1Department of Physiology, The Auditory Laboratory, The University of Western Australia, Nedlands, Western Australia 6907, Australia; and 2Department of Otolaryngology, Guangdong Provincial People's Hospital, Guangzhou, People's Republic of China
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ABSTRACT |
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Zhang, Si Yi, Donald Robertson, Graeme Yates, and Alan Everett. Role of L-Type Ca2+ Channels in Transmitter Release From Mammalian Inner Hair Cells I. Gross Sound-Evoked Potentials. J. Neurophysiol. 82: 3307-3315, 1999. Intracochlear perfusion and gross potential recording of sound-evoked neural and hair cell responses were used to study the site of action of the L-type Ca2+ channel blocker nimodipine in the guinea pig inner ear. In agreement with previous work nimodipine (1-10 µM) caused changes in both the compound auditory nerve action potential (CAP) and the DC component of the hair cell receptor potential (summating potential, or SP) in normal cochleae. For 20-kHz stimulation, the effect of nimodipine on the CAP threshold was markedly greater than the effect on the threshold of the negative SP. This latter result was consistent with a dominant action of nimodipine at the final output stage of cochlear transduction: either the release of transmitter from inner hair cells (IHCs) or the postsynaptic spike generation process. In animals in which the outer hair cells (OHCs) had been destroyed by prior administration of kanamycin, nimodipine still caused a large change in the 20-kHz CAP threshold, but even less change was observed in the negative SP threshold than in normal cochleae. When any neural contamination of the SP recording in kanamycin-treated animals was removed by prior intracochlear perfusion with TTX, nimodipine caused no significant change in SP threshold. Some features of the data also suggest a separate involvement of nimodipine-sensitive channels in OHC function. Perfusion of the cochlea with solutions containing Ni2+ (100 µM) caused no measurable change in either CAP or SP. These results are consistent with, but do not prove, the notion that L-type channels are directly involved in controlling transmitter release from the IHCs and that T-type Ca2+ channels are not involved at any stage of cochlear transduction.
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INTRODUCTION |
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A variety of in vivo and in vitro recordings have
shown convincingly that the membranes of cochlear hair cells of both
mammalian and nonmammalian vertebrates possess voltage-gated calcium
channels and that these are important in regulating excitation of the
auditory nerve fibers (Issa and Hudspeth 1994;
Kollmar et al. 1997a
,b
; Robertson and Johnstone
1979
; Siegel and Relkin 1987
; Tucker and Fettiplace 1995
; Zidanic and Fuchs
1995
). Such channels are known to have a variety of important
roles. As in other secretory cells, when located close to the release
sites for synaptic vesicles, they have been presumed to play a crucial
role in controlling neurotransmitter release from the hair cells
(Martinez-Dunst and Fuchs 1997
; Roberts and
Hudspeth 1990
; Robertson and Johnstone 1979
;
Siegel and Relkin 1987
). In addition, voltage-gated
calcium channels can regulate the conductance of the basolateral wall of the hair cells, principally through the action of
Ca2+-activated potassium channels (Art and
Fettiplace 1995
; Dulon et al. 1998
; Issa
and Hudspeth 1994
; Kimitsuki et al. 1998) and may also contribute to the regulation of other
Ca2+-dependent aspects of hair cell function, including
adaptation and slow contractile processes (Assad and Corey
1992
; Ulfendahl 1987
; Zenner et al.
1985
).
A wide variety of voltage-gated Ca2+ channel subtypes have
now been described that differ in their voltage dependence,
conductance, and kinetic characteristics and in their pharmacological
profile with respect to specific antagonists (for review, see
Hofman et al. 1994; Miljanich and Ramachandran
1995
; Tsien et al. 1991
). For a number of
reasons it is of interest to know which of these subtypes is involved
in each of the various stages of the complex process of cochlear
transduction and activation of the primary afferent neurons.
Such knowledge could ultimately lead to the development of drugs
targeted to different stages of transduction, and these could have
utility in the treatment of a variety of inner ear disorders. With
particular reference to the mammalian inner hair cells (IHCs), which
constitute the major source of afferent neural outflow from the
cochlea, it is known that different primary afferent neurons emanating
from the same hair cells have different spontaneous firing rates,
thresholds, adaptation characteristics, and dynamic ranges
(Kiang et al. 1982; Muller and Robertson
1990; Winter et al. 1990), and one hypothesis is
that this diversity may be related to variations in a presynaptic
property such as the type of voltage-gated Ca2+ channel
that controls transmitter release.
Recent molecular biological studies have revealed the presence of mRNAs
coding for subunits of L-type channels in both mammalian and
nonmammalian inner ear tissues (see, for example, Green et al.
1996, Kollmar et al. 1997a
,b
). Furthermore,
channels with kinetic and pharmacological characteristics typical of
L-type channels (slow inactivation and susceptibility to the
dihydropyridine class of blockers, such as nimodipine) have been
demonstrated to be present in the hair cells of nonmammalian
vertebrates. An intensive study of the effects of various blockers in
chick cochlear hair cells (Zidanic and Fuchs 1995
)
indicates that this is the only channel type present in this species.
In hair cells of the frog sacculus and semicircular canal, however, the
presence of at least one other subtype (possibly N-type) in addition to
L-type channels has been suggested (Prigioni at al.
1992
; Su et al. 1995
). A variety of studies in
bullfrog, chick, and turtle have shown that voltage-gated calcium
channels are largely found at discrete sites in the hair cell membrane
that colocalize with presynaptic dense bodies (the presumed sites of
neurotransmitter release). However, although the circumstantial
evidence is compelling, none of these studies on isolated hair cells
has provided direct evidence for the control of neurotransmitter
release by the L-type or any other specific subtype of Ca2+ channel.
In the intact mammalian cochlea, study of this question is complicated
by the presence of two distinct sets of receptor hair cells with
different but coupled roles in the transduction process. Briefly, the
outer hair cells (OHCs) generate an AC receptor potential, the cochlear
microphonic, and control the overall mechanoelectrical gain of the
cochlea through their "active process" or "cochlear amplifier."
The inner hair cells (IHCs) on the other hand, generate a DC receptor
potential, the summating potential (SP), and provide the final
excitation of most of the primary afferent neural output of the organ
(for review see Patuzzi and Robertson 1988, Yates et al. 1992
).
In a study in the guinea pig cochlea, Bobbin et al. (1990)
provided good evidence that L-type channels are involved in
various aspects of cochlear responses to sound. These workers
demonstrated a series of complex changes in gross receptor potentials
after perfusion with nimodipine (a specific L-type channel antagonist), that were consistent with an effect on the OHCs. However, they also
showed a reduction in the maximum amplitude of the compound action
potential (CAP) of the auditory nerve, suggesting an additional effect
on the neural output from the IHCs (a decrease in gain alone mediated
by OHC effects would not be expected to reduce the maximum neural
output to high-level acoustic stimuli).
In the present study, we used gross recording techniques in the intact organ to pursue the specific question of the site of action of nimodipine and a T-type blocker Ni2+. We provide evidence to support the notion that L-type channels are involved in transmitter release from the IHCs, whereas T-type channels probably do not have a role in cochlear function.
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METHODS |
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Experiments were performed on young pigmented guinea pigs (260-425 g) of either sex. All procedures conformed to the Code of Practice of the National Health and Medical Research Council of Australia and were approved by the Animal Experimentation Ethics Committee of The University of Western Australia.
Animals were anesthetized according to previously published procedures
(Lowe and Robertson 1995). A small hole drilled in the
basal turn scala tympani was used for insertion of a perfusion pipette
(~80 µm tip diam), which was sealed in place with a small bead of
Sylgard attached to the shank of the pipette. The perfusion outlet hole
was made in the cochlear apex, and great care was taken to prevent
bleeding from this hole to eliminate the possibility of blockage of the
perfusion by blood clots. Perfusion was performed at a rate of 3 µlmin
1 for 10 min. The control artificial
perilymph comprised (in mM) 137 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2, 1 NaH2PO4, 12 NaHCO3, and 10 glucose. The solution pH was
adjusted to 7.4 at 37°C with the addition of 1 M NaOH.
Drugs were dissolved in the artificial perilymph without modifying the solution composition. Nimodipine (Alomone Laboratories) was dissolved first in DMSO, the final concentration of which in the perfusion solution ranged from 0.1 to 0.01% when nimodipine concentration ranged from 10 to 1 µM. TTX was obtained from Calbiochem-Behring, and all other salts and chemicals were obtained either from Sigma or BDH Chemicals.
The perfusion pipette served also to house a fine, etched platinum wire
electrode for recording sound-evoked cochlear potentials. Figure
1 illustrates the perfusion and recording
arrangement. The reference electrode was a chlorided silver wire
inserted in the animals' neck muscles. Gated tone bursts (10-ms
duration with 1-ms rise-fall times, repetition rate 10 s1) were delivered through a closed, calibrated
sound delivery system. The sound-evoked potentials measured from the
recording electrode were amplified and filtered before display on an
oscilloscope screen and storage on computer disk.
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A number of measures of the sound-evoked potentials were obtained and used in subsequent analysis. Direct observation from an oscilloscope screen was used to measure the visual detection threshold of the first negative wave (N1) of the nerve fiber CAP and the negative DC receptor potential (SP) to tone bursts ranging in frequency from 2 to 30 kHz (Fig. 1C). Note that the sign of the SP is traditionally denoted by its sign in the scala media, which is generally the reverse of that measured in the scala tympani.
Repeated blind measures of the visual detection thresholds for the CAP
and the SP at a variety of sound frequencies indicated that these were
reliable to within ±3 dB for any one observer (see also
Johnstone et al. 1979; Rajan et al.
1991
). Also measured were the amplitude of the CAP
(N1-P1 peak-to-peak
amplitude) and the SP (DC shift from baseline at 8 ms after tone onset)
at a range of stimulus intensities. These measures were derived from off-line analysis of computer-generated averaged responses to 10 stimulus presentations.
A separate group of animals had their OHCs destroyed throughout much of
the basal cochlear turn by administration of the ototoxic antibiotic
kanamycin. Animals were injected intraperitoneally with a solution of
kanamycin sulfate (Sigma) dissolved in saline at a dose of 400 mg/kg
for 10 days (Dallos and Harris 1978; Dallos and
Wang 1974
). They were then allowed to recover for ~2 weeks before final experimentation. At the end of the experiments, cochleae were fixed by perfusion with 2.5% glutaraldehyde in 0.1 M phosphate buffer, postfixed with 1% osmium tetroxide in phosphate buffer, and
dissected, and surface mounts were prepared for inspection of missing
hair cells.
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RESULTS |
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Control perfusions
Perfusion of the scala tympani with artificial perilymph caused very small changes in all measures of CAP and SP responses. Typically, there was a depression of ~3 dB in the threshold measures in the first 1-2 minutes of perfusion, but these rapidly recovered. We interpret the early and reversible change in thresholds to be a response to minor hydrostatic pressure changes in the cochlea at the onset of perfusion. When the mean visual detection thresholds of the CAP and SP to a 10-kHz tone burst were compared before and at the end of a 10-min perfusion the changes were <2 dB (n = 25 for CAP and n = 20 for SP). These changes were within measurement error and were not considered statistically significant (P > 0.05 for both CAP and SP). Input-output functions derived from averaged response waveforms at a range of intensities also showed no significant change after 10 min of artificial perilymph perfusion (Fig. 2, A-C).
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Perfusions with artificial perilymph containing 0.1% DMSO (the solvent used for nimodipine) also had no significant effects on CAP and SP thresholds. Similar to the standard artificial perilymph perfusion, the average change in thresholds at the end of a 10-min perfusion was <2 dB (n = 5) for both electrophysiological responses.
Nickel perfusions
In five animals, the cochlea was perfused with artificial perilymph containing 100 µM Ni2+. The effects of these perfusions did not differ significantly from control perfusions, showing no significant difference in the 10-kHz CAP and SP visual detection thresholds or in their respective input-output functions at the end of a 10-min perfusion. (Fig. 3, A-C).
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Nimodipine perfusions: normal cochleas
In contrast to the results obtained with both artificial
perilymph and artificial perilymph containing 100 µM
Ni2+ or 0.1% DMSO, perfusion with artificial
perilymph containing 0.01-0.1% DMSO and nimodipine at concentrations
ranging from 1 to 10 µM, respectively, caused marked changes in CAP
and SP visual detection thresholds and input-output curves. For
concentrations of nimodipine 3 µM, the effects were generally
reversible over a period of tens of minutes after the cessation of
perfusion, and this reversibility could be accelerated by reperfusion
with control perilymph. At concentrations of >3 µM, reversibility
was very slow and could not always be demonstrated. Figure
4 shows the effects on CAP input-output
curves for two frequencies of tone burst, 10 kHz and 20 kHz. Similar to
the results obtained by Bobbin et al. (1990)
for 10-kHz
stimulation, there was a shift to the right in the curves and a
reduction in the maximum CAP amplitude for both frequencies.
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To quantify the relationship between CAP and SP threshold changes, we
used 20-kHz tone burst stimulation in preference to 10 kHz, because the
recording electrode was placed near the high-frequency end of the
cochlea in the basal turn scala tympani immediately adjacent to the
round window. It is known that whereas CAP recordings of tone bursts
are relatively unbiased by electrode location (Johnstone et al.
1979), the SP recording is more localized and, particularly near threshold, reflects to a large extent the cellular responses in
the immediate vicinity of the recording electrode (Cheatham and
Dallos 1984
). Therefore, a 10-kHz tone would faithfully probe the neural sensitivity of the 10-kHz region of the cochlea, but the SP
recording would be contaminated by cellular responses from more basal
regions in the immediate vicinity of the recording electrode.
The results in Fig. 5, A-D, show that the shift in visual detection threshold produced by nimodipine is strikingly less for the SP than for the CAP. The average CAP threshold changes were 16, 30, and 49 dB in 1, 3, and 5 µM nimodipine, respectively. This contrasted with changes of only 10, 12, and 15 dB in the respective average SP thresholds. An action of nimodipine restricted to the OHC active process would affect the overall cochlear gain and would therefore be expected to cause equivalent changes in CAP and SP threshold. The result shown in Fig. 5 is therefore not consistent with a pure OHC effect. Rather, the proportionately greater effect on the CAP threshold suggests a dominant action of nimodipine on the final stage of nerve fiber excitation (the IHC-afferent nerve fiber synapse).
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Salicylate perfusions
It was important to test the validity of our technique of
measuring corresponding CAP and SP thresholds as a means of dissecting the possible sites of action of nimodipine. We therefore chose to
repeat the basic measurements with intracochlear perfusion of
salicylate, a drug known to affect the OHC active process selectively (Fitzgerald et al. 1993; Stypulkowski
1990
). Three animals were perfused with artificial perilymph
containing 10 mM salicylate, and the SP and CAP thresholds were
measured for 20-kHz acoustic stimuli before and during the perfusion.
It can be clearly seen from the data presented in Fig.
6 that in contrast to the nimodipine result, CAP and SP changes caused by salicylate were essentially identical over a wide range. This result shows that the technique can
distinguish between drug actions that affect the OHC active process and
those that affect other stages of the transduction chain.
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Nimodipine perfusions: kanamycin animals
Although nimodipine caused a markedly greater change in CAP than
SP threshold, which points to a dominant action on stages of
transduction after the OHC active process, there was still a
significant SP threshold change caused by all concentrations of
nimodipine. In an attempt to determine the cause of this change in
another group of animals, the OHCs over a restricted region of the
cochlea were destroyed by prior administration of kanamycin. Figure
7 shows examples of the CAP thresholds
from 2 to 30 kHz (so-called CAP audiograms) obtained in animals
administered kanamycin, compared with the average CAP audiograms in
normal animals. There was considerable variability in the effects of
kanamycin, but in all cases, there was a marked elevation of the CAP
threshold at high frequencies. This correlated with the degeneration of OHCs confirmed in subsequent histology (results not shown). It is well
established from previous work that CAP sensitivity losses of 50 dB
are to be expected in regions of the cochlea with total loss of OHCs
(Patuzzi and Rajan 1992
).
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We managed to obtain six animals in which the threshold loss at 20 kHz was ~40-60 dB (absolute threshold ~80 dB sound pressure level). The average threshold change at 20 kHz was 49 dB. Despite these greatly elevated thresholds caused by OHC destruction, when nimodipine perfusions were performed in these animals, CAP and SP threshold changes were still observed. Only a small number of perfusions using 5 µM nimodipine were performed, and the data shown in Fig. 8, A-C, are for perfusions with 1-3 µM only. As shown in Fig. 8, CAP threshold changes were comparable to those produced by the same concentrations of nimodipine in normal animals. On the other hand, the SP threshold changes were significantly less. For example, perfusion with 3 µM nimodipine caused mean CAP and SP threshold changes of 30.4 and 12 dB, respectively, whereas in kanamycin-treated animals, the threshold changes were 29 dB for the CAP and only 7 dB for the SP. When data from 1- and 3-µM perfusions were pooled (n = 10), the difference between CAP threshold changes in the normal and kanamycin groups were not significant (P > 0.95), whereas the differences in SP threshold changes were important (P < 0.05). This result suggests, first, that the CAP threshold changes caused by nimodipine are not dependent on intact OHCs and, second, that a substantial portion of the SP threshold change seen during nimodipine perfusion in the normal cochlea is somehow caused by an action of nimodipine on the OHCs.
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In an attempt to deduce how much of the small nimodipine-induced
SP threshold change that still remained after OHC destruction was
caused by contamination of the SP recording by neural, rather than hair
cell activity, we performed experiments in five kanamycin animals using
perfusion with TTX (1 µM) to eliminate action potentials by blockage
of voltage-gated Na+ channels (Narahashi
1974). The success of TTX in blocking all neural activity was
assessed by the complete disappearance of the CAP. Subsequent
nimodipine perfusion (3-µM concentration) caused almost no change in
the SP thresholds or input-output curves. The mean SP threshold change
caused by nimodipine after TTX was only 2 dB (n = 5). This was within measurement error, and, when compared with the mean
SP threshold change when TTX was not present, the difference was highly
significant (P < 0.001)
The effects of nimodipine on SP thresholds in the various experimental groups are compared in Fig. 9, which shows the average 20-kHz SP threshold change observed after 10 min of 3 µM nimodipine perfusion (n = 5 for each group of animals). SP threshold changes caused by nimodipine were greatest in normal animals, were reduced substantially by loss of OHCs, and were insignificant when neural activity was blocked by TTX.
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DISCUSSION |
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These results show that under particular conditions, an L-type Ca2+ channel blocker can cause changes in the gross neural potentials (CAP) in the cochlea that are more or less dissociated from the major DC receptor potential (SP). We interpret this result to mean that this blocker is acting on the IHC-afferent nerve fiber synapse. Figure 10 shows schematically the logic behind this conclusion. As the figure attempts to show, an effect of nimodipine on the OHCs alone would be expected to cause proportional alterations in SP and CAP thresholds through an effect on the gain of mechanical drive to the IHCs. This expected result was obtained using salicylate, which is well known to affect the OHC active process. On the other hand, an action restricted to the final stages of neural excitation at the base of the IHCs should lead to a greater change in CAP than SP thresholds (some small effect on SP thresholds might still be expected, as shown in the Fig. 10, if there were a second-order effect on OHCs as well). A result approximating this latter prediction was seen when a 20-kHz tone was used, and agreement with the prediction was even better when OHCs were removed by the use of kanamycin. Finally, it was shown by the use of TTX that the small residual SP threshold change caused by nimodipine in the absence of OHCs was probably due to contamination of the SP response by neural activity.
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We believe these results constitute evidence that L-type Ca2+ channels exert a major and direct controlling influence on one of the final stages of transduction in the inner ear. Given the association of voltage-gated Ca2+ channels with transmitter release sites in other hair cell systems, we feel it is most likely that these L-type channels control the influx of Ca2+ at specific presynaptic release sites located around the basolateral pole of the inner hair cells.
The possibility should be considered that nimodipine may produce these
effects in other ways than through blockage of voltage-gated Ca2+ channels at the presynaptic locus of vesicle
exocytosis from the IHCs. For example, it is possible that the action
could be exerted postsynaptically on the primary afferent dendrites.
However, the fact that TTX virtually eliminates the CAP suggests that
voltage-gated Ca2+ channels in the auditory
dendrites do not contribute a major component to postsynaptic currents
recorded by our techniques. The possibility should also be considered
that nimodipine may alter the standing current through the IHCs,
perhaps by changing the large DC endocochlear potential (EP) known to
be present in the scala media (Sellick and Johnstone
1972). Bobbin et al. (1990) have reported that
nimodipine perfusion, if it produces any change in the EP, does so by
only a few millivolts (a mean decrease in EP of 3 mV). This would have
a trivial effect on the overall electrochemical driving force of some
130-140 mV across the IHC transduction channels but could have a
significant effect on evoked transmitter release depending on the
precise relationship between IHC membrane potential and exocytosis.
Sewell (1984)
has reported that single afferent fiber
acoustic thresholds change by ~1 dB for every millivolt change in EP,
which suggests that any change in gross thresholds caused by such an
effect of nimodipine would be probably within measurement error.
Alternatively, nimodipine may affect the IHC mechanoelectrical
transduction process or the IHC membrane potential. The most plausible
mechanism for such an effect would be an indirect one, through
Ca2+-activated K+ channels.
Although such channels have been extensively described in nonmammalian
hair cells, the available evidence suggests that if they exist in
mammalian inner hair cells, they are not gated by
Ca2+ entry through the plasma membrane
(Dulon et al. 1995; Kros and Crawford
1990
). We believe that these possibilities are inconsistent with our findings in kanamycin-treated animals, because in the kanamycin-treated cochlea poisoned with TTX, the SP observed should be
purely derived from the IHCs; therefore, the absence of effect of
nimodipine on the SP indicates that it has little or no action at the
level of IHC transduction.
Despite our contention that the results presented in this study are
consistent with involvement of L-type channels at the IHC-afferent
nerve fiber synapse, L-type channels may also be involved in OHC
function. Certainly, dihydropyridine-sensitive channels have been
demonstrated in OHCs using other methods (Chen et al.
1995). Some aspects of our own data are also consistent with
this notion. For example, the 20-kHz SP threshold change caused by
nimodipine was substantially reduced when the OHCs were destroyed in
this region of the cochlea. This implies that nimodipine may alter the
OHC gain as well as act at the IHC synapse. Second, we found in one
group of normal animals, by using 10-kHz stimulation, that TTX
perfusion did not eliminate the effect of nimodipine on the SP (results
not shown). Thus, neural contamination of the SP recording cannot fully
account for the observable SP threshold change caused by blockage of
L-type channels when the OHCs are present. Finally, although we have
not reported the results in detail in this article, we have confirmed
the observation by Bobbin et al. (1990)
that certain
concentrations of nimodipine can cause a reversal in the polarity of
the SP at some sound pressure levels. This was most evident for 10-kHz
stimulation at sound pressures ~80 dB SPL and in the presence of 5 µM nimodipine. We observed it only rarely and inconsistently for
20-kHz stimulation. A possible explanation for this reversal of SP is
that nimodipine may cause the OHCs to generate a substantial SP of
opposing polarity to the IHCs. The possibility that OHCs may generate
an abnormal and reversed-polarity SP in the presence of nimodipine
provides a possible explanation for the fact that SP changes caused by
nimodipine are less in kanamycin-treated animals compared with normal
subjects, whereas the CAP threshold changes are the same. The SP
threshold change in the presence of OHCs may not reflect a gain change
caused by nimodipine, but the addition of negative and positive
contributions to the SP from the two sets of receptor cells. If this is
correct, it would add even more weight to the notion that nimodipine
produces changes in neural sensitivity (CAP threshold) by an action on the IHC-afferent synapse.
There are a number of outstanding questions with regard to the identity
of the channels involved both in IHCs and OHCs. The IHC resting
potential is between 40 and
50 mV (Russell and Sellick 1978
) and the maximum depolarization in response to nondamaging levels of sound has been reported to be ~10 mV (Patuzzi and
Sellick 1983
). Thus, the least negative value of membrane
potential expected in these cells is well below the normal operating
range of classic L-type channels (Tsien et al. 1991
),
and therefore, although the channels are, similar to classic L-type
channels, blocked by nimodipine, they are clearly unusual in their
voltage dependence. Recent molecular biological studies on calcium
channels in chick (Kollmar et al. 1997a
,b
) and mouse
(Green et al. 1996
) suggest strongly that alternative splicing of mRNAs encoding for one or more of the subunits of the
L-type channel may be an explanation for this unusual
voltage-sensitivity of the nimodipine-sensitive channels in the inner
ear. A similar situation may exist in vertebrate photoreceptors in
which membrane potentials of
35 to
40 mV lead to constant
calcium-dependent release of transmitter in the dark (Schmitz
and Witkovsky 1997; Wilkinson and Barnes 1996).
From gross potential measurements we cannot say that only L-type
channels are involved in the IHC. There is a diversity of single
primary afferent fiber characteristics, in spontaneous rates,
thresholds, and adaptation characteristics, and we cannot exclude the
possibility that a small subgroup of primary afferents has their
excitation controlled by other subtypes of presynaptic calcium channel.
For example, highly sensitive neurons with rapid adaptation may be
controlled by hair cell channels with characteristics more akin to
T-type than L-type. In the present experiments, we attempted to test
this notion using the selective T-type channel blocker
Ni2+, but could see no effect on either CAP or
SP, even though the concentrations used were well in excess of those
known to block T-type channels in other systems (Fox et al.
1987). This result, however, is largely inconclusive, because
an Ni2+-sensitive subgroup of neurons may be
insufficient in number to contribute significantly to the gross neural
response. Furthermore, we have not yet tried perfusion with
specific blockers of other types of Ca2+ channels, such as
the various conotoxins and agatoxins, which are selective for N-, P-,
and Q-type channels. Such future experiments seem important in view of
the evidence suggesting that N-type channels may operate in addition to
L-type channels in hair cells of the frog sacculus and crista
ampullaris (Prigioni et al. 1992
; Su et al.
1995
). In addition, the gross potential measurements reported
in the current study cannot tell us whether L-type channels control the
spontaneous release of transmitter from IHCs and the background firing
rates of the auditory afferent neurons, although perfusion with
nonspecific Ca2+ channel blockers (Robertson and
Johnstone 1979
) have clearly shown that the spontaneous
discharge of single auditory afferents is under the control of
Ca2+ channels. Further experiments will be required to
address these latter issues.
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ACKNOWLEDGMENTS |
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The authors thank G. Bennett for animal care and solution preparation and R. Patuzzi for helpful discussion.
This research was supported by grants from the National Health and Medical Research Council, The Australian Research Council, and The University of Western Australia. S. Y. Zhang was the recipient of an AusAid grant.
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FOOTNOTES |
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Address for reprint requests: D. Robertson, The Auditory Laboratory, Dept. of Physiology, The University of Western Australia, Nedlands, WA 6907, Australia.
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.
Received 7 June 1999; accepted in final form 10 August 1999.
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REFERENCES |
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