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CaV3.2 calcium channels control NMDA receptor-mediated transmission: a new mechanism for absence epilepsy

Wang et al. show that CaV3.2 channels control NMDA-sensitive glutamatergic receptor (R)-mediated transmission and subsequent NMDA-R-dependent plasticity of AMPA-R-mediated transmission at rat central synapses. Human childhood absence epilepsy (CAE)-linked hCaV3.2(C456S) mutant channels have a higher channel open probability, induce more calcium influx, and enhance glutamatergic transmission.

absence epilepsy (CAE) (Chen et al. 2003;Liang et al. 2006;Heron et al. 2007), which is the most common form of pediatric epilepsy (Snead 1995;Tenney and Glauser 2013). More perplexingly, ∼50% of CAE patients did not respond to ethosuximide, a T-type calcium channel antagonist and first-line drug used to treat CAE (Coulter et al. 1989;Huguenard and Prince 1994), and the majority of the nonresponsive patients carry Ca V 3.2 channel mutations (Glauser et al. 2010). These findings establish the functional significance of Ca V 3.2 channels but raise the fundamental questions of what the primary function of Ca V 3.2 channels is and how CAE-linked Ca V 3.2 mutations may be epileptogenic.
At central synapses, NMDA transmission plays a decisive role in controlling AMPA transmission strength (Kessels and Malinow 2009; Stornetta and Zhu 2011;Huganir and Nicoll 2013). It is also clear that NMDA transmission strength itself is regulated at synapses, albeit the detailed mechanisms and dynamics of this regulation are much less explored (Lau and Zukin 2007;Hunt and Castillo 2012;Paoletti et al. 2013). Since synaptic NMDA transmission is central to fundamental cognitive functions, including sensory perception and behavior adaption, it is not surprising to see that dysregulation of NMDA transmission can lead to various neurological, mental, and psychiatric disorders, including addiction, Alzheimer's disease, autism, depression, pathological pain, and schizophrenia (Lau and Zukin 2007;Russo et al. 2010;Hunt and Castillo 2012;Paoletti et al. 2013;Monteggia and Zarate 2015). However, what controls NMDA transmission strength remains unknown.
We report here an investigation of regulation and function of Ca V 3.2 channels in multiple distinct types of rat central neurons in intact circuits and intact brains. Using simultaneous multiple patch clamp recordings and/or multiple two-photon imaging techniques (Wang et al. 2015), we found that, in sharp contrast to the other Ttype calcium channels (i.e., Ca V 3.1 and Ca V 3.3 channels), Ca V 3.2 channels did not contribute to either modulation of membrane properties or production of lowthreshold calcium spikes in central neurons. Instead, functional Ca V 3.2 channels primarily incorporated into synapses by replacing existing synaptic Ca V 3.2 channels and served to control the strength of NMDA transmission. This Ca V 3.2 channel-dependent regulation of NMDA transmission required synaptic activity, activation of Ca V 3.2 channels, and calcium influx. Consistent with these findings, expression of human CAE-linked mutant hCa V 3.2(C456S) channels in rats resulted in replacement of endogenous Ca V 3.2 channels with higher open probability mutant channels that led to 2-to 4-Hz spike and wave discharges (SWDs) and absence-like epilepsy characteristic of CAE patients. The SWDs and absence-like epilepsy were suppressed by AMPA receptor (AMPA-R) and NMDA-R antagonists but not T-type calcium channel antagonists. These results reveal a surprising role of Ca V 3.2 channels in regulation of synaptic NMDA transmission strength and establish the first genetic model for CAE patients carrying Ca V 3.2 channel mutations.

Ca V 3.2 channels regulate glutamatergic transmission at various synapses
To examine whether expression of Ca V 3.2 channels affect synaptic transmission in intact brains, we virally expressed hCa V 3.2(WT)-GFP, hCa V 3.2(C456S)-GFP, or Ca V 3.2ct-GFP in L5 pyramidal neurons and TRN neurons in vivo for ∼18 h and then measured the effects on the synaptic and intrinsic responses of these neurons in acutely prepared slices ( Fig. 2A). Compared with the nearby control nonexpressing neurons, L5 pyramidal neurons expressing hCa V 3.2(C456S)-GFP had enhanced AMPA and NMDA responses, whereas L5 pyramidal neurons expressing Ca V 3.2ct-GFP had reduced AMPA and NMDA responses ( Fig. 2B,C). Expression of hCa V 3.2(WT)-GFP in L5 pyramidal neurons had no effect on AMPA and NMDA responses (Fig. 2C). Similar to L5 pyramidal neurons, compared with the nearby nonexpressing neurons, TRN neurons expressing hCa V 3.2(C456S)-GFP had enhanced AMPA and NMDA responses, whereas TRN neurons expressing Ca V 3.2ct-GFP had reduced AMPA and NMDA responses (Fig. 2D,E). Expression of hCa V 3.2(WT)-GFP in TRN neurons had no effect on AMPA and NMDA responses (Fig. 2E). L5 pyramidal neurons expressing hCa V 3.2(WT)-GFP, hCa V 3.2(C456S)-GFP, or Ca V 3.2ct-GFP in vivo for ∼18 h had the same membrane properties, including RMP, input resistance, membrane time constant, and average spiking frequency, compared with nearby nonexpressing neurons (Supplemental Fig. S4). Similarly, expression of hCa V 3.2(WT)-GFP, hCa V 3.2(C456S)-GFP, or Ca V 3.2ct-GFP in TRN neurons in vivo for ∼18 h had no significant effect on the basic membrane properties, including RMP, input resistance, membrane time constant, burst firing, and average spiking frequency (Supplemental Fig. S5). Together, the results suggest that Ca V 3.2 channels regulate glutamatergic transmission in central neurons.
TRN neurons receive glutamatergic synaptic input from both thalamic and cortical afferent fibers. To determine whether Ca V 3.2 channels enhance AMPA and NMDA responses at thalamic and/or cortical synapses, we simultaneously expressed mCherry-tagged channelrhodopsin-2 (ChR2-mCherry) in the thalamic ventrobasal nucleus (VB) or barrel cortex and hCa V 3.2-GFP or Ca V 3.2ct-GFP in the TRN in vivo ( Fig. 2A). Neurons expressing ChR2-mCherry were confined within the VB and barrel cortex after viral delivery in these areas, respectively (Supplemental Fig. S6). After ∼18 h expression of ChR2-mCherry in the VB, we optogenetically activated the thalamic ChR2-mCherry-expressing terminals in the TRN and recorded ensuing EPSCs from TRN neurons in acutely prepared slices. In response to optogenetic stimulation of the thalamic terminals, TRN neurons expressing hCa V 3.2(C456S)-GFP had enhanced optogenetically evoked AMPA and NMDA responses, while TRN neurons expressing Ca V 3.2ct-GFP had reduced optogenetically evoked AMPA and NMDA responses (Fig. 2F,G). Similarly, after ∼72 h of expression of ChR2-mCherry in the barrel cortex, we optogenetically activated the cortical ChR2-mCherry-expressing terminals in the TRN and recorded evoked EPSCs from TRN neurons in vitro. In response to optogenetic stimulation of the cortical terminals, TRN neurons expressing hCa V 3.2(C456S)-GFP had enhanced optogenetically evoked AMPA and NMDA responses, while TRN neurons expressing Ca V 3.2ct-GFP had reduced optogenetically evoked AMPA and NMDA responses (Fig. 2F,G). These results indicate that Ca V 3.2 channels regulate AMPA and NMDA responses at both the thalamic and cortical synapses of TRN neurons. To independently support these findings, we quantified the distribution of AMPA-Rs and NMDA-Rs at thalamic synapses and cortical synapses of the TRN using pre-embedding immunogold labeling. In the TRN, excitatory afferent inputs originating from the thalamus and cortex form the characteristic L terminals and D terminals, respectively, and could thus be unambiguously identified (Ohara 1988). Expression of hCa V 3.2(C456S) increased, whereas expression of Ca V 3.2ct decreased, the amount of GluA2/3-immunoreactive silver-gold particles in the postsynaptic density (PSD) of the thalamic and cortical synapses in the TRN (Fig. 3A,B). Similarly, expression of hCa V 3.2(C456S) increased, whereas expression of Ca V 3.2ct decreased, the amount of GluN1-immunoreactive silver-gold particles in the PSD of the thalamic and cortical synapses in the TRN (Fig. 3C,D). These results suggest that Ca V 3.2 channels function to regulate synaptic incorporation of AMPA-Rs and NMDA-Rs. As a control, expression of hCa V 3.2(WT) had no effect on the amounts of GluA2/3-and GluN1-immunoreactive silver-gold particles in the PSD of the thalamic and cortical synapses in the TRN (Fig. 3). Collectively, the electrophysiological, optogenetic, and immunoelectron microscopic experiments suggest that the Ca V 3.2 channelmediated regulation of glutamatergic transmission is a general phenomenon at central synapses.
We then examined evoked EPSCs in CA1 neurons in the normal bath solution. In the presence of 50 μM Ni 2+ or 1 μM TTA-P2 in culture medium, CA1 neurons expressing hCa V 3.2(C456S)-GFP or Ca V 3.2ct-GFP had the same AMPA and NMDA responses compared with nearby control nonexpressing neurons (Fig. 4C,D). These results suggest that pharmacological blockade of Ca V 3.2 channel activity blocks the potentiation of AMPA and NMDA responses in hCa V 3.2(C456S)-GFP-expressing neurons and occludes the depression of AMPA and NMDA responses in Ca V 3.2ct-GFP-expressing neurons. These results suggest that activation of Ca V 3.2 channels is required for the Ca V 3.2 channel-dependent regulation of glutamatergic transmission.
We wished to know whether synaptic activity drives the activation of Ca V 3.2 channels to regulate glutamatergic transmission, since low-threshold calcium channels, including Ca V 3.2 channels, can be activated by synaptic input at the RMP. To test this idea, we included 12 mM MgCl 2 or 4 µM TTX, which depresses or blocks synaptic transmission (Zhu et al. 2000;McCormack et al. 2006), in culture medium during expression of hCa V 3.2 (C456S)-GFP and Ca V 3.2ct-GFP in CA1 neurons. We then examined the evoked EPSCs in these neurons in the normal bath solution. In the presence of high Mg 2+ or TTX in culture medium, CA1 neurons expressing hCa V 3.2(C456S)-GFP or Ca V 3.2ct-GFP had the same AMPA and NMDA responses compared with nearby nonexpressing neurons (Supplemental Fig. S7). These results suggest that synaptic activity activates Ca V 3.2 channels to regulate glutamatergic transmission.
Because the above-reported data showed that activation of Ca V 3.2 channels did not significantly alter the basic membrane properties, we speculated that the Ca V 3.2 channel-dependent regulation of glutamatergic transmission might be mediated by calcium influx via the channels. To test this possibility, we cultured CA1 neurons in medium in which calcium ions were replaced with barium ions, which are equally or more efficient charge carriers for Ca V 3.2 channels (McRory et al. 2001). Substitution of calcium ions with barium ions prevented the hCa V 3.2(C456S)-induced potentiation and the Ca V 3.2ct-induced depression in CA1 neurons (Fig.  4C,D). Collectively, these results suggest that synaptic activity activates Ca V 3.2 channels and induces calcium influx that potentiates NMDA responses prior to AMPA responses.
To directly verify that activation of Ca V 3.2 channels induces calcium influx, we imaged Ca 2+ transients in CA1 neurons expressing hCa V 3.2-GFP and Ca V 3.2ct-GFP in cultured hippocampal slices. Simultaneously, we loaded the green Ca 2+ -sensitive indicator Fluo-5F and the red Ca 2+ -insensitive indicator Alexa 594 into expressing and neighboring nonexpressing CA1 neuron pairs and recorded the back-propagating action potential (bAP)-evoked fluorescence changes in nearby pairs of spines and their parent dendrites of these neurons (Fig. 5A,B). We found that the spines, but not the parent dendrites, of neurons expressing hCa V 3.2(C456S)-GFP had significantly increased bAP-evoked Ca 2+ transients (Δ[Ca] bAP ) compared with those of nonexpressing neurons (Fig. 5C). In contrast, the spines, but not the parent dendrites, of neurons expressing Ca V 3.2ct-GFP had significantly reduced Δ[Ca] bAP (by ∼50%) compared with those of nonexpressing neurons (Fig. 5C). As controls, the spines and parent dendrites of neurons expressing hCa V 3.2(WT)-GFP had the same Δ[Ca] bAP compared with those from nonexpressing neurons (Fig. 5C). To confirm that synaptic Δ[Ca] bAP is mediated by T-type calcium channels, we carried out additional pharmacological experiments. Including 1 μM TTA-P2 or 50 μM Ni 2+ in the bath solution blocked the hCa V 3.2(C456S)-induced increase in Δ[Ca] bAP and occluded the Ca V 3.2ct-induced decrease in Δ[Ca] bAP at the spines (Fig. 5C). These results confirm that activation of Ca V 3.2 channels contributes to local calcium influx at the synapses.
To define the contribution of Ca V 3.2 channels to the T-type calcium channel-mediated calcium influx at the synapses, we imaged Ca 2+ transients in CA1 neurons in acutely prepared slices after ∼18 h of in vivo expression of hCa V 3.2-GFP and Ca V 3.2ct-GFP. In these experiments, we included L-, N-, P/Q-, and R-type channel antagonists, 10 μM nimodipine, 1 μM CTX-MVIIC, and 0.3 μM SNX-482 in the bath solution to isolate T-type calcium currents. In the presence of nimodipine, CTX-MVIIC, and SNX-482, the spines, but not the parent dendrites, of neurons expressing hCa V 3.2(C456S)-GFP had significantly increased Δ[Ca] bAP . Moreover, after the isolation of T-type calcium currents, the spines, but not the parent dendrites, of neurons expressing Ca V 3.2ct-GFP had substantially reduced Δ[Ca] bAP (by ∼75%) (Fig. 5C). Collectively, these results show that endogenous Ca V 3.2 channels are the primary contributors to the T-type channel-mediated calcium influx at synapses.
We also analyzed Ca 2+ transients at single spines using a second approach that does not require counting the failure of Δ[Ca] bAP , which can be very low in some spines, and thus allowed the analysis to include all data points (Yasuda et al. 2003). We plotted the ratio of the squared coefficients of variance of the trial-to-trial fluctuation versus the ratio of mean amplitude of fluorescence in all nearby spine pairs of expressing and neighboring nonexpressing CA1 neuron pairs (Fig. 5F). The plot shows that enhanced calcium influx in the spines of neurons expressing hCa V 3.2(C456S)-GFP was quantitatively consistent with an increase in p but no change in N and calcium influx per opening q. Moreover, the results indicate that no change in calcium influx in the spines of neurons expressing hCa V 3.2(WT)-GFP was quantitatively consistent with no change in p, N, and q (Fig. 5F). Finally, we virally expressed hCa V 3.2-GFP with an additional HA tag in its extracellular domain (hCa V 3.2-HA-GFP) in cortical L5 pyramidal neurons in vivo and then processed the tissues for anti-HA immunohistochemistry in a nonpermeabilized condition. Both hCa V 3.2(C456S)-HA-GFP and hCa V 3.2 (WT)-HA-GFP appeared in dendrites, with some delivered to the surface in spines. Quantitative analysis showed no difference in anti-HA Texas red immunofluorescence intensity at the spines of neurons expressing hCa V 3.2 (C456S)-HA-GFP and hCa V 3.2(WT)-HA-GFP (Supplemental Fig. S8), suggesting the same synaptic surface expression of hCa V 3.2(C456S)-HA-GFP and hCa V 3.2(WT)-HA-GFP channels. Collectively, these results consistently support the conclusion that human CAE-linked Ca V 3.2 (C456S) mutant channels enhance synaptic calcium influx by replacing endogenous calcium channels with higher open probability mutant channels without changing calcium influx per opening.

CAE-linked hCa V 3.2(C456S) mutant channels induce absence-like epilepsy
To test whether CAE-linked gain-of-function mutations on human Ca V 3.2 channels may promote seizures associated with absence epilepsy, we virally expressed recombinant hCa V 3.2-GFP in the rat barrel cortex, one of the primary sites known to initiate absence epilepsy (Steriade 2005;Huguenard and McCormick 2007). After ∼18 h of expression, we simultaneously monitored the animal's electroencephalography (EEG) with a wireless transmitter and monitored its behavior with a video camera (Fig. 6A). We found that >60% of animals expressing hCa V 3.2 (C456S)-GFP exhibited 2-to 4-Hz SWDs (Fig. 6B,C), characteristic of CAE patients (Snead 1995;Chen et al. 2003). On average, animals expressing hCa V 3.2(C456S)-GFP were approximately fourfold to sixfold more likely to have 2-to 4-Hz SWDs and generated 2-to 4-Hz SWDs approximately sixfold to eightfold more frequently compared with animals expressing hCa V 3.2(WT)-GFP or GFP alone (Fig. 6B,C). Animals that had sham surgeries displayed no 2-to 4-Hz SWDs during the recording period. Several animals expressing hCa V 3.2(C456S)-GFP exhibited long and robust epileptic activity that lasted ∼20-40 sec and was accompanied by behavioral arrests typical of absence seizures (n = 6) ( Fig. 6B; Supplemental Movie S1). None of the animals expressing hCa V 3.2(WT)-GFP or GFP alone or animals with sham surgery displayed any absence-like behavioral arrest or seizure (see Supplemental Movie S2). These results indicate that the expression of hCa V 3.2(C456S) mutant proteins enhances the susceptibility of absence-like epilepsy.
Approximately 50% of CAE patients do not respond to ethosuximide, an effective T-type calcium channel antagonist and first-line drug used to treat absence epilepsy (Coulter et al. 1989;Huguenard and Prince 1994), and many of the nonresponsive patients have mutations in Ca V 3.2 calcium channels (Glauser et al. 2010). We speculated that the absence epilepsy in these patients may result from the Ca V 3.2 channel mutation-induced potentiation of glutamatergic transmission and thus renders the patients to be nonresponsive to T-type calcium channel antagonists once the alteration in glutamatergic transmission has taken place. To test this idea, we administered T-type channel and/or glutamatergic receptor antagonists intraperitoneally in animals expressing hCa V 3.2(C456S)-GFP prior to EEG and behavior examination (Fig. 6B,C). Administration of a single dose of TTA-P2 in animals after 18 h of expression of hCa V 3.2(C456S)-GFP had no effect on the occurrence of 2-to 4-Hz SWDs. To validate whether the hCa V 3.2(C456S)-mediated glutamatergic potentiation is responsible for SWDs, we administered TTA-P2 in animals every 4 h (four doses total) during the expression of hCa V 3.2(C456S)-GFP to prevent the hCa V 3.2(C456S)-induced potentiation of glutamatergic transmission. This dosing paradigm significantly depressed the occurrence of 2-to 4-Hz SWDs in animals expressing hCa V 3.2 (C456S)-GFP (Fig. 6B,C). Consistent with the proposed epileptogenic mechanism involved in the potentiated glutamatergic transmission, we found that the four-dose, but not one-dose, TTA-P2 administration paradigm blocked the hCa V 3.2(C456S)-enhanced glutamatergic transmission in cortical L5 pyramidal neurons (Supplemental Fig.  S9). On the other hand, administration of one dose of the drug cocktail of MK 801, an NMDA-R antagonist, and GYKI 52466, an AMPA-R antagonist, in animals after 18 h of expression of hCa V 3.2(C456S)-GFP was sufficient to significantly depress the occurrence of 2-to 4-Hz SWDs. Adding additional TTA-P2 in the glutamatergic antagonist drug cocktail only slightly improved the ameliorating effect on the occurrence of 2-to 4-Hz SWDs. None of the animals expressing hCa V 3.2(C456S)-GFP that were administrated with the glutamatergic antagonist drug cocktail or glutamatergic antagonist and T-type calcium channel antagonist drug cocktail displayed any absencelike behavioral arrest or seizure (see Supplemental Movie S3). These results lead us to conclude that hCa V 3.2 (C456S) mutant channels can increase the susceptibility of absence-like epilepsy by potentiating glutamatergic transmission.

Discussion
In this study, we investigated the physiological and pathological functions of Ca V 3.2 T-type low-threshold calcium channels in multiple distinct types of central neurons. Our results indicate that the general function of Ca V 3.2 channels is to control NMDA-R-mediated transmission at central synapses, and dysregulation of Ca V 3.2 channels enhances susceptibility for human CAE.
The profound interest in the regulation of NMDA-Rmediated transmission stems from the fact that NMDA transmission contributes to fundamental neural functions, such as synaptic integration and amplification (Lavzin et al. 2012;Smith et al. 2013;Augustinaite et al. 2014;Grienberger et al. 2014;Palmer et al. 2014;Crandall et al. 2015), synaptic plasticity of AMPA-R-mediated transmission (Kessels and Malinow 2009;Stornetta and Zhu 2011;Huganir and Nicoll 2013), and behavior adaption (Cichon and Gan 2015). It is generally believed that synaptic trafficking of NMDA-Rs is more closely regulated than that of AMPA-Rs, particularly at mature synapses, and dysregulation of NMDA-R-mediated transmission has been implicated in a number of neurological, mental, and psychiatric disorders, including addiction, Alzheimer's disease, autism, depression, pathological pain, and schizophrenia (Lau and Zukin 2007;Russo et al. 2010;Hunt and Castillo 2012;Paoletti et al. 2013;Monteggia and Zarate 2015). Consistent with this idea, we show here that the numbers of functional Ca V 3.2 calcium channels, the regulators of NMDA transmission, are tightly regulated at the synapses; newly expressed Ca V 3.2 channels only replace existing Ca V 3.2 channels at the synapses in central neurons to maintain a constant number of synaptic Ca V 3.2 channels. The tight regulation of functional Ca V 3.2 channels provides a mechanism for maintaining the relatively stable numbers of NMDA-Rs at the synapses. Our analysis reveals that the amount of Ca V 3.2 channel-mediated calcium influx at the synapses plays a central role in controlling the strength of NMDA transmission via the regulation of synaptic expression of NMDA-Rs. We further show that synaptic activity activates Ca V 3.2 channels to regulate NMDA transmission strength at the synapses (Supplemental Fig. S7), thereby extending the previous work that synaptic activity activates T-type currents (Magee and Johnston 1997). By analogy, we speculate that certain signaling molecules, including G-protein-coupled receptors, which can modulate the activity of Ca V 3.2 channels (Yasuda et al. 2003;DePuy et al. 2006;Yao et al. 2006), may also mold synaptic NMDA responses.
How exactly postsynaptic Ca V 3.2 channel-mediated calcium transients signal synaptic trafficking of NMDA-Rs remains largely unknown, although calcium influx at the synapses is believed to be essential for the regulation of NMDA transmission (Lau and Zukin 2007;Hunt and Castillo 2012;Paoletti et al. 2013;Cerpa et al. 2015). In addition, recent findings have implicated that several molecules may relay signaling downstream from synaptic calcium influx to regulate NMDA transmission (Setou et al. 2000;Jeyifous et al. 2009;Udagawa et al. 2012;Plattner et al. 2014;Wang and Zhu 2014;Wei et al. 2014;Sheng et al. 2015). Some of the signaling molecules (e.g., PKA, PKC, JNK, Fyn, and Cdk5) could potentially control phosphorylation of transcription and translation factors, sorting, transporting, and scaffolding proteins as well as NMDA-Rs to signal synaptic delivery and/or removal of NMDA-Rs. Obviously, compared with synaptic regulation of AMPA-Rs, our understanding of the dynamics of trafficking and function of NMDA-Rs has advanced much more slowly, and many questions remain unanswered (Lau and Zukin 2007;Hunt and Castillo 2012;Paoletti et al. 2013). Our discovery of the powerful effect of Ca V 3.2 channels on NMDA transmission provides a technically feasible approach to address these fundamental questions in the near future. This study also identifies Ca V 3.2 channels as a new potential drug target for treating the disorders resulting from the dysregulation of NMDA transmission.

Pathological function of Ca V 3.2 channels
Human CAE is the most common form of pediatric epilepsy (Tenney and Glauser 2013). A number of absence epilepsy animal models have been reported. However, with the exception of the γ-butyrolactone-based pharmacological model (Schofield et al. 2009;Zaman et al. 2011), all of the commonly used animal models, including genetic animal models, failed to produce the ∼2to 4-Hz SWDs characteristic of CAE patients (Snead 1995;Chen et al. 2003;Tenney and Glauser 2013). Therefore, it is still challenging to define the genetic nature and epileptogenic mechanism of human CAE. Here, we showed that expression of hCa V 3.2(C456S) alone in rats was sufficient to markedly enhance the incidence of ∼2to 4-Hz SWDs and behavioral arrests or seizures closely resembling patients with CAE (Snead 1995;Chen et al. 2003;Tenney and Glauser 2013). These results provide the first evidence indicating CACNA1H to be a primary gene responsible for CAE susceptibility. Our analysis shows that expression of hCa V 3.2(C456S) results in the replacement of endogenous Ca V 3.2 channels with higher open probability mutant channels without altering calcium influx per opening. This replacement increases the local calcium influx at synapses and leads to the up-regulation of glutamatergic transmission and enhancement of the susceptibility of absence-like epilepsy. These findings reveal a new epileptogenic mechanism for CAE and explain the clinical conundrum that CAE patients carrying Ca V 3.2 channel mutations do not respond to ethosuximide.
As a fairly common form of generalized seizure, the network mechanisms of SWDs have been extensively investigated for more than seven decades (Avoli 2012;Luttjohann and van Luijtelaar 2015). Accumulating evidence indicates that SWDs are generated within the corticothalamocortical circuitry and that the dynamic interaction of local microcircuits within this neuronal network leads to the "full-blown" SWDs associated with absence epilepsy (McCormick and Contreras 2001;Steriade 2005;Huguenard and McCormick 2007;Luttjohann and van Luijtelaar 2015). The very early studies of absence epilepsy (also known as petit mal epilepsy) reported that SWDs, as in human absence epilepsy, could initiate and sustain after a brief electric stimulation in the thalamus in animals (Lewy and Gammon 1940;Lindsley et al. 1949). Later, EEG recordings in humans showed that SWDs occurred in the thalamus prior to the cortex (Williams 1953). These results suggest a possible thalamic origin of SWDs. In the following 50 years, results supporting either the cortical or thalamic origin of SWDs have been reported (Avoli 2012). In the most recent studies, investigators employed the simultaneous EEG and/or functional magnetic resonance imaging (fMRI) recordings at multiple cortical and thalamic sites and saw the precursor activity in specific cortical areas prior to SWD onset in animal models (Meeren et al. 2002;David et al. 2008) as well as in human patients (Tyvaert et al. 2009;Bai et al. 2010). These findings provide the most compelling evidence supporting that the cortex is actually the initiator of SWDs, although technical limitations prevented the simultaneous monitoring of every thalamic and cortical area or subareas potentially involved in SWDs in these studies (Luttjohann and van Luijtelaar 2015). Here, we introduced an alternative genet-ic approach that may address network mechanisms in a complementary way. We showed that the selective expression of human CAE-linked Ca V 3.2(C456S) mutant proteins in the rat barrel cortex is sufficient to enhance the susceptibility of 2-to 4-Hz SWDs characteristic of human CAE patients. The results validate the possibility of making CAE-linked genetic modifications in one or multiple thalamic and/or cortical areas in animal models and thus open a new avenue to dissect the contributions of local microcircuits in the thalamic nuclei and/or cortical areas in the generation, maintenance, and termination of SWDs. For example, it would be interesting and feasible to test whether expression of hCa V 3.2(C456S) in TRN alone may induce the ictogenic process and whether, in both TRN and cortical areas, as likely occurs in the human condition, it may synergistically reduce the ictogenic threshold. Together, the complementary research approaches will facilitate delineation of the genetic, cellular, and neuronal network mechanisms of SWDs associated with human CAE, which are essential for developing new effective therapeutic options to treat this neurological disorder (Maheshwari and Noebels 2014;Paz and Huguenard 2015).

Cultured and acute slice preparation
Organotypic hippocampal cultured slices were prepared from postnatal 6-to 7-d-old (P6-P7) male and female Sprague Dawley rats, wild-type mice, and Ca V 3.2 knockout (Orestes et al. 2009) mice bred congenically on the C57BL/6 background, following our previous studies (Qin et al. 2005;Lim et al. 2014). All procedures for animal surgery and maintenance were performed following protocols approved by the animal care and use committee of the University of Virginia and in accordance with National Institutes of Health guidelines. The hippocampi were dissected out in ice-cold HEPES-buffered Hanks' solution (pH 7.35) under sterile conditions, sectioned into 400-µm slices on a tissue chopper, and explanted onto a Millicell-CM membrane (0.4-µm pore size; Millipore). The membranes were then placed in 750 µL of MEM culture medium containing 30 mM HEPES, 20% heat-inactivated horse serum, 1.4 mM glutamine, 16.25 mM D-glucose, 5 mM NaHCO 3 , 1 mM CaCl 2 , 2 mM MgSO 4 , 1 mg/mL insulin, and 0.012% ascorbic acid (pH 7.28, osmolarity 320). Cultured slices were maintained at 35°C in a humidified incubator (ambient air enriched with 5% CO 2 ). Acute thalamic, cortical, and hippocampal slices were prepared from P14-P26 male and female Sprague Dawley rats. Animals were deeply anesthetized by xylazine-ketamine and decapitated. The brain containing the thalamus, barrel cortex, and/or hippocampus was quickly removed and placed into cold (0°C-4°C) oxygenated physiological solution containing 125 mM NaCl, 2.5 mM KCl, 1.25 mM NaH 2 PO 4 , 25 mM NaHCO 3 , 1 mM MgCl 2 , 25 mM dextrose, and 2 mM CaCl 2 (pH 7.4). To obtain the thalamic slices containing the TRN, horizontal slices 350 μm thick were cut from the brain blocks with their ventral surface glued down to the cutting chamber of a microslicer. For acute cortical preparations, parasagittal slices 350 μm thick were cut from the brain blocks. For acute hippocampal preparations, parasagittal slices 350 μm thick were cut from the brain blocks. These slices were kept at 37.0°C ± 0.5°C in oxygenated physiological solution for ∼0.5-1 h before recording.

Constructs and recombinant protein expression
To avoid the potential compensatory and other nonspecific effects that may be associated with chronic overexpression of Ca V-3.2 channels (Simms and Zamponi 2014) and long-lasting changes in glutamatergic transmission (Kessels and Malinow 2009;Stornetta and Zhu 2011), the rapid Sindbis and lentiviral expression systems were used in all experiments. Because the regulation of protein trafficking in cultured neurons can be very different from those in cultured slices and intact brains (Kessels and Malinow 2009;Stornetta and Zhu 2011), all expressions were made in neurons in cultured slices and intact brains. GFP-and/or HAtagged human Ca V 3.2 wild-type constructs (Supplemental Fig.  S10)-hCa V 3.2(WT)-GFP and hCa V 3.2(WT)-HA-GFP-were gifts from Dr. Emmanuel Bourinet (Dubel et al. 2004). Mutant C456 → S constructs were generated from hCa V 3.2(WT)-GFP and hCa V 3.2(WT)-HA-GFP using Quick Change XL site-directed mutagenesis kit (Stratagene). The wild-type and mutant hCa V 3.2 constructs were subcloned into a Sindbis viral construct. The C terminus of rat Ca V 3.2 was first subcloned into pEGFP-C1 or pRFP-C1 (Clontech Laboratories) and then into the Sindbis viral construct. ChR2-mCherry was subcloned into Sindbis and lentiviral constructs. In some experiments, GFP was replaced with CFP, mCherry, or mOrange (gifts from Dr. Roger Tsien). For in vitro expression, neurons were infected after 6-14 d in culture. For in vivo expression, unless stated otherwise, P13-P25 animals were initially anesthetized with an intraperitoneal (i.p.) injection of 10 mg/kg ketamine and 2 mg/kg xylaxine and then placed in a stereotaxic frame with a hole of ∼1 mm × 1 mm drilled above the right barrel cortex. A glass pipette was used to penetrate into the thalamic VB, TRN, barrel cortex, and/or hippocampus according to stereotaxic coordinates, and ∼50 nL of viral solution was delivered by pressure injection. The injection produced the restricted expression of ChR2 in either the VB or the barrel cortex (Supplemental Fig. S6). The titer of the viral solution was adjusted for both in vitro and in vivo electrophysiology experiments to express Ca V 3.2 constructs in only a few neurons (typically, one to five neurons per slice/brain) (Supplemental Fig. S1A). For behavioral experiments, three parallel strips of ∼3 mm × 0.75 mm were drilled above the right barrel cortex, and eight to nine injections of ∼6 µL of high-titer viral solution were delivered in the entire barrel cortex. Experiments were typically performed at 18 h ± 3 h after Sindbis viral infection or 72 h ± 6 h after lentiviral infection.

Immunostaining and immunoelectron microscopy
Immunostaining of the HA tag was carried out as previously described (Zhu et al. 2000). To reduce the variation of immunostaining intensity, hCa V 3.2(WT)-HA-GFP and hCa V 3.2(C456S)-HA-GFP were expressed in the barrel cortical L5 in the different hemispheres of the same animals, and the tissue sections containing the expressing neurons were processed simultaneously. To evaluate the surface expression of hCa V 3.2-HA-GFP, detergents were omitted in all incubations. Antibody incubations were in PBS for 48 h at 4°C (primary anti-HA antibody, 1:500; Covance, 16B12) or overnight (secondary antibody and avidin-Texas red). Pre-embedding immunogold labeling was carried out following the procedures of previous studies (Erisir and Harris 2003;Kielland et al. 2009). In brief, 1 µg/mL rabbit anti-GluA2/3 (Millipore, AB1506) was used to detect AMPA-Rs because of the nonrectified AMPA responses seen in CA1 and L5 pyramidal neurons (Zhu 2009;Lim et al. 2014) and TRN neurons (values for rectification: R VB stim = 1.51 ± 0.06 and R cortex stim = 1.54 ± 0.08; n = 19 from nine animals), an indicator of incorporation of GluA2 subunits in the majority of AMPA-Rs. A mouse monoclonal anti-GluN1 (Luo et al. 1997) was used to detect the obligatory GluN1 subunits of NMDA-Rs. Sample sections were rinsed in PBS and incubated with antibodies and 1 nm of gold-conjugated goat anti-rabbit or mouse IgG (1:50; GE Healthcare Bio-Sciences) for 2 h followed by citrate buffer rinse and silver intensification for ∼8-10 min. Immunolabeled sections were fixed with 1% OsO 4 for 1 h and flat-embedded between two pieces of Aclar (Ted Pella, Inc.), and the resin was left to polymerize for 48 h at 60°C. With high-titer viruses, ∼70%-85% of neurons, identified by GFP fluorescence, could be infected near the injection sites (Supplemental Fig.  S11). Only the highly infected areas were excised and resectioned into 80-nm ultrathin sections using an ultramicrotome. Ultrathin sections were examined with a transmission electron microscope (Japan Electron Optic, JEOL-1010). Nonspecific immunogold labeling was low, as indicated by very low counts of silver-enhanced gold particles in presynaptic regions. The thalamic and cortical synapses of TRN neurons could be unambiguously classified into two groups based on their characteristic presynaptic terminals. Specifically, the thalamic synapses were contacted by L terminals that had a large terminal with loosely packed spherical vesicles and typically contained multiple mitochondria, whereas the cortical synapses were contacted by D terminals that had a small terminal with densely packed spherical vesicles and contained no or single mitochondria (Ohara 1988). We counted all silver-enhanced gold particles within 500 nm from the postsynaptic membrane (approximately one to six per synapse) and classified those within the concentric rings of −6.25 to 31.25 nm from the postsynaptic membrane into the synaptic pool and those of 31.25-106.25 nm into the deliverable pool (cf. Kielland et al. 2009).

Electrophysiology
Using our recently developed stable multiple patch clamp recording system (Jiang et al. 2013;Lee et al. 2014b;Wang et al. 2015), simultaneous and/or sequential multiple whole-cell recordings were obtained from two to five nearby infected and noninfected TRN neurons or cortical L5 or hippocampal CA1 pyramidal neurons under visual guidance using fluorescence and transmitted light illumination. For cultured slices, bath solution (29°C ± 1.5°C) contained 119 mM NaCl, 2.5 mM KCl, 4 mM CaCl 2 , 4 mM MgCl 2 , 26 mM NaHCO 3 , 1 mM NaH 2 PO 4 , 11 mM glucose, 0.1 mM picrotoxin (PTX), 0.01 mM bicuculline, and 0.002 mM 2chloroadenosine (pH 7.4) gassed with 5% CO 2 /95% O 2 . For experiments in which slices were maintained in culture medium with additional 100 µM DL-APV, 5 µM DCKA, 50 μM Ni 2+ , 1 μM TTA-P2, 10 mM Mg 2+ , or 2 μM TTX, these reagents were included in the culture medium during expression but not in the bath solution during recording. For acute slices, 2-chloroadenosine was excluded; 2 mM CaCl 2 and 1 mM MgCl 2 were used instead in the bath solution. PTX was excluded when GABA responses were examined. Whole-cell recordings were made with up to five Axopatch-200B patch clamp amplifiers (Molecular Devices). Junction potentials were not corrected. Synaptic responses were evoked by bipolar electrodes with single voltage pulses (200 μsec, up to 20 V). Synaptic AMPA and NMDA responses at −60 mV and +40 mV were averaged over 90 trials.

Two-photon imaging and optogenetics
Two-photon imaging and electrophysiology recordings were simultaneously performed using a custom-built microscope operated by a custom-written IGOR Pro 6 program (WaveMetrics) (Wang et al. 2015). To minimize the Ca 2+ signal variances resulting from the different location of synapses and loading time of indicators (Sabatini and Svoboda 2000;Yasuda et al. 2003), neighboring expressing and nonexpressing CA1 neuron pairs were broken in simultaneously to load 200 μM green calcium-sensitive indicator Fluo-5F and 20 μM red calcium-insensitive indicator Alexa 594. After ∼15-30 min of loading of indicators, fluorescence of nearby spine pairs of neighboring expressing and nonexpressing CA1 neuron pairs was simultaneously imaged with 500-Hz line scanning that intersected both the spines and their parent dendrites. Alexa 594 and Fluo-5F were excited by a femtosecond Ti:Sapphire laser (Chameleon Ultra, Coherent) at a wavelength of 880 nm. Calcium channels were activated by the single bAPs evoked with somatic current injections in expressing and nonexpressing neurons. Changes in fluorescence were quantified as increases in green (Fluo-5F) fluorescence from baseline divided by resting red (Alexa fluor 594) fluorescence (ΔG/R) and averaged for ∼50-200 trials. No optical fluctuation analysis of calcium channels at single spines was performed in neurons expressing Ca V 3.2ct due to no calcium transients being detected in many of the spines imaged (n = 8 out of 18). Optogenetic activation of the ChR2-expressing axonal terminals originating from VB neurons or cortical neurons in the TRN was made with a 473-nm blue laser (OBIS, Coherent). The OBIS laser was coupled to the two-photon laser light path and delivered onto the specimen using the same objective. The laser power at the specimen was set at 0.1-15 mW, and 10-msec light pulses were generated with a fast shutter (Uniblitz UHS1, Vincent Associates).

EEG recording and behavior monitoring
After viral expression of hCa V 3.2-GFP or GFP in the right side of the barrel cortex, a wireless EEG transmitter (Epoch in vivo recording system, Ripple) was implanted above the cortex of the animals ( Fig. 6A; Supplemental Movie 1). EEG activity was recorded at 18 h ± 3 h after surgery and viral expression, and animal behavior was simultaneously monitored by a video camera for a period of 6 h ± 2 h (from approximately 12 a.m.-6 a.m.). TTA-P2 (10 mg/ kg; supplied by Dr. Victor Uebele at Merck), 0.05 mg/kg MK 801, and/or 20 mg/kg GYKI 52466 (cf. Rajasekaran et al. 2012) were administered via i.p. injections prior to EEG and behavior recording. In some experiments, to prevent Ca V 3.2(C456S)-GFP from potentiating glutamatergic synaptic transmission in expressing animals, four doses of TTA-P2 spaced by 4 h were administered, with the first dose of TTA-P2 administered at 4 h after the viral injection, considering ∼6-8 h required for viral expression of recombinant proteins. These drugs were dissolved in saline, and saline alone was used as a control. The behavioral data were analyzed blind of control and experiment groups.

Statistical analysis
Statistical results were reported as mean ± SEM. Statistical significances of the means (P ≤ 0.05, two-sided) were determined using Wilcoxon and Mann-Whitney rank sum nonparametric or χ 2 tests for paired and unpaired samples, respectively.