Cochlear outer hair cell horizontal top connectors mediate mature stereocilia bundle mechanics

Abstract

Outer hair cell (OHC) stereocilia bundle deflection opens mechanoelectrical transduction channels at the tips of the stereocilia from the middle and short rows, while bundle cohesion is maintained owing to the presence of horizontal top connectors. Here, we used a quantitative noncontact atomic force microscopy method to investigate stereocilia bundle stiffness and damping, when stimulated at acoustic frequencies and nanometer distances from the bundle. Stereocilia bundle mechanics were determined in stereocilin-deficient mice lacking top connectors and with detached tectorial membrane (Strc ^-/-/Tecta ^-/- double knockout) and heterozygous littermate controls (Strc ^+/-/Tecta ^-/-). A substantial decrease in bundle stiffness and damping by ~60 and ~74% on postnatal days P13 to P15 was observed when top connectors were absent. Additionally, we followed bundle mechanics during OHC top connectors development between P9 and P15 and quantified the observed increase in OHC bundle stiffness and damping in Strc ^+/-/Tecta ^-/- mice while no significant change was detected in Strc ^-/-/Tecta ^-/- animals.

Fig. 1. Noncontact acoustic FM-AFM for quantitative and physiological measurement of stereociliary bundle mechanical properties.

(A) The schematic illustrates the organ of Corti subdivided into three regions: base, middle, and apex. Sensory hair cells (three OHC rows and one IHC row) are illustrated in green. The highlighted apical turn region is the location where all experiments were performed. (B) The diagram illustrates the noncontact acoustic FM-AFM method to measure the stereociliary hair bundle stiffness of OHCs and IHCs. It is based on the phenomenon that, when an acoustically vibrating AFM cantilever with an attached micrometer-sized bead in a liquid environment approaches within nanometer distances from the sample surface, a hydrodynamic coupling interaction will cause a frequency shift (Δf) to the cantilever oscillation that can be easily measured with the sensitive AFM detection (laser diode and photodetector) system. The photodetector records the displacement of the AFM cantilever with nanometer sensitivity. In the diagram, A is the oscillation amplitude, f is the cantilever/bead drive frequency, R is the microbead radius, h_m is the minimum gap height, and β is the angle between the reticular lamina and the basilar membrane.

Fig. 2. Methodology validation by determining the spring constant of AFM tipless microcantilevers with known value.

(A) Phase-contrast image showing the AFM cantilever with a 10-μm sphere (white dashed circle) and a tipless AFM cantilever with known stiffness. Scale bar, 45 μm. (B) Phase-frequency curves depicting frequency shifts acquired by keeping a constant π/2 phase (dotted line) when the acoustically vibrating cantilever with microsphere is moved from 1 μm to 50 nm from the tipless cantilever. (C) Determined AFM tipless cantilever spring constant using standard thermal fluctuations and novel noncontact FM-AFM methods. Data are presented as means ± SEM; NS indicates nonsignificant differences in comparison with controls, P = 0.14 (by unpaired two-tailed Student’s t test with Welch’s correction); numbers of measurements pooled were five tune curves for the thermal method and eight phase-frequency curves for the noncontact FM-AFM.

Fig. 3. Visualization of viable cochlear apical turn hair cells during noncontact FM-AFM measurements.

(A and B) Phase-contrast images showing the localization of the AFM cantilever with a 10-μm sphere (white dashed circle) over the OHCs. (A) Representative confocal image of the apical turn cells labeled with Calcein-AM confirming tissue viability. Scale bar, 45 μm. (B) Representative confocal image of the apical turn sensory epithelium transducing OHCs and IHCs labeled with 5 μM FM1-43, confirming hair cells’ proper MET channel functionality. Note that, with the 3-min incubation time used, cytoplasmic labeling of the sensory hair cells indicates that FM1-43 enters the cells mostly through their MET channels and not significantly through endocytosis (33). Scale bar, 25 μm. Images (A and B) were collected at the apical turn from different Strc^+/−/Tecta^−/− mice at P14.

Fig. 4. Deletion of stereocilin markedly reduces the OHC stereociliary bundle stiffness, whereas IHC bundle stiffness is unaffected.

(A and B) Phase-frequency curves obtained for P14 Strc^+/−/Tecta^−/− (A) or Strc^−/−/Tecta^−/− (B) OHCs from the apical turn of the cochlea. Acquisitions were done with a constant π/2 phase (dotted line) when the acoustically vibrating cantilever with a 10-μm microsphere was moved from 1 μm to within 50 nm from the top of the tall stereocilia row. (C and D) In the absence of stereocilin, the OHC hair bundle stiffness is markedly reduced (C), but the IHC bundle stiffness is unaffected (D). Data are presented as means ± SEM; *** indicates significant differences in comparison with heterozygous littermate controls, P < 0.05 (by unpaired two-tailed Student’s t test with Welch’s correction); NS indicates nonsignificant differences in comparison with controls, P > 0.05 (by unpaired two-tailed Student’s t test with Welch’s correction); the indicated number of cells used was from seven animals for Strc^+/−/Tecta^−/− (n = 40) and seven animals for Strc^−/−/Tecta^−/− (n = 46) OHCs and from five animals for Strc^+/−/Tecta^−/− (n = 14) and four animals for Strc^−/−/Tecta^−/− (n = 13) IHCs.

Fig. 5. OHC and IHC hair bundle viscous damping parameter.

(A) Changes in the slope of the phase-frequency curve acquired on an apical turn OHC stereocilia bundle from P14 Strc^+/−/Tecta^−/− (blue circles) and Strc^−/−/Tecta^−/− (red squares) mice. When the acoustically oscillating 10-μm sphere is moved closer to the sensory hair bundles, changes in dΦ/df measured at f_π/2 are negligible. (B and C) In the absence of stereocilin, the damping parameter is significantly reduced in OHCs (B) but unaffected in IHCs (C). Note that cochlear IHC bundles have elevated fluid-like behavior compared to OHCs, since their hydrodynamic frictional drag is reduced compared to OHCs. Data are presented as means ± SEM; *** indicates significant differences in comparison with heterozygous littermate controls, P < 0.05 (by unpaired two-tailed Student’s t test with Welch’s correction); NS indicates nonsignificant differences in comparison with controls, P > 0.05 (by unpaired two-tailed Student’s t test with Welch’s correction); the indicated number of cells used was from seven animals for Strc^+/−/Tecta^−/− (n = 40) and seven animals for Strc^−/−/Tecta^−/− (n = 46) OHCs and from five animals for Strc^+/−/Tecta^−/− (n = 14) and four animals for Strc^−/−/Tecta^−/− (n = 13) IHCs.

Fig. 6. Developmental changes in apical turn OHC stereociliary bundle stiffness and damping from P9 to P15.

Apical turn OHC bundle stiffness (A) and bundle damping (B) in control Strc^+/−/Tecta^−/− mice and Strc^−/−/Tecta^−/− mice lacking horizontal top connectors. At P9 and P10, the bundle stiffness values show no significant differences, whereas the viscous damping shows significant difference at P10. From P12 onward, bundle stiffness and damping markedly increase in Strc^+/−/Tecta^−/− OHCs. Data are presented as means ± SEM; *** indicates significant differences, P < 0.05 (by unpaired two-tailed Student’s t test with Welch’s correction); NS indicates nonsignificant differences, P > 0.05 (by unpaired two-tailed Student’s t test with Welch’s correction); number of measurements pooled per postnatal day (animals, cells) for Strc^+/−/Tecta^−/− OHCs [P9 (2, 4), P10 (1, 4), P11 (1, 8), P12 (1, 8), P14 (4, 32), and P15 (2, 8)] and Strc^−/−/Tecta^−/− OHCs [P9 (1, 11), P10 (2, 16), P12 (1, 6), P13 (1, 6), P14 (3, 17), and P15 (3, 23)].