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The Journal of Physiology logoLink to The Journal of Physiology
. 2010 Sep 20;588(Pt 22):4347–4363. doi: 10.1113/jphysiol.2010.192864

Comparison of synaptic transmission and plasticity between sensory and cortical synapses on relay neurons in the ventrobasal nucleus of the rat thalamus

Ching-Lung Hsu 1, Hsiu-Wen Yang 3,4, Cheng-Tung Yen 1, Ming-Yuan Min 1,2
PMCID: PMC3008844  PMID: 20855435

Abstract

Relay neurons in the ventrobasal nucleus of the thalamus transmit somatosensory information to the cerebral cortex and receive sensory and cortical (feedback) synaptic inputs via, respectively, medial lemniscal (ML) and corticothalamic (CT) fibres. Here, we report that calcium-permeable AMPA receptors are expressed at CT synapses, but not ML synapses, and that the NMDA receptor (NMDAR)-mediated/non-NMDAR-mediated synaptic current ratio is significantly larger at CT synapses than at ML synapses. Moreover, NMDAR-dependent LTP and L-type voltage-gated calcium channel-dependent LTD are readily induced at CT synapses, but not ML synapses. In particular, LTD of CT synaptic transmission is induced by spiking of postsynaptic relay neurons in continuous mode, but not burst mode, in current-clamp recordings. These results show that the strength of the cortical input to thalamic relay neurons is selectively subjected to use-dependent modification, which could be a mechanism for regulation of thalamocortical–corticothalamic interactions and the underlying sensory processing.

Introduction

The ventrobasal nucleus (VBN) of the thalamus serves as a major gateway for relaying somatosensory information to the cerebral cortex (Jones, 2007). Thalamocortical (TC) relay neurons fire action potentials either in continuous or burst mode, depending on their membrane potential (Vm). During depolarization, T-type voltage-gated calcium channels (T-VGCCs) are inactivated and TC relay neurons fire continuously, while during hyperpolarization, T-VGCCs are activated and TC relay neurons fire in burst mode (Jahnsen & Llinas, 1984; Suzuki & Rogawski, 1989; Bal & McCormick, 1993; Sherman, 2001). Continuous firing is commonly recorded when animals are awake and alert and the sensory information is usually faithfully relayed, whereas burst firing is recorded when the animals are asleep and sensory information is not relayed (McCormick & Feeser, 1990; Le Masson et al. 2002; Sherman, 2005).

Regulation of firing mode switching in TC relay neurons and thus of the transfer of sensory information has been intensively studied, the focus being on the regulation of intrinsic membrane properties that govern the activation of T-VGCCs and other ion channels involved in firing mode switching. However, a significant role for synaptic drive in firing mode switching has been suggested (Ergenzinger et al. 1998; Castro-Alamancos & Calcagnotto, 2001; Hughes et al. 2002; Sillito & Jones, 2002). TC relay neurons in the VBN receive synaptic inputs from somatosensory afferents through the medial lemniscal (ML) tract and strong cortical feedback through corticothalamic (CT) tracts (Liu et al. 1995; Sherman & Guillery, 2002). It has been estimated that in the lateral geniculate nucleus sensory inputs account for only 10% of synapses on TC relay neurons, whereas cortical inputs account for more than 50% (Erisir et al. 1997; Latawiec et al. 2000). Of the cortical inputs, 50% directly synapse on TC relay neurons, while the other 50% exert their influence through GABAergic interneurons located in the reticular thalamic nucleus (Liu et al. 1995; Sherman & Guillery, 2002). Because of the predominant number of CT synaptic contacts on relay neurons, cortical feedback signals are generally regarded as an essential modulator of the properties of thalamic spike transfer (Murphy & Sillito, 1987; Ergenzinger et al. 1998; Krupa et al. 1999; Sillito & Jones, 2002; Sherman, 2005).

Despite the presence of massive CT feedback on thalamic relay neurons, the function of this system remains unclear. Since there are intense interactions between the cerebral cortex and TC relay neurons and since the extent of this interaction is important for sensory processing in the brain, use-dependent changes in the synaptic efficacy may provide a means by which the extent of TC–CT interactions and the underlying sensory processing are regulated. In this study, we systematically studied the synaptic plasticity properties of CT synapses and compared them to those of sensory inputs to thalamic relay neurons, namely the synapses between ML fibres and TC relay neurons.

Methods

Preparation of thalamic slices

The use of animals in this study was in accordance with the guidelines of the Ethics Committee for Animal Research of the National Taiwan University. Sprague–Dawley rats of both sexes aged 13–24 days were used. The rats were anaesthetized with isoflurane and decapitated with a small-animal guillotine and their brains rapidly removed and placed in ice-cold artificial cerebrospinal fluid (ACSF). The ACSF contained (in mm): 119 NaCl, 2.5 KCl, 1.3 MgSO4, 26.2 NaHCO3, 1 NaH2PO4, 2.5 CaCl2, and 11 glucose; the pH was adjusted to 7.4 by gassing with 95% O2–5% CO2. Initially, coronal slices were cut and used. However, we then discovered that it was much easier to simultaneously recruit excitatory postsynaptic currents (EPSCs) of ML and CT synapses in a TC relay neuron in horizontal slices, and the horizontal preparation was adopted for most parts of this study. To prepare horizontal slices, the two halves of the brain were separated and the half-brain glued to the stage of the vibroslicer via its ventral surface. A block of agar was glued behind the medial surface of the half-brain to provide support during slicing. Slices with a thickness of 300 μm containing the VBN, internal capsule (IC), and ML tract (see Fig. 1A) were cut using a vibroslicer (Zero 1, DSK, Osaka, Japan). To allow recovery, the slices were kept in oxygenated ACSF (95% O2–5% CO2) at room temperature (24–25°C) for at least 90 min before recording was started.

Figure 1. Recording of CT EPSCs and ML EPSCs in VBN relay neurons.

Figure 1

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A, photograph of a horizontal forebrain slice used in this study. The slice was fixed with 4% paraformaldehyde. Two schematic drawings show the positions of the bipolar stimulating electrode for eliciting CT EPSCs and ML EPSCs. IC, internal capsule; ML, medial lemniscal tract; VBN, ventrobasal nucleus; L, lateral; R, rostral; scale bar: 500 μm. B, a biocytin-filled relay neuron located in the VBN. The arrows indicate the axon. Scale bar: 25 μm. C, voltage responses of a VBN relay neuron following injection of a depolarizing (upper traces) or hyperpolarizing (lower traces) current. Ca shows the recording with the membrane potential (Vm) held at ∼ −55 mV. Note the continuous firing mode (upper trace) and the rebound burst-firing (indicated by the arrow) evoked by termination of the current injection (lower trace). Cb shows recordings with Vm held at ∼ −70 mV. Note the burst firing mode indicated by the arrows. D and E, stimulus–response relationship of the CT EPSC (D) and ML EPSC (E). The upper traces show superimposed synaptic responses upon increasing stimulation of the IC (D) or ML (E). The lower panels show the summarized results from 6 neurons for the CT EPSCs and 5 neurons for the ML EPSCs. Note the continuous increase in synaptic responses and the all-or-none responses upon increasing stimulation of the IC or ML, respectively. F and G, synaptic responses to paired-pulse stimulation (IPI = 50 ms) of CT (F) or ML (G) inputs. Note the paired-pulse facilitation of CT EPSCs and the paired-pulse depression of ML EPSCs. The responses shown in F and G were evoked from the same relay neuron.

Electrophysiology

Slices were transferred to an immersion-type recording chamber mounted on an upright microscope (BX50WI, Olympus Optical, Tokyo, Japan), equipped with an infrared differential interference contrast microscopic video. The VBN, IC, and ML tract were clearly identified under low magnification in the horizontal slices (see Fig. 1A). Neurons in the VBN were recorded under visual guidance with patch pipettes pulled from borosilicate glass (1.5 mm outer diameter, 0.32 mm wall thickness; G150F-4, Warner Instruments, Hamden, CT, USA). The patch electrodes had a resistance of 3–8 MΩ when filled with a solution consisting of (in mm): 131 potassium gluconate, 20 KCl, 10 Hepes, 2 EGTA, 8 NaCl, 2 ATP, 0.3 GTP, and 6.7 biocytin; the pH was adjusted to 7.2 by KOH and the osmolarity to 300–305 mosmol l−1. For the experiments shown in Figs 1 and 2, potassium gluconate was replaced with caesium gluconate (114.7 mm) and KCl with CsCl (17.5 mm) and the pH was adjusted with CsOH. In the experiments in which repeated depolarizing pulse commands were needed for induction of long-term depression (LTD), 5 mm QX-314 was included to internally block voltage-gated sodium channels. Recordings were made with an Axopatch 1D amplifier (Molecular Devices, Sunnyvale, CA, USA) at room temperature. For voltage-clamp recordings, the whole-cell conductance (GW) and series resistance (Rs) were continuously monitored by applying a voltage pulse of 3 mV; Rs was usually <20 MΩ and was not compensated for. For current-clamp recordings, the input resistance (Rn) was continuously monitored by applying a current pulse of 30 pA and the bridge was balanced by adjusting the Rs compensation of the amplifier. Data were discarded when Rs or Rn varied by more than 20% from its original value during the recording. All signals were low-pass-filtered at the corner frequency of 2 kHz, then digitized at 10 kHz using a Micro 1401 interface (Cambridge Electronic Design, Cambridge, UK). Data were collected using Signal software (Cambridge Electronic Design).

Figure 2. Differences in the ionotropic glutamate receptor content at CT and ML synapses.

Figure 2

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A, both CT EPSCs and ML EPSCs (black traces) were blocked by application of 10 μm DNQX (grey traces). All traces shown are averaged from 10 sweeps. B, I–V relationship of ML EPSCs (Ba) and CT EPSCs (Bb) evoked in relay neurons. The dotted lines are linear fits to the currents recorded with a negative Vm. C, application of 10 μm PhTx-433, a selective CaP-AMPAR blocker, significantly attenuates CT EPSCs (Cb), but not ML EPSCs (Ca), recorded in the same relay neuron. The insets are recordings before application of the drug (control), during application of the drug (PhTx), and during wash out (wash); each trace was averaged from 6 sweeps. D, summarized results showing a significant difference in rectification index (RI) (left), blockade by PhTx-433 (middle), and blockade by NAS (right) between CT EPSCs (grey bars) and ML EPSCs (black bars). E, difference in the peak current ratio of NMDAR- to non-NMDAR-mediated components of CT EPSCs and ML EPSCs. The ML EPSC (left) and CT EPSC (right) shown were evoked from the same relay neuron. F, summarized results showing the significant difference in the peak current ratio of NMDAR- to non-NMDAR-mediated components between ML (black bar) and CT (grey bar) synapses. The data include those from one experiment in which the CT EPSCs and ML EPSCs were evoked in the same relay neuron.

Stimulation of CT and ML synapses

After transfer to a recording chamber, the slices were continuously perfused with ACSF containing 0.1 mm picrotoxin and 1 μm strychnine to isolate EPSCs or excitatory postsynaptic potential (EPSP) activity. A bipolar stainless steel electrode (FHC, St Bowdoin, ME, USA) was placed locally in the VBN or the IC for CT EPSC recruitment; no significant differences were observed between the CT EPSCs evoked by these two conditions (see Supplemental Table 1). Another stimulating electrode was placed in the ML tract to evoke ML EPSCs. We intentionally tried to simultaneously recruit CT EPSCs and ML EPSCs in the same TC relay neuron in each series of experiments, except for those shown in Figs 3, 4, and 6–8. In some cases, only CT EPSCs or ML EPSCs were evoked; these data were also pooled for statistical comparison. EPSCs were evoked at 0.067–0.125 Hz; in experiments in which CT EPSCs and ML EPSCs were simultaneously recruited, alternate stimulations were made to the IC (or locally) and ML tracts, the stimulation rate to both inputs being 0.067–0.125 Hz. Unless specified otherwise, Vm values of the recorded TC relay neurons were clamped (or held) at −70 mV. The estimated liquid junction potential was ∼10 mV and was left uncorrected. For the recording of EPSP activity in current-clamp mode, data were accepted if the EPSP activity had smooth rising and exponentially decaying phases. If EPSPs showed any signs of contamination, possibly contributed by T-VGCC- and/or N-methyl-d-aspartate receptor (NMDAR)-mediated activity (Landisman & Connors, 2007) and characterized by the appearance of a slow component on top of the fast EPSPs, the data were discarded.

Figure 3. NMDAR-dependent LTP is induced at CT synapses, but not at ML synapses, on relay neurons.

Figure 3

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A, time courses of the evoking of ML EPSCs (Aa) and CT EPSCs (Ab). The horizontal bar indicates the pairing of postsynaptic depolarization to 0 mV and presynaptic stimulation at 1 Hz for 90 s. Note that LTP was induced at CT synapses, but not at ML synapses, after pairing. The insets are averaged responses over 10 sweeps of recordings at baseline (black) and 23 min after pairing (grey). B, summarized results from 7 experiments for ML EPSCs (open circles) and 20 experiments for CT EPSCs (filled circles). The pairing protocol is shown in the inset. The upper panel shows the time course of synaptic potentiation, while the lower panel shows GW. C–E, LTP of CT EPSCs is blocked when APV is added to the bath to block NMDARs (C) or is not induced when postsynaptic depolarization alone (D) or presynaptic stimulation alone (E) is applied. The insets are averaged responses over 10 sweeps of recordings at baseline (black) and 23 min after pairing (grey) in a typical experiment. The lower panels in C–E show the time course of GW during recording; no significant variation in GW was found in this series of experiments.

Figure 4. Conjunctive pairing of depolarizing voltage pulses with presynaptic stimulation does not induce LTP of CT EPSCs.

Figure 4

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A, representative experiment showing no LTP of CT EPSCs after conjunctive paring of postsynaptic depolarizing pulses with presynaptic stimulation (indicated by the horizontal bar). The insets are the CT EPSC recorded before (black) and after (grey) application of paired pre- and postsynaptic simulations, averaged over 10 sweeps of recordings. B, summarized results from 11 experiments showing no LTP after application of paired pre- and postsynaptic simulations, the protocol of which is shown in the inset. The 1 Hz conjunctive pairing stimulation was repeated for 90 cycles. The lower panel shows the time course of GW during recording; no significant variation in GW was found.

Figure 6. LTD of CT EPSCs is L-VGCC-dependent.

Figure 6

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A and B, summarized results showing blockade of LTD of CT EPSCs upon intracellular loading of the postsynaptic relay neurons with BAPTA (A) or bath-application of nimodipine (B), showing the dependence of LTD induction on L-VGCCs. C, application of depolarizing pulses stepping from −50 to 0 mV induces LTD of CT EPSCs. D, LTP of CT EPSCs is induced by conjunctive pairing of postsynaptic depolarizing pulses with presynaptic stimulation after blockade of L-VGCCs. In the experiments in A–D, application of depolarizing pulses or paired pre-and postsynaptic stimulations is indicated by the horizontal bars, and their respective protocols are shown in the upper insets. 90 pulses were applied for LTD induction in the experiments shown in A–C, and 90 paired cycles were applied for LTP induction in the experiment shown in D. The insets to the right of each plot are EPSC activity recorded before (black) and after (grey) application of the LTP/LTD induction protocols, averaged over 10 sweeps of recordings. The lower panels in A–D show the time course of GW during recording; no significant variation in GW was found in this series of experiments.

Figure 8. L-VGCC-dependent LTD of CT synaptic transmission is induced in current-clamp mode in a spiking pattern-dependent manner.

Figure 8

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A, representative experiment showing induction of LTD of CT EPSPs in a relay neuron. After a period of baseline recording, LTD of CT EPSPs was induced by continuous spiking of the relay neuron elicited by injection of 90 cycles of depolarizing current pulses with Vm held at ∼ −55 mV (indicated by the horizontal bar). The protocol of current injection (500 pA given at 1 Hz with a 50% duty-cycle) and the resulting spiking pattern of the postsynaptic relay neuron are shown in the lower traces. B, representative experiment showing that no LTD of CT EPSPs is induced by burst spiking of the relay neuron elicited by injection of 90 cycles of short current pulses with Vm held at ∼ −70 mV (indicated by the horizontal bar). The protocol of current injection (2 nA given at 1 Hz with a 0.2% duty-cycle) and the resulting spiking patterns of the postsynaptic relay neuron are shown in the lower traces. The insets in A and B are EPSP activities recorded before (black) and after (grey) application of current pulses, averaged over 10 sweeps of recordings. C, summarized results from 9 experiments performed as in A (black circles). Note that significant LTD of CT EPSPs was observed. The grey squares show the summarized results from 10 experiments in which nimodipine was applied to the bath. Note the blockade of LTD of CT EPSPs, showing the dependence of LTD induction on L-VGCC activation. D, summarized results from 8 experiments performed as in B. Note that no LTD of CT EPSPs is observed. The lower panel in C and D shows the time course of Rn during recording; no significant variation in Rn was found in this series of experiments.

Drugs

The chemicals used for the ACSF and internal solution were purchased from Merck (Darmstadt, Germany). QX-314 bromide was purchased from Alomone Laboratories (Jerusalem, Israel). dl-2-Amino-5-phosphonopentanoic acid (APV), 6,7-dinitroquinoxaline-2,3-dione (DNQX), nimodipine, and spermine tetrahydrochloride were purchased from Tocris Cookson (Bristol, UK). MgATP, NaGTP, BAPTA, picrotoxin, strychnine, philanthotoxin-433 (PhTx-433), and 1-naphthylacetyl spermine (NAS) were purchased from Sigma (St Louis, MO, USA).

Data analysis and statistics

The weighted decay time constant was calculated from the time constants of the double exponential fit to the decay phase of EPSCs. To measure the relative contributions of NMDAR- and non-NMDAR-mediated components to EPSC activity, a recording was first made with Vm clamped at −90 mV to sample non-NMDAR-mediated responses, then 10 μm DNQX was applied to block them and a recording was made with Vm clamped at +60 mV to displace Mg2+ from NMDARs and NMDAR-mediated responses were sampled, and, finally, 100 μm APV was applied to confirm that the recorded activity was mediated by NMDARs. NMDAR- and non-NMDAR-mediated currents were averaged over 10–40 individual sweeps and their peak amplitudes measured and the ratio taken. To measure the rectification index (RI) of non-NMDAR-mediated EPSCs in relay neurons, 100 μm APV was added to the bath to block NMDARs and EPSCs were evoked with Vm clamped at various levels, ranging from −90 to +60 mV in increments of 30 mV. Non-NMDAR-mediated synaptic currents were averaged over 10 individual sweeps for each clamped voltage and the peak amplitude measured. The RI was defined as the ratio of the measured current amplitude to the predicted value at +60 mV (extrapolated from the linear fit of the I–V relationship from −90 to 0 mV). For long-term potentiation (LTP) or LTD experiments, a stable baseline was recorded for at least 5 min, followed by an additional ∼25 min of recording after applying the LTP/LTD induction protocols. Synaptic responses were normalized to the averaged value of the baseline responses, and the normalized synaptic responses recorded ∼20–25 min after LTP/LTD induction were averaged and used for statistical comparison. All data are presented as the mean ± s.e.m. The non-parametric Mann–Whitney–Wilcoxon U test was used for the statistical comparison of parameters between CT EPSCs and ML EPSCs and the Wilcoxon matched-pairs sign-ranks test was used to compare responses before and after LTP/LTD induction or between simultaneously-recruited CT EPSCs and ML EPSCs. The statistical significance of the correlation analyses was assessed by the non-parametric Spearman's rank-order correlation test. The criterion for significance was P < 0.05.

Histochemistry

In all experiments, the recorded neuron was filled by passive diffusion of biocytin from the patch pipette during the recording period. After recording, the brain slices were fixed overnight at 4°C with 4% paraformaldehyde in 0.1 m phosphate buffer (PB, pH 7.4), rinsed several times with PB, and subjected to histochemical staining without further sectioning. Briefly, the slices were treated with 3% H2O2 in PB for 30 min, then with phosphate-buffered saline containing 0.03% Triton X-100 (PBST). They were then incubated with 2% bovine serum albumin and 10% normal goat serum in PBST for 1 h at room temperature, followed by overnight incubation at 4°C with avidin-biotinylated horseradish peroxidase complex in PBST (1:100 dilution; Vector Laboratories, Burlingame, CA, USA). Finally, bound horseradish peroxidase was visualized using 3,3-diaminobenzidine (DAB) as the chromogen. The staining results for the biocytin-filled neurons were examined and photographed under bright-field microscopy.

Results

Recording of ML EPSCs and CT EPSCs in TC relay neurons in the VBN

All of the recorded neurons were located in the VBN (Fig. 1A), as confirmed by post hoc histochemistry visualizing the injected biocytin. They had a polygonal soma, six to 10 large primary dendrites, and the bushy appearance of dendritic arborization (Fig. 1B). They all displayed Vm-dependent dual (continuous and burst) firing modes, with the watershed for mode switching at ∼ −60 mV (Fig. 1C). Since all of these physiological and morphological properties are consistent with previous reports (Peschanski et al. 1984; Jahnsen & Llinas, 1984) and there are no GABAergic interneurons in the VBN in rats (Spacek & Lieberman, 1974), we presumed that the recordings were made from TC relay neurons.

EPSCs were evoked in the TC relay neurons by positioning one stimulating electrode in the IC or the local VBN to evoke CT EPSCs and another in the ML tract to evoke EPSCs of the sensory inputs. We systemically compared EPSCs evoked by local stimulation and IC stimulation and found no significant difference in any of the examined characteristics (see Supplemental Table 1). We therefore considered that the EPSCs evoked by both IC and local stimulation were of cortical origin and refer to them as CT EPSCs. In response to an increased stimulation intensity, CT EPSCs showed a monotonic increase in amplitude (Fig. 1D). On the other hand, a substantial portion of the EPSCs evoked by stimulation of the ML tract showed all-or-none responses (Fig. 1E). Because all-or-none responses are a unique feature of sensory inputs to relay neurons (Castro-Alamancos & Calcagnotto, 1999; Castro-Alamancos, 2002; Miyata & Imoto, 2006), we considered that the EPSCs that were evoked by ML tract stimulation and showed all-or-none responses were from sensory inputs; these are referred to hereafter as ML EPSCs. EPSCs that failed to meet these two criteria, namely ML tract stimulation and all-or-none responses, were discarded without further analysis. Occasionally, all-or-none responses were evoked with the electrode positioned in the local VBN, which might have resulted from the stimulation of fibres from the ML or from layer V cortical neurons (Reichova & Sherman, 2004; Landisman & Connors, 2007). Since these EPSCs could not be clearly identified, these data were also discarded.

We first compared the dynamic characteristics and short-term plasticity of CT EPSCs and ML EPSCs. Significant differences were observed in onset latency, the 10–90% rise time, and the decay time constant (Table 1). In addition, upon application of paired-pulse stimulation (PPS) with an inter-pulse interval of 50 ms, CT EPSCs showed paired-pulse facilitation (Fig. 1F and Table 1), while ML EPSCs showed depression (Fig. 1G and Table 1). We also noted that, in response to PPS, while the latency of the first EPSC was shorter than that of the second by 0.21 ± 0.08 ms for ML EPSCs, the latency of the first was longer than that of the second by 0.28 ± 0.07 ms for CT EPSCs (Table 1); in other words, CT EPSCs showed conduction supernormality, while ML EPSCs showed conduction subnormality (Landisman & Connors, 2007). These differences in dynamic characteristics and short-term plasticity between CT EPSCs and ML EPSCs are consistent with previous reports (Castro-Alamancos & Calcagnotto, 1999; Castro-Alamancos, 2002; Miyata & Imoto, 2006; Landisman & Connors, 2007), and suggest that the CT EPSCs and ML EPSCs recorded were truly from cortical and sensory inputs.

Table 1.

Comparison of the dynamic characteristics and short-term plasticity of corticothalamic excitatory postsynaptic currents (CT EPSCs) and medial lemniscal (ML) EPSCs

CT EPSCs ML EPSCs SR# Statistics*
OL (ms) 4.0 ± 0.4 2.7 ± 0.2 7 P < 0.001
n = 18 n = 26
RT (ms) 2.7 ± 0.2 1.0 ± 0.1 6 P < 10−5
n = 16 n = 15
Dτ (ms) 10.7 ± 0.7 8.1 ± 0.7 6 P < 0.05
n = 16 n = 15
PPR 2.4 ± 0.2 0.4 ± 0.1 8 P < 10−8
n = 20 n = 26
LS (ms) 0.28 ± 0.07 −0.21 ± 0.08 5 P < 10−4
n = 15 n = 17
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OL, onset latency; RT, 10%–90% rise time; Dτ, weighted decay time constant; PPR, paired-pulse ratio; LS, latency shift (defined as the latency of the 1st EPSC minus the latency of the 2nd EPSC). The data are expressed as the mean ± s.e.m.

#

SR, number of experiments in which CT EPSCs and ML EPSCs were simultaneously recruited in a TC relay neuron.

*

Non-parametric Mann–Whitney–Wilcoxon test.

Differences in ionotropic glutamate receptor content between ML and CT synapses in the VBN

Both CT EPSCs and ML EPSCs were glutamatergic and consisted of non-NMDAR- and NMDAR-mediated components (Fig. 2A and E). The I–V relationship of the non-NMDAR-mediated component of ML EPSCs showed clear linearity, with a rectification index (RI) of 0.93 ± 0.09 (Fig. 2Ba, D). In contrast, CT EPSCs showed significant inward-rectification, with an RI of 0.61 ± 0.05 (Fig. 2Bb, D). There was a significant difference in RI between ML EPSCs and CT EPSCs (P < 0.01) (Fig. 2D), suggesting that CT synapses might contain the calcium-permeable α-amino-3-hydroxyl-5-methyl-isoxazolepropionic acid receptor (CaP-AMPAR) (Iino et al. 1990; Bowie & Mayer, 1995), while ML synapses do not. In support of this argument, we found that CT EPSCs were significantly attenuated by 10 μm PhTx-433 or 250 μm NAS, two selective CaP-APMAR blockers (Washburn & Dingledine, 1996; Koike et al. 1997), while ML EPSCs were not affected. As shown in Fig. 2C and D, application of PhTx-433 blocked ML EPSCs by 0.6 ± 6.0% and CT EPSCs by 44.6 ± 15.4% (n = 6; P < 0.05), while application of NAS blocked ML EPSCs by 4.5 ± 4.9% (n = 4) and CT EPSCs by 46.3 ± 5.5% (n = 6; P < 0.05). The relative contributions of NMDAR-mediated and non-NMDAR-mediated synaptic currents to glutamatergic transmission also showed a significant difference between CT EPSCs and ML EPSCs (Fig. 2E and F). The peak current ratio of NMDAR- to non-NMDAR-mediated components of CT EPSCs was 2.62 ± 0.61 (n = 14), significantly larger than the value of 0.43 ± 0.05 for ML EPSCs (n = 10; P < 10-4).

Expression of NMDAR-LTP and L-VGCC-LTD at CT, but not ML, synapses

Since NMDARs (Bliss & Collingridge, 1993; Bear & Malenka, 1994) and CaP-AMPARs (Lei et al. 2003; Ho et al. 2007) are essential for various types of synaptic plasticity in the brain, we examined whether the expression of LTP and LTD by these two synapses also differed. The standard conjunctive pairing of postsynaptic depolarization to 0 mV with 1 Hz presynaptic stimulation for 90 s was used for LTP induction (Gustafsson et al. 1987). This protocol did not result in LTP of ML EPSCs (95 ± 9% of the baseline; n = 7), but readily resulted in LTP of CT EPSCs (146 ± 11% of the baseline; n = 20, P < 10−4) and a persistent increase in GW (141 ± 12% of the baseline, P < 10−4) (Fig. 3A and B). The LTP of CT EPSCs was NMDAR dependent, as it was blocked when 50 μm APV was added to the bath to block NMDARs (97 ± 8% of the baseline; n = 9) (Fig. 3C). Interestingly, the increase in GW produced by the pairing protocol was also blocked by APV (110 ± 6% of the baseline) (Fig. 3C, lower panel), showing that the effect was also NMDAR dependent (see also Fan et al. 2005). Neither depolarization alone (110 ± 8% of the baseline; n = 4) nor 1 Hz presynaptic stimulation alone (101 ± 10% of the baseline; n = 5) induced LTP of CT EPSCs (Fig. 3D and E), demonstrating the properties of associativity and input specificity for LTP induction as previously reported (Bliss & Collingridge, 1993).

T-VGCCs are largely expressed in relay neurons and contribute to their unique feature of dual firing modes (Sherman, 2001). It was, therefore, of interest to know whether T-VGCCs also contributed to the induction of LTP of CT EPSCs. Since T-VGCCs in thalamic relay neurons undergo rapid inactivation (Kuo & Yang, 2001), it was unlikely that they would be sufficiently activated by the steady-state depolarization of the LTP induction protocol. Accordingly, we tried a modified protocol in which repeated depolarizing pulses (stepping from −70 to 0 mV at 1 Hz with a 50% duty cycle) were used to pair presynaptic stimulations, with no delay between the pre- and postsynaptic stimulations in each paired cycle. This protocol allowed the simultaneous activation of T-VGCCs and NMDARs, as Vm consistently returned to −70 mV and removed the inactivation of T-VGCCs before the start of the next paired cycle. Interestingly, application of this modified protocol for 90 cycles failed to induce LTP of CT EPSCs (98 ± 14% of the baseline, n = 11) (Fig. 4). It was possible that the modified protocol was not adequate for LTP induction or that some, as yet unidentified, form of LTD was simultaneously developed (Cummings et al. 1996; Lei et al. 2003; Ho et al. 2007) and counteracted LTP expression. To test these possibilities, we examined whether LTD of CT EPSCs could be induced by applying 90 depolarizing pulses alone to postsynaptic relay neurons without presynaptic stimulation. As shown in Fig. 5, LTD of CT EPSCs was readily induced (73 ± 4% of the baseline, n = 26; P < 10−4) (Fig. 5Aa and B), while the same protocol failed to induce significant LTD of ML EPSCs (110 ± 6% of the baseline, n = 9) (Fig. 5Ab and B). It is unlikely that LTD was caused by a running down of synaptic activity because the data shown in Fig. 5B include a series of experiments (n = 6) in which CT EPSCs and ML EPSCs were simultaneously recruited in the same relay neurons. A significant difference in the expression of LTD between CT EPSCs and ML EPSCs was found in these six experiments (P < 0.05). Unlike the induction of NMDAR-dependent LTP, no significant change in GW was observed after induction of LTD of CT EPSCs (Fig. 5C).

Figure 5. Application of repeated depolarizing pulses induces LTD at CT synapses, but not at ML synapses, on relay neurons.

Figure 5

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A, representative experiment showing the time courses of the CT EPSCs (Aa) and ML EPSCs (Ab) evoked in the same relay neuron. After a period of baseline recording, the application of depolarizing pulses (indicated by the horizontal bar) induced LTD of the CT EPSCs, but not the ML EPSCs. The insets are EPSC activity recorded before (black) and after (grey) application of depolarizing pulses, averaged over 7 sweeps of recordings. B, summarized results showing induction of LTD of the CT EPSCs (filled circles), but not the ML EPSCs (open circles) after application of 90 depolarizing pulses, the protocol of which is shown in the inset. Note that the data include 6 experiments in which the 2 inputs were simultaneously stimulated in the same relay neurons. The lower panel shows the time course of GW during recording; no significant variation in GW was found.

LTD of CT EPSCs is L-VGCC dependent

Induction of LTD of CT EPSCs was blocked by intracellular loading of 10 mm BAPTA, a calcium chelator (94 ± 5% of the baseline; n = 4) (Fig. 6A). Moreover, it was also blocked by addition of 10 μm nimodipine, a selective L-type VGCC (L-VGCC) blocker, to the bath (100%± 4% of the baseline; n = 11) (Fig. 6B). These results show that induction of LTD of CT EPSCs was L-VGCC dependent. To assess the possible role of T-VGCCs in LTD induction, we applied depolarizing pulses stepping from −50 to 0 mV, conditions under which T-VGCCs were inactivated, as no low-threshold spike was observed when the neurons were held at > −55 mV in current-clamp mode (Fig. 1C). As shown in Fig. 6C, LTD of CT EPSCs was still induced (63 ± 6% of the baseline; n = 12, P < 0.001). Because the above results implied that application of the modified conjunctive paring protocol also induced LTD, which masked LTP expression, LTD suppression by blocking L-VGCCs with nimodipine was expected to uncover LTP expression. As shown in Fig. 6D, addition of 10 μm nimodipine to the bath resulted in induction of significant LTP (118 ± 7% of the baseline; n = 14; P < 0.05) by the modified pairing protocol, although the magnitude was smaller than that of the LTP induced by the standard protocol (but not statistically different; P > 0.05).

LTD of CT EPSCs is independent of CaP-AMPARs

Ho et al. (2007) reported an L-VGCC-dependent LTD that involves the downregulation of CaP-AMPARs at developing mossy fibre synapses in the hippocampus. Since CT, but not ML, synapses were found to contain CaP-AMPARs, we tested whether expression of L-VGCC-dependent LTD at CT synapses involved the downregulation of CaP-AMPARs, thereby accounting for the expression of L-VGCC-dependent LTD at CT, but not at ML, synapses. We first examined the RI of the I–V relationship for non-NMDARs of CT EPSCs before and after induction of LTD. The recordings were performed with addition of 100 μm APV to ACSF to block NMDARs. As shown in Fig. 7A and B, the RI measured during baseline recording was 0.56 ± 0.08 (n = 6), which was not significantly different from the value measured after LTD induction (RI = 0.61 ± 0.06), showing that LTD expression did not alter the non-NMDAR content of CT synapses. It should be noted that an LTD of 62 ± 8% of the baseline was still induced when NMDARs were blocked in this series of experiments, showing that induction of LTD of CT EPSCs did not depend on NMDAR activation (Fig. 7A). We next examined whether induction of LTD of CT EPSCs required CaP-AMPARs. As shown in Fig. 7C, LTD was induced by the addition of 10 μm PhTx-433 to the ACSF to block CaP-AMPARs (64 ± 3% of the baseline, n = 8), showing that induction of LTD of CT EPSCs did not depend on CaP-AMPAR activation. Taken together, the above results showed that neither the expression nor the induction of LTD of CT EPSCs required CaP-AMPARs.

Figure 7. Induction and expression of L-VGCC-dependent LTD of CT EPSCs do not involve CaP-AMPARs.

Figure 7

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A, representative experiment showing the measurement of the RI of the I–V relationship of non-NMDAR mediated synaptic currents before and after induction of LTD of the CT EPSCs (Aa). The insets are EPSCs recorded before (left) and after (right) application of depolarizing pulses, averaged over 5–8 sweeps of recordings. The outward and inward currents show recordings made with Vm clamped at +50 and −70 mV, respectively. The summarized results from 8 experiments show the LTD of CT EPSCs (Ab) when recordings were made with APV added to the ACSF. The lower panel shows the time course of GW during recording; no significant variation in GW was found. B, summarized results showing no change in the RI before (baseline) and after (LTD) LTD induction. The data are the results for 6 cells, with individual results being shown as open circles with dotted lines and the average as filled circles with a continuous line. C, representative experiment showing induction of LTD at CT synapses with CaP-AMPARs blocked by addition of PhTx-433 to the ACSF (Ca). The insets are EPSCs recorded before (black) and after (grey) application of depolarizing pulses, averaged over 10 sweeps of recordings. Summarized results from 8 experiments show LTD was still induced with blockade of CaP-AMPARs (Cb). The lower panel in Cb shows the time course of GW during recording; no significant variation in GW was found.

L-VGCC-dependent LTD of CT EPSPs can be induced by continuous, but not burst, spiking of postsynaptic relay neurons

In current-clamp mode, we found there were QX-314-resistant spikes that were mediated by L-VGCCs (Supplemental Fig. 1). This raised the possibility that physiological spiking might be able to activate L-VGCCs sufficiently to induce LTD of CT EPSPs in relay neurons. We thereby tested whether LTD of CT EPSPs could be induced in a relatively physiological context by testing the effects of two different physiologically relevant firing patterns on CT EPSPs recorded in relay neurons. As shown in Fig. 8A, a stable baseline of CT EPSPs was first recorded, followed by the introduction of continuous spiking of the postsynaptic relay neuron elicited by injection of depolarizing current pulses (500 pA, 50% duty-cycle at 1 Hz) for 90 s with Vm held at ∼ −55 mV and temporary cessation of presynaptic stimulation, after which presynaptic stimulation was resumed and a slowly developing LTD of CT EPSPs was observed. Figure 8C shows the summarized results from nine experiments, which show that significant LTD of CT EPSPs (82 ± 7% of the baseline; P < 0.05) was induced by continuous spiking of postsynaptic relay neurons. Although the magnitude was smaller, it did not differ significantly from the LTD induced by depolarizing pulses using voltage-clamp recording. Furthermore, its induction could be prevented by blocking L-VGCCs. As shown in Fig. 8C (grey squares), the LTD of CT EPSPs was blocked by 10 μm nimodipine (actually, there was a slight potentiation of CT EPSPs following blockade of L-VGCCs with a value of 114 ± 5% of the baseline, n = 10; P < 0.05). However, it was possible that this small LTD might have resulted from small changes in Rn, so we measured this parameter to test this possibility. Rn was measured every 10 s and the values measured 20–25 min after LTD induction protocol averaged and normalized to the value measured during baseline recording. As shown in Fig. 8C (lower panel), the averaged Rn for recordings at 20–25 min after LTD induction in control medium or in the presence of nimodipine was 94 ± 3% or 97 ± 4% of the baseline, respectively, and the difference was not significant. The range of the mean spiking frequency of the relay neurons was 35–85 Hz and no significant correlation was seen between the mean frequency of spiking and the magnitude of the induced LTD. In contrast, when repetitive burst spiking was induced, it failed to induce significant LTD of CT EPSPs (116 ± 18% of the baseline; n = 8) (Fig. 8D). Burst spiking was elicited by injection of a current pulse with a short duration (2 ms) and high intensity (2 nA) at 1 Hz for 90 cycles, with Vm at ∼ −70 mV (Fig. 8B, D). These results show that LTD of CT synapses can be induced by physiological spiking of relay neurons in a spiking pattern-dependent manner.

Discussion

In addition to previously reported differences (see below), in this study, we found that the ionotropic glutamate receptor content and the expression of NMDAR-dependent LTP and L-VGCC-dependent LTD differed between sensory and cortical inputs on thalamic relay neurons in the VBN. Of particular interest was the observation that induction of L-VGCC-dependent LTD was spiking pattern dependent. These results suggest that cortical feedback to thalamic relay neurons can be dynamically tuned in a use-dependent manner and that the spiking pattern of thalamic relay neurons plays an indispensable role in some part of this regulation.

Previous studies have demonstrated several pathway-specific properties of cortical and sensory inputs onto relay neurons. First, the onset latency of ML EPSCs is shorter than that of CT EPSCs (Castro-Alamancos, 2002; Landisman & Connors, 2007), and conduction supernormality, which has been proposed to be specific to CT axons originating from layer VI pyramidal neurons of the cerebral cortex (Li et al. 2003; Landisman & Connors, 2007), is observed in CT EPSCs, but not ML EPSCs. Our recording of conduction supernormality was ∼0.3 ms, a value consistent with that reported by Landisman & Connors (2007). The second previously identified difference between ML and CT synapses is that the rise and decay of ML EPSCs are faster than those of CT EPSCs (Miyata & Imoto, 2006). Since CT inputs synapse on more distal dendritic compartments of relay neurons than ML inputs (Bourassa et al. 1995), these differences in EPSC shape might simply be ascribed to the differential dendritic cable filtering effects of CT and ML EPSCs. In addition, other factors, such as the existence of kainate receptor-mediated components in CT EPSCs, but not ML EPSCs, may further contribute to the difference in EPSC shape between the two synapses (Miyata & Imoto, 2006). Finally, the most well-known differences between the two synapses are the stimulus–response relationship and short-term plasticity, with CT EPSCs showing a linear stimulus–response relationship and paired-pulse facilitation, while ML EPSCs show an all-or-none response and paired-pulse depression (Castro-Alamancos & Calcagnotto, 1999; Castro-Alamancos, 2002; Reichova & Sherman, 2004; Miyata & Imoto, 2006). Since all of the above features were confirmed in the present study, we are confident that our recordings of CT and ML EPSC(P)s in relay neurons were truly from cortical and sensory inputs.

The other important difference between CT and ML synapses found in this study was the ionotropic glutamate receptor content, which may account for the difference in ability to express long-term synaptic plasticity between the two synapses. In line with the observation made in mouse thalamic slices (Miyata & Imoto, 2006), we found that the ratio of NMDAR-mediated to non-NMDAR-mediated EPSCs at CT synapses was significantly larger than that at ML synapses. This difference might explain the readily inducible NMDAR-dependent associative LTP at CT synapses, but not at ML synapses. However, Castro-Alamancos & Calcagnotto (1999) reported that the homosynaptic LTP of CT synaptic input induced with presynaptic stimulation at 10 Hz is NMDAR independent. This discrepancy between their report and our results may stem from the different protocols used for LTP induction (homosynaptic vs. associative) and/or different species of animals used (mice vs. rats). NMDAR-LTP of CT synapses was induced with conjunctive pairing of prolonged postsynaptic depolarization to 0 mV with 1 Hz presynaptic stimulation, but not with repeated depolarizing pulses used to pair presynaptic stimulations. Since the application of repeated depolarizing pulses alone could result in LTD of CT EPSCs, a straightforward interpretation of this observation is that LTP and LTD were simultaneously induced by conjunctive pairing of repeated depolarizing pulses with presynaptic stimulations. This argument is supported by our results showing induction of LTP using depolarizing pulses paired with presynaptic stimulations when L-VGCCs were blocked by nimodipine. However the magnitude of the LTP was smaller than that induced by prolonged postsynaptic depolarization and mechanisms other than the proposed simple model of simultaneous LTP and LTD cancelling each other out should be considered.

Pairing of postsynaptic depolarization to 0 mV with presynaptic stimulation of the CT pathway not only induced LTP of CT EPSCs, but also induced a parallel increase in GW, which was not observed when NMDARs were blocked (Fig. 3C), when depolarization (Fig. 3D) or CT pathway stimulation (Fig. 3E) was applied alone, or when stimulation of the CT pathway was paired with depolarizing pulses (Fig. 4B). Furthermore, it was not observed when postsynaptic depolarization to 0 mV was paired with ML pathway stimulation (Fig. 3B, open circles). These observations suggest that the increase in GW was NMDAR dependent and also had the properties of associativity and input specificity. In pyramidal neurons in the hippocampal CA1 area, in parallel with synaptic potentiation, activation of NMDARs by paired pre- and postsynaptic activities also activates HCN1 channels, which mediate hyperpolarization-activated cationic currents (Ih), and thus reduce neuronal excitability (Fan et al. 2005). It is possible that the increase in GW in the present study was due to an increase in the Ih. Since such a reduction in cellular excitability, together with synaptic potentiation, could provide a feedback mechanism for normalizing neuronal output firing and enhancing synaptic specificity (Fan et al. 2005), it will be important to determine whether a similar mechanism exists in cortical inputs on thalamic relay neurons in the VBN. Unlike the NMDAR-dependent LTP, the induction of LTD of CT EPSCs by repeated voltage pulses was not associated with a significant change in GW (Fig. 6, 7).

Our argument for the presence of CaP-AMPARs at CT synapses, but not at ML synapses, is based on the observations that the CT EPSCs showed inward rectification and were sensitive to CaP-AMPAR-specific polyamine toxins. Neither of these features was observed for ML EPSCs. Ho et al. (2007) reported a form of L-VGCC-dependent LTD that involves the downregulation of CaP-AMPAR expression at developing mossy fibre synapses in the hippocampus. Although CT synapses also contained CaP-AMPARs, our results showed that neither the induction nor the expression of L-VGCC-dependent LTD required CaP-AMPARs. Accordingly, the selective expression of CaP-AMPARs at CT synapses, but not at ML synapses, could not account for the difference between the two synapses in the ability to express L-VGCC-dependent LTD, and the physiological role of CaP-AMPARs in the functions of CT synapses requires further investigation. It has been reported that some forms of LTD can be induced by the co-activation of L-VGCCs and either group I metabotropic glutamate receptors (Lin et al. 2006; Adermark & Lovinger, 2007; Naie et al. 2007) or type 1 cannabinoid receptors (Auclair et al. 2000; Diana & Marty, 2004; Chevaleyre et al. 2006), and the differential expression of these receptors may account for the expression of L-VGCC-dependent LTD at CT, but nor at ML, synapses.

L-VGCCs in TC relay neurons have been reported to undergo fast calcium-dependent inactivation during a long period of membrane depolarization (Kammermeier & Jones, 1998; Meuth et al. 2002). This may explain why L-VGCCs appeared to be more efficiently activated by repeated depolarizing voltage pulses than by steady-state depolarization, as application of steady-state depolarization to postsynaptic relay neurons alone did not induce significant LTD (Fig. 3D). In the current-clamp recordings, a train of L-VGCC-mediated spikes was induced upon application of a depolarizing current injection with Vm held at ∼40 mV (see Supplemental Fig. 1). Sufficient calcium influx might occur during this kind of continuous spiking activity to result in LTD of synaptic inputs of cortical origin on TC relay neurons. In contrast, only a small number of overshooting spikes were generated when TC relay neurons were driven to fire in burst mode with Vm held at ∼ −70 mV (data not shown). Under these conditions, the influx of calcium might have been too small to induce any significant change in the strength of the synaptic inputs on TC relay neurons. This spiking-pattern-dependent modification of synaptic inputs of cortical origin is interesting and functionally important. Provided cortical feedback can change the status of spiking of relay neurons (Murphy & Sillito, 1987; Ergenzinger et al. 1998; Krupa et al. 1999; Sillito & Jones, 2002), the altered spiking activity of TC relay neurons would modify the strength of cortical inputs onto relay neurons. In other words, this spiking-pattern-dependent induction of LTD of cortical inputs provides a means by which cortical modulation of the transfer of somatosensory information to the cortex can be dynamically tuned.

In conclusion, the present results suggest that peripheral sensory information, carried by ML synapses, is faithfully and effectively conducted to the VBN, and that the use-dependent change in their synaptic strengths is limited. In contrast, LTP and LTD are readily inducible at CT synapses. In vivo, TC relay neurons commonly have a high, noisy synaptic conductance caused by corticothalamic barrages (Steriade, 2001). Wolfart et al. (2005) showed that spike transfer from the thalamus to the cortex can be controlled by background synaptic activity in thalamic relay neurons. The induction and expression of use-dependent LTP and LTD of CT synaptic transmission might provide a mechanism for adjusting the extent of background synaptic barrages from cortical feedback on TC relay neurons, and, thus, for regulating spike transfer to the cortex. Of particular interest is the observation that this adjustment of plasticity is state dependent, as shown by our findings that induction of CT LTP was masked by expression of CT LTD by manipulations in which L-VGCCs were activated. Furthermore, CT LTD could only be induced by a continuous-spiking, and not a burst-spiking, pattern. Thus, long-term CT-TC relay neuron plasticity might be selectively operating when an animal is awake and conscious.

Acknowledgments

This work was supported by grants from the National Science Council, Taiwan (NSC95-2320-B-002-067 and NSC96-2311-B-002-025-MY2).

Glossary

Abbreviations

AP

action potential

CaP-AMPAR

calcium-permeable AMPA receptor

CT

corticothalamic

EPSC

excitatory postsynaptic current

GW

whole-cell conductance

IC

internal capsule

L-VGCC

L-type voltage-gated calcium channel

ML

medial lemniscus

NMDAR

NMDA receptor

PPS

paired-pulse stimulation

Rn

input resistance

Rs

series resistance

TC

thalamocortical

VBN

ventrobasal nucleus

T-VGCC

T-type voltage-gated calcium channel

VGCC

voltage-gated calcium channel

Vm

membrane potential

Author contributions

C-L.H. collected and analysed the data. C-T.Y. helped in developing the use of appropriate thalamic slices and wrote part of the manuscript. H-W.Y. and M-Y.M. designed experiments and wrote the manuscript. All authors approved the final version for publication.

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