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1 Neurophysiologie Cellulaire, CNRS, UMR 6150
2 Neurobiologie des Canaux Ioniques, INSERM, U641, IFR Jean Roche, Faculté de Médecine, Boulevard Pierre Dramard, 13916, Marseille Cedex 20, France
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Abstract |
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(Received 19 August 2005;
accepted after revision 5 October 2005;
first published online 6 October 2005)
Corresponding author P. Delmas: Laboratoire de Neurophysiologie Cellulaire, CNRS UMR 6150, IFR Jean Roche, Faculté de Médecine, Boulevard Pierre Dramard, 13916, Marseille Cedex 20, France. Email: delmas.p{at}jean-roche.univ-mrs.fr
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Introduction |
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CG cells respond repetitively to excitatory inputs conveyed by mossy fibres with complex discharge patterns. Electrical behaviour such as spike bursting, oscillations and resonance have been revealed by pharmacological manipulation (D'Angelo et al. 1998, 2001). More recently, sensory stimulation of the vibrissae in anaesthetized rats has been shown to generate bursts of action potentials in CG cells, with maximal frequencies as high as 200 Hz (Chadderton et al. 2004). Likewise, single extracellular stimulation of rat cerebellar parallel fibres unexpectedly triggers a doublet or a burst of action potentials in CG cells, a mechanism that may be involved in the induction of long-term depression at the parallel fibre–Purkinje cell synapse (Isope & Barbour, 2002; Isope et al. 2004) and that suggests that complex, as yet unresolved, ion channel dynamics underlie granule cell firing patterns.
An important step towards understanding the molecular determinants of the excitability of granule cells is to determine the properties of Na+ currents, as they control both subthreshold activity and action potential electrogenesis. Although substantial knowledge has been accumulated about voltage-dependent K+ and Ca2+ currents in granule cells (Cull-Candy et al. 1989; Pearson et al. 1995; Shibata et al. 2000; D'Angelo et al. 2001), Na+ currents have not been explored thoroughly. Among the 10 different subunits encoding Na+ channel subtypes (Goldin, 1999), in situ hybridization and immunodetection have demonstrated that CG cells of adult rodents express Nav1.2 and Nav1.6, and possibly Nav1.1 (Westenbroek et al. 1989; De Miera et al. 1997; Felts et al. 1997; Schaller & Caldwell, 2000). A major challenge then is to relate these different channel subunits to the Na+ currents observed in CG cells and to establish their distribution.
Functionally, Na+ channels recorded in mature granule cells are tetrodotoxin (TTX)-sensitive with fast activation/inactivation kinetics (Hockberger et al. 1987; Cull-Candy et al. 1989; D'Angelo et al. 1994; Stewart et al. 1995; Carlier et al. 2000). However, these conclusions are largely based on CG cell recordings in which the electronically remote regenerative currents were inadequately controlled, making it difficult to characterize the Na+ channel isoforms electrically. Moreover, based on current clamp recordings, D'Angelo et al. (1998) have suggested a role for a persistent Na+ current in sustaining subthreshold depolarizing potentials in CG cells, although direct voltage-clamp evidence for this current has yet to be presented.
Accordingly, our aim was to answer three main questions. First, which Na+ channel subunits are expressed in differentiated CG cells, and what is their specific subcellular location? Second, what are the properties of the Na+ currents in CG cells when these currents can be voltage clamped adequately? Third, are the different components of the Na+ current attributable to channel heterogeneity and/or subcellular location? We show that CG cells display different distributions of Nav1.2 and Nav1.6 isotypes in soma, AIS and dendrites, giving rise to functionally compartmentalized Na+ currents. Some of these results have appeared in abstract form (Osorio et al. 2004).
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Methods |
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Animal use followed guidelines established by the European Animal Care and Use Committee (86/609/CEE). Cerebellar granule cells were cultured according to previously described procedures (Levi et al. 1984) with some modifications. Briefly, primary cultures were prepared from decapitated 7-day-old Wistar rats (P7). Immediately after death, cerebella were dissected out and treated with trypsin (0.02%) in Hank's balanced salt solution (HBSS, Sigma, St Louis, MO, USA) for 15 min at 37°C. The tissue was then washed several times in Neurobasal medium (Gibco-Invitrogen, Grand Island, NY, USA) supplemented with 10% fetal bovine serum and gently triturated using fire-polished Pasteur pipettes. The homogenate was centrifuged at 800 r.p.m. (50 g) for 2 min and the pellet was resuspended in plating medium that consisted of: Neurobasal medium supplemented with B27 (20 µl ml–1, Gibco), glutamine (2 mM), penicillin (50 i.u. ml–1), streptomycin (50 µg ml–1) and KCl (24 mM). After a brief centrifugation at 800 r.p.m., cells were plated in Petri dishes coated with poly-L-lysine (0.05 mg ml–1) at a density of 1 x 105 cells cm–2. Cells were incubated at 37°C in an atmosphere of 5% CO2 humidity. After 72 h, 2 µM AraC (Sigma) was added to avoid glial cell proliferation. Half of the culture medium was changed every 4 days.
Immunocytochemistry and imaging
Granular cell cultures were fixed using 4% paraformaldehyde in phosphate-buffered saline (PBS). After several washes in PBS, non-specific binding was reduced by preincubating cells in blocking buffer containing 3% bovine serum albumin and 0.1% Triton X-100 in PBS. Primary antibodies were incubated in blocking buffer for 3 h at room temperature. Secondary antibodies were incubated in blocking buffer for 1 h at room temperature. Cells were then washed in PBS and mounted in Mowiol (Colbiochem, Merck, Darmstadt, Germany). Blocking controls for non-specific staining were performed by 1 h preincubation of the primary antibodies with a large molar excess of the corresponding immunizing peptides.
Immunostaining of HEK293 cells stably expressing the human Nav1.1 sodium (Clare et al. 2000; Mantegazza et al. 2005) channel was performed as described above.
Primary antibodies used and dilutions were: ß-III tubulin (monoclonal SDL.3D10, Sigma) 1/1000; serine-rich domain of ankyrin-G node (Bouzidi et al. 2002) 1/500; MAP2 (monoclonal HM-2, Sigma) 1/400; PanNav (monoclonal K58/35, Sigma) 1/200; Nav1.1 (polyclonal ASC-001, Alomone, Jerusalem Israel) 1/100; Nav1.2 (polyclonal ASC-002, Alomone, and polyclonal 06-633, Upstate Biotechnology, Lake Placid, NY, USA) 1/100; Nav1.6 (polyclonal ASC-009, Alomone) 1/100 and ankyrin-B (monoclonal, Oncogene) 1/50. Alexa Fluor 488- and Alexa Fluor 546-conjugated goat secondary antibodies (1/200 to 1/400, Molecular Probes, Eugene, OR, USA) were used to detect rabbit polyclonal antibodies and mouse monoclonal antibodies, respectively.
Images were acquired using an optical Nikon microscope (Nikon, Tokyo, Japan) equipped with a digital Nikon Coolpix camera or a TCS SP2 laser-scanning confocal microscope (Leica Microsystems, Mannheim, Germany) and later exported into Photoshop (Adobe Systems, San Jose, CA, USA) for final processing. Images comparing peptide-blocked and unblocked antibody staining were acquired and digitally processed in an identical manner.
Reverse transcriptase-PCR
Total RNA was extracted from three cultures of CG cells at DIV 7 with TRI Reagent (Sigma) following the manufacturer's protocol. Reverse transcription reactions with primers for rat Nav1.2, Nav1.3, Nav1.6 and GAPDH (glyceraldehyde-3-phosphate dehydrogenase) were performed as described by Alessandri-Haber et al. (2002). PCR samples corresponding to an input of 0, 0.1 and 1 ng of reverse-transcribed cDNA were analysed on 1% agarose gels after 34 and 38 cycles of amplification.
Patch-clamp recordings
Whole-cell patch-clamp recordings were conducted at room temperature on CG cells kept 7–15 days in vitro (DIV 7–15). Patch pipettes were pulled from thick-walled borosilicate glass capillaries (Harvard Apparatus, Edenbridge, UK) and had a resistance of 8–12 M
. The small soma size of CG cells (< 10 µm) prevented stable recordings when using pipettes with lower resistance. All recordings were made using an Axopatch 200B amplifier (Axon Instruments, Foster City, CA, USA), low-pass filtered at 2 kHz and digitized at 25 kHz. Transient and leakage currents were digitally subtracted using a standard P/n protocol (n= 6), unless otherwise stated (e.g. voltage ramp protocol). The series resistance measured in the whole-cell mode was 12–20 M and was compensated by 60–75%. The intracellular solution used to achieve a standard Na+ gradient consisted of (mM): 100 CsCl, 30 CsF, 10 NaCl, 1 MgCl2, 0.5 CaCl2, 10 Hepes, 10 EGTA, 4 Mg-ATP and 0.4 Na-GTP (Coste et al. 2004). The bathing solution contained the following (mM): 120 NaCl, 3 KCl, 2 CaCl2, 1 MgCl2, 11 glucose, 10 Hepes, 0.5 CdCl2, 20 TEA-Cl and 1 4-aminopyridine (4-AP). In inverse Na+ gradient (Numann et al. 1991; Carlier et al. 2000), the intracellular solution consisted of (mM): 20 CsCl, 110 NaCl, 1 MgCl2, 0.5 CaCl2, 10 Hepes, 10 EGTA, 4 Mg-ATP and 0.4 Na-GTP and the extracellular solution was (mM): 210 sucrose, 10 NaCl, 3 KCl, 2 CaCl2, 1 MgCl2, 11 glucose, 10 Hepes, 0.5 CdCl2, 20 TEA-Cl and 1 4-AP. pH was adjusted to 7.35 with either NaOH or CsOH (302–305 mosmol l–1). In Na+-free external solution, the concentration of sucrose was increased to 220 mM. Input resistance and resting potential of CG cells were measured in current clamp mode with standard extracellular solution (without CdCl2, 4-AP and TEA) and an intrapipette solution consisting of (mM): 125 potassium gluconate, 5 NaCl, 1 MgCl2, 0.5 CaCl2, 10 Hepes, 10 EGTA, 4 Mg-ATP and 0.4 Na-GTP.
Local application of TTX
Local application of TTX, Na+-free saline or kainate was achieved by pressure ejection from a small patch pipette (
1 µm i.d.) positioned at 15 µm from the site of interest and directly in front of a large suction pipette (15–20 µm i.d.) (see Fig. 7A). Using small micropipettes and fast flow exchange rates, the maximum spread was estimated to be 5–10 µm in diameter as measured when sucrose was perfused in normal Na+. The fact that local application of TTX to proximal dendritic sites (7–15 µm away from the cell body) had little or no effect on the somatically recorded whole-cell current indicated that there was little, if any, diffusion of the applied solutions.
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Data were analysed using pCLAMP 8.02 (Axon Instruments) and Prism 4.0 (GraphPad, San Diego, CA, USA) software. Under conditions of inverse Na+ gradients, Na+ currents were isolated from residual contaminating outward currents by subtracting TTX-insensitive currents (except in Fig. 7A).
Conductance–voltage curves were calculated from the peak current according to the equation GNa=INa/(V–ENa) where V is the test pulse potential and ENa the reversal potential extrapolated from the current–voltage (I–V) curve. It should be noted that though ENa (obtained by fitting I–V relationships in inverse Na+ gradient) was close to the theoretical ENa, Na+ currents were typically isolated by subtraction of currents remaining in 1 µM TTX. The activation curve (G–V) was fitted using the Boltzmann function: G/Gmax= 1/(1 + exp(V0.5–V/k)), where G/Gmax is the normalized conductance, V0.5 is the potential of half-maximum activation and k is the steepness factor. Inactivation curves were constructed from normalized currents and fitted according to the Boltzmann function: I/Imax= 1/(1 + exp(V–V0.5/k)), where V is the conditioning pulse potential and V0.5 is the membrane potential at which half of the channels are inactivated. The time course of entry or recovery from inactivation was determined using the Chebyshev method to fit current traces with a single exponential equation: Y=Ymax(1 – exp(–t/
)), where is the time constant and Y represents the fraction of I recovery (It/Ipre) or the normalized current (I/Imax). For the study of slow inactivation, which required long-lasting recordings, the measured INa was corrected for run-down (estimated periodically from a holding potential (Vh) of –100 mV). The concentration–inhibition curves for TTX were fitted with the Hill equation of the form Y=Ymax[TTX]nH/(IC50nH+[TTX]nH), where Y is the percentage inhibition (e.g. 100 xI/I[TTX]= 0), IC50 the TTX concentration that produces half-maximal inhibition and nH the Hill coefficient. Results are presented as mean ±S.E.M. and n represents the number of cells examined. Statistical analysis was performed using unpaired or paired t tests, P < 0.05 being considered significant.
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Results |
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CG cells isolated at P7 were studied after 7–15 days in vitro, at a stage where they exhibit typical features of functionally mature cells, namely high input resistance (1 ± 0.1 G), negative resting potential (–66 ± 2 mV, n= 7), overshooting action potentials and spontaneous synaptic inputs, often in the form of bursting patterns (Gallo et al. 1987; Hockberger et al. 1987; Galdzicki et al. 1991; Becherer et al. 1997).
From DIV 7 onwards, CG cells had elongated thin neurites and developed a bipolar or multipolar morphology as revealed by staining for the neurone-specific marker ß-III tubulin (Fig. 1A). Axon initial segments were identified by the presence of ankyrin-G (Boiko et al. 2003), overlapping ß-III tubulin-positive processes (Fig. 1A). Using low density cultures, it was possible to assign an individual AIS to a particular CG cell and to specify the characteristics of each AIS. Ankyrin-G labelling typically began 11.5 ± 1 µm away from the soma and extended to 16.6 ± 1 µm (n= 52) (Fig. 1). Superimposition of ankyrin-G and MAP2 staining showed that the two proteins did not colocalize. MAP2 labelling was restricted to the soma and to one or more processes, identified as dendrites, while in some instances staining was also observed in the axon hillock (Fig. 1B) as already reported in hippocampal neurones (Cáceres et al. 1986). The axonal nature of the neurites harbouring ankyrin-G-positive segments was verified by dual staining with the anti-MAP2 antibody and an antibody directed against an axon-specific epitope of ankyrin-B (data not shown).
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-subunit isoforms of neuronal Na+ channels. Double labelling with PanNav and ankyrin-G antibody revealed intense immunostaining for Nav channels that coincided with ankyrin-G labelling at the AIS, as shown by confocal analysis of the extensive neuritic network (Fig. 2A) as well as single neurones (Fig. 2B). Examination of adjacent confocal planes also revealed faint PanNav staining in CG cell bodies and along axons and dendrites (Fig. 2B). PanNav-labelled segments did not immediately border CG cell bodies, indicating that Na+ channels do not cluster at high density in the axon hillock and instead concentrate in the AIS (Fig. 2B). Pre-adsorption of the PanNav antibody led to virtually complete loss of staining in the AIS, leaving some background staining in the cell body (Fig. 2C).
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-subunit antibodies and PanNav, using PanNav immunoreactivity to mark AIS based on the results presented above. Staining with Nav1.1-specific antibody at DIV 8 produced a dim signal in the soma, which was not extinguished by preincubating the primary antibody with the immunizing peptide (Fig. 3A). Similar results were obtained at DIV 14, indicating that there was no specific staining for Nav1.1, though the Nav1.1 antibody specifically recognized a protein of 250 kDa in Western blots (data not shown) and labelled HEK cells stably expressing human Nav1.1 (Fig. 3A, inset). From DIV 7 to DIV 14, anti-Nav1.2 antibody (ASC-002) consistently labelled a short proximal segment of neurites that overlapped with PanNav immunoreactivity, indicative of an accumulation of Nav1.2 at the AIS. Out of 315 initial segments identified by PanNav staining at DIV 8, 77% were colocalized with bright Nav1.2 staining (Figs 3B and 4C). This percentage remained stable over time in culture, with 85% of co-labelled PanNav/Nav1.2 AIS at DIV 14 (see Fig. 4C). It should be noted that confocal through-focus series did not detect, or barely detected, specific Nav1.2 staining along distal axons and dendrites. This pattern of Nav1.2 staining with clustering at AIS was confirmed using a second Nav1.2 antibody (06-633, see Methods).
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In agreement with immunochemistry, reverse transcription-PCR from CG cells cultured for 7 days consistently identified mRNA transcripts for Nav1.2 and Nav1.6 subunits (Fig. 5). Because the Nav1.3 isoform is often present at early stages of development in multiple brain structures, we also tested for the expression of Nav1.3 mRNA in our culture system using specific primers for rNav1.3 (Alessandri-Haber et al. 2002). Nav1.3 transcripts were not detected using cDNA templates that typically produced amplicons for Nav1.2 and Nav1.6 (Fig. 5).
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Whole-cell Na+ currents in cultured CG cells were isolated using external Cd2+ (500 µM), 4-AP (1 mM) and TEA (20 mM) and Cs+-based intracellular solutions. Under these conditions, CG cells displayed a fast activating–inactivating inward current that was fully suppressed by isosmotically substituting external Na+ by impermeant molecules (see below) and by bath-applying TTX. The TTX dose–response curve gave an IC50 of 13.2 ± 1 nM with a Hill coefficient of 0.8 ± 0.1 (data not shown).
Although cultured CG cells have small capacitance (4–7 pF) and are electrotonically compact, they possess long thin neurites, which make them not amenable to good time-clamp control. Thus, under regular or even reduced Na+ gradient, depolarizing voltage steps activated non-graded, rapidly activating Na+ currents, which manifested as late or all-or-none currents (left and middle panels in Fig. 6A). To gain better control of the transmembrane voltage, Na+ currents were examined using an inverse Na+ gradient (i.e. 110 mM[Na+]i and 10 mM[Na+]o), rendering the Na+ currents non-regenerative. Under these conditions, outward Na+ currents evoked by a standard activation protocol were smoothly graded with increasing depolarization and free of notches suggesting adequate control of the membrane potential (Fig. 6A, right panel). Outward Na+ currents were then isolated from outward contaminating currents by subtracting TTX-insensitive currents (Fig. 6B). Currents recorded in inverse Na+ gradient have rapid activation kinetics over the whole activation range and inactivate quickly and completely within a few milliseconds. Outward Na+ currents had TTX sensitivity indistinguishable from those recorded in standard Na+ and were fully blocked by 1 µM TTX (Fig. 6B). The current–voltage relationship of normalized TTX-sensitive currents plotted in Fig. 6C (n= 11) shows that currents activated at –40 mV and gradually reached maximal activation at around +10 mV. The extrapolated reversal potential for Na+ ions was estimated to be –60 mV, which agrees well with the predicted Nernst potential for Na+ ions in inverse gradient (ENa=–60.4 mV). It should be noted that I–V relationships of TTX-sensitive Na+ currents determined at DIV 7–8 or DIV 12–14 were indistinguishable, both in terms of voltage dependence and maximum peak currents (see below), indicating that redistribution of Nav1.6 subunits at DIV 7–14 was not correlated with detectable changes in current properties.
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1 µm i.d.), the tip of which was positioned at 15 µm from the site of interest and directly in front of a large suction pipette (15 µm i.d.) (Fig. 7A). This was exclusively done on CG cells that displayed a bipolar morphology with visually identified neurites comparable to those illustrated in Fig. 1. Local application of TTX at the expected site of the AIS or distally (up to 50 µm) had no effect on outward Na+ currents in 11 out of 14 neurites tested (2 neurites tested per cell; Fig. 7A). In the three remaining neurites, a partial block of 10, 32 and 52% was seen (Fig. 7B). In marked contrast, application of TTX onto the soma suppressed the Na+ current in all cells tested giving an average block of 79 ± 3% (n= 7; Fig. 7B). Total outward current block was never attained by local TTX application onto the soma whereas application of TTX via the general perfusion system fully inhibited Na+ currents. This indicates that 20% of the recorded current in inverse gradient actually resulted from channels located outside the CG cell bodies.We repeated the protocol of Fig. 7A by applying TTX locally to neuritic or somatic sites of the same cell in standard Na+ gradient (Fig. 7C). Application of TTX onto the soma blocked a lower fraction of the whole-cell Na+ current (42 ± 4%, n= 18), as expected from recruiting fast transient Na+ currents from poorly clamped portions of the neurites. Nevertheless, effects of TTX on the two neurites were not identical since application had either very small effects (2.1 ± 1%, range 0–10%, n= 11) or yielded substantial block (66 ± 6%, range 50–70%). Thus, when TTX could be applied successfully to both neurites of the same cell (n= 5), only one neurite was found to be clearly sensitive to TTX and identified a posteriori as the axon. An alternative approach was also used, which involved local application of a Na+-free external solution. This essentially replicated the results obtained with TTX since local application of Na+-free solution on the soma blocked on average 52 ± 4% of the whole Na+ current (n= 11). Likewise, puffing Na+-free solution on neurites at the expected site of the AIS produced either substantial (40 ± 5%) or very little (4.8 ± 2%) Na+ current block depending on the neurites. Taken together, these data indicate that Na+ currents recorded in an inverse Na+ gradient result primarily from somatic Na+ channels, whereas in a standard Na+ gradient both somatic and axonal channels contribute to the whole-cell Na+ current.
Voltage dependence of activation and inactivation of somatic Na+ currents
We have analysed the biophysical properties of the somatic Na+ current to help assign them to their cloned counterparts. Whole-cell Na+ currents in inverse Na+ gradient were typically elicited by depolarizing steps from a holding potential of –80 mV preceded by a short prepulse to –100 mV to remove fast inactivation (Fig. 8A, left panel). The voltage dependence of activation was measured by plotting the normalized conductance against membrane potential and fitting the curve with the Boltzmann equation (Fig. 8B). The membrane potential at half-maximal activation (V0.5) was –23.4 ± 0.4 mV and the slope (k) of the curve was estimated to be 7 ± 0.5 mV (n= 11; DIV 7–15). The voltage dependence of fast inactivation was further investigated by holding the cells at prepulse potentials between –110 and –35 mV before stepping to the test potential (typically +20 mV) (Fig. 8A, right panel). Relatively short preconditioning pulses of 100 ms were used in this protocol to prevent entry of Na+ channels into slow inactivation (see below). Data fitted to a single Boltzmann function gave a midpoint of fast inactivation of –61.8 ± 0.2 mV and a slope factor of 5.5 ± 0.2 mV (n= 11) (Fig. 8B). Note that when cells at DIV 7–9 or DIV 12–15 were analysed separately, V0.5 values for activation and fast inactivation were not significantly different (activation: V0.5=–21.5 ± 0.5 mV versus–24.5 ± 0.6 mV; fast inactivation: V0.5=–61 ± 0.5 mV versus–62.3 ± 0.4 mV, respectively). The same holds for the Na+ current amplitude (280 ± 16 pA, n= 59 and 332 ± 27 pA, n= 48, respectively; P= 0.08; test step to +20 mV).
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fast of 3.4 and 7 ms and slow of 3 and 2 s, respectively) (Fig. 9B).
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30 Hz (data not shown). Detection of a persistent Na+ current
Based on current clamp studies, a persistent inward current (INaP) has been hypothesized in CG cells (D'Angelo et al. 1998). In a first series of experiments, we sought evidence for the presence of persistent Na+ currents under conditions of inverse Na+ gradients using slow depolarizing voltage ramps rising at a rate of 43 mV s–1 (3 s duration). These slow ramps were selected because they usually allowed full inactivation of the fast-inactivating Na+ current described above. Figure 11A shows currents evoked by such protocols in a representative CG cell, both in control conditions and in the presence of TTX (1 µM). Isolation of TTX-sensitive Na+ currents by offline digital subtraction showed no persistent components, consistent with the lack of INaP using the pulse protocol (cf. Figs 6B and 8A).
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An inhomogeneous voltage clamp is known to cause a potential gradient in remote electrotonic regions of neurones, which produces steady-state axial current flow into the soma, thereby increasing the persistent current measured somatically. To test this, we determined the reversal membrane potential of kainate (250 µM)-induced currents applied locally on the neurites at increasing distance from the soma (up to 80 µm, n= 4–9). No significant deviation of the reversal potential (range from –1.8 to +2.1 mV) was observed with distance from the cell body. Although these data argue that distal neurites are adequately clamped under steady-state conditions, they do not decisively prove that the persistent Na+ current is well clamped.
Because the persistent current was not observed under inverse Na+ gradient, where Na+ currents reflect primarily activity of somatic channels, we examined the issue of the localization of Na+ channels that generate INaP. Local application of TTX onto the soma did not cause significant block of INaP (–16.4 ± 2 pA and –13.8 ± 3 pA in control and upon TTX application, respectively; P= 0.1, paired t test; n= 5), whereas it did inhibit total transient Na+ current by 52 ± 5% in the same cells (Fig. 11E). In all these cells, TTX (1 µM) applied at the end of the experiment via the general perfusion system (i.e. superfusing both soma and neurites) fully abolished INaP suggesting that the persistent Na+ current results from channels mostly located in cell processes.
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Discussion |
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Differential distribution of Na+ channel subunits in cultured CG cells
From day 7 onwards, CG cells in our culture system had developed polarity as reflected by the segregation of axonal- and dendritic-specific cytoskeletal markers to morphologically distinct compartments. The formation of AIS was demonstrated by clustering of ankyrin-G, a prerequisite for the correct targeting and retention of multiple proteins at the AIS (Zhou et al. 1998; Bennett & Chen, 2001; Jenkins & Bennett, 2001). Using dual staining with a PanNav antibody, we showed that Na+ channels accumulate at the AIS, showing colocalization with ankyrin-G domains. It should be noted that immunoreactivity for PanNav was clearly observed only at sites of relatively high Na+ channel density, namely the AIS and the cytoplasm of the cell body, where they may represent newly synthesized pools of channels. Hence, though the amount of functional channel protein is likely to be related to the amount of cytoplasmic channel protein, Na+ channel immunodetection should not be taken as reflecting channels inserted into the plasma membrane, but rather subcellular regions of intense synthesis/trafficking or with limited diffusion (e.g. the AIS).
In line with immunodetection, Nav1.2 and Nav1.6 mRNAs were detected in cerebellar cultures by RT-PCR. Though study of cultured cells may not be indicative of the in vivo condition, our results are generally consistent with previous immunohistochemistry and in situ hybridization studies on rat cerebellum (Westenbroek et al. 1989; Felts et al. 1997; Schaller & Caldwell, 2000), indicating that the expression of Nav subunits in cultured CG cells provides a reasonable qualitative match to that in situ. Although Nav1.2 and Nav1.6 are simultaneously expressed in cultured CG cells, they show some differences in their subcellular expression pattern. Thus, at DIV 7–8, Nav1.2 was concentrated at AIS and colocalized with ankyrin-G and PanNav, suggesting that the high density of Na+ channels at these sites is mainly due to Nav1.2 subunits. At this stage, Nav1.6 was not confined to AIS but diffusely distributed throughout the cell body, dendrites and axons; accumulation of Nav1.6 at most AIS was evident by DIV 12. The rearrangement of Nav1.6 at AIS does not seem to result from an overall increase in expression since the appearance of Nav1.6 clusters was not paralleled by significant changes in maximum current amplitudes (–605 ± 35 pA at DIV 7–10, n= 46 versus–570 ± 55 pA at DIV 12–15, n= 13; P= 0.6) but rather reflects the new ability of initial segments to sequester Nav1.6. This sequential distribution in which Nav1.2 precedes Nav1.6 at AIS resembles the developmental expression of these subunits at AIS of Purkinje neurones and retinal ganglion cells (Jenkins & Bennett, 2001; Boiko et al. 2003) and in nodes of Ranvier (Boiko et al. 2001; Kaplan et al. 2001). Moreover, Nav1.6 accumulation was not associated with a reduction in Nav1.2 as we did not notice any appreciable change in the number of Nav1.2 clusters; however, whether the two subunits are spatially segregated within the initial segment could not be directly tested due to the lack of appropriate Na+ channel antibodies.
The differential distribution of Nav1.2 and Nav1.6 subunits also implies that within the non-conserved structural domains of these two highly related proteins are signals that direct channels to their specific subcellular compartments. Ankyrin-G is clearly required for clustering of Na+ channels at AIS, as Na+ channels failed to accumulate at AIS of ankyrin-G knockout mice (Zhou et al. 1998). Recently, it has been shown that the cytoplasmic loop connecting domains II and III (AIS motif) is an important determinant conferring compartmentalization of Nav1.2 at the AIS in rat hippocampal neurones (Garrido et al. 2003). Given that the AIS motif identified in Nav1.2 is highly conserved in Nav1.6, our findings suggest that the AIS signal may be necessary but not sufficient to localize Nav1.6 at AIS. A corollary therefore is that signals for ankyrin-G-based targeting of Nav1.6 is likely to involve additional Nav1.6-interacting protein(s) downstream of ankyrin-G in the pathway to formation of the AIS specialized domain.
Properties of rapidly inactivating and persistent Na+ currents
The biophysical properties of the Na+ current could be satisfactorily investigated by using inverse Na+ gradients, rendering the currents non-regenerative. The biophysical signature of the rapidly inactivating TTX-sensitive Na+ current in inverse gradient was undistinguishable from those of Nav1.2 or Nav1.6 when coexpressed with auxiliary ß subunits with normal Na+ gradient. With respect to the voltage dependence of activation and inactivation (fast and slow components), no major differences could be seen between the somatic Na+ current of CG cells and recombinant Nav1.2 or Nav1.6 or in acutely dissociated neurones (Li et al. 1992; West et al. 1992; Sarkar et al. 1995; Xie et al. 1995, 2001; Smith et al. 1998; Toib et al. 1998; Herzog et al. 2003). Likewise, the somatic current has fast repriming kinetics that ranged from 2 to 7 ms between –120 and –80 mV, which are comparable to those of Nav1.2 or Nav1.6 but are 2- to 10-fold faster than those of Nav1.3 and Nav1.7 (Cummins et al. 2001; Herzog et al. 2003). Taken together, our results suggest that the somatic Na+ current is dominated by fast gating channels, consistent with the contribution of Nav1.2 and/or Nav1.6 subunits, though we were unable to decipher the specific roles played by each subtype in the soma.
In addition to fast gating Na+ channels that play a critical role in transmitting high frequency action potentials, we observed that CG cells also exhibit a non-inactivating steady-state (‘persistent’) Na+ current. An important property of this current is its activation at voltages near the resting potential, suggesting a possible role in determining action potential threshold and in maintaining repetitive activity. Even though the window current of Na+ channels (see Fig. 8B) does appear to be capable of making a modest contribution to the persistent current at negative potentials, it is minute and its maximum amplitude would reach only 0.14% of the total Na+ conductance (as estimated from inverted Na+ currents), which is clearly incompatible with the size of the persistent current. Importantly, the persistent current was not observed in inverse Na+ gradient, where Na+ currents are thought to reflect activity of somatic channels, nor was it blocked by TTX applied onto the soma in normal Na+ gradient, suggesting that ‘persistent’ channels might be localized in the cell processes.
This interpretation, however, may be biased by poor neuritic voltage control, which can lead to the appearance of pseudo-persistent currents in the cell body (White et al. 1995). We have shown that CG cells are electronically compact at steady state (e.g. voltage can be controlled in remote neuritic segments), which makes them amenable to relatively ‘uniform’ space clamp. Naturally, the situation varies considerably under transient and steady-state conditions; that is injection of current from an eccentrically (somatic) placed patch electrode limits the rate at which the membrane capacitance of axons can be charged, thereby limiting the voltage clamp speed. This, along with high levels of ion channel expression in axons, results in poor control of fast Na+ currents in axons. In addition, the kainate experiments, although indicative of good voltage control of distal portions of neurites, do not provide evidence that remote neurites are clamped adequately while voltage-dependent Na+ currents are flowing, raising the possibility that steady-state axial current may be the prime source of INaP. Taken together, our present data therefore cannot determine whether INaP is caused by uncontrolled window Na+ currents occurring in unclamped portions of the neurones or by the activity of non-inactivating channels located in remote compartments (or both).
Previous studies have provided evidence for the preferential distribution of non-inactivating Na+ channels in the dendrites of hippocampal (Masukawa et al. 1991) and dorsal horn (Safronov et al. 1997) neurones. The existence of a persistent Na+ current of non-somatic origin has just been recently suggested in granular cells of rat cerebellar slices (Magistretti et al. 2004). The fact that Nav1.6 is commonly linked to the presence of persistent Na+ currents in a variety of neurones (Raman et al. 1997; Maurice et al. 2001), together with our finding that Nav1.6 is the main isoform expressed in dendrites of CG cells, support the proposal that Nav1.6 may contribute to the persistent currents. However, a better definition of the Na+ channel subtype(s) that generates INaP awaits further experiments.
In conclusion, our study establishes which Nav isoforms are expressed in CG cells and defines the subcellular distribution of each subtype. It also reveals the existence of functionally compartmentalized Na+ currents. These specialized Na+ channels, which include persistent and fast gating channels, are key determinants of CG cell excitability, making them reliable in terms of following sustained high-frequency stimulation and repetitive firing. Our model system therefore should prove useful for future studies addressing the molecular basis of Nav channel clustering as well as the functions of compartmentalized Na+ channels that shape the excitability of CG cells.
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3). Na+ currents evoked at –20 mV (C: control; W1 and W2: washouts 1 and 2). Scale bars: 2 ms, 200 pA.
) were plotted against membrane potential and fitted to single Boltzmann functions. Half-activation voltage for activation and fast inactivation are indicated. Each data point is the mean ±S.E.M. of 11 CG cells.
t= 1–200 ms) followed by a test step to +20 mV (Vt). For the family of current traces shown, recovery was studied at –100 mV and 
; –100 mV, 
). The interpulse duration was lengthened by 0.2, 1 or 10 ms between sweeps. The recovery subsequent to the initial delay can be fitted by a monoexponential function at –120 mV, which gave a time constant of 2.5 ms and by double exponential functions at –100 and –80 mV, which gave fast and slow time constants of 3.4 ms and 3 s (4% of Itotal) and 7 ms and 2 s (14% of Itotal), respectively. Note that because calculated slow time constants are longer that the actual duration of the voltage protocol, they should be considered as tentative. Each data point represents the mean ±S.E.M. of 11–12 cells.
