Astrocytes regulate heterogeneity of presynaptic strengths in hippocampal networks

Edited by Mu-ming Poo, Institute of Neuroscience and Key Laboratory of Primate Neurobiology, State Key Laboratory of Neuroscience, Chinese Academy of Sciences Center for Excellence in Brain Science, Shanghai Institutes for Biological Sciences, Shanghai, China, and approved March 28, 2016 (received for review December 1, 2015)
April 26, 2016
113 (19) E2685-E2694

Significance

We addressed the basic mechanisms underlying synapse heterogeneity, and we identified a form of regulation that serves to increase the variations in the efficacy with which neurons communicate with each other through synapses. We demonstrate that this process requires astrocytes, which, previously, have been thought to play mostly a passive role in maintaining neuronal functions. The cellular mechanism that regulates synaptic efficacy requires astrocyte membrane depolarization, activation of astrocyte NMDA receptors, and astrocyte calcium signaling. The fundamental nature of the regulation is underscored by the preservation of the mechanism from acute brain slices down to dissociated cultures that lack the native topology of brain networks.

Abstract

Dendrites are neuronal structures specialized for receiving and processing information through their many synaptic inputs. How input strengths are modified across dendrites in ways that are crucial for synaptic integration and plasticity remains unclear. We examined in single hippocampal neurons the mechanism of heterosynaptic interactions and the heterogeneity of synaptic strengths of pyramidal cell inputs. Heterosynaptic presynaptic plasticity that counterbalances input strengths requires N-methyl-d-aspartate receptors (NMDARs) and astrocytes. Importantly, this mechanism is shared with the mechanism for maintaining highly heterogeneous basal presynaptic strengths, which requires astrocyte Ca2+ signaling involving NMDAR activation, astrocyte membrane depolarization, and L-type Ca2+ channels. Intracellular infusion of NMDARs or Ca2+-channel blockers into astrocytes, conditionally ablating the GluN1 NMDAR subunit, or optogenetically hyperpolarizing astrocytes with archaerhodopsin promotes homogenization of convergent presynaptic inputs. Our findings support the presence of an astrocyte-dependent cellular mechanism that enhances the heterogeneity of presynaptic strengths of convergent connections, which may help boost the computational power of dendrites.
An enduring challenge in neurobiology is to understand how neurons set the strengths of their numerous synapses to efficiently process and store different information while maintaining network homeostasis. Electrophysiology and imaging approaches have revealed that synapses display a high degree of functional heterogeneity, even for those sharing the same axon or dendrite (13). The observation that synaptic strengths are heterogeneous, in turn, suggests that synapses can operate independently from one another. Accordingly, many studies have demonstrated the input-specificity of Hebbian and also of homeostatic forms of synaptic plasticity, where synaptic changes are restricted to inputs whose activity is altered (46). Nevertheless, such a synapse-autonomous behavior could potentially compromise the global network homeostasis by biasing the overall activity toward excitation or depression, and to overcome this issue, it has been proposed that distinct inputs cooperate by coordinating their relative strengths through heterosynaptic interactions (79). In support of the idea that synapses behave as interdependent rather than isolated functional units, the restriction of synaptic strength changes to active inputs has been demonstrated to break down at times, with the induction of synaptic plasticity in the stimulated input accompanying either synaptic depression or potentiation of the nonstimulated inputs (1013). In a highly studied plasticity paradigm of long-term potentiation (LTP) at hippocampal Schaffer collateral–CA1 synapses, tetanic stimulation that induces LTP is often accompanied by presynaptic long-term depression (LTD) of nonstimulated Schaffer collateral–CA1 synapses (11, 13, 14). Heterosynaptic LTD might be advantageous for sharpening the difference between the strengths of active and inactive inputs received by the postsynaptic CA1 neuron, and by counterbalancing potentiation, heterosynaptic LTD might help promote the stability of the overall CA3–CA1 connection by placing limits on excitation.
Among the proposed mechanisms for mediating heterosynaptic interactions, the astrocyte network has recently emerged as a key regulator (15, 16). Astrocytes display fine processes that are proximal to synapses, and they can detect and integrate local synaptic activity (1721). Moreover, astrocytes are coupled to each other through gap junctions, and by forming a network, they are thought to be capable of modulating the efficacy of a population of synapses by coordinately releasing diffusible gliotransmitters such as glutamate, endocannabinoids, ATP or d-serine that target synaptic receptors (15, 16). In the hippocampus, astrocytes respond to Schaffer collateral stimulations (14, 2224), and they mediate tetanus-induced heterosynaptic presynaptic LTD at CA1 synapses through purinergic signaling (14, 2527). The involvement of astrocytes in heterosynaptic depression of nonstimulated inputs raises the intriguing possibility that astrocytes might actively play a role in balancing synaptic strengths between different inputs. Whether such astrocyte-dependent regulation is a basic mechanism that controls bidirectional heterosynaptic interactions in spontaneously active networks and if such a regulation is implemented at the level of single inputs are unclear.
Here, using electrophysiology and optical methods, we addressed the basic cellular principle governing the distinctness of individual synaptic inputs. Specifically, we examined whether and how two comparable inputs from a same neuron type show coordinated regulation of their synaptic strengths when they share the target neuron. In both hippocampal dissociated cultures and acute slice preparation, we find that presynaptic strengths of inputs received by a postsynaptic neuron are highly heterogeneous. The differences in presynaptic strengths are controlled and maintained by astrocyte Ca2+ signaling requiring NMDA receptors (NMDARs) and L-type voltage-gated Ca2+ channels (L-VGCCs). This mechanism is shared at least in part, by that of activity-dependent heterosynaptic interactions in which the presynaptic strength of the nonstimulated input is counterbalanced relative to the presynaptic strength of the stimulated input, and such a plasticity mechanism could further augment the variations in presynaptic strengths. Importantly, under basal conditions, the presynaptic strengths of convergent inputs become rapidly correlated on blocking NMDARs or L-VGCCs or by buffering Ca2+ in astrocytes, and such correlation is also observed when GRIN1 is conditionally deleted in astrocytes. Furthermore, hyperpolarizing astrocytes by light activation of archaerhodopsin (ArchT) promotes the correlation of presynaptic input strengths over the time scale of minutes. These findings reveal an important contribution of activity-dependent astrocyte signaling in diversifying the strengths of convergent presynaptic connections received by the postsynaptic neuron.

Results

Activity-Dependent Coordinated Modulation of Presynaptic Strengths in Hippocampal Networks.

To explore whether synaptic inputs operated in isolation or in coordination with one another and to determine how this could impact synapse heterogeneity, we examined the extent to which activity-dependent synaptic changes were restricted to stimulated inputs. To this end, we compared two independent monosynaptic inputs onto a single postsynaptic neuron. We reasoned that a fundamental mechanism should operate not only in a native brain circuit but it should be recapitulated in a simplified neuronal network formed in culture. We took advantage of low density dissociated hippocampal neurons cocultured with glia, in which unitary connections formed between identified neurons can be stimulated and monitored through multiple whole-cell patch-clamp recordings with relative ease. Triple patch-clamp experiments (Fig. 1A) were restricted to cases in which the two presynaptic neurons showed no direct synaptic coupling to each other. To induce synaptic changes at a given input, we tested a variety of stimulation paradigms that were used previously in dissociated cultures similar to ours (2831); we finally opted for a low-frequency stimulation protocol (1 Hz for 3 min; Materials and Methods) that efficiently induced long-lasting changes in excitatory postsynaptic current (EPSC) amplitude in ∼80% of the recordings [n = 17 of 21; no change in n = 4 of 21 (+0.99 ± 1.7%); Fig. 1 B and C]. Interestingly, the direction of plasticity was variable, and the stimulation could produce either potentiation (+25.9 ± 4.4%, n = 6 of 21 recordings) or depression (−29.9 ± 3.8%, n = 11 of 21) of the EPSC amplitude in the stimulated input (referred as “homosynapses”; Fig. 1 B and C). Furthermore, EPSC amplitude changes were inversely correlated with paired-pulse ratio (PPR) and CV−2 (Fig. 1 B and E and Fig. S1A), two parameters related to neurotransmitter release probability (pr) (28, 3234). When the same stimulation paradigm was given to a cell pair in which the presynaptic neuron expressed VGLUT-pHluorin, a reporter of synaptic vesicle exo-endocyosis, the readily releasable vesicle pool that was proportional to pr (1) also showed potentiation (two of six pairs), depression (one of six pairs), or no change (three of six pairs), which further confirmed the presynaptic origin of plasticity (Fig. S2). This observation was consistent with presynaptic expression of plasticity that had been previously reported for hippocampal synapses (28, 30, 33, 3537).
Fig. 1.
Presynaptic interactions between convergent inputs in dissociated hippocampal cultures. (A) (Left) Experimental scheme. One of the two convergent inputs is repeatedly stimulated (homosynapse; 180 action potentials at 1 Hz), whereas the other input is unstimulated (heterosynapse). (Right) DIC image of 12 DIV cultured hippocampal neurons during a triple recording. (Scale bar, 40 μm.) (B) Average homosynaptic EPSC traces to a paired stimuli (50-ms interstimulus interval) before and after the repeated stimulation. (C and D) Time course of homosynaptic (C; n = 21 cells) and heterosynaptic (D; n = 21 cells) EPSC amplitude changes induced by the repeated stimulation sorted into those showing potentiation [dark circles: n = 6 (C); n = 6 (D)], no change [medium circles: n = 4 (C); n = 8 (D)], or depression [light circles: n = 11 (C); n = 7 (D)]. Individual recordings were sorted into three groups according to the comparison of the baseline with the poststimulation period using the Wilcoxon test. Data are expressed as mean ± SEM. (E and F) Plots of the changes in PPR vs. mean EPSC amplitude at homosynapses (E: r2 = 0.33, **P = 0.0061) and heterosynapses (F: r2 = 0.48, ***P = 0.0005). Linear regression (black lines) and 95% CI (gray shade) are indicated. (G) Comparison of the extent changes in homosynaptic vs. heterosynaptic EPSC amplitudes (r2 = 0.18, P = 0.0563). (H) Comparison of the extent changes in homosynaptic vs. heterosynaptic PPR (r2 = 0.04, P = 0.3940).
Fig. S1.
Presynaptic interactions between convergent inputs in dissociated hippocampal cultures. (A and B) Plots of the extent changes in CV−2 vs. EPSC amplitudes normalized to the baseline at homosynapses (A) and heterosynapses (B) (Kruskall–Wallis test followed by Dunn’s multiple comparison test, *P < 0.05, **P < 0.01).
Fig. S2.
Optical monitoring of presynaptic plasticity in cultured neurons. (A) Experimental scheme. A presynaptic neuron expressing VGLUT1-pH (green) and a postsynaptic partner filled with an AlexaFluor dye (red) are patch-clamped; 40-APs, 20-Hz stimulation of the presynaptic neuron (green bar) is applied to assess the readily releasable pool (RRP) size before and 20 min after the plasticity induction stimulation. (B) Alexa dye-filled neuron (red) contacted by VGLUT1-pH–positive puncta (green). Arrowheads indicate putative synapses. (Scale bars, 30 and 5 μm.) (C) An example of VGLUT1-pH fluorescence change triggered by a 40-APs, 20-Hz stimulation at indicated boutons (arrowheads in B). (D) Traces showing the time course of fluorescence intensity triggered by the stimulation (arrows) to mobilize the RRP given before and 20 min after the 1-Hz, 3-min stimulation. Gray traces correspond to different boutons from the same input. The dark trace represents the average. (E–G) Summary (Left) and scatter plot (Right) comparing the RRP size before and after the 1-Hz, 3-min stimulation for three different outcome examples: potentiation (E; n = 16 boutons, paired t test, P < 0.0001), depression (F; n = 18 boutons, paired t test, P = 0.0011), and no change (G; n = 9 boutons, paired t test, P = 0.2646).
Importantly, EPSC amplitude change in the stimulated input was often accompanied by a stable EPSC amplitude change in the unstimulated input (referred as “heterosynapses”), suggesting that the expression of synaptic plasticity was not confined to the stimulated input (Fig. 1D). Therefore, this stimulation paradigm allowed us to study the properties of heterosynaptic interactions involving changes in synaptic strength. As for homosynapses, heterosynaptic EPSC amplitude changes were highly heterogeneous (potentiation: +28.4 ± 4.0%, n = 6 of 21 recordings; depression: −22.4 ± 5.9%, n = 7 of 21; no change: 0.7 ± 5.2%, n = 8 of 21; Fig. 1D) and correlated with changes in the presynaptic parameters PPR and CV−2 (Fig. 1F and Fig. S1B). Surprisingly, the direction and the magnitude of homo- and heterosynaptic changes appeared nonetheless uncorrelated to each other (Fig. 1G), indicating that the active and inactive inputs interacted nonuniformly.
Given that the heterosynaptic change was induced by stimulating the homosynaptic input, we asked if there was a homosynaptic parameter that was related to the direction and the magnitude of the heterosynaptic change. We found that the extent changes in heterosynaptic EPSC amplitude or PPR were correlated to the initial PPR of the homosynaptic input but not to the homosynaptic PPR change (Figs. 1H and 2 A and B) nor to the initial homosynaptic amplitude (Fig. S3). The apparent independence of the heterosynaptic changes to the homosynaptic amplitude suggests that the heterosynaptic change does not depend on the number of activated synapses. Notably, the heterosynaptic PPR change showed a significant inverse relationship to the initial homosynaptic PPR (heterosynaptic PPR = −0.47 × initial homosynaptic PPR + 0.45; r2 = 0.23, P = 0.0292; Fig. 2B), in that stimulating a high pr input was more likely to decrease pr of nonstimulated convergent inputs and vice versa. Such an activity-dependent coordinate modulation could play a potential role in sharpening the differences in the efficacies of active and inactive inputs received by the target neuron.
Fig. 2.
Presynaptic interactions between convergent inputs depend on NMDARs, astrocytes, and L-VGCCs but not on postsynaptic Ca2+. (A) Comparison of the percent change in heterosynaptic mean EPSC amplitude vs. the initial homosynaptic PPR in the control condition (r2 = 0.20, P = 0.0418). (B–F) Comparison of the heterosynaptic PPR change vs. the initial homosynaptic PPR under indicated conditions: control untreated (B: r2 = 0.23, *P = 0.0292); BAPTA in the postsynaptic neuron (C: r2 = 0.75, *P = 0.0111); AP5 (D: r2 = 0.05, P = 0.5431); fluoroacetate (E: r2 = 0.004, P = 0.8661); and nifedipine (F: r2 = 0.82, **P = 0.0019). Linear regression (black lines) and 95% CI (gray shade) are shown.
Fig. S3.
Lack of correlation between heterosynaptic changes and initial homosynaptic EPSC amplitude. (A) Comparison of the percent change in heterosynaptic mean EPSC amplitude vs. the initial homosynaptic EPSC amplitude in the control condition in dissociated cultures (r2 = 0.15, P = 0.0827). (B) Comparison of the heterosynaptic PPR change vs. the initial homosynaptic EPSC amplitude in the same set of experiments (r2 = 0.002, P = 0.8524).

Heterosynaptic Presynaptic Modulation Requires NMDARs and Astrocytes but Not Postsynaptic Ca2+.

To gain insights into the mechanism by which stimulating the homosynaptic input produced the PPR change at the nonstimulated input, we first tested the involvement of postsynaptic Ca2+, which was implicated in some forms of heterosynaptic plasticity (12, 38). However, after infusing the postsynaptic neuron with 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid (BAPTA; 30 mM) through the patch pipette, stimulating the homosynaptic input still produced changes in the heterosynaptic EPSC amplitude and PPR to extents comparable to the control condition (range of PPR change for control: −0.48 to 1.03, CV = 19.8, n = 21; for postsynaptic BAPTA: −0.72 to 0.32, CV = 21.8, n = 7; P = 0.7761, F test; Fig. 2C and Fig. S4), and the heterosynaptic PPR change remained inversely related to the basal homosynaptic PPR (heterosynaptic PPR = −1.43 × initial homosynaptic PPR + 1.04; r2 = 0.75, P = 0.0111; Fig. 2 B and C). Therefore, heterosynaptic presynaptic plasticity in our culture system appears not to require postsynaptic Ca2+.
Fig. S4.
Activity-dependent heterosynaptic interactions in cultures depend on NMDARs, astrocytes, and L-VGCCs but not on postsynaptic Ca2+. (A) Plot of the change in PPR vs. % change in EPSC amplitude at homosynapses when the postsynaptic neuron is filled with BAPTA (n = 10 cells, r2 = 0.71, P = 0.0022) or in presence of AP5 (n = 10 cells, r2 = 0.68, P = 0.0032), fluoroacetate (n = 10 cells, r2 = 0.43, P = 0.0389), or nifedipine (n = 8 cells, r2 = 0.92, P = 0.0002). (B) Plot of the change in PPR vs. % change in EPSC amplitude at homosynapses when the postsynaptic neuron is filled with BAPTA (n = 7 cells, r2 = 0.68, P = 0.0221) or in presence of AP5 (n = 10 cells, r2 = 0.25, P = 0.1389), fluoroacetate (n = 10 cells, r2 = 0.86, P = 0.0001), or nifedipine (n = 8 cells, r2 = 0.38, P = 0.1045). Linear regression (black lines) and 95% CI (gray shade) are indicated.
We next examined a role for NMDARs and astrocytes that had been previously shown to mediate heterosynaptic LTD at hippocampal CA3–CA1 synapses (14, 2527, 39). Despite the apparent lack of requirement for postsynaptic Ca2+, the heterosynaptic PPR change was fully blocked by the NMDAR antagonist d-2-amino-5-phosphonovaleric acid (AP5; 50 μM) (range of PPR change for control: −0.48 to 1.03, CV = 19.8, n = 21; for AP5: −0.05 to +0.21, CV = 1.7, n = 10; P = 0.0002, F test) and unrelated to initial homosynaptic PPR (heterosynaptic PPR = 0.03 × initial homosynaptic PPR + 0.01, r2 = 0.05, P = 0.5431; Fig. 2D and Fig. S4), whereas the basal PPR itself was not affected by the AP5 treatment (control: 0.92 ± 0.07, AP5: 1.20 ± 0.17; Fig. S5A); this suggested a possible involvement of NMDARs expressed in cells other than the postsynaptic neuron. Also, incubating cultures for 30 min with fluoroacetate (5 mM), an inhibitor of the Krebs cycle that preferentially compromised glial cells (3941), not only attenuated the heterosynaptic PPR change (range of PPR change for control: −0.48 to 1.03, CV = 19.8, n = 21; for fluoroacetate: −0.33 to 0.22, CV = 8.79, n = 10; P = 0.07, F test), but the PPR change was no longer related to the initial homosynaptic PPR (heterosynaptic PPR = −0.01 × initial homosynaptic PPR + 0.04; r2 = 0.004, P = 0.8661; Fig. 2E). Fluoroacetate did not significantly alter the mean basal PPR in our conditions (control: 0.92 ± 0.07, n = 21, fluoroacetate: 1.14 ± 0.21, n = 10; Fig. S5A). These results suggest that NMDARs and astrocytes mediate activity-dependent heterosynaptic interactions in cultured hippocampal networks.
Fig. S5.
Blocking NMDARs, L-VGCCs, or astrocyte activity does not affect the average basal PPR in hippocampal cultures and acute slices. (A) Summary of basal PPR in cultures in control, AP5, postsynaptic BAPTA, fluoroacetate, or nifedipine (n = 21, 10, 7, 10, or 8 cells, respectively; one-way ANOVA followed by Holm–Sidak’s multiple comparison test, P > 0.05). (B) Summary of basal PPR in acute slices in control, AP5, postsynaptic BAPTA, nifedipine, or nimodipine (n = 29, 11, 11, 17, or 11 cells, respectively; Kruskal–Wallis test followed by Dunn’s multiple comparison test). (C) Summary of basal PPR in acute slices when astrocytes are perfused intracellularly via a patch pipette containing control internal solution, MK-801, BAPTA, QX-314, or D890 (n = 12, 11, 12, 11, or 10 cells, respectively; one-way ANOVA followed by Holm–Sidak’s multiple comparison test, P > 0.05). (D) Plots of PPR before (Pre) and after (Post) incubation of acute slices with control (Left; n = 8, paired t test, P = 0.7811), AP5 (Center; n = 10, paired t test, P = 0.0893), or nimodipine (Right; n = 6, Wilcoxon test, P = 0.6875).
We also tested a possible role for L-VGCCs. The L-VGCC antagonist nifedipine (10 µM) did not prevent the bidirectional change in heterosynaptic PPR per se (range of PPR change for control: −0.48 to 1.03, CV = 19.8, n = 21; for nifedipine: −0.28 to 0.74, CV = 5.73, n = 8; P = 0.7398, F test; Fig. 2F and Fig. S4). Strikingly, however, nifedipine reversed the polarity of the inverse relationship between the heterosynaptic PPR change and the initial homosynaptic PPR into a positive one (heterosynaptic PPR = 0.67 × initial homosynaptic PPR – 0.85, r2 = 0.82, P = 0.0019; Fig. 2F). This observation suggested that in the presence of nifedipine stimulating an input with high pr was likely to potentiate nonstimulated inputs and vice versa. Thus, although L-VGCCs were not essential for homo- or heterosynaptic plasticity, blocking their activity had the apparent effect of equalizing presynaptic strengths between homosynapses and heterosynapses in active networks. L-VGCC activity could contribute to the cellular process by which inactive inputs adapt to the strength of active inputs.
Given that NMDARs and astrocytes mediate heterosynaptic plasticity in cultured networks similarly to tetanus-induced heterosynaptic LTD in brain slices (14, 27), we wondered if the apparent lack of requirement for postsynaptic Ca2+ and the dependence on L-VGCCs of the polarity of heterosynaptic plasticity that we observed in cultured networks could be extended to brain slices. We tested this possibility by examining heterosynaptic LTD at CA3–CA1 synapses in acute hippocampal slices (14). Two independent Schaffer collateral inputs were stimulated with extracellular electrodes (>200 μm apart) to evoke EPSCs in CA1 pyramidal neurons (Fig. 3 A and B). Applying three 100-Hz–1-s tetanus at 30-s intervals to one of the two inputs elicited homosynaptic LTP of EPSC amplitude (+62.9 ± 9.8%, n = 6; Fig. 3C), whereas the EPSC amplitude of the nonstimulated input was stably decreased (−16.6 ± 8.5%, n = 6; Fig. 3C). Unlike the homosynaptic LTP, the heterosynaptic LTD displayed a decrease in CV−2 and an increase in PPR (Fig. 3D), which supported for a decreased pr as the basis for the synaptic depression, in agreement with previous reports (14, 25, 26). Importantly, dialyzing the postsynaptic neuron with BAPTA fully blocked the homosynaptic LTP (−8.8 ± 7.7%; Fig. 3E) in accord with its requirement for postsynaptic NMDARs (42). In contrast, the heterosynaptic LTD still occurred (−17.4 ± 4.9%, n = 10), and CV−2 and PPR showed changes similarly to those observed in the absence of BAPTA (Fig. 3 E and F). Therefore, like heterosynaptic plasticity in cultured neurons, heterosynaptic presynaptic LTD in acute slices did not require postsynaptic Ca2+. Interestingly, when tetanic stimulation was applied in the presence of nifedipine or another L-VGCC inhibitor, nimodipine (10 μM), although homosynaptic LTP was elicited (nifedipine: +62.7 ± 29.3%, n = 5; nimodipine: +78.9 ± 28.7%, n = 8), heterosynaptic LTD was blocked, and no statistically significant changes in PPR and CV−2 were observed (Fig. 3 G and H and Fig. S6). Rather, there was a tendency for an increase in the heterosynaptic EPSC amplitude (nifedipine: +6.4 ± 8.9%, n = 5; nimodipine: +9.5 ± 14.5%, n = 8), which was accompanied by a decrease in PPR in the majority of the recorded cells (nifedipine: five of five cells; nimodipine: four of eight cells). This increase in EPSC amplitude was reminiscent of the effect of nifedipine in reversing the direction of heterosynaptic plasticity in dissociated cultures.
Fig. 3.
L-VGCCs control heterosynaptic LTD in acute hippocampal slices independently of postsynaptic Ca2+. (A) (Left) Experimental scheme in acute hippocampal slices: two independent Shaffer collateral inputs are stimulated with extracellular electrodes (S1 and S2), and corresponding responses are recorded in a CA1 neuron. A tetanic stimulation (3 × 100 Hz) of one of the inputs elicits homosynaptic LTP along with heterosynaptic LTD of the nonstimulated input. (Right) Photomicrograph of hippocampal area CA1 showing the stimulating and recording electrode positions: So, stratum oriens; Sp, stratum pyramidale; Sr, stratum radiatum. (Scale bar, 60 μm.) (B) Average EPSC traces evoked by a pairwise cross-stimulation of the two independent inputs show a lack of short-term plasticity. (C, E, and G) Time course of homosynaptic (light circles) and heterosynaptic (dark circles) EPSC amplitudes before and after the tetanic stimulation in control (C, n = 6 cells) or with intracellular postsynaptic BAPTA (B, n = 10 cells) or bath applied nifedipine (E, n = 5 cells). Postsynaptic BAPTA but not nifedipine blocks the homosynaptic LTP (postsynaptic BAPTA, **P = 0.0015; nifedipine, P = 0.9171; two-way ANOVA followed by Fisher’s LSD test). In contrast, nifedipine but not postsynaptic BAPTA blocks the heterosynaptic LTD (nifedipine, *P = 0.0474; postsynaptic BAPTA, P = 0.9518; two-way ANOVA followed by Fisher’s LSD test). Data are expressed as mean ± SEM. (D, F, and H) (Left) Representative heterosynaptic EPSC traces of PPR and plots comparing heterosynaptic PPR before and after the tetanus in the three conditions (paired t test: control, *P = 0.0381; postsynaptic BAPTA, **P = 0.0043; nifedipine, *P = 0.0286). (Right) Plots of heterosynaptic CV−2 analysis of heterosynaptic EPSCs before and after the tetanus in the three conditions (paired t test: control, *P = 0.0378; postsynaptic BAPTA, *P = 0.0199; nifedipine: P = 0.6993).
Fig. S6.
L-VGCCs control heterosynaptic LTD in acute hippocampal slices independently of postsynaptic Ca2+. (A and B) Time course of homosynaptic (light circles) and heterosynaptic (dark circles) EPSC amplitude changes relative to the baseline before and after the tetanic stimulation in nifedipine when the postsynaptic neuron is also filled with BAPTA (A; n = 10 cells) or in nimodipine alone (B; n = 8 cells). Heterosynaptic LTD is blocked in both conditions (Wilcoxon signed rank test: nifedipine + postsynaptic BAPTA, P = 0.3750; nimodipine, P = 0.7422). Homosynaptic LTP is blocked with postsynaptic BAPTA + nifedipine but not with nimodipine (Wilcoxon signed rank test; postsynaptic BAPTA + nifedipine, P = 0.9219; nimodipine, *P = 0.0391). Data are expressed as mean ± SEM. (C and D) (Left) Representative EPSCs traces of PPR from heterosynapses and corresponding plots of PPR before and after the tetanus for recordings in A and B (paired t test: postsynaptic BAPTA + nifedipine, P = 0.3309; nimodipine, P = 0.6420). (Right) Plots showing the heterosynaptic CV−2 of EPSC amplitudes before and after the tetanus for recordings in A and B (paired t test: postsynaptic BAPTA + nifedipine, P = 0.3246; nimodipine, P = 0.8699).
Taken together, these results suggest that the mechanism of expression of heterosynaptic presynaptic plasticity is shared between hippocampal dissociated cultures and acute slices, and although independent of postsynaptic Ca2+, it involves NMDARs and astrocytes. Moreover, the finding that the polarity of heterosynaptic plasticity is reversed by L-VGCC inhibitors suggests that the heterosynaptic plasticity might regulate the disparity of presynaptic strengths between inputs.

Astrocyte NMDARs and L-VGCCs Promote Differences in Basal PPR of Convergent Inputs.

In the above experiments, the counterbalancing of presynaptic strengths between convergent inputs was observed on deliberately stimulating one of the inputs to elicit homo- and heterosynaptic plasticity. We wondered whether the heterosynaptic coordination is exclusive to the induction of synaptic plasticity, or alternatively, stimulation could have simply exaggerated the presynaptic regulation that normally occurs under basal activity levels. If the latter were the case, one would expect to observe differences in the basal presynaptic strengths of convergent inputs, where the heterogeneity is regulated by astrocytes, NMDARs, and also by L-VGCCs. We next sought to test this possibility by comparing the basal presynaptic strengths of convergent inputs in cultures and acute slices.
Comparisons of PPR under basal conditions revealed that the PPR was heterogeneous and uncorrelated between two presynaptic neurons targeting a common postsynaptic neuron in culture and for two independent Schaffer collateral inputs onto a CA1 neuron in acute slices (Fig. 4 A and B and Fig. S7 A and B). In both cultures and acute slices, bath applying AP5 (50 μM) or nifedipine (or nimodipine, 10 μM) in the absence of any conditioning stimulation decreased the PPR difference between the convergent inputs, and basal PPR became more correlated across experiments (Fig. 4 C–E and Fig. S7 C and D). Moreover, in cultures, 30-min treatment with fluoroacetate (5 mM) that inhibited heterosynaptic plasticity also promoted the correlation of PPR between inputs (Fig. S7E). Whereas none of the drug treatments affected the average PPR compared with the control condition (control, 1.90 ± 0.12, n = 29; AP5, 1.86 ± 0.11, n = 11; nifedipine, 1.74 ± 0.10, n = 17, nimodipine, 2.21 ± 0.11, n = 11; Fig. S5 B and D), strikingly, in all drug treatment conditions, the PPR difference between the two inputs (the PPR disparity) was decreased by more than 50% in slices (control: 0.59 ± 0.13, n = 29; AP5: 0.23 ± 0.06, n = 9; nifedipine: 0.19 ± 0.06, n = 17; nimodipine: 0.23 ± 0.07, n = 11) and in cultures (control: 0.66 ± 0.17, n = 10; AP5: 0.20 ± 0.06, n = 9; fluoroacetate: 0.18 ± 0.05, n = 9; nifedipine: 0.28 ± 0.06, n = 7; Fig. 4F and Fig. S7F). Therefore, conditions that compromised heterosynaptic plasticity resulted in increased correlation of basal PPR. The increase in correlation occurred relatively rapidly, in that nimodipine application in acute slices decreased the PPR disparity between the two inputs to its full extent within 10–15 min (−48.2 ± 9.3%, n = 6; Fig. 4G). Altogether, these results suggest the existence of a cellular process involving astrocytes, NMDARs, and L-VGCCs that maintain variations in presynaptic strengths under basal conditions, which appears to be a basic process that is not strictly dependent on the native hippocampal circuit but is reproduced in a simplified culture network, and that this process is embedded in the mechanism of heterosynaptic plasticity.
Fig. 4.
NMDARs and l-VGCCs activity in astrocytes decorrelate presynaptic strengths of convergent inputs in hippocampal slices. (A) Scheme in acute hippocampal slices. (B–E) Scatter plots comparing basal PPR of the two independent inputs normalized to the fist EPSC amplitude for control (n = 29 cells, r2 = 0.06, P = 0.1822), AP5 (n = 9 cells, r2 = 0.26, P = 0.1998), nifedipine (n = 17 cells, r2 = 0.57, ***P = 0.0004), and nimodipine (n = 11 cells, r2 = 0.67, **P = 0.0022). (Inset) Average traces from representative recordings. (F) Summary of the average basal PPR difference for the same conditions (one-way ANOVA followed by Holm–Sidak’s multiple comparison test, *P < 0.05, **P < 0.01). Data are expressed as mean ± SEM. (G) Time course of PPR disparity between two inputs when nimodipine is added to the bath (n = 6) or not (n = 7) (two-sample t test, *P < 0.05, **P < 0.01). (H) (Upper) Experimental scheme as in A with concurrent patch-clamping of astrocytes that are intracellularly coupled via GAP junctions. (Lower) Fluorescence image of an experiment showing a CA1 neuron filled with a green dye and the intracellular spread of a red dye across the astrocyte syncytium. (Scale bar, 30 μm.) (I–M) Scatter plots comparing basal PPR of the two independent inputs when astrocytes are dialyzed with control intracellular solution (n = 12 cells, r2 = 0.16, P = 0.1994), MK-801 (n = 11 cells, r2 = 0.63, **P = 0.0038), BAPTA (n = 12 cells, r2 = 0.75, ***P = 0.0003), D890 (n = 9 cells, r2 = 0.66, **P = 0.0077), or QX-314 (n = 11 cells, r2 = 0.90, ****P < 0.0001). (Inset) Average traces from representative recordings. (N) Summary of the average basal PPR difference for the same conditions (one-way ANOVA followed by Holm–Sidak’s multiple comparison test, *P < 0.05, **P < 0.01). Data are expressed as mean ± SEM. (Scale bar for insets of normalized PPR traces, 25 ms.)
Fig. S7.
NMDAR and L-VGCC activity in astrocytes decorrelate presynaptic strengths of convergent inputs in hippocampal cultures. (A) Experimental scheme for comparing two independent convergent inputs in triplets of neurons in cultures. Monosynaptic currents recorded in the postsynaptic neuron by stimulating each of the two presynaptic neurons are compared. (B–E) Scatter plots and average traces from representative cells comparing basal PPR of the two independent inputs for control (n = 10 cells, r2 = 0.04, P = 0.5834), AP5 (n = 9 cells, r2 = 0.6960, **P = 0.0052), nifedipine (n = 7 cells, r2 = 0.88, ***P = 0.0016), and fluoroacetate (n = 9 cells, r2 = 0.81, **P = 0.001). (F) Summary of the average basal PPR difference for the same conditions (one-way ANOVA followed by Holm–Sidak’s multiple comparison test, *P < 0.05, **P < 0.01). Data are expressed as mean ± SEM.
We next sought to obtain further insights into how astrocytes regulated the presynaptic strength heterogeneity and the relationship between astrocytes, NMDARs, and L-VGCCs in this mechanism. Having confirmed that the basic properties of heterosynaptic coordination of presynaptic strengths were shared between cultures and acute slices, we examined more detailed mechanisms primarily in acute hippocampal slices. We first determined a requirement for astrocyte Ca2+ signaling by whole-cell patch clamping an astrocyte in the area CA1 with a pipette containing BAPTA (30 mM) and waiting for >15 min. During this time, Alexa dye that was also included in the patch pipette spread across the astrocyte network that suggested of extensive gap junctional coupling (Fig. 4H); in turn, this observation indicated that BAPTA could also spread across the astrocyte network. Recording from a CA1 neuron close to the patched astrocyte, a comparison of the PPR of two independent Schaffer collateral inputs showed an ∼50% reduction in the PPR disparity compared with control recordings in which an astrocyte was patched without BAPTA for the same duration (control: 0.42 ± 0.11, n = 12; BAPTA: 0.20 ± 0.04, n = 12; Fig. 4 I, J, and N). Thus, astrocyte Ca2+ plays a role in decorrelating convergent presynaptic strengths.
We next sought to clarify the cellular location of NMDARs and L-VGCCs with respect to their involvement in the astrocyte-dependent mechanism of presynaptic strength regulation. Previous reports have suggested that both NMDARs and L-VGCCs are expressed in hippocampal astrocytes (4348, but see ref. 49). Therefore, we tested whether NMDARs and L-VGCCs on astrocytes contributed to the observed PPR disparity. Analogous to the BAPTA experiment described above, the channels were blocked intracellularly by perfusing drugs into astrocytes via the patch pipette. We tested the following inhibitors: MK-801 (1 mM), a noncompetitive antagonist of NMDARs that was shown to block astrocyte NMDARs intracellularly (50); D890 (2 mM), a membrane impermeant verapamil derivative that blocks L-VGCCs (51, 52); and QX-314 (10 mM), a lidocaine derivative that has been shown to block voltage-gated channels including L-VGCCs (53). All three inhibitors reduced the PPR disparity between two convergent inputs by >50% compared with the control (control: 0.42 ± 0.11, n = 12; MK-801: 0.12 ± 0.03, n = 12; D890: 0.16 ± 0.06, n = 9; QX-314: 0.11 ± 0.03, n = 11; Fig. 4 K–N).
Collectively, these results implicate a Ca2+-dependent signaling mechanism involving NMDAR and L-VGCC activities in astrocytes in decorrelating presynaptic strengths of convergent inputs.

Activation of Astrocyte NMDARs Depolarizes Astrocyte Membrane and Opens L-VGCCs.

To further clarify the contribution of NMDARs and L-VGCCs expressed in astrocytes to the observed regulation of PPR, we investigated whether astrocytes could directly respond to NMDA application and if this response required NMDAR or L-VGCC activity specifically in astrocytes. We performed whole-cell patch-clamp recordings from astrocytes in the area CA1 of hippocampal slices (Fig. 5A). Astrocytes showed a linear I-V relationship with a low resting membrane potential under I-clamp (−85.2 ± 0.89 mV, n = 12; Fig. 5B), and a dye included in the patch pipette efficiently diffused throughout the astrocyte network (Fig. 4H). On bath applying NMDA and glycine (20 µM each for 3 min) in the presence of TTX (0.5 µM) to block action potentials (Fig. 5A), we observed a large and slow depolarization of the astrocyte membrane potential (peak depolarization, 30.67 ± 0.59 mV, n = 12; Fig. 5 C and D), which was in agreement with previous studies (46, 50). In addition, observation of the NMDA response despite of the low resting astrocyte membrane potential and in the presence of 1.5 mM extracellular Mg2+ was reminiscent of the weak Mg2+ block found for NMDARs expressed by cortical astrocytes (54). Lowering extracellular Ca2+ (from 2 to 0.25 mM) or dialyzing astrocytes with MK-801 (1 mM) for 15 min before applying NMDA+glycine significantly reduced the peak depolarization by ∼45% and ∼65%, respectively (low Ca2+: 16.52 ± 4.36 mV, n = 4; intracellular MK-801: 11.63 ± 2.97 mV, n = 10); moreover, blocking L-VGCCs with bath applied nifedipine (10 μM) or nimodipine (10 μM), or intracellularly loading D890 (2 mM) or QX-314 (10 mM) also attenuated the peak depolarization by 25–45% (nifedipine: 17.12 ± 3.36 mV, n = 8; nimodipine: 21.02 ± 3.11 mV, n = 9; D890: 22.82 ± 2.2 mV, n = 9; QX-314: 23.59 ± 1.85 mV, n = 9; Fig. 5 C and D). Together, these results indicate that NMDAR activation in astrocytes triggers astrocyte membrane depolarization that is strongly dependent on Ca2+-influx, and the depolarization, in turn, triggers voltage-gated conductances, including those mediated by L-VGCCs.
Fig. 5.
Astrocyte NMDAR activation depolarizes astrocyte membrane and promotes PPR decorrelation. (A) Experimental scheme in acute hippocampal slices. TTX is present throughout, and astrocyte membrane potential change is measured from whole-cell patch-clamp recordings. (B) Astrocyte identification based on the typical linear I-V relationship (Right) and the morphology visualized by infusing a dye via the patch pipette (Left), which spreads across other cells (arrows) of the syncytium. (C) Representative traces of astrocyte whole-cell patch clamp recordings showing membrane depolarization induced by bath applied NMDA+glycine in control, in low extracellular Ca2+, in nifedipine or nimodipine, or on infusing astrocytes intracellularly with MK-801, D890, or QX-314. (D) Summary data of the effect of different blockers on astrocyte depolarization induced by NMDA+glycine (one-way ANOVA followed by Holm–Sidak’s multiple comparison test, *P < 0.05, ***P < 0.001, ****P < 0.0001). (E) (Left) Experimental strategy for GRIN1 genetic deletion in CA1 astrocytes. AAV DJ/8 carrying Cre-mCherry under hGFAP promoter is microinjected into the hippocampal area CA1 of GRIN1 floxed mice. (Right) Confocal section showing specific expression of Cre (red, arrows) in GFAP-expressing astrocytes (green) but not in NeuN-expressing neurons (blue). (F) (Left) Average traces from patch-clamp recordings of Cre-mCherry (Cre, red) or GFP-expressing (Control, black) astrocytes showing depolarization induced by NMDA+glycine in presence of CNQX and CdCl2. (Right) Summary data of the average peak depolarization in EGFP or Cre-mCherry expressing astrocytes (unpaired t test, P = 0.0006). (G) Scatter plots comparing basal PPR of two independent inputs when astrocytes express EGFP (Control, n = 23 cells, r2 = 0.16, P = 0.1223) or Cre-mCherry (Cre, n = 23 cells, r2 = 0.47, P = 0.0003). (Inset) Average traces from representative recordings. (Scale bar, 20 ms.) (H) Summary of the average basal PPR difference (Left; unpaired t test, P = 0.0188) and average PPR (Right; unpaired t test, P = 0.1494) for the same conditions. Data are expressed as mean ± SEM.

GluN1 Expressed in Astrocytes Promotes the Differences in PPR of Convergent Inputs.

We next used a genetic approach to confirm the contribution of astrocyte NMDARs in controlling the PPR disparity between convergent inputs. To impair NMDAR activity specifically in astrocytes, we used mice homozygous for the floxed GRIN1 gene encoding GluN1, the obligatory NMDAR subunit (55). Adeno-associated virus (AAV DJ/8) carrying either GFP or Cre-mCherry under the shortened version of the human glial fibrillary acidic protein (hGFAP) promoter (56) was microinjected bilaterally into the hippocampal area CA1 of 5- to 6-wk-old GRIN1 floxed mice. Two weeks after the injection, Cre-mCherry was expressed in GFAP-positive cells that showed a highly ramified morphology typical of astrocytes but not in cells labeled for NeuN, a neuronal marker (Fig. 5E; 0.7% of mCherry positive cells colabeled with NeuN; n = 138 cells from two slices). The robust neuronal expression of NMDARs might interfere with assessing the efficacy of GluN1 conditional KO by a biochemical or immunolabeling approach. Therefore, we used a functional assay by whole-cell patch-clamping astrocytes in hippocampal slices prepared from virus injected mice and monitoring membrane depolarization induced by NMDA+Gly application as described above except, in addition to TTX, Cd2+ (100 µM), and CNQX (10 µM) were also present in the bath to block VGCCs and AMPARs, respectively. In slices from mice injected with the Cre AAV, peak membrane depolarization induced by NMDA+Gly was reduced by 60% relative to recordings from control AAV slices (control: 10.4 ± 1.4 mV, n = 14; Cre: 4.4 ± 1.7 mV, n = 14; Fig. 5F); the decrease was similar to the extent inhibition achieved by intracellular perfusion of MK-801 into astrocytes via the patch electrode (Fig. 5 C and D). Under these conditions, recordings from a CA1 neuron showed that PPR disparity of convergent Schaffer collateral inputs was reduced by ∼50% (control: 0.39 ± 0.07, n = 23; Cre: 0.19 ± 0.03, n = 23; Fig. 5 G and H). These observations provide further support to the key role for astrocyte NMDA receptors in promoting basal synaptic strength heterogeneity.

Optogenetic Hyperpolarization of Astrocytes Correlates Basal PPR of Convergent Inputs.

Taken together, the above experiments suggested that directly activating astrocyte NMDARs induced depolarization of the astrocyte membrane that was needed to activate L-VGCCs, which, along with NMDARs, were required for the observed decorrelation of basal presynaptic strengths. If this were the case, then preventing astrocyte membrane depolarization should reverse the decorrelation (similar to the L-VGCC blockade), and the presynaptic strengths of convergent inputs should become more similar. Given the highly ramified structure of astrocytes and their low input resistance that made controlling their membrane potential difficult by standard electrophysiology methods, we took an optogenetic approach and used light-induced hyperpolarization of astrocyte membrane potential using ArchT (57) to test the effect on the PPR differences between convergent inputs.
AAV (DJ/8) carrying either GFP or ArchT-GFP under the full-length hGFAP promoter was microinjected bilaterally into the hippocampal area CA1 of 3-wk-old mice. After 6 d, GFP or ArchT-GFP was reliably expressed in GFAP-positive cells (Fig. 6 A and B). The expression was highly specific to astrocytes in that cells double labeled for GFP and NeuN were limited to 0.8% of neurons in area CA1 (5 of 666 NeuN-positive cells showed colabeling with GFP; n = 4 slices). Using acute hippocampal slices prepared from virus-injected animals, we performed whole-cell patch-clamp recordings from ArchT-GFP–expressing astrocytes. Light stimulation (Materials and Methods) produced a reversible hyperpolarization of the astrocyte membrane potential under I-clamp (from −78.36 ± 0.35 to −102.51 ± 3.58 mV, n = 7; Fig. 6C) and outward currents were detected in V-clamp (Fig. S8) similarly to a recent study that expressed ArchT in astrocytes in the cerebellum (58).
Fig. 6.
Optogenetic hyperpolarization of astrocytes produces the correlation of PPR of convergent inputs reversibly. (A) Confocal section showing specific expression of EGFP in GFAP-expressing astrocytes (red) but not in NeuN-expressing neurons (blue). (B) Astrocyte-specific expression of ArchT-GFP (green) throughout the GFAP-positive processes (red) with stereotyped astrocyte morphology. (C) Membrane potential hyperpolarization induced by light (n = 7 cells, paired t test, ***P = 0.0005) and representative membrane potential trace acquired in I-clamp in acute hippocampal slices. (D) Time course of the EPSC amplitude changes recorded from CA1 neurons near astrocytes infected with EGFP (n = 6 cells) or ArchT-GFP (n = 7 cells) in acute slices before, during, and after 10-min light exposure (two-sample t test with Welch correction, *P = 0.00826). [Scale bars for representative traces (Upper), 50 pA, 20 ms.] (E) Time course of PPR disparity for the recordings in D (two sample-t test with Welch correction, **P = 0.00609). (F and G) Scatter plots comparing basal PPR of two independent inputs normalized to the first EPSC amplitude in control (Left) or with light exposure (Right) when nearby astrocytes are infected with EGFP (F: no light, n = 17 cells, r2 = 0.000003, P = 0.9950; light, n = 9 cells, r2 = 0.01, P = 0.7897) or ArchT-GFP (G: no light, n = 11 cells, r2 = 0.0007, P = 0.9392; light, n = 7 cells, r2 = 0.89, **P = 0.0015). (Inset) Representative PPR traces. (Scale bar, 40 ms.)
Fig. S8.
Photocurrents in ArchT-expressing astrocytes. (A) ArchT-expressing astrocytes show characteristic linear current-voltage relationship (n = 6 cells). (B) Representative traces showing instantaneous and slow photocurrents evoked by light. (C) Summary data of instantaneous and slow photocurrents amplitudes (n = 4 cells).
We then monitored EPSC amplitude and PPR of two independent inputs received by a CA1 neuron and tested the effect of a continuous, 10-min light activation of astrocyte ArchT. Light stimulation, which hyperpolarized astrocytes, rapidly and reversibly increased the EPSC amplitude relative to the baseline (ArchT: +27.8 ± 11.8%, n = 7; control: −6.01 ± 6.04%, n = 6; Fig. 6D). This increase in EPSC amplitude indicated that basal depolarization of astrocytes provided an inhibitory tone on excitatory synaptic transmission. Remarkably, compared with the baseline, light stimulation decreased the PPR disparity between the two inputs in slices expressing ArchT but not in GFP control (at time 35 min, control: +33.3 ± 20.4%, n = 6; ArchT: −64.6 ± 20.5%, n = 7; Fig. 6 E–G). Moreover, reminiscent of the time course of the effect of blocking L-VGCCs (Fig. 4G), on light stimulation of ArchT, the PPR disparity was decreased by greater than 50% over several minutes; this effect was reversible as the level of disparity returned to the baseline 20–25 min after shutting off the light (Fig. 6E). Therefore, our finding strongly implicates astrocyte membrane depolarization in the mechanism that promotes the decorrelation of PPR.

Discussion

Here we identified an astrocyte-dependent cellular process that serves to enhance the heterogeneity of presynaptic strengths by increasing the PPR disparity between inputs targeting the same neuron. This mechanism shares its key properties with the mechanism of heterosynaptic plasticity that we have also studied here, which counterbalances PPR of the nonstimulated input relative to that of the stimulated input. Furthermore, this astrocyte-dependent presynaptic regulation is observed in acute hippocampal slices and is recapitulated in dissociated cultures, a simplified system without the native topological organization of the hippocampal circuit. That the features of this presynaptic regulation are conserved in different conditions—i.e., between the basal state and during synaptic plasticity—and across different experimental preparations underscores the fundamental nature of the underlying mechanism.
The mechanism for generating presynaptic strength heterogeneity, which also supports heterosynaptic plasticity, may play an important role in the developing nervous system. For instance, this mechanism may represent a powerful way to orchestrate synapse competition where the relative strengths of competing terminals biases the winner by favoring stabilization of strong synapses at the expense of weaker synapses (18, 59, 60). Specifically, the mechanism we describe here could promote a two-step process where first it will help create differences in input strengths to ensure variability; subsequently, it will facilitate the selective stabilization of the strongest input (59, 60). Importantly, the astrocyte-dependent mechanism that controls the disparity or the decorrelation of PPR we report here is not limited to the developing brain. Acute hippocampal slices from adult mice, which have been used for GluN1 conditional KO and ArchT experiments, show the dependence on astrocyte NMDARs and membrane depolarization for maintaining the PPR decorrelation (Figs. 5 E–H and 6). Therefore, the mechanism that promotes the presynaptic strength heterogeneity is likely to function in concert with the various rules of synaptic plasticity that operates in the adult hippocampal circuit. For example, decorrelated presynaptic strengths may boost the circuit responsiveness under sparse activity conditions (61) or facilitate the reorganization of correlated networks that is associated with learning (62). Moreover, its dysregulation might culminate in diseased states; for instance, unrestrained correlation may nucleate synchronization of activity across synaptic networks, which is a hallmark feature of epileptic disorders (63).
Mounting evidence supports a role for astrocytes in regulating synaptic transmission and synaptic plasticity (15, 64). In general, astrocyte influence on neurons has been thought to be global, a view established by the prominent slow Ca2+ transients that spread through the astrocyte network. However, recent studies have identified fast and local Ca2+ signals that regulate synapse function in astrocyte processes, which are in physical proximity of synapses (17, 1921). Our finding of the role for astrocytes in controlling presynaptic strengths is compatible with such locally generated signals. Astrocytes must be capable of deciphering signals associated to different inputs, and the decoding mechanism could engage local Ca2+ signals whose magnitudes might be related to the strength of synaptic inputs, as recently reported at the neuromuscular junction (18).
We find that the decorrelation of presynaptic strengths requires membrane depolarization, NMDARs and L-VGCCs, and Ca2+ signaling, all within astrocytes. Whereas the expression of NMDARs and L-VGCCs in astrocytes has been debated, our study is in line with previous studies reporting of the presence of these channels in astrocytes (4348, 50). With respect to astrocyte NMDARs, we find that applying NMDA in the presence of TTX to prevent synaptic transmission directly triggers astrocyte membrane depolarization, and this depolarization is compromised by intracellularly infusing MK-801 or deleting GRIN1 in astrocytes. That we could obtain astrocyte-dependent NMDAR responses reliably in the presence of 1.5 mM extracellular Mg2+ even though astrocytes have low resting membrane potential is compatible with the presence of NR3 containing NMDARs with reduced Mg2+ block reported for cortical astrocytes (54). Furthermore, a function for astrocyte NMDARs in modulating presynaptic release is supported by observations in which intracellularly infusing MK-801 or conditionally deleting GRIN1 in astrocytes reduces the PPR disparity between convergent inputs. Similarly, the expression of L-VGCCs in astrocytes is supported by the decrease in PPR disparity of convergent inputs on intracellularly delivering D890 into astrocytes.
How are these astrocyte-specific components—membrane depolarization, activation of NMDARs and L-VGCCs, and astrocyte Ca2+ signaling—orchestrated to impose the presynaptic strength differences between inputs? Whereas we sampled, for every experiment, a set of two presynaptic inputs that converge onto the postsynaptic target neuron, the postsynaptic neuron receives synaptic inputs from numerous other afferents. One would then expect that the presynaptic strengths of other nonsampled inputs might be similarly decorrelated. Given such a situation, it is doubtful that an astrocyte mechanism exists that can specifically compare and counterbalance the strengths between a pair of presynaptic inputs. Rather, the decorrelated presynaptic strengths might emerge out of a local control of single synapses that is embedded within a global form of synapse regulation. Following experimental observations support the involvement of per synapse basis regulation. First, in heterosynaptic plasticity experiments, we find that, although the heterosynaptic EPSC amplitude and PPR changes are inversely related to the initial PPR of the stimulated input (Fig. 2 A and B), they do not show a relationship to the initial EPSC amplitude of the stimulated input (Fig. S3). This lack of relationship to the basal EPSC amplitude indicates that the presynaptic strength change at the nonstimulated input does not necessarily depend on the total number of active, stimulated synapses, and suggests that the presynaptic control could be executed independently of the coincident activation of many synapses. Second, the preservation of the astrocyte-dependent decorrelation of PPR in dissociated cultures indicates that the underlying mechanism does not rely on the unique topological organization of hippocampal area CA1 astrocytes and pyramidal neurons (65). A regulatory process that operates on the basis of individual synapses that functions in concert with a global form of regulation is compatible with a simple system in which the spatial organization of the synaptic connections is variable. Although a single astrocyte in area CA1 is likely to contact more than 105 synapses (65), given the extensive gap junctional coupling of astrocytes, the spatial extent to which the present form of regulation operates remains to be addressed.
The mechanism that ensures presynaptic strength heterogeneity we describe here highlights a form of homeostatic function for the postsynaptic neuron. That is, by compensating high pr synaptic inputs by weakening of other synaptic inputs received by the postsynaptic neuron, it provides a boundary to the total excitatory drive. This compensatory activity functions over the time scale of minutes as illustrated by the rapid loss of decorrelation on blocking L-VGCCs or hyperpolarizing of astrocyte membrane by ArchT (Figs. 4G and 6 E and G). Notably, the overall average presynaptic strength received by the neuron is not altered under conditions when the decorrelation of presynaptic strengths between convergent inputs is decreased (Fig. S5). This lack of change in average synaptic strength suggests that the mechanism that decorrelates presynaptic strengths itself is not setting the limits for the range of excitatory drive but functions in compliance with the homeostat to produce the variability.
Based on our findings, we propose the following scheme for initiating the decorrelation of presynaptic strengths (Fig. S9). We postulate that astrocytes impart a suppressive tone on excitatory synaptic transmission (27, 66, 67); this could be mediated via the tonic release of inhibitory gliotransmitters such as ATP that is hydrolyzed extracellularly into adenosine or endocannabinoids (15, 64, 68). Glutamate released from active synapses (with a high pr) activates NMDARs on the surface of a nearby astrocyte process, causing a local depolarization of the astrocyte membrane. The membrane depolarization in turn, opens L-VGCCs that engage intracellular Ca2+ signaling within astrocytes. We propose that the change in global Ca2+ signaling serves to strengthen the tonic inhibitory tone on excitatory synapses, whereas locally, the active synapses are protected from the inhibition. The model involves a global signal that acts in concert with a local signal, and it takes into account the astrocyte membrane depolarization, NMDARs and L-VGCCs on astrocytes, and astrocyte Ca2+ signaling that are required for the observed PPR decorrelation. Notably, the model also explains the reversal of the polarity of heterosynaptic depression by the L-VGCC inhibitor, which compromises the global Ca2+ signaling that is proposed to stimulate the inhibitory tone on excitatory synapses. That astrocytes in general exert an inhibitory tone on excitatory synapses is supported by the ArchT experiments in which light-induced hyperpolarization of the astrocyte membrane potential is accompanied by a rapid increase of the EPSC amplitude (Fig. 6D) (66). At present we do not know the nature of the protective signal that spares active synapses from the inhibitory tone. The apparent protection from inhibition could at least in part result from a local positive feedback regulation by the active synapse (20) and likely involving astrocyte Ca2+ signaling that shows a highly complex spatial and temporal dynamics (15, 64, 69). Future studies warrant a careful examination of the various synaptically evoked signals to tease apart those crucial for controlling the presynaptic strength decorrelation.
Fig. S9.
Working model for the astrocyte-dependent decorrelation of presynaptic strengths. (A) Illustration highlights astrocyte-derived inhibitory tone (black curved arrows) acting on synapses received by a CA1 pyramidal neuron (blue). In the model, high release probability synapses (red) are, however, protected from the inhibition. (B) Glutamate released from a high pr synapse activates NMDARs on the surface of a nearby astrocyte process, causing a local depolarization of the astrocyte membrane. This in turn activates L-VGCCs that engage intracellular Ca2+ signaling within astrocytes. In our model, the change in global Ca2+ signaling strengthens the tonic inhibitory tone on excitatory synapses, whereas locally, the high pr synapses are protected from the inhibition. See Discussion for detail.

Materials and Methods

Cell Culture Preparations and Transfection.

Hippocampal cultures were prepared from P0–P1 rats and plated at low density onto an astrocyte monolayer. The cultures were maintained as described previously (1) and used for electrophysiology experiments at days in vitro (DIV) 9–14. To probe synaptic vesicles dynamics, neurons were transfected with a plasmid encoding VGLUT1-pHluorin (kindly provided by Robert Edwards, University of California, San Francisco) at DIV 6 using Lipofectamine 2000 (Invitrogen). Cultures were used for imaging/electrophysiology experiments at DIV 9–14.

In Vivo Virus Injections.

Infection of astrocytes in vivo was performed as described previously (70). For ArchT experiments, full-length hGFAP promoter constructs were used, and 700 nL AAV-DJ/8 virus solutions were bilaterally injected into brains of 3-wk-old C57BL/6J mice. Targeting coordinates for dorsal hippocampus area CA1 were +1.9 mm anteroposterior from bregma, ±1.7 mm mediolateral, and +1.6 mm dorsoventral. Concentrations of recombinant AAV, determined by real-time quantitative PCR, were 1.0 × 1013 viral particles (vp)/mL for both AAV-DJ/8–hGFAP-GFP and AAV-DJ/8–hGFAP-ArchT-EGFP. For GluN1 conditional KO experiments, Cre and control AAV constructs were based on a shortened hGFAP promoter (56), and virus solutions were injected into brains of 5- to 6-wk-old mice homozygous for the floxed gene encoding GluN1 (55). The stereotaxic coordinates were as follows: X (anteroposterior from bregma), +1.9 mm; Y (mediolateral), ±1.9; Z (dorsoventral), +1.4 mm; and X, +3 mm; Y, ±2.7; Z, +1.4 mm in both hemispheres. Recombinant AAV concentrations were 9.0 × 1012 vp/mL for AAV-DJ/8-GFAP104-EGFP and 5.9 × 1012 vp/mL for AAV-DJ/8-GFAP104-nlsCre-mCherry, respectively.

Hippocampal Slice Preparations.

Transverse hippocampal slices were obtained from young (P14–P21) male Sprague–Dawley rats. For experiments involving astrocyte expression of control GFP or ArchT, acute hippocampal slices were prepared from 4-wk-old mice at least 6 d after virus injection, and for conditional GluN1 KO experiments, slices were made from 7- to 8-wk-old mice at 2 wk after virus injection (above). For details, see SI Materials and Methods.

Electrophysiology.

Patch-clamp recordings from dissociated cultures and acute hippocampal slices were performed at room temperature using Axopatch 200B and Multiclamp 700B amplifiers (Axon Instruments). For details, see SI Materials and Methods.

Live Cell Imaging.

For estimating the RRP size at single boutons in dissociated cultures, VGLUT1-pHluorin (VGLUT1-pH)–transfected neuron was patch-clamped along with a postsynaptic neuron filled with 100 μM AF 594 dye. The presynaptic neuron was stimulated at 20 Hz for 2 s (100 mV, 1- to 2-ms step depolarization) under V-clamp. Time-lapse VGLUT1-pH images were acquired at 1Hz on an iXon EMCCD camera (Andor Technology) driven by Metamorph software (Universal Imaging). ΔF/F for identified active boutons was measured after subtracting local background, where F is the initial fluorescence. This measurement was repeated before and 20 min after the 1-Hz, 3-min stimulation of the presynaptic neuron.

Immunohistochemistry.

Brain sections from mice injected with AAV were prepared and processed for immunohistochemistry using standard procedures. For details, see SI Materials and Methods.

Light Stimulation.

The light at excitation wavelength for red fluorescence probes from Spectra X light engine (Lumencor) was passed through a neural density filter and a 560/40-nm filter, and delivered through a 40×, 0.8 N.A. water-immersion objective. Under this condition, the power density measured at 560 nm was 2 mW/mm2.

Statistics.

For normally distributed data (as determined by the d’Agostino–Pearson normality test), differences were tested using the paired or unpaired two-tailed Student t test or one-way ANOVA. The Mann–Whitney test, the Wilcoxon test, or the Kruskal–Wallis test was used when criteria for normality were not met. Graphpad Prism software was used for statistical analysis. Data are expressed as mean ± SEM.

SI Materials and Methods

Electrophysiological Recordings in Dissociated Cultures.

Whole-cell patch-clamp recordings were carried out from cultures placed on the stage of an Olympus IX71 inverted microscope or an Olympus BX51 upright microscope at room temperature and using Axopatch 200B and Multiclamp amplifiers (Axon Instruments). The recording chamber was continuously perfused with an artificial cerebrospinal fluid (ACSF) containing (in mM) 130 NaCl, 2.5 KCl, 2.2 CaCl2, 1.5 MgCl2, 10 d-glucose, 10 Hepes, and 0.1 picrotoxin (pH 7.35, osmolarity adjusted to 290 mOsm). The micropipettes were made from borosilicate glass capillaries, with a resistance in the range of 3–5 MΩ. The intracellular solution contained 100 mM K-gluconate, 17 mM KCl, 5 mM NaCl, 5 mM MgCl2, 10 mM Hepes, 0.5 mM EGTA, 4 mM ATPK2, and 0.5 mM GTPNa (pH 7.3, osmolarity adjusted to 280 mOsm).
For multiple perforated patch-clamp recordings, the pipettes were tip-filled with internal solution and then back filled with internal solution containing 150 μg/mL amphotericin B. For some recordings, internal solution contained BAPTA (30 mM) to buffer intracellular Ca2+ in whole-cell configuration. To assess connectivity among neurons in simultaneous multiple recording experiments, each neuron was stimulated by 1- to 2-ms step depolarization from −80 to +20 mV in V-clamp mode while identifying the neurons responding with EPSCs. Monosynaptic connections were identified by a short latency between the stimulation artifact and EPSC onset (<10 ms). To induce long-lasting changes in the synaptic strength of a given input, we applied brief depolarization at 1 Hz for 3 min in one of the presynaptic neurons under V-clamp to elicit action potentials while holding the postsynaptic neuron in I-clamp mode. Series resistance (Rs) was always lower than 30 MΩ and left uncompensated. Recordings in which Rs changed by more than 20% were discarded. PPRs were calculated from average traces obtained from a minimum of 15 traces.

Hippocampal Slice Preparations.

Transverse hippocampal slices were obtained from young (P14–P21) male Sprague–Dawley rats. Rats were anesthetized with isoflurane and decapitated. The brain was rapidly removed and placed in ice-cold dissection buffer containing (in mM) 252 sucrose, 3 KCl, 7 MgCl2, 0.5 CaCl2, 1.25 NaH2PO4, 26 NaHCO3, and 25 glucose. The cerebellum was removed with a 20°–30° angle between dorso-ventral and caudo-rostral directions and the resulting flat cut surface was glued onto a precooled vibratome plate; 300/400-µm sections were collected in the ice-cold dissection buffer using a Pelco vibrating microtome Series 1000 or Leica VT1200S and maintained at 32 °C for 1–2 h in ACSF containing (in mM) 125 NaCl, 2.5 KCl, 26 NaHCO3, 1.25 NaH2PO4, 2 CaCl2, 1 MgCl2, and 25 glucose. Dissection buffer and ACSF were bubbled with 95% O2/5% CO2. For experiments to test the effect of ArchT or Cre-mediated KO of GluN1 and control-EGFP in astrocytes via in vivo infection with AAV viruses, acute hippocampal slices were prepared from C57BL/6J mice or GRIN1 floxed homozygous mice that were infected with viruses at 3 or 5–6 wk old, respectively, and recovered from the stereotaxic surgery for at least 6 d up to 17 d.

Electrophysiological Recordings in Acute Hippocampal Slices.

Whole-cell patch-clamp recordings were carried out from CA1 neurons in acute hippocampal slices placed on the stage of an Olympus BX51 or a Nikon Eclipse FN1 upright microscope at room temperature and using Axopatch 200B and Multiclamp 700B amplifiers (Axon Instruments). The recording chamber was continuously perfused with ACSF bubbled with 95% O2/5% CO2 containing (in mM) 125 NaCl, 2.5 KCl, 26 NaHCO3, 1.25 NaH2PO4, 2 CaCl2, 1 MgCl2, and 25 glucose. For recordings in acute slices from P14–P21 rats, the internal solution was the same as for cultured neurons, and mEPSCs were recorded as described for cultures. For recordings in acute slices from mice injected with AAV virus, pipette solutions (285 mOsm with pH 7.25) contained 130 mM CsMeSO4, 8 mM NaCl, 4 mM Mg-ATP, 0.3 mM Na-GTP, 0.5 mM EGTA, 10 mM Hepes 10, and 6 mM QX-314. For comparing EPSCs from two independent Schaffer collateral pathways, two bipolar stimulation electrodes in borosilicate theta glass were filled with ACSF and placed in the stratum radiatum on each side of a recorded CA1 neuron (∼200 μm apart). Independence of the two pathways was assessed by measuring EPSC amplitudes after crossed paired-pulse stimulation in which the stimulation of one pathway preceded the stimulation of the other by an interpulse interval of 50 ms. For tetanization-induced heterosynaptic depression, three trains at 100 Hz for 1 s at 30-s intervals were given to one of the pathways (homosynapse). Rs was always lower than 30 MΩ and left uncompensated. Recordings in which Rs changed by more than 20% were discarded. EPSCs and PPR measurements were performed as for dissociated cultures.
Single astrocytes from cultures or slices were patch clamped with an internal solution containing 130 mM K-gluconate, 10 mM Hepes, 4 mM MgCl2, 4 Na2-ATP, 0.4 Na3-GTP, 10 Na-creatine, and 500 μM Alexa Fluor 350 or 250 μM Alexa Fluor 488, and held for at least 10–15 min before recording. Astrocytes were identified based on the lack of electrical excitability, a low membrane potential (< −75 mV), and a low input resistance (<10 MΩ). Moreover, they displayed an extensive network coupling as revealed by the diffusion of Alexa Fluor dyes in neighboring astrocytes. Calcium in the astrocytic network was buffered by including 30 mM BAPTA in the internal solution and decreasing K-gluconate accordingly; 10 mM QX-314, 2 mM D890, or 1 mM MK-801 was included in the internal solution to block VGCCs, L-VGCCs, and NMDARs, respectively.
To investigate whether the viral vectors expressing Cre into GRIN1 floxed mice effectively knocked out NMDARs in astrocytes, we prepared acute hippocampal slices from mice infected with AAV-DJ/8-GFAP104-nlsCre-mCherry or AAV-DJ/8-GFAP104-EGFP and monitored astrocyte membrane depolarization to bath applied NMDA + glycine (20 μM each). Recordings were made in the presence of a pharmacological mixture that included 100 μM CdCl2, 10 μM CNQX, 100 μM picrotoxin, and 0.5 μM TTX to minimize the contribution of VGCCs and synaptic effects that could confound the astrocyte NMDAR-mediated components.

Immunohistochemistry.

A week after virus injection, mice were transcardially perfused with saline followed by 4% paraformaldehyde (PFA). After an overnight postfixation in PFA, brains were washed in PBS and cut to 300-μm sections using a vibratome (Leica VT1200). Floating sections were then permeablized and blocked by 0.3% Triton X-100 and 1% goat serum albumin for 2 h at room temperature. Primary antibody incubations were then performed overnight at 4 °C in the presence of 0.3% Triton X-100 and 0.3% goat serum albumin. Secondary antibodies diluted in incubation buffer as desired were applied for 2 h at room temperature. Primary antibodies used for immunofluorescence labeling experiments with astrocyte and neuronal markers were rabbit anti-GFAP (1:500; Abcam), and mouse monoclonal anti-NeuN (1:200; Millipore). The expression of NeuN, GFAP, and GFP or mCherry was detected by goat anti-mouse Alexa 633 (1:1,000), goat anti-rabbit Alexa 555 (1:1,000), or goat anti-rabbit Alexa 488 (1:1,000). For double immunofluorescence labeling for ArchT-EGFP and GFAP, primary rabbit polyclonal anti-EGFP (1:500) and mouse-monoclonal anti-GFAP (1:200) were visualized by goat anti-mouse Alexa 555 (1:500) and goat anti-rabbit Alexa 488 (1:500). All of the secondary antibodies were purchased from Invitrogen.
The specificity of astrocyte targeting by hGFAP promoters used in AAV vectors was confirmed by immunolabeling for GFAP or NeuN with virally expressed EGFP signal in sections prepared from PFA-fixed brains, or with Cre-mCherry signal in acute hippocampal slices fixed with PFA. EGFP signal was amplified by anti-EGFP labeling whereas Cre-mCherry signal did not require amplification by antibody labeling. Images were captured on a Zeiss 780 confocal microscope using a 20×, 1.0 N.A. water immersion objective (1,024 × 1,024 pixels, 424.68 × 424.68 μm). For samples obtained from mice infected with pAAV-GFAP-EGFP, regions of interest (ROIs) were manually drawn around NeuN-labeled somata and checked for EGFP coexpression (four slices, three to four fields of view/slice, 666 somata). For samples infected with pAAV-GFAP-Cre-mCherry, ROIs were manually drawn around Cre-labeled puncta and checked for NeuN colabeling (two slices, three fields of view/slice, 138 puncta).

Acknowledgments

We thank Ayumu Konno, Hirokazu Hirai, Hajime Hirase, Eunmi Hwang, and Joshua Johansen for DNA constructs; Thomas McHugh for DNA constructs and GRIN1 floxed mice; Olivier Thoumine, Aude Panatier, Hajime Hirase, Yasunori Hayashi, and Taro Toyoizumi for discussions; David Elliott, Izumi Kono, and Mizuki Kurosawa for expert technical assistance; and Rachel Wong, Yasunori Hayashi, and Charles Yokoyama for comments on an earlier version of the manuscript. This work was supported by the Medical Research Council UK, European Union 7th Framework Program Grant HEALTH-F2-2009-241498 (“EUROSPIN” project), RIKEN Brain Science Institute, JSPS Core-to-Core Program, Grants-in-Aid for Scientific Research (15H04280) from the MEXT, and the Brain/MINDS from the Japan AMED.

Supporting Information

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Information & Authors

Information

Published in

Go to Proceedings of the National Academy of Sciences
Proceedings of the National Academy of Sciences
Vol. 113 | No. 19
May 10, 2016
PubMed: 27118849

Classifications

Submission history

Published online: April 26, 2016
Published in issue: May 10, 2016

Keywords

  1. synapse heterogeneity
  2. synaptic strength
  3. astrocyte
  4. hippocampal neuron
  5. heterosynaptic plasticity

Acknowledgments

We thank Ayumu Konno, Hirokazu Hirai, Hajime Hirase, Eunmi Hwang, and Joshua Johansen for DNA constructs; Thomas McHugh for DNA constructs and GRIN1 floxed mice; Olivier Thoumine, Aude Panatier, Hajime Hirase, Yasunori Hayashi, and Taro Toyoizumi for discussions; David Elliott, Izumi Kono, and Mizuki Kurosawa for expert technical assistance; and Rachel Wong, Yasunori Hayashi, and Charles Yokoyama for comments on an earlier version of the manuscript. This work was supported by the Medical Research Council UK, European Union 7th Framework Program Grant HEALTH-F2-2009-241498 (“EUROSPIN” project), RIKEN Brain Science Institute, JSPS Core-to-Core Program, Grants-in-Aid for Scientific Research (15H04280) from the MEXT, and the Brain/MINDS from the Japan AMED.

Notes

This article is a PNAS Direct Submission.

Authors

Affiliations

Mathieu Letellier3 [email protected]
Brain Science Institute, RIKEN, Saitama 351-0198, Japan;
Previous address: Medical Research Council Laboratory for Molecular Cell Biology and Cell Biology Unit, University College London, London WC1E 6BT, United Kingdom.
Present address: Interdisciplinary Institute for Neuroscience, University of Bordeaux, UMR 5297, 33077 Bordeaux, France.
Yun Kyung Park
Brain Science Institute, RIKEN, Saitama 351-0198, Japan;
Thomas E. Chater4
Brain Science Institute, RIKEN, Saitama 351-0198, Japan;
Peter H. Chipman4
Brain Science Institute, RIKEN, Saitama 351-0198, Japan;
Sunita Ghimire Gautam
Brain Science Institute, RIKEN, Saitama 351-0198, Japan;
Tomoko Oshima-Takago
Brain Science Institute, RIKEN, Saitama 351-0198, Japan;
Brain Science Institute, RIKEN, Saitama 351-0198, Japan;
Saitama University Brain Science Institute, Saitama University, Saitama 338-8570, Japan;
Department of Life Sciences, Graduate School of Arts and Sciences, University of Tokyo, Tokyo 153-8902, Japan
Previous address: Medical Research Council Laboratory for Molecular Cell Biology and Cell Biology Unit, University College London, London WC1E 6BT, United Kingdom.

Notes

3
To whom correspondence may be addressed. Email: [email protected] or [email protected].
Author contributions: M.L. and Y.G. designed research; M.L., Y.K.P., T.E.C., P.H.C., S.G.G., and T.O.-T. performed research; M.L., Y.K.P., T.E.C., and P.H.C. analyzed data; and M.L. and Y.G. wrote the paper.
4
T.E.C. and P.H.C. contributed equally to this work.

Competing Interests

The authors declare no conflict of interest.

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    Astrocytes regulate heterogeneity of presynaptic strengths in hippocampal networks
    Proceedings of the National Academy of Sciences
    • Vol. 113
    • No. 19
    • pp. 5137-E2759

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