Deletion of the NR2A subunit prevents developmental changes of NMDA-mEPSCs in cultured mouse cerebellar granule neurones
1 Department of Physiology and Biophysics, Georgetown University School of Medicine, Washington, DC, USA
Abstract
We investigated the role N-methyl-D-aspartate (NMDA) receptor subunits play in shaping excitatory synaptic currents in cultures of cerebellar granule cells (CGCs) from NR2A knockout (NR2A–/–) and wild-type (+/+) mice. Cultures were maintained in a condition that facilitates the occurrence of functional synapses, allowing us to record NMDA-miniature excitatory postsynaptic currents (mEPSCs) in addition to NMDA receptor-mediated whole-cell currents at three ages in vitro. Whole-cell NMDA current density decreased with development in both strains though currents from NR2A–/– neurones demonstrated greater sensitivity to CP101 606, an NR2B subunit specific blocker. Sensitivity to Mg2+ blockade decreased with age in vitro in +/+ but not in NR2A–/– CGCs. Immunocytochemistry revealed that dendrites and somas displayed distinct NR1 and NR2A subunit clusters which became increasingly colocalized in +/+ neurones. Qualitatively the overall NR2B subunit staining pattern was similar in +/+ and NR2A–/– neurones throughout development, suggesting that the NR2B subunit distribution is not mediated by the NR2A subunit. In addition, staining with markers for excitatory synapses showed that expression of NR2A subunit (but not NR2B) increases at both synaptic and extrasynaptic sites in +/+ neurones during development. In parallel, NMDA-mEPSCs were faster in +/+ compared with NR2A–/– neurones at all time points studied, suggesting that the NR2A subunit begins to replace NR2B-rich NMDA receptors even at early stages of development. Many NR2A–/– neurones were devoid of NMDA-mEPSCs at the later time point, and transfection of the NR2A subunit in these neurones restored fast decay and the occurrence of NMDA-mEPSCs. Taken together, our results indicate that the NR2A subunit is mainly responsible for the developmental changes observed in the maturation of excitatory synapses.
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Introduction
N-methyl-D-aspartate (NMDA) receptors, a subclass of ionotropic glutamate receptors, mediate excitatory synaptic transmission in the mammalian CNS. A functional receptor complex is generated by the assembly of NR1 (8 members), NR2 (4 members) and possibly NR3 (2 members) subunits into a quaternary array (see Cull-Candy et al. 2001 for review). Given that individual subunits convey distinct physiological and pharmacological properties to the receptor complexes, a wide variety of receptors with distinct functional characteristics may be generated. The NMDA receptors undergo a developmentally regulated shift in their underlying subunit composition in the CNS. Specifically, the NR2B subunit (NR2B) predominates in receptor complexes at earlier stages of synaptic development, while the NR2A subunit predominates at later stages (Cull-Candy et al. 2001). Such a shift gives rise to developmental changes in synaptic physiology and has been the subject of intense investigation. The cerebellum provides an ideal model system to study changes in functional properties of native NMDA receptors with development and with subunit ablation (Cull-Candy et al. 2001). Studies in cerebellar slices have shown that the decay time course of spontaneous and evoked NMDA-excitatory postsynaptic currents (EPSCs) becomes faster throughout development (Takahashi et al. 1996; Rumbaugh & Vicini, 1999; Cathala et al. 2000; Losi et al. 2002). In addition, the decay time course of NMDA-EPSCs in NMDA receptor NR2A–/– (knockout) mice was slower than in wild-type (+/+) mice (Takahashi et al. 1996). These results suggested that receptors containing the NR2A subunit determine the fast kinetics of the NMDA-EPSCs. However, since the decay time of NMDA-EPSCs decreases with development in both +/+ and NR2A–/– mice (Takahashi et al. 1996), additional mechanisms must underlie the developmental changes of NMDA-EPSC kinetics. Excitatory synaptic currents from NR2A–/– mice have also been studied in development in superior collicular neurones in slices (Townsend et al. 2003). In these neurones, development of excitatory synapses was accompanied by the disappearance of miniature NMDA-EPSCs (NMDA-mEPSCs) in NR2A–/– but not in +/+ mice, suggesting that NR1/NR2B receptors expressed early in development become restricted to perisynaptic sites where they participate primarily in evoked currents. In an apparent contrast to these findings, NMDA receptors contributed to quantal EPSCs only at immature synapses in developing granular neurones in rat cerebellar slices (Cathala et al. 2003). Thus, the role of the NR2A subunit in the developmental redistribution of NMDA receptors remains unclear.
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When maintained in appropriate growing conditions, functional excitatory synapses are seen in primary cerebellar granule cell (CGC) cultures with the occurrence of NMDA-mEPSCs due to spontaneous vesicular release at individual synaptic sites (Chen et al. 2000; Losi et al. 2002). Beyond the fact that they offer ideal voltage-clamp control (Silver et al. 1992), cultured CGCs are well suited to address questions relating to the organization of pre- and postsynaptic elements through the use of immunocytochemistry. We combined these two approaches to study developmental changes of synaptic and extrasynaptic NMDA receptors in cultured CGCs, and to examine further the link between synaptic function at excitatory synapses and the underlying receptor composition. We show that the developmental changes reported in synaptic physiology in vivo also occur in vitro.
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Methods
Cerebellar granule cell culture and transfection
Primary cultures of cerebellar granule neurones were prepared from postnatal day 5–7 NR2A–/– or +/+ mice. The generation of NR2A–/– (1) mice is described in Takahashi et al. (1996). Mouse pups were killed by decapitation in agreement with the guidelines of the Georgetown University Animal Care and Use Committee. The cerebella were then removed, dissociated with trypsin (0.25 mg ml–1, Sigma, St Louis, MO, USA) and plated in 35 mm Nunc dishes at a density of 1.1 x 106 cells ml–1 on glass coverslips (Fisher Scientific, Pittsburgh, PA, USA) coated with poly L-lysine (10 μg ml–1, Sigma). The cells were cultured in basal Eagle's medium supplemented with 10% bovine calf serum, 2 mM glutamine, and 100 μg ml–1 gentamicin (all from Invitrogen Corporation Carlsbad, CA, USA), and maintained at 37°C in 5% CO2. The final concentration of KCl in the culture medium was adjusted to 25 mM. At 5 days in vitro (DIV), the medium was replaced with low (5 mM) K+ medium (MEM supplemented with 5 mg ml–1 glucose, 0.1 mg ml–1 transferrin, 0.025 mg ml–1 insulin, 2 mM glutamine, 20 μg ml–1 gentamicin (Invitrogen) and cytosine arabinofuranoside 10 μM, Sigma) as previously described (Chen et al. 2000; Losi et al. 2002).
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Primary cultures of mouse CGCs were transfected at DIV5 using a modification of the calcium phosphate precipitation technique (Chen & Okayama, 1987). Briefly, a glass coverslip with CGCs was transferred to a well in a four-well plate with 500 μl transfection medium composed of MEM medium (Cat no. 12370-037, Invitrogen, Carlsbad, CA, USA) with pH adjusted to 7.85 by 5 M NaOH. Then 30 μl DNA/Ca2+ mixture containing cDNA and 2 mM CaCl2 was added and incubated for 30 min at room temperature. After two washes with the transfection medium, the original culture medium was returned and neurones were maintained at 37°C in 5% CO2. EGFP plasmid (pEGFP, Clontech, Palo Alto, CA, USA) was also transfected to allow visualization of successfully transfected cells. Each coverslip was transfected with 0.3 μg pEGFP, and 1 μg of NR2A subunit plasmids. The NR2A construct has been previously described in Vicini et al. (1998).
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Electrophysiology
Coverslips with CGCs were placed on the stage of an inverted microscope (TM2000, Nikon) equipped with fluorescent and phase contrast optics. All recordings were performed at room temperature (24–26°C) from neurones maintained for 6–14 days in vitro. Continuously perfused extracellular solution contained (mM): NaCl (145), KCl (5), MgCl2 (1), CaCl2 (1), Hepes (5), glucose (5), phenol red (0.25 mg l–1) and D-serine (10 μM) (all from Sigma) adjusted to pH 7.4 with NaOH. Osmolarity was adjusted to 325 mosmol l–1 with sucrose. Electrodes were pulled in two stages on a vertical pipette puller to a resistance of 5–8 M, from borosilicate glass capillaries (Wiretrol II, Drummond, Broomall, PA, USA), and filled with recording solution containing (mM): K-gluconate (145), Hepes (10), ATP.Mg (5), GTPNa (0.2), and BAPTA (10), adjusted to pH 7.2 with KOH. Whole-cell voltage-clamp recordings from CGCs were made at –60 mV and performed at room temperature using an Axopatch 200 or an Axopatch-1D amplifier (Axon Instruments, Union City, CA, USA). A transient current response to a hyperpolarizing 10 mV pulse was used to assess access resistance and capacitance throughout the recordings. Currents were filtered at 1 kHz with an 8-pole low-pass Bessel filter (Frequency Devices, Haverhill, MA, USA), digitized at 5–10 kHz using an IBM-compatible microcomputer equipped with Digidata 1322 A data acquisition board and pCLAMP9 software (both from Axon Instruments). Off-line data analysis, curve fitting, and figure preparation were performed with Clampfit 9 software. NMDA-mEPSCs were pharmacologically isolated using bicuculline metabromide (25 μM), TTX (0.5 μM) and NBQX (5 μM) (all from Sigma) in a Mg2+-free solution. Half-maximal inhibition (IC50) of NMDA current by MgCl2 was obtained by using the following equation to fit the data:
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where IMg is the inhibition of the response at a given MgCl2 concentration and n is the Hill slope. NMDA-mEPSCs' decay was fitted using Clampfit 9 (Axon Instruments) from averages of several events selected with Minianalysis (Synaptosoft, Decatur, GA, USA).The decay phase of currents was fitted using a simplex algorithm for least squares exponential fitting routine with a double exponential equation of the form
where Ix is the peak current amplitude of a decay component and x is the corresponding decay time constant. To allow for easier comparison of decay times between experimental conditions, the two decay time components were combined into a weighted time constant
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All data are expressed as mean ± standard error of the mean, P-values represent the results of analysis of variance (ANOVA) for multiple comparisons or two-tailed unpaired Student's t tests.
Immunocytochemistry
Cultured neurones were fixed with methanol for 10 min at –20°C and subsequently permeabilized with 0.25% Triton X-100 for 3–5 min at room temperature, followed by extensive washing using phosphate-buffered saline (PBS). Coverslips were then either blocked for 1 h at room temperature or for 40 min at 37°C using 10% BSA (Sigma) in PBS, followed by incubation of the primary antibody in PBS containing 3% BSA for 1 h at 37°C. For double labelling, appropriate combinations of primary antibodies were incubated together. In some cases, NR2A or NR2B antibodies were incubated overnight at room temperature on an orbital shaker. The coverslips were then washed extensively, and the second primary antibody of interest was incubated in PBS containing 3% BSA, and 0.02% sodium azide for 1 h at 37°C. After primary incubation, the cells were treated with three, 5 min washes in PBS and subsequently incubated with secondary antibodies for 1 h at 37°C. Primary antibodies used were: rabbit anti-NR2B subunit (1: 400), mouse anti-NR1(1 μg ml–1) (gifts from Dr Wolfe, Georgetown University), rabbit anti-NR2A subunit (1.5 μg ml–1) (a gift from Dr Watanabe, Osaka Japan), rabbit antisynapsin (1 μg ml–1), rabbit anti-GluR2/3 (2 μg ml–1) (Chemicon, Temecula, CA, USA), and mouse antivGlut-1 (0.5 μg ml–1) and antivGlut-2 (1 μg ml–1) (Synaptic System, Goettingen, Germany). Secondary antibodies used were: goat antirabbit and/or antimouse Alexa 488-conjugated 1: 400 (Molecular Probe, Eugene, OR, USA), or Cy3-conjugated 1: 1000 (Jackson ImmunoResearch laboratories, West Grove, PA, USA).
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Imaging and analysis
Stained neurones were imaged on an Axioskop FS microscope (Zeiss, Germany) equipped with a 63x, 0.9 NA, Achroplan, water-immersion objective or with a Nikon E600 microscope (Nikon, Japan) equipped with a 60x, 1.0 NA. objective. Digital images were acquired with a CFW-1310 (Scion Corporation, Frederick, MD, USA), 10-bit (1024 grey scale intensity level) CCD digital camera, 1360 x 1024 pixel array. For a given antibody stain, images were acquired using identical parameters. Eight-bit images were analysed blindly with MetaMorph (Universal Imaging, Downingtown, PA, USA) after background subtraction of camera noise, and flatfield division to level the intensity. Images were pseudocoloured for presentation with Adobe Photoshop 7.0 (Adobe, San Jose, CA, USA).
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Antibody-positive clusters were defined as clusters of fluorescence that were at least twice the background fluorescence of the image. The cluster size limitation was set by the autoregion selection (0.5 x 0.5 μm) by using MetaMorph software. Receptor cluster colocalization was quantified over a length of 100 μm in 2–3 dendrites evaluated in at least eight selected neurones in three distinct cultures. NR1 and NR2 subunit-positive puncta were considered to be colocalized if clusters were touching or overlapping (centres of the puncta less than 1 μm), while colocalization of vGlut and the NR2 subunit was defined as having the centres less than 1.6 μm.
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To control variability across culture, we used mice from the same litter in each of the two strains for each culture preparation and plated the cells at the same density. In each of three sets of cultures, we selected coverslips of cells displaying similar density at the various ages studied. Coverslips of CGCs from both +/+ and NR2A–/– strains were fixed simultaneously at the two distinct DIV time points studied and the set was stained at the same time using methods as described above. To quantify changes in cluster fluorescence, we measured the average pixel intensity of each cluster along a dendritic segment. We normalized changes in fluorescence intensity to the intensity seen with the specific antibody in CGCs at DIV7 in each experimental group, to compare fluorescence intensity between distinct set of cultures. All data are expressed as mean ± standard error of the mean, P-values represent the results of two-tailed unpaired Student's t tests.
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Results
NMDA receptors in developing CGCs in cultures from +/+ and NR2A–/– mice
We used a focal application of saturating doses of NMDA (200 μM) to examine the expression of functional NMDA receptors in developing CGCs in culture. Whole-cell currents derived in this manner represent the contribution of both synaptic and non-synaptic receptors. These currents were subsequently normalized to cell capacitance to generate current density measurements. We studied CGCs at two distinct developmental ages, DIV6–8 and DIV13–14, from cultures derived from +/+ and NR2A–/– mice. As illustrated in Fig. 1A and B, the current density in response to saturating NMDA application is significantly reduced, as cultures mature. This reduction was observed in CGCs from both genotypes, however, no significant difference was observed between age-matched +/+ and NR2A–/– CGCs, suggesting that the overall number of surface receptors is identical between genotypes and furthermore, that the regulation of surface receptors is reliant on a homeostatic mechanism that is independent of the NR2A subunit presence.
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A, whole-cell currents were elicited in CGCs from NR2A–/– and +/+ mice by application of a saturating concentration of NMDA (200 μM). Application duration is indicated by solid bars above traces. B, summary of the results from the experiments shown in A, indicating a similar developmental reduction in current density in the NR2A–/– and +/+ CGCs during development. Data derived from >30 cells from four different cultures. C, representative traces from whole-cell recordings of currents elicited by 200 μM NMDA in +/+ and NR2A–/– CGCs at DIV8 in the presence or absence of CP101 606 (10 μM). D, summary of the data obtained from experiments as in C, showing that the whole cell responses from NR2A–/– CGCs are more sensitive to the NR1/NR2B blocker than +/+ CGCs. Data are shown as percentage reduction of NMDA whole-cell current by CP101 606 (10 μM) normalized to control conditions derived from >6 cells in each experimental group. **Significant difference compared to DIV6–8 (P < 0.01, two-tailed, unpaired Student's t test); *Significant difference compared to +/+ at the same age (P < 0.05; two-tailed, unpaired Student's t test); error bars: S.E.M.
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We then investigated the relative contribution of NR1/NR2B receptors to whole-cell currents using the selective antagonist CP 101 606 (Mott et al. 1998). Figure 1C shows examples of whole-cell currents elicited by NMDA in +/+ and NR2A–/– CGCs in the presence or absence of CP101 606 (10 μM). The percentage reduction of NMDA current density by CP101 606 was significantly greater in NR2A–/– than in +/+ CGCs in both the DIV8 and DIV13 groups (Fig. 1D), implying that a difference in the receptor subunit composition exists between the two strains. In addition, CP101 606 reduction was significantly larger at DIV8 than at DIV13 for +/+ but not NR2A–/– CGCs (Fig. 1D), consistent with an increased incorporation of the NR2A subunits into receptor complexes of +/+ CGCs. Of note, the percentage reduction of NMDA current density by CP101 606 in NR2A–/– CGCs at all ages tested was less than the expected decrease seen with recombinant NR1/NR2B receptors (80%, Mott et al. 1998), suggesting the presence of a residual CP101 606-insensitive current.
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The Mg2+ sensitivity of NMDA current in CGCs decreases with development in both +/+ and NR2A–/– mice (Cathala et al. 2000) as a result of the increased expression of the NR2C subunit (Kuner & Schoepfer, 1996). This prompted us to investigate the extent of dose-dependent inhibition by MgCl2 on the whole-cell NMDA current of CGCs. As seen in Fig. 2A and B, the Mg2+ inhibition was less in +/+ CGCs at DIV14 than in any other experimental group. +/+ CGCs show a rightward shift in the Mg2+ sensitivity at the later developmental stage while no such shift was observed for the NR2A–/– CGCs. Such a finding is consistent with an increase of NR2C subunits incorporating into functional receptor complexes at +/+ CGCs, while the complement of NR2A–/– surface receptors remains unchanged.
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A, whole-cell currents elicited in CGCs from +/+ mice at DIV7 (left) and DIV14 (right) by application of a saturating NMDA dose (200 μM) in the presence of MgCl2 at different concentrations as indicated. Application duration is indicated by solid bars above traces. B, summary of the results obtained from experiments as in A, illustrating a rightward shift in Mg2+ blockade in +/+ but not in NR2A–/– CGCs with age in vitro. Data derived from >20 cells in two sets of cultures. IC50s are presented in parentheses.*Significantly different in comparison to DIV7+/+ (P < 0.05); error bars: S.E.M.
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To support the electrophysiological findings and to study the distribution of NMDA receptor subunits in CGCs, we performed immunostaining of cultures from the two genotypes at the same developmental time points selected for whole-cell recordings. As seen qualitatively in example fluorescence images (Fig. 3A and B), NR2A subunit staining increased as the +/+ CGCs mature. In addition, dendrites and somas displayed distinct NR1 and NR2A subunits clusters, which were increasingly colocalized (Fig. 3A and B; arrows). We used quantitative immunocytochemistry (see Methods) to analyse NMDA receptor clusters per 10 μm of dendrite (Fig. 4A) as well as the extent of colocalization between NR1 and NR2 subunit clusters (Fig. 4B). In the case of the NR2A subunit, the cluster density increased with development (Fig. 4A) as did the average fluorescence intensity (Table 1). Immunostaining with NR2A antibodies of CGCs from NR2A–/– mice (Fig. 3C and D; arrowheads) revealed few clusters (<1.5 per 10 μm) that were poorly colocalized with the NR1 subunit (<14%), and that did not change with development, reflecting the high specificity of the NR2A antibody we have used throughout the study (see also Fig. 3A and B for comparison).
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A–D, example images illustrating an increase in NR2A staining intensity as the neurones mature, that is concomitant with a quantitative increase in NR1/NR2A colocalization. Large panels show granule cells double labelled for NR1 (green) and NR2A (red) subunits using fluorescently tagged antibodies. Exposure and thresholding for all images in the figure are identical, allowing for a qualitative appreciation of the findings. Fluorescent images are shown overlaying the matching Nomarski DIC images. Dendritic regions indicated by the white boxes in the larger panels are magnified to the right, illustrating the colocalization of NR1 and NR2A receptor subunit clusters (arrows). The separate channels are shown individually in the magnified panels (top, middle) and as an overlaid composite (bottom). Neurones from both +/+ (A and B) and NR2A–/– (C and D) mice are shown at DIV7 (A and C) and DIV14 (B and D). Arrowheads indicate clusters which are not colocalized. Note the very faint, background staining for NR2A in the NR2A–/– CGCs (C and D), illustrating the specificity of the antibody used. Scale bar 10 μm and 3 μm for insets. E–H, example fluorescence images illustrating that a decrease in NR2B staining intensity is correlated with culture age but not genotype. Legend as in A–D, but granule cells are double labelled for NR2B (red) and NR1 (green) subunits. Neurones from both +/+ (E and F) and NR2A–/– (G and H) mice are shown at DIV7 (E and G) and DIV14 (F and H).
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A, a comparison of the linear cluster density measurements (clusters(10 μm of dendrite)–1) of NR2A, NR2B or NR1 subunits in CGCs at DIV7 and DIV14 from both genotypes. Images were analysed with Metamorph Software. Antibody-positive clusters were defined as clusters of fluorescence that were at least twice the background fluorescence of the image. B, summary of quantitative results examining the colocalization of NR1 and NR2 subunits. Colocalization was based on the mean centres of fluorescent clusters being separated by <1 μm. Data were derived from at least 24 dendritic segments, in at least 18 neurones per subunit tested and 3 distinct culture preparations in each group. **Significantly different in comparison to DIV7 (P < 0.01); significantly different in comparison to +/+ at the same age (P < 0.01); error bars: S.E.M.
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In contrast to the NR2A staining pattern, we observed a decrease in NR2B staining with culture age in +/+ CGCs as well as a decrease in the colocalization between NR1 and NR2B subunits in both genotypes, trends which are shown qualitatively in Fig. 3E–H (arrows). Through quantitative assessment, we found that the NR2B subunit cluster density and the percentage of NR1 clusters opposed to NR2B clusters decreased significantly with development in both +/+ and NR2A–/– CGCs (Fig. 4A and B). Additionally, the average fluorescence intensity of NR2B subunit clusters (Table 1) decreased with development in +/+ CGCs though, significantly, it remained unchanged in NR2A–/– CGCs, a result which is consistent with the sensitivity of whole-cell current to the NR2B-specific blocker, CP101 606.
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Analysis of NR1 subunit staining revealed that, unlike the NR2 subunits, no changes in cluster density occurred with development or genotype (Fig. 4A). Of interest, NR2A–/– CGCs had significantly less NR1 cluster fluorescence intensity (Table 1) than their wild-type counterpart at DIV7, though the levels became similar to those of +/+ CGCs at DIV14.
Taken together, these results show that there are few developmental changes in the expression of the NR1 subunit in either genotype, while the NR2A subunit increases in +/+ CGCs and the NR2B subunit decreases in both genotypes. These findings indicate that a clear enrichment of NR1/NR2A and/or NR1/NR2A/NR2B receptor heteromers occurs in the +/+ population during development, agreeing with the reduction in sensitivity to CP101 606 seen with whole-cell recordings (Fig. 1D). In fact, the presence of the NR2A subunit in +/+ CGCs at DIV7 is already remarkable, as shown by immunocytochemistry and by the fact that the CP101 606 sensitivity of current recorded in these cells is significantly decreased compared to age-matched NR2A–/– CGCs.
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Developmental changes of synaptic NMDA receptors in CGCs from NR2A–/– mice
To investigate the properties of NMDA receptors located at excitatory synapses in developing CGCs, we recorded NMDA-mEPSCs in the presence of TTX, NBQX and with a solution lacking Mg2+ as previously described in detail (Losi et al. 2002; Prybylowski et al. 2002). Sample traces of NMDA-mEPSCs recorded at DIV7 in both genotypes are illustrated in Fig. 5A. While amplitude (Fig. 5C), and frequency of occurrence (range 0.01–0.2 Hz) did not change between wild-type and NR2A–/– CGCs at DIV6–7, the weighted time constant of decay (w) was significantly faster in +/+ CGCs (Fig. 5D) at all time points examined when compared to NR2A–/– CGCs. In addition, the occurrence of NMDA-mEPSCs was significantly reduced in cells from NR2A–/– mice as development progressed (Fig. 5E), although in the CGCs that did exhibit NMDA-mEPSCs, the frequency of occurrence was not different from their wild-type counterparts.
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A and B, representative traces from NMDA-mEPSCs recorded from +/+ and NR2A–/– CGCs at DIV7 (A) and DIV14 (B). NMDA-mEPSCs were pharmacologically isolated using bicuculline metabromide (25 μM), TTX (0.5 μM) and NBQX (5 μM) in a Mg2+-free solution. Each panel illustrates example traces of NMDA-mEPSCs on the left with their equally scaled fitted averages (superimposed in grey) and an indication of the respective weighted time constant (w) on the right. C and D, summary of the amplitude and kinetics from the analysis of synaptic currents in CGCs illustrating that the NMDA-mEPSCs become larger and faster with development in +/+ but not in NR2A–/– cells. Data derived from >15 cells in each experimental group from at least four different culture preparations. E, summary of the percentage of CGCs that exhibit NMDA-mEPSCs in +/+ and NR2A–/– groups at different ages showing the considerable reduction in events for NR2A–/– CGCs at later time points. Data derived from >40 cells in each experimental group from at least four different culture preparations.**Significant difference in comparison to DIV6–7 (P < 0.01); *significant difference in comparison to DIV6–7 (P < 0.05); significant difference in comparison to +/+ at the same age (P < 0.01); error bars: S.E.M.
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In CGCs from +/+ mice, significant increases in NMDA-mEPSCs' amplitude were observed in both DIV8–9 and >DIV11 cultures (Fig. 5C) compared to CGCs at DIV6–7. Of interest, w showed a significant decrease only at >DIV11, suggesting that the synaptic receptor number increases at DIV8–9 while the subunit composition changes only at the later time point (>DIV11). Changes in current amplitude were not seen in cells from NR2A–/– mice at any of the time points studied. However, like the +/+ CGCs, a significant decrease in w was observed (Fig. 5D) in the older cultures (>DIV11) compared to the other age groups.
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When recording in the presence of CP101 606 (10 μM), we observed a marked reduction in the NMDA-mEPSCs' frequency in NR2A–/– GCCs at both developmental time points, while a non-significant reduction was seen in the +/+ CGCs (Fig. 6A and B), suggesting differences in subunit composition of synaptic receptors between strains at all ages considered. NMDA-mEPSC peak amplitude was not affected by CP101 606 in +/+ CGCs while w was decreased by 23%. Further analysis of these current parameters in the NR2A–/– group was not possible since the marked reduction in NMDA-mEPSCs' frequency prevented a large enough sample size from being created.
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A, example traces of NMDA-mEPSCs recorded from +/+ and NR2A–/– CGCs at DIV7 and DIV12 in the presence and the absence of 10 μM CP101 606, a specific antagonist of NR1/NR2B receptors. B, summary of the action of 10 μM CP101 606 on the frequency of NMDA-mEPSCs recorded from CGCs at different ages in culture and in distinct genotypes. Data derived from >8 cells in each experimental group from two culture preparations. Data on NMDA-mEPSCs amplitude and decay are summarized in the Results section. *Significant difference in comparison to +/+ at the same age (P < 0.05). C, summary of the action of 300 μM spermine on the amplitude (left) and weighted decay time constant (right) of NMDA- mEPSCs recorded from CGCs at different DIV and in distinct genotypes. Data derived from >10 cells in each experimental group from two culture preparations. *Significant difference compared to control (P < 0.05; two-tailed, paired Student's t test); error bars: S.E.M.
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To gain further insight into the subunit composition of synaptic NMDA receptors in developing CGCs, we investigated the action of spermine on NMDA-mEPSCs from these neurones. Spermine has been shown to have a variety of actions on NMDA receptors such as glycine-dependent and -independent enhancement of channel activity. Such actions are selective for NMDA receptors that contain the NR2B subunit complexed together with NR1–1a, the exon5-lacking variant of the NR1 subunit (Rock & MacDonald, 1995 for review). In addition, spermine speeds up NR1–1a/NR2B channel deactivation in response to ‘synaptic-like’ agonist applications (Rumbaugh et al. 2000) and produces a voltage-dependent NMDA channel blockade that is independent of subunit composition (Rock & MacDonald, 1992).
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As shown in the summary data in Fig. 6C, spermine (300 μM) had quite distinct actions depending on culture age and genotype. In CGCs at DIV7 from both genotypes, spermine significantly decreased the weighted decay time constant, consistent with the presence of NR1–1a/NR2B complexes. In cells from NR2A–/– mice however, spermine produced an additional significant increase of the NMDA-mEPSCs' amplitude which may be a further indication that the two genotypes differ in their receptor subunit proportions (see Discussion). No significant effects were measured on NMDA-mEPSC frequency of occurrence (data not shown). In CGCs at DIV13, spermine failed to affect amplitude, frequency or decay of NMDA-mEPSCs in either genotype, although a trend to a reduced peak amplitude was noted in +/+ CGCs.
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To investigate whether the wild-type phenotype could be rescued, we transfected cDNA encoding for the NR2A subunit in NR2A–/– CGCs at DIV5, together with a vector-containing sequence encoding for EGFP (pEGFP) as a marker of successful transfection. In NR2A–/– CGCs recorded at DIV7–9, NMDA-mEPSCs' amplitude and w were not significantly different from those of currents recorded from +/+ CGCs transfected only with pEGFP (Fig. 7A and B) or from wild-type non-transfected CGCs (Fig. 5C and D), but were significantly different from those of pEGFP-transfected NR2A–/– CGCs. Most importantly, the number of cells with NMDA-mEPSCs was significantly larger than age-matched untransfected cells (Fig. 7B), suggesting that NR2A transfection restored the proper synaptic receptor complement to the NR2A–/– CGCs.
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A, NMDA-mEPSCs recorded from –/– CGCs at DIV7, which have been transfected with a plasmid that contains either pEGFP alone (top panels, pEGFP) or pEGFP together with NR2A subunit (bottom panels, NR2A and pEGFP). To the right, each panel also illustrates the respective fitted averages of NMDA-mEPSCs (superimposed in grey) with an indication of the respective weighted time constant of decay (w). B, summary of the amplitude (left) and kinetics (middle) illustrating that the NMDA-mEPSCs become larger and faster when NR2A–/– CGCs are transfected with the NR2A subunit. Data derived from >11 cells. Summary of the percentage of cells (right) with NMDA-mEPSCs (% CGCs) in +/+ and NR2A–/– groups. Data derived from >25 cells in each experimental group. **Significant difference in comarison to pEGFP-transfected CGCs (P < 0.01); error bars: S.E.M.
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To further investigate the results from electrophysiological recordings of NMDA-mediated synaptic currents, we double-labelled CGCs in cultures from the two genotypes, combining antibodies for the NMDA receptor subunits with those for vesicular glutamate transporters, presynaptic markers specific for excitatory synapses (vGlut-1 and vGlut-2, Bellocchio et al. 2000; Takamori et al. 2000; Studler et al. 2002; Wojcik et al. 2004). As previously reported (Studler et al. 2002), we found that vGlut-1-positive glutamatergic terminals in cultured CGCs rapidly increase with development in vitro (data not shown). However, CGCs also express the type 2 transporter (vGlut-2) early in development, which is gradually replaced by vGlut-1 (Wojcik et al. 2004). To accommodate the possibility that slight differences in the degree of development in our cultures may lead to disparate vGlut-1 staining patterns, we combined the antibodies to both transporters in immunocytochemical staining studies for use as presynaptic markers (designated as vGlut). The number of dendrites with excitatory synapses was limited in CGCs of both genotype at DIV7, while a large number of synapses that often made aggregates on neuritic plexi (Leao et al. 2000) were seen at DIV14. However, in individual dendrites that were innervated (Fig. 8), the linear density of vGlut-positive boutons was not significantly different between groups being 5.3 ± 0.3 in +/+ and 5.7 ± 1.4 in NR2A–/– CGCs at DIV7 (n = 80), and 5.3 ± 0.1 and 5.6 ± 0.5, respectively, at DIV14 (n = 80). Consistent with the results in Figs 3 and 4, the NR2A subunit staining increased with development in +/+ CGCs, while that of the NR2B subunit did not (Fig. 8A–D). The colocalization of vGlut terminals and glutamate receptor subunit clusters can be seen in the magnified images shown in Fig. 8A–D (arrows). The quantified data set from this study (Fig. 8E left) shows that, in +/+ CGCs, the percentage of vGlut-positive terminals located on the dendrites of CGCs facing an NR2A subunit significantly increased in one week. In contrast, a developmental decrease in the proportion of excitatory terminals opposed to NR2B-containing sites was observed in +/+ CGCs but not in N2A–/– CGCs. As expected by the reported disappearance of silent synapses in these cultures with development (Losi et al. 2002), the portion of vGlut excitatory terminals facing the AMPA receptor GluR2/3 subunit doubled with development in cultures from both genotypes (Fig. 8E left).
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A–D, large panels show examples of +/+ CGC cultures double labelled for vGlut-1 and vGlut-2 (vGlut, green) and NR2 (red) subunits using fluorescently tagged antibodies at DIV7 (A and C) and DIV14 (B and D). Images are shown with the matching Nomarski DIC image. Dendritic regions indicated by the white boxes in the larger panels are magnified to the right to illustrate the matching of vGlut and NR2 receptor subunit clusters. The separate channels are shown individually in the magnified panels (top, middle) or overlaid (bottom). Exposure and thresholding for all images in the figure are identical, allowing for a qualitative appreciation of the increase in synapses facing NR2A subunit clusters and decrease in those facing the NR2B clusters in +/+ CGCs with age in vitro. In all panels, arrows indicate synaptic receptors while arrowheads indicate extrasynaptic receptors. Scale bar 10 μm and 6 μm for insets. E, summary of data obtained from colocalization experiments using a combination of vGlut1 and 2 (VGlut) as a marker of excitatory synapses. The left panel compares the percentage of vGlut-positive boutons that face NR2 or Glur2/3 subunit clusters in CGCs (%synapses versus subunit) from different genotypes and at different ages in culture, with an indication of statistical significance with respect to wild-type (**) and DIV7 ( ). The right panel compares the percentage of NR2 subunit clusters that face vGlut positive boutons (% subunits versus synapses) in CGCs from different genotypes and at different ages in culture with an indication of statistical significance (**) with respect to DIV7. Colocalization was based on the mean centres of fluorescent clusters being separated by <1.6 μm. Data for NR2 subunit from >40 cells in three separate cultures and for Glur2/3 subunit from >12 cells in one culture. **Significantly different in comparison to +/+ at the same age (P < 0.01); significantly different in comparison to DIV7 (P < 0.01); error bars: S.E.M.
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When the synaptic localization of NR2A and NR2B subunits was investigated, few changes were seen during development in the proportion of receptors located at synapses for any of the subunits studied in +/+ CGCs (Fig. 8E right). Together with the results indicating changes in the extent of colocalization between vGlut puncta and the NR2A or NR2B subunits (Fig. 8E left), this suggest that expression of the NR2A subunit (but not NR2B) increase at both synaptic and extrasynaptic sites in +/+ neurones during development. Furthermore, the fluorescence intensity of synaptic NR2A subunit clusters in +/+ CGCs increased by 46 ± 8% between DIV7 and DIV14, while it decreased by 24 ± 5% for NR2B subunit clusters, paralleling our observations for total cluster intensity (Table 1). In contrast to these observations, the NR2B subunit showed an increase in the proportion of receptors located at synapses during development in NR2A–/– CGCs (Fig. 8E right), while the fluorescence intensity of these clusters remained unchanged. Such observations are again consistent with +/+ CGCs possessing a greater proportion of synapses that contain NR2A complexes rather than NR2B complexes later in development, while NR2B-containing complexes are retained at synapses in the absence of the NR2A subunit.
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Discussion
We examined the role played by the NR2A subunit in the developmental organization of synaptic receptor composition by using a mouse line devoid of this subunit. With a combination of immunostaining of pre- and postsynaptic elements along with electrophysiological recordings of NMDA receptor-mediated whole-cell and synaptic currents, we were able to demonstrate that the NR2A subunit is mainly responsible for the developmental changes observed in the maturation of synaptic currents: the weighted decay time of NMDA-mEPSCs is considerably reduced later in development, while the amplitude is increased; the sensitivity to Mg2+ blockade and to spermine is reduced. These results are consistent with an observed increase over the same time period in NR2A subunit cluster density, intensity and colocalization with NR1 subunit, and a concomitant decrease in these parameters for the NR2B subunit distribution. Furthermore the NR2A subunit serves a critical organizational role in the formation of functional synaptic NMDA receptors since NR2A–/– CGCs show a remarkable decrease in cells that exhibit mEPSCs. Significantly, this trend is completely reversed in NR2A–/– CGCs that have been transfected with the NR2A subunit, such that the amplitude, decay time and the occurrence of NMDA-mEPSCs are restored to the +/+ level.
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Further evidence for the importance of the NR2A subunit in orchestrating the NMDA receptor composition at synapses is demonstrated by the significant effects CP101 606 has on the frequency, and spermine has on the amplitude of NMDA-mEPSCs in NR2A–/– at DIV7 but not in +/+ CGCs (Fig. 6B and C). Since the action of these drugs is linked to the presence of NR1–1a/NR2B heteromeric assemblies, these are likely to be the main receptor type expressed at synapses in NR2A–/– CGCs at DIV7. The lack of effect by these agents on +/+ CGCs may be explained by the greater NR2A subunit contribution to synaptic currents in these cells. This indicates a difference in the synaptic receptor population depending on the presence or absence of the NR2A subunit.
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The use of NR2A–/– neurones allowed for the unmasking of the NR2A contribution to functional receptor complexes at early stages of development. NMDA-mEPSCs in +/+ CGC cultures display the same developmental changes in decay kinetics as reported for CGCs in slices (Losi et al. 2002), emphasizing the crucial role increased NR2A subunit expression plays in regulating synaptic decay. Here we show that, at all developmental time points examined, the +/+ CGCs have currents exhibiting faster rates of decay compared with those currents measured from NR2A–/– CGCs. This suggests that in our culture system, NR2A subunits are incorporated into NR2B-rich NMDA receptor complexes, even at the early stages of synapse formation.
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Recordings from CGCs in cerebellar slices from both +/+ and NR2A–/–mice show that the decay time of NMDA-EPSCs decreased with development (Takahashi et al. 1996). This finding was also observed in our cultures since NMDA-mEPSCs became faster with development in both +/+ and NR2A–/– CGCs, albeit to very different extents (Fig. 5D). Previous findings (Rumbaugh & Vicini. 1999) have shown that spermine can shorten the deactivation of currents recorded in response to rapid application of glutamate from recombinant NR1/NR2B receptors containing the NR1 splice variant which lacks the exon 5 cassette (NR1–1a). Responses from recombinant receptors containing the NR1 variant with the exon 5 cassette or the NR2A subunit however, are not affected. Conceivably, the reported developmental increase in the expression of the NR1 variant containing exon 5 (Prybylowski et al. 2000) may underlie the increase in speed of NMDA-EPSCs and the decreased spermine sensitivity of the weighted decay time with age in vitro in NR2A–/– mice (Fig. 6C).
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Our results also report changes in whole-cell NMDA current density and in the colocalization of NR1 with the NR2A or NR2B subunit. In contrast to the NR2A subunit, the NR2B subunit cluster density, fluorescence intensity and the extent of colocalization with the NR1 subunit all decrease with development in +/+ CGCs, suggesting that NR1/NR2B or NR1/NR2A/NR2B receptors are gradually lost from synaptic and non-synaptic pools, perhaps being replaced with NR1/NR2A-containing complexes in these cells. NR1 subunit cluster density did not decrease significantly with development in either genotype, although changes in fluorescence intensity were observed. A decrease in whole-cell NMDA current density with development was observed in both +/+ and NR2A–/– CGCs, a possible result of the loss in NR2B subunit that, in +/+ CGCs, cannot be compensated for by the increase in NR2A subunit expression.
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We also investigated the developmental regulation of the NR2C subunit in culture by examining the sensitivity of NMDA currents to Mg2+ in CGCs from the four experimental groups. The Mg2+ sensitivity displayed a rightward shift with age in vitro in +/+ but not in NR2A–/– CGCs (Fig. 2). In cerebellar slices, Cathala et al. (2000) showed that a similar shift in Mg2+ sensitivity at the mossy fibre–granule cell synapse was probably due to the developmental increase of NR2C subunit in the cerebellum. Such a change may also underlie the Mg2+ sensitivity shift that we now report here in +/+ CGCs. An increased NR2C subunit expression has already been reported for CGCs in culture (Vallano et al. 1996). The absence of the Mg2+ sensitivity shift that we see in the NR2A–/– CGCs may reflect a lack of change in NR2C expression level with development, in contrast to what has been reported in cerebellar slices (Cathala et al. 2000). Immunocytochemical analysis with NR2C subunit antibody was not satisfactory enough for quantitative analysis, although qualitatively the developmental profile of the staining matched the electrophysiological data (data not shown). We do not currently know the reason for the discrepancy between our in vitro results and the reported in vivo findings. We can only speculate that since NR2C expression has been reported to be dependent on neuregulin and NMDA receptor activity in cerebellar slice cultures (Ozaki et al. 1997), the decreased occurrence of synaptic activity in the NR2A–/– may have prevented the Mg2+ sensitivity shift.
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The presence of NR2C or NR2D subunits might also account for the residual CP101 606-insensitive current observed in NR2A–/– CGCs both at DIV8 and DIV13 (Fig. 1D), since heteromeric assemblies of these subunits would generate a current that is not blocked by CP101 606.
The majority of NMDA receptors in NR2A–/– CGCs must comprise NR1/NR2B subunits, since the robust sensitivity of whole-cell NMDA current to the NR2B-selective blocker remains unchanged throughout development. This suggests a compensatory expression of this subunit in the NR2A–/– CGCs or that the majority of NR2A subunits in +/+ CGCs are complexed with NR2B subunits. Western blot analysis showed no developmental differences in the decline of NR2B subunit expression in the cerebellum of both +/+ and NR2A–/– mice, suggesting the lack of compensation of NR2A by the NR2B subunit (Takahashi et al. 1996), and supporting the occurrence of NR1/NR2A/NR2B complexes in +/+ CGCs.
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Further evidence to suggest that the NR2A subunit may be expressed early in a heterotrimeric configuration is provided by our immunocytochemical studies. At DIV7, the NR2A subunit is expressed at levels far below those seen at DIV14, but above the background staining level seen in NR2A–/– cells, suggesting the presence of functional receptors. Taking into account that at DIV7, 90% of NR1 subunits colocalize with NR2B, while 50% colocalize with NR2A, at least some, if not all, of the functional NR2A receptors must be in complexes with NR1/NR2B subunits. Conceivably then, our in vitro findings reflect the in vivo situation proposed for developing cortical neurones (Stocca & Vicini, 1998), in which NR2A subunits are thought to be expressed early as part of a heterotrimeric NMDA receptor complex and only later become distinct NR1/NR2A receptors. However, the fact that NMDA current density at DIV8 in +/+ CGCs was affected significantly less than that in NR2A–/– CGCs by CP101 606, suggests that, even at early stages of development in vitro, the NR2A subunit may already be incorporated into all functional receptors and not just into synaptic ones.
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The immunocytochemistry data also showed a decrease in the colocalization between NR1 and NR2B with development in both +/+ and NR2A–/– CGC (Fig. 4B). This decrease could be explained in wild-type CGCs by the increase in NR1 subunit colocalized with NR2A and/or other NR2 subunits. However, in NR2A–/– CGCs, the presence of the NR1 subunit without a matching NR2B subunit must indicate either the presence of another subunit, such as NR2C or NR2D, or the occurrence of homomeric NR1 subunit clusters on the membrane surface or in subsurface pools.
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Similar to what has been reported in superior collicular neurones in slices (Townsend et al. 2003), development of excitatory synapses in culture was accompanied by the disappearance of NMDA-mEPSCs in NR2A–/– but not in +/+ CGCs. These data reinforce the hypothesis that NR1/NR2B receptors are expressed early in development but become excluded from the synapse at later developmental stages (Townsend et al. 2003). On the other hand, our immunohistochemical data showed an increase in synaptic localization of this subunit in NR2A –/–CGCs as development progressed (Fig. 8E right). Thus the lower occurrence of NMDA-mEPSCs in the NR2A–/– CGCs may rather be due to the overall loss of NR2B subunit clusters (Fig. 4A and B) at DIV14. An alternative possibility is that the rapid presynaptic glutamate release is not sufficient to fully activate NR2B-containing receptors due to the slow activation rate reported for this receptor (Chen et al. 2001). However, since activation rate is temperature dependent, one would expect the increase/appearance of NMDA-mEPSCs with temperature. This was not the case in at least five CGCs from NR2A–/– mice tested at 33°C (data not shown). An alternative possibility is that the NR2B subunits found at synapses in NR2A–/– CGCs at DIV14 are not incorporated into functional complexes or may be localized to intracellular pools.
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The ideal voltage-clamp control offered by cerebellar granule cells (Silver et al. 1992), combined with culture conditions that facilitate the formation of functional excitatory synapses (Chen et al. 2000; Losi et al. 2002), provided us with the necessary means to investigate in vitro the changes that occur during development of functional cerebellar synapses in vivo. We present several lines of evidence to indicate that the reported acceleration of NMDA-mediated synaptic currents and the increase of NR2A-containing complexes during development of +/+ CGCs in vivo is reproduced in vitro: (1) the decreased sensitivity with development to the NR2B antagonist CP101 606, (2) the increased NR1 and NR2A colocalization and (3) increased fluorescence intensity of NR2A subunit clusters. The increase in NMDA-mEPSCs' amplitude seen with development in +/+ CGCs has not been reported to occur in vivo (Losi et al. 2002). It is possible that in vitro the loss of the NR2B subunit is less dramatic than that occurring in vivo, allowing for the combined presence of the NR2A and NR2B subunit to enhance NMDA-mEPSC size. Alternatively, the number of NR2C or NR2B subunits may be much higher in vivo, allowing a greater proportion of these receptors to exist compared to our in vitro situation.
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The NR2A subunit not only imparts physiological differences to synaptic receptors during development, but furthermore serves a critical organizational role in the formation of functional synaptic NMDA receptors. However, this subunit does not necessarily participate in the regulation of all NMDA receptor subunits, since changes in the NR2B subunit expression occur independently as originally demonstrated in the cerebellum of developing NR2A–/– mice (Takahashi et al. 1996). Thus we provide a model system which will allow further investigation into the mechanisms that control the developmental changes of excitatory synaptic function.
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References
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Abstract
We investigated the role N-methyl-D-aspartate (NMDA) receptor subunits play in shaping excitatory synaptic currents in cultures of cerebellar granule cells (CGCs) from NR2A knockout (NR2A–/–) and wild-type (+/+) mice. Cultures were maintained in a condition that facilitates the occurrence of functional synapses, allowing us to record NMDA-miniature excitatory postsynaptic currents (mEPSCs) in addition to NMDA receptor-mediated whole-cell currents at three ages in vitro. Whole-cell NMDA current density decreased with development in both strains though currents from NR2A–/– neurones demonstrated greater sensitivity to CP101 606, an NR2B subunit specific blocker. Sensitivity to Mg2+ blockade decreased with age in vitro in +/+ but not in NR2A–/– CGCs. Immunocytochemistry revealed that dendrites and somas displayed distinct NR1 and NR2A subunit clusters which became increasingly colocalized in +/+ neurones. Qualitatively the overall NR2B subunit staining pattern was similar in +/+ and NR2A–/– neurones throughout development, suggesting that the NR2B subunit distribution is not mediated by the NR2A subunit. In addition, staining with markers for excitatory synapses showed that expression of NR2A subunit (but not NR2B) increases at both synaptic and extrasynaptic sites in +/+ neurones during development. In parallel, NMDA-mEPSCs were faster in +/+ compared with NR2A–/– neurones at all time points studied, suggesting that the NR2A subunit begins to replace NR2B-rich NMDA receptors even at early stages of development. Many NR2A–/– neurones were devoid of NMDA-mEPSCs at the later time point, and transfection of the NR2A subunit in these neurones restored fast decay and the occurrence of NMDA-mEPSCs. Taken together, our results indicate that the NR2A subunit is mainly responsible for the developmental changes observed in the maturation of excitatory synapses.
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Introduction
N-methyl-D-aspartate (NMDA) receptors, a subclass of ionotropic glutamate receptors, mediate excitatory synaptic transmission in the mammalian CNS. A functional receptor complex is generated by the assembly of NR1 (8 members), NR2 (4 members) and possibly NR3 (2 members) subunits into a quaternary array (see Cull-Candy et al. 2001 for review). Given that individual subunits convey distinct physiological and pharmacological properties to the receptor complexes, a wide variety of receptors with distinct functional characteristics may be generated. The NMDA receptors undergo a developmentally regulated shift in their underlying subunit composition in the CNS. Specifically, the NR2B subunit (NR2B) predominates in receptor complexes at earlier stages of synaptic development, while the NR2A subunit predominates at later stages (Cull-Candy et al. 2001). Such a shift gives rise to developmental changes in synaptic physiology and has been the subject of intense investigation. The cerebellum provides an ideal model system to study changes in functional properties of native NMDA receptors with development and with subunit ablation (Cull-Candy et al. 2001). Studies in cerebellar slices have shown that the decay time course of spontaneous and evoked NMDA-excitatory postsynaptic currents (EPSCs) becomes faster throughout development (Takahashi et al. 1996; Rumbaugh & Vicini, 1999; Cathala et al. 2000; Losi et al. 2002). In addition, the decay time course of NMDA-EPSCs in NMDA receptor NR2A–/– (knockout) mice was slower than in wild-type (+/+) mice (Takahashi et al. 1996). These results suggested that receptors containing the NR2A subunit determine the fast kinetics of the NMDA-EPSCs. However, since the decay time of NMDA-EPSCs decreases with development in both +/+ and NR2A–/– mice (Takahashi et al. 1996), additional mechanisms must underlie the developmental changes of NMDA-EPSC kinetics. Excitatory synaptic currents from NR2A–/– mice have also been studied in development in superior collicular neurones in slices (Townsend et al. 2003). In these neurones, development of excitatory synapses was accompanied by the disappearance of miniature NMDA-EPSCs (NMDA-mEPSCs) in NR2A–/– but not in +/+ mice, suggesting that NR1/NR2B receptors expressed early in development become restricted to perisynaptic sites where they participate primarily in evoked currents. In an apparent contrast to these findings, NMDA receptors contributed to quantal EPSCs only at immature synapses in developing granular neurones in rat cerebellar slices (Cathala et al. 2003). Thus, the role of the NR2A subunit in the developmental redistribution of NMDA receptors remains unclear.
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When maintained in appropriate growing conditions, functional excitatory synapses are seen in primary cerebellar granule cell (CGC) cultures with the occurrence of NMDA-mEPSCs due to spontaneous vesicular release at individual synaptic sites (Chen et al. 2000; Losi et al. 2002). Beyond the fact that they offer ideal voltage-clamp control (Silver et al. 1992), cultured CGCs are well suited to address questions relating to the organization of pre- and postsynaptic elements through the use of immunocytochemistry. We combined these two approaches to study developmental changes of synaptic and extrasynaptic NMDA receptors in cultured CGCs, and to examine further the link between synaptic function at excitatory synapses and the underlying receptor composition. We show that the developmental changes reported in synaptic physiology in vivo also occur in vitro.
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Methods
Cerebellar granule cell culture and transfection
Primary cultures of cerebellar granule neurones were prepared from postnatal day 5–7 NR2A–/– or +/+ mice. The generation of NR2A–/– (1) mice is described in Takahashi et al. (1996). Mouse pups were killed by decapitation in agreement with the guidelines of the Georgetown University Animal Care and Use Committee. The cerebella were then removed, dissociated with trypsin (0.25 mg ml–1, Sigma, St Louis, MO, USA) and plated in 35 mm Nunc dishes at a density of 1.1 x 106 cells ml–1 on glass coverslips (Fisher Scientific, Pittsburgh, PA, USA) coated with poly L-lysine (10 μg ml–1, Sigma). The cells were cultured in basal Eagle's medium supplemented with 10% bovine calf serum, 2 mM glutamine, and 100 μg ml–1 gentamicin (all from Invitrogen Corporation Carlsbad, CA, USA), and maintained at 37°C in 5% CO2. The final concentration of KCl in the culture medium was adjusted to 25 mM. At 5 days in vitro (DIV), the medium was replaced with low (5 mM) K+ medium (MEM supplemented with 5 mg ml–1 glucose, 0.1 mg ml–1 transferrin, 0.025 mg ml–1 insulin, 2 mM glutamine, 20 μg ml–1 gentamicin (Invitrogen) and cytosine arabinofuranoside 10 μM, Sigma) as previously described (Chen et al. 2000; Losi et al. 2002).
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Primary cultures of mouse CGCs were transfected at DIV5 using a modification of the calcium phosphate precipitation technique (Chen & Okayama, 1987). Briefly, a glass coverslip with CGCs was transferred to a well in a four-well plate with 500 μl transfection medium composed of MEM medium (Cat no. 12370-037, Invitrogen, Carlsbad, CA, USA) with pH adjusted to 7.85 by 5 M NaOH. Then 30 μl DNA/Ca2+ mixture containing cDNA and 2 mM CaCl2 was added and incubated for 30 min at room temperature. After two washes with the transfection medium, the original culture medium was returned and neurones were maintained at 37°C in 5% CO2. EGFP plasmid (pEGFP, Clontech, Palo Alto, CA, USA) was also transfected to allow visualization of successfully transfected cells. Each coverslip was transfected with 0.3 μg pEGFP, and 1 μg of NR2A subunit plasmids. The NR2A construct has been previously described in Vicini et al. (1998).
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Electrophysiology
Coverslips with CGCs were placed on the stage of an inverted microscope (TM2000, Nikon) equipped with fluorescent and phase contrast optics. All recordings were performed at room temperature (24–26°C) from neurones maintained for 6–14 days in vitro. Continuously perfused extracellular solution contained (mM): NaCl (145), KCl (5), MgCl2 (1), CaCl2 (1), Hepes (5), glucose (5), phenol red (0.25 mg l–1) and D-serine (10 μM) (all from Sigma) adjusted to pH 7.4 with NaOH. Osmolarity was adjusted to 325 mosmol l–1 with sucrose. Electrodes were pulled in two stages on a vertical pipette puller to a resistance of 5–8 M, from borosilicate glass capillaries (Wiretrol II, Drummond, Broomall, PA, USA), and filled with recording solution containing (mM): K-gluconate (145), Hepes (10), ATP.Mg (5), GTPNa (0.2), and BAPTA (10), adjusted to pH 7.2 with KOH. Whole-cell voltage-clamp recordings from CGCs were made at –60 mV and performed at room temperature using an Axopatch 200 or an Axopatch-1D amplifier (Axon Instruments, Union City, CA, USA). A transient current response to a hyperpolarizing 10 mV pulse was used to assess access resistance and capacitance throughout the recordings. Currents were filtered at 1 kHz with an 8-pole low-pass Bessel filter (Frequency Devices, Haverhill, MA, USA), digitized at 5–10 kHz using an IBM-compatible microcomputer equipped with Digidata 1322 A data acquisition board and pCLAMP9 software (both from Axon Instruments). Off-line data analysis, curve fitting, and figure preparation were performed with Clampfit 9 software. NMDA-mEPSCs were pharmacologically isolated using bicuculline metabromide (25 μM), TTX (0.5 μM) and NBQX (5 μM) (all from Sigma) in a Mg2+-free solution. Half-maximal inhibition (IC50) of NMDA current by MgCl2 was obtained by using the following equation to fit the data:
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where IMg is the inhibition of the response at a given MgCl2 concentration and n is the Hill slope. NMDA-mEPSCs' decay was fitted using Clampfit 9 (Axon Instruments) from averages of several events selected with Minianalysis (Synaptosoft, Decatur, GA, USA).The decay phase of currents was fitted using a simplex algorithm for least squares exponential fitting routine with a double exponential equation of the form
where Ix is the peak current amplitude of a decay component and x is the corresponding decay time constant. To allow for easier comparison of decay times between experimental conditions, the two decay time components were combined into a weighted time constant
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All data are expressed as mean ± standard error of the mean, P-values represent the results of analysis of variance (ANOVA) for multiple comparisons or two-tailed unpaired Student's t tests.
Immunocytochemistry
Cultured neurones were fixed with methanol for 10 min at –20°C and subsequently permeabilized with 0.25% Triton X-100 for 3–5 min at room temperature, followed by extensive washing using phosphate-buffered saline (PBS). Coverslips were then either blocked for 1 h at room temperature or for 40 min at 37°C using 10% BSA (Sigma) in PBS, followed by incubation of the primary antibody in PBS containing 3% BSA for 1 h at 37°C. For double labelling, appropriate combinations of primary antibodies were incubated together. In some cases, NR2A or NR2B antibodies were incubated overnight at room temperature on an orbital shaker. The coverslips were then washed extensively, and the second primary antibody of interest was incubated in PBS containing 3% BSA, and 0.02% sodium azide for 1 h at 37°C. After primary incubation, the cells were treated with three, 5 min washes in PBS and subsequently incubated with secondary antibodies for 1 h at 37°C. Primary antibodies used were: rabbit anti-NR2B subunit (1: 400), mouse anti-NR1(1 μg ml–1) (gifts from Dr Wolfe, Georgetown University), rabbit anti-NR2A subunit (1.5 μg ml–1) (a gift from Dr Watanabe, Osaka Japan), rabbit antisynapsin (1 μg ml–1), rabbit anti-GluR2/3 (2 μg ml–1) (Chemicon, Temecula, CA, USA), and mouse antivGlut-1 (0.5 μg ml–1) and antivGlut-2 (1 μg ml–1) (Synaptic System, Goettingen, Germany). Secondary antibodies used were: goat antirabbit and/or antimouse Alexa 488-conjugated 1: 400 (Molecular Probe, Eugene, OR, USA), or Cy3-conjugated 1: 1000 (Jackson ImmunoResearch laboratories, West Grove, PA, USA).
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Imaging and analysis
Stained neurones were imaged on an Axioskop FS microscope (Zeiss, Germany) equipped with a 63x, 0.9 NA, Achroplan, water-immersion objective or with a Nikon E600 microscope (Nikon, Japan) equipped with a 60x, 1.0 NA. objective. Digital images were acquired with a CFW-1310 (Scion Corporation, Frederick, MD, USA), 10-bit (1024 grey scale intensity level) CCD digital camera, 1360 x 1024 pixel array. For a given antibody stain, images were acquired using identical parameters. Eight-bit images were analysed blindly with MetaMorph (Universal Imaging, Downingtown, PA, USA) after background subtraction of camera noise, and flatfield division to level the intensity. Images were pseudocoloured for presentation with Adobe Photoshop 7.0 (Adobe, San Jose, CA, USA).
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Antibody-positive clusters were defined as clusters of fluorescence that were at least twice the background fluorescence of the image. The cluster size limitation was set by the autoregion selection (0.5 x 0.5 μm) by using MetaMorph software. Receptor cluster colocalization was quantified over a length of 100 μm in 2–3 dendrites evaluated in at least eight selected neurones in three distinct cultures. NR1 and NR2 subunit-positive puncta were considered to be colocalized if clusters were touching or overlapping (centres of the puncta less than 1 μm), while colocalization of vGlut and the NR2 subunit was defined as having the centres less than 1.6 μm.
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To control variability across culture, we used mice from the same litter in each of the two strains for each culture preparation and plated the cells at the same density. In each of three sets of cultures, we selected coverslips of cells displaying similar density at the various ages studied. Coverslips of CGCs from both +/+ and NR2A–/– strains were fixed simultaneously at the two distinct DIV time points studied and the set was stained at the same time using methods as described above. To quantify changes in cluster fluorescence, we measured the average pixel intensity of each cluster along a dendritic segment. We normalized changes in fluorescence intensity to the intensity seen with the specific antibody in CGCs at DIV7 in each experimental group, to compare fluorescence intensity between distinct set of cultures. All data are expressed as mean ± standard error of the mean, P-values represent the results of two-tailed unpaired Student's t tests.
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Results
NMDA receptors in developing CGCs in cultures from +/+ and NR2A–/– mice
We used a focal application of saturating doses of NMDA (200 μM) to examine the expression of functional NMDA receptors in developing CGCs in culture. Whole-cell currents derived in this manner represent the contribution of both synaptic and non-synaptic receptors. These currents were subsequently normalized to cell capacitance to generate current density measurements. We studied CGCs at two distinct developmental ages, DIV6–8 and DIV13–14, from cultures derived from +/+ and NR2A–/– mice. As illustrated in Fig. 1A and B, the current density in response to saturating NMDA application is significantly reduced, as cultures mature. This reduction was observed in CGCs from both genotypes, however, no significant difference was observed between age-matched +/+ and NR2A–/– CGCs, suggesting that the overall number of surface receptors is identical between genotypes and furthermore, that the regulation of surface receptors is reliant on a homeostatic mechanism that is independent of the NR2A subunit presence.
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A, whole-cell currents were elicited in CGCs from NR2A–/– and +/+ mice by application of a saturating concentration of NMDA (200 μM). Application duration is indicated by solid bars above traces. B, summary of the results from the experiments shown in A, indicating a similar developmental reduction in current density in the NR2A–/– and +/+ CGCs during development. Data derived from >30 cells from four different cultures. C, representative traces from whole-cell recordings of currents elicited by 200 μM NMDA in +/+ and NR2A–/– CGCs at DIV8 in the presence or absence of CP101 606 (10 μM). D, summary of the data obtained from experiments as in C, showing that the whole cell responses from NR2A–/– CGCs are more sensitive to the NR1/NR2B blocker than +/+ CGCs. Data are shown as percentage reduction of NMDA whole-cell current by CP101 606 (10 μM) normalized to control conditions derived from >6 cells in each experimental group. **Significant difference compared to DIV6–8 (P < 0.01, two-tailed, unpaired Student's t test); *Significant difference compared to +/+ at the same age (P < 0.05; two-tailed, unpaired Student's t test); error bars: S.E.M.
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We then investigated the relative contribution of NR1/NR2B receptors to whole-cell currents using the selective antagonist CP 101 606 (Mott et al. 1998). Figure 1C shows examples of whole-cell currents elicited by NMDA in +/+ and NR2A–/– CGCs in the presence or absence of CP101 606 (10 μM). The percentage reduction of NMDA current density by CP101 606 was significantly greater in NR2A–/– than in +/+ CGCs in both the DIV8 and DIV13 groups (Fig. 1D), implying that a difference in the receptor subunit composition exists between the two strains. In addition, CP101 606 reduction was significantly larger at DIV8 than at DIV13 for +/+ but not NR2A–/– CGCs (Fig. 1D), consistent with an increased incorporation of the NR2A subunits into receptor complexes of +/+ CGCs. Of note, the percentage reduction of NMDA current density by CP101 606 in NR2A–/– CGCs at all ages tested was less than the expected decrease seen with recombinant NR1/NR2B receptors (80%, Mott et al. 1998), suggesting the presence of a residual CP101 606-insensitive current.
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The Mg2+ sensitivity of NMDA current in CGCs decreases with development in both +/+ and NR2A–/– mice (Cathala et al. 2000) as a result of the increased expression of the NR2C subunit (Kuner & Schoepfer, 1996). This prompted us to investigate the extent of dose-dependent inhibition by MgCl2 on the whole-cell NMDA current of CGCs. As seen in Fig. 2A and B, the Mg2+ inhibition was less in +/+ CGCs at DIV14 than in any other experimental group. +/+ CGCs show a rightward shift in the Mg2+ sensitivity at the later developmental stage while no such shift was observed for the NR2A–/– CGCs. Such a finding is consistent with an increase of NR2C subunits incorporating into functional receptor complexes at +/+ CGCs, while the complement of NR2A–/– surface receptors remains unchanged.
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A, whole-cell currents elicited in CGCs from +/+ mice at DIV7 (left) and DIV14 (right) by application of a saturating NMDA dose (200 μM) in the presence of MgCl2 at different concentrations as indicated. Application duration is indicated by solid bars above traces. B, summary of the results obtained from experiments as in A, illustrating a rightward shift in Mg2+ blockade in +/+ but not in NR2A–/– CGCs with age in vitro. Data derived from >20 cells in two sets of cultures. IC50s are presented in parentheses.*Significantly different in comparison to DIV7+/+ (P < 0.05); error bars: S.E.M.
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To support the electrophysiological findings and to study the distribution of NMDA receptor subunits in CGCs, we performed immunostaining of cultures from the two genotypes at the same developmental time points selected for whole-cell recordings. As seen qualitatively in example fluorescence images (Fig. 3A and B), NR2A subunit staining increased as the +/+ CGCs mature. In addition, dendrites and somas displayed distinct NR1 and NR2A subunits clusters, which were increasingly colocalized (Fig. 3A and B; arrows). We used quantitative immunocytochemistry (see Methods) to analyse NMDA receptor clusters per 10 μm of dendrite (Fig. 4A) as well as the extent of colocalization between NR1 and NR2 subunit clusters (Fig. 4B). In the case of the NR2A subunit, the cluster density increased with development (Fig. 4A) as did the average fluorescence intensity (Table 1). Immunostaining with NR2A antibodies of CGCs from NR2A–/– mice (Fig. 3C and D; arrowheads) revealed few clusters (<1.5 per 10 μm) that were poorly colocalized with the NR1 subunit (<14%), and that did not change with development, reflecting the high specificity of the NR2A antibody we have used throughout the study (see also Fig. 3A and B for comparison).
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A–D, example images illustrating an increase in NR2A staining intensity as the neurones mature, that is concomitant with a quantitative increase in NR1/NR2A colocalization. Large panels show granule cells double labelled for NR1 (green) and NR2A (red) subunits using fluorescently tagged antibodies. Exposure and thresholding for all images in the figure are identical, allowing for a qualitative appreciation of the findings. Fluorescent images are shown overlaying the matching Nomarski DIC images. Dendritic regions indicated by the white boxes in the larger panels are magnified to the right, illustrating the colocalization of NR1 and NR2A receptor subunit clusters (arrows). The separate channels are shown individually in the magnified panels (top, middle) and as an overlaid composite (bottom). Neurones from both +/+ (A and B) and NR2A–/– (C and D) mice are shown at DIV7 (A and C) and DIV14 (B and D). Arrowheads indicate clusters which are not colocalized. Note the very faint, background staining for NR2A in the NR2A–/– CGCs (C and D), illustrating the specificity of the antibody used. Scale bar 10 μm and 3 μm for insets. E–H, example fluorescence images illustrating that a decrease in NR2B staining intensity is correlated with culture age but not genotype. Legend as in A–D, but granule cells are double labelled for NR2B (red) and NR1 (green) subunits. Neurones from both +/+ (E and F) and NR2A–/– (G and H) mice are shown at DIV7 (E and G) and DIV14 (F and H).
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A, a comparison of the linear cluster density measurements (clusters(10 μm of dendrite)–1) of NR2A, NR2B or NR1 subunits in CGCs at DIV7 and DIV14 from both genotypes. Images were analysed with Metamorph Software. Antibody-positive clusters were defined as clusters of fluorescence that were at least twice the background fluorescence of the image. B, summary of quantitative results examining the colocalization of NR1 and NR2 subunits. Colocalization was based on the mean centres of fluorescent clusters being separated by <1 μm. Data were derived from at least 24 dendritic segments, in at least 18 neurones per subunit tested and 3 distinct culture preparations in each group. **Significantly different in comparison to DIV7 (P < 0.01); significantly different in comparison to +/+ at the same age (P < 0.01); error bars: S.E.M.
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In contrast to the NR2A staining pattern, we observed a decrease in NR2B staining with culture age in +/+ CGCs as well as a decrease in the colocalization between NR1 and NR2B subunits in both genotypes, trends which are shown qualitatively in Fig. 3E–H (arrows). Through quantitative assessment, we found that the NR2B subunit cluster density and the percentage of NR1 clusters opposed to NR2B clusters decreased significantly with development in both +/+ and NR2A–/– CGCs (Fig. 4A and B). Additionally, the average fluorescence intensity of NR2B subunit clusters (Table 1) decreased with development in +/+ CGCs though, significantly, it remained unchanged in NR2A–/– CGCs, a result which is consistent with the sensitivity of whole-cell current to the NR2B-specific blocker, CP101 606.
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Analysis of NR1 subunit staining revealed that, unlike the NR2 subunits, no changes in cluster density occurred with development or genotype (Fig. 4A). Of interest, NR2A–/– CGCs had significantly less NR1 cluster fluorescence intensity (Table 1) than their wild-type counterpart at DIV7, though the levels became similar to those of +/+ CGCs at DIV14.
Taken together, these results show that there are few developmental changes in the expression of the NR1 subunit in either genotype, while the NR2A subunit increases in +/+ CGCs and the NR2B subunit decreases in both genotypes. These findings indicate that a clear enrichment of NR1/NR2A and/or NR1/NR2A/NR2B receptor heteromers occurs in the +/+ population during development, agreeing with the reduction in sensitivity to CP101 606 seen with whole-cell recordings (Fig. 1D). In fact, the presence of the NR2A subunit in +/+ CGCs at DIV7 is already remarkable, as shown by immunocytochemistry and by the fact that the CP101 606 sensitivity of current recorded in these cells is significantly decreased compared to age-matched NR2A–/– CGCs.
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Developmental changes of synaptic NMDA receptors in CGCs from NR2A–/– mice
To investigate the properties of NMDA receptors located at excitatory synapses in developing CGCs, we recorded NMDA-mEPSCs in the presence of TTX, NBQX and with a solution lacking Mg2+ as previously described in detail (Losi et al. 2002; Prybylowski et al. 2002). Sample traces of NMDA-mEPSCs recorded at DIV7 in both genotypes are illustrated in Fig. 5A. While amplitude (Fig. 5C), and frequency of occurrence (range 0.01–0.2 Hz) did not change between wild-type and NR2A–/– CGCs at DIV6–7, the weighted time constant of decay (w) was significantly faster in +/+ CGCs (Fig. 5D) at all time points examined when compared to NR2A–/– CGCs. In addition, the occurrence of NMDA-mEPSCs was significantly reduced in cells from NR2A–/– mice as development progressed (Fig. 5E), although in the CGCs that did exhibit NMDA-mEPSCs, the frequency of occurrence was not different from their wild-type counterparts.
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A and B, representative traces from NMDA-mEPSCs recorded from +/+ and NR2A–/– CGCs at DIV7 (A) and DIV14 (B). NMDA-mEPSCs were pharmacologically isolated using bicuculline metabromide (25 μM), TTX (0.5 μM) and NBQX (5 μM) in a Mg2+-free solution. Each panel illustrates example traces of NMDA-mEPSCs on the left with their equally scaled fitted averages (superimposed in grey) and an indication of the respective weighted time constant (w) on the right. C and D, summary of the amplitude and kinetics from the analysis of synaptic currents in CGCs illustrating that the NMDA-mEPSCs become larger and faster with development in +/+ but not in NR2A–/– cells. Data derived from >15 cells in each experimental group from at least four different culture preparations. E, summary of the percentage of CGCs that exhibit NMDA-mEPSCs in +/+ and NR2A–/– groups at different ages showing the considerable reduction in events for NR2A–/– CGCs at later time points. Data derived from >40 cells in each experimental group from at least four different culture preparations.**Significant difference in comparison to DIV6–7 (P < 0.01); *significant difference in comparison to DIV6–7 (P < 0.05); significant difference in comparison to +/+ at the same age (P < 0.01); error bars: S.E.M.
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In CGCs from +/+ mice, significant increases in NMDA-mEPSCs' amplitude were observed in both DIV8–9 and >DIV11 cultures (Fig. 5C) compared to CGCs at DIV6–7. Of interest, w showed a significant decrease only at >DIV11, suggesting that the synaptic receptor number increases at DIV8–9 while the subunit composition changes only at the later time point (>DIV11). Changes in current amplitude were not seen in cells from NR2A–/– mice at any of the time points studied. However, like the +/+ CGCs, a significant decrease in w was observed (Fig. 5D) in the older cultures (>DIV11) compared to the other age groups.
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When recording in the presence of CP101 606 (10 μM), we observed a marked reduction in the NMDA-mEPSCs' frequency in NR2A–/– GCCs at both developmental time points, while a non-significant reduction was seen in the +/+ CGCs (Fig. 6A and B), suggesting differences in subunit composition of synaptic receptors between strains at all ages considered. NMDA-mEPSC peak amplitude was not affected by CP101 606 in +/+ CGCs while w was decreased by 23%. Further analysis of these current parameters in the NR2A–/– group was not possible since the marked reduction in NMDA-mEPSCs' frequency prevented a large enough sample size from being created.
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A, example traces of NMDA-mEPSCs recorded from +/+ and NR2A–/– CGCs at DIV7 and DIV12 in the presence and the absence of 10 μM CP101 606, a specific antagonist of NR1/NR2B receptors. B, summary of the action of 10 μM CP101 606 on the frequency of NMDA-mEPSCs recorded from CGCs at different ages in culture and in distinct genotypes. Data derived from >8 cells in each experimental group from two culture preparations. Data on NMDA-mEPSCs amplitude and decay are summarized in the Results section. *Significant difference in comparison to +/+ at the same age (P < 0.05). C, summary of the action of 300 μM spermine on the amplitude (left) and weighted decay time constant (right) of NMDA- mEPSCs recorded from CGCs at different DIV and in distinct genotypes. Data derived from >10 cells in each experimental group from two culture preparations. *Significant difference compared to control (P < 0.05; two-tailed, paired Student's t test); error bars: S.E.M.
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To gain further insight into the subunit composition of synaptic NMDA receptors in developing CGCs, we investigated the action of spermine on NMDA-mEPSCs from these neurones. Spermine has been shown to have a variety of actions on NMDA receptors such as glycine-dependent and -independent enhancement of channel activity. Such actions are selective for NMDA receptors that contain the NR2B subunit complexed together with NR1–1a, the exon5-lacking variant of the NR1 subunit (Rock & MacDonald, 1995 for review). In addition, spermine speeds up NR1–1a/NR2B channel deactivation in response to ‘synaptic-like’ agonist applications (Rumbaugh et al. 2000) and produces a voltage-dependent NMDA channel blockade that is independent of subunit composition (Rock & MacDonald, 1992).
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As shown in the summary data in Fig. 6C, spermine (300 μM) had quite distinct actions depending on culture age and genotype. In CGCs at DIV7 from both genotypes, spermine significantly decreased the weighted decay time constant, consistent with the presence of NR1–1a/NR2B complexes. In cells from NR2A–/– mice however, spermine produced an additional significant increase of the NMDA-mEPSCs' amplitude which may be a further indication that the two genotypes differ in their receptor subunit proportions (see Discussion). No significant effects were measured on NMDA-mEPSC frequency of occurrence (data not shown). In CGCs at DIV13, spermine failed to affect amplitude, frequency or decay of NMDA-mEPSCs in either genotype, although a trend to a reduced peak amplitude was noted in +/+ CGCs.
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To investigate whether the wild-type phenotype could be rescued, we transfected cDNA encoding for the NR2A subunit in NR2A–/– CGCs at DIV5, together with a vector-containing sequence encoding for EGFP (pEGFP) as a marker of successful transfection. In NR2A–/– CGCs recorded at DIV7–9, NMDA-mEPSCs' amplitude and w were not significantly different from those of currents recorded from +/+ CGCs transfected only with pEGFP (Fig. 7A and B) or from wild-type non-transfected CGCs (Fig. 5C and D), but were significantly different from those of pEGFP-transfected NR2A–/– CGCs. Most importantly, the number of cells with NMDA-mEPSCs was significantly larger than age-matched untransfected cells (Fig. 7B), suggesting that NR2A transfection restored the proper synaptic receptor complement to the NR2A–/– CGCs.
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A, NMDA-mEPSCs recorded from –/– CGCs at DIV7, which have been transfected with a plasmid that contains either pEGFP alone (top panels, pEGFP) or pEGFP together with NR2A subunit (bottom panels, NR2A and pEGFP). To the right, each panel also illustrates the respective fitted averages of NMDA-mEPSCs (superimposed in grey) with an indication of the respective weighted time constant of decay (w). B, summary of the amplitude (left) and kinetics (middle) illustrating that the NMDA-mEPSCs become larger and faster when NR2A–/– CGCs are transfected with the NR2A subunit. Data derived from >11 cells. Summary of the percentage of cells (right) with NMDA-mEPSCs (% CGCs) in +/+ and NR2A–/– groups. Data derived from >25 cells in each experimental group. **Significant difference in comarison to pEGFP-transfected CGCs (P < 0.01); error bars: S.E.M.
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To further investigate the results from electrophysiological recordings of NMDA-mediated synaptic currents, we double-labelled CGCs in cultures from the two genotypes, combining antibodies for the NMDA receptor subunits with those for vesicular glutamate transporters, presynaptic markers specific for excitatory synapses (vGlut-1 and vGlut-2, Bellocchio et al. 2000; Takamori et al. 2000; Studler et al. 2002; Wojcik et al. 2004). As previously reported (Studler et al. 2002), we found that vGlut-1-positive glutamatergic terminals in cultured CGCs rapidly increase with development in vitro (data not shown). However, CGCs also express the type 2 transporter (vGlut-2) early in development, which is gradually replaced by vGlut-1 (Wojcik et al. 2004). To accommodate the possibility that slight differences in the degree of development in our cultures may lead to disparate vGlut-1 staining patterns, we combined the antibodies to both transporters in immunocytochemical staining studies for use as presynaptic markers (designated as vGlut). The number of dendrites with excitatory synapses was limited in CGCs of both genotype at DIV7, while a large number of synapses that often made aggregates on neuritic plexi (Leao et al. 2000) were seen at DIV14. However, in individual dendrites that were innervated (Fig. 8), the linear density of vGlut-positive boutons was not significantly different between groups being 5.3 ± 0.3 in +/+ and 5.7 ± 1.4 in NR2A–/– CGCs at DIV7 (n = 80), and 5.3 ± 0.1 and 5.6 ± 0.5, respectively, at DIV14 (n = 80). Consistent with the results in Figs 3 and 4, the NR2A subunit staining increased with development in +/+ CGCs, while that of the NR2B subunit did not (Fig. 8A–D). The colocalization of vGlut terminals and glutamate receptor subunit clusters can be seen in the magnified images shown in Fig. 8A–D (arrows). The quantified data set from this study (Fig. 8E left) shows that, in +/+ CGCs, the percentage of vGlut-positive terminals located on the dendrites of CGCs facing an NR2A subunit significantly increased in one week. In contrast, a developmental decrease in the proportion of excitatory terminals opposed to NR2B-containing sites was observed in +/+ CGCs but not in N2A–/– CGCs. As expected by the reported disappearance of silent synapses in these cultures with development (Losi et al. 2002), the portion of vGlut excitatory terminals facing the AMPA receptor GluR2/3 subunit doubled with development in cultures from both genotypes (Fig. 8E left).
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A–D, large panels show examples of +/+ CGC cultures double labelled for vGlut-1 and vGlut-2 (vGlut, green) and NR2 (red) subunits using fluorescently tagged antibodies at DIV7 (A and C) and DIV14 (B and D). Images are shown with the matching Nomarski DIC image. Dendritic regions indicated by the white boxes in the larger panels are magnified to the right to illustrate the matching of vGlut and NR2 receptor subunit clusters. The separate channels are shown individually in the magnified panels (top, middle) or overlaid (bottom). Exposure and thresholding for all images in the figure are identical, allowing for a qualitative appreciation of the increase in synapses facing NR2A subunit clusters and decrease in those facing the NR2B clusters in +/+ CGCs with age in vitro. In all panels, arrows indicate synaptic receptors while arrowheads indicate extrasynaptic receptors. Scale bar 10 μm and 6 μm for insets. E, summary of data obtained from colocalization experiments using a combination of vGlut1 and 2 (VGlut) as a marker of excitatory synapses. The left panel compares the percentage of vGlut-positive boutons that face NR2 or Glur2/3 subunit clusters in CGCs (%synapses versus subunit) from different genotypes and at different ages in culture, with an indication of statistical significance with respect to wild-type (**) and DIV7 ( ). The right panel compares the percentage of NR2 subunit clusters that face vGlut positive boutons (% subunits versus synapses) in CGCs from different genotypes and at different ages in culture with an indication of statistical significance (**) with respect to DIV7. Colocalization was based on the mean centres of fluorescent clusters being separated by <1.6 μm. Data for NR2 subunit from >40 cells in three separate cultures and for Glur2/3 subunit from >12 cells in one culture. **Significantly different in comparison to +/+ at the same age (P < 0.01); significantly different in comparison to DIV7 (P < 0.01); error bars: S.E.M.
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When the synaptic localization of NR2A and NR2B subunits was investigated, few changes were seen during development in the proportion of receptors located at synapses for any of the subunits studied in +/+ CGCs (Fig. 8E right). Together with the results indicating changes in the extent of colocalization between vGlut puncta and the NR2A or NR2B subunits (Fig. 8E left), this suggest that expression of the NR2A subunit (but not NR2B) increase at both synaptic and extrasynaptic sites in +/+ neurones during development. Furthermore, the fluorescence intensity of synaptic NR2A subunit clusters in +/+ CGCs increased by 46 ± 8% between DIV7 and DIV14, while it decreased by 24 ± 5% for NR2B subunit clusters, paralleling our observations for total cluster intensity (Table 1). In contrast to these observations, the NR2B subunit showed an increase in the proportion of receptors located at synapses during development in NR2A–/– CGCs (Fig. 8E right), while the fluorescence intensity of these clusters remained unchanged. Such observations are again consistent with +/+ CGCs possessing a greater proportion of synapses that contain NR2A complexes rather than NR2B complexes later in development, while NR2B-containing complexes are retained at synapses in the absence of the NR2A subunit.
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Discussion
We examined the role played by the NR2A subunit in the developmental organization of synaptic receptor composition by using a mouse line devoid of this subunit. With a combination of immunostaining of pre- and postsynaptic elements along with electrophysiological recordings of NMDA receptor-mediated whole-cell and synaptic currents, we were able to demonstrate that the NR2A subunit is mainly responsible for the developmental changes observed in the maturation of synaptic currents: the weighted decay time of NMDA-mEPSCs is considerably reduced later in development, while the amplitude is increased; the sensitivity to Mg2+ blockade and to spermine is reduced. These results are consistent with an observed increase over the same time period in NR2A subunit cluster density, intensity and colocalization with NR1 subunit, and a concomitant decrease in these parameters for the NR2B subunit distribution. Furthermore the NR2A subunit serves a critical organizational role in the formation of functional synaptic NMDA receptors since NR2A–/– CGCs show a remarkable decrease in cells that exhibit mEPSCs. Significantly, this trend is completely reversed in NR2A–/– CGCs that have been transfected with the NR2A subunit, such that the amplitude, decay time and the occurrence of NMDA-mEPSCs are restored to the +/+ level.
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Further evidence for the importance of the NR2A subunit in orchestrating the NMDA receptor composition at synapses is demonstrated by the significant effects CP101 606 has on the frequency, and spermine has on the amplitude of NMDA-mEPSCs in NR2A–/– at DIV7 but not in +/+ CGCs (Fig. 6B and C). Since the action of these drugs is linked to the presence of NR1–1a/NR2B heteromeric assemblies, these are likely to be the main receptor type expressed at synapses in NR2A–/– CGCs at DIV7. The lack of effect by these agents on +/+ CGCs may be explained by the greater NR2A subunit contribution to synaptic currents in these cells. This indicates a difference in the synaptic receptor population depending on the presence or absence of the NR2A subunit.
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The use of NR2A–/– neurones allowed for the unmasking of the NR2A contribution to functional receptor complexes at early stages of development. NMDA-mEPSCs in +/+ CGC cultures display the same developmental changes in decay kinetics as reported for CGCs in slices (Losi et al. 2002), emphasizing the crucial role increased NR2A subunit expression plays in regulating synaptic decay. Here we show that, at all developmental time points examined, the +/+ CGCs have currents exhibiting faster rates of decay compared with those currents measured from NR2A–/– CGCs. This suggests that in our culture system, NR2A subunits are incorporated into NR2B-rich NMDA receptor complexes, even at the early stages of synapse formation.
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Recordings from CGCs in cerebellar slices from both +/+ and NR2A–/–mice show that the decay time of NMDA-EPSCs decreased with development (Takahashi et al. 1996). This finding was also observed in our cultures since NMDA-mEPSCs became faster with development in both +/+ and NR2A–/– CGCs, albeit to very different extents (Fig. 5D). Previous findings (Rumbaugh & Vicini. 1999) have shown that spermine can shorten the deactivation of currents recorded in response to rapid application of glutamate from recombinant NR1/NR2B receptors containing the NR1 splice variant which lacks the exon 5 cassette (NR1–1a). Responses from recombinant receptors containing the NR1 variant with the exon 5 cassette or the NR2A subunit however, are not affected. Conceivably, the reported developmental increase in the expression of the NR1 variant containing exon 5 (Prybylowski et al. 2000) may underlie the increase in speed of NMDA-EPSCs and the decreased spermine sensitivity of the weighted decay time with age in vitro in NR2A–/– mice (Fig. 6C).
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Our results also report changes in whole-cell NMDA current density and in the colocalization of NR1 with the NR2A or NR2B subunit. In contrast to the NR2A subunit, the NR2B subunit cluster density, fluorescence intensity and the extent of colocalization with the NR1 subunit all decrease with development in +/+ CGCs, suggesting that NR1/NR2B or NR1/NR2A/NR2B receptors are gradually lost from synaptic and non-synaptic pools, perhaps being replaced with NR1/NR2A-containing complexes in these cells. NR1 subunit cluster density did not decrease significantly with development in either genotype, although changes in fluorescence intensity were observed. A decrease in whole-cell NMDA current density with development was observed in both +/+ and NR2A–/– CGCs, a possible result of the loss in NR2B subunit that, in +/+ CGCs, cannot be compensated for by the increase in NR2A subunit expression.
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We also investigated the developmental regulation of the NR2C subunit in culture by examining the sensitivity of NMDA currents to Mg2+ in CGCs from the four experimental groups. The Mg2+ sensitivity displayed a rightward shift with age in vitro in +/+ but not in NR2A–/– CGCs (Fig. 2). In cerebellar slices, Cathala et al. (2000) showed that a similar shift in Mg2+ sensitivity at the mossy fibre–granule cell synapse was probably due to the developmental increase of NR2C subunit in the cerebellum. Such a change may also underlie the Mg2+ sensitivity shift that we now report here in +/+ CGCs. An increased NR2C subunit expression has already been reported for CGCs in culture (Vallano et al. 1996). The absence of the Mg2+ sensitivity shift that we see in the NR2A–/– CGCs may reflect a lack of change in NR2C expression level with development, in contrast to what has been reported in cerebellar slices (Cathala et al. 2000). Immunocytochemical analysis with NR2C subunit antibody was not satisfactory enough for quantitative analysis, although qualitatively the developmental profile of the staining matched the electrophysiological data (data not shown). We do not currently know the reason for the discrepancy between our in vitro results and the reported in vivo findings. We can only speculate that since NR2C expression has been reported to be dependent on neuregulin and NMDA receptor activity in cerebellar slice cultures (Ozaki et al. 1997), the decreased occurrence of synaptic activity in the NR2A–/– may have prevented the Mg2+ sensitivity shift.
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The presence of NR2C or NR2D subunits might also account for the residual CP101 606-insensitive current observed in NR2A–/– CGCs both at DIV8 and DIV13 (Fig. 1D), since heteromeric assemblies of these subunits would generate a current that is not blocked by CP101 606.
The majority of NMDA receptors in NR2A–/– CGCs must comprise NR1/NR2B subunits, since the robust sensitivity of whole-cell NMDA current to the NR2B-selective blocker remains unchanged throughout development. This suggests a compensatory expression of this subunit in the NR2A–/– CGCs or that the majority of NR2A subunits in +/+ CGCs are complexed with NR2B subunits. Western blot analysis showed no developmental differences in the decline of NR2B subunit expression in the cerebellum of both +/+ and NR2A–/– mice, suggesting the lack of compensation of NR2A by the NR2B subunit (Takahashi et al. 1996), and supporting the occurrence of NR1/NR2A/NR2B complexes in +/+ CGCs.
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Further evidence to suggest that the NR2A subunit may be expressed early in a heterotrimeric configuration is provided by our immunocytochemical studies. At DIV7, the NR2A subunit is expressed at levels far below those seen at DIV14, but above the background staining level seen in NR2A–/– cells, suggesting the presence of functional receptors. Taking into account that at DIV7, 90% of NR1 subunits colocalize with NR2B, while 50% colocalize with NR2A, at least some, if not all, of the functional NR2A receptors must be in complexes with NR1/NR2B subunits. Conceivably then, our in vitro findings reflect the in vivo situation proposed for developing cortical neurones (Stocca & Vicini, 1998), in which NR2A subunits are thought to be expressed early as part of a heterotrimeric NMDA receptor complex and only later become distinct NR1/NR2A receptors. However, the fact that NMDA current density at DIV8 in +/+ CGCs was affected significantly less than that in NR2A–/– CGCs by CP101 606, suggests that, even at early stages of development in vitro, the NR2A subunit may already be incorporated into all functional receptors and not just into synaptic ones.
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The immunocytochemistry data also showed a decrease in the colocalization between NR1 and NR2B with development in both +/+ and NR2A–/– CGC (Fig. 4B). This decrease could be explained in wild-type CGCs by the increase in NR1 subunit colocalized with NR2A and/or other NR2 subunits. However, in NR2A–/– CGCs, the presence of the NR1 subunit without a matching NR2B subunit must indicate either the presence of another subunit, such as NR2C or NR2D, or the occurrence of homomeric NR1 subunit clusters on the membrane surface or in subsurface pools.
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Similar to what has been reported in superior collicular neurones in slices (Townsend et al. 2003), development of excitatory synapses in culture was accompanied by the disappearance of NMDA-mEPSCs in NR2A–/– but not in +/+ CGCs. These data reinforce the hypothesis that NR1/NR2B receptors are expressed early in development but become excluded from the synapse at later developmental stages (Townsend et al. 2003). On the other hand, our immunohistochemical data showed an increase in synaptic localization of this subunit in NR2A –/–CGCs as development progressed (Fig. 8E right). Thus the lower occurrence of NMDA-mEPSCs in the NR2A–/– CGCs may rather be due to the overall loss of NR2B subunit clusters (Fig. 4A and B) at DIV14. An alternative possibility is that the rapid presynaptic glutamate release is not sufficient to fully activate NR2B-containing receptors due to the slow activation rate reported for this receptor (Chen et al. 2001). However, since activation rate is temperature dependent, one would expect the increase/appearance of NMDA-mEPSCs with temperature. This was not the case in at least five CGCs from NR2A–/– mice tested at 33°C (data not shown). An alternative possibility is that the NR2B subunits found at synapses in NR2A–/– CGCs at DIV14 are not incorporated into functional complexes or may be localized to intracellular pools.
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The ideal voltage-clamp control offered by cerebellar granule cells (Silver et al. 1992), combined with culture conditions that facilitate the formation of functional excitatory synapses (Chen et al. 2000; Losi et al. 2002), provided us with the necessary means to investigate in vitro the changes that occur during development of functional cerebellar synapses in vivo. We present several lines of evidence to indicate that the reported acceleration of NMDA-mediated synaptic currents and the increase of NR2A-containing complexes during development of +/+ CGCs in vivo is reproduced in vitro: (1) the decreased sensitivity with development to the NR2B antagonist CP101 606, (2) the increased NR1 and NR2A colocalization and (3) increased fluorescence intensity of NR2A subunit clusters. The increase in NMDA-mEPSCs' amplitude seen with development in +/+ CGCs has not been reported to occur in vivo (Losi et al. 2002). It is possible that in vitro the loss of the NR2B subunit is less dramatic than that occurring in vivo, allowing for the combined presence of the NR2A and NR2B subunit to enhance NMDA-mEPSC size. Alternatively, the number of NR2C or NR2B subunits may be much higher in vivo, allowing a greater proportion of these receptors to exist compared to our in vitro situation.
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The NR2A subunit not only imparts physiological differences to synaptic receptors during development, but furthermore serves a critical organizational role in the formation of functional synaptic NMDA receptors. However, this subunit does not necessarily participate in the regulation of all NMDA receptor subunits, since changes in the NR2B subunit expression occur independently as originally demonstrated in the cerebellum of developing NR2A–/– mice (Takahashi et al. 1996). Thus we provide a model system which will allow further investigation into the mechanisms that control the developmental changes of excitatory synaptic function.
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