Age-related changes in the biophysical and morphological characteristics of mouse cochlear outer hair cells

Ethics Statement

All animal work was performed at the University of Sheffield (UK) and licensed by the Home Office under the Animals (Scientific Procedures) Act 1986. Procedures and were approved by the University of Sheffield Ethical Review Committee. Auditory functional measurements were conducted under anesthesia using ketamine (100 mg/kg body weight, Fort Dodge Animal Health) and xylazine (10 mg/kg, Rompun 2%, Bayer), which were administered with intraperitoneal injection as previously described (Ingham et al. 2011). At the end of the procedure, mice were either culled by schedule 1 or recovered from anesthesia with intraperitoneal injection of atipamezole (1 mg/kg). Mice under recovery from anaesthesia were returned to their cage, placed on a thermal mat, and monitored for 2 to 4 hr. Cages were returned to the holding racks once the mice were able to move well and respond to external stimuli. Wild-type C57BL/6NTac, C57BL/6J and C3H/HeJ, and the transgenic C57BL/6NTac^Cdh23+ mice (Mianné et al. 2016) were used for the experiments.

Tissue preparation

In vitro recordings were performed from apical-coil OHCs (~9-12 kHz: see Results) of acutely dissected organs of Corti dissected from mice of 1-18 months of age. Males and females were killed by cervical dislocation and the inner ear placed in a Petri dish containing ice-cold extracellular solution composed of (in mM): 135 NaCl, 5.8 KCl, 1.3 CaCl₂, 0.9 MgCl₂, 0.7 NaH₂PO₄, 5.6 D-glucose, 10 HEPES-NaOH. Sodium pyruvate (2 mM), amino acids and vitamins were added from concentrates (Thermo Fisher Scientific, UK). The pH was adjusted to 7.5 (osmolality ~308 mmol kg^-1). The dissection procedure begins with the removal of the muscles and connective tissue surrounding the inner ear. The “cleaned” inner ear is then moved to a new Petri dish containing fresh solution (i.e. free from debris). By holding the whole inner ear from the vestibular system with a pair of forceps, the bone covering the part of the apical-coil cochlear turn (around the 8-20 kHz: see Ceriani et al. 2019) is removed using a second pair of forceps (Dumont #3 or #4, which are further polished using a sharpening stone). This initial step is aimed at exposing the middle bony-spiral structure containing the cochlear nerve, which is then gently cut with very fine forceps, a procedure that allows the removal of the entire bone covering the apical cochlear coil. When the apical coil is exposed, it is very important to keep the solution clean (i.e. adding fresh ice-cold solution) and avoiding to touch the delicate organ of Corti, which is where the hair cells are located. This is then followed by the removal of the bone covering the site of the apical-coil of the organ or Corti in one single step. The final step requires the use of very fine forceps to cut the auditory nerve and gently remove the apical portion of the organ of Corti, which is then placed in the recoding chamber using a small stainless steel spoon (about 4 mm in diameter, custom made). The tip size of the forceps used for this procedure changes depending on the thickness of the bone, which changes with age and between strains; usually we have 3-4 sets of forceps to cover the full range of thicknesses. The same procedure was applied to all dissections performed in this study irrespective of the age of the animal. In the recoding chamber, the isolated organ of Corti was immobilized using a nylon mesh fixed to a stainless steel ring and viewed using an upright microscope (Olympus BX51, Japan; Leica, DMLFS, Germany; Nikon, Germany). Hair cells were observed with Nomarski Differential Interface Contrast (DIC) optics (x63 or x60 water immersion objective) and x15 eyepieces.

Single-cell electrophysiology

For whole-cell basolateral K⁺ current recordings, experiments were performed as previously described (Corns et al. 2018; Ceriani et al. 2019). The pipette intracellular solution contained (in mM): 131 KCl, 3 MgCl₂, 1 EGTA-KOH, 5 Na₂ATP, 5 HEPES-KOH, 10 Na-phosphocreatine (pH was adjusted with 1M KCl to 7.28; osmolality was 294 mmol kg⁻¹). Membrane currents and voltage responses were investigated at room temperature (20-24 °C) using the Optopatch amplifier (Cairn Research Ltd, UK). Data acquisition was controlled by pClamp software using Digidata 1440A boards (Molecular Devices, USA). Recordings were low-pass filtered at 2.5 kHz (8-pole Bessel), sampled at 5 kHz and stored on data-drives for off-line analysis (Origin: OriginLab, USA). Membrane potentials in whole-cell recordings were corrected for the residual series resistance Rs after compensation (usually 70-90%) and the liquid junction potential (LJP) of –4 mV measured between electrode and bath solutions.

The presence of electromotile activity in OHCs was estimated by measuring nonlinear (voltage-dependent) capacitance using whole-cell patch clamp recordings. In order to block most of the ion channels in hair cells, the pipette intracellular solution contained (in mM): 125 CsCl, 3 MgCl₂, 1 EGTA-CsOH, 5 Na₂ATP, 5 HEPES-CsOH, 5 tetraethylammonium (TEA), 5 4-amynopyridine (4-AP) (pH was adjusted with CsOH to 7.28; osmolality was 280 mmol kg^-1). Real-time changes in nonlinear membrane capacitance (C _N-L) were investigated using the capacitance track-in-mode of the Optopatch amplifier (Cairn Research Ltd, UK) during the application of a 4 kHz sine wave of 13 mV RMS. From the holding potential of –84 mV, hair cells were subjected to a voltage ramp from –154 mV to +96 mV over 2s. The capacitance signal from the Optopatch amplifier was filtered at 250 Hz and sampled at 5 kHz.

Mechanoelectrical transduction (MET) currents in young adult and aged mice were recorded from OHCs Tecta^EGFP/EGFP; Tectb^-/-double knockout mice (Tecta/Tectb^-/-), in which the tectorial membrane is detached from the OHC hair bundles. MET currents were elicited using a fluid jet from a pipette driven by a 25 mm diameter piezoelectric disc (Corns et al. 2014; Corns et al. 2018). The fluid jet pipette tip had a diameter of 8-10 μm and was positioned at ~8 μm from the hair bundles to elicit a maximal MET current. Mechanical stimuli were applied as steps or 50 Hz sinusoids. The resting MET current becomes evident when the MET channel shut off in the inhibitory phase of the stimulus: the holding current minus the current present during inhibitory bundle deflection (Corns et al. 2014).

Auditory brainstem responses

Auditory brainstem responses (ABRs) were recorded from male and female mice between 1 and 18 months of age. Recordings were performed in a soundproof chamber (MAC-3 Acoustic Chamber, IAC Acoustic, UK) as previously described (Ingham et al. 2011). Briefly, stimuli were delivered to the ear by calibrated loudspeakers (MF1-S, Multi Field Speaker, Tucker-Davis Technologies, USA) placed 10 cm from the animal’s pinna. Sound pressure was calibrated with a low-noise microphone probe system (ER10B+, Etymotic, USA). Experiments were performed using BioSigRZ software (Ingham et al. 2011) driving an RZ6 auditory processor (Tucker-Davis Technologies). Response thresholds were estimated from the resulting ABR waveforms and defined as the lowest sound level where any recognisable feature of the waveform was visible. Each set of ABR data was independently assessed by at least 3 members of the laboratory without prior knowledge of the mouse strain, age or sex. Responses were measured for click and stimulus pure tones of frequencies between 3 and 42 kHz. Stimulus sound pressure levels were typically 0-95 dB SPL, presented in steps of 5 dB. The brainstem response signal was averaged over 256 repetitions. Tone bursts were 5 ms in duration with a 1 ms on/off ramp time, and were presented at a rate of 42.6/sec.

Distortion product otoacoustic emissions

OHC function was assessed in vivo by measuring the distortion product otoacoustic emissions (DPOAEs). Recordings were performed in a soundproof chamber (MAC-3 Acoustic Chamber, IAC Acoustic, UK). DPOAEs were recorded at 2f1-f2 in response to primary tones f1 and f2, where f2/f1 = 1.2. The f2 level (L2) was set from 20 to 80 dB with 10 dB increments, and the f1 level (L1) was set equal to L2. Frequency pairs of tones between f2=6.5 kHz and f2=39.2 kHz were presented directly into the ear canal by means of a metal coupler connected to two calibrated loudspeakers (MF1-S, Multi Field Speaker, Tucker-Davis Technologies, USA). The emission signals were recorded by a low-noise microphone (ER10B+: Etymotic Research Inc, USA) connected to the coupler. Experiments were performed using BioSigRZ software driving an RZ6 auditory processor (Tucker-Davis Technologies). The DPOAE thresholds were defined by the minimal sound level, where the DPOAEs were above the standard deviation of the noise. The determined DPOAE thresholds were plotted against geometric mean frequency of f1 and f2. Stimulus sound pressure levels were typically 0-80 dB SPL, presented in steps of 10 dB. The response signal was averaged over 500 repetitions.

Immunofluorescence microscopy

Dissected inner ears from mice were fixed with 4% paraformaldehyde in phosphate-buffered saline (PBS, pH 7.4) for 20 minutes at room temperature. Cochleae were microdissected, rinsed three times for 10 minutes in PBS, and incubated for 1 hour at room temperature in PBS supplemented with 5% normal goat or horse serum and 0.5% Triton X-100. The samples were then incubated overnight at 37°C with the primary antibody in PBS supplemented with 1% of serum. Primary antibodies were: mouse anti-myosin 7a (1:1000, Developmental Studies Hybridoma Bank, #138-1C), rabbit anti-myosin 7a (1:200, Proteus Biosciences, #25-6790), rabbit anti-prestin (1:5000, kindly provided by Robert Fettiplace), rabbit anti-SK2 (1:500, Sigma-Aldrich, P0483) goat anti-choline acetyltransferase (ChAT, 1:500, Millipore, AB144P), and mouse anti-CtBP2 (1:200, Biosciences, #612044). All primary antibodies were labelled with species appropriate Alexa Fluor or NorthernLights secondary antibody for 1 hour at 37°C. Samples were then mounted in VECTASHIELD. The z-stack images were captured with a Nikon A1 confocal microscope equipped with Nikon CFI Plan Apo 60X Oil objective, which is part of the Light Microscope Facility at the University of Sheffield. Image stacks were processed with Fiji Image Analysis software. The number of synaptic ribbon (CtBP2) and SK2 puncta was estimated from the z-stack images of the immunolabelled proteins using Fiji ImageJ. For the SK2 puncta, the threshold for staining identification was manually set on a per-sample basis at a level at which puncta of the expected size, shape and intercellular location were clearly distinct from the background. Within each mouse strain, 1 month and 15 month old mice were fixed and processed for imaging simultaneously, while 6 month old mice were done separately. Images were taken within a few days of fixing the samples. To identify the presence of colocalized SK2 and ChAT puncta we used the blob finder function within Arivis Vision4D 3.1 software on Z-projected images. Briefly, images were processed with a Gaussian filter, followed by an automatic seed finding and a watershed algorithm. A threshold at < 1 μm was set to define the juxtaposition of SK2 and ChAT puncta and the results were confirmed by visual inspection. Each set of experiments was performed in at least 3 mice.

RNA isolation, cDNA synthesis, RT-qPCR and gene sequencing

Cochleae were dissected from mice at 1 and 15 months of age and frozen until processing. Total RNA was isolated using the RNAqueous-4PCR Total RNA Isolation Kit (Invitrogen, AM1914) and RNA quantity and quality were verified with the Nanodrop. Equal amounts of cDNA were synthesised using the High-Capacity RNA-to-cDNA Kit (Applied Biosystem, 4387406). Following DNase I treatment, quantitative real-time PCRs were performed using QuantStudio 12K Flex Real-Time PCR System and PowerUp SYBR Green Master Mix (Applied Biosystem, A25776). Every sample was run in triplicate in 386-well plate (StarLab, B2323-3840), with inclusion of appropriate no-RT and no-template controls. The following gene-specific primers were used: Slc26a5 (5’-CTCTTTGAGTCGTTACCCCAG-3’ and 5’-GTAATCAGTCCGTAGTCCAAGC-3’); Ocm (5’-TGGAGATGGGAAGATTGGG-3’ and 5’-GGACTTGGTAAATTAGTTCGGG-3’); Cdh23 (5’-GCACCCACATTCCACAAC-3’ and 5’-AGTCTAGCTCCTGAATCACG-3’), Pcdh15 (5’-GTTTCTGCCTTGTGTTCTTGTG-3’ and 5’TGTTTCGGTCCTGATCAATGG-3’). Expression levels were normalised using the housekeeping gene Hprt (5’-GCTTGCTGGTGAAAAGGAC-3’ and 5’-AGATTCAACTTGCGCTCATC-3’) and analysed using the 2 -ΔΔCT method.

In order to investigate whether Tecta/Tectb double knockout mice (Tecta/Tectb^-/-) harbour the Cdh23^ahl allele (Johnson et al. 1997; Noben-Trauth et al. 2003), we performed DNA sequencing. Genomic DNA from animals was extracted using NaOH. The targeted region was PCR amplified using GoTaq Master Mixes (M7123, Promega) and primers 5’-TCAGGCTCCCCTGCTTCTAT-3’ and 5’-CCAAGCTGTTCCTGGTAGCT-3’. PCR products were purified and sequenced by Sanger sequencing using the DNA sequencing service from Core Genomic Facility, University of Sheffield.

Scanning electron microscopy (SEM)

For SEM, cochleae were excised from aged C57BL/6N mice (15 months). They were fixed by intralabyrinthine perfusion using a fine hypodermic needle through the round window with 2.5% vol/vol glutaraldehyde in 0.1M sodium cacodylate buffer containing 2 mM calcium chloride (pH 7.4) and then immersed in this fixative for 2 hrs. They were stored in fixative diluted 1/10^th in buffer and subsequently dissected by removing the bone from the apical coil to expose the organ of Corti and then immersed in 1% osmium tetroxide in the cacodylate buffer for 1 h. For osmium impregnation, which avoids gold coating, cochleae were incubated in solutions of saturated aqueous thiocarbohydrazide (20 mins) alternating with 1% osmium tetroxide in buffer (2 h) twice (the OTOTO technique (Furness & Hackney, 1986). They were dehydrated through an ethanol series and critical point dried using CO2 as the transitional fluid, then mounted on specimen stubs using conductive silver paint (Agar Scientific, Stansted, UK) and examined in a Hitachi S4500 field emission SEM operated at 5 kV accelerating voltage.

Toluidine blue staining

After glutaraldehyde fixation, cochleae were washed 3 times in 0.1 M sodium cacodylate buffer pH 7.2 and post-fixed in 1% osmium tetroxide in 0.1 M sodium cacodylate buffer for 3 h at room temperature. Samples were then washed 3 times in sodium cacodylate buffer and decalcified in 0.5 M EDTA pH 8.0 containing 0.1% glutaraldehyde for 3 days at 4°C. Samples were then washed briefly in water, dehydrated through an ascending ethanol series, equilibrated in propylene oxide and embedded in epoxy (TAAB 812) resin. Blocks were cured at 60°C for 24 h and trimmed with a glass knife after which semi-thin 1 micron thick sections were cut on a Reichert Ultracut E ultramicrotome using a histo-grade Diatome diamond knife. Sections were dried onto glass slides and stained briefly with Toluidine blue before viewing on a Zeiss Axioplan 2 wide-field microscope. Images were captured with a Spot RT digital camera.

Statistical analysis

Statistical comparisons of means were made by Student’s two-tailed t-test or for multiple comparisons, analysis of variance (one-way and two-way ANOVA followed by Tukey’s or Sidak’s post-test). P < 0.05 was selected as the criterion for statistical significance. Only mean values with a similar variance between groups were compared. Mean values are quoted in text and figures as means ± S.D. For DPOAE experiments, due to the presence of “not found” values (i.e. above the upper threshold limit of our equipment), the non-parametric aligned ranks transformation 2-way ANOVA (Mann-Whitney U-test for pairwise comparisons with Bonferroni adjusted P values) statistical test was used; data are quoted as median, and first and third quartiles.

Age-related changes in hearing function

We initially determined the hearing loss in the four mouse strains, which were kept under the same environmental conditions, in order to provide a more comprehensive understanding of the progression of the pathology. Auditory brainstem responses (ABRs), which indicate the activity of the afferent auditory pathway downstream of IHCs, were made using the same experimental set-up. At 12-14 months of age, good hearing C3H mice had ABR thresholds for click and pure-tone evoked ABRs (Fig. 1A,B) comparable to those published previously (Trune et al. 1996). ABRs threshold to clicks were significantly different among strains (P < 0.0001, one-way ANOVA). Compared to C3H mice, aged-matched 6J and 6N strains showed significantly increased ABR thresholds for clicks (6J, P < 0.001; 6N, P <0.05: Tukey’s post-test, one-way ANOVA: Fig. 1A). 6N-Repaired mice had ABR click stimulus thresholds comparable to C3H mice (P > 0.05), but significantly lower values than those measured in the co-isogenic 6N strain (P < 0.01) and 6J mice (P < 0.001: Tukey’s post-test, one-way ANOVA) (Fig. 1A).ABR thresholds for pure-tones were not significantly different between C3H and 6N-Repaired (P = 0.140), or between 6N and 6J (P = 0.979, Tukey’s post-test, two-way ANOVA). However, they were increased in the early-onset hearing loss (6N and 6J) compared to C3H and 6N-Repaired (P < 0.0001, Tukey’s post-test for all four strain combinations, two-way ANOVA) (Fig. 1B). All four mouse strains showed similar ABR thresholds only at 3 kHz (P = 0.3168) and 6 kHz (P = 0.1376: Tukey’s post-test, one-way ANOVA).

In order to investigate the specific contribution of OHCs to these hearing phenotypes, we performed distortion product otoacoustic emissions (DPOAEs) (Fig. 2 and 3), which are readouts of cochlear amplification caused by the displacement of OHC stereociliary bundles during sound-induced motion of the cochlear partition. We initially compared the DPOAE thresholds between males and females at 13-14 months of age, and we found no significant differences in all four strains (C3H, P = 0.3853; 6N, P = 0.5136; 6N-Repaired, P = 0.1988; 6J, P = 0.1956, aligned ranks transformation two -way ANOVA). Because no statistical significance was detected between strain-matched males and females, data from males and females were pooled together to investigate possible inter-strain differences in DPOAE thresholds (Fig. 2). We found no significant difference between C3H and 6N-Repaired mice (P = 0.5737) or between 6N and 6J mice (P = 0.1800, aligned ranks transformation 2-way ANOVA). However, DPOAE thresholds were significantly raised in both 6N and 6J mice compared to those recorded from C3H and 6N-Repaired mice (P < 0.0001 for all four combinations, aligned ranks transformation two-way ANOVA). These results also indicate that OHC defects are partially responsible for the different ABR thresholds between early-onset hearing loss and good hearing mice (Fig. 1). We then used male mice to investigate age-related changes in DPOAE thresholds (6-24 kHz range: Fig. 3A-F). As expected, the C3H and 6N-Repaired mice showed no significant change in DPOAE thresholds between 1 and 17 months of age (C3H, P = 0.4791, Fig. 3A, D; 6N-Repaired, P = 0.8290, Fig. 3B, E), while early-onset hearing loss mice exhibited highly increased DPOAE thresholds (6N, P < 0.0001, Fig. 3C, F, aligned ranks transformation 2-way ANOVA). At the 12 kHz region, which was used for the in vitro measurements described below, we found that DPOAE values were significantly different between strains (P < 0.0001, aligned ranks transformation two -way ANOVA). At 14-17 months, DPOAE thresholds were highly elevated in 6N compared to both C3H and 6N-Repaired strains (P = 0.0108; P = 0.0002, respectively), but not significantly different between C3H and 6N-Repaired mice (P = 0.5808, post-test from aligned ranks transformation two -way ANOVA: see Methods). These findings are in agreement with the extensive literature describing the progressive changes in DPOAE threshold in early-onset hearing loss and good hearing mice (e.g. Spongr et al. 1997; Jimenez et al. 1999; Zheng et al. 1999).

The biophysical properties of OHCs in ageing mice

In order to investigate possible changes in the biophysical properties of OHCs in ageing mice, we performed electrophysiological recordings from apical (~9-12 kHz) OHCs of 12-13 month old male and female mice. Mature mouse OHCs express a negatively activating K⁺ current called I _K,n, which is carried by KCNQ4 channels (Kcnq4 gene: Kubisch et al. 1999) and represents the major K⁺ current present in these cells (Marcotti & Kros, 1999). Using whole-cell patch clamp, we recorded the K⁺ currents from OHCs by applying a series of depolarizing voltage steps in 10 mV increments from –124 mV (holding potential was –84 mV). We found that a large I _K,n was present in the OHCs of females and males from all four mouse strains (Fig. 4A-Dshows female OHCs). The peak current-voltage (I-V) relationship of the total K⁺ current (I _K: Fig. 4E) in OHCs was not significantly different in females and males of all four mouse strains (C3H: P = 0.6558; 6N-Repaired: P = 0.3563; 6N: P = 0.7571; 6J: P = 0.2578, two-way ANOVA). The size of the total K⁺ current (Fig. 4F), which was measured at 0 mV for consistency with previously published work in adult OHCs (Marcotti & Kros, 1999), was similar across all four mouse strains and sex (P = 0.9218, two-way ANOVA). We then investigated two crucial biophysical properties of OHCs, their membrane capacitance (Cm), which provides an indication of the cell surface area, and their resting membrane potential (Vm). Both were comparable between strains and sex (Cm: P = 0.1990, Fig. 4G; Vm: P = 0.4705, Fig. 4H, two-way ANOVA). The resting V _m of OHCs was measured under whole-cell current clamp conditions and using 1.3 mM extracellular Ca²⁺, which also includes the effect of the very small resting MET current. MET channel blockers were not used because they normally affect the basolateral membrane conductances (e.g. Kenyon et al. 2017).

Following this initial characterization, we sought to investigate whether the biophysical characteristics of OHCs changed with age. Considering that we did not find any specific sex differences in aged mice in terms of hearing ability (Fig. 2) or OHC biophysics (Fig. 4), the following experiments are reported by combining the data from males and females. The size of IK decreased significantly with age in each mouse strain (C3H: P = 0.0096; 6N-Repaired: P = 0.0018; 6N: P = 0.0292; 6J: P = 0.0007, one-way ANOVA: Fig. 5A-C), but was not significantly different between strains (P = 0.1267, all strains in the 1-13 month range; P = 0.1484, three strains in the 1-17 month range, two-way ANOVA). Within each strain and over the full age range investigated, the C _m of OHCs was also significantly reduced with age (P < 0.0001 in each of the four strains, one-way ANOVA: Fig. 5D), with OHC surface being reduced by ~40% in 6J, 6N and 6N-Repaired and ~25% in C3H mice by 15-17 months. This reduction was not significantly different between strains (P = 0.1243, all strains in the 1-13 month range; P = 0.4959, three strains in the 1-17 month range, two-way ANOVA). We have also found that C _m at 6 months was already significantly smaller than that at 1 month in 6J, 6N and C3H mice (P < 0.0001), but not in 6N-Repaired mice (P = 0.0727, Tukey’s post-test, one-way ANOVA). When the size of IK in OHCs was normalized to their respective C _m (i.e. current density), it was no longer significantly different for any of the four mouse strains (C3H: P = 0.1125; 6N-Repaired: P = 0.7086; 6N: P = 0.4292; 6J: P = 0.1577, one-way ANOVA, Fig. 5E), and also between strains (four strains in the 1-13 month age range: P > 0.0537; three strains in the 1-17 month range P > 0.3055, two-way ANOVA). Despite the reduced size of IK, the V _m of OHCs remained stable until at least 16-17 months of age in all mouse strains investigated (C3H: P = 0.0622; 6N-Repaired: P = 0.9826; 6N: P = 0.5555; 6J: P = 0.1125, one-way ANOVA, Fig. 5F), and also in between strains (four strains in the 1-13 month age range: P = 0.2861; three strains in the 1-17 month range P = 0.7763, two-way ANOVA).

Electromotility is maintained with ageing

One of the characteristic signatures of mature OHCs is the expression of the motor protein prestin (gene name: Slc26a5), which drive their somatic motility and thus generate DPOAEs. Therefore, we investigated whether prestin was maintained in ageing OHCs. We found that prestin was expressed in ageing OHCs from all four mouse strains until at least 16 months of age (6N and 6N-repaired: Fig. 6A-D). Since prestin is also used as an OHC marker, it allowed us to investigate any change in the number of OHCs present in the equivalent apical coil region used to perform the above electrophysiological experiments. Myosin 7a antibodies (e.g. Fig. 8) were also used as an additional hair cell marker to avoid underestimating the loss of OHC number with age due to the reduced expression of prestin. We found that the number of OHCs, which was measured over a distance of 140 μm in the ~9-12 kHz region, remained unchanged between 1 and 6 months of age in C3H (P = 0.9999), 6N (P = 0.2510) and 6N-Repaired mice (P = 0.9981, Tukey’s post-test, one-way ANOVA). However, by 14-16 months of age the number of OHCs significantly decreased compared to one month in all four strains although (C3H: P = 0.0019, 6N-Repaired: P = 0.0018, 6N: P < 0.0001, 6J: P < 0.0001, one-way ANOVA; Fig. 6E-H).

Although aged OHCs expressed the motor protein prestin (Fig. 6), we sought to test whether it was able to induce voltage-dependent changes in OHC motility (Zheng et al. 2000; Liberman et al. 2002). OHC electromotile activity can be assessed by either measuring OHC movement visually, or by using the non-linear (voltage-dependent) capacitance change (C _N-L) under whole-cell patch clamp, which is an electrical signature of electromotility (e.g. Abe et al. 2007; Jeng et al. 2020). In the present study we preferred to use C _N-L because it is particularly difficult to measure visual OHC motility on a precise horizontal plane and because displacement can be misleading through its dependence on intracellular turgor pressure, which is variable. We found that the maximum size of C _N-L recorded in OHCs decreased significantly between 1 month and aged 6N (P < 0.0013), 6J (P = 0.0160) and 6N-Repaired mice (P < 0.0017, Tukey’s post-test, one-way ANOVA), but was similar among strains (P = 0.6120, two-way ANOVA) (Fig. 7A-C). When the C _N-L was normalized to the OHC membrane capacitance, there was no significant difference among all data groups (P = 0.0589, one-way ANOVA) or between strains (P = 0.2306, two-way ANOVA) (Fig. 7D). These data indicate that the electromotile activity is maintained in aged OHCs, but reduced in amplitude most likely due to the cells becoming smaller and losing some prestin in their lateral membrane (Fig. 5D). We have also investigated whether the expression level of prestin was affected by ageing by performing quantitative PCR experiments with RNA obtained from the cochleae of 1 and 15-17 month old 6N and 6N-Repaired mice, showed that the expression of prestin (Slc26a5) was significantly reduced in both aged mouse strains (6N, P = 0.0117; 6N-Repaired, P = 0.0248, t-test) (Fig. 7E). We also found that Ocm, which encodes for the main Ca²⁺binding protein oncomodulin expressed in OHCs (Simmons et al. 2010), was significantly reduced with age (6N, P = 0.0110; 6N-Repaired, P = 0.0028, t-test) Fig. 7F) in both strains. Although these qPCR experiments are intrinsically biased by the fact that OHC loss along the length of the cochlea is likely to be different among mouse strains (e.g. Hequembourg & Liberman, 2001; Francis et al. 2003; Zachary & Fuchs, 2018; Sha et al. 2008; Ohlemiller et al. 2010), the results show a similar reduction in prestin level between the early-onset (6N) and late-onset (6N-Repaired) hearing loss mice, suggesting that its expression level is likely to be similarly affected by ageing.

OHC function also depends upon the tectorial membrane (TM), which is an acellular structure attached to the stereociliary bundles of the OHCs that gradually degrades with age especially in C57BL/6J mice (Goodyear at al., 2019). The presence and structure of the TM was investigated using light microscopy and Toluidine blue staining. In both 6N and 6N-Repaired mice, the TM was attached medially to the spiral limbus, and laterally sitting atop the OHC stereociliary bundles (Fig. 7G,H), indicating a similar gross structural organization in 17 month old mice. However, a recent ultrastructure study has shown that the tectorial membrane progressively detaches from the spiral limbus in very old CBA mice (Bullen et al. 2020), indicating some possible strain differences.

Afferent and efferent synapses are reduced in aged OHCs

We evaluated the number of afferent synapses on OHCs from 6N, 6J, 6N-Repaired and C3H mice using an anti-CtBP2 antibody to label the presynaptic ribbon protein RIBEYE. We found that CtBP2 puncta were present at the OHC presynaptic site in the different mouse strains (Fig. 8A,B for 6N and 6N-Repaired mice). The number of CtBP2 puncta in OHCs, over a distance of 140 μm in the ~9-12 kHz region, was significantly different when comparing strains tested at 1, 6 and 15 months (C3H, 6N and 6N-Repaired: P = 0.0025) or all four strains tested at 1 and 15 months (P = 0.0054, two-way ANOVA). Within each strain, the number of CtBP2 puncta decreased significantly in all strains (C3H and 6N-Repaired: P < 0.0001, 6N: P = 0.0090, one-way ANOVA; 6J: P < 0.0001, t-test, Fig. 8C-F).

Mature OHCs are mainly innervated by the cholinergic medial olivocochlear (MOC) neurons (Liberman, 1980; Maison et al. 2003), which release the neurotransmitter acetylcholine (ACh) (Simmons et al. 1996). The role of the MOC system is to modulate OHC electromotility and thus mechanical amplification in the cochlear partition (Guinan, 1996). The effects of efferent activity on OHCs is inhibitory because the release of ACh causes Ca²⁺ influx through α9α10-nAChRs, which leads to the activation of a hyperpolarizing current through small-conductance Ca²⁺ activated K⁺ channels (SK2: Oliver et al. 2000; Marcotti et al. 2004; Lioudyno et al. 2004; Katz et al. 2004). We investigated whether the efferent innervation was retained in the aged cochlea of all four mouse strains by immunolabelling with a choline acetyl transferase (ChAT) antibody, which labels the presynaptic efferent terminals, and with the postsynaptic SK2 antibody. We found that juxtaposed SK2 channels (in OHCs) and ChAT-immunoreactivity (efferent terminals) were still present in most of the remaining OHCs of 15 month old mice (Fig. 9A,B for 6N and 6N-Repaired mice). The number SK2 puncta per OHC, within a cochlear region of 140 μm of the apical coil (9-12 kHz), was not significantly reduced between 1 and 6 months (C3H: P = 0.4595; 6N: P = 0.5089; 6N-Repaired: P = 0.2419, Tukey’s post-test, two-way ANOVA). However, between 1 and 15 months, the number of SK2 puncta was significantly reduced 6N, 6N-Repaired and 6J (P = 0.0005, P = 0.0030, P < 0.0001, respectively, Tukey’s post-test, one-way ANOVA) but not in C3H mice (P = 0.6580) (Fig. 9C). We also found that although most of the SK2 channels in OHCs were juxtaposed with ChAT staining (Fig. 9D) no significant age-related changes in all strains compared between 1 and 15 months: P = 0.7668, two-way ANOVA), some OHCs were observed with only efferent terminals (Fig. 9A,B, arrowheads) or SK2 channels (Fig. 9A,B, arrows).

Mechanoelectrical transduction is reduced in OHCs from ageing mice

Another key biophysical property of hair cells is their ability to convert sound into an electrical signal, which is performed by the mechanoelectrical transducer (MET) apparatus. The gross morphology of the OHC stereociliary bundles appeared normal in the apical coil of aged 6N mice (Fig. 10A; see also Bullen et al. 2020), despite their elevated ABR and DPOAE thresholds (Figs. 1-3). We, therefore, investigated whether other properties of the MET apparatus had changed in ageing mice. Quantitative PCR revealed that the expression of Cdh23 was significantly reduced at 15 months of age in both 6N and 6J mice (P = 0.0119 and P < 0.0001, respectively, t-test, Fig. 10B). A reduction in Cdh23 expression also occurred in the 6N-Repaired strain (P = 0.0073), which have overall better hearing than both 6N and 6J mice, and like CBA (Sha et al. 2008; Sergeyenko et al. 2013) are unlikely to lose many OHCs along the cochlea at old ages. On the other hand, Pcdh15 (protocadherin-15), which together with cadherin 23 forms the stereocilia tip links required for gating the MET channels (Richardson et al. 2011), was significantly increased in 6N and 6J (P = 0.0375 and P = 0.0057, respectively) but not in 6N-Repaired mice (P = 0.0891, t-test, Fig. 10B). These data indicate that despite the normal morphology of the stereociliary bundles, the function of the MET apparatus is likely to change in the ageing cochlea, which could be investigated by recording the MET current. However, recording the MET current in OHCs is usually limited to pre-hearing ages because of the difficulties associated with making measurements from adult hair cells. Furthermore, the stereociliary bundles of OHCs are easily damaged during the physical removal of the attached tectorial membrane. In order to provide some information on possible changes in the MET current with age, we attempted a few recordings from mice (Tecta/b^-/-: see Methods), in which the tectorial membrane is not attached to the OHC stereociliary bundles and as such its removal does not disrupt the classical V-shape bundle structure (Fig. 10C, D). As for the 6J and 6N strains, we found that these Tecta/b^-/- mice, which are primarily on the 6N background, harbour the Cdh23^ahl allele. The OHCs of 11 month old Tecta/b^-/- mice were all present in the 9-12 kHz cochlear region, and their basolateral current profile was comparable to that of 1 month old OHCs (Fig. 10E, F), but with a significantly reduced K⁺ current (P = 0.0373, t-test, Fig. 10G), as also shown for the other mouse strains (Fig. 5C). Using these Tecta/b^-/- mice, we showed that a large MET current was present in OHCs from both 1 month and 11 month old mice (Fig. 10H-I), although in aged cells it was significantly reduced compared to 1 month (P = 0.0003, t-test, Fig. 10J). Despite the different MET current size, the open probability of MET channels at rest and in 1.3 mM Ca²⁺was comparable between the two ages (P = 0.9403, Fig. 10K). These data suggest the intriguing possibility that changes in the MET current with age could be a mechanism involved in the progression of age-related hearing loss in ageing mice, at least in those harbouring the Cdh23^ahl allele as previously suggested (Johnson et al. 1997; Noben-Trauth et al. 2003; Johnson et al. 2017).