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The Journal of Neuroscience, July 1, 1999, 19(13):5275-5292
The Electrical Properties of Auditory Hair Cells in the Frog
Amphibian Papilla
Michael S.
Smotherman and
Peter M.
Narins
Department of Physiological Science, The University of California,
Los Angeles, California 90095-1527
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ABSTRACT |
The amphibian papilla (AP) is the principal auditory organ of the
frog. Anatomical and neurophysiological evidence suggests that this
hearing organ utilizes both mechanical and electrical (hair cell-based)
frequency tuning mechanisms, yet relatively little is known about the
electrophysiology of AP hair cells. Using the whole-cell patch-clamp
technique, we have investigated the electrical properties and ionic
currents of isolated hair cells along the rostrocaudal axis of the AP.
Electrical resonances were observed in the voltage response of hair
cells harvested from the rostral and medial, but not caudal, regions of
the AP. Two ionic currents, ICa and
IK(Ca), were observed in every
hair cell; however, their amplitudes varied substantially along the
epithelium. Only rostral hair cells exhibited an inactivating potassium
current (IA), whereas an inwardly
rectifying potassium current (IK1)
was identified only in caudal AP hair cells.
Electrically tuned hair cells exhibited resonant frequencies from 50 to
375 Hz, which correlated well with hair cell position and the tonotopic
organization of the papilla. Variations in the kinetics of the outward
current contribute substantially to the determination of resonant
frequency. ICa and
IK(Ca) amplitudes increased with resonant
frequency, reducing the membrane time constant with increasing resonant
frequency. We conclude that a tonotopically organized hair cell
substrate exists to support electrical tuning in the rostromedial
region of the frog amphibian papilla and that the cellular mechanisms
for frequency determination are very similar to those reported for
another electrically tuned auditory organ, the turtle basilar papilla.
Key words:
hair cells; hearing; electrical tuning; electrical
resonances; frequency discrimination; frogs; Rana pipiens
pipiens; amphibian papilla; calcium currents; potassium
currents
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INTRODUCTION |
The frog amphibian papilla (AP)
processes acoustic stimuli within a frequency range of 100 to ~1250
Hz (Feng et al., 1975 ; Lewis et al., 1982a; Ronken, 1991). It is
tonotopically organized (Lewis et al., 1982a), with the lowest
frequencies being encoded by hair cells in the rostral patch and
progressively higher frequencies being transduced more caudally.
Electrical resonances in the membrane potential, which may constitute
an electrical tuning mechanism (Crawford and Fettiplace, 1981), have
been reported for rostral and medial hair cells of the AP in
vitro, with the frequency of membrane oscillations closely
overlapping the known tonotopic map of the AP (Roberts et al., 1986;
Pitchford and Ashmore, 1987). Hair cell recordings from the caudal AP
have not been reported. The degree to which hair cell electrophysiology
is position-dependent within the frog AP offers insight into the
physiology of hearing in a comparatively primitive terrestrial ear.
The frog AP is considered analogous to the primary auditory organs of
fish, reptiles, and birds (Lewis and Narins, 1998). In the goldfish
sacculus and the chick basilar papilla (BP), two populations of hair
cells have been classified based on their position within the
auditory epithelium and the collection of basolateral membrane ionic
currents they possess (Fuchs et al., 1988; Fuchs and Evans, 1990;
Sugihara and Furukawa, 1989; Eatock et al., 1993; Murrow, 1994), and in
the turtle BP hair cell, electrical properties have been shown to vary
tonotopically (Crawford and Fettiplace, 1981; Art and Fettiplace, 1987;
Goodman and Art, 1996a,b). In these cases, the waveform of the hair
cell receptor potential is typically driven by a voltage-dependent
calcium current, ICa, and either a
calcium-dependent potassium current,
IK(Ca), or a voltage-dependent potassium
current, IK. In the turtle BP and frog sacculus
(Crawford and Fettiplace, 1981; Hudspeth and Lewis, 1988; Goodman and
Art, 1996a), these currents have been shown to modulate the behavior of
the hair cell basolateral membrane as an electrical resonant
filter. This "electrical tuning" (Crawford and Fettiplace,
1981; Art and Fettiplace, 1987; Hudspeth and Lewis, 1988) represents
the contribution of the hair cell to the spectral acuity of the
auditory afferent nerve fiber it contacts. Other ionic currents, such
as IA, Ih,
and IK1, have been identified in
vertebrate auditory hair cells, but their functional significance remains poorly understood.
This study characterizes the electrical properties of hair cells
located in the rostral, medial, and caudal regions of the leopard frog
AP. We define separate rostral and caudal populations of AP hair cells
based on their excitability and their compliment of ionic currents. We
found that, despite the presence throughout the AP of those currents
known to mediate electrical tuning (ICa and
IK(Ca)), electrical tuning per se was not
evident throughout the entire amphibian papilla. For those hair cells
exhibiting electrical resonances, we found that the distribution of
resonant frequencies was consistent with a tonotopic gradation in the
magnitude and kinetics of the underlying ionic currents.
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MATERIALS AND METHODS |
Dissociation of hair cells. Amphibian papillae were
dissected out of pithed and decapitated adult northern leopard frogs
(Rana pipiens pipiens), sectioned by eye into three parts
(approximately rostral, medial, and caudal thirds) (Fig.
1), which were treated separately for 20 min at room temperature with 500 µg/ml papain (purchased from
either Calbiochem, San Diego, CA, or Sigma, St. Louis, MO; both equally
effective) dissolved in a dissociation solution containing (in
mM): NaCl 120, KCl 5, CaCl2 0.1, D-glucose 3, and HEPES 10, pH 7.2. Each segment was then
transferred to a dissociation solution with bovine serum albumin
(500 µg/ml), replacing papain for 30-45 min at <10°C. Papillar
sections were then transferred to a 0.3 ml recording dish containing
dissociation solution alone, and hair cells were gently scraped free
with a tungsten needle. The hair cells settled but did not adhere
firmly to the sylgard base of the recording chamber. The dissociation solution was then replaced via a continuous perfusion system with a
perilymph-like (Bernard et al., 1986) standard external recording solution (Table 1). For the purpose of
assessing the potentially detrimental effects of papain on hair cell
electrical properties, such as those reported for frog saccular hair
cells (Armstrong and Roberts, 1998), a series of experiments was
performed on hair cells dissociated without the use of papain. For
these experiments, hair cells were incubated in a zero calcium-added
solution (rather than 0.1 mM calcium) buffered with 2 mM EGTA for a period of 40-60 min at <10°C. This method
resulted in a lower hair cell yield but usually provided normal-looking
hair cells. We present evidence here that papain did not appear to
either remove or reduce the amplitude of any ionic currents present in
AP hair cells, and generally we observed no difference in the
electrical properties of hair cells isolated with or without papain
used as a dissociative agent.
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Figure 1.
Map of the AP sensory epithelium. The arrangement
of sensory hair cells within the chamber of the leopard frog AP
is typical of most ranid frogs. The epithelium is located on the inner
dorsal surface of the AP. The AP nerve branchlet exits the VIII nerve
and passes around the AP ventrally and laterally before entering the
dorsolateral edge of the AP. The outline of the sensory epithelium is
visible through a dissecting microscope once the ventral wall of the AP
chamber is excised. Dotted lines indicate where the AP
was cut into three sections before hair cell dissociation. The
approximate range of frequencies encoded by each section is taken from
Lewis et al. (1982b).
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The recording chamber was placed on the stage of an inverted microscope
(Diaphot; Nikon, Tokyo, Japan) equipped with a 40× objective with
phase contrast optics. The positions of the infusion and vacuum
pipettes within the recording chamber were adjusted to allow for the
most rapid flow while minimizing cell movement. It was generally true
that after 15 min the flow speed could be substantially increased
without washing away the target hair cells. Electrode tip junction
potentials (13 mV, pipette negative) were added as in Fenwick et al.
(1982). A series of ionic and pharmacological agents was incorporated
into the recording solution (Table 1). The exchange of recording
solutions was achieved via continuous gravity perfusion of the entire
recording chamber at a rate of ~1 ml/min. The exchange was allowed to
equilibrate for at least 3 and typically 5 min before experiments
continued and was maintained until no further change was observed in
the ionic currents. Recordings were stable and consistent enough to
allow several solution exchanges and reversals.
Whole-cell recordings. Currents and voltages were recorded
with the conventional whole-cell tight-seal patch-clamp technique (Hamill et al., 1981). Borosilicate glass pipettes were pulled with a
Narishige two-stage vertical pipette puller (model PP-83; Narishige,
Tokyo, Japan) to tip diameters of ~1 µm. Electrode resistances
typically ranged from 2 to 10 M . Series resistances (RS) during recordings ranged from 6 to
20 M and were compensated 60-95% during voltage-clamp recordings
using the compensation circuitry of the amplifier. Cell capacitances
ranged from 4 to 20 pF. Cell capacitances and uncompensated series
resistances implied a range of voltage-clamp time constants between 1.2 and 140 µsec, although minor compensation-induced transients limited most experiments to >50 µsec settling times. Cell capacitance and
series resistance values were read from the compensation dials of the
amplifier and found to be within 5% of those calculated by fitting a
single exponential function to a capacity transient and estimating
Cm from the area under the curve
(Cm = Q/ V, where Q is charge and V was a 5 mV step) and
RS from Cm and the time constant of the curve (RS =  /Cm). Series resistance and cell capacitance compensation were updated continuously throughout all
experiments. During long-lasting experiments, cell capacitance values
taken from the amplifier typically increased 10-20% over an hour.
This increase in cell capacitance has been observed previously in frog
saccular hair cells and appears to be accounted for by calcium-triggered exocytosis (Parsons et al., 1994). Linear leak subtraction (real-time analog or digital) was generally not included as
part of the experimental protocol. Leak substraction and corrections for voltage errors caused by RS were
performed off-line during data analysis. Typical leak conductances
measured at ±10 mV at the holding potential of approximately  60 mV
were 0.2-2.0 nS in rostral hair cells and 3.0-8.0 nS in caudal hair
cells. Averaging was performed for a subset of cells but not generally
included in the experimental protocol. The Axopatch 200A (Axon
Instruments, Foster City, CA) was used for all current-clamp and
voltage-clamp experiments. For current-clamp experiments, the Axopatch
200A has separate fast and slow current-clamp modes. Oscillatory
voltage responses recorded in the slow mode (which incorporates a
stabilizing circuit) frequently had excessive or unrealistically high
quality factors (Qe values), suggesting
the presence of a contaminating electrical feedback from the amplifier.
Switching to the fast current-clamp mode eliminated any evidence of
feedback and sharply reduced these Qe values to
within values typically reported for electrically tuned hair cells in
the literature. Therefore, the fast current-clamp mode was the standard
recording mode for studying membrane oscillations. The frequency
response of both current-clamp modes was examined empirically. For a
constant amplitude, swept-frequency sinusoidal input to the amplifier,
the gain of the system in the fast mode was essentially flat (±2 dB)
up to 2 kHz, well above the highest characteristic frequency (CF)
reported for the auditory afferents innervating the AP (Ronken, 1991).
The frequency response of the amplifier in the slow current-clamp mode
closely resembled that for the fast mode and exhibited a broad peak (+3
dB) at ~1 kHz. Stimuli were generated and data were sampled with a
12-bit digital-to-analog and analog-to-digital converter (Digidata
1200; Axon Instruments) and controlled by the data acquisition software package pClamp 5.5 (Axon Instruments). Sampling intervals were tailored
to the kinetics of the study; in general, however, voltage-clamp data
were sampled at 10 or 20 µsec intervals, and current-clamp data were
sampled at 50-100 µsec intervals. Voltage and current waveforms were
low-pass filtered by the amplifier with a 2 kHz cutoff frequency.
Experiments were performed at room temperature (19-20°C).
Data analysis. Current-clamp and voltage-clamp data were
stored digitally and later analyzed off-line using the pClamp 5.5 Clampfit program (Axon Instruments). For determining activation and
deactivation time constants, exponential curves were fit using a
least-squares algorithm in pClamp. Boltzmann curves were fit using the
program SigmaPlot 4.0 (Jandel Scientific, Corte Madera, CA), using the
Marquardt-Levenberg algorithm. Boltzmann functions presented in this
paper are of the form:
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(1)
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for activation plots and a similar appropriately modified
function for inactivation plots, where
I/Imax is the relative current (Imax is the largest amplitude tail current),
V1/2 is the midpoint of activation
(V1/2(a)) [or inactivation
(V1/2(in))] in millivolts, Vm is the membrane potential in millivolts, and
k (the Boltzmann constant) is a slope factor. Tail current amplitudes
at step offset were estimated by fitting a single exponential function
to the tail current and extrapolating back to the time of step offset. Steady-state current amplitudes were determined by averaging the last
50 msec of a 125 msec voltage step. Peak currents were determined by
eye during computer analysis. Resonant frequency was calculated as the
inverse of the mean period between successive oscillatory peaks, either
at the onset or offset of a constant current pulse. We define the
resonant frequency ( ) of a hair cell at 53 mV, which for most
cells coincides with the membrane potential at which oscillatory
quality (Qe) peaked and by the addition
of a standing current or through adjustments in current step amplitude could be the designated potential for onset or offset oscillations. Onset or offset oscillations for a given cell will have identical resonant frequencies if recorded (separately) around a common membrane
potential. Qe was determined as by Crawford and
Fettiplace (1981), using the equation Qe = [( )2 + 0.25]1/2, where is the
resonant frequency at 53 mV and is the time constant of the decay
of the oscillatory peaks. A small standing current was used when
necessary to raise the standing potential of a cell to 53 mV before
the application of a depolarizing current pulse, thus allowing for
offset oscillations to occur at approximately 53 mV in all cells for
purposes of comparison. Results are presented as mean ± SD unless
otherwise noted.
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RESULTS |
Current-clamp recordings
Resting potentials
Based on hair cell resting potentials, two populations of hair
cells are recognizable in the AP: a rostral population resting at
approximately 60 mV and a caudal population resting at approximately 75 mV. Hair cells were examined in external recording solutions that
contained either 5 or 2 mM K+. The 2 mM [K+]ext condition is
closest to the reported frog perilymph composition (~2.5
mM K+) (Bernard et al., 1986). Medial
hair cells could be divided into subgroups based on the presence or
absence of an inward rectifier current
(IK1), which has been shown in leopard
frog saccular hair cells to contribute predictably to the in
vitro hair cell resting potential (Holt and Eatock, 1995). In 5 mM [K+]ext, rostral
hair cells and medial hair cells without IK1 had resting potentials of approximately 60 mV [rostral, 60.4 ± 1.7 (mean ± SE); n = 11; medial, 61.0 ± 2.6; n = 7), which was not significantly different from
cells recorded in 2 mM
[K+]ext (rostral, 63.3 ± 1.5;
n = 26; medial, 62.5 ± 1.0; n = 31). Caudal hair cells (all of which have
IK1) and medial hair cells with
IK1 had resting potentials of approximately 70
mV in 5 mM [K+]ext (medial
with IK1, 70.2 ± 2.4;
n = 6; caudal, 71.7 ± 1.0; n = 23) and approximately 75 mV in 2 mM
[K+]ext (medial with
IK1, 76.5 ± 2.7;
n = 8; caudal, 74.6 ± 1.9; n = 23). Although the observed difference in resting potentials is
reflective of a differential distribution of
IK1, there is reason to suspect that
these potentials are not accurate representations of the in
situ condition. Isolated hair cells may be resting at abnormally
negative potentials as a result of the absence of a resting transducer
current, because the transduction apparatus is known to be disrupted by
zero-calcium conditions (Assad et al., 1991), such as those used in our
dissociation procedure. Contradictions in in situ and
in vitro resting potentials have been identified in turtle
BP hair cells; although IK1 is selectively distributed among hair cells of the turtle BP (Goodman and Art, 1996b),
measurements of in situ resting potentials have not
uncovered comparable differences in hair cell resting potentials
(Crawford and Fettiplace, 1981).
Voltage responses and electrical resonances
We examined the voltage responses (Fig.
2) of hair cells isolated from the
rostral, medial, and caudal regions of the AP. We found that electrical
resonances in the membrane potential could be induced by small
depolarizing current pulses (>20 pA) in all cells isolated from the
rostral region and in approximately two-thirds of the cells located
medially, whereas very few of the caudal cells responded with
oscillations (Table 2). The range of
resonant frequencies observed in rostral and medial hair cells was
50-375 Hz at 53 mV. The few caudal hair cells that resonated (9 of
29 in 2 mM [K+]ext)
had resonant frequencies ranging from 160 to 270 Hz. Electrical resonances were studied in external recording solutions that contained either 2 or 5 mM [K+]ext.
The quality of electrical resonances is considerably greater in the
lower potassium concentration, increasing the likelihood that
measurable resonances would be observed and also enhancing our ability
to accurately measure the voltage-dependent properties of electrical
resonance. Cells were classified as oscillatory if any depolarizing
current pulse could induce voltage oscillations with a
Qe >1.0 within a physiologically reasonable
range of membrane potentials (±10 mV) centered around the threshold of
activation for the voltage-dependent calcium current (55 mV). For
rostral and medial hair cells, this range coincides reasonably with the typical in vitro resting potential, but for caudal cells,
the resting potential is typically 15-20 mV negative to the threshold for gCa. To remove this difference, caudal cells
were first stimulated from their zero-current membrane potential with
current pulse amplitudes chosen to produce steady-state membrane
potentials at or slightly positive to 55 mV (Fig.
2B); they were next stimulated with smaller amplitude
current pulses from that holding voltage, which was artificially
maintained with a standing depolarizing current (Fig. 2C).
Most caudal cells responded with a brief peak depolarization that
quickly decayed to a steady-state plateau, regardless of holding
potential, which resembled an overdamped oscillatory voltage response.
Oscillatory and nonoscillatory cell types appeared to be separated into
approximately rostral and caudal halves of the AP.
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Figure 2.
Variations of the voltage response of isolated AP
hair cells. Current-clamp recordings of the voltage response to a
depolarizing current pulse for representative rostral and caudal hair
cells. A, Rostral hair cells always exhibited
oscillatory responses, and many exhibited membrane potential
oscillations at rest. Here, a rostral hair cell is first depolarized by
a 20 pA constant current, which is later reduced to a 10 pA current
before being turned off. Oscillations occurred at both the onset and
transition of the current pulse because the steady-state (direct
current) potential was above the threshold for
ICa. Both resonant frequency and
Qe were higher at the onset of the 20 pA
pulse than at its offset (230 Hz; Qe of 13;
steady-state voltage of 52 mV; vs 181 Hz;
Qe of 8, Vss of
55 mV). B, Caudal cells typically respond to
depolarizing current pulses with a peak that rapidly decays to a
plateau, sometimes resembling an overdamped oscillatory response. In
this caudal cell, a 150 pA stimulus applied at the zero-current
potential of 78 mV of the cell depolarized the membrane potential to
a steady-state potential of 53 mV (the potential at which
Qe of electrical resonances would typically
be highest for oscillatory rostral hair cells). C, The
same caudal hair cell with its membrane potential artificially
maintained at approximately 55 mV before a 20 pA depolarizing
stimulus. Medial hair cell voltage responses could be characterized as
one of these two categories (Table 2). Rostral hair cell, 9 pF,
3 M uncompensated series resistance (usr).; caudal hair cell, 4 pF,
1.8 M usr.
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Electrical resonances observed in rostromedial hair cells of the AP
appeared qualitatively similar to those reported for hair cells of the
bullfrog sacculus (Hudspeth and Lewis, 1988) and turtle basilar papilla
(Art and Fettiplace, 1987). The frequency and quality factor
(Qe) of oscillations induced by a current
pulse are voltage-sensitive. Figure 3
shows that onset oscillation frequency increases as progressively
larger depolarizing current pulses were applied to the cell. This is
presumably caused by the increased calcium entry with greater
depolarization resulting in a more rapid opening of the
calcium-dependent potassium channels. Qe is
sensitive to the amplitude of the whole-cell currents and, in
particular, the ratio of the inward to the outward currents, which
changes with membrane potential. As a result, Qe
is typically highest just above the threshold for
gCa (Fig. 3C). We typically observed
resting oscillations in hair cells whose zero-current membrane
potential was close to or positive to the threshold for ICa. These "spontaneous" oscillations were
also observed if the membrane potential of the cell was depolarized
positive to 55 mV by a steady current.
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Figure 3.
Voltage-dependent properties of
electrical resonance. A, An example of electrical
resonances recorded in a rostral hair cell. Depolarizing current pulses
of increasing amplitude will increase the onset resonant frequency but
diminish the quality factor (Qe) of
the onset oscillations. Offset oscillation frequency remains unchanged.
This cell is not included in B and C.
B, As input current is increased, the steady-state
membrane potential around which the onset oscillations occur is
increasingly depolarized, resulting in higher frequency electrical
resonance. C, The quality of electrical resonances
(Qe) is found to be maximal within 2 or 3 mV of the threshold for the voltage-dependent calcium current. The
hair cells represented here (same cells as in B) had
maximal onset Qe values of ~20; however, a
small standing current could be used in each of the cells shown to
generate continuous oscillations near the resting potential, which
corresponds to an infinite Qe. Open
circles, Medial hair cell, 8 pF, 245 Hz at peak
Qe of 20; Vh of
54 mV; filled circles, medial hair cell, 10 pF, 155 Hz
at peak Qe of 13.7;
Vh of 54 mV; filled
diamonds, rostral hair cell, 16 pF; 80 Hz at peak
Qe of 21; Vh of
53 mV. Curves in B were drawn by eye,
and curves in C are interpolated.
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Characterization of the ionic currents
Overview of the ionic currents
We identified and characterized the four largest ionic currents
found in AP hair cells. ICa,
IK(Ca), IA,
and IK1 were all found to be expressed in
significant amplitudes in a large number of hair cells and provided
sufficient grounds for the classification of two hair cell subtypes
within the AP: a rostral population likely to possess
IA, and a caudal population likely to
possess IK1. We encountered evidence of other
ionic currents that were of substantially smaller amplitudes and/or in
such limited numbers that a full characterization of them could not be
included in this report. In particular, we present evidence that at
least one additional voltage-dependent potassium current,
IK, appears to be expressed in hair cells
throughout the AP (although not apparent in every hair cell). The
distribution and pharmacology of IK remains
under investigation. We also saw evidence of a second inward rectifying
potassium current, Ih, in a very small
subset of cells. The presence of the four primary ionic currents was assessed for all three regions (rostral, medial, and caudal) of the AP
from which cells were isolated. Given the apparent distribution of two
hair cell subtypes, an extensive ionic and pharmacological characterization of the four ionic currents is presented for rostral and caudal hair cells, and medial hair cells included in the analysis were classified as rostral or caudal "types" based on whether they
possessed IA or IK1. In
the following sections, we will present ionic and pharmacological data
comparing results derived from rostral and caudal hair cells.
Oscillatory and nonoscillatory hair cells have different ionic
current components
In voltage-clamp mode, three ionic currents were identifiable
under control conditions. Figure 4
demonstrates how representative rostral and caudal hair cells were
tested for IK(Ca),
IA, and IK1. All
AP hair cells exhibited a noninactivating outward current, IK(Ca), after depolarization to membrane
potentials positive to 55 mV (Fig.
4A,C). For 97% of the rostral hair
cells and 58% of the medial hair cells (Table 2), a hyperpolarizing
prepulse before depolarization recruited an additional slowly
inactivating (inact of ~70-150 msec)
voltage-dependent outward potassium current, IA
(Fig. 4B). Only 16% of the hair cells isolated from
the caudal region possessed an inactivating outward current; most
caudal hair cells responded as shown in Figure 4D.
Hyperpolarizing 90% of the caudal hair cells evoked a large and
fast-activating voltage-dependent current that reversed polarity at
EK, becoming an inward current negative
to EK. This inward potassium current,
IK1, was also found in 20% of the medial
hair cells but in only 10% of the hair cells isolated from the rostral
patch. Table 2 compares the distribution of oscillatory and
nonoscillatory hair cells (determined as described above) with the
distribution of the two voltage-dependent potassium currents
(IA and IK1).
There is a strong correlation between oscillatory hair cells and the
presence of IA and between nonoscillatory hair cells and IK1. However, our data also show that
voltage oscillations are neither dependent on nor exclusive to either
current; 4 of 34 hair cells from the medial AP that possessed
IK1 also exhibited typical electrical
resonances, whereas, conversely, 9 of 34 oscillatory medial hair cells
did not display IA. We can now separate hair cells into two populations in the AP based on position, resting potential, voltage response, IA distribution,
and IK1 distribution.
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Figure 4.
Voltage-dependent activation of ionic
currents. Two voltage-clamp protocols were used to rapidly assess the
presence of three different potassium currents
(IK(Ca),
IA, and
IK1) in a rostral (A,
B) and a caudal (C, D)
hair cell. In the first protocol, hair cells were voltage-clamped to
potentials between 133 and +7 mV from a holding potential of 73 mV,
evoking both inward and outward voltage-dependent currents. This could
establish the presence of IK1 and a net
outward current (A, C). The net outward
current could be dissected by taking advantage of the different
inactivating properties of IA and
IK(Ca) in the second protocol
(B, D); hair cells were preconditioned
with 2 sec steps in membrane potential ranging from 133 to 33 mV
before a 400 msec depolarizing step to 0 mV (note that different
sampling rates were used during the conditioning and depolarizing
steps). Rostral and caudal hair cells are readily separated by the
presence of either IA
(B) or IK1
(C). An additional slowly activating inward
rectifying potassium current resembling Ih
(Holt and Eatock, 1995) was observed in a few rostral hair cells during
prolonged hyperpolarization (B).
A, B, Rostral hair cell, 12 pF; 2.4 M
usr; zero-current membrane potential of 61 mV; C,
D, caudal hair cell, 7 pF; 2.8 M usr; zero-current
membrane potential of 78 mV.
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Pharmacological dissection of the outward current
As stated above, the outward current could be separated into
inactivating and noninactivating components. From a holding potential of 63 mV, depolarization evoked predominantly the noninactivating outward current in both rostral and caudal hair cells. To minimize the
contribution of IA to the outward current,
experiments were typically performed from the most positive holding
potential just below the threshold of activation of the
voltage-dependent currents, which was determined empirically during the
experiment and varied between 65 and 55 mV among cells.
To confirm that the noninactivating outward current was a potassium
current, we examined the reversal potential of the noninactivating outward current tail current. Tail current reversal potentials were
measured in normal external solution composed of either 2 or 5 mM [K+]ext. Tail currents
were found to reverse polarity at 76.0 ± 6.5 mV
(n = 3) in 5 mM
[K+]ext (EK of
81 mV) or at 96.0 ± 1.6 mV (n = 3) in 2 mM [K+]ext
(EK of 104 mV), supporting the assumption that
the outward current is principally a potassium current.
To establish the pharmacological identity of the outward current, we
examined its sensitivity to several pharmacological agents or
experimental conditions known to affect potassium channels in other
hair cells. Figure 5 demonstrates those
results which most strongly suggest that, for both rostral and caudal
hair cells, the noninactivating outward current passes through the
large-conducting, BK-type, calcium-dependent potassium channel.
First, the outward current was routinely sensitive to low
concentrations of tetraethylammonium chloride (TEA) (Sigma). Within the
range of 60 to 0 mV, 10 mM TEA eliminated 100% of the
outward current in 18 of 21 hair cells (12 rostral, 9 caudal), with a
mean reduction of 99.3 ± 0.4% (mean ± SD), and typically
revealed a voltage-dependent inward current (ICa). TEA also eliminated the large
outward tail currents, leaving only the fast transient capacitive
currents combined with the fast ICa tail
currents. Two micromolar TEA was less effective, blocking 74.6 ± 7.5% (n = 21; 11 rostral, 10 caudal) of the outward current. In turtle BP hair cells, 2 mM TEA was found to
block >95% of the BK-type channels (Goodman and Art, 1996a), which
leads to the suggestion that IK(Ca) might not be
the only steady-state outward current present in AP hair cells. Whether
or not an additional outward current would be exposed by TEA depends on
the relative sizes and waveforms of the inward calcium current and the
remaining outward current. In Figure
6A, it can be seen in
both a rostral and caudal hair cell that, although 10 mM
TEA technically eliminated all outward current, an additional outward
component is present with the evoked inward current. Adding 1 mM 4-AP reliably eliminated this component, revealing a
waveform more typical of an uncontaminated inward calcium current.
Next, we examined the sensitivity of the outward current to the
scorpion toxin iberiotoxin (Ibtx) (Tocris, Ballwin, MO), which has been
reported to be a highly selective BK-type potassium channel blocker
(Galvez et al., 1990). In six rostral hair cells and seven caudal hair
cells, 20 nM Ibtx eliminated all outward currents between
60 and 0 mV (Fig. 5); an inward current was revealed over this same
range of membrane potentials in five of six rostral cells but in only
two of seven caudal cells (Fig. 6B), again suggesting
that an additional outward current is present in some hair cells. We
also tested the effects of another scorpion toxin, charybdotoxin (Chtx)
(Sigma) on the outward current. Chtx has also been identified as a
selective blocker of BK-type potassium channels, although it may not be
as selective as Ibtx (Kaczorowski et al., 1996). Chtx concentrations as
low as 10 nM were rapidly effective in completely removing
the noninactivating outward current, negative to 0 mV, in five of six
rostral hair cells and four of five caudal hair cells. Like Ibtx, Chtx
usually revealed an inward current in rostral hair cells (five of five) but not in caudal hair cells (two of four). In rostral hair cells, we
were able to establish that Chtx was selectively removing
IK(Ca) and not IA;
Chtx did not alter the amplitude, kinetics, or voltage-dependence of
IA.
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Figure 5.
The outward current is predominantly
IK(Ca) in rostral and caudal hair cells. The
sensitivity of the outward current to agents or conditions expected to
block the calcium-dependent potassium current
IK(Ca) is presented here. TEA (2 mM) and 20 nM Ibtx both block the outward
current and expose an inward current, Ica.
Zero-calcium in the external solution or the addition of 5 mM barium to the external bath caused the replacement of
the outward current with a large inward current. The zero-calcium
condition also resulted in a hyperpolarizing shift in the threshold of
the voltage-activated current. Barium is also observed to block the
inward current present in caudal hair cells below 80 mV. TEA: Rostral
cell, 16 pF; 2 M usr; 77 Hz; caudal cell, 7.8 pF; 1.5 M usr.
Ibtx: Rostral cell, 14 pF; 3 M usr; 90 Hz; caudal cell, 8 pF; 2.5 M usr. Zero-calcium: Rostral cell, 14 pF; 2.4 M usr; 133 Hz;
caudal cell, 7.8 pF; 2.6 M usr. Barium: rostral cell, 10.5 pF; 3.6 M usr; 225 Hz; caudal cell, 6.5 pF; 2.8 M usr. All rostral cells
shown here exhibited IA but not
IK1 and conversely so for caudal
cells.
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Figure 6.
Dissection of the outward current.
A, As shown in Figure 4, TEA could reliably replace the
outward current with an inward current, yet evidence remains for the
presence of an additional small outward current. Here, a rostral and a
caudal hair cell are shown being depolarized to 20 mV from a holding
potential of 60 mV (which should eliminate most
IA in rostral hair cells). In the presence
of 10 mM TEA, a slowly inactivating component
(IK) remains visible within the
inward current. This component is removed by the further addition of 1 mM 4-AP. Although the slowly inactivating component appears
pharmacologically similar to IA, it
is likely to be a separate voltage-dependent potassium current (rostral
cell, 12 pF; 1.8 M usr; caudal cell, 7 pF; 2.1 M usr).
B, In many other rostral hair cells, selective
elimination of IK(Ca) (shown here being
eliminated by 20 nM Ibtx) revealed only an
Ica. This rostral cell possessed a large
IA (data not shown), which does not
appear to contaminate the inward current being recorded
(Vstep of 20 mV;
Vh of 60 mV). While recording a caudal
hair cell under similar conditions, Ibtx eliminated the outward current
but did not expose an inward current, which suggests that some outward
current remains. C, Activation of the outward current
was determined by measuring the peak tail current amplitude at the end
of depolarizing voltage steps. Data expressed as percentages of the
maximum tail current amplitude. Curves are Boltzmann functions fit to
the data. For rostral hair cells, filled circles are the
mean data (n = 30) collected from tail currents
recorded in control solution from a holding potential of 60 mV (error
bars indicate SD, shown only where error exceeds size of
symbol). Open circles reflect
rostral hair cell tail current measurements obtained in the presence of
1 mM 4-AP (n = 12), which was used to
remove any IA or
IK contributing to the tail currents. The
results show that either IA and/or
IK are contributing to the net outward
current in control solutions in rostral hair cells. For caudal hair
cells, IK1 contaminated outward tail
currents in control solution, making it necessary to pharmacologically
separate IK(Ca) for analysis. Open
circles indicate mean measurements of the TEA-sensitive or
Ibtx-sensitive component of tail currents obtained for caudal hair
cells. Tail currents were collected in control solution and in the
presence of either TEA (n = 7) or Ibtx
(n = 5), and then tails recorded in the presence of
TEA-Ibtx were digitally subtracted from the control tails, which
removed the tail current contributions of both
IK and IK1 (which
is 4-AP-insensitive). Boltzmann constants: control
(filled circles),
V1/2(a) of 41.6; k = 7.3; rostral
(open circles), V1/2(a) of
50.4; k = 2.7; caudal (open squares),
V1/2(a) of 51.0; k = 2.7.
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The dependence of the noninactivating outward current on calcium was
tested several ways. It was found that reducing external calcium
concentrations from 4 to 0.1 mM eliminated >90% of the outward current (n = 6). If calcium were eliminated
from the external solution completely (zero-calcium added plus 2 mM EGTA), the outward current was replaced by a very large
inward current in 10 of 10 rostral hair cells and 10 of 10 caudal hair
cells (Fig. 5). This effect has been described previously (Art et al.,
1993) and is a result of the inward calcium current being transformed
in the absence of calcium into a nonselective inward current.
Furthermore, it was found that if calcium were replaced with equimolar
cadmium, the noninactivating outward current was completely eliminated in 10 of 11 rostral hair cells and seven of eight caudal hair cells.
The A-current evoked in rostral hair cells remained in the presence of
cadmium. Adding 500 µM cadmium to the normal external solution could also reliably eliminate 100% of the noninactivating inward and outward currents at holding potentials between 60 and 0 mV
in rostral hair cells (n = 6). In caudal hair cells, cadmium did not block the inward rectifier
IK1; thus, a small outward current was
recognizable between 30 mV and Ek in these cells. Finally, replacing external calcium with 5 mM barium
(Fig. 5) eliminated the outward current and produced a large inward current (~2.5 times the size of the control
ICa) in 12 of 12 rostral hair cells and
nine of nine caudal hair cells. Barium is known to pass more readily
than calcium ions through L-type calcium channels and, in doing so,
generates a large inward current. Barium did not block
IA in rostral hair cells but in caudal hair
cells barium blocked the inward rectifier
IK1.
The amplitude of IK(Ca) appeared to vary
substantially both between rostral and caudal hair cells and between
hair cells within each region. We observed a large difference in the
mean amplitude of IK(Ca) between rostral and
caudal hair cells; rostral cells possessed a mean
IK(Ca) at 20 mV of 2.73 ± 1.10 nA
(corrected for inward current amplitudes measured in TEA-4-AP),
whereas mean values for caudal hair cells were much lower at 0.82 ± 0.12 nA. The range of current amplitudes observed in rostromedial
hair cells (0.5-4.5 nA at 20 mV) fully encompassed the much lower range of outward current amplitudes in caudal hair cells (0.4-2.2 nA).
We will show below that the amplitude of the outward current was
closely coupled to resonant frequency in rostral and medial hair cells,
but we found no consistent relationship between outward current
amplitude and cell size in the caudal AP.
The activation range of the noninactivating outward current was
assessed for rostral and caudal hair cells by examining peak tail
current amplitudes. In Figure 6C, the filled
circles represent the mean (n = 30) tail
current amplitudes after depolarizing steps that were initiated from an
interpulse holding potential of 60 mV. Open circles
represent the same data collected for 12 rostral hair cells but in the
presence of 4-AP. Removal of the 4-AP-sensitive component of the tail
currents caused the Boltzmann curve to steepen. Tail currents measured
in the presence of 4-AP should specifically reflect the activation of
IK(Ca): for this set of tail currents, V1/2(a) of 50.4 ± 6.0 mV; the Boltzmann
constant k = 2.7 (and for the control set,
V1/2(a) of 41.6 ± 3.5 mV; k = 7.3),
which are similar to values reported for IK(Ca)
in other hair cells (Art and Fettiplace, 1987; Hudspeth and Lewis,
1988). The observed shift in the Boltzmann curve correlates with a 12%
decline in the mean Imax (comparing the 4-AP
mean to the control mean). The 4-AP-sensitive component of the tail
current may reflect the contaminating presence of
IA (because the holding potential used falls
within the region of overlapping activation and inactivation ranges of IA, although >95% of
IA should be inactive from a 60 mV holding potential), or it may reveal the presence of an additional
4-AP-sensitive potassium current. For caudal hair cells, tail currents
are contaminated by the activation of the inward rectifier
IK1 after repolarization of the membrane
potential. To eliminate this effect, we recorded outward current tail
currents in control solution and then in the presence of either TEA or
Ibtx. Tail currents recorded in the presence of TEA or Ibtx were then
digitally subtracted from the control tail currents, and thereby
revealed the TEA-sensitive (or Ibtx-sensitive) tail currents. We were
able to estimate tail current amplitudes using this method, and the
open squares in Figure 6D represent
the means of a total 12 caudal hair cells analyzed in this way. The
results show that the primary components of the outward current in both
rostral and caudal hair cells follow identical voltage-dependent
activation patterns.
For both rostral and caudal hair cells, the noninactivating outward
current appears to pass primarily through BK-type calcium-dependent potassium channels. It was generally true that an additional outward component was observed in both steady-state and tail current
measurements. Although some of this may by accredited to A-current
contamination, there are multiple lines of evidence to support the
presence of a second voltage-dependent potassium current in some cells,
active from normal hair cell resting potentials, which we will refer to
for now as IK. In particular, the
TEA-insensitive outward current observed in caudal cells not having an
appreciable A-current and the effect of 4-AP on tail currents, which
(because of the length of the depolarizing step) should not have
included significant A-current, imply the presence of an
IK in both rostral and caudal hair cells.
Furthermore, an IK was recently reported in frog
saccular hair cells (Armstrong and Roberts, 1998) and frog BP hair
cells (Smotherman and Narins, 1999), both of which share a very similar physiology with AP hair cells. We have not completed our investigation of this current, and we have not definitively separated this current from IA, but if IK
does exist in AP hair cells, it does not appear to be restricted to
either rostral or caudal hair cells.
Characterization of IA
In rostral hair cells, the inactivating component of the outward
current (Figs. 4B,
7A) was sensitive to low
concentrations of 4-AP. The prepulse protocol (Fig. 4, Protocol
2) was used to recruit and measure the amplitude and
voltage-dependence of the A-current before the addition of 1 mM 4-AP. The inactivating component of the outward current
was then routinely eliminated (95.8 ± 0.6%; n = 15) by the presence of 1 mM 4-AP added externally. From a holding potential of 120 mV, the combined addition of 10 mM TEA and 1 mM 4-AP was sufficient to
completely remove all outward currents, exposing the noninactivating
inward calcium current. The effect of 10 mM TEA on
IA was a reduction in the peak amplitude of the
inactivating component of 8.5 ± 6.5% (n = 10;
mean peak amplitude was 448 pA at 0 mV).
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Figure 7.
Isolation, activation, and inactivation of
IA. Although
IK(Ca) could be blocked by several methods,
external cadmium was exceptionally useful for exposing
IA because it eliminated both
IK(Ca) and ICa.
As described in Results, external cadmium also shifted the
activation and inactivation ranges of IA by
approximately +40 mV, thereby removing the need for hyperpolarizing
steps before depolarization and activation of
IA. In A, an example of
IA isolated by the application of cadmium is
presented. Here, IA is evoked by steps to
30 mV (4), 10 mV (3),
+10 mV (2), and +30 mV (1),
from a holding potential of 73 mV. B, Percent maximum
A-current amplitude at 0 mV was measured after a series of
hyperpolarizing prepulses (x-axis) for 2 sec.
Filled circles represent the average of 15 cells (errors
bars reflect SD) studied in control solution, and the data were fit
with the Boltzmann function shown (Eq. 1);
V1/2(in) of 81.2 mV; k = 7.2. Open circles represent the average of five cells studied
in the presence of Chtx. For a Boltzmann fit to this set of data,
V1/2(in) of 79.7 mV; k = 6.8, supporting the assumption that Chtx has no effect on the voltage dependence of IA.
Error bars and Boltzmann curve not shown for Chtx data.
C, Activation (filled circles) and
inactivation (open circles) curves for a cell studied in
the presence of Chtx, which completely blocked
IK(Ca). Inactivation was determined as
described in Figure 3. Activation was determined from peak tail
currents at 60 mV after brief depolarizing steps that ended at or
shortly after IA had reached its peak
outward current. Boltzmann curves fit to both activation and
inactivation data sets; for activation,
V1/2(a) of 45.6 mV; k = 7; and for
inactivation, V1/2(in) of 76.5 mV; k = 5.2. Rostral hair cell, 11 pF; 2.2 M usr.
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Some studies of the A-current were performed using CdCl2 as
a blocker of both the inward calcium current and the outward
calcium-dependent potassium current. Figure 7A presents an
example of IA recorded in 2 mM
cadmium. It was discovered that, although this was a useful method for
isolating IA, cadmium caused a large
(approximately +40 mV) shift in all of the voltage-dependent properties
of IA. Cadmium produced a reversible
depolarization of the V1/2(in) from 78.6 to
40.4 mV (Boltzmann fits to mean of five cells studied in both control
and 2 mM Cd2+), and caused a concomitant
shift of similar magnitude in the activation threshold for both the
steady-state currents and peak tail currents. Recordings in cadmium did
not appear to change the time course of activation, inactivation, or
deactivation of the A-current.
Figure 7B illustrates the mean inactivation curves for
IA recorded in control solution and in Chtx. The
V1/2(in) for control cells (mean data set,
n = 15) was 81.2 mV, which is very close to the
initial value reported by Hudspeth and Lewis (1988) in bullfrog
saccular hair cells. We saw no systematic shift in the inactivation
range of IA during the course of our
experiments. The mean threshold of activation for
IA as determined by eye from Boltzmann curves
fit to peak tail currents recorded in Chtx was 61.2 ± 3.5 mV
(n = 15); the mean V1/2(a) for
the same set was 44.7 ± 3.0 mV (n = 15). Figure
7C illustrates the activation and inactivation range for a
single cell studied in the presence of Chtx (1 µM). At
approximately the measured resting potential of rostral hair cells,
there is a narrow region of overlap between the activation and
inactivation range of IA, suggesting that
this current could contribute to both the resting potential and the receptor potential of the cell. The time courses of activation (Tpeak; time to peak) and inactivation
(inact) were voltage-sensitive, both becoming
faster with increasing depolarization. Tpeak
varied only slightly between cells, typically ranging between 18 and 23 msec at 20 mV (19.7 ± 4.5 ms; n = 28). At 20
mV, inact was observed to vary between 70 and 120 msec
among all cells studied, with a mean of 110.3 ± 27.2 msec
(n = 28).
An analysis of tail current amplitudes confirmed that
IA is a potassium current. Tail currents of
IA were analyzed in the presence of Chtx
(n = 2) or CdCl2 (n = 2).
In 5 mM external potassium, the mean tail current reversal
potential was 75.2 ± 2.3 mV (n = 4).
The calcium current
Most of the analysis of ICa kinetics and
voltage dependence presented here was performed with a 110 mM CsCl-based internal pipette solution (Table 1), which
always revealed a noninactivating inward current after depolarization
of the cell. For both rostral and caudal hair cells, the mean threshold
of activation (estimated by eye from I-V curves) was
55.8 ± 0.8 mV (n = 59); the mean V1/2(a) was 42.2 ± 2.8 mV (Boltzmann
fits to peak tail current measurements, n = 9), and the
peak inward current occurred at a potential of 23.0 ± 1.2 mV
(n = 59). The time course of activation was fit with a
single exponential curve over the first 5 msec of the record. The
activation time constant at 23 mV varied between 0.5 and 1.1 msec.
Deactivating tail currents were best fit with a double exponential
curve (Zidanic and Fuchs, 1995), with the fast component (accounting
for most of the inward tail current) having a time constant between 0.2 and 0.5 msec (approaching the limitations of the voltage clamp).
Measuring the inward current I-V curve in caudal hair cells
was complicated by the presence of the inward rectifying potassium
conductance IK1, which contributed a
sizable inward current at the standard holding potential (60 mV) when
the cell was infused with Cs+. For these cells,
external barium (replacing calcium) was used to establish that the
inward current in caudal hair cells exhibited a voltage dependence and
range of kinetic properties essentially identical to rostral hair cells
under otherwise similar recording conditions.
For both rostral (n = 18) and caudal (n = 15) hair cells, the amplitude of the inward current was sensitive to
external calcium concentrations (Fig.
8A). The inward current
was rapidly blocked by micromolar concentrations of cadmium (Fig.
8A). These channels passed barium ions more readily
than calcium ions (rostral, n = 12; caudal,
n = 9) (Fig. 8F); currents recorded
in 5 mM [Ba2+]ext were
approximately 2.5 times greater than those recorded in 4 mM
[Ca2+]ext. Calcium and barium currents
exhibited a steady decline in amplitude, typically decreasing by at
least 20% over the course of a 15 min experiment. As mentioned above,
a zero-calcium external solution resulted in a large nonselective
inward current appearing in place of the inward calcium current. The
threshold of activation for this current was ~20 mV negative to the
threshold of ICa, and the steady-state
current reversed polarity close to 0 mV. These features are consistent
with the effects of low calcium reported by Art et al. (1993) on L-type
calcium channels in turtle BP hair cells in which it was shown that, in
the absence of calcium, these calcium channels become nonselective ion
channels and their activation threshold is hyperpolarized. We applied
the calcium channel antagonist nifedipine (50 µM) while
recording the nonselective inward current in five rostral hair cells
and observed a mean blockage of 65 ± 20% of the peak inward
current. This partial blockage is consistent with the effects of
nifedipine on calcium currents in other hair cells (Fuchs et al.,
1990). These observations support the assumption that this is the
L-type calcium current commonly reported in other auditory hair cells
(Art and Fettiplace, 1987; Hudspeth and Lewis, 1988; Zidanic and Fuchs,
1995). To further test this assumption, we applied the L-type calcium
channel agonist Bay K 8644 (Sigma) to both rostral and caudal hair
cells (n = 3 and 2, respectively). In every case, Bay K
8644 (0.1 µM) caused a pronounced increase in the
steady-state amplitude of the inward current (135 ± 22%;
n = 5). As can be seen in Figure 8C, the
corresponding peak tail current amplitude is enlarged and relaxation of
the tail current is prolonged, which is consistent with the reported action of Bay K 8644 on L-type calcium channels (Fuchs et al., 1990).
Furthermore, it can be observed that Bay K 8644 changes the shape of
the calcium I-V curve (Fig. 8, compare B,
D), producing a much narrower voltage activation range and
hyperpolarizing the potential at which the peak inward current occurs
by almost 25 mV. We conclude that, for rostral and caudal hair cells,
the inward current activated by depolarization is predominantly an
L-type calcium current. We did not test for the presence of other
calcium channels, such as the N-type channels reported in frog saccular hair cells (Su et al., 1995). Furthermore, it should be noted that,
because of their smaller size and greater sensitivity to the
detrimental effects of patch-clamp recording, the pharmacology of
caudal hair cell calcium currents is less certain than that of rostral
hair cells. Yet, the essentially identical kinetics, voltage
dependence, and ionic selectivities of the calcium currents in rostral
and caudal hair cells support the conclusions that these currents are
at least very similar for all AP hair cells.
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Figure 8.
Ionic sensitivity and pharmacology of
the inward calcium current. A, The inward current
revealed by a CsCl internal pipette solution could be modulated by
changes in [Ca+2]ext. The control
solution used in most experiments contained 4 mM calcium.
Increasing the external concentration to 20 mM caused an
approximately fourfold increase in the size of the inward current. In
contrast, the addition of 50 µM cadmium to the control
solution caused a rapid but reversible block of the inward current.
B, I-V curve derived for the same cell
in 4 and 20 mM
[Ca+2]ext, from a holding
potential of 73 mV. Oscillatory medial hair cell, 11 pF; 9 M; 80%
src. C, Bay K 8644 (5 µM), an
L-type calcium channel agonist, caused an increase in the steady-state
and peak tail-current amplitudes, which is attributed to an extension
in the single channel mean open time. The tail current becomes
substantially slower with Bay K 8644. This drug not only increases the
steady-state amplitude over the entire voltage range
(D) but produces a much steeper activation slope
(the peak inward current is shifted from 23 to 48 mV) and appears
to hyperpolarize the threshold of activation. Medial hair cell, 10 pF;
16 M; 80% series resistance compensation (src). E,
The mean steady-state I-V curves for rostral
(n = 10) and caudal (n = 10) hair cells recorded in zero external calcium. Error bars are shown
where they exceed the size of the symbols. The
steady-state reversal potentials were 8 (rostral) and 19 mV
(caudal). F, The mean steady-state I-V
curves for rostral (n = 12) and caudal
(n = 9) hair cells recorded in 5 mM
external barium. The steady-state reversal potentials were +36
(rostral) and +25 mV (caudal).
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Although the calcium current could be exposed either through the
infusion of cesium or the external application of 10 mM TEA together with 1 mM 4-AP, it was found that there was a
substantial difference in the amplitude of the mean inward current
generated by the two methods. For example, for rostral hair cells, the
mean inward current recorded at 20 mV using an internal cesium
pipette solution was 328 ± 76 pA (mean ± SD;
n = 21), with a mean current density of 25 ± 6 pA/pF, whereas using TEA-4-AP with a standard internal solution
produced mean inward currents of 430 ± 85 pA (n = 14), with a mean current density of 33 ± 6 pA/pF. There appeared to be a difference in the means and overall range of inward currents observed between rostral and caudal hair cells. Using TEA-4-AP to
expose the inward current, the mean peak inward current for caudal hair
cells was 120 ± 34 pA (n = 9) and ranged from 60 to 180 pA. For rostral cells, the mean peak inward current was 430 ± 85 pA (n = 14) and ranged from 160 to 634 pA.
Calcium currents measured in cesium-loaded hair cells also follow this
trend; for rostral hair cells, the mean peak inward current was
350 ± 103 pA (n = 12), whereas for caudal hair
cells, it was 110 ± 43 pA (n = 9).
IBa in caudal hair cells was 314 ± 150 pA
(n = 9; ranging from 134 to 600 pA), which was
approximately half the mean value found in rostral hair cells, 625 ± 340 pA (n = 9; ranging from 400 pA to 1.1 nA). It
remains possible that sampling error contributed to these differences
and that subregional variations in current amplitudes are the source of
the observed amplitude variations. As we will show below, current
amplitudes were seen to vary with position (and frequency) within the
rostromedial region of the AP, but we have not yet uncovered evidence
of position-related gradients in the amplitudes of
ICa and IK(Ca) in caudal
hair cells.
Calcium current reversal potentials (Fig. 8), were consistently
negative to the presumptive ECa (>+80 mV) but
were similar (±10 mV and spanning the same range) for both Cs-exposed
and TEA-4-AP-exposed inward currents. These reversal potentials,
typically 25-30 mV in 4 mM
[Ca2+]ext, are comparable with
those reported in chick hair cells (Fuchs et al., 1990) and frog
saccular hair cells (Hudspeth and Lewis, 1988) under similar recording
conditions. Barium currents, recorded in 5 mM
[Ba2+]ext plus 1 mM 4-AP,
exhibited reversal potentials in the range of 20-40 mV, but the mean
reversal potential for caudal cells was 11 mV more negative than the
mean for rostral hair cells (Fig. 8F). This pattern
was also observed regarding the reversal potential of the nonselective
inward current recorded in zero-calcium; the mean reversal potential
for caudal hair cells was 11 mV more negative than the mean for rostral
hair cells. Although we have not yet accounted for this experimental
difference, one likely explanation might be the contaminating presence
of an additional IK, which, although
small, may contribute to a larger percentage of the total membrane
conductance in caudal hair cells than in rostral hair cells.
The inward rectifying K+ current
As described above, hair cells isolated from the caudal half of
the AP are likely to possess the inward rectifying potassium current
IK1 (Figs. 4C,
9). This current begins to activate below a threshold of approximately 30 mV (derived from fit by eye to steady-state I-V curves recorded in external cadmium;
n = 7), and the steady-state current has a reversal
potential near EK (Fig. 9B). The
speed of activation increases with increasing hyperpolarization, and
the current exhibits mild inactivation with hyperpolarizations to
potentials negative to 120 mV. The time course of activation could be
fit by a single exponential curve. The mean activation time constant at
100 mV was 5.6 ± 1.1 msec (n = 28; recorded in
5 mM [K+]ext).
After return to a holding potential of 50 mV (in 4 mM CdCl2), tail currents were best fit with a double
exponential, with fast (0.38 ± 0.10 msec; n = 7)
and slow (2.45 ± 0.35 msec) time constants. The reversal
potential of the steady-state current closely followed
EK; in 2 mM
[K+]ext, the steady-state
current was zero at a mean potential of 97.3 ± 4.2 mV
(n = 8), whereas in 5 mM
[K+]ext, the steady-state
current was zero at a mean potential of 76.5 ± 2.5 mV
(n = 28). IK1 was insensitive to
external cadmium but was highly sensitive to barium (in control
solution the mean IK1 for caudal cells at 120
mV was 1.3 ± 0.3 nA; n = 22) (Fig. 8F). Barium (500 µM) applied
externally resulted in a rapid and complete block of
IK1. Based on its kinetics, its voltage
dependence, and its sensitivity to barium but not cadmium, we conclude
that this current is the same IK1 described in
hair cells of the leopard frog sacculus (Holt and Eatock, 1995) and at
least very similar to the IIR of turtle BP hair
cells (Goodman and Art, 1996b).
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Figure 9.
The inward-rectifying potassium current
IK1. A, In caudal hair cells
and some medial hair cells, hyperpolarization of the membrane potential
evoked a large and fast-activating current, which reversed polarity
around EK. With greater levels of
hyperpolarization, this current exhibited some inactivation, as seen
here in the step to 135 mV. B, This steady-state
I-V curve for IK1 was
determined for a caudal cell bathed in 2 mM
CdCl2, which would reliably remove all other ionic
currents active within this voltage range. The current first appears as
an outward (positive) current at potentials negative to 30 mV and
becomes inward (negative) below 80 mV.
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The distribution of IA and IK1
implies overlapping populations
The distributions of IK1 and
IA in AP hair cells presented in Table 2 make it
clear that these two populations are not entirely separate but rather
appear to comingle within the medial portion of the AP. Figure
10 shows the percent distribution of
IA and IK1 as a function
of cell size in the AP (included as a measure of the likelihood of a
hair cell of a particular size possessing either of these currents but
distinct from variations in current amplitude). Two separate but
overlapping populations emerge from this figure. Surprisingly, each
current appears to be present along the full length of the epithelium.
Experimental records support the observation that the largest cells
possessing IK1 (14 and 15 pF) were indeed very
tall hair cells (>60 µm) and not simply bloated short hair cells,
making it very unlikely that these cells came from anywhere but the
rostral patch of the AP.
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Figure 10.
Distribution of IA and
IK1. Using the protocols described in Figure
4, the presence or absence of IA and
IK1 (independent of amplitude) was
determined for cells of different sizes in the AP. An ionic current was
considered present if its amplitude could be separated from the passive
leak current and if it exhibited both time- and voltage-dependent
kinetics matching those described for either
IA or IK1. Bars
represent the percentage of cells at each size found to possess a
current, independent of amplitude. Some cells possess both currents.
For IA, n = 124;
mean number of cells per bin, nine cells (minimum of 6); for
IK1, n = 168;
mean number of cells per bin, 13 cells (minimum of 8). The sample size
(n) is included above each bar.
Cell size, given as the total membrane area calculated from whole-cell
capacitance measurements, is directly proportional to cell height and
has been shown to correspond to rostrocaudal position in the AP (see
Results).
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Factors influencing resonant frequency
Resonant frequency varies with cell morphology
To establish that hair cells are arranged tonotopically, we first
examined the relationship between cell morphology and whole-cell capacitance. Simmons et al. (1994) demonstrated that AP hair cell bodies become progressively shorter moving rostrocaudally, declining from >60 µm in rostral hair cells to <20 µm caudally, with most of the change occurring within the rostral half of the AP. Notably, they identified the hair cells at the center of the rostral region (and
not the rostral-most hair cells) as the tallest. We also found the
tallest hair cells were those isolated from the center of the rostral
patch, and these had whole-cell capacitances as large as 20 pF.
Consistent with a relationship between cell size and frequency, Lewis
et al. (1982b) showed, while reporting an overall rostrocaudal
frequency gradient in the primary fibers innervating the AP, that the
lowest frequencies were found not at the rostral-most position but more
toward the center of the rostral patch. Together, these reports and our
observations suggest that frequency is not mapped strictly
rostrocaudally in the AP but instead may follow a more "radial"
organization, with the lowest frequencies originating near the center
of the rostral patch. The majority of rostral hair cells exhibited
whole-cell capacitances that ranged from 12 to 18 pF. Medial hair cells
had values between 8 and 12 pF, and caudal hair cells ranged from 4 to
8 pF. Using measurements taken from photographs of previously recorded
hair cells, we plotted cell body length versus the measured whole-cell
capacitance (Fig.
11A). Changes in cell
body length are accurately reflected by cell capacitance measurements.
The dashed line indicates the predicted cell capacitance for
hair cell bodies of different lengths (not including estimates of
bundle capacitance), using the mean cell body cross-sectional width of 14 µm and assuming a standard value of membrane capacitance of 1 µF/cm2. In Figure 11A, most of
the data points are slightly below the predicted line, which is most
likely the result of the compensation circuitry of the amplifier
consistently underestimating the true whole-cell capacitance
(whole-cell capacitance and series resistance compensations were never
100%). In addition to cell length differences, there were also
noticeable variations in cell shape, which would affect the accuracy of
the predicted length-capacitance relationship; some rostral hair cells
were gourd-shaped, whereas others resembled simple cylinders. Shape
variations disappeared as cells became shorter. In other preparations
(most notably the turtle), a large gradient in stereovillar bundle
length precludes the use of whole-cell capacitances as an indicator of
cell size. In the AP, however, the only reported gradient in bundle
height is mediolateral and not rostrocaudal (Lewis and Li, 1975; Lewis,
1977); thus, a rostrocaudal gradient in cell capacitance is expected to
occur independent of hair bundle architecture. We did not estimate the
contributions of the hair bundle to the overall membrane capacitance,
although we believe that this is one important source of the observed
variations in cell capacitance within each region of the AP. We
conclude that the cell capacitance measurements obtained at the
beginning of each experiment are useful and reasonably accurate
indicators of cell body length and therefore of the original position
of the hair cells along the rostrocaudal axis of the AP.
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Figure 11.
Resonant frequency changes with cell size.
A, Whole-cell capacitance measurements are linearly
related to hair cell body length. The dashed line is the
predicted relationship between cell length and membrane capacitance,
based on the mean AP hair cell cross-sectional diameter (14 µm) and
the standard for membrane capacitance 1 µF/cm2.
B, Whole-cell capacitance (in picofaradays) is plotted
against the resonant frequency of each hair cell at a common
steady-state membrane potential of approximately 53 mV for cells
isolated with papain (filled circles). For most
cells, this represented the highest Qe
oscillations, which were generated with small depolarizing current
pulses. In some cases, the value included was taken from offset
oscillations depending on the zero-current resting potential of the
cell. Data were best fit using a least-squares algorithm by an equation
of the form y = 7700 · x 1.6; r2 = 0.74; n = 53. Open circles represent
data collected from hair cells isolated without using papain as a
dissociative agent.
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Figure 11B shows the variation in resonant
frequencies with cell size in the rostral half of the AP. Despite the
scatter in the data, this plot supports the conclusion that cell size
decreases as resonant frequency increases and, therefore, that resonant frequency covaries with rostrocaudal position in the AP. This evidence
of a tonotopic distribution of resonant frequencies underlies the
assumption that electrical tuning contributes to frequency selectivity
in the AP.
Offset resonant frequencies recorded at a common membrane potential of
approximately 53 mV varied from 50 Hz in the largest hair cells to
375 Hz in medial hair cells. Onset resonant frequencies were recorded
to 475 Hz. This range and distribution of resonant frequencies closely
overlies the range of resonant frequencies recorded intracellularly in
the AP by Pitchford and Ashmore (1987) and the known tonotopic map of
the rostromedial AP (Lewis et al., 1982b). Afferent fibers innervating
the medial AP have best frequencies at least as high as 600 Hz,
approximately twice the highest offset resonant frequencies observed in
isolated hair cells. A similar inconsistency exists in the turtle
cochlea in which the upper limit of resonant frequencies recorded from
hair cells in situ is approximately double that reported for
isolated hair cells (Art and Fettiplace, 1987). The explanation
suggested by Goodman and Art (1996a) for the turtle BP is that a
reduction in resonant frequencies of isolated hair cells occurs as a
result of a consistent reduction in the total number of ion channels
during the dissociation procedure (Goodman and Art, 1996a). Armstrong
and Roberts (1998) demonstrated that the use of papain as a
dissociative agent appeared to increase the range of resonant
frequencies observed in frog saccular hair cells. Given the potential
pitfalls of enzymatic digestion, we isolated hair cells without the use
of papain and investigated their electrical properties. In Figure
11B, the open circles represent
"papain-free" hair cells. The relationship between cell capacitance
and resonant frequency in the AP appeared to be unaltered by the use of papain.
Outward current kinetics mediate resonant frequency
In oscillatory AP hair cells, the majority of the outward
current evoked from the cell resting potential is the calcium-dependent potassium current IK(Ca). In the turtle basilar
papilla, resonant frequency is strongly correlated with the kinetics of
IK(Ca) (Art and Fettiplace, 1987). Similarly, we
found that the kinetics of the outward current changed systematically
with resonant frequency in oscillatory AP hair cells. Figure
12A provides three
examples of offset resonant frequency of a hair cell (at 53 mV;
current clamp) and its corresponding outward tail current (at 53 mV; voltage clamp). Hair cells with lower resonant frequencies had slower
net outward tail currents, reflecting longer channel mean open times.
The outward tail current was predominantly
IK(Ca), but as stated earlier, on
average, 12% of the peak tail current in rostral hair cells could be
eliminated by the addition of 4-AP, which was most likely caused by
IA contamination. Unlike Goodman and Art
(1996a), however, no frequency-related variations in the relative
contribution IK(Ca) to the outward tail current
were observed. Figure 12B plots resonant frequency
() of a hair cell measured at a common steady-state membrane
potential of 53 mV versus the outward tail current time constant
(d) at a holding potential of 60 mV after a 200 msec step to 20 mV. In both the turtle basilar papilla (Crawford and
Fettiplace, 1981; Art and Fettiplace, 1987) and the frog amphibian
papilla, the relationship between and d appears well
described by the function
|
(2)
|
which may be expressed as
|
(3)
|
where k is an empirically determined coefficient (Art
et al., 1986; Art and Fettiplace, 1987; Wu et al., 1995) representing the relationship between conductance and membrane potential and also
includes membrane capacitance. The data presented in Figure 12B were well fit by Equation 3. The relationship
becomes linear when resonant frequency is plotted against the inverse
square root of the time constant (Fig. 12C). Also included
in Figure 12C is the relationship between and
d in turtle BP hair cells, taken from Art and Fettiplace
(1987). The mild difference between the turtle and frog lines can
probably be accounted for by (1) differences in the external calcium
and potassium concentrations used, and (2) the holding potential at
which d was measured; as mentioned above, we studied
d at 60 mV and plotted it against at 53 mV,
whereas the data from the turtle were collected at the resting
potential of each cell and therefore appear to have been typically in
the range of 40 to 50 mV (Art and Fettiplace, 1987, their Figs. 1, 3-5). Because d becomes smaller with
hyperpolarization (Art and Fettiplace, 1987), our tail current time
constants are expected to be slightly less than those reported for the
turtle over the same range of resonant frequencies and would therefore also have larger d1/2 values.
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Figure 12.
Resonant frequency changes with the
time constant of the outward current. A, Offset resonant
frequencies (recorded in current-clamp mode) oscillating at a common
membrane potential of approximately 53 mV are presented on the
left, along with their corresponding net outward tail
currents on the right (recorded in voltage-clamp mode)
at 53 mV after a 200 msec voltage step to 30 mV. Holding potentials
in current-clamp mode were maintained at approximately 53 mV with
small standing currents. Offset oscillations followed a 100 pA current
pulse. The time constant of outward current relaxation
(d) is progressively faster in higher frequency
hair cells. Top, 100 Hz; Qe
of 6.5; d of 6.0 msec; middle, 170 Hz;
Qe of 2.5; d of 3.1 msec;
bottom, 240 Hz; Qe of 3.6;
d of 1.8 msec. B, Resonant frequency
(o) at 53 mV is plotted against
d at 60 mV after a 200 msec step to 30 mV. The data
were well fit by an equation of the form d of
k/o2;
(k = 24133.6; r2 = 0.88;
n = 33). C, Here, the inverse square
root of the time constant is plotted against resonant frequency. Data
fit with a straight line (y = 0.004x + 0.24; r2 = 0.82). Open circles represent data collected from hair
cells isolated without using papain as a dissociative agent. The
dashed line represents the relationship reported for the
turtle BP, taken from Art and Fettiplace, 1987. The turtle line is
lower primarily because those tail currents were measured at potentials
10 to 20 mV positive to the 60 mV used in our study of the AP.
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Again for the purposes of comparison, we examined the relationship
between and d in papain-free hair cells, represented by the open circles in Figure 12C. Although
the relationship between the outward current kinetics and resonant
frequency appears virtually identical in the papain-free hair cells,
some observations are worth mentioning. Our highest resonant
frequencies and smallest d values were recorded from
papain-dissociated hair cells, and the lowest resonant frequencies were
recorded from papain-free hair cells, supporting the possibility that
papain may be shifting the range of frequencies observed upward.
However, the net effect on the overall range of frequencies and time
constants observed is, if anything, small.
ICa and IK(Ca) amplitudes vary
with resonant frequency
The electrical resonances in AP hair cells are driven by the
interactions of the L-type, voltage-dependent calcium current ICa and the calcium-dependent potassium current
IK(Ca). After first recording the net outward
current amplitude (in voltage-clamp mode) and the electrical resonance
properties of the cell (in current-clamp mode), we used two separate
methods to isolate and measure the amplitude of
ICa. In our first attempt to investigate the
relationship between IK(Ca) and
ICa amplitudes and resonant frequency, we
applied a combination of TEA and 4-AP to block
IA and IK(Ca). Art and
Fettiplace (1987) reported that this method produced unreliable
measures of calcium current amplitude in turtle hair cells, and indeed,
we found conflicting results using this combination of potassium
channel blockers. There was a strong correlation between outward
current amplitude and resonant frequency (r2 = 0.82; n = 14), but in the presence of TEA-4-AP, inward
current amplitudes were not observed to change significantly with
resonant frequency. To address the possibility that the TEA-4-AP
cocktail was interfering with calcium current measures, we instead
measured barium current amplitudes passing through the presumed
voltage-dependent calcium channel
(gCa). Barium has been shown to
pass through hair cell calcium channels more readily than calcium but
does not activate IK(Ca) (Art and Fettiplace,
1987; Fuchs et al., 1990; Zidanic and Fuchs, 1995). During initial
experiments, it became clear that external barium alone was
insufficient to guarantee the elimination of all outward currents; in
low-frequency cells in particular, there was consistent evidence of
either IA or IK
contamination. Therefore, barium currents were recorded in 5 mM [Ba2+]ext to which 10 mM 4-AP was added. This combination appeared to provide the
most reliable measure of inward current amplitudes; we found no
evidence of inactivating outward currents under these conditions.
Under these conditions, peak IBa was observed to
increase smoothly with resonant frequency in oscillatory hair cells
(Fig. 13A) over the recorded
frequency range of ~300 Hz. Furthermore, plotting this distribution
alongside the data presented for hair cells of the turtle basilar
papilla (Fig. 13A, turtle data taken from Art and
Fettiplace, 1987) demonstrates a close similarity between the two
preparations in this regard.
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Figure 13.
Current amplitudes increase with resonant
frequency. A, Peak inward current was determined for
oscillatory hair cells in an external medium containing 5 mM [Ba+2]ext and 10 mM 4-AP, after first recording their resonant frequency at
53 mV in control solution (standard internal pipette solution). Peak
barium currents occurred at approximately a mean potential of 23 ± 3 mV and were observed to be larger in hair cells exhibiting higher
resonant frequencies. The data are plotted on a semi-log scale along
with data for isolated hair cells of the turtle basilar papilla
(open diamonds) taken from Art and Fettiplace (1987) to
emphasize the similarities in IBa amplitude
distribution exhibited by these two preparations. Both lines drawn by
eye; solid line, frog AP; dotted line,
turtle BP. B, Outward conductance amplitude, corrected
for inward current amplitude, was calculated and plotted against the
measured resonant frequency at 53 mV. For the cells included from
A, inward barium currents (20 mV) were divided by 2.5 (see Results) to provide an estimate of the calcium current amplitude,
and this value was subtracted from the measured steady-state outward
current at 20 mV to produce an estimate of the total outward current
amplitude (filled circles). In cells in which the
calcium current was isolated using a TEA-4-AP cocktail (open
circles), the calcium current amplitude at 20 mV was
subtracted directly from the outward current amplitude at 20 mV. Both
sets of data produce a similar description of the rise in
gK(Ca) amplitude with resonant frequency.
Conductances rather than currents are presented here to correct for
differences in external K+ concentrations; barium
current experiments were performed in 5 mM
[K+]ext, and the TEA-4-AP
experiments were performed in 2 mM
[K+]ext. Curve fit by eye.
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As mentioned above, outward current amplitude was also observed to
increase with resonant frequency in oscillatory hair cells. Figure
13B plots gK(Ca) amplitude versus
resonant frequency. The conductances included in Figure 13B
were believed to be almost exclusively gK(Ca)
based on the observation that the values included here were
noninactivating steady-state currents and were mostly blocked by TEA,
Ibtx, or barium; however, it cannot be ruled out that some
voltage-dependent K+ conductances contributed to
these values.
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DISCUSSION |
Two hair cell subtypes in the amphibian papilla
There are two experimentally distinctive populations of hair cells
in the amphibian papilla: an oscillatory-type electrically tuned hair
cell that dominates the low- to mid-frequency auditory range of the AP,
and a nonoscillatory cell type that dominates the mid- to
high-frequency region of the AP. Those hair cells exhibiting prominent
electrical resonances were more likely to possess
IA than IK1,
whereas the opposite was true for those hair cells, predominantly the
caudal ones, not exhibiting high Qe resonances.
A similarly bisected epithelial pattern in hair cell biophysics was
also reported for the goldfish sacculus (Sugihara and Furukawa, 1989)
and chick BP (Fuchs and Evans, 1990), but the quality and frequency
range of the resonances observed in those two preparations fall far
short of the resonances described in turtle BP hair cells. The frog AP
is unique then, in that it displays an epithelial organization similar
to both fish and birds, and yet also exhibits most of the
specializations of the electrically tuned turtle hair cells. With
mounting evidence that the AP also possesses mechanical tuning
mechanisms (Lewis et al., 1982a; Hillery and Narins, 1984; Stiebler and
Narins, 1990), the AP becomes an ideal organ for investigating the
evolution of electrical and mechanical tuning in terrestrial vertebrates.
Our observation that rostral, but not caudal, hair cells are
electrically tuned is consistent with neurophysiological data. Electrical tuning in the rostral half of the AP is supported by the
highly temperature-sensitive tuning properties of the afferent fibers
innervating the rostral but not caudal region (CFs exhibiting Q10(temp) values at approximately 2.0) (Stiebler
and Narins, 1990; Van Dijk et al., 1990), which implies that the
temperature-sensitive ion channels are contributing substantially to
frequency selectivity (Smolders and Klinke, 1984; Schermuly and Klinke,
1985; Wu et al., 1995). Other differences in the response properties of
rostral and caudal AP afferent fibers are consistent with two separate auditory processing mechanisms. Ronken (1991) demonstrated that AP
afferent fibers could be divided into two populations based on
W10 dB (tuning curve bandwidth 10 dB above
threshold at CF) and Q10 dB (the frequency at
the lowest threshold divided by W10 dB). Fibers with CFs below ~600 Hz have higher Q10
dB values and narrower W10 dB values.
Low-frequency fibers have been reported to have lower thresholds and
higher spontaneous rates (Frishkopf and Goldstein, 1963; Narins, 1987),
and only low-frequency fibers are suppressible, exhibiting both one-
and two-tone suppression (Feng et al., 1975; Christensen-Dalsgaard and
Jorgensen, 1996).
Several lines of anatomical evidence support the recognition of two
hair cell subtypes in the frog AP. Simmons et al. (1994) described two
morphologically distinct groups of rostral and caudal hair cells in the
leopard frog AP. Wever (1973) also divided the AP into anterior
(rostral) and posterior (caudal) regions based on cell and cell nuclei
morphology. Innervation patterns in the AP also suggest two
populations. Thicker afferent fibers innervate the rostral AP, and
these terminate on more hair cells than caudal afferents (Lewis et al.,
1982b; Simmons et al., 1992). Efferent fibers only extend to the
rostral region of the AP (Simmons et al., 1992).
Our failure to record electrical resonances in caudal AP hair cells may
be the result of a preferential sensitivity to the enzymatic
dissociation procedure. Hair cell electrical properties are almost
surely modified by the dissociation procedure (Art and Fettiplace,
1987; Goodman and Art, 1996a; Armstrong and Roberts, 1998).
Additionally, patch-clamp recording conditions become compromised as
the size of the target cell is reduced, making it more likely that
caudal cells will exhibit degraded electrical properties. Furthermore,
the considerable resting conductance
(IK1) in caudal hair cells has the
potential to diminish Qe and could in effect "squelch" electrical resonances in isolated caudal hair cells. Thus, the absence of electrical resonances cannot by itself eliminate the potential for caudal hair cells to contribute to acoustic tuning.
The role of IA in auditory hair cells
In the AP, IA is generally restricted to
oscillatory hair cells in the low-frequency range of the AP. The
IA described here appears identical to the
IA described in chick BP hair cells (Murrow, 1994), and its distribution is qualitatively similar as well. In both
the frog AP (Simmons et al., 1992) and the chick BP (Murrow and Fuchs,
1990; Murrow, 1994), IA is present in
oscillatory-type hair cells that receive efferent innervation. Efferent
innervation has been shown to hyperpolarize frog saccular hair cells by
as much as 20 mV (Ashmore and Russell, 1982). Just such a
hyperpolarization would release nearly 100% of
IA from its steady-state inactivation, and after
succeeding depolarizations, the addition of the A-current would
increase the outward current, diminishing the amplitude of the receptor
potential. Similar to what was proposed for the chick (Murrow and
Fuchs, 1990), we suspect IA may act in
conjunction with efferent inhibition to either suppress or modulate the
time course of the receptor potential in hair cells of the rostral AP.
Contributions of IK1 to the
receptor potential
Two potential contributions of the inward rectifier to the
electrical response properties of hair cells have been described recently. First, the presence of IK1 tends to
hyperpolarize hair cell resting potentials in vitro (Holt
and Eatock, 1995); it is unknown whether or not this effect occurs
in vivo. Second, the relaxation of
IK1 at the initiation of the voltage response
might provide positive feedback during the depolarizing phase of the voltage response. In the turtle, this was shown to enhance the quality
of electrical tuning at the lowest frequencies (Goodman and Art,
1996b), but in that preparation, IK1 disappears
well below the frequency range at which it is found in the AP. In the frog AP, we find that IK1 is generally
restricted to highest frequency range of the epithelium. The
distribution of IK1 in the AP is more
reminiscent of the chick BP than the turtle BP. In both the frog AP and
chick BP (Murrow, 1994), IK1 is found in
nonoscillatory hair cells and is generally restricted to those cells
that do not possess IA. The kinetics of
IK1 (typical deactivation time constants exceed
2.0 msec at 55 mV) are slow enough to make it potentially detrimental
to electrical tuning at frequencies above 500 Hz. Additionally, the
presence of IK1 in the frog sacculus (Hudspeth
and Lewis, 1988; Holt and Eatock, 1995), caudal AP, and BP (Smotherman
and Narins, 1999) make it difficult to envision a consistent electrical
tuning function for IK1 in situ. The
role of IK1 in frog auditory hair cells remains
an enigma.
Resonant frequency is determined by multiple factors
The range of resonant frequencies observed in the frog AP can be
accounted for by three factors: whole-cell capacitance, current amplitudes, and outward current kinetics. Reducing the total area of
basolateral membrane while increasing conductance amplitudes (and
conductance density) produces lower input impedances and smaller
membrane time constants, which speeds up the rate at which the membrane
potential can fluctuate. The inward calcium current is increased, and
the outward calcium-dependent potassium current becomes larger and
faster to produce higher resonant frequencies. The result of these
combined cellular changes is an array of hair cells that exhibit
resonant frequencies ranging from 50 to ~400 Hz. For these hair
cells, the predominant outward current flows through the BK-type
calcium-dependent potassium channel, and a thorough investigation in
the turtle cochlea has provided us with considerable insight into the
molecular mechanisms by which the kinetics of this channel are
manipulated (Art et al., 1995; Wu et al., 1995). It appears that the
role of this ion channel is similar for the frog and turtle in
mediating electrical tuning, and the pattern of expression of this ion
channel plays a large part in establishing the map of resonant
frequencies within each sensory epithelium. The range of resonant
frequencies observed appears similar for the frog and turtle, yet the
acoustic frequency range of the AP is twice that of the turtle BP.
Analytical modeling based on the properties of ion channels in turtle
hair cells (Wu et al., 1995) has been used to demonstrate that it would
be possible to extend electrical tuning up to several kilohertz, but
the necessary physiological parameters have not yet been uncovered in
any preparation.
Effect of papain on hair cell electrical properties
Armstrong and Roberts (1998) reported that papain digestion caused
an alteration of the electrical resonance properties in frog saccular
hair cells. The alterations were the result of papain (1) selectively
removing a voltage-dependent potassium current (IK) in some cells, and (2) speeding up
the kinetic properties of IK(Ca). In our
investigations, we did not observe a significant IK in AP hair cells dissociated without papain,
and we have recently characterized an IK in
papain-dissociated hair cells of the frog BP (Smotherman and Narins,
1999). We did not test the effects of papain on
IK(Ca) kinetics directly. Papain is not expected to change the relationship between the outward tail-current
time-constant and resonant frequency, but it might cause a shift in the
range of resonant frequencies observed. Our results could not rule out the suggestion that papain might speed up
IK(Ca), but we saw little change in the
overall range of resonant frequencies, and the correlation between cell
size and resonant frequency was papain-insensitive.
| |
FOOTNOTES |
Received Jan. 29, 1999; revised March 31, 1999; accepted April 19, 1999.
This work was supported by National Institute on Deafness and Other
Communications Disorders Grant DC00222 to P.M.N. and a Hyde Fellowship
from the University of California, Los Angeles Department of
Physiological Science to M.S.S. We thank Drs. D. D. Simmons, C. Bertolotto, A. P. Purgue, W. M. Yamada III, E. R. Lewis,
and F. Bezanilla for their technical assistance and many helpful
discussions. All experiments comply with the National Institutes of
Health Principles of Animal Care Publication 86-23 and
all current United States laws.
Correspondence should be addressed to Peter M. Narins, Department of
Physiological Science, University of California, Los Angeles, 405 Hilgard Avenue, Los Angeles, CA 90095-1527.
| |
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