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1Center for Neuroscience and 2Program in Communication Science, University of California Davis, Davis, California; 3Department of Nephrology, Xijing Hospital, The Fourth Military Medical University, Xi'an, China; 4Department of Otorhinolaryngology, University of Pennsylvania, Philadelphia, Pennsylvania; 5Department of Physiology and Biophysics, University of Miami School of Medicine, Miami, Florida; and 6School of Biological Sciences, University of Sussex, Brighton, United Kingdom
Submitted 26 June 2008; accepted in final form 12 August 2008
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ABSTRACT |
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INTRODUCTION |
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; Bao et al. 2003; Fuchs et al. 1990; Hudspeth and Lewis 1988; Kollmar et al. 1997; Michna et al. 2003; Rodriguez-Contreras and Yamoah 2001, 2003; Rodriguez-Contreras et al. 2002; Schnee and Ricci 2003; Zidanic and Fuchs 1995). The role of the Cav1.3 in hair cells is underpinned by the evidence that Cav1.3-deficient (Cav1.3–/–) mice are deaf, demonstrating a significant reduction in whole cell Ca2+ current in inner hair cells (IHCs) (Dou et al. 2004; Platzer et al. 2000). However, a residual Ca2+ current remains in IHCs and outer hair cells (OHCs) of Cav1.3–/– mice. In these mice no apparent vestibular disorder was observed (Dou et al. 2004), suggesting that cochlear and vestibular hair cells may express multiple VGCCs. Moreover, data from Cav1.3–/– mice strongly suggest that non-Cav1.3 channel currents suffice to maintain normal cellular morphology and function in vestibular sensory organs (Dou et al. 2004). Recent reports have demonstrated the fleeting expression of T-type currents in the chicken basilar papilla, during hair-cell development and regeneration (Levic et al. 2007). These findings are essential to uncover the mechanism of hair-cell regeneration and restoration of hearing. Thus identification of the diverse Ca2+ channel subtypes in hair cells is key to our understanding of the multiple Ca2+-dependent functions.
Previous studies have used pharmacological, immunohistological, and functional techniques to identify non-Cav1.3 channels in hair cells. These studies have suggested the presence of Cav2.2 channels in hair cells of the frog saccule (Rodriguez-Contreras and Yamoah 2001; Su et al. 1995). Similarly, the Cav2.3 channels have been identified in hair cells within the vestibule (Martini et al. 2000). Furthermore, earlier recordings from mammalian vestibular hair cells showed that a portion of the Ca2+ channel current was derived from a transient low-voltage–activated (LVA) current, unlike a typical Cav2.x current (Rennie and Ashmore 1991). Additionally, a recent report has identified T-type currents in rat OHCs (Inagaki et al. 2008). This observation, together with another recent report of a transient Ca2+ current in the chicken basilar papilla (albeit transitory), suggests that Cav3.x channels are possible candidates of a hair-cell non-Cav1.3 channel subtype (Levic et al. 2007). The role of T-type channels in tonically active hair cells remains undetermined. Even more perplexing is the fact that the non-Cav3.1 currents in hair cells do not always exhibit exemplary fast inactivation of T-type currents (see Levic et al. 2007). Here, we have cloned a distinct inner ear Cav3.1 channel, derived from alternative splicing of the mouse brain Cav3.1 isoform. Functional expression of the channel yielded currents with the characteristic T-type current profile. Unlike most multi-ion channels, however, the T-type channel did not exhibit an anomalous mole fraction effect (AMFE) and the underlying mechanisms for its aberrant property were discussed. Moreover, we have demonstrated the differential expression of the Cav3.1 channel, during development in the sensory and nonsensory epithelia of the mouse cochlea and vestibule.
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METHODS |
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Mice were housed and killed using approved protocols by the University of California, Davis, IACUC committee on Animal Research Services. Total RNA was isolated from microdissected mouse cochlear and utricular tissues, using the RNeasy Mini kit (Qiagen). The first strand of complementary (c)DNA was synthesized from 1 µg of total RNA using Arrayscript reverse transcriptase (Ambion) and random primers by incubating initially at room temperature for 5 min and then at 42°C for 2 h. The reaction was terminated by heating at 70°C for 15 min. Finally, the cDNAs were treated with RNaseH for 20 min at 37°C before further use. PCR was performed using primer pairs corresponding to known mouse Ca2+ channel sequences as subsequently indicated. For Cav1.3, sense: GCT CAA TGG CAG TGT GTG TC and antisense: GTC TGG CTC CTC GTC ACTG (239–258 and 350–332 of AK018426 [GenBank] , respectively), and for Cav3.1 sense: AGG CCA AGA GTT CCT TTG AC and antisense: AGC CGA CTT GCC ATT ACAG (3611–3630 and 3705–3723 of NM-009783.1, respectively). The PCR reaction was initiated by denaturation at 94°C for 1 min, followed by 50 cycles of amplification consisting of 30 s at 94°C, 30 s at 52°C, and 45 s at 72°C. The resultant PCR product was separated on a 2% agarose gel and purified using a Qiaquick gel extraction column (Qiagen). The purified PCR products were sequenced to elucidate the sequence identities.
Molecular cloning of full-length Cav3.1 channels
A mouse cochlea 
ZAP-cDNA library of DBA mice was screened using multiple probes. The probes were PCR-amplified fragments; probes P1, P2, P3, and P4 corresponding to the regions of nucleotides 32–951, 2276–3010, 3248–3942, and 6165–6890 for the mouse Cav3.1 cDNA (NM_009783.1), respectively. The probes were labeled by random priming with digoxigenin-UTP and approximately 1.6 million individual clones of the cDNA library were screened following the manufacturer's instructions (Roche Molecular Biochemicals, Indianapolis, IN). Sequences of clones obtained from the cDNA library were determined by double-strand sequencing. For expression studies, the entire open reading frame of the Cav3.1 channel was cloned into a pNLR-XV vector, in which the channel was flanked by the 5'- and 3'-untranslated regions of a Xenopus β-globin gene (Nie et al. 2004). From the resulting expression plasmid, cRNAs of the Cav3.1 channel were transcribed in vitro using T7 RNA polymerase and injected into stage V–VI oocytes as described previously (Nie et al. 2004).
Immunohistochemistry
A rabbit polyclonal antibody for Cav3.1 (Alomone Labs, Jerusalem, Israel) generated against the cytoplasmic N-terminus of the T-type Ca2+ channel Cav3.1 was used. Embryos were harvested from timed pregnant 129Sv females at E12, E15, and E18. The embryo (E12) or whole heads (E15 and E18) were fixed for 2 h at room temperature in 4% paraformaldehyde in 0.1 M phosphate buffer (PB). Inner ear tissues were also harvested from postnatal day 0 (P0), P3, P6, P8, P12, P14, P18, P21, and adult 129Sv mice. At least three animals were examined for each time point. For P0 mice, entire heads were immersed in 4% paraformaldehyde for 2 h at room temperature. For the remainder of the ages, the sedated (Avertin [2,2,2-tribromethanol], 300 µg/gm body weight, administered intraperitoneally) mice were transcardially perfused with phosphate-buffered saline (PBS) followed by 4% paraformaldehyde in 0.1 M PB. The temporal bones were isolated and the cochlea was perfused via the round window, then immersed in the fixative for 60 min. Following fixation the cochleae were decalcified (0.12 M EDTA, pH 7.0, 24 h, 23°C), dehydrated in a graded ethanol series, embedded in paraffin (Paraplast), and sectioned (5 µm) in the mid-modiolar plane. Sections were deparaffinized, rehydrated, and equilibrated in PBS with 5% nonimmune normal goat serum (NGS, Vector Laboratories, Burlingame, CA) for 30 min and reacted (1:250) overnight at 4°C with anti-Cav3.1 primary antibody. The PBS-rinsed sections were incubated (30 min, 23°C) in biotinylated goat anti-rabbit IgG (1:200) then rinsed and treated with Vectastain ABC reagent (Vector Labs) for 30 min. Sites of bound primary antibody were visualized by development in a 3,3'-diaminobenzidine (DAB)–H2O2 substrate medium (Fast DAB tablets; Sigma Chemical, St. Louis, MO). Images were captured using an Olympus BH-2 microscope fitted with a SPOT RT-KE CCD camera and image-analysis software (Diagnostic Instruments, Sterling Heights, MI). Final figures were assembled using Adobe Photoshop and Illustrator software (Adobe Systems, San Jose, CA).
Electrophysiological recordings
Two-electrode voltage-clamp experiments were carried out with the oocyte clamp amplifier (OC-725C; Warner Instruments, Hamden, CT). The microelectrodes were filled with 3 M KCl. Oocytes were bathed in a solution that contained (in mM) 40 NaCl, 56 N-methyl-D-glucamine (NMG), 2 KCl, 2–65 BaCl2, 5 HEPES (N-[2-hydroxyethyl] piperazine-N'-[2-ethanesulfonic acid]), and 1 niflumic acid to block endogenous chloride currents in oocytes (pH adjusted to 7.4 with NaOH). To maintain tonicity of the external solution (280 mosmol), the concentration of NaCl/NMG was adjusted accordingly, as the BaCl2/CaCl2/SrCl2 concentration was increased from 2 to 65 mM. The current was activated using different voltage-clamp protocols as described in RESULTS. Liquid junction potentials were measured and corrected as described previously (Rodriguez-Contreras et al. 2002).
Standard patch-clamp recording techniques were used to record single-channel currents from Xenopus oocytes, which expressed a robust current (2–3 nA). An Axopatch 200B amplifier was used (Molecular Devices, Union City, CA). The amplitude histogram at a given test potential was generated. Leak-subtracted current recordings were idealized with a half-height criterion. Patch electrodes contained (in mM): 70 NMG-Cl, 40 KCl, 20 BaCl2/CaCl2, 5 HEPES, and 10 D-glucose, pH 7.4 (KOH). The bath solution contained (in mM): 140 KCl, 4 NaCl, 1 CaCl2, 0.5 MgCl2, 5 HEPES, and 10 D-glucose, pH 7.4 (KOH). The Q-software was used for single-channel analysis, as described previously (Rodriguez-Contreras and Yamoah 2001). Reagents were purchased from Sigma Chemical, unless specified otherwise. Experiments were carried out at room temperature (
21°C). Analyses were carried out using custom-written software, linked to Origin software (MicroCal, Northampton, MA). We expressed pooled data as means ± SD.
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RESULTS |
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The expression of non-Cav1.3 channels in the mouse inner ear was first examined by RT-PCR using Cav3.1 channel-specific primers. Messenger RNA of Cav3.1 channels was detected in both the cochlea and utricle of CBA mice (Fig. 1A). However, the levels of Cav3.1 messenger RNA were qualitatively low compared with that of Cav1.3 channels. Motivated by this evidence, together with the existence of a transient LVA Ca2+ current in mammalian hair cells (Inagaki et al. 2008; Rennie and Ashmore 1991) and recent evidence demonstrating the presence of T-type Ca2+ currents in the developing and regenerating hair cells, we cloned a full-length Cav3.1 channel from the mouse cochlea. Using a cDNA library, we used multiple probes, which correspond to highly conserved regions of the known mouse Cav3.1 cDNA. In all, 25 clones were obtained from library screening and 6 of them were fully sequenced. Moreover, the remaining 19 were sequenced and found to be redundant. The full-length cDNA sequence of the inner ear Cav3.1 channel was determined from the assembly of overlapping clones. In addition, we performed long-range PCR to confirm the right connection of different clones. As a result, the Cav3.1 cDNA obtained from the mouse inner ear was 7281 base pairs in length (Accession number DQ317412), derived from alternative splicing that excluded exon14, exon25A, exon34, and exon35 (Fig. 1, B–D). This splice variant of mouse Cav3.1 has the same exon utilization as that of the human Cav3.1 isoform 11 (GenBank accession number NM_198382.1). Furthermore, the splice variant had not been identified in mice. The splice variant encodes a putative protein of 2265 amino acids (aas) with a typical T-type channel structure: four functional domains of six transmembrane segments and a pore region. The Cav3.1 isoform shares 94% sequence similarity with its human counterpart. As in many other cases, the highest sequence similarities appear at the transmembrane domains and the pore-forming regions (98%), whereas the lowest was at the intracellular N- and C-termini (84%). The functional impact of alternative splicing on the property of other isoforms of Cav3.1 channels has been reported previously (Chemin et al. 2002a). Thus we examined the properties of the newly identified Cav3.1 channels in the inner ear.
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The inner ear Cav3.1 channel was subcloned into the pNLR-XV vector and expressed in Xenopus oocytes. Features of the Cav3.1 channel current were determined using a two-electrode voltage-clamp approach (Fig. 2). In contrast to saline-injected oocytes, oocytes injected with about 25 nM Cav3.1 mRNA yielded inward Ba2+ currents (Fig. 2A). The Ba2+ current of the inner ear Cav3.1 channel showed voltage-dependent decay and the magnitude of the current increased as the external Ba2+ concentration was raised (Fig. 2, A and B). Moreover, the current appeared to saturate at Ba2+ concentrations >30 mM, which was also the case for Ca2+ and Sr2+ currents (Fig. 2, B and C). Data on the relations between current magnitudes and external divalent cation concentrations were fitted with a Langmuir isotherm. The estimated apparent KD values were markedly similar (Fig. 2C), as compared with the reported KD of Cav1.3 in hair cells and chick cilary ganglion neurons (Church and Stanley 1996; Rodriguez-Contreras et al. 2002). It should be noted that physiological Ca2+ concentration (1–2 mM) lies within the most sensitive range of the saturation curve (Fig. 2C), suggesting that the channel operates at the optimal concentration of Ca2+. The shift in the peak voltage and voltage-dependent activation of the current, as seen in the current–voltage (I–V) relation, was consistent with surface charge screening effects resulting from increased divalent cations (Hagiwara and Ohmori 1982; Hagiwara and Takahashi 1967; Rodriguez-Contreras and Yamoah 2003; Zhou and Jones 1995). These results were in accord with previous reports of T-type Ca2+ currents (Huguenard 1996; Huguenard et al. 1993; Perez-Reyes 1998, 2003).
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; Hille 2001). The half-maximal activation voltage and the slope factors of the steady-state activation curves for Ba2+, Ca2+, and Sr2+ currents were –33.2 ± 0.6, –35.5 ± 0.6, and –38.5 ± 0.6 mV; and 6.4 ± 0.5, 6.3 ± 0.6, and 5.6 ± 0.5 mV (n = 17), respectively (Fig. 3C). Using a standard steady-state inactivation protocol, a 500-ms conditioning pulse was applied to different activation potentials between –80 and 0 mV, which was then followed by a deactivation gap (2 ms) at the holding potential (–110 mV) and a test pulse to –20 mV. Current traces similar to the one shown in the inset (Fig. 3D) were used to plot the curves. Ba2+, Ca2+, and Sr2+ currents were half-inactivated at –52.1 ± 0.6, –53.8 ± 0.6, and –56.8 ± 0.5 mV; and the curves had slope factors (in mV) of 5.4 ± 0.5, 4.7 ± 0.4, and 5.5 ± 0.5 (n = 14), respectively. As shown (Fig. 3D), superimposition of the activation (dotted lines) and inactivation (solid lines) curves predicted a window current resulting from overlap of the curves, which ranged from about –60 to –40 mV. Most notably, the peak of the window current of the Sr2+ currents (about –50 mV) was 
5 mV more negative than the Ba2+ and Ca2+ currents.
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values) of deactivation. Then the deactivation values, the activation values of Ba2+ current traces, and the relation between step voltages were examined (Fig. 4). One provided the best fit to the tail currents. The early phase of the tail currents (0.3 ms) was ignored during curve fitting to remove possible contamination by the capacitive transients. Similar protocols and analyses were used to generate the voltage dependence of activation time constants for Ca2+ and Sr2+ currents (Fig. 4). In another series of experiments, the time dependence of recovery from and development of inactivation of Ba2+ currents at different voltages were determined by use of double-pulse protocols (Fig. 5). Whereas the kinetics of recovery from inactivation were best described with at least two time constants, one time constant was sufficient to fit the profile of the development of inactivation of Ba2+ currents (Fig. 5, A and B). Similar results were obtained for Sr2+ currents (data not shown).
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). The properties and pharmacology of the inner ear Cav3.1 channel currents will require further molecular and functional characterization. Meanwhile, single-channel recordings were obtained from oocytes injected with Cav3.1 mRNA. To estimate the values of the voltage steps in the cell-attached configurations, oocytes were bathed in solutions containing high K+ to clamp the membrane potential at about 0 mV. The patch pipette contained 20 mM Ba2+ or Ca2+. As shown in Fig. 8A, the resulting single-channel fluctuations displayed inward currents that were invariably transient at test potentials ranging from –110 to –30 mV, from a holding potential of –120 mV. An example of the amplitude histograms used to generate the unitary current amplitude is shown in Fig. 8B. The estimated single-channel conductances (in pS) from the regression line of the I–V relationship (Fig. 8, C and D) were 10.7 ± 0.9 (n = 5) and 13.3 ± 1.1 pS (n = 6), for Ba2+ and Ca2+, respectively.
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We investigated the expression of Cav3.1 in the mouse inner ear using immunohistochemistry and immunoelectron microscopy approaches. Positive, moderate-intensity staining of Cav3.1 was observed in the neuroepithelial tissues of both the cochlea and vestibule (Figs. 9A and 10A) in the adult mouse. In the cristae ampullaris of the semicircular canals and the macula of the utricle and saccule, the immunoreactivity was most intense in the apical portions of the hair cells and their supporting cells (Fig. 9, B and C). Both type I and type II vestibular hair cells showed light reactivity in the cytosol with increasing reactivity at the apical portion underlying the cuticular plate. However, as best shown in Fig. 9B, the nerve chalice surrounding the lower portion of the type I vestibular hair cell lacks reactivity, although the supporting cells appeared slightly reactive for Cav3.1. At the light microscope level, the supporting cell can be differentiated from type II hair cells by the presence of the dark reticular laminar material just beneath the apical surface of the supporting cell. The transitional zone adjacent to the vestibular hair cells and the dark cell region were positive for Cav3.1.
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Immunoreactivity for Cav3.1 was first detected in the embryonic inner ear at embryonic day 18 (E18; Fig. 9, D and E). Virtually all of the epithelial cells lining the membranous labyrinth of the vestibular portion of the inner ear express Cav3.1. Interestingly, the intensity of the reactivity was greater in the nonsensory regions than that in the immature neurosensory epithelium. A light-moderate expression of Cav3.1 was noted in the epithelial cells determined to become the transitional cells, as well as in dark cells and the membrane separating endolymph and perilymph. Whereas in the immature hair cells and supporting cells of the cristae ampullaris (Fig. 9D) and macule of the saccule (Fig. 9E) and utricle (not shown), the reactivity was barely above background. However, the immature hair cells of the cochlea (Fig. 10B) show the same level of reactivity as do the immature support cells of the organ of Corti and the outer sulcus epithelium. The cuboidal epithelial cells, which demarcate the future stria vascularis, lack reactivity. As the hair cells of both the vestibule and cochlea mature, a light, sometimes punctate reactivity for Cav3.1 can be detected throughout the cytosol from E18 to P6 (Figs. 9, F and G and 10C). At P6, the OHCs of the cochlea display Cav3.1 labeling primarily along the plasmalemma (Fig. 10D), whereas the other hair-cell types of the inner ear continue to show cytosolic staining (Figs. 9, H and I and 10D).
The differences in the degree and location of Cav3.1 immunopositivity between the vestibule and cochlea as well as between sensory and nonsensory tissues continue with increasing maturation of the inner ear. At P8, a moderate reactivity occurs in the perinuclear region of the type I and type II vestibular hair cells, as well as at the apically located cuticular plate. The supporting cells between the hair cells continue to display light-moderate reactivity throughout the cytosol with increasingly greater reactivity at their apical reticular membrane (Fig. 9, J and K). However, reactivity decreases in the transitional zone and in the dark cells so that they show negative Cav3.1 reactivity by P8 (not shown). The nerve chalice surrounding the type I hair cell was also negative for staining. With increasing age beyond P8, the intensity of the reactivity of the Cav3.1 at the reticular membrane of the supporting cell and hair cells increases, and until P18 it is equivalent to that seen in the adult mouse (Fig. 9, B and C).
The immature hair cells of the P8 cochlea show a moderate level of reactivity (Fig. 10D, inset) throughout the cytosol of the IHC and in the subnuclear region of the OHCs. At this age reactivity is also noted in the apical "cup" region of the Deiter's cell under the base of the OHC, as well as in the epithelial cells of the newly formed inner sulcus. In addition, the fibrocytes of the spiral limbus and ligament begin to show immunopositivity. Expression of Cav3.1 continues to be noted in the nonsensory cells (Hensen's and Claudius’) of the organ of Corti as well as the outer sulcus cells. However, as the cochlear tissues continue to mature, reactivity in these cells declines and is virtually at the level of background by P21. During this same time period (P8–P21), the level of immunoreactivity in the fibrocytes of the spiral ligament and limbus increases to exceed that of the cochlear hair cells. At P2, the pattern of Cav3.1 immunoreactivity in the cochlea assumes that shown in Fig. 11A for the mature, adult cochlea.
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DISCUSSION |
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; Llinás and Yarom 1981) and pacemaker cells (Hagiwara et al. 1988), as well as Ca2+-dependent signaling, including cell proliferation (Rodman et al. 2005a,b), differentiation (Bertolesi et al. 2003; Strobeck et al. 1999), and apoptosis (Wang et al. 1999). We have identified and isolated a novel Cav3.1 channel isoform from the mouse inner ear: a splice variant that excludes exon14, exon25, exon34, and exon35, but includes exon26 and exon38 (Fig. 1). Previous studies have shown that Cav3.1 channels are subject to extensive alternative RNA splicing. So far, >30 distinct splice variants have been identified in the fetal and adult human brain (Emerick et al. 2006; Mittman et al. 1999; Monteil et al. 2000; Murbartian et al. 2004; Yunker and McEnery 2003), suggesting a fundamental mechanism to confer the diversity of T-type Ca2+ currents. However, the import of alternative splicing is less understood in mice; only two full-length splice variants of Cav3.1 channels have been reported to date (Mittman et al. 1999; Murbartian et al. 2004; Yunker and McEnery 2003). The inner ear Cav3.1 has distinct utilization of exons from those of the Cav3.1 channels found in mouse brain (BC057399
[GenBank]
and AJ012569
[GenBank]
).
One of the Cav3.1 channels (BC057399
[GenBank]
) excludes exon8 and encodes a nonfunctional channel lacking half of the pore-forming helix in the first functional domain (IS6). Compared with the other mouse Cav3.1 channels (AJ012569
[GenBank]
), the inner ear Cav3.1 has short I–II and II–III linkers because of the exclusion of exon14 and exon25A (Fig. 1). Exon14 introduces an additional 23 aas into the II–III loop of the channel. Splice variants with this insert display faster inactivation kinetics compared with channels without. Moreover, Cav3.1 channels including exon25A, which encodes an extra stretch of 7 aas in the intracellular III–IV loop, activate at more negative membrane potentials and show a faster activation rate compared with that of other Cav3.1 channels (Chemin et al. 2001). As predicted based on their structure, the inner ear and other mouse Cav3.1 channels differ in their molecular and electrophysiologic properties.
The primary structure of the inner ear Cav3.1 is distinct from that of other Cav3.1 channels at several important functional domains. The lack of exon14 excludes a consensus phosphorylation site for protein kinase C in the intracellular II–III loop, which may serve as an important regulatory site (Chemin et al. 2001). Similarly, the inner ear Cav3.1 channel also lacks a casein kinase II phosphorylation site in the intracellular III–IV loop, encoded by exon34. Again, these differences reinforce the potential for differential regulatory mechanisms between the inner ear and other Cav3.1 channels. Although the biological significance of these splice variants is not completely understood, the differences in their physiological functions, together with their tissue-specific distribution, may contribute to differential regulation and modulation of biological processes, resulting in tissue-specific intracellular Ca2+ handling.
The kinetic phenotype of the inner ear Cav3.1 channels in heterologous expression system, with respect to the fast inactivation, was distinct from typical Ca2+ currents recorded from developing hair cells (Levic et al. 2007). Moreover, the results were not surprising since recapitulation of the native cell current properties invariably requires auxiliary subunits (Dolphin et al. 1999; Lacerda et al. 1994; Leuranguer et al. 1998). The sensitivity of the channel to nimodipine may explain why the current has been missed in previous studies in hair cells (Rodriguez-Contreras and Yamoah 2003).
Ca2+ influx from T-type Ca2+ currents gives rise to many scenarios for developing and regenerating hair cells (Levic et al. 2007). Membrane excitability is promoted when the low-threshold features of the current amplify the depolarizing inputs. Inactivation of this current may generate burst firing patterns from tonic inputs. As a result, Ca2+ entry driven by a large tail current follows to facilitate a robust change of intracellular Ca2+ during activity. Additionally, steady Ca2+ influx will occur due to a window current caused by the overlap of the steady-state inactivation and activation relations at a potential range near the resting potential of hair cells. This steady inward flow of Ca2+ will result in a further boost of intracellular Ca2+ (Barish and Mansdorf 1991). Last, Ca2+ current inactivation may be self-limiting, creating a feedback control to prevent Ca2+ overload. Functional studies have shown that T-type currents can mediate phasic changes in intracellular Ca2+, which may prove to be an important requirement for the synthesis and release of neurotrophins during hair-cell development and regeneration (Eatock and Hurley 2003). Patterned electrical activity and the ensuing changes in Ca2+ transients, opposed to steady firing activity, preferentially stimulate the transcription and secretion of neurotrophins (Balkowiec and Katz 2002). Moreover, electrical activity tightly regulates the expression of neurotrophin receptors, giving insight into a mechanism by which neurotrophins may selectively affect electrically active neurons. In particular, there is a solid link between the release of brain-derived neurotrophic factor and neurotrophin 3 and the specificity of hair-cell innervation in the cochlea of the inner ear. A base-to-apex preference gradient is created when neurotrophins define the innervation pattern of type II afferents on OHCs (Farinas et al. 2001).
Tissue-specific localization of T-type Ca2+ channels has prompted the suggestion that the channel subtypes play unique roles that cannot be compensated by other channels (Talley et al. 1999). Because mibefradil inhibits cell proliferation in vascular tissues and there appears to be robust expression of T-type current as cells transition from G0 to G1 and S-phase, the possibility that T-type channels could be involved in regulating cell proliferation has been raised (Bertolesi et al. 2003). Localization of the Cav3.1 immunoreactivity to nonsensory cells of the cochlea, vestibule, and fibrocytes of the spiral ligament and limbus is striking. The correlation between the expression of Cav3.1 and the propensity of nonsensory cells to proliferate (Holley 2005) is also remarkable and it is conceivable that the Cav3.1 channels in cells in the inner ear may be an index for cell division. Indeed, a recent report has demonstrated the presence of a T-type current in developing hair cells (Levic et al. 2007), which is consistent with the hypothesis that Cav3.1 may be important for cellular differentiation. Also important, the robust expression of Cav3.1 and reappearance in regenerating hair cells is in further agreement with the concept that expression of T-type currents begets cellular development. T-type currents are frequently observed in the early development of cells and their density, amplitude, and properties change over time as seen in embryonic dorsal root ganglia, retinal Müller cells, hippocampal neurons, and thalamocortical cells (Bringmann et al. 2000; Desmadryl et al. 1998; Pirchio et al. 1990; Yaari et al. 1987). Moreover, in some cell types, T-type current density decreases with age until a mature stage is reached. At that time the current density is reduced substantially, giving way to the expression of high-voltage–activated (HVA) Ca2+ currents (Chameau et al. 1999). Indeed, it has been reported that T-type currents may regulate the expression of HVA Ca2+ currents (Chemin et al. 2002b), raising the possibility of mechanisms for coordination of T-type and HVA Ca2+ channel expression in hair cells.
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GRANTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Address for reprint requests and other correspondence: E. N. Yamoah, Center for Neuroscience, Program in Communication Science, University of California, Davis, 1544 Newton Ct., Davis, CA 95618 (E-mail: enyamoah{at}ucdavis.edu)
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), Ca2+ (
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) currents at various potentials.
