The conditional allele of Grin1 (Grin1^fl, JAX #018825, The Jackson Laboratory, Bar Harbor, ME, United States) (Zhang et al., 2013), SERT-Cre [Tg(Slc6a4-cre)ET124Gsat, Mutant Mouse Resource and Research Center (MMRRC)] (Gong et al., 2007), Rosa26-tdTomato reporter (Rosa^Ai14, JAX #007914, The Jackson Laboratory) (Madisen et al., 2010), conditional allele of Mtor (Mtor^fl, JAX #011009, The Jackson Laboratory) (Risson et al., 2009), Gria3 knockout (B6-129P2-Gria3^tm1-Dgen, MMRRC), and Gria4 knockout (B6-129P2-Gria4^tm1-Dgen, MMRRC) were housed in the Research Facility at The Jackson Laboratory. All the strains were on a C57BL/6J background. Both male and female mice were used in the experiments. Genotyping were determined by PCR using tail genomic DNA and allele-specific primers. For the SERT-Cre strain, the primers were forward 5′-CAACAGAGCTCTCAGTCTTGTC-3′ and reverse 5′-TGGTGTACGGTCAGTAAATTGG-3′. For Gria3 knockout strain, the primers were wild type forward 5′-GGGTGGGATTAGATAAATGCCTGCTCT-3′, mutant forward 5′-GGTCACGAGGTTCTTCATTGTTGTC-3′, and common reverse 5′-AGCTGATATAGCTGTTGCTCCACTC-3′. For Gria4 knockout strain, the primers were forward 5′-TGTCACAGCAAAACTGTTGGCAGT-3′, and reverse 5′-GGGTGG GATTAGATAAATGCCGCTC-3′. Primer sequences for all the other strains can be found at the website of The Jackson Laboratory. The Jackson Laboratory Animal Care and Use Committee approved all animal protocols.

Mice were anesthetized with tribromoethanol and decapitated. Brains were dissected quickly and sagittal sections 300 μm thick were prepared with a vibratome (Leica VT1200) and kept in artificial cerebral spinal fluid (ACSF) containing (in mM): 124 NaCl, 3.0 KCl, 1.5 CaCl₂, 1.3 MgCl₂, 1.0 NaH₂PO₄, 26 NaHCO₃, and 20 glucose, saturated with 95% O₂ and 5% CO₂ at room temperature (21–23°C). We used 1.5 mM Ca²⁺ in ACSF because a previous study showed that the concentration of Ca²⁺ in the cerebral spinal of young rats is about 1.5 mM (Jones and Keep, 1988).

For patch clamp recording, a slice was transferred to a submersion type chamber where it was continuously perfused with ACSF saturated with 95% O₂ and 5% CO₂ at 30–32°C. Neurons in the VPm were viewed with a 40x objective and Cre⁺ neurons were identified by epifluorescence of tdTomato. Whole-cell recordings were obtained from the soma with a Multiclamp 700B amplifier (Molecular Devices, Sunnyvale, CA, United States). The pipette solution contained (in mM): 120 K-gluconate, 10 KCl, 4 ATP-Mg, 0.3 GTP-Na, 0.5 EGTA, and 10 HEPES (pH 7.2, 270–280 mOsm with sucrose). For current clamp recording, the series resistance (R_s) was fully compensated. For voltage clamp recording, R_s, usually between 8 and 12 Mω, was continuously monitored but not compensated. Data was discarded when R_s changed by more than 30% during the experiment. Experiments were conducted using AxoGraph X (AxoGraph Scientific). Data were filtered at 4 kHz and digitized at 20 kHz. Data analysis was performed using AxoGraph X.

Intrinsic properties of neurons were analyzed using methods described previously (Zhang, 2004; Zhang and Arsenault, 2005). Resting membrane potentials were measured within 20 s of break in. Input resistance and time constant were calculated from voltage responses to 600-ms current steps of -100 pA. For the analyses of AP rheobase and F-I curves, current steps of 1 s in duration were applied once every 10 s, with increments of 10 or 20 pA. To minimize the effect of resting potential variation on rheobase and F-I curve, cells were held at -60 mV between current steps. In some experiments, synaptic blockers picrotoxin, kynurenic acid, and 6,7-Dinitroquinoxaline-2,3-dione (DNQX; all from Tocris) were applied through bath perfusion.

Voltage-dependency of I_h activation was analyzed using a voltage-clamp protocol similar to that described in a previous study (Kanyshkova et al., 2009). This protocol takes into consideration voltage-dependent changes in the onset rate of I_h. Initially held at -50 mV, the membrane potential of the cell was stepped to increasingly hyperpolarizing levels for several seconds to reach steady-state activation, then to -100 mV. The activation curve of I_h was established by measuring the tail currents at 40 ms after stepping to -100 mV. The normalized I_h, p(V), was calculated using the equation:

where I_min and I_max are the amplitudes of tail currents for pre-pulses at -50 and -150 mV, respectively. The activation curves of I_h were fitted to a Boltzmann equation:

where V_h is the membrane potential of half-maximal activation, and k is the slope factor.

Recordings of EPSCs at VPm relay synapses were performed as described previously (Arsenault and Zhang, 2006; Wang and Zhang, 2008). The pipette solution contained (in mM): 110 Cs methylsulfate, 20 TEA-Cl, 15 CsCl, 4 ATP-Mg, 0.3 GTP, 0.5 EGTA, 10 HEPES, 4.0 QX-314 and 1.0 spermine (pH 7.2, 270–280 mmol kg^-1 with sucrose). A concentric bipolar electrode (FHC, Bowdoin, ME, United States) was placed in the medial lemniscus, and stimuli were applied at 0.1 Hz. GABAergic transmission was blocked by picrotoxin (100 μM) in the bath.

Neurons were labeled during whole-cell recording with 0.3% biocytin in the patch pipette. Slices were fixed in 4% paraformaldehyde in 0.1 M phosphate buffer (PB) overnight and processed as described previously (Zhang and Deschenes, 1997; Zhang, 2004). Briefly, slices were treated with 0.3% H₂O₂ in 10% MeOH in 0.1 M PB to inactivate endogenous peroxidase. After wash, slices were processed for biocytin histochemistry using the ABC Elite Standard Kit (Vector Laboratories, Burlingame, CA, United States), 3,3-diaminobenzidine, and nickel ammonium sulfate. Cells were reconstructed and analyzed with the Neurolucida System (MicroBrightfield, Williston, VT, United States).

Mice were anesthetized with tribromoethanol and perfused with 4% paraformaldehyde in 0.1 M PB. Brains were post fixed over night. Sagittal sections of 70 μm thickness were cut on a vibratome. Sections were incubated with primary antibodies for 48 h at 4°C. The primary antibodies were NeuN (Millipore, MAB377, mouse, 1:1000), p-S6 (Ser240/244) (Cell Signaling, Beverly, MA, United States #5364, rabbit, 1:1000), p-Akt (Thr308) (Cell Signaling, #2965, rabbit, 1:1000), and RFP (Rockland, Limerick, PA, United States #200-301-379, mouse, 1:1000). For p-Akt staining, sections were pretreated with pepsin (Sigma, 0.5 mg/ml in 0.2 N HCl) for 5 min at 37°C before incubation with primary antibody. Secondary antibodies were conjugated with Alexa fluor (Life Technologies, 1:500). Sections were mounted in ProLong Diamond (Life Technologies). Confocal stacks were taken with a 63x (NA 1.4) objective on a Leica SP5 microscope. The pixel size was 80.2 nm. The z-step size was 130 nm and each stack had 10 steps.

Data were analyzed using ImageJ. Changes in cell body size were quantified by measuring soma area. Confocal stacks of thalamic neurons immunostained for NeuN were z-projected at maximum intensity. Cre⁺ neurons were identified by the expression of the reporter protein tdTomato. Cell bodies of individual neurons were outlined using the polygon selection tool and soma area was analyzed. The same method was used to quantify immunofluorescence of p-Akt and p-S6. The mean fluorescence intensity of immunofluorescence staining was measured for the soma.

Statistical analyses were performed with JMP software (SAS, Cary, NC, United States). Data were presented either as median with interquartile range and the minimum and maximum (box and whisker) or as mean ± SEM. Statistical differences were determined using non-parametric Wilcoxon Rank Sum Test (two groups) or Krustal–Wallis Test (three or more groups). P-values were provided in the results or figure legends.

N-methyl-D-aspartate receptors are ubiquitously expressed in the brain during development. Pharmacological block or genetic deletion of NMDARs from a large number of neurons disrupts activities of neuronal circuits. To distinguish cell-autonomous vs. non-autonomous effects, we performed mosaic deletion of NMDARs in the thalamus of newborn mice by using the conditional allele of the NMDAR subunit GluN1 (Grin1^fl) (Zhang et al., 2013) and the transgenic serotonin transporter Cre strain (SERT-Cre) (Gong et al., 2007; Zhang et al., 2013). Our previous study showed that in SERT-Cre; Grin1^fl/fl mice, about 50% of VPm neurons express Cre and lack NMDARs, and the loss of NMDAR function occurs between P7 and P10 (Zhang et al., 2013). The expression of Cre in VPm neurons was random. To identify neurons with or without Cre expression, we used the Rosa26-tdTomato reporter (Rosa^Ai14) (Madisen et al., 2010) to express the fluorescent marker tdTomato in Cre⁺ neurons (Figure 1A). We first recorded VPm neurons in acute slices obtained from SERT-Cre; Grin1^fl/fl; Rosa^Ai14 mice at P14-16, the age at which VPm neurons reach maturity in synaptic connectivity (Arsenault and Zhang, 2006; Wang et al., 2011). We applied NMDA to the recorded neurons to confirm the absence or presence of NMDARs (Figures 1B,C). Intrinsic excitability was analyzed with whole-cell patch clamp recording.

Deletion of NMDARs dramatically altered excitability of VPm neurons at P14-16 (Figures 1D,E). Compared with neighboring control neurons with NMDARs (Grin1^fl/fl; Cre^-), mutant neurons deficient of NMDARs (Grin1^fl/fl; Cre⁺) had lower rheobases for firing action potential (AP), and steeper slopes of the frequency to current (F-I) curves (Figures 1F,G). Mutant neurons showed a small reduction in AP amplitude (Figure 1H; 67.5 ± 1.1, n = 22 for mutant vs. 71.4 ± 1.0 mV, n = 27 for control; ^∗∗p = 0.005, Wilcoxon test) and a small increase in AP half width (0.42 ± 0.01 vs. 0.37 ± 0.01 ms; ^∗∗p = 0.002, Wilcoxon test), but no change in AP rise time (0.13 ± 0.01 ms vs. 0.12 ± 0.01 ms; p = 0.12, Wilcoxon test). There was no difference between mutant and control neurons in after hyperpolarization (-9.2 ± 0.5 mV for mutant vs. -9.7 ± 0.4 mV for control neurons, p = 0.31, Wilcoxon test). There was no significant difference between mutant and control neurons in resting membrane potential, or AP threshold (Figures 1I,J). Mutant neurons had significantly higher input resistance and lower whole-cell capacitance than control neurons (Figures 1K,L). Mutant neurons showed a small reduction in the time constant (14.0 ± 0.1 ms, n = 22 for mutant vs. 16.3 ± 0.1 ms, n = 27 for control; ^∗p = 0.012, Wilcoxon test).

Previous studies have shown that SERT is transiently expressed in all VPm neurons during early life (Lebrand et al., 1998; Narboux-Neme et al., 2008), so it is unclear why only a fraction of VPm neurons in the SERT-Cre strain express Cre. To exclude the possibility that SERT-Cre is expressed in a subpopulation of VPm neurons with distinct intrinsic excitability, we recorded Cre⁺ neurons in the VPm of SERT-Cre; Grin1^fl/+; Rosa^Ai14 mice at P14-16. All Cre⁺ neurons (Grin1^fl/+; Cre⁺) in these Grin1 heterozygous mice had normal NMDAR function. We did not find any difference in intrinsic excitability between Grin1^fl/+; Cre⁺ neurons (Figures 1F,G,I–L, in green) and Grin1^fl/fl; Cre^- neurons, indicating that in the presence of NMDARs, Cre⁺ and Cre^- neurons are indistinguishable in membrane property and excitability. Together, these findings suggest that the deletion of NMDARs alters excitability of VPm neurons through cell-autonomous mechanisms.

The experiments described above were conducted in the absence of any synaptic blockers. Deletion of NMDARs may alter spontaneous synaptic responses, which in turn, affect neuronal excitability. To examine this possibility, we performed recordings in the presence of glutamatergic blockers DNQX and kynurenic acid, and the GABAergic blocker picrotoxin. The differences in excitability between mutant and control neurons persisted in the presence of these synaptic blockers (Supplementary Figures S1A–E). These results show that the effects of NMDAR deletion on excitability in thalamic neurons are independent of spontaneous synaptic transmission.

Thalamic relay neurons fire bursts of APs when stimulated from hyperpolarized membrane potentials. We analyzed rebound bursts following hyperpolarizing current steps. Both mutant and control neurons showed rebound bursts when hyperpolarized to -70 mV (Figure 2A). There was no difference in the number of APs in a burst between mutant (3.1 ± 0.3, n = 11 cells) and control (2.8 ± 0.3, n = 12 cells; p = 0.39, Wilcoxon test). However, the average firing frequency was significantly reduced in mutant neurons (Figure 2B).

Deletion of NMDARs may cause a delay in the maturation of neuronal excitability. To examine this possibility, we analyzed VPm neurons at P10-11 and P20-21. The effects of NMDAR deletion on excitability and membrane properties were already present at P10-11. Neurons deficient of NMDARs showed a decrease in rheobase (Figure 3A) and an increase in the slope of the F-I curve (Figure 3B), together with an increase in input resistance (Figure 3C) and a decrease in whole-cell capacitance (Figure 3D). These effects are comparable to those observed at P14-16 and P20-21 (Figures 3A–D). These results suggest that NMDARs are required for postnatal development of intrinsic excitability in thalamic neurons.

Our previous study showed that deletion of NMDARs down-regulates synaptic AMPARs in VPm neurons (Zhang et al., 2013). The amplitudes of AMPAR-mediated EPSCs were reduced by 80% in VPm neurons deficient of NMDARs. To determine whether the effects of NMDAR deletion on intrinsic excitability are mediated by changes in AMPAR function, we used mice deficient of GluA3 (Gria3) and GluA4 (Gria4), two major AMPAR subunits for the thalamus. We have previously shown that deletion of both Gria3 and Gria4 reduces AMPAR-mediated EPSCs by 80% in VPm neurons without any changes in NMDAR-mediated EPSCs (Wang et al., 2011). Consistent with our previous finding, AMPAR-mediated EPSCs were dramatically reduced in VPm neurons from mice deficient of GluA3 and GluA4 (Figure 4A). However, intrinsic excitability of VPm neurons was not affected by the deletion of both GluA3 and GluA4 (Figure 4B). There was no difference between mutant and control neurons in the rheobase, slope of the F-I curve, input resistance, or whole-cell capacitance (Figures 4C–F). These findings suggest that the effects of NMDAR deletion on intrinsic excitability are not caused by the reduction in AMPAR functions.

Neuronal morphology is an important determinant of cellular excitability. The reduction of whole-cell capacitance in NMDAR-deficient neurons indicates a decrease in cell size. We further examined the effect of NMDAR deletion on cell body size at P16-18. NeuN staining was used to label neuronal somata and soma area was quantified. In the VPm of mosaic mutant mice (SERT-Cre; Grin1^fl/fl; Rosa^Ai14), Cre⁺ mutant neurons (Figure 5A, left panel, cell body encircled by while dash line) had smaller soma size than neighboring Cre^- control neurons (cell body encircled by yellow dash line) (Figure 5B, red vs. gray). To determine whether SERT-Cre expression by itself may affect soma size, we also analyzed soma area in the VPm of heterozygous control mice (SERT-Cre; Grin1^fl/+; Rosa^Ai14) at P16-18 (Figure 5A, right panel). There was no difference in soma area between Cre⁺ and Cre^- in Grin1 heterozygous mice (Figure 5B, green vs. blue). Importantly, there was no difference between control neurons (Cre^-) in Grin1 homozygous mice and neurons (Cre⁺ or Cre^-) in Grin1 heterozygous mice (Figure 5B). These results demonstrate that NMDARs regulate cell size in the thalamus through cell-autonomous mechanisms.

To determine the effects of NMDAR deletion on dendritic morphology, we labeled VPm neurons from SERT-Cre; Grin1^fl/fl; Rosa^Ai14 mice at P14-16 with biocytin in the whole-cell recording pipettes. Dendritic branches of individual VPm neurons were reconstructed and analyzed (Figure 5C). The total dendritic length was significantly shorter in mutant neurons (Figure 5D). Mutant neurons also had few dendritic branches than control neurons (Figure 5E). We performed Sholl analysis of dendritic morphology. The distribution of Sholl intersections was significantly different between mutant and control neurons (Figure 5F). The majority of Sholl intersections were much closer to the cell body for mutant than control neurons. These results suggest that NMDAR signaling regulates the growth of dendritic branches of thalamic neurons.

Hyperpolarization-activated cation channels have important functions in the regulation of neuronal excitability and these channels are up-regulated during early life in many parts of the brain including the thalamus (Kanyshkova et al., 2009). We first analyzed voltage sag generated by I_h in VPm neurons. Control and mutant neurons were hyperpolarized by current steps from -60 to -75 mV (Figure 6A). Mutant neurons had smaller voltage sag than control neurons (Figures 6A,B), indicating a down-regulation of HCN channel function in mutant neurons.

Next, we analyzed I_h in VPm neurons using voltage clamp recording. Compared with control neurons, mutant neurons had much smaller I_h (Figures 6C,D). In contrast, Grin1 deletion did not significantly affect the kinetics of I_h activation (720.5 ± 28.5 ms, n = 20 cells for mutant vs. 651 ± 27.8 ms, n = 19 cells for control, p = 0.074, Wilcoxon test) or inactivation (240.0 ± 5.8 ms vs. 232.2 ± 6.0 ms, p = 0.49, Wilcoxon test). To determine whether NMDAR deletion alters voltage dependency of HCN channels, we analyzed the current-voltage (I-V) curve for steady-state I_h using tail current measurement (Figure 6E). The I-V relationship of steady-state I_h was not significantly different between mutant and control neurons (Figure 6F). The density of I_h, estimated using the maximal tail current, was significantly reduced in mutant neurons (7.9 ± 0.5 pA/pF, n = 17 cells for mutant vs. 9.7 ± 0.5 pA/pF, n = 15 cells for control, ^∗p = 0.034, Wilcoxon test). Together, these results suggest that deletion of NMDARs reduces the number of HCN channels in thalamic neurons.

To determine whether HCN channels are implicated for the changes in intrinsic excitability in NMDAR-deficient neurons, we examined the effects of blocking HCN channels on excitability of VPm neurons in wild type (WT) mice. Bath application of the specific I_h blocker ZD 7288 (20 mM) significantly enhanced intrinsic excitability of these cells (Figure 6G). Similar to the effects of NMDAR deletion, the blockade of I_h reduced the rheobase of spiking and increased the slope of the F-I curve of WT neurons (Figures 6H,I). In the presence of ZD7288, the rheobase and slope of the F-I curve of WT neurons were comparable to those of GluN1-deficient neurons (0.562 ± 0.037 vs. 0.607 ± 0.048 Hz/pA for F-I slope, p = 0.76, Wilcoxon test; 69.2 ± 4.8 vs. 61.6 ± 8.4 pA for rheobase, p = 0.24, Wilcoxon test). The blockade of I_h in WT neurons also caused an increase in input resistance (109.5 ± 6.9 vs. 172.6 ± 8.2 MW, n = 14 cells; ^∗∗∗p < 0.001, Wilcoxon test). These results suggest that NMDAR signaling regulates intrinsic excitability of thalamic neurons at least partially through I_h.

Deletion of NMDARs disrupts many molecular pathways. Because of the reduction in cell size, we hypothesized that the mTOR pathway, known to play a critical role in cell growth (Hay and Sonenberg, 2004), is implicated in NMDAR-mediated regulation of intrinsic excitability. The PI3K/Akt pathway is upstream of mTOR activation. We first analyzed Akt phosphorylation in the VPm of SERT-Cre; Grin1^fl/fl; Rosa^Ai14 mice by measuring immunofluorescence of phosphorylated Akt (p-Akt) in single cells. We found that mutant neurons had significantly lower p-Akt signal than neighboring control neurons (Figures 7A,B). This result indicates that deletion of NMDARs down-regulated Akt function in thalamic neurons.

A key downstream effector of the mTOR signaling is the ribosomal protein S6. Activation of mTOR phosphorylates S6 through the S6 kinase (Hay and Sonenberg, 2004). We analyzed S6 phosphorylation in the VPm of SERT-Cre; Grin1^fl/fl; Rosa^Ai14 mice with immunofluorescence using an antibody against phosphorylated S6 (p-S6). Mutant neurons had much lower p-S6 signal than neighboring control neurons (Figures 7C,D). Together, our results show that deletion of NMDARs down-regulates Akt/mTOR signaling in VPm neurons.

Next, we examined the effects of Mtor deletion on intrinsic excitability. We used the SERT-Cre and the conditional Mtor allele (Risson et al., 2009) to perform mosaic deletion of Mtor in the VPm, and analyzed intrinsic properties of mutant (Mtor^fl/fl; Cre⁺) and control neurons (Mtor^fl/fl; Cre^-) in SERT-Cre; Mtor^fl/fl; Rosa^Ai14 mice aged P14-16. Deletion of Mtor mimicked the effects of NMDAR deletion in VPm neurons (Figures 8A–J). Mutant neurons showed hyperexcitability with lower rheobase (Figure 8C), steeper slope of the F-I curve (Figure 8D), higher input resistance (Figure 8E), and lower whole-cell capacitance (Figure 8F). Mutant neurons deficient also exhibited a more depolarized resting potential than control neuron (Figure 8G) but no change in AP threshold (Figure 8H). As in the case of NMDAR-deficient neurons, neurons deficient in mTOR had smaller I_h (Figure 8I), but no change in the I-V relationship of I_h activation (Figure 8J). We also analyzed changes in cell size in fixed sections from SERT-Cre; Mtor^fl/fl; Rosa^Ai14 mice at P17. Soma areas of mutant neurons were significantly smaller than those of control neurons (196.9 ± 2.9 μm², n = 103 vs. 249.6 ± 5.6 μm², n = 66; ^∗∗∗∗p < 0.0001, Wilcoxon test). To examine the possibility that the deletion of Mtor down-regulates NMDARs, we analyzed NMDAR-mediated EPSCs in VPm neurons in SERT-Cre; Mtor^fl/fl; Rosa^Ai14 mice aged P15-16. There was no difference between mutant and control neurons in NMDAR-mediated EPSCs (Supplementary Figure S2), indicating that the deletion of Mtor does not alter NMDAR function in these cells. Together, these results support the role of mTOR signaling in NMDAR-mediated regulation of intrinsic excitability.