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1H T-type calcium channels in excitability of mouse embryonic primary vestibular neurones
1 Institut des Neurosciences de Montpellier, INSERM U583, Hôpital Saint Eloi, 80 rue Augustin Fliche, 34091 Montpellier cedex 5, France
2 Institut de Génomique Fonctionnelle, CNRS UMR 5203, INSERM U661, 141 rue de la Cardonille, 34094 Montpellier cedex 5, France
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Abstract |
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(Received 26 April 2005;
accepted after revision 15 June 2005;
first published online 16 June 2005)
Corresponding author G. Desmadryl: INSERM U583, INM, Hôpital Saint Eloi, 80 rue Augustin Fliche, 34091 Montpellier cedex 5, France. Email: desmad{at}univ-montp2.fr
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Introduction |
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To date, three different T-channel subunit isoforms, designated as Cav3.1/
1G, Cav3.2/1H and Cav3.3/1I, have been cloned and characterized (Cribbs et al. 1998; Perez-Reyes et al. 1998; Lee et al. 1999a; Monteil et al. 2000a,b). These subunits vary in their electrophysiological properties such as activation, inactivation and deactivation kinetics, as well as recovery properties following short-term inactivation (Klockner et al. 1999; Lee et al. 1999a; McRory et al. 2001a,b; Chemin et al. 2002a). These isotypes also display distinct sensitivity to Ni2+ (Lee et al. 1999b), show differences in their distribution in the nervous system (Talley et al. 1999) and, as expected, differ in their roles in physiology (for review see Perez-Reyes, 2003). Further, Cav3.2 subunit expression has been correlated with skeletal muscle (Bijlenga et al. 2000) and neuronal (Chemin et al. 2002b; Mariot et al. 2002) differentiation.
In the mouse embryonic peripheral vestibular system, large T-type calcium currents (T-currents) were recorded in neurones isolated between embryonic day 14 (E14) and E17 (Chambard et al. 1999). The proportion of neurones with a large T-current decreased drastically between E17 and postnatal day 4 (P4) (Desmadryl et al. 1997; Chambard et al. 1999). This change in T-channel expression coincides with the neurite outgrowth which starts at E13 and E14 and reaches a maximum around E17 (Nordemar, 1983), when the first synaptic contacts with the sensory cells are observed (Anniko, 1983; Mbiene et al. 1988; Desmadryl & Sans, 1990). Because the presence of a large T-current at E17 is transient and occurs at a critical period of the development and maturation of vestibular neurones, we have analysed its properties and function in acutely isolated E17 vestibular neurones. Here, we demonstrate that their electrophysiological and pharmacological properties are similar to those of the T-current generated by the recombinant Cav3.2 subunit, and that these T-channels play an important role in the excitability of developing vestibular neurones.
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Methods |
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Pregnant Swiss mice (CERJ, Le Genest, France) were killed by inhalation of CO2 followed by cervical dislocation in accordance with French and European guidelines. The age of the E17 fetuses was determined according to the vaginal plug day (E1). The gravid uterus was collected in sterile phosphate-buffered saline (PBS, Invitrogen) supplemented with glucose (33 mM). The embryos were killed by decapitation. For each experiment about 20 vestibular ganglia were dissected and collected in sterile PBS and enzymatic dissociation was performed by treatment with 0.15% EDTA-trypsin at 37°C for 6 min. The enzymatic reaction was stopped by addition of 10% fetal calf serum (Invitrogen). The culture medium contained Neurobasal Medium supplemented with 2% B-27 (Invitrogen), 25 µM glutamate and 0.5 mM glutamine. Mechanical dissociation was performed with fire-polished Pasteur pipettes of decreasing diameters. Ganglia were carefully triturated and neurones were plated onto 35 mm culture dishes (Nunc) previously coated with 10 µg ml–1 polyornithine. Electrophysiological experiments were performed between 2 and 6 h after dissociation.
Cav3 transfection of HEK-293 cells
The culture and transfection of human embryonic kidney (HEK-293) cells were performed as previously described (Chemin et al. 2002a). The following cDNAs encoding for human 1G (Monteil et al. 2000a), human 1I-a (Monteil et al. 2000b) and human 1H (Cribbs et al. 1998) were mixed with reporter cDNAs (GFP or CD8) using a 10: 1 ratio. Two days after transfection, cells were re-plated at low confluence and T-currents were recorded as previously described (Chemin et al. 2002a).
Electrophysiological recordings
The patch-clamp technique was used in the whole-cell configuration to record ionic currents under voltage-clamp conditions and action potentials (APs) under current-clamp conditions using an Axopatch 200B amplifier (Axon Instruments, Foster City, CA, USA). Calcium currents were recorded using the following extracellular solution (mM): TEA-Cl 120, Hepes 10, glucose 10, and CaCl2 2, pH 7.35 (with TEA-OH). Recording pipettes (2–3 M
) were pulled from microhaematocrit tubes (Modulohm I/S, Herlev, Denmark) and filled with the following solution (mM): CsCl 130, Hepes 25, glucose 10, EGTA 10, Mg-ATP 3, Na-GTP 1, pH 7.35 (with CsOH). To record APs, the extracellular solution contained (mM): NaCl 135, Hepes 10, glucose 10, MgCl2 1, KCl 5, and CaCl2 2, pH 7.35 (with NaOH). The pipette solution contained (mM): KCl 135, Hepes 10, glucose 10, NaCl 5, EGTA 5, ATP-Mg 3, GTP-Na 1, pH 7.35 (with KOH). The osmolarity of the all buffers used in this study was 310 mosmol l–1. Series resistances were in the range of 5–9 M corrected to 85% when necessary. The membrane capacitance could be charged with a time constant of 100 µs. No online linear leakage compensation was performed. Current signals were filtered at 10 kHz, digitized and stored. The mechanical response of T-channels was studied using a velocity-controlled perfusion system (Microlab 500B/C Series Diluter, Hamilton Company, Reno, NV, USA). Calibrated pipettes with 30 µm diameter tips were placed at a distance of 1 mm from the recorded cells to deliver extracellular medium with various velocities. Application of Ni2+ was performed by the successive application of various concentrations of Ni2+ delivered by a slow flow gravity over a 100–120 s period.
Data analysis
All experimental parameters, such as the holding and test potentials, were controlled with an IBM PC equipped with a Tecmar Labmaster analog interface (Axon Instruments). Cell stimulation, data acquisition, and analysis were performed using pCLAMP software (v6, Axon Instruments). T-current activation curves were obtained from current–voltage (I–V) relationships using the equation gmax
=
Ipeak/(Vm
–
VR) where Ipeak is the maximum current, Vm the voltage command and VR the apparent calcium reversal potential. Since the vestibular neurones also displayed large high-voltage-activated (HVA) calcium currents (Desmadryl et al. 1997) we only could estimate the value of VR at 40 mV, using an extrapolation of the ascending I–V curve (n
= 8). Conductance values were normalized and fitted with the standard Boltzmann equation G/Gmax
= 1/(1 + exp((V1/2ac
–
Va)ka) where G is the conductance at Va potential, Gm the maximum conductance, V1/2ac the midpoint of the activation curve and Ka the activation slope factor. Steady-state inactivation curves were obtained using a double-pulse protocol in which a –40 mV test pulse was preceded by a 5 s conditioning pulse at various voltages from –110 to –20 mV. Peak current at various potentials was normalized to the peak T-current amplitude measured from –110 mV and steady-state inactivation was plotted versus the conditioning potential. Data were fitted with a Boltzmann equation of I/Imax
= 1/(1 + exp((Vi
–
V1/2inac)ki) where I is the peak current at Vi potential, Imax the maximum current for the –110 mV conditioning pulse, V1/2inac the half-inactivation potential and Ki the inactivation slope factor. Time constants (
) of activation and inactivation were best fitted with a single-exponential function of the form I(t) =
A exp(–t/) where A was the current peak and the exponential time constant. Kinetics of recovery from short inactivation were calculated with a double-exponential equation of the form I(t) =
A1 exp(–t/1) +
A2 exp(–t/2) + C, where A1 and A2 were the amplitudes of each component, 1 and 2 the time constants. The Ni2+ concentration–effect relationship was fitted with a Hill equation of the form I(x) =
Axn/(IC501n
+
xn) where A was the maximum response and n the Hill coefficient. Spike and afterdepolarization potential (ADP) analyses were performed using pCLAMP software (v8, Axon Instruments). The AP peak values and the half-widths were determined. Rise and decay times were measured from 10% to 90% from the bottom and the top of the events. The ADP amplitudes and half-repolarization times were measured. Pooled data are given as mean ±
S.D. Statistical significances were determined using Student's t test.
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Results |
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Based on the amplitude of the T-current, we previously reported that E17 primary vestibular neurones could be divided into two subpopulations (Desmadryl et al. 1997; Chambard et al. 1999). In the present study, we have focused our attention on the electrophysiological properties of the neurones with large T-current amplitudes (I > 200 pA in 2 mM Ca2+), which represents 75% of the recorded cells (Fig. 1A). Calcium currents were recorded from a holding potential (Vh) of –100 mV. The average capacitance of this neurone population was 12.3 ± 3.4 pF (n = 60) and the mean amplitude of the T-current measured at –40 mV was 495 ± 255 pA (n = 36). A ramp protocol was also used to visualize LVA and HVA calcium currents (Fig. 1B). Using this protocol, the LVA/T-current presented a maximum peak amplitude at –38.4 ± 5.5 mV (n = 22). The second peak that reflected the activation of HVA currents showed a maximum at 1.8 ± 2.3 mV (n = 24) (Fig. 1B). Figure 1C shows I–V relationships of the global current, elicited by a series of 300 ms step commands from Vh –100 mV, for the peak and the sustained currents, the latter being measured between 250 and 300 ms after the pulse onset (n = 20). T-current activated between –60 and –50 mV, with peak current around –40 mV, while HVA activated above –20 mV. The voltage dependency of activation and inactivation properties of the T-current are illustrated in Fig. 1D where smooth curves correspond to Boltzmann fits with V1/2ac = –43.4 ± 1.1 mV (slope 6.0 ± 0.9 mV, n = 23) and V1/2inac = –58.2 ± 0.3 mV, (slope 3.1 ± 0.3 mV n = 5). These activation and inactivation parameters suggest a window current between –65 and –45 mV.
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ac) and inactivation (inac) were 10.0 ± 3.6 ms (n
= 20) and 32.9 ± 11.1 ms (n
= 19), respectively. To analyse T-current deactivation, cells were held at –100 mV, stepped to –40 mV for 8 ms and then hyperpolarized from –60 to –140 mV (Fig. 2D). T-current deactivation was best fitted by a monoexponential function. The time constant of deactivation (deac) was 5.28 ± 1.0 ms at –60 mV and 0.52 ± 0.09 ms at –140 mV (n
= 12) as illustrated in Fig. 2E.
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rec) of 571 ± 140 ms (n
= 22). However, a two-exponential function improved the fit of the relationships with rec of 117 ± 36 ms and 1394 ± 255 ms (n
= 22) for, respectively, the fast and the slow components. Recovery from inactivation for the T-current present in E17 vestibular neurones was reminiscent of that described for the Cav3.2 subunit (Chemin et al. 2002a).
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It was reported that neuronal T-channels are affected by mechanical stimulation (Bouskila & Bostock, 1998). Because we routinely observed that a significant reduction of the T-current amplitude occurred when the primary vestibular neurones were subject to superfusion of extracellular medium, we have analysed the changes in T-current amplitude when stimulating the neurones with a fast perfusion device (see Methods). These experiments revealed that T-current amplitude in vestibular neurones was significantly reduced during perfusion (Fig. 4A) while HVA calcium currents were unaffected (Fig. 4B). Current amplitude reduction was velocity dependent and could reach up to 40% (average value 30 ± 17%, n = 24) for the highest velocity (2 µl s–1). Voltage ramps applied before and after the perfusion stimulation verified the absence of shift in the activation properties (Fig. 4B). When the perfusion velocity was set to 1 µl s–1, current amplitude reached a minimum in 78 ± 14 s (n = 20) after the beginning of the extracellular medium application. No change in either I–V relationships (evaluated on voltage ramps) or time to peak of activation were observed before and after the perfusion (P > 0.2, n = 19). The inactivation rates appeared to be slightly slower after perfusion (P = 0.01, n = 17). As illustrated in Fig. 4C, long-term recordings subsequent to the extracellular medium application (1 µl s–1) revealed that the recovery in current amplitude, i.e. return to the initial T-current amplitude, was obtained in about 7 min (ranging from 5 to 15 min, n = 4).
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Sensitivity to Ni2+
The analysis of sensitivity to Ni2+ of the T-current in primary vestibular neurones was performed under gentle superfusion of the recording chamber. A constant low rate of flow of extracellular medium was first applied to prevent any decrease in current amplitudes due to mechanical inhibition. When the T-current amplitude was stabilized in control perfusion, Ni2+ was applied at various concentrations. Figure 5A illustrates the T-current inhibition during 10 µM Ni2+ application, whereas HVA calcium currents were not changed (Fig. 5A, inset). Since our results showed a high Ni2+ affinity, the reversibility analysed at 0.3 µM Ni2+ concentration revealed a complete reversal after wash (n = 5; Fig. 5B). Figure 5C shows the percentage of the T-current block as a function of the Ni2+ concentration from 1 nM to 10 mM. The curve fitted using a one-term Hill equation gave an IC50 of 13.4 µM.
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For current-clamp recordings in E17 primary vestibular neurones, APs were elicited from each neurone's resting potential (–55.0 ± 3.7 mV, n = 34) using a short depolarizing pulse (1.5 ms, 100–450 pA). The minimum current values required to trigger the APs and their thresholds were not significantly changed after 30 µM Ni2+ application (Table 1). The AP was characterized by a single Na+-dependent spike that could be followed by an ADP (see Fig. 6A and C) or, in 30% of the neurones, by an afterhyperpolarization (AHP) (Fig. 6D). It is important to stress here that, in our recording conditions, no train of consecutive APs was ever seen, even with long pulses (10 s). The ADP amplitudes ranged between 8 and 33 mV above the resting potential (Table 1), with a half-repolarization time lying between 4.1 and 21.9 ms (Table 1). In 11 recorded neurones, after AP recordings, neurones were switched to voltage-clamp configuration in the presence of 300 nM TTX, 60 mM TEA and 5 mM 4-AP in the extracellular medium in order to block Na+ and reduce K+ conductances, as described earlier (Chabbert et al. 1997, 2001). In voltage-clamp conditions, when the current-clamp recordings had revealed the presence of an ADP (Fig. 6A) a large T-current was always observed, with a mean amplitude measured during a voltage ramp protocol of 325 ± 383 pA (n = 5) (Fig. 6B). Conversely, no large T-currents (I < 60 pA) were recorded in any neurones that did not exhibit ADPs (n = 6, data not shown), but in which AHPs were present. More importantly, gentle addition of 10 µM Ni2+ to the bath medium of neurones reduced the ADP component (Fig. 6C) in all recorded neurones (n = 13). The shapes of Na+-dependent APs were not significantly different before and after 30 µM Ni2+ application when ADPs were recorded (Table 1; Fig. 6C). Application of 30 µM Ni2+ to neurones presenting an AHP had no effect on either the AP profiles or the post-potential component (Fig. 6D). Finally, when present, a significant reduction of the ADP components following APs was also obtained when a rapid perfusion of the extracellular medium was delivered to ADP-presenting neurones (n = 8, data not shown).
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-conotoxin GVIA (500 nM) and -agatoxin IVA (300 nM) (n
= 5), respectively (Fig. 6E). When Na+ channels were blocked (300 nM TTX) and K+ conductances were reduced (60 mM TEA and 5 mM 4-AP) subsequent voltage-clamp recordings revealed a very large T-current (Fig. 6F, I
1400 pA, for the illustrated neurone). The Ca2+ spikes remained stable when Na+-dependent spikes were blocked with 300 nM TTX but were abolished by 30 µM Ni2+ (Fig. 6G). Together, the Ni2+ sensitivity and mechanical sensitivity of the post-spike components in primary vestibular neurones strongly suggested that the occurrences following the AP were generated by the Cav3.2 subunit activation.
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Discussion |
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1H subunit. Although, the present study performed on native peripheral vestibular neurones did not molecularly identify the subunit carrying the large T-current, its high sensitivity to Ni2+, its mechanosensitivity (similar to that obtained with the Cav3.2 subunit transfected in HEK-293 cells), as well as its electrophysiological properties, reveal that the T-channels recorded in vestibular neurones are comparable in most aspects to cloned Cav3.2 channels (for a review see Perez-Reyes, 2003). Therefore, our results strongly support the concept that the Cav3.2 subunit is involved in the physiology of vestibular neurones. This study also demonstrates that this T-current plays a critical role in the excitability of the vestibular neurones at E17, when ontogenesis of the peripheral vestibular system takes place. Ni2+ sensitivity of native and Cav3.2 T-channels
High sensitivity to Ni2+ is a critical distinctive feature of the recombinant Cav3 channels since both Cav3.1 and Cav3.3 channels present a 20-fold lower sensitivity to Ni2+ than Cav3.2 (Lee et al. 1999b). Our data show that the block of T-current can be observed at a concentration as low as 100 nM and was reversible. Concentration dependence of the blockade of T-current by Ni2+ reveals an IC50 of 13.4 µM comparable to the IC50 of 12 µM reported for cloned Cav3.2 calcium channels (Lee et al. 1999b) and is in the range of the IC50 of 3 µM found in sensory neurones (Dubreuil et al. 2004). Such high sensitivity to Ni2+ of the T-current is a strong pharmacological argument for the claim that the Cav3.2 channel subunit carries the large T-current present in developing primary vestibular neurones.
Mechanical sensitivity of native and Cav3.2 T-channels
Another novel aspect of this study is to demonstrate the sensitivity of the native and recombinant Cav3.2 channels to the perfusion flux. In vestibular neurones short-term application of extracellular medium produces a decrease in the T-current amplitude, and in current-clamp experiments suppresses the ADP component, suggesting a direct effect of the flux on the T-channels. These responses could be reversed several minutes after the end of the perfusion flux. We also observed a comparable mechanical inhibition of T-current on recombinant Cav3.2 channels expressed in HEK-293 cells while no such modulation was found with Cav3.1 and Cav3.3 channels. Similarly, Calabrese et al. (2002) reported that recombinant Cav3.3 channels were insensitive to stretch stimuli. An identical mechanosensitivity of T-current induced by a stream of bath solution has been reported in rat anterior pituitary cells (Ben-Tabou et al. 1994) as well as in rat sensory neurones (Bouskila & Bostock, 1998). This phenomenon was also reversible within several minutes. Since the Cav3.2 isotype appears to be the principal T-channel subunit in sensory neurones (Talley et al. 1999), our study reconciles data obtained in vestibular (sensory) neurones and in transfected HEK-293 cells. The mechanisms by which decreases in T-current occurred during fast flow perfusion are unknown. Stream flux could induce changes in cell volume and then modulate the activity of stretch channels as reported in magnocellular neurosecretory cells (Oliet & Bourque, 1993; Bourque & Oliet, 1997). This could be mediated by a combination of force transmission through cytoskeletal and biochemical constituents (for review see Papadaki & Eskin, 1997). However, the latter hypothesis could not explain why among the three T-type Cav3 subunits expressed in HEK-293 cells only Cav3.2 is affected by the fast flow perfusion.
T-channels in vestibular neurones share properties of Cav3.2 subunit
The electrophysiological parameters that we have described here are complementary features to differentiate among T-type Cav3 subunits. The T-current in vestibular neurones showed fast activation and inactivation kinetics similar to those of Cav3.1 and Cav3.2, while the Cav3.3 current displayed activation and inactivation kinetics six times slower (Klockner et al. 1999; Lee et al. 1999a; McRory et al. 2001a,b; Chemin et al. 2002a). Therefore, a role for the Cav3.3 channels in vestibular neurones can be excluded. Deactivation kinetics is an another criterion for identifying T-channel subunits since Cav3.1 and Cav3.2 currents deactivate slowly, as compared to the Cav3.3 current (Klockner et al. 1999; Lee et al. 1999a; McRory et al. 2001a,b). Our data show that the T-current deactivates with kinetics close to those of currents induced by Cav3.1 or Cav3.2 subunits (Klockner et al. 1999; Kozlov et al. 1999; Chemin et al. 2002a). However, the deactivation rate appeared to be faster than that reported for Cav3.2 and was closer to those of the Cav3.1 subunit (Klockner et al. 1999; Chemin et al. 2002a). Such discrepancies could reflect differences among human and rodent T-channels as previously reported in rat brain T-channels subunits expressed in HEK-293 cells (McRory et al. 2001a,b). Another difference from the cloned Cav3.2 channel was the 15 mV more positive half-steady-state inactivation voltage found in primary vestibular neurones compared to that reported in cloned channels. This could depend on disparities recorded in heterologous cell expression systems and those present in between channels native neurones that may imply specific regulations unidentified to date.
Activation, inactivation and deactivation kinetics indicate that native T-channels present in E17 vestibular neurones are likely to correspond to either Cav3.1 or Cav3.2 isotypes. Interestingly, the demonstration that recovery from short inactivation of the Cav3.2 current was a significantly slower process than that of Cav3.1 and Cav3.3 current recovery (Klockner et al. 1999; Chemin et al. 2002a) offers the possibility of distinguishing between Cav3.1 or Cav3.2 isotypes by an electrophysiological approach. The time constants of recovery from inactivation found in vestibular neurones were very slow and comparable to those reported in Cav3.2-transfected HEK-293 cells (Chemin et al. 2002a). However, the fits of the short inactivation recovery were improved by using two exponentials as previously reported for native T-currents in neurones (Bossu & Feltz, 1986; Takahashi et al. 1991), clonal pituitary cells (Herrington & Lingle, 1992) and skeletal muscle fibres (Berthier et al. 2002) expressing the Cav3.2 subunit. Our results indicate that the recovery following short inactivation could occur in two separate phases as reported by Berthier et al. (2002).
Role of T-current in AP profile
Our results suggest that T-channels could be implicated in AP repolarization components by inducing an ADP or, in some instances, Ca2+ spikes. The involvement of the Cav3.2 subunit was further confirmed since ADP could be reduced during the application of 10 µM Ni2+. At this concentration, the ADP was not totally removed since the T-current diminished by about 40%. Because the HVA calcium currents were affected by higher Ni2+ concentration (and probably other conductances as calcium-activated currents) we were unable to carry out concentration-dependent blockage of ADP and completely abolish this component. This contribution of T-channels in central neurones has been reported and already attributed to Cav3.2 channels since a low concentration of Ni2+ blocked ADP and suppressed intrinsic burst firing (Su et al. 2002; Dubreuil et al. 2004). In rat dorsal root ganglion neurones, T-current was shown to generate ADP that contributes to burst firing (Lovinger & White, 1989; Dubreuil et al. 2004). Moreover, in rat sensory neurones, changes in the AP repolarization phase induced by the flow of bath solution have also been reported and attributed to N-type and T-channels (Bouskila & Bostock, 1998).
Physiological significance
Based on our present data, we suggest that Cav3.2 channels are functionally expressed in E17 embryonic vestibular neurones when neuronal growth and synaptogenesis between afferents and sensory hair cells occur (Desmadryl & Sans, 1990), and therefore may play a key role in these events. The involvement of Cav3.2 in morphological differentiation has been identified in the neuroblastoma–glioma NG108-15 cell line where these channels contribute to both electrical behaviour and neuritogenesis (Chemin et al. 2002b). This Cav3.2 channel activity should contribute to a transient increase in [Ca2+]i which is likely to participate in the vestibular ontogenesis by controlling axon growth and guidance, as described for embryonic spinal cord development (Gu & Spitzer, 1993; Gomez & Spitzer, 1999). Since the afferent patterns of vestibular hair cells are complex (Baird et al. 1988; Fernandez et al. 1990; Goldberg, 1991), it will therefore be relevant to investigate in detail how Cav3.2 channels modulate the ontogenesis of primary vestibular afferents and the related synaptogenesis in sensory hair cells. The role of the mechanical sensitivity of Cav3.2, which could be involved in the development of the vestibular system, should be explored further since dendritic spines are able to generate momentary contractions in relation to [Ca2+]i transients evoked by action potentials (Korkotian & Segal, 2001). Further investigations could now include the exploration of the vestibular development in Cav3.2 knockout mice (Chen et al. 2003).
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, peak current; •, sustained current). D, steady-state activation (•) and inactivation (
