From the Sections of Otolaryngology and Neurobiology, Yale University School of Medicine, New Haven, Connecticut 06510
The effects of turgor pressure-induced membrane tension on junctional coupling of Hensen cell isolates from the inner ear were evaluated by input capacitance or transjunctional conductance measurement techniques. Turgor pressure was altered by changing either pipette pressure or the osmolarities of extracellular solutions. Both positive pipette pressure and extracellular applications of hypotonic solutions, which caused cell size
to concomitantly increase, uncoupled the cells as indicated by reduced input capacitance and transjunctional conductance. These changes were, in many cases, reversible and repeatable. Intracellular application of 50 µM H-7, a
broad-based protein kinase inhibitor, and 10 mM BAPTA did not block the uncoupling effect of positive turgor
pressure on inner ear gap junctions. The transjunctional conductance at a holding potential of
80 mV was 53.6 ± 5.8 nS (mean ± SEM, n = 9) and decreased ~40% at a turgor pressure of 1.41 ± 0.05 kPa. Considering the coincident kinetics of cell deformation and uncoupling, we speculate that mechanical forces work directly on gap
junctions of the inner ear. These results suggest that pathologies that induce imbalances in cochlear osmotic pressure regulation may compromise normal cochlear homeostasis.
Key words:
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INTRODUCTION |
The supporting cells of the organ of Corti are structurally and electrically coupled together by gap junctions
(Jahnke, 1975; Gulley and Reese, 1976; Iurato et al.,
1976; Hama and Saito, 1977; Santos-Sacchi and Dallos,
1983; Kikuchi et al., 1995). Such gap junctional coupling among the supporting cells provides for electrical and metabolic uniformity; cochlear homeostasis is believed to rely on intercellular coupling (Santos-Sacchi,
1985, 1986, 1991; Kikuchi et al., 1995).
Gap junction channels are distinguished from other
ionic channels since the integration of two aligned
hemichannels from adjacent cells is required for normal function. In early work, hypertonic solutions,
which cause cell and tissue shrinkage, were found to uncouple gap junctions in several different preparations (Barr et al., 1965, 1968; Goodenough and Gilula,
1974; Loewenstein et al., 1967). More recently, hypotonic treatments, which cause cell swelling, were determined to either increase (Kimelberg and Kettenmann, 1990) or decrease (Ngezahayo and Kolb, 1990) gap
junctional coupling. These effects could have been due
to a variety of factors, including direct mechanical influences, changes in nonjunctional resistance, and
modulation of intracellular factors that are known to
uncouple cells. In the study of Ngezahayo and Kolb
(1990), where junctional conductance was studied directly, the decrease in coupling was abolished by 5 mM
EGTA in nominally Ca2+-free internal solutions, and
was linked to the activity of PKC. In the present report,
we used the whole-cell voltage clamp technique to examine the effects of turgor pressure on junctional coupling of isolated pairs or small groups of cochlear supporting cells. Both input capacitance (Santos-Sacchi,
1991; Bigiani and Roper, 1995) and transjunctional
conductance measures were used to gauge intercellular
communication. We report that data obtained with both techniques indicate that positive intracellular
pressure, which is known to induce membrane tension,
uncouples gap junctions of supporting cells in Corti's
organ.
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METHODS |
Detailed experimental methods can be found in previous reports
(Santos-Sacchi, 1991; Sato and Santos-Sacchi, 1994). In brief, isolated supporting cells or cell aggregates were freshly obtained from the organ of Corti of the guinea pig cochlea by shaking for 5-15 min in nominally Ca2+-free Leibovitz medium containing
1 mg/ml trypsin. To reduce the voltage-dependent ionic currents from nonjunctional membrane during double voltage
clamp experiments, cells were perfused with an ionic blocking solution containing (mM): 100 NaCl, 20 TEA, 20 CsCl, 1.25 CoCl2,
1.48 MgCl2, 10 HEPES, pH 7.2, 300 mosM. In initial experiments,
a modified Leibovitz medium was used for measurement of input
capacitance (Cin)1 with a single pipette voltage clamp containing
(mM): 136.9 NaCl, 5.37 KCl, 1.25 CaCl2, 1.48 MgCl2, 10 HEPES,
pH 7.2, 300 mosM. Pipette solutions were composed of (mM):
140 KCl, 10 EGTA or BAPTA, 2 MgCl2, and 10 HEPES, pH 7.2. For double voltage clamp recording, 140 mM KCl was replaced with
140 mM CsCl. Patch electrodes had initial resistances of 2.5-4
M
, corresponding to 1-2 µm in diameter. Series resistance (Rs)
after whole cell configuration was estimated from the current in
response to 10-mV steps (Huang and Santos-Sacchi, 1993). In single Hensen cells, where Rs could be unequivocally determined after whole cell configuration, the average value was 7.16 ± 0.43 M
(mean ± SEM, n = 48). Cells were typically held at
80 mV,
within the Hensen cell's linear current-voltage range (Santos-Sacchi, 1991). Currents were filtered at 10 kHz with a four-pole
Bessel filter (Axon Instruments, Foster City, CA). Intracellular pressure was modified either through the patch pipette with a syringe connected to the Teflon® tubing attached to the patch pipette holder or by changing osmolarity with "Y-tube" bath perfusion. Pipette pressure was monitored via a T-connector to a pressure monitor (World Precision Instruments, Inc., Sarasota, FL).
All experiments were video tape recorded and performed at
room temperature.
Since the input capacitance can be measured by a single pipette voltage clamp and is correlated with junctional conductance (Santos-Sacchi, 1991; Bigiani and Roper, 1995), it can be
conveniently used to study gap junctional coupling under conditions of less cellular damage than the double voltage clamp technique. Input capacitance, in conjunction with input resistance
(Rin), was continually measured on line to monitor junctional
coupling. Cin and Rin were determined from the transient charge
and steady state current, respectively, induced by small (
10
mV) test pulses with duration of 18× the clamp time constant at
the holding potential; measures were made at ~1-3 Hz (Santos-Sacchi, 1991).
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(1)
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(2)
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where
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(3)
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Qin is the charge moved, Vtest is the voltage of the test pulse, Ic is
the capacitive current induced by the test pulse, and
I
is the
current difference between the steady state current induced by
the test pulse and the holding current at the holding potential.
For the double voltage clamp, each cell in a cell pair was separately voltage clamped using 200A and 200B patch clamps (Axon Instruments). Both cells were clamped at the same holding potentials and a test pulse (10 mV, 10 ms) superimposed only on
cell 1. The transjunctional current (Ij) is equal to the current difference (
I2) in cell 2 caused by the test pulses applied to cell 1. The transjunctional conductance (Gj) can be calculated by:
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(4)
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where Vtest is the test pulse voltage applied to cell 1. Data collection and analysis were performed with an in-house developed windows-based whole-cell voltage clamp program, jClamp (http: //), using a Digidata 1200 board (Axon Instruments). In some experiments, Gj was measured online at 2-4 Hz and the corresponding
video images of recorded cells were digitally captured every 5-10 s
under software (jClamp) control. The captured images were
printed at ~1,700× and the plane cell areas calculated. To gauge
membrane stress, area strain (
A/A0) was calculated, where
A is
the change of cell area after pressure or osmotic treatment and
A0 is the original cell area.
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RESULTS |
Hensen cells can be easily distinguished from other inner ear supporting cells by their prominent lipid vacuoles. The number of cells comprising isolates of Hensen
cells can be determined under the light microscope,
and corresponds to the isolate's Cin since Hensen cells
are well coupled electrically. Although the size of
Hensen cells is variable, the distributions of Cin for one,
two, and three Hensen cells, whose numbers were visually confirmed, were quite distinct (Fig. 1 A). At the
holding potential of
80 mV, the peaks of the isolate
distributions were clearly separated at 31.03 ± 0.86, 64.75 ± 1.5, and 103.9 ± 3.05 pF, corresponding to
one, two, and three cell contributions, respectively.
The number of cells within isolates can also be confirmed using uncoupling agents, such as CO2, octanol,
or, as we now find, positive turgor pressure, to uncouple the cells. When cells fully uncoupled, Cin reached
single cell capacitance levels (e.g., Figs. 2 and 3). The
correlation of Cin with degree of cell coupling is illustrated by real measures of Cin in a coupled two-cell electrical model (Fig. 1 B). Cin of the electrical model was a
monotonic function of transjunctional resistance or
conductance, indicating the validity of Cin as an indicator of cell coupling.

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Fig. 1.
(A) The distributions of Cin of Hensen cell (HC) isolates. The cell numbers (1, 2, or 3) in the isolates were determined
under the light microscope, and the Cin was obtained at the holding potential 80 mV. Each bar represents the number of isolates
within a bin width of 10 pF. The lines plotted over each histogram
represent the fitted Gaussian distribution for the three isolate
groups. (B) Cin was measured for a coupled two-cell electrical
model as transjunctional resistance (Rj) was changed. Rs, 4.7 M ;
Rm, 500 M ; Cm, 33 pF. (inset) Coupled two-cell electrical model.
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Fig. 2.
Osmolarity-induced turgor pressure increase uncouples Hensen cells. Both bath solution and electrode solution were
300 mosM. (A) When an extracellular 150 mosM solution was perfused (indicated by the horizontal bar) the cell pair swelled (see
insets). Concomitantly, Cin decreased to half value due to cell uncoupling. Rs, 6.5 M . (insets) Captured cell images before and during the perfusion of the hypo-osmotic solution. The white scale bar
represents 20 µm. (B) Decreases of Cin due to hypo-osmotic challenge were reversible and repeatable. In this case, the cells finally
fully uncoupled and Cin remained at the single cell capacitance
level. Rs, 3.8 M .
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Fig. 3.
Hensen cells uncouple as turgor pressure is directly increased via the patch pipette. (A)
In this example, increased pipette pressure
caused Cin to decrease to single cell levels after an
initial delay (possibly due to pipette plugging),
but returned to the original level soon after pipette pressure was released. Rs, 9.76 M . (B) A
Hensen cell pair uncoupled after prolonged application of pressure. Cin decreased from 71.2 pF
to a single cell level of 34.7 pF. Rs, 8.5 M .
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Positive turgor pressure induced either by osmolarity
changes or directly via the patch pipette decreased Cin
of cell pairs or three-cell groups (Figs. 2 and 3), but did
not reduce single cell capacitance (Fig. 4). This indicates that positive turgor pressure uncouples gap junctions between adjacent Hensen cells.

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Fig. 4.
Changes of turgor pressure caused either directly via the patch pipette (A) or by perfusion of hypo-osmotic solutions (B) do not
decrease the measured capacitance in a single Hensen cell. (insets) Captured cell images before and during applications of positive intracellular pressure. Cell swelling was visible as turgor pressure increased. The white scale bar represents 20 µm. Rs: (A) 8.4 M , (B) 4.4 M .
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In Fig. 2 A, bath application of hypo-osmotic solution
(150 mosM) caused a Hensen cell pair to swell (insets)
and decreased Cin of the pair to single cell levels. The
uncoupling induced by increased turgor pressure is reversible since return to normal osmolarity solution often restored initial Cin values; subsequent reperfusion
with hypo-osmotic solution remained effective as an uncoupling stimulus (Fig. 2 B). In single cells, while the
same hypo-osmotic treatment caused cell swelling, Cin
remained stable (Fig. 4 B).
Fig. 3 illustrates the uncoupling effect of cell turgor
pressure change induced by patch pipette pressure. As
turgor pressure was directly increased to ~1.2 kPa via
the patch pipette, Cin decreased to almost single cell
levels (after an initial delay possibly due to pipette plugging), and immediately began to return when the pressure was released (Fig. 3 A). The cells could be permanently uncoupled during the application of prolonged,
continuous positive pressure (Fig. 3 B). The uncoupling effect of positive turgor pressure was found in 40 of 42 cell pairs, or three-cell groups. As with osmolarity
change, direct application of positive turgor pressure
via the patch pipette also did not decrease the measured capacitance in single Hensen cells despite cell
swelling (Fig. 4 A, insets).
Although Cin can be easily measured by single pipette
voltage clamp to gauge the degree of cell coupling,
transjunctional conductance cannot be measured directly since transjunctional voltage and current are unknown. Additionally, a quantitative estimate of degree
of coupling based on Cin is not easily established since
Cin is a nonlinear function of transjunctional conductance (see Fig. 1 B and DISCUSSION). To further investigate the uncoupling effect of positive turgor pressure
on gap junctions in Hensen cells, the transjunctional
conductance was directly assessed with a double voltage
clamp technique, and corresponding changes of the
cell plane surface areas (
A/A0) (i.e., an indicator of
membrane strain) were simultaneously measured.
Figs. 5 and 6 illustrate the results of such experiments. Cell areas increased in concert with decreases of
transjunctional conductance as positive turgor pressure
was delivered to the cells. The changes in cell area were
observable before gap junctional uncoupling and occurred faster than Gj decay (Figs. 5 and 6). However,
unlike pressure changes induced by pipette pressure, hypo-osmotic shocks produced changes in Gj and cell
areas that were quite fast. With extracellular perfusion
of a 150-mosM solution, the time constant for Gj decay
was 9.53 s in Fig. 6 B, and the average value was 5.1 ± 1.86 s (n = 6). In Fig. 6 B, the rise time constant of
membrane strain was 4.43 s. The average rise time constant of membrane strain is estimated to be close to or less than that of the average Gj decay since in most
cases the swelling fully occurred within the 5-10-s video
capture rate. In most, but not all, cases, it was noted
that after membrane tension stabilized, transjunctional
conductance likewise stabilized (Fig. 5). The correlated
and reciprocal changes in Gj and membrane strain
(
A/A0) were reversible and repeatable (Fig. 6 A),
strongly indicating that Gj decreases were relative to increases of membrane strain; i.e., membrane tension. It
should be noted that the latency to Gj change after
A/
A0 change is possibly due to the absence of significant
membrane stress during the initial cell inflation, which
clearly (based on the magnitude of cell enlargement)
was accompanied by membrane unfolding.

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Fig. 5.
Positive turgor pressure-induced reduction in transjunctional conductance is correlated with increases in membrane
strain ( A/A0). Transjunctional
conductance was measured with
a double voltage clamp technique, and membrane strains
were calculated from captured
images (see METHODS). Each cell
in a Hensen cell pair was separately held at 40 mV. The continuous treatments used to increase intracellular pressure began at the arrows. (A) Positive
pipette pressure induced an increase in membrane strain and
concomitantly reduced Gj. (B)
Perfusion of 150 mosM solution
also induced an increase in membrane strain and concomitantly
reduced Gj. The pipette solution
contained 50 µM H-7.
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Fig. 6.
(A) Hypo-osmotic solutions induced reversible, concomitant changes in transjunctional conductance and membrane
strain. Both cells were held at 40 mV. Repeated applications of
150 mosM solution are indicated by the horizontal bars. (B) The
time course of the uncoupling effect caused by hypo-osmotic
shock is compared with the increase of membrane strain. The beginning of treatment is indicated by an arrow and continued during the observed period. Thick solid lines represent single exponential fits. Membrane strain appears to have increased before the
decrease of Gj. Pipettes contained 50 µM H-7.
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Uncoupling of Hensen cell gap junctions by membrane stress was not inhibited by using pipette solutions
containing 50 µM H-7 (dihydrochloride; Calbiochem
Corp., La Jolla, CA), a broad-based serine/threonine
kinase inhibitor (Boulis and Davis, 1990) (Figs. 5 B and
6 B). These data imply that the uncoupling effect of
positive turgor pressure on inner ear gap junctions is
independent of protein kinases, and that the effect is
different from previous observations that cell volume
changes induced uncoupling of gap junctions via the
PKC pathway (Ngezahayo and Kolb, 1990). Nevertheless, cell swelling induced by hypo-osmotic shocks has
been linked to increases of another uncoupling agent,
intracellular Ca2+ (Hoffmann and Simonsen, 1989; Suzuki et al., 1990). However, uncoupling by Ca2+, which
occurs at millimolar intracellular concentrations in
Hensen cells (Sato and Santos-Sacchi, 1994), can be
ruled out since pipette solutions contained 10 mM
BAPTA, a fast highly selective calcium chelating reagent, and extracellular and intracellular solutions were nominally Ca2+ free. Considering all evidence, the
observed uncoupling effect of positive turgor pressure
on inner ear gap junctions, which is fast (within seconds), correlated with changes of membrane strain,
and independent of protein kinases and Ca2+, is likely
to occur via direct mechanical effects on the plasmalemma; i.e., membrane tension.
The effect of membrane tension on gap junctional
conductance was further studied by increasing turgor
pressure in cell 1 and measuring Ij in cell 2 at different
membrane potentials (Fig. 7). Gap junctional conductance in Hensen cells at a holding potential of
80 mV
was 52.9 ± 12.1 nS (n = 51). As the turgor pressure in
cell 1 was increased, Ij decreased (Fig. 7 A). The junctional conductance at different membrane potentials
reduced in parallel when the turgor pressure was increased. In those cell pairs where turgor pressure alterations were successfully applied without losing the cells,
Gj at
80 mV holding potential decreased 38.3 ± 9.5%
from 50.5 ± 14 nS (n = 9) at a turgor pressure of 1.41 ± 0.05 kPa. The Vm dependence of Gj is also visible in
Fig. 7. In this case, as the cells were depolarized, Gj decreased (Fig. 7 B). Other Vm dependencies of transjunctional conductance were also found, including Vm
insensitivity and an increase with depolarization. Pressure did not alter voltage-dependent behaviors.

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Fig. 7.
Turgor pressure does not affect membrane potential
(Vm or Vi-o) dependence of gap junctions in Hensen cells. (A) Voltage stimulus protocols for each cell and current trace recorded in
cell 2 are plotted (only five traces are shown for clarity). Each cell
in a Hensen cell pair was separately voltage clamped at the same
holding potential of 80 mV. Voltage steps from 140 to 70 mV
for 100 ms in 10-mV increments were simultaneously delivered
into both cells except for 10 mV, 10-ms test pulses superimposed
on cell 1 only. Transjunctional current (Ij) is measured in cell 2. The turgor pressure of cell 1 was directly changed by the patch pipette. Three current traces from cell 2 at different pressures were
zeroed and superimposed in the middle. (B) Positive turgor pressure decreased Gj at all membrane potentials. Note Gj decrease as
the cells were depolarized. Rs at 0 kPa, 12.5 M ; at 1.2 kPa, 12.7 M ; at 2 kPa, 12.2 M .
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DISCUSSION |
We provide evidence, based on input capacitance and
double voltage clamp measures, that junctional coupling is sensitive to positive turgor pressure-induced
membrane tension. Turgor pressure has been used to
induce membrane tension in a wide variety of cells, including the outer hair cell (OHC), where it has been shown that motility and motility-related gating current
characteristics are directly altered (Iwasa, 1993; Gale
and Ashmore, 1994; Kakehata and Santos-Sacchi, 1995).
Membrane tension (possibly acting via cytoskeletal interactions) is also known to gate stretch-activated ionic
channels (Yang and Sachs, 1989), which have been observed in outer hair cells (Ding et al., 1991; Iwasa et al.,
1991). It is possible that membrane tension also alters
gating characteristics of supporting cell gap junctions.
We show, however, that unlike stretch channels, inner
ear gap junctional conductance decreases with membrane stress. Recently, it has been postulated that gap junction channels possess two distinct gating mechanisms, namely, a voltage gating mechanism and a
chemical gating mechanism (Bukauskas et al., 1995;
Bukauskas and Peracchia, 1997; Bukauskas and Weingart, 1994). Chemical uncoupling agents, such as CO2,
H+, and Ca2+, may act on sensor elements from the cytoplasmic side. Supporting cell coupling has been
shown to be sensitive to a variety of chemical uncoupling agents (Santos-Sacchi, 1985; 1991), and we now
report that supporting cell coupling is voltage dependent as well. The existence of voltage-dependent gap
junctional conductance may account in part for previous reports of temperature-induced depolarization on
supporting cell coupling ratios (Santos-Sacchi, 1986).
Interestingly, junctional voltage dependence is unaffected by concomitant tension-induced junctional conductance change, possibly indicating that an independent tension gating mechanism may exist.
Gap junctions consist of two aligned transmembrane
hemichannels (connexons), one from each cell (Revel
et al., 1984; Goodenough et al., 1988; Bennett et al.,
1991). Each of these hemichannels is formed by six
connexin subunits (Kumar and Gilula, 1996; Perkins et
al., 1997). Our data indicate that membrane stress acts
on inner ear gap junctions in a manner independent of
Ca2+, pH, and protein kinases. The rapid and reversible nature of the uncoupling also indicates that the
mechanism is not due to some sort of mechanical destruction of the channels. While there may be other unknown links between membrane stress and junctional conductance, it is conceivable that tension may gate
gap junction channels by a conformational change in
connexon structure, possibly causing only the stressed
membrane's hemichannel to close.
Gap junction connexins represent a family of homologous proteins that have differing voltage gating characteristics (Harris et al., 1981; Spray et al., 1981; Bennett et al., 1991; Dahl, 1996). Using immunocytochemistry and transmission electron microscopy, Cx26 was
found in gap junctions of the rat (Kikuchi et al., 1995) and gerbil (Forge et al., 1997) organ of Corti. More recently, Cx26, Cx30, Cx32, and Cx43 have been localized to supporting cell regions of the rat cochlea (Lautermann et al., 1997). Such diversity of connexins within
the organ may provide for a variety of junctional communication characteristics; for example, rectifying junctional conductance. Indeed, in addition to our direct
observation that voltage-dependent junctional communication exists in the supporting cells, we have preliminary evidence that junctional rectification occurs. Directional flow of ions mediated by rectified gap junctions may be crucial for normal cochlea homeostasis
(see below).
Since the mid 1980's, gap junctional coupling has
usually been studied with double voltage clamp. However, input capacitance and resistance reflect the degree of electrical coupling and can be conveniently
measured using a single voltage clamp (Santos-Sacchi, 1991; Sato and Santos-Sacchi, 1994; Bigiani and Roper,
1995). Based on a coupled two-cell model (see Fig. 1 B,
inset), and assuming that the individual cells have the
same input impedance, the following equations are obtained (Bigiani and Roper, 1995),
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(5)
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(6)
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where Rs and Rm are electrode series resistance and
nonjunctional membrane resistance, respectively, and
Cm is single cell capacitance (see Fig. 1 B, inset). Since
Rm is not readily available from recordings, we can solve
Eqs. 5 and 6 to remove Rm. Rj can be finally expressed:
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(7)
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Cin, Rin, and Rs are readily obtained from recordings.
Fig. 8 illustrates the measurement of these parameters
during an uncoupling event, and the bottom panel
shows the estimated Gj based on those data. Changes in
estimated Gj mirror pressure-induced changes in Cin. It
should be noted that Rs changes can also produce
changes in Cin and Rin. For example, to obtain the observed maximum change in Cin, an order of magnitude
increase of Rs would be required in this case. In our experiments, changes solely in Rs required to produce a
comparable change in Cin were not observed. Series resistance remained constant, being 7.79 ± 0.49 M
(n = 7) for cell pairs that were well coupled and 6.34 ± 1.13 M
after those same cells were uncoupled with positive
pipette pressure.

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Fig. 8.
Transjunctional conductance is estimated by Cin and
Rin in single pipette voltage clamp. (top) Cin decreased and Rin simultaneously increased as turgor pressure increased. Cin and Rin
followed changes in pressure. Rs, 8.73 M . (bottom) Gj is estimated
from Cin and Rin measurements from the cell pair data based on
the coupled two-cell model. Cm was set as half of Cin, 27.5 pF, at
zero applied pressure, and Rs was 8.5 M . The change in estimated Gj corresponds well to changes in Cin. The magnitude of Gj
is larger than the largest obtained under double voltage clamp
(~500 nS), and may be due to the fact that cell impedances are
not actually identical as required in the model.
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Finally, how might the turgor pressure dependence
of junctional coupling in the organ of Corti affect cochlear function? In vivo, the organ of Corti, comprising
hair cells and supporting cells, is bathed in two different media, high K+ endolymph apically and low K+ perilymph basally. Since the receptor current through hair
cells is predominantly carried by K+, an accumulation
of K+ within the perilymphatic space along the basolateral region of the hair cells is unavoidable. This would
potentially depolarize hair cells with disastrous consequences for both forward and reverse sensory transduction. In the mammal, forward transduction (gating of
stereociliar transduction channels) relies on the large
driving force present across the hair cell's apical plasma
membrane. Voltage gradients across the apical membranes of inner and outer hair cells (i.e., endolymphatic potential minus membrane potential) range
from 125 to 150 mV, and drive the K+-based receptor
currents. Reduction of this gradient (e.g., by membrane depolarization) will reduce the magnitude of receptor potentials and synaptic output. Reverse transduction is a phenomenon that is restricted to the outer
hair cell and is believed to provide for the enhanced
high frequency selectivity and sensitivity enjoyed by
mammals. OHCs, which are additionally mechanically
active, possess lateral membrane motors that are driven
by voltage (Santos-Sacchi and Dilger, 1988); the cell's
mechanical response provides feedback into the basilar
membrane, thereby enhancing the stimulus to the primary receptor cells, the inner hair cells (for review see
Ruggero and Santos-Sacchi, 1997). Not only will depolarization of the OHC alter the driving force for the
mechanical response, but the function relating mechanical response to voltage will be shifted along the
voltage axis as well, resulting in an altered gain for the
"cochlear amplifier" (Santos-Sacchi et al., 1998). Some
mechanisms must prevent such an undesirable scenario. A nutritive and K+ sinking role for gap junctions
in the avascular organ of Corti has been proposed
(Santos-Sacchi, 1985, 1991; Santos-Sacchi and Dallos,
1983). Inner ear supporting cells have been shown to
"share" plasmalemmal voltage-dependent conductances
due to the high degree of cell coupling (Santos-Sacchi,
1991). The magnitude and stability of their resting potentials is pronounced (close to
100 mV), and likely
depends on cell coupling since isolated cell resting input conductance is only ~1 nS. At the normal resting
potential of this cellular syncytium, an inward rectifier
appears continuously activated and may result in K+ removal from perilymphatic spaces. It should be noted
that the large perilymphatic fluid spaces may provide
little support in sinking K+ or directing its movement,
since hair cell regions that are likely to experience K+
elevations are not directly exposed to those spaces. Inner hair cells are closely surrounded by supporting
cells, and the region of the OHCs that possesses voltage-dependent conductances (e.g., outward K+) is restricted to the basal pole of the cell (Santos-Sacchi et al., 1997), which is surrounded by a Deiters cell cup.
Recently, Kikuchi et al. (1995) provided morphological
evidence detailing epithelial and connective tissue gap
junctional systems within the cochlea that may complete the mechanism responsible for recycling K+ from
the perilymphatic space near hair cells to the K+-rich
endolymph via the stria vascularis. The maintenance of
normal fluid space architecture within the inner ear requires fine osmotic control; imbalances can lead to serious auditory and vestibular problems (e.g., Meniere's
disease). While at present we do not know the normal
physiological significance of tension-dependent gap
junctional communication, it is likely that fluid balance disorders in the inner ear will affect gap junctional
communication, thus compromising sensory function
by indirectly modifying hair cell activity.
Address correspondence to Joseph Santos-Sacchi, Ph.D., Professor,
Surgery (Otolaryngology), BML 244, Yale University School of Medicine, 333 Cedar St., New Haven, CT 06510. Fax: 203-737-2245; E-mail:
joseph.santos-sacchi{at}yale.edu
Original version received 12 January 1998 and accepted version received 24 July 1998.
Portions of this work were previously published in abstract form
(Zhao, H.B., and J. Santos-Sacchi. 1997. This work was supported by National Institute on Deafness and Other Communication Disorders grant DC00273 to J. Santos-Sacchi.
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