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NEUROSCIENCE |
1 Department of Biomedical Science, University of Sheffield, Sheffield S10 2TN, UK
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Abstract |
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(Received 18 September 2007;
accepted after revision 11 December 2007;
first published online 3 January 2008)
Corresponding author W. Marcotti: Department of Biomedical Science, University of Sheffield, Sheffield S10 2TN, UK. Email: w.marcotti{at}sheffield.ac.uk
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Introduction |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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P12: Mikaelian & Ruben, 1965; Woolf & Ryan, 1984) afferent synaptic transmission is driven by spontaneous Ca2+-dependent action potentials (Glowatzki & Fuchs, 2002; Marcotti et al. 2003b; Johnson et al. 2005), which are likely to influence both the refinement of downstream neural circuits and intrinsic IHC development (Johnson et al. 2007) in analogy with other developing systems (Zhang & Poo, 2001; Moody & Bosma, 2005). Since voltage-gated Ca2+ channels play such a crucial role in immature and adult IHCs, any tonotopic and/or developmental differences in their biophysical properties are likely to have a significant impact on function. It is now well established that the majority (
90%) of the Ca2+ current in hair cells of both mammals (Platzer et al. 2000; Knirsch et al. 2007) and non-mammals (turtle: Schnee & Ricci, 2003; chick: Fuchs et al. 1990; Spassova et al. 2001; frog: Rodriguez-Contreras & Yamoah, 2001) is carried by L-type Ca2+ channels containing the CaV1.3 subunit (Kollmar et al. 1997; Platzer et al. 2000). The nature of the remaining 10% of non-L-type Ca2+ channels is still debatable since N-, R- and T-types have been described in hair cells of various vertebrates (Martini et al. 2000; Rodriguez-Contreras & Yamoah, 2001). In vertebrates, the cochlear sensory neuroepithelium is tonotopically organized such that the characteristic frequency of HCs (the frequency at which they respond best) gradually changes with their position along the organ. In the non-mammalian cochlea the characteristic frequency of HCs is intrinsically dictated by the interplay between the relative number and kinetic properties (Wu et al. 1995) of both Ca2+ and BK channels (Art & Fettiplace, 1987; Fuchs et al. 1988) as well as local Ca2+ buffering kinetics (Ricci et al. 2000), which vary as a function of position along the cochlea. This inherent electrical tuning in lower vertebrate hair cell is not found in IHCs of the adult mammalian cochlea where their characteristic frequency is thought to be determined by the active mechanical amplification of the cochlea partition via the outer hair cells (Dallos, 1992). Although numerous studies have investigated Ca2+ channels in IHCs of the mammalian cochlea (Platzer et al. 2000; Marcotti et al. 2003b; Johnson et al. 2005) there is still an absence of information regarding any change in their biophysical properties as a function of the frequency position along the cochlea. However, a recent study using immunogold staining has shown that tonotopic differences exist at least in the concentration of Ca2+ buffering proteins in mammalian cochlear hair cells (Hackney et al. 2005).
The aim of this paper was to investigate the kinetic properties and Ca2+-dependent inactivation of ICa in low- and high-frequency IHCs of the gerbil cochlea during development. This information would provide the first electrophysiological evidence for a position-dependent specialization intrinsic to the IHCs, the function of which could be to fine-tune the responses of these auditory receptors. All ICa recordings, apart from those designed to investigate its temperature dependence, were performed in near physiological conditions (body temperature and using 1.3 mM extracellular Ca2+) to ensure a more realistic estimation of the channels biophysical properties.
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Methods |
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Apical-coil and basal-coil inner hair cells (IHCs, n
= 133) were studied in acutely dissected organs of Corti from Mongolian gerbils during development (from postnatal day 3 (P3) to P69, where the day of birth is P0). The reason for choosing the gerbil, instead of the more commonly used mouse, was due to their extended low-frequency hearing range that would emphasize any tonotopic differences (gerbil: 0.1–50 kHz; mouse: 2–100 kHz, Greenwood, 1990).
Gerbils were killed by cervical dislocation in accordance with UK Home Office regulations. The cochleae were dissected in normal extracellular solution (mM): 135 NaCl, 5.8 KCl, 1.3 CaCl2, 0.9 MgCl2, 0.7 NaH2PO4, 5.6 D-glucose, 10 Hepes-NaOH. Sodium pyruvate (2 mM), amino acids and vitamins (Eagle's MEM) were also added from concentrates (Fisher Scientific, Loughborough, UK). The pH was adjusted to 7.5 (osmolality 308 mosmol kg–1). The dissected cochleae were transferred to a microscope chamber, immobilized under a nylon mesh attached to a stainless steel ring, and continuously perfused with a peristaltic pump using the above extracellular solution. The organs of Corti were viewed using an upright microscope (Leica DMLFS, Germany) with Nomarski optics (x63 objective). In situ recordings were made by exposing the basolateral surfaces of IHCs using a suction pipette (tip diameter about 4 µm), which was filled with extracellular solution. The approximate position of IHCs along the cochlea was recorded as fractional distance from the extreme base. Adult IHCs were positioned at a fractional distance of between 0.9 and 0.95 (apex) and between 0.1 and 0.2 (base), corresponding to a frequency range of 250 Hz and 420 Hz in apical and 20 kHz and 37 kHz in basal cells (Müller, 1996). Immature IHCs were recorded from a similar fractional distance to that of adult cells.
Electrical recording
Patch clamp recordings were performed near body temperature (34–37°C) using an Optopatch (Cairn Research Ltd, Faversham, UK) and an Axopatch 200B (Molecular Devices, USA) amplifier. A few experiments were conducted at room temperature (21–23°C) in order to determine the effect of temperature on the size of the Ca2+ current. The temperature dependence of ICa was described by its temperature coefficient (Q10), which was calculated from the van't Hoff equation:
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| (1) |
Patch pipettes were made from soda glass capillaries (Harvard Apparatus Ltd, Edenbridge, UK) and had a typical resistance in extracellular solution of 2–3 M
. In order to reduce the electrode capacitance, patch electrodes were coated with surf wax (Mr Zogs Sex Wax; Zog Industries, Carpenteria, CA, USA). Patch electrodes were positioned on the IHC membrane using a PatchStar micromanipulator (Scientifica Ltd, Uckfield, UK). The patch pipette filling solution used for most of the whole-cell recordings was (mM): 106 caesium glutamate, 20 CsCl, 3 MgCl2, 1 EGTA-CsOH, 5 Na2ATP, 0.3 Na2GTP, 5 Hepes-CsOH, 10 Na2-phosphocreatine (pH 7.3, 294 mosmol kg–1). For a few experiments different concentrations (0.1, 0.3, 1 and 5 mM) of the fast Ca2+ buffer BAPTA (Roche Diagnostic, UK) were used instead of 1 mM EGTA (Fluka, UK) in the above intracellular solution. In the BAPTA solutions the concentration of CsCl was adjusted to keep the osmolality constant. For the current clamp experiment, used to record the action potential applied to cells as a voltage protocol in Fig. 9, the pipette filling solution contained (mM): 131 KCl, 3 MgCl2, 1 EGTA-KOH, 5 Na2ATP, 5 Hepes-KOH, 10 sodium phosphocreatine (pH 7.3, 292 mosmol kg–1). Data acquisition was performed using pCLAMP software with a Digidata 1320A/1440A interface (Molecular Devices). Depending on the protocols used, data were sampled at 5, 10, 20 or 100 kHz and filtered at 2.5, 5 or 10 kHz (8-pole Bessel) and stored on computer for off-line analysis (Origin; OriginLab Corp., Northampton, MA, USA; Mini Analysis Program: Synaptosoft Inc., Decatur, GA, USA). Current recordings were corrected off-line for the linear leak conductance (gleak) typically calculated between –84 mV and –74 mV (immature IHCs: 2.6 ± 0.1 nS, n
= 65, P3–P9; adult IHCs 1.5 ± 0.1 nS, n
= 68, P25–P69). Holding currents are plotted as zero current. Membrane potentials were corrected for the voltage drop across the uncompensated residual series resistance (Rs: 4.7 ± 0.1 M, n
= 126, compensation 0–80%) and for a liquid junction potential (LJP), measured between electrode and bath solutions, according to Neher (1992). The LJP was –11 mV for the caesium glutamate and –4 mV for the KCl intracellular solution. The average voltage-clamp time constant (product of Rs and membrane capacitance Cm) was 46 ± 1.0 µs (n
= 126). For the experiments aimed at investigating the activation kinetics of ICa the average voltage-clamp time constant was 32 ± 3.0 µs (n
= 30).
Extracellular superfusion
All voltage-clamp experiments were performed in the presence of 30 mM TEA and 15 mM 4-AP (Fluka, UK) in order to block the K+ currents (immature: IK,neo and IK1, Marcotti et al. 2003a; adult: bEK,f and IK,s, Marcotti et al. 2004a). Moreover, the K+ channel blockers apamin (300 nM: Calbiochem, UK) and linopirdine (80–100 µM: Tocris Bioscience, Bristol, UK) were also used to block ISK2 (in immature IHCs, Marcotti et al. 2004b) and IK,n (in adult IHCs, Marcotti et al. 2003a), respectively. When the concentration of added K+-channel blockers was > 1 mM, or when 1.3 mM Ca2+ was replaced by either 5 mM Ca2+ or Ba2+, NaCl was adjusted to keep the osmolality constant. Solutions containing drugs were applied through a multibarrelled pipette positioned close to the preparation.
Statistical analysis
Statistical comparisons of means were made by Student's two-tailed t test or, for multiple comparisons, analysis of variance, usually one-way ANOVA followed by Tukey's test. Two-way ANOVA, followed by the Bonferroni's test, was used to compare data sets from apical and basal regions of the cochlea. Mean values are quoted ± S.E.M. where P < 0.05 indicates statistical significance. In some of the figures statistical significance is indicated by asterisks.
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Results |
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The Ca2+ current (ICa) was isolated from the total membrane current by blocking all the K+ currents present in IHCs using a cocktail of K+ channel blockers (see Methods). A typical example of ICa, in the presence of 1.3 mM extracellular Ca2+, recorded from apical-coil (Ac) and basal-coil (Bc) IHCs of the immature cochlea is shown in Fig. 1A and B,
respectively. The average current–voltage (I–V) curves measured at the peak ICa, for both apical and basal immature gerbil IHCs, are shown in Fig. 1C. I–V curves for individual cells were fitted using the following equation:
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| (2) |
is the potential at which the conductance is half-activated and S is the slope factor that defines the voltage sensitivity of current activation. The maximum size of ICa (measured at –15 mV) was found to be similar between apical (–408 ± 96 pA n
= 6) and basal (–365 ± 37 pA n
= 5) immature IHCs. ICa recorded from both apical and basal IHCs activated at around –70 mV (defined as 1% of gmax). No significant differences were observed in V and S between the two cochlear regions.
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is the time constant of activation and 
was fixed as 2 (Zidanic & Fuchs, 1995; Marcotti et al. 2003b; Johnson et al. 2005), which is consistent with a Hodgkin–Huxley model with two opening gating particles (Hodgkin & Huxley, 1952). In some apical IHCs (about 50%), the onset of ICa at more hyperpolarized potentials appeared slow and possibly multicomponent (Fig. 1A), although this was not investigated further. Although the average size of ICa was similar between the two cell populations (Fig. 1C) the activation time constants (Fig. 1F) were significantly faster in basal than in apical IHCs (P < 0.0001 – two-way ANOVA; post test indicated significance at P < 0.001 between –61 mV and –46 mV).
The Ca2+ current recorded in adult apical (300 Hz, P28) and basal (30 kHz, P30) gerbil IHCs is shown in Fig. 2A and B,
respectively. The maximum size of ICa measured at –11 mV in adult apical IHCs (Fig. 2C: –155 ± 22 pA, n
= 6, P27-P30) was found to be slightly smaller than that of basal cells (–181 ± 21 pA, n
= 7, P27–P30) but the difference was not quite significant (P
= 0.05). The activation of ICa in both apical and basal adult IHCs occurred positive to –65 mV. Half-maximal activation and slope factor were not significantly different between apical and basal IHCs (Fig. 2C). Surprisingly, V activation was significantly more negative (about 10 mV) in immature than in adult IHCs (P < 0.001 for both cochlear regions: one-way ANOVA). Figure 2D shows the direct comparison of the peak ICa as a function of development and cochlear position. While the maximum size of ICa did not change significantly along the length of the cochlea within either age group, it was much smaller in adult compared to immature IHCs (P < 0.001), as previously observed in mouse IHCs (Marcotti et al. 2003b; Johnson et al. 2005, 2007). The activation time course of ICa in adult IHCs is shown on an expanded time scale in Fig. 2E and F. Similar to immature IHCs, the ICa activation time constants in adult basal IHCs were significantly faster than those of low-frequency cells (Fig. 2G: P < 0.0001 – two-way ANOVA; post test at P < 0.001 between –56 mV and –41 mV).
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T used: 14°C). In adult IHCs, the maximum size of ICa measured at room temperature (–116 ± 13 pA, n
= 5, Fig. 3C) was also significantly smaller compared to that recorded at body temperature (P < 0.0001). In adult IHCs the size of ICa changed with temperature with a Q10 of 1.36, indicating that somehow Ca2+ channels (most likely their open channel probability) become less temperature dependent with maturation. These results are similar to recent findings from mouse P14 IHCs (Q10
= 1.1 in whole cell: Nouvian, 2007).
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In order to obtain a realistic representation of ICa inactivation it was important to completely block the small conductance Ca2+-activated K+ current SK2 (using 300 nM apamin: Marcotti et al. 2004b) since its presence in whole-cell recordings gives a false impression of a pronounced Ca2+ current inactivation (Schnee & Ricci, 2003; Marcotti et al. 2003b). Figure 4A
shows some examples of inward Ca2+ currents recorded over a period of 600 ms in apical (middle panel) and basal (bottom panel) adult gerbil IHCs (P27–P30). Inactivation curves (Fig. 4D) were obtained by measuring the peak ICa during a 100 ms test step to –11 mV (apical IHC: Fig. 4B; basal IHC: Fig. 4C) following a series of 600 ms conditioning steps (Fig. 4A). The fits through the data were obtained using a modified first-order Boltzmann equation:
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(Fig. 4F, middle panel) and the slope factor (Fig. 4F, right panel) did not change significantly as a function of both development and cochlear region, the degree of ICa inactivation (Fig. 4F, left panel) was larger in basal (58.0 ± 4.5%
n
= 6) compared to apical (44.8 ± 2.2%
n
= 6, P < 0.02) adult IHCs.
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1 min) were used for these experiments and Ba2+ was mainly superfused onto IHCs after Ca2+, possibly leading to a smaller Ba2+ current due to a partial current run-down. Inactivation curves (Fig. 5E) were obtained by measuring the peak current (5 mM Ca2+: Fig. 5C, mM Ba2+: Fig. 5D) obtained using the same voltage protocol described in Fig. 4A–C. The extent of inactivation in the presence of 5 mM Ba2+ (18%: Fig. 5F) was significantly reduced compared to both 5 mM Ca2+ (43%, P < 0.01) and 1.3 mM Ca2+ (50%; P < 0.001), suggesting the presence of both Ca2+- and voltage-dependent inactivation processes. No significant difference was observed between 1.3 mM and 5 mM Ca2+ (Fig. 5F), although the inactivation–voltage relation in the latter had a more pronounced U-shaped appearance that resembled that of the I–V curve (Fig. 5B).
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fast) but not the slow (Fig. 6D: slow) time constant was slightly affected by substituting EGTA with BAPTA as intracellular Ca2+ buffer (P < 0.02, one-way ANOVA). These results suggest that the source of Ca2+ responsible for Ca2+ channel inactivation is likely to be very colocalized.
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Neurotransmitter release at the IHC ribbon synapse is likely to be regulated by a few L-type Ca2+ channels (Brandt et al. 2005). The readily releasable pool (RRP), which represents the number of vesicles docked at the active zones (von Gersdorff et al. 1996; Moser & Beutner, 2000), is recruited when IHCs are stimulated using relatively short depolarizing voltage steps (up to around 100 ms at body temperature and using 1.3 mM extracellular Ca2+: Johnson et al. 2005, 2007). Since the release of the RRP could be affected by ICa inactivation, we investigated how Ca2+ channels would respond to repeated stimulation. For this set of experiments, both immature and adult gerbil IHCs were depolarized using a series of voltage steps to –11 mV (50 ms in duration) every 150 ms (Fig. 7A). An example of ICa recordings, using the above protocol, from immature (P7) and adult (P25) IHCs is shown in Fig. 7B. In adult IHCs (Fig. 7C), repeated stimulation caused an initial decline in size of ICa (8% in apical and 11% in basal IHCs with the second stimulus) followed by a very similar shallow reduction, reaching a maximum inactivation (after 25 repetitions) of 20% in apical and 23% in basal cells. This result suggests that low- and high-frequency adult IHCs are likely to exhibit a similar recovery of ICa from inactivation. The slight variation in the degree of ICa inactivation between apical and basal IHCs (Fig. 7C) is likely to be due to their different steady-state inactivation (Fig. 4D). When the same voltage protocol was applied to immature IHCs the initial ICa inactivation increased from 12% and 7% (with the second stimulus) to 55% and 36% after 25 repetitions in apical and basal cells, respectively (Fig. 7D). The extent of this inactivation was significantly greater in apical than in basal IHCs (P < 0.0001: Fig. 7D), suggesting a more rapid recovery from ICa inactivation in the latter.
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= 106.0 ± 8.7 ms, n
= 5) showed a significantly slower recovery from inactivation than basal cells (
= 48.2 ± 2.5 ms, n
= 4, P < 0.001).
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Discussion |
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300 Hz) and basal (high-frequency: 30 kHz) regions of the cochlea using near physiological recording conditions (body temperature and 1.3 mM extracellular Ca2+). Although most of this information is well established in lower vertebrate auditory organs (Zidanic & Fuchs, 1995; Schnee & Ricci, 2003), the existence of tonotopic differences in the biophysical properties of ICa in mammalian IHCs has never been investigated before. The information gained by this study will provide important insights into whether Ca2+ channels could be involved in the fine tuning of mammalian IHCs. CaV1.3 Ca2+ channels in developing mammalian IHCs
IHCs are the primary sensory receptors of the mammalian cochlea responsible for signalling the perception of sound to the central nervous system via the vast majority (90–95%) of afferent auditory nerve fibres entering the cochlea (Ryugo, 1992). Before the onset of hearing, which in most rodents occurs at around P10–P12, immature IHCs fire spontaneous Ca2+-dependent action potentials (Marcotti et al. 2003a, 2003b; Johnson et al. 2007), which are likely to be responsible for the normal morphological and physiological maturation of the cochlea (Johnson et al. 2007). Upon maturation, IHCs respond to incoming sound with rapid and graded receptor potentials (Palmer & Russell, 1986) that are transmitted onto auditory afferent fibres. IHC depolarization, induced by either spontaneous action potential activity or incoming sound, causes an influx of Ca2+ into cells via the opening of voltage-gated Ca2+ channels. Since Ca2+ channels are clustered and colocalized with IHC presynaptic active zones (Roberts et al. 1990; Tucker & Fettiplace, 1995) their activation triggers the release of neurotransmitter from ribbon synapses (Parsons et al. 1994; Moser & Beutner, 2000). In addition to this synaptic activity, Ca2+ channels are also responsible for regulating IHC membrane excitability (Marcotti et al. 2004a,b). More than 90% of the total Ca2+ current expressed in IHCs is carried by L-type 1D (CaV1.3) Ca2+ channels (Platzer et al. 2000; Koschak et al. 2001), suggesting that the properties of ICa could be similar in IHCs irrespective of position or age. CaV1.3 channels are also expressed in the electromotile outer hair cells (mouse: Knirsch et al. 2007) although their relative number and/or single channel conductance is likely to be smaller than in IHCs.
Here we show that in contrast to lower vertebrates (Schnee & Ricci, 2003) the size of ICa did not change significantly along the cochlea in both immature and adult IHCs (Figs 1 and 2). However, as previously reported in mouse IHCs (Beutner & Moser, 2001; Marcotti et al. 2003b; Johnson et al. 2005), a substantial reduction of ICa occurred with development (Fig. 2D). Changing the recording conditions from body (35–37°C) to room (21–23°C) temperature caused a significant reduction in the size of ICa in both immature (< P12: Q10 = 1.9) and adult (> P20: Q10 = 1.4) gerbil IHCs. These results are similar to recent observations in 2-week-old mouse IHCs (Nouvian, 2007) where a reduction in the size of ICa at room temperature was observed (Q10 = 1.1, whole-cell recordings; Q10 = 1.3, perforated patch). The significantly greater temperature dependence of ICa in immature IHCs (Fig. 3D), suggests that some differences exist in the properties of Ca2+ channels between the two stages of development. The larger Ca2+ current at physiological temperature is likely to be caused by an increased open channel probability (Acerbo & Nobile, 1994).
At body temperature, the activation time course of ICa was significantly faster in IHCs located in the high-frequency region of the gerbil cochlea throughout development. This scenario is further complicated if we consider that ICa in adult IHCs (Fig. 2E–G) showed overall faster activation kinetics than that of immature cells (Fig. 1D–F). The activation time constant of ICa in gerbil basal IHCs (Figs 1F and 2G) was comparable to that recorded in mouse apical IHCs (physiological 1.3 mM extracellular Ca2+: Marcotti et al. 2003b; Johnson et al. 2005; 2 mM Ca2+: Nouvian, 2007) although no developmental difference was observed in the latter (Johnson et al. 2005). Although we did not investigate the activation kinetics of ICa at room temperature, previous studies on mouse IHCs have clearly reported that lowering the temperature from 37°C to room temperature slowed its activation time constant by about half (Marcotti et al. 2003b; Nouvian, 2007). A further difference between ICa in immature and adult IHCs was the 10 mV depolarizing shift in the voltage range of ICa activation (V: immature IHCs: –33 mV; adult: –23 mV). The reason for this developmental shift, assuming that it occurs in vivo, is currently unknown, although the activation of ICa in IHCs at both stages is likely to be within the range of the cell resting membrane potential (Fig. 4D and E). A similar shift with IHC maturation has recently been reported in rat IHCs (Knirsch et al. 2007).
Inactivation of CaV1.3 Ca2+ channels in gerbil IHCs
Recent investigations have shown that ICa in auditory hair cells inactivates, although the degree of inactivation appears to vary depending on the cell investigated (Marcotti et al. 2003b; Schnee & Ricci, 2003; Lee et al. 2007). Ca2+ channel inactivation provides a negative feedback mechanism to adjust the available Ca2+ current and is used to regulate a variety of physiological processes in both neuronal and non-neuronal cells (von Gersdorff & Matthews, 1996). Inactivation of the Ca2+ channel can be induced by either Ca2+ or voltage (Budde et al. 2002). Our results show that the extent of Ca2+ channel inactivation was largely reduced by Ba2+ (Fig. 5), indicating that Ca2+-dependent inactivation (CDI) is predominant in gerbil IHCs. This was also supported by the fact that the relation between ICa inactivation and voltage (Fig. 5E) in the presence of high Ca2+ roughly resembles that of its U-shaped I–V curve (Fig. 5B; Brehm & Eckert, 1978). Similar results have previously been described in lower vertebrates (Schnee & Ricci, 2003; Lee et al. 2007). Furthermore, ICa also showed voltage-dependent inactivation (VDI), since the inactivation of the Ba2+ current (Fig. 5E) through the Ca2+ channels increased with voltage and tended to saturate at positive potentials. A comparable degree of Ca2+ current inactivation was observed between immature (apical: 50%; basal: 47%) and adult (apical: 45%; basal: 58%) IHCs, even though the size of ICa in the latter was about a third. However, high-frequency adult IHCs showed a significantly larger Ca2+ channel inactivation than low-frequency cells (Fig. 4E), suggesting possible differences in the channel itself or in the control of local Ca2+ homeostasis between the two cochlear locations. This steady-state inactivation of ICa is likely to have a greater effect on the physiological voltage responses of adult IHCs (especially on the DC receptor potentials of basal cells) than those of immature cells (low-frequency action potentials). Possible roles for CDI in adult IHCs could be to regulate the number of available Ca2+ channels at the ribbon synapses and/or prevent Ca2+ loading. Less likely is its role in afferent fibre adaptation since this occurs on a faster time scale and is most likely caused by a reduced release of synaptic vesicle over time (Goutman & Glowatzki, 2007).
CDI can be brought about by the increase of cytosolic Ca2+ near the Ca2+ channel caused by current through the channel itself, adjacent Ca2+ channels, Ca2+ release from intracellular stores or the accumulation of Ca2+ following long-lasting cell depolarization. In order to determine how close the Ca2+ source was to the Ca2+ channel, and therefore attempt to discriminate between the above possibilities, we used different concentrations of intracellular BAPTA (Fig. 6) instead of EGTA. Our results show that 5 mM BAPTA did not significantly affect the CDI, suggesting that in gerbil IHCs the Ca2+ source for CDI is likely to be within 18 nm of each Ca2+ channel (Naraghi & Neher, 1997; Neher, 1998). These findings differ from those described in lower vertebrates where 1 mM BAPTA was sufficient to affect CDI in chick auditory hair cells (Lee et al. 2007), indicating a more distant coupling (about 40 nm) between Ca2+ channels and the Ca2+ source. Although we have not tested the possible direct modulation of CDI by Ca2+ stores (Lee et al. 2007) it is likely that the influx of Ca2+ from the opening of a single voltage-gated Ca2+ channel could cause CDI in gerbil IHCs (as seen in ventricular myocytes: Imredy & Yue, 1992) via the interaction of the Ca2+-sensing protein calmodulin with the channel -subunit (Peterson et al. 1999). This could explain the comparable degree of inactivation observed between immature and adult IHCs.
Repetitive maximal stimulation of ICa revealed another important difference between immature and adult IHCs, with a less extensive reduction in the size of ICa in adult cells over time (Fig. 7). This suggests that ICa in both apical and basal adult IHCs is able to sustain the repeated release of vesicles from the readily releasable pool (von Gersdorff et al. 1996; Moser & Beutner, 2000), which is essential for sound encoding. Although ICa inactivation is in theory less vital in immature IHCs (see above) we found position-dependent differences in its recovery from inactivation (Fig. 8C). The functional implication of this was that the maximum action potential frequency supported in apical IHCs is likely to be lower than that in basal cells (Fig. 9), suggesting that a difference in the spontaneous firing rate could exist along the mammalian cochlea. A similar action potential frequency in apical IHCs has been previously reported in the mouse (4 Hz: Marcotti et al. 2003b; Johnson et al. 2007). It is possible that this intrinsic limit in the firing activity prevents large influx of Ca2+ into developing IHCs, which could alter their normal functional development (Johnson et al. 2007). Alternatively, it could be responsible for creating a tonotopic signal that guides the organization of the auditory system before the onset of sensory-driven activity (Stellwagen & Shatz, 2002).
Our results indicate that although > 90% of ICa in mammalian IHCs is carried by CaV1.3 Ca2+ channels their activation and inactivation properties change with development and cochlear position pointing towards some differences in their structure or modulation. Splice variants of the 1 subunit (Kollmar et al. 1997) together with differences in the accessory subunits (Catterall, 2000) could be responsible for the diverse biophysical properties observed in this study. Moreover, Ca2+-dependent inactivation of ICa in IHCs could be modulated by a subtype of calmodulin-like Ca2+-binding protein (CaBP: Yang et al. 2006). Additional CaBPs such as calbindin D28k, calretinin and parvalbumin, generally assumed to play a role in intracellular Ca2+ buffering and known to be expressed in IHCs (Hackney et al. 2005), also function as Ca2+ sensors (Berggard et al. 2002). Evidence also points towards a role for calbindin D28k in modulating CDI of L-type Ca2+ channels (Lee et al. 2006). It would be informative to assess whether there is a differential expression of these proteins between low- and high-frequency adult gerbil IHCs.
In this study we have shown that the biophysical properties of ICa changed as a function of development. We suggest that these differences are likely to fulfil the different functional requirements of prehearing (spontaneous action potential activity) and mature (rapid and graded receptor potentials) IHCs. In addition, we also provide the first evidence for the existence of tonotopic differences in ICa throughout development, indicating that CaV1.3 Ca2+ channels are likely to be finely tuned by the expression of splice variants of the 1 subunit, different auxiliary subunits and/or specific Ca2+ binding proteins.
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) Imax
=
–166 pA, Iconst
= 0.56, V
) Imax
=
–148 pA, Iconst
= 0.43, V
) Imax
=
–274 pA, Iconst
= 0.50, V
