Excess plasma membrane and effects of ionic amphipaths on
mechanics of outer hair cell lateral wall
Noriko
Morimoto,
Robert M.
Raphael,
Anders
Nygren, and
William E.
Brownell
Department of Otorhinolaryngology and Communicative
Science, Baylor College of Medicine, Houston, Texas 77030
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ABSTRACT |
The interaction between the outer
hair cell (OHC) lateral wall plasma membrane and the underlying
cortical lattice was examined by a morphometric analysis of cell images
during cell deformation. Vesiculation of the plasma membrane was
produced by micropipette aspiration in control cells and cells exposed
to ionic amphipaths that alter membrane mechanics. An increase of total
cell and vesicle surface area suggests that the plasma membrane
possesses a membrane reservoir. Chlorpromazine (CPZ) decreased the
pressure required for vesiculation, whereas salicylate (Sal) had no
effect. The time required for vesiculation was decreased by CPZ,
indicating that CPZ decreases the energy barrier required for
vesiculation. An increase in total volume is observed during
micropipette aspiration. A deformation-induced increase in hydraulic
conductivity is also seen in response to micropipette-applied fluid jet
deformation of the lateral wall. Application of CPZ and/or Sal
decreased this strain-induced hydraulic conductivity. The impact of
ionic amphipaths on OHC plasma membrane and lateral wall mechanics may
contribute to their effects on OHC electromotility and hearing.
membrane reservoir; electromotility; membrane bending; hearing; hydraulic conductivity
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INTRODUCTION |
THE OUTER HAIR
CELL (OHC) contributes to fine-tuning in the mammalian inner ear
through its unique membrane potential-dependent motility. OHC
electromotility is a shape change that results from the direct
conversion of electrical potential into a mechanical force
(6). It does not depend on calcium or intracellular stores of ATP (7, 30). The mechanism underlying electromotility is associated with the plasma membrane and its lipid environment. This
membrane motor transmits force via the OHC cytoskeleton to produce cell
deformation. Cellular micro- and nanomechanics are integral to the
mechanism of OHC force production (9). Theoretical models
predict that passive mechanical properties of the OHC should affect
electromotility (29, 52, 72, 73). The protein prestin was
recently isolated and demonstrated to be involved in the motor mechanism (36, 63, 78) along with small anions
(44). These findings provide support for the hypothesis
that electromotility is associated with the conformational change of an
integral membrane protein (13, 29, 62). Hence, the
mechanical properties of the plasma membrane (PM) should influence the
function of the protein and therefore electromotility. Although the
motor is often specified to be an area motor, any out-of-plane bending
of the membrane (as commonly occurs in biological membranes) would
greatly decrease the efficiency of such a mechanism. Ionic amphipaths readily insert into membranes and are known to alter the curvature and
mechanics of erythrocytes (66) as well as OHC mechanics (15, 24, 32, 37, 46, 59, 67, 74, 77). These molecules may
be used to probe the effects of membrane curvature on OHC function.
The OHC lateral wall is a trilaminate structure composed of the PM,
cortical lattice (CL), and subsurface cisterna (SSC). Electron
micrographs reveal that the outermost layer PM is rippled (15,
70) whereas the membrane of the outermost layer of the SSC
presents a flat, crisp bilayer appearance (15, 60). The CL, sandwiched between the SSC and the PM, is arranged in cytoskeletal microdomains of parallel actin filaments cross-linked with spectrin (21, 25-27). Bridging the distance between the actin
filaments and the PM are structures of unknown composition called
pillars (2, 3, 21). The OHC is a cellular hydrostat with a
positive turgor pressure. This pressure maintains the cell shape and is required for the full expression of the electromotile response (5, 31, 61, 67). The regulation of cell volume and
cytoplasmic pressure is particularly important for the OHC, and
previous studies explored the effective water permeability of the OHC
PM (8, 11, 56).
Micropipette aspiration is a technique that measures cell mechanical
properties and has been applied to lipid vesicles and red blood cells
(4, 18). This technique has also been applied to the OHC
to measure the "stiffness parameter" of the lateral wall (37,
42, 69). Because of the orthotropic nature of the cell, this
stiffness parameter is a combination of its axial and radial stiffness
(71), both of which have components originating from the
membrane and cytoskeleton. These experiments revealed that as
the pressure was increased, vesiculation of the PM occurred (69). Oghalai et al. (42), using fluorescence
labeling specific for the PM, F-actin, and SSC, showed that the
vesicles contained only the PM label. In the study reported here, we
used the micropipette aspiration technique and morphometrically
analyzed the vesiculation process. The results demonstrate the
existence of excess PM in the OHC. The existence of membrane reservoirs
in fibroblasts has been demonstrated with tether formation experiments,
in which narrow membranous tubes are pulled from the surface of a cell (57). We have applied this technique and demonstrated
excess membrane in OHCs (1, 35). A membrane reservoir is
important in the function of endothelial cells and neutrophils
(64). Membrane unfolding is also believed to play a role
in other cells, such as the response of alveolar epithelial cells to
mechanical forces in the lung (76).
We also investigated the effect of altering membrane mechanics with
ionic amphipaths. The anionic amphipath salicylate (Sal) diminishes OHC
electromotility (67), decreases OHC axial stiffness, and
decreases OHC force production (24). The lateral wall
stiffness parameter is also affected by Sal (37). In this
report, we extend previous studies of the effect of Sal on membrane
mechanics by exploring how Sal affects the vesiculation process. In
addition, we investigated the effect of the cationic amphipath
chlorpromazine (CPZ), because of its effects on the red blood cell as
well as our observation that it alters lipid lateral diffusion in the PM of the OHC (43) and translates the voltage-displacement
function for OHC electromotility toward depolarizing potentials
(38). Both drugs reduced the aspiration-associated
increase in water permeability observed in OHCs. The findings presented
here indicate that Sal and CPZ change the mechanical properties of OHC
membranes consistent with their effects on OHC electromotility and
hearing. A preliminary version of these results was presented
previously (40).
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MATERIALS AND METHODS |
OHC isolation.
Albino guinea pigs weighing 200-400 g were decapitated by
guillotine. The temporal bones were taken, and the middle ear bullae were opened. The otic capsule was removed, and the spiral ligament was
peeled off to expose the organ of Corti. The organ of Corti was
dislodged from the cochlea and harvested in a standard medium. OHCs
were isolated from the organ of Corti by gentle trituration without
enzyme (11). All OHCs were plated onto a glass coverslip cemented to the bottom of a plastic chamber. Isolated cells were selected for study on the basis of standard morphological criteria within 3 h of animal death. Healthy cells displayed a
characteristic birefringence and a transparent cytoplasm, the nucleus
was in the base of the cell, and cells possessed a uniformly
cylindrical shape without regional swelling. Cells were excluded if
Brownian motion of intracellular particles was observed.
Solutions.
The standard medium was composed of HEPES-buffered extracellular
solution supplemented with (in mM) 135 NaCl, 4 KCl, 1 MgCl2, 2 CaCl2, 10 HEPES, and 10 glucose. The
solution had an osmolarity of 285-290 mosM and a pH of 7.4 and was
at room temperature (22-24°C). The hypotonic solution
consisting of the standard medium and distilled water was adjusted to
an osmolarity of 250 mosM and a pH of 7.4.
For some experiments, 10 mM sodium salicylate (S-3007, Sigma, St.
Louis, MO) and/or 0.1 mM CPZ (C-8138, Sigma) were added to the
extracellular solution. Each drug was dissolved in the extracellular
solution at twice the desired concentration. A volume of the 2× drug
solution was added to an equal volume of the control solution
containing OHCs. Each experiment was started within 10 min of
application of drugs and was finished in 40 min.
Experimental protocols and data collection.
A chamber containing cells was placed onto the stage of an inverted
microscope (Zeiss Axiovert 135TV), and the cells were imaged using an
oil-immersion ×100 objective lens. A Dage MTI Newvicon video camera
was used to capture images. Either negative or positive pressure
applied via a small-diameter pipette was used to deform the OHC lateral
wall. A negative pressure was applied during micropipette aspiration. A
positive pressure was applied to a pipette containing hypoosmotic
extracellular medium (250 mosM) to produce a fluid jet. Both protocols
were followed after application of 10 mM Sal, 0.1 mM CPZ, or both. The
response of the OHC was monitored on a television screen and recorded
with a videocassette recorder (Panasonic AG1960, S-VHS). The time and date were superimposed and recorded on the video image. The level of
the water column and other experimental details were recorded on one of
the audio channels.
Micropipette aspiration.
Aspiration micropipettes were fabricated from borosilicate glass
capillary (LG16, 1.65-mm OD, 1.1-mm ID; Dagan, Minneapolis, MN) with a
programmable micropipette puller (BB-CH-PC; Mecanex, Geneva,
Switzerland). The inner diameter of the tip was ~3 µm and was
fire-polished with a microforge (Microforge De Fonbrune; Aloe
Scientific, St Louis, MO).
The micropipette was connected to a water column via a polyethylene
tube that was filled with the extracellular solution and 0.2 mg/ml
bovine serum albumin (Sigma). The micropipette was mounted on a
joystick-controlled manipulator (Zeiss) and advanced at a shallow angle
(<20° against the stage) at the center of the field of view. The
zero pressure point of the water column was referenced to the level of
the microscope stage and was defined as the water level at which no
flow in or out of the micropipette was observed.
Once a cell was located, the micropipette was brought into contact with
the side of the OHC midway on its lateral wall. Negative pressure was
applied in a stepwise manner as measured from the null point until
vesiculation occurred. Initially, the entire trilaminate lateral wall
is aspirated into the pipette. Eventually, the PM separates from the
underlying CL and only the PM continues to elongate. The PM pinches off
and forms a vesicle that is composed solely of PM (42).
Several vesicles were removed from each cell by aspiration. The maximum
number of vesicles that can be removed varies from cell to cell. More
than 20 vesicles have been removed from long OHCs (69).
When a cell-specific number of vesicles have been removed (presumably
when all the excess PM has been exhausted), the membrane in the
micropipette ruptures and there is a precipitous loss of cell volume as
indicated by the collapse of the cell. At this point, the cell is often
sucked into the micropipette and lost.
Morphometric analysis.
A cell was subjected to morphometric analysis before and after vesicles
were pulled. Analysis was made only if there was no Brownian motion of
internal organelles. Single frames from the S-VHS tape were selected
and digitized with NIH Image. Each image was traced five times per cell
with a MATLAB program (Mathworks). The program was similar to an
earlier morphometric analysis program (11). The surface
area and volume of the cell are approximated as a sum of cylindrical
segments, each with surface area
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(1)
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and volume
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(2)
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Where
x and ri are the
height and radius, respectively, of segment i. The whole
cell was traced before aspiration. Each vesicle that pinched off during
the aspiration process was traced. The cell was traced again after
release of three, six, and nine vesicles. The resolution of the
digitized images was 5.5 pixels/µm. The surface area and volume,
length, and radius of the cell (defined as the average radius for
segments in the middle 2/3 of the length of the cell) were calculated
assuming radial symmetry about the long axis (Fig.
1A). The surface area and
volume of the vesicle were calculated by dividing the vesicle into
three sections. The center part was modeled as a cylinder, and both
ends were modeled as hemispheres (Fig. 1B). An error
analysis (see APPENDIX A) supports the conclusions based on
these measurements.

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Fig. 1.
Morphometric analysis of the outer hair cell (OHC) and
vesicle. A: segmentation of an OHC image. The surface area
and volume, length, and radius of the cell were calculated assuming
radial symmetry about the long axis. B: vesicle analysis.
The surface area and volume of the vesicle were calculated by dividing
the vesicle into 3 sections. The center part was assumed to be a
cylinder, and both ends of the cap were assumed to be a part of a
sphere.
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Micropipette fluid jet protocol.
Micropipettes with an inner diameter of ~5 µm were fabricated in
the same manner as aspiration micropipettes. The micropipette was
connected to a water column, and the entire fluid system was filled
with hypotonic extracellular solution. Once a cell was located, the tip
of the pipette was brought within 10 µm of the cell. The water column
was raised up to +20 cmH2O, and a jet of water on one side
of the OHC was produced. A jet of hypoosmotic medium (250 mosM)
produces a local dimpling of the lateral wall, and then the cell starts
swelling. The associated volume increase as a function of time was
analyzed to determine the local hydraulic conductivity (8,
56).
The whole cell was traced before application of the fluid jet. The cell
was traced after 5, 10, 15, and 20 s and the length, radius,
surface area, and volume were calculated with the MATLAB program
described in Morphometric analysis.
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RESULTS |
Cell shape changes during micropipette aspiration.
The effects of removing PM vesicles on the geometry of the OHC are
summarized in Fig. 2A, which
shows the length and the radius of the cell during aspiration. Control
OHCs maintained a constant radius but decreased their length by 15%
during the aspiration process. Exposure to either Sal or CPZ did not
result in an immediate change in cell shape. However, these drugs did
affect the behavior of the cell during the aspiration process. When
either Sal or CPZ was applied, the length decrease was ~30% and the
radius increase was ~10% more than that of control cells. When Sal
and CPZ were applied together, the cell's shape change resembled that
of control cells. Figure 2B shows the OHC length and radius
change accompanied by cell swelling in response to the hypoosmotic
fluid jet. A length decrease of ~5% and a radius increase of 10%
were observed for all groups.

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Fig. 2.
Change of OHC length and radius. A: mean
length and radius of cells after removal of 3, 6, and 9 vesicles during
aspiration. B: mean length and radius of cells during fluid
jet protocol. Values are means ± SE (*P < 0.01, repeated-measures ANOVA). Sal, salicylate; CPZ, chlorpromazine.
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Surface area change.
The total surface area for the OHC and the sum of the removed vesicles
was compared with the surface area of the cell before the aspiration
process was initiated. The total mean surface area gradually increased
~7% for all groups (Fig.
3A). Removed original surface
area was compared with total vesicle surface area (Fig. 3B).
In almost all cases, the ratios of removed surface area from the OHC
over the formed vesicle surface area are <1. This ratio should be 1 if
membrane area is conserved. The total removed vesicle surface area
exceeds the original apparent surface area of the cell, indicating that
the intact OHC PM possesses excess PM. This excess membrane is likely
stretched during aspiration and covers the vesicle when vesiculation
occurs. There are no apparent drug effects on the control surface area
of the PM. To observe whether the drugs affected the size of the
vesicles, the surface area of each vesicle was compared (Fig.
3C) and was positively correlated with the pipette diameter
unrelated to drug application. The results of the hypoosmotic fluid jet
experiment confirm that the apparent surface area remains constant
despite volume change (Fig. 3D) as previously reported
(8). There are also no detectable drug effects on the
apparent surface area, as is the case with aspiration.

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Fig. 3.
Change of OHC surface area. A: total surface area
increases for all groups during aspiration protocol. Values on the
abscissa are the sum of the OHC surface area after aspiration of
n (ordinate) vesicles plus the total area of the aspirated
vesicles, divided by the OHC surface area before aspiration. Values are
means ± SE. B: surface area removed from the original
OHC surface area divided by total vesicle surface area. Vesicle surface
area is larger than removed apparent surface area because these ratios
are <1. The number above each bin in the histogram is n.
Error bars indicate SE. C: surface area for each vesicle is
proportional to the pipette diameter unrelated to the drug effects.
D: surface area remains constant although the cell's volume
increases dramatically during fluid jet protocol. Values are means ± SE.
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Drug-induced mechanical property changes.
Figure 4A presents the
pressure required to remove the first vesicle in the aspiration
protocol. CPZ decreased this pressure, whereas Sal did not change it.
Even less pressure was required when both Sal and CPZ were present. The
mean time to form subsequent vesicles was also measured (Table
1). The mean time between vesicles was
greatest for control and Sal-treated cells; CPZ and Sal + CPZ
reduced the time >50%. Because there were no differences in size
among the vesicles, it appears that CPZ facilitates vesicle formation
by decreasing the work required for vesiculation.

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Fig. 4.
A: pressure required to remove 1 vesicle. B:
effective lateral wall tension to form the vesicle. The effective
tension is calculated with Eq. B3. Error bars indicate SE
(*P < 0.001, unpaired t-test). The same
cells were used for both A and B. The number of
cells was 134 control, 220 Sal, 192 CPZ, and 189 Sal + CPZ.
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The relevant mechanical property for describing these drug effects is
the tension within the lateral wall. To interpret the data in this
context, we converted the aspiration pressure to an effective tension
(see APPENDIX B). The results in Fig. 4B
demonstrate the effect of drugs on the calculated tension required to
form a vesicle. Sal increased and CPZ decreased the effective tension
required to form a vesicle.
Drug effects on hydraulic conductivity of PM.
The total volume of the OHC and sum of the removed vesicles was
compared with the volume of the cell before the aspiration process was
initiated (Fig. 5A). Cytosol
was lost during aspiration for all groups. The volume loss for control
cells was ~13%, whereas CPZ-treated cells lost <5%. Figure
5B shows the proportion of cytosol leakage for each group.
After six vesicles were removed, the amount of cytosol lost through
leakage was half as large in Sal- and CPZ-treated cells as in control
cells. There were no significant differences in the vesicle volume
regardless of the drug applied. The strain-induced volume increase
associated with perfusion with a hypoosmotic fluid jet was measured and
plotted in Fig. 5C. The slope (Lp) of
each plot is a measure of the hydraulic conductivity of the PM
(8, 56). The plot for the control cells has a slope that
is steeper than those for drug-treated cells.

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Fig. 5.
A: total volume change throughout aspiration. Total pre-
and postexperiment OHC and vesicle volume are compared. When CPZ was
applied, volume was decreased less than those of control cells. Values
are means ± SE (*P < 0.05, repeated-measures
ANOVA). B: volume loss during aspiration. Removed original
volume from OHC and vesicle volume is compared during aspiration.
Removed original volume is more than twice greater than vesicle volume
(*P < 0.01, P < 0.05, unpaired
t-test), which indicates cytosol leakage. Bars indicates SE.
Top number of each histogram shows n. C: volume
change during fluid jet. Volumes for OHC before and after hypoosmotic
fluid jet protocol are compared. The slope (Lp;
10 14 mPa 1s 1) of each plot is
a measure of the cell's hydraulic conductivity (R > 0.9). The slope for control cells is steeper than those of drug-treated
cells. Values are means ± SE of 7-9 independent
determination. CPZ- and Sal + CPZ-treated cells have significant
difference with control cells (*P < 0.05, repeated-measures ANOVA).
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DISCUSSION |
The major observations of this study are that the OHC PM possesses
excess membrane and that Sal and CPZ alter the cell's mechanical properties such as lateral wall tension and PM hydraulic conductivity. The excess surface area measured for nine vesicles (Fig. 3A)
was slightly more than the excess represented by membrane folding as
measured in an ultrastructural study on the OHC (75).
Excess membrane or a membrane reservoir has also been observed while pulling membrane tethers from an OHC with optical tweezers (1, 35). The tether experiments reveal a strong attachment force between the membrane and the CL. After the membrane separates from the
CL, the force required to pull a tether falls to a nearly constant
value and tethers of >50 µm in length are routinely pulled. The
force plateau demonstrates that no additional force is needed during
this phase of tether elongation. If stretching of the membrane and/or
underlying structures were involved, additional force would be
required. Long membrane tethers that require no additional force for
their formation are consistent with a membrane reservoir (57).
The folded PM visible in transmission electron microscopy is an obvious
source of excess membrane, but it is also possible that internal
membranes contribute to the membrane reservoir. Micropipette aspiration
experiments indicate that membrane insertion from internal stores does
not occur before vesiculation. Micropipette aspiration studies on the
kinetics of amphiphilic insertion reveal membrane insertion by a
clearly visible increase in the projection length within the pipette at
a constant aspiration pressure (41). In our experiments,
the OHC projection length does not creep at constant pressure
(42, 69), indicating that a deformation-induced membrane
insertion does not occur before vesiculation. If membrane insertion did
occur after vesiculation, the most likely source of internal membranes
would be the SSC, and we demonstrated previously (42) that
SSC membranes do not insert into PM during the formation of the first
five or six vesicles. This finding is consistent with an earlier study
showing that uptake of extracellularly applied horseradish peroxidase
is not observed in the lumen of the SSC, indicating that the SSC
belongs to a separate membrane pool than that of the endoplasmic
reticulum-Golgi-PM (68). In addition, there has never been
a report of vesicle fusion to the OHC PM for any location along the
lateral wall where the intimate association between the CL and the PM
might interfere with membrane fusion. There is evidence for membrane
turnover at both the base and apex of the cell, where vesicle fusion
has been shown ultrastructurally (68), and membrane
recycling has been observed through imaging pinocytotic uptake of
fluorescent-labeled PM (42). The insertion of additional
membrane from internal membrane stores most likely occurs at these
locations. The new membrane must then diffuse along the length of the
lateral wall. For the fast, physiologically relevant changes in cell
length and membrane tension associated with OHC function, an
interaction with an internal membrane pool is probably not important.
Hence, the most direct interpretation of our experiments is that after
the attachment to the CL is broken, the overlying membrane is free to
unfold and this is responsible for the experimentally observed excess
membrane. Additional pipette aspiration studies that simultaneously
monitor membrane capacitance and/or confirm changes in membrane folding
by transmission electron microscopy could further probe the magnitude,
origin, and functional significance of the OHC membrane reservoir.
The membrane reservoir is likely to be associated with nanoscale
membrane curvature. Sal and CPZ are ionic amphipaths of opposite charge
that alter PM curvature (Fig. 6). The
theory of chemically induced bending moments (19) predicts
that membrane bending results from an increase in the area of one
leaflet relative to the other. The differential area expansion causes a
bending in the direction of the layer into which the amphipath inserts.
The tendency of an amphipath to intercalate in one layer or the other depends on its molecular shape and charge. Sal is expected to preferentially intercalate in a positively charged leaflet, whereas CPZ
will prefer a negatively charged leaflet. In red blood cells, Sal
causes outward membrane buckling (crenation) (34) whereas CPZ causes inward buckling (cupping) (58, 65, 66). The
surface charge of the OHC membranes has not been directly measured, but negatively charged liposomes adhere to the OHC, suggesting that its
outer membrane is positively charged (51).

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Fig. 6.
Partition of ionic amphipaths in the PM. Sal intercalates
into the positively charged outer leaflet of the PM and expands its
area, causing the membrane to bend outward, whereas CPZ intercalates
into the negatively charged inner leaflet, resulting in an inward
bending of the membrane. CL, cortical lattice.
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Excess PM and OHC shape.
The membrane contained in the ripples observed in electron microscopic
studies (15) is likely to be the source of the vesicle membrane. During aspiration, it is likely that the rippling is lost,
particularly as the vesicles form without the support of the CL
(42). In contrast, during the volume increase resulting from the hypoosmotic fluid jet there is no change in apparent surface
area (Fig. 3D), suggesting that the reduction in osmotic pressure resulting from water entry and an increased wall tension serve
to maintain the apparent surface area of the lateral wall constant in
the face of increased cell volume. A membrane reservoir has
implications for the mechanism of electromotility. Excess membrane area
and the associated nanoscale rippling would decrease the efficiency of
a mechanism based on area change or the development of in-plane
strains. However, nanoscale rippling is consistent with the recently
proposed membrane-bending motor model of electromotility (52). Moreover, the voltage- and tension dependence of
lateral diffusion in the PM can be interpreted on the basis of
nanoscale rippling (43).
Alteration of mechanical properties of PM.
Figure 7 shows two steps of the
vesiculation process. The first step occurs when the PM breaks away
from the CL-SSC complex (Fig. 7A). The second step is when
the PM stretches and elongates until a vesicle pinches off (Fig.
7B). As the PM stretches, it flows around the pillar
attachment points. The nature of the PM's attachment to the pillars is
unknown. Electron microscopic studies show that the pillars directly
insert into the membrane (22). Integral membrane proteins
are thought to prefer regions of the cell with a specific membrane
curvature (12, 14, 33, 48). The connection between the
pillars and the PM may require a PM with a local negative curvature
(membrane bending into the cell). The drawings in Fig. 6 representing
normal and Sal-treated cells show negative PM curvature at the point of
pillar contact. If CPZ is acting on the nanoscale as it does on the
microscale in red blood cells, then CPZ may reduce this negative
curvature and possibly result in it becoming locally positive (as shown
in Fig. 6). The change in curvature may contribute to the
reduction in force that is required to form a vesicle. The dramatic
decrease in the mean time for vesicle formation also supports the
possibility that the pillar interacts primarily with the inner leaflet
of the PM and that CPZ interferes with that interaction.

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Fig. 7.
Vesiculation process model. A: 1st step. The
PM breaks its tether with the CL. The pressure required for this
process most likely reflects the force of attachment between the PM and
the pillars. B: 2nd step. Only the PM is stretched and
elongating, with additional membrane flowing around the pillars into
the growing "tongue"; eventually, the PM pinches off to form a
vesicle.
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The second step of vesiculation must also involve consideration of the
tension in the lateral wall. In APPENDIX B, we calculate
the tension from the measured aspiration pressure (Eq. B3). This approach reveals that Sal and CPZ have opposing effects on the tension required to form a vesicle (see Fig.
4B). The differences between the drug treatments are
consistent with other observations showing that crenators increase and
cup formers (such as CPZ) decrease differential membrane tension
(45). Theoretical models (39) and
experimental observations (15, 16, 28) of membrane
vesiculation resulting from ionic amphipaths predict a pivotal role for
differential membrane tension. Drug-induced changes in membrane
mechanics may play a role in effects of amphipaths on electromotility.
Sal reduces OHC electromotility (15, 67), whereas CPZ has
no effect on the magnitude of electromotility but shifts its operating
voltage (38). In pure membranes, Sal disorders the lipid
component of the bilayer (23) and reduces membrane
mechanical strength (50). Sal also alters the
deformability of the red blood cell (10). Both Sal and CPZ
alter membrane fluidity in the OHC (43). CPZ and other
amphipathic cup-forming agents decrease, whereas crenating agents
increase, transmembrane currents in COS cells (45). The
ability of Sal to alter membrane properties such as curvature, tension,
and fluidity is in harmony with its ability to reversibly attenuate OHC
electromotility and produce a reversible hearing loss. The ability of
CPZ to shift the nonlinear capacitance may be due to a change in
differential area elasticity (Ref. 49; Raphael RM, Popel
AS, and Brownell WE, unpublished observation).
Sal and CPZ decrease strain-dependent hydraulic conductivity.
The OHC normally has low water permeability (11, 55, 56);
however, there is an increase in water permeability when the PM is
deformed (8). The observed loss of volume on aspiration and increase with the water jet are consistent with a strain-induced change in hydraulic conductivity. A possible mechanism for the increase
in water permeability is the formation of small pores in the stretched
membrane. Membranes naturally form defects that permit the movement of
lipids between the leaflets and movement of nonpermeable solutes
(54); the addition of mechanical tension could increase
the probability of defect formation. We showed previously
(8) that stretching the membrane leads to formation of
pores that admit mono- and disaccharides but not raffinose, implying a
pore size
4 nm. CPZ reduces the strain-induced increase in hydraulic
conductivity. This may be related to its ability to decrease membrane
tension (45) or antagonize the energy for pore formation,
which depends on differential membrane tension. The minor change in
radius of OHCs during aspiration of vesicles for control cells (Fig.
2A) reflects the loss of cytosol and associated reduction in
cell turgor because of the strain-induced increase in hydraulic
conductivity. The loss is reduced in the drug-treated cells.
Role of cytoskeleton in electromotility.
Electromotility depends on the connection between the cytoskeleton and
the PM. Our results indicate that CPZ loosens this connection when the
cell is aspirated under high pressures. The fact that CPZ does not
affect the magnitude of electromotility implies that CPZ does not have
a significant effect on the connectivity of the layers during normal
electromotile generated strains. The intact OHC is predisposed to
change length rather than diameter because the stiffness of the
cytoskeleton is greater circumferentially than it is longitudinally
(27). However, Sal- and CPZ-treated cells under aspiration
change their shape (both length and radius), which suggests a
contribution of the membranes to the elastic properties of the lateral
wall (59). However, treatment with Sal and CPZ alone does
not change the resting shape of OHCs. This implies that intercalation
of these amphiphiles into the membrane does not generate enough force
to overcome the cytoskeletal factors controlling cell shape. This also
implies that although Sal and CPZ may affect the association of the PM
and the CL, making them easier or harder to separate, these molecules
do not affect the integrity of the cytoskeleton itself. Although it is
possible that cytoskeletal remodeling occurs during the large
deformations in the pipette that also contribute to the forces, the
differential effect of Sal and CPZ are likely the result of effects on
the membrane.
In conclusion, we have shown that amphipathic drugs change the
mechanical properties of the OHC PM. Our results on the shape change
during aspiration, the tension required for vesiculation, and the time
to form a vesicle are rationalized in terms of differential tension.
The existence of excess PM acting as a membrane reservoir coupled with
changes in mechanical properties induced by amphipaths indicate that
membrane curvature may be intricately linked to the molecular mechanism
of electromotility.
 |
APPENDIX A |
Sources of Errors in Surface Area and Volume
Measurements
In measuring cell surface area and volume, we incur both random
errors (an individual measured value deviates from its expected value
according to some probability distribution) and systematic errors (the
expected value deviates from the true value). With this classification,
four sources of measurement errors can be identified: 1)
random errors due to errors incurred in tracing the cell/vesicle
outline; 2) systematic errors caused by cell bending;
3) systematic errors at the apical and basal ends of the
cell; and 4) systematic errors incurred if the cross section of the cell is not circular.
Error types 2-4 are all caused by discrepancies between the actual
shape and the cylindrical shape of the segments assumed by the
measurement algorithm. Given that vesicles are removed from the central
portion of the cell, type 3 errors are likely to be the same before and
after vesicle removal in a given cell. Type 3 errors are therefore
unlikely to affect before/after comparisons and can be ignored for the
purpose of this paper. The presence of type 4 errors has been addressed
in a previous publication (11) where the cross-sectional
profile was found to be circular by confocal microscopy. The
strain-dependent increase in hydraulic conductivity leads to volume
increases for both the aspiration and fluid jet experiments. Our
morphometric analysis revealed a systematic increase in radius (Fig.
2B). Microscope focus was adjusted during the experiment to
maintain the focal z-plane at the maximum cell diameter. If
an increase in cell diameter were the result of the OHC flattening out,
the focal plane would be closer to the glass substrate, which never
occurred. The observed elevation in the focal plane was consistent with
the OHC maintaining a circular profile.
Random (type 1) errors increase the number of observations necessary to
establish a significant difference between two groups. Large random
errors therefore limit our ability to resolve small differences but do
not introduce spurious differences between groups. Systematic errors
(type 2), on the other hand, have the potential to introduce such
spurious differences into the results and must be considered carefully.
Random errors.
With a Taylor series expansion, one can show that if the relative error
(standard deviation
expected value) of the measured radius
ri is
, the relative error in
Si and Vi will be
and 2
, respectively. However, because the total surface area and volume
are computed as the sum of n cylindrical segments, the error
in these parameters will be
/
and 2
/
, respectively. Thus, if n = 100, a 10% measurement error in radius will translate into 1% and 2% errors in total surface area and volume, respectively. Repeating the
measurement five times and taking the average of the results further
reduces the errors by a factor of
.
A similar argument can be made for the surface area and volume of the
cylindrical midportion of a vesicle. However, here, the length
x is also a measured parameter with a measurement error,
and the approximate errors will be 2
and 3
, respectively. Assuming that the midportion accounts for the majority of the vesicle
area and volume, we can use this as a rough estimate of the relative
error in these parameters. Thus, assuming a measurement error of 10%
for the length and radius, the measurement error in vesicle area and
volume may be as high as 20% and 30%, respectively. Again, repeating
the measurement five times and taking the average of the results
reduces the errors by a factor of
to ~9% and 13%,
respectively. The impact of this error is further reduced by summing
the vesicle surface area and volume in multiples of three vesicles and
comparing the sum with either the initial surface and area of the cell
or the total change (cell +
vesicles) up to that point.
Systematic errors due to bending.
From formulas for the surface area and volume of a circular torus
(47), we can compute the surface area and volume of a segment.
|
(A1)
|
and
|
(A2)
|
where the angle describing a segment of torus is
(this angle
is zero if the cell is not bent), rt1 is the
inner radius of the toroidal segment, rt2
is the radius to the center of the toroidal segment, and
rt3 is the outer radius. The surface area and
volume of the cylindrical approximation in Fig. 1 are
|
(A3)
|
and
|
(A4)
|
The error incurred by using the approximations can thus be
written as
|
(A5)
|
where sinc(x) = sin(x)/x
and a positive error corresponds to an overestimation of the area or
volume. Note that the value of the sinc function is one for a zero
angle, i.e., the error in the estimate approaches zero as the angle
approaches zero. If a "knee" in a cell with a large bending angle
is represented by n segments each having a bending angle
=
/n, the total error will be
|
(A6)
|
For example, a 45° (
/4 radian) bend approximated by five
segments would yield a total error of ~0.5%. This error is minor compared with the random errors, as can be appreciated by examination of Table 2, where measurements were made
from a drawing representing the projection of a model cell and the
calculated values compared with the true quantities represented by the
model.
A determination of measurement was obtained by subjecting a model image
of a cylindrical cell to the exact same analysis as the OHC images.
Model images were generated having bending angles of 0° (straight) or
buckled at 25° and 30°. The model images were traced 20 times with
the MATLAB program, and the error was calculated (Table 2). In our
experiments, some of the cells were bent up to 150°, but this was not
correlated with drug application. The error for the model image was
within 2% for length and radius and within 5% for surface area and
volume. Therefore, the magnitude of possible errors does not alter the
conclusions based on our measurements.
 |
APPENDIX B |
Theoretical Considerations for Calculation of Apparent
Tension
To evaluate the effect of ionic amphipaths on cell mechanical
properties, the aspiration pressure in the pipette must be related to
the tension. We adopt the approach used for micropipette-aspirated red
blood cells (18, 20). The lateral wall is treated as a continuum and is described mechanically by shell theory. Because the
thickness of the lateral wall is 100 nm and the radius of the cell is 4 µm, the lateral wall can adequately be treated as a thin shell so
long as it is realized that the calculated force resultants will be
effective parameters that are distributed between the membrane and the
cytoskeleton. Assuming that the transverse shear component does not
vary with position on the surface, in-plane force resultants (tensions)
can simply be related to hydrostatic pressure differences scaled by the
radius of curvature (the classic law of Laplace) (17, 20).
In the analysis of the micropipette experiment, assuming tension in the
pipette (
p) is isotropic, force balance in the pipette
predicts that
|
(B1)
|
where Pp equals aspiration pressure in the pipette
(in mN/m2), Pi equals the pressure inside the
cell, and Rp equals inner radius of the
micropipette (in µm). The pressure inside the cell is an
unknown. To eliminate it, we apply the law of Laplace to the portion of
the cell outside the micropipette. The outer portion of the cell is
anisotropic: the membrane tension in the longitudinal direction is
|
(B2)
|
where Po is the external pressure and
Rc equals the average radius of the cell.
Equating
p and
l and combining Eqs.
B1 and B2 gives an equation for estimating the tension
in the lateral wall
|
(B3)
|
where
P = Pp
Po.
This tension must be viewed as an "apparent" tension and must not
be interpreted as the tension residing in the plasma membrane. The OHC
has a complex trilamellar lateral wall and is best modeled as an
orthotropic cylindrical shell (72), meaning that there may
be a contribution from the circumferential tension. In applying Eq. B3, we have assumed that this apparent tension "turns
the corner" of the pipette and remains the same in the far field. In
reality, there is likely to be a tension gradient as the membrane
leaves the pipette that would affect the absolute magnitude of the
calculated tension. However, we only claim the validity of this
analysis up to the point at which the vesicle separates from the
membrane. After this event, the mechanical analysis becomes more
complicated. Nevertheless, our approach provides a framework for the
interpretation of experiments on drug effects (see Fig. 4B).
 |
ACKNOWLEDGEMENTS |
We thank A. S. Popel and J. S. Oghalai for helpful comments.
 |
FOOTNOTES |
This work supported by National Institute of Deafness and Other
Communications Disorders Grants DC-00354, DC-02775, and National Research Service Award DC-00363 and National Science Foundation Grant 9871994.
Present addresses: N. Morimoto, Dept. of Otolaryngology, National
Children's Hospital, 3-35-31 Taishido Setagaya, Tokyo 154-8509, Japan
(E-mail: nmorimoto{at}nch.go.jp); R. M. Raphael, Department of
Bioengineering, MS142, Rice University, Houston, TX 77251-1892 (E-mail:
rraphael{at}rice.edu); A. Nygren, Dept of Physiology and Biophysics,
University of Calgary, 3300 Hospital Dr NW, Calgary, AB, Canada T2N 4N1
(E-mail: nygren{at}ucalgary.ca).
Address for reprint requests and other correspondence:
W. E. Brownell, Dept. of Otorhinolaryngology and
Communicative Sci., NA 505, Baylor College of Medicine, Houston TX
77030 (E-mail: brownell{at}bcm.tmc.edu;
http://www.bcm.tmc.edu/oto/research/cochlea/index.html).
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
First published December 12, 2001;10.1152/ajpcell.00210.2001
Received 7 May 2001; accepted in final form 10 December 2001.
 |
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