DPP6 regulation of dendritic morphogenesis impacts hippocampal synaptic development

DPP6 is important for filopodia formation and stability

We found that labeled-DPP6 localizes to dendritic filopodia and spines (Supplementary Fig. S1), suggesting a potential role in synaptic development. To investigate this possibility, we first measured dendritic filopodia formation in hippocampal neurons cultured from P0 WT and DPP6 knockout (DPP6-KO) mice. The density of immature dendritic filopodia at DIV3 in cultured hippocampal neurons obtained from DPP6-KO mice was significantly decreased, to about 45% of that found for WT (Figure 1A–B; p<0.05, unpaired Student’s t-test). In a second experiment, we expressed either GFP or GFP-tagged DPP6 (DPP6-GFP) in P0 DPP6-KO mouse hippocampal neurons and counted the number of immature dendritic filopodia at DIV3. DPP6-GFP expressing neurons showed a significantly increased density of immature dendritic filopodia compared to the GFP control demonstrating a specific role for DPP6 in regulating filopodia number (Figure 1C–D; p<0.05, unpaired Student’s t-test). We also measured the density of filopodia in identified axons (tau-1 positive) in cultured neurons at DIV3. Unlike our results for immature dendritic filopodia, we found no significant difference in the number of axonal filopodia between DPP6-KO and WT neurons (WT=1.81±0.16 filopodia/20 μm axon length, n=64; DPP6-KO=1.40±0.13 filopodia/20 μm axon length, n=85, p>0.05, unpaired Student’s t-test). Although the density of filopodia in identified dendrites of DPP6-KO cultured neurons is significantly decreased compared with WT, we did not find a significant difference between the density of dendritic branches of DIV3 WT and DPP6-KO cultured neurons (WT=0.08±0.02 branches/20 μm, n=44; DPP6-KO=0.07±0.01 branches/20 μm, n=77, p>0.05, unpaired Student’s t-test).

These results, showing a role for DPP6 in regulating filopodia number, could indicate either an effect on filopodia formation, the stability of filopodia once formed, or both. To determine this, we performed live imaging experiments to monitor filopodia in DIV3 hippocampal neurons from WT and DPP6-KO mice (Figure 2). First we measured the mean lifetime of existing filopodia over 1h at 30 sec intervals. Figure 2A–B shows that filopodia in WT neurons are very stable, with ~ 75% of 78 total filopodia lasting the full 60 min of time-lapse recording. In DPP6-KO neurons, however, only ~57% of 80 total filopodia lasted the full hour. Next, we measured the effect of transfecting either GFP or DPP6-GFP into DPP6-KO P0 hippocampal neurons. Results showed that filopodia in DPP6-GFP expressing neurons were more stable (~70% survival for 1h) compared to the GFP control (~59% survival, Figure 2C–D). This rescue of filopodia stability in DPP6-KO neurons to WT levels by DPP6-GFP expression suggests a fundamental role for the DPP6 protein in maintaining filopodia in developing hippocampal neurons.

In these live imaging studies, we also counted the number of new filopodia initiated during the 1h recording (Figure 2E–F). We found DPP6-KO neurons to produce about 60% fewer filopodia than WT (p<0.05, unpaired Student’s t-test). In DPP6-GFP expressing DPP6-KO hippocampal neurons, we found that DPP6-GFP increased the initiation of filopodia about 3-fold compared to the GFP control, nearly restoring the rate of filopodia formation to WT levels (Figure 2G–H; p<0.05, unpaired Student’s t-test). Taken together, the live imaging measurements suggest that the decreased density of filopodia in DPP6-KO hippocampal neurons is due to a combination of both decreased stability and initiation of filopodia by DPP6.

Spine density is reduced in developing DPP6-KO neurons

Since filopodia can transform into spines ²², we looked for a corresponding effect on spine number at a later developmental stage. To detect and count spines in dendrites, cultures were immunostained for MAP2 to highlight the dendritic arbor, and PSD-95 to visualize spines at post-synaptic sites (see Methods). Figure 3A shows images of cultured WT and DPP6-KO hippocampal neurons at DIV14. The density of the spines (number per 20 μm length) was reduced in DPP6-KO neurons ~30% compared to those from WT mice (Figure 3B; p<0.05, unpaired Student’s t-test). As found for dendritic filopodia (Figures 1 and 2), expression of DPP6-GFP in DPP6-KO mouse P0 hippocampal neurons enhanced spine density compared with GFP controls when measured at DIV 14 (Figure 3C–D; p<0.05, unpaired Student’s t-test). To determine the functional impact of spine decrease in DPP6-KO neurons, we measured miniature excitatory synaptic currents (mEPSCs) in DIV14 cultured WT and DPP6-KO neurons. While no differences between the two groups were found for mEPSC amplitude, mEPSC frequency was significantly decreased in DPP6-KO neurons (Figure 3E–G, p<0.05, unpaired Student’s t-test). Therefore, in addition to regulating filopodia number early in development, DPP6 subsequently impacts spine formation and synapse number in hippocampal neurons.

Developmental effects of DPP6 knockdown

To see if DPP6’s role in regulating dendritic spine formation impacted synaptogenesis, we used RNAi against DPP6 in hippocampal neuron cultures over a period of rapid synapse formation and a decrease in dendritic filopodia density. E18 rat hippocampal neurons were transfected with either RNAi targeting DPP6 (“siDPP6”) ²³ or control RNAi (siCtrl) at plating. As a control, we measured the effect of siDPP6 expression on filopodia number in neurons cultured from DPP6-KO mice. Neither siDPP6 nor siCtrl affected the number of filopodia in DPP6-KO neurons (DPP6-KO+sictrl=1.49±0.24 filopodia/20 μm, n=12; DPP6-KO+siDPP6=1.35±0.26 filopodia/20 μm, n=14, p>0.05, unpaired Student’s t-test).

At DIV7, 14 and 21 cultured neurons were fixed and immunostained with MAP2 to highlight the dendritic arbor, and with phalloidin to highlight actin-based protrusions. To measure the density of synapses, we co-labeled neurons with the presynaptic marker synaptophysin and the post-synapse marker PSD-95. Puncta where synaptophysin and PSD-95 labeling overlapped were counted as synapses. To determine the effect of DPP6 knockdown on the formation of filopodia, spines and synaptogenesis, we detected and counted the density of filopodia, spines and synapses in maturing WT hippocampal neurons and transfected neurons either with siCtrl-GFP or siDPP6-GFP. WT hippocampal neuronal cultures displayed a progressive decrease in the density of filopodia between DIV7 and DIV21, and a corresponding increase in the density of spines and synapses during maturation (Supplementary Fig. S2). In siDPP6 expressing neurons, we found the density of filopodia was significantly reduced by ~42% at DIV7 and ~30% at DIV14 compared to siCtrl (Figure 4A–B,E; p<0.05, unpaired Student’s t-test). The number of filopodia was negligible in both groups by DIV21 (Figure 4C–E). We also found siDPP6 to cause a corresponding decrease in spine density by ~40% at DIV14, and ~50% at DIV21 compared to siCtrl (Figure 4B–C,F; p<0.05, unpaired Student’s t-test) while the density of synapses was reduced by ~20% at DIV14 and 40% in DIV21 neurons expressing siDPP6 compared to siCtrl (Figure 4C,G; p<0.05, unpaired Student’s t-test). Accordingly mEPSC frequency was reduced in siDPP6 expressing neurons compared to control (siCtrl=105.7±28.1 events/min, n=20; siDPP6=43.3±9.0 events/min, n=17, p<0.05, unpaired Student’s t-test). No difference in mEPSC amplitude was found however (siCtrl=15.4±1.3 pA, n=20; siDPP6=13.8±1.5 pA, n=17, p>0.05, unpaired Student’s t-test). Moreover, expression of DPP6-GFP in mature DPP6-KO neurons increased the number of spines compared to control DPP6-KO neurons expressing only GFP controls (DPP6-KO+GFP=5.26±0.23 spines/20 μm, n=45; DPP6-KO+DPP6-GFP=7.11±0.31 spines/20 μm, n=38, p>0.05, unpaired Student’s t-test). These results show that DPP6 plays an important role in regulating the development of synapses in hippocampal neurons.

DPP6 effects are independent of Kv4.2

DPP6 has an important role in regulating Kv4.2 dendritic expression and function ¹³. To see if the effects of DPP6 loss on synaptic development involved its role as a Kv4.2 auxiliary subunit, we repeated the above experiments using DIV3 Kv4.2-KO cultured neurons. However, unlike for DPP6-KO neurons, results showed that the density of filopodia in axon and immature dendrites are not significantly different between Kv4.2-KO and WT neurons (Figure 5; for dendritic filopodia, WT=2.33±0.24 filopodia/20 μm, n=44; Kv4.2-KO=2.11±0.19 filopodia/20 μm, n=45, p>0.05; for filopodia in axon, WT=1.81±0.16 filopodia/20 μm, n=64; Kv4.2-KO=1.87±0.32 filopodia/20 μm, n=58, p>0.05, unpaired Student’s t-tests). We also observed no differences in the number of immature dendritic branches (WT=0.08±0.02 branches/20 μm, n=44; Kv4.2-KO=0.09±0.01branches/20 μm, n=45, p>0.05, unpaired Student’s t-test). At DIV14, we found that the number of spines are unchanged in Kv4.2-KO neurons compared with WT (WT=7.66±0.29 spines/20 μm, n=49; Kv4.2-KO=6.88±0.27 spines/20 μm, n=47, p>0.05, unpaired Student’s t-test) and neither mEPSC frequency (WT=58.2±11.8 events/min, n=25; Kv4.2-KO=43.3±8.9 events/min, n=25, p>0.05, unpaired Student’s t-test) nor amplitude (WT=16.4±0.9 pA, n=25; Kv4.2-KO=15.7±1.2 pA, n=25, p>0.05, unpaired Student’s t-test) differed. Therefore, we conclude that the effect of DPP6 on filopodia formation and stability occurs independent of its role as a Kv4.2 auxiliary subunit.

Consequences of DPP6-deficiency in adult neurons

To determine if the developmental effects described above are functionally relevant into adulthood, we used biocytin staining to isolate single CA1 pyramidal neurons from both 8 week old WT and DPP6-KO mice. Compared with WT controls, pyramidal cells from adult DPP6-KO mice exhibited a significant reduction in spine density in both apical and basal dendritic locations (Figure 6A–B). When the data were subdivided into distinct parts of the apical dendrite, it was apparent that DPP6-KO mouse spine density decreased dramatically (~52%) in distal dendrites (~240 μm) compared with WT (n=10 for DPP6-KO, n=10 for WT, p<0.05, unpaired Student’s t-test), whereas only a small reduction (~15%) in spine number was observed in proximal dendrites (~40 μm, n=10 for DPP6-KO, n=10 for WT).

To determine if this loss of spines leads to functional impairments, we performed electrophysiological recordings from WT and DPP6-KO adult acute hippocampal slices. Genetic loss of DPP6 resulted in a significant decrease of mEPSC frequency in both P14 young mice (Figure 6C–D; WT n=8; DPP6-KO n=7; p<0.05, unpaired Student’s t-test) and adult mice compared with WT (Figure 6C–D; WT n=6; DPP6-KO n=7; p<0.05, unpaired Student’s t-test). mEPSC amplitude was not significantly affected by DPP6 loss at either developmental stage (Figure C–D; P14 amplitude for WT n=8, DPP6-KO n=7; Adult amplitude for WT n=6, DPP6-KO n=7; p>0.1). No differences were found between genotypes in paired-pulse facilitation experiments (Supplementary Fig. S3) indicating a postsynaptic effect of DPP6 loss.

Taken together, the reduction of mEPSC frequency but not amplitude, along with the lack of PPF change suggests that DPP6-KO mice have fewer functional excitatory synapses in CA1 neuron dendrites. We therefore investigated dendritic morphology in biocytin stained hippocampal CA1 pyramidal neurons in both WT and DPP6-KO mice (Figure 6E–G). Sholl analysis revealed a simpler dendritic branching pattern in neurons from DPP6-KO mice compared to WT controls (Figure 6F–G). We found in DPP6-KO mice, less dendritic intersections at locations greater than 40 μm from the soma throughout the dendritic tree compared to WT (Figure 6G; n=10 for DPP6, n=10 for WT, p<0.05, unpaired Student’s t-test). In DPP6-KO neurons, apical dendrite trunk length was not different from WT (WT=232.18 μm, DPP6-KO=225.1 μm, n=10, p>0.05), but the total apical dendritic length of DPP6-KO is significantly decreased (WT=781.6 μm, DPP6=488.6 μm, n=10, p<0.05, unpaired Student’s t-test). The decrease in functional excitatory synapses in DPP6-KO CA1 neurons compared with WT may then be accounted by their sparse dendritic arbor as well as their reduced spine density per unit length of remaining apical dendrites. This finding differs from that reported above for DIV3 DPP6-KO neurons where we did not find a difference from WT in the number of dendritic branches at this young stage, indicating a developmental impact of DPP6 on branch complexity. Above (Figure 5), we determined that DPP6 acts independent of its role as a Kv4.2 auxiliary subunit in developing neurons, as filopodia formation and stability were not affected in Kv4.2-KO neurons. Consequently, in adult mice we also found no effect of Kv4.2 KO on dendritic branching compared to WT (Supplementary Fig. S4).

Mechanism of DPP6 effects on filopodia

Our results indicate that DPP6 plays a role in regulation of synaptic development and function including filopodia formation and stability. Protrusion of filopodia involves the reorganization of the actin cytoskeleton. In begin to understand the molecular mechanisms underlying DPP6’s effects on synaptic development, we immunopreciptated DPP6 from lysates of hippocampal neurons to identify DPP6 interacting proteins (Figure 7A). Mass spectrometry (MS) of binds that were immunoprecipated from WT but not DPP6-KO neurons led to the identification of Myosin-X as a DPP6 interacting proteins. Myosin-X is an unconventional myosin known to have important roles in the formation and elongation of filopodia ²⁴. Myosin-X is also thought to transport cargo molecules within filopodia and provides a motor-base link between integrins and the cytoskeleton ²⁵.

For further confirmation a DPP6-Myosin-X interaction, we found that endogenous DPP6 and Myosin-X co-immunopreciptate in lysates from mouse brain (Figure 7B). DPP6 and Myosin-X also co-immunopreciptated in when transfected into COS7 cells (Figure 7C). Since one of the most intriguing features of Myosin-X is its localization at the tips of filopodia ^{26, 27}, we also tested if DPP6 is present in this location. As shown in Figure 8D, DPP6 and Myosin-X are both present at the tips of filopodia (Figure 7D). HEK cells, co-expressing the membrane protein marker, membrane-mCherry, along with DPP6-GFP showed clear co-localization suggesting membrane expression of DPP6-GFP (Supplementary Fig. S5).

DPP6 induces filopodia formation through cell adhesion

As with other DPPs the extracellular domain of DPP6 consists of a β propeller and an α/β hydrolase domain ¹⁵. β propellers provide excellent platforms for protein–protein interactions, and in DPPIV (also termed CD26) the extracellular domain binds to components of the extracellular matrix like collagen, fibronectin, and integrin β1 ^{16, 17, 28}. DPP6 has approximately 50% similarity to CD26 ²¹ and also has a cysteine-rich domain that may bind to components of the ECM function in cell adhesion. We investigated the effects of DPP6 transfection on adhesion of COS7 cells to the culture substratum poly-L-lysine. We found that COS7 co-expressed with DPP6 enhanced cell adhesion about 2-fold compared to the control cells transfected with vector only (Figure 8 A–B; p<0.05, unpaired Student’s t-test), indicating that DPP6 promotes cell adhesion to the substratum.

Integrins form a large family of heterodimeric (α/β subunits) membrane-spanning ECM receptors ^{29, 30} which bind to an arginine-glycine-aspartate (RGD) motif found in many ECM proteins. Since this short peptide sequence is able to interact with integrin receptors on the cell surface by itself, the RGD motif has frequently been introduced into various polymeric materials to enhance their cell adhesive abilities for use in tissue engineering. To determine whether DPP6 plays a role in cell adhesion by binding to RGD motifs in ECM proteins, we mimicked the effects of ECM on integrin signaling in cultured hippocampal neurons by applying an RGD-containing peptide and then examined its effects on filopodia formation. Cultured P0 DPP6-KO hippocampal neurons expressing either GFP or DPP6-GFP were treated at DIV3 with either control RAD or RGD-containing peptides (250 μM, 60 min). The neurons were then fixed and their actin filaments visualized with TRITC-phalloidin. We counted the number of filopodia per dendrite length and found that, upon RGD treatment, the density of filopodia is slightly but not significantly increased in DPP6-KO neurons compared to the control RAD peptides treatment (Figure 8E–F p>0.1). However, with the same RGD treatment in DPP6-GFP expressing DPP6-KO neurons, filopodia were significantly increased by ~40% compared to the control RAD (p<0.05, unpaired Student’s t-test).

These results are compatible with that of Shi and Ethell ³¹ who showed that RGD treatment induced elongation of existing dendritic spines and promoted formation of new filopodia, suggesting that the RGD peptide acts as a substrate to enhance stability. ECM–DPP6 interactions thus likely support filopodia formation and stability through binding to the RGD motif of the ECM to enhance cell adhesion. We performed an ECM binding assay to look for potential ECM protein partners for DPP6. Results presented in Figure 8C–D show such an interaction of DPP6, specifically with fibronectin. We did not, however, find evidence for DPP6 interaction with collagen type 1, 4 or laminin.

Constructs

DPP6-GFP or DPP6-mCherry constructs was prepared by PCR on construct rat pcDNA3-DPP6²¹ and subcloned into the pEGFP-N1 or pmCherry-N1 vector (Clontech, Mountain View, CA). Myosin X-GFP constructs are previously described and characterized (Berg and Cheney, 2002). Additional details, including construction specifics and primer sequences, are given in the Supplementary Methods.

Rat and mouse hippocampal neuron culture and transfection

For details on the preparation of primary hippocampal neuron cultures from E18–19 WT, DPP6-KO and Kv4.2-KO mice or Sprague-Dawley rats, please see Supplementary Methods. These methods were approved by NICHD’s Animal Care and Use Committee.

Immunocytochemistry

Cultured hippocampal neurons were fixed with freshly prepared 4% paraformaldehyde plus 4% sucrose in PBS. Neurons were blocked with 10% normal goat serum in PBS and permeabilized with 0.2% Triton X-100 PBS for 10min, immunostained with rabbit anti-MAP2 (1:1000, Chemicon), mouse anti-PSD95 (1:200, NeuroMab), mouse anti-Synaptophysin (1:500, Millipore), mouse anti-Tau-1 (1:1000, Millipore) for 1 h at 37°C, washed with PBS and incubated with Alexa Fluor 488-conjugated goat anti-rabbit (1:500, Invitrogen), Alexa Fluor 555 conjugated goat anti-mouse or Alexa Fluor 633 goat anti-rabbit for 1 h at 37°C. Some neurons were counterstained with TRITC-phalloidin (1:500, Invitrogen) to visualize actin filaments. Cells were imaged with a Zeiss Axioplan 2 microscope or a Zeiss LSM510 confocal microscope (Carl Zeiss, Germany) using a 63X objective with a 1.4 numerical aperture. Imaging and Measurements are described in full in Supplementary Information.

Time-lapse light microscopy

To make time-lapse sequences, WT or DPP6-KO P0 mouse ¹³ hippocampal neurons, in some cases expressing DPP6-GFP or GFP as a control, were cultured for 3–4 days on 25mm round coverslips. Time series were acquired in Differential Interference Contrast (DIC) mode using 0.5% transmission of the 561 nm laser on the transmission path of a Zeiss Live Duo scan microscope. The neurons were maintained at 37° C with 5% CO₂ in a humidified environment. The cells were exposed to light only during image acquisition (5–7 s/image). We used a 63x/1.40 Oil DIC M27 Plan-Apochromat objective. Time-lapse recordings of dendrites were carried out for 60 min during which time images were collected at a rate of one image every 30 seconds. Filopodia detected during the recording period and which remained for the duration were counted as new filopodia. All images were analyzed using MetaMorph software.

Staining and imaging analysis

8-week-old male WT C57BL/6 and DPP6-KO mouse brain slices (250 μm thick) were prepared using a vibrating tissue slicer as described for the whole-cell electrophysiological recording methods listed below. 2 mg biocytin was added to 1 ml internal solution containing (in mM): 20 potassium chloride, 125 potassium gluconate, 10 HEPES, 4 NaCl, 0.5 EGTA, 10 phosphocreatine, 4 ATP, 0.3 TrisGTP (pH 7.2 with NaOH) and was passively perfused into CA1 neurons. Somatic patches were maintained for 20 min to let biocytin diffuse throughout the neuron. Access (5~10 MΩ) was carefully monitored during all injections. After slowly withdrawing injection pipettes, slices were rapidly transferred to PBS containing 4% PFA. Tissue was fixed for 24 hours before IHC staining. Biocytin filled slices were single labeled with Alexa-488 conjugated streptavidin (1:500, Invitrogen) overnight at 4°C after being permeabilized with 0.3% Triton-X 100 in PBS. Following 4X washes in PBS, slices were then cryoprotected using a 25% sucrose/PBS solution, and subsectioned into 70 μm thick slices using a freezing microtome, washed in PBS at 4°C for 2–3 hr and mounted on gelatin-coated slides using Mowiol mounting medium (Calbiochem, La Jolla, CA).

Confocal image stacks of stained cells were performed at the NICHD Microscopy & Imaging Core facility using a Zeiss LSM 510 Inverted Meta with 63x oil objective lens. Branching complexity was analyzed by Sholl analysis ³⁷. Neurons were traced with the center of the soma as a focal point. Sholl measurements were obtained by quantifying the number of dendritic intersections crossing each 20 μm radius from soma. To calculate spine density, all dendrites located at a specific distance (40, 120, 240 μm from soma) were counted, and the number of spines along the length was counted to give a measure of spine/20 μm length.

Co-immunoprecipitation and Western Blotting

To confirm an interaction between DPP6 and Myosin-X, we performed co-IP experiments either in native, detergent-solubilized mouse brain extracts or COS7 cells co-transfected with various DPP6 and Myosin-X constructs for 24–48 h. Mouse brain or cells were lysed in lysis buffer: 150 mM NaCl, 20 mM Tris-HCl, 1% NP40, 0.5% SDS and protease inhibitor mixture (Roche, Indianapolis, IN). Anti-DPP6 (2 μg/500 μg protein, Abcam, Cambridge, MA), IgG (Invitrogen) as nonspecific control was then added to the lysate. The mixture was then incubated and rotated at 4°C overnight. The antibody-antigen complex was immobilized by adsorption onto 50 μl of immobilized protein A (Pierce, Rockford, IL) and incubated for 2 h at RT. The protein-bead mixtures were washed 6x with lysis buffer. The beads were resuspended in reducing SDS sample buffer and analyzed on 3–8% Tris-acetate SDS polyacrylamide gels. The separated proteins were immuonoblotted using DPP6 (1:2000) or Myosin-X antibody (1:4000, Cat# HPA024223, Sigma) and visualized by Alexa Fluor 680 secondary antibody (1:10,000, Invitrogen) and Alexa Fluor 800 secondary antibody (1:10,000, Rockland, Gilbertsville, PA). Immunoreactivity was detected with the Odyssey infrared imaging system (LI-COR Biosciences, Lincoln, Nebraska).

Cell adhesion and ECM binding assay

Adhesion of COS7 cells to a substratum was assayed as described ^{38, 39}. Ninety-six-well plates were coated with 0.1 mg/ml poly-L-lysine overnight at 4 °C were used for cell adhesion assay; ECM-coated 96-well plates (BD BioCoat, Bedford, MA) were used for binding assay. Plates were blocked with DPBS containing 2% BSA for 2 h. Transfected cells were harvested with enzyme-free cell dissociation buffer (Invitrogen), and then plated in DMEM with 1% BSA at a density of 60,000 cells/well. Cells were allowed to attach to the wells for 60 min at 37 °C, then non-adherent cells were washed away. The remaining adherent cells were fixed with 5% glutaraldehyde for 30 min and stained with 5 mg/ml crystal violet in 20% methanol for 30 min, washed with PBS, and extracted with 0.2% Triton X-100 for 30min. Microplates were colorimetrically evaluated at 576 nm.

RGD or RAD treatment

DIV3 mouse hippocampal neurons were treated with 250 μM RGD-containing peptide [Gly-Arg-Gly-Asp-Thr-Pro (GRGDTP)] (EMD Biosciences, San Diego, CA) or the control RAD-containing peptide [Gly-Arg-Ala-Asp-Ser-Pro (GRADSP)] (EMD Biosciences) for 1 h at 37°C with 5% CO₂, then neurons were fixed and immunostained with TRITC-phalloidin and density of filopodia quantified.

Electrophysiology

6–8-week-old male C57BL/6 WT, Kv4.2-KO and DPP6-KO mice were anesthetized by isoflurane prior to decapitation according to methods approved by NICHD’s Animal Care and Use Committee. Acute brain slices (250 μm thick) were prepared using a vibrating tissue slicer as described previously ¹³. Cultured hippocampal neurons were recorded on DIV 14–16. During recording, cultured neurons were bathed in extracellular solution containing (in mM) 145 NaCl, 3 KCl, 10 HEPES, 8 dextrose, 2 CaCl₂, 2 MgCl₂. Somatic whole-cell recordings were performed at 32~33°C for slices and 20°C for cultured neurons, and were made from visually identified CA1 pyramidal neurons using DIC optics. Recording pipettes had tip resistances of 3~5 MΩ. Series resistance varied between 6~15 MΩ, and was carefully monitored throughout the experiments. No compensation for series resistance was employed. Neurons showed a resting membrane potential lower than −60mV. Recordings in which the resting potential changed by more than ±5mV of the initial value were excluded from analysis. All electrophysiological data were recorded using a Multiclamp 700B amplifier. Signals were digitized at 10 kHz with a Digidata 1440 A and filtered at 2 kHz for mEPSCs and at 4 kHz for PPF recordings.

For recording mEPSCs, recording pipettes were filled with intracellular solution containing (in mM) 100 Cs-gluconate, 5 MgCl₂, 0.6 EGTA, 8 NaCl, 40 HEPES, 2 NaATP, 0.3 trisGTP (pH 7.2 with KOH). TTX (1 μM), Bicuculline (20 μM) were included in the external solution during all mEPSC recordings. After whole-cell formation, a 5 min recovery period elapsed before data collection. Membrane potential was held at −65 mV. Each neuron was recorded continuously for 10 min. Synaptic currents were analyzed with the Mini Analysis program (Synaptosoft Inc.). Only events showing a rise time and decay time less than 2 ms and 10 ms, respectively, and amplitude greater than 8 pA, were analyzed. For PPF recordings, the patch electrode solution contained the same internal solution listed above for mEPSC recordings. Experiments were performed in 20 μM bicuculline. A 50 ms interval elapsed between the 1^st and 2^nd stimulus pulse. Paired-pulse ratios are the ratio of the amplitude of the second EPSC to the first. All PPF recordings were analyzed using Clampfit 10.1 and Microsoft Excel. Statistical significance was evaluated using Student’s t-test (unpaired, two tails).