| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
1Departments of Molecular Physiology and Biophysics, and 2Pharmacology and 3Vanderbilt Kennedy Center for Research on Human Development, Vanderbilt University Medical Center, Nashville, Tennessee
Submitted 16 August 2006; accepted in final form 11 January 2007
 |
ABSTRACT |
|---|
|
|
|
INTRODUCTION |
|---|
|
; Del Rio et al. 1997; Frotscher et al. 2003). Reelin binds to two members of the low-density lipoprotein receptor (LDLR) family, apolipoprotein E receptor 2 (apoER2), and very-low-density lipoprotein receptor (VLDLR) and results in phosphorylation of disabled 1 (dab1) and activation of Src family protein tyrosine kinases (D'Arcangelo et al. 1999; Hiesberger et al. 1999; Howell et al. 1997, 1999). Recent studies have shown that reelin signaling may also play an important role in the adult brain function. For example, bath perfusion of recombinant reelin onto hippocampal slices enhances tyrosine phosphorylation of N-methyl-D-aspartate receptor (NMDAR) subunits, increases NMDAR-mediated whole cell currents, and elevates the magnitude of long-term potentiation (Beffert et al. 2005; Chen et al. 2005; Weeber et al. 2002). On the other hand, genetic ablation of either apoER2 or VLDLR leads to impaired memory function in mice (Weeber et al. 2002).
It has been recently shown that reelin is required for normal development of dendritic structures (Niu et al. 2004), maturation of somatic NMDARs (Sinagra et al. 2005), and modulation of glutamate-induced NMDAR activities (Chen et al. 2005). These findings also raise an interesting question of whether reelin signaling affects maturation of hippocampal glutamatergic function in vivo. It is well established that NMDAR function is dependent on its subunit composition. Two major forms of NR2 subunits in adult hippocampus, NR2A and NR2B, are developmentally regulated (Chavis and Westbrook 2001; Monyer et al. 1994; Sheng et al. 1994), subjected to activity- and behavior-induced changes (Barria and Malinow 2005; Carmignoto and Vicini 1992; Quinlan et al. 2004), and contribute to CA1 synaptic plasticity (Malenka and Nicoll 1993; Nicoll and Malenka 1995). NR2A has faster kinetics and gradually increases during the first several postnatal weeks in mice (Monyer et al. 1994). Moreover, both NR2A and NR2B can be tyrosine phosphorylated in responses to synaptic activity (Lau and Huganir 1995; Lu et al. 1998; Salter and Kalia 2004; Yu et al. 1997) and reelin signaling (Beffert et al. 2005; Chen et al. 2005). Therefore changes of NMDAR activity through developmental alteration of subunit composition and/or tyrosine phosphorylation by reelin could have a dramatic impact on synaptic strength and plasticity.
The maturation of CA1 synapses is also reflected by an increased AMPAR response at synapses that initially contain only NMDARs (silent synapses) (Isaac et al. 1995; Kullmann 1994; Liao et al. 1995). In addition, CA1 synaptic strength and plasticity could be accomplished by altered AMPAR subunits phosphorylation (Barria et al. 1997; Lee et al. 2000) and/or trafficking to postsynaptic sites following synaptic activity (Hayashi et al. 2000; Shi et al. 1999; Song and Huganir 2002). In this study, we provide evidence that reelin-induced signaling facilitates the developmental maturation of hippocampal glutamatergic function in the postnatal hippocampus.
|
|
METHODS |
|---|
|
The B6C3Fe strain mice that carry either a single or both alleles of RELN gene were obtained from the Jackson Laboratory. These mice were mated to produce progenies of various genotypes that were determined by PCR, as previously described (Qiu et al. 2005). Mice at postnatal days 6–7 or embryonic days 16–17 were used for slice cultures and hippocampal neuronal cultures, respectively. Animal care and use protocol was according to National Institutes of Health guidelines and approved by the Institutional Animal Care and Use Committee of Vanderbilt University.
Hippocampal slice cultures and drug treatment
Hippocampal slice cultures were prepared using the protocol described by Stoppini et al. (1991). Briefly, horizontal slices were made from 6- to 7-day-old litters and were cultured on Millicell culture plate inserts (Millipore, Billerica, MA) in DMEM/F12 medium supplemented with 10% FBS. Drugs were applied after 3 days in culture and replenished every 3 days thereafter. Recombinant reelin was produced by using HEK293 cells that were stably transfected with the full-length reelin construct pCrl vector (D'Arcangelo et al. 1997). The expression of GST-RAP fusion protein was obtained by using DH5
cells carrying the GST-RAP vector construct [provided by Dr. G. Bu (Zhuo et al. 2000)]. The preparation and purification of reelin and GST-RAP, preparation of mock conditioned medium was essentially as previously described (Sinagra et al. 2005). The chemicals including TTX, 6-cyano-7-nitroquinoxalene-2,3-dione (CNQX), D-AP5, and bicuculline were purchased from Tocris-Cookson (Ellisville, MO). The Src inhibitor PP1 was purchased from Calbiochem (San Diego, CA).
Electrophysiology
The cultured slices were transferred to the recording chamber and perfused with artificial cerebrospinal fluid (ACSF; containing in mM: 125 NaCl, 2.5 KCl, 1.25 NaH2PO4, 26 NaHCO3, 1.2 MgCl2, 2.0 CaCl2, and 10 glucose, pH 7.3–7.4, and bubbled with 95% O2-5% CO2). 20 µM bicuculline was included in the perfusate. Slices were incubated in the recording chamber at room temperature for 
1.5 h before recording.
Spontaneous or synaptically evoked whole cell responses of CA1 pyramidal neurons were recorded at room temperature (22–24°C) using an EPC10 double amplifier (Heka Elektronik, Germany) with the aid of DIC microscopy (Leica DM-LSFA). The patch electrodes (4–7 M
) were filled with a solution containing (in mM) 117 Cs gluconate, 10 tetraethylammonium-Cl, 10 HEPES, 8 NaCl, 5 QX314-Cl, 4 Mg2+-ATP, 2.5 CsCl, 0.3 Na3GTP, and 0.2 EGTA, and adjusted to pH 7.2, 280–290 mOsm. CA1 pyramidal neurons were voltage clamped at –65 mV in whole cell configuration for 10 min to achieve stable holding currents. Series resistance (8 –15 M) was monitored by applying 5-ms hyperpolarizing voltage steps. Experiments were terminated if serial resistance or input resistance changed by >15%. To measure AMPAR-mediated spontaneous miniature EPSCs (mEPSCs), slices were preincubated with 20 µM bicuculline and 1 µM TTX for 30 min, and recordings were carried out in the continued presence of these chemicals.
Monosynaptically evoked whole cell responses, as verified by their high synchrony across varying stimulus intensities, were recorded at either –65 or +40 mV. Electrical stimulus was driven by a Master-8-cp stimulator (AMPI) and delivered through a bipolar tungsten electrode (Frederick Haer, Bowdoinham, ME) to Schaffer collateral fibers. To obtain NMDAR-mediated whole cells currents, neurons were clamped at +40 mV in the presence of 10 µM CNQX, and the response was confirmed by washing in 100 µM D-AP5 on termination of experiments. Electric signals were digitized either by the EPC10 or Digidata 1322A at 10 kHz and filtered at 2 kHz, stored on a PC computer, and analyzed off-line.
To determine the proportion of silent synapse and synaptic failures of CA1 pyramidal neurons in cultured hippocampal slices, a minimum stimulation protocol was adopted (Liao et al. 1995). CA1 pyramidal neurons were voltage clamped at –65 mV, and stimulus intensity was gradually increased from zero to obtain a minimum stimulus intensity that elicits a mixture of responses and failures. This stimulus intensity was kept constant and synaptic responses were recorded for 15 min at 0.2 Hz. Cells were then clamped at +40 mV to record the mixed AMPAR- and NMDAR-mediated EPSCs. D-AP5 (100 µM) was washed in at the end of the experiment to ensure the failure rates of AMPAR-mediated EPSCs at –65 and +40 mV were the same, only those cells showed same failure rates were included in analysis.
Cell surface protein biotinylation and Western blotting
Cultured slices treated with reelin or mock for 6–7 days or nontreated controls were biotinylated in ACSF with 1 mg/ml sulfo-NHS-S-S-biotin for 30 min at 4°C, using a Pinpoint cell surface protein isolation kit (Pierce, Rockford, IL), similar to that described previously for biotinylation in slices (Broutman and Baudry 2001). CA1 regions were dissected out and sonicated in RIPA buffer [contains 50 mM Tris–HCl, pH 7.4, 10 mM EDTA, 100 µM leupeptin, 1 mM pepstatin, 10 µg/ml aprotinin, 10 µg/ml bacitracin, 100 µM phenylmethylsulfonyl fluoride (PMSF), 1 mM sodium orthovanadate and 1% Nonidet P40]. Half volume of the tissue lysate was used for probing total proteins and the other half was mixed with immobilized neutravidin beads (Pierce) and rotated for 2 h at 4°C. The beads were then washed three times in RIPA buffer and, along with the total protein fraction, treated with 2 x Laemmli buffer supplemented with 50 mM DTT. Both the total protein and biotinylated proteins were separated by SDS-PAGE by running through 9% SDS-polyacrylamide gels. The proteins were then transferred to PVDF membranes (Immobilon-P, Millipore) and incubated with antibodies against GluR1, NR1, NR2A or NR2B diluted in 5% nonfat dry milk and 1% BSA for 2 h at RT. The membranes were then washed extensively in 0.01M PBS-Tween 20 and incubated for 1 h at room temperature with HRP-conjugated secondary antibodies (Promega, Madison, WI) and developed using an enhanced chemiluminescence method. Rabbit anti-NR2A and anti-NR2B antibodies were purchased from Upstate Biotechnology. Rabbit anti-GluR1, anti-NR1 and monoclonoal NR1 antibodies were obtained from Chemicon (Temecula, CA). The final dilution of antibodies was between 1:1,000 and 1:5,000. The optical density of immunoreactive bands was quantified by densitometry using Quantity One software (Bio-Rad, Hercules, CA).
Hippocampal neuronal culture, immunocytochemistry, and image analysis
Hippocampal neuronal cultures were prepared from 16- to 17-day embryonic mice. Briefly, hippocampi from fetal brains were collected and digested in Hank's balance salt solution containing 0.5 mg/ml papain at 37°C for 20 min. In the meanwhile, the tails of the embryos were collected for genotype analysis. The hippocampi were then separated into individual cells using a glass micropipette. The cells were seeded on poly-D-lysine-coated 12-mm glass coverslips at a density of 5,000 cells/cm2 and grown in Neurobasal medium supplemented with 2% B27. Every 5 days after plating, half of the medium was replaced with fresh plating medium containing 10 µM cytosine arabinoside. At day 14–20, neurons were fixed in 4% paraformaldehyde in PBS for 20 min followed by prechilled (–20°C) methanol for 10 min. The neurons were permeabalized for 20 min in 0.2% Triton-X 100, and were blocked with 0.01M PBS containing 10% normal donkey serum and 1% BSA, incubated overnight with primary antibodies (rabbit anti-GluR1 and mouse anti-NR1, both 1:500 dilutions), followed by a 2-h incubation with Alexa Fluor 488-conjugated donkey anti-rabbit secondary IgG (Invitrogen, Carlsbad, CA) and CY3-conjugated donkey anti-mouse IgG (Jackson Immunoresearch, West Grove, PA).
Digital images were taken with a Zeiss LSM510 confocal microscope using a 40x oil-immersion lens (numerical aperture: 1.3). All images from the same experiment were acquired with identical settings for laser power, photomultiplier gain, offset and a fixed pinhole size of one Airy unit. Images were encoded to analyze blindly using MetaMorph software (Molecular Devices, Sunnyvale, CA). Thresholds for GluR1 and NR1 immunostaining were chosen such that all visually recognizable punctate labeling were included for analysis. For colocalization analysis, regions were defined around the thresholded puncta in one channel (NR1, red), and these regions were used to create a mask and then overlaid on the second channel (GluR1, green). Thresholded puncta defined by at least partially overlapping regions were considered colocalizing. 10 neurons were analyzed per group and the sections of 50–100 µm of all the primary dendrites were counted and averaged to determine the number of GluR1, NR1, or colocalized puncta. Each experiment was repeated in four cultures.
Data analysis
The amplitude of whole cell EPSCs were defined as the averaged 2–4 ms value around the peak minus the stable holding current immediately proceeding the stimulus artifact and was quantified by averaging 12 consecutive traces during a 1-min bin. To determine the time constant of EPSCNMDA decay, the fallen phase of averaged EPSCNMDA was fitted with a single exponential decay using Igor Pro software. Although NMDAR whole cell current can be best fitted by multi-exponential kinetics (Kampa et al. 2004), a single-exponential function nevertheless provides an accurate estimation on the decay time constant, especially in early developmental stages (Barth and Malenka 2001). The peak current amplitude was used to quantify AMPA/NMDA ratio and ifenprodil sensitivity. Detection and analysis of mEPSCs were done using Clampfit 9.0, and each detected event was confirmed by visual inspection.
To estimate the proportion of silent synapses and quantify synaptic failure rates, the amplitude distribution histograms of minimum stimulation-evoked EPSC at either –65 or +40 mV were plotted and a smooth curve was drawn using the Spline function provided in GraphPad Prism 4.0 software. The integral of the peak centered at 0 pA represents failures of transmission and matches the noise amplitude distribution. The failure rates were measured by using the fraction of responses with amplitude less than zero and then timed by two based on the symmetry of noise (Liao et al. 1995). Failure rates were also visually inspected in a subset of cells to ensure that this approach provided an accurate estimation. The decrease of failure rate at +40 mV (mixed AMPAR + NMDAR response) compared with that in –65 mV (AMPAR response) indicate the presence of silent synapses, whose numbers are proportional to the differences of failure rates (Liao et al. 1995).
For statistical analysis, data were represented by means ± SE. An unpaired Student's t-test, one-way ANOVA (with post hoc Student Neuman-Keuls test when overall significance was achieved) or two-way ANOVA analysis were performed when data passed normality test. For the comparison of cumulative histogram of mEPSC amplitudes, Kolmogorov-Smirnov test was used. Significance was assigned for all tests at P < 0.05.
|
|
RESULTS |
|---|
|
Reelin binds to its lipoprotein receptors, apoER2 and VLDLR, and initiates a signaling cascade that is transmitted by Dab1 and Src (Bock and Herz 2003; Hiesberger et al. 1999; Howell et al. 1999; Trommsdorff et al. 1999). In adult hippocampal slices, this signaling leads to increased NMDAR tyrosine phosphorylation, increased NMDAR-mediated currents, and elevated magnitude of LTP (Beffert et al. 2005; Chen et al. 2005; Weeber et al. 2002). It is not known whether reelin signaling affects glutamatergic transmission during postnatal hippocampal development. We therefore tested this using hippocampal slice cultures prepared from 6- to 7-day-old mice. Either 5 nM reelin or reelin in combination with the functional lipoprotein receptors antagonist GST-RAP (20 µM) (Herz et al. 1991) or Src inhibitor PP1 (10 µM) (Hanke et al. 1996) were applied to slice cultures at day 3. CA1 pyramidal neurons were then recorded for spontaneous miniature activities or synaptically evoked whole cell responses at 6–7 days after drug application. To quantify reelin effects on spontaneous mEPSCs, 
2,000 consecutive events pooled from 8 to 10 CA1 pyramidal neurons derived from six to eight slices were analyzed for each group. It was found that reelin treatment markedly increased spontaneous mEPSC amplitude, an effect that was blocked by either GST-RAP or PP1 (Fig. 1, A–C compared with mock treated group, P < 0.01 for reelin; P > 0.05 for reelin + GST-RAP and reelin + PP1. Kolmogorov-Smirnov test). No significant change in mEPSC frequency was seen after reelin treatment [Fig. 1D, F(3,30) = 2.43, P > 0.05]. Therefore chronic reelin treatment significantly increased spontaneous AMPAR-mediated responses in CA1 pyramidal neurons.
|
; Kullmann 1994; Liao et al. 1995). In addition, the subunit composition of NMDARs at murine CA1 synapses normally changes over the two to three postnatal weeks (Monyer et al. 1994) with a developmental switch from NR2B to NR2A subunits. We first compared the relative current ratio of AMPAR and NMDAR under mock- and reelin-treated conditions. The AMPA/NMDA current ratio of mock-treated slices was significantly lower than that from slices treated with reelin [Fig. 2B, mock treated, 0.84 ± 0.07; reelin treated, 1.73 ± 0.12. F(3,35) = 10.4, P < 0.01], an effect that was not observed when reelin was co-applied with GST-RAP or PP1. These results indicate that chronic reelin treatment enhances AMPAR function and facilitates glutamatergic transmission in the developing hippocampus.
|
) of EPSCNMDA was also significantly smaller after reelin treatment [Fig. 2D, reelin treated, 93.6 ± 10.5 ms; mock treated, 185.8 ± 17.2 ms, F(3,25) = 8.7, P < 0.01]. Pharmacological blockade of reelin signaling with GST-RAP or PP1 abolished reelin effects on both ifenprodil sensitivity and EPSCNMDA decay , suggesting an accelerated switch to faster NMDAR kinetics occurred in the developing hippocampus as a result of reelin signaling. Differential effects of reelin treatment on surface levels of AMPAR and NMDAR subunits
Because of the increased AMPA component of synaptic response, altered of EPSCNMDA kinetics and ifenprodil sensitivity after chronic reelin treatment, we asked whether the amounts of AMPAR and NMDAR subunits were changed by reelin in CA1. Both total and surface levels of GluR1, NR1, NR2A, and NR2B were probed by Western blotting. We first examined whether GluR1, an AMPAR subunit that is increasingly expressed during developmental maturation (Petralia et al. 1999; Wu et al. 1996) and subjected to regulated trafficking during synaptic plasticity (Hayashi et al. 2000; Shi et al. 1999), was increased on CA1 cells surfaces. Figure 3 shows, as expected, that reelin treatment significantly increased levels of surface GluR1 [F(2,11) = 15.6, P < 0.001] as well as total GluR1 [F(2,11) = 11.2, P < 0.01] compared with mock-treated groups, indicating regulated expression and surface insertion of AMPARs at least partly account for the increased mEPSCAMPA and AMPA/NMDA current ratio after chronic reelin treatment. No changes of either surface or total NR1 levels were observed. In comparison, both total and surface NR2A showed significant increase after reelin treatment compared with mock treatment [total protein, F(2,14) = 9.75, P < 0.01; surface protein, F(2,11) = 44.9, P < 0.001]. Moreover, both total and surface NR2B showed significant decrease [total protein, F(2,11) = 4.1, P < 0.05; surface protein, F(2,11) = 22.6, P < 0.001] after reelin treatment.
|
If synapses acquire more AMPARs at postsynaptic sites as a result of reelin treatment, then the number of silent synapses, which contain only NMDARs and therefore are quiescent at normal resting membrane potentials, should decrease. This scenario could also explain increased AMPA/NMDA ratio and mEPSCAMPA amplitudes after reelin treatment. We further tested this hypothesis by comparing the proportion of silent synapses in CA1 neurons in mock- and reelin-treated slices by using a minimum stimulation protocol (see METHODS). In both mock- and reelin-treated slices, neurons clamped at –65 mV displayed a consistently higher failure rate than that obtained when clamped at +40 mV, indicating the existence of silent synapses (Fig. 4, A and E). The failure rates were further quantified and frequency distributions of synaptic EPSC amplitudes were analyzed. Histogram of EPSCs revealed that reelin treatment increased clustering of EPSCs at larger amplitudes at both –65 and +40 mV holding potentials (Fig. 4, D and H). When the failure rates obtained at –65 and +40 mV holding potentials were compared (Fig. 4, I–K), mock- and reelin-treated cells showed a 41.4 ± 5.4 and 5.8 ± 1.9% reduction in failure rates at +40 mV, respectively (n = 7–8, P < 0.001, t-test), indicating reelin treatment significantly reduced the proportion of silent synapses compared with mock treatment. In addition, at –65 mV, reelin-treated cells showed 39.2% ±4.4% less failures compared with mock-treated cells (reelin, 59.5 ± 5.3%; mock treated, 20.3 ± 4.7%. n = 6, P < 0.01, t-test); this suggests facilitated synaptic transmission by AMPAR at normal resting membrane potentials after reelin treatment.
|
|
Recent findings have indicated that reelin is required for normal development of dendritic structure and maturation of somatic NMDARs in cultured hippocampal neurons (Niu et al. 2004; Sinagra et al. 2005). We hypothesized that reelin signaling also facilitates the formation and function of glutamatergic synapses through cell autonomous mechanisms and tested this hypothesis in cultured low-density embryonic hippocampal neurons. In agreement with the previous findings (Niu et al. 2004), we observed that in cultures obtained from reeler embryos, neurons generally exhibited stunted neurite growth and less number of ramifications at 14 days after plating, which could be corrected by supplying 5 nM reelin in the culture medium from day 3 after plating (Fig. 6, A–C). To quantify the potentially functional glutamatergic synapses, we labeled both NR1 and GluR1 proteins using immunofluorescence techniques. In wild-type neurons, primary dendrites showed extensive punctate labeling of NR1 and GluR1 (Fig. 6A), which represented potential sites for synapses. In addition, a considerable proportion of NR1 positive puncta also contained GluR1. In contrast to those puncta that show only NR1 immunoreactivity (therefore are silent at resting membrane potentials), acquisition of GluR1 in these silent synapses is believed to render them capable of fast synaptic transmission. In neurons obtained from reeler embryos, less number of NR1 and GluR1 clustering was observed (Fig. 6B), and this deficit was corrected by supplying reelin in the culture medium (Fig. 6C).
|
2,000 mEPSCs pooled from 8 to 10 neurons for each group). This difference was eliminated when 5 nM reelin was supplemented in the culture medium in reeler cultures (Z = 0.26, P > 0.05, compared with WT neurons, Kolmogorov-Smirnov test). In addition, mEPSC frequency was significantly lower in cultured reeler neurons compared with that from WT neurons. Supplement of reelin in the reeler culture fully restores mEPSC frequency to the WT levels (number of mEPSC/s, WT, 2.35 ± 0.24; reeler, 0.72 ± 0.33; reeler + reelin, 2.11 ± 0.37; F(2,19) = 4.9, P = 0.026, data not shown). Therefore these results suggest that reelin signaling facilitates glutamatergic synapse formation and function through cell autonomous mechanisms that do not require neuronal activities under in vivo conditions. |
|
DISCUSSION |
|---|
|
; D'Arcangelo et al. 1995; Goffinet 1995; Howell et al. 1997, 1999). Recently, emerging evidence also suggests that normal reelin-lipoprotein receptor-mediated signaling also plays an important role in neuronal maturation, synaptic plasticity, and memory formation in adult brain (Beffert et al. 2005; D'Arcangelo 2005; Herz and Chen 2006; Nimpf and Schneider 2000; Weeber et al. 2002). We have previously shown that application of reelin to acutely prepared hippocampal slices enhances tetanus-induced LTP, and the modulatory role of reelin on LTP and memory is dependent on apoER2 and its alternatively spliced intracellular domain encoded by exon 19 (Beffert et al. 2005; Weeber et al. 2002). In this study, we provide evidence that reelin signaling facilitates the developmental maturation of hippocampal glutamatergic function.
Recent studies have demonstrated that reelin signaling is required for normal development of several structural and functional aspects of hippocampal neurons. For example, hippocampal neurons derived from reeler embryos showed stunted neurite growth due to mechanisms involving apoER2/VLDLR and dab1, a deficit that can be rescued by adding reelin to the culture medium (Niu et al. 2004). However, from this study it is not clear whether a more elaborated neurite network is correlated with a more mature and functional glutamatergic synapse. A recent functional study by Sinagra et al. (2005) has shown that reelin signaling is required for normal developmental switch of somatic NMDAR subunits from NR2B to NR2A in cultured hippocampal neurons. However, this finding may not necessarily reflect the in vivo conditions and offers limited knowledge on reelin effects on synaptic NMDARs, which play a determinant role in CA1 synaptic plasticity (Zalutsky and Nicoll 1990). Moreover, maturation and subunit switch of NMDARs in vivo is critically dependent on neuronal circuitry activity (Lu and Constantine-Paton 2004; Monyer et al. 1994; Nakayama et al. 2005; Quinlan et al. 1999), a condition that cannot be satisfied in dispersed neuronal cultures. To address these potential questions, we examined effects of reelin on the developmental maturation of glutamatergic function using both hippocampal slice cultures and embryonic neuronal cultures. When slice cultures are prepared from young rodents, neural circuitries of the hippocampal formation are known to mature and to maintain a surprising three-dimensional, organotypic organization for many weeks in vitro (Gahwiler et al. 1997; Zimmer and Gahwiler 1984).
We have first examined reelin effects on spontaneous synaptic transmission by analyzing AMPAR-mediated mEPSCs, which represent postsynaptic responses to single quanta of neurotransmitter releases. Reelin application for 6–7 days in cultured slices significantly increases amplitude of mEPSCs, an effect that is dependent on apoER2/VLDLR, suggesting reelin signaling leads to enhanced AMPAR function in CA1. We next examined effects of reelin signaling on synaptic responses mediated by NMDARs and AMPARs. In agreement with previous findings (Sinagra et al. 2005), reelin facilitated a functional switch from NR2B to NR2A, as shown by increased surface expression of NR2A and decreased surface expression of NR2B, reduced ifenprodil blockade of synaptic EPSCNMDA and decreased of EPSCNMDA decay current. Importantly, these effects were dependent on apoER2/VLDLR and Src, as they were blocked by either GST-RAP or PP1 (Fig. 2). Decreased of NMDAR decay through NR2A acquisition during maturation has been described in many CNS regions (Carmignoto and Vicini 1992; Flint et al. 1997; Hestrin 1992). Reduced ifenprodil sensitivity and faster of EPSCNMDA indicate more NR2A subunits are present at synaptic sites in reelin-treated slice cultures. Because NMDAR subunit composition determines surface delivery of AMPAR subunits (Kim et al. 2005), differentially couples with intracellular biochemical events (Barria and Malinow 2005; Kim et al. 2005), and may differentially affect CA1 synaptic plasticity (i.e., LTP or LTD) (Bartlett et al. 2006; Morishita et al. 2006), an accelerated switch to NR2A by reelin could have a profound impact on synaptic strength and plasticity.
We next examined whether reelin signaling leads to increased efficacy of synaptic transmission by comparing spontaneous miniature events, synaptically evoked AMPA/NMDA current ratios, synaptic failures and estimated relative abundance of silent synapses. As CNS synapses mature, those containing only NMDARs acquire AMPARs in an activity-dependent manner (Isaac et al. 1995; Kullmann 1994; Liao et al. 1995). We have found that both mEPSCAMPA amplitude and AMPA/NMDA current ratio were significantly increased after chronic reelin treatment. These effects were also dependent on apoER2/VLDLR and Src. The increased AMPAR responses might be due to increased levels of surface expression GluR1 subunit (Fig. 3). However, it is not clear whether this increased GluR1 expression is mainly synaptic and whether other AMPAR subunits, such as GluR2 and GluR4 (Kolleker et al. 2003; Shi et al. 2001; Zhu et al. 2000), also contribute to enhanced AMPAR function. Moreover, in the presence of GST-RAP or PP1, there was a significant reduction in NMDAR subunit switch and an increased synaptic failure rate compared with untreated slice cultures undergoing normal in vitro development (Fig. 5). These results indicate that a constitutive level of reelin signaling is required to maintain the physiological maturation process in CA1. Taken together, our data show that prolonged reelin treatment during development facilitates maturation of CA1 glutamatergic function through an enhanced AMPAR-mediated synaptic transmission and a functional switch from NR2B to NR2A. It is interesting to note that reduced silent synapses should result in increased mEPSCAMPA frequency, which was not observed in our cultured slices. This may be due to decreased presynaptic release probability, structural changes of synapse and/or reduced number of total synapse formed in CA1 neurons in the presence of reelin. Moreover, reduced NR2B contents and enhanced AMPAR currents as a result of reelin signaling should not favor LTP induction. This apparent discrepancy with our previous report that reelin enhances LTP in acute hippocampal slices (Weeber et al. 2002) may be due to different tissue preparations and the timing of reelin signaling under these two distinct conditions.
Our immunocytochemistry data using cultured low-density hippocampal neurons provide strong connection between reelin signaling with glutamatergic synapse formation and support a cell autonomous rather than activity-dependent mechanism for reelin signaling. During development, newly formed synapses contain exclusively NMDARs (silent) and progressively acquire AMPARs in an activity-dependent manner (Liao et al. 1999; Wu et al. 1996). In the absence of reelin, impaired neurite outgrowth occurs (Niu et al. 2004). We have further shown that neurons from reeler cultures exhibit less glutamate receptor-mediated activity, significantly reduced numbers of GluR1 and NR1 clusters and colocalized puncta that contains both GluR1 and NR1. Moreover, in the absence of reelin, more NR1 puncta did not show GluR1 immunoreactivity. In comparison, neurons derived from wild-type embryos or reeler cultures supplemented with recombinant reelin showed increased number of NR1 and GluR1 clustering and a larger proportion of double immunoreactivity for GluR1 and NR1 (therefore reduced number of silent synapses). By employing this morphological correlate of silent synapses, we provide direct evidence that reelin signaling through cell autonomous mechanisms facilitates maturation of glutamatergic function through increased clustering of GluR1 and NR1 and reduces the number of silent synapses by promoting AMPAR synaptic insertion. However, the molecular mechanisms for increased AMPAR insertion remain unclear. One possible mechanism may be that enhanced tonic NMDAR activity after reelin signaling and Src activation leads to increased acquisition of AMPARs and spontaneous activity at NMDAR-containing synapses over a prolonged developmental process similar to that described previously (Zhu and Malinow 2002). This potential mechanism may function in the maturation process of cultured hippocampal slices as well.
In light of our observation that increased reelin signaling accelerates maturation of CA1 glutamatergic synapses, it would be intriguing to know whether a decreased level reelin signaling may manifest the opposite effects. Complete loss of reelin in the reeler mouse disrupts many laminated brain structures and precludes a meaningful electrophysiological or behavioral testing in these animals. A partial loss (50%) of reelin in heterozygote mice results in a significant impairment of memory performance and long-term plasticity (Qiu et al. 2005; Tueting et al. 1999; Yabut et al. 2005). It is not clear whether the phenotypes found in heterzygote reeler mouse are primarily due to deficits in network connections or due to insufficient signaling caused by reelin haploinsufficiency. Our data using slice and neuronal cultures support the later in that chronic reelin application facilitates maturation of glutamatergic responses that is dependent on reelin receptors and the intracellular signaling component Src.
Based on the findings of this study, it would be tempting to speculate that a reduced level of reelin signaling may have broader implications in distinct brain regions from development to senescence. It is especially intriguing considering some recent interesting findings that aberrant reelin signaling may play an etiological role in some neurodegenerative or neuropsychiatric diseases such as Alzheimer's disease (Botella-Lopez et al. 2006), schizophrenia, lissencephaly, bipolar disease, and autism (for a review, see Qiu et al. 2006). The novel reelin signaling events uncovered by this study could modify glutamatergic transmission, plasticity, and, potentially, memory and cognition in a profound manner as to affect the pathogenesis and outcome of these neurological disorders.
|
|
GRANTS |
|---|
|
|
|
NOTE ADDED IN PROOF |
|---|
|
|
|
ACKNOWLEDGMENTS |
|---|
|
|
|
FOOTNOTES |
|---|
Address for reprint requests and other correspondence: E. J. Weeber, Dept. of Molecular Physiology and Biophysics, 754 Robinson Research Bldg., Vanderbilt University Medical Center, Nashville, TN 37232-0615 (E-mail: Edwin.j.weeber{at}vanderbilt.edu)
|
|
REFERENCES |
|---|
|
Barria A, Muller D, Derkach V, Griffith LC, Soderling TR. Regulatory phosphorylation of AMPA-type glutamate receptors by CaM-KII during long-term potentiation. Science 276: 2042–2045, 1997.
Barth AL, Malenka RC. NMDAR EPSC kinetics do not regulate the critical period for LTP at thalamocortical synapses. Nat Neurosci 4: 235–236, 2001.[CrossRef][Web of Science][Medline]
Bartlett TE, Bannister NJ, Collett VJ, Dargan SL, Massey PV, Bortolotto ZA, Fitzjohn SM, Bashir ZI, Collingridge GL, Lodge D. Differential roles of NR2A and NR2B-containing NMDA receptors in LTP and LTD in the CA1 region of two-week old rat hippocampus. Neuropharmacology 52: 60–70, 2007.[CrossRef][Web of Science][Medline]
Beffert U, Weeber EJ, Durudas A, Qiu S, Masiulis I, Sweatt JD, Li WP, Adelmann G, Frotscher M, Hammer RE, Herz J. Modulation of synaptic plasticity and memory by Reelin involves differential splicing of the lipoprotein receptor Apoer2. Neuron 47: 567–579, 2005.[CrossRef][Web of Science][Medline]
Bock HH, Herz J. Reelin activates SRC family tyrosine kinases in neurons. Curr Biol 13: 18–26, 2003.[CrossRef][Web of Science][Medline]
Botella-Lopez A, Burgaya F, Gavin R, Garcia-Ayllon MS, Gomez-Tortosa E, Pena-Casanova J, Urena JM, Del Rio JA, Blesa R, Soriano E, Saez-Valero J. Reelin expression and glycosylation patterns are altered in Alzheimer's disease. Proc Natl Acad Sci USA 103: 5573–5578, 2006.
Broutman G, Baudry M. Involvement of the secretory pathway for AMPA receptors in NMDA-induced potentiation in hippocampus. J Neurosci 21: 27–34, 2001.
Carmignoto G, Vicini S. Activity-dependent decrease in NMDA receptor responses during development of the visual cortex. Science 258: 1007–1011, 1992.
Chavis P, Westbrook G. Integrins mediate functional pre- and postsynaptic maturation at a hippocampal synapse. Nature 411: 317–321, 2001.[CrossRef][Medline]
Chen Y, Beffert U, Ertunc M, Tang TS, Kavalali ET, Bezprozvanny I, Herz J. Reelin modulates NMDA receptor activity in cortical neurons. J Neurosci 25: 8209–8216, 2005.
D'Arcangelo G. Apoer2: a reelin receptor to remember. Neuron 47: 471–473, 2005.[CrossRef][Web of Science][Medline]
D'Arcangelo G, Homayouni R, Keshvara L, Rice DS, Sheldon M, Curran T. Reelin is a ligand for lipoprotein receptors. Neuron 24: 471–479, 1999.[CrossRef][Web of Science][Medline]
D'Arcangelo G, Miao GG, Chen SC, Soares HD, Morgan JI, Curran T. A protein related to extracellular matrix proteins deleted in the mouse mutant reeler. Nature 374: 719–723, 1995.[CrossRef][Medline]
D'Arcangelo G, Nakajima K, Miyata T, Ogawa M, Mikoshiba K, Curran T. Reelin is a secreted glycoprotein recognized by the CR-50 monoclonal antibody. J Neurosci 17: 23–31, 1997.
Del Rio JA, Heimrich B, Borrell V, Forster E, Drakew A, Alcantara S, Nakajima K, Miyata T, Ogawa M, Mikoshiba K, Derer P, Frotscher M, Soriano E. A role for Cajal-Retzius cells and reelin in the development of hippocampal connections. Nature 385: 70–74, 1997.[CrossRef][Medline]
Flint AC, Maisch US, Weishaupt JH, Kriegstein AR, Monyer H. NR2A subunit expression shortens NMDA receptor synaptic currents in developing neocortex. J Neurosci 17: 2469–2476, 1997.
Frotscher M, Haas CA, Forster E. Reelin controls granule cell migration in the dentate gyrus by acting on the radial glial scaffold. Cereb Cortex 13: 634–640, 2003.
Gahwiler BH, Capogna M, Debanne D, McKinney RA, Thompson SM. Organotypic slice cultures: a technique has come of age. Trends Neurosci 20: 471–477, 1997.[CrossRef][Web of Science][Medline]
Goffinet AM. Developmental neurobiology. A real gene for reeler. Nature 374: 675–676, 1995.[CrossRef][Medline]
Hanke JH, Gardner JP, Dow RL, Changelian PS, Brissette WH, Weringer EJ, Pollok BA, Connelly PA. Discovery of a novel, potent, and Src family-selective tyrosine kinase inhibitor. Study of Lck- and FynT-dependent T cell activation. J Biol Chem 271: 695–701, 1996.
Hayashi Y, Shi SH, Esteban JA, Piccini A, Poncer JC, Malinow R. Driving AMPA receptors into synapses by LTP and CaMKII: requirement for GluR1 and PDZ domain interaction. Science 287: 2262–2267, 2000.
Herz J, Chen Y. Reelin, lipoprotein receptors and synaptic plasticity. Nat Rev Neurosci 7: 850–859, 2006.[CrossRef][Web of Science][Medline]
Herz J, Goldstein JL, Strickland DK, Ho YK, Brown MS. 39-kDa protein modulates binding of ligands to low density lipoprotein receptor-related protein/alpha 2-macroglobulin receptor. J Biol Chem 266: 21232–21238, 1991.
Hestrin S. Developmental regulation of NMDA receptor-mediated synaptic currents at a central synapse. Nature 357: 686–689, 1992.[CrossRef][Medline]
Hiesberger T, Trommsdorff M, Howell BW, Goffinet A, Mumby MC, Cooper JA, Herz J. Direct binding of Reelin to VLDL receptor and ApoE receptor 2 induces tyrosine phosphorylation of disabled-1 and modulates tau phosphorylation. Neuron 24: 481–489, 1999.[CrossRef][Web of Science][Medline]
Howell BW, Hawkes R, Soriano P, Cooper JA. Neuronal position in the developing brain is regulated by mouse disabled-1. Nature 389: 733–737, 1997.[CrossRef][Medline]
Howell BW, Herrick TM, Cooper JA. Reelin-induced tryosine phosphorylation of disabled 1 during neuronal positioning. Genes Dev 13: 643–648, 1999.
Isaac JT, Nicoll RA, Malenka RC. Evidence for silent synapses: implications for the expression of LTP. Neuron 15: 427–434, 1995.[CrossRef][Web of Science][Medline]
Kampa BM, Clements J, Jonas P, Stuart GJ. Kinetics of Mg2+ unblock of NMDA receptors: implications for spike-timing dependent synaptic plasticity. J Physiol 556: 337–345, 2004.
Kim MJ, Dunah AW, Wang YT, Sheng M. Differential roles of NR2A- and NR2B-containing NMDA receptors in Ras-ERK signaling and AMPA receptor trafficking. Neuron 46: 745–760, 2005.[CrossRef][Web of Science][Medline]
Kolleker A, Zhu JJ, Schupp BJ, Qin Y, Mack V, Borchardt T, Kohr G, Malinow R, Seeburg PH, Osten P. Glutamatergic plasticity by synaptic delivery of GluR-B(long)-containing AMPA receptors. Neuron 40: 1199–1212, 2003.[CrossRef][Web of Science][Medline]
Kullmann DM. Amplitude fluctuations of dual-component EPSCs in hippocampal pyramidal cells: implications for long-term potentiation. Neuron 12: 1111–1120, 1994.[CrossRef][Web of Science][Medline]
Lau LF, Huganir RL. Differential tyrosine phosphorylation of N-methyl-D-aspartate receptor subunits. J Biol Chem 270: 20036–20041, 1995.
Lee HK, Barbarosie M, Kameyama K, Bear MF, Huganir RL. Regulation of distinct AMPA receptor phosphorylation sites during bidirectional synaptic plasticity. Nature 405: 955–959, 2000.[CrossRef][Medline]
Liao D, Hessler NA, Malinow R. Activation of postsynaptically silent synapses during pairing-induced LTP in CA1 region of hippocampal slice. Nature 375: 400–404, 1995.[CrossRef][Medline]
Liao D, Zhang X, O'Brien R, Ehlers MD, Huganir RL. Regulation of morphological postsynaptic silent synapses in developing hippocampal neurons. Nat Neurosci 2: 37–43, 1999.[CrossRef][Web of Science][Medline]
Lu W, Constantine-Paton M. Eye opening rapidly induces synaptic potentiation and refinement. Neuron 43: 237–249, 2004.[CrossRef][Web of Science][Medline]
Lu YM, Roder JC, Davidow J, Salter MW. Src activation in the induction of long-term potentiation in CA1 hippocampal neurons. Science 279: 1363–1367, 1998.
Malenka RC, Nicoll RA. NMDA-receptor-dependent synaptic plasticity: multiple forms and mechanisms. Trends Neurosci 16: 521–527, 1993.[CrossRef][Web of Science][Medline]
Monyer H, Burnashev N, Laurie DJ, Sakmann B, Seeburg PH. Developmental and regional expression in the rat brain and functional properties of four NMDA receptors. Neuron 12: 529–540, 1994.[CrossRef][Web of Science][Medline]
Morishita W, Lu W, Smith GB, Nicoll RA, Bear MF, Malenka RC. Activation of NR2B-containing NMDA receptors is not required for NMDA receptor-dependent long-term depression. Neuropharmacology 52: 71–76, 2007.[CrossRef][Web of Science][Medline]
Nakayama K, Kiyosue K, Taguchi T. Diminished neuronal activity increases neuron-neuron connectivity underlying silent synapse formation and the rapid conversion of silent to functional synapses. J Neurosci 25: 4040–4051, 2005.
Nicoll RA, Malenka RC. Contrasting properties of two forms of long-term potentiation in the hippocampus. Nature 377: 115–118, 1995.[CrossRef][Medline]
Nimpf J, Schneider WJ. From cholesterol transport to signal transduction: low density lipoprotein receptor, very low density lipoprotein receptor, and apolipoprotein E receptor-2. Biochim Biophys Acta 1529: 287–298, 2000.[Medline]
Niu S, Renfro A, Quattrocchi CC, Sheldon M, D'Arcangelo G. Reelin promotes hippocampal dendrite development through the VLDLR/ApoER2-Dab1 pathway. Neuron 41: 71–84, 2004.[CrossRef][Web of Science][Medline]
Petralia RS, Esteban JA, Wang YX, Partridge JG, Zhao HM, Wenthold RJ, Malinow R. Selective acquisition of AMPA receptors over postnatal development suggests a molecular basis for silent synapses. Nat Neurosci 2: 31–36, 1999.[CrossRef][Web of Science][Medline]
Qiu S, Korwek KM, Pratt-Davis AR, Peters M, Bergman MY, Weeber EJ. Cognitive disruption and altered hippocampus synaptic function in Reelin haploinsufficient mice. Neurobiol Learn Mem 85: 228–242, 2006.[CrossRef][Web of Science][Medline]
Qiu S, Korwek KM, Weeber EJ. A fresh look at an ancient receptor family: emerging roles for low density lipoprotein receptors in synaptic plasticity and memory formation. Neurobiol Learn Mem 85: 16–29, 2006.[CrossRef][Web of Science][Medline]
Quinlan EM, Lebel D, Brosh I, Barkai E. A molecular mechanism for stabilization of learning-induced synaptic modifications. Neuron 41: 185–192, 2004.[CrossRef][Web of Science][Medline]
Quinlan EM, Philpot BD, Huganir RL, Bear MF. Rapid, experience-dependent expression of synaptic NMDA receptors in visual cortex in vivo. Nat Neurosci 2: 352–357, 1999.[CrossRef][Web of Science][Medline]
Salter MW, Kalia LV. Src kinases: a hub for NMDA receptor regulation. Nat Rev Neurosci 5: 317–328, 2004.[CrossRef][Web of Science][Medline]
Sheng M, Cummings J, Roldan LA, Jan YN, Jan LY. Changing subunit composition of heteromeric NMDA receptors during development of rat cortex. Nature 368: 144–147, 1994.[CrossRef][Medline]
Shi S, Hayashi Y, Esteban JA, Malinow R. Subunit-specific rules governing AMPA receptor trafficking to synapses in hippocampal pyramidal neurons. Cell 105: 331–343, 2001.[CrossRef][Web of Science][Medline]
Shi SH, Hayashi Y, Petralia RS, Zaman SH, Wenthold RJ, Svoboda K, Malinow R. Rapid spine delivery and redistribution of AMPA receptors after synaptic NMDA receptor activation. Science 284: 1811–1816, 1999.
Sinagra M, Verrier D, Frankova D, Korwek KM, Blahos J, Weeber EJ, Manzoni OJ, Chavis P. Reelin, very-low-density lipoprotein receptor, and apolipoprotein E receptor 2 control somatic NMDA receptor composition during hippocampal maturation in vitro. J Neurosci 25: 6127–6136, 2005.
Song I, Huganir RL. Regulation of AMPA receptors during synaptic plasticity. Trends Neurosci 25: 578–588, 2002.[CrossRef][Web of Science][Medline]
Stoppini L, Buchs PA, Muller D. A simple method for organotypic cultures of nervous tissue. J Neurosci Methods 37: 173–182, 1991.[CrossRef][Web of Science][Medline]
Trommsdorff M, Gotthardt M, Hiesberger T, Shelton J, Stockinger W, Nimpf J, Hammer RE, Richardson JA, Herz J. Reeler/Disabled-like disruption of neuronal migration in knockout mice lacking the VLDL receptor and ApoE receptor 2. Cell 97: 689–701, 1999.[CrossRef][Web of Science][Medline]
Tueting P, Costa E, Dwivedi Y, Guidotti A, Impagnatiello F, Manev R, Pesold C. The phenotypic characteristics of heterozygous reeler mouse. Neuroreport 10: 1329–1334, 1999.[Web of Science][Medline]
Weeber EJ, Beffert U, Jones C, Christian JM, Forster E, Sweatt JD, Herz J. Reelin and ApoE receptors cooperate to enhance hippocampal synaptic plasticity and learning. J Biol Chem 277: 39944–39952, 2002.
Wu G, Malinow R, Cline HT. Maturation of a central glutamatergic synapse. Science 274: 972–976, 1996.
Yabut O, Renfro A, Niu S, Swann JW, Marin O, D'Arcangelo G. Abnormal laminar position and dendrite development of interneurons in the reeler forebrain. Brain Res In press.
Yu XM, Askalan R, Keil GJ, 2nd, Salter MW. NMDA channel regulation by channel-associated protein tyrosine kinase Src. Science 275: 674–678, 1997.
Zalutsky RA, Nicoll RA. Comparison of two forms of long-term potentiation in single hippocampal neurons. Science 248: 1619–1624, 1990.
Zhu JJ, Esteban JA, Hayashi Y, Malinow R. Postnatal synaptic potentiation: delivery of GluR4-containing AMPA receptors by spontaneous activity. Nat Neurosci 3: 1098–1106, 2000.[CrossRef][Web of Science][Medline]
Zhu JJ, Malinow R. Acute versus chronic NMDA receptor blockade and synaptic AMPA receptor delivery. Nat Neurosci 5: 513–514, 2002.[Web of Science][Medline]
Zhuo M, Holtzman DM, Li Y, Osaka H, DeMaro J, Jacquin M, Bu G. Role of tissue plasminogen activator receptor LRP in hippocampal long-term potentiation. J Neurosci 20: 542–549, 2000.
Zimmer J, Gahwiler BH. Cellular and connective organization of slice cultures of the rat hippocampus and fascia dentata. J Comp Neurol 228: 432–446, 1984.[CrossRef][Web of Science][Medline]
This article has been cited by other articles:
|
|
 |
|
  H.-S. Hoe, K. J. Lee, R. S. E. Carney, J. Lee, A. Markova, J.-Y. Lee, B. W. Howell, B. T. Hyman, D. T. S. Pak, G. Bu, et al. Interaction of Reelin with Amyloid Precursor Protein Promotes Neurite Outgrowth J. Neurosci., June 10, 2009; 29(23): 7459 - 7473. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Niu, O. Yabut, and G. D'Arcangelo The Reelin Signaling Pathway Promotes Dendritic Spine Development in Hippocampal Neurons J. Neurosci., October 8, 2008; 28(41): 10339 - 10348. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
L. Groc, D. Choquet, F. A. Stephenson, D. Verrier, O. J. Manzoni, and P. Chavis NMDA Receptor Surface Trafficking and Synaptic Subunit Composition Are Developmentally Regulated by the Extracellular Matrix Protein Reelin J. Neurosci., September 19, 2007; 27(38): 10165 - 10175. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
TOP
ABSTRACT
INTRODUCTION
in the right panel) in comparison with the putative functional synapse that contains both NR1 and GluR1 (
). Note that reeler cultures show less number of NR1 and GluR1 puncta and relatively more synapses contain only NR1 (B). C: reelin supplement in reeler cultures restores the number of dendritic NR1 and GluR1 puncta. Scale bars, 20 µm. D: quantification of the number of NR1 and GluR1 puncta and the proportion of putative silent synapses. Reeler cultures showed significantly reduced number of NR1 [F(2,17) = 4.7, *P = 0.04, one-way ANOVA] and GluR1 [F(2,17) = 5.2, *P = 0.019, one-way ANOVA], while adding reelin in the culture medium eliminated the genotypic difference. Similar results were obtained for the reelin effects on the proportion of silent synapses (E). F: reeler cultures showed significantly reduced amplitude of mEPSCs [Z = 2.28, P = 0.02, compared with wild-type (WT) neurons, Kolmogorov-Smirnov test], which can be prevented by supplying reelin in the culture (Z = 0.26, P > 0.05, compared with WT neurons). Representative recording traces were displayed above the plot. Scale bar, 20 pA, 500 ms.