Ethics statement

Mouse studies were carried out in accordance with UK Home Office regulations and the UK Animals (Scientific Procedures) Act of 1986 (ASPA) under UK Home Office licences, and the study was approved by the King’s College London and Wellcome Trust Sanger Institute Ethical Review Committees. Mice were culled using methods approved under these licences to minimize any possibility of suffering.

Mice

The mouse lines carrying the two alleles of Klhl18 used in this study originated from the Wellcome Sanger Institute Mouse Genetics Project, both generated and maintained on a C57BL/6N genetic background. One was a spontaneous missense mutation, Klhl18^lowf (Fig 1A), that occurred in a colony carrying a targeted mutation of Mab21l4 (also known as 2310007B03Rik). The mutation was detected through observation of occasional mice in this colony with raised ABR thresholds to low frequency stimuli that did not segregate with the targeted mutation in Mab21l4. The mutation was identified as a point mutation in Klhl18 by positional cloning (g.9:110455454C>A), predicted to cause a Val55Phe amino acid change in the BTB domain of the protein [2]. The second was a targeted mutant allele (Klhl18^{tm1a(KOMP)Wtsi}, referred to here as Klhl18^tm1a, Fig 1B) carrying a promoter-driven knockout-first allele, with a large cassette inserted between exons 5 and 6, which interferes with transcription leading to knockdown of expression [4,15]. The inserted cassette contains a β-galactosidase/LacZ reporter gene. Further details can be found at www.mousephenotype.org. Colonies were maintained by heterozygous intercrosses or homozygotes crossed with heterozygotes. Both Klhl18 mutant lines produce homozygous offspring that are viable and fertile, and hearing impairment was inherited in a recessive manner. Both males and females were included as no difference in auditory function was noted between them. These mutant mice are available through the European Mouse Mutant Archive (EMMA).

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Fig 1. Schematic representations of the Klhl18^lowf and Klhl18^tm1a alleles.

A. Klhl18^lowf is formed by a single base pair change on Chr9, at position 110455454, in exon 3 of the gene, with the wildtype C replaced with the mutant A. This produces an amino acid substitution in the resultant protein with the wildtype valine (conserved across species) being replaced by phenylalanine at position 55 (see [2] for further information). B. Klhl18^tm1a was generated at the Wellcome Sanger Institute by insertion of a large DNA cassette before the critical exon (exon 6) of the gene, which interrupts transcription of the gene into mRNA. The cassette is composed of a number of components; En2 SA (engrailed2 splice acceptor), IRES (internal ribosome entry site), lacZ (beta-galactosidase reporter gene), pA (polyadenylation site), hBactP (beta-actin promotor part) and neo (neomycin resistance gene), which are flanked by FRT (Flp-FRT) and loxP recombination sites to produce the promoter-driven Knockout First, reporter-tagged insertion with conditional potential allele (tm1a) of Klhl18 (see [15] for further information).

https://doi.org/10.1371/journal.pone.0258158.g001

Genotyping

Klhl18^lowf mice were genotyped by PCR amplification followed by restriction enzyme digest of the PCR product. The Klhl18^lowf mutation is a C>A missense mutation in exon 10 of the gene [2]. We amplified genomic DNA in this region (forward primer sequence: GCACAATGGTAGGGGTTCAG; reverse primer sequence, GCAGTGTCGCTCAATATTTGTCTTTGTATTCTCTTTGGCCCACAGATTGGGGACCACAAGTTCAGTGCTCACCAG). The PCR primers used together with the point mutation generated a Bgl II restriction site (AGATCT) in the mutant PCR product while the corresponding wildtype allele sequence (AGATCC) was not recognised. BgI II was used to digest the PCR product yielding 2 sequences of 78 bp and 127 bp from the mutant allele and one sequence of 205 bp from the wildtype allele, and these products were identified by gel electrophoresis. In some cases, genotypes were obtained by sequencing the PCR product amplified from a separate pair of primers (forward sequence GAAGTGTAAGGGTGTGGGGG, reverse sequence CTTGCCCGTCTGTCATCCC). The reverse primer was used for sequencing.

Klhl18^tm1a mice were genotyped using a common forward primer (CCTGTGACAAGCAGTCTGAAGG), a wildtype reverse primer (TGCTAGGGAGTGAATCTAGGGC) and a mutant-specific reverse primer (CasR1 TCGTGGTATCGTTATGCGCC). The resulting band sizes were 524 bp for the wildtype product and 384 bp for the mutant product. Primers specific for the Neomycin resistance gene in the introduced cassette were also used to detect the presence or absence of the inserted DNA of the mutant allele (Fig 1) [4,15].

Anaesthesia

In experiments where mice were tested longitudinally at different ages, mice were anaesthetised by intra-peritoneal injection of 100 mg/kg ketamine (Ketaset, Fort Dodge Animal Health) and 10 mg/kg xylazine (Rompun, Bayer Animal Health) and recovery was promoted using 1 mg/kg atipamezole (Antisedan, Pfizer). For terminal experiments, mice were anaesthetised with intra-peritoneal urethane (0.1 ml/10 g bodyweight of a 20% w/v solution of urethane in water).

Auditory Brainstem Response (ABR) recordings

Brainstem auditory evoked potentials were measured using the method described in detail previously [16,17]. Anaesthetised mice were placed in a sound-attenuating chamber (IAC Acoustics Limited) and subcutaneous recording needle electrodes (NeuroDart; Unimed Electrode Supplies Ltd, UK) were inserted on the vertex and overlying the left and right bullae. Responses were recorded to free-field calibrated broadband click stimuli (10 μs duration) and tone pips (5 ms duration, 1 ms onset and offset ramp) at frequencies of 6, 12, 18, 24 and 30 kHz, at levels ranging from 0–95 dB SPL (in 5 dB steps) at a rate of 42.6 stimuli per second. Stimuli were generated via custom software on a RZ6 multifunction processor (Tucker Davis Technologies, TDT) and presented via a FF1 loudspeaker (TDT). Evoked responses were amplified, digitized, and bandpass filtered between 300–3000 Hz, using custom software and TDT hardware (RZ6 processor, RA4LI low impedance headstage, RA4PA preamplifier). Thresholds of ABRs were defined as the lowest stimulus level to evoke a visually identifiable waveform (S1 Fig). Evoked ABR waveforms were analysed offline and the latency and amplitude of ABR waves 1–4 were plotted as a function of sound level.

Distortion Product Otoacoustic Emissions (DPOAE) recordings

We made measurements of the 2f1-f2 DPOAE component in mice across a range of ages, either as a terminal experiment in different cohorts of mice, or as part of longitudinal experiments in the same animals at increasing ages. In all cases, measurements were made inside a sound-attenuating chamber (IAC Ltd) with the mouse positioned on a heating blanket. In terminal experiments, urethane-anaesthetised mice had their left pinna and cartilaginous ear canal removed before a hollow conical speculum was positioned to give an unobstructed view of the tympanic membrane. The DPOAE measurement probe (see later), with a small rubber gasket close to its tip was then positioned within the speculum. For longitudinal experiments, ketamine/xylazine-anaesthetised mice were placed in a prone position and the head was tilted approximately 45° such that the left ear was uppermost. The DPOAE probe assembly was positioned vertically, such that the probe tip was sitting just behind the tragus, with the probe pointing down towards the opening of the ear canal. The DPOAE probe assembly used here was comprised of a pair of EC1 electrostatic drivers (TDT) coupled to the guide tubes of an ER10B+ low noise DPOAE system (Etymotic Inc) via 5 cm plastic tubes.

Stimuli were generated and DPOAE responses recorded using a RZ6 multifunction processor (TDT), under the control of TDT BioSigRZ software (TDT). Continuous f1 and f2 tone stimuli were generated and presented via different EC1 drivers within the DPOAE probe. Frequencies for f2 were set to match the ABR tone-pip frequencies used (i.e. 6, 12, 18, 24 and 30 kHz). The f2 tone was presented at a frequency 1.2 x f1, and a level 10 dB SPL lower than f1. Sound pressure levels of the f2 stimulus ranged from -10 dB to 65 dB in 5 dB steps. ER10B+ microphone signals recording during stimulus presentation were digitised at a sampling rate of 195312.5 Hz on the RZ6 processor for online Fast Fourier Transformation (FFT) to yield a power spectrum containing the f1 and f2 stimulus components and the main 2f1-f2 DPOAE component of interest in this study (S2 Fig).

From each FFT trace recorded, a number of parameters were calculated; the 2f1-f2 DPOAE amplitude, the mean noise-floor amplitude (as the average of the 20 spectral lines surrounding the DPOAE frequency) and two-times the standard deviation (SD) of the noise-floor mean. These values are plotted across stimulus level to produce plots from which the threshold of the DPOAE was defined as the lowest stimulus level when the DPOAE amplitude exceeded the 2 SDs above the recording noise-floor. From these data, we calculated mean DPOAE growth functions for each f2 stimulus frequency for control and mutant mice and mean DPOAE thresholds for these frequencies in control and mutant mice.

Endocochlear Potential (EP) recordings

The positive potential within the scala media of Klhl18^lowf mice was measured in urethane-anaesthetised mice as described previously [18,19]. A reference electrode (Ag-AgCl pellet) was positioned under the skin of the neck. A small hole was made in the basal turn lateral wall and the tip of a 150 mM KCl-filled glass micropipette was inserted into the scala media. The EP was recorded as the differential potential between the tip of the glass electrode and the reference electrode.

Round Window Response (RWR) recordings

Measurements of evoked potentials detected at the round window of the cochlea were made in Klhl18^lowf mice aged 6 weeks. The methods used here were modified from those of Harvey and Steel [20]. In urethane-anaesthetised mice, following insertion of a tracheal cannula and placement in a custom-built head-holder, a subdermal needle reference electrode was inserted on the midline of the neck. A further subdermal needle electrode was inserted in the skin at the base of the tail to serve as a ground electrode. After removing the pinna, a small opening was made in the bulla and the exposed tip of a Teflon-coated silver wire was positioned on the round window membrane.

Auditory stimuli were generated using custom software and RZ6 multifunction processor (TDT) and presented free-field via a MF1 magnetic loudspeaker (TDT) positioned 15 cm away from and directly opposite the exposed ear canal. Stimuli used were 20 ms duration tone pips, with a 1 ms ramp at the onset and offset, of the same frequencies used in the previous in vivo measurements (6, 12, 18, 24 and 30 kHz), with a fixed starting phase of 90°. Tone pips were presented from 0–95 dB SPL, at a repetition rate of 10.65 stimuli/second, with the onset of the tone delayed by 5 ms after the onset of recording. For each sound level, 256 presentations of the tone pip contributed to the generation of an averaged evoked RWR. Signals detected at the round window were amplified (x1000 gain) and filtered (1 Hz high pass, 50 kHz low pass) using a DP311 differential amplifier (Warner Instruments). The amplified filtered signal was sampled at 97656.25‬ Hz by the RZ6 processor (TDT) and under software control, 80 ms snippets of the signal were used to form an averaged RWR to each combination of tone frequency and sound level.

Analysis of electrophysiological responses from the round window of Klhl18 mice

The RWRs recorded were analysed offline (S3 Fig) using custom-written scripts in Matlab (v2019a, The Mathworks Inc., Natick MA, USA). Each RWR (S3A Fig) was filtered into narrow bands to extract different components of the signal. A low pass filter (corner frequency, Fc, 3000 Hz) was used to extract a combined cochlear nerve Compound Action Potential (CAP) and Summating Potential (SP) response (S3Bi Fig). The SP was measured as the mean amplitude of the waveform calculated over the 19–24 ms time window (equivalent to the 14–19 ms section of the tone burst stimulus). The SP could be either positive or negative in sign, relative to the zero baseline of the response (S3Bii Fig). The amplitude of SP measured across stimulus level (from 0–95 dB) was plotted to form an input-output function (S3Biii Fig). A bandpass filter (high pass Fc, 300 Hz; low pass Fc, 3 kHz) was used to extract the cochlear nerve CAP (S3Ci Fig). The CAP was formed of significant negative and positive peaks to its waveform (labelled N and P, respectively in S3Ci and S6-1Cii Figs). For each stimulus level, the N-P peak-to-peak amplitude was calculated and along with the N and P latencies following the onset of the tone pip were plotted to form input-output functions for these parameters (S3Ciii and S3Civ Fig). A narrow bandpass filter centred at stimulus frequency (with low and high pass Fc’s of ± 100 Hz) was used to isolate the Cochlear Microphonic (CM) (S3Di Fig). This filtered response was trimmed to a time window equivalent to the 2–18 ms section of the tone burst stimulus, and this steady-state section of the CM response was subject to Fast Fourier Transformation. The resultant power spectrum (S3Dii Fig) was used to determine the amplitude of the CM component. The amplitude of the CM was converted to a dB scale, using 1 μV as a reference amplitude. The dB (re 1 μV) CM amplitude measured across stimulus level was plotted to form an input-output function of CM growth (S3Diii Fig).

Innervation and synaptic labelling with confocal microscopy imaging

Inner ears from 6-week old mice were fixed in 4% paraformaldehyde for 2 hours and decalcified in 0.1 M EDTA overnight at room temperature. Following fine dissection in PBS, the organ of Corti was permeabilised in 5% Tween20 in PBS for 40–60 minutes and incubated in blocking solution (0.45% Triton X-100, 10% normal horse serum in PBS) for 2 hours. Following blocking, the samples were incubated overnight at room temperature with shaking using primary antibodies in 0.36% Triton, 6% normal horse serum in PBS. For synaptic labelling, primary antibodies were mouse anti-GluR2 (1:200, MAB397, Emd Millipore) and rabbit anti-Ribeye (1:500, 192 103, Synaptic Systems). For neuron labelling, primary antibodies were mouse anti-CtBP2 (1:400, BD Transduction Laboratories 612044) to label pre-synaptic ribbons and IHC nuclei and chicken anti-Neurofilament-Heavy (1:800, Abcam ab4680) to label unmyelinated neural dendrites. Samples were then washed with PBS and then incubated with secondary antibodies for 45–60 minutes in the dark. For synapses, secondary antibodies were either donkey anti-mouse IgG Alexa Fluor594 (A21203, ThermoFischer Scientific) or goat anti-mouse IgG2a Alexa Fluor488 (A21131, ThermoFischer Scientific) and goat anti-rabbit IgG Alexa Fluor546 (A11035, ThermoFischer Scientific). For neuron labelling, secondary antibodies were donkey anti-mouse Alexa Fluor594 (1:500, A-21203, Molecular Probes) and goat anti-chicken Alexa Fluor488 (1:300, A11039, Life technologies). Samples were then washed in PBS before mounting in ProLong Gold mounting medium with DAPI and stored at 4°C in the dark. Specimens were viewed using a Zeiss Imager 710 confocal microscope interfaced with ZEN 2010 software with a x63 objective and numerical aperture of 1.4. The whole length of the organ of Corti was examined and images collected at best-frequency regions based upon the frequency-place map of Müller and colleagues [21]. Z-stacks were taken at 0.25 μm intervals and maximum intensity projection images were generated. Brightness and contrast were normalised for the dynamic range in all images. The number of ribbon synapses per IHC was quantified by manually counting the co-localised Ribeye and GluR2 puncta in the confocal maximum projection images and dividing the sum by the number of DAPI-labelled IHC nuclei. An average of six IHCs per image was considered for ribbon synapses counting using the cell-counter plugin in Fiji software.

Scanning electron microscopy

Cochlear samples from Klhl18^lowf mice aged 3 weeks and 4 weeks old were fixed in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer with 3 mM calcium chloride, dissected to expose the organ of Corti, then processed by a standard osmium tetroxide-thiocarbohydrazide (OTOTO) protocol [22]. After dehydration, samples were subjected to critical point drying, mounted and viewed using a Jeol JSM-7800F Prime Schottky field emission scanning electron microscope. An overview of the cochlea was imaged to allow calculation of percentage distances along the cochlear duct to superimpose the frequency-place map [21], allowing subsequent imaging of consistent locations across different specimens. Images were assessed by three independent viewers who were blinded to genotype.

Experimental design and statistical analysis

We used heterozygotes as controls in most of the electrophysiological experiments because we did not see any obvious difference between heterozygotes and wildtype littermates. There was no significant effect of sex on any of the physiological recordings in control or mutant mice of either Klhl18^lowf or Klhl18^tm1a lines (S4A–S4F Fig; S1 Table), so both males and females were combined into a single group for their age and genotype. As the auditory phenotype changes with age of the homozygous mutants, we use littermates as controls and carefully age-matched mice in different age groups; 2 weeks (Post-natal day, P, 14 +/- 0 days); 3 weeks (P21 +/- 0 days); 4 weeks (P28 +/- 1 day); 6 weeks (P42 +/- 2 days); 8 weeks (P56 +/- 4 days); 14 weeks (P98 +/- 4 days).

All statistical comparisons were made using the analysis routines within Graphpad Prism (v8.4.2). Auditory Brainstem Response, Distortion Product Otoacoustic Emissions and Round Window Response data were analysed using a Mixed-effect Model, with either Sidak’s or Tukey’s multiple comparisons tests. Endocochlear Potential values were compared using a Kruskal-Wallis One-Way ANOVA. Counts of synaptic components were compared using a two-tailed unpaired t-test, with Welch’s correction. All tests were performed with alpha = 0.05. Statistical significance was determined at the level of p < 0.05.

Klhl18 mutant mice showed progressive increases in ABR thresholds with age, particularly for low frequency stimuli

The ABR is a convenient electrophysiological approach that can be recorded non-invasively, allowing us to track any changes in auditory responses in the same mouse as it ages. In heterozygous control mice, ABR thresholds improved from 2 weeks old, through 3 weeks old to 4 weeks old (Fig 2A–2C), indicating maturation of auditory sensitivity. At 2 weeks old, ABR thresholds of Klhl18^lowf/lowf mutants were slightly but statistically significantly elevated, compared to age-matched heterozygous littermate controls (Fig 2A; S1 Table). At 3 weeks old, ABR thresholds of mutant and control mice are generally comparable (Fig 2B). From the age of 4 weeks (Fig 2C–2F), ABR thresholds for all stimuli were significantly higher in mutants than in age-matched heterozygous controls (See S1 Table). As the mice aged beyond 4 weeks, control mice maintain low ABR thresholds but there was a progressive elevation of thresholds from 4 weeks old to 14 weeks in the Klhl18^lowf mutant mice, most pronounced at 6–18 kHz (Fig 2C–2F). A similar pattern of age-related threshold changes is seen in Klhl18^tm1a mutant mice (S5 Fig).

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Fig 2. Age-related changes in ABR thresholds of Klhl18^lowf mice.

A summary of ABR thresholds estimated from control (all heterozygotes) and mutant (homozygotes) are shown for Klhl18^lowf mice aged 2 weeks (A, n = 13 control, n = 12 mutant), 3 weeks (B, n = 8 control, n = 19 mutant), 4 weeks (C, n = 17 control, n = 9 mutant), 6 weeks (D, n = 17 control, n = 10 mutant), 8 weeks (E, n = 10 control, n = 6 mutant) and 14 weeks (F, n = 10 control, n = 6 mutant). Thresholds from individual mutant mice are indicated by grey circles. Mean threshold (±SD) are plotted for control mice as black down-triangles and for mutant mice as red up-triangles. Ck: Click stimulus. The results of a mixed-effects model statistical analysis between control and mutant thresholds, followed by Sidak’s multiple comparisons test was used to examine the difference between thresholds for each stimulus are shown in S1 Table. Significant differences are indicated here by *.

https://doi.org/10.1371/journal.pone.0258158.g002

Klhl18 mutant mice showed abnormal ABR waveform shapes

In addition to determining the threshold for detection of an ABR, we also examined the waveform features. Group averaged ABR waveforms from Klhl18^lowf mice for selected age groups and frequency stimuli are plotted in Fig 3. As the ABR waveform changes with increasing stimulus level (amplitude increases and latency decreases and the stimulus increases above threshold), we plot mean waveforms at both a fixed dB SPL (65dB SPL; Fig 3A–3I) and also at 20 dB sensation level (dB SL; dB above threshold; Fig 3J–3R). For mice aged 2 weeks old, waveforms evoked by click stimuli and 12 kHz and 24 kHz tone pips (Fig 3A–3C and 3J–3L) showed that Klhl18 mutant mice had smaller amplitudes than controls. At 3 weeks old, when ABR thresholds were generally comparable across genotypes, the averaged waveforms for these stimuli were more similar (Fig 3D–3F and 3M-3O); the click-evoked response was smaller in mutants, but the tone-evoked responses had similar amplitudes in control compared with mutant mice. These mean waveform amplitudes from 2–3 weeks old mice were in contrast to those from older mice. At 6 weeks old, when mutant mice exhibited a clear increase in thresholds, the averaged waveforms of mutants evoked by click, 12 kHz and 24 kHz stimuli were much smaller than those in control mice (Fig 3G–3I and 3P–3R). Similar observations were obtained from Klhl18^tm1a mutant mice (S6 Fig).

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Fig 3. Mean ABR waveforms of Klhl18^lowf mice at 65dB SPL and 20 dB SL.

Group averaged ABR waveforms, evoked by 65 dB SPL stimuli, are plotted for mutant mice (red lines) and heterozygous littermate controls (black lines) aged 2 weeks (A-C), 3 weeks (D-F) and 6 weeks (G-I) in response to click stimuli (A,D,G), 12 kHz stimuli (B,E,H) and 24 kHz stimuli (C,F,I). Group averaged ABR waveforms, evoked by stimuli presented at 20 dB above threshold (20 dB sensation level, dB SL), are plotted for mice aged 2 weeks (J-L), 3 weeks (M-O), 6 weeks (P-R) in response to click stimuli (J,M,P), 12 kHz stimuli (K,N,Q) and 24 kHz stimuli (L,O,R). Group sizes were, at 2 weeks old, n = 13 control mice, n = 12 mutants; at 3 weeks old, n = 8 control mice, n = 19 mutants; at 6 weeks old, n = 17 controls, n = 10 mutants.

https://doi.org/10.1371/journal.pone.0258158.g003

ABR wave 1 showed reduced amplitude and longer latency in Klhl18 mutant mice

In order to quantify the abnormal features of the ABR waveform, ABR wave 1 amplitudes (measured as wave P1 –N1 peak to peak amplitude) and P1 latencies were plotted as a function of stimulus level (Fig 4; See also S1 Fig). The mild threshold elevation in 2-week old Klhl18^lowf mutants was reflected in a shift to the right of growth curves (Fig 4A–4C and 4J–4L). In mutant mice aged 3 weeks old, wave 1 amplitude was similar to controls (Fig 4D–4F). However, P1 latency maintained a slight increase in mutants compared to age-matched heterozygous controls (Fig 4M–4O). By 6 weeks old, when ABR thresholds were clearly raised in mutants, much larger changes in ABR wave 1 amplitude and latency were noted (Fig 4G–4I and 4P–4R). Similar patterns of amplitude and latency changes were observed in Klhl18^tm1a mutant mice (S7 Fig). Amplitudes and latencies of ABR wave 1 in Klhl18^tm1a heterozygotes were similar to their wildtype littermates (S8 Fig).

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Fig 4. ABR wave 1 amplitude and latency input-output functions in Klhl18^lowf mice.

Mean ABR wave 1 P1-N1 peak-to-peak amplitudes (±SD) as a function of stimulus level for mutant mice and heterozygous littermate controls aged 2, 3 and 6 weeks old are plotted in A-C (n = 13 controls, n = 12 mutants), D-F (n = 8 controls, n = 19 mutants) and G-I (n = 17 controls, n = 10 mutants), respectively. Results are plotted for click stimuli (A,D,G), 12 kHz tones (B,E,H) and 24 kHz tones (C,F,I). Data from control mice are plotted as black down-triangles. Data from mutant mice are plotted as red up-triangles. Mean ABR wave P1 latency (±SD) as a function of stimulus level is plotted for the same mice aged 2, 3 and 6 weeks old are plotted in J-L, M-O and P-R, respectively. Results are plotted for click stimuli (J,M,P), 12 kHz tones (K,N,Q) and 24 kHz tones (L,O,R). The results of a mixed-effects model statistical analysis between control and mutant thresholds are shown in S1 Table. Sidak’s multiple comparisons test was used to examine the difference between control and mutant data for each stimulus. Significant differences are indicated here by *.

https://doi.org/10.1371/journal.pone.0258158.g004

Normal DPOAE responses in Klhl18 mutant mice

Auditory Brainstem Responses reflect auditory nerve and brainstem responses, but DPOAEs are useful additional measures because they reflect outer hair cell function [23]. At 2 weeks old, there was a small statistically significant reduction in DPOAE amplitudes and elevation in DPOAE thresholds (for f2 frequencies of 12–30 kHz) in Klhl18^lowf mutants, similar to the small differences in ABR responses recorded at this age (Fig 5A–5F). In mice aged 3 weeks old, DPOAE responses were the same in Klhl18^lowf mutants as in heterozygous littermate controls, again mimicking the ABR thresholds at 3 weeks old (Fig 5G–5L; no significant interaction of either DPOAE amplitude or threshold with genotype; see S1 Table). However, in contrast to the significant elevation in ABR thresholds by the age of 6 weeks (Fig 3), there was no difference between DPOAE responses of control and mutant mice at this age (Fig 5M–5R; no significant interaction of either DPOAE amplitude or threshold with genotype; see S1 Table). Furthermore, no differences were found in DPOAEs in Klhl18^tm1a mutant mice up to 6 weeks old (S9 Fig).

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Fig 5. DPOAE growth and thresholds in Klhl18^lowf mice.

DPOAE results from mutant mice and heterozygous littermate controls aged 2, 3 and 6 weeks old are plotted in A-F (n = 13 controls, n = 12 mutants), G-L (n = 8 controls, n = 19 mutants) and M-N (n = 9 controls, n = 13 mutants), respectively. Data from control mice are plotted as black down-triangles. Data from mutant mice are plotted as red up-triangles. The mean (±SD) amplitude of the 2f1-f2 DPOAE (dB SPL) is plotted as a function of f2 stimulus level (dB SPL) for f2 frequencies of 6 kHz (A,G,M), 12 kHz (B,H,N), 18 kHz (C,I,O), 24 kHz (D,J,P) and 30 kHz (E,K,Q). Mean threshold (±SD) of the 2f1-f2 DPOAE (derived from individual growth functions, e.g. shown in S2 Fig) and plotted in F, L and R for mice aged 2 weeks, 3 weeks and 6 weeks respectively. In addition to the mean data, thresholds from individual mutant mice are plotted as grey open circles. Results from a mixed-effects model statistical analysis between control and mutant data are shown in S1 Table. Sidak’s multiple comparisons test was used to examine the difference between control and mutant data for each stimulus. Significant differences are indicated here by *.

https://doi.org/10.1371/journal.pone.0258158.g005

Endocochlear Potential (EP) was normal in Klhl18^lowf mutant mice

Some mutations lead to reduced endocochlear potential which can cause hearing impairment [18], so we measured EP in 6 weeks old Klhl18^lowf mutant and littermate control mice. EP was 122.4 ± 5.4 mV (mean ± SD) in 6 wildtype mice (aged 42.0 ± 0 days), 121.2 ± 5.3 mV in 6 heterozygote mice (aged 42.3 ± 1.2 days) and 123.4 ± 5.6 mV in 7 homozygous mutant mice (aged 42.1 ± 0.9 days). There was no significant difference in EP between mice of these 3 genotypes (S1 Table).

Round Window Responses (RWRs) match ABR and DPOAE responses

Additional information about cochlear function can be obtained by recording responses using an electrode placed on the round window of the cochlea, which is closer to the source of the electrical activity. We analysed three different responses extracted from recordings from the round window in 6-week old Klhl18^lowf mutants and heterozygous littermate controls: Compound Action Potential (CAP), Cochlear Microphonic (CM) and Summating Potential (SP) (Fig 6). Extraction of the different potentials is illustrated in S3 Fig. Examples of RWRs and their components are shown in S10 and S11 Figs for a Klhl18^+/+ wildtype mouse and a Klhl18^lowf/lowf mutant mouse, respectively, for 18kHz tone stimuli.

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Fig 6. Round window response measurements from 6-weeks old Klhl18^lowf mice.

RWRs were obtained from 10 wildtype (Klhl18^+/+, aged 41.8 ± 0.6 days), 7 heterozygote (Klhl18^+/lowf aged 42.6 ± 1.1 days) and 10 homozygote mutant (Klhl18^lowf/lowf aged 41.3 ± 1.2 days) mice. CAP latency (of wave N) and amplitude (N-P amplitude) are plotted in A-E and F-J, respectively, for potentials measured in response to tones of 6, 12, 18, 24 and 30 kHz. Data are plotted as mean ± SD for wildtype control mice (black down-triangles) and homozygote mutant mice (red up-triangles). K. Mean ± SD of the CAP threshold is plotted against stimulus frequency for wildtype control mice (black down-triangles) and homozygote mutant mice (red up-triangles). Open circles and grey lines indicate CAP thresholds for individual mutant mice. L-P plot mean (± SD) CM amplitude (dB re 1 μV) against stimulus level (dB SPL) for wildtype mice (black down-triangles) and homozygote mutant mice (red up-triangles). We were unable to completely remove stimulus artifact from the RWRs recorded in our system. To account for this, we assessed the magnitude of the stimulus artifact across frequency and sound level, from analysis of measurements taken after the mouse was removed the system and the recording electrodes placed in 150 mM KCl. The biological CM was orders of magnitude greater in amplitude than the stimulus artifact and thus it was distinguishable from the artifact. The artifact estimated across all experiments, across stimulus frequency and level are plotted as grey lines (with SD error bars). Q. Mean ± SD of the estimated CM threshold is plotted against stimulus frequency for wildtype control mice (black down-triangles) and homozygote mutant mice (red up-triangles). Open circles and grey lines indicate CM thresholds for individual mutant mice. Ri-Vi plot mean (± SD) Summating Potential amplitude (μV) against stimulus level (dB SPL) for wildtype mice (black down-triangles) and homozygote mutant mice (red up-triangles). Rii-Vii plot SP input-output functions (IOFs) for individual control (black lines) and mutant (red lines) mice. Data from Klhl18^+/lowf mice are shown in S12 Fig. Results of a mixed-effects model statistical analysis between wildtype, heterozygote (S12 Fig) and homozygote data are shown in S1 Table. Sidak’s multiple comparisons test was used to examine the difference between control and mutant data for each stimulus. Significant differences are indicated here by *.

https://doi.org/10.1371/journal.pone.0258158.g006

Compound action potentials (CAP), reflecting auditory nerve activity [e.g. 24], had thresholds and amplitudes very similar to those seen for ABRs, with low frequencies more severely affected than high frequencies in mutants (compare Fig 6K with Fig 2D, and Fig 6G and 6I with Fig 4H and 4I). Latencies of CAP responses were longer in mutants than in controls (Fig 6A–6E) but the shift in latency at each frequency matched the shift in threshold so the latencies appear to be normal when adjusted for hearing level.

Cochlear microphonics (CM), reflecting mostly outer hair cell activity [e.g. 25], showed no significant difference in thresholds or amplitudes in Klhl18^lowf/lowf compared with Klhl18^+/+ mice (Fig 6L–6Q, S1 Table). This matched our findings from DPOAEs, which were also normal in the mutants.

Summating potentials (SP) are a dc shift in the round window recording sustained for the duration of a toneburst and are thought to reflect hair cell depolarisation, particularly of basal turn IHCs [25,26]. They can be negative or positive in polarity, with negative SP usually detected at lower frequencies and higher stimulus intensities. We found these responses to be variable between individual mice, whatever their genotype (Fig 6Rii–6Vii). Despite the variability, SP amplitudes were significantly different in mutants at 6 and 12 kHz, while there was no difference in SP at higher frequencies (See Fig 6 and S1 Table). No significant differences between wildtypes (Klhl18^+/+) and heterozygotes (Klhl18^+/lowf) were found in any of the round window measures (Figs 4–6 and S1 Table).

Normal innervation of the cochlea

The presence of abnormal ABRs and CAPs together with normal DPOAE and CM responses and normal EPs suggests that Klhl18 mutant mice exhibit an IHC defect or neural hearing loss, so we looked in more detail at the neural elements of the organ of Corti. We used neurofilament immunolabelling in whole mount organ of Corti preparations to examine the dendrites of cochlear neurons in 6-week old Klhl18^lowf mutants and heterozygous littermate controls in regions corresponding to 6, 18 and 30 kHz characteristic frequencies (Fig 7). Confocal imaging showed a normal pattern of innervation in all regions examined. To examine synaptic organization in Klhl18^lowf mice, we immunolabelled GluR2 (a post-synaptic glutamate receptor subunit) and Ribeye (a component of presynaptic ribbon synapses) in the organ of Corti of 6 week old mice (Fig 8). We found no significant differences (see S1 Table) in numbers of ribbons or postsynaptic densities or co-localised labelled puncta (indicating synapses) in these mutants compared with heterozygous littermate controls, either in regions exhibiting raised ABR thresholds (12 kHz region) or relatively normal ABR thresholds (24 kHz region).

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Fig 7. Auditory nerve innervation of the organ of Corti at 6 weeks old.

Representative confocal microscopy images of the organ of Corti at the 6 kHz (A,B), 18 kHz (C,D) and 30 kHz (E,F) regions of the cochlea from a sample of 7 ears from 5 Klhl18^+/lowf mice (A,C,E), and 8 ears from 6 Klhl18^lowf/lowf mice (B,D,F). CtBP2-positive regions (IHC nuclei and ribbon synapses) are immuno-stained in red; Neurofilament-positive regions (nerve fibres) are immuno-stained in green. The scale bar shown in (E) represents 10μm.

https://doi.org/10.1371/journal.pone.0258158.g007

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Fig 8. IHC synapses in Klhl18 mutant mice at 6 weeks old.

Representative confocal microscopy images of IHCs at the 12 kHz (A,B) and 24 kHz (C,D) regions of the cochlea from a sample of 7 Klhl18^+/lowf mice (A,C), and 7 Klhl18^lowf/lowf mice (B,D). A row of IHC nuclei can be seen across the upper portion of panels A-D, stained blue with DAPI. Below this row, clusters of immuno-stained puncta are visible. Green puncta show immuno-positive staining for the post-synaptic receptor subunit GluR2. Red puncta show immuno-positive staining for Ribeye, a protein component of the ribbon synapse apparatus found in inner hair cells. Yellow staining represents overlap between GluR2 and Ribeye (colocalised) staining and is presumed to represent a functional synapse between the inner hair cells and the auditory nerve. Scale bars indicate 5 μm. (E,F) Counts of Ribeye-positive, GluR2-positive and colocalised Ribeye and GluR2-positive puncta are plotted for the 12 kHz region (E) and the 24 kHz region (F). Data are plotted as bars representing the mean count per IHC (error bars indicate the SD) for control (Klhl18^+/lowf, blue bars) mice and homozygous mutant mice (Klhl18^lowf/lowf, red bars).

https://doi.org/10.1371/journal.pone.0258158.g008

Tapering of tips of stereocilia in Klhl18^lowf mutants

The stereocilia bundle at the top of each hair cell houses the mechano-electrical transduction apparatus and so is essential for normal auditory function. Hearing impairment often results from abnormal or damaged stereocilia, so we examined the apical surface of the organ of Corti by scanning electron microscopy to look for any signs of hair cell degeneration or other defects. Homozygotes, heterozygotes and wildtypes were studied at two ages (3 weeks and 4 weeks old), corresponding to stages when ABR thresholds were normal (3 weeks old) and when thresholds had started to increase in homozygotes (4 weeks old). In general, the organization of the sensory epithelium, the general patterning of inner and outer hair cells within the epithelium, and the appearance of all the associated supporting cell types were normal in Klhl18^lowf mutant mice. There was no evidence seen of hair cell degeneration in the mutants at these ages, although we did not analyse the lower basal turn.

However, we did observe stereocilia abnormalities in the mutants that were most pronounced in the apical turn. Surprisingly we found similar defects at both 3 and 4 weeks old. The most severe defects were found in IHCs which showed frequent tapering of the tallest stereocilia towards their tips (examples shown in Fig 9A–9H). This feature was variable between hair bundles and also between individual stereocilia within a single bundle (see for example Fig 9D, stereocilia on right side of image). The shorter two rows of stereocilia had a more normal appearance. Outer hair cell stereocilia of mutants looked close to normal, but some bundles appeared slightly more irregular than in wildtypes and there were more often missing individual stereocilia in the shortest row in the mutants (Fig 10A–10H). In both inner and outer hair cells, stereocilia were arrayed in the cuticular plate in the normal V-shaped arrangement for OHCs or crescent for IHCs, and the angle of the V varied along the cochlear duct in a normal way in mutants, with a wider angle of the V in OHCs towards the base. Stereocilia were also arranged in rows of graded heights as usual, but with more irregularity in heights within each row in mutant bundles. All of these hair cell defects in IHCs and OHCs showed a gradient along the length of the cochlear duct with more severe defects towards the apex of the cochlea, but the abnormal IHC hair bundles extended further towards the base than abnormal OHC bundles. This tendency to more severe defects in the apex fits with the predominance of ABR threshold increases at low frequencies.

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Fig 9. Scanning electron microscopy of typical stereocilia bundles of inner hair cells in Klhl18 mutant mice in middle and apical turns.

Representative images are shown from (A-D) Klhl18^lowf/lowf homozygotes and (E-H) littermate wildtype mice. (A, E) 3 weeks old mice at the 12 kHz best-frequency region; (B, F) 3 weeks old mice at the 24 kHz best-frequency region; (C, G) 4 weeks old mice at the 12 kHz best-frequency region; (D, H) 4 weeks old mice at the 24 kHz best-frequency region. (A-G) show views from the modiolar side of the bundle, but (H) shows a view from the pillar side to illustrate the normal shape of the tallest row of stereocilia. Scale bar (on panel H) represents 1 μm.

https://doi.org/10.1371/journal.pone.0258158.g009

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Fig 10. Scanning electron microscopy of typical stereocilia bundles of outer hair cells in Klhl18 mutant mice in mid- and apical turns.

Representative images are shown from (A-D) Klhl18^lowf/lowf homozygotes and (E-H) littermate wildtype mice. (A, E) 3 weeks old mice at the 12 kHz best-frequency region; (B, F) 3 weeks old mice at the 24 kHz best-frequency region; (C, G) 4 weeks old mice at the 12 kHz best-frequency region; (D, H) 4 weeks old mice at the 24 kHz best-frequency region. Scale bar (on panel H) represents 1 μm.

https://doi.org/10.1371/journal.pone.0258158.g010

Interestingly, we observed some minor tapering of the distal ends of IHC stereocilia in the extreme apical 10% of the cochlear duct even in wildtypes. Heterozygotes showed an intermediate degree of IHC stereocilia tapering especially visible at 4 weeks old at the 3 and 6kHz best-frequency regions.

Expression of Klhl18 in the organ of Corti

To investigate the expression of Klhl18 in the inner ear, we used single cell RNAseq data from the mouse inner ear at embryonic day(E) 16, post-natal day (P) 1, P7 [27] and P15 [28], accessed via the gEAR portal (https://umgear.org/). Klhl18 expression was seen in IHCs, OHCs, pillar cells and Deiters’ cells at one or more of these stages. It was not seen at any stage in outer pillar cells, Hensen cells, or inner phalangeal cells (Fig 11). To visualise the expression, we chose 6 marker genes for comparison (Myo7a for hair cells, Fgf8 for inner hair cells, Slc26a5 for outer hair cells, Sox2 for non-sensory cells, S100b for inner pillar cells and Hes5 for Deiters’ cells) and normalised expression to the housekeeping gene Hprt. Klhl18 is expressed in both inner and outer hair cells at low but increasing levels from P1 to P15, and in Deiters’ cells from P15, while its expression in inner pillar cells at P1 and P7 is very low. Its temporal and spatial expression pattern most closely match Myo7a, although it is expressed at much lower levels.

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Fig 11. Expression of Klhl18 in key cell types during development.

Expression of Klhl18 (red), Myo7a (green), Fgf8 (yellow), Slc26a5 (blue), Sox2 (purple), S100b (teal) and Hes5 (grey) at stages from E16 to P15 in the mouse inner ear. Single cell RNAseq data from [27,28]. Expression levels were normalised to Hprt expression, indicated by a horizontal black line at y = 1 on each plot. n/a indicates data not available for this age.

https://doi.org/10.1371/journal.pone.0258158.g011