Neuronal differentiation involves both morphological and electrophysiological changes, which depend on calcium influx. Voltage-gated calcium channels (VGCCs) represent a major route for calcium entry into neurons. The recently cloned low-voltage-activated T-type calcium channels (T-channels) are the first class of VGCCs functionally expressed in most developing neurons, as well as in neuroblastoma cell lines, but their roles in neuronal development are yet unknown. Here, we document the part played by T-channels in neuronal differentiation. Using NG108–15, a cell line that recapitulates early steps of neuronal differentiation, we demonstrate that blocking T-currents by nickel, mibefradil, or the endogenous cannabinoid anandamide prevents neuritogenesis without affecting neurite outgrowth. Similar results were obtained using antisense oligodeoxynucleotides directed against the α1H T-channel subunit. Furthermore, we describe that inhibition of α1H T-channel activity impairs concomitantly, but independently, both high-voltage-activated calcium channel expression and neuritogenesis, providing strong evidence for a dual role of T-channels in both morphological and electrical changes at early stages of neuronal differentiation.
- T-type calcium channel
- HVA calcium channel
- neuroblastoma NG108–15 cell line
Neuroblast differentiation into neurons is associated with morphological and electrical changes. Major morphological changes are the appearance of axonal and dendritic extensions, collectively called neurites. The electrical changes mainly result from the expression of various voltage-gated channels, and many studies have indicated a correlation between calcium current expression and morphological differentiation (Bader et al., 1983). Voltage-gated calcium channels constitute a central component of signal transduction cascades (Berridge, 1998), and neuronal differentiation is altered when calcium influx is inhibited (Gu and Spitzer, 1997). In most neurons, including hippocampal, motor, and sensory neurons, low-voltage-activated T-type calcium currents (T-currents) appear first, whereas high-voltage-activated (HVA) calcium currents (especially L- and N-types) appear later in conjunction with neurite extension and dominate the total calcium influx in mature neurons (Nowycky et al., 1985; Yaari et al., 1987; Gottmann et al., 1988;McCobb et al.,1989). This sequence of events is also observed in a variety of neuroblastoma cell lines, including NG108–15 cells (Nirenberg et al., 1983; Hamprecht et al., 1985; Docherty et al., 1991).
Although the presence of T-currents at early stages of neuronal development suggests that they are involved in neuronal differentiation (Frischknecht and Randall, 1998), the functional significance of their early expression in neuroblasts remains to be determined. Cellular models that recapitulate early steps of neuronal differentiation allow such an investigation in a more defined way than in vitrocultures of undifferentiated neuroblasts collected from early embryos, which cannot be manipulated as a homogenous–synchronized cell population. It has been demonstrated that the neuroblastoma–glioma NG108–15 cell line, which expresses classical T-type calcium channels (Randall and Tsien, 1997), is a suitable model for investigating the mechanisms involved in neuronal development and differentiation, especially for the transition from neuroblast to neuron (Nirenberg et al., 1983; Hamprecht et al., 1985; Docherty et al., 1991). NG108–15 cells display a synchronized neuronal differentiation when cultivated in the presence of cAMP (Nirenberg et al., 1983), and differentiated NG108–15 cells exhibit well characterized morphological, electrophysiological, and pharmacological properties that are similar to neurons, including neurite outgrowth, synapse formation, and HVA calcium channel expression (Kleinman et al., 1988; Han et al., 1991;Kasai and Neher, 1992; Taussig et al., 1992). Using this cellular model, we describe in the present study that inhibition of T-channel activity impairs concomitantly, but independently, HVA channel expression and neuritogenesis, indicating a crucial role of T-channels encoded by the α1H subunit in both morphological and electrical changes during early stage of neuronal differentiation.
MATERIALS AND METHODS
Cell culture and assessment of neurite outgrowth. The NG108–15 cell line was used between 15 and 40 passages and was cultured as described previously (Chemin et al., 2001b). Cells were seeded at 250 cells/mm2 in 35 mm Petri dishes, and differentiation was induced by decreasing fetal calf serum (Eurobio, Les Ulis, France) in the medium to 1% and by adding 1 mm dibutyryl cAMP (Sigma, St. Louis, MO). Fourteen hours after plating, cells were examined under the microscope, and multiple random fields were examined. Neurite formation was quantified by scoring the percentage of cells possessing neurites over the total number of cells. Neurite extension (Table1) was evaluated by counting cells with short neurites (length, ≤1 cell body diameter), medium-sized neurites (length, >1 cell body diameter and <2 cell body diameter), and long neurites (length, >2 cell body diameter). Cell clumps containing more than five cells were not considered in the counting. These experiments were conducted using a double-blind strategy to avoid errors attributable to subjective judgment. Dunnett's multiple comparison test was used for statistical comparisons, and the values were expressed as mean ± SEM of n independent experiments.
Whole-cell patch-clamp recordings. Macroscopic currents were recorded by the whole-cell patch-clamp technique at room temperature using an Axopatch 200B amplifier (Axon Instruments, Foster City, CA). Data were acquired on a personal computer using the pClamp6 software suite (Axon Instruments). Current recordings were filtered at 5 kHz. Capacitance and R s were compensated by 85–95% using the whole-cell parameters of the Axopatch 200B amplifier. The extracellular solution contained (in mm): 2 CaCl2, 160 tetraethylammonium (TEA)-Cl, and 10 HEPES, pH 7.4 with TEA-OH. Pipettes made of borosilicate glass had a typical resistance of 1–3 MΩ when filled with a solution containing (in mm): 110 CsCl, 10 EGTA, 10 HEPES, 3 Mg-ATP, and 0.6 Na-GTP, pH 7.2 with CsOH. Whole-cell currents were analyzed as described previously (Chemin et al., 2001a), and Student'st tests or one-way ANOVA combined with a Student–Newman–Keuls post hoc test (for multiple comparisons) were used and considered significant with *p < 0.05, **p < 0.01, and ***p < 0.001 as indicated in the table and in figures. Results are presented as the mean ± SEM, and n is the number of cells used.
Bromodeoxyuridine labeling. For bromodeoxyuridine (BrdU) labeling, cells were plated at 250 cells/mm2 on 12 mm glass coverslips. After 3 hr, 10 μm BrdU (Roche Products, Hertforshire, UK) was added to the medium for 45 min. BrdU-treated cells were rinsed with PBS and fixed at room temperature for 10 min in a 3.7% paraformaldehyde solution (Sigma). Immunostainings and Hoechst 33258 nuclear dye (Sigma) labeling were performed as described previously (Chemin et al., 2000). Control experiments were performed in the absence of BrdU incorporation (data not shown). Digital images were acquired on a microscope (Leica, Nussloch, Germany) and further analyzed using Adobe Photoshop 4.0 (Adobe Systems, San Jose, CA). Percentage of proliferative cells was defined as the ratio of BrdU-positive cells over Hoechst-labeled cells (using multiple random microscope fields). One-way ANOVA combined with a Student–Newman–Keuls post hoc test were used to compare the different values and were considered significant atp < 0.05. The values were expressed as mean ± SEM, and n is the number of independent experiments.
Reverse transcription-PCR and Southern blotting. RNA from undifferentiated and differentiated NG108–15 cells, as well as from rat and mouse brains, were isolated using Trizol (Invitrogen, Gaithersburg, MD) according to the protocol of the manufacturer. Poly(A+) RNA was separated using an mRNA purification kit (Dynal, Great Neck, NY). Reverse transcription (RT) was performed with superscript II reverse transcriptase primed with random hexamers using the superscript first-strand synthesis system for RT-PCR (Invitrogen). PCR primers were designed for the analysis of the three T-channel α1 subunits. The α1G primers 5′-GCTCTTTACTTCATCGCCCTC-3′ (forward) and 5′-CCTCATCATTGTCATCATCCCC-3′ (reverse) generated a 795 bp fragment. The α1H primers 5′-GGACGGACACAACGTGAG-3′ (forward) and 5′-GTTCCAGTTGATGCAGGC-3′ (reverse) generated a 459 bp fragment. The α1I primers 5′-ATGCTGGTGATCCTGCTGAAC-3′ (forward) and 5′-GCACGCGGTTGATGGCTTTGAG-3′ (reverse) generated a 300 bp fragment. Southern blotting was performed using standard methods (Sambrook et al., 1989). Briefly, 10 pmol of oligonucleotides matching an internal sequence of each PCR product (α1G, 5′-GCCAAGAGTTCCTTTGACCT-3′; α1H, 5′-GGAACAACAACCTGACCTTC-3′; and α1I, 5′-TGCAAGATCCTGCAGGTCTT-3′) were labeled with [γ-32P]ATP using T4 polynucleotide kinase. Membranes were hybridized in Express-Hyb buffer (Clontech, Cambridge, UK) overnight (42°C) and further revealed by autoradiography. Negative controls for RT-PCR were obtained using reverse transcriptase reaction of mRNA samples in which random hexamers were omitted (negative RT; see Fig.2), and positive controls were made using mRNA obtained from rat brain, as well as from mouse brain.
Transfection protocols. Antisense (AS) oligodeoxynucleotides against α1G mRNA [5′-CCTCATCATTGTCATCATCC-3′ (AS-α1G)], AS against α1H subunit mRNA [5′-GTTCCAGTTGATGCAGGC-3′ (AS-α1H)], and AS against α1I subunit mRNA [5′-GCACGCGGTTGATGGCTTTG-3′ (AS-α1I)] were used in transfection experiments. A scramble oligodeoxynucleotide 5′-GTAGCATGATCGGTGCTC-3′ (Scramble in figure legends) was used as control. Two hours before transfection, cells were plated at ∼50% confluence in 35 mm Petri dishes. A standard transfection procedure was performed using Fugene 6 transfection reagent (Roche Products) with 500 nm/dish of AS solution. Three days later, cells were plated at 250 cells/mm2 in 35 mm Petri dishes, and differentiation was induced as described above.
Calcium currents in undifferentiated and differentiated NG108–15 cells
In undifferentiated conditions, NG108–15 cells were preferentially organized in clusters and displayed no neurite extension (Fig. 1 A). In this condition, T-type calcium currents were observed in 95% of the cells (T-current density, 1.5 ± 0.3 pA/pF; n = 40 cells) (Fig. 1 C,E), whereas HVA currents were mostly absent or very weak (HVA current density, 0.22 ± 0.11 pA/pF; n = 40 cells). Three to 6 d after differentiation, cells exhibited neurites (Fig. 1 B), and, consequently, membrane capacitance was larger [53.7 ± 4.3 pF (n = 40 cells) and 91.3 ± 12.1 pF (n = 30 cells) for undifferentiated and differentiated cells, respectively]. After differentiation, the total calcium current density was significantly increased because of the expression of HVA currents (HVA current density, 4.9 ± 0.9 pA/pF; n= 30 cells) (Fig. 1 D,F). In differentiated cells, the HVA current composition was evaluated using specific pharmacological agents (Fig. 1 F,inset). Calcium currents were strongly sensitive to ω-conotoxin-GVIA (percentage of block, 43 ± 9%;n = 12 cells) and to nitrendipine (percentage of block, 34 ± 7%; n = 12 cells), which block N- and L-type currents, respectively. A smaller fraction of the calcium current was blocked by ω-agatoxin-IVB, a blocker of P/Q channels (percentage of block, 11 ± 4%; n = 12 cells). Also, a fraction of this current (R-type, 12 ± 7%;n = 12 cells) was insensitive to all of the previously described blockers, as well as to 30 μm nickel and SNX-482 (Newcomb et al., 1998). In contrast, no change in T-current density was observed after differentiation (1.5 ± 0.2 pA/pF;n = 30 cells), and the electrophysiological properties of T-currents were similar in undifferentiated and differentiated cells (data not shown), suggesting that differentiation does not affect T-channel expression in NG108–15 cells.
Molecular characterization of the α1H nickel-sensitive T-currents
To resolve the molecular nature of the T-channels expressed in NG108–15 cells, we performed RT-PCR experiments followed by Southern blotting analysis (see Materials and Methods). A strong RT-PCR signal for the α1H subunit mRNA was found in both undifferentiated and differentiated NG108–15 cells, whereas no α1G and only small amounts of α1I mRNA were detected (Fig.2 A). AS oligodeoxynucleotides were designed against each T-channel subunit mRNA to knock-down T-channel expression. The percentage of transfection was ∼70%, as measured with AS coupled to FITC. AS-α1H significantly decreased the T-current amplitude in undifferentiated (I/I control, 0.45 ± 0.06; n = 15 cells) as well as in differentiated NG108–15 cells (I/I control, 0.4 ± 0.1; n = 12 cells; data not shown), whereas AS-α1G, AS-α1I, and a scramble AS had no effect (Fig. 2 B). Moreover, we observed no difference in the electrophysiological characteristics between the residual T-currents after AS-α1H transfection and the control T-currents, including steady-state activation and inactivation properties and activation and inactivation kinetics (data not shown), further indicating that only α1H channels are functionally expressed in NG108–15 cells. In addition, micromolar concentrations of nickel ions (Ni2+) inhibited T-currents in undifferentiated and differentiated NG108–15 cells [IC50, 4.1 ± 0.2 μm (n = 20 cells) and 3.8 ± 0.4 μm (n = 20 cells), respectively] (Fig. 2 C). For a concentration of 30 μm Ni2+ that completely blocked T-currents (percentage of block, 96 ± 1%;n = 20 cells), HVA currents were only weakly affected (percentage of block, 12 ± 4%; n = 20 cells), even after 30 min of Ni2+ perfusion, and this inhibition was fully reversible (Fig. 2 D).
T-Currents promote differentiation and neuritogenesis
We next investigated whether T-currents could promote neuritogenesis and neurite extension. After only 14 hr in differentiating conditions, we observed a very large increase in the number of cells displaying neurites [from 6.2 ± 0.6% (n = 8 experiments) to 65 ± 2% (n = 30 experiments), for undifferentiated and differentiated cells, respectively]. In contrast, differentiation occurring in the presence of 30 μmNi2+ dramatically reduced neuritogenesis. Figure 3 shows that treatment with the T-channel blockers Ni2+ and mibefradil significantly decreased the number of cells with neurites [percentage of cells with neurites/control, 65 ± 2% (n = 20 experiments) and 78 ± 3% (n = 15 experiments), respectively], whereas HVA blockers had no effect. Similarly, 1 μm methanandamide, the nonhydrolyzable analog of the endocannabinoid anandamide, recently identified as an endogenous cannabinoid (CB) receptor ligand that also directly inhibits T-currents (Chemin et al., 2001c), decreased the number of cells with neurites in the presence of 100 nm of the CB1 receptor antagonist SR141716A (percentage of cells with neurites/control, 41 ± 6%; n = 8 experiments). Reducing free external Ca2+ with EGTA (see figure legend) inhibited neuritogenesis (percentage of cells with neurites/control, 45 ± 5%; n = 8 experiments), and no additional effect of Ni2+ was observed in these conditions (Fig. 3). Similarly, reducing free internal Ca2+ with 10 μm BAPTA-AM inhibited neuritogenesis (percentage of cells with neurites/control, 17 ± 1%;n = 5 experiments), and no additional effect of Ni2+ was observed in these conditions (Fig. 3). Conversely, increasing free internal Ca2+ with 100 nmionomycin increased neuritogenesis (percentage of cells with neurites/control, 115 ± 1%; n = 5 experiments). In this later case, it is worth noting that Ni2+ inhibited significantly neuritogenesis (Fig. 3). Finally, AS-α1H specifically decreased the number of cells exhibiting neurites (percentage of cells with neurites/control, 74 ± 4%; n = 16 experiments) (Fig. 3). Interestingly, neither AS-α1H nor Ni2+ affected neurite length (Table 1), suggesting that T-currents are important for initiation of neuritogenesis but are not involved in neurite extension. In contrast, removing cAMP from the differentiating medium affected neuritogenesis (percentage of cells with neurites/control, 60 ± 10%;n = 8 experiments; data not shown), as well as neurite extension (Table 1).
Considering that α1H T-currents appear to have an early role in NG108–15 differentiation, we also explored whether T-currents could control the proliferation of NG108–15 cells after 4 hr of differentiation. For this purpose, we performed labeling with BrdU, a thymidine analog that is incorporated specifically into cells in S-phase (Fig. 4). After 4 hr of differentiation, the percentage of BrdU-positive cells decreased (percentage of positive cells/undifferentiated cells, 57 ± 4%;n = 9 experiments), indicating that differentiation was initiated. In contrast, in the presence of 30 μm Ni2+, cell proliferation was not significantly reduced compared with undifferentiated cells (percentage of positive cells/undifferentiated cells, 92 ± 7%; n = 9 experiments). Overall, these data suggest an important role of T-currents in triggering the onset of differentiation, leading to morphological changes.
Block of T-currents promotes inhibition of HVA calcium current expression
We next investigated whether T-currents could also contribute to changes in the expression of HVA calcium channels, a hallmark of neuronal differentiation. Interestingly, a strong correlation between HVA and T-current densities was observed (r = 0.8;p < 0.001; n = 80 cells) (Fig.5 A), suggesting that T-currents may regulate expression of HVA channels. The correlation between HVA and T-current densities was abolished with 30 μm Ni2+ treatment during differentiation (r = 0.14; p > 0.05; n = 50 cells) (Fig. 5 B). In this case (Fig. 5 C,D), a significant decrease in HVA currents was observed, because HVA current density was 5.1 ± 1.9 pA/pF (n = 20 cells) in control cells and 1.1 ± 0.3 pA/pF (n = 28 cells) in Ni2+-treated cells after Ni2+ washout (p < 0.001). In addition, we observed no change in the pharmacological properties of the HVA current after Ni2+treatment (n = 15 cells; data not shown), suggesting that functional expression of each type of HVA channels was similarly inhibited. In contrast, T-current density was not affected by chronic Ni2+ treatment [1.21 ± 0.22 pA/pF (n = 20 cells) and 1.12 ± 0.38 pA/pF (n = 28 cells) for control and Ni2+-treated cells, respectively] (Fig.5 C), indicating that inhibition of HVA currents was not attributable to inadequate removal of Ni2+ions from the medium. More importantly, transfection of AS-α1H significantly decreased HVA currents (HVA current density/control, 0.36 ± 0.16 pA/pF; n = 12 cells), similar to chronic Ni2+ treatment (Fig.5 D). Because the inhibition of HVA current expression could be a consequence of the decrease in the number of cells displaying neurites, we next analyzed HVA current expression with respect to the neurite length. Chronic Ni2+ treatment decreased neuritogenesis without affecting neurite extension, and ∼45% of the cells displayed long neurites after 4–6 d of differentiation. In control cells, HVA current density increased with the neurite length [HVA current density, 0.9 ± 0.4 pA/pF (n = 10 cells) and 9.2 ± 2.9 pA/pF (n = 10 cells) in cells with short and long neurites, respectively] (Fig. 5 E), whereas T-current density did not [1.5 ± 0.3 pA/pF (n = 10 cells) in cells with short neurites and 1.4 ± 0.3 pA/pF (n = 10 cells) in cells with long neurites; data not shown]. In contrast, no enhancement of HVA currents was observed in Ni2+-treated cells [HVA current density, 1.1 ± 0.3 pA/pF (n = 15 cells) and 1.2 ± 0.5 pA/pF (n = 13 cells) in cells with short and long neurites, respectively] (Fig. 5 E). Altogether, these data demonstrate that T-currents are important for the expression of HVA calcium conductances independently from the induction of neuritogenesis.
T-type channels are the first voltage-gated calcium channels expressed in developing neurons. Although it is widely admitted that calcium influx plays a crucial role in neuronal differentiation, the role of T-channels in this process is unknown. Using the neuronal model NG108–15 cell line, we provide new findings indicating that T-currents encoded by the α1H subunit contribute to morphological and electrical changes occurring during neuronal differentiation. We report that T-currents are involved in the onset of the differentiation process, leading to the arrest of cell proliferation and the induction of neuritogenesis. In addition, the data reveal that T-currents are involved in the expression of HVA calcium currents, indicating a crucial role of T-channels in neuronal differentiation.
Our study demonstrates that NG108–15 cells exhibit both low-voltage-activated T-currents and HVA currents and that these two classes of channels are differentially modulated during neuronal differentiation. Undifferentiated NG108–15 cells have no neurites and display calcium currents of small amplitude that are entirely T-currents. Differentiation of NG108–15 cells affected neither the density nor the electrophysiological and pharmacological properties of T-currents. In contrast, we found a striking change in the functional expression of HVA currents during differentiation, which is in agreement with previous studies (Freedman et al., 1984; Kasai and Neher, 1992; Lukyanetz, 1998). The increase in HVA current expression in differentiating NG108–15 cells occurs in concert with neuritogenesis, similar to that observed in many neurons (Nowycky et al., 1985; Yaari et al., 1987; Gottmann et al., 1988; McCobb et al.,1989). The presence of T-currents in both undifferentiated NG108–15 cells and every cell with neurites designates T-channels as potential actors in neuronal differentiation. The recent characterization of three genes coding for T-type α1 subunits [α1G (CaV3.1), α1H (CaV3.2), and α1I (CaV3.3)] (Cribbs et al., 1998;Perez-Reyes et al., 1998; Klugbauer et al., 1999; Lee et al. 1999a;Williams et al., 1999; Monteil et al., 2000a,b; McRory et al., 2001) has enabled the molecular investigation of T-channel functions. Both in undifferentiated and differentiated NG108–15 cells, T-currents are very sensitive to Ni2+ and have characteristics similar to those described in neurons (Carbone and Lux, 1984; Armstrong and Matteson, 1985; Nowycky et al., 1985; Fox et al., 1987; Randall and Tsien, 1997). Overall, RT-PCR experiments and Ni2+ sensitivity of T-currents (Lee et al., 1999b) both indicate that these channels in NG108–15 cells comprise the α1H subunit. The use of pharmacological agents and antisense strategies allowed us to demonstrate that α1H T-currents contribute to morphological and electrical changes during neuronal differentiation. During differentiation, NG108–15 cells rapidly lose their ability to proliferate, and this can be prevented if α1H T-currents are blocked. More importantly, blockade of T-current decreases the number of cells expressing neurites. Such a reduction in neuritogenesis depends on the Ca2+ influx and is observed with a variety of T-channel blockers, including anandamide, which directly block T-currents independently of CB receptors (Chemin et al., 2001c). Interestingly, our data indicate that there is no correlation between neurite length and T-current density and that the block of T-currents does not affect neurite outgrowth. Because appearance of HVA currents and neuritogenesis are concomitant (and possibly associated) events, T-channel blockade could also affect HVA channel expression. Indeed, we found a strong correlation between T-current and HVA current amplitudes, and the chronic block of T-currents inhibits expression of HVA currents. Nevertheless, inhibition of HVA current expression cannot simply be explained by the decrease in neuritogenesis because it is observed even in cells with long neurites. Conversely, inhibition of HVA current expression does not account for the inhibition of neuritogenesis because pharmacological blockade of HVA currents does not affect the number cells expressing neurites. Altogether, these data demonstrate that α1H T-currents play a central role in the early onset of morphological differentiation, as well as in the maturation of calcium conductances of the NG108–15 cell line.
To our knowledge, we are aware of no other study that either disproves or conclusively demonstrates a role played by T-channels in neuronal differentiation. NG108–15 is a cholinergic cell line (Hamprecht et al., 1985; Docherty et al., 1991) in which T-current properties are similar to peripheral neurons (Carbone and Lux, 1984), which also express the α1H subunit (Talley et al., 1999). Two other cholinergic cell lines, SN56 (Kushmerick et al., 2001) and N1E-115 (Lievano et al., 1994), also exhibit Ni2+-sensitive (α1H-related) T-currents that precede neuritogenesis and HVA channel expression, suggesting that α1H T-channels could play a specific role in neuronal differentiation of the cholinergic system. Ni2+-sensitive T-currents are also present at early stages in the peripheral nervous system, including dorsal root ganglion neurons (Gottmann et al., 1988) and motor neurons (McCobb et al.,1989; Mynlieff and Beam, 1992). Similarly, dot blot analysis of human brain mRNA showed that the α1H subunit is expressed at higher level during fetal development (A. Monteil, P. Lory, and J. Nargeot, unpublished results), and Ni2+-sensitive T-currents have been recorded in floor plate cells of the developing CNS (Frischknecht and Randall, 1998). Therefore, in the light of the results described here, the implication of α1H T-channels in neuronal differentiation should be analyzed in the peripheral nervous system, as well as in the CNS, in regions such as the hippocampus (Yaari et al., 1987). Ni2+-sensitive α1H T-channels are likely to mediate differentiation of a variety of cell types, because it was shown recently that α1H T-channels promote differentiation (fusion) of human myoblasts (Bijlenga et al., 2000). In addition, in the human prostate cancer epithelial LNCaP cells, there is an elevated expression of α1H T-channels during cell differentiation, which is likely to facilitate neurite-like lengthening (Mariot et al., 2002). An important question that now needs to be addressed is how α1H T-channels contribute to differentiation and HVA channel expression. For skeletal muscle differentiation, it has been shown that α1H T-window currents could increase resting intracellular Ca2+ in fusing myoblasts (Bijlenga et al., 2000), a property that was also found for recombinant α1H T-channels overexpressed in HEK293 cells (Chemin et al., 2000). Although our data demonstrate that entry of Ca2+ through T-channels plays a crucial role in neuronal differentiation, we did not find any significant difference in intracellular Ca2+ concentration when comparing NG108–15 cells treated or not with Ni2+using the Ca2+ indicator fura-2 (data not shown). Interestingly, although a rise of intracellular calcium seems crucial for T-channel-induced neuritogenesis (as assessed by the use of BAPTA-AM and ionomycin), block of T-channels in ionomycin-treated cells still inhibited neurite emergence. These data might be explained by the presence, close to T-channels, of Ca2+buffer proteins that are capable of selectively transducing this Ca2+ signal, thus allowing neuritogenesis. Calcineurin and calmodulin are involved in neuronal differentiation (Goshima et al., 1993; Chang et al., 1995; Lautermilch and Spitzer, 2000) and are highly expressed in NG108–15 cells (Komeima et al., 2000; Higashida et al., 2001). It will therefore be of great interest to examine whether Ca2+ influx via α1H T-channels can modulate differentiation processes through these pathways.
This work was supported by Centre National de la Recherche Scientifique, the Association pour la Recherche contre le Cancer, and the Ligue contre le Cancer. We thank Drs. M. Mangoni, E. Bourinet, C. Altier, S. Barrère, D. Fisher, S. Jarvis, and S. Dubel for helpful discussions and comments on this manuscript. We are grateful to Dr. F. A. Rassendren for technical help with Southern blot techniques, Dr. G. Dayanithi (Institut National de la Santé et de la Recherche Médicale U432) for the gift of SNX-482, and Dr. I. A. Lefevre for critical reading of this manuscript.
Correspondence should be addressed to Philippe Lory, Institut de Génétique Humaine, Centre National de la Recherche Scientifique, Unité Propre de Recherche 1142, 141 rue de la Cardonille, F-34396 Montpellier cedex 05, France. E-mail:.