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1The Neurosciences Institute; and 2The Scripps Research Institute, San Diego, California
Submitted 1 March 2006; accepted in final form 4 July 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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), and is a major cause of autistic-like behaviors. The syndrome is produced by the transcriptional silencing of a single gene on the X chromosome, termed Fmr1, that encodes a protein, the Fragile X mental retardation protein (FMRP), which can act as a translational suppressor in dendrites (for review, see Bagni and Greenough 2005; Bear et al. 2004; O'Donnell and Warren 2002). Efforts to understand its neural basis have focused on mouse models in which Fmr1 transcription and translation have been disrupted (Yan et al. 2004). A major finding of these studies is that neurons in the neocortex of Fmr1 knockout (KO) mice exhibit a preponderance of abnormally long and thin dendritic spines (Bagni and Greenough 2005; Comery et al. 1997). This finding parallels data from human Fragile X patients (Irwin et al. 2000) and suggests that altered plasticity is crucial to the disorder (Huber et al. 2002; Larson et al. 2005; Li et al. 2002; Zhao et al. 2005). Many lines of evidence indicate that spine properties, including dynamic changes in morphology, play important roles in neuronal plasticity (Calverley and Jones 1990; Hayashi and Majewska 2005; Segal 2005; Yuste and Bonhoeffer 2004).
Spine abnormalities in the developing neocortex of Fmr1 KO mice appear to have a distinctive time course. An in vivo imaging study of layer V pyramidal neurons in somatosensory cortex concluded that differences in spine density and shape are most pronounced in the early postnatal period and diminish steadily throughout the first month of life (Nimchinsky et al. 2001). This conclusion was supported by a different study that found that spine abnormalities had largely disappeared by the end of the first postnatal month (only to reemerge as animals matured into adulthood) (Galvez and Greenough 2005). The temporal pattern is interesting because the early postnatal period is a time of intense synaptogenesis and a time marked by widespread, stereotyped changes in spine morphology and motility (Blue and Parnavelas 1983; Lendvai et al. 2000; Micheva and Beaulieu 1996; Nimchinsky et al. 2001). It is also a time when activity-dependent plasticity begins to be important for shaping neuronal circuits (Desai et al. 2002; Fox and Wong 2005; Stern et al. 2001; Turrigiano and Nelson 2004).
These considerations led us to ask whether early postnatal plasticity is impaired or otherwise altered in the neocortex of Fmr1 KO mice. There are many kinds of plasticity and even more experimental protocols for eliciting plasticity. We chose to focus on two that have previously been demonstrated in developing rodent neocortex (using rats), that can be induced by stimulation protocols consistent with natural spike trains, and that encompass two broad classes of neuronal plasticity: intrinsic and synaptic (Malenka and Bear 2004; Zhang and Linden 2003). The first of these, long-term potentiation of intrinsic excitability (LTP-IE), depends on activation of the group 1 metabotropic glutamate receptor mGluR5 and is mediated by changes in intrinsic membrane currents (Ireland and Abraham 2002; Sourdet et al. 2003). The second, spike timing-dependent plasticity (STDP), depends on activation of N-methyl-D-aspartate (NMDA) receptors and is mediated by changes in excitatory synaptic currents (Dan and Poo 2004; Sjöström et al. 2001).
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Early postnatal (P10-18) male C57Bl/6J-Fmr1-tm1Cgr mice (Neuromice.org) and wild-type littermates were used in these studies. For experiments involving a comparison between wild-type and KO mice, animals were chosen at random, and all experiments and data analyses were done blind to the animal's genotype (except for some of those to measure NMDA/AMPA ratios). After slice preparation on each experimental day, the tail was preserved for use in later genotyping.
Electrophysiological recordings
In all experiments, whole cell patch recordings were made from layer V pyramidal neurons in somatosensory cortex. Comparisons are between recordings made in tissue prepared from KO animals and from littermate wild-type animals. In every comparison, recordings were distributed between at least three animals in each condition (KO or wild-type). Tests for statistical significance were made using the unpaired Student's t-test unless otherwise specified. Average data are presented as means ± SE.
Brain slices were prepared and electrophysiological recordings made essentially as previously described (Desai and Walcott 2006). Briefly, coronal sections of 350 µm thickness containing primary somatosensory cortex were cut on a vibratome. Tissue was allowed to equilibrate for 
1 h before the start of experiments and was used for 
7 h after preparation. For recordings, slices were transferred one at a time to a recording chamber mounted on a fixed-stage upright microscope and were perfused with warmed (31°C), oxygenated artificial cerebrospinal fluid (ACSF). Layer V pyramidal neurons were identified at x400 magnification using infrared DIC optics and an infrared-sensitive camera. Whole cell patch recordings were obtained with pulled glass micropipettes (5–6 M
, 1- to 2-µm tip diameter), normally containing a potassium gluconate-based internal solution (see solution formulae in the following text). Liquid junction potentials (5 mV for the normal internal solution) were left uncorrected. Recordings were accepted if input resistances were >100 M, series resistances were <20 M, and membrane potentials were more negative than –55 mV. Current- and voltage-clamp recordings were made with a patch-clamp amplifier; bridge and capacitance compensation was used as appropriate. Extracellular stimulation was delivered through concentric bipolar stimulating electrodes driven by stimulus isolation units. Signals were filtered at 4 kHz and digitized at 10 kHz.
LTP-IE
LTP-IE was elicited by following the stimulation protocol of Sourdet et al. (2003). A stimulating electrode was placed in layer II/III, and the stimulation intensity was adjusted to produce an excitatory postsynaptic potential (EPSP) in the recorded layer V neuron of roughly 5-mV amplitude. Kynurenate (2 mM) and bicuculline (20 µM) were then added to the bath solution to block ionotropic synaptic transmission. A family of current steps was injected through the recording electrode into the postsynaptic neuron and a test current (500 ms long) that produced two to four spikes was selected. To monitor excitability, this test current was injected once every 10 s for the remainder of the recording, and the number of spikes it produced was counted. After a stable baseline (10 min) had been established, metabotropic glutamate receptors were stimulated for 4 min by delivering 10-Hz stimuli through the extracellular electrode; the postsynaptic neuron continued to fire during this induction period, in response to a test current injected once every 10 s. Afterward, during a test period of 30–50 min, the response of the neuron to current steps was again measured. Series resistance, input resistance, and resting membrane potential were monitored throughout. If series resistance changed by >20%, input resistance by >10%, or resting potential by >5 mV, recordings were discarded. When measuring the effect of the test current, a DC current was superimposed to keep the membrane potential to within 1 mV of its original value.
STDP
STDP was elicited according to the extracellular stimulation protocol of Sjöström et al. (2001). A whole cell patch recording was obtained from a layer V pyramidal neuron after a stimulating electrode had been placed in lower layer IV or upper layer V. (The electrode was placed more proximally than in the LTP-IE experiments so as to conform to the protocol of Sjöström et al.) The stimulation intensity was set so as to produce EPSPs of 3- to 5-mV amplitude. A number of criteria were employed to make sure that only monosynaptic EPSPs were examined: recordings that showed synaptic responses with long latency (>5 ms), excessive jitter (>0.5 ms), or any hint of inhibitory contamination (probed by voltage clamping the postsynaptic neuron to –50 mV) were discarded. Also, synaptic responses for each neuron were characterized by both the peak current recorded in voltage clamp, at a holding potential of –70 mV, and the initial EPSP slope recorded in current clamp; these were interleaved. Input resistance, series resistance, and membrane potential were periodically checked; if either resistance changed by >20% or the membrane potential by >6 mV, recordings were discarded. Throughout each recording, the postsynaptic response was tested every 10 s. After a stable baseline (6–8 min) had been established, potentiation or depression was induced by pairing presynaptic stimulation with postsynaptic spiking. The postsynaptic spikes were produced by injecting brief (5 ms) current pulses. To induce potentiation, presynaptic stimulation was given 10 ms before each postsynaptic spike; to induce depression, it was given 20 ms after each postsynaptic spike. In both cases, a total of 75 pairings were used. These were divided into 15 trains of five pairings each. The trains were delivered at 0.1 Hz; the pairings within each train were delivered at 5 Hz.
mEPSC and NMDA/AMPA measurements
Miniature excitatory postsynaptic currents (mEPSCs) were measured in the presence of tetrodotoxin (0.2 µM), 2-amino-5-phosphonopentanoic acid (AP-5; 50 µM), and bicuculline (20 µM). Neurons were voltage clamped to –70 mV, and spontaneous currents were measured for 1–5 min. Miniature currents were detected using in-house software. The detection thresholds for amplitude and 10–90% rise time were >5 pA and <2 ms, respectively.
NMDA/AMPA ratios were measured using a cesium-based internal solution and an ACSF containing 20 µM glycine. EPSCs were evoked by stimulating lower layer IV or upper layer V, while neurons were voltage clamped at –90, +50 mV, and one or more potentials in between. At least five responses were averaged at each holding potential. Responses that showed evidence of inhibitory or polysynaptic contamination were rejected. Also rejected were recordings judged to have inadequate voltage clamp, namely responses that reversed >15 mV from 0 mV. Following Myme et al. (2003), the current measured at –90 mV was assumed to be mediated entirely by AMPA receptors. Its peak was used to establish the 1-ms time window for measuring the AMPA current at +50 mV. The NMDA current at +50 mV was measured over a 1-ms time window centered 45 ms after the stimulus artifact. The ratio of the two measurements at +50 mV was taken as the NMDA/AMPA ratio.
Solutions
The standard ACSF used to prepare slices and in most recordings contained (in mM) 124 NaCl, 3 KCl, 1.25 NaH2PO4, 26 NaHCO3, 2 MgCl2, 2 CaCl2, and 10 dextrose. The osmolality was adjusted to 
310 mosM with dextrose, and the pH buffered to 7.4 by bubbling with 95% O2-5% CO2. The ACSF used in STDP experiments was similar, but the CaCl2 concentration was increased to 2.5 mM, the MgCl2 concentration decreased to 1 mM, and the osmolality adjusted to 320 mosM. The standard internal solution contained (in mM) 110 K-gluconate, 10 KCl, 10 (Na)phosphocreatine, 10 HEPES, 4 (Mg)ATP, 0.3 (Na)GTP, 0.5 EGTA, and 0.1% wt/vol biocytin. The internal solution was adjusted with KOH to pH 7.4 and with sucrose to 290–300 mosM. In STDP experiments, the calcium buffer was omitted. To measure NMDA-AMPA ratios, the internal solution contained (in mM) 115 cesium methanesulfonate, 20 cesium chloride, 5 QX-314, 10 (Na)phosphocreatine, 10 HEPES, 4 (Mg)ATP, 0.3 (Na)GTP, 5 EGTA, and 0.1% wt/vol biocytin.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Intrinsic electrical properties
The neurons used in this study were characterized by a location in layer V, medium-to-large triangular somata, and prominent apical dendrites that extended into the uppermost layers (Fig. 1A). There were no obvious morphological differences between neurons in wild-type and KO slices. (The differences in dendritic spines that have been reported would not have been observable using our microscopy techniques.) Intrinsic electrical properties were probed by injecting a family of DC current steps (–100 to 200 pA, 500 ms) through the patch pipette (Fig. 1B). All neurons could adequately be described as "regular spiking": suprathreshold steps elicited one or more spikes at low-to-moderate firing rates (0–30 Hz), which showed mild spike frequency adaptation. None were intrinsically bursting, consistent with previous reports in rodent neocortex at this age (Franceschetti et al. 1998).
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Most efforts to understand activity-dependent plasticity have been concerned with synaptic plasticity. This makes sense given the enormous body of evidence that demonstrates that synaptic properties are plastic on several different time scales and in multiple ways. Yet a growing number of reports, involving work in several different brain areas, has established a persuasive case that the intrinsic properties of individual neurons, how they integrate synaptic input and produce action potentials, are also plastic and regulated by activity (Daoudal and Debanne 2003; Zhang and Linden 2003). Indeed, some of the very same experimental protocols (tetanic stimulation and burst firing, spike pairing, activity blockade) that have been used to study synaptic plasticity also give rise to intrinsic plasticity.
Our focus here was on LTP-IE, a phenomenon that has been studied in both neocortex and hippocampus. More than one experimental protocol—involving synaptic stimulation, pharmacological stimulation, burst firing, or some combination—has been given this name (Cudmore and Turrigiano 2004; Ireland and Abraham 2002; Sourdet et al. 2003; Xu et al. 2005). Although the protocols are distinct, they are closely related, and it seems likely that the same underlying biophysical processes are at work. In these studies, we adopted the electrical stimulation protocol of Sourdet et al. (2003), which is illustrated in Fig. 2A. Intrinsic excitability was assayed by injecting a fixed test current at the soma and then counting the resulting number of action potentials. When mGluR receptors were activated for a short period (4 min), by electrically stimulating synaptic afferents after ionotropic transmission had been blocked, intrinsic excitability increased and remained elevated for 40 min (Fig. 2B). (Sourdet et al. 2003) showed that LTP-IE in somatosensory-motor cortex depends specifically on activation of synaptic mGluR5 receptors and is mediated by a reduction in medium afterhyperpolarization currents.
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). Postsynaptic firing alone, without concurrent synaptic stimulation, resulted in no net change in excitability (103 ± 3%, n = 6). Consistent with the report in rat neocortex, we found in our mouse slices that a similar increase in intrinsic excitability could be produced by a direct application of the mGluR agonist ACPD (50 µM, 3–4 min, data not shown). LTP-IE is normal in KO animals
LTP-IE in neurons from Fmr1 KO mice was indistinguishable from that in neurons from wild-type littermates. An increase in excitability was apparent immediately on termination of mGluR stimulation, and could be as large as 200%. Excitability remained elevated and stable for as much as 40 min after LTP-IE induction, which is as long as we measured it (Fig. 3A). Responses were quite variable with some recordings showing no change in numbers of spikes elicited by the test current and others showing a doubling in numbers, but the variability was the same in KO and wild-type recordings (Fig. 3B). Likewise, the average amount of potentiation was no different between the two conditions. (Fig. 3B, inset). This was true regardless of the time point at which potentiation was measured. In Fig. 3B, we averaged data at a late time point (25 min after induction) to emphasize the long-lasting aspect of the potentiation, but differences were not statistically significant at any time point. Neither resting potential nor resting input resistance was altered after LTP-IE induction.
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Synaptic transmission
Previous studies using older animals have found that baseline synaptic transmission is normal (Li et al. 2002; Zhao et al. 2005). However, in principle, this might not be true at earlier developmental time points, which are characterized by dramatic synaptic change. We used two measures to check synaptic transmission: AMPA-mediated mEPSCs and NMDA/AMPA ratios.
The measurement of mEPSCs offers a convenient way of characterizing, in a global way, the synaptic inputs onto a given neuron, as they represent the postsynaptic response to the spontaneous, random release of vesicles by all of the neuron's presynaptic partners. We recorded mEPSCs at a holding potential of –70 mV in the presence of TTX to prevent spiking and pharmacological blockers of NMDA and GABAA currents. An example recording is show in Fig. 4A (top) along with the average mEPSC recorded in that neuron. As expected, there was a large variability in mEPSCs measurements, with some currents as small as 5 pA (the detection cutoff) and others as large as 50 pA. Even so, wild-type and KO recordings were remarkably similar. There were no significant differences in average mEPSC amplitude or charge (wild-type, 9.6 ± 0.5 pA, 85 ± 5 fC, n = 9; KO, 10.9 ± 0.7 pA, 93 ± 3 fC, n = 13). Nor were there differences in the distributions of these quantities (Fig. 4A, bottom). Other parameters which showed no difference between conditions included: 10–90% rise time, decay time constant (estimated by fitting a single exponential), and mEPSC frequency (wild-type, 1.14 ± 0.07 ms, 6.18 ± 0.63 ms, 7.2 ± 1.9 Hz, n = 9; KO, 1.15 ± 0.03 ms, 6.09 ± 0.32 ms, 5.4 ± 0.7 Hz, n = 13).
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To estimate NMDA currents, we measured the NMDA/AMPA ratio of EPSCs evoked by electrical stimulation of proximal fibers (Myme et al. 2003). No bicuculline or picrotoxin was included in the recording solution because, with extracellular stimulation, these tended to produce epileptic behavior. Instead we simply discarded recordings that showed evidence of inhibitory contamination (i.e., any component that reversed near the chloride reversal potential). We similarly discarded recordings we suspected of containing polysynaptic components, using several criteria: multiple peaks, large trial-to-trial jitter, or long decaying phases. NMDA/AMPA ratios were measured in the manner of (Myme et al. 2003). As illustrated in Fig. 4B (left), the EPSC measured at a holding potential of –90 mV was assumed to be mediated entirely by an AMPA current; the time at which it peaked was used to measure the AMPA amplitude from the EPSC measured at a holding potential of +50 mV. The NMDA amplitude was measured from the +50 mV EPSC at a time point 45 ms after the stimulus was given; the assumption here was that, by this time, the AMPA current had decayed away entirely and only an NMDA current was left. The ratio of the two numbers was taken to be the NMDA/AMPA ratio.
Using this definition, we obtained values for the NMDA/AMPA ratio at the low end of those reported previously (Myme et al. 2003); our individual values ranged from 0.4 to 0.8, as opposed to an average in the literature of close to 1. This was probably a consequence of two things: our definition of the ratio (specifically, how late in the decaying phase we measured the NMDA component) and the difficulty of clamping layer V pyramidal neurons at very depolarized potentials. However, what was clear from our data were that the average ratio was the same whether recorded in neurons from KO animals or those from wild-type animals (Fig. 4B, right).
STDP in mouse neocortex
We examined how Fmr1 deletion affects synaptic plasticity in early development by studying STDP, which has emerged in recent years as a leading model for activity-dependent long-term plasticity (Dan and Poo 2004). In the STDP protocol, the temporal order of pre- and postsynaptic spikes determines whether a synapse is potentiated or depressed (Fig. 5A). Under most conditions, when a presynaptic spike is produced (or, equivalently, a presynaptic stimulation given) before a postsynaptic spike, the synapse is strengthened; when the presynaptic spike or stimulus comes after the postsynaptic spike, the synapse is depressed. The time windows for potentiation and depression at cortical synapses are normally on the order of 10–50 ms wide.
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in layer V rat visual cortex. Under this protocol, postsynaptic spiking (produced by a brief current injection) was paired with stimulation of proximal afferents. The stimulation strength in all cases was set so as to elicit initial EPSP amplitudes of 3–5 mV. As when measuring NMDA/AMPA ratios, recordings that showed evidence of inhibitory or polysynaptic contamination were not used. We found that when pairings where delivered in 5-Hz trains, the protocol worked robustly and without modification in somatosensory slices from wild-type mice. Synapses remained potentiated or depressed for 40 min after induction, which is as long as our recordings lasted (Fig. 5, B and C). STDP: potentiation is impaired, but depression is robust
In comparing STDP in wild-type and KO mice, two timing intervals were used consistently. To produce synaptic long-term potentiation (LTP), presynaptic stimulation was delivered 10 ms before a postsynaptic spike was elicited (
t = +10 ms, Fig. 5B). To produce synaptic long-term depression (LTD), the stimulation was delivered 20 ms after the postsynaptic spike (t = –20 ms, Fig. 5C). (To distinguish these timing-dependent forms of LTP and LTD from rate- and depolarization-based forms, we will refer to them as tLTP and tLTD, respectively.)
A difference in plasticity between wild-type and KO animals was starkly apparent when we tried to potentiate synaptic currents. Whereas potentiation in the wild-type case was strong (albeit variable), it was entirely absent in the KO case (Fig. 6A). In tissue from KO mice, we attempted to elicit tLTP in nine slices, which were distributed among five animals. In no case did synaptic amplitude increase by >20%. In only three cases did it increase by >15%. In fact, an equal number of attempts resulted in slight decreases in synaptic amplitude. On average, running the STDP protocol with a positive timing interval (t = +10 ms) produced a net synaptic change in KO slices of only 5 ± 4%, which is not distinguishable from zero. On the other hand, the potentiation protocol worked as well in wild-type mouse slices as has previously been reported in rat neocortical slices, with 9 of 14 recordings showing 40% potentiation (Feldman 2000; Froemke and Dan 2002; Sjöström et al. 2001). The time course of potentiation in wild-type recordings was also similar to that in previous reports: synaptic currents were found to be potentiated immediately at the end of the pairing protocol and remained elevated for >40 min afterward. In KO recordings, synaptic currents were elevated to a degree immediately after pairing, but reverted to baseline within 
6 min (Fig. 6A).
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; Sjöström et al. 2001). Again there was significant variability, as expected given results in rat slices (Sjöström et al. 2001). However, there were no significant differences between the two conditions whether measured at a late time point (as in Fig. 6B, right) or by averaging over the whole postinduction period (P = 0.15). The time course of the effect was similar between conditions (Fig. 6B, left) as was the distribution of individual data points (Fig. 6B, right). mGluR5 dependence of STDP
Much current interest in FXS stems from the hypothesis that FMRP acts as a regulator of group 1 mGluR-activated translational processes (Bear et al. 2004). There have been several reports of a link between mGluR activation and neocortical LTP and LTD (Eckert and Racine 2004; Huemmeke et al. 2002; Morris et al. 1999; Sawtell et al. 1999), but these have varied in their conclusions, possibly reflecting differences in brain area or synapse type. Moreover, the method by which plasticity is induced can affect the results (Huemmeke et al. 2002), presumably because different induction protocols trigger different biochemical processes at different rates.
We therefore examined whether STDP in layer 5 pyramidal neurons depends on activation of mGluR5 receptors by using the selective antagonist 2-methyl-6-(phenylethynyl)-pyridine (MPEP). mGluR5 is the dominant group 1 mGluR in the neocortex, particularly in pyramidal neurons (Blue et al. 1997; Lopez-Bendito et al. 2002). All of these experiments were done in slices from wild-type mice. MPEP at 10 µM was added to the bath solution 5–10 min before each recording began and was present throughout the recording. Using a timing interval of t = +10 ms, we attempted to induce potentiation in the presence of MPEP, and were successful (Fig. 7, A and C). The average amount of potentiation obtained was comparable to the amount previously obtained in control ACSF (32 ± 10 vs. 39 ± 7%, P = 0.59). On the other hand, when we used a timing interval of t = –20 ms to induce depression, we were unsuccessful (Fig. 7, A and C). Of seven attempts, none produced depression of more than –20%. The average amount of depression was significantly reduced from the amount obtained in control ACSF (–6 ± 3 vs. –24 ± 5%, P < 0.01). On the basis of these data, we conclude that tLTP does not depend on mGluR5 activation but tLTD does.
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30 min before tLTD induction and throughout each recording. As shown in Fig. 7, B and C, we found that depression could be robustly induced even when protein synthesis was blocked. The average amount of depression in the presence of anisomycin was not significantly different from the amount in control ACSF (–26 ± 8 vs. –24 ± 5%, P = 0.8). tLTD is therefore mGluR5 dependent but not translation dependent. |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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STDP as a model of developmental synaptic plasticity
Many forms of plasticity—e.g., synaptic, intrinsic, Hebbian, homeostatic—are active during early postnatal development and are (presumably) important for the correct formation and sculpting of neocortical circuits (Cohen-Cory 2002). We made a specific choice of experimental model to use in investigating synaptic plasticity, namely STDP. This was a natural choice because STDP has emerged in recent years as (arguably) the leading model of sensory developmental plasticity (Feldman and Brecht 2005; Fox and Wong 2005). The principal reason is that it involves spike trains that are a closer match to trains recorded in vivo than those used in most rate-based protocols (Carandini and Ferster 2000; Celikel et al. 2004). Also STDP has other attractive features: it allows an explicit role for spike timing, indirectly helps to implement the Hebbian idea of causality, and is buttressed by theoretical work suggesting its temporal asymmetry leads to the creation of stable networks (Abbott and Nelson 2001).
Our finding that tLTP is impaired in KO animals is probably not exclusive to our choice of experimental model. Previous studies, which used rate-based or depolarization-based induction protocols, also found that neocortical LTP was impaired or absent (Larson et al. 2005; Li et al. 2002; Zhao et al. 2005). This was true even though these studies differed from ours in several other ways: older animals, different synaptic pathways, and different parts of neocortex. Together, all of this work suggests that impaired neocortical LTP is a quite general feature of FXS. It should be noted, though, that in olfactory cortex, abnormalities in LTP were observed in animals older than 6 mo but not in those between 3 and 6 mo (Larson et al. 2005). Given the reported time course of spine abnormalities in somatosensory cortex (Galvez and Greenough 2005; Nimchinsky et al. 2001), it will be important to investigate whether abnormal LTP has a similar age dependence.
Another way the present study differs from previous reports on neocortical synaptic plasticity in Fmr1 KO mice is that we also examined tLTD. To our knowledge, ours is the first examination of neocortical LTD in this mouse model. That the effect of Fmr1 deletion on depression was different from on potentiation is interesting and perhaps reflects the fact that these timing-dependent forms of plasticity are mediated by different biophysical mechanisms. Using STDP protocols on layer V pyramidal neurons, Sjöström and colleagues demonstrated that tLTD is expressed presynaptically and is independent of postsynaptic NMDA receptors but does depend on retrograde endocannabinoid signaling and presynaptic NMDA receptors (Sjöström et al. 2003). By contrast, tLTP may have both pre- and postsynaptic components and depends on both postsynaptic calcium influx and postsynaptic NMDA receptors. These findings have recently received additional support from an STDP study using layer II/III pyramidal neurons (Bender et al. 2006). That study also concluded that tLTD depends on endocannabinoid signaling, presynaptic or other nonpostsynaptic NMDA receptors, and changes in vesicle release probability, whereas tLTP does depend on postsynaptic NMDA receptors. That tLTP, but not tLTD, is abnormal in the Fmr1 KO points toward a postsynaptic location for the defect in plasticity, consistent with FMRP's localization at postsynaptic sites (Antar et al. 2004; Bagni and Greenough 2005) and spine shape abnormalities.
Although our data do not indicate a significant change in tLTD, it is important to consider the possibility that the balance between potentiation and depression has been shifted in KO animals, away from the former and toward the latter (Abraham et al. 2001; Savi
et al. 2003) This is a possibility even if tLTP and tLTD are not expressed in the same way (e.g., presynaptic versus postsynaptic). For example, Sjöström et al. (2001) showed that when pairing between pre- and postsynaptic spikes is done at high frequency (>40 Hz), tLTD never results, even when a negative timing interval is used. They argued that the fact that at high frequency, a presynaptic spike will always be preceded by one postsynaptic spike and followed by another within a narrow time window (<25 ms) means that tLTP and tLTD are simultaneously induced and interact in a nonlinear fashion. To cite another example, Bender et al. (2006) found that blocking tLTP by selectively blocking postsynaptic NMDA receptors uncovered a tLTD that had previously been occluded by the tLTP. (Although, it should be noted that our data indicate that in Fmr1 KO animals tLTP is absent and not replaced by tLTD.) As these examples demonstrate, the interaction between tLTP and tLTD can be quite complicated and depends on the spike trains used to elicit plasticity (Dan and Poo 2004). The absence of tLTP in Fmr1 KO slices might open up more opportunities for tLTD to be expressed, an idea that might be explored in future, for example, by pairing with different spike trains.
mGluR theory
An obvious question is whether and how these findings relate to the mGluR theory of the biochemical basis of FXS (Bear et al. 2004). Much current work on Fragile X has been inspired by the suggestion that loss of FMRP alters synaptic processes triggered by group 1 mGluRs. Termed the "mGluR theory," this idea predicts that in the absence of FMRP, exaggerated mGluR-induced mRNA translation leads to synaptic changes that are the proximal causes of symptoms in FXS (Bear 2005). Since its publication 2 years ago, it has accommodated disparate data within a simple theoretical framework and offered the hope of devising rational therapeutic strategies (Antar et al. 2004; Aschrafi et al. 2005; Chuang et al. 2005; Koekkoek et al. 2005; McBride et al. 2005).
We chose to examine LTP-IE, in part, because it depends on activation of mGluR5 receptors, which are the principal group 1 mGluR type in neocortex. We found no exaggeration or abnormality in LTP-IE in Fmr1 KO mice. Does this conflict with the mGluR hypothesis? The answer is no. Evidence to date suggests that only those mGluR-induced events requiring mRNA translation are altered in FXS in accord with the notion that FMRP functions as a translational suppressor in dendrites (Bear et al. 2004). In recordings from rat somatosensory cortex, we found that we were able to elicit LTP-IE even in the presence of 25 µM anisomycin, a protein synthesis inhibitor (our unpublished observations). Given that LTP-IE does not require translation, exaggerated or otherwise abnormal LTP-IE is not a requirement of the mGluR theory in its strong form.
What the mGluR theory would predict generally for neocortical synaptic plasticity is not clear. There have been numerous reports that LTP and LTD, normally elicited by high- or low-frequency stimulation, depend on mGluR activation and/or translation (Anwyl 1999; Cohen et al. 1998; Harney et al. 2006; Holscher 2002; Miura et al. 2002; Morris et al. 1999; Raymond et al. 2000; Thompson et al. 2005). Some of these reports have not been easy to reconcile with each other and most have focused on plasticity in the hippocampal formation. Hippocampal and neocortical plasticity are closely related, of course, but there are good reasons to think they are different phenomena. In particular, whereas impaired neocortical LTP has previously been reported in mature Fmr1 KO mice (Larson et al. 2005; Li et al. 2002; Zhao et al. 2005), hippocampal LTP has been reported to be normal (Godfraind et al. 1996; Larson et al. 2005; Li et al. 2002; Paradee et al. 1999).
Using the mGluR5 antagonist MPEP, we found that, in layer V, the potentiation part of STDP does not depend on mGluR5 activation but the depression part does. A similar observation regarding tLTD was made very recently in layer II/III (Bender et al. 2006). [Note that this recent result differs from one previously obtained in layer II/III using low-frequency stimulation (Sawtell et al. 1999), illustrating the fact that timing- and frequency-dependent plasticity are not fully equivalent.] tLTD in neocortex involves postsynaptic endocannabinoid release and subsequent retrograde activation of presynaptic CB1 receptors (Bender et al. 2006; Chevalyre et al. 2006; Duguid and Sjöström 2006; Sjöström et al. 2003). Group 1 mGluRs are known to stimulate endocannabinoid synthesis, a process that involves activation of phospholipase C and IP3-mediated release of calcium from internal stores (Chevalyre et al. 2006). It follows that MPEP's effect on tLTD likely results from a reduction in activity-evoked endocannabinoid synthesis. This would not involve de novo protein synthesis, consistent with our finding that anisomycin does not impair tLTD.
A simple relationship between STDP and the mGluR theory can therefore be ruled out, as neither potentiation nor depression depends directly on mGluR-dependent translational processes for their induction—unlike, for example, mGluR-dependent LTD in the hippocampus (Huber et al. 2000). However, the mGluR idea cannot be excluded entirely because mGluR-dependent translation might still affect STDP indirectly by affecting the basal state on which it acts. Plasticity at excitatory synapses is thought to depend critically on the shapes of dendritic spines, and these in turn can be modified by mGluR-dependent translation (Vanderklish and Edelman 2002). Also, ongoing mGluR-dependent translation might create proteins that are useful for plasticity and do not need to be synthesized during the plasticity protocol itself (Nosyreva and Huber 2006).
Plasticity and development
A major unanswered question is how does the absence of FMRP impair LTP formation? Given its role in translational regulation, loss of FMRP might be expected to have an effect on the process of consolidation, which requires local mRNA translation (Vanderklish and Edelman 2005). The abnormal spine shapes associated with FXS also suggest this may be the case. A convergence of studies on the relationship among spine shape, the number of AMPA receptors in the postsynaptic density, and mGluR-induced translation suggests that mGluR-induced translation responses can produce an elongated and thinner spine shape that is less efficacious because it has less space for AMPA receptors (Huber et al. 2000; Matsuzaki et al. 2001, 2004; Takumi et al. 1999; Vanderklish et al. 2002). This process may underlie the consolidation of forms of LTD that are expressed postsynaptically (Vanderklish and Edelman 2005). Moreover, it has been shown that mGluR-induced translation primes LTP consolidation (Raymond et al. 2000), potentially through the synthesis of proteins involved in spine remodeling, such as PSD95. Overexpression of PSD95 has been shown to enhance synaptic AMPA receptor levels, enlarge dendritic spines, and result in potentiation that occludes electrically induced LTP (Ehrlich and Malinow 2004; Stein et al. 2003). A recent report established that in the Fmr1 KO, PSD95 synthesis is reduced (Todd et al. 2003). Although it is tempting to speculate that loss of FMRP reduces LTP through translation-dependent changes in spine morphology that affect AMPA receptor content, it remains possible that perturbations in spine shape alter plasticity through a number of other mechanisms (Bloodgood and Sabatini 2005; Segal 2005; Tsay and Yuste 2004). Our results would not exclude any of these possibilities. Although mEPSC measurements might in principle be used as a constraint, in practice, this is very difficult to do; other models that result in altered synapse shape (e.g., PSD95 or Rho overexpression, NRBI KO) have produced inconsistent effects on mEPSC amplitudes and distributions (Segal 2005).
An even harder question is how impaired LTP affects the development of sensory neocortex. If the defect is localized to the developmental time point we investigated, the mature circuit may not exhibit gross abnormalities. Some basic features of sensory systems seem to be in place even before birth and well before traditional "critical periods" (Katz and Crowley 2002). Moreover, the existence of multiple forms of plasticity—some of which, as we have demonstrated, are unimpaired by the absence of FMRP—suggests that compensatory mechanisms might play a role. In this regard, it will be important in future studies to investigate LTP in the Fmr1 KO mouse using a single protocol in a single brain area throughout developmental time.
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Address for reprint requests and other correspondence: N. S. Desai, The Neurosciences Fine Institute, 10640 John J. Hopkins Dr., San Diego, CA 92121 (E-mail: desai{at}nsi.edu)
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