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W4220996879 abstract "Article Figures and data Abstract Editor's evaluation Introduction Results Discussion Materials and methods Data availability References Decision letter Author response Article and author information Metrics Abstract Dravet syndrome (DS) is a neurodevelopmental disorder due to pathogenic variants in SCN1A encoding the Nav1.1 sodium channel subunit, characterized by treatment-resistant epilepsy, temperature-sensitive seizures, developmental delay/intellectual disability with features of autism spectrum disorder, and increased risk of sudden death. Convergent data suggest hippocampal dentate gyrus (DG) pathology in DS (Scn1a+/-) mice. We performed two-photon calcium imaging in brain slice to uncover a profound dysfunction of filtering of perforant path input by DG in young adult Scn1a+/- mice. This was not due to dysfunction of DG parvalbumin inhibitory interneurons (PV-INs), which were only mildly impaired at this timepoint; however, we identified enhanced excitatory input to granule cells, suggesting that circuit dysfunction is due to excessive excitation rather than impaired inhibition. We confirmed that both optogenetic stimulation of entorhinal cortex and selective chemogenetic inhibition of DG PV-INs lowered seizure threshold in vivo in young adult Scn1a+/- mice. Optogenetic activation of PV-INs, on the other hand, normalized evoked responses in granule cells in vitro. These results establish the corticohippocampal circuit as a key locus of pathology in Scn1a+/- mice and suggest that PV-INs retain powerful inhibitory function and may be harnessed as a potential therapeutic approach toward seizure modulation. Editor's evaluation Recent work has shown that one of the major dogmas in epilepsy – that interneuron deficits underlie Dravet syndrome and maybe other epileptic encephalopathies – is overly simplistic. This manuscript makes a significant step forward with novel findings using ex vivo and in vivo experiments providing strong evidence that changes in excitatory connections in the corticohippocampal circuit contribute to mechanisms that drive epilepsy in Dravet syndrome. https://doi.org/10.7554/eLife.69293.sa0 Decision letter Reviews on Sciety eLife's review process Introduction Pathogenic variants in SCN1A, which encodes the voltage-gated sodium channel α subunit Nav1.1, cause a spectrum of epilepsies including Dravet syndrome (DS) (Claes et al., 2001), the most common developmental and epileptic encephalopathy. DS is characterized by treatment-resistant epilepsy, developmental delay/intellectual disability, features of or formal diagnosis of autism spectrum disorder, and motor dysfunction (hypotonia, ataxia, gait impairment) (Villas et al., 2017). DS-associated SCN1A variants are due to loss of function leading to haploinsufficiency of Nav1.1. The heterozygous Scn1a mutant (Scn1a+/-) mouse is a well-established preclinical model of DS that recapitulates key phenotypic features of the human condition (Mistry et al., 2014). When expressed on a 50:50 129S6:C57BL/6J genetic background, these mice exhibit spontaneous seizures beginning at approximately post-natal day (P) 18, and high rates of sudden unexpected death in epilepsy (SUDEP) (Mistry et al., 2014). These mice also exhibit temperature-sensitive seizures, akin to seizures triggered in the setting of fever or hyperthermia in human patients with DS, which represents a key experimental advantage of this mouse model as it readily facilitates study of inducible but naturalistic seizures in vivo (Tran et al., 2020). The prevailing theory as to how reduction in sodium current leads to epilepsy in DS is the so-called ‘interneuron hypothesis’, which posits that haploinsufficiency of Nav1.1 results in selective deficits in inhibitory interneuron excitability based on the relative reliance of this cell class on Nav1.1 for action potential generation. Nav1.1 is prominently expressed in parvalbumin-expressing GABAergic inhibitory interneurons (PV-INs) in neocortex (Ogiwara et al., 2007), as well as by somatostatin (SST-INs) and vasoactive intestinal peptide-expressing interneurons (VIP-INs). Electrophysiological recordings from acutely dissociated hippocampal neurons from Scn1a+/- mice found decreased sodium current density in bipolar-shaped presumptive GABAergic interneurons, but not in pyramidal cells (Bechi et al., 2012; Yu et al., 2006); impaired excitability of inhibitory interneurons – PV-INs as well as SST and VIP-INs – has been demonstrated in acute brain slices from Scn1a+/- mice (Favero et al., 2018; Goff and Goldberg, 2019; Ogiwara et al., 2007; Tai et al., 2014). Consistent with this, human DS patient-derived induced pluripotent stem cells (iPSCs) differentiated to form developing interneurons with biophysical deficits, whereas derived excitatory neurons were normal (Sun et al., 2016). This ‘interneuron hypothesis’ is further supported by the fact that the DS phenotype is recapitulated by selective loss of Nav1.1 exclusively in GABAergic interneurons (Cheah et al., 2012; Dutton et al., 2013; Ogiwara et al., 2013; Rubinstein et al., 2015). However, multiple lines of evidence suggest that the pathophysiology of epilepsy in Scn1a+/- mice is more complex than impairment of inhibition. First, the net circuit effect of VIP-INs is disinhibition, so the abnormal excitability of this population (Goff and Goldberg, 2019) should in fact increase inhibition in neocortical circuits. Second, contradictory data from the Scn1a+/- mouse models (Mistry et al., 2014) and from human-derived iPSCs (Jiao et al., 2013; Liu et al., 2013) suggest that subsets of excitatory neurons in DS may in fact be hyperexcitable. Third, recent work found that impaired action potential generation present at early developmental time points in neocortical PV-INs normalized by P35 (Favero et al., 2018), whereas DS mice continue to exhibit epilepsy, cognitive impairment, and SUDEP. Finally, several in vivo studies have failed to find a clear decrease in baseline interneuron activity in Scn1a+/- mice (De Stasi et al., 2016; Tran et al., 2020). Hence, mechanisms of seizure generation and maintenance of chronic epilepsy in DS remain unclear despite this now growing literature characterizing deficits in Scn1a+/- mice at a single cell level (Cheah et al., 2012; Dutton et al., 2013; Favero et al., 2018; Goff and Goldberg, 2019; Ogiwara et al., 2007; Tai et al., 2014). This may be in part due to the lack of data linking cellular deficits to circuit-level abnormalities. Convergent data suggest that the dentate gyrus (DG) may be a key locus of pathology and seizure generation in Scn1a+/- mice. Although Scn1a+/- mice exhibit multifocal epilepsy, temperature-induced seizures have been shown to prominently emanate from the temporal lobe (Liautard et al., 2013), and focal hippocampal Nav1.1 reduction is sufficient to confer temperature-sensitive seizure susceptibility in conditional Scn1a+/- mice (Stein et al., 2019). Although the DG receives a strong excitatory input from entorhinal cortex (Amaral et al., 2007), inhibition within the DG typically regulates population activity such that granule cells (GCs) are only sparsely activated under physiologic conditions both in reduced preparations in vitro and in experimental animals in vivo (Chawla et al., 2005; Diamantaki et al., 2016; Dieni et al., 2013; Ewell and Jones, 2010; Lee et al., 2016; Liu et al., 2014; Neunuebel and Knierim, 2012; Senzai and Buzsáki, 2017; Yu et al., 2013). However, seizure-evoked immediate early gene activation is most apparent in the DG granule cell layer in Scn1a+/- mice (Dutton et al., 2017), suggesting an underlying circuit-level hyperexcitability. In this study, we demonstrate a profound hyperexcitability of the corticohippocampal circuit in young adult Scn1a+/- mice that is not present at epilepsy onset using two-photon calcium imaging and cellular and synaptic physiology in an acute slice preparation. We find this circuit dysfunction is likely driven by excessive excitation, as opposed to impaired inhibition. We then extend these findings in vivo using optogenetics in awake, behaving mice during temperature-sensitive seizures to demonstrate a role for this corticohippocampal dysregulation in ictogenesis. Finally, we selectively inhibit DG PV-INs (via chemogenetics) to exacerbate seizures in vivo, and recruit PV-INs (via optogenetics) to decrease evoked DG granule cell response to perforant path input in brain slice in vitro. These findings highlight the corticohippocampal circuit as a critical locus of pathology in DS and suggest that PV-INs retain powerful regulation of excitability within the circuit. Results Response of dentate gyrus to entorhinal cortical input is selectively impaired in young adult Scn1a+/- mice Emerging data suggest an important role for the hippocampus and, more specifically, the dentate gyrus (DG), in DS pathology (Dutton et al., 2013; Liautard et al., 2013; Stein et al., 2019). We therefore hypothesized that the circuit function of DG – filtering input from the entorhinal cortex to limit propagation of activity through the limbic system – might be impaired in Scn1a+/- mice. We used two photon calcium imaging (2 P imaging) to achieve large-scale quantification of granule cell (GC) activation in response to stimulation of the perforant path (PP), the excitatory projection from entorhinal cortex that constitutes the major input to GCs. We first injected an adeno-associated virus (AAV) encoding the calcium indicator GCaMP7s (Dana et al., 2019) under control of a pan-neuronal promoter hSyn1 into DG of Scn1a+/- mice and age-matched wild-type littermate controls (Figure 1A). We prepared acute hippocampal-entorhinal cortex (HEC) slices (Xiong et al., 2017), which preserve the PP projection from entorhinal cortex to hippocampus within the slice (Figure 1B). We then performed 2 P imaging of evoked calcium transients in cells within the granule cell layer (GCL) in response to electrical stimulation of the PP (Figure 1C–D). Figure 1 Download asset Open asset Two-photon calcium imaging of perforant path-evoked dentate gyrus activation in acute brain slice. (A) Mice were injected with AAV9-hSyn-jGCaMP7s-WPRE into DG. (B) Acute slices are cut at a 15° angle off-axial to maximize connectivity. Shown is the location of a stimulation electrode (stim) in the perforant path (PP), and the imaging field in DG (green box). (C) GCaMP labeling of GCs in the granule cell layer (GCL; dashed lines) between the hilus (H) and molecular layer (ML). (D) Example GC response to PP stimulation (arrow), displayed as change in fluorescence over baseline (dF/F0). (E) Representative image showing a quantum dot (Andrásfalvy et al., 2014)-labeled pipette tip (red) used for cell-attached recording of GCaMP-expressing GCs (green). (F) Representative data from a single GC firing 0, 1, 3, or 13 action potentials (left) in response to PP stimulation, while calcium transients (right) were simultaneously recorded. Scale bar represents 75 ms (left) and 5 s (right). Vertical dashed line indicates stimulus onset. (G) Action potentials versus calcium transient magnitude for GCs from WT (blue) and Scn1a+/- (red) mice. Data were fit using a linear model, with no significant difference between genotypes (solid / dashed line = best fit / 95% confidence interval). n = 7 (cells), 5 (mice). (H) Results for 0 and 1 action potentials. Threshold for action potential detection (green bar) was defined as p = 0.001 from a normal distribution fit to the dF/F0 values for 0 action potentials. (I) Observed action potentials versus action potentials derived from deconvolution (R2 = 0.83; p < 0.0001). Figure 1—source data 1 Quantification of spikes. https://cdn.elifesciences.org/articles/69293/elife-69293-fig1-data1-v2.xlsx Download elife-69293-fig1-data1-v2.xlsx To characterize GCaMP7s as a reporter of action potentials in GCs within this experimental paradigm, we performed simultaneous cell-attached recording and calcium imaging of GCaMP-expressing cells within the GCL in Scn1a+/- and wild-type mice. We used a quantum dot-labeled pipette tip (Andrásfalvy et al., 2014) for 2P-guided targeted electrophysiological recording during simultaneous calcium imaging (Figure 1E). This allowed us to correlate calcium transients within individual cells against a gold-standard quantitative measure of action potentials (APs). Action potentials were evoked via PP stimulation. However, given the normally sparse activation of DG in response to PP stimulation in wild-type mice, we bath-applied picrotoxin (PTX), a GABAA receptor antagonist that is known to induce widespread PP-driven recruitment of GCs (Dengler et al., 2017), such that nearly all GCs were activated by PP stimulation. We found that the number of APs and the magnitude of the calcium signal were highly correlated (Figure 1F–G; R2 = 0.769). This linear relationship between AP and calcium signal was the same in wild-type and Scn1a+/- mice for single APs and across a measured range (0–14 APs; Figure 1G). We next evaluated the sensitivity and specificity of GCaMP7s in detecting single APs in the absence of PTX as GABAA-mediated responses in dentate GCs can be depolarizing even in the adult DG (Chiang et al., 2012), and could therefore cause subthreshold calcium transients which may mimic spikes (Stocca et al., 2008). We fit a normal distribution to the dF/F0 values associated with 0 APs (with an AP defined as an increase in calcium signal (dF/F0) of at least three times the standard deviation above the mean noise), set a global threshold defined by this data (p = 0.001), and verified that subthreshold responses (n = 9) could be resolved from single APs (n = 10; Figure 1H). Finally, we deconvolved the dF/F0 signals (Pnevmatikakis et al., 2016) to extract the number of action potentials from the calcium transients and verified the deconvolution algorithm on the ‘ground truth’ AP dataset (R2 = 0.83; p < 0.0001; Figure 1I). Thus, GCaMP7s reports APs in GCs similarly in both genotypes, with single-AP resolution, validating 2 P calcium imaging in brain slice as a method for comparing large-scale DG excitability between wild-type and Scn1a+/-. In order to quantify the extent of DG activation in response to entorhinal cortex input, we stimulated the PP while performing 2 P imaging of the evoked responses across hundreds of GCs simultaneously (Figure 2). We tested early postnatal (P14-21; at/around epilepsy onset) and young adult (P50-100; chronic phase) mice. We delivered either a single pulse or a train of 4 pulses at 20 Hz across a range of stimulation intensities, similar to prior studies of this circuit in acute brain slices (Dengler et al., 2017; Ewell and Jones, 2010; Yu et al., 2013). We then identified activated GCs within the imaging field in response to each stimulation condition, defining activation as an increase in calcium signal (dF/F0) at least three times the standard deviation above the mean noise (Figure 2, columns 1–4) and/or if one or more action potentials were extracted using deconvolution (Figure 2, columns 5–6). To determine if the responses across stimulation intensities differed by genotype, we employed a statistical approach based on mixed-effects modeling, to account for potential variation between individual imaging fields in a slice, slices from a given mouse, and/or between mice. (See Figure 2—figure supplement 1 for data including all individual imaging fields, which are omitted for clarity from Figure 2.) Figure 2 with 4 supplements see all Download asset Open asset Selective impairment of dentate gyrus function in young adult Scn1a+/- mice. Proportion of activated GCs (A–C) and magnitude of activation (D) in response to PP stimulation (early postnatal P14-21, columns 1 and 2; young adult P48-91, columns 3 and 4), with deconvolved data for the young adult time point (columns 5 and 6). Wild-type in blue; Scn1a+/- in red. Data were analyzed using a mixed model to account for potential variability between animal, slice, field, and cell. Proportion of responsive GCs was calculated (A) relative to all GCaMP-expressing cells, or restricting analysis (B) to only GCs that respond to PP stimulation in the presence of 100 µM picrotoxin or (C) to only GCs that respond to the maximal stimulation delivered. (D) Magnitude of activated GC responses to PP stimulation, expressed as natural log dF/F0 (columns 1–4) or estimated spikes based on deconvolution (columns 5–6; size of data points reflects number of cells at that value). Dots: raw data from all imaging fields or cells; dark lines: average of fits; shaded: 95% confidence intervals of average fits. Stars indicate significant differences in curve fits: *, p < 0.05; **, p < 0.01; ***, p < 0.001. For early postnatal mice, n for the experiments in rows A, C, and D was, for Scn1a+/- and WT, respectively: 991 and 1,210 (cells), 51 and 58 (fields), 31 and 29 (slices), 11 and 10 (mice); for row B: 372 and 445 (cells), 16 and 17 (both fields and slices), 5 and 5 (mice). For the young adult mice, n for the experiments in rows A, C, and D: 1236 and 1167 (cells), 17 and 17 (both fields and slices), 8 and 6 (mice); for row B: 1109 and 1073 (cells), 16 and 14 (both fields and slices), 6 and 7 (mice). Figure 2—source data 1 Quantified imaging data, reported by cell. https://cdn.elifesciences.org/articles/69293/elife-69293-fig2-data1-v2.xlsx Download elife-69293-fig2-data1-v2.xlsx Figure 2—source data 2 Quantified imaging data, reported by field. https://cdn.elifesciences.org/articles/69293/elife-69293-fig2-data2-v2.xlsx Download elife-69293-fig2-data2-v2.xlsx Figure 2—source data 3 Quantified imaging data, summarizing field response to PTX. https://cdn.elifesciences.org/articles/69293/elife-69293-fig2-data3-v2.xlsx Download elife-69293-fig2-data3-v2.xlsx We first calculated the proportion of activated GCs relative to all identifiable GCaMP7s-expressing cells, fit the data from each genotype (as a binomial distribution fitted with a probit link function), and compared the fit of the curves between genotypes (‘All cells’ in Figure 2A). We validated these data in two different ways to account for the possibility that some GCs might be unresponsive due to deafferentation during the slice preparation, as opposed to this being due to physiologic circuit inhibition. First, we excluded from analysis all GCs that failed to respond to maximal PP stimulation in the presence of PTX (‘Responds to PTX’ in Figure 2B). Note that most ( > 85%) GCs did respond under these conditions (Figure 2—figure supplement 2), demonstrating a high level of connectivity within the HEC slice; hence, results were similar whether considering all cells or only PTX-responsive cells. Second, we excluded from analysis those GCs that failed to respond to the highest amplitude PP stimulation delivered in the absence of PTX (‘Responds to max stim’ in Figure 2C). Finally, in Figure 2D, we quantified the magnitude of the evoked calcium signal (dF/F0; columns 1–4) or estimated the action potential numbers calculated via deconvolution (columns 5–6) and used a mixed-effects model to compare across genotypes, using a normal distribution fitted with a logarithmic link function. Note that we included in this analysis only GCs activated under each specific condition (‘Responds to each stim’); that is we quantified how many spikes occurred per active cell. We additionally analyzed the data including all GCs activated under highest amplitude PP stimulation (‘Responds to max stim’; Figure 2—figure supplement 3) to quantify the total activation of the population: that is how many spikes occurred, averaged across cells. We saw enhanced activation at the early postnatal timepoint for both genotypes (Figure 2, columns 1–2), with only a subtle difference between genotypes. Consistent with the known increase in sparsity of GC activation that occurs with development (Yu et al., 2013), the wild-type activity pattern was markedly decreased at the young adult relative to the early postnatal timepoint (Figure 2, columns 3–6). However, the Scn1a+/- activation pattern remained markedly enhanced at the young adult relative to the early postnatal timepoint, resulting in a robust, statistically significant difference between genotypes at this later timepoint. For instance, proportional Scn1a+/- GC activation at the young adult timepoint was over threefold larger than that of wild-type (0.51 ± 0.08 vs 0.16 ± 0.00; p < 0.001) in response to 4 × 400 µA pulses (subset of data in Figure 2A 4), and dF/F0 was increased by ~130% (p < 0.001; Figure 2D 4). Hence, there was a profound abnormality of Scn1a+/- DG circuit function at the young adult timepoint, with a markedly higher proportion of GCs activated by PP input, as well as an increase in the number of action potentials fired in those GCs, versus age-matched wild-type controls, across stimulation paradigms and a broad range of intensities. Mechanisms of dentate gyrus circuit dysfunction in young adult Scn1a+/- mice Increased PP activation of GCs could be due to increased intrinsic GC excitability, dysfunction of feed-forward inhibition, or increased synaptic excitatory drive onto GCs. To investigate the mechanism of DG circuit dysfunction in young adult Scn1a+/- mice, we first assessed the intrinsic excitability of GCs in Scn1a+/- mice versus controls. Whole-cell current-clamp recordings of GCs from Scn1a+/- mice and age-matched wild-type littermate controls (at both the early postnatal and young adult timepoints) showed no differences across a range of measures of intrinsic excitability, properties of individual action potentials (APs), and repetitive AP firing (Figure 2—figure supplement 4; Table 1). Table 1 Properties of early postnatal and young adult DG GCs from Scn1a+/- and wild-type mice. MeasurementEarly postnatalYoung adultGenotype variation (ANOVA)Scn1a+/-WTp-valueScn1a+/-WTp-valuep-Valuen cells (mice)19 (3)21 (3)16 (7)13 (4)Age (days)18 ± 219 ± 10.8855 ± 658 ± 20.75Vm (mV)–78.0 ± 1.6–77.6 ± 1.70.99–78.1 ± 1.6–78.8 ± 2.10.990.95Rm (MΩ)379 ± 19405 ± 430.89237 ± 20217 ± 190.970.89Time Constant8.3 ± 1.49.1 ± 1.20.967.2 ± 0.88.1 ± 1.50.960.51Rheobase (pA)65 ± 869 ± 80.99134 ± 16119 ± 190.860.66AP Threshold (mV)–36.2 ± 1.7–34.1 ± 2.80.89–37.2 ± 1.5–36.8 ± 1.60.990.56AP Amplitude (mV)80.9 ± 2.484.2 ± 1.80.6481.4 ± 1.683.8 ± 2.50.880.19AP Peak (mV)44.7 ± 1.446.1 ± 2.20.9444.2 ± 1.447.0 ± 2.10.770.27AP Rise Time (ms)0.54 ± 0.020.53 ± 0.030.970.57 ± 0.020.59 ± 0.020.940.87AP Halfwidth (ms)0.76 ± 0.020.80 ± 0.040.690.77 ± 0.030.86 ± 0.040.340.05AHP Amplitude (mV)15.2 ± 1.115.7 ± 0.70.9813.9 ± 1.013.8 ± 1.1> 0.990.81AHP time (ms)3.08 ± 0.342.99 ± 0.270.991.91 ± 0.142.16 ± 0.090.930.09Sag (percent)3.2 ± 0.34.0 ± 0.50.713.5 ± 0.63.8 ± 0.90.980.32Max instantaneous (Hz)140 ± 10160 ± 80.44184 ± 13161 ± 80.490.91Max steady-state (Hz)57 ± 365 ± 40.4166 ± 463 ± 60.970.50 Table 1—source data 1 Data summary for all DG GCs electrophysiological data. https://cdn.elifesciences.org/articles/69293/elife-69293-table1-data1-v2.xlsx Download elife-69293-table1-data1-v2.xlsx Single-cell electrophysiology data from acute brain slices and acutely dissociated neurons prepared from various Scn1a+/- mouse lines from multiple laboratories has repeatedly identified interneuron dysfunction (Cheah et al., 2012; Dutton et al., 2017; Ogiwara et al., 2007; Richards et al., 2018; Rubinstein et al., 2015; Tai et al., 2014), and in particular dysfunction of parvalbumin-expressing fast-spiking GABAergic interneurons (PV-INs) (Dutton et al., 2013; Rubinstein et al., 2015; Tai et al., 2014). Therefore, we next tested the intrinsic properties of DG PV-INs in Scn1a+/- mice versus wild-type littermate controls based on the involvement of these cells in DS pathogenesis as well as the fact that the GC response to PP input is known to be powerfully regulated by feedforward inhibition mediated by DG PV-INs (Ewell and Jones, 2010; Lee et al., 2016). We prepared acute brain slices from early postnatal and young adult Scn1a+/- mice and wild-type littermates expressing tdTomato (tdT) under Pvalb-specific Cre-dependent control, as described previously (Favero et al., 2018). PV-INs in DG were thus identified by endogenous tdT expression visualized with epifluorescence and characteristic location at the GCL:hilus border. Early postnatal Scn1a+/- PV-INs exhibited profoundly impaired firing in response to depolarizing current steps (Figure 3A–C). The findings at the young adult timepoint were more subtle: young adult Scn1a+/- PV INs fired normally at the onset of a depolarizing current step and reached identical maximal steady-state firing frequencies but exhibit gradual spike height accommodation that ultimately progresses to spike failure with prolonged and large-amplitude (~2.5-fold rheobase) current injections (Figure 3D–F). Comparing across timepoints, while PV-INs from young adult Scn1a+/- mice continue to exhibit a reduced maximal instantaneous firing frequency relative to age-matched wild-type (Figure 3G), there was normalization of maximal steady-state firing frequency as well as across multiple other metrics of intrinsic excitability and properties of individual action potentials (Figure 3H–J; Table 2). Table 2 Properties of early postnatal and young adult DG PV-INs from Scn1a+/- and wild-type mice. MeasurementEarly postnatalYoung adultGenotype variation (ANOVA)Scn1a+/-WTp-valueScn1a+/-WTp-valuep-Valuen cells (mice)13 (5)10 (5)18 (5)19 (4)Age (days)19 ± 119 ± 10.7269 ± 366 ± 20.56Vm (mV)–57.9 ± 1.7–61.5 ± 2.30.65–57.6 ± 1.8–57.2 ± 1.80.990.42Rm (MΩ)94 ± 1153 ± 60.007 (**)105 ± 6101 ± 60.970.005 (**)Time Constant5.3 ± 0.84.7 ± 1.00.916.4 ± 0.35.1 ± 0.30.260.10Rheobase (pA)555 ± 96802 ± 1200.10489 ± 35463 ± 340.990.10AP Threshold (mV)–35.7 ± 3.4–45.8 ± 3.10.04 (*)–50.4 ± 1.4–49.0 ± 1.60.960.07AP Amplitude (mV)64.2 ± 2.161.8 ± 4.20.9275.1 ± 1.776.9 ± 1.90.930.91AP Peak (mV)28.5 ± 3.416.1 ± 4.00.008 (**)24.7 ± 31.327.9 ± 1.30.690.05AP Rise Time (ms)0.63 ± 0.150.50 ± 0.070.610.67 ± 0.080.50 ± 0.050.410.06AP Halfwidth (ms)0.54 ± 0.030.43 ± 0.040.01 (*)0.35 ± 0.010.37 ± 0.010.860.048 (*)AHP Amplitude (mV)10.7 ± 1.57.2 ± 3.50.5811.4 ± 1.19.7 ± 1.20.850.14AHP time (ms)1.60 ± 0.111.12 ± 0.100.001 (**)0.98 ± 0.050.93 ± 0.030.920.0008 (***)Sag (percent)11.1 ± 1.726.4 ± 6.60.007 (**)9.0 ± 1.117.3 ± 2.30.100.0001 (***)Max instantaneous (Hz)181 ± 8277 ± 190.002 (**)237 ± 15297 ± 130.006 (**)< 0.0001 (****)Max steady-state (Hz)139 ± 12213 ± 200.006 (**)189 ± 11191 ± 120.990.008 (**) Table 2—source data 1 Data summary for all DG PV-IN electrophysiological data. https://cdn.elifesciences.org/articles/69293/elife-69293-table2-data1-v2.xlsx Download elife-69293-table2-data1-v2.xlsx Figure 3 Download asset Open asset Profound impairment of spike generation in DG PV-INs from early postnatal Scn1a+/- mice, with partial normalization by young adulthood. (A) Example current clamp recordings of DG PV-INs from WT (blue) and Scn1a+/- (red) mice at the juvenile timepoint, demonstrating early spike failure in Scn1a+/- PV-INs. Scale bar 20 mV / 100ms. For juvenile GC PV-INs, (B) current/frequency (I-f) plot and (C) I-F plot with current normalized to Rheobase for each cell. (D) Example current clamp recordings from WT (blue) and Scn1a+/- (red) DG PV-INs at the young adult timepoint, demonstrating progressive spike-height accommodation in Scn1a+/- PV-INs in response to a prolonged depolarizing current step. Scale bar as in A. For young adult GC PV-INs, (E) current/frequency (I-F) plot and (F) I-F plot with current normalized to Rheobase for each cell. Scn1a+/- PV-INs have significantly lower instantaneous firing frequency at both timepoints (G) but the steady state firing frequency normalizes by the young adult timepoint (H). Scn1a+/- PV-INs display larger input resistance (I) and action potential half-width (J) at the early postnatal timepoint only. For B–F, line and shaded areas represent mean and SEM, and bars indicate significance calculated using one-way ANOVA and post-hoc tests with Bonferroni correction. For G–J, significance is determined by Tukey’s multiple comparisons test. *, p < 0.05; **, p < 0.01; ***, p < 0.001. n and mouse ages are as per Table 2. See also Table 2—source data 1. Given near-normalization of PV-IN firing properties at the young adult timepoint, we reasoned that PV-IN dysfunction was unlikely to underlie the larger circuit deficit, which worsens, rather than improves, across development (Figure 2). To further confirm that the subtle deficits seen in individual PV-IN firing properties (Figure 2) were not responsible for the GC hyperactivation observed in young adult Scn1a+/- mice, we performed an additional set of experiments using Hm1a, a peptide toxin that acts as an Nav1.1-specific activator (Osteen et al., 2016) and has been shown to correct the abnormalities seen in PV-INs in Scn1a+/- mice (Goff and Goldberg, 2019). We found that bath-application of Hm1a to brain slices prepared from Scn1a+/- mice corrected the PV-IN deficits apparent in response to large and prolonged current injections, whereas Hm1a had no impact on firing of PV-INs from wild-type mice, consistent with prior literature (Richards et al., 2018; Figure 4A–D). However, Hm1a had no effect on the large-scale evoked activation of GCs – as measured via 2 P imaging – in either genotype (Figure 4E–H). This result suggests that the subtle identified deficits in Scn1a+/- PV-IN spike generation and impairment in repetitive firing does not underlie the observed large-scale circuit impairment at the young adult timepoint. Figure 4 Download asset Open asset Hm1a enhances fast-spiking discharge properties in DG PV-INs from Scn1a+/- (but not wild-typ" @default.
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W4220996879 date "2022-01-19" @default.
W4220996879 modified "2023-09-26" @default.
W4220996879 title "Author response: Corticohippocampal circuit dysfunction in a mouse model of Dravet syndrome" @default.
W4220996879 doi "https://doi.org/10.7554/elife.69293.sa2" @default.
W4220996879 hasPublicationYear "2022" @default.
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