Experiments were performed on 3–6 weeks (pre-symptomatic animals) or 4–6 months-old (after seizure onset) homozygous Syn II knockout (Syn II^−/−) mice generated by homologous recombination and age-matched C57BL/6J wild-type (WT) animals. All experiments were carried out in accordance with the guidelines established by the European Community Council (Directive 2010/63/EU of September 22nd, 2010) and were approved by the Italian Ministry of Health. Adequate measures were always taken to minimize animal pain or discomfort.

Mice were anesthetized with isofluran by inhalation, decapitated and the brain dissected out in ice cold cutting solution containing (mM): 87 NaCl, 25 NaHCO₃, 2.5 KCl, 1.25 NaH₂PO₄, 0.5 CaCl₂, 7 MgCl₂, 25 glucose, and 75 sucrose saturated with 95% O₂ and 5% CO₂. Horizontal, 400 μm thick, cortico-hippocampal slices were cut using a Microm HM 650 V vibratome equipped with a Microm CU 65 cooling unit (Thermo Fisher Scientific, Waltham MA, USA). Slices were cut at 2°C in the bath solution. After cutting, slices were left to recover for 45–60 min at 35°C and for another hour at room temperature in artificial cerebrospinal fluid (aCSF) containing in mM: 125 NaCl, 25 NaHCO₃, 25 glucose, 2.5 KCl, 1.25 NaH₂PO₄, 2 CaCl₂, and 1 MgCl₂ (bubbled with 95% O₂–5% CO₂). The same solution was used for the perfusion of slices during recordings.

Whole-cell recordings were performed with a Multiclamp 700B/Digidata1440A system (Molecular Devices, Sunnyvale, CA) using an upright BX51WI microscope (Olympus, Tokyo, Japan). For all experiments we used a high-K gluconate intracellular solution containing (in mM): 126 K gluconate, 4 NaCl, 1 MgSO₄, 0.02 CaCl₂, 0.1 BAPTA, 15 Glucose, 5 HEPES, 3 ATP, and 0.1 GTP. The pH was adjusted to 7.3 with KOH and osmolarity was adjusted to 290 mosmol/l using sucrose. Patch-pipette resistance was 3–6 MΩ when filled with intracellular solution. Somatic access resistance (R_a) was continuously monitored, and cells with unstable R_a (20% changes) or with values larger than 15 MΩ were excluded from the analysis.

Mossy cells were identified by their shape, which has a triangular appearance in an infrared differential interference contrast image, size (mossy cells are clearly larger that surrounding interneurons) and location (deep hilus). Mossy cells were also filled with AlexaFluor 568 (40 μM in the pipette solution) to make the specific complex spines (named thorny excrescences) on the proximal dendrites visible in epifluorescence imaging (see Figure 4C). For cells that could not be clearly visualized, electrophysiological features, such as firing frequency adaptation during positive current injection, reduced afterhyperpolarization, broad action potentials, and high frequency/large amplitude spontaneous excitatory postsynaptic currents (sEPSPs) in the presence of bicuculline/CGP 55845, were used to identify them. Intrinsic cell properties were calculated from recordings performed in the presence of 50 μM D-APV, 10 μM CNQX, 5 μM CGP 55845, and 30 μM bicuculline. For the recording of miniature excitatory postsynaptic currents (mEPSCs), aCSF cointaining 5 μM CGP 55845, 30 μM bicuculline, and 0.3 μM TTX was used. All experiments were performed at a holding potential (V_h) of -70 mV in the presence of 30 μM bicuculine and 5 μM CGP 55845 (all from Tocris Bioscience, Ellisville, MO).

Granule neurons were selected based on their oval shape and their middle position in the granule layer. Since dentate granule neurons can be in different stages of maturation we recorded only mature neurons in which R_m < 300 MΩ (Liu et al., 1998).

In experiments where extracellular stimulation was performed, a monopolar stimulation electrode (a glass pipette filled with aCSF) was placed in the hilus, close to the granule cell layer and connected to an external stimulator (A-M Systems, Sequim WA).

All data were acquired with Clampex and analyzed offline with Clampfit 10.2 (Molecular Devices, Sunnyvale CA, USA), MiniAnalysis (Synaptosoft, Decatur GA, USA), Excel and GraphPad.

In the current-clamp mode, 30 current steps lasting 1 s, starting from −100 pA in 10 pA increments, were applied to both granule and mossy cells. To precisely determine the resting membrane potential (V_m) value, this was continuously recorded for 1 min, plotted in a histogram, and fitted with a Gaussian curve. V_m was taken as the mean of the distribution. To determine the start (V_s) of an action potential (AP), phase plane plots were constructed and V_s was considered the voltage point where dV/dt exceeded 10 mV/ms. The AP amplitude was measured as the difference between the maximum voltage reached during the overshoot minus V_s. The afterhyperpolarization (AHP) value was considered as the point where dV/dt was equal to zero, after the repolarization phase. The rheobase was estimated as the smallest current step that elicited an AP. For calculating the input resistance (R_in), the steady state voltage values from the first step above and below zero current (−10 and +10 pA), plus the zero current trace were plotted against the injected current and fitted with a linear regression. R_in was taken as the slope of the fitting line. The coefficient of determination R² was always higher than 0.95.

For the analysis of mEPSCs, 30 s of each recording were imported into MiniAnalysis (Synaptosoft Inc., New Jersey). Events' peaks were identified manually because of the presence of multipeak events due to the high frequency of spontaneous events. Subsequently, the software automatically calculated amplitude, rise, and decay. Minimum amplitude threshold was set to 8 pA. For the analysis of mEPSC kinetics, a set of individual 40–60 events was chosen in which the rise and the decay phase were very clear. The events were all aligned at the point of 50% rise and averaged for further calculations. The decay phase was best fitted with a monoexponential equation on the 90–10% decay phase. The cumulative distributions of amplitudes and frequencies were constructed by pooling all values, and analyzed with a Kolmogorov–Smirnov test for distributions.

The APS-MEA, extensively described elsewhere (Ferrea et al., 2012), consists of a microelectrode array chip and an amplification system designed to provide simultaneous extracellular recordings from 4096 electrodes at a sampling rate of 7.7 kHz. Each square pixel measures 21 × 21 μm, and the array is integrated with an electrode pitch (center-to-center) of 42 μm. Pixels are arranged in a 64 × 64 array configuration, yielding an active area of 7.22 mm², with a pixel density of 567 pixels/mm². The three on-chip amplification stages provide a global gain of 60 dB, with a 0.1–5 kHz band-pass filter. This bandwidth is suitable for recording both slow LFP signals and fast APs. The acquisition is controlled by the BrainWave software (3Brain Gmbh, Switzerland). For stimulation experiments of the DG, a monopolar stimulation electrode placed on the medial perforant path and connected to an external stimulator (A-M Systems, Sequim WA) was used. Clear evoked responses separated from the stimulation artifact were obtained using stimulation intensities between 100 and 500 μA (for 30 μs). Large-scale field recordings were acquired using BrainWave and analyzed offline with MatlabR2010a, Clampfit 10.2 and ImageJ. For the analysis of the peak amplitude, 3 representative channels (named “pixels”) from the molecular layer and 3 from the hilus were exported in Matlab (MathWorks, http://www.mathworks.it/) and then imported in Clampfit for analysis. The maximum amplitude of the evoked events on each channel was measured, averaged (10 events/experiment) and normalized to the control amplitude. For the analysis of the mean amplitude for the entire area, an image at the point of highest response for the respective area was imported in ImageJ and the mean intensity was calculated after normalization of the maximum intensity to the maximum voltage of the same response.

WT and Syn II^−/− mice were deeply anesthetized with 20% urethane (0.1 ml/10 gm) and perfused transcardially with 0.1 M phosphate buffer containing 4% paraformaldehyde (pH 7.4). Brains were subsequently postfixed overnight in paraformaldehyde solution, and then washed in PBS, infiltrated with a 30% sucrose PBS solution for cryoprotection and frozen in OCT. Ten μm thick horizontal sections were cut using a Leica CM3050 S Cryostat and collected on SuperFrost slides. Sections were incubated 1 h in blocking buffer (2% NGS, 1% BSA, 0.1% CFG, 0.1% Triton X-100, 0.05% Tween in PBS), then for 2 h in blocking buffer containing primary antibodies. After several washes in PBS, sections were incubated with blocking buffer containing fluorochrome-conjugated secondary antibodies (Invitrogen, 1:500), washed and mounted using Prolong Gold antifade reagent with DAPI staining (Invitrogen) for fluorescence microscopy observation on a Leica SP5 confocal laser scanning fluorescence microscope. The mean fluorescence intensity ratio between Syn I or Syn II and calretinin were calculated with the software ImageJ. Antibodies used were: Anti-Synapsin 1 (Mouse, 1:500, SYSY #106 001; Rabbit, 1:200 G-177), Anti-Synapsin 2 (Mouse, 1:200, clone 19.21), Anti-panSynapsins (Rabbit, 1:500 G143), Anti-Calretinin (Guinea Pig, 1:1000, SYSY# 214 104).

Acute slices were fixed by immersion in 1.3% glutaraldehyde in 66 mM sodium cacodylate buffer. Subsequently, slices were post-fixed with 1% OsO4 in 1.5% K₄Fe(CN)₆ in 0.1 M sodium cacodylate, en bloc stained with 0.5% uranyl acetate, dehydrated through a series of graded ethanol solutions, washed in propylene oxide and flat embedded in Embed 812 between two Aclar sheets. After 48 h of polymerization at 60°C, a small region corresponding to the DG was excised, glued with cyanoacrylate glue on blocks of resin and cut with a Leica EM UC6 ultramicrotome. Ultrathin sections (thickness: 70–90 nm) were collected on Formvar carbon coated copper grids. Grids were observed in a JEOL JEM-1011 microscope operating at 100 kV using an ORIUS SC1000 CCD camera (Gatan). Total and docked SV densities were calculated using the software ImageJ.

All data are expressed as means ± s.e.m. All statistics were performed with GraphPad Prism. For comparison between WT vs. Syn II^−/− experiments, two-tailed unpaired Student's t-test was used. For multiple comparisons, one or two-way analysis of variance (ANOVA) with Bonferroni post-hoc test or Kruskal-Wallis test followed by Dunn's post-hoc test was used, depending on the type of data. The level of significance was set at p < 0.05.

Our first step was to investigate the response of different areas of the DG to the stimulation of the perforant path in horizontal brain slices containing the hippocampus and rhinal cortices from pre-symptomatic (3–6 weeks) Syn II^−/− mice. To this aim, we employed a high-resolution Active Pixel Sensor microelectrode array system (APS-MEA, 4096 electrodes: see Materials and Methods) (Ferrea et al., 2012). Field postsynaptic potentials (fPSPs), evoked by the stimulation of the perforant path with an extracellular electrode, were detected in the granule layer of the DG and they further propagated to the hilus (Figure 1A). The mean amplitude of the response, calculated over the entire activated region in the granule cell layer, was similar in WT and Syn II^−/− slices (153.7 ± 15.2 μV for WT vs. 135.6 ± 10.6 μV for Syn II^−/−, n = 13 slices for each genotype; two-tailed unpaired Student's t-test, p = 0.342) (Figures 1B,D). On the other hand, the mean amplitude of the response in the hilar region was significantly reduced in Syn II^−/− slices (143.0 ± 19.4 μV for WT vs. 87.13 ± 10.7 μV for Syn II^−/−, n = 12 slices for each genotype; two-tailed unpaired Student's t-test, p = 0.013) (Figures 1C,D). The stimulus amplitude was set to 400–500 μA based on previous input-output curves performed for 3 selected electrodes in each of the two regions. At all stimulation intensities, a reduced amplitude of the responses in the Syn II^−/− hilar region with respect to the WT was observed (Figures 1E,F). The recorded signal on several neighboring electrodes on the APS-MEA correlated with the fine anatomy of the DG and its polarity corresponded to current sinks in the dendritic-granule layer (negative) and to current sources in the hilus (positive) (Figure 1D). Moreover, the propagation time of the evoked fPSP, measured between the peaks of the response from one representative electrode in the granule and hilar regions, was significantly longer in Syn II^−/− than in WT slices (2.8 ± 0.78 ms for WT vs. 5.6 ± 0.59 ms for Syn II^−/−, n = 6/5 slices; two-tailed unpaired Student's t-test, p = 0.032) (Figure 1G), suggesting the presence of functional impairments of the hilar region.

The hilus of the DG contains hilar mossy cells and inhibitory interneurons, whose activity modulate the excitability of granule neurons (Scharfman and Myers, 2012), by forming a regulatory loop. We have previously shown that the plasticity of the young Syn II^−/− excitatory and inhibitory synapses upon train stimulation of the perforant path does not significantly differ from that of WT synapses (Medrihan et al., 2013). Since we observed a reduced signal in the hippocampal hilus, we reasoned that a sustained train of stimuli would reveal possible dysfunctions of the granule layer-hilus network function. Thus, we stimulated the perforant path with a train of 20 Hz for 5 s and measured the ratio between the first and the last evoked fPSP response of the train over the entire granule cell layer area (Figures 2A,B). To our surprise, the depression induced by a train of stimuli was significantly higher in WT than in Syn II^−/− slices (0.45 ± 0.03 for WT vs. 0.73 ± 0.08 for Syn II^−/−, n = 4/5 slices; two-tailed unpaired Student's t-test, p = 0.033) (Figure 2C). Based on previous results showing that perforant path plasticity is unaltered in Syn II^−/−, these results point toward a reduced feedback inhibition of the granule cell layer.

Next, we confirmed the presence of Syn II in the DG. Acute brain slices from 6 weeks old WT mice were immunolabeled for endogenous Syns I and II and for the mossy cell marker calretinin (Blasco-Ibanez and Freund, 1997) (Figure 3A). Since in some species calretinin is also a marker for GABAergic interneurons (Scharfman and Myers, 2012), we co-immunolabeled for GABA and noticed that calretinin immunoreactive hilar cells are negative for GABA (data not shown), suggesting that they are indeed hilar mossy cells. Both Syn I and Syn II puncta were abundantly observed in the hilar region of the DG, where mossy fiber terminals are localized. Remarkably, Syn II, but not Syn I, positive puncta were present in high density in the inner molecular layer, colocalizing with the hilar cell terminals. We therefore measured the mean intensity ratio between either Syn isoform and calretinin in the inner molecular layer of the DG (0.09 ± 0.007 for Syn I/calretinin and 0.61 ± 0.11 for Syn II/calretinin, n = 3 mice; two-tailed unpaired Student's t-test, p = 0.034) and in the hilar region (0.195 ± 0.0994 for Syn I/calretinin and 0.229 ± 0.1157 for Syn II/calretinin, n = 3 mice; two-tailed unpaired Student's t-test, p = 0.837) (Figures 3B–D). To further prove the specific presence of Syn II at mossy cell terminals, we stained brain slices from Syn I^−/− and Syn II^−/− mice respectively for calretinin (red) and all Syn isoforms (pan-synapsin antibody, green). While in the Syn I^−/− slices the pan-synapsin staining is visible in the IML, it was completely absent in the same region of the Syn II^−/− slices (Figure 3E, bottom panels).

Since we noticed an impaired function of the hilar region of the DG (Figure 1) and Syn II is highly expressed at the mossy fiber terminals (Figure 3), we next investigated the synaptic input received by hilar mossy cells from granule neurons. First, we used electron microscopy to morphologically evaluate the number and spatial distribution of SVs within the mossy fiber terminals in the hilar region of the DG of pre-symptomatic Syn II^−/− and WT mice (Figure 4A). The total density of SVs in Syn II^−/− synapses was significantly lower than that of WT synapses (198.7 ± 13.7, n = 44 terminals/3 mice and 89.27 ± 10.8 SVs/μm², n = 51 terminals/3 mice, for WT and Syn II^−/− respectively; two-tailed unpaired Student's t-test, p = 0.0003; Figures 4A,B). However, the number of docked SVs did not differ between genotypes (9.42 ± 0.6, n = 19 terminals/3 mice and 9.70 ± 0.7 SVs/μm, n = 19 terminals/3 mice, for WT and Syn II^−/− respectively; two-tailed unpaired Student's t-test, p = 0.782) (Figures 4A,B), as previously reported for Syn deletions at various synapses (Gitler et al., 2004; Medrihan et al., 2013).

To functionally analyze these synapses, we recorded mEPSCs from pre-symptomatic Syn II^−/− hilar mossy cells after blocking GABA receptors and Na⁺ channels with bicuculline (30 μM), CGP55845 (5 μM) and TTX (0.3 μM). To distinguish mossy cells from the surrounding hilar neurons, we filled the patch pipette with AlexaFluor568-containing intracellular solution. This enabled the detection of moss-resembling spines (called “thorny excrescences”) present on the proximal dendrites of these cells (Figure 4C). As reviewed before (Henze and Buzsaki, 2007; Scharfman and Myers, 2012), hilar mossy cells are highly excitable, receiving massive input from the mossy fibers, and their mEPSCs have an unusually large amplitude and frequency in comparison with other central synapses (Figure 4D). Both the amplitude and the frequency distributions of mEPSCs were significantly shifted toward smaller values in Syn II^−/− hilar mossy neurons (n = 11 neurons/7 mice for WT and 6 neurons/3 mice for Syn II^−/−; Kolmogorov–Smirnov test, p < 0.001) (Figure 4E). Such a leftward shift of frequencies indicates the presence of a presynaptic involvement. Although the distribution of mEPSC amplitudes in Syn II^−/− was also left-shifted, it was not associated with a change in the kinetic parameters of the response (rise-time 10–90%: 0.83 ± 0.04 vs. 0.71 ± 0.05 ms, p = 0.168; decay τ: 4.39 ± 0.35 vs. 3.87 ± 0.23 ms, p = 0.374; two-tailed unpaired Student's t-test) (Figure 4F), suggesting an overall decreased excitatory presynaptic input on Syn II^−/− hilar mossy cells from young mice.

Extracellular fPSPs are the summation of a series of events, notably synaptic activity and synchronous firing of APs by groups of neurons (Buzsaki et al., 2012). Thus, in the next experiment, we evaluated the firing rate of hilar mossy neurons from pre-symptomatic Syn II^−/− in the current clamp configuration. In the presence of specific antagonists that fully block synaptic activity, mossy cells were injected with 30 current steps, lasting 1 s and ranging from −100 to +200 pA, in 10 pA increments (Figures 5A,B). The firing rate of mossy neurons was lower in Syn II^−/− slices with respect to WT recordings (Figure 5B) and was accompanied by a significant increase in the rheobase (45.0 ± 6.7, n = 11 neurons/5 mice for WT vs. 83.3 ± 8.8 for Syn II^−/−, n = 6 neurons/4 mice; two-tailed unpaired Student's t-test, p = 0.003) (Figure 5C, left). Input resistance, a parameter correlated with the firing rate, was also significantly reduced in Syn II^−/− (396.0 ± 45.2 MΩ, n = 11 neurons/5 mice for WT vs. 248.7 ± 23.2 for Syn II^−/−, n = 6 neurons/4 mice; two-tailed unpaired Student's t-test, p = 0.031) (Figure 5C, right). On the contrary, recording from granule cells revealed no differences in the firing rates of WT and Syn II^−/− neurons (Figures 5D,E), with no genotype-dependent difference in either input resistance or rheobase (data not shown). Other intrinsic membrane properties (resting and threshold potential, AP amplitude, half-amplitude width, after-hyperpolarization current) were similar for the two genotypes in both mossy and granule neurons (data not shown). These results indicate that the reduced fPSPs in the hilar region of pre-symptomatic Syn II^−/− mice (Figure 1) are associated with reduced excitability of Syn II^−/− mossy cells, but not granule cells.

To verify if the reduced excitability of hilar mossy cells in pre-symptomatic Syn II^−/− slices persists after the initiation of epileptic seizures in these mice, we repeated the experiments from Figures 4, 5 on adult (4–6 months old) Syn II^−/− mouse slices. As in pre-symptomatic mice, both the amplitude and the frequency distributions of mEPSCs were significantly shifted toward lower values in Syn II^−/− hilar mossy neurons (n = 4 neurons/3 mice for WT and 6 neurons/3 mice for Syn II^−/−; Kolmogorov–Smirnov test, p < 0.001) (Figures 6A,B). The smaller amplitude distribution of Syn II^−/− cells was not accompanied by any change in the kinetic parameters of the response with respect to the WT (rise-time 10–90%: 1.36 ± 0.2 vs. 1.08 ± 0.1 ms, p = 0.296; decay τ: 5.58 ± 1.1 vs. 5.63 ± 0.6 ms, p = 0.967; two-tailed unpaired Student's t-test) (Figure 6C). Moreover, the firing rate of mossy cells was lower in adult Syn II^−/− (Figures 6D,E), with a significant increase in rheobase (60.0 ± 5.7, n = 3 neurons/3 mice for WT vs. 85.0 ± 8.6 pA for Syn II^−/−, n = 8 neurons/3 mice; two-tailed unpaired Student's t-test, p = 0.043) (Figure 6F, left) and a decrease in input resistance (422.0 ± 12.5 MΩ, n = 3 neurons/3 mice for WT vs. 269.0 ± 38.0 for Syn II^−/−, n = 8 neurons/3 mice; two-tailed unpaired Student's t-test, p = 0.042) (Figure 6F, right). These results show that the cellular phenotype of Syn II^−/− mice appears long before the appearance of an overt epileptic phenotype.

The axons of hilar mossy cells project into the inner molecular layer of the DG, where they make excitatory synapses directly with granule cells, or with GABA interneurons, leading to disynaptic inhibition of granule cells (Scharfman and Myers, 2012; Jinde et al., 2013). Since Syn II is abundantly and specifically expressed in the mossy cell terminals of the inner molecular layer (Figure 3), the next step was to evaluate the net effect of the mossy cell output on granule cell activity. To this aim, we patched granule neurons from the granule cell layer and stimulated the axons of mossy cells in the region below the granule layer (Figure 7A). When granule neurons are voltage-clamped at -80 mV, close to the reversal potential of Cl⁻, the stimulation should result in an evoked excitatory inward response non-contaminated by inhibition. On the contrary, when the clamped voltage is shifted to 0 mV, the Cl⁻ drive will be predominant, and the stimulation should elicit a net inhibitory, outward response representing the fast-forward inhibition resulting from the intermediate activation of GABA interneurons (Figure 7B). Single stimulation did not reveal any difference between the amplitude of both eEPSC and eIPSCs in WT and pre-symptomatic and adult Syn II^−/− slices (n = 16 neurons/6 mice for WT, 10 neurons/4 mice for young Syn II^−/− and 13 neurons/3 mice for adult Syn II^−/−; One-Way ANOVA followed by the Bonferroni's multiple comparison test, p = 0.344 and 0.751 for eEPSCs and eIPSC, respectively) (Figures 7C,D). Instead, the application of a 40 Hz tetanic stimulation revealed that depression was significantly increased at inhibitory synapses in both young and adult Syn II^−/− granule neurons (Figure 7F), while it was similar between genotypes at excitatory synapses (Figure 7E). We quantified this effect by measuring the ratio between the evoked excitatory and inhibitory responses (E/I ratio) on the same granule cell. Single pulse stimulation of the mossy cell axon produced postsynaptic inhibitory and excitatory currents whose ratio was similar between genotypes (p = 0.938; One-Way ANOVA followed by the Bonferroni's multiple comparison test) (Figure 7G, left). However, when the E/I ratio was measured for the last 10 responses in the train, it was significantly increased in both pre-symptomatic and symptomatic Syn II^−/− slices, with a decrease of inhibitory responses (E/I ratio = 0.3 ± 0.04 for WT, n = 6 neurons/3 mice; 2.6 ± 0.6 for young Syn II^−/−, n = 7 neurons/4 mice; 1.8 ± 0.6 for adult Syn II^−/−, n = 6 neurons/3 mice; p = 0.0011; Kruskal–Wallis test followed by the Dunn's multiple comparison test) (Figure 7G, right). This change in the ratio between excitation and inhibition may be responsible for the hyperexcitability of the DG under conditions of sustained high frequency synaptic input, as seen in Figure 2.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.