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The Journal of Neuroscience, January 1, 1998, 18(1):48-58
Fluorescence-Imaged Microdeformation of the Outer Hair Cell Lateral Wall
John S. Oghalai1, Alpen A. Patel1, Takashi Nakagawa1, 2, and William E. Brownell1 1 Bobby R. Alford Department of Otorhinolaryngology and Communicative Sciences, Baylor College of Medicine, Houston, Texas 77030, and 2 Department of Otorhinolaryngology, Faculty of Medicine, Kyushu University, Fukuoka 812, Japan
ABSTRACT
Top
Abstract
Introduction
Outer hair cell (OHC) electromotility appears to be central to mammalian hearing and originates within its lateral wall. The OHC lateral wall is a unique trilaminate structure consisting of the plasma membrane (PM), the cortical lattice (CL), and the subsurface cisternae (SSC). We selectively labeled and imaged the lateral wall components in the isolated guinea pig OHC under confocal microscopy. The PM was labeled with a voltage-sensitive dye, di-8-ANEPPS; the SSC was labeled with the sphingomyelin precursor, NBD-C6-ceramide; and F-actin in the CL was labeled with conjugates of phalloidin. Interactions among the three layers were evaluated with the micropipette aspiration technique. The PM was tethered to the CL and SSC until, at a critical deformation pressure, the PM separated, allowing visualization of the extracisternal space, and ultimately formed a vesicle. After detaching, the stiffness parameter of the PM was 22% of that of the intact lateral wall. We conclude that the lateral wall PM is more compliant than the CL/SSC complex. The data clarify the structural basis for electromotile force coupling in the OHC lateral wall.
Key words: cochlea; inner ear; hearing; cytoskeleton; cell stiffness; stiffness parameter; micropipette aspiration; patch-clamp technique; confocal microscopy; fluorescence labeling; biophysics
INTRODUCTION
The cochlear outer hair cell (OHC) is necessary for the exquisite sensitivity of mammalian hearing because of its unique electromotile property that amplifies the cochlear traveling wave (Dallos and Corey, 1991
; Ruggero and Rich, 1991
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Figure 1. The outer hair cell (OHC) lateral wall. The OHC (lower left) is a cylindrical epithelial cell, with stereocilia at the apex and the nucleus at the base. The stereocilia are rooted in the cuticular plate (CP). The lateral wall is a trilaminate structure composed of the plasma membrane (PM), the cortical lattice (CL), and the subsurface cisternae (SSC). The PM has been visualized by electron microscopy and noted to have a high density of membrane particles. The CL is composed of microdomains of parallel actin filaments, which are cross-linked by spectrin. Although the actin orientation can vary between microdomains by >90°, the average orientation is nearly circumferential, with a mean angle to the transverse axis of 9-15° (Holley and Ashmore, 1988a
When isolated from the organ of Corti, the electromotile response demonstrates an increasing amount of displacement with increasing distance from the point of OHC fixation. This pattern of displacement has been postulated to originate from multiple, evenly distributed unit motors along the lateral wall (Holley and Ashmore, 1988b
How do motors based in the lateral wall direct their forces to change cell length? The nature of force coupling pathways among the three layers of the lateral wall is central to this question. We used the technique of fluorescence-imaged microdeformation (Discher et al., 1994
MATERIALS AND METHODS
OHC isolation. Albino guinea pigs of either sex weighing 200-300 gm and having a normal startle response to a handclap were decapitated. The temporal bones were taken and the middle ear bullae opened. The otic capsule was removed, and the spiral ligament was peeled off to expose the organ of Corti. The modiolus with the intact organ of Corti was removed from the temporal bone.
Two different experimental preparations were used. For OHCs that were labeled with di-8-ANEPPS, NBD-C6-ceramide, or rhodamine phalloidin, the respective staining protocols were performed in situ (described below). After staining, the organ of Corti was rinsed with extracellular solution consisting of (in mM): 135 NaCl, 4 KCl, 1 MgCl2, 2 CaCl2, 10 HEPES, 10 glucose, and 0.1% bovine serum albumin (Sigma, St. Louis, MO). The extracellular solution had an osmolality of 285-290 mOsm/kg. Mechanical trituration without enzyme was performed to dissociate the OHCs (Brownell et al., 1985
All OHCs were plated onto the glass bottom of an uncoated microwell Petri dish (MatTek, Ashland, MA). Isolated cells were selected for study on the basis of standard morphological criteria within 4 hr of animal death. Under the light microscope, healthy cells display a characteristic birefringence, a uniformly cylindrical shape without regional swelling, a basally located nucleus, and no Brownian motion of subcellular cytoplasmic particles (Shehata et al., 1991
Di-8-ANEPPS labeling. Di-8-ANEPPS is a fluorescent molecule with a nonpolar region that inserts into membranes and a polar region that is responsible for the fluorescence. This dye stains the PM of the OHC very well and does not internalize significantly (Fluhler et al., 1985
NBD-C6-ceramide labeling. NBD-C6-ceramide is a labeled precursor of sphingomyelin. In more conventional cells, it preferentially labels the Golgi apparatus (Lipsky and Pagano, 1985
20°C. An aliquot was diluted with extracellular solution to a final concentration of 10 nmol/ml of NBD-C6-ceramide and 0.34 mg/ml defatted bovine serum albumin. The organ of Corti was incubated in this solution at room temperature for 30 min and then rinsed with the standard extracellular solution.
Rhodamine phalloidin labeling. Phalloidins bond specifically to F-actin, and fluorescent conjugates have been shown to label the CL in the OHCs (Carlisle et al., 1988
Texas Red-X phalloidin labeling. To stain actin while maintaining the integrity of the PM (in contrast to the rhodamine phalloidin staining procedure), we also performed an intracellular dialysis technique, using a tight-seal whole-cell ruptured patch configuration. One vial (300 U) of Texas Red-X phalloidin (T-7471, Molecular Probes) was dissolved in 100 µl of methanol. Aliquots were dried and stored at
. Seals were formed at the base of the OHC by using an Axopatch 200B amplifier and pCLAMP6 software (Axon Instruments, Foster City, CA). The cells were held at their zero current potential; only cells with potentials negative to
Confocal microscopy. Data were gathered by a scanning confocal system (MRC-600, Bio-Rad, Hercules, CA) with a krypton/argon laser, configured on an Axiovert 35 microscope (Zeiss, Oberkochen, Germany). Images were obtained with a 100× oil immersion objective (numerical aperture 1.30; Plan-NEOFLUAR, Zeiss). Fluorescent images of cells labeled with di-8-ANEPPS and NBD-C6-ceramide were collected, using an excitation wavelength of 488 nm. Images of cells labeled with fluorescent phalloidins were collected at an excitation wavelength of 568 nm. Emission wavelengths were collected, using long-pass filters with corner wavelengths of 515 nm for the membrane dyes and 585 for the actin dyes. Images equivalent to those normally seen by light microscopy were collected with the transmitted light path of the system, using Nomarski optics. In Figure 5d, the fluorescent signal was so low that we had to open the confocal aperture all the way to visualize the dye staining. This greatly increased the z-plane thickness. In all other confocal images a more typical aperture setting was used, and the thickness of the z-plane sections was ~1.0 µm.
Micropipette aspiration and stiffness parameter calculations. The micropipette aspiration technique (Fig. 2) was used to produce a controlled deformation of the lateral wall of the OHC. Aspiration micropipettes with internal diameters of ~3 µm were fabricated from borosilicate glass capillary tubes (LG16; outer diameter, 1.65 mm, inner diameter, 1.1 mm; Dagan, Minneapolis, MN) with an automated glass pipette puller (BB-CH-PC, Switzerland) and microforge (Microforge De Fonbrune, Aloe Scientific, St. Louis, MO). The micropipette was connected to a water column via polyethylene tubing, and both were filled with extracellular solution. The micropipette was mounted on a joystick-controlled electronic micromanipulator (Zeiss) and advanced to the center of the field of view. The level of the water column was referenced to the microscope stage (i.e., the zero pressure point).
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Figure 2. The micropipette aspiration technique. This technique involves bringing a glass pipette with an internal diameter of ~3 µm in contact with the lateral wall of the cell. As negative pressure is applied, a tongue of lateral wall is pulled inside the pipette. The tongue length is measured at different aspiration pressures and used to determine the stiffness parameter (see Fig. 7). With increasing suction pressure, the individual components of the lateral wall can be visualized (see Fig. 8).
Once a healthy cell was located, the micropipette was brought into contact with the midlateral wall of the OHC. Negative pressure was applied in a stepwise manner as measured from the null point. This was performed by sequentially decreasing the pressure by 1 cm of H2O (1 cm of H2O = 0.09807 nN/µm2) and then waiting 30 sec. This was repeated until
The stiffness parameter (SP) is calculated from the tongue length versus aspiration pressure data (Evans, 1973
where
P is the change in aspiration pressure applied to the lateral wall through the micropipette,
RESULTS
Labeling of the plasma membrane
Figure 3 shows an OHC stained with di-8-ANEPPS under low-pressure deformation. The transmitted image (Fig. 3a) shows the lateral wall tongue being aspirated into the micropipette. The apex of the cell was out of the plane of focus. The fluorescent image (Fig. 3b) demonstrates labeling of the basolateral surfaces of the cell. Di-8-ANEPPS also stained the apex and stereocilia (data not shown). There was no staining of intracellular structures. The tongue lengths in both images are the same; note that the tip of the tongue has a rounded shape. With higher aspiration pressures, the tongue length increases until the most distal portion detaches from the cell, forming a vesicle. An example of a vesicle is demonstrated in Figure 3c. Its fluorescent image (Fig. 3d) demonstrates that the wall of the vesicle is labeled. Because di-8-ANEPPS is known to label the PM, the PM is a component of the vesicle (n = 10).
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Figure 3. Di-8-ANEPPS labeling of the PM. a, Transmitted image of an OHC undergoing micropipette aspiration (
Labeling of the subsurface cisternae
Figure 4 demonstrates an OHC that was labeled with NBD-C6-ceramide. The transmitted image (Fig. 4a) shows the tongue of lateral wall aspirated inside the micropipette. Two layers of the lateral wall are visible within the pipette: the outer layer is indicated PM, and the inner layer is indicated SSC. The fluorescent image (Fig. 4b) shows labeling of the lateral wall and of some intracellular structures, including the nuclear membrane. The length of the tongue in Figure 4b corresponds to that of the inner layer in Figure 4a. This figure demonstrates that this NBD-C6-ceramide staining procedure labels the SSC, but not the PM. This image also shows that the PM can be separated from the SSC. After separation an inner tongue could be seen only in the transmitted image in approximately one-half of the cells; however, by fluorescence labeling, an inner tongue was visible in every cell. The SSC tongue in the fluorescent image has a flat tip, as opposed to the round tip of the PM tongue in the transmitted image. Figure 4c is the transmitted image of a vesicle pulled from a NBD-C6-ceramide-stained cell. Because the simultaneous fluorescent image (Fig. 4d) does not show labeling of the vesicle, SSC membranes are not a component of the vesicle (n = 11).
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Figure 4. NBD-C6-ceramide labeling of the SSC. a, Transmitted image of an OHC undergoing micropipette aspiration (
Labeling of the cortical lattice
Figure 5, a and b, demonstrates a cell that was permeabilized with saponin and subsequently labeled for F-actin with rhodamine phalloidin. The transmitted image (Fig. 5a) shows the poor condition of the cell, which is to be expected after treatment with saponin. However, a tongue of the lateral wall could be aspirated into the pipette. The fluorescent image (Fig. 5b) shows a large amount of labeling at the apex as well as faint labeling along the lateral walls. This labeling pattern demonstrates the difference in the density of F-actin in the cuticular plate and stereocilia versus the CL. The labeled tongue is visible and is at the same location as the tip of the tongue seen in the transmitted image. Aspiration pressure was increased in these cells until they ruptured (typically
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Figure 5. Fluorescent phalloidin labeling of the CL. a, Transmitted image of an OHC after treatment with saponin and staining with rhodamine phalloidin (
Figure 5, c and d, demonstrates a cell that was labeled intracellularly for F-actin with Texas Red-X phalloidin. This technique did not require the use of saponin, so the integrity of the PM was preserved. Simultaneous aspiration and patch pipettes were used. First, aspiration of the lateral wall was performed to hold a tongue at constant pressure, usually approximately
Dual labeling of the plasma membrane and the subsurface cisternae
The cell in Figure 6 was stained with both di-8-ANEPPS and NBD-C6-ceramide. A series of fluorescent images demonstrates the sequence of separation and vesiculation. In Figure 6a, the PM and CL/SSC complex were indistinguishable from each other. The layers were bonded together, and both had a round tip. The lines radiating from the base of the tongue are folds in the cell membranes caused by the applied suction pressure. They disappeared as the manipulation progressed (Fig. 6b-f), perhaps because of an increase in cell turgor pressure. These folds were not visible in Figures 3, 4, 5. In Figure 6b, the PM separated from the CL/SSC complex. At this point the PM continued to have a round tip, whereas the tip of the SSC flattened out. In Figure 6c-e, the PM tongue continued to elongate, whereas the SSC tongue remained in nearly the same location. Finally, in Figure 6f, the PM tongue broke free to form a vesicle, leaving its base to reseal. This process can continue, leading to the release of multiple vesicles. For example, we have aspirated up to 26 vesicles from a single cell.
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Figure 6. Dual labeling of the PM and SSC. This figure captures the sequence of separation and vesiculation during micropipette aspiration of the OHC lateral wall. a, The PM and the SSC overlapped initially (
Stiffness parameters
Both PM and CL/SSC complex tongue lengths were measured from OHCs stained with di-8-ANEPPS and NBD-C6-ceramide as the aspiration pressure was increased. All data points from 11 cells are plotted (Fig. 7). The dotted and solid lines represent the PM and the CL/SSC complex, respectively, both before and after separation. The reciprocal of the slope is the SP of the structure. These lines are the average of the slopes and y-intercepts of lines fit to each of the data points of the 11 cells. The scatter apparent in the figure is attributable to variation in the y-intercept points (i.e., the starting tongue lengths) of different cells; however, the slope (1/SP) varied little (see data below). The inset is an example of the data recorded from one cell.
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Figure 7. Tongue length versus aspiration pressure. These data were taken from dual-labeled OHCs (n = 11). The circles represent PM tongue lengths, and the triangles represent SSC tongue lengths. The point of separation is represented by the large circle. The dotted and solid lines correspond to the PM and CL/SSC complex, respectively. The four lines were created by averaging the slopes and y-intercepts of linear fits to each of the 11 cells both before and after the point of separation. The corresponding stiffness parameters (SP) are the reciprocal of the slopes of each line and are defined in units of nN/µm. The inset is an example of data taken from one cell.
The PM and CL/SSC complex overlapped with low aspiration pressures. The average SP of the intact trilaminate wall was 0.79 ± 0.18 nN/µm (mean ± SD). The point of separation had an x value of 1.67 ± 0.30 nN/µm (mean ± SD), corresponding to a pressure of
On the other hand, the CL/SSC complex tongue length did not increase as rapidly with pressure increments after separation as it did before separation. Although the PM tongue maintained a round shape throughout the aspiration process, the flat tips of the CL and SSC tongues after separation (more clearly seen in Figs. 4b, 5d) imply that there was no pressure gradient across them. So, after separation, the negative pressure applied by the micropipette acted to pull directly on the PM alone. This means that the reciprocal of the slope of the line indicating the CL/SSC complex (Fig. 7) is not representative of its SP. Hence, subtraction of the SP of the PM from the SP of the trilaminate wall was performed to determine this value (0.79-0.17 = 0.62 nN/µm). So the PM is responsible for 22% of the SP (0.17/0.79) of the trilaminate wall, and the remaining two layers comprise the remaining 78% (0.62/0.79). The lower SP of the PM indicates that it is more compliant than the CL/SSC complex.
To verify that the dyes did not affect lateral wall mechanics, we compared SP and separation pressures of labeled cells versus controls. SP of unlabeled controls (n = 11), as well as OHCs stained with di-8-ANEPPS (n = 10) or NBD-C6-ceramide (n = 11), were not significantly different (p > 0.1) both before and after separation (data not shown). Additionally, the separation pressures were not significantly different (p > 0.1; data not shown).
DISCUSSION
Selective labeling and the extracisternal space
Previous reports have demonstrated fluorescent labeling of structures within the OHC lateral wall (Carlisle et al., 1988
With the application of negative pressure, a tongue of lateral wall was pulled into the micropipette. Initially, the trilaminate structure was bonded together. As the suction pressure was increased, the PM separated from the lateral wall, representing disruption of the tethering between the PM and the CL/SSC complex. The fluid compartment between the two layers is the extracisternal space. Not only were we able to determine that the vesicle is composed of PM and does not contain actin from the CL or membranes from the SSC, but we could further dissect their structural contribution to wall mechanics by quantitating their deformation.
Component contributions to lateral wall stiffness
Our findings can be compared with other experiments in which the stiffness of the lateral wall has been altered. Although the absolute values of stiffness measurements may vary, depending on the type of experimental deformation (Spector et al., 1997b
However, our observation that the lateral wall stiffness is dominated by the CL/SSC complex contrasts with the conclusions of Holley and Ashmore (1988a)
The role of the cytoskeleton and pillars
Figure 8 is a representative diagram of the separation process. Although the PM can be detached from the lateral wall, the CL and SSC remain behind and are primarily responsible for the SP of the lateral wall. The phenomenon of separation proves that the PM is tethered to the underlying lateral wall and that this tethering can be broken. SP of the OHC trilaminate lateral wall is anywhere from 2 to 76 times higher than in other cells (Sit et al., 1997
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Figure 8. The phenomenon of separation. Initially, the bonded trilaminate lateral wall is pulled into the pipette (as seen in Fig. 2). After separation (a), the tip of the CL/SSC complex flattens out, whereas the PM continues to elongate. b, The vesicle consists of only PM and does not contain actin from the CL or membranes from the SSC. The remaining PM reseals over the lateral wall and begins to elongate again. Although represented as remaining with the actin of the CL in this diagram, the actual fate of the pillars is unknown (see Discussion).
The pillars are probably responsible for this tethering. The maximal tethering force depends both on their longitudinal strength as well as on their anchoring strength into the PM and the CL (Spector et al., 1996
Implications for electromotile force coupling
One hypothesis to explain OHC force coupling is the cytoskeletal spring model proposed by Holley and Ashmore (1990)
Our findings do not support these hypotheses. Because the PM appears to be more compliant than the CL, it would seem that membrane-based motors should just push the bilayer phospholipids out of the way as they change conformation, rather than directing their force purposefully. This makes it unlikely that the motors exert a force when increasing in surface area. However, it is possible that they could still exert a force when decreasing in surface area, by pulling on the lipid bilayer. A contractile force within the PM probably could lead to changes in cell shape, because the PM can withstand the pulling force of micropipette aspiration to the point at which bending of the entire cell occurs (Figs. 3, 4; see also Sit et al., 1997
This extensile force might be passive resting tension from within the framework of the CL, associated with cell turgor pressure (Brownell et al., 1985
Interestingly, although the pillars clearly are tethered to the PM, the motors do not have to be fixed in place. This leaves the intriguing possibility that the motors could be mobile within the PM lipid bilayer (as in the fluid mosaic model), sensing transmembrane potential and changing in surface area correspondingly. Meanwhile, passive tension, acting to elongate the OHC, "fits" cell length to the available amount of PM surface area. There are many more particles in the PM (the postulated molecular motors) than pillars (Forge, 1991
FOOTNOTES Received July 23, 1997; revised Oct. 9, 1997; accepted Oct. 10, 1997.
This project was supported by research grants from the Deafness Research Foundation (to J.S.O.) and Grants DC00354 and DC02775 (to W.E.B.) from the National Institute on Deafness and Other Communication Disorders. We are grateful to Drs. B. R. Alford, A. Bullen, R. A. Eatock, H. A. Jenkins, S. Popel, R. Raphael, T. Ratnanather, P. Saggau, A. Spector, and M. Zhi. N. Gavia and C. Shope provided technical support. Illustrations were done by S. Carmichael.
Correspondence should be addressed to Dr. John S. Oghalai, Bobby R. Alford Department of Otorhinolaryngology and Communicative Sciences, Baylor College of Medicine, One Baylor Plaza, Houston, TX 77030.
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