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Abstract

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix

Synaptic plasticity at corticostraital synapses is proposed to fine tune movment and improve motor skills. We found paired-pulse plasticity at corticostriatal synapses reflected variably expressed short-term facilitation blended with a consistent background of longer-lasting depression. Presynaptic modulation via neuotransmitter receptor activation was ruled out as a mechanism for long-lasting paired-pulse depression by examining the effect of selective receptor antagonists. EPSC amplitude and paired-pulse plasticity, however, was influenced by block of D2 dopamine receptors. Block of glutamate transport with l-transdicarboxylic acid (PDC) reduced EPSCs, possibly through a mechanism of AMPA receptor desensitization. Removal of AMPA receptor desensitization with cyclothiazide reduced the paired-pulse depression at long-duration interstimulus intervals (ISIs), indicating that AMPA receptor desensitization participates in corticostriatal paired-pulse plasticity. The low-affinity glutamate receptor antagonist cis-2,3-piperidine dicarboxylic acid (PDA) increased paired-pulse depression, suggesting that a presynaptic component also exists for long-lasting paired-pulse depression. Low Ca2+–high Mg2+ or BAPTA-AM dramatically reduced the amplitude of corticostriatal EPSCs and both manipulations increased the expression of facilitation and, to a lesser extent, they reduced long-lasting paired-pulse depression. EGTA-AM produced a smaller reduction in EPSC amplitude and it did not alter paired-pulse facilitation, but in contrast to low Ca2+ and BAPTA-AM, EGTA-AM increased long-lasting paired-pulse depression. These experiments suggest that facilitation and depression are sensitive to vesicle depletion, which is dependent upon changes in peak Ca2+ (i.e. low Ca2+–high Mg2+ or BAPTA-AM). In addition, the action of EGTA-AM suggests that basal Ca2+ regulates the recovery from long-lasting paired-pulse depression, possibly thourgh a Ca2+-sensitive process of vesicle delivery.

Basal ganglia-related behaviours are defined in large part by the functional and anatomical link existing between the cortex and the striatum. An additional mechanism potentially involved in fine-tuning striatal-related behaviour is the plasticity expressed at corticostriatal synapses (Calabresi et al. 1996). Changes in plasticity at these synapses are also implicated in behavioural problems seen in ageing, age-related disease and certain addictive disorders (Grace, 1995; Calabresi et al. 1996; Ou et al. 1997; Canales et al. 2002; Akopian & Walsh, 2006).

Ipsilateral layer V cortical neurons and, to a lesser extent, contralateral layer II–III neurons send axons to the striatum (McGeorge & Faull, 1987; Kolb et al. 1992; Reiner et al. 2003). The primary target for these cortical projections is the most common cell type found in the striatum, medium spiny (MS) projection neurons. Cortical input retains much of its somatotopic and functional organization through dorsoventral, medio-lateral and rostro-caudal extensions of the striatum (McGeorge & Faull, 1987; Deniau et al. 1996). MS neurons also receive glutamatergic input from the thalamus, serotonergic input from the raphe and dopaminergic input from the substantia nigra (Beckstead et al. 1979; Priestley et al. 1981; Deschenes et al. 1995). In addition, MS cells receive synapses from cholinergic and GABAergic interneurons and synapses made by axonal collaterals from neighbouring MS cells (Chang & Kita, 1992; Bennett & Bolam, 1994; Kawaguchi et al. 1995; Tepper et al. 2004).

Excitatory postsynaptic potentials (EPSPs) evoked by activation of corticostriatal synapses are mediated primarily by AMPA receptors, with a smaller contribution coming from NMDA receptors when cells are depolarized (Akopian & Walsh, 2002). Delivery of stimuli to the corpus callosum recruits many corticostriatal synapses, which express varied plasticity at interstimulus intervals (ISIs) shorter than 100 ms (Mori et al. 1994a,b; Akopian et al. 2000). In vitro brain slice analysis has also shown anatomical and developmental differences in short- and long-term synaptic plasticity at these synapses (Choi & Lovinger, 1997; Partridge et al. 2000; Smith et al. 2001). Each corticostriatal synapse appears to uniquely blend the processes of facilitation and depression to transmit information, which is in contrast to the uniformity in plasticity often seen at other central nervous system (CNS) synapses. For example, the climbing fibre synapses in the cerebellum have proven to be a model system for studying short-term synaptic depression (Dittman & Regehr, 1998; Xu-Friedman & Regehr, 2003), while hippocampal synapses are well suited for studying short-term forms of synaptic facilitation (Dittman et al. 2000). The present study investigates how plasticity changes as a function of the ISI between paired activation of cortical afferents, the potential modulation of the plasticity by other transmitter systems in the striatum, and the role played by corticostriatal synapse handling of Ca2+.

Methods

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix

Experiments were performed on Fischer 344 rat pups postnatal days (P) P9–19. Pregnant Fischer 344 rats were purchased from Harlan laboratories. Pups were anaesthetized by placing them in a desiccator containing halothone vapours (in a hood) and they were killed by decapitation in accordance with a protocol approved by the University of Southern California's Institutional Animal Care and Use Committee (IACUC). Brains were removed and blocked in cold low-sodium sucrose-substituted saline (see below). Coronal sections (200 μm thick) of the striatum were cut in ice-cold low-sodium sucrose-substituted saline (Camden vibroslicer, WPI). Slices were then stored in artificial cerebral spinal fluid (aCSF) at room temperature (23°C) for at least 1 h prior to recording. All solutions were continuously oxygenated with 95% O2–5% CO2. Slices were then transferred to a submerged brain slice-recording chamber perfused with oxygenated aCSF kept at a recording temperature of 23°C.

aCSF consisted of 124 mm NaCl, 1.3 mm MgSO4, 3 mm KCl, 1.25 mm NaH2PO4, 25 mm NaHCO3, 2.4 mm CaCl2, and 10 mm glucose. Sucrose saline was identical except NaCl was reduced to 90 mm and 105 mm sucrose was added (modified from Aghajanian & Rasmussen, 1989). These efforts were taken to minimize excitotoxic damage that could occur during brain removal, blocking and making of brain slices. The pH of all oxygenated solutions was 7.4. All experiments were performed with 30 μm bicuculline methiodide (BIC–aCSF; Sigma). BIC was used to block γ-amino butyric acid-A (GABAA) receptor-mediated inhibition in an attempt to isolate excitatory synaptic events.

The following drugs were used in different phases of the study: adenosine; α-methyl-4-carboxyphenylglycine (MCPG); (RS)-α-methyl-4-sulphonophenylglycine (MSPG); 1S,3R-1-aminocyclopentane-1,3-dicarboxylic acid (t-ACPD); baclofen; 1,2-bis(2-aminophenoxy) ethane-N,N,N′,N′-tetraacetic acid (BAPTA) (AM ester, BAPTA-AM); cis-2,3-piperidine dicarboxylic acid (PDA); 8-cyclopentyl-1,3-dipropylxanthine (DPCPX); 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX); cyclothiazide; N-ethylmaleimide (NEM); ethylene glycol-bis (2-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA) (AM ester, EGTA-AM); raclopride; saclofen; l-sulpiride; theophylline; l-transdicarboxylic acid (PDC) (all were purchased from Sigma). P-[3-aminopropyl]-P-diethoxymethylphosphinic acid (CGP 35348) was a gift from Novartis (formerly Sandoz).

Whole-cell voltage clamp and electrical stimulation

Whole-cell voltage clamp methods were used to examine corticostriatal synaptic plasticity to reduce possible activation of postsynaptic conductances, which can contribute to plasticity under current clamp conditions (Akopian & Walsh, 2002). Whole-cell recordings from visualized neurons were obtained using a fixed stage microscope and water immersion lenses (Zeiss Axioscop, Germany). Patch electrodes were pulled on a Flaming/Brown P-87 Micropipette Puller (Sutter Instruments) and backfilled with internal solutions. The internal electrode solution consisted of (mm): 120 caesium gluconate; 2 MgCl2; 0.5 EGTA; 10 Hepes; 10 TEA; 3 QX-314; 3 Na-ATP; pH 7.2; (270–280 mosmol l−1). Electrode resistance was constantly monitored in voltage clamp mode using the Clampex data acquisition software and an Axopatch-1D patch clamp amplifier (Axon Instruments). Series resistance was monitored throughout the experiment by measuring the instantaneous current response to a −5 mV voltage step from −70 mV. A gravity-fed array of inflow tubes of ∼100 μm inner diameter and an outflow tube attached to a vacuum reservoir provided solution flow. The ground electrode consisted of a salt bridge constructed from glass electrode filled with agar.

Large tipped whole-cell electrodes filled with 140 mm NaCl were positioned 100–500 μm from the recording electrode at the border between the striatum and the overlying corpus callosum. All recordings were taken from the dorsomedial aspect of the striatum, in coronal sections. Constant current stimuli (10–100 μA) were delivered using durations less than 0.1 ms through an 8365 stimulus isolator unit (WPI, Sarasota, FL, USA). Stimulation intensity was set to evoke approximately the same amplitude synaptic response in each cell (i.e. 200 pA). Each cell received the same sequence of paired-pulse stimulation using ISIs, which enhance facilitation and isolate synaptic depression. Paired-pulse plasticity obtained from stimuli delivered at an ISI of 50 ms was used as an index of facilitation and paired-pulse plasticity evoked at an ISI of 500 ms was used as an index of synaptic depression. These two ISIs were delivered every 30 s, alternating ISI 50 ms to ISI 500 ms before, after and during application of experimental drugs. In some experiments the total distribution of ISIs consisting of ISIs of 50 ms, 100 ms, 250 ms, 500 ms, 1 s, 5 s and 10 s were examined. Pharmacological analyses were performed by comparing drug effects to values obtained in the same cell in aCSF alone and statistical analyses were therefore performed using paired t tests.

Spontaneous EPSCs (sEPSCs) were recorded for 2 min before and after the addition of raclopride. Average sEPSC amplitude and average sEPSC frequency was calculated automatically by pCLAMP software for the 2 min sampling period. Cumulative frequency histograms were generated for sEPSC amplitude and sEPSC frequency for each cell. Average cumulative frequency histograms were also generated for sEPSC amplitude and frequency by grouping all cells studied.

Results

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix

Our electrophysiological investigation of corticostriatal synaptic function focused on cortical contacts made with medium-sized spiny projection neurons (MS) located in the dorsomedial aspect of the striatum. MS neurons are the overwhelming majority of neurons found in the striatum (around 95%) (Lucas & Harlan, 1995). All striatal MS neurons are GABAergic, but differences exist in the co-expression of other neuronal markers (i.e. tachykinins and opiate peptides), the formation of cell islands called patches and in the projections each cell type makes (Graybiel & Chesselet, 1984; Lucas & Harlan, 1995). This study was not designed to account for differences in synaptic physiology associated with striatal expression of these markers. However, previous work by Kawaguchi et al. 1989) did not reveal any difference in medium spiny neuron synaptic physiology based upon the distribution of striatal patch or matrix markers. We were sensitive to potential medial-to-lateral differences in expression of paired-pulse plasticity and limited this source of variation by recording from striatal neurons located in the rostral, dorsomedial part of the striatum (Partridge et al. 2000; Smith et al. 2001).

The simple protocol of pairing two action potentials, or in the case of our preparation, two stimuli to an afferent pathway, mobilized processes dependent upon the temporal separation of the two stimuli. Paired-pulse depression became apparent at corticostriatal synapses at ISIs of 10 s or less (Fig. 1A). This form of synaptic depression was consistently observed at every set of corticostriatal synapse examined. The level of depression grew in magnitude to peak at an ISI of 500 ms (Fig. 1B). The 500 ms ISI was thus used as the testing interval for evaluating long-lasting paired-pulse depression in all experiments and it produced an average paired-pulse ratio (PPR) of 72.2 ± 0.82% (n= 133, s.e.m.) (Fig. 1C). The 500 ms peak was not a true peak, however, since facilitation could be triggered at shorter duration ISIs. Our readout of synaptic release, the evoked EPSC, thus reflected the summed interaction of what appeared to be independent processes of paired-pulse facilitation and depression (Fig. 1). When facilitation was present, it was most clearly seen at ISIs of 100 ms or less, but its expression varied considerably between cells studied (Fig. 1C). The average PPR measured at an ISI of 50 ms was 90.5 ± 1.62% (n= 133, s.e.m.). As a first approximation to test for the independence between the processes governing paired-pulse plasticity evoked at ISIs of 50 versus 500 ms we compared the variance of the PPR measured at these ISIs and found a clear difference (d.f. 1,218; Levene statistic, 28.156, P= 0.0001) (Fig. 1C). Regressions were also performed between the amplitude of the first EPSC and the PPR (both at 50 and 500 ms ISI), but no relationship was found.

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Figure 1. Paired-pulse plasticity at corticostriatal synapses A, superimposed paired responses for the ISIs indicated. Cells were voltage clamped at the resting membrane potential of −70 mV. B, paired-pulse ratio (EPSC2/EPSC1) is plotted as a function of ISI (the ISI is plotted on a logarithmic scale). Each time point reflects the average of 3 consecutive pairs separated by an inter-trial interval of 30 s. •, the average obtained from 25 cells. ⋄ and ○, individual examples plotted to illustrate the range of paired-pulse plasticity observed at ISIs shorter than 1 s. C, frequency histograms of paired-pulse ratio measured at ISIs of 50 and 500 ms. These two intervals were selected to represent peaks of facilitation (50 ms) and depression (500 ms) at corticostriatal synapses. The large number of cells in this sample comes from including the many ‘controls’ accumulated during each phase of pharmacological testing. The paired-pulse ratio bin size for the abscissa is 5%. The average paired-pulse ratio measured at 50 ms was 89.9% (range: 48.7–141.1%) and at 500 ms was 71.9% (range: 50.9–94.3%). D, frequency histograms separating cells into three categories of PPR at the 50 ms ISI: depression (PPR > 95%), no change (95% < PPR < 105%) and facilitation (PPR > 105%). Histograms are illustrated for all cells (n= 155) and then separated based upon postnatal age. P9–P11 (n= 29), P12–P15 (n= 65) and P16–P19 (n= 61).

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Tang et al. (2001) found that corticostriatal paired-pulse plasticity measured at an ISI of 50 ms was relatively constant from P10–P20, but after P20 the incidence of paired-pulse potentiation increased. Our sample of cells were younger than the P20 transition described by Tang et al. (2001), ranging from P9 to P19 with a mean age of 15.2 ± 0.2 days (n= 155). The change in the sample size reflects the incorporation of additional cells where we did not study the 500 ms ISI. Much like Tang et al. (2001), we found little change in the percentage of cells expressing facilitation or depression over the P9–P19 age range of our study. Cells were categorized as showing synaptic depression (PPR < 95%), no change (95% < PPR < 105%) or facilitation (PPR > 105%). When all ages were combined we found 66% showed depression, 17% showed no change and 17% showed facilitation (n= 155). In the P9–P11 group, 62% showed depression, 10% showed no change and 28% showed facilitation (n= 29). In the P12–P15 group, 66% showed depression, 15% showed no change and 19% showed facilitation (n= 65). In the P16–P19 group, 69% showed depression, 21% showed no change and 10% showed facilitation (n= 61) (Fig. 1D).

Role of neuromodulation

Presynaptic cortical neuron cell bodies are distant from striatal recording sites and their input to striatal neurons is not laminar. Corpus callosum stimulation is used to approximate corticostriatal stimulation, but current spread from the stimulation site is likely to activate other, modulatory components of striatal circuitry. The long time-course of paired-pulse depression (up to an ISI of 10 s, Fig. 1) suggested that modulatory neurotransmitters acting via G-proteins could be involved in its expression. To investigate this possibility we screened the action of a number of modulators with receptors in the striatum as well as a G-protein inhibitor.

Each experiment began by testing whether the putative modulator (i.e. adenosine) altered synaptic function. If the agonist affected synaptic transmission, we examined the ability of selective receptor antagonists to block the action of the agonist. These two steps established ‘modulator candidacy’. Once the antagonist efficacy was demonstrated, we applied the antagonist alone to see if it had any effect on the paired-pulse depression. These three steps (agonist, agonist + antagonist, antagonist alone) were carried out in parallel in separate cells to avoid artifacts associated with changes in receptor sensitivity resulting from multiple exposures to agonists or antagonists. Each experiment examined EPSC amplitude produced by a single stimulus and the paired-pulse plasticity produced by paired activation at ISIs of 50 and 500 ms. Intervals of 50 and 500 ms were chosen since these intervals produced peaks of facilitation and depression as measured in this study (Fig. 1). Treatment effects on synaptic function were examined using a paired statistical format, since the same set of synapses was examined before and after each drug exposure.

Adenosine (50 μm) (Fig. 2) caused a significant reduction in the amplitude of corticostriatal EPSCs and it increased the paired-pulse plasticity observed at 50 and 500 ms ISIs (n= 6, paired t test, P < 0.05). The action of adenosine was blocked by the selective A1 adenosine receptor antagonist DPCPX (0.5 μm) as well as by the non-selective adenosine receptor antagonist theophylline (100 μm; n= 3; Fig. 2B). Addition of either adenosine antagonist alone, however, did not affect the paired-pulse plasticity measured at 50 or 500 ms (Fig. 2C; n= 5). Figure 2C was generated by normalizing the post-drug effect on synaptic function to the pre-drug (control) measures of EPSC amplitude and paired-pulse plasticity (PPR for ISIs of 50 and 500 ms) for each cell studied (treatment-induced change).

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Figure 2. Adenosine receptor activation does not contribute to the paired-pulse plasticity created by activation of corticostriatal synapses in vitro A, representative current traces illustrating adenosine-mediated reductions of EPSC amplitude and increases in the paired-pulse ratio (PPR) measured at 50 and 500 ms ISIs. B, bar graph illustrating how adenosine (50 μm) reduced EPSC amplitudes to 26.9 ± 3.1% of control (P < 0.003, n= 6). PPR at an ISI of 50 ms changed from 91.6 ± 8.4% to 237 ± 28.2% of control (P < 0.002) and the PPR at an ISI of 500 ms changed from 71.8 ± 3.3% to 116.6 ± 10.4% of control (P < 0.02, n= 6). C, treatment induced change in EPSC amplitude and PPR (50 and 500 ms). Measures of EPSC amplitude and PPR for 50 and 500 ms were obtained before and after each adenosine-related treatment. Post-treatment measures were normalized to pretreatment measures and illustrated as the percentage change in each value created by the treatment (EPSC amplitude and PPR for 50 and 500 ms ISIs). Each cell was exposed to a single drug application condition (i.e. theophylline alone, adenosine + theophylline, etc.) As shown in B, adenosine had a clear effect on EPSC amplitude and the PPR at 50 and 500 ms. The adenosine effects were blocked with the non-selective antagonist theophylline (100 μm) and the selective A1 receptor antagonist DPCX (0.5 μm). Addition of either antagonist alone did not affect EPSC amplitude or paired-pulse plasticity.

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The GABAB receptor agonist baclofen (5 μm) also reduced corticostriatal EPSC amplitudes (P < 0.002) and increased the PPR measured at 50 ms (P < 0.02) and 500 ms (P < 0.03) ISIs (Fig. 3A and B; paired t tests, n= 5). The selective GABAB receptor antagonists saclofen (500 μm) and CGP 35348 (10 μm) blocked these actions of baclofen (5 μm) (Fig. 3B; n= 3). To test for possible GABAB receptor activation in the expression of striatal paired-pulse plasticity we applied either saclofen (500 μm) or CGP 35348 alone and found no effect on paired-pulse plasticity at either 50 or 500 ms ISI (Fig. 3B; n= 4). Interestingly, both saclofen and CGP 35348 caused a reduction in the amplitude of EPSCs, possibly due to a non-specific effect as reported earlier by Nisenbaum et al. (1992) (P < 0.03).

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Figure 3. GABAB receptor activation does not contribute to the paired-pulse plasticity created by activation of corticostriatal synapses in vitro A, representative current traces illustrating baclofen-mediated reductions of EPSC amplitude and increases in the PPR measured at 50 and 500 ms ISIs. Baclofen (5 μm) reduced EPSC amplitude to 10.7 ± 1.7% of control (P < 0.002, n= 5). PPR at an ISI of 50 ms changed from 87.9 ± 8.6% to 186 ± 30.4% (P < 0.02) and the PPR at an ISI of 500 ms changed from 74.8 ± 3.6% to 132.0 ± 16.5% (P < 0.03, n= 5). B, GABAB receptor-related treatment outcomes on EPSC amplitude and PPR at 50 and 500 ms ISIs are plotted as a percentage of these measures obtained prior to the indicated pharmacological treatment. Baclofen effects were partially blocked by the selective GABAB receptor antagonists saclofen (500 μm) and CGP 35348 (500 μm). Addition of saclofen (P < 0.05, n= 5) or CGP 35348 (P < 0.05, n= 3) alone both reduced the amplitude of EPSCs, but they did not affect paired-pulse plasticity.

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A similar profile was found for mGLuRs. The selective metabotropic glutamate receptor (mGluR) agonist t-ACPD (5 μm) reduced corticostriatal EPSCs (P < 0.02) and increased the PPR at 50 ms (P < 0.03) and 500 ms (P < 0.03) ISIs (paired t test, n= 6). The selective mGluR receptor antagonist MCPG (500 μm) eliminated t-ACPD's change in the PPR, but it did not fully block t-ACPD's reduction of the EPSC (P < 0.02, n= 5) (Hashimoto & Kano, 1998). Addition of MCPG alone, however, did not affect EPSC amplitude or the PPR measured at ISIs of 50 and 500 ms (n= 8) (Fig. 4). We also tested the selective mGluR2,3 antagonist MSPG (100 μm) and found that this also did not fully block the action of t-ACPD (n= 5) (Fig. 4).

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Figure 4. Analysis of mGluR modulation of corticostriatal EPSCs Addition of the mGluR agonist t-ACPD (5 μm) reduced the amplitude of the EPSC (P < 0.01) and increased the PPR at 50 ms (P < 0.02) and 500 ms (P < 0.03) (n= 5). Addition of t-ACPD (5 μm) in combination with the mGluR antagonist MCPG (500 μm) partially blocked the effects of t-ACPD (n= 5). Addition of MCPG alone (500 μm) had no effect (n= 5). The selective mGluR2/3 receptor antagonist MSPG (100 μm) did not have an effect by itself and it was ineffective in blocking the action of t-ACPD.

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We next examined the action of dopamine on our readout of corticostriatal synaptic function. Dopamine (20 μm) caused a reduction in EPSC amplitude (P < 0.02), but did not cause a significant change in the PPR at 50 and 500 ms (n= 5) (Fig. 5). The selective D2-dopamine receptor antagonist l-sulpiride (20 μm) only partially blocked the dopamine-mediated reduction in the EPSC (n= 5). Interestingly, addition of l-sulpiride (20 μm) alone caused a small but insignificant reduction in EPSC amplitude and it increased the PPR at the 50 ms ISI (P < 0.05, paired t test, n= 5); a similar trend was seen at 400 ms (Fig. 4). This latter experiment was repeated with another D2 receptor antagonist, raclopride (3 μm), and the identical trend was observed (n= 6) (Fig. 5). Raclopride (3 μm) reduced the evoked EPSC amplitude (P < 0.05, paired t test) and it increased the PPR at 50 ms (P < 0.02) and 500 ms (P < 0.05, paired t test, n= 6). We also tested the effect of raclopride block of D2 receptors on sEPSCs. Raclopride did not affect sEPSC amplitude and on average, it did not affect sEPSC frequency. Average sEPSC amplitude was 11 ± 0.86 pA and sEPSC frequency was 2.9 ± 0.6 events s−1 before raclopride and 11.4 ± 1.4 pA and 2.5 ± 0.6 events s−1 after raclopride (3 μm) (n= 5). Individually, however, sEPSC frequency was decreased in three cells, increased in one cell and no change was seen in one other cell (n= 5) (Fig. 5B).

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Figure 5. D2 dopamine receptor contribution to the paired-pulse plasticity created by activation of corticostriatal synapses in vitro A, bar graphs represent the percentage change in EPSC amplitude and PPR for 50 and 500 ms ISIs created by dopamine-related treatments. Control values for EPSC amplitude and PPR were obtained in each cell before and after each treatment and plotted as percentage of control (each treatment referenced to its own control). Dopamine (20 μm) reduced the EPSC amplitude (n= 5). Addition of the D2 receptor antagonist sulpiride in combination with dopamine did not fully block the effect dopamine had on EPSC amplitude (n= 5). Addition of sulpiride alone caused an increase in the 50 ms ISI PPR (P < 0.05, n= 5). A similar trend was seen with addition of the selective D2 receptor antagonist raclopride (3 μm), which reduced EPSC amplitude and increased the PPR at 50 and 500 ms (P < 0.05, paired t tests; n= 6). B, cumulative frequency histograms for sEPSC frequency. Ba, example of a raclopride (3 μm) induced increase in sEPSC frequency. Raclopride changed the average sEPSC frequency from 1.53 to 2.55 events s−1. Bb, example of raclopride induced decrease in sEPSC frequency. Raclopride changed the average sEPSC frequency from 3.97 to 1.68 events s−1. Bc, average cumulative histograms for sEPSC frequency before and after addition of raclopride (n= 5).

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While our receptor antagonists did not support a role for the above modulators, the long-lasting nature of the paired-pulse depression suggested it might still be mediated by G-protein activation. To investigate this possibility we examined the effect of G-protein inhibition by NEM (200 μm). Much like previously reported in the striatum by Tang & Lovinger (2000), we found the G-protein inhibition by NEM increased EPSC amplitudes (P < 0.04) and reduced the PPR at 50 ms (P < 0.004) and 500 ms (P < 0.02, paired t test, n= 4) (Fig. 6A and B). We also found the dramatic decrease in EPSC amplitude and increase in 50 ms ISI created by adenosine (see Fig. 2) was blocked by pre-treatment of the slice with NEM (Fig. 6). The adenosine (50 μm) + NEM (200 μm) bar graphs were generated by comparing the response in adenosine + NEM with prior responses obtained in NEM alone.

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Figure 6. G-protein activation modulates corticostriatal paired-pulse plasticity A, the G-protein activator NEM was added to brain slices. Representative current traces are shown for NEM-mediated increases in EPSC amplitude and decreases in the paired-pulse ratio (PPR) measured at 50 and 500 ms ISIs. B, NEM (200 μm) increased EPSC amplitude (P < 0.03) and reduced the PPR at 50 ms (P < 0.003) and 500 ms (P < 0.012) (n= 4). Pretreatment of slices with NEM was effective in blocking the change in EPSC created by adenosine. The effect of adenosine alone is shown for comparison. The adenosine + NEM responses are compared with those responses obtained first in NEM alone (adenosine + NEM/NEM).

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AMPA receptor desensitization

Corticostriatal EPSCs are mediated almost entirely by the activation of AMPA receptors when cells are examined at holding potentials of −70 mV (Akopian & Walsh, 2002). AMPA receptors activated by glutamate undergo desensitization. This property could contribute to paired-pulse depression if glutamate clearance is incomplete before the second release of neurotransmitter. To test this hypothesis we blocked glutamate uptake with PDC (100 μm) (Hashimoto & Kano, 1998). PDC caused a reduction in the amplitude of the corticostriatal EPSC, possibly through the process of AMPA receptor desensitization (P < 0.0001, paired t test, n= 8). It affected the first and second EPSC of each pair equally and thus did not affect the PPR (Fig. 7B). Another method of addressing AMPA receptor desensitization is to reduce desensitization with cyclothiazide (Yamada & Tang, 1993). Cyclothiazide increased the duration of corticostriatal EPSCs, and it caused a significant increase in the PPR at 500 ms (n= 6, paired t test, P < 0.04). No change in PPR was seen at 50 ms (Fig. 7A and B).

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Figure 7. Glutamate receptor desensitization contributes to paired-pulse depression at corticostriatal synapses A, representative current traces before and after addition of the AMPA receptor desensitization reducing compound cyclothiazide (100 μm). Cyclothiazide increased corticostriatal EPSC duration, which is consistent with the ability of cyclothiazide to remove desensitization of glutamate receptors. Cyclothiazide also reduced the paired-pulse depression seen at an ISI of 500 ms (P < 0.05, n= 6). The second EPSC evoked at an ISI of 50 ms started from an elevated baseline due to the removal of desensitization during the first response of the pair. Control responses are superimposed for comparison. B, the glutamate uptake inhibitor PDC (100 μm) reduced the EPSC amplitude (P < 0.03), but it did not affect the PPR at 50 ms and 500 ms (n= 8).

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Evidence supporting a presynaptic component to long-lasting paired pulse depression

The low affinity glutamate receptor antagonist PDA is displaced more readily from AMPA receptors by higher concentrations of glutamate. Conversely, when glutamate is lower, PDA is more effective at blocking AMPA receptors (Hashimoto & Kano, 1998). We added PDA (1 mm) to our brain slices and found it reduced the amplitude of corticostriatal EPSCs to 45.30 ± 4.96% of control (P < 0.02, paired t test, n= 5). PDA had no effect on the PPR measured at an ISI of 50 ms. The control 50 ms PPR was 99.85 ± 12.6% and after PDA the 50 ms PPR was 97.96 ± 8.55%. PDA did reduce the PPR at the 500 ms ISI from 73.91 ± 5.77% to 62.76 ± 6.063% (P < 0.004, paired t test, n= 5) (Fig. 8). By contrast, application of the high-affinity competitive AMPA receptor antagonist CNQX (1 μm) blocked the first and second EPSC of the pair equivalently and there was no change in the PPR (n= 5). CNQX (1 μm) reduced the EPSC amplitude to 11.5 ± 3.5% (n= 5) of control.

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Figure 8. Increased block of the 500 ms ISI paired EPSP by the low-affinity AMPA receptor antagonist PDA A, bath application of 1 mm PDA decreased EPSC amplitude to 45.30 ± 4.96% (P < 0.02, n= 5) and the PPR evoked at an ISI of 500 ms from 73.91 ± 5.77% to 62.76 ± 6.06% (P < 0.003, n= 5). B, representative EPSCs evoked with the 500 ms ISI pairing before (a) and after (b) bath application of PDA. The first EPSC amplitudes are normalized in c to illustrate the increased block of the second EPSC by PDA. APV (50 μm) was included to eliminate NMDA receptor activation, since PDA is a weak NMDA agonist. The identical experiment performed with the high-affinity antagonist CNQX (1 μm) blocked the first and second EPSC of the pair equivalently and no change in paired-pulse plasticity was observed.

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Ca2+ sensitivity of corticostriatal paired-pulse plasticity

To further examine the presynaptic role in determining corticostriatal paired-pulse plasticity we altered Ca2+ availability in corticostriatal terminals. The Ca2+ sensitivity of paired-pulse facilitation and depression were examined by exposing corticostriatal synapses to a low Ca2+–high Mg2+ external solution, BAPTA-AM or EGTA-AM (Fig. 9A and B). All experiments were performed using a ‘paired statistical design’ where control and post-Ca2+ treatment measures were obtained sequentially from the same set of synapses. ISIs of 50 and 500 ms were tested to represent paired-pulse facilitation and depression, respectively. Changing the extracellular solution from 2.4 mm Ca2+–1.3 mm Mg2+ to 0.5 mm Ca2+–3.2 Mg2+ reduced the corticostriatal EPSC to 33.9% of control (P < 0.015; paired t test, n= 5) and the PPR at an ISI of 50 ms increased from 92.6 ± 7.7% to 160 ± 10.1% (P < 0.015; paired t test, n= 5). By contrast, the paired-pulse plasticity evoked at an ISI of 500 ms was less sensitive to the change in extracellular Ca2+ with the PPR going from 80.1 ± 4.1% to 105.3 ± 10.1% (P= 0.076, paired t test, n= 5) (Fig. 9B). BAPTA-AM (50 μm) reduced the EPSC amplitude to 32.4% of control (P < 0.00001, paired t test, n= 8) and the PPR measured at an ISI of 50 ms increased from 89.3 to 116.4% (P < 0.015, paired t test, n= 8) and the PPR measured at an ISI increased from 74 to 104% (P < 0.03, paired t test, n= 8). EGTA-AM (50 μm) reduced the amplitude of corticostriatal EPSCs to 77% of control (P < 0.04, paired t test, n= 8), but it did not cause a significant difference in the PPR measured at 50 ms where the PPR changed from 80 ± 3% to 88 ± 4% (P= 0.09, paired t test, n= 8). By contrast, EGTA-AM produced the opposite effect from that seen with low Ca2+ or BAPTA when it reduced the PPR measured at 500 ms from 78.1% to 61% (P < 0.002, paired t test, n= 8) (Fig. 9B).

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Figure 9. Ca2+ differentially regulates paired-pulse facilitation and depression The Ca2+ sensitivity of paired-pulse facilitation and depression were examined by exposing corticostriatal synapses to a low Ca2+–high Mg2+ external solution, BAPTA-AM or EGTA-AM. All experiments were performed using a ‘paired statistical design’ where control and post-Ca2+ treatment measures were obtained sequentially from the same set of synapses. ISIs of 50 and 500 ms were tested to represent paired-pulse facilitation and depression, respectively. A, representative current traces for changes in corticostriatal paired-pulse plasticity occurring in response to changing the extracellular solution from 2.4 mm Ca2+–1.3 mm Mg2+ to 0.5 mm Ca2+–3.2 Mg2+ (maintains osmolarity), application of BAPTA-AM (50 μm), and application of EGTA-AM (50 μm). B, EPSC amplitude and paired-pulse plasticity measured after each Ca2+ manipulation are ‘normalized’ to the value measured prior to perfusion with the experimental solution. The low-Ca2+ solution reduced the corticostriatal EPSC to 33.9% of control (P < 0.015; paired t test, n= 5) and the PPR at an ISI of 50 ms increased from 92.6 ± 7.7% to 160 ± 10.1% (P < 0.015; paired t test, n= 5). By contrast, the paired-pulse plasticity evoked at an ISI of 500 ms was less sensitive to the change in extracellular Ca2+ with the PPR going from 80.1 ± 4.1% to 105.3 ± 10.1% (P= 0.076, paired t test, n= 5). BAPTA-AM (50 μm) reduced the EPSC amplitude to 32.4% of control (P < 0.00001, paired t test, n= 8) and the PPR measured at an ISI of 50 ms increased from 89.3% to 116.4% (P < 0.015, paired t test, n= 8) and the PPR measured at an ISI increased from 74% to 104% (P < 0.03, paired t test, n= 8). EGTA-AM (50 μm) reduced the amplitude of corticostriatal EPSCs to 77% of control (P < 0.04, paired t test, n= 8), but it did not cause a significant difference in the PPR measured at 50 ms where the PPR changed from 80 ± 3% to 88 ± 4% (P= 0.09, paired t test, n= 8). By contrast, EGTA-AM reduced the PPR measured at 500 ms from 78.1% to 61% (P < 0.002, paired t test, n= 8). The effect of EGTA at the 500 ms ISI was completely opposite to the increase in PPR produced by either BAPTA or low extracellular Ca2+.

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Discussion

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix

Corticostriatal synaptic plasticity varies with stimulation protocols, the anatomical site of activation and pharmacological manipulation of brain slices (Calabresi et al. 1992; Lovinger & Choi, 1995; Choi & Lovinger, 1997; Partridge et al. 2000; Smith et al. 2001). We now show that a simple form of short-term synaptic plasticity, paired-pulse plasticity, is determined at corticostriatal synapses by the relative contributions made by the two independent processes of synaptic facilitation and depression. We found the expression of facilitation was quite variable, possibly reflecting the relative contributions made by different types of excitatory synapses found in the striatum (Mori et al. 1994a,b). By contrast, longer-lasting paired-pulse synaptic depression was consistent and it blended with facilitation to determine paired-pulse plasticity. Exogenous application of modulators demonstrated their potential for influencing both facilitation and depression. However, experiments using selective receptor antagonists indicated that the plasticity produced by paired stimulation of the corpus callosum in vitro is intrinsic to the activated corticostriatal terminals. Ca2+ was shown to play an important and differential role in governing the expression of both facilitation and depression at these synapses.

Modulation of presynaptic terminals

The striatum receives excitatory glutamatergic input via corticostriatal afferents, which retain much of the anatomical organization found in the cortex (McGeorge & Faull, 1987; Deniau et al. 1996). Potential modulation of corticostriatal synapses also comes from dopaminergic input from the substantia nigra, seratoninergic input from the raphe nucleus and intrinsic GABAergic and cholinergic interneurons (Graybiel, 1986; Soghomonian et al. 1987; Jimenez-Castellanos & Graybiel, 1989; Vertes, 1991; Tepper et al. 2004; for review see van Domburg & ten Donkelaar, 1991). There is also evidence for modulation by adenosine and cannabinoids in the striatum (Lovinger & Choi, 1995; Gerdeman & Lovinger, 2001; Ronesi & Lovinger, 2005; for review see Tepper et al. 2004). The long time-course of paired-pulse depression we report for corticostriatal synapses is in line with the long-lasting action of many modulators that act through a G-protein mechanism (Fig. 1) (Bannister et al. 2002; for review see Nicoll, 2004). Our method of stimulating the corpus callosum to activate corticostriatal afferents in vitro runs the risk of current spreading from the stimulating electrodes to activate other afferent pathways, which could influence corticostriatal synapses. Alternatively, activation of corticostriatal fibres may trigger a feed-forward polysynaptic form of modulation. To rule out these possibilities we screened a number of candidate modulators including dopamine, adenosine, acetylcholine, the modulatory actions of glutamate through mGluRs and GABA through GABAB receptors. In agreement with previous studies, we found these modulators or neurotransmitters reduced the amplitude of corticostriatal EPSCs and this change in amplitude was associated with increases in the paired-pulse ratio at both 50 and 500 ms ISIs (Lovinger & Choi, 1995; Lovinger & McCool, 1995; Weiler et al. 1984; Villar & Walsh, 1999).

The only exception was dopamine. However, our results did not produce a clear picture for dopamine's role in modulating the function of corticostriatal terminals. Prior work has shown dopamine to reduce the release of glutamate from corticostriatal terminals via a D2 receptor-mediated mechanism (Bamford et al. 2004). We found dopamine reduced the amplitude of the EPSC, but did not affect the PPR at ISIs of 50 or 500 ms. In addition, the D2 receptor antagonist l-sulpiride did not fully block the dopamine-mediated reduction in the evoked EPSC. Application of the selective D2 receptor antagonists l-sulpiride and raclopride alone also produced effects counter to the hypothesis of dopamine acting presynaptically to reduce release. Raclopride reduced the amplitude of evoked EPSCs and, as predicted for a presynaptic mechanism, it increased the PPR at 50 and 500 ms. A similar trend was observed with l-sulpiride; however, only the increase in PPR at 50 ms was significant (Fig. 5). Analysis of raclopride modulation of sEPSC frequency and amplitude did not help to clarify the effect. Raclopride reduced sEPSC frequency in three cells, increased sEPSC frequency in one cell and had no impact in another. On average, raclopride did not impact sEPSC frequency (Fig. 5). The variability in the dopamine and dopamine antagonist outcomes we observed is not new. Previous studies have shown only select populations of corticostriatal terminals respond to D2 receptor pharmacology (Flores-Hernandez et al. 1997; Bamford et al. 2004). Indeed, ultrastructural analysis suggests only 9% of corticostriatal terminals possess D2 receptors (Wang & Pickel, 2002). A more consistent presynaptic effect for D2 receptors is seen for D2 autoreceptors found on dopaminergic nigrostriatal terminals, where block of D2 receptors causes a dramatic increase in dopamine overflow (Wu et al. 2002). The increase in extrasynaptic dopamine created by block of autoreceptors may also influence corticostriatal synaptic terminal function, even in the presence of D2 antagonists.

Responses to adenosine and baclofen were blocked in large part by their appropriate receptor antagonist and application of receptor-specific antagonists alone did not alter long-lasting paired-pulse depression, indicating these extrinsic modulators (adenosine and GABA) are not involved in the long-lasting paired-pulse depression examined in this study. The results were not as clear for mGluRs, where both the non-specific mGluR antagonist MCPG and the selective mGluR2/3 antagonist MSPG had marginal effects on the response to the mGluR agonist t-ACPD. MCPG or MSPG alone did not alter corticostriatal paired-pulse plasticity.

We also examined the effect of G-protein inhibition on corticostriatal neurotransmission by applying the non-selective G-protein inhibitor NEM. As reported previously by Tang & Lovinger (2000), we found that NEM increased the amplitude of corticostriatal EPSCs, while reducing the paired-pulse ratio (Fig. 6). NEM was also effective in blocking the response to adenosine (Fig. 6). These data suggest NEM is inhibiting G-proteins, since other studies have shown that adenosine acts on presynaptic terminals to reduce neurotransmitter release through a G-protein mechanism (Gubitz et al. 1996; for review see Dunwiddie & Masino, 2001). The change in EPSC amplitude and PPR created by NEM alone suggests that a background level of G-protein activity could limit neurotransmitter release at corticostriatal synapses in vitro, as suggested by Tang & Lovinger (2000).

NEM could be acting via other mechanisms as well, since it is a strong alkalizing agent that targets -SH groups found in many proteins (Maruhashi et al. 1984; Brimecombe et al. 1999). NEM is also an inhibitor of the ATPase NSF and this action increases the readily releasable pool of neurotransmitter (Lonart & Südhof, 2000). NEM inhibition of NSF could thus increase the response evoked by the first action potential of the pair by increasing the number of core complexes used during vesicle fusion, which could also contribute to temporary synaptic depression (Lonart & Südhof, 2000).

Postsynaptic mechanism for paired-pulse depression

Another important use-dependent process shown to impact synaptic communication at glutamatergic synapses is synaptic depression resulting from receptor desensitization. We used the 500 ms ISI to study synaptic depression because shorter duration ISIs recruited the process of facilitation, which masked the underlying process of depression. We found that block of glutamate uptake with PDC caused a reduction in the amplitude of the corticostriatal EPSC, possibly via AMPA receptor desensitization or increased AMPA receptor occupancy. However, much like Hashimoto & Kano (1998) found for climbing fibre synapses, we found PDC affected the first and second EPSC of each 500 ms pair equally. The PDC finding is in contrast to the reduction in paired-pulse depression created by cyclothiazide at the 500 ms ISI (Fig. 7). Cyclothiazide has been shown to be effective at reducing AMPA receptor desensitization at other synapses (Yamada & Tang, 1993), and our results indicate that AMPA receptor desensitization may contribute to the paired-pulse depression seen at longer pairing intervals. Xu-Friedman & Regehr (2003) demonstrated that AMPA receptor desensitization has a time constant of recovery of 30 ms at climbing fibre synapses. Our cyclothiazide results indicate that, while the process of desensitization may recover fairly rapidly, it is still active enough to affect responses evoked at the pairing interval of 500 ms. Cyclothiazide has also been shown to reduce synaptic depression at the calyx of Held by blocking a Ca2+-sensitive process, but this action operates at ISIs < 150 ms and should not be active at the 500 ms pairing interval we used to study depression (Bellingham & Walmsley, 1999). Cyclothiazide also slows AMPA receptor deactivation, but slowing deactivation should reduce the second EPSP of the pair via the combined action of electrotonic shunting and a reduction of the net current generated by binding of glutamate to partially active AMPA receptors (Rammes et al. 1998). Cyclothiazide produced the opposite effect at corticostriatal synapses.

Another postsynaptic glutamate receptor-based mechanism, which could contribute to long-lasting paired pulse depression at CNS synapses, is receptor occupancy (Harrison & Jahr, 2003). Kinney et al. (1997) found that the granule cell to brush cell synapse in the cerebellum displayed a slower component of paired-pulse depression, with a time constant of 800 ms that was associated with the slow removal of glutamate from the synaptic cleft.

Presynaptic mechanisms controlling paired-pulse plasticity

Use-dependent short-term plasticity expressed at corticostriatal synapses reflects a blend of facilitation and depression. A large body of evidence supports changes in the release properties by presynaptic terminals as a mechanism to explain short-term facilitation and depression (for review see Zucker & Regehr, 2002). We found that the low-affinity glutamate receptor antagonist PDA was more effective at blocking the second depressed EPSC evoked with long-duration ISIs, which supports a presynaptic mechanism of reduced glutamate release in response to long-duration pairing intervals (Fig. 8A and B). Hashimoto & Kano (1999) reported identical results for the climbing fibre to Purkinje cell synapse. Corticostriatal synapses express variable facilitation, acting at short ISIs and long-lasting paired-pulse depression. Thus, little paired-pulse plasticity is seen on average at short ISIs and predictably no pairing difference was observed after PDA at these short ISIs. By contrast, the high-affinity AMPA receptor antagonist CNQX was more effective at blocking the EPSC and it did not alter the 500 ms ISI PPR seen previously in aCSF alone.

We further examined the presynaptic nature of corticostriatal paired-pulse plasticity by varying Ca2+ availability through addition of a low Ca2+ solution or by exposing brain slices to the AM-esters of BAPTA or EGTA. Lowering the Ca2+/Mg2+ ratio of the external solution is a common method employed at many synapses to investigate the relationship between Ca2+ influx and presynaptic forms of synaptic plasticity (Hashimoto & Kano, 1999; Kreitzer & Regehr, 2000; Scheuss et al. 2002). Low Ca2+–high Mg2+ solutions reduced the size of corticostriatal EPSCs and it increased the PPR at the 50 ms ISI (Fig. 9). The same result was obtained when synapses were exposed to BAPTA-AM. BAPTA-AM has rapid binding kinetics, which enables buffering of rapid and large Ca2+ transients. EGTA-AM, which has much slower binding kinetics, was far less effective in reducing the amplitude of corticostriatal EPSCs and it did not affect paired-pulse plasticity at the short-duration sampling ISI of 50 ms. The increase in paired-pulse potentiation created by reducing Ca2+ transients indicates that closely paired action potentials enhance the Ca2+ signal seen by vesicles during the second action potential to create facilitation when vesicle depletion is not a limiting factor.

Many presynaptic Ca2+-related mechanisms are proposed to explain the expression of short-term synaptic facilitation, including differential Ca2+ channel expression, a change in Ca2+ availability, and differential contribution from Ca2+ binding proteins (Ali & Nelson, 2006; for review see Katz & Miledi, 1968; Zucker & Regehr, 2002; Burnashev & Rozov, 2005). Felmy et al. (2003) demonstrated that Ca2+ remaining from prior activity sums with new Ca2+ to enhance release, but that this mechanism only accounts for 30% of observed facilitation. They speculated that saturation of cytosolic buffers during the first action potential of a pair would be sufficient to explain the remaining degree of facilitation (Felmy et al. 2003). Indeed, the mobile Ca2+ buffer calbindin-D28k has been shown to play a clear role in mediating facilitation at inhibitory cortical synapses (Blatow et al. 2003). However, neocortical pyramidal cell synapses onto cortical interneurons express either facilitation or depression, which appears to be dependent on residual Ca2+ and not influenced by endogenous buffers (Rozov et al. 2001). Atluri & Regehr (1996) proposed the existence of a high-affinity Ca2+ binding site that retains Ca2+ when a second action potential arrives and interacts with the vesicle sensor to enhance release. Neuronal calcium sensor 1 (NCS-1) is a candidate molecule (Sippy et al. 2003), but as pointed out by Zucker (2003), NCS-1 could be acting either as a high-affinity Ca2+ sensor or as a saturable buffer to create facilitation. We did not examine paired-pulse plasticity at ISIs shorter than 50 ms, but analysis and modelling of neonatal CA3–CA1 synapses indicates variation in vesicle release probability can also influence paired-pulse plasticity creating either depression or facilitation at the short ISI of 20 ms (Hanse & Gustafsson, 2001).

We next examined the Ca2+ dependence of long-lasting paired-pulse depression at corticostriatal synapses. We found EGTA-AM slowed the recovery from paired-pulse depression, indicating it is Ca2+ dependent. One mechanism to explain this EGTA effect is that it keeps Ca2+ from speeding the recovery of Ca2+ channels from inactivation (Lee et al. 1999, 2000). However, similar experiments performed at the calyx of Held did not reveal significant Ca2+ current inactivation even when activated at a frequency of 100 Hz (Wang & Kaczmarek, 1998). Further analysis of these large synapses revealed that synaptic depression develops from depletion of vesicles and that the delivery of new vesicles is Ca2+ dependent (Wang & Kaczmarek, 1998). This mechanism could explain the EGTA-mediated enhancement of paired-pulse depression we observed at corticostriatal synapses. It has been suggested that the time course of refilling the releasable pool reflects the kinetics of two separate pools of vesicles, expressing slow and rapid recovery, and that the delivery of vesicles in the faster pool requires the accumulation of Ca2+ and its binding to calmodulin (Wu & Borst, 1999; Sakaba & Neher, 2001). Dittman & Regehr (1998) also found EGTA-AM (100 μm) slowed the recovery from long-lasting paired pulse depression and, much like Wang & Kaczmarek (1998), they concluded that the mechanism was related to the role played by Ca2+ in facilitating the delivery of vesicles to the readily releasable pool of vesicles. The calyx synapses in the auditory brainstem also show synaptic depression from depletion of Ca2+ in the synaptic cleft, but this mechanism has a time constant of 60 ms, which is much faster then the depression we observed. Furthermore, this mechanism operates when the postsynaptic cell experiences a large sustained depolarization, which did not occur in this study (Borst & Sakmann, 1999).

Conclusion

The underpinnings of short-term corticostriatal synaptic plasticity are an intrinsic interaction between facilitation and depression. This template varies considerably between synapses, with possible functional links between striatal anatomy and extrinsic modulatory influences that may influence presynaptic function (Spencer & Murphy, 2000; Partridge et al. 2000; Smith et al. 2001; Tang et al. 2001). The paired-pulse depression observed at corticostriatal synapses is clearly defined by its consistency and its slow time-course of recovery. Paired-pulse depression showing similar slow kinetics of recovery has been reported at many synapses throughout the CNS, suggesting that this form of plasticity may serve as a backbone for patterning of synaptic efficacy at CNS synapses (Dittman & Regehr, 1998; Wu & Borst, 1999; Sakaba & Neher, 2001; Fitzpatrick et al. 2001; Cragg, 2003). Facilitation is also intrinsic to activated corticostriatal synapses, but its expression is quite variable. Together these two opposing forms of short-term plasticity mould striatal responsiveness to cortical drive.

References

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix
  • Aghajanian GK & Rasmussen K (1989). Intracellular studies in the fa facial nucleus illustratiny a simple new method for obtaining viable motoneurons in adult rat brain slices. Synapse 3, 331338.
  • Akopian G, Musleh W, Smith R & Walsh JP (2000). Functional state of presynaptic terminals influences the expression of short- and long-term plasticity at corticostriatal synapses. Synapse 38, 271280.
  • Akopian G & Walsh JP (2002). Corticostriatal paired-pulse potentiation produced by voltage-dependent increases in NMDA receptor and L-type Ca2+ channel activation. J Neurophysiol 87, 157165.
  • Akopian G & Walsh JP (2006). Reduced expression of short- and long-term facilitation at aged corticostriatal synapses. Synapse 60, 223238.
  • Ali AB & Nelson C (2006). Distinct Ca2+ channels mediate transmitter release at excitatory synapses displaying different dynamic properties in rat neocortex. Cerebral Cortex 16, 386393.
  • Atluri PP & Regehr WG (1996). Determinants of the time course of facilitation at the granule cell to Purkinje cell synapse. J Neurosci 16, 56615671.
  • Bamford NS, Zhang H, Schmitz Y, Wu NP, Cepeda C, Levine MS, Schmauss C, Zakharenko SS, Zablow L & Sulzer D (2004). Heterosynaptic dopamine neurotransmission selects sets of corticostriatal terminals. Neuron 42, 65366563.
  • Bannister RA, Melliti K & Adams BA (2002). Reconstituted slow muscarinic inhibition of neuronal (Ca(v)1.2c), L-type Ca2+ channels. Biophys J 83, 32563267.
  • Beckstead RM, Domesick VB & Nauta WJ (1979). Efferent connections of the substantia nigra and ventral tegmental area in the rat. Brain Res 175, 191217.
  • Bellingham MC & Walmsley B (1999). A novel presynaptic inhibitory mechanism underlies paired pulse depression at a fast central synapse. Neuron 23, 159170.
  • Bennett BD & Bolam JP (1994). Synaptic input and output of parvalbumin-immunoreactive neurons in the neostriatum of the rat. Neurosci 62, 707719.
  • Blatow M, Caputi A, Burnashev N, Monyer H & Rozov A (2003). Ca2+ buffer saturation underlies paired pulse facilitation in calbindin-D28k-containing terminals. Neuron 38, 7988.
  • Borst JG & Sakmann B (1999). Depletion of calcium in the synaptic cleft of a calyx-type synapse in the rat brainstem. J Physiol 521, 123133.
  • Brimecombe JC, Potthoff WK & Aizenman E (1999). A critical role of the N-methyl-D-aspartate (NMDA) receptor subunit (NR) 2A in the expression of redox sensitivity of NR1/NR2A recombinant NMDA receptors. J Pharmacol Exp Ther 291, 785792.
  • Burnashev N & Rozov A (2005). Presynaptic Ca2+ dynamics, Ca2+ buffers and synaptic efficacy. Cell Calcium 37, 489495.
  • Calabresi P, Maj R, Pisani A, Mercuri NB & Bernardi G (1992). Long-term synaptic depression in the striatum: Physiological and pharmacological characterization. J Neurosci 12, 42244233.
  • Calabresi P, Pisani A, Mercuri NB & Bernardi G (1996). The corticostriatal projection: From synaptic plasticity to basal ganglia disorders. Trends Neurosci 1, 1924.
  • Canales JJ, Capper-Loup C, Hu D, Choe ES, Upadhyay U & Graybiel AM (2002). Shifts in striatal responsivity evoked by chronic stimulation of dopamine and glutamate systems. Brain 125, 23532363.
  • Chang HT & Kita H (1992). Interneurons in the rat striatum: relationships between parvalbumin neurons and cholinergic neurons. Brain Res 574, 307311.
  • Choi S & Lovinger DM (1997). Decreased probability of neurotransmitter release underlies striatal long-term depression and postnatal development of corticostriatal synapses. Proc Natl Acad Sci U S A 94, 26652670.
  • Cragg SJ (2003). Variable dopamine release probability and short-term plasticity between functional domains of the primate striatum. J Neurosci 23, 43784385.
  • Deniau JM, Menetrey A & Charpier S (1996). The lamellar organization of the rat substantia nigra pars reticulata: Segregated pattterns of striatal afferents and relationship to the topography of corticostriatal projections. Neurosci 73, 761781.
  • Deschenes M, Bourassa J & Parent A (1995). Two different types of thalamic fibers innervate the rat striatum. Brain Res 701, 288292.
  • Dittman JS, Kreitzer AC & Regehr WG (2000). Interplay between facilitation, depression, and residual calcium at three presynaptic terminals. J Neurosci 20, 13741385.
  • Dittman JS & Regehr WG (1998). Calcium dependence and recovery kinetics of presynaptic depression at the climbing fiber to purkinje cell synapse. J Neurosci 18, 61476162.
  • Dunwiddie TV & Masino SA (2001). The role and regulation of adenosine in the central nervous system. Ann Rev Neurosci 24, 3155.
  • Felmy F, Neher E & Schneggenburger R (2003). Probing the intracellular calcium sensitivity of transmitter release during synaptic facilitation. Neuron 37, 801811.
  • Fitzpatrick JS, Akopian G & Walsh JP (2001). Short-term plasticity at inhibitory synapses in rat striatum and its effects on striatal output. J Neurophysiol 85, 20882099.
  • Flores-Hernandez J, Galarraga E & Bargas J (1997). Dopamine selects glutamatergic inputs to neostriatal neurons. Synapse 25, 185195.
  • Gerdeman G & Lovinger DM (2001). CB1 cannabinoid receptor inhibits synaptic release of glutamate in rat dorsolateral striatum. J Neurophysiol 85, 468471.
  • Grace AA (1995). The tonic/phasic model of dopamine system regulation: its relevance for understanding how stimulant abuse can alter basal ganglia function. Drug Alcohol Dependence 37, 111129.
  • Graybiel AM, Baughman RW & Eckenstein F (1986). Cholinergic neuropil of the striatum observes striosomal boundaries. Nature 323, 625627.
  • Graybiel AM & Chesselet MF (1984). Compartmental distribution of striatal cell bodies expressing [Met]enkephalin-like immunoreactivity. Proc Natl Acad Sci U S A 81, 79807984.
  • Gubitz AK, Widdowson L, Kurokawa M, Kirkpatrick KA & Richardson PJ (1996). Dual signalling by the adenosine A2a receptor involves activation of both N- and P-type calcium channels by different G proteins and protein kinases in the same striatal nerve terminals. J Neurochem 67, 374381.
  • Hanse E & Gustafsson B (2001). Paired-pulse plasticity at the single release site level: an experimental and computational study. J Neurosci 21, 83628369.
  • Harrison J & Jahr CE (2003). Receptor occupancy limits synaptic depression at climbing fiber synapses. J Neurosci 23, 377383.
  • Hashimoto K & Kano M (1998). Presynaptic origin of paired-pulse depression at climbing fibre-Purkinje cell synapses in the rat cerebellum. J Physiol 506, 391405.
  • Jimenez-Castellanos J & Graybiel AM (1989). Compartmental origins of striatal efferent projections in the cat. Neurosci 32, 297321.
  • Katz B & Miledi R (1968). The role of calcium in neuromuscular facilitation. J Physiol 195, 481492.
  • Kawaguchi Y, Wilson CJ, Augood SJ & Emson PC (1995). Striatal interneurones: chemical, physiological and morphological characterization. Trends Neurosci 18, 527535.
  • Kawaguchi Y, Wilson CJ & Emson PC (1989). Intracellular recording of identified neostriatal patch and matrix spiny cells in a slice preparation preserving cortical inputs. J Neurophysio 62, 10521068.
  • Kinney GA, Overstreet LS & Slater NT (1997). Prolonged physiological entrapment of glutamate in the synaptic cleft of cerebellar unipolar brush cells. J Neurophysiol 78, 13201333.
  • Kolb B, Gibb R & Van Der Kooy D (1992). Cortical and striatal structure and connectivity are altered by neonatal hemidecortication in rats. J Comp Neurol 322, 311324.
  • Kreitzer AC & Regehr WG (2000). Modulation of transmission during trains at a cerebellar synapse. J Neurosci 20, 13481357.
  • Lee A, Scheuer T & Catterall WA (2000). Ca2+/calmodulin-dependent facilitation and inactivation of P/Q-type Ca2+ channels. J Neurosci 20, 68306838.
  • Lee A, Wong ST, Gallagher D, Li B, Storm DR, Scheuer T & Catterall WA (1999). Ca2+/calmodulin binds to and modulates P/Q-type calcium channels. Nature 399, 155159.
  • Lonart G & Südhof TC (2000). Assembly of SNARE core complexes prior to neurotransmitter release sets the readily releasable pool of synaptic vesicles. J Biol Chem 275, 2770327707.
  • Lovinger DM & Choi S (1995). Activation of adenosine A1 receptors initiates short-term synaptic depression in rat striatum. Neurosci Let 199, 912.
  • Lovinger DM & McCool BA (1995). Metabotropic glutamate receptor-mediated presynaptic depression at corticostriatal synapses involves mGLuR2 or 3. J Neurophysiol 73, 10762083.
  • Lucas LR & Harlan RE (1995). Cholinergic regulation of tachykinin- and enkephalin-gene expression in the rat striatum. Mol Brain Res 30, 181195.
  • McGeorge AJ & Faull RL (1987). The organization and collateralization of corticostriate neurones in the motor and sensory cortex of the rat brain. Brain Res 423, 318324.
  • Maruhashi J, Oomura Y, Kato M & Kusano K (1984). Three types of chemical modification-effects induced by various chemical reagents on the glutamate receptors in molluscan neurons. Jap J Physiol 34, 10491064.
  • Mori A, Takahashi T, Miyashita Y & Kasai H (1994a). Two distinct glutamatergic synaptic inputs to striatal medium spiny neurones of neonatal rats and paired-pulse depression. J Physiol 476, 217228.
  • Mori A, Takahashi T, Miyashita Y & Kasai H (1994b). Quantal properties of H-type glutamatergic synaptic input to the striatal medium spiny neurons. Brain Res 654, 177179.
  • Nicoll RA (2004). My close encounter with GABAB receptors. Biochem Pharmacol 68, 16671674.
  • Nisenbaum ES, Grace AA & Berger TW (1992). Functionally distinct subpopulations of striatal neurons are differentially regulated by GABAergic and dopaminergic inputs II. In vitro analysis. Neurosci 48, 579593.
  • Ou X, Buckwalter G, McNeill TH & Walsh JP (1997). Age-related changes in short-term synaptic plasticity intrinsic to excitatory striatal synapses of the rat. Synapse 27, 5768.
  • Partridge JG, Tang K-C & Lovinger DM (2000). Regional and postnatal heterogeneity of activity-dependent long-term changes in synaptic efficacy in the dorsal striatum. J Neurophysiol 84, 14221429.
  • Priestley JV, Somogyi P & Cuello AC (1981). Neurotransmitter-specific projection neurons revealed by combining PAP immunohistochemistry with retrograde transport of HRP. Brain Res 220, 231240.
  • Rammes G, Swandulla D, Spielmanns P & Parsons CG (1998). Interactions of GYKI 52466 and NBQX with cyclothiazide at AMPA receptors: experiments with outside-out patches and EPSCs in hippocampal neurones. Neuropharmacol 37, 12991320.
  • Reiner A, Jiao Y, Del Mar N, Laverghetta AV & Lei WL (2003). Differential morphology of pyramidal tract-type and intratelencephalically projecting-type corticostriatal neurons and their intrastriatal terminals in rats. J Comp Neurol 457, 420440.
  • Ronesi J & Lovinger DM (2005). Induction of striatal long-term synaptic depression by moderate frequency activation of cortical afferents in rat. J Physiol 562, 245256.
  • Rozov A, Burnashev N, Sakmann B & Neher E (2001). Transmitter release modulation by intracellular Ca2+ buffers in facilitating and depressing nerve terminals of pyramidal cells in layer 2/3 of the rat neocortex indicates a target cell-specific difference in presynaptic calcium dynamics. J Physiol 531, 807826.
  • Sakaba T & Neher E (2001). Calmodulin mediates rapid recruitment of fast-releasing synaptic vesicles at a calyx-type synapse. Neuron 32, 11191131.
  • Scheuss V, Schneggenburger R & Neher E (2002). Separation of presynaptic and postsynaptic contributions to depression by covariance analysis of successive EPSCs at the calyx of Held synapse. J Neurosci 22, 728739.
  • Sippy T, Cruz-Martin A, Jeromin A & Schweizer FE (2003). Acute changes in short-term plasticity at synapses with elevated levels of neuronal calcium sensor-1. Nature Neurosci 6, 10311038.
  • Smith R, Musleh W, Akopian G, Buckwalter G & Walsh JP (2001). Regional differences in the expression of corticostriatal synaptic plasticity. Neurosci 106, 95101.
  • Soghomonian JJ, Doucet G & Descarries L (1987). Serotonin innervation in adult rat neostriatum. I. Quantified regional distribution. Brain Res 425, 85100.
  • Spencer JP & Murphy KP (2000). Bi-directional changes in synaptic plasticity induced at corticostriatal synapses in vitro. Exp Brain Res 135, 497503.
  • Tang K-C & Lovinger DM (2000). Role of pertussis toxin-sensitive G-proteins in synaptic transmission and plasticity at corticostriatal synapses. J Neurophysiol 83, 6069.
  • Tang K-C, Low MJ, Grand DK & Lovinger DM (2001). Dopamine-dependent synaptic plasticity in striatum during in vivo development. Proc Natl Acad Sci U S A 98, 12551260.
  • Tepper JM, Koos T & Wilson CJ (2004). GABAergic microcircuits in the neostriatum. Trends Neurosci 27, 662669.
  • Van Domburg PH & Ten Donkelaar HJ (1991). The human substantia nigra and ventral tegmental area. A neuroanatomical study with notes on aging and aging diseases. Adv Anat Embryol Cell Biol 121, 131132.
  • Vertes RP (1991). A PHA-L analysis of ascending projections of the dorsal raphe nucleus in the rat. J Comp Neurol 313, 643668.
  • Villar FAS & Walsh JP (1999). Modulation of long-term synaptic plasticity at corticostriatal synapses. Neurosci 90, 10311041.
  • Wang H & Pickel VM (2002). Dopamine D2 receptors are present in prefrontal cortical afferents and their targets in patches of the rat caudate-putamen nucleus. J Comp Neurol 442, 392404.
  • Wang LY & Kaczmarek LK (1998). High-frequency firing helps replenish the readily releasable pool of synaptic vesicles. Nature 394, 384388.
  • Weiler MH, Misgeld U & Cheong DK (1984). Presynaptic muscarinic modulation of nicotinic excitation in the rat neostriatum. Brain Res 296, 111120.
  • Wu LG & Borst JG (1999). The reduced release probability of releasable vesicles during recovery from short-term synaptic depression. Neuron 23, 821832.
  • Wu Q, Reith ME, Walker QD, Kuhn CM, Carroll FI & Garris PA (2002). Concurrent autoreceptor-mediated control of dopamine release and uptake during neurotransmission: an in vivo voltammetric study. J Neurosci 22, 62726281.
  • Xu-Friedman MA & Regehr WG (2003). Ultrastructural contributions to desensitization at cerebellar mossy fiber to granule cell synapses. J Neurosci 23, 21822192.
  • Yamada KA & Tang C-M (1993). Benzothiazides inhibit rapid glutamate receptor desensitization and enhance glutamatergic synaptic currents. J Neurosci 13, 39043915.
  • Zucker RS (2003). NCS-1 stirs somnolent synapses. Nature Neurosci 6, 10061008.
  • Zucker RS & Regehr WG (2002). Short-term synaptic plasticity. Annu Rev Physiol 64, 355405.

Appendix

  1. Top of page
  2. Abstract
  3. Methods
  4. Results
  5. Discussion
  6. References
  7. Appendix

Acknowledgements

National Institutes of Health Grants AG12679 and AG21937 supported this work.