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. Author manuscript; available in PMC: 2016 Feb 24.
Published in final edited form as: Cell Rep. 2016 Jan 7;14(3):471–478. doi: 10.1016/j.celrep.2015.12.061

An NMDA receptor-dependent mechanism underlies inhibitory synapse development

Xinglong Gu 1, Liang Zhou 1, Wei Lu 1
PMCID: PMC4765167  NIHMSID: NIHMS756089  PMID: 26774487

Summary

In the mammalian brain GABAergic synaptic transmission provides inhibitory balance to glutamatergic excitatory drive and controls neuronal output. The molecular mechanisms underlying the development of GABAergic synapses remain largely unclear. Here we report that NMDA-type ionotropic glutamate receptors (NMDARs) in individual immature neurons are the upstream signaling molecules essential for GABAergic synapse development, which requires signaling via Calmodulin binding motif in the C0 domain of the NMDAR GluN1 subunit. Interestingly, in neurons lacking NMDARs, while GABAergic synaptic transmission is strongly reduced, the tonic inhibition mediated by extrasynaptic GABAA receptors is increased, suggesting a compensatory mechanism for the lack of synaptic inhibition. These results demonstrate a crucial role for NMDARs in specifying the development of inhibitory synapses, and suggest an important mechanism for controlling the establishment of the balance between synaptic excitation and inhibition in the developing brain.

Introduction

Neural circuit function relies on precise information transfer between neurons through chemical synapses, which are either excitatory or inhibitory. Glutamate is the predominant excitatory neurotransmitter and mainly acts on AMPA-type and NMDA-type glutamate receptors (AMPARs and NMDARs) to mediate excitatory synaptic transmission. On the other hand, although GABA (gamma-aminobutyric acid) acting on GABAA-type ionotropic receptors (GABAARs) can elicit membrane depolarization in developing neurons due to higher intracellular Cl− concentration, GABA is the chief inhibitory neurotransmitter in the adult brain (Ben-Ari et al., 2007). In mature neurons, GABAergic inhibitory transmission balances glutamatergic excitatory input and controls neuronal excitability. The excitatory (E)/inhibitory (I) balance is established during development and delicately maintained in mature neurons, a process that is essential for cognition and behavior (Akerman and Cline, 2006; Cline, 2005; Dorrn et al., 2010; Maffei et al., 2004; Tao and Poo, 2005). When the development of chemical synapses is perturbed, the E/I balance can be impaired, which can result in devastating neurological and neuropsychiatric diseases, such as autism, schizophrenia and epilepsy (Chao et al., 2010; Cline, 2005; Dudek, 2009; Lisman, 2012; Rubenstein, 2010). Therefore, it is crucial to understand the regulatory mechanisms underlying the development of both excitatory and inhibitory synapses.

The molecular and cellular mechanisms underlying the development of excitatory glutamatergic synapses have been extensively investigated. In contrast, much less is known about the regulation of inhibitory GABAergic synapse development. Accumulating evidence demonstrates that neuronal activity regulates the development of inhibitory GABAergic synapses. Indeed, chronic and global blockade of TTX-sensitive neuronal activity triggered homeostatic reduction of neural inhibition and decreased inhibitory synapse density in developing neurons (Hartman et al., 2006; Kilman et al., 2002; Rutherford et al., 1997; Seil and Drake-Baumann, 1994). Surprisingly, however, selective suppression of neuronal activity in individual developing neurons had no effect on the development of inhibitory synapses (Hartman et al., 2006), indicating that at the level of individual neurons, neuronal activity is not essential for the development of inhibitory synapses. AMPARs and NMDARs are functionally expressed in embryonic neurons before glutamatergic synaptogenesis (Ben-Ari et al., 2007). Pharmacological studies with global inhibition of ionotropic glutamate receptor activities or genetic manipulation of glutamate receptors in developing neurons indicate that glutamate receptor activities regulate GABAergic synapse development (Aamodt et al., 2000; Gaiarsa, 2004; Hartman et al., 2006; Henneberger et al., 2005; Lu et al., 2013; Marty et al., 2000; Rosato-Siri et al., 2002). However, the precise role of glutamate receptors in inhibitory synapse development has been unclear. Here we employed a single-cell molecular replacement approach to demonstrate that at the level of individual developing neurons, signaling via the CaM-binding motif in the C0 domain of the NMDAR GluN1 subunit underlies the establishment of GABAergic transmission.

Results

GABAergic synapse development requires ionotropic glutamate receptors

To investigate the role of glutamate receptors in GABAergic synapse development, we utilized a quadruple conditional knockout mouse line in which three genes encoding AMPAR subunits (GluA1, A2 and A3) and the gene encoding the obligatory NMDAR GluN1 subunit are all conditional alleles (Gria1-3f/fGrin1f/f) (Lu et al., 2013). We in utero electroporated plasmids to sparsely express Cre fused to mCherry or GFP in hippocampal progenitor cells in E14.5 Gria1-3f/fGrin1f/f embryos to inactivate conditional alleles (Figure S1A and S1B) and established dissociated neuronal cultures at ~E18. We estimated that Cre-positive neurons accounted for less than 1% of the neurons in our cultures (data not shown), and thus the manipulation of glutamate receptor expression in these neurons should have little effect on overall neuronal network activity, allowing us to study the cell-autonomous role of ionotropic glutamate receptors in GABAergic synapse development. In our cultures GABAAR-mediated miniature inhibitory postsynaptic currents (mIPSCs) were rarely detected at DIV 3–4 and started to emerge at ~DIV 6 (data not shown). Thus, these nascent GABAergic synapses at DIV 6–7 should represent inhibitory synapses formed at early developmental stages.

Analysis of GABAergic miniature inhibitory postsynaptic currents (mIPSCs) in DIV6 (6 days in vitro) immature neurons in cultures revealed that there was approximately a 90% reduction of mIPSC frequency and a significant decrease of mIPSC amplitude in Cre-positive neurons (Figures 1A, left, and S1D; Table S1). The effect of loss of AMPARs and NMDARs on mIPSCs was not transient and not limited to neurons at early developmental stages, but was persistent as similar deficits in GABAergic transmission were observed in mature neurons at DIV15 (Figures 1A, right, and S1E; Table S1). GABAergic deficits in neurons without AMPARs and NMDARs were not due to general neuronal developmental deficits, as general dendritic development was not significantly changed (Figure 1B). To corroborate the data collected in vitro, we performed electrophysiological analysis in CA1 pyramidal neurons in acute hippocampal slices prepared from Gria1-3f/fGrin1f/f mice that were electroporated in utero with Cre-mCherry plasmids at E14.5. GABAergic transmission was significantly impaired, indicating that proper development of GABAergic transmission in vivo requires AMPARs and/or NMDARs at the level of individual neurons (Figures 1C, S1F and S1G; Table S1). In addition, there was no change of paired pulse ratio, suggesting that GABA release probability is not altered (Figure S1H). Furthermore, immunocytochemical analysis demonstrated strong reductions of immunostaining of vGAT (vesicular GABA transporter) and gephyrin/neuroligin 2, the pre- and post-synaptic markers for GABAergic synapses, respectively, at both somatic and dendritic regions (Figures 1D, 1E and S1C), indicating that decrease of GABAergic transmission in Cre-expressing neurons represented a reduction of GABAergic synapse numbers. Finally, there was no change of GABA-evoked GABAAR-mediated whole-cell currents, indicating that functional expression of GABAARs on neuronal surface does not depend on glutamate receptors (Figure 1F). However, the GABAAR-mediated tonic currents were significantly increased, suggesting that extrasynaptic GABAARs are enhanced (Figure 1G). Taken together, these data show that the establishment of GABAergic transmission in immature neurons requires AMPARs and/or NMDARs.

Figure 1. GABAergic synapse development requires ionotropic glutamate receptors.

Figure 1

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(A) Reduced mIPSC frequency and amplitude in Cre-positive cultured neurons. Frequency: DIV6, control (Cnt), 100 ± 14.5, n = 16; Cre, 10.8 ± 3.4, n = 15; p < 0.001; DIV 15, Cnt, 100 ± 16.2; Cre, 24.1 ± 4.5; p < 0.001 and n =12 for each. Amplitude: DIV 6, Cnt, 100 ± 5.2 n =16; Cre, 81 ± 7.2, n = 15; DIV 15, Cnt, 100 ± 5.4; Cre, 79.6 ± 6.0 and n = 12 for each, p < 0.05) from Gria1-3f/fGrin1f/f. Scale bar represents 20 pA and 1 s.

(B) Sholl analysis of dendrites from control and Cre-positive cultured Gria1-3f/fGrin1f/f neurons (Cnt, n = 15; Cre, n = 13; for dendrite length, p = 0.97; for the number (#) of intersections, p = 0.92). Arrow indicates Cre-GFP in the nucleus. Scale bar represents 50 μm.

(C) Decreased mIPSC frequency in Cre-positive CA1 pyramidal neurons in acute hippocampal slices from P10 (frequency: Cnt, 100 ± 9.1, n = 11; Cre, 59.9 ± 5.7, n = 10; p < 0.01. amplitude: Cnt, 100 ± 3.1, n = 11; Cre, 102.7 ± 3.0, n = 10; p = 0.56) and P17 (frequency, Cnt, 100 ± 22.8, n = 7; Cre, 23.9 ± 9.8, n = 6; p < 0.05; amplitude, Cnt, 100 ± 4.2, n = 7; Cre, 105.0 ± 10.4, n = 6; p = 0.64) Gria1-3f/fGrin1f/f mice. Scale bar represents 20 pA and 1 s.

(D–E) Immunostaining of MAP2 (blue) and vGAT (D, green) or gephyrin (E, green) in hippocampal cultures, showing reduced gephyrin (dendrites: Cnt, 4.4 ± 0.6, n = 24; Cre, 1.0 ± 0.2, n = 26; p < 0.001; soma: Cnt, 26.3 ± 6.7, n = 6; Cre, 5.4 ± 0.92, n = 8; p < 0.01) and vGAT (dendrites: Cnt, 2.2 ± 0.3, n = 19; Cre, 0.73 ± 0.18, n = 17; p < 0.001; soma: Cnt, 16.8 ± 3.2, n = 10; Cre, 5.3 ± 2.1, n = 10, p < 0.01) puncta at both somatic regions (indicated by number 1) and dendritic regions (indicated by number 2) in Cre-positive cultured neurons. Note that Cre-mCherry was localized to neuronal nucleus (red). Scale bar represents 10 μm.

(F) There was no change of GABAAR-mediated GABA-evoked whole-cell currents (Cnt, −2374 ± 284 pA; Cre, −2129 ± 349 pA; n = 9 for each; p = 0.59). Scale bar represents 1, 000 pA and 2 s.

(G) Tonic inhibitory current in control and Cre-positive Gria1-3f/fGrin1f/f neurons at DIV 6–7 (Cnt, −3.8 ± 1.7 pA, n = 11; −11.7 ± 2.4 pA, Cre, n = 10, p < 0.05). Scale bar represents 10 pA and 10 s.

All data are presented as mean ± SEM. See also Figure S1A–S1H and Table S1.

Ca2+ influx through NMDARs is essential for development of GABAergic synapses

Electrophysiological analysis of DIV6 neurons in cultures showed that genetic deletion of AMPARs in Gria1-3f/f neurons had no effect on GABAergic mIPSCs (Figures 2A and S1I; Table S1). In contrast, strong reduction of mIPSC frequency was observed in Grin1f/f neurons expressing Cre (Figures 2B, and S1J; Table S1), suggesting that NMDARs, but not AMPARs, are important for the development of GABAergic transmission. Indeed, both vGAT and gephyrin immunostaining were reduced in Cre-positive Grin1f/f neurons (Figures S1K and S1L). In addition, the GABAAR-mediated tonic currents were significantly increased in Cre-positive Grin1f/f neurons (Figure S1M). Furthermore, mIPSC deficits in Gria1-3f/fGrin1f/f neurons expressing Cre were completely rescued by co-expression of GluN1-1a-IRES-GFP (hereafter referred to as GluN1-1a) (Figure 2C), one of the dominant GluN1 isoforms expressed in the brain (Laurie and Seeburg, 1994). In contrast, a GluN1-1a mutant at a channel pore residue (N616Q) that has been shown to dramatically reduce Ca2+ influx through the receptor (GluN1-1aN616Q) (Single et al., 2000) failed to fully rescue GABAergic transmission in Cre-expressing Gria1-3f/fGrin1f/f cells both in vitro and in vivo (Figures 2C, S2A, S2B and S3; Table S1), indicating that Ca2+ influx through NMDARs is important for GABAergic synapse development.

Figure 2. NMDARs are both necessary and sufficient for GABAergic synapse development in individual neurons.

Figure 2

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(A and B) Strong reduction of mIPSC frequency in cultured neurons lacking NMDARs (B, frequency: Cnt, 100 ± 23.9; Cre, 28.3 ± 16.9, p < 0.05; amplitude: Cnt, 100 ± 15.6; Cre, 89.0 ± 10.7, p = 0.57; n = 9 for both), but not in neurons lacking AMPARs (A, frequency: Cnt, 100 ± 25.3, n = 21; Cre, 106.8 ± 22.9, n = 18, p = 0.84; amplitude: Cnt, 100 ± 5.1, n = 21; Cre, 87.3 ± 5.1, n = 18, p = 0.09).

(C) Expression of GluN1-1a (frequency: 103.2 ± 16.8, p = 0.35; amplitude: 109.2 ± 5.6, p = 0.70; n = 25) rescued mIPSC deficits in Cre-positive cultured neurons from Gria1-3f/fGrin1f/f. In contrast, the GluN1 mutant with impaired Ca2+ permeability (GluN1-1aN616Q) failed to rescue mIPSC deficits (frequency, 28.0 ± 5.1, p < 0.001; amplitude, 104.1 ± 8.4, p = 0.68, n = 14).

(D and E) Overexpression of the GluN2A(N+1S) mutant increased GABAergic mIPSC frequency in cultured neurons (D, frequency: Cnt, 100 ± 22.1, n = 21; GluN2A(N+1S), 161.3 ± 20.1, n = 22; p < 0.05. amplitude: Cnt, 100 ± 8.0, n = 21; Cre, 103.8 ± 5.1, n = 22; p = 0.68), and in acute hippocampal slices (E, frequency: Cnt, 100 ± 22.1, n = 11; GluN2A(N+1S), 192.6 ± 40.9, n = 8; p < 0.05; amplitude, Cnt, 100 ± 6.7, n = 11; GluN2A(N+1S), 119.2 ± 16.2, n = 8, p = 0.24). Scale bar represents 20 pA and 1 s.

All data are presented as mean ± SEM. See also Figures S1I–S1M, S2 and S3 and Table S1

We also wondered whether molecular manipulations that enhanced NMDAR activity in individual neurons would promote GABAergic synapse development. To test this, we overexpressed a voltage-independent GluN2A mutant (GluN2A(N+1S)). NMDARs containing this mutant have reduced Mg2+ blockade, and thus are active at resting membrane potentials upon binding to agonists (Wollmuth et al., 1998) (Figures S2C and S2D). GABAergic transmission, but not GABA-evoked GABAAR-mediated whole-cell currents, was significantly enhanced in immature neurons expressing GluN2A(N+1S) (Figures 2D, 2E; Figures S2E, S2F; Table S1), suggesting that promoting NMDAR activity in individual neurons is sufficient to enhance GABAergic transmission.

The GluN1 C0 domain is required for inhibitory synapse development

The GluN1 Carboxyl-tail (C-tail) contains protein-protein interaction sites that are important for NMDAR-dependent signaling (Lau and Zukin, 2007). To investigate the role of the GluN1 C-tail, we expressed the GluN1-1aΔC mutant lacking the entire C-tail together with Cre. Although GluN1-1aΔC could rescue the majority of NMDAR-mediated synaptic currents (Figure S3), it failed to restore GABAergic transmission, indicating that the GluN1 C-tail is crucial for controlling GABAergic synaptic development by NMDARs (Figure 3B). Which domain(s) in the C-tail functions in GABAergic synapse formation? There are four alternative splice variants in the GluN1 C-tail (GluN1-1a, 2a, 3a, and 4a) (Figure 3A). Importantly, all splice isoforms restored NMDAR-mediated synaptic transmission in Gria1-3f/fGrin1f/f neurons that also expressed Cre (Figure S3). Interestingly, all four variants rescued GABAergic transmission (Figure 3A and Table S1), suggesting that the domain common to all isoforms, the C0 domain, is important. Indeed, deletion of the C0 domain (GluN1-1aΔC0), but not deletion of either C1 or C2 (or C2′) (i.e., GluN1-2a and GluN1-4a), abolished the rescue (Figures 3A and 3B). Furthermore, the GluN1 mutant that only contained the C0 domain in the C-tail (GluN1-1aΔC1C2) was sufficient to restore mIPSC deficits (Figure 3B). Taken together, these findings indicate that the GluN1 C0 domain plays an important role in regulating GABAergic synapse development.

Figure 3. The GluN1 C0 domain is required for GABAergic synapse development.

Figure 3

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(A) All four GluN1 splice isoforms in the C-tail rescued mIPSC deficits in Cre-positive cultured neurons from Gria1-3f/fGrin1f/f. Frequency: GluN1-1a, 103.2 ± 16.8, n = 25, p = 0.35; GluN1-2a, 95.5 ± 27.2, n = 12, p = 0.86; GluN1-3a, 103.8 ± 25.4 n = 11, p = 0.89; GluN1-4a, 78.7 ± 17.3, n = 16, p = 0.37. Amplitude: GluN1-1a, 109.2 ± 5.6, n = 25, p = 0.70; GluN1-2a, 97.6 ± 7.8, n = 12, p = 0.83; GluN1-3a,104.9 ± 12.3, n = 11, p = 0.67; GluN1-4a, 101.5 ± 8.8, n = 16, p = 0.88. Scale bar represents 20 pA and 1 s.

(B) The CaM-binding region in the GluN1 C0 domain is important for GABAergic synapse development. Frequency: GluN1-1aΔC, 31.2 ± 7.0, n = 14, p < 0.001; GluN1-1aΔC1C2, 104.1 ± 26.2, n = 14, p = 0.82; GluN1-1aΔC0, 31.1 ± 9.6, n = 11, p < 0.001; GluN1-1aΔ854–863, 20.0 ± 5.9, n = 11, p < 0.001; GluN1-1aQMQL/EEEE, 39.2 ± 9.3, n = 10, p < 0.01. Amplitude: GluN1-1aΔC, 105.5 ± 10.6, n = 14, p = 0.63; GluN1-1aΔC1C2, 123.8 ± 10.8, n = 14, p = 0.13; GluN1-1aΔC0, 109.6 ± 12.2, n = 11, p = 0.44; GluN1-1aΔ854–863, 88.8 ± 11.3, n = 11, p = 0.37; GluN1-1aQMQL/EEEE, 108.6 ± 8.6 n = 10, p = 0.50.

All data are presented as mean ± SEM. See also Figures S3 and S4A–D, and Table S1

Ca2+/CaM binding to the GluN1 C0 domain controls inhibitory synapse development

The GluN1 C0 domain contains a Ca2+-dependent Calmodulin (CaM) binding motif that is important for the regulation of NMDAR function (Ehlers et al., 1996; Krupp et al., 1999; Wyszynski et al., 1997; Zhang et al., 1998). We made a series of mutants to test a potential role of Ca2+-dependent interaction between the C0 domain and CaM in GABAergic synapse development. A GluN1-1a mutant (GluN1-1aΔ854–863) harboring a 10 residue-deletion in the C-terminus of the C0 domain that mediates the C0 domain interaction with both CaM and α-actinin 2 (Krupp et al., 1999; Wyszynski et al., 1997) failed to recover GABAergic synapse deficits in Gria1-3f/fGrin1f/f neurons expressing Cre (Figures 3B and S4A–S4C; Table S1). A more specific mutant in which four residues in the C0 domain (amino acids 847–850) were changed to glutamate (GluN1-1aQMQL-EEEE), disrupting the interaction with CaM (Figure S4D) (Zhang et al., 1998), was unable to restore mIPSC deficits (Figure 3B).

To further explore the role of CaM binding to the C0 domain in the development of GABAergic synapses, we replaced the last 24 residues in the C0 domain with a 24-amino acid domain from CalcineurinA1 that mediates CalcineurinA1 binding to CaM in a Ca2+-dependent manner (GluN1-1a(Calcineurin)) (Rumi-Masante et al., 2012) (Figure S4E and S4F). Significantly, co-expression of GluN1-1a(Calcineurin) and Cre in Gria1-3f/fGrin1f/f neurons fully rescued mIPSC deficits (Figure 4; Table S1). In contrast, a GluN1-1a mutant (GluN1-1a(MyosinV)) harboring a similar replacement in the C0 domain with the MyosinV CaM binding domain that binds to CaM in a Ca2+-independent manner (Martin and Bayley, 2004) (Figure S4E and S4F) did not rescue the deficits (Figure 4; Table S1). In addition, a GluN1-1a mutant (GluN1-1a(CRP1)) in which the last 24 residues in the C0 domain were replaced by 24 amino acids from a cysteine-rich protein 1 (CRP1) that mediates the interaction between CRP1 and α-actinin 2 (Harper et al., 2000) (Figure S4E–S4G) failed to rescue mIPSC deficits (Figure 4; Table S1). Importantly all of these GluN1 mutants could recover NMDA mEPSCs (Figure S3). Collectively these data support an important role for Ca2+-dependent binding of CaM to the GluN1 C0 domain in GABAergic synapse development.

Figure 4. Ca2+-dependent CaM binding to the C0 domain is critical for GABAergic synapse development.

Figure 4

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GluN1-1a mutants that harbored the Ca2+-dependent CaM binding motif, but not the Ca2+-independent CaM binding motif in the C0 domain region rescued mIPSC frequency deficits in Cre-expressing Gria1-3f/fGrin1f/f neurons. Frequency: GluN1-1a(Calcineurin), 115.8 ± 23.6, n = 14, p = 0.24; GluN1-1a(MyosinV), 32.7 ± 7.8, n = 10, p < 0.01; GluN1-1a(CRP1), 34.1 ± 9.0, n = 18, p < 0.01; Amplitude: GluN1-1a(Calcineurin), 112.0 ± 7.0, n = 14, p = 0.21; GluN1-1a(MyosinV), 101.9 ± 9.2, n = 10, p = 0.86; GluN1-1a(CRP1), 88.0 ± 5.8, n = 18, p = 0.37. Scale bar represents 20 pA and 1 s. All data are presented as mean ± SEM. See also Figures S3 and S4E–G, and Table S1

Discussion

Our data demonstrate that single-cell genetic deletion of NMDARs, but not AMPARs, in embryonic hippocampal neurons leads to a strong reduction of GABAergic transmission. On the other hand, promoting NMDAR activity in single postsynaptic neurons is sufficient to enhance GABAergic synaptic transmission. Mechanistically, NMDAR-dependent GABAergic synapse development requires Ca2+ influx through the receptors and the C0 domain in the carboxyl-terminus of the NMDAR GluN1 subunit. More importantly, our data indicate that calmodulin acts as the NMDA receptor downstream molecule for the regulation of GABAergic synapse development. Thus, in addition to the well-established role of NMDARs in synaptic plasticity at glutamatergic synapses (Malenka and Bear, 2004), we have now demonstrated that the NMDAR acts as an essential signaling molecule for controlling GABAergic synapse development in immature neurons.

It is worth noting that when the number of GABAergic synapses decreases in neurons lacking NMDARs, the extrasynaptic GABAAR-mediated tonic inhibition increases (Figures 1G and S1M). It is possible that increased tonic inhibition may serve as a compensatory mechanism for the lack of synaptic phasic inhibition. Thus, in addition to its role in GABAergic synapse development, the NMDAR may play a role in the regulation of the balance of tonic and phasic inhibition in neurons.

It is well-established that GABA acts as an excitatory neurotransmitter in developing brain (Ben-Ari et al., 2007). Indeed, it has been shown that activation of GABAARs on neuronal surface by ambient GABA in immature neurons provides membrane depolarization necessary for NMDAR activation (Ben-Ari et al., 2007). NMDARs also have a higher affinity for glutamate (Patneau and Mayer, 1990), and thus ambient glutamate and depolarization provided by GABA action in developing brain have been shown to trigger tonic activation of NMDARs before glutamatergic synaptogenesis (Ben-Ari et al., 2007; LoTurco et al., 1991). Thus, it is likely that in early developing neurons excitatory activity of GABAARs on neuronal surface facilitates NMDAR activation, which in turn, through signaling via the CaM binding motif in the C0 domain of the NMDAR GluN1 subunit, regulates GABAergic synapse development. Interestingly, it has been reported that in developing neurons NMDARs strongly colocalize with GABAARs at GABAergic synapses, providing neuroanatomical evidence for the regulation of GABAergic synapse development by NMDARs (Cserep et al., 2012; Gundersen et al., 2004; Szabadits et al., 2011). In addition, the mechanism for the NMDAR-dependent GABAergic synapse development is difficult to interpret through a homeostatic or compensatory process. This is because activities of both GABAARs and NMDARs are depolarizing in immature neurons (Ben-Ari et al., 2007), and because several GluN1 mutants restored NMDAR-mediated synaptic transmission, but failed to rescue GABAergic mIPSCs (Figures 2 and 3). Finally, although synaptogenic adhesion molecules, transcription factors and signaling molecules have been identified that play important roles in inhibitory synapse development (Bloodgood et al., 2013; Fazzari et al., 2010; Gottmann et al., 2009; Huang and Scheiffele, 2008; Kuzirian et al., 2013; Lin et al., 2008; Siddiqui and Craig, 2011; Sudhof, 2008; Takahashi et al., 2012; Terauchi et al., 2010; Woo et al., 2013; Yim et al., 2013), possible functional interaction between NMDAR signaling and these molecules in the regulation of GABAergic synapse development remains unclear. Recent studies in mature neurons show that NMDARs can regulate GABAergic synapses through calcineurin- (Bannai et al., 2009; Muir et al., 2010), CaMKII- (Flores et al., 2015; Marsden et al., 2010; Petrini et al., 2014) or nitric oxide synthase-dependent mechanisms (Nugent et al., 2007). It would be interesting to examine the role of these signaling pathways in NMDAR-dependent GABAergic synapse development in early immature neurons in the future.

Experimental Procedures

Mouse genetics

Animal housing was conducted according to the ACUC guidelines at NINDS, NIH. Gria1-3f/fGrin1f/f, Gria1-3f/f and Grin1f/f mice were generated as described previously (Lu et al., 2013; Lu et al., 2011; Lu et al., 2009; Tsien et al., 1996).

Plasmids

pCAGGS-GluN1-1a-IRES-GFP plasmid was a gift from Roger Nicoll’s lab at UCSF. pCAGGS-GluN2A-IRES-GFP vector was a gift from Katherine Roche’s lab at NINDS, NIH. cDNAs encoding mouse GluN1-2a, -3a, -4a, all GluN1-1a mutants and the GluN2A mutant were generated by overlapping PCR and cloned into the pCAGGS-IRES-GFP/mCherry expression plasmid.

Antibodies

The following primary antibodies were used: anti-vGAT, anti-Gephyrin, anti-MAP2, anti-Myc, anti-Calmodulin and anti-GST.

In Utero electroporation and neuronal culture

In Utero electroporation was performed to introduce the plasmids into neuronal progenitors as described in Supplemental Experimental Procedures. Hippocampi from E17.5–E18.5 (three days after in utero electroporation) mouse embryos were dissected and cultured on coverslips.

Electrophysiology

Electrophysiological recordings were performed in dissociated hippocampal neuronal cultures and in 300 μm acute hippocampal slices. Recordings were performed artificial cerebrospinal fluid containing (in mM) NaCl 119, KCl 2.5, NaHCO3 26, Na2PO4 1, glucose 11, CaCl2 2.5, MgCl2 1.3, and recovered at 32°C with appropriate drugs. The intracellular solution for GABA IPSC recording contained (in mM) CsMeSO4 70, CsCl 70, NaCl 8, EGTA 0.3, HEPES 20, MgATP 4 and Na3GTP 0.3. The intracellular solution for AMPA and NMDA EPSC recording contained (in mM) CsMeSO4 135, NaCl 8, HEPEs 10, Na3GTP 0.3, MgATP 4, EGTA 0.3, QX-314 5, and spermine 0.1. Osmolality was adjusted to 285–290 mOsm and pH was buffered at 7.25–7.35.

Immunocytochemistry

Hippocampal primary dissociated cultures at DIV 6–7 were fixed, permeabilized and incubated with indicated antibodies. Neurons were then mounted and imaged under a Zeiss LSM 510 confocal microscope. For puncta analysis, images from 3–5 dendrites (35 μm in length) per neuron from at least ten neurons per experiment were collected and quantified by counting the number of puncta per 10 μm dendrites or per soma with ImageJ software (NIH).

Statistics

All data were presented as mean ± s.e.m (standard error of mean). Direct comparisons between two groups were made using two-tailed Student’s t-test with Welch’s correction when the standard deviation (SD) is significantly different. Multiple group comparisons were made using one-way analysis of variance (ANOVA) with post hoc Fisher’s LSD test. For the Sholl analysis, two-way ANOVA analysis was performed. The significance of the shift in cumulative probability distributions was assessed by Kolmogorov-Smirnov test. Statistical significance was defined as p < 0.05, 0.01 or 0.001 (indicated as *, ** or ***, respectively). p values ≥0.05 were considered not significant.

Supplementary Material

Supplemental data

Acknowledgments

We appreciate Xia Mao for mouse colony management. We are grateful to Chris McBain, Jeffrey Diamond, Katherine Roche, Claire Le Pichon, Mary Anne Hutchison and Jun Li for critical comments on the manuscript. The authors also thank Cindy Clark, NIH Library Writing Center, for manuscript editing assistance. This research was supported by the Intramural Research Program of NIH, NINDS (W.L).

Footnotes

Author Contributions X.G., L.Z., and W.L. designed and performed whole-cell electrophysiological experiments. X.G. performed immunocytochemical assays. L.Z. performed biochemical assays. W.L. wrote the manuscript and all authors read and commented on the manuscript.

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