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The Journal of Neuroscience, September 29, 2004, 24(39):8606-8620; doi:10.1523/JNEUROSCI.2660-04.2004
 Previous Article | Next Article 
Cellular/Molecular
ATP Excites Interneurons and Astrocytes to Increase Synaptic Inhibition in Neuronal Networks
David N. Bowser and
Baljit S. Khakh
Medical Research Council Laboratory of Molecular Biology, Cambridge CB2 2QH, United Kingdom
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Abstract
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We investigated the role of extracellular ATP at astrocytes and inhibitory GABAergic interneurons in the stratum radiatum area of the mouse hippocampus. We show that exogenously applied ATP increased astrocyte intracellular Ca2+ levels and depolarized all calbindinand calretinin-positive interneurons in the stratum radiatum region of mouse hippocampus, leading to action potential firing and enhanced synaptic inhibition onto the postsynaptic targets of interneurons. Electrophysiological, pharmacological, and immunostaining studies suggested that the effect of ATP on interneurons was mediated by P2Y1 receptors, and that the depolarization of interneurons was caused by the concomitant reduction and activation of potassium and nonselective cationic conductances, respectively. Electrical stimulation of the Schaffer collaterals and perforant path, as well as local stimulation within the stratum radiatum, evoked increases in intracellular Ca2+ in astrocytes. Facilitation of GABAergic IPSCs onto interneurons also occurred during electrical stimulation. Both the stimulation-evoked increases in astrocyte Ca2+ levels and facilitation of GABAergic IPSCs were sensitive to antagonists of P2Y1 receptors and mimicked by exogenous P2Y1 receptor agonists, suggesting that endogenously released ATP can activate P2Y receptors on both astrocytes and interneurons. Overall, our data are consistent with the hypothesis that ATP released from neurons and astrocytes acts on P2Y1 receptors to excite interneurons, resulting in increased synaptic inhibition within intact hippocampal circuits.
Key words: purinergic; interneuron; astrocyte; inhibition; IPSP; hippocampus; P2Y; P2X
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Introduction
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Increasing evidence suggests that astrocytes actively participate in synaptic and neuronal function as well as fulfill their classical supportive roles in the brain (Fields and Stevens, 2000 ; Haydon, 2001; Fields and Stevens-Graham, 2002; Newman, 2003a). Astrocytes release gliotransmitters into the extracellular space that may then activate receptors on nearby neurons (Araque et al., 2001). ATP functions as an extracellular signaling molecule (Zimmermann, 1994; Burnstock, 2004), and recent work suggests that ATP is also a gliotransmitter. Once released from a point source, ATP may trigger a wave of astrocyte excitation that can travel hundreds of micrometers (Guthrie et al., 1999; Haydon, 2001; Newman, 2001a,b, 2003a). Recently, glial ATP has been shown to decrease the excitability of neurons in the retina (Newman, 2003b) and mediate presynaptic modulation in cultured hippocampal neurons (Koizumi et al., 2003; Zhang et al., 2003). An important unresolved issue, however, is whether endogenously released glial ATP actually regulates the excitability of brain neurons in networks with functionally preserved connectivity (Haydon, 2001). In this report, we present evidence to suggest that endogenously released ATP can affect the function of nearby astrocytes and interneurons within the stratum radiatum (s.r.) and stratum lacunosum-moleculare (s.l-m.) regions of the mouse hippocampus.
A role for extracellular ATP as a signaling molecule in the nervous system was first proposed over 30 years ago (Burnstock, 1972). Extracellular ATP acts on two families of receptors, P2X and P2Y receptors. P2X receptors are ATP-gated nonselective cation channels that are found throughout the body (Khakh, 2001), whereas P2Y receptors are seven transmembrane domain receptors that couple to G-proteins (Lazarowski et al., 2003) and in some cases directly to channels (O'Grady et al., 1996; Lee et al., 2003). P2Y receptors comprise a family of at least eight members; they couple to Gq and Gi proteins and thus activate phospholipase C and inhibit adenylyl cyclase (Lazarowski et al., 2003). In relation to other transmitter receptor families, the functions of P2Y and P2X receptors in the brain are not well understood, and one important goal is to decipher how ATP affects brain neurons through the interplay of P2X and P2Y receptor signaling.
Inhibitory GABAergic interneurons have small range projections within the hippocampus and constitute a diverse group of neurons (McBain and Fisahn, 2001). S.r. interneurons in the CA1 region of the hippocampus receive inhibitory innervations from other local interneurons and form GABAergic synapses onto output pyramidal neurons to mediate feedforward IPSPs. By receiving excitatory input from CA3 neurons and evoking inhibition of interneurons and output CA1 pyramidal neurons, s.r. interneurons profoundly affect network activity within circuits (McBain and Fisahn, 2001). Astrocytes are abundant in the hippocampus; their processes profusely interleave between synapses in the s.r. (Ventura and Harris, 1999). However, the possibility that astrocytes may regulate interneuron excitability has remained unexplored. In the present study, we provide evidence for a novel ATP-mediated excitatory response in interneurons, which draws on neurons and astrocytes, leading to increased synaptic inhibition in interneuron networks.
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Materials and Methods
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Preparation of brain slices and electrophysiological recording. The methods used have been described previously (Khakh et al., 2003). Briefly, young [postnatal day 13 (P13) to P18] C57 mice were killed in accordance with home office and local procedures. Coronal slices of hippocampus (200 or 300 µm) were cut (model 1000 Plus; Vibratome, St. Louis, MO) and submerged at room temperature in artificial CSF (aCSF) comprising the following (in mM): 126 NaCl, 2.5 KCl, 1.3 MgCl2,10 D-glucose, 2.4 CaCl2, 1.24 NaH2PO4, and 26 NaHCO3 saturated with 95% O2 and 5% CO2. Experiments were performed at room temperature or 34°C as indicated. Experiments were performed with potassium gluconate or chloride-based internal solution comprising the following (in mM): 120 K-gluconate (or Cl-), 10 KCl, 1 MgCl2, 0.03 CaCl2, 0.1 EGTA, 1 ATP, 0.2 GTP, 10 HEPES, 4 glucose, pH 7.25. The resistance of the pipettes was  4-9M . Cells were visualized with infrared optics (900 nm; Olympus, Southall, UK) on an upright microscope (BX51; Olympus). Puffs of ATP were applied under visual control using a Picospritzer II (General Valve, Fairfield, NJ). In our previous work, we applied agonists directly to the bath (Khakh et al., 2003) over tens of seconds, but in the present study, we used a fast bath application system (RV-8; Warner Instruments, Hamden, CT), ensuring that oxygenated solutions were applied to the brain slice chamber within 500 msec for complete bath change in 1.5 sec. Synaptic currents were recorded in the presence of 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX; 10 µM) to block AMPA/kainate receptors. ATP-induced inward currents were recorded in CNQX (10 µM), AP-5 (10 µM), bicuculline (10 µM), and tetrodotoxin (TTX; 1 µM). The adenosine receptor antagonist cyclopentyl-1,3-dipropylxanthine (DPCPX; 10-100 µM) was always present in the bathing medium and ruled out P1 receptor involvement. The concentrations of agonists and antagonists used are higher than those used in biochemical studies of P2Y receptors because of breakdown and/or penetration barriers that are inherent with brain slice recording (North, 2002). Inward currents were recorded at -60 mV using pCLAMP 9 software, Multiclamp 900A amplifier, and Digidata 1322A (Axon Instruments, Foster City, CA).
Paired recordings. The whole-cell patch-clamp configuration (-60 mV) was attained for postsynaptic interneurons and pyramidal neurons with a KCl-based pipette solution containing 1 µM N-ethyl bromide quaternary salt of lidocaine to block action currents. Presynaptic interneurons were then patched with a K-gluconate-based pipette solution and held in current-clamp bridge mode. Depolarizing current (100-400 pA, 2-4 msec) was injected in the presynaptic neuron to evoke action potentials and to determine whether it was connected to the putative postsynaptic neuron. If there was no postsynaptic current after 50 trials, the pair was deemed unconnected, the presynaptic neuron pipette was withdrawn, and another presynaptic neuron was sought. This procedure was repeated until connected pairs were found. Consistent with previous work, s.r. interneuron-interneuron connectivity was found in 14 of 67 pairs (20.8%), and s.r. interneuron-CA1 pyramidal neuron connectivity was found in 11 of 132 pairs (8.3%) (Bertrand and Lacaille, 2001).
Confocal imaging of astrocyte activity. Brain slices were loaded at room temperature in the dark with 5 µM Fluo-4-AM (Molecular Probes, Invitrogen, Paisley, UK) in aCSF for 60 min and then transferred to dye-free aCSF for at least 30 min before experimentation to allow for cleavage of the AM ester group (Porter and McCarthy, 1996). Live astrocytes were predominantly loaded with the fluorescent dye with these conditions. This was confirmed by simultaneous acquisition of infrared differential interference contrast optics (IR-DIC) images of astrocytes in the same area. In some cases, individual interneurons were loaded through the patch pipette with Alexa-488 (50 µM) or biocytin (0.5%) to approximate neuron shape. Cells and slices were imaged using a Bio-Rad (Ringstead, UK) MRC-600 laser scanning confocal scan head with attached solid state 5 mW 488 nm laser light source (Laser 2000). Emitted green fluorescence was colleted through a 515 long-pass filter. Bio-Rad CoMOS software was used for image acquisition.
Immunofluorescent labeling in fixed sections. Young (P13-P18) C57 mice were perfused through the heart under deep anesthesia in accordance with home office and local procedures. The perfusate comprised 20 ml of 0.1 M PBS, followed by 50 ml of 4% paraformaldehyde in 0.1 M PBS (herein called fixative). Blocks of hippocampus, 1 mm thick, were placed in fixative and left overnight at 4°C and then cryopreserved in 20% sucrose (0.1 M PBS) overnight before cryosectioning (Leica CM3050S cryostat; Leica, Milton Keynes, UK). Free-floating 30 µm sections of hippocampus were then extensively washed in PBS, and nonspecific protein binding was blocked by a subsequent incubation in 3% normal goat serum (NGS) for 2 hr. Sections were then incubated in the following primary antibodies: rabbit anti-parvalbumin polyclonal antibody (1:2000, 23°C, 2 hr; Abcam, Cambridge UK), rabbit anti-calbindin polyclonal antibody (1:1000, 23°C, 2 hr; Chemicon, Hampshire, UK), rabbit anti-P2Y1 polyclonal antibody (1:200, 4°C overnight; Abcam), mouse anti-calretinin monoclonal antibody (1:50, 23°C, 12 hr; Abcam), and mouse anti-glial fibrillary acidic protein (GFAP) monoclonal antibody (1:1000, 23°C, 2 hr; Abcam). All antibodies were prepared in 1% NGS and 0.3% Triton X-100 in 0.1 M PBS. After extensive washes in 0.1 M PBS, sections were exposed to an appropriate secondary (anti-rabbit or anti-mouse IgG) antibody conjugated to either Alexa-488 or Alexa-647 (Molecular Probes) for 30 min at 23°C. Sections were then placed onto electrostatically charged slides and mounted underneath coverslips with Vectashield (Vector Laboratories, Peterborough, UK). Sections were imaged with a Bio-Rad Radiance confocal microscope equipped with a 488 nm emitting argon laser and a 637 nm emitting red diode enabling simultaneous acquisition of double fluorescently labeled sections.
Classification of interneurons. In a specific set of experiments, 0.5% biocytin was included in the intracellular solution during whole-cell recording from interneurons in 200 µm coronal sections of hippocampus. In some cases, interneurons were actively filled with biocytin by current injection for up to 20 min (+200 pA for 1 sec at 0.5 Hz). After recording and biocytin filling, interneurons were "depatched" by applying small amounts of positive pressure while slowly withdrawing the pipette to minimize cell damage and leaking of biocytin. Sections were then returned to oxygenated buffer for 1-2 hr to enable diffusion of biocytin to distant processes and then placed in fixative at 4°C overnight. The next day, sections were processed for immunoreactivity for the calcium-binding proteins calbindin or calretinin or for the astrocyte marker GFAP as described above for 30 µm fixed sections. After thorough washes in 0.1 M PBS, sections were incubated in a solution containing streptavidin-conjugated to Alexa-488 (dilution, 1:1000; Molecular Probes) as well as appropriate fluorescent secondary antibodies for visualization of calbindin, calretinin, or GFAP immunoreactivity as described above. Sections were placed on electrostatically charged slides and mounted underneath coverslips with Vectashield (Vector Laboratories). Sections were imaged with a Bio-Rad Radiance confocal microscope (see above). In many cases, we collected several image series going deep into the slice (Z-stacks) using a 100x oil immersion lens (Nikon, Tokyo, Japan). These images were later used to determine the shape and dendritic orientation of the biocytin-filled interneuron using Bio-Rad software, NIH ImageJ, and CorelDraw12 (Corel, Maidenhead, UK).
Chemicals. All chemicals used were from Tocris Cookson (Bristol, UK) or Sigma (Poole, UK). Fluorescent dyes were from Molecular Probes. The chemicals are pyridoxal-phosphate-6-azophenyl-2',4'-disulfonic acid (PPADS), CNQX, L-(+)-2-amino-5-phosphonopentanoic acid (L-AP-5), 2-methylthio-ATP (2MeSATP), 2-methylthio-ADP (2MeSADP), adenosine 5'-o-(2-thiodiphosphate) (ADP S), RS-3,5-dihydroxyphenylglycine (DHPG), and 2-methyl-6-(phenylethynyl)pyridine hydrochloride (MPEP).
Analysis. Synaptic currents were analyzed using MiniAnalysis program 5.6.22 (Synaptosoft, Decatur, GA), Strathclyde Electrophysiology Software (John Dempster, University of Strathclyde, Glasgow, Scotland, UK), pCLAMP9 (Axon Instruments), Origin 6.1 (OriginLab, Northampton, MA), and Graphpad Instat 3.06 for Windows (GraphPad Software, San Diego, CA). Postsynaptic unitary IPSC (uIPSC) latency was defined as the time between the maximum slope of the presynaptic action potential rising phase and the initiation of the uIPSC (defined as a 5% deviation from baseline). Confocal and epifluorescence imaging data were analyzed using Image J, custom written macros for the MRC-600 and CorelDraw12. Data in the text and graphs are shown as mean ± SEM from n determinations as indicated. Two tailed t tests were used for most statistical analyses with significance declared at p < 0.05. The Kolmogorov-Smirnov two-sample test was used to compare cumulative probability curves of sIPSC amplitudes and mIPSC interevent intervals with significance declared at p < 0.05.
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Results
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Unless otherwise stated, all of the experiments reported in this study are in the presence of the adenosine receptor antagonist DPCPX (10 µM), ruling out adenosine receptor involvement. We start by reporting the effects of exogenous ATP on single interneurons, astrocytes, and inhibitory transmission. We go on to study the effects of endogenous ATP on astrocyte signaling and synaptic inhibition in interneurons.
ATP evokes slow inward currents and depolarizations of stratum radiatum interneurons
S.r. interneurons were discriminated from CA1 pyramidal neurons with the aid of IR-DIC on the basis of anatomical location, characteristic somatic shape, and ability to fire high-frequency action potentials (see next section for interneuron classification). ATP, or the more stable analog ATP S, failed to evoke inward currents when puffed briefly to voltage-clamped s.r. interneuron somata (Khakh et al., 2003) (100 µM;10-500 msec; 2.6 ± 1.7 pA change in holding current at -60 mV; n = 12) (Fig. 1A). However, rapid bath application of ATPS (see Materials and Methods) evoked peak inward currents of -36.0 ± 6.7 pA (20-80% rise time; 25.7 ± 5.2 sec; n = 24) (Fig. 1B). These slow ATPS-evoked currents showed little desensitization for agonist applications of 180 sec, and slow-incomplete recovery such that the residual current remaining 5 min after washout was 36.6 ± 3.1% of that at the peak (n = 24) (Fig. 1C). In contrast to the results with ATPS, glutamate readily evoked reversible inward currents whether it was puffed briefly onto the soma to activate AMPA channels (Fig. 1B) or applied to the bath when ionotropic glutamate receptors were blocked (Fig. 1C) (-271.4 ± 44.1 pA for puff application, n = 12; -103.8 ± 31.6 pA for bath application, n = 8). These experiments suggest that, although glutamate activates receptors on interneuron somata, ATP may activate receptors accessible only when agonist was applied more globally and rapidly to the slice. Alternatively, or in addition, ATPS may activate receptors on nearby astrocytes to cause the release of ATP or other substances (Newman, 2003a) that then act on the interneurons to evoke the slow inward current (Fig. 1C). We address these possibilities below.
In current-clamp recordings, ATPS caused a significant (+10.7 ± 0.4 mV) depolarization of interneurons from resting levels (-65.2 ± 1.5 mV; n = 10) (Fig. 1D). In 8 of 10 neurons, the depolarization was sufficient to cause interneurons to fire action potentials (Fig. 1D). The size of the ATPS-evoked depolarization was of similar magnitude when action potentials were blocked (+10.4 ± 0.6 mV in the presence of TTX; n = 7; p > 0.05) (Fig. 1E). In the other two (of 10) cases, where the depolarization was small (<5 mV), ATPS increased the excitability of interneurons such that the interneurons fired more action potentials at lower levels of depolarizing current (Fig. 1F). We also performed extracellular cell-attached recordings from interneurons at the more physiological temperature of 34°C. All neurons that responded to glutamate with an increase in action potential firing (14.7 ± 3.8 Hz) responded likewise to ATPS (62.9 ± 9.9 Hz; n = 9). Collectively, these experiments indicate that although the slow response to ATPS is modest under voltage-clamp conditions (Fig. 1B), it is of sufficient size to excite interneurons.
ATP excites calbindin- and calretinin-positive stratum radiatum interneurons
All interneurons in s.r.-s.l-m. responded to ATP with responses similar to those described above (266 interneurons from 180 mice over a 1 year period) (Fig. 1). We next sought to further classify these interneurons within the framework proposed on the basis of morphological, biochemical, and electrophysiological properties (Parra et al., 1998). We started by performing an initial survey of calcium-binding protein expression (parvalbumin, calretinin, or calbindin) in 30 µm sections of juvenile mouse hippocampus (Fig. 2A). Consistent with recent mouse studies (Matyas et al., 2004), interneurons within s.r.-s.l-m. were predominantly calbindin and calretinin immunoreactive, whereas parvalbumin immunoreactivity was limited to stratum oriens (s.o.) and s.r. areas close to the pyramidal cell layer (Fig. 2A). We consider it unlikely that the electrophysiological responses described in this study (Fig. 1) are from parvalbumin-expressing interneurons, because we recorded only from s.r. interneurons distal to the pyramidal cell layer and near the s.l-m. border. We next tested the hypothesis that the neurons reported in this study are positive for calbindin and calretinin and attempted to relate the size of the ATP-evoked currents with electrophysiological and morphological properties of interneurons. We recorded peak and residual ATP-evoked currents from 44 interneurons (Figs. 1, 2C) and compared these to the location of interneuron somata (Fig. 2B), dendritic orientations (Fig. 2B), resting membrane potentials (supplemental material, available at www.jneurosci.org), firing patterns (Fig. 2C), action potential half widths (Fig. 2C), and expression of calbindin (Fig. 2C) (supplemental material, available at www.jneurosci.org) or calretinin (supplemental material, available at www.jneurosci.org). For our most complete data set, 68% of interneurons with ATP-evoked currents were positive for calbindin, whereas 32% were negative (supplemental material, available at www.jneurosci.org). Moreover, in another set of experiments, we attempted to relate ATP-evoked currents and calretinin immunoreactivity. We found that 41% of interneurons with ATP-evoked currents were positive for calretinin, whereas 59% were negative (supplemental material, available at www.jneurosci.org). Thus, of the interneurons that are depolarized by ATP, 60-70% are calbindin positive and 30-40% are calretinin positive. Within these classes, defined by expression of the calcium-binding proteins calbindin and calretinin, ATP-evoked currents occurred in all neurons regardless of soma location, dendritic orientation, firing pattern, resting membrane potential, or action potential half width (supplemental material, available at www.jneurosci.org) (Fig. 2). The basic properties of interneurons reported in this study are consistent with past work (Parra et al., 1998; Matyas et al., 2004).
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Figure 2. Characterization of CA1 s.r.-s.l-m. interneurons. A, Immunofluorescence micrographs of neurons immunoreactive for parvalbumin, calbindin, or calretinin. Scale bar, 50 µm. B, Montage of confocal Z-stacks flattened top resent an overview of a biocytin-filled interneuron reconstructed on the right with predominantly horizontal dendrites (black) and axonal arborization confined to s.r.-s.l-m. (gray) (see supplemental material, available at www.jneurosci.org, cell 6, for additional details of this interneuron). Scale bar, 50 µm. C, Properties of the interneuron presented in B. i, Immunofluorescence image of biocytin (green) and calbindin (red) immunoreactivity, with overlay in yellow. Note that several calbindin-positive interneurons are present in this image. Scale bar, 25 µm. ii, Current in response to 100 µM ATPS. iii, Action potential generated by a 5 msec currentpulse of + 150 pA. iv, Irregular action potential discharge pattern generated during depolarizing current injection. s.p., Stratum pyramidale.
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The slow ATP-evoked response increases synaptic inhibition
If interneurons fire more readily in response to ATP (Fig. 1F), then does this have an impact on their postsynaptic targets within the hippocampal circuit? To address this, we determined whether ATPS could increase GABAergic synaptic events onto s.r.-s.l-m. interneurons and CA1 pyramidal neurons. Interneurons were voltage clamped with a KCl-based intracellular solution (ECl = -1.3 mV) to allow the detection of inward GABAA receptor-mediated IPSCs at -60 mV (Fig. 3A). In 21 of 26 neurons, application of ATPS (100 µM) evoked an inward current (-28.9 ± 4.2 pA) and an increase in both the frequency (2.7 ± 0.5 to 9.6 ± 2.0 Hz) and amplitude of spontaneous IPSCs (sIPSCs; in all cases, the amplitude distributions shifted to larger peaks) (Fig. 3B). The sIPSCs evoked by ATPS were blocked by bicuculline and TTX (data not shown). These experiments suggest that when s.r.-s.l-m. interneurons are excited by ATPS, they fire action potentials and release GABA onto their postsynaptic targets, thus increasing inhibition within the network. To further explore this possibility, we recorded from connected pairs of s.r.-s.l-m. interneurons (Fig. 3C). The probability of a single evoked action potential causing a uIPSC (-75.5 ± 11.0 pA) in the postsynaptic interneuron was 0.24 ± 0.04 (n = 14 pairs; 100 trials per pair). ATPS-evoked spontaneous action potential firing in eight of nine presynaptic interneurons and 28% (probability, 0.28 ± 0.09; n = 8) of these triggered uIPSCs in the postsynaptic interneurons (Fig. 3C) (-75.5 ± 11.4 pA).
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Figure 3. ATP increases synaptic inhibition onto stratum radiatum interneurons and CA1 pyramidal neurons. A, ATPS caused an inward current as well an increase in the frequency (freq.) and amplitude of spontaneous IPSCs in 21 of 26 interneurons (traces show 3 superimposed 3 sec sweeps). B, Left graph, Average data for changes in sIPSC frequency in the absence of TTX. Right graph, Cumulative probability (Prob.) plots of IPSC amplitude before and during ATPS. C, The diagram illustrates the recording set up, with one s.r. interneuron as the presynaptic cell held in current clamp and another as the postsynaptic cell held in voltage clamp. Top panel, Membrane potential recording from a presynaptic s.r. interneuron showing spontaneous action potentials during the application of ATPS. Bottom panel, Membrane current recording from a postsynaptic s.r. interneuron showing IPSCs. D, ATPS caused no inward current but increased in the frequency and amplitude of sIPSCs in CA1 pyramidal neurons (traces show 3 superimposed 3 sec sweeps). Postsynaptic uIPSCs showed a short latency (2.7 ± 0.1 msec; n = 14 pairs) and displayed 20-80% rise times of 1.7 ± 0.1 msec and decay time constants of 13.4 ± 0.7 msec, respectively (n = 14 pairs). E, Left graph, Average data for changes in sIPSC frequency in the absence of TTX. Right graph, Cumulative probability plots of IPSC amplitude before and during ATPS. F, The diagram illustrates the recording set up, with an s.r. interneuron as the presynaptic cell held in current clamp and a CA1 pyramidal neuron as the postsynaptic cell held in voltage clamp. Top panel, Membrane potential recording from a presynaptic s.r. interneuron showing spontaneous action potentials during the application of ATPS. Bottom panel, Membrane current recording from a postsynaptic CA1 pyramidal neuron showing IPSCs. Unitary IPSCs onto pyramidal neurons displayed 20-80% rise times of 1.0 ± 0.2 msec and decay time constants of 11.6 ± 1.1 msec (n = 8 pairs). s.r.-s.p. pairs had an average uIPSC latency of 4.1 ± 0.3 msec in control conditions and 4.2 ± 0.3 msec in ATPS (n = 5 pairs; Fig. 3g).
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We next tested whether the s.r. interneurons that are excited by ATPS synapse exclusively onto other interneurons or additionally onto output CA1 pyramidal cells. Bath-applied ATPS failed to evoke inward currents in CA1 pyramidal neurons (Khakh et al., 2003) (Fig. 3D) (n = 8) but dramatically increased the frequency and amplitude of sIPSCs in a TTX-sensitive manner (seven of eight neurons) (Fig. 3E). We recorded from connected pairs of SR interneurons and CA1 pyramidal neurons to determine whether the ATPS-evoked increase in sIPSC frequency onto pyramidal neurons (Fig. 3F) arises from s.r. or other interneurons within the slice. The probability of a spontaneous action potential causing a uIPSC in the postsynaptic pyramidal neuron was 0.57 ± 0.01, and in the presence of ATPS, this was 0.60 ± 0.02. However, uIPSCs triggered by presynaptic ATPS-evoked action potentials were larger than those evoked by action potentials as a result of depolarizing current injection into the presynaptic neurons (-81.4 ± 7.0 pA vs -55.1 ± 4.7 pA; p < 0.05); the GABAA synaptic conductance was increased from 0.9 ± 0.1 to 1.4 ± 0.1 nS (n = 8). Thus, ATPS increased inhibition onto output pyramidal neurons in two ways, by increasing the frequency of IPSCs and by triggering IPSCs of larger synaptic conductance. The increased synaptic conductance may be caused by presynaptic effects of ATP such as facilitation or to postsynaptic effects, which may have increased GABAA receptor-mediated responses in pyramidal neurons. We hope to discriminate between these two possibilities in future experimental work. Together, our data from interneuron-interneuron and interneuron-pyramidal neuron pairs suggest that although the effect of ATP on single interneurons is excitatory (Fig. 1), its effect at the level of the network is dominated by heightened synaptic inhibition (Fig. 3).
A role for P2Y1 receptors
The ATPS-evoked inward current in interneurons developed slowly with a 20-80% rise time of 25.7 ± 5.2 sec (Fig. 1C). Moreover, membrane conductance was unaltered (1.6 ± 0.2 nS in control to 1.6 ± 0.2 nS in ATPS; n = 23) during the slow inward currents, whereas P2X channels increase neuronal membrane conductance (North, 2002). The lack of effect on membrane conductance combined with strong membrane potential depolarization (Fig. 1) recalls similar data with muscarinic agonists and suggested the activation of metabotropic receptors (Shen and North, 1992; McQuiston and Madison, 1999). To determine the nature of the receptors responsible for the responses described in Figures 1, 2, 3, 4, 5, 6, we exploited the known pharmacological properties of ATP receptors. We used two measures of receptor activation, the amplitude of the agonist-evoked inward current in single neurons and percentage of increases in the frequency of sIPSCs onto single SR interneurons. We began by examining the relative effects of agonists known to activate distinct types of ATP receptors. Each putative agonist was applied for 3 min, washed out for 7 min, and then 100 µM ATPS was applied to verify that the interneuron was responsive to this control agonist. Inward currents similar to those evoked by ATPS were observed for ATP (1 mM; pH 7.4), 2MeSATP (100 µM), 2MeSADP (100 µM), and ADPS (100 µM). In contrast, UTP (1 mM; pH 7.4),  meATP (100 µM), and adenosine (1 mM in the absence of DPCPX) were without effect (Fig. 4A,B). Overall, this agonist profile argues against P1 and P2X receptors and implies the involvement of P2Y1, P2Y11, or P2Y12 receptors (von Kugelgen and Wetter, 2000). Consistent with this, the responses were blocked by the P2 receptor antagonist PPADS (30 µM) and the P2Y1 antagonist 2'-deoxy-N 6-methyladenosine 3',5'-bisphosphate (MRS2179; 10 µM) (Moro et al., 1998). A similar pharmacological profile was found when we measured the increase in sIPSCs arriving onto interneurons, which probably provides a better measure of increased synaptic inhibition across the network (Fig. 4C,D).
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Figure 4. Pharmacological properties of ATP responses in SR interneurons. A, Exemplar traces showing whole-cell voltage-clamp currents at -60 mV first to a test nucleotide (ATP) and then to ATPS (100 µM). B, Average data for each test agonist: ATPS (100 µM; n = 22), ATP (1 mM; pH 7.4; n = 8), 2MeSADP (100 µM; n = 8), ADPS (100 µM; n = 16), meATP (100 µM; n = 10), UTP (100 µM; n = 11), and adenosine (100 µM; n = 7). Data are also shown for ATPS-evoked currents in slices incubated in MRS2179 (30 µM; n = 19) and PPADS (10 µM; n = 6). C, Spontaneous IPSCs were recorded in voltage-clamp mode at -60 mV (KCl intracellular solution) first to a test nucleotide agonist and then to test ATPS. An example experiment is shown in C where the frequency of sIPSCs is normalized to baseline. D shows the average peak sIPSC frequency increase for each agonist tested: ATPS (100 µM; n = 21), ATP (1 mM; pH 7.4; n = 5), 2MeSADP (100 µM; n = 7), ADPS (100 µM; n = 16), meATP (100 µM; n = 8), UTP(100 µM;n = 11), and adenosine (100 µM; n =6). Data are also shown for ATP S-induced responses in slices incubated in MRS2179 (30 µM; n = 9) and PPADS (10 µM; n = 11). The asterisk indicates significant responses. E, Confocal images of P2Y1 expressing interneurons in CA1. Scale bar, 50 µm. Note the highly fluorescent distinct somata are confined to interneurons in s.r., s.l-m., and s.o. The more diffuse staining in the pyramidal cell layer is probably not specific and has been observed by others (Matyas et al., 2004).
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Figure 5. Electrophysiological properties of P2Y1 receptor-mediated inward currents. A, The trace shows average ADPS-evoked inward currents recorded from interneurons at -60 mV (n = 12). The points i, ii, and iii indicate the time when current-voltage relationships presented in B-E were determined. Current-voltage (I-V) relationships were determined using voltage ramps from -100 to +40mV (9.3 mV/sec) before (i), at the peak of the ATP S current (ii), and after washout of ATPS(iii) as indicated in A. The resulting I-V relationships for a single experiment are shown in B. Subtraction of I-V relationships indicates the average I-V relationships of the peak and residual currents (n = 10; C-E). Inset in D is an expanded I-V region from -100 to -80 mV to show reversal potentials for each individual neuron. F, Average ADPS-evoked currents recorded from interneurons under control conditions (F) and then in the presence of group I mGluR antagonists MPEP and LY367385 (100 µM; G). The points i, ii, and iii in G indicate where current-voltage relationships presented inI and J were determined. H, Normalized traces shown inF and G. I, Current-voltage relationships from ramps for an ADPS-evoked current in the presence of mGluR blockers. J, Subtraction of I-V relationships at i and ii indicates the average I-V relationship of the peak current, with reversal of the current close to 0 mV (n = 8).
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Figure 6. Astrocyte Ca2+ dynamics. A, Fluorescence micrographs of an acute hippocampal slice loaded with fluo-4-AM (see Materials and Methods) before (i), during (ii), and after (iii) ADPS. iv indicates the location of the astrocytes that were monitored (numbered 1-5) and hippocampal cell layers as well as the location of the recording pipette (the patched interneuron was below the confocal section). Scale bar, 50 µm. B, Traces showing changes in intracellular Ca2+ for five representative astrocytes indicated in the panels in A. C, Average fluorescence trace from nine astrocytes and the corresponding interneuron current in this experiment. D, Traces showing spontaneous Ca2+ oscillations before and after ADP S for two astrocytes on an expanded time scale. The asterisk indicates a transient peak in intracellular Ca2+ concentration. E, Number of oscillations in the 200 sec before and 200 sec after ADPS for 33 astrocytes. F, Coefficient of variance was also measured before and after ADPS as a measure of astrocyte activity. In F, astrocytes with increased oscillations are shown in black, and those with decreased oscillations are shown in gray.
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We detected P2Y1 immunoreactivity in numerous CA1 interneurons in s.o., s.r., and s.l-m. (Fig. 4E), which was similar to the pattern observed for parvalbumin, calbindin, and calretinin expressing neurons (Fig. 2A). No P2Y1 staining was observed in CA1 pyramidal neurons, which is consistent with the lack of ATP-evoked currents in these cells (Khakh et al., 2003) (Fig. 3D). Higher magnification views clearly showed that the somata of interneurons in s.r. and s.l-m. are immunopositive for P2Y1 (Fig. 4E). Interestingly, within the s.r., there were numerous finer structures that were also immunopositive for P2Y1 receptors (Fig. 4E, arrows); these may represent the processes of interneurons and/or astrocytes. Overall, these pharmacological and antibody labeling experiments provide evidence for functional P2Y1 receptors on interneurons (Moore et al., 2000; Moran-Jimenez and Matute, 2000).
The peak and residual ATP-evoked inward currents are carried by two types of conductance
We used the more specific agonist ADPS to gain insight into the ionic basis of the P2Y1 receptor-mediated inward current (Fig. 4B) and either applied slow voltage ramps from -100 to +40 mV (over 15 sec) or 10 mV voltage steps over the same voltage range before ADPS (Fig. 5Ai) at the peak of the inward current (Fig. 5Aii) and after ADPS washout for 10 min at the residual phase of the current (Fig. 5Aiii,Biii). All data shown in Figure 5 are from voltage ramps, but Table 1 includes average data for ramps and steps. The current-voltage relationship at the peak of the ADPS-evoked inward current (ii minus i) was complex, with no apparent reversal potential over the range -100 to +40 mV (Fig. 5C), suggesting that the peak response may consist of at least two underlying currents with differing reversal potentials but overlapping activation ranges (Shen and North, 1992; McQuiston and Madison, 1999). We calculated the current-voltage relationship of the slow plateau current (Fig. 5D, iii minus i) and found the reversal potential to be -89.6 ± 1.5 mV (n = 10) close to the Nernst potential for K+ ions at -99.8 mV. This reversal potential is hard to see from the average graph (Fig. 5D), because the current was small and the reversal potentials varied between cells. To ameliorate this, we plotted the ramp current-voltage relationships for all neurons on an expanded scale (Fig. 5D, inset); it is clear that all of the currents do indeed reverse polarity, and the average data are presented in Table 1. This suggests that the ATPS-evoked residual current was attributable to suppression of a K+ conductance (McQuiston and Madison, 1999). We determined the current-voltage relationship of the peak current (Fig. 5E, ii minus iii). The most salient features were as follows: (1) an inward current at -60 mV, (2) little change in slope conductance in the range of -100 to -20 mV, (3) strong outward rectification, and (4) a reversal potential (-14.5 ± 3.4 mV), suggestive of a nonselective cationic current. Together, these electrophysiological properties argue against P2X channels (North, 2002) and instead imply activation of a nonselective cationic current (average current-voltage relationships for 10 cells are presented in Fig. 5C-E). Consistent with this, addition of the broad-spectrum cation channel blockers Cd2+ (30 µM; n = 8) and La3+ (10 µM; n = 8) reduced the amplitude of the inward current by 63.7 ± 4.4% (n = 8) and 72.7 ± 5.4% (n = 8), respectively. Interestingly, intracellular dialysis of interneurons with GDPS (100 µM; n = 6) through the patch pipette to block G-protein-mediated signaling did not reduce the ATPS-evoked inward current (-30.3 ± 3.2 pA vs -36.0 ± 6.7 pA). This suggests that G-proteins are not involved, and P2Y1 receptors may directly couple to ion channels, as predicted from detailed heterologous expression studies (O'Grady et al., 1996; von Kugelgen and Wetter, 2000; Lee et al., 2003). Overall, the experiments presented in Figure 5 and Table 1 suggest that ADPS activation of P2Y1 receptors depolarizes interneurons by activating a nonselective cationic conductance and suppressing a K+ conductance.
The residual component of the ADPS-evoked current was mediated by glutamate and accompanied by persistent increases in astrocyte Ca2+ oscillations
We considered the possibility that a component of the ATP-evoked responses recorded from interneurons may be secondary to astrocyte activation and the release of glutamate (Parpura and Haydon, 2000; Newman, 2003b; Liu et al., 2004). The responses described in this study are not caused by kainate-AMPA-NMDA channels because they were not affected by CNQX and AP-5 (10 µM). We explored the possibility that metabotropic glutamate receptors (mGluRs) may contribute to part of the ATPS-evoked inward current (Figs. 1B, 5A), because group I mGluRs are expressed in hippocampal interneurons (van Hooft et al., 2000) and astrocytes are known to release glutamate in response to elevated Ca2+ levels (Pasti et al., 2001). Consistent with previous work (Mannaioni et al., 2001), bath application of the group I mGluR agonist DHPG (10 µM) activated inward currents in interneurons (-46.6 ± 7.9 pA; n = 7) that could be blocked with mGluR antagonists MPEP and (S)-(+)--amino-4-carboxy-2-methylbenzeneacetic acid (LY367385) (100 µM; antagonists of mGluR5 and mGluR1a receptors, respectively). This current, like the residual ADPS-evoked current (Fig. 5D), is caused by suppression of a K+ conductance (Krause et al., 2002). The mGluR blockers also reduced the peak ADPS-evoked current by 20% in all cases, although this did not reach significance (n = 12) (Fig. 5F,G). However, the ADPS-evoked residual current was abolished when metabotropic glutamate receptors were blocked (Fig. 5G,H). This is further evidenced by voltage-ramp experiments (Fig. 5I). With mGluR blocked the peak current-voltage relationship shows a reversal potential (-3.2 ± 1.0 mV; n = 8) indicative of a nonselective cationic current and a shape almost identical to that calculated for the peak current by subtraction (Fig. 5, compare E and J). In short, when mGluRs were blocked, the ADPS-evoked inward current displayed no residual component, and there was no evidence for suppression of a K+ conductance by ADPS. We suggest the residual current (Figs. 1B, 5A), which is dominated by suppression of a K+ conductance (Fig. 5D) and persists even when the agonist has been washed from the slice (Figs. 1B, 5A), is primarily attributable to ADPS-evoked glutamate release acting on group I metabotropic glutamate receptors (Krause et al., 2002). Because interneurons have dendritic arbors, we cannot rule out the possibility that the reported reversal potential estimates may be contaminated to some extent by unavoidably inadequate voltage clamp of distal processes. The data should be interpreted with these considerations in mind.
Astrocytes are potential sources of endogenous glutamate release in response to P2Y receptor activation. But are astrocytes actually excited by P2Y receptor agonists in our experiments in brain slices? To address this, we applied ATPS or ADPS (100 µM) (Fig. 6B), monitored interneuron transmembrane current, and imaged Ca2+ levels in astrocytes in the s.r.-s.l-m. (Fig. 6 A-C) (movie in supplemental material, available at www.jneurosci.org). Some astrocytes displayed spontaneous Ca2+ oscillations, whereas others fluoresced only in ADPS, confirming that functional P2Y receptors are expressed in astrocytes (Fam et al., 2000; Koizumi et al., 2003). On average, the ADPS-evoked increase in astrocyte Ca2+ levels displayed rise times of 11.3 ± 1.8 sec, whereas the peak current recorded concomitantly from interneurons was slower (61.5 ± 3.7 sec) (Fig. 6C). Thus, at the peak of the astrocyte Ca2+ increase, the inward current had developed by only 31.2 ± 4.7%, whereas at the time of the peak current, the astrocyte Ca2+ level had already decayed by 54.1 ± 6.5% (n = 33). Moreover, there was no correlation between the magnitude of the peak ADPS-evoked current and the peak  F/F in astrocytes (r = 0.4), implying that the two responses were independent. How then do P2Y1 receptor agonists cause release of glutamate that mediates the residual current (Fig. 5H) recorded from interneurons? Interestingly, we observed persistent increases in the number of Ca2+ oscillations within astrocytes when the P2Y1 receptor agonists had been washed from the bath. This is evident in astrocytes 1, 3, 4, and 5 in Figure 6B, and two other examples are shown on an expanded scale in Figure 6D before and 5 min after ADPS. Quantification revealed that the vast majority of astrocytes (88%) within the s.r.-s.l-m. showed increased Ca2+ oscillations (Fig. 6E), which was reflected as a significant increase in the coefficient of variation (SD divided by mean) in astrocyte Ca2+ levels (Fig. 6F). The oscillatory responses in astrocytes are not likely to be a secondary consequence of neuron activation, because the experiments were performed in the presence of TTX. We propose that the residual ATPS-evoked current recorded from interneurons that persists even when ATPS has been removed is because of persistent increases in Ca2+ oscillations within astrocytes that release glutamate (Parpura and Haydon, 2000; Newman, 2003b) to activate metabotropic glutamate receptors on interneurons.
Evidence for endogenous activation of P2Y receptors on astrocytes
Astrocytes can be activated by numerous stimuli as they functionally express receptors for the transmitters glutamate, acetylcholine, noradrenaline, serotonin, and ATP (Haydon, 2001). In vitro electrical stimulation of Schaffer collateral (SC) afferents leads to excitation of astrocytes in the CA1 region (Porter and McCarthy, 1996). This effect has been partly attributed to glutamate release, which acts on metabotropic glutamate receptors of neighboring astrocytes. SC terminals have also been shown to release ATP (Wieraszko et al., 1989; Pankratov et al., 1998). We thus explored the possibility that ATP may activate nearby astrocytes. We used similar conditions to those already reported for SC stimulation (Porter and McCarthy, 1996) but also stimulated the perforant path (PP) axons from entorhinal cortex and recorded calcium signals from CA1 astrocytes in the same s.r.-s.l-m. area (Fig. 7A). Astrocytic calcium responses were evoked by trains of stimuli (300-800 µA; 30 Hz; 3 sec) similar to those used by researchers to induce calcium responses and long-term synaptic potentiation in hippocampal pyramidal neurons (400-1000 µA; 50 Hz; 2 sec) (Regehr et al., 1989; Regehr and Tank, 1990) and calcium responses in astrocytes (200-400 µA; 50 Hz; 2 sec) (Porter and McCarthy, 1996). Stimulating electrodes were placed between 300 and 750 µm from the confocal imaging site in all eight slices examined. In five slices, we first stimulated the PP and then repositioned the electrode to stimulate the SC (Fig. 7A) (the order was reversed for three slices). As shown in the fluorescence micrographs (Fig. 7B) and the overlaid astrocyte Ca2+ transients (Fig. 7C), only a proportion of the cells activated by SC stimulation were also activated by PP stimulation (six and four astrocytes, respectively, for the exemplar experiment in Fig. 7C). This was true of all eight slices investigated (73.8 ± 12.5% of SC-stimulated astrocytes responded to PP stimulation). These differences suggest that excitation of astrocytes may be stronger for the SC as opposed to the PP pathway (Fig. 7C,D). We next determined whether local stimulation within s.r. could evoke similar responses in astrocytes. A glass monopolar electrode was positioned in the center of the confocal field of view, and astrocyte calcium was monitored while the length of the stimulation train was gradually increased from 1 to 80 stimuli (10-40 µA; 30 Hz) (Fig. 8A,B). This experiment was repeated in six slices, and the results of two experiments are shown in Figure 8B. Astrocytes closest to the electrode were activated by lower numbers of stimuli than those further from the point of stimulation, for which activation was also delayed (Fig. 8B) (also see below). All loaded astrocytes could be activated with 80 stimuli at 30 Hz. We chose to use 100 stimuli at 30 Hz in subsequent pharmacological experiments to ensure reproducible astrocyte activation.
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Figure 7. Astrocyte activation by stimulation (stim) of the Schaffer collaterals and the peforant path. A, Schematic diagram of the hippocampus showing glutamatergic SC and PP axons. A concentric wire electrode was placed in stratum radiatum to stimulate SC or PP axons as indicated, whereas Ca2+ transients were recorded in astrocytes. B, Fluorescence micrographs of fluo-4-loaded astrocytes in s.r. without stimuli, with SC stimuli, and with PP stimuli. Numbers in the left panel indicate astrocytes monitored to obtain recordings shown below in C. D, Bar graph indicating number of astrocytes that responded to SC and PP stimulation from eight slices.
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Figure 8. Astrocytes are activated by vesicular release during local electrical stimulation. A, Confocal images of fluo-4-loaded astrocytes in hippocampal slices. A glass electrode is placed in the center of the field of view (stim), and trains of stimuli (number indicated on image) are delivered to the slice at 30 Hz. The concentric circles are 50 µm apart. The final panel identifies astrocytes monitored and the number of micrometers they are from the electrode. Astrocytes appear to be confined to the bottom left of the image as the confocal section is taken across an uneven hippocampal slice where fluo-4 loading is often confined to superficial astrocytes. B, Representative recordings of astrocyte Ca2+ transients from two slices with increasing numbers of electrical stimuli. The calcium transients in astrocytes of slice 2 are from the experiment presented in A. C, Ten superimposed traces showing the change in intracellular Ca2+ for astrocytes under control conditions (i) and in the presence of 1 µM TTX (ii), 10 µM cadmium (iii), and 20 µg/ml botulinum toxin E (iv). D, Average traces for treatments indicated in C (TTX: n = 36, 9 slices; cadmium: n = 46, 6 slices; botulinum toxin E: n = 35, 5 slices).
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How does local electrical stimulation within s.r. cause astrocyte Ca2+ responses? Within a 200 µm square field of view, 95% of the astrocyte responses (56 of 59 astrocytes; nine slices) were blocked by TTX (1 µM). Moreover, astrocyte responses were abolished (46 of 46 cells; six slices) when voltage-gated Ca2+ channels were blocked with cadmium (10 µM) (Fig. 8C,D). Finally, very few astrocytes (Fig. 8C) responded during electrical stimulation in slices that had been preincubated in botulinum neurotoxin E to cleave synaptosomal-associated protein of 25 kDa (SNAP-25) (BoToxE, 20 µg/ml, 4 hr), constituting a blockade of the response by >93% (Fig. 8C,D). Because (1) the majority of CA1 s.r. astrocytes do not express TTX-sensitive Na+ channels (Sontheimer and Waxman, 1993), (2) there is no evidence to indicate that s.r. astrocytes express voltage-gated Ca2+ channels, and (3) astrocytes mostly express SNAP-23 rather than SNAP-25 (Sadoul et al., 1997; Wilhelm et al., 2004), these data imply that the trigger for astrocyte responses during electrical stimulation is exocytic release of transmitter from SC and/or PP nerve terminals. These experiments do not address the possibility that astrocytes may also release transmitter by exocytosis but rather support the hypothesis that the initiating stimulus for the responses described in Figure 8 is release from nerve terminals.
If astrocytes are activated through release from nerve terminals, then which transmitter is responsible? In a series of experiments, we applied trains of stimuli in the center of the field of view (313 by 208 µm) and imaged Ca2+ changes in s.r. astrocytes (Fig. 9) (movies in supplemental material, available at www.jneurosci.org). Under these conditions, astrocyte Ca2+ increases were measured in surrounding astrocytes (Fig. 8A) up to 200 µm from the stimulating electrode. As alluded to earlier, the latency between the time of the stimulus and the peak of the Ca2+ increase in astrocytes were smaller for astrocytes close to the electrode (Fig. 9Ci) and longer for astrocytes up to 200 µm away (Fig. 9Cii), indicative of a signal that spreads out from the site of stimulation (all data points are shown in Fig. 10A, control and averages are shown in B). This is reminiscent of the spread in ATP levels from a point source in retinal slices and cultured astrocytes (Guthrie et al., 1999; Newman, 2001b; Koizumi et al., 2003). We next tested the hypothesis that the spread of astrocyte activation evoked by electrical stimulation in the center of the field of view was attributable to ATP acting on astrocyte P2Y receptors and/or glutamate acting on metabotropic glutamate receptors. The peak F/F for astrocyte Ca2+ increases in the antagonists or under control conditions was similar (Fig. 9E) but significantly briefer in the presence of PPADS (10 µM) and MRS2179 (30 µM) than in control (Fig. 9D,E). Furthermore, whereas in control slices, Ca2+ changes could be measured in astrocytes 200 µm from the stimulating electrode (Fig. 10A,B) (supplemental material, available at www.jneurosci.org), in the presence of PPADS and MRS2179, astrocytes only responded in a region half this size at 100 µm (Figs. 9Bi,ii, 10A,B) (supplemental material, available at www.jneurosci.org). Similar data were gathered from slices treated with extracellular apyrase (10 U/ml), a nucleotidase that is expected to hydrolyze any released ATP (Fig. 9D). The residual astrocyte responses that remain in the presence of P2Y1 receptor antagonists are likely caused by activation of group I metabotropic glutamate receptors (Porter and McCarthy, 1996). Consistent with this, treatment of slices with the group I mGluR antagonists MPEP and LY367358 (100 µM) significantly reduced the peak F/F (41 astrocytes, 7 slices) (Fig. 9D,E), and cotreatment with PPADS completely blocked evoked astrocyte activation in 95% of cells (3 of 55 astrocytes from 9 slices still responded). From these data, we suggest that the triggering stimulus to astrocytes is likely to be vesicular release of glutamate and ATP from presynaptic terminals in s.r. This is readily seen in the representative traces in Figure 9D, averaged data in Figure 9E, and was further quantified by integrating the area under the Ca2+ transients (Fig. 10D). In the most straightforward explanation, the astrocyte signal appears to be propagated by a mixture of glutamate acting on astrocytes within 100 µm of the stimulus and by ATP to more distant astrocytes (200 µm) (Fig. 10E).
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Figure 9. Astrocyte Ca2+ transients evoked by electrical stimulation (stim). A, B, Confocal images of fluo-4-AM-loaded hippocampal slices 30 sec before field stimulation and 10 and 30 sec after stimulation under control conditions (A) and in the presence of 30 µM PPADS (B). The position of the stimulating electrode is shown on a grid of 25 µm spacing. C, Example Ca2+ traces for two astrocytes (i, ii indicated in A, B) and the distances from the stimulating electrode under control conditions and in the presence of PPADS. D, Ten superimposed traces showing change in intracellular Ca2+ for astrocytes in control (i), in the presence of mGluR antagonists (ii), PPADS (iii), and both mGluR antagonists and PPADS (iv). Note that the transients are briefer in the presence of PPADS and completely blocked in the presence of both PPADS and the mGluR antagonists. Superimposed traces are also shown for slices treated with 20 U/ml apyrase (v) and for experiments conducted at 34°C (vi). Average astrocyte transients for the indicated treatments are shown in E.
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Figure 10. Properties of astrocyte Ca2+ transients evoked by electrical stimulation. Pooled astrocyte data for control (9 slices, 54 astrocytes), PPADS treated (9 slices, 29 astrocytes), MRS2179 treated (4 slices, 17 astrocytes), mGluR antagonist treated (7 slices, 41 astrocytes), and control experiments conducted at 34°C (6 slices, 34 astrocytes). A, Scatter plot of calcium peak response latency (time from beginning of spike train to peak of Ca2+ transient) against distance from stimulus region. B, Averaged data for 50 µm bins. C, Peak astrocyte F/F against distance from stimulus region. D, Integrated astrocyte responses evoked by electrical stimulation. E, Summary diagram of the results shown in A-E. In control slices, astrocytes within a radius of 200 µm show an increase in Ca2+ during electrical stimulation; however, when P2Y receptors were blocked, this radius was reduced to 100 µm, implying that endogenously released ATP activates P2Y receptors to allow astrocyte excitation to spread to large distance scales.
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Endogenous activation of P2Y receptors on astrocytes increases synaptic inhibition
If endogenous ATP activates astrocyte P2Y receptors over 200 µm distance scales (Figs. 9, 10), then does this affect inhibitory synaptic activity onto interneurons? To address this, we monitored the frequency of spontaneous IPSCs in 30 sec windows before and after electrical stimulation using conditions identical to those that caused excitation of astrocytes (Figs. 9, 10). We expressed the data as the ratio of the number of IPSCs before and after stimulation (the facilitation ratio) (Fig. 11 A, B) under control conditions and during pharmacological treatments to affect ATP signaling. The facilitation ratio was significantly reduced when P2Y receptors were blocked with PPADS at 23°C (Fig. 11 Bi) (1.06 ± 0.04 to 0.84 ± 0.04; n = 10; p < 0.05) and at the more physiological temperature of 34°C (Fig. 11 Bii) (1.11 ± 0.06 to 0.91 ± 0.05; n = 7; p < 0.05). We also used complimentary approaches to block endogenous ecto-nucleotidases with 6-N,N-diethyl---dibromomethylene-D-ATP (ARL67156 and therefore presumably increase extracellular ATP levels and used apyrase (20 U/ml) to cleave any released ATP. In experiments conducted at 34°C, ARL67156significantly increased the facilitation ratio by 25% (Fig. 11Biii) (0.95 ± 0.02 to 1.20 ± 0.05; n = 8; p < 0.05). In experiments at 34°C, apyrase decreased the facilitation ratio in 9 of 12 interneurons investigated (Fig. 11Biv). Although the pooled data from all 12 interneurons treated with apyrase did not reach statistical significance (p > 0.05), statistical significance was obtained (from 1.11 ± 0.01 to 1.02 ± 0.01; n = 9; p < 0.05) by removing the two experiments where the facilitation ratio increased. The variability in the effect of apyrase may be explained by the fact that it is a protein, for which penetration may vary between slices and also because apyrase is expected to generate ADP, a potent agonist of P2Y1 receptors (Zimmermann, 1999). Although all of our experiments were performed with adenosine receptors blocked, we also determined whether adenosine had any role to play in the responses described in Figure 11. We found no significant difference in the facilitation ratio after 15 min incubations in 10 µM DPCPX (1.08 ± 0.04 to 1.06 ± 0.04; p > 0.05; n = 8). We propose that 20% of the IPSCs impacting on interneurons after a period of electrical stimulation were caused by ATP release from astrocytes acting on interneuron P2Y1 receptors to increase their excitability (Figs. 1, 3).
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Figure 11. Electrical stimulation increases GABAergic sIPSCs onto interneurons. A, Interneurons were voltage clamped at -60mV with a KCl-based internal solution allowing visualization of inward GABAergic synaptic events. A train of 100 pulses (7-15 µA; indicated by the gap in the trac | |
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Abstract
Introduction
