC activity would be expected to participate in regulation of neuronal excitability in the postsynaptic cell. While studies suggest that eCBs interact significantly with GABAergic transmission (Wilson & Nicoll, 2001; Sigel et al., 2011), glutamate transmission has also been shown to be affected by neuronal CB1Rs (Takahashi & Linden, 2000; Gerdeman & Lovinger, 2001; Hajos et al., 2001; Huang et al., 2001; Melis et al., 2004; HajDahmane & Shen, 2005; Nemeth et al., 2008), and a recent study suggests a heightened glutamate transmission originating from activation of astrocytic CB1Rs in the brainstem (K€ oszeghy et al., 2014). As we did not discern an effect of WIN-2 on the frequency or amplitude of EPSCs in the majority of cells specifically examined for alterations in glutamate transmission, our data do not support the conclusion that processes underlying a suppression or enhancement of glutamate input following activation of CB1Rs would occur to any great degree within the LDT. However, as we did record a WIN-2-mediated reduction in EPSC frequency in two LDT cells, explicit studies examining this issue need to be conducted to confirm this point, especially in the light of reports that WIN-2-induced changes in excitatory transmission were less sensitive than alterations in inhibitory transmission (Hajos & Freund, 2002; Ohno-Shosaku et al., 2002). Decrease in excitability of postsynaptic LDT neurons Activation of CB1Rs has strong actions on reducing the excitability of LDT neurons as WIN-2 exposure led to an increase in rheobase, reflecting the necessity of injection of larger currents to induce action potential firing. The specificity of this action to CB1R activation was indicated by partial reversal of this effect by AM251. The reversal potential of the WIN-2-induced current and blockade by TEA suggests that the CB1R-induced conductance was mediated by a potassium channel, which inhibited firing of LDT cells via membrane hyperpolarisation and an action on shunting of current. eCBs have been shown in other cell types to modulate the intrinsic excitability of neurons by alterations in membrane resistance (Glickfeld


3648 N. Soni et al. & Scanziani, 2005). Low threshold spiking neocortical neurons were found to exert a self-inhibition via activity-dependent release of eCBs acting to stimulate a potassium conductance which induced membrane hyperpolarisation sufficient to alter the intrinsic excitability of these cells (Bacci et al., 2004). We did not in the present study identify the specific potassium channel activated by WIN-2 beyond determination of its TEA sensitivity; however, CB1R activation has been shown to induce activity of the inwardly rectifying K+ (GIRK) current in other neuronal regions (Henry & Chavkin, 1995), and the presence of GIRK channels on cholinergic LDT neurons (Luebke et al., 1993; Leonard & Llinas, 1994), which mediate membrane hyperpolarisation following stimulation of other G-protein receptors (Luebke et al., 1992, 1993; Leonard & Llinas, 1994; Kohlmeier & Kristiansen, 2010), suggests that activation of the CB1R could exert actions on this channel. However, a precise identification of the TEA-sensitive potassium conductance activated subsequent to CB1R stimulation awaits future investigation. How do the presynaptically-mediated decrease in synaptic inhibition and reduction in postsynaptic membrane excitability interact at a cellular level? Our findings indicate that application of WIN-2 results in an inhibition in excitability of LDT cells which is in contrast to the presynaptically-mediated action which would be expected to lead to enhancement of postsynaptic neuronal excitability. The interaction of these two opposing actions at the cellular level is probably complex, precluding confident predictions of resultant outcomes on the net functioning and output of the postsynaptic cell. However, our data suggest that exposure to a CB1R agonist would profoundly influence synaptic transmission and membrane excitability. Selective activation of the reduction in IPSCs in the absence of postsynaptic inhibition would be expected to result in an increased excitatory response to subsequent depolarisation, thereby producing a CB1R shift in bias towards excitation (Qian & Sejnowski, 1990; Gauck & Jaeger, 2000). If the CB1R-mediated reduction in IPSCs directed to the postsynaptic cell occurs concurrently with the inhibition in postsynaptic excitability, the decrease in IPSCs may be unlikely to override the membrane inhibition and lead to cell firing or otherwise significantly regulate the strength of cell output, a conclusion supported by our findings that bath application of WIN-2 led to decreased firing. However, co-activation of these processes could have a modulatory action including a reduction in membrane shunting, leading to a limit in the duration of inhibition of the cell, altering the synchrony of firing across ensembles of LDT neurons. Distribution of CB1Rs relative to exposure of agonist would determine whether actions are present concurrently or occur in isolation. Distinct actions of CB1Rs on influencing firing and on altering synaptic release have previously been attributed to distinct synaptic and nonsynaptic sites of action (Tsou et al., 1998; Ong & Mackie, 1999; Fawley et al., 2014), suggesting the possibility that receptors located distally to neuronal synapses mediate the decrease in excitability and that these receptors would be activated when levels of CB1R agonist are high which, if produced locally, may occur when LDT firing is high, thereby serving as a brake on excitability. Our findings with AM251 lend some support to this conclusion as we found evidence of an endogenous activation of presynaptic receptors in the brain slice, but did not see an effect of AM251 on firing rates, suggesting receptors mediating postsynaptic cellular excitability were not activated by endogenously available eCBs. Besides differences in localisation of receptor, subtle molecular differences in receptors, such as splice variants of the CB1R which may show ana-

tomical heterogeneity, could confer differential responses to agonists, allowing for further distinction in postsynaptic outcomes dependent on specificity of agonist (Shire et al., 1995; Straiker et al., 2012). Further studies need to be conducted to examine the properties and anatomical localisation of CB1Rs within the LDT. Endogenous cannabinoids within the LDT As blockade of CB1Rs with AM251 resulted in an increase in the frequency of sIPSCs, this suggests that eCBs were being produced and released within the slice. An intriguing question, therefore, is what are the mechanisms underlying endogenous production of agonists of CB1Rs in the LDT? Production of eCBs have been shown to occur following depolarisation of cells by activation of metabotropic glutamate receptors (mGluRs; Glitsch et al., 1996; Morishita et al., 1998; Varma et al., 2001; Puente et al., 2011), metabotropic excitatory muscarinic ACh receptors (mAChRs; Kim et al., 2002; Zhao & Tzounopoulos, 2011) and receptors for the arousal-promoting peptide orexin (Haj-Dahmane & Shen, 2005) which are all present in the LDT (Baghdoyan et al., 1994; Marcus et al., 2001; Burlet et al., 2002; Kohlmeier et al., 2012, 2013) and, when activated, induce depolarisation of postsynaptic LDT cells (Burlet et al., 2002; Kohlmeier et al., 2012, 2013), which might be sufficient to induce production of endogenous eCBs. While ongoing activation in the LDT brain slice of cholinergic or orexinergic transmission has not been noted, a significant degree of glutamate activity originating in large part from local glutamate-containing cells remains in the LDT brain slice even though connections are severed by this preparation (Kohlmeier et al., 2012). Although we conducted most of the IPSC studies in the presence of blockers of ionotropic glutamate receptors, endogenous glutamate released from terminals in the slice may have acted at postsynaptic mGluRs. As activation of mGluRs has been shown in other cell types to result in production of eCBs, activation of mGluRs by glutamate released locally in the LDT within the slice may have been sufficient to activate CB1Rs, thereby reducing inhibitory transmission, a process opposed by AM251 application. Taken together, as we now find functional evidence for the presence of CB1Rs in the LDT and endogenous activation of these receptors in the brain slice, future experiments are being conducted to find out whether activation of metabotropic receptors demonstrated to be present on LDT neurons and shown in other types to mediate increases in eCBs, as well as serve as targets for enhanced glutamate from CB1R stimulation on astrocytes (K€ oszeghy et al., 2014), impacts on cellular LDT functioning via actions at the CB1R. Further, if eCBs are produced endogenously within the LDT in vivo as seen in the brain slice, it is worth considering that presence of eCBs have been shown to activate CB1R-independent mechanisms such as modulation of functional properties of voltage-gated ion channels including Ca2+ channels (Shimasue et al., 1996; Oz et al., 2000), Na+ channels (Nicholson et al., 2003), a variety of types of K+ channels (Oliver et al., 2004; Sade et al., 2006) and ligand-gated ion channels such as nicotinic ACh (Oz et al., 2003, 2004), and glycine receptors (Lozovaya et al., 2005, 2011). Therefore, while actions noted in the present study appeared to be specific to involvement of activation of CB1Rs, endogenous presence of eCBs in the LDT could stimulate a host of actions beyond CB1R-mediated effects. Functional significance of eCB-mediated reduced excitability of LDT neurons In this study, we found that CB1R activation had strong actions on synaptic activity and cellular excitability of LDT neurons,


Functional presence of CB1Rs in the LDT 3649 which suggests the eCB system could play a role in behaviours controlled in part by this nucleus, such as behavioural state and motivated behaviour. While it was not determined that these actions were exclusive to cholinergic LDT neurons, nor that all cholinergic cells would be similarly affected, the presence of bNOS in recovered cells suggested that this effect extended to cholinergic LDT cells and would be present in at least a subpopulation of this cellular phenotype in this nucleus. An important role for ACh and for the cholinergic neurons of the reticular core, which includes ACh-containing neurons of the LDT along with activity of adjacent ACh-containing pedunculopontine tegmental cells, in stimulating cortical activation and generating rapid eye movement sleep has been recognised (Pare et al., 1988; Webster & Jones, 1988; el Mansari et al., 1989; Jones, 1990, 1993; Semba et al., 1990; Steriade, 1993; Santucci et al., 1996; Steriade & McCarley, 2005). Further, projections to the ventral tegmental area and nucleus accumbens originating from cholinergic and glutamatergic cells in this nucleus play a role in drug addiction-associated behaviours (Forster & Blaha, 2000; Omelchenko & Sesack, 2006; Ishibashi et al., 2009; Lammel et al., 2012; Dautan et al., 2014). As reductions in cellular excitability were evoked by bath application of a CB1R agonist, even in the presence of concurrent reductions in inhibitory synaptic activity, it could be speculated that widespread exposure to the agonist would induce in a subset of cholinergic cells the demonstrated postsynaptic action leading to reductions in action potential firing. Accordingly, the decrease in firing of cholinergic neurons upon stimulation of CB1Rs noted in the present study could be expected to reduce output of ACh to target regions of the LDT, which is consistent with behavioural actions of exogenous application of CB1R agonists. For example, exogenous application of cannabinoids, such as marijuana and hashish, are well known to affect arousal levels, cognitive ability and sleep architecture suggestive of alterations of cholinergic tone (Feinberg et al., 1975; Bolla et al., 2002, 2008; Huang et al., 2007; Goonawardena et al., 2011). Recent evidence suggests the presence in the LDT of a mixed population of glutamate and GABAergic neurons exhibiting state-dependent firing, suggesting that they are also involved in modulation of arousal (Boucetta et al., 2014). If actions extend to these cells, the effects of CB1R agonists on these non-cholinergic LDT cells could also play a role in LDT function. Although highly speculative, inhibition of firing of cholinergic LDT neurons, and glutamatergic LDT cells if effects extend to these cells as well, might contribute to the reduced addiction potential shown by this illicitly-used substance when compared to other drugs of abuse such as nicotine, which have been shown to induce robust increases in firing of cholinergic and non-cholinergic LDT cells which would be expected to lead to enhancements of transmission from this nucleus to midbrain target regions (Ishibashi et al., 2009; Christensen et al., 2014). The reduction in inhibitory synaptic transmission noted following CB1R stimulation could be an action invoked by endogenous production of eCBs, perhaps subsequent to activation of metabotropic receptors. The in vivo production could be discrete, limiting the extent of activation by acting at distinct presynaptic sites. The reduction in inhibitory synaptic activity would be expected to shape the excitability and output of the cell. As LDT neurons exhibit behavioural state-related firing (Hallanger et al., 1987; Hallanger & Wainer, 1988; el Mansari et al., 1989; Boucetta & Jones, 2009; Boucetta et al., 2014), if their firing leads to production of eCBs, this production could vary across behavioural state and play a role in initiation and/or maintenance of that behavioural state. Based on findings that CB1Rs can control release of ACh

(Gifford & Ashby, 1996; Gessa et al., 1997; Acquas et al., 2000, 2001; Kathmann et al., 2001), a role for eCBs in generation of state has been previously proposed (Murillo-Rodriguez et al., 1998; Murillo-Rodriguez, 2008). While evidence of inhibition of ACh release has been presented, rises noted in the hippocampus and cortical regions via a CB1R-mediated mechanism (Acquas et al., 2000, 2001) have led to the suggestion that CB1R-mediated actions in the pons, stimulated by endogenous production of eCBs, could lead to activation of cholinergic LDT neurons, leading to brainstem rises in ACh and thereby, via activation of the caudal reticular formation, participate in generation of the sleep states (Murillo-Rodriguez et al., 2011). The presynaptically invoked CB1R-mediated decrease in inhibition seen in the present study could play a role in CB1Rmediated increases in release of ACh, along with glutamate, when LDT activity is high via a feed-forward mechanism. Further studies are being conducted examining the source of endogenous production of eCBs in the LDT of the brain slice, as well as to further elucidate cellular effects of endogenously- and exogenously-mediated CB1R stimulation within the LDT, in order to gain a more complete understanding of the role played by eCB transmission and CB1Rs in processes governed by this nucleus.

Acknowledgements This work was supported by the Drug Research Academy (DRA) at the University of Copenhagen, Copenhagen, Denmark and by the Philip Morris External Research Program, USA. The authors acknowledge Mr Jason Allen Teem for his excellent technical assistance in immunohistochemistry and for preparation of brain slices. We are also thankful to Professor Chunaram Choudhary (The Novo Nordisk Foundation Center for Protein Research, University of Copenhagen, Copenhagen) for providing facilities to perform molecular biology studies. We would also like to thank the reviewers and editors at EJN for their helpful suggestions that resulted in a substantial improvement of the discussion of this manuscript. The authors declare they have no conflicts of interest, financial or otherwise, regarding this manuscript.

Abbreviations ACh, acetylcholine; AM251, 1-(2,4-dichlorophenyl)-5-(4-iodophenyl)-4-methylN-(1-piperidyl) pyrazole-3-carboxamide; AP-5, D(2)-2-amino-5-phosphonopentanoic acid; bNOS, brain nitric oxide synthase; CB1R, cannabinoid type-1 receptor; cDNA, complementary DNA; ChAT, choline acetyltransferase; CNQX, 6-cyano-7-nitroquinoxaline-2,3-dione; eCB, endocannabinoid; EPSC, excitatory postsynaptic current; gDNA, genomic DNA; K-S, Kolmogorov– Smirnov; LDT, laterodorsal tegmental nucleus; mEPSC, miniature EPSC; mIPSC, miniature inhibitory postsynaptic current; RT-PCR, real-time PCR; sEPSC, spontaneous EPSC; sIPSC, spontaneous IPSC; TEA, tetraethylammonium; TTX, tetrodotoxin; WIN-2, WIN55,212-2 [(R)-(+)-[2,3-dihydro-5methyl-3-(4-morpholinylmethyl)pyrrolo[1,2,3-de]-1,4-benzoxazin-6-yl]-1naphthalenylmethanone mesylate].

References Acquas, E., Pisanu, A., Marrocu, P. & Di Chiara, G. (2000) Cannabinoid CB (1) receptor agonists increase rat cortical and hippocampal acetylcholine release in vivo. Eur. J. Pharmacol., 401, 179–185. Acquas, E., Pisanu, A., Marrocu, P., Goldberg, S.R. & Di Chiara, G. (2001) Delta9-tetrahydrocannabinol enhances cortical and hippocampal acetylcholine release in vivo: a microdialysis study. Eur. J. Pharmacol., 419, 155–161. Acuna-Goycolea, C., Obrietan, K. & van den Pol, A.N. (2010) Cannabinoids excite circadian clock neurons. J. Neurosci., 30, 10061–10066. Akerman, S., Kaube, H. & Goadsby, P.J. (2004) Anandamide is able to inhibit trigeminal neurons using an in vivo model of trigeminovascularmediated nociception. J. Pharmacol. Exp. Ther., 309, 56–63. Allen Mouse Brain Atlas (2012) Allen Institute for Brain Science. Allen Mouse Brain Atlas [Internet] Available http://mouse.brain-map.org/.


3650 N. Soni et al. Bacci, A., Huguenard, J.R. & Prince, D.A. (2004) Long-lasting self-inhibition of neocortical interneurons mediated by endocannabinoids. Nature, 431, 312–316. Baghdoyan, H.A., Mallios, V.J., Duckrow, R.B. & Mash, D.C. (1994) Localization of muscarinic receptor subtypes in brain stem areas regulating sleep. NeuroReport, 5, 1631–1634. Bolla, K.I., Brown, K., Eldreth, D., Tate, K. & Cadet, J.L. (2002) Doserelated neurocognitive effects of marijuana use. Neurology, 59, 1337– 1343. Bolla, K.I., Lesage, S.R., Gamaldo, C.E., Neubauer, D.N., Funderburk, F.R., Cadet, J.L., David, P.M., Verdejo Garcia, A. & Benbrook, A.R. (2008) Sleep disturbance in heavy marijuana users. Sleep, 31, 901–908. Boucetta, S. & Jones, B.E. (2009) Activity profiles of cholinergic and intermingled GABAergic and putative glutamatergic neurons in the pontomesencephalic tegmentum of urethane-anesthetized rats. J. Neurosci., 29, 4664–4674. Boucetta, S., Cisse, Y., Mainville, L., Morales, M. & Jones, B.E. (2014) Discharge profiles across the sleep-waking cycle of identified cholinergic, GABAergic, and glutamatergic neurons in the pontomesencephalic tegmentum of the rat. J. Neurosci., 34, 4708–4727. Burlet, S., Tyler, C.J. & Leonard, C.S. (2002) Direct and indirect excitation of laterodorsal tegmental neurons by Hypocretin/Orexin peptides: implications for wakefulness and narcolepsy. J. Neurosci., 22, 2862– 2872. Caulfield, M.P. & Brown, D.A. (1992) Cannabinoid receptor agonists inhibit Ca current in NG108-15 neuroblastoma cells via a pertussis toxin-sensitive mechanism. Brit. J. Pharmacol., 106, 231–232. Christensen, M.H., Ishibashi, M., Nielsen, M.L., Leonard, C.S. & Kohlmeier, K.A. (2014) Age-related changes in nicotine response of cholinergic and non-cholinergic laterodorsal tegmental neurons: Implications for the heightened adolescent susceptibility to nicotine addiction. Neuropharmacology, 85, 263–283. Clements, J.R. & Grant, S. (1990) Glutamate-like immunoreactivity in neurons of the laterodorsal tegmental and pedunculopontine nuclei in the rat. Neurosci. Lett., 120, 70–73. Compton, D.R., Gold, L.H., Ward, S.J., Balster, R.L. & Martin, B.R. (1992) Aminoalkylindole analogs: cannabimimetic activity of a class of compounds structurally distinct from delta 9-tetrahydrocannabinol. J. Pharmacol. Exp. Ther., 263, 1118–1126. Cornwall, J., Cooper, J.D. & Phillipson, O.T. (1990) Afferent and efferent connections of the laterodorsal tegmental nucleus in the rat 2. Brain Res. Bull., 25, 271–284. Dautan, D., Huerta-Ocampo, I., Witten, I.B., Deisseroth, K., Bolam, J.P., Gerdjikov, T. & Mena-Segovia, J. (2014) A major external source of cholinergic innervation of the striatum and nucleus accumbens originates in the brainstem. J. Neurosci., 34, 4509–4518. Devane, W.A., Hanus, L., Breuer, A., Pertwee, R.G., Stevenson, L.A., Griffin, G., Gibson, D., Mandelbaum, A., Etinger, A. & Mechoulam, R. (1992) Isolation and structure of a brain constituent that binds to the cannabinoid receptor. Science, 258, 1946–1949. Diana, M.A., Levenes, C., Mackie, K. & Marty, A. (2002) Short-term retrograde inhibition of GABAergic synaptic currents in rat Purkinje cells is mediated by endogenous cannabinoids. J. Neurosci., 22, 200–208. Fawley, J.A., Hofmann, M.E. & Andresen, M.C. (2014) Cannabinoid 1 and transient receptor potential vanilloid 1 receptors discretely modulate evoked glutamate separately from spontaneous glutamate transmission. J. Neurosci., 34, 8324–8332. Feinberg, I., Jones, R., Walker, J.M., Cavness, C. & March, J. (1975) Effects of high dosage delta-9-tetrahydrocannabinol on sleep patterns in man. Clin. Pharmacol. Ther., 17, 458–466. Forster, G.L. & Blaha, C.D. (2000) Laterodorsal tegmental stimulation elicits dopamine efflux in the rat nucleus accumbens by activation of acetylcholine and glutamate receptors in the ventral tegmental area. Eur. J. Neurosci., 12, 3596–3604. Gaoni, Y. & Mechoulam, R. (1964) Isolation, structure, and partial synthesis of an active constituent of Hashish. J. Am. Chem. Soc., 86, 1646. Gatley, S.J., Gifford, A.N., Volkow, N.D., Lan, R. & Makriyannis, A. (1996) 123I-labeled AM251: a radioiodinated ligand which binds in vivo to mouse brain cannabinoid CB1 receptors. Eur. J. Pharmacol., 307, 331– 338. Gauck, V. & Jaeger, D. (2000) The control of rate and timing of spikes in the deep cerebellar nuclei by inhibition. J. Neurosci., 20, 3006–3016. Gerdeman, G. & Lovinger, D.M. (2001) CB1 cannabinoid receptor inhibits synaptic release of glutamate in rat dorsolateral striatum 5. J. Neurophysiol., 85, 468–471.

Gessa, G.L., Mascia, M.S., Casu, M.A. & Carta, G. (1997) Inhibition of hippocampal acetylcholine release by cannabinoids: reversal by SR 141716A. Eur. J. Pharmacol., 327, R1–R2. Gifford, A.N. & Ashby, C.R. Jr. (1996) Electrically evoked acetylcholine release from hippocampal slices is inhibited by the cannabinoid receptor agonist, WIN 55212-2, and is potentiated by the cannabinoid antagonist, SR 141716A. J. Pharmacol. Exp. Ther., 277, 1431–1436. Glickfeld, L.L. & Scanziani, M. (2005) Self-administering cannabinoids 5. Trends Neurosci., 28, 341–343. Glitsch, M., Llano, I. & Marty, A. (1996) Glutamate as a candidate retrograde messenger at interneurone-Purkinje cell synapses of rat cerebellum. J. Physiol., 497(Pt 2), 531–537. Goonawardena, A.V., Plano, A., Robinson, L., Platt, B., Hampson, R.E. & Riedel, G. (2011) A pilot study into the effects of the CB1 cannabinoid receptor agonist WIN55,212-2 or the antagonist/inverse agonist AM251 on sleep in rats. Sleep Disord., 2011, 178469. Guo, J. & Ikeda, S.R. (2004) Endocannabinoids modulate N-type calcium channels and G-protein-coupled inwardly rectifying potassium channels via CB1 cannabinoid receptors heterologously expressed in mammalian neurons. Mol. Pharmacol., 65, 665–674. Haj-Dahmane, S. & Shen, R.Y. (2005) The wake-promoting peptide orexinB inhibits glutamatergic transmission to dorsal raphe nucleus serotonin neurons through retrograde endocannabinoid signaling. J. Neurosci., 25, 896–905. Hajos, N. & Freund, T.F. (2002) Pharmacological separation of cannabinoid sensitive receptors on hippocampal excitatory and inhibitory fibers. Neuropharmacology, 43, 503–510. Hajos, N., Ledent, C. & Freund, T.F. (2001) Novel cannabinoid-sensitive receptor mediates inhibition of glutamatergic synaptic transmission in the hippocampus. Neuroscience, 106, 1–4. Hallanger, A.E. & Wainer, B.H. (1988) Ascending projections from the pedunculopontine tegmental nucleus and the adjacent mesopontine tegmentum in the rat. J. Comp. Neurol., 274, 483–515. Hallanger, A.E., Levey, A.I., Lee, H.J., Rye, D.B. & Wainer, B.H. (1987) The origins of cholinergic and other subcortical afferents to the thalamus in the rat. J. Comp. Neurol., 262, 105–124. Henry, D.J. & Chavkin, C. (1995) Activation of inwardly rectifying potassium channels (GIRK1) by co-expressed rat brain cannabinoid receptors in Xenopus oocytes. Neurosci. Lett., 186, 91–94. Herkenham, M., Groen, B.G., Lynn, A.B., De Costa, B.R. & Richfield, E.K. (1991) Neuronal localization of cannabinoid receptors and second messengers in mutant mouse cerebellum. Brain Res., 552, 301–310. Hoffman, A.F. & Lupica, C.R. (2000) Mechanisms of cannabinoid inhibition of GABA(A) synaptic transmission in the hippocampus. J. Neurosci., 20, 2470–2479. Hohmann, A.G., Briley, E.M. & Herkenham, M. (1999) Pre- and postsynaptic distribution of cannabinoid and mu opioid receptors in rat spinal cord. Brain Res., 822, 17–25. Huang, C.C., Lo, S.W. & Hsu, K.S. (2001) Presynaptic mechanisms underlying cannabinoid inhibition of excitatory synaptic transmission in rat striatal neurons. J. Physiol., 532, 731–748. Huang, H., Acuna-Goycolea, C., Li, Y., Cheng, H.M., Obrietan, K. & van den Pol, A.N. (2007) Cannabinoids excite hypothalamic melanin-concentrating hormone but inhibit hypocretin/orexin neurons: implications for cannabinoid actions on food intake and cognitive arousal. J. Neurosci., 27, 4870– 4881. Ishibashi, M., Leonard, C.S. & Kohlmeier, K.A. (2009) Nicotinic activation of laterodorsal tegmental neurons: implications for addiction to nicotine. Neuropsychopharmacology, 34, 2529–2547. Jia, H.G., Yamuy, J., Sampogna, S., Morales, F.R. & Chase, M.H. (2003) Colocalization of gamma-aminobutyric acid and acetylcholine in neurons in the laterodorsal and pedunculopontine tegmental nuclei in the cat: a light and electron microscopic study. Brain Res., 992, 205–219. Jones, B.E. (1990) Immunohistochemical study of choline acetyltransferaseimmunoreactive processes and cells innervating the pontomedullary reticular formation in the rat. J. Comp. Neurol., 295, 485–514. Jones, B.E. (1993) The organization of central cholinergic systems and their functional importance in sleep-waking states. Prog. Brain Res., 98, 61–71. Kathmann, M., Weber, B., Zimmer, A. & Schlicker, E. (2001) Enhanced acetylcholine release in the hippocampus of cannabinoid CB(1) receptor-deficient mice. Brit. J. Pharmacol., 132, 1169–1173. Katona, I., Sperlagh, B., Sik, A., Kafalvi, A., Vizi, E.S., Mackie, K. & Freund, T.F. (1999) Presynaptically located CB1 cannabinoid receptors regulate GABA release from axon terminals of specific hippocampal interneurons 28. J. Neurosci., 19, 4544–4558.


Functional presence of CB1Rs in the LDT 3651 Kim, J., Isokawa, M., Ledent, C. & Alger, B.E. (2002) Activation of muscarinic acetylcholine receptors enhances the release of endogenous cannabinoids in the hippocampus. J. Neurosci., 22, 10182–10191. Kohlmeier, K.A. & Kristiansen, U. (2010) GABAergic actions on cholinergic laterodorsal tegmental neurons: implications for control of behavioral state. Neuroscience, 171, 812–829. Kohlmeier, K.A., Soja, P.J. & Kristensen, M.P. (2006) Disparate cholinergic currents in rat principal trigeminal sensory nucleus neurons mediated by M1 and M2 receptors: a possible mechanism for selective gating of afferent sensory neurotransmission. Eur. J. Neurosci., 23, 3245–3258. Kohlmeier, K.A., Ishibashi, M., Wess, J., Bickford, M.E. & Leonard, C.S. (2012) Knockouts reveal overlapping functions of M(2) and M(4) muscarinic receptors and evidence for a local glutamatergic circuit within the laterodorsal tegmental nucleus. J. Neurophysiol., 108, 2751–2766. Kohlmeier, K.A., Christensen, M.H., Kristensen, M.P. & Kristiansen, U. (2013) Pharmacological evidence of functional inhibitory metabotrophic glutamate receptors on mouse arousal-related cholinergic laterodorsal tegmental neurons. Neuropharmacology, 66, 99–113. Kovacs, A., Bıro, T., Sz€ucs, P., Vincze, J., Hegyi, Z., Antal, K€oszeghy, A., M. & Pal, B. (2014) Endocannabinoid signaling modulates neurons of the pedunculopontine nucleus (PPN) via astrocytes. Brain Struct. Funct., doi: 10.1007/s00429-014-0842-5. [Epub ahead of print]. Kreitzer, A.C. & Regehr, W.G. (2001) Cerebellar depolarization-induced suppression of inhibition is mediated by endogenous cannabinoids. J. Neurosci., 21, RC174. Kuster, J.E., Stevenson, J.I., Ward, S.J., D’Ambra, T.E. & Haycock, D.A. (1993) Aminoalkylindole binding in rat cerebellum: selective displacement by natural and synthetic cannabinoids. J. Pharmacol. Exp. Ther., 264, 1352–1363. Lammel, S., Lim, B.K., Ran, C., Huang, K.W., Betley, M.J., Tye, K.M., Deisseroth, K. & Malenka, R.C. (2012) Input-specific control of reward and aversion in the ventral tegmental area. Nature, 491, 212–217. Leonard, C.S. & Llinas, R. (1994) Serotonergic and cholinergic inhibition of mesopontine cholinergic neurons controlling REM sleep: an in vitro electrophysiological study. Neuroscience, 59, 309–330. Lodge, D.J. & Grace, A.A. (2006) The laterodorsal tegmentum is essential for burst firing of ventral tegmental area dopamine neurons. Proc. Natl. Acad. Sci. USA, 103, 5167–5172. Lozovaya, N., Yatsenko, N., Beketov, A., Tsintsadze, T. & Burnashev, N. (2005) Glycine receptors in CNS neurons as a target for nonretrograde action of cannabinoids. J. Neurosci., 25, 7499–7506. Lozovaya, N., Mukhtarov, M., Tsintsadze, T., Ledent, C., Burnashev, N. & Bregestovski, P. (2011) Frequency-dependent cannabinoid receptorindependent modulation of glycine receptors by endocannabinoid 2-AG. Front. Mol. Neurosci., 4, 13. Luebke, J.I., Greene, R.W., Semba, K., Kamondi, A., McCarley, R.W. & Reiner, P.B. (1992) Serotonin hyperpolarizes cholinergic low-threshold burst neurons in the rat laterodorsal tegmental nucleus in vitro. Proc. Natl. Acad. Sci. USA, 89, 743–747. Luebke, J.I., McCarley, R.W. & Greene, R.W. (1993) Inhibitory action of muscarinic agonists on neurons in the rat laterodorsal tegmental nucleus in vitro. J. Neurophysiol., 70, 2128–2135. Mackie, K. & Hille, B. (1992) Cannabinoids inhibit N-type calcium channels in neuroblastoma-glioma cells. Proc. Natl. Acad. Sci. USA, 89, 3825–3829. el Mansari, M., Sakai, K. & Jouvet, M. (1989) Unitary characteristics of presumptive cholinergic tegmental neurons during the sleep-waking cycle in freely moving cats. Exp. Brain Res., 76, 519–529. Marcus, J.N., Aschkenasi, C.J., Lee, C.E., Chemelli, R.M., Saper, C.B., Yanagisawa, M. & Elmquist, J.K. (2001) Differential expression of orexin receptors 1 and 2 in the rat brain. J. Comp. Neurol., 435, 6–25. Matsuda, L.A., Lolait, S.J., Brownstein, M.J., Young, A.C. & Bonner, T.I. (1990) Structure of a cannabinoid receptor and functional expression of the cloned cDNA. Nature, 346, 561–564. Mechoulam, R., Shani, A., Edery, H. & Grunfeld, Y. (1970) Chemical basis of hashish activity. Science, 169, 611–612. Melis, M., Pistis, M., Perra, S., Muntoni, A.L., Pillolla, G. & Gessa, G.L. (2004) Endocannabinoids mediate presynaptic inhibition of glutamatergic transmission in rat ventral tegmental area dopamine neurons through activation of CB1 receptors. J. Neurosci., 24, 53–62. Mieda, M., Hasegawa, E., Kisanuki, Y.Y., Sinton, C.M., Yanagisawa, M. & Sakurai, T. (2011) Differential roles of orexin receptor-1 and -2 in the regulation of non-REM and REM sleep. J. Neurosci., 31, 6518–6526. Mieda, M., Tsujino, N. & Sakurai, T. (2013) Differential roles of orexin receptors in the regulation of sleep/wakefulness. Front. Endocrinol., 4, 57.

Miles, R., Toth, K., Gulyas, A.I., Hajos, N. & Freund, T.F. (1996) Differences between somatic and dendritic inhibition in the hippocampus. Neuron, 16, 815–823. Morishita, W., Kirov, S.A. & Alger, B.E. (1998) Evidence for metabotropic glutamate receptor activation in the induction of depolarization-induced suppression of inhibition in hippocampal CA1. J. Neurosci., 18, 4870–4882. Murillo-Rodriguez, E. (2008) The role of the CB1 receptor in the regulation of sleep. Prog. Neuropsychopharmacol. Biol. Psychiat., 32, 1420– 1427. Murillo-Rodriguez, E., Sanchez-Alavez, M., Navarro, L., Martinez-Gonzalez, D., Drucker-Colin, R. & Prospero-Garcia, O. (1998) Anandamide modulates sleep and memory in rats. Brain Res., 812, 270–274. Murillo-Rodriguez, E., Poot-Ake, A., Arias-Carrion, O., Pacheco-Pantoja, E., Fuente-Ortegon Ade, L. & Arankowsky-Sandoval, G. (2011) The emerging role of the endocannabinoid system in the sleep-wake cycle modulation. Cent. Nerv. Syst. Agents Med. Chem., 11, 189–196. Musella, A., Sepman, H., Mandolesi, G., Gentile, A., Fresegna, D., Haji, N., Conrad, A., Lutz, B., Maccarrone, M. & Centonze, D. (2014) Pre- and postsynaptic type-1 cannabinoid receptors control the alterations of glutamate transmission in experimental autoimmune encephalomyelitis. Neuropharmacology, 79, 567–572. Neher, E. (1992) Correction for liquid junction potentials in patch clamp experiments. Method. Enzymol., 207, 123–131. Nelson, C.L., Wetter, J.B., Milovanovic, M. & Wolf, M.E. (2007) The laterodorsal tegmentum contributes to behavioral sensitization to amphetamine. Neuroscience, 146, 41–49. Nemeth, B., Ledent, C., Freund, T.F. & Hajos, N. (2008) CB1 receptordependent and -independent inhibition of excitatory postsynaptic currents in the hippocampus by WIN 55,212-2. Neuropharmacology, 54, 51–57. Nicholson, R.A., Liao, C., Zheng, J., David, L.S., Coyne, L., Errington, A.C., Singh, G. & Lees, G. (2003) Sodium channel inhibition by anandamide and synthetic cannabimimetics in brain. Brain Res., 978, 194–204. Nutt, D., King, L.A., Saulsbury, W. & Blakemore, C. (2007) Development of a rational scale to assess the harm of drugs of potential misuse. Lancet, 369, 1047–1053. Nutt, D.J., King, L.A., Phillips, L.D. & Independent Scientific Committee on Drugs (2010) Drug harms in the UK: a multicriteria decision analysis. Lancet, 376, 1558–1565. Oakman, S.A., Faris, P.L., Kerr, P.E., Cozzari, C. & Hartman, B.K. (1995) Distribution of pontomesencephalic cholinergic neurons projecting to substantia nigra differs significantly from those projecting to ventral tegmental area 4. J. Neurosci., 15, 5859–5869. Ohno-Shosaku, T., Tsubokawa, H., Mizushima, I., Yoneda, N., Zimmer, A. & Kano, M. (2002) Presynaptic cannabinoid sensitivity is a major determinant of depolarization-induced retrograde suppression at hippocampal synapses. J. Neurosci., 22, 3864–3872. Oliver, D., Lien, C.C., Soom, M., Baukrowitz, T., Jonas, P. & Fakler, B. (2004) Functional conversion between A-type and delayed rectifier K+ channels by membrane lipids. Science, 304, 265–270. Omelchenko, N. & Sesack, S.R. (2006) Cholinergic axons in the rat ventral tegmental area synapse preferentially onto mesoaccumbens dopamine neurons. J. Comp. Neurol., 494, 863–875. Ong, W.Y. & Mackie, K. (1999) A light and electron microscopic study of the CB1 cannabinoid receptor in primate brain. Neuroscience, 92, 1177– 1191. Oz, M. (2006a) Receptor-independent actions of cannabinoids on cell membranes: focus on endocannabinoids. Pharmacol. Therapeut., 111, 114–144. Oz, M. (2006b) Receptor-independent effects of endocannabinoids on ion channels. Curr. Pharm. Design., 12, 227–239. Oz, M., Tchugunova, Y.B. & Dunn, S.M. (2000) Endogenous cannabinoid anandamide directly inhibits voltage-dependent Ca(2+) fluxes in rabbit T-tubule membranes. Eur. J. Pharmacol., 404, 13–20. Oz, M., Ravindran, A., Diaz-Ruiz, O., Zhang, L. & Morales, M. (2003) The endogenous cannabinoid anandamide inhibits alpha7 nicotinic acetylcholine receptor-mediated responses in Xenopus oocytes. J. Pharmacol. Exp. Ther., 306, 1003–1010. Oz, M., Zhang, L., Ravindran, A., Morales, M. & Lupica, C.R. (2004) Differential effects of endogenous and synthetic cannabinoids on alpha7-nicotinic acetylcholine receptor-mediated responses in Xenopus Oocytes. J. Pharmacol. Exp. Ther., 310, 1152–1160. Pare, D., Smith, Y., Parent, A. & Steriade, M. (1988) Projections of brainstem core cholinergic and non-cholinergic neurons of cat to intralaminar and reticular thalamic nuclei. Neuroscience, 25, 69–86.


3652 N. Soni et al. Pitler, T.A. & Alger, B.E. (1994) Depolarization-induced suppression of GABAergic inhibition in rat hippocampal pyramidal cells: G protein involvement in a presynaptic mechanism 5. Neuron, 13, 1447– 1455. Puente, N., Cui, Y., Lassalle, O., Lafourcade, M., Georges, F., Venance, L., Grandes, P. & Manzoni, O.J. (2011) Polymodal activation of the endocannabinoid system in the extended amygdala. Nat. Neurosci., 14, 1542– 1547. Qian, N. & Sejnowski, T.J. (1990) When is an inhibitory synapse effective?. Proc. Natl. Acad. Sci. USA, 87, 8145–8149. Roberto, M., Cruz, M., Bajo, M., Siggins, G.R., Parsons, L.H. & Schweitzer, P. (2010) The endocannabinoid system tonically regulates inhibitory transmission and depresses the effect of ethanol in central amygdala. Neuropsychopharmacology, 35, 1962–1972. Sade, H., Muraki, K., Ohya, S., Hatano, N. & Imaizumi, Y. (2006) Activation of large-conductance, Ca2+-activated K+ channels by cannabinoids. Am. J. Physiol.-Cell Ph., 290, C77–C86. Salio, C., Fischer, J., Franzoni, M.F., Mackie, K., Kaneko, T. & Conrath, M. (2001) CB1-cannabinoid and mu-opioid receptor co-localization on postsynaptic target in the rat dorsal horn. NeuroReport, 12, 3689–3692. Salio, C., Fischer, J., Franzoni, M.F. & Conrath, M. (2002) Pre- and postsynaptic localizations of the CB1 cannabinoid receptor in the dorsal horn of the rat spinal cord. Neuroscience, 110, 755–764. Santucci, V., Storme, J.J., Soubrie, P. & Le Fur, G. (1996) Arousal-enhancing properties of the CB1 cannabinoid receptor antagonist SR 141716A in rats as assessed by electroencephalographic spectral and sleep-waking cycle analysis. Life Sci., 58, PL103–PL110. Schultz, W. (1998) Predictive reward signal of dopamine neurons. J. Neurophysiol., 80, 1–27. Semba, K., Reiner, P.B. & Fibiger, H.C. (1990) Single cholinergic mesopontine tegmental neurons project to both the pontine reticular formation and the thalamus in the rat. Neuroscience, 38, 643–654. Shimasue, K., Urushidani, T., Hagiwara, M. & Nagao, T. (1996) Effects of anandamide and arachidonic acid on specific binding of (+) -PN200-110, diltiazem and (-) -desmethoxyverapamil to L-type Ca2+ channel. Eur. J. Pharmacol., 296, 347–350. Shire, D., Carillon, C., Kaghad, M., Calandra, B., Rinaldi-Carmona, M., Le Fur, G., Caput, D. & Ferrara, P. (1995) An amino-terminal variant of the

central cannabinoid receptor resulting from alternative splicing. J. Biol. Chem., 270, 3726–3731. Sigel, E., Baur, R., Racz, I., Marazzi, J., Smart, T.G., Zimmer, A. & Gertsch, J. (2011) The major central endocannabinoid directly acts at GABA(A) receptors. Proc. Natl. Acad. Sci. USA, 108, 18150–18155. Steriade, M. (1993) Cholinergic blockage of network- and intrinsically generated slow oscillations promotes waking and REM sleep activity patterns in thalamic and cortical neurons. Prog. Brain Res., 98, 345–355. Steriade, M. & McCarley, R. (2005) Neuronal Activities in Brainstem and Basal Forebrain Structures Controlling Waking and Sleep States Brain Control of Wakefulness and Sleep. Springer, US, pp. 381–416. Straiker, A., Wager-Miller, J., Hutchens, J. & Mackie, K. (2012) Differential signalling in human cannabinoid CB1 receptors and their splice variants in autaptic hippocampal neurones. Brit. J. Pharmacol., 165, 2660–2671. Takahashi, K.A. & Linden, D.J. (2000) Cannabinoid receptor modulation of synapses received by cerebellar Purkinje cells. J. Neurophysiol., 83, 1167–1180. Tsou, K., Brown, S., Sanudo-Pena, M.C., Mackie, K. & Walker, J.M. (1998) Immunohistochemical distribution of cannabinoid CB1 receptors in the rat central nervous system. Neuroscience, 83, 393–411. Varma, N., Carlson, G.C., Ledent, C. & Alger, B.E. (2001) Metabotropic glutamate receptors drive the endocannabinoid system in hippocampus. J. Neurosci., 21, RC188. Vincent, S.R. & Kimura, H. (1992) Histochemical mapping of nitric oxide synthase in the rat brain. Neuroscience, 46, 755–784. Vincent, S.R., Satoh, K., Armstrong, D.M. & Fibiger, H.C. (1983) NADPHdiaphorase: a selective histochemical marker for the cholinergic neurons of the pontine reticular formation. Neurosci. Lett., 43, 31–36. Wang, H.L. & Morales, M. (2009) Pedunculopontine and laterodorsal tegmental nuclei contain distinct populations of cholinergic, glutamatergic and GABAergic neurons in the rat. Eur. J. Neurosci., 29, 340–358. Webster, H.H. & Jones, B.E. (1988) Neurotoxic lesions of the dorsolateral pontomesencephalic tegmentum-cholinergic cell area in the cat. II. Effects upon sleep-waking states. Brain Res., 458, 285–302. Wilson, R.I. & Nicoll, R.A. (2001) Endogenous cannabinoids mediate retrograde signalling at hippocampal synapses. Nature, 410, 588–592. Zhao, Y. & Tzounopoulos, T. (2011) Physiological activation of cholinergic inputs controls associative synaptic plasticity via modulation of endocannabinoid signaling. J. Neurosci., 31, 3158–3168.


Copyright of European Journal of Neuroscience is the property of Wiley-Blackwell and its content may not be copied or emailed to multiple sites or posted to a listserv without the copyright holder's express written permission. However, users may print, download, or email articles for individual use.

The role of the laterodorsal tegmental nucleus of the rat in experimental seizures.

Afferent and efferent connections of the laterodorsal tegmental nucleus in the rat.

Ultrastructure of cholinergic neurons in the laterodorsal tegmental nucleus of the rat: interaction with catecholamine fibers.

The role of the laterodorsal tegmental nucleus in methamphetamine conditioned place preference and locomotor activity.

Critical role of cholinergic transmission from the laterodorsal tegmental nucleus to the ventral tegmental area in cocaine-induced place preference.

Cannabinoid CB1 receptors mediate the effects of dipyrone.

Extracellular characteristics of putative cholinergic neurons in the rat laterodorsal tegmental nucleus.

Firing of 'possibly' cholinergic neurons in the rat laterodorsal tegmental nucleus during sleep and wakefulness.

Effects of selective toxic lesions of cholinergic neurons of the laterodorsal tegmental nucleus on experimental seizures.

Cardiorespiratory anomalies in mice lacking CB1 cannabinoid receptors.

A restricted population of CB1 cannabinoid receptors with neuroprotective activity.

Basolateral amygdala CB1 cannabinoid receptors mediate nicotine-induced place preference.

Cannabinoid CB1 receptors in the dorsal hippocampus and prelimbic medial prefrontal cortex modulate anxiety-like behavior in rats: additional evidence.

Increased latencies to initiate cocaine self-administration following laterodorsal tegmental nucleus lesions.

Cannabinoid Receptors CB1 and CB2 Modulate the Electroretinographic Waves in Vervet Monkeys.

Blockade of cannabinoid CB1 and CB2 receptors does not prevent the antipruritic effect of systemic paracetamol.

Cannabinoid CB1 receptors activation and coactivation with D2 receptors modulate GABAergic neurotransmission in the globus pallidus and increase motor asymmetry.

Glutamate-like immunoreactivity in neurons of the laterodorsal tegmental and pedunculopontine nuclei in the rat.

Evidence for the presence of N-methyl-D-aspartate receptors in the ventral tegmental area of the rat: an electrophysiological in vitro study.

Pharmacological Blockade of Cannabinoid CB1 Receptors in Diet-Induced Obesity Regulates Mitochondrial Dihydrolipoamide Dehydrogenase in Muscle.

Cannabinoid type-1 receptors in the paraventricular nucleus of the hypothalamus inhibit stimulated food intake.

Clinical Significance of Cannabinoid Receptors CB1 and CB2 Expression in Human Malignant and Benign Thyroid Lesions.

CB1 cannabinoid receptors are involved in neuroleptic-induced enhancement of brain neurotensin.

Effects of pro-inflammatory cytokines on cannabinoid CB1 and CB2 receptors in immune cells.