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NMDA receptor (NMDAR) activation is required for many forms of learning and memory as well as sensory system receptive field plasticity, yet the relative contribution of pre- and postsynaptic NMDARs over cortical development remains unknown. Here we demonstrate a rapid developmental loss of functional presynaptic NMDARs in the neocortex. Presynaptic NMDARs enhance neurotransmitter release at synapses onto visual cortex pyramidal cells in young mice (< postnatal day 20; P20), but they have no apparent effect after the onset of the critical period for receptive field plasticity (>P21). Immuno-electron microscopy revealed that the loss of presynaptic NMDAR function is likely due in part to a 50% reduction in the prevalence of presynaptic NMDARs. Coincident with the observed loss of presynaptic NMDAR function, there is an abrupt change in the mechanisms of timing-dependent long-term depression (tLTD). Induction of tLTD before the onset of the critical period requires activation of pre- but not postsynaptic NMDARs, while the induction of tLTD in older mice requires activation of postsynaptic NMDARs. By demonstrating that both pre- and postsynaptic NMDARs contribute to the induction of synaptic plasticity, and that their relative roles shift over development, our findings define a novel, and perhaps general, property of synaptic plasticity in emerging cortical circuits.
Synaptic connections in sensory cortices such as the primary visual cortex are initially sculpted in an experience-independent manner, allowing rough cortical maps and receptive field properties to emerge in the absence of sensory experience (Rakic, 1977; Mower et al., 1985; Stryker and Harris, 1986; Horton and Hocking, 1996; Crowley and Katz, 1999; Feller and Scanziani, 2005). Visual experience begins sculpting receptive field properties at eye opening (Smith and Trachtenberg, 2007). This is followed by a “critical period” where ocular dominance plasticity is particularly robust and receptive field properties are additionally refined in an experience-dependent manner (Fox and Zahs, 1994; Berardi et al., 2000; Sengpiel and Kind, 2002; Hensch, 2004). Activation of the NMDA-type glutamate receptors (NMDARs) is required for many experience-dependent forms of plasticity as well as some forms of activity-dependent plasticity that do not rely on sensory experience (Bear and Rittenhouse, 1999; Iwasato et al., 2000; Malenka and Bear, 2004). Thus, an important goal is to establish the precise role of NMDARs in key forms of cortical plasticity, and whether this changes over development.
NMDARs were traditionally thought to exert their influences postsynaptically, and their presynaptic existence has been largely ignored. A surprising finding is that NMDARs are anatomically expressed presynaptically (Aoki et al., 1994; Charton et al., 1999) and these presynaptic receptors are involved in both neurotransmission and plasticity (Berretta and Jones, 1996; Woodhall et al., 2001; Casado et al., 2002; Humeau et al., 2003; Bardoni et al., 2004; Duguid and Smart, 2004; Mameli et al., 2005; Duguid and Sjöström, 2006; Lien et al., 2006; Yang et al., 2006). For example, tLTD between layer (L) 5 pyramidal cells in the visual cortex requires the simultaneous activation of presynaptic NMDARs and cannabinoid receptors (Sjöström et al., 2003). This tLTD is expressed presynaptically as a reduction in the probability of neurotransmitter release. Presynaptic NMDARs are also involved in neurotransmission in L2/3 of visual cortex (Li and Han, 2006) and tLTD induction at L2/3 in barrel cortex of young rodents (Bender et al., 2006b). While growing evidence suggests an important role for presynaptic NMDARs early in development (Lien et al., 2006), it remains unknown whether presynaptic NMDARs are regulated in a laminar or developmental fashion.
Here we examined the laminar and developmental profile of presynaptic NMDAR function in the visual cortex, a well-studied model for sensory map plasticity (Fox and Zahs, 1994; Katz and Shatz, 1996; Bear and Rittenhouse, 1999; Berardi et al., 2000; Sengpiel and Kind, 2002; Hensch, 2004; Taha and Stryker, 2005; Hofer et al., 2006). We demonstrate that while presynaptic NMDARs enhance spontaneous neurotransmission onto pyramidal neurons in L2/3, L4, and L5 early in development, there is an abrupt loss of this function at the onset of the critical period. Furthermore, presynaptic, but not postsynaptic, NMDARs are required for the induction of tLTD at the L4→L2/3 synapse during the pre-critical period. At the onset of the critical period, however, a dramatic loss of presynaptic NMDARs appears to trigger the emergence of a postsynaptic requirement of NMDARs for tLTD.
C57BL/6 mice were purchased from Charles River laboratories and used between P7 and P84. Mice were maintained on a 12:12 light:dark cycle and fed ad libitum. All experiments were performed under the animal care guidelines for Tufts University School of Medicine, the University of Virginia, and the University of North Carolina at Chapel Hill.
Mice were anesthetized with pentobarbital sodium (40 mg/kg, i.p.) and decapitated upon disappearance of corneal reflexes. Brains were rapidly removed and immersed in ice-cold dissection buffer (composition in mM: 87 NaCl, 2.5 KCl, 1.25 NaH2PO4, 25 NaHCO3, 75 sucrose, 10 dextrose, 1.3 ascorbic acid, 7 MgCl2, and 0.5 CaCl2) bubbled with 95% O2 and 5% CO2. The visual cortex was rapidly dissected and 350 µM coronal slices were prepared using a vibrating microtome (Leica VT100S). Slices were allowed to recover for 20 min in a submersion chamber at 35°C filled with warmed ACSF (124 mM NaCl, 3 mM KCl, 1.25 mM Na2PO4, 26 mM NaHCO3, 1 mM MgCl2, 2 mM CaCl2, and 20 mM D-glucose, saturated with 95% O2, 5% CO2; ~315 mOsm and pH ~7.25) and then kept at room temperature until use. For recordings, visual cortex slices were placed in a submersion chamber, maintained at 30°C, and perfused with oxygenated ACSF.
Patch pipettes were pulled from thick-walled borosilicate glass. Open tip resistances were 3–6 MΩ when pipettes were filled with an internal solution typically containing (in mM) 20 KCl, 100 (K)Gluconate, 10 HEPES, 4 (Mg)ATP, 0.3 (Na)GTP, 10 (Na)Phosphocreatine with pH adjusted to 7.25 and osmolarity adjusted to ~300 mOsm with sucrose or ddH2O. For isolating NMDAR-mediated currents, internal solution contained (in mM): 102 cesium gluconate, 5 TEA-chloride, 3.7 NaCl, 20 HEPES, 0.3 sodium guanosine triphosphate, 4 magnesium adenosine triphosphate, 0.2 EGTA, 10 BAPTA, and 5 QX-314 chloride (Alomone Labs, Jerusalem, Israel). Cells were voltage-clamped in the whole-cell configuration using a patch-clamp amplifier (Multiclamp 700A; Molecular Devices), and data were acquired and analyzed using pCLAMP 9.2 software (Molecular Devices). For current-voltage (I–V) curves of isolated NMDAR responses, voltage was adjusted for the large (16 mV), empirically determined junction potential of the cesium-based internal solution. Changes in series resistance were monitored throughout the experiment by giving a test pulse and measuring the amplitude of the capacitive current. Only cells with series resistance < 30 MΩ were included for analysis. No series resistance compensation was applied. Input resistance was monitored throughout the experiment by measuring the amplitude of the steady-state current, filtered at 2 kHz, evoked from a test pulse. Only cells with <30% change in Rinput, Rseries, and Iholding were included for analysis. EPSCs were evoked from a stimulating electrode (two-conductor cluster electrodes with 75 µM tip separation, FHC Inc., Bowdoin, ME.). NMDAR currents were pharmacologically isolated by modifying the standard ACSF to contain 4 mM Mg2+, 4 mM Ca2+, 1 µM glycine, 50 µM picrotoxin, and 40 µM DNQX or 20 µM CNQX.
Excitatory miniature excitatory postsynaptic currents (mEPSCs) were recorded in the presence of blockers for voltage-gated sodium channels (tetrodotoxin, TTX; 200 nM) and GABAA receptors (picrotoxin; 50 µM) as well as 1 µM glycine. We recorded AMPA receptor-mediated mEPSCs at negative holding potentials (−80 mV) to block the postsynaptic NMDAR currents, and measured mEPSC amplitude and frequency before, during, and after bath application of the NMDAR antagonist D-APV (50µM) or D,L-APV (100 µM). D,L-APV is abbreviated as APV unless otherwise noted. Comparisons were made for (1) the last 4 minutes of a 10 minute baseline period, (2) the last 4 minutes of an 8–10 minute window following application of 100 µM APV, and (3) the last 4 minutes following drug washout lasting 15–20 minutes. We also performed experiments of the same duration without drug application as a demonstration of the stability of the recordings. Events were identified by their rapid rise time (<3 ms), and were detected using an automatic template detection program (pCLAMP, Molecular Devices) (Clements and Bekkers, 1997). The detection threshold remained constant for the duration of each experiment. All events were manually verified, and only events with a monotonic rise time and exponential decay were included in the analysis. Normalized frequency and amplitudes were used for mEPSC data analysis. Over 150 events, with an average of ~700 events, were analyzed for each data point for each cell.
The internal recording solutions used for these experiments consisted of (in mM): 100 (K)Gluconate, 20 KCl, 4 (Mg)ATP, 10 phosphocreatine, 0.3 GTP, 10 Hepes, with pH adjusted to 7.25 and osmolarity adjusted to 290–295 mOsm. Some solutions also contained 0.4% biocytin and/or MK-801 (0.5–1 mM). Picrotoxin (50 µM) was included in the bath solution for a subset of recordings. We chose to bath apply picrotoxin rather than applying it focally. While the bath application of picrotoxin during recording led to a high attrition rate for experiments due to polysynaptic activity, particularly in older mice, the bath application of picrotoxin assured a response free of inhibition. Experiments recorded in the presence or absence of picrotoxin in the younger age group were combined, as they yielded similar results, consistent with previous observations that the properties of spike timing-dependent plasticity are similar with and without the blockade of inhibition (Feldman, 2000; Froemke and Dan, 2002). In the P23-30 age group, however, tLTD could only be induced in the presence of picrotoxin to block inhibition. Therefore, at this age group experiments with and without picrotoxin were analyzed separately. L2/3 pyramidal cells were recorded in current-clamp, and weak stimulation was delivered to L4, which makes a particularly strong and vertically organized projection to L2/3 (Burkhalter, 1989). Extracellular stimulation produced a monophasic and fixed latency response, which we interpreted as predominately coming from L4 or other vertical inputs. We feel that it is unlikely that we are activating local axon collaterals, as we never evoked antidromic action potentials at the low stimulation intensities used in these studies. Baseline stimulation was delivered once every 15–18 seconds. After a 10–15 minute stable baseline, action potential (AP) and excitatory postsynaptic potential (EPSP) pairings were delivered 75–100 times (@ ~0.2Hz) with a postsynaptic action potential produced by brief (<5 ms) depolarization followed 5–25 ms later by an EPSP evoked by L4 stimulation. After pairing, stimulation was delivered for 30 minutes at baseline frequencies. Short-term plasticity (i.e. the rate of synaptic depression) was compared before and after pairing by stimulating 6 pulses at 30 Hz. Because slices in the older mice were susceptible to polysynaptic activity evoked by 30 Hz stimulation in picrotoxin, we only induced 30 Hz stimulation for brief periods (rather than every 18 seconds) before and after the tLTD induction in these older mice.
A change in the amount of synaptic depression between the baseline responses and the responses after the induction of tLTD was quantified using a short-term depression (STD) index described by Sjöström et al. (2003). The change in the amount of synaptic depression observed in a train of six EPSPs evoked at 30 Hz was compared between the average responses in the first 10 minutes (baseline) and the last 10 minutes (post-induction). Net amplitudes for each EPSP were used. The STD index is calculated as
Mice were given an overdose of Nembutal, and perfused transcardially with heparinized Tyrode solution for 3 minutes, followed by a mixture of 4% paraformaldehyde (dissolved in 0.1M phosphate buffer at pH 7.4) and 0.5% gluteraldehyde, (EM grade; Electron Microscopy Sciences) for 15 minutes. After perfusions, brains were kept within the skull and postfixed overnight in the same fixative. A vibrating microtome was used to cut 60 µM sections coronally through the visual cortex. Sections were treated with 1% NaBH4 to terminate the cross-linking actions of the fixatives, and stored free-floating at 4°C in 0.01M phosphate buffered saline (PBS) containing 0.05% sodium azide. To prepare for pre-embedding immunocytochemistry, sections were rinsed and treated in 1% bovine serum albumin (BSA) in PBS for 30 minutes, and they were incubated in a 1 µg/ml dilution of polyclonal rabbit anti-NR1 (Chemicon, CA) in PBS with 1% BSA and 0.05% sodium azide, for three days at room temperature. The sections were then rinsed and incubated in a biotinylated goat anti-rabbit secondary (Vector Labs, CA) for 2 hours, followed by 2 hours incubation in HRP-conjugated avidin-biotin complex (ABC; Vector Labs, CA). Immunoreactivity was visualized using diaminobenzidine (DAB; 0.05%) and H2O2 (0.001%). Deletion of primary antibodies eliminated all specific staining discernible at the EM level.
Immunostained sections were fixed with 2% glutaraldehyde in PB for 10 minutes, followed by 1% osmium tetroxide for 1 hour. Sections were then dehydrated in a series of alcohols (50% ETOH, 70% containing 4% uranyl acetate, 90%, and 100% twice), infiltrated with liquid resin (Embed 812, EMS, PA), and placed between two sheets of clear acetate (Aclar; Ted Pella, CA). The resin was allowed to polymerize 1–2 days at 60°C. Sections were drawn with the aid of camera lucida, and the areas to be analyzed by EM were cut and placed on flat surfaces of Beem capsule caps. These capsules then were filled with resin and left in a 60°C oven, until the resin in the capsule polymerized. Trapezoids that contained a strip of cortex from the pial surface to the white matter were prepared; these were reexamined on the light microscope to note landmarks for laminar borders. This ensured reliable laminar analysis, as described below. Ultrathin sections were cut using an ultramicrotome (Leica UMC), and these were oriented near-parallel to the surface of vibratome sections. This approach allows ultrathin sections to be taken from the area where the antibody penetration is maximal. Sections were examined on a JEOL JEM1010 microscope.
To determine cortical layer borders on the resin-embedded sections, capsule-embedded sections were drawn using camera lucida, and certain landmarks, such as the border of cell-sparse layer 1, and the position of tissue within the resin block, were marked. The drawings from each block were then compared to ultrathin sections obtained from those blocks, and the position of L2/3 was determined within the resin trapezoid. For quantitative EM analysis, the synapse was used as the main counting unit. From each immunostained brain, at least 50 adjacent but non-overlapping images were captured at 10,000X magnification, using a 16Mpixel CCD camera (SIA). The images were examined at 35- 60,000X final magnification using Image-Pro Express software. Systematic sweeps were used to locate synapses on each image. Then, the experimenter judged if the pre- or postsynaptic elements of the synapse contained any label, and evaluated the type of synaptic contact (symmetric or asymmetric). A second experimenter who was blinded to the experimental conditions reevaluated the images to confirm the quantitative analysis.
Identification of synapses and the DAB label was performed with the following considerations. A synaptic terminal was identified by the presence of at least one synaptic vesicle in contact with a plasma membrane, at least three or more vesicles within the same profile, and the parallel alignment of the postsynaptic plasma membrane with that of the terminal. A postsynaptic density was deemed DAB positive, if it contained an accumulation of black DAB chromagen at the postsynaptic membrane. The chromagen accumulation was invariably darker and irregularly shaped in contrast to the regularly shaped morphology of the unlabeled, gray postsynaptic densities. Profiles that contained DAB accumulation that was not in contact with the postsynaptic membrane were deemed unlabeled. The criterion to classify a presynaptic terminal as DAB positive was the presence of any discernible black DAB accumulation anywhere in the terminal. Typically, DAB accumulation either uniformly filled the terminal, or appeared as patches of label attached to the presynaptic membrane or nonsynaptic membranes. Excluding primary antibody from tissue processing following our regular visualization protocol eliminated all specific DAB labeling from our material. From the counts of synapses that are unlabeled or displayed labeling in the presynaptic terminal, or at the postsynaptic density, we calculated (1) the prevalence of presynaptic labeling (N presynaptic label ×100 / N all synapses), (2) the prevalence of postsynaptic labeling (N postsynaptic label × 100 / N all synapses), and (3) the ratio of presynaptic / postsynaptic label. The number of synapses that were encountered in each brain ranged from 159–420. The difference in the number of synapses examined from each brain was an outcome of the presence of different number of synapses within a predetermined number of images captured from each brain. Every synapse from all images was included in the analysis. Two brains each at P16 and P27 were analyzed. The experimenters analyzing the data were blind to either the expected outcome or the ages of the animals.
Unless otherwise noted, all drugs were purchased from Sigma (St. Louis, MO).
Means are reported as ±S.E.M, unless specified otherwise. The nonparametric Kruskal-Wallis test was used to determine statistical significance of cumulative probability histograms. Student’s t tests for were used at the p < 0.05 significance level, except where significance was corrected to p < 0.0045 for multiple comparisons using the Bonferroni method (Figure 3).
To probe for the presence of functional presynaptic NMDARs in visual cortical pyramidal cells, we used established protocols in slices from mice aged P7-P84 (Berretta and Jones, 1996; Sjöström et al., 2003; Bender et al., 2006a; Yang et al., 2006). In one strategy, postsynaptic NMDARs are first blocked with strong hyperpolarization. Then, the subsequent effects of bath applying the NMDAR antagonist APV (100 µM) are measured to reveal the effect that the remaining, presumably presynaptic, NMDARs have on spontaneous neurotransmitter release (Figure 1A). To validate this approach, we first pharmacologically isolated NMDAR-mediated EPSCs and examined the current-voltage (I–V) relationship at the L4→L2/3 synapse. L4 stimulation was adjusted to evoke ~100 pA response at +24 mV. The synaptically evoked NMDAR currents exhibited strong rectification generated from Mg2+ block (Mayer et al., 1984; Nowak et al., 1984), showing that <4% of the current remained at −80 mV compared to what would be expected from a linear I–V relationship (Figure 1B). Although strong hyperpolarization clearly blocks evoked NMDAR currents, we also wanted to verify that it blocked NMDAR currents driven by spontaneous neurotransmitter release. Our findings demonstrate that APV has no effect on the amplitude or kinetics of mEPSCs recorded at −80 mV in the presence of TTX (200 nM) (Figure 1C), suggesting that the NMDAR component of the mEPSCs was either absent or so small that is was below the detection level. To demonstrate that functional postsynaptic NMDARs can be detected in mEPSC recordings in the absence of Mg2+ block, we showed that APV blocked a long duration current recorded at −60 mV in low Mg2+ (0.1 mM)(Figure 1C). These observations demonstrate that hyperpolarization is an effective means for blocking postsynaptic NMDAR currents.
Postsynaptic NMDARs can also be blocked by including 0.5–1 mM MK-801 in the postsynaptic recording pipette (iMK-801) (Berretta and Jones, 1996; Bender et al., 2006b; Yang et al., 2006). Notably, the postsynaptic block of NMDARs with iMK-801 does not act by spillover into the extracellular medium (see Bender et al., 2006b and results in Figure 5). By examining the input output (I–O) relationship of L2/3 cells recorded at +24 mV with and without iMK-801, we demonstrate that iMK-801 blocks more than 96% of the NMDAR current evoked at 20 µA, which was our average stimulation intensity for further experiments utilizing this technique. On average we evoked 38±12.8 pA NMDAR responses under control conditions (n=6) but only 1.4±1.8 pA responses with iMK-801 (n=7; p=0.0076), similar to the >90% block of NMDAR currents that has been observed previously using iMK-801 (Berretta and Jones, 1996; Bender et al., 2006b). Importantly, iMK-801 was even more effective at eliminating postsynaptic NMDAR currents at holding potentials near rest (Figure 1E). With iMK-801 and using a stimulation intensity of 30 µA, the NMDAR current measured at −65 mV is completely blocked (0.22±5.42 pA; n=4) compared to control (−35.13±22.78 pA; n=5).
To probe for functional presynaptic NMDARs in L2/3 at P16, we first blocked postsynaptic NMDARs with hyperpolarization to −80 mV. We then measured the effects of bath applying APV on the frequency and amplitude of mEPSCs recorded in TTX. Miniature EPSCs arise from spontaneous neurotransmitter release, and their frequency typically depends on presynaptic properties, whilst their amplitude depends on postsynaptic properties. Under these conditions, APV reversibly attenuated the frequency of mEPSCs in L2/3 pyramidal cells (Figure 2A–C ; p=0.002, Kruskal-Wallis test), without affecting their amplitude or kinetics (Figure 1C and Figure 2C). Notably, control cells recorded for the same duration, but without APV application, showed no change in mEPSC frequency or amplitude (see Figure 2D). In a subset of experiments, D-APV was used to confirm that the decrease in frequency was due to a block of NMDARs and not a non-specific effect of L-APV on metabotropic glutamate receptors (Thoreson and Ulphani, 1995; Lieske and Ramirez, 2006). Taken together these observations indicate that presynaptic NMDARs tonically enhance the frequency of spontaneous release in young animals.
The observation of presynaptic NMDARs in L2/3 visual cortex of young mice prompted us to establish whether there are laminar or developmental differences in their function. We assayed presynaptic NMDARs in 4 age groups: before eye-opening (P7-P11), after eye-opening (P13-P20), during the peak of the critical period for ocular dominance plasticity (P21-P27)(Gordon and Stryker, 1996), and in adulthood (P74-P84). We found that the reduction in mEPSC frequency by APV was not limited to L2/3 at P16, as APV similarly reduced mEPSC frequency in L2/3, L4, and L5 in mice from P7-11 and P13-P20 (Figure 3A and Table 1; p’s < 0.0045, paired t-tests with a Bonferroni correction). However, there was a striking absence of an effect by APV in L2/3, L4, and L5 in older mice (P21-P27 and P74-P84; Figure 3A and Table 1; p’s > 0.05), arguing that functional presynaptic NMDARs are lost. Several lines of evidence indicate that the reduction in mEPSC frequency in young mice was due to a presynaptic change in neurotransmitter release and was not mediated by a postsynaptic mechanism (e.g. rundown or reduced postsynaptic AMPA receptor number). (1) The reduction in mEPSC frequency by APV was reversible (Figure 2B,Figure 3B; frequency in D-APV or APV = 75.3 ± 5.0 % baseline, p=0.0003; frequency after APV washout = 99.6 ± 5.0 % baseline, p=0.93 n=15). (2) We did not observe a significant reduction in mEPSC frequency in control cells that were recorded for a similar duration (Figure 3B). (3) There was no significant change in mEPSC amplitude or kinetics by APV application in all lamina at all ages studied (Figure 2D, Figure 3C,D; Table 1). Collectively, our data show that there is a developmental loss in the ability of presynaptic NMDARs to facilitate neurotransmitter release in the visual cortex. This might partially explain previous observations in cats that the NMDAR contribution to visual responses diminishes around critical period onset, at least in layers 4 and 5 (Fox et al., 1989).
One parsimonious explanation for the developmental loss of presynaptic NMDAR function is that fewer synapses contain presynaptic NMDARs in older mice. To test this, we used EM to examine the pre- and postsynaptic prevalence of asymmetric (excitatory) synapses in L2/3 containing the NR1 subunit, which is obligatory for NMDAR function (Figure 4). We examined a total of 1073 synapses from two brains each at P16 and P27 (Table 2). While the percent of synapses containing NR1 at the postsynaptic density was similar in P16 and P27 brains (77% and 75% at P16, and 84% and 75% at P27), there was a 50% developmental reduction in NR1 at presynaptic sites (Figure 4E). Specifically, a majority (>60%) of presynaptic terminals contained NR1 in each of two brains at P16 (n=240 and 420 synapses examined), while less than 30% of the presynaptic sites were labeled in two P27 brains (n=254 and 159 synapses examined). Postsynaptic NR1, on the other hand, was above 75% at both ages, revealing approximately a 50% developmental reduction in the ratio of pre- versus postsynaptic localization of NR1 (Table 2). It is unlikely that age-dependent differences in the quality of either the perfusions or antibody penetration into the tissue confounded this analysis, because pre- and postsynaptic profiles in each brain shared the same factors that may influence the outcomes of the immunocytochemistry (such as caliber of pre- and postsynaptic profiles, size of extracellular space, and astrocytic processes). Similarly, it is unlikely that the reduction in presynaptic NR1 prevalence is a consequence of reduced antibody penetration and generally light presynaptic labeling, combined. This is because it is in fact easier to detect DAB label in a terminal than in a postsynaptic profile due to presence of gray postsynaptic density in the latter. The observation that the prevalence of postsynaptic NR1 was unchanged during development suggests that no drastic changes have occurred in immunolabeling quality between the two ages examined. Thus, the anatomical reduction in presynaptic NMDARs correlates with our physiological data demonstrating a dramatic developmental loss of functional presynaptic NMDARs over this time course, suggestive of a causal relationship.
What is the functional significance of this developmental loss of presynaptic NMDARs to synaptic plasticity during development? Presynaptic NMDARs in young rodents have been implicated in tLTD (Sjöström et al., 2003; Bender et al., 2006b), a form of synaptic plasticity thought to be important for the acquisition of receptive field properties (Dan and Poo, 2006). Thus, we predicted that the developmental loss of presynaptic NMDARs in L2/3 would either abolish the ability to induce tLTD or that there would be a different induction mechanism for tLTD later in development. To test these possibilities, we used a standard tLTD protocol by repeatedly pairing a postsynaptic action potential (AP) in a L2/3 pyramidal cell closely followed (5–25 ms) by an excitatory postsynaptic potential (EPSP) evoked by L4 stimulation. In young mice (P13-P17), AP-EPSP pairings produced strong tLTD (Figure 5A,B; 43.6 ± 5.6 % baseline slope, n=13, p<0.007). Similarly, robust tLTD could be induced when the NMDAR antagonist MK-801 was included in the recording electrode (iMK-801) to block postsynaptic NMDARs (Berretta and Jones, 1996; Bender et al., 2006b) (Figure 5C; 36.2 ± 9.1 % baseline, n=9, p=0.017). However, bath application of APV prevented induction of tLTD in young mice (Figure 5D; 95.3±9.0 % baseline, n=13, p=0.80), suggesting that the synaptic weakening relied on activation of pre- and not postsynaptic NMDARs. The absence of LTD in the presence of bath APV could be due to either a direct block of LTD induction or it could be due to an occlusion of LTD from an APV-mediated suppression of neurotransmission (Sjöström et al., 2003; Bender et al., 2006b). Either way, the block by APV demonstrates an important role for presynaptic NMDARs in tLTD induction. Notably, the iMK-801 experiments also serve as an additional control showing that iMK-801 is not acting by spillover into the extrasynaptic medium, otherwise iMK-801 would have blocked tLTD in a manner similar to APV.
To determine if the tLTD in young mice was expressed as a presynaptic reduction in neurotransmitter release, we analyzed synaptic depression in a high-frequency stimulus train of 6 pulses at 30 Hz before and after tLTD induction. The short-term depression (STD) index was used as a measure of changes in synaptic depression (see Methods). In the STD index, a negative number indicates that a manipulation reduces the rate of short-term depression. Because higher rates of short-term depression are generally associated with a higher initial probability of neurotransmitter release, a negative STD index indicates that a manipulation is likely to have lowered the probability of neurotransmitter release. By analyzing the STD index on the same cells shown in Figure 5, we observed that the rate of synaptic depression during stimulation was reduced after AP-EPSP pairings under control conditions (Figure 6A,B; STD index= −0.30±0.11, n=13) and when the recording electrode contained MK-801 (Figure 6C; STD index= −0.24±0.09, n=9). One control cell was excluded from LTD and STD analysis for technical and statistical reasons. Technically, the magnitude of LTD in this particular cell was so large that we were unable to detect an EPSP above noise level. Statistically, the data point failed the Grubb’s outlier test when we tried to analyze it to the best of our abilities. When AP-EPSP pairings were made during the bath application of APV to block all NMDARs, there was no change in the STD index (Figure 6D; STD index= −0.005±0.09, n=13). The STD index under conditions where tLTD was successfully induced (control and iMK-801 groups) was significantly different from bath APV experiments where tLTD was not induced (p <0.03). Therefore, in young mice, we conclude that the induction of tLTD in L2/3 requires the activation of presynaptic NMDARs, and this tLTD is expressed as a reduction in the probability of neurotransmitter release.
In contrast to the failure of iMK-801 to block tLTD in younger mice, postsynaptic blockade of NMDARs with iMK-801 in older mice (P23-30) prevented the induction of tLTD in L2/3 pyramidal cells (Figure 7C; n=7 p=0.79). Notably, in control conditions, AP-EPSP pairings significantly reduced synaptic strength in the visual cortex of these older mice (Figure 7A and B; 67.7 ± 7.0 % baseline, n=5, p=0.018). Collectively, these data demonstrate that tLTD can be induced in the L4→L2/3 pathway in the visual cortex of mice aged P23-30, but the induction at this older age requires postsynaptic NMDARs. These results, however, do not rule out the possibility that presynaptic NMDARs also contribute to tLTD at this older age. Our observations that postsynaptic NMDARs are required for tLTD induction in critical period mice is consistent with a previous study using a low-frequency pairing protocol to induce LTD at this age (Crozier et al., 2007). Notably, that study also demonstrated that timing-dependent LTD in younger mice did not require postsynaptic NMDARs, and our data suggest that the differential involvement of postsynaptic NMDARs in LTD is likely due to age rather than induction protocol.
Interestingly, we found a developmental shift in the capacity for tLTD induction that depended on the presence of intact inhibition in the slice. We found that, in P13-20 animals, the magnitude of tLTD was similar in the presence or absence of picrotoxin to block inhibition. Because the amount and quality of the tLTD did not differ between the two conditions in this young age group, these data were combined. In the older group, however, we found that tLTD could not be induced in the absence of picrotoxin (Figure 7D; 95.7 ± 21.0 % baseline, n=8, p=0.30). The basis for this effect is currently unknown, but it could be a consequence of the maturation of inhibition at critical period onset (Katagiri et al., 2007) and/or the onset of a shunting inhibition (Hirsch, 1995).
To determine if the tLTD in older mice was also expressed as a presynaptic reduction in neurotransmitter release, we analyzed the STD index and observed that the rate of synaptic depression during stimulation was decreased after AP-EPSP pairings under control conditions (Figure 8; STD index= −0.72±0.42, n=11). Therefore, in older mice, while the induction of tLTD in L2/3 requires the activation of postsynaptic NMDARs, it is expressed, at least in part, by a decrease in neurotransmitter release. Though a change in STD index after tLTD induction suggests a presynaptic tLTD expression mechanism, we cannot rule out the possibility that the tLTD is being simultaneously expressed postsynaptically (Lisman and Raghavachari, 2006).
A vast area of neuroscience research is dedicated to the study of NMDAR-mediated synaptic plasticity (Malenka and Bear, 2004). To date, most research has concentrated on postsynaptic NMDARs, while the role and even the very existence of presynaptic NMDARs have been controversial or largely ignored. However, presynaptic NMDARs are important to consider, as studies in different cell types, pathways, brain regions, species, and ages have found varying degrees of involvement of presynaptic and postsynaptic NMDARs in synaptic plasticity (Sjöström et al., 2003; Malenka and Bear, 2004; Froemke et al., 2005; Bender et al., 2006b; Duguid and Sjöström, 2006). Our study focused on the function of presynaptic NMDARs and their putative role in LTD, specifically within the developing mouse visual cortex. Consistent with observations in other cortical layers and other regions of the brain (Sjöström et al., 2003; Malenka and Bear, 2004; Froemke et al., 2005; Bender et al., 2006b; Duguid and Sjöström, 2006), we demonstrate that presynaptic NMDARs are present and contribute to the spike timing-dependent induction of LTD in neocortical L2/3 of young animals. This tLTD is expressed, at least in part, as a presynaptic reduction in neurotransmitter release. We also provide three key findings demonstrating that the role of presynaptic NMDARs in synaptic transmission and plasticity is highly age dependent. (1) The tonic function of presynaptic NMDARs is lost in development, at least in the assays used here. (2) There is a developmental reduction in the prevalence of presynaptic terminals containing the obligatory NR1 subunit. (3) The developmental reduction of presynaptic NMDARs may be compensated for by the emergence of a postsynaptic role for NMDARs in the induction of tLTD. Our data thus highlight the changing functions of presynaptic NMDARs over development and provide the first evidence for a developmental switch in the involvement of NMDARs in timing-dependent plasticity.
The loss of functional presynaptic NMDARs has a clear mechanistic importance for the induction of tLTD, but what is the cellular basis for this loss? An obvious possibility, which we tested, is that the prevalence of presynaptic NMDARs decreases with development. Here we show the first anatomical data to indicate both that NMDARs are present presynaptically at the majority of excitatory synapses in L2/3 early in development (P16) and that their prevalence is reduced by 50% over the next ten days. The developmental decrease in presynaptic NMDAR prevalence correlates temporally with a loss in their contribution to neurotransmitter release, suggesting a causal relationship. However, there may be additional contributing factors to the loss of presynaptic NMDAR function, as our anatomical data also indicate that presynaptic NMDARs remain at approximately 30% of the synapses even when we no longer observe their effect on mEPSC frequency. At least five possibilities exist for why we observe a complete loss of functional presynaptic NMDARs in these older mice despite our anatomical evidence that presynaptic NMDARs remain at a small subset of synapses. First, the number of NMDARs remaining at individual boutons may be below a threshold needed to have a measurable impact on neurotransmitter release. Second, presynaptic NMDARs may undergo a developmental change in their subunit composition rendering them less effective. Because functional presynaptic NMDARs in both the visual cortex (Sjöström et al., 2003) and the entorhinal cortex (Woodhall et al., 2001) are thought to contain NR2B subunits, the developmental increase in the relative expression of NR2A to NR2B NMDAR subunits in the visual cortex (Nase et al., 1999; Quinlan et al., 1999a; Quinlan et al., 1999b) may contribute to the loss of presynaptic NMDAR function. Third, the inputs that maintain presynaptic NMDARs later in development may not participate in the spontaneous release of neurotransmitter. Fourth, presynaptic NMDARs may become functionally uncoupled during development from the neurotransmitter release machinery. A final possibility is that the remaining presynaptic NMDARs may lose their efficacy due to a developmental reduction in a co-factor that is necessary for them to exert their influences. Although we did not observe a functional role for presynaptic NMDARs at later stages of development, an adult functionality for presynaptic NMDARs may become apparent in assays other than the ones used in this study.
Our data add to recent literature suggesting that presynaptic NMDARs are regulated in a developmental and activity-dependent manner (Mameli et al., 2005; Yang et al., 2006). For example, the neurosteroid pregnenolone increases the probability of glutamate release through presynaptic NMDARs in the CA3→CA1 hippocampal synapse, and this effect is thought to be lost at P5 due to a decrease in NR2D-containing NMDARs (Mameli et al., 2005). A similar but much more gradual reduction in presynaptic NMDAR function occurs in layer 5 of the rat entorhinal cortex, where their function is pronounced at ~5 weeks of age and only modest at ~5 months of age (Yang et al., 2006). Notably, in the entorhinal cortex, presynaptic NMDAR function can also be regulated in an activity-dependent manner, as the induction of seizures in adults restores juvenile levels of presynaptic NMDAR function (Yang et al., 2006). The combined data demonstrate that presynaptic NMDAR function can be bidirectionally regulated to alter neurotransmitter release. The developmental loss of presynaptic NMDARs may be a general feature of neural development, and the time course for this loss could be highly dependent on brain region. We also stress that the present study was performed in mice and that there may be important differences between species. Further studies are needed to explore these possibilities.
Why would the ability of presynaptic NMDARs to enhance neurotransmitter release diminish with development? We suggest that presynaptic NMDARs may be needed to support high neurotransmitter release probabilities. This idea is consistent with the general observations that the probability of neurotransmitter release is much greater early in development when presynaptic NMDARs are presumably present than at later stages of development when they are more likely to be absent (Bolshakov and Siegelbaum, 1995; Choi and Lovinger, 1997; Reyes and Sakmann, 1999; Silver et al., 2003; Volgushev et al., 2004). A high probability of neurotransmitter release in immature synapses may be needed for terminals to make their initial postsynaptic connections and to maintain neurotransmission when postsynaptic receptors are fewer in number (Rumpel et al., 2004). The developmental loss of the ability of presynaptic NMDARs to support neurotransmitter release may also help explain the general observation that synapses switch from depressing to facilitating during development in many brain regions (Pouzat and Hestrin, 1997; Reyes and Sakmann, 1999).
Presynaptic NMDARs can be activated from evoked glutamate release, but our study and others demonstrate that they can also be activated tonically. This suggests the receptors are activated by ambient levels of glutamate. Indeed, low-levels of ambient glutamate have been observed (Meldrum, 2000). In vitro, not only can ambient glutamate tonically activate NMDARs (Sah et al., 1989), but ambient glutamate can also activate kainate receptors (Lauri et al., 2006) and metabotropic glutamate receptors (Bandrowski et al., 2003). Interestingly, glia have recently been shown to provide a major source of ambient and evoked glutamate (Jourdain et al., 2007; Le Meur et al., 2007). Notably, the NMDAR glycine binding sites on presynaptic NMDARs appear to be saturated, unlike postsynaptic NMDARs, suggesting that presynaptic NMDARs are primed to be activated by glutamate (Li and Han, 2006). Currently it is unknown if presynaptic NMDARs require depolarization for their activation. Due to the high input resistance of presynaptic terminals, even modest excitatory currents locally at terminals might be sufficient to cause depolarization-induced removal of Mg2+ from NMDARs (Jourdain et al., 2007). Another possibility is that presynaptic NMDARs may not require depolarization to be activated; presynaptic NMDARs containing NR2C, NR2D, or NR3A all show less voltage sensitivity than other NMDAR subtypes (Monyer et al., 1992; Cull-Candy et al., 2001; Sasaki et al., 2002).
A common theme appears to be emerging that the induction mechanisms of synaptic plasticity adjust over development to adapt to ongoing changes in presynaptic and postsynaptic properties (Yasuda et al., 2003; Frenkel and Bear, 2004; Nosyreva and Huber, 2005; Yashiro et al., 2005; He et al., 2006; Jo et al., 2006). For example, there is a developmental switch in the relative presynaptic versus postsynaptic expression of metabotropic glutamate receptor-mediated LTD in the hippocampus (Nosyreva and Huber, 2005). Thus, our findings add to a growing body of literature that the properties of synaptic plasticity must be studied within a developmental context. Our data also raise the exciting possibility that presynaptic NMDARs might regulate plasticity during a pre-critical period early in development but that postsynaptic NMDARs may be more critically involved in experience-dependent synaptic plasticity later in development. Given the observations that presynaptic NMDARs exist early in development in the hippocampus (Mameli et al., 2005), entorhinal cortex (Yang et al., 2006), and visual cortex (present study) but not at later stages of maturation, we suggest that the development loss of presynaptic NMDAR function may be an important and general property of early circuit formation.
We thank J. de Marchena, M. Ehlers, R. Froemke, M. Henson, P. Manis, A. Roberts, O. Palmer, and P.J. Sjöström for constructive advice on experiments and the manuscript. This work was supported by the National Science Foundation (RC), the Zucker Research Center for Women Scholars and Charlton Award (YW), a UNC Career Development Award (BDP), NARSAD (BDP), and the Whitehall Foundation (BDP).