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Department of Neuroscience, University of Connecticut Health Center, Farmington, Connecticut
Submitted 23 July 2007; accepted in final form 17 September 2007
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ABSTRACT |
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INTRODUCTION |
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). Postsynaptic depolarization and subsequent calcium influx triggers the release of eCBs that can act as fast retrograde signaling molecules that activate presynaptic type 1 cannabinoid (CB1) receptors. Activation of these receptors can transiently inhibit the release of GABA or glutamate in processes termed depolarization-induced suppression of inhibition (DSI) or excitation (DSE). DSI and DSE are found in many different regions of the brain, including, but not limited to, cerebellum (Kreitzer and Regehr 2001; Llano et al. 1991), hippocampus (Ohno-Shosaku et al. 2001; Pitler and Alger 1992; Wilson and Nicoll 2001), and neocortex (Fortin and Levine 2007; Trettel and Levine 2003). In the neocortex, brief trains of action potentials (APs) in pyramidal neurons triggers DSI of a subset of inhibitory synapses, but it is not known whether inhibitory synapses onto interneurons also express CB1 receptors and whether interneurons themselves can release eCBs onto afferent terminals.
The neocortex is comprised of six discrete layers that contain two major groups of neurons: pyramidal neurons (PNs), which constitute 70–80% of the neuronal population, and local circuit neurons, or interneurons (INs), which constitute the remaining 20–30% of the total population of neurons. PNs are glutamatergic neurons that have relatively stereotyped anatomical, biochemical, and electrophysiological properties, and these cells provide the sole output of the cortex to other brain areas. Local circuit interneurons have varied morphologies, express different calcium-binding proteins (calretinin, calbindin and/or parvalbumin), and although typically GABAergic, they can also express a number of different neuropeptide co-transmitters, e.g., somatostatin, vasoactive intestinal peptide, neuropeptide Y, and cholecystokinin (DeFelipe 2002; Markram et al. 2004). These INs play an important role in regulating and synchronizing the activity of PNs. In particular, a well-defined subset of GABAergic basket cells that co-express the neuropeptide cholecystokinin has been shown to express the CB1 receptor (Bodor et al. 2005; Galarreta et al. 2004; Marsicano and Lutz 1999; Tsou et al. 1999).
Although eCBs and CB1 receptor activation play a role in regulating afferent inhibition onto PNs, it is not clear whether they serve a similar function at interneuron-interneuron synapses. In the hippocampus, it has been shown that INs in the s. radiatum and the s. oriens do receive cannabinoid-sensitive inputs (Hoffman et al. 2003) but fail to show DSI in response to postsynaptic depolarization (Hoffman et al. 2003; Patenaude et al. 2005), although a recent report suggests that a subset of these INs may show DSI (Ali 2007). In cortical layer 5 low-threshold-spiking (LTS) cells, however, prolonged depolarization induces an autocrine hyperpolarization mediated by activation of somatodendritic CB1 receptors, suggesting that these INs can release eCBs, although effects on synaptic inputs were not examined (Bacci et al. 2004). In a recent paper (Beierlein and Regehr 2006), it was demonstrated that interneurons in the cerebellum, specifically stellate cells and basket cells, can mobilize eCBs to modulate excitatory synaptic inputs. These cells showed DSE and synaptically evoked suppression of excitation, which required postsynaptic elevations of free intracellular Ca2+, was blocked by CB1 receptor antagonists, and was lacking in CB1-deficient mice. Thus at least in some situations, local circuit neurons can mobilize eCBs to regulate afferent inputs. In this paper, we set out to compare the cannabinoid sensitivity of inhibitory inputs to the different cell types in layer 2/3 and to determine whether interneurons in layer 2/3 can mobilize endocannabinoids and induce DSI.
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METHODS |
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All experiments were performed on postnatal day 14–21 Swiss CD-1 mice (Charles River, Wilmington, MA) using protocols that were approved by University of Connecticut Health Center Animal Care Committee. Briefly, animals were killed by rapid decapitation without anesthesia, and whole brains were removed and immersed in ice-cold "cutting and incubating solution" composed of (in mM) 125 NaCl, 2.5 KCl, 1.25 NaH2PO4, 25 NaHCO3, 0.5 CaCl2, 4 MgCl2, 4 MgSO4, 4 lactic acid, 2 pyruvic acid, 20 glucose, 0.4 ascorbic acid, and 0.25 kynurenic acid equilibrated with 95% O2-5% CO2 (pH 7.3, 310 ± 5 mmol/kg). Transverse slices (300–350 µm) containing auditory and somatosensory cortex (Paxinos and Franklin 2001) were cut with a Dosaka EM DTK-1000 vibratome (Kyoto, Japan) and incubated at room temperature. After 50–60 min, slices were transferred to a recording chamber (room temperature) fixed to the stage of an Olympus BX51WI upright microscope equipped with a x40 water-immersion objective lens (0.8 NA). During recordings, slices were continuously perfused at 2 ml/min with oxygenated bath solution consisting of (in mM) 125 NaCl, 2.5 KCl, 1.25 NaH2PO4, 25 NaHCO3, 2 CaCl2, 2 MgCl2, and 15 glucose (pH 7.3, 310 ± 5 mmol/kg); pH was equilibrated by continuous bubbling with 95% O2-5% CO2.
Chemicals
Unless otherwise stated, all drugs were from Sigma-Aldrich (St Louis, MO). All drugs were delivered by bath perfusion. Stock solutions of the cannabinoid agonist WIN55,212-2, and the CB1 receptor antagonists SR141716A (RTI International, Research Triangle Park, NC) and 1-(2,4-dichlorophenyl)-5-(4-iodophenyl)-4-methyl-N-1-piperidinyl-1H-pyrazide-3-carboxamide (AM251; Tocris Bioscience, Ellisville, MO) were dissolved in 100% DMSO at 5 x 103 times their final concentration and stored at –20°C. This gave a final DMSO concentration of 0.02%, which by itself had no effect on synaptic transmission (Trettel and Levine 2002). AM404 (Tocris Bioscience, Ellisville, MO) was dissolved in Tocrisolve and carbamylcholine chloride (carbachol) was made in water and stored at 0.01 M.
Electrophysiology and Ca2+ imaging
Whole cell patch recordings were obtained from layer 2/3 PNs and INs identified using infrared differential interference contrast video microscopy and AP firing patterns. Patch electrodes were pulled from borosilicate glass capillaries using a Flaming/Brown P-97 micropipette puller (Sutter Instruments, Novato CA). For whole cell recordings of spontaneous inhibitory postsynaptic potentials (sIPSPs) in current-clamp mode, the pipette solution contained (in mM) 65 K-gluconate, 85 KCl, 10 HEPES, 1 EGTA, 0.1 CaCl2, 4 Na2-ATP, 0.3 Na-GTP, and 10 Tris-phosphocreatine, pH 7.3 (290 ± 5 mmol/kg). The membrane potential (Vm) was set at –70 mV after determining baseline resting Vm with no current injection. Input resistance (Ri) was measured with a –50-pA hyperpolarizing current injection for 500 ms. Frequency accommodation was measured as the percent drop in instantaneous frequency between the first and last interspike intervals in response to a 500-ms depolarizing current injection. Hyperpolarization sag potential was measured using a –250-pA hyperpolarizing current injection for 500 ms. The Nernst equilibrium potential for chloride (ECl) using our internal and external solutions was approximately –10 mV, thus sIPSPs were depolarizing and recorded as upward deflections. The ionotropic glutamate receptor antagonists 6,7-dinitroquinoxaline-2,3-dione (DNQX; 10 µM) and 3-(2-carboxypiperazin-4-propyl-1-phosphonic acid (CPP; 3 µM) were added to the bath solution to isolate inhibitory activity. All electrical events were filtered at 2.9 kHz and digitized at >6 kHz using a HEKA ITC-16 digitizer built into an EPC-9 amplifier (Heka Elektronic, Darmstadt, Germany). Neurons were rejected from analysis if the holding current to maintain a Vm of –70 mV increased by more than 50 pA, Ri changed by >15% during the course of an experiment, or Ri fell <100 M
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For recording miniature inhibitory postsynaptic currents (mIPSCs), 1 µM tetrodotoxin (TTX) was added to the bath solution to block voltage-dependent Na+ channels. The ionotropic glutamate receptor antagonists DNQX (10 µM) and CPP (3 µM) were used to isolate inhibitory activity. The pipette solution contained (in mM) 20 KCl, 100 K-gluconate, 10 HEPES, 1 EGTA, 0.1 CaCl2, 4 Na2-ATP, 0.3 Na-GTP, and 10 Tris-phosphocreatine, pH 7.3 (290 ± 5 mmol/kg). INs were chosen based on morphology and the presence of a sag potential when hyperpolarized with a –250-pA current injection. For these voltage-clamp recordings, series resistance (Rs) was compensated to 
60% at 10–100 µs lag. Neurons were rejected from analyses if Rs was >25 M at the time of break-in or >10.5 M after compensation, if Ri changed by >15% during the course of an experiment, or if Ri fell <100 M.
Clampfit (Molecular Devices, Sunnyvale, CA) was used to detect and analyze mIPSCs and sIPSPs. We quantified synaptic activity by integrating the area under individual sIPSPs, an index that takes into account event frequency, amplitude and duration. DSI was calculated as the percent suppression in area measured from 5 to 15 s after depolarization. Group data are presented as means ± SE. Tests of statistical significance were based on Student's paired t-test unless otherwise noted.
For Ca2+ imaging experiments, the cell-impermeant form of Oregon Green 488 BAPTA-1 (100 µM, Molecular Probes, Carlsbad, CA) was dissolved in the internal pipette solution. Recording of Ca2+ fluorescence was carried out 15–20 min after breaking into whole cell mode to allow equilibration of the dye. Changes in Ca2+ fluorescence were detected by a cooled charge-coupled device camera (IMAGO-QE, TILL Photonics, Munich, Germany) at a sampling rate of 20 Hz. Intracellular changes in postsynaptic [Ca2+]i are expressed in terms of 
F/F0, where F0 is the average background-subtracted fluorescence intensity when the cell was at rest and F is the change in fluorescence intensity immediately after the depolarizing stimuli protocol. Fluorescence intensity was measured in an 
3 x 3 µm region within the soma. A similar size region of interest was used to detect changes in background fluorescence.
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RESULTS |
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PNs and INs in neocortical layer 2/3 were identified morphologically and classified by AP firing patterns in response to 500-ms current injection pulses ranging from –250 to +250 pA. As shown in Fig. 1, PNs typically displayed a regular firing pattern with amplitude and frequency adaptation, and INs were classified as FS, IS, or RSNP. RSNP cells (Fig. 1D) were the most common and displayed a regular firing pattern with some degree of amplitude and frequency adaptation but were able to fire at higher frequencies than PNs. They also showed a noticeable transient deflection (or sag) when hyperpolarized, which was also seen in other IN types and to a lesser extent in PNs. IS cells (Fig. 1B) fired irregularly in response to depolarizing current pulses and their APs exhibited a noticeable fast afterhyperpolarization (AHP). FS interneurons had the largest and fastest AHPs and discharged at a much higher rate than any other interneuron type; the example FS cell shown in Fig. 1C is firing at 160 Hz. The physiological properties of PNs and the different classes of INs are shown in Table 1. In general, INs had a more depolarized resting membrane potential, shorter spike duration, and more pronounced afterhyperpolarization compared with PNs.
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100 µm from the soma as it approaches the superficial cortical layer 1. The axon can also be seen heading toward deeper cortical layers. IS cells, which were found mostly near the border with layer 1, typically had elongated somas and bipolar processes (Fig. 1B). Both FS and RSNP cells were typically multipolar. Some FS cells, like the one shown in Fig. 1C, had a chandelier-like morphology with two long dendrites emerging from the soma and pointing, almost parallel, toward layer 1 while the axon emerges from the opposite direction and split close to the soma. The example shown for the RSNP cell (Fig. 1D) is a typical multipolar cell with processes splitting either immediately or within 10–20 µm from the soma. Layer 2/3 interneurons receive cannabinoid-sensitive inhibitory inputs
We next investigated whether layer 2/3 INs receive inhibitory inputs that can be suppressed by cannabinoids. In these experiments, we utilized bath application of the muscarinic agonist CCh, which causes a marked increase in the frequency of sIPSPs recorded from cortical PNs. This CCh-induced activity in PNs is potently suppressed by CB1R activation (Trettel et al. 2004). We obtained similar results with PNs in the present study as shown in the top trace in Fig. 2. Interestingly, we found that bath-applied CCh (10 µM) also enhanced sIPSP activity in all three classes of interneurons (Fig. 2), and the magnitude of the effect was similar across all cell types. Quantifying synaptic area to account for increases in both frequency and amplitude of sIPSPs, we found that activity was increased 582 ± 123% in PNs (n = 29), 410 ± 66% in IS cells (n = 18), 622 ± 139% in FS cells (n = 25), and 598 ± 140% in RSNP cells (n = 25). All cell types also showed sustained depolarization of the resting membrane potential in response to CCh. As shown in Fig. 2, depolarization was greatest in IS cells (15.8 ± 1.4 mV) and often led to AP firing. The depolarization was smaller but also significant in the other cell types (PN: 5.9 ± 0.7 mV; FS: 4.5 ± 1.1 mV; RSNP: 7.8 ± 1.6 mV). The depolarizing effects of CCh likely result from direct effects on PNs as well as indirect effects due to the increase in sIPSPs, which are depolarizing under our experimental conditions.
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Lack of interneuron DSI
We next investigated whether INs can synthesize and release eCBs by examining whether DSI occurs at IN-IN synapses. First, we confirmed the presence of DSI at inhibitory synapses on PNs, as previously reported. Figure 4A shows a typical example of DSI in a layer 2/3 PN after a 20-AP/40-Hz train, and the blockade of DSI by a CB1 receptor antagonist. Overall, as shown in Fig, 4B, sIPSP area in PNs was transiently suppressed by an average of 40.7 ± 6.9% (n = 9, P < 0.05). DSI was blocked by pretreatment with the CB1 receptor antagonist AM251 or SR141716A (6 µM; 12.0 ± 12.9%, n = 5).
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800 APs and frequencies as high as 100 Hz. Figure 5A shows examples of traces obtained from each class of IN (IS: top; RSNP: middle; and FS: bottom). Each trace illustrates a 25-s pre-DSI period (baseline), followed by an AP train of 400 APs at 80 Hz (omitted for clarity), and the period immediately after the AP train. The group data shown in Fig. 5B indicates the amount of suppression from each class of IN using a 400-AP/80-Hz train. Note that for the FS cells, the small but consistent suppression that was observed under baseline conditions (12.6 ± 8.0%, n = 9) was unchanged in the presence of the CB1 receptor antagonist AM251 (11.4 ± 3.2%, n = 5), indicating that this transient suppression was not CB1 receptor-mediated. Thus in layer 2/3, only PNs appeared to be capable of mobilizing cannabinoids and inducing DSI in response to AP trains.
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; Trettel and Levine 2003; Wilson and Nicoll 2001). It is possible that the lack of interneuron DSI in the present study is due to an insufficient rise in intracellular calcium. In fact, interneurons have been shown to have a high [Ca2+]i buffering capacity compared with PNs (Baimbridge et al. 1992). We therefore monitored the change in intracellular calcium in PNs and INs in response to DSI induction protocols, using the membrane impermeable fluorescent calcium dye, Oregon Green 488 BAPTA-1 (100 µM). After breaking into the cell and allowing 10–20 min for the dye to equilibrate between the pipette and the cell, we collected fluorescence images before, during, and after various DSI induction protocols (trains of 20, 50, 100, or 200 APs at 40 or 80 Hz). In PNs, the magnitude of the Ca2+ signal doubled in response to a train of 20 APs at 40 Hz, a protocol that was sufficient to induce reliable DSI (Fig. 6A). The different classes of INs all showed a smaller calcium increase in response to that particular protocol, but increasing the frequency and duration of the AP train led to relative increases in the Ca2+ signal comparable to that seen in PNs (Fig. 6, B–E). As shown in Fig. 6F, a minimum of 100 APs was able to achieve a relative increase in the Ca2+ signal in all classes of INs similar to that seen with DSI induction in PNs, but as shown in the preceding text, these protocols were still unable to induce DSI in INs. Thus the lack of DSI at IN-IN synapses does not appear to be due to an insufficient rise in intracellular Ca2+.
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DISCUSSION |
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; Trettel et al. 2004). In the present study, we examined the role of cannabinoids in modulating inhibitory inputs to three distinct classes of physiologically identified neocortical INs. We found that all three classes of interneurons, FS, IS, and RSNP, received inhibitory inputs that could be activated by muscarinic stimulation, and these inhibitory inputs were suppressed by CB1 receptor activation to an extent similar to that seen in PNs. This cannabinoid-mediated suppression was a presynaptic effect, indicating that CB1 receptors are expressed at inhibitory synapses onto INs as well as PNs. Of the cell types recorded in layer 2/3, it is likely that the IS cells provide some of the cannabinoid-sensitive input to INs. IS cells were the only cells that showed a marked depolarization in the presence of CCh sufficient to increase AP discharge. In addition, Galarreta et al. (2004) demonstrated that layer 2/3 INs (with multipolar and/or bitufted morphologies) that express CB1 receptors exhibit electrophysiological characteristics of IS cells. For those reasons, we believe that it is most likely that the IS cells are responsible for the increase in sIPSPs induced by CCh application.
In marked contrast to PNs, there was no eCB-mediated DSI of inhibitory inputs onto INs, even with prolonged high-frequency spike trains. One of the requirements for activity-induced DSI is a steep increase in free intracellular calcium concentration ([Ca2+]i). Estimates of the level of calcium needed to induce DSI range from 4 µM in hippocampal CA1 pyramidal neurons (Wang and Zucker 2001) to 15 µM for the Purkinje cells in the cerebellum (Brenowitz and Regehr 2003), although retrograde inhibition may also be triggered by lower calcium levels (Glitsch et al. 2000). In the present studies, we found that the relative calcium increase in INs is less than seen in PNs for the same activity trains. INs express higher levels of calcium binding proteins such as calbindin D28k, parvalbumin, and calretinin, and therefore they generally have a high [Ca2+]i buffering capacity compared with the PNs (Baimbridge et al. 1992). However, the lack of DSI did not appear to result from an insufficient rise in intracellular calcium because prolonged high-frequency spike trains in INs resulted in relative calcium increases comparable to what is seen in PNs during DSI induction. It should be noted, however, that absolute calcium concentrations were not monitored, and baseline calcium levels may differ between INs and PNs. These results are in agreement with a study in the hippocampus, where it was reported that INs in the s. oriens and s. radiatum also receive cannabinoid-sensitive inputs but fail to show DSI (Hoffman et al. 2003). A more recent study, however, suggests that DSI may occur between cholecystokinin-expressing INs in the hippocampus (Ali 2007). In the neocortex, although it has been shown that layer 5 LTS cells can release endocannabinoids to modulate self-excitability (Bacci et al. 2004), it is not clear whether these INs can also produce DSI or DSE.
The lack of DSI at IN-IN synapses suggests that either layer 2/3 INs are not capable of producing endocannabinoids or releasing them at relevant synaptic sites or that insufficient stimulation was delivered to mobilize eCBs from these cells. INs might be lacking some of the components of the cellular enzymatic cascade required for eCB synthesis. It is also possible that there is a lack of co-localization between the site of eCB release by INs and the presynaptic terminals expressing CB1 receptors. A related explanation could be the presence of more efficient endocannabinoid re-uptake at IN-IN synapses compared with IN-PN synapses, although the inability of the reuptake inhibitor AM404 to unmask DSI in interneurons suggests that this is probably not the case. It may be that depolarization alone is not a sufficient trigger for eCB mobilization in these INs, and some combination of neuronal activity plus metabotropic stimulation (other than muscarinic) or NMDA receptor activation is required. However, if INs do not release eCBs, could there be some other functional relevance for CB1 receptor expression at IN-IN synapses? One possibility is that these receptors are activated by spillover of eCBs from neighboring PNs when PNs display high levels of prolonged activity. Under these conditions, activation of CB1 receptors at IN-IN synapses would disinhibit IN activity, and the resulting increase in inhibitory input to PNs could act as a negative feedback mechanism to dampen PN excitability.
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GRANTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: E. S. Levine, Dept. of Neuroscience, MC-6125, University of Connecticut Health Center, 263 Farmington Ave., Farmington, CT 06030 (E-mail: eslevine{at}neuron.uchc.edu)
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TOP
ABSTRACT
INTRODUCTION
) or in the added presence of the type 1 cannabinoid receptor (CB1) antagonist AM251 or SR141716A (6 µM; 
). Synaptic activity was integrated over a 1-min period occurring 5 min after onset of WIN application compared with a 1-min baseline period just prior to WIN application. C: group data for the effect of WIN on the frequency and amplitude of miniature inhibitory postsynaptic currents recorded from interneurons in the presence of 1 µM tetrodotoxin. The number above each bar denotes the number of cells tested. *, P < 0.05. Inset: example of the cumulative amplitude distribution of mIPSCs before and during WIN exposure. Note the decreased occurrence of large-amplitude events in the presence of WIN.
, occurrence of the AP train, which is blanked for clarity and shown in inset. Scale bars, 10 mV, 10 s. Scale bars for inset, 50 mV, 200 ms. B: normalized group data illustrating the magnitude and time course of DSI in the absence (
, n = 9) or presence (
