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. 2004 Oct 28;562(Pt 1):257–269. doi: 10.1113/jphysiol.2004.074211

Low threshold calcium currents in rat cerebellar Purkinje cell dendritic spines are mediated by T-type calcium channels

Philippe Isope 1,2, Timothy H Murphy 1,2,3,4
PMCID: PMC1665489  PMID: 15513942

Abstract

The functional role of low voltage activated (LVA) calcium channels in the cerebellar Purkinje cell dendritic tree is not completely understood. Since the localization of these channels will influence their possible roles in dendritic integration and induction of plasticity, we set out to characterize the LVA calcium current in Purkinje cell dendrites in acute cerebellar slices of young rats. Using a combination of electrophysiological recordings and two-photon laser scanning microscopy, we show that LVA calcium current recorded at the soma can be correlated with voltage-dependent calcium transients in Purkinje cell dendritic spines. Blocking sodium and potassium conductances allowed us to isolate and characterize a fast inactivating inward current activated positive to −55 mV. Activation and steady-state inactivation kinetics, voltage-dependent deactivation kinetics, and pharmacological experiments (using ω-agatoxin-IVA, mibefradil and nickel) show that this current is carried by T-type calcium channels. Furthermore, the LVA calcium transient observed in the dendritic spines of the Purkinje cell is well correlated with the current recorded at the soma, suggesting that T-type calcium channels are the main component of the LVA calcium input in spines. The fast rising phase of the calcium transient in spines and the absence of delay between the onset in the spine and the parent dendrite show that T-type calcium channels are present both in spines and dendrites of the Purkinje cell.

The two main afferents of the cerebellar cortex, the mossy fibres and the climbing fibres, converge on the Purkinje cell: the mossy fibre system via granule cells, the climbing fibre by a strong multisite contact on the Purkinje cell dendritic tree. One Purkinje cell receives 150 000–175 000 synaptic inputs onto dendritic spines from the parallel fibres, the granule cell axons (Harvey & Napper, 1991). In vivo experiments showed that the parallel fibre–Purkinje cell synapse is a major site of information storage in the cerebellum (Ito, 1984; Kim & Thompson, 1997; Hesslow & Yeo, 1998). In order to understand how the cell discriminates, integrates and stores information from these two inputs it is necessary to understand both somatic and dendritic integration of these signals. Dendritic integration will depend on the dendritic morphology, electrotonic properties and the type and distribution of voltage-gated channels (Rall et al. 1992). In the Purkinje cell, immunohistochemical and electrophysiological studies have demonstrated a wide variety of voltage-dependent potassium channels and calcium channels (Chung et al. 2000; Trimmer & Rhodes, 2004), among them low voltage activated (LVA; T-type/Cav3) and high voltage activated calcium channels (HVA; P/Q-type/Cav2.1, N-type/Cav2.2, L-type/Cav1, R-type/Cav2.3). Determination of the exact subunit composition of the LVA calcium channel in the Purkinje cell will require further studies since discrepancies exist between protein and mRNA expression for LVA calcium channel subtypes in the cerebellar molecular layer. For example, only mRNA for α1G was demonstrated in the molecular layer (Talley et al. 1999) but there was strong immunostaining for both α1G and α1I subunits (Yunker et al. 2003).

High voltage activated calcium channels (HVA), mainly the P-type channel, have been implicated in the generation of calcium spikes in the Purkinje dendritic tree after climbing fibre and strong parallel fibre stimulation (Llinas et al. 1989; Mintz et al. 1992; Watanabe et al. 1998), but the exact influence of LVA calcium channels is still controversial. It has been suggested that they might be involved in boosting P-type calcium spike generation (Llinas et al. 1992; Watanabe et al. 1998) or in the specific generation of low threshold calcium spikes (Cavelier et al. 2002). Calcium imaging techniques have shown that stimulation of a bundle of parallel fibres can induce a voltage-dependent calcium current restricted to dendritic spiny branchlets (Eilers et al. 1995) or even to individual spines (Denk et al. 1995; Wang et al. 2000). However, the link with expression of specific calcium channel functional classes in the Purkinje cell dendrites is still unclear.

Accordingly, we characterized and localized LVA calcium channel activity in the fine processes of the Purkinje cell dendritic tree. We used a combination of two-photon laser scanning microscopy and electrophysiology to study calcium input directly at the spiny parallel fibre–Purkinje cell synapses and correlated it with the current recorded at the soma. We show that T-type calcium channels are present both in dendritic spines and the dendritic shaft of the Purkinje cell and are in an ideal position to either boost calcium spiking, or serve as their own subthreshold means of calcium signalling.

Methods

Slice preparation

Wistar rats, 9–14 days old, were anaesthetized using halothane and decapitated. All animals used in this project were cared for in accordance with recommendations of the Canadian Council on Animal Care and the regulations and policies of the University of British Columbia Animal Care Facility and the University Animal Care Committee. The head was chilled over ice while the cerebellum was dissected out. After removal of the lateral hemispheres, the vermis was cooled and sliced in a solution containing (mm): 120 NaCl, 3 KCl, 26 NaHCO3, 1.25 NaH2PO4, 0.8 CaCl2, 4 MgCl2, 20 glucose, 1 kynurenate and 0.1 picrotoxin. Then 200 μm sagittal slices were prepared and kept in bubbled (95% O2–5% CO2) bicarbonate-buffered saline (BBS) solution that contained (mm): 120 NaCl, 3 KCl, 26 NaHCO3, 1.25 NaH2PO4, 2 CaCl2, 1 MgCl2, 20 glucose and 1 kynurenate. After slicing, the slices were maintained at 32°C for 0.5–1 h and then allowed to cool to room temperature.

Recordings

All recordings were done at room temperature (20–25°C). Slices were transferred in a recording chamber and superfused first with the same BBS solution supplemented with picrotoxin (0.1 mm). Cells were visualized with an Olympus microscope (BX50) using a ×60 water immersion objective (0.9 NA, LUMPlanfl) using red light and a CCD camera (XC-ST70, Sony). Whole-cell patch clamp recordings in voltage clamp mode were obtained using an Axopatch 200A and series resistance (Rs) compensation (40–75% of 2–10 MΩ typically, and at least 50% in experiments with solution 2 where Rs was < 5 MΩ, see below). Pipettes (2–3 MΩ resistance when filled with the internal solution) were pulled using thin-wall borosilicate glass capillaries (Warner Instruments). Cells with leak current above 800 pA were discarded. The pipette solution contained (mm): 140 mm CsCl, 5 mm TEACl, 0.5 mm MgCl2, 10 mm HEPES, 4 MgATP, 0.5 Na3GTP, 0.2 mm Oregon green 488 BAPTA 1 (OGB-1; Molecular Probes) pH adjusted to 7.3 with CsOH at 300 mosmol l−1. Recordings were neither corrected for the liquid junction potential, estimated to be 3 mV, nor for the potential drop across uncompensated series resistance, estimated to be below 5 mV. Synaptic currents in Purkinje cells were filtered at 1–5 kHz and digitized at 50 kHz. Acquisition of the data was performed with pCLAMP 8 software using the P/4 subtraction procedure (when using solution 1, see below) or the P/8 leak subtraction procedure (when using solution 2, see below). Reverse polarity pulses were used for the leak subtraction procedure. Analyses were done with IGOR graphing and analysis environment (Wavemetrics) using a procedure originally written by Boris Barbour (Ecole Normale Supérieure) and adapted by P. Isope.

For experiments combining electrophysiology and imaging, calcium currents were recorded using a solution containing (mm): 115 NaCl, 3 KCl, 26 NaHCO3, 1.25 NaH2PO4, 15 glucose, 2 CaCl2, 1 MgCl2, 0.5 tetrodotoxin, 2 TEACl, 0.1 4-aminopyridine (4-AP), 0.1 picrotoxin, 1 CsCl, 0.5 kynurenate (solution 1) and 0.1 nickel in four cells. In order to improve voltage clamping, seven cells were also recorded in low calcium concentration solution. The solution was the same as that above except (mm): 1 CaCl2, 3 MgCl2, 0 KCl and 0.050 ZD7288, 0.1 nickel, 0.003 nifedipine (solution 2).

LVA calcium currents were isolated by preincubation of the slices with 200 nmω-agatoxin IVA for 45 min. A closed perfusion (5–10 ml) was also used to maintain a high concentration of toxin during the experiments. Five cells were also studied with solution 1 after preincubation with a cocktail of ω-agatoxin IVA (200 nm) + conotoxin MVIIC (3 μm) and nifedipine (3 μm) in the bath in order to test contamination from other types of HVA calcium conductances.

Mibefradil (6 μm) and nickel (100 μm) were used as T-type and R-type channel antagonists (see Results and Discussion). All drugs were purchased from Sigma-Aldrich except 4-AP and ZD7288 which were from Tocris.

Imaging

Cells were loaded with OGB-1 for 15 min before imaging. We used a custom-built two-photon laser scanning microscope with an Olympus objective (×60, 0.9 NA) and a Ti–sapphire laser (Coherent) emitting at λ= 800 nm. In a first set of experiments an MRC 600 Biorad system was used for scanning and acquisition was done by a National Instruments card set driven by custom software. The setup was subsequently updated by a custom-made scanning system controlled by software written in IGOR Pro (Wavemetrics) by Jamie Boyd and Kerry Delaney (Simon Fraser University).

Fluorescence from spines and their parent dendrite was measured by scanning repeatedly (250 Hz) along a line that intersected both structures or by using high-speed frame scanning (up to 60 Hz) of a small area of the dendritic tree in a time series mode. All the analyses were carried out using IGOR Pro (Wavemetrics) and means are always given ±s.d.

Results

Properties of the low threshold calcium current

In order to isolate calcium currents in the Purkinje cell, we blocked most of the potassium conductances using a CsCl-based internal solution, with TEACl both in the internal solution (5 mm) and in the extracellular solution (2 mm). TTX (0.5 μm) and kynurenate (0.5–1 mm) were also added in the bath to block action potential and glutamatergic transmission, respectively. Calcium currents were recorded using two types of solutions (see Methods): solution 1 (2 mm CaCl2) for the combination of imaging and electrophysiology, and solution 2 (1 mm CaCl2) for electrophysiology only. We chose to experiment with young rats (postnatal day (P) 9–P14; P10–P12 for electrophysiology only) to improve the quality of voltage clamp recordings.

LVA calcium currents were elicited by depolarizing voltage steps (100–200 ms) applied to Purkinje cells from slices systematically preincubated for 45 min with ω-agatoxin IVA (200 nm), a P/Q-type calcium channel antagonist. Inactivation of the LVA calcium channels was relieved by a hyperpolarizing prepulse to −105 mV (300 ms) preceding depolarizing voltage steps (Fig. 1A, left panel). Using solution 1 (2 mm CaCl2) LVA calcium current threshold was −55 mV and reached maximal amplitude in the vicinity of −30 mV (mean maximum amplitude was 2.3 ± 0.9 nA, n = 9; Fig. 1A). The mean 20–80% rise time was voltage dependent: from 11.7 ± 4.2 ms at −50 mV to 6.4 ± 0.7 ms at −40 mV (n = 7). The mean inactivation decay time constant was also voltage dependent: from 26.1 ± 18 ms at −50 mV to 16.44 ± 2.13 at −40 mV (n = 7). In order to determine activation properties, estimates of the relative conductance (G/Gmax) were obtained from peak current–voltage relations after correction for change in driving force. The reversal potential used in the correction was estimated by extrapolation of the peak current–voltage plot (mean Vrev= 47.2 ± 7.6 mV; n = 8; see Fig. S1 in Supplementary Material). For each cell, the relative conductance was plotted against depolarizing voltage steps and fitted by a Boltzmann equation (see Fig. 1 legend). The mean half-conductance potential was −45.6 ± 3.7 mV and the slope factor was 2.6 ± 1.6 mV (n = 9; Fig. 1A and C). Steady-state inactivation of the LVA calcium current was estimated by fitting the relative amplitude of the current as a function of the hyperpolarizing prepulse with the Boltzmann equation. The mean half-steady-state inactivation potential was −77.2 ± 7 mV and the slope factor was 5.4 ± 1.6 mV (n = 7; Fig. 1B and C).

Figure 1. Voltage dependence of activation and inactivation of the LVA calcium current.

Figure 1

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A, activation of the LVA calcium current. To remove inactivation a prepulse to −105 mV (300 ms) was applied before depolarizing voltage increments (100 ms). Holding potential was −70 mV. Left panel, traces for steps to −65, −55, −45 and −40 mV, respectively. Right panel, the normalized conductance for the cell shown in the left panel is plotted as a function of membrane potential and fitted with a Boltzmann equation: 1/(1 + exp((V1/2–V)/k)); see also Fig. S1 in Supplementary Material), with half-maximal activation at −44.4 mV and a slope factor of 4.1 mV. B, steady-state inactivation of the LVA calcium current. A test potential (100 ms) at −40 mV was preceded by incremental hyperpolarizing pulses (300 ms) from −100 mV. Holding potential was −60 mV. Left panel, traces for hyperpolarizing steps to −85, −75, −70 and −60 mV. Right panel, the mean normalized amplitude of the current for the cell shown in the left panel is expressed as a function of the membrane potential and described with a Boltzmann equation, with half-maximal activation at −74 mV and a slope factor of 5.3 mV. C, summary data. Kinetic parameters for experiments using solution 1 (black columns, n = 9), solution 2 (grey columns, n = 7) and solution 1 supplemented with conotoxin MVIIC (3 μm) and nifedipine (3 μm; open columns, n = 4). Note that only the V1/2 for activation is significantly different when calcium concentration was decreased to 1 mm CaCl2.

Since incorrect voltage clamping might affect activation and inactivation kinetics in the Purkinje cell, we set out to minimize possible voltage escape due to large dendritic calcium currents. Seven cells were recorded in a low calcium concentration solution (solution 2, 1 mm CaCl2) supplemented with a specific Ih antagonist (ZD7288, 50 μm), 0.1 mm nickel and 0.003 mm nifedipine. Furthermore, only rats that were 10–12 days old were used. Under these conditions, the threshold for the LVA calcium current was −45 mV and the mean maximum amplitude was 0.9 ± 0.6 nA (n = 7). The mean half-conductance potential was −35.3 ± 4.75 mV (n = 7; slope factor 3.75 ± 1.35 mV) and the mean half-steady-state inactivation potential was −68.3 ± 4.5 mV (n = 5; slope factor 5 ± 0.5 mV; Fig. 1C, Table 1 and Fig. S2 in Supplementary Material). Under those conditions the V1/2 for activation is significantly different (see Discussion). These results suggest that under agatoxin IVA block, the LVA calcium current recorded is mediated by T-type calcium channels (see Table 1 for a comparison with other preparations).

Table 1.

Kinetic properties of the LVA Ca2+ current recorded compared to other preparations

Type of preparation [Divalent ion] (mm) Threshold(mV) V1/2,activation(mV) V1/2,inactivation(mV) References
Cerebellar slices
 High calcium 2 Ca2+ −55 −46.6 −77.2 This article
 Low calcium 1 Ca2+ −45 −35.3** −68.3 This article
Oocytes
 Cav3.1 +α2δ2 40 Ba2+ −50 −39 — Li et al. 2004
HEK293 cell
 Cav3.1 10 Ca2+ −50 −32.3 −62 Huang et al. 2004
 Cav3.1 20 Ca2+ −30 −24.9 −54.9 Hobom et al. 2000
 Cav3.1 +α2δ2 20 Ca2+ −30 −22.3 −50.4 Hobom et al. 2000
NG108-15 cell 10 Ba2+ −60 −26.9 −53.4 Randall & Tsien, 1997
Isolated Purkinje cell 10 Ca2+ −60 −40 −79 Kaneda et al. 1990
Organotypic cerebellar slices 0.5 Ca2+ −60 −51 −86 Mouginot et al. 1997
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**

P < 0.001 (Mann-Whitney U test). For an exhaustive review of the kinetic parameters, see Perez-Reyes (2003)

Specificity of the LVA calcium current

Since other types of calcium channels have been demonstrated in Purkinje cell dendrites, we tested for a possible contamination of the LVA calcium current by HVA current. P/Q-type calcium channel contamination is unlikely because: (1) in three experiments, additional agatoxin IVA (200 nm) was added in the closed perfusion system and no additional block was observed, suggesting that the preincubation allowed a complete block of the P/Q-type calcium current, and (2) P-type calcium currents show little inactivation (Mintz et al. 1992) whereas the LVA calcium current recorded here shows a rapid inactivation.

Four cells were recorded using solution 1 (2 mm CaCl2) after preincubation with ω-agatoxin IVA (200 nm) + conotoxin MVIIC (3 μm) and application of nifedipine (3 μm) to assess whether N- or L-type channels contaminated the LVA calcium current. Under these conditions no significant difference in the biophysical properties of the LVA calcium current or in imaging experiments (see Fig. 1C and Fig. S3 in Supplementary Material) were observed. The mean half-maximal conductance voltage was −46 ± 9.6 mV (n = 4; slope factor 4.3 ± 1.55 mV) and the mean half-steady-state inactivation was −85 ± 4.5 mV (n = 5; slope factor 5 ± 0.5 mV). This suggests that current from N-type and L-type calcium channels under the conditions we used is negligible since blockade of these channels did not change the properties of the LVA calcium current recorded. Furthermore, under ω-agatoxin IVA (200 nm) block, application of mibefradil (6 μm; n = 6), a T-type and R-type calcium channel antagonist, almost completely abolished the LVA calcium current (mean block 95 ± 3.8%; see also Fig. 4C–E).

Figure 4. The major component of the LVA calcium transient observed in Purkinje cell dendritic spines is mediated by T-type calcium channel.

Figure 4

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A, calcium channel recruitment in spine when hyperpolarization prepulse (300 ms) precedes a depolarizing step (100 ms) at −40 mV. Upper traces, mean variation of fluorescence in 3 spines of the same branchlet correlated with current traces recorded at the soma. Current traces are expanded; the shaded box corresponds to the duration of the depolarizing pulse. Note that at −70 mV only a small fraction of the T-type calcium channel pool is available. Experiment includes 100 μm nickel in the bath. Fluorescence traces shown are individual sweeps filtered using a binomial filter. B, calcium channel recruitment in spines by incremental depolarizing steps (100 ms) following hyperpolarizing prepulses (300 ms) to −100 mV. Same spines as in A. C, mibefradil (6 μm) inhibition of the LVA calcium transient in an individual spine correlates with inhibition of the LVA calcium current recorded at the soma. Black line, LVA calcium transient and current elicited by a depolarizing pulse at −45 mV following hyperpolarizing pulse at −105 mV. Grey line, same protocol during application of 6 μm mibefradil in the bath. Fluorescence traces shown are individual sweeps filtered using a binomial filter. D, correlation between LVA calcium current block and LVA Ca2+ transient block by mibefradil in 6 cells. Normalized variation of fluorescence was measured by averaging the first 100 ms after the onset of the response compared to the maximal value observed before mibefradil application. Normalized current amplitude is the peak current amplitude compared to the maximal amplitude before mibefradil application. E, summary data. Left columns, percentage of inactivation of calcium current (black column) and calcium transient in spines (grey column) when there is no hyperpolarizing prepulse in the protocol of activation. Right columns, maximal block of calcium current (black column) and calcium transient in spines (grey column) after mibefradil (6 μm) application in the bath. The peak calcium current and the mean variation of fluorescence during the first 100 ms after the stimulation were used to determine the percentage of inactivation/maximal block.

Immunohistochemical studies have shown that the α1E subunit, associated with the R-type channel, is also expressed in Purkinje cells (Yokoyama et al. 1995). Because some partial overlap in the kinetics and pharmacological properties exist between the T-type and R-type channels (Randall & Tsien, 1997), we tested whether the LVA current recorded could be contaminated by an R-type calcium current. Deactivation kinetics of R-type channels has been shown to be very fast (τdeactivation < 0.25 ms upon repolarization to −60 mV and decreasing by e-fold per 65 mV of hyperpolarization; Randall & Tsien, 1997) compared to the deactivation kinetics of the T-type current, which is slow and highly voltage dependent (τdeactivation > 4 ms upon repolarization to −60 mV and decreasing by e-fold per 36 mV of hyperpolarization; Randall & Tsien, 1997). In order to determine the deactivation kinetics of the LVA calcium current in the Purkinje cell, prepulses to −105 mV were followed by 20–30 ms activation pulses, and deactivation was measured upon repolarization to varying potentials. We found that deactivation followed a slow time constant with τdeactivation= 8.8 ± 1.4 ms upon repolarization to −60 mV, and was highly voltage dependent – decreasing e-fold per 34 mV of hyperpolarization – in agreement with Randall & Tsien (1997; Fig. 2A and B). It is conceivable that the dendritic filtering could also explain the slow kinetics of deactivation that we observed; however, the high voltage dependence of the deactivation (e-fold per 34 mV) suggests that T-type calcium channels are the major component of the recorded LVA calcium current.

Figure 2. Specificity of the LVA calcium current.

Figure 2

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A and B, deactivation of the LVA calcium current. A, typical recordings illustrating deactivation kinetics of the LVA calcium current. After a hyperpolarizing prepulse to −105 mV (300 ms), a test potential at −50 mV (20 ms) was followed by repolarizing pulses (100 ms) ranging from −50 mV to −100 mV. Holding potential was −70 mV. Traces for repolarization steps to −50, −65, −75 and −90 mV are shown. Note, the progressive increase in speed of tail current decay with more negative repolarization levels. B, voltage dependence of the deactivation time constant. Pooled data for LVA calcium current (n = 10). Smooth curve is an exponential function of voltage (e-fold per 34 mV). C, effect of nickel application (100 μm) on LVA calcium current (mean inhibition was 35 ± 15%, n = 4).

Finally, results from Tottene et al. (1996, 2000) suggest that in the absence of α1H subunits, differential sensitivity to nickel can be used to discriminate between T- and R-type channels (see Discussion). Consistent with this, we found that application of 100 μm nickel in the bath blocks only 35 ± 15% (n = 4) of the LVA calcium current (Fig. 2C). Together these data argue that T-type calcium channels mediate the LVA calcium current recorded at the soma of the Purkinje cell in acute slices between P9 and P14.

Correlation of the low threshold calcium current and the calcium transient in dendrites

We then set out to localize LVA calcium input in the Purkinje cell dendritic tree using the two-photon laser scanning microscopy in combination with whole-cell voltage clamp (Fig. 3). Cells were filled with a high affinity calcium probe, Oregon green BAPTA 1 (OGB-1), that exhibits significant fluorescence at resting calcium levels, permitting us to view the structure of the dendritic tree (Fig. 3A). Figure 3C shows an average of 50 frames recorded at 40 Hz during a step depolarization at the soma. Both LVA calcium current at the soma and level of fluorescence in spines and dendrites can be recorded (Fig. 3D). The variation of fluorescence recorded in spines is not an artefact resulting from optical blurring of signals emanating from the dendrites, because extracellular regions immediately adjacent to dendritic spines showed no significant changes in fluorescence during step depolarizations (not shown).

Figure 3. Calcium transient observed in the Purkinje cell dendrite and spines is correlated with the LVA calcium current recorded at the soma.

Figure 3

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A, Purkinje neurone (P10) filled with Oregon green BAPTA 1 through the patch pipette. Maximal intensity projection of a stack of 120 scans every 0.5 μm in Z-axis. B, expansion of the outlined box in A identifying spines (sp1 and sp2). C, expanded view of the white dashed box outlined in B. Box illustrates regions of interest where fluorescence was determined. sp1 and sp2 are spines shown in B. d1 and d2 are parent dendrites of sp1 and sp2, respectively. The image is an average of 50 frames at a scan rate of 38 Hz. D, variation of fluorescence observed in boxes shown in C correlated with the current recorded at the soma. Fluorescence traces are averages of 2 sweeps. Bottom, voltage clamp protocol. Experiment includes 100 μm nickel in the bath.

We show that the LVA calcium current recorded at the soma is correlated with a rise in dendritic calcium concentration (Fig. 3D). In order to further characterize the calcium transient we set out to manipulate the holding potential at the soma and simultaneously monitor the variation of fluorescence induced by step depolarizations in a small area of the Purkinje cell dendritic tree using a fast scanning (40–50 Hz) time series. In all cells tested (n = 5, 24 spines), incremental hyperpolarized prepulses allowed a larger recruitment of channels as indicated by the current recorded at the soma and the increase in OGB-1 fluorescence observed in dendritic spines (Fig. 4A and E). When no hyperpolarizing prepulse was given (from −70 mV) before the step depolarization both the current and the transient were small, suggesting that at holding potentials near the resting membrane potential, only a fraction of the calcium channels are available for activation. We also show that incrementally depolarizing steps correlated with an increase in dendritic spine OGB-1 fluorescence (Fig. 4B). These results show that changes in OGB-1 fluorescence measured in the dendritic spines were well correlated with the current, strongly suggesting that T-type calcium channels mediated the calcium transient. Furthermore, application of mibefradil (6 μm) under ω-agatoxin IVA block completely abolished the calcium transient in spines in four cells and reduced it by nearly 60% in two other cells (Fig. 4C–E; n = 6). As mentioned above, preincubation with ω-agatoxin IVA (200 nm) + conotoxin MVIIC (3 μm) and application of nifedipine (3 μm) failed to modify the properties of the calcium transient (n = 4; see Fig. S3 in Supplementary Material).

Although the voltage-dependent properties of the calcium transient observed in spines and dendrites strongly suggest that it is mediated mostly by T-type calcium channels, we also addressed the possibility of a putative R-type contaminant in the dendritic calcium signal by applying nickel (100 μm) in the bath. As mentioned above, the current recorded at the soma in 100 μm nickel was reduced only by 35 ± 15% (n = 4). Under nickel application, the calcium transient could still be observed both in the dendrite and spine (Figs 3, and 4A and B, were carried out with 100 μm nickel in the bath). These findings show that calcium transients observed in the dendritic tree of the Purkinje cell are mainly mediated by T-type calcium channels composed of α1G and/or α1I subunits.

T-type calcium channels are present in dendritic spines

In order to rule out the possibility that the calcium transient recorded in spines is due to diffusion from the parent dendrites we monitored the onset kinetics of the calcium transient in spines and the parent dendritic shaft (Figs 5 and 3). Linescan mode (scan of only one pixel line) using a line crossing the spine and the parent dendrite allowed us to record calcium transients at 250 Hz (Fig. 5A). This experiment showed that: (1) the mean time constant of the calcium transient rise time was 16.57 ± 11 ms (n = 8) in spines and 16.62 ± 7.1 ms (n = 8) in dendrites (Fig. 5B and C), and (2) the spine calcium transient occurred concurrently with the dendritic calcium transient (Fig. 5D and see also Fig. 3D). These results suggest that it is unlikely that the calcium transient observed in dendritic spines is due solely to diffusion from the dendritic shaft (see Discussion).

Figure 5. Coincident LVA calcium transient in spine and dendritic shaft.

Figure 5

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A, simultaneous LVA calcium transient onset in spine and parent dendrite. Upper panel, average image identifying a dendritic spine and parent dendrite (arrowheads). Dashed line identifies the line repeatedly scanned during the experiment. Middle panel, variation of fluorescence in the spine (grey line) and in the parent dendrite (black line) during a depolarizing pulse to −40 mV (protocol below). The sampling rate was 250 Hz. Fluorescence traces are averages of 4 sweeps and a binomial smoothing filter was applied. B, LVA calcium transient onset fitted by a monoexponential function. First point of the fit is the closest point from the intersection between baseline (dashed line) fluorescence and onset. C, plot of the time constant of the LVA calcium transient onset in spine versus parent dendrite. D, left panel, average ΔF/F for 7 spine (thin line) and parent (thick line) dendrite pairs (4 cells; 50 sweeps) observed in linescan mode. Note the absence of delay between the onset in the spine and parent dendrite and the fast rising time. Right panel, enlargement of the dashed box at an expanded time scale.

Discussion

We have shown here that LVA calcium current elicited in Purkinje cells from acute slices was correlated with dendritic calcium transients. The presence of T-type calcium channels has already been demonstrated in proximal dendrites (Mouginot et al. 1997) and in secondary dendritic branches (Watanabe et al. 1998), but we now show strong evidence for the presence of T-type calcium channels in the smallest integrative compartment of the dendrite, the spine.

Clamping issues

Since incorrect voltage clamping can affect the kinetics of the current recorded we attempted to optimize our conditions and protocols in order to reduce escape from voltage clamp. For example, we used young animals, mostly P9–P12 with relatively smaller dendritic trees. For imaging studies, we used step depolarization protocols that did not produce currents in excess of 2.5 nA. The caesium chloride internal solution and potassium channel blocker used (5 mm TEACl in internal solution, 2 mm TEACl and 0.1 mm 4-AP in the bath) should improve cell compactness and clamp speed.

To confirm that voltage clamp errors induced by large currents were not significantly altering our conclusions, in a second series of experiments, we recorded from cells under low calcium concentration (1 mm CaCl2) and an Ih blocker in order to decrease the calcium current and increase electrical compactness of the cell. Under such conditions, it is noteworthy that activation and inactivation parameters were found to be closer to those in heterologous expression systems (see Table 1 and Perez-Reyes, 2003). A significant shift (about 10 mV) toward more depolarized potential is observed, but only for the activation parameter, suggesting, as already stated by Destexhe et al. (1998) and Perez-Reyes (2003), that the differences were probably due to a better voltage clamp in low calcium concentration (1 mm CaCl2). However, during imaging studies (in 2 mm CaCl2), small depolarization steps were used, reducing the voltage escape in the distal dendrites. Also, the calcium transients observed were well correlated with small depolarizations in the vicinity of the resting membrane potential, confirming that they were mediated by T-type calcium channels.

Another clamping issue is the assumption that spiny dendrites accurately follow somatic voltage commands. Modelling studies have shown that the Purkinje cell is remarkably compact in the steady state (Shelton, 1985; Hausser & Roth, 1997; Roth & Hausser, 2001), but voltage transients are heavily filtered. We therefore postulate that the holding potential and the long steps of voltage during prepulses and steady depolarization are accurate, but the voltage transients are filtered and delayed. Our imaging studies show that small hyperpolarizing or depolarizing steps generated at the soma of the Purkinje cell correlate linearly with changes in fluorescence (ΔF/F) in spines (Fig. 4A, B and D), indicating that voltage steps can be transmitted to distal parts of the dendritic tree. We concede that dendritic filtering might influence the deactivation/inactivation time constants of the calcium currents. However, we found a high voltage dependency in the deactivation kinetics (e-fold per 34 mV) that is consistent with T-type, but not R-type, deactivation (see Randall & Tsien, 1997).

In some traces, the LVA calcium current does not decay to baseline during a 100 ms pulse (Fig. 4A and B). This feature was already observed in Randall & Tsien (1997; see their Fig. 1) and Mouginot et al. (1997; see their Figs 3B and 6B). Several hypotheses could explain this sustained component in our experiments. (1) There may be an error in the leak subtraction procedure due to the activation of unblocked Ih channels during the hyperpolarizing prepulse to −105 mV. (2) Another T-type subunit such as α1I/Cav3.3 may be activated (Yunker et al. 2003). The α1I/Cav3 subunit has much slower kinetics than α1G/Cav3.1, the major component of the T-type calcium current, and can show a sustained-like decay (Monteil et al. 2000b). α1I also has an activation curve that is shifted toward more depolarized values. This could explain why the sustained current is not always present. (3) There may be a contaminating calcium current since we did not systematically use the cocktail including ω-agatoxin IVA–conotoxin MVIIC–nifedipine. This contamination would not necessarily appear in spines, because mibefradil application (Fig. 4C and D) completely blocked calcium transients in 4 of the 6 cells tested.

Pharmacological discrimination of T-type versus R-type calcium channel

Even if we cannot completely exclude contamination of the LVA calcium transient observed in spines by R-type calcium channels, this hypothesis is unlikely for several reasons. Tottene et al. (1996, 2000) have defined three different types of native R-type calcium current in cerebellar granule cells: Ra or G1, Rb or G2 and Rc or G3. In their experiments using gene-specific knock-down by antisense oligonucleotide, Ra-Rb-Rc current is formed by the α1E subunit that is also expressed in the Purkinje cell. The Rb subunit shares some kinetic properties with T-type channels, because its activation curve is shifted towards more negative values than the other R-type channels. Fortunately both Ra and Rb are fully blocked by 30–50 μm nickel (Forti et al. 1994; Tottene et al. 1996, 2000). The Rc subunit is poorly blocked by nickel (IC50= 153 μm), but its activation range is shifted towards positive potentials as compared to Rb with a half-maximal activation potential (V1/2) of −4 mV and a threshold of −25 mV. For Ra and Rb isoforms of the R-type channel, the sensitivity to nickel is 6–10 times higher than the reported sensitivity of the T-type channels containing α1G and α1I subunits in HEK cells (Lee et al. 1999; Monteil et al. 2000a, b) and in Purkinje cells (Mouginot et al. 1997) thus inconsistent with the LVA calcium current we observed. Nickel is a high affinity blocker of the α1H channel, but this subunit is not expressed in Purkinje cells (Talley et al. 1999). In the absence of α1H subunits, differential sensitivity to nickel can be used to discriminate between T- and R-type channels in Purkinje cells.

We postulate that 100 μm nickel would have blocked both Ra- and Rb-type calcium currents. Also, the high threshold of activation for the Rc isoform makes a significant contamination unlikely, particularly at potentials between −60 mV and −40 mV at which T-type calcium channels are not fully activated and then reducing the potential for escape from voltage clamp induced by the large inward currents within the dendritic tree. Most of our imaging data were obtained with depolarizations to potentials ranging between −50 and −40 mV (see Figs 3 and 4). Furthermore, another study by Wilson et al. (2000), using α1E knock-out mice, suggests that the Rc-type calcium current could be mediated by the α1A subunit which induces slowly inactivating calcium currents that contrast strongly with the current we have recorded.

In our study, only 35 ± 15% of LVA calcium current was blocked by 100 μm nickel, in agreement with studies which characterized T-type channels (Mouginot et al. 1997), but in apparent contrast to Watanabe et al. (1998) in which the same concentration of nickel blocked a transient calcium input in dendrites under ω-agatoxin IVA block. However, the protocol used and a subsequent modelling study (Miyasho et al. 2001) suggest that the major component of that transient calcium rise in distal dendrites might have been mediated by the R-type calcium channel. In Watanabe et al. (1998), the more physiological conditions and the low-resolution imaging system might have lead to undetectable T-type calcium currents/transients.

Calcium transients in dendritic spine

In order to demonstrate that the calcium transients observed in dendritic spines are due to voltage-dependent channels located on the spine, we needed to rule out buffered calcium or dye-bound calcium diffusion from the dendrite. The onset of the calcium transient in spines occurs with a time-to-peak of less than 20 ms (mean τonset= 14 ± 8.8 ms; n = 10). This rapid elevation occurs with the same slope in both the dendrite and spine. In addition, no delay between the spine and dendrite was observed, making a dendritic source of spine calcium unlikely. Data from hippocampal spines in which clear failures of spine calcium channel openings can be resolved indicate that diffusion of dendrite-derived calcium would lead to spine calcium transients that were significantly delayed and slower compared to the dendrite (Sabatini & Svoboda, 2000). With regard to dye-bound calcium diffusion, experiments from Majewska et al. (2000) have shown a time constant of 671 ms for recovery of calcium green fluorescence in spines after photobleaching. Since a dye of similar structure was used in our study, we postulate that the time for equilibration of dendrite-derived diffusing calcium-bound dye would be too long to account for the increase in calcium in spines to be attributed to calcium-bound indicator diffusion. In addition, experiments on the diffusional mobility of parvalbumin, one of the highly mobile calcium-binding proteins in Purkinje cells, demonstrated an apparent diffusion coefficient of 43 μm2 s−1 (Schmidt et al. 2003) which is inconsistent with the kinetics of the calcium transient in spines. Therefore, we conclude that our results are most consistent with a model in which T-type channels are present on both spines and dendrites.

Several studies have previously shown synaptic depolarization-dependent calcium transients in Purkinje cell dendritic spines (Lev-Ram et al. 1992; Miyakawa et al. 1992; Denk et al. 1995; Eilers et al. 1995; Miyata et al. 2000; Wang et al. 2000), but none of them have identified the type(s) of calcium channels involved. Denk et al. (1995) showed that synaptic inputs induce two kinds of calcium transients in individual spines. In some spines, hyperpolarizing the soma blocked the calcium transient in the spine, suggesting that it was mediated by voltage-dependent calcium channels, but in other spines hyperpolarization increased the size of the calcium transient. The authors postulated the presence of calcium-permeable AMPA receptors in that particular case. However, it is thought that AMPA receptors in Purkinje cells contain the GluR2 subunit, which would make them impermeable to calcium (Petralia et al. 1997). Another hypothesis consistent with our data would be that the hyperpolarizing holding potential deinactivated T-type calcium channels in spines, and that the synaptic input was strong enough to drive the membrane potential above −60 mV. A similar explanation could be proposed to explain the persistence of calcium transients in isolated spines after hyperpolarizing pulses in Fig. 2 of Wang et al. (2000).

Physiological implications

Our results were obtained in immature animals (P9–P14), but it is interesting to consider the implications of T-type calcium channel expression in the adult. Although we did not study older animals, we observed that the size of the T-type calcium current increased with the age of the rat (between P9 and P14; data not shown). Obviously, this effect might only reflect the increase in the dendritic surface in the Purkinje cell between P9 and P14, but immunochemistry has also shown that the levels of α1G expression in the dendritic tree of the Purkinje cells in rodent increase during postnatal development (Yunker et al. 2003), suggesting that the localization observed here might indeed persist in adults.

Our findings are consistent with previous results showing that the HVA calcium transient is mainly mediated by P/Q-type calcium channels (Llinas et al. 1989; Usowicz et al. 1992; Bindokas et al. 1993; Watanabe et al. 1998) and that the LVA calcium transient can be unmasked when P/Q-type channels are turned off (Watanabe et al. 1998). Our results are in agreement with the hypothesis that a boosting of P/Q-type calcium spikes in the dendrites can be mediated by regenerative LVA calcium spikes (Llinas et al. 1992; Watanabe et al. 1998) or that regenerative LVA calcium spikes can occur independently of P/Q-type spikes in the dendritic tree (Cavelier et al. 2002).

Recent studies showed that the number of AMPA receptors at the parallel fibre–Purkinje cell synapse can be low, even null (Zhao et al. 1997) and most of the synapses do not show any detectable current at the soma (Isope & Barbour, 2002). Our study suggests that even small synaptic inputs (below the threshold for a regenerative calcium spike generated by P-type channels) might generate a voltage-dependent calcium current via T-type calcium channels, allowing presumed silent synapses to be involved in forms of calcium-dependent plasticity. Together with findings reporting the presence of P/Q-type calcium channels in spines (Kulik et al. 2004), two levels of voltage-dependent calcium currents might exist in the same spine. We can then speculate that in spines having a parallel fibre input inducing relatively small depolarization only, T-type channels could be activated, whereas climbing fibre stimulation or a coincidence between parallel fibre and climbing fibres could trigger the opening of both P-type and T-type channels. This selectivity could be involved in determining the sign of the plasticity at the parallel fibre–Purkinje cell synapse. Parallel fibre input alone would promote LTP (Lev-Ram et al. 2002, 2003), while coincidence between climbing fibre and parallel fibre inputs would trigger LTD (see Ito, 2001 for review).

Acknowledgments

We would like to thank Anne Feltz, Boris Barbour and Yo Otsu for helpful discussion, Jamie Boyd and Kerry Delaney for assistance with two-photon hardware and software, and Karen Brebner. This study was supported by a CIHR grant to T.H.M. (MT12675). T.H.M. is a CIHR investigator and a Michael Smith Foundation for Health Research (MSFHR) senior scholar. P.I. was funded by Fondation pour la Recherche Medicale (France) and by Neurosciences Graduate Program (Canada) and is a MSFHR Post Doctoral Fellow.

Supplementary material

The online version of this paper can be accessed at: DOI: 10.1113/jphysiol.2004.074211 http://jp.physoc.org/cgi/content/full/jphysiol.2004.074211/DC1 and contains supplementary material consisting of three figures and legends entitled:

Figure S1. Estimation of the relative conductance

Figure S2. Example of activation of the LVA calcium current in solution 2 (1 mm CaCl2)

Figure S3. Example of imaging experiment after preincubation with ω-agatoxin IVA (200 nm) + conotoxin MVIIC (3 μm) and application of nifedipine (3 μm) in the bath.

This material can also be found at: http://www.blackwellpublishing.com/products/journals/suppmat/tjp/tjp623/tjp623sm.htm

Supplemental Data
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