Synaptic activation of kainate receptors gates presynaptic ... - Nature

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Synaptic activation of kainate receptors gates presynaptic CB1 signaling at GABAergic synapses

© 2010 Nature America, Inc. All rights reserved.

Joana Lourenço1–4, Astrid Cannich1,4, Mario Carta3,4, Françoise Coussen3,4, Christophe Mulle3–5 & Giovanni Marsicano1,4,5 Glutamate can control inhibitory synaptic transmission through activation of presynaptic kainate receptors. We found that glutamate released by train stimulation of Schaffer collaterals could lead to either short-term depression or short-term facilitation of inhibitory synaptic transmission in mouse CA1 pyramidal neurons, depending on the presence of cannabinoid type 1 (CB1) receptors on GABAergic afferents. The train-induced depression of inhibition (t-Di) required the mobilization of 2-arachidonoylglycerol through postsynaptic activation of metabotropic glutamate receptors and [Ca 2+] rise. GluK1 (GluR5)dependent depolarization of GABAergic terminals enabled t-Di by facilitating presynaptic CB 1 signaling. Thus, concerted activation of presynaptic CB1 receptors and kainate receptors mediates short-term depression of inhibitory synaptic transmission. In contrast, in inhibitory connections expressing GluK1, but not CB1, receptors, train stimulation of Schaffer collaterals led to short-term facilitation. Thus, activation of kainate receptors by synaptically released glutamate gates presynaptic CB 1 signaling, which in turn controls the direction of short-term heterosynaptic plasticity. Balanced interaction between excitatory and inhibitory neurotransmission is required for the proper functioning of neural networks. A growing body of evidence suggests that endogenously released glutamate can control this interaction by presynaptic mechanisms modulating both inhibitory and excitatory synaptic transmission 1. Kainate receptors (KARs) are ionotropic glutamate receptors that are known to regulate the activity of synaptic networks by a variety of pre- and postsynaptic mechanisms 2,3. KARs activated by synaptically released glutamate can induce short-term inhibition of evoked inhibitory postsynaptic currents (eIPSCs) in CA1 pyramidal cells 4. In contrast, paired recordings involving specific GABAergic interneuron subtypes have shown that endogenous activation of KARs can also lead to a facilitation of inhibitory transmission5. The reasons for these apparent discrepancies are unknown. The endocannabinoid system (ECS), formed by metabotropic cannabinoid receptors and by their endogenous lipid ligands (endocannabinoids)6, is important for the regulation of synaptic transmission and plasticity at both excitatory and inhibitory synapses, mainly through the retrograde activation of presynaptic CB1 receptors7,8. The activity of the ECS can be directly promoted by both ionotropic and metabotropic glutamate receptors (NMDA type and mGluRs, respectively), generally through a postsynaptic mobilization of endocannabinoids and the consequent retrograde activation of presynaptic CB1 receptors9–11. Notably, presynaptic ionotropic NMDA glutamate receptors can also cooperate with the ECS in the regulation of synaptic transmission in the neocortex12,13.

The neurotoxin kainate has been shown to stimulate the synthesis of endocannabinoids in cultured neurons14 and the ECS was found to be neuroprotective against kainate-induced epileptiform seizures15,16. However, very little is known concerning functional interactions between KARs and the ECS in active synaptic networks under physio logical conditions. Here, we analyzed the mechanisms that regulate inhibitory transmission by endogenously released glutamate. We found that the concerted actions of the ECS and KARs induced short-term depression of inhibitory synaptic transmission, which was reverted to facilitation at synapses devoid of CB1 receptors. RESULTS GluK1 KARs mediate t-Di To explore the mechanisms of regulation of eIPSCs by synaptically released glutamate, we used an experimental setup that was similar to one described previously4 (Fig. 1a). In the presence of GYKI53655 (50 µM) and d-AP5 (50 µM), which block AMPA and NMDA receptors, and CGP 55845 (5 µM), which blocks presynaptic GABAB receptors17, a short train of stimulation (10 pulses at 200 Hz) to the Schaffer collateral fibers transiently reduced eIPSCs recorded in CA1 pyramidal neurons (Fig. 1 and Supplementary Fig. 1). This form of t-Di was fully blocked by application of 20 µM CNQX (control, −15 ± 2%; CNQX, −2 ± 2%; n = 5, P = 0.0004, paired t test; Fig. 1b,c), confirming that it is mediated by KARs4. LY382884 (10 µM), a selective antagonist of the GluK1 KAR sub unit, completely blocked t-Di (control, −12 ± 2%; LY382884, −2 ± 1%; n = 4, P = 0.0478, paired t test; Fig. 1d). t-Di was normally expressed

1INSERM

U862 NeuroCentre Magendie, Endocannabinoids and Neuroadaptation, Bordeaux, France. 2PhD Programme in Experimental Biology and Biomedicine, Center for Neuroscience and Cell Biology, University of Coimbra, Coimbra, Portugal. 3Laboratoire Physiologie Cellulaire de la Synapse, Centre National de la Recherche Scientifique UMR 5091, Bordeaux, France. 4University of Bordeaux, Bordeaux, France. 5These authors contributed equally to this work. Correspondence should be addressed to C.M. ([email protected]). Received 8 September; accepted 8 December 2009; published online 17 January 2010; doi:10.1038/nn.2481

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in slices from wild-type mice (−15 ± 2%, n = 11; Fig. 1e) and from GluK2−/− (also known as Grik2−/−) mice (−10 ± 3%, n = 10, P > 0.05 as compared to wild-type mice; Fig. 1e). In contrast, t-Di was abolished in GluK1−/− (also known as Grik1−/−) and GluK1−/−; GluK2−/− slices (GluK1−/−, +1 ± 1%, n = 11; GluK1−/−; GluK2−/−, +3 ± 2%;, n = 3; P < 0.001 for both comparisons with wild type, one-way ANOVA followed by Dunnett’s post-hoc test; Fig. 1e and see Supplementary Fig. 1 for raw data). In the hippocampus, GluK1 KARs are almost exclusively expressed in GABAergic interneurons18,19. Thus, our



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Figure 1 GluK1-KARs mediate t-Di. CNQX a S2 distal b Baseline Train Recovery 150 (a) Schematic illustration of the experimental CTR 125 setting showing the positions of the recording 100 pipette and stimulating electrodes in the S1 proximal 75 Schaffer hippocampus. IPSCs were evoked by a CNQX collateral 50 stimulating glass electrode (S1 proximal) 25 25 pA positioned in the stratum radiatum 100 µm 0 from to the recorded CA1 pyramidal neuron 50 ms 10 pulses at 200 Hz (gray). Schaffer collateral axons were stimulated with a bipolar electrode positioned at the Time (s) c d e boundary between CA3 and CA1 hippocampal 10 10 10 Baseline Train Recovery regions (S2 distal). Solid circle represents *** *** *** –/– 0 * GluK2 a GABAergic interneuron. (b) Left, a train 0 0 conditioning protocol (10 stimuli at 200 Hz) –10 25 pA –10 –10 applied at the S2 electrode induced depression 50 ms –20 of eIPSCs in control conditions (upper traces, –/– GluK1 –20 –20 CTR), which was blocked by the application of –30 CNQX (20 µM, lower trace). Right, individual –30 –30 10 pA normalized responses in control conditions 10 pulses at 200 Hz 50 ms and on KAR blockade (CNQX) of a single representative cell. Open circles represent the effect of a conditioning train given 200 ms before each of ten consecutive eIPSCs and solid circles represent control eIPSCs in the absence of the conditioning train. (c) Bar graph of ∆eIPSCs after conditioning train stimulation in control conditions (gray bar, filled symbols) and after CNQX treatment (white bar, open symbols). (d) Bar graph of ∆eIPSCs in control conditions (gray bar, filled symbols) and after treatment with the GluK1-KAR specific antagonist LY382884. (e) t-Di was normal in isogenic C57BL/6 control mice (black bar) and in GluK2−/− mice, but it was absent in slices from GluK1−/− mice and GluK1−/−; GluK2−/− double mutants. Left, representative traces. Right, summary bar graph. Data are single values and/or mean ± s.e.m. * P < 0.05 and *** P < 0.001 as compared with respective control conditions (see Supplementary Table 1 for detailed statistics).

results strongly suggest that the activation of GluK1 KARs in hippocampal GABAergic interneurons is necessary for t-Di. The ECS is required for the expression of t-Di Because CB1 receptor activation modulates GABAergic transmission7, we examined whether the ECS interacts with KARs to induce t-Di. t-Di was abolished by the CB1 receptor–specific antagonist SR141716A in wild-type mice (vehicle, −15 ± 1%; SR141716A, + 4 ± 5%; n = 6, P = 0.0126, paired t test; Fig. 2a,b) and was absent in CB1−/− (also known as Cnr1−/−) mice (wild type, −12 ± 1%, n = 5; CB1−/−, +7 ± 5%, n = 8; P = 0.0134, t test; Fig. 2a,c). To confirm that CB1 receptors involved in t-Di are expressed on GABAergic interneurons to regulate GABA release7,8, we analyzed t-Di in slices from conditional mutant mice lacking CB1 receptors in either GABAergic (GABA-CB1−/−) or cortical glutamatergic neurons (Glu-CB1−/−) and their respective wild-type littermate controls16,20 (see Online Methods). t-Di was fully abolished in GABA-CB1−/− mice (wild type, −15 ± 3%, n = 5; GABA-CB1−/−, +7 ± 4%, n = 6; P = 0.0031; Fig. 2a,c,d), but was normally expressed in Glu-CB1−/− mice Figure 2 CB1 receptors are required for the expression of t-Di. (a) t-Di was present in slices from C57BL/6 (CTR), CB1+/+ and GABA-CB1+/+ mice (upper traces), but it was impaired by treatment with the CB1 antagonist SR141716A, and in slices from CB1−/− and GABACB1−/− mice, respectively (lower traces). (b) Bar graph of ∆eIPSCs in control conditions (gray bar, filled symbols) and after treatment with SR141716A. (c) Bar graph representing ∆eIPSCs in slices from wild-type control littermates (black bars), global CB1−/− mice, and conditional mutants lacking CB1 receptor expression in GABAergic (GABA-CB1−/−) or in cortical glutamatergic neurons (Glu-CB1−/−). (d) Individual normalized responses of single representative cells. Open circles represent the effect of a conditioning train given 200 ms before each of ten consecutive eIPSCs and solid circles represent control eIPSCs in the absence of the conditioning train. Left, GABA-CB1+/+ mice. Right, GABA-CB1−/− mice. Data are single values and mean ± s.e.m. * P < 0.05 and ** P < 0.01 as compared with respective controls (see Supplementary Table 1 for detailed statistics).

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(wild type, −15 ± 2%, n = 7; Glu-CB1−/−, −10 ± 2%, n = 4; P > 0.1; Fig. 2c and see Supplementary Fig. 1 for raw data). CB1-expressing interneurons belong to the cholecystokinin-containing family of basket cells, strongly innervating not only the cell bodies, but also dendrites of pyramidal neurons21,22. Consistently, we found strong CB1 protein expression in GABAergic terminals of the stratum radiatum in both wild-type (Supplementary Fig. 2) and CaMKCB1−/− mice (see Online Methods), in which the CB1 receptor is deleted in all forebrain principal neurons15,16 (Supplementary Fig. 2). Depolarization-induced suppression of inhibition (DSI), a classical form of ECS-dependent short-term plasticity, relies on a global somatodendritic depolarization7,8,23. Accordingly, it occurred independently of the positioning of the stimulating electrode (Supplementary Fig. 2). In contrast, t-Di occurred only when IPCSs were evoked in the stratum radiatum at approximately 100 µm from the pyramidal cell layer (Supplementary Fig. 2). Thus, our data suggest that t-Di requires the spatial proximity of inhibitory and Schaffer collaterals glutamatergic synapses.

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Figure 3 t-Di requires postsynaptic mobilization of 2-AG. (a,b) The Ca2+ chelator BAPTA (20 mM) in the recording pipette blocked t-Di. Individual normalized responses of a single representative cell in presence of postsynaptic BAPTA (symbols as in Fig. 1b) are shown in a. A bar graph representing t-Di in control conditions (black bar and scattered solid circles) or in presence of BAPTA (white bar and scattered open circles) is shown in b (see Supplementary Fig. 10 for the complete experiment). (c) Postsynaptic application of the DAGL inhibitor THL (5 µM, white bar and scattered open circles) fully blocked t-Di, which was unaltered by vehicle control infusion ( 0.2 for both drugs as compared to control; Fig. 3d). However, when used in combination, LY3675385 and MPEP fully abolished t-Di (+1 ± 2%, n = 10, P < 0.001 as compared with control, one-way ANOVA followed by Dunnett’s post hoc test; Fig. 3d). In addition, t-Di was not altered by the noncompetitive mGluR type 1 antagonist BAY36-7620 (10 µM) alone, but it was fully blocked by the combination of BAY367620 and MPEP, as well as by the nonspecific group I mGluR competitive antagonist methyl-4-carboxyphenylglycine (MCPG, 1 mM) (Supplementary Fig. 5). Electron microscopy studies have revealed that mGluR1 and 5 are mostly expressed at the postsynaptic level in the hippocampus26. To assess whether postsynaptic G proteins are necessary for the expression of t-Di, we blocked this pathway by including the nonhydrolysable GDP analog guanosine 5′-O(2-thiodiphosphate) (GDP-βS, 500 µM) in the patch pipette. This treatment fully blocked t-Di (+1 ± 3%, n = 6, P < 0.001 as compared with control; Fig. 3d). Taken together, these results strongly suggest that 2-AG mobilization during t-Di relies on an mGluR-dependent [Ca2+] rise in the postsynaptic pyramidal neurons.


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GluK1-dependent depolarization increases CB1 efficacy Glutamate released during the conditioning train might depolarize interneurons through GluK1 KARs, thereby ‘setting’ an adequate presynaptic activity state, which has been proposed to be important for the inhibitory effects of CB1 (for example, see ref. 27). To test the possibility that GluK1 KARs contribute to t-Di by a mechanism linked to depolarization, we first increased the [K+] in the artificial cerebrospinal fluid (ACSF) that we used (to 7.5 mM). A nonsignificant trend to increase t-Di amplitude was induced by elevating extracellular [K+] from 2.5 to 7.5 mM in untreated wild-type slices (2.5 mM [K+], −15 ± 1%, n = 17; 7.5 mM [K+], −23 ± 3%, n = 5; P > 0.05; Fig. 4a,b). Notably, 7.5 mM [K+] rescued t-Di in GluK1−/− mice and in wildtype slices treated with LY382884 (GluK1−/− 7.5 mM [K+], −15 ± 5%, n = 4; wild type LY382884 7.5 mM [K+], −11 ± 4%, n = 6; P > 0.05 as compared to wild-type 2.5 mM [K+], one-way ANOVA followed by Dunnett’s post hoc test; Fig. 4a,b). The depression of eIPSCs in high [K+] still depended on CB1 receptors, as application of SR141716A fully blocked t-Di in GluK1−/− mice in these conditions (GluK1−/− 7.5 mM [K+] SR141716A, +0.5 ± 2%, n = 6, P < 0.001 as compared with wild-type 2.5 mM [K+], one-way ANOVA followed by Dunnett’s post hoc test; Fig. 4a,b). Increased [K+] might compensate for the lack of GluK1 KARs simply by increasing the synthesis of endo cannabinoids through postsynaptic [Ca2+] rise. However, currentclamp recordings revealed that 7.5 mM [K+] induced only a modest depolarization of pyramidal neurons (change in membrane potential from 2.5 to 7.5 mM [K+] = +13.7 ± 1.1 mV, n = 5), which is unlikely to mobilize sufficient amounts of endocannabinoids per se. Furthermore, 200

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+ Figure 4 GluK1 KAR–dependent 10 a Wild type 7.5 mM K GluK1–/– b depolarization enhances the efficacy of Baseline Train Baseline Train CB1 signaling. (a) Representative traces *** *** *** 0 CTR of t-Di in 7.5 mM [K +] in untreated wild-type slices (CTR), slices from GluK1−/− mice, –10 and in wild-type slices treated with the 10 pA 10 pA –20 GluK1 antagonist LY382884 (10 µM) and 50 ms 50 ms the CB1 receptor antagonist SR141716A LY382884 SR141716A –30 (5 µM), respectively. (b) Bar graph +/+ +/+ –/– –/– +/+ +/+ –/– Genotype GluK1 representing the effects of the conditioning + 2.5 7.5 2.5 7.5 2.5 7.5 7.5 [K ] in mM 10 pA 10 pA train on eIPSCs in control conditions of LY382884 – – – – + + – + 50 ms 50 ms normal extracellular [K ] (2.5 mM) and in high SR141716A – – – – – – + + [K ] (7.5 mM) under different conditions. 150 nM WIN55,212-2 1 s at 0 mV DSI (c) Plot of the effect of WIN55,212-2 10 c 150 d 125 + 2.5 mM K (150 nM) on the average normalized eIPSC 0 + 125 7.5 mM K 100 amplitudes in 2.5 mM [K +] (solid circles) and –10 100 7.5 mM [K+] (open circles) in the presence 75 –20 of mGluRs antagonists and intracellular 75 –30 BAPTA to prevent postsynaptic synthesis of 50 50 –40 + endocannabinoids. (d,e) Modulation of DSI 2.5 mM K + 25 –50 25 7.5 mM K by increased interneuronal activity. (d) Left, * summary plot of eIPSCs before and after 0 600 1,200 1,800 0 20 40 60 80 100 the depolarization step in 2.5 mM [K +] Time (s) Time (s) DSI 10 (solid circles) and 7.5 mM [K +] (open circles) DSI Post-DSI Pre-DSI e in the presence of mGluRs antagonists. 0 1 s at 0 mV Right, summary bar graph of DSI amplitudes –10 in control conditions (black bar) and in –20 the presence of 7.5 mM [K +] (white bar). –30 (e) Left, representative traces of eIPSCs of 1 s at 0 mV –40 DSI induced by a step of depolarization + train 25 pA –50 alone (top traces) or by a step of ** 50 ms depolarization preceded by a conditioning train (bottom traces). Right, summary bar graph of DSI amplitudes in control conditions (black bar), after train-conditioning (white bar) and after train-conditioning in the presence of the GluK1-KAR antagonist LY382884 (gray bar). Data are mean ± s.e.m. * P < 0.05, ** P < 0.01 and *** P < 0.001 as compared with control conditions (see Supplementary Table 2 for detailed statistics).

increased [K+] enhanced the efficacy of the CB1 agonist WIN 55,212-2 in the presence of the mGluRs antagonists and intracellular BAPTA to blunt endocannabinoid synthesis. At 150 nM, WIN 55,212-2 decreased eIPSCs by 33 ± 5% (n = 5; Fig. 4c), as compared with 52 ± 1% (n = 3) at a concentration of 1 µM. Increasing [K+] to 7.5 mM facilitated the effect of the nonsaturating concentration of WIN 55,212-2 (150 nM WIN 55,212-2 in 7.5 mM [K+], −50 ± 5% of baseline, n = 5, P < 0.05 as compared with normal ACSF, t test; Fig. 4c). In addition, 7.5 mM [K+] potentiated DSI induced by a short depolarization step, in the presence of the mGluRs antagonists (DSI 1s in 2.5 mM [K+], −29.8 ± 5%, n = 7; DSI 1 s in 7.5 mM [K+], −43.5 ± 4%, n = 8; P = 0.0423, t test; Fig. 4d). These data strongly suggest that the increase of [K+] enhances presynaptic CB1 signaling, probably by depolarizing pre synaptic terminals. Notably, t-Di–inducing trains in normal ACSF mimicked the effect of high [K+] on DSI in a GluK1-dependent manner (DSI 1 s, −25 ± 4%, n = 8; DSI 1 s + train, −46 ± 2%, n = 8, P < 0.01 as compared with DSI 1 s; DSI 1 s + train + LY382884, −29 ± 2%, n = 4, P > 0.05 as compared to DSI 1 s, one-way ANOVA followed by Dunnett’s post hoc test; Fig. 4e). The role of GluK1 KARs in this context might merely be to increase glutamate release enhancing mGluRs signaling and eventually endocannabinoid mobilization. However, increasing ambient glutamate by blocking glutamate uptake with dl-threo-b-benzyloxyaspartic acid (TBOA,100 µM) did not compensate for the absence of GluK1 KARs activation in t-Di (Supplementary Fig. 6). Thus, increase of extracellular glutamate does not appear to be the mechanism by which GluK1-KARs induce t-Di. As expected, KARs are not involved VOLUME 13 | NUMBER 2 | february 2010 nature NEUROSCIENCE

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Figure 5 t-Di is present in unitary connections 5 s at 0 mV a b Wild type DSI sensitive (+) 160 expressing CB1 receptors. (a) Schematic S2 distal Pre-DSI DSI Post-DSI illustration of the experimental setup showing 120 the positions of the recording pipettes and 80 stimulating electrode in CA1 subfield. IPSCs Schaffer 20 mV collateral 50 ms were evoked in pyramidal cells (gray) by 40 Wild-type DSI (+) triggering spikes in connected interneurons Wild-type DSI (–) 25 pA 0 CB1–/– (black). Schaffer collateral axons were 50 ms 0 20 40 60 80100120 stimulated with a bipolar electrode (S2 distal), Time (s) as in Figure 2. (b) Left, representative traces Baseline Train Recovery c of a DSI-sensitive paired connection. Upper Wild type traces show spikes triggered in the presynaptic d e ## # DSI sensitive 25 25 *** *** interneuron and the lower traces show the ** uIPSC in the connected pyramidal cell. Right, 0 0 summary plot of IPSCs evoked at a frequency of 0.33 Hz during DSI (5 s at 0 mV). Black circles –25 –25 Wild type indicate DSI-sensitive pairs from wild-type DSI sensitive 20 mV mice (DSI (+), n = 4), gray circles indicate + LY382884 –50 –50 50 ms DSI-insensitive pairs from wild-type mice (DSI (−), n = 6) and white circles indicate pairs from CB1−/− mice (n = 6). Pairs showing >7% 25 pA depolarization-induced decrease of uIPSCs 50 ms 10 pulses at 200 Hz were defined as being DSI sensitive. (c) t-Di was present in DSI-sensitive pairs (upper traces), where it is blocked by the GluK1 KAR antagonist LY382884 (10 µM, lower traces). Gray traces represent superimposed single trace recordings and the lower black traces are of average recordings. (d) Bar graph representing the effect of the GluK1 antagonist LY382884 on unitary t-Di in DSI-sensitive connected pairs from wild-type mice. (e) Bar graph of the changes in uIPSC amplitude following the conditioning train in DSI-sensitive and DSI-insensitive connected pairs from wild-type and from CB1−/− mice. ** P < 0.01 and *** P < 0.001 as compared to controls. # P < 0.05 and ## P < 0.01 as compared with no effect (one-sample t test) (see Supplementary Table 2 for detailed statistics).

Differential expression of t-Di at unitary connections t-Di occurs at physiological temperature (33–34 °C; Supplementary Fig. 7) and can be induced by physiologically relevant patterns of stimulation29 (see Supplementary Fig. 7). The occurrence of t-Di as a result of bulk stimulation of GABAergic afferents is a very consistent phenomenon, but it is quite limited in amplitude (inhibition of about 15%, see also ref. 4). The limited amplitude of t-Di might be a result of the averaging of synaptic connections, which are affected differently by the conditioning train. To address this question, we carried out paired recordings from GABAergic interneurons in the stratum radiatum and their pyramidal cell targets (Fig. 5a). In the hippo campus, CB1 receptors are expressed only in subsets of GABAergic interneurons30–32. Therefore, we indirectly tested the connected pairs after each experiment for the presence of functional CB 1 receptors on their GABAergic terminals (see ref. 33) by applying a DSI protocol (Fig. 5b). Both GABAergic interneurons that were sensitive to DSI induction (defined as depolarization-induced reduction of unitary IPSCs greater than 7% of baseline) and those that were not showed distinct patterns of spike frequency adaptation. DSI-sensitive neurons (that is, presumably expressing CB1 receptors at their terminals) adapted, whereas DSI-insensitive neurons did not (DSI-sensitive

adaptation coefficient, 0.4 ± 0, n = 9; DSI-insensitive adaptation coefficient, 0.7 ± 0, n = 7; P < 0.0001; Supplementary Fig. 8). As expected, DSI did not occur in any of the pairs from CB1−/− mice (Fig. 5b). Notably, in all of the DSI-sensitive pairs from wild-type mice (DSI, −77 ± 7%; Fig. 5c,d), the application of the t-Di–inducing train markedly depressed unitary IPSCs (uIPSCs, −38 ± 1%, n = 8, P < 0.0001, one-sample t test; Fig. 5c–e and see Supplementary Fig. 8 for raw data). Conversely, t-Di of uIPSCs was not present in any of the DSI-insensitive pairs from wild-type mice (+11 ± 1%, n = 7, P < 0.001 compared with DSI sensitive; Fig. 5c,e) or in pairs from CB1−/− mice (+10 ± 3%, n = 6, one-way ANOVA followed by Dunnett’s post hoc test, P < 0.001 compared to DSI sensitive; Fig. 5e; see Supplementary Fig. 8 for raw data). In wild-type pairs, the selective GluK1 antagonist LY382884 fully blocked t-Di of uIPSCs (control, −35 ± 1%; LY382884, +2 ± 2%; n = 4, P = 0.0013, paired t test; Fig. 5d). The differential effects of the conditioning train in separate unitary connections helps to explain the limited extent of t-Di observed in conditions of bulk stimulation of GABAergic afferents, where both types of connections are probably combined. GluK1 KARs might increase glutamatergic inputs onto GABAergic interneurons34. However, blockade of GluK1 KARs did not alter paired-pulse facilitation of glutamatergic inputs of both adapting and nonadapting interneurons (Supplementary Fig. 8), further suggesting that the function of GluK1 KARs during t-Di is not to increase glutamate release. In our current-clamp recordings, interneurons did not show any spontaneous firing and the train stimulation did not alter their membrane potential in either the presence or absence of GluK1 KAR signaling (Supplementary Fig. 8). Paired recordings also allowed us to directly test whether pre synaptic depolarization increases per se the efficiency of CB1 signaling. Subthreshold depolarization of dentate gyrus granule cells has been shown to reach the presynaptic terminals of mossy fibers35. We applied a similar subthreshold depolarization step (from −90 to −50 mV) to CA1 interneurons paired with pyramidal neurons. This


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in the induction of DSI28 (Supplementary Fig. 6). However, local application of glutamate via light-induced uncaging in the presence of the general blocker of mGluRs LY341495 (100 µM, to avoid glutamate-induced endocannabinoid mobilization) enhanced DSI in a GluK1-dependent manner (Supplementary Fig. 6). Thus, DSI is enhanced to a similar extent by train stimulation of Schaffer collaterals and by exogenous activation of GluK1, independently of postsynaptic mGluRs. Altogether, these data strongly suggest that GluK1-induced depolarization does not enhance endocannabinoid release per se, but it does increase the efficacy of both endogenous (t-Di and DSI) and exogenous (WIN 55,212-2) activation of pre synaptic CB1 signaling.

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The ECS controls the direction of GluK1-dependent plasticity In CB1−/− mice and DSI-insensitive pairs from wild-type mice, we observed a significant potentiation of uIPSCs induced by train stimulation of Schaffer collaterals (CB1−/−, P = 0.0189; wild type DSI insensitive, P = 0.0007; one-sample t test; Fig. 5e). This strongly suggests that endogenous release of glutamate facilitates GABA release by activation of presynaptic GluK1 KARs at single synapses that do not coexpress CB1 receptors. Therefore, we tested whether the simultaneous retrograde endocannabinoid signaling at CB1-containing synapses would convert this facilitation into a depression (Fig. 6). We applied the GluK1 antagonist LY382884 in DSI-insensitive connected pairs from wild-type mice and from CB1−/− mice. This treatment fully blocked the potentiation induced by the train in wild-type DSI-insensitive pairs (control, +13 ± 5%; LY382884, −2 ± 3%; n = 4, P = 0.0031, paired t test; Fig. 6a,c) and in CB1−/− mice (CB1−/−, +15 ± 2%; CB1−/− + LY382884, −3 ± 1%; n = 3, P = 0.0262, paired t test; Fig. 6d). To directly confirm that glutamate released during the conditioning train is able to increase GABA release at synapses devoid of CB1, we applied the CB1 antagonist SR141716A at DSI-sensitive pairs. This treatment not only abolished t-Di (control, −39 ± 9%; SR141716A, +12 ± 3%; n = 4, P = 0.0046, paired t test; Fig. 6b,e), but also reverted the effect of the conditioning train on uIPSCs from depression to facilitation (SR141716A, P = 0.0335, one-sample t test; see also Supplementary Table 2). Facilitation of eIPSCs was also observed in bulk stimulation conditions when the ECS was genetically or pharmacologically blocked (see Figs. 2 and 3). Despite the variability of these data (probably as a result of the heterogeneity of the stimulated fibers), this phenomenon reached statistical significance when BAPTA or THL were applied in the postsynaptic cells (see Fig. 3). Notably, this effect of postsynaptic BAPTA was absent in GluK1−/− mice (Supplementary Fig. 10), further supporting its dependence on GluK1 KAR activation. These results indicate that GluK1 KARs facilitate eIPSCs in the absence of ECS control and suggest that GluK1-containing GABAergic interneurons in the hippocampus belong to two distinct populations, those that express CB1 receptors and those that do not. To confirm this 202

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manipulation, combined with to a DSI-inducing protocol (1 s at 0 mV), did not change the uIPSC amplitudes of DSI-insensitive pairs (Supplementary Fig. 9), but it consistently increased the amplitude of DSI in DSI-sensitive pairs (DSI −90 mV, −51.4 ± 12.9%; DSI −50 mV, −62.9% ± 10.7%; n = 5, P = 0.0148, paired t test; Supplementary Fig. 9). Notably, this potentiation was obtained in the presence of CNQX and it was inversely proportional to the amplitude of DSI (Supplementary Fig. 9). These data further confirm that presynaptic depolarization is able to increase the efficacy of CB1 signaling.

a



Figure 6 In the absence of ECS signaling, GluK1 KARs facilitate inhibitory transmission. (a) In DSI-insensitive pairs, train stimulation of Schaffer collaterals induced facilitation of uIPSCs (upper traces), which was fully blocked by the GluK1 KAR antagonist LY382884 (10 µM, lower traces). (b) In DSI-sensitive pairs, the application of the CB 1 antagonist SR141716A reverted the effect of the train stimulation from depression (upper traces) to facilitation (lower traces). Gray traces represent superimposed single trace recordings and the lower black traces are of average recordings. (c,d) Bar graphs summarizing the effect of the GluK1 KAR antagonist LY382884 on DSI-insensitive pairs from wild-type mice (c) and on pairs from CB1−/− mice (d). (e) Bar graph showing that unitary t-Di induced by the conditioning train in DSI-sensitive pairs from wild-type mice was reverted to a facilitation in the presence of SR141716A. Data are single values and mean ± s.e.m. * P < 0.05 and ** P < 0.01 as compared with controls. # P < 0.05 as compared with no effect (one-sample t test) (see Supplementary Table 2 for detailed statistics).

assumption, we carried out highly sensitive double fluorescent in situ hybridization (D-FISH) using specific probes directed against CB1 (ref. 32) and GluK1 mRNAs18, respectively. Approximately 50–60% of GluK1-positive GABAergic interneurons contained high levels of CB1 mRNA, whereas the remaining ones contained GluK1, but not CB1 mRNA (Supplementary Fig. 11). DISCUSSION We found that release of glutamate by a conditioning train of Schaffer collaterals induced short-term inhibition of eIPSCs (t-Di) that relied on the ECS and required the coordinated activation of presynaptic GluK1-KARs, that the GluK1-dependent depolarization facilitated CB1 signaling, enabling the inhibition of GABA release at connections co-expressing CB1 receptors, and that GluK1 KARs facilitated inhibitory transmission at inhibitory connections that did not coexpress CB1 receptors. Altogether, our data indicate that GluK1 KARs gate pre synaptic CB1 signaling and that the concerted actions of GluK1 KARs and the ECS determine the direction of GABAergic transmission regulation by synaptically released glutamate. t-Di, a form of ECS-dependent heterosynaptic plasticity t-Di is a form of short-term heterosynaptic plasticity of GABAergic transmission that is induced by the conditioning activity of gluta matergic afferents. We propose the following mechanistic explanation for t-Di (Supplementary Fig. 12). Synaptically released glutamate activates postsynaptic mGluR1 and mGluR5 to trigger intracellular [Ca2+] increase in CA1 pyramidal cells, leading to activation of DAGL and to mobilization of the endocannabinoid 2-AG. Simultaneous synaptic release of glutamate activates presynaptic GluK1-containing KARs, inducing a moderate depolarization of the GABAergic axon terminal. This depolarization facilitates presynaptic CB1 receptors signaling and leads to inhibition of GABA release from interneurons co-expressing both receptors. GluK1 KARs are present in somato-dendritic compartments of hippo campal interneurons36–38. The fact that we did not detect any significant VOLUME 13 | NUMBER 2 | february 2010 nature NEUROSCIENCE


a r t ic l e s somato-dendritic depolarization of interneurons in paired recordings during the conditioning train strongly suggests that somato-dendritic GluK1-KARs are not involved in t-Di. Nevertheless, our experiments with slightly increased [K+] indicate that a certain degree of depolarization is necessary to induce t-Di. Elevated [K+] might potentiate t-Di or DSI simply by enhancing endocannabinoid synthesis. However, we can discard this possibility for several reasons. First, only very high levels of [K+] (40 mM) have been shown to induce the synthesis of endocannabinoids, probably via massive neuronal depolarization39. In our conditions, elevating [K+] to 7.5 mM induced a modest depolarization, which is unlikely to mobilize substantial amounts of endocannabinoids. Second, synaptic release of glutamate elicited by a conditioning train, local application of glutamate, increased [K+] and subthreshold depolarization of presynaptic interneurons all increased the amplitude of DSI to a similar extent, suggesting that these manipulations act in a similar manner. Third, increased [K+] facilitated the inhibitory effect of an exogenous synthetic CB1 agonist in the presence of BAPTA and mGluRs antagonists to blunt postsynaptic endocannabinoid synthesis. Overall, the most parsimonious explanation of our data is that, after a conditioning train, GluK1 KARs depolarize the presynaptic terminals at which CB1 receptors mediate depression of GABA release. Our data suggest that the amount of endocannabinoids released from post synaptic pyramidal cells with t-Di protocols is not sufficient to activate presynaptic CB1 receptors in the absence of the facilitatory action of presynaptic KARs. As opposed to other forms of ECS-dependent synaptic plast icity9,33,40, t-Di depends on both postsynaptic mGluR signaling and [Ca2+] rise. However, these two pathways have been proposed to converge onto phospholipase C activity, acting as a coincidence detector to mobilize 2-AG7. Thus, it is reasonable to propose that t-Di represents an additional example of this mechanism, applied to endogenous actions of glutamate. Endocannabinoid release induced by postsynaptic depolarization as in DSI is relatively global and should affect all GABAergic connections with CB1 receptors. In contrast, the concerted action of presynaptic CB1 receptors and KARs requires anatomical proximity between the glutamatergic and GABAergic afferents involved. Following the conditioning train, the retrograde depression of GABA release depends on local glutamatergic synaptic and retrograde endocannabinoid signaling. Therefore, the concerted action of the ECS and GluK1KARs impart synapse specificity to short-term plasticity induced by endogenously released glutamate.

In previous studies, endogenous glutamate acting on KARs has been reported as either inhibiting4 or facilitating GABAergic transmission in CA1 pyramidal cells5. Our data help to explain this apparent discrepancy, suggesting that the direction of the effect of KARs is highly dependent on synaptic connection specificity and, in particular, on the presynaptic presence of CB1 receptors. A previous study5 found that endogenous glutamate potentiates inhibitory transmission from nonadapting interneurons41. Consistently, our data demonstrate that only inhibitory connections from adapting interneurons are under the control of the ECS. Thus, at least two sets of GABAergic connections onto CA1 pyramidal cells exist, in which KARs can either potentiate or inhibit synaptic transmission depending on the expression and activity of CB1 receptors. As a consequence, the net effect of a conditioning train applied to the Schaffer collaterals by bulk stimulation of GABAergic afferents4 probably represents the average output of differentially regulated connections. There is growing evidence that CB1 receptors are able to physically interact with other proteins42 and the possibility of a direct crosstalk with GluK1 KARs cannot be excluded. In addition, GluK1 KARs have been proposed to mediate metabotrobic functions in GABAergic interneurons2. However, GluK1 KAR blockade and genetic deletion can be rescued by presynaptic depolarization to induce CB 1 potentiation, indicating that t-Di is instead linked to the classical ionotropic functions of GluK1 KARs. Moreover, we found no evidence of physical interaction between these two receptors in heterologous expression systems (Supplementary Fig. 13). Presynaptic activity and priming signals have been proposed to be important for endocannabinoid action at different types of synapses12,13,27,43–45. For example, increasing the firing rate of pre synaptic hippocampal interneurons to >20 Hz markedly decreases the magnitude of DSI and the action of CB1 receptor agonists27. The mechanisms of this modulation are presently unknown, but could be linked to presynaptic Ca2+ levels possibly associated with changes in membrane potential27. Overall, these data suggest that a window of presynaptic activity for proper retrograde endocannabinoid signaling might exist, above a minimum threshold,, but below a maximal limit27.

Bidirectional plasticity of GABAergic transmission The genetic deletion or pharmacological blockade of CB1 receptors, as well as the blockade of endocannabinoid synthesis, not only abolished t-Di, but also induced a GluK1-dependent facilitation of eIPSCs. For example, DSI-sensitive paired connections showing strong t-Di facilitated uIPSCs on treatment with the CB1 antagonist SR141716A. Moreover, in untreated wild-type mice, the DSI-insensitive paired connections (that is, those that probably did not express CB1 receptors) responded with a similar GluK1-mediated potentiation of uIPSCs. These results clearly indicate that GluK1 KAR activation by endogenous release of glutamate increased the efficacy of GABAergic transmission in the absence of CB1 receptors (Supplementary Fig. 12). The results of our mRNA distribution study support the idea that two populations of GluK1-expressing interneurons exist and are distinguished by the presence or lack of CB1 receptors. Because of the absence of suitable antibodies, it is not possible to directly check for the anatomical localization of GluK1 protein on presynaptic terminals and to confirm the colocalization with CB1 receptors at the ultrastructural level.

Functional implications of t-Di Synaptically released glutamate during trains of stimulation to Schaffer collaterals increases ECS-mediated depression, thus promoting excitation of postsynaptic pyramidal cells. Conversely, high rates of firing of presynaptic interneurons containing CB1 receptors limit depression by the ECS, thereby keeping inhibitory connections active27,44. These complementary processes might help GABAergic transmission to sense and adapt to changes in the overall network activity27,46. It has been suggested that the hippocampal GABAergic network is critical for the control of network oscillations and temporal dynamics47. Our data suggest that the presence of CB1 receptors and/or GluK1 KARs on these interneurons might strongly regulate these mechanisms. The firing of CA3 pyramidal cells, as during exploratory behavior, induces a direct glutamatergic excitatory response and an indirect GABAergic response mediated by hippocampal interneurons in CA1 pyramidal cells (feedforward inhibition and feedback inhibition)46. Notably, CB1-expressing cells in the hippocampus seem to be particularly important for mediating feedback inhibition46. t-Di induces a considerable depression (~40% of inhibition) of GABAergic transmission at selected synaptic connections that depends on the activation of glutamatergic afferents. Thus, this associative phenomenon clearly has the potential to modulate feedback inhibition in CA1 pyramidal cells. It will be worthwhile to investigate the effects of trains of afferent stimulation triggering t-Di on the extent of feedback inhibition. In addition, this new form of short-term plasticity calls for a reappraisal of the conditions needed


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a r t ic l e s to induce CB1-dependent long-term depression of GABAergic synaptic responses (I-LTD) in the hippocampus. I-LTD was originally described in the presence of KAR antagonists33. Our data suggest that the threshold for I-LTD induction might be lowered by facilitation of CB1 signaling on KARs activation by synaptically released glutamate. Overall, our results suggest that interactions between presynaptic ionotropic glutamate receptors and metabotropic cannabinoid receptors control the direction of inhibitory synaptic transmission, a process that might be important for fine-tuning network activity and behavior. Methods Methods and any associated references are available in the online version of the paper at http://www.nature.com/natureneuroscience/.


Note: Supplementary information is available on the Nature Neuroscience website. Acknowledgments We thank the members of the laboratories of G. Marsicano and C. Mulle for fruitful discussions, P. Pinheiro for help during the experiments and A. Vimeney, D. Gonzales and the Genotyping Facility of the NeuroCentre Magendie for mouse genotyping. We are grateful to K. Mackie (Indiana University) for providing the CB1 antiserum. We thank J.P. Mazat, D. Commenges, C. Schwierz and L. Rosinus for help with statistical analyses. We thank K. Nave, J. Rubenstein, M. Ekker and G. Schütz for the use of Cre-expressing mouse lines. We also thank F. Chaouloff and N. Rebola for comments on the manuscript. This work was supported by an AVENIR grant of the Institut National de la Santé et de la Recherche Médicale (INSERM) in partnership with the Foundation Bettencourt-Schueller (G.M.), by the Agence National de la Recherche (ANR-06-NEUR-043-01 to G.M. and ANR-05-NEUR-033-01 to C.M.), by the Conseil Régional d’Aquitaine (G.M. and C.M.), by the Fundação para a Ciência e a Tecnologia, Portugal (J.L.), the Centre National de la Recherche Scientifique (C.M.) and European Commission Coordination Action Network of European Neuroscience Institutes (ENINET) (LSHM-CT-2005-19063 to G.M.). AUTHOR CONTRIBUTIONS J.L. conducted the electrophysiological experiments, A.C. carried out the anatomical studies, M.C. performed the glutamate uncaging experiments and F.C. carried out the biochemical assays. J.L., C.M. and G.M. designed the experiments and wrote the manuscript. Published online at http://www.nature.com/natureneuroscience/. Reprints and permissions information is available online at http://www.nature.com/ reprintsandpermissions/. 1. Pinheiro, P.S. & Mulle, C. Presynaptic glutamate receptors: physiological functions and mechanisms of action. Nat. Rev. Neurosci. (2008). 2. Lerma, J. Kainate receptor physiology. Curr. Opin. Pharmacol. 6, 89–97 (2006). 3. Pinheiro, P. & Mulle, C. Kainate receptors. Cell Tissue Res. 326, 457–482 (2006). 4. Min, M.Y., Melyan, Z. & Kullmann, D.M. Synaptically released glutamate reduces gamma-aminobutyric acid (GABA)ergic inhibition in the hippocampus via kainate receptors. Proc. Natl. Acad. Sci. USA 96, 9932–9937 (1999). 5. Jiang, L., Xu, J., Nedergaard, M. & Kang, J. A kainate receptor increases the efficacy of GABAergic synapses. Neuron 30, 503–513 (2001). 6. Piomelli, D. The molecular logic of endocannabinoid signaling. Nat. Rev. Neurosci. 4, 873–884 (2003). 7. Kano, M., Ohno-Shosaku, T., Hashimotodani, Y., Uchigashima, M. & Watanabe, M. Endocannabinoid-mediated control of synaptic transmission. Physiol. Rev. 89, 309–380 (2009). 8. Alger, B.E. Retrograde signaling in the regulation of synaptic transmission: focus on endocannabinoids. Prog. Neurobiol. 68, 247–286 (2002). 9. Maejima, T., Hashimoto, K., Yoshida, T., Aiba, A. & Kano, M. Presynaptic inhibition caused by retrograde signal from metabotropic glutamate to cannabinoid receptors. Neuron 31, 463–475 (2001). 10. Ohno-Shosaku, T. et al. Endocannabinoid signaling triggered by NMDA receptormediated calcium entry into rat hippocampal neurons. J. Physiol. (Lond.) 584, 407–418 (2007). 11. Varma, N., Carlson, G.C., Ledent, C. & Alger, B.E. Metabotropic glutamate receptors drive the endocannabinoid system in hippocampus. J. Neurosci. 21, RC188 (2001). 12. Bender, V.A., Bender, K.J., Brasier, D.J. & Feldman, D.E. Two coincidence detectors for spike timing–dependent plasticity in somatosensory cortex. J. Neurosci. 26, 4166–4177 (2006). 13. Sjöström, P.J., Turrigiano, G.G. & Nelson, S.B. Neocortical LTD via coincident activation of presynaptic NMDA and cannabinoid receptors. Neuron 39, 641–654 (2003). 14. Cadas, H., Gaillet, S., Beltramo, M., Venance, L. & Piomelli, D. Biosynthesis of an endogenous cannabinoid precursor in neurons and its control by calcium and cAMP. J. Neurosci. 16, 3934–3942 (1996).

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15. Marsicano, G. et al. CB1 cannabinoid receptors and on-demand defense against excitotoxicity. Science 302, 84–88 (2003). 16. Monory, K. et al. The endocannabinoid system controls key epileptogenic circuits in the hippocampus. Neuron 51, 455–466 (2006). 17. Frerking, M., Petersen, C.C. & Nicoll, R.A. Mechanisms underlying kainate receptor–mediated disinhibition in the hippocampus. Proc. Natl. Acad. Sci. USA 96, 12917–12922 (1999). 18. Bureau, I., Bischoff, S., Heinemann, S.F. & Mulle, C. Kainate receptor–mediated responses in the CA1 field of wild-type and GluR6-deficient mice. J. Neurosci. 19, 653–663 (1999). 19. Paternain, A.V., Herrera, M.T., Nieto, M.A. & Lerma, J. GluR5 and GluR6 kainate receptor subunits coexist in hippocampal neurons and coassemble to form functional receptors. J. Neurosci. 20, 196–205 (2000). 20. Monory, K. et al. Genetic dissection of behavioral and autonomic effects of Delta(9)tetrahydrocannabinol in mice. PLoS Biol. 5, e269 (2007). 21. Tsou, K., Brown, S., Sanudo-Pena, M.C., Mackie, K. & Walker, J.M. Immunohistochemical distribution of cannabinoid CB1 receptors in the rat central nervous system. Neuroscience 83, 393–411 (1998). 22. Pawelzik, H., Hughes, D.I. & Thomson, A.M. Physiological and morphological diversity of immunocytochemically defined parvalbumin- and cholecystokinin-positive interneurones in CA1 of the adult rat hippocampus. J. Comp. Neurol. 443, 346–367 (2002). 23. Wilson, R.I. & Nicoll, R.A. Endogenous cannabinoids mediate retrograde signaling at hippocampal synapses. Nature 410, 588–592 (2001). 24. Bacci, A., Huguenard, J.R. & Prince, D.A. Long-lasting self-inhibition of neocortical interneurons mediated by endocannabinoids. Nature 431, 312–316 (2004). 25. Ali, A.B. Presynaptic Inhibition of GABAA receptor-mediated unitary IPSPs by cannabinoid receptors at synapses between CCK-positive interneurons in rat hippocampus. J. Neurophysiol. 98, 861–869 (2007). 26. Luján, R., Nusser, Z., Roberts, J.D., Shigemoto, R. & Somogyi, P. Perisynaptic location of metabotropic glutamate receptors mGluR1 and mGluR5 on dendrites and dendritic spines in the rat hippocampus. Eur. J. Neurosci. 8, 1488–1500 (1996). 27. Földy, C., Neu, A., Jones, M.V. & Soltesz, I. Presynaptic, activity-dependent modulation of cannabinoid type 1 receptor-mediated inhibition of GABA release. J. Neurosci. 26, 1465–1469 (2006). 28. Mulle, C. et al. Subunit composition of kainate receptors in hippocampal interneurons. Neuron 28, 475–484 (2000). 29. Buzsáki, G. Theta oscillations in the hippocampus. Neuron 33, 325–340 (2002). 30. Freund, T.F. & Katona, I. Perisomatic inhibition. Neuron 56, 33–42 (2007). 31. Katona, I. et al. Presynaptically located CB1 cannabinoid receptors regulate GABA release from axon terminals of specific hippocampal interneurons. J. Neurosci. 19, 4544–4558 (1999). 32. Marsicano, G. & Lutz, B. Expression of the cannabinoid receptor CB1 in distinct neuronal subpopulations in the adult mouse forebrain. Eur. J. Neurosci. 11, 4213–4225 (1999). 33. Chevaleyre, V. & Castillo, P.E. Heterosynaptic LTD of hippocampal GABAergic synapses: a novel role of endocannabinoids in regulating excitability. Neuron 38, 461–472 (2003). 34. Sun, H.Y., Bartley, A.F. & Dobrunz, L.E. Calcium-permeable presynaptic kainate receptors involved in excitatory short-term facilitation onto somatostatin interneurons during natural stimulus patterns. J. Neurophysiol. 101, 1043–1055 (2009). 35. Alle, H. & Geiger, J.R. Combined analog and action potential coding in hippocampal mossy fibers. Science 311, 1290–1293 (2006). 36. Cossart, R., Esclapez, M., Hirsch, J.C., Bernard, C. & Ben-Ari, Y. GluR5 kainate receptor activation in interneurons increases tonic inhibition of pyramidal cells. Nat. Neurosci. 1, 470–478 (1998). 37. Frerking, M., Malenka, R. & Nicoll, R. Synaptic activation of KARs on hippocampal interneurons. Nat. Neurosci. 1, 479–486 (1998). 38. Christensen, J.K., Paternain, A.V., Selak, S., Ahring, P.K. & Lerma, J. A mosaic of functional kainate receptors in hippocampal interneurons. J. Neurosci. 24, 8986–8993 (2004). 39. Di Marzo, V. et al. Formation and inactivation of endogenous cannabinoid anandamide in central neurons. Nature 372, 686–691 (1994). 40. Chevaleyre, V. & Castillo, P.E. Endocannabinoid-mediated metaplasticity in the hippocampus. Neuron 43, 871–881 (2004). 41. Kang, J., Jiang, L., Goldman, S.A. & Nedergaard, M. Astrocyte-mediated potentiation of inhibitory synaptic transmission. Nat. Neurosci. 1, 683–692 (1998). 42. Hudson, B.D., Hebert, T.E. & Kelly, M.E. Ligand and heterodimer directed signaling of the CB1 cannabinoid receptor. Mol. Pharmacol. published online, doi:10.1124/ mol.109.060251 (16 October 2009). 43. Varma, N. et al. Presynaptic factors in the regulation of DSI expression in hippocampus. Neuropharmacology 43, 550–562 (2002). 44. Heifets, B.D., Chevaleyre, V. & Castillo, P.E. Interneuron activity controls endocannabinoid-mediated presynaptic plasticity through calcineurin. Proc. Natl. Acad. Sci. USA 105, 10250–10255 (2008). 45. Edwards, D.A., Zhang, L. & Alger, B.E. Metaplastic control of the endocannabinoid system at inhibitory synapses in hippocampus. Proc. Natl. Acad. Sci. USA 105, 8142–8147 (2008). 46. Glickfeld, L.L. & Scanziani, M. Distinct timing in the activity of cannabinoidsensitive and cannabinoid-insensitive basket cells. Nat. Neurosci. 9, 807–815 (2006). 47. Klausberger, T. & Somogyi, P. Neuronal diversity and temporal dynamics: the unity of hippocampal circuit operations. Science 321, 53–57 (2008).

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ONLINE METHODS Animals. Experimental procedures followed the recommendations of the Centre National de la Recherche Scientifique Ethics Committee and the French Ministry of Agriculture and Forestry concerning animal care (authorization number A33093). C57BL/6 mice were purchased from Janvier or were bred at the NeuroCentre Magendie. GluK1−/−, GluK2−/− and GluK1−/−; GluK2−/− double mutants were obtained and genotyped as described previously28. These mice were backcrossed on a C57BL6 genetic background (for 11 generations)48, and wild-type C57BL6 mice were used as controls. Full CB1 receptor mutant mice (CB1−/−) were obtained and genotyped as described49. Conditional mouse mutants lacking CB1 receptors from cortical glutamatergic neurons (Glu-CB1−/−), from GABAergic neurons (GABA-CB1−/−) or from forebrain pyramidal neurons (CaMK-CB1−/−) were obtained using the cre-loxP system and maintained and genotyped as described previously15,16,20. Specifically, CB1f/f;Dlx5/6-Cre (here referred as GABA-CB1−/− mice) were obtained by crossing mice carrying a loxP-flanked (‘floxed’) CB1 allele (CB1-floxed mice, CB1f/f ) with transgenic mice expressing Cre under the control of the I56i and I56ii intergenic enhancer sequences of the Dlx5 and Dlx6 genes (Dlx5/6-Cre mice) to generate selective deletion of CB1 receptors in GABAergic neurons16. CB1f/f;NEX-Cre (here referred as Glu-CB1−/− mice) were obtained by crossing CB1f/f mice with transgenic mice expressing Cre under the control of the regulatory sequences of the NEX gene (Nex-Cre mice) to generate selective deletion of CB1 receptors in glutamatergic cortical neurons20. CB1f/f;CaMKIIα-Cre mice (here referred as CaMK-CB1−/− mice) were obtained by crossing CB1f/f mice with mice expressing Cre recombinase under the control of the regulatory sequences of the Ca2+/calmodulin-dependent kinase II-α (Camk2a) gene (CaMKIIα-Cre mice) to generate mice lacking CB1 receptors in all principal neurons of the forebrain but maintaining their expression in cortical GABAergic interneurons15,16. Given the mixed genetic background of these CB1 mutant mice15,16,20, littermate controls were used. Drugs. Drugs were obtained from Sigma, Tocris and Caymann Chemical. SR141617A was kindly provided by US National Institutes of Mental Health Chemical Synthesis and Drug Supply Program. GYKI53655 was synthesized on demand by ABX GmbH. To pharmacologically block GluK1 KARs, we used LY382884 (a kind gift from D. Lodge, Eli Lilly) or the commercially available compound UBP302. Electrophysiology. Parasagittal hippocampal slices (320 µm thick) were obtained from 15- to 21-d-old mice that were killed by cervical dislocation. Slices were transferred to a recording chamber, in which they were continuously superfused with oxygenated (95% O2 and 5% CO2) ACSF containing 125 mM NaCl, 2.5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 1.25 mM NaH2PO4, 26 mM NaHCO3 and 16 mM glucose (pH 7.4). For the paired recording experiments, NaCl was replaced by an equimolar concentration of choline in the cutting solution. Slices were used within 6 h of cutting. Whole-cell voltage-clamp recordings were made at 24–25 °C (except for experiments in Supplementary Fig. 7, which were made at 33–34 °C) from CA1 pyramidal cells under infrared differential interference contrast imaging. Unless stated otherwise, the intracellular solution contained 145 mM CsCl, 10 mM HEPES, 5 mM EGTA, 2 mM MgCl2, 2 mM CaCl2, 2 mM Na2ATP, 5 mM phospho-creatine and 0.33 mM GTP (pH 7.2). Neurons were voltage-clamped at −70 mV, unless otherwise indicated. d-AP5 (50 µM), GYKI53655 (50 µM) and CGP 55845 (5 µM) were present in the superfusate of all experiments unless otherwise indicated. The access resistance of the cells was 20% over the course of the experiment. Recordings were made using either an EPC 9.0 or an EPC 10.0 amplifier (HEKA Elektronik) and were filtered at 0.5–1 kHz, digitized at 1–5 kHz and stored on a personal computer for additional analysis (IGOR PRO 5.0, Wavemetrics). Unless stated otherwise, IPSCs were evoked via a glass microelectrode positioned in the stratum radiatum 100 µm from CA1 pyramidal cell layer. For t-Di, a second distal bipolar stimulation was positioned at the boundary between CA1 and CA3 subfields (if not stated otherwise, see Supplementary Fig. 7) to stimulate Schaffer collaterals. After reaching stable baseline (10 consecutive eIPSCs separated by 30 s, not differing more than approximately 10%), t-Di and/or DSI protocols were applied. DSI was induced by a 5-s (5 s at 0 mV) or 1-s (1 s at 0 mV) voltage step to 0 mV applied to the postsynaptic pyramidal neuron. t-Di was induced with a conditioning train of 10 stimuli at 200 Hz

doi:10.1038/nn.2481

given to Schaffer collaterals or by theta burst stimulation (5 × 5 pulses at 100 Hz, interval time 200 ms; Supplementary Fig. 7). Each stimulus preceded one eIPSC of 200 ms (total of ten). The changes in eIPSC amplitude following the conditioning train were calculated by averaging ten consecutive responses separated by 30 s. Glutamate was applied to visually identified somata and apical dendrites of CA1 pyramidal neurons by an ultraviolet flash photolysis (Xenon flash lamp) of MNI-caged l-glutamate (100 µM, Tocris). The efficiency of the caged compound was analyzed by determining EPSCs amplitude in the absence of AMPA receptor blockade. AMPA receptor activation gave rise to a large EPSC of several pA of amplitude (data not shown). During the application of caged glutamate, a total amount of 10 ml of ACSF containing the caged compound was continuously re-circulated and oxygenated. This solution was not used for more than 1 h. One pulse light of 20 ms was applied at the beginning of a depolarization step of 1 s at 0 mV DSI (see Supplementary Fig. 6). Paired recordings between GABAergic interneurons and CA1 pyramidal cells were made with a double amplifier EPC 10.0. Interneurons in the stratum radiatum close to stratum pyramidale were visually identified and recorded in the current-clamp mode with a patch pipette containing 140 mM potassium gluconate, 10 mM HEPES, 5 mM EGTA, 3 mM MgCl2, 10 mM phospho-creatine and 0.2 mM GTP. The spiking pattern of the interneurons was determined, after achieving whole-cell configuration, by current injection trials consisting of a 300-ms square pulse. Adaptation coefficient was analyzed at maximum spiking frequency and measured as the ratio between firing frequency during the last 100 ms and instantaneous firing frequency between the first two spikes triggered by the pulse50. In all connected pairs (n = 36 of 325), DSI (5 s at 0-mV depolarization step) was tested to check for the presence of functional ECS signaling. In the presynaptic subthreshold depolarization experiment (Supplementary Fig. 9), mild DSI-inducing stimulations (1 s 0 mV) were applied to the post synaptic pyramidal neurons while alternatively holding the interneuronal membrane potential at −90 mV or −50 mV. The holding potential of the postsynaptic pyramidal neuron was not changed during the experiment. The potential of −50 mV was chosen on the basis of the observation that neuronal firing of interneurons in adaptation ramp tests never occurred before a thres hold potential of −43 to −40 mV (data not shown). No change was observed in baseline amplitudes of uIPSC on changes of interneuronal membrane potential (data not shown). Immunohistochemistry. Mice were anesthetized with pentobarbital and killed by transcardial perfusion with 4% paraformaldehyde in phosphate-buffered saline (pH 7.4). The brains were post-fixed for 1 h, transferred to 30% sucrose and incubated at 4 °C overnight. Brains were frozen and kept at −80 °C. We cut 30-µm cryostat sections and stored them in cryoprotection solution (25% (vol/vol) glycerin and 25% (vol/vol) ethylene glycol in phosphate-buffered saline). The sections were incubated as previously described16 with a polyclonal antibody to CB1 (1:1,000, L15, a kind gift of K. Mackie) at 4 °C overnight, followed by incubation with a biotinylated antibody to rabbit (ABC Kit, Vector Laboratories). The sections were stained with diaminobenzydine (Sigma-Aldrich Chemie GmbH), washed, mounted on slides and air dried overnight. They were mounted with Eukitt (O. Kindler GmbH) and analyzed in bright field with a Olympus DP20 stereomicroscope and a Leica CTR 2000 microscope. In situ hybridization. Probes and tissues were prepared as described using DIGlabeled riboprobes to mouse CB1 receptor15,32 and FITC-labeled riboprobes to rat GluK1 (ref. 18). For signal amplification, we used the TSA Plus System Cyanine 3/Fluorescein (PerkinElmer). Blocking buffer TNB and wash buffer TNT were prepared according to manufacturer’s protocols. Slides were analyzed by epifluorescence microscopy at 40× (Leica). Statistical analysis. Supplementary Tables 1–3 detail the statistical parameters and analyses that we used. Values are presented as mean ± s.e.m. of n experiments. DSI was calculated by comparing the mean amplitude (pA) of the first three IPSCs after the voltage step to the mean of last five IPSCs immediately preceding the step and expressed as relative values10. In experiments involving train stimulations, raw data (amplitudes, pA) were obtained from blocks of ten sweeps alone or preceded by the Schaffer collaterals stimulation. Train effect was

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x − x1 expressed as a relative value obtained as ∆ of eIPSCs = 100 × 2 , where x1 x1

48. Sachidhanandam, S., Blanchet, C., Jeantet, Y., Cho, Y.H. & Mulle, C. Kainate receptors act as conditional amplifiers of spike transmission at hippocampal mossy fiber synapses. J. Neurosci. 29, 5000–5008 (2009). 49. Marsicano, G. et al. The endogenous cannabinoid system controls extinction of aversive memories. Nature 418, 530–534 (2002). 50. Ascoli, G.A. et al. Petilla terminology: nomenclature of features of GABAergic interneurons of the cerebral cortex. Nat. Rev. Neurosci. 9, 557–568 (2008).


is the mean of raw IPSC amplitudes before train stimulation and x2 is the mean of raw IPSC amplitudes after each train stimulation. For DSI and t-Di, statistical analyses were conducted on relative values (∆). To analyze the effect of the train, we used the one-sample t test on relative data compared with the hypothetical value of 0 (no effect of train). Group differences were analyzed using the following methods. Paired t test was used to compare conditions that allowed recordings of the same neuron before and after drug treatments (for example, Fig. 1c,d). Unpaired t test was used to compare two independent groups of neurons (for example, Fig. 2c).

One-way ANOVA was used to compare several independent groups. When the ANOVA resulted in a significant general group effect, Dunnett’s post hoc test was used to compare different groups versus the control group, always applying family-wise 95% confidence levels, (for example, Figs. 1e and 4b).

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doi:10.1038/nn.2481