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. 2020 Jun 17;119(2):314–325. doi: 10.1016/j.bpj.2020.06.011

Interactions between Passive and Active Vibrations in the Organ of Corti In Vitro

Talat Jabeen 1, Joseph C Holt 2,4,5, Jonathan R Becker 3, Jong-Hoon Nam 1,3,4,∗
PMCID: PMC7376138  PMID: 32579963

Abstract

High sensitivity and selectivity of hearing require an active cochlea. The cochlear sensory epithelium, the organ of Corti, vibrates because of external and internal excitations. The external stimulation is acoustic pressures mediated by the scala fluids, whereas the internal excitation is generated by a type of sensory receptor cells (the outer hair cells) in response to the acoustic vibrations. The outer hair cells are cellular actuators that are responsible for cochlear amplification. The organ of Corti is highly structured for transmitting vibrations originating from acoustic pressure and active outer hair cell force to the inner hair cells that synapse on afferent nerves. Understanding how the organ of Corti vibrates because of acoustic pressure and outer hair cell force is critical for explaining cochlear function. In this study, cochleae were freshly isolated from young gerbils. The organ of Corti in the excised cochlea was subjected to mechanical and electrical stimulation that are analogous to acoustic and cellular stimulation in the natural cochlea. Organ of Corti vibrations, including those of individual outer hair cells, were measured using optical coherence tomography. Respective vibration patterns due to mechanical and electrical stimulation were characterized. Interactions between the two vibration patterns were investigated by applying the two forms of stimulation simultaneously. Our results show that the interactions could be either constructive or destructive, which implies that the outer hair cells can either amplify or reduce vibrations in the organ of Corti. We discuss a potential consequence of the two interaction modes for cochlear frequency tuning.

Significance

The function of the mammalian cochlea is characterized by sharp tuning and a high level of amplification. Both tuning and amplification are achieved mechanically through the action of cellular actuators in the sensory epithelium. According to widely accepted theory, cochlear tuning is achieved by “selectively amplifying” acoustic vibrations. This study presents a set of data suggesting that the cochlear actuators can both amplify and reduce vibrations to enhance cochlear tuning.

Introduction

The cochlea encodes sounds into neural signals according to frequency and amplitude. As sound waves enter the cochlea through the middle ear bones, interscala pressure waves propagate from the base to the apex of the cochlea (1). The traveling waves peak at different locations such that low-frequency waves propagate farther toward the apex than high frequency waves. As a result, a specific location of the cochlea responds best to a specific frequency, called the tonotopy. Cochlear tonotopy is closely related to the mechanical gradient along the cochlear length, which is dominated by the basilar membrane (2,3). An auditory epithelium, called the organ of Corti (OoC), sits on the basilar membrane and is covered by the tectorial membrane to form the OoC complex. Relative motion (shear) between the OoC and the tectorial membrane results in activation/inhibition of the mechanoreceptive cells—the inner and the outer hair cells (4). The outer hair cells elongate or contract as their transmembrane potential changes (5). Vibrations of the OoC complex are then modulated by mechanical feedback from the outer hair cells so that small sounds at best frequency are amplified (6). As such, the outer hair cells contribute to the key functions of the cochlea—tuning and amplification of sounds.

OoC vibrations are more complicated than it was thought before the finding of outer hair cell electromotility. The classical understanding of OoC mechanotransduction is based on kinematics, treating the OoC as a rigid body (7, 8, 9). The classical model explains how the stereocilia are deflected by the relative motion between the OoC and the tectorial membrane. This mechanism of stereocilia deflection underlies the foundation of theoretical models (10, 11, 12, 13). The rigid-body approximation, however, is being relaxed as more experimental data are revealing complex relative motions among OoC fine structures. For example, the top and the bottom surfaces of the OoC (often represented by the reticular lamina and the basilar membrane) have been shown to vibrate with considerable phase difference. This relative motion among OoC structures is dependent on the level and frequency of stimulation (14,15). Outer hair cell electromotility affects the reticular lamina more than the basilar membrane (16, 17, 18, 19). Although there is relatively good agreement that outer hair cell electromotility is essential for cochlear amplification and tuning, how those functions are achieved remains controversial. For example, some argue that cochlear amplification is achieved by longitudinal accumulation of energy (17,20,21), whereas others consider that cochlear amplification is essentially local (19). Whereas the outer hair cells are considered to provide power for cochlear amplification, some researchers argue that the outer hair cells instead attenuate intense sounds (22).

The subject of this study is the modulation of OoC vibration patterns due to outer hair cell motility. We used excised cochlear tissues from young gerbils to investigate how OoC vibrations due to mechanical and electrical stimulation interact. We captured the deformation of individual outer hair cells using optical coherence tomography (OCT). Our results demonstrate that vibration patterns generated by the two stimuli can either constructively or destructively interact. The consequences of these interactions are discussed.

Materials and Methods

Tissue preparation

Cochleas harvested from young Mongolian gerbils (2–5 wk old) were used for experiments according to the institutional guidelines of the University Committee on Animal Resources at the University of Rochester. Gerbils were deeply anesthetized with isoflurane, after which the cochlea was acutely isolated and placed in a petri dish containing the following: 145 mM sodium gluconate, 7 mM NaCl, 3 mM KCl, 5 mM NaH2PO4, 0.1 mM MgCl2, 5 mM D-glucose, 0.1 mM CaCl2, and 5 mM HEPES. The acidity and the osmolarity of solutions used in this study were adjusted to pH 7.35 and 300 mOsm, respectively. In the petri dish, the cochlea was further reduced to a single cochlear turn. This study targeted 1-mm, mid-to-apical sections centered at 8 mm from the basal end (red colored section in Fig. 1 A). Basal and apical turns were removed using forceps and sharp blades. Bones between scalae were removed from both basal and apical sides to expose the cochlear epithelium from the section of interest (Fig. 1 B). Apical and basal openings of the remaining cochlear coil were sealed using cyanoacrylate glue (orange colored in Fig. 1, C and D). The reduced cochlear turn was then transferred to a microfluidic chamber initially containing the same perilymph-like solution. Cyanoacrylate sealant was applied along the circumference of the cochlear turn (Fig. 1 C). It took 60–80 min for tissue preparation.

Figure 1.

Figure 1

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Preparation of excised cochlear turn. (A) Excised gerbil cochlea. The apical and the basal turns are removed to expose the target location (red section). (B) Excised cochlear turn placed in microfluidic chamber. (C) Schematic of prepared tissue in the microfluidic chamber. Cochlear section was exaggerated for illustration. The excised cochlear turn is sealed with glue to separate two fluid spaces. The tissue is stimulated mechanically through a stimulating port (red arrow) and/or electrically through a pair of electrodes. Resulting vibrations are measured using optical coherence tomography. (D) An enlarged view of the organ of Corti (OoC) in the chamber from (C).

Microfluidic chamber

The microfluidic chamber was fabricated using a stereolithography printer (Moai; Peopoly, Los Angeles, CA). Mechanical stimulation was delivered by vibrating an opening covered with elastic membrane with a piezoelectric actuator (red arrow in Fig. 1 C). Another opening was provided for pressure release. There was a pair of inlet-outlet ports to refresh the solution in the chamber (data not shown). Electrical stimulation across the OoC was applied through a pair of electrodes (Fig. 1 C). After the cochlear tissue was attached and sealed to the chamber, bottom and top fluids were replaced with an artificial endolymph (145 mM KCl, 0.1 mM CaCl2, 4 mM HEDTA, 10 mM K-HEPES, 8 mM D-glucose, 2 mM sodium pyruvate) and a second perilymph (145 mM sodium gluconate, 7 mM NaCl, 3 mM KCl, 5 mM NaH2PO4, 0.1 mM MgCl2, 5 mM D-glucose, 1 mM CaCl2, 5 mM HEPES), respectively.

Stimulation and measurements

Transepithelial electrical stimulation was applied through a pair of Ag/AgCl electrodes. Alternating current (typically between 50 and 200 μA) was supplied by a current source (CS580; Stanford Research Systems, Sunnyvale, CA) or a custom-built current generator. The voltage across the tissue was monitored. Typical electrical impedance between two electrodes was 4–6 kΩ. Mechanical stimuli were delivered using a piezoelectric actuator (PC4WM; Thorlabs, Newton, NJ) driven by a high-voltage amplifier (E−505.00; Physik Instrumente, Sausalito, CA). The piezo stack actuator tip vibrated with an amplitude of 30 nm at 0.5 V of driving voltage. At 1 kHz, this stimulation level generated ∼1 Pa of pressure beneath the slit. To measure hydrodynamic pressure subjected to cochlear tissue, a pressure transducer was placed at a symmetric position to the tissue. Fabrication and calibration procedure of the pressure transducer is described in the Supporting Materials and Methods. Stimulus functions were generated using MATLAB code (The MathWorks, Natick, MA) that controls a data acquisition board (PCI-6353; National Instruments, Austin, TX). For vibration measurements, a commercial optical OCT imaging system (Ganymede; Thorlabs) was used. The system uses a light source with a 900-nm center wavelength, and its scanning unit has an A-scan rate of 100 kHz. The system was modified to use a 20× objective (numerical aperture 0.4; Mitutoyo, Kanagawa, Japan) to enhance optical resolution. Full-width at half-maximum resolution determined by imaging a 0.4-μm bead was 2.5 and 3.0 μm along the lateral and optical directions, respectively. The imaging system was driven with a MATLAB program to operate its M-scan mode for vibration measurements. For validation of OCT vibration measurements, a piezoelectric actuator was vibrated under the OCT system and then under a laser interferometry system (MMA 300 system; Polytec, Irvine, CA). Measurement results from the two vibrometry systems were compared. The difference between two measurement systems was less than 5% within 0.1–20 kHz frequency range. Between 0.3 and 3 kHz, the root mean-square noise of OCT vibration measurements was 0.5 nm. Most experiment protocols ran a set of measurements either at different scanning points (spatial sweep), at different frequencies (frequency sweep), at different phases between stimulations (phase sweep), etc. To avoid potential artifacts due to measurement sequence, the order of events was randomized.

Viability of tissue

Per the purpose of this study, the viability of prepared tissue is most explicitly represented with the level of outer hair cell electromotility. All presented data were obtained when the outer hair cell electromotility was greater than 10 nm per 100 μA and when the tectorial membrane was considered attached. Besides electromotility, two morphological features served as good indicators for the state of each preparation. First, the tectorial membrane attachment to the OoC was particularly vulnerable to surgical or chemical insults. Therefore, a stable attachment was a good indicator for a preparation’s viability. The optical resolution of 3 μm is sufficient to resolve the expected dimension of subtectorial space, 5–10 μm (23). There exist uncertainties regarding the tectorial membrane attachment. For instance, the tectorial membrane could be partly detached. The gap above the third-row outer hair cells often looked greater than that of the other two rows. Second, outer hair cell shape was sensitive to incomplete separation between the two fluid compartments. In these instances, OoC deformation due to the swelling outer hair cells was evident within 10 min of artificial endolymph application. We declared a proper seal when leakage between the two fluid spaces was <0.16 μL/min at slit pressures of 20 Pa, which was estimated by observing the level change of a 2-mm water column (Marnell et al., 2018). Leakage tended to be all or nothing (there was no case of modest leakage like 1 μL/min). When it leaked, it was massive, and potassium-rich endolymph damaged OoC cells in minutes. When these two conditions were positive at early stages, the electromotility stayed near or above 10 nm per 100 μA current for over 180 min post-cochlear isolation. All presented data were obtained when 1) the tectorial membrane was attached to the OoC, 2) leakage between two fluid spaces was negligible, and 3) outer hair cell electromotility was above 10 nm per 100 μA current.

Results

Fine OCT imaging helps to discern tissue status

Our preparation combined with OCT provided high resolution imaging of OoC microstructures. Imaging signal was stronger for the reticular lamina, outer hair cells, pillar cells, Hensen’s cells, basilar membrane fiber layers, and the bottom layer of tectorial membrane, whereas it was weaker in Deiters’ cells and the body of the tectorial membrane (Fig. 2 A). Micrometer-level anatomical characteristics could be effectively resolved, including the gaps between the three rows of outer hair cells, the subtectorial space, and nerve fibers passing through the tunnel of Corti. Such resolution enabled us to observe the deformation of individual outer hair cells throughout the experiment. These images were helpful in monitoring the status of preparation, especially when the tissue was deformed because of stress or damage. For example, as outer hair cells swelled, they shrank along their length and the reticular lamina deflected downward, whereas the Hensen’s cell outside the third-row outer hair cells bulged upward. As this deformation progressed, the tectorial membrane became detached from the OoC at its lateral extreme (Fig. 2 B). Micromechanics seemed local. Even if a radial section showed swollen outer hair cells with little electromotility, the outer hair cells in other sections of the same preparation could be in good shape with strong electromotility. In our preparation, the basilar membrane was tilted between 10 and 25° from the horizontal plane (Fig. 2 A). In this orientation, the outer hair cells were roughly aligned to the optical axis, which was helpful in measuring their length change (electromotility).

Figure 2.

Figure 2

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Status of tectorial membrane attachment. Vibrations of the tectorial membrane (TL), reticular lamina (RL), and Deiters’ cell apex (DC) due to mechanical stimulations were measured. The status of tectorial membrane attachment was assessed from the size of the sub-TM gap. (A) Tissue shape at the beginning of a preparation 65 min from animal death. (B) The same preparation as (A) 3 h later. The sub-TM gap was widened toward its lateral edge. (C and D) The relative motion of TM w.r.t. RL was measured at different stimulating frequencies. Thick curves and shaded spans represent mean and one standard deviation, respectively. (E) Vibration phase of TM w.r.t. RL versus sub-TM gap size. (F) Vibration phase of DC w.r.t RL versus sub-TM gap size. In (E) and (F), the phase value was obtained from the frequency range between 0.3 and 0.5 kHz. To see this figure in color, go online.

The tectorial membrane was considered attached to the OoC when the subtectorial gap size measured at the second-row outer hair cells is less than 8 μm. In this article, no data were used when the tectorial membrane was considered detached except those used for comparison in Fig. 2. The images in Fig. 2, A and B represent examples of intact and detached tectorial membrane. The image of Fig. 2 B was acquired 3 h past Fig. 2 A. In a series of experiments, relative motion between the tectorial membrane and the reticula lamina was measured. To observe the relationship between tectorial membrane status and mechanical response, from 11 preparations, 13 cases of frequency response to mechanical stimulation were analyzed. In 5 cases, the tectorial membrane was considered attached (symbol ●, in Fig. 2, E and F) and six were judged detached (Fig. 2 C). In two cases (asterisks), the tectorial membrane “reattached” to the OoC at the time of measurement—the subtectorial gap had been >8 μm for some time span, but the gap decreased to <8 μm at the time of measurement. When tectorial membrane was intact (attached), the vibration amplitude ratio between the tectorial membrane (TM) and the reticula lamina (RL) was ∼0.9 and nearly flat over measured frequency range (0.3 and 0.3 kHz, Fig. 2 C). With detached tectorial membrane, the ratio was smaller at low frequencies (0.6 at 0.3 kHz) but became similar to the intact case for frequency range >1 kHz. For both attached and detached cases, tectorial membrane vibrations led reticula lamina’s, but it was greater when detached (φTM-RL of 23 vs 13° at 0.3 kHz). The phase difference decreased toward zero as the frequency increased (Fig. 2 D). At frequencies <0.5 kHz, there was positive correlation between φTM-RL and the gap size (R2 = 0.57), but not at frequencies >1 kHz (R2 < 0.0001). The OoC vibrated in phase regardless of tectorial membrane status (Fig. 2 F). The phase difference between the reticula lamina and the Deiters’ cell apex remained close to zero (<5°) despite stimulating frequency and the subtectorial gap size. When detached tectorial membrane reattached to the OoC (asterisks, in Fig. 2, E and F), the relative vibrations were indistinguishable from intact cases. To summarize, the status of tectorial membrane attachment could be ascertained from subtectorial space. Detachment of the tectorial membrane modestly affected OoC mechanics, and the effect was greater at low frequencies.

Micromechanics without traveling waves

Acoustic energy propagates as a compressive wave in the air at the speed of 340 m/s in the air and 1500 m/s in the water. In the cochlea, acoustic energy propagates much more slowly at a group velocity of less than a few meters per second (22). Cochlear traveling wave is formed by interaction between the basilar membrane with graded stiffness and scalar fluids. Longitudinal coupling to form cochlear traveling waves is achieved primarily by the scalar fluids (i.e., many cochlear models reproduce reasonable traveling waves without structural longitudinal coupling). Although our preparation preserved structural mechanics (basilar membrane), it modified “scalar” fluid mechanics (more specifically, fluid boundary conditions represented by the scalar cavity). In the microchamber, applied fluid pressure is considered uniform over exposed cochlear section, which minimizes the effect of longitudinal coupling provided by the scalar fluid.

Traveling waves that characterize cochlear mechanics were not observed in our preparation (Fig. 3). When the stimulating port was vibrated at a constant amplitude (30 nm), the vibration amplitude peaked between 0.5 and 1 kHz (Fig 3 A). At the target location (x = 8 mm from the basal end), the best responding frequency of gerbil cochlea ranges between 1.5 kHz (when sensitive) and 1.1 kHz (when insensitive). Over the measured frequency range (0.3–3 kHz), the phase changed by less than a cycle (1.5π to 2.0 π rad, Fig 3 B). The pattern of pressure was similar to displacement but had greater amplitude at frequencies >1 kHz and smaller phase change along frequency (Fig 3, C and D). When the displacement is referenced to the pressure (Fig 3, E and F), the response curves looked more like a simple resonator with minimal overshoot. To conclude, this preparation minimized the longitudinal coupling by scalar fluid mechanics. The observations made in this study should be considered as OoC micro (local) mechanics. The characteristic frequency of this preparation may be determined by the apical (low-frequency) end of the section rather than the middle of the section because in multi-degree-of-freedom systems, low-frequency components dominate.

Figure 3.

Figure 3

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Frequency response to mechanical stimulation. The stimulating port was vibrated at constant amplitude (30 nm) at different frequencies between 0.3 and 3 kHz. Three sets of measurements from different preparations were distinguished by different symbols. (A and B) Vibration amplitude and phase measured at the reticular lamina. (C and D) Pressure amplitude and phase measured by the pressure transducer beneath the slit. (E and F) Displacement amplitude and phase with respect to pressure.

Two vibration patterns

Vibration measurements were made from regions 7–9 mm from the basal end of the gerbil cochlea, which corresponds to the frequency range of 1–3 kHz. Note that our excised preparation compromised natural mechanical and electrical boundary conditions, including the pressure delivery along the length of the scalae. Therefore, instead of examining the relationship between input stimulation and tissue vibration, we analyzed the relative motion between microstructures within the OoC. Similar to other studies (14,17,24), the reticular lamina and the basilar membrane were chosen for analysis because they represent the top and bottom of the OoC and because optical signals were stronger at these locations. We elected to use a calcium level of 0.1 mM in the upper chamber (scala media space), which is higher than reported physiological levels. This choice was a compromise between two considerations: to decrease the mechanotransduction current with higher than the physiological calcium level (25) so that we could better separate the mechanical and electrical responses and to maintain the shape of the tectorial membrane with a calcium level not too far from physiological conditions (26).

Vibration patterns due to mechanical and electrical stimulations were distinctly different (Fig. 4). When mechanically stimulated, the reticular lamina and the basilar membrane vibrated in phase, and the vibrations of the reticular lamina were smaller than those of the basilar membrane (Fig. 4 B). This trend regarding the relative motion between the top and bottom of the OoC was highly consistent across different stimulating frequencies (symbol ● in Fig. 4, D and E). The ratio in vibration amplitude between the top and bottom of the OoC (between “RL” and “BM” in Fig. 4 A) was between 0.5 and 1. Our observations of passive mechanics are in line with the classical kinematic model (7) in that the ratio between the top and bottom of the OoC is ∼1. Our measurements also agree with reported ratios between the reticular lamina and the basilar membrane made in other in vivo observations (14,15,24). When electrically stimulated, however, the reticular lamina vibrated at a different phase from the basilar membrane (>60°, Fig. 4 E). The relative displacement between the top and bottom of the OoC depended on stimulating frequency (square in Fig. 4, D and E).

Figure 4.

Figure 4

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OoC vibrations due to mechanical and electrical stimulation. (A) B-scan image of the OoC. Red vertical dashed line represents the optical axis where vibrations were measured. The horizontal distance between the curve from the vertical line (green curve) indicates signal strength along the optical depth. Two void circles indicate the measurement points presented in (B)–(E). (B) Vibrations of the reticular lamina (blue) and the basilar membrane (red) due to mechanical stimulation. (C) Vibrations due to electrical stimulation (100 μA at 1 kHz). (D and E) Relative motion (amplitude and phase) of the reticular lamina with respect to the basilar membrane at different stimulating frequencies. The measurement location was 9 mm from the basal end. The cross symbols (×) indicate data points with poor signal at which signal/noise ratio <8 dB. To see this figure in color, go online.

Spatial patterns of OoC vibration were also different in response to the two types of stimulation (Fig. 5). These plots were obtained from 50 to 40 scan lines across the radial span of the OoC for mechanical and electrical stimulation cases, respectively, corresponding to a spacing of 3.0 and 3.6 μm between scan lines. For each scan, a sinusoidal stimulation (20-ms ramp time and 100 cycles at plateau) was applied, and the resulting vibrations were measured. Data pixels with signal/noise ratios >8 dB are represented with colored dots. Mechanical vibration patterns show that the vibration amplitude is greater in the middle of the OoC and decreases toward the lateral edges (Fig. 5 A). This pattern reflects the anatomy where ligaments hold the medial and lateral ends of the basilar membrane. The peak displacement (the brightest spot in the amplitude plot) occurred near the joint between arcuate and pectinate zones of the basilar membrane and at the center of the OoC near the outer hair cells, in agreement with previous observations (16,27). When the focal plane was at the level of the outer hair cells, the tectorial membrane did not have strong optical signals, but occasionally its top or bottom surface produced signals. We did not observe a phase difference between the tectorial membrane and the reticular lamina.

Figure 5.

Figure 5

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Two vibration patterns in the OoC. The amplitude and phase measurements of 50 and 40 A-scan lines are shown on top of corresponding B-scan image. The top and bottom row panels show amplitude and phase of vibrations, respectively. (A) Amplitude and phase of vibrations due to mechanical stimulation. The stimulating port was vibrated at 1 kHz with a 30-nm amplitude. (B) Amplitude and phase of vibrations due to electrical stimulation. Stimulating current was applied at 2 kHz with a 100-μA amplitude. The top panels show the red curves, which indicate vibrating shapes along the broken lines. Color bar units are nanometers. The bottom panels show the asterisk symbols, which indicate the extremities of three outer hair cells. The color rings at the left bottom corner indicate the color scale of phase angle, e.g., between the dark blue and yellow spots, there is a phase difference of 180°. Scale bars, 25 μm.

Image resolution was fine enough to resolve the motility of individual outer hair cells. For the case shown in Fig. 5 B, the phase difference between the top and bottom of the three rows of outer hair cells (asterisk symbols) were 162, 149, and 60°. With a current amplitude of 100 μA, the amplitude of electromotility was 28, 22, and 8 nm for the first-, second-, and third-row cells, respectively. This trend of greater motility for the first- and second-row cells versus the third-row cell was observed in majority of cases (Fig. 6, A and B). Despite such observations, we cannot exclude potential experimental artifacts such as faster deterioration of the third-row outer hair cells.

Figure 6.

Figure 6

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Electromotility of individual outer hair cells. Electromotility of outer hair cells was measured over time and frequency. Time zero is defined as the onset of experimental dissection. (A) Motility of individual cells from eight cochleae are presented. The lines indicate the trend of exponential decay for the three rows of outer hair cells distinguished by different symbols and colors (see legends). (B) Mean motility of the three rows of outer hair cells over the first hour of measurement (<2.25 h). The error bar indicates standard deviation (n = 19, 16, 7 for OHC1, 2, and 3, respectively). (C) Motility from different cochleae are distinguished by different symbols. (D) Sodium salicylate decreased outer hair cell electromotility. Shaded vertical columns indicate the span of sodium salicylate application. The different symbols indicate independent trials. (E and F) Amplitude and phase of motility w.r.t electrical stimulation at different stimulating frequencies. Data are from three different cochleae. To see this figure in color, go online.

Outer hair cell electromotility

The electromotility of outer hair cells was defined as a length change in the cell body per transepithelial current. The electrical resistance between endolymphatic and perilymphatic electrodes was ∼5 kΩ. The first measurements of individual preparations were made typically 1.2 h after cochlear isolation. The time-course of outer hair cell electromotility decay varied across preparations (Fig. 6 A). This variation is ascribed to different factors, including the extent of surgical insults, chemical conditions, different measurement locations, and level of stimulation. In most cases, motility was greater than 100 nm/mA for 3 h or longer. Larger and longer electrical stimulations tended to exhaust the preparation quicker, whereas modest level of mechanical stimulation (peak displacement at the reticular lamina <100 nm) did not appear to affect tissue conditions. The motility of individual hair cells is presented in Fig. 6 A. When analyzed for early hour groups (within 1 h from measurement onset), the third-row outer hair cells had smaller motility compared to the other rows (Fig. 6 B). This difference may be attributed in part to the trend for quicker swelling of the third-row outer hair cells. Considering the trend of motility decay over time (lines in Fig. 6 A), the electromotility of outer hair cells from our measurement location (8 mm from the basal end of the gerbil cochlea) is expected to be greater than 1000 nm/mA. In Fig. 6 C, each data point represents the average motility of measurable outer hair cells within observed radial section (mostly outer hair cell row 1 or 2). Sodium salicylate, a blocker of outer hair cell electromotility (28), reduced the motility (Fig. 6 D). In a case with 15 min of exposure to 10 mM sodium salicylate (square), the motility decreased quickly (>90% drop within an hour) and did not recover. In another trial (symbol ●), the tissue sample was subjected to sodium salicylate at three 2-min exposures with 30-min intervals. After the first application, the motility reduced by 80% and then recovered to the half of the pretreatment motility. In subsequent applications, however, both inhibition and recovery were smaller. Outer hair cell electromotility decreased by less than an order of magnitude as stimulating frequency increased from 200 to 3200 Hz (Fig. 6, E and F). The phase of motility, defined as the phase of elongation with respect to top-to-bottom chamber current, decreased approximately by 90° over the frequency range, from 180 to 90°. That is, as the frequency increases, the motility changed from conductive to capacitive, which is in line with the Evans-Dallos model (29). Because we have not investigated other cochlear locations, it is unclear whether this trend is true regardless of location.

Our measurements are consistent with previously reported observations. Karavitaki and Mountain (21,30) reported radial displacements of 400 and 2000 nm/mA at 120 Hz from the apical and middle turn locations of the gerbil cochlea. Note that we present outer hair cell elongation instead of reticular lamina displacement. The measurements in Zha et al.’s (31) are analogous to our measurements in that they estimated outer hair cell length changes from the differential motion between reticular lamina and basilar membrane. In their study, outer hair cells in basal locations (characteristic frequency (CF) = 20 kHz) of the gerbil cochlea deformed up to a few nanometers. Regarding the sodium salicylate treatment, a previous study showed 50% of reduction in outer hair cell motility after 5 min application of 10 mM sodium salicylate, and the motility was fully recovered within minutes (32).

Vibrations due to simultaneous stimulation

Mechanical and electrical stimulation were applied simultaneously to observe how vibrations due to different sources interact (Fig. 7). A modest level of stimulation was applied to minimize quick exhaustion of the tissue. The chosen response level (10–30 nm vibration amplitude at the reticular lamina) is within the physiological range (e.g., (33)), and well above the noise floor of our measurements. Three different phases between the mechanical and electrical stimulation were tested (φME = 0, 90, and 180°). Reflecting the phase variation in the horizontal direction of the electrically stimulated case, the spatial pattern of vibration amplitude was affected by φME. Among the three measured cases (φME = 0, 90, and 180°, Fig. 7), when φME = 0, the reticular lamina vibrated least. When φME = 180°, the reticular lamina vibrated greatest. Notably, the lateral wall of the OoC (the rectangles in Fig. 7 A) vibrated differently from the rest of OoC.

Figure 7.

Figure 7

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OoC vibration patterns due to simultaneous stimulations. Mechanical and electrical stimulations (1 kHz sinusoids) were applied simultaneously, but with different phases between the two stimulations (φME) so that electrical stimulation lags the mechanical stimulation by (A) 0°, (B) 180°, and (C) 90°. The upper and the lower row panels represent the vibrations in amplitude and phase, respectively. The scale bars shown only in the right column as the same scales were use in the other columns.

The interaction between vibrations due to different stimulating methods was investigated further. The OoC vibrations were measured at 12 different phases between mechanical and electrical stimulations (φME = 0, 30, 60, …, 360°) for nine different frequencies between 0.3 and 1.5 kHz (Fig. 8). To decide on the level of the two stimulations, frequency responses due to mechanical or electrical stimulation alone were measured (columns A and B). When the stimulating port was vibrated with the same amplitude (mechanical stimulations), the OoC vibration peaked at 0.8 kHz, and the phase between the stimulation and the tissue vibration changed by 0.5–1 cycle over the test frequency range. For electrical stimulation, the reticular lamina response was relatively flat over the tested frequency range, whereas the basilar membrane response decayed as the frequency increased. Then, mechanical and electrical stimulations were applied simultaneously. When one stimulation overwhelms the other, it is not ideal for observing interactions between the two vibration patterns. Therefore, we adjusted the levels of mechanical stimulation so that the reticula laminar vibrated 10 nm across the frequency range. When this equalized mechanical stimulation and the electrical stimulation were applied simultaneously, a vibration amplitude of ∼20 nm was expected (10 nm from each stimulations).

Figure 8.

Figure 8

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Constructive and destructive interactions between two vibrations. Measurements were made at two points of the OoC: the reticular lamina (RL) and the basilar membrane (BM). (A) Frequency response to mechanical stimulations. (B) Frequency response to electrical stimulations. (C) The top of the plot displays simultaneous stimulations. Electrical stimulation lags mechanical stimulation by φME. The bottom of the plot shows, depending on φME, responses due to the two stimulations add up (constructive) or cancel each other (destructive). (D) Normalized vibration amplitude at the RL (C) and BM (σquare) for different φME-values. Results at three frequencies are shown. The constructive φME is defined at the peak of fitted sinusoidal curve (arrows). (E) Constructive φME versus stimulating frequency for the reticular lamina and the basilar membrane. The filled and void symbols represent two sets of data obtained from different locations of a preparation.

There existed a specific value of φME at which the resulting response because of the simultaneous stimulations was constructive or destructive (add up, or cancel each other, Fig. 8 C). As expected from the summation of two sinusoidal curves, the constructive φME is 180° apart from the destructive φME. The vibration amplitudes were obtained at different φME-values, whereas other conditions including measurement point, stimulating amplitudes, and frequency remained the same. The data points of response amplitude versus φME were fitted with a sinusoidal curve from which the constructive φME was obtained (Fig. 8 D). The constructive φME varies over stimulating frequency (Fig. 8 E). As was implicated in Figs. 5 B and 7, the interaction was not uniform across the OoC. For the case presented in Fig. 8, the reticular lamina and basilar membrane had similar constructive φME near 1.2 kHz, or both the top and bottom surfaces of the OoC will constructively respond when stimulated with φME ≈ −150° at 1.2 kHz. On the contrary, at 0.3 kHz, constructive φME of the reticular lamina and the basilar membrane are ∼180° apart. That is, at 0.3 kHz, when one side of the OoC is under constructive interaction, the other side of the OoC will be under destructive interaction.

Different modes of interaction between mechanically and electrically evoked OoC vibrations can affect frequency selection at a cochlear location. An example of frequency selection through constructive and destructive interaction is demonstrated in Fig. 9. Two φME-values were chosen that were 180° apart (−120 and 60°). Because of interactions between the two stimulations, the vibration amplitude of reticular lamina varied between 10 and 30 nm depending on frequency (top panel of Fig. 9 A). Near 0.8 kHz, the constructive and destructive interaction contrasted most prominently. The difference between the two curves (φME = −120 and 60°) is reminiscent of a tuning curve (bottom panel of Fig. 9 A). Consistent with the difference between the reticular lamina and the basilar membrane responses in Fig. 8 B, the frequency responses measured at the two locations were different. That is, a condition that amplifies the reticular vibration could reduce the basilar membrane vibrations.

Figure 9.

Figure 9

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Synthesized frequency response. (A) Response of the reticular laminar to simultaneous stimulation. (B) Response of the basilar membrane to simultaneous stimulation. Mechanical stimulation level was adjusted to have a flat vibration amplitude of 10 nm at the reticular lamina. To see this figure in color, go online.

Discussion

Active motility of outer hair cells is required for both sensitivity and selectivity of hearing. Apparently, uniform amplification over frequency (Fig. 10 C) will not enhance frequency tuning. Therefore, the prevailing view on cochlear tuning is that the active feedback from outer hair cells “selectively” amplifies vibrations at the best frequency, specific to a location (34,35). The mechanism of this selective amplification has been a central theme in hearing research. In theory, an actuator in a vibrating system could be used for either amplification or reduction, if the timing of actuation varies depending on frequency. The outer hair cells can operate more proactively for frequency tuning by using both amplification and reduction depending on frequency (Fig. 10 D).

Figure 10.

Figure 10

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Consequence of destructive interaction. (A) Passive vibrations. The OoC vibrates in phase. The relative motion between the tectorial membrane and the reticular lamina results in hair bundle deflection, which activates mechnotransduction. (B) Active vibrations. Because of the change in transmembrane potential, the outer hair cells elongate or contract. When the outer hair cells are motile, fine structures of the OoC vibrate out of phase. (C) When active and passive vibrations interact constructively regardless of frequency, the responses are amplified, but tuning quality remains similar. (D) Tuning can be enhanced if destructive interaction occurs away from the CF. To see this figure in color, go online.

Two characteristics of cochlear physics make cochlear physics remarkable and the same time complicated—outer hair cells’ electromechanical feedback and its longitudinal coupling through traveling waves. To investigate micromechanical aspects of OoC, this study reduced the two characteristics. First, the feedback loop of cochlear mechanics due to outer hair cell motility was snapped. Because the stria vascularis did not function, there was no source for endocochlear potential. As we put endolymph into the top chamber that was similar to intracellular fluid, there was little electrical potential to drive outer hair cell motility through mechanotransduction current. This reduction removed another characteristic of cochlear mechanics—nonlinear amplification. Outer hair cell motility was not a result of hair bundle deflection (mechanotransduction current). That is, outer hair cell motility is independent of the OoC vibrations. Instead, outer hair cell motility was induced by a transepithelial current source. Second, there were few signs of traveling waves in our preparation. In intact cochlea, acoustic pressures propagate along the cochlear scale that look like a bundle of long slender tubes. In our preparation, pressure is delivered through a wide opening beneath a piece of sensory epithelium. As a consequence, our cochlear tissue does not generate obvious traveling waves (Fig. 3). Because traveling waves and active feedback were reduced in this preparation, the focus of this study was on OoC micromechanics. Taking advantage of these reductions, we could isolate the passive and active vibration modes of OoC micromechanics and investigate their interactions.

In this study, different timings of outer hair cell actuation were tested to examine the idea of destructive interaction between two vibration patterns (Figs. 6 and 7). Before testing the interactions, we characterized the two vibration patterns independently to determine how the OoC vibrates when it is mechanically or electrically stimulated. When mechanically stimulated (passive, Fig. 10 A), the vibration pattern was in line with the classical understanding of OoC kinematics. That is, the entire OoC vibrated in phase and the maximal displacement occurred near the location of the outer hair cells and Deiters’ cells (Fig. 5 A). Relative motion between the top and the bottom surface of OoC remained the same over the tested frequency range (Fig. 4). In contrast, when electrically stimulated (active, Fig. 10 B), OoC microstructures vibrated out of phase (Fig. 5 B). Our observed electrical responses compare better with previous measurements with sensitive (active) cochleae that showed the phase difference between the top and the bottom sides of the OoC (14,17,24). In addition to phase variation along the depth of the OoC, our results show apparent phase variation along the radial direction (Fig. 5 B). We demonstrated that, depending on the timing (phase) of actuations, there can be either constructive or destructive interactions between the two vibration modes (Fig. 9). If the actuation timing varies over the frequency, there can be constructive or destructive interactions depending on stimulating frequency. Such constructive or destructive interaction could enhance cochlear tuning (Fig. 10 D). We showed an example of destructive interaction under controlled conditions. To better observe the interaction between vibrations due to different stimulation types, stimulation levels were chosen so that two types of stimulation resulted in similar vibration amplitude. This condition may represent the situations when there is minimal amplification. That is, the results in Fig. 9 may be better compared with the responses in the tail region of traveling waves in natural cochlea.

The destructive interaction may occur under physiological conditions, considering that the timing of outer hair cell actuation varies over frequency. For example, the phase between the reticular lamina and the basilar membrane varies over frequency (14,15,36). Ren and his colleagues have discussed the idea of constructive interaction based on the phase difference between the reticular lamina and the basilar membrane (17). Our measurements (Fig. 7) are in line with their view—“in-phase vibrations of the reticular lamina and basilar membrane result in constructive interference” (17). Dong and Olson (37) showed that the phase between the extracellular potential near the outer hair cells with respect to the basilar membrane motion varies over frequency exceeding 90°. It was argued that the phase shift over 90° is an indication of selective amplification (38). Our argument of tuning shares similar reasoning as Olson and her colleagues in that outer hair cell motility away from the best responding location does not amplify.

New experimental attempts, to our knowledge, were made in this study to better observe OoC mechanics. It may be worthy to discuss the opportunities and challenges of our approach. First, the microchamber was designed to stimulate isolated cochlear tissue both acoustically and electrically. Previous studies using microchambers were referred to, including Chan and Hudspeth (39), and Karavitaki and Mountain (21,30). Besides adopting the merits of those studies (endolymph-perilymph separation and calibrated electrical and mechanical stimulation), simultaneous application of mechanical and electrical stimulation was newly attempted in this study. This approach was instrumental in isolating and synthesizing different vibration modes within the OoC. Second, we measured individual outer hair cell deformation in situ using OCT. Experiments with isolated outer hair cells have been providing biophysical information such as the electromotile properties of the outer hair cell membrane (40, 41, 42, 43). The OoC vibrations due to outer hair cell motility have been providing insights on how outer hair cells contribute to cochlear amplification (32,39,44). To our knowledge, the change in the length of individual outer hair cells in situ has not been measured before. This was achieved by combining OCT imaging with our microchamber preparation. Compared to the laser interferometry that has long been the gold standard in cochlear vibration measurements, OCT was shown to be efficient in scanning vibrations over a cross section or a volume of cochlear tissue (18,24). We achieved an enhanced spatial resolution as compared to existing studies thanks to the removal of bones along the optical path and incorporation of a higher numerical aperture objective (numerical aperture of 0.4).

There are aspects that have not been fully exploited regarding our experimental approach. First, we did not take advantage of the electrochemical separation across the OoC. Although we have tested both perilymph-perilymph and endolymph-perilymph conditions, we are not ready to discuss the effects of different chemical conditions, primarily because of the lack of acquired data. The change from perilymph to endolymph in the top fluid space was usually done within 30 min after aggressive surgical procedures including the removal of cochlear bones and the cleaving of the stria vascularis. Therefore, it was difficult to tell whether any change in the early state was due to the tissue’s settling from surgical procedures or whether it was because of the replacement of the top fluid. Besides the effect of endolymph-perilymph, we could have performed different chemical assays such as different calcium levels or the application of mechanotransduction channel blockers, but they have not been examined yet. In this study, we dedicated a 2- to 3-h experimental window to explore OoC mechanics by measuring OoC vibration patterns due to different types of stimulation. The other aspect that will be helpful but was not implemented in this work is vibration measurement in different directions. For example, the vibrations were measured along one optical axis. We chose the direction approximately parallel to the length of the outer hair cells. As a result, the basilar membrane was tilted ∼30° from the optical plane. Had we measured at different orientations of the preparation, we could have distinguished radial and transverse vibrations, as was done by Lee et al. (36). The radial and longitudinal motions at the base of the outer hair cell were shown to be significant when the OoC was electrically stimulated (21,30). This study presents the displacement along the axis of the outer hair cell, which is closer to the transverse rather than radial axis.

Author Contributions

T.J., J.C.H., and J.-H.N. conceived the study. T.J., J.R.B., and J.-H.N. performed experiments. J.-H.N. drafted the manuscript and the figures. T.J., J.-H.N., and J.C.H. edited the manuscript. J.R.B. calibrated experimental devices including the OCT system.

Acknowledgments

The authors thank Dr. Sebastian Schaefer in Thorlabs for guiding us to customize the OCT system.

This work was supported by National Institutes of Health R01 DC014685.

Editor: Steven Rosenfeld.

Footnotes

Supporting Material can be found online at https://doi.org/10.1016/j.bpj.2020.06.011.

Supporting Material

Document S1. Supporting Materials and Methods and Figs. S1 and S2
mmc1.pdf (405.4KB, pdf)
Document S2. Article plus Supporting Material
mmc2.pdf (3.5MB, pdf)

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Document S1. Supporting Materials and Methods and Figs. S1 and S2
mmc1.pdf (405.4KB, pdf)
Document S2. Article plus Supporting Material
mmc2.pdf (3.5MB, pdf)

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