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Sensory transduction is required for normal development and maturation of cochlear inner hair cell synapses

  1. John Lee
  2. Kosuke Kawai
  3. Jeffrey R Holt  Is a corresponding author
  4. Gwenaëlle SG Géléoc
  1. Speech and Hearing Bioscience & Technology Program, Division of Medical Sciences, Harvard University, United States
  2. Department of Otolaryngology, Boston Children’s Hospital and Harvard Medical School, United States
  3. Department of Neurology, Boston Children’s Hospital and Harvard Medical School, United States
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Cite this article as: eLife 2021;10:e69433 doi: 10.7554/eLife.69433

Abstract

Acoustic overexposure and aging can damage auditory synapses in the inner ear by a process known as synaptopathy. These insults may also damage hair bundles and the sensory transduction apparatus in auditory hair cells. However, a connection between sensory transduction and synaptopathy has not been established. To evaluate potential contributions of sensory transduction to synapse formation and development, we assessed inner hair cell synapses in several genetic models of dysfunctional sensory transduction, including mice lacking transmembrane channel-like (Tmc) 1, Tmc2, or both, in Beethoven mice which carry a dominant Tmc1 mutation and in Spinner mice which carry a recessive mutation in transmembrane inner ear (Tmie). Our analyses reveal loss of synapses in the absence of sensory transduction and preservation of synapses in Tmc1-null mice following restoration of sensory transduction via Tmc1 gene therapy. These results provide insight into the requirement of sensory transduction for hair cell synapse development and maturation.

Editor's evaluation

Deafness is often caused by a defect in mechanotransduction. Lately it has become clear that synaptopathy, a defect at the first synapse in the auditory pathway, also causes hearing loss. Here the authors show that synapses can be lost following a loss of hair cell mechanotransduction, but that restoration of mechanotransduction can prevent the synaptic loss. These results are important for understanding hearing loss and restoration.

https://doi.org/10.7554/eLife.69433.sa0

Introduction

Acoustic overexposure and aging can cause significant loss of synaptic connections between inner hair cells (IHCs) and the afferent fibers of spiral ganglion neurons (SGNs), which function to relay information to the brain via the eighth cranial nerve (Kujawa and Liberman, 2009; Kujawa and Liberman, 2006; Sergeyenko et al., 2013). Loss of IHC-SGN synapses, known as cochlear synaptopathy, is evident following noise exposure producing both temporary threshold shifts (TTS) and permanent threshold shifts (PTS), with rapid loss of up to 40–50% of synapses and gradual degeneration of SGNs even in the absence of hair cell death (Kujawa and Liberman, 2009). Aging mice show similar synaptopathy, with a steady loss of synapses and a delayed but similar decrease in the number of SGNs that precedes age-related hair cell loss (Sergeyenko et al., 2013).

Damage to stereocilia, the mechanosensory microvilli, is also observed in aging and noise-exposed cochleas. Following PTS-inducing noise exposure, irreversible disarray, fusion, and loss of IHC stereocilia are visible even in the absence of hair cell death (Wang et al., 2002). Similarly, IHC stereocilia in aging mice undergo a number of changes including fusion, elongation, and internalization without obvious hair cell loss (Bullen et al., 2019). Stereocilia defects compromise the integrity of the mechanosensory transduction apparatus, resulting in absent or impaired sensory transduction and thus contribute to diminished hearing sensitivity (Pickles et al., 1987; Assad et al., 1991).

Despite the concurrence of mechanosensory insult and cochlear synaptopathy in noise exposure and aging, impaired sensory transduction has not been implicated as a mechanism contributing to the loss of hair cell synapses. To begin to address the contributions of sensory transduction to the development and maturation of IHC-SGN synapses, we evaluated IHC ribbon synapses across multiple developmental timepoints in five genetic models with disrupted sensory transduction, including mice lacking Tmc1, Tmc2, or both (Tmc1Δ/Δ, Tmc2Δ/Δ, Tmc1Δ/Δ;Tmc2Δ/Δ), Beethoven mice which carry a dominant mutation in Tmc1 (Bth) and Spinner mice (Tmiesr), which carry a mutation in Tmie. TMC proteins form the pore of hair cell transduction channels (Pan et al., 2018) and TMIE is a necessary component of the hair cell mechanosensory transduction complex (Zhao et al., 2014). Mutations in these proteins cause transduction dysfunction and hearing loss in mice and humans (Kurima et al., 2003; Vreugde et al., 2002; Kawashima et al., 2011; Naz et al., 2002; Mitchem et al., 2002; Zhao et al., 2014).

Our analyses reveal an unanticipated and complex relationship between sensory transduction, synaptogenesis, and synaptopathy. In mouse models with genetic disruption of sensory transduction, we find that IHC synapses undergo exuberant synaptogenesis during the first postnatal week and that the number of synapses declines drastically over the following few weeks. We also investigated whether restoration of sensory transduction, using an established Tmc1 gene therapy strategy capable of targeting nearly 100% of IHCs (Lee et al., 2020; Wu et al., 2021), preserved normal synaptic development and maturation. We report that Tmc1 gene therapy in Tmc1Δ/Δ mice preserves synapses in mature mice, and that synapse counts were correlated with recovery of ABR thresholds. Together, these data provide insight into synaptic changes associated with sensory transduction and suggest that dysfunction of sensory transduction may contribute to cochlear synaptopathy.

Results

Tmc deletion alters synapse development and maturation

To determine whether initial ribbon formation was influenced by the presence of functional mechanosensory transduction channels, the number of CtBP2-positive puncta per IHC was quantified in five wild-type (WT) and six Tmc1Δ/Δ;Tmc2Δ/Δ mice at postnatal day 2 (P2) from cochlear regions corresponding to 8, 11.3, 16, 22.6, and 32 kHz. Counts of CtBP2-positive puncta were similar in both groups of mice and to those previously reported for WT C57/BL6 mice at P2 (Huang et al., 2012). No significant differences in counts were observed between WT and Tmc1Δ/Δ;Tmc2Δ/Δ groups across all frequency regions (Figure 1, Supplementary file 1), suggesting the initial formation of presynaptic ribbon precursors is independent of the acquisition of sensory transduction, which begins at the base at P0 and progresses tonotopically toward the apex by P4 (Lelli et al., 2009).

CtBP2+ puncta counts in Tmc1Δ/Δ;Tmc2Δ/Δ mice lacking sensory transduction are similar to those in wild-type (WT) mice at postnatal day 2 (P2).

(A) Representative 3D projections of confocal z-stacks of P2 WT and Tmc1Δ/Δ;Tmc2Δ/Δ (B) inner hair cells (IHCs) from the 16 kHz region. Scale bar: 7 µm. The tissue was immunostained with anti-Myosin7a (blue) and anti-CtBP2 (red). (C) To determine a mean number of ribbon precursors/IHC, the total number of CtBP2+ puncta from 8 to 10 IHCs was counted for each frequency region. Individual points represent counts from one mouse. Data from WT (black; n = 5) and Tmc1Δ/Δ; Tmc2Δ/Δ (red; n = 5–6) groups are illustrated. Bold lines indicate mean ± SD.

Figure 1—source data 1

CtBP2+ puncta counts in Tmc1Δ/Δ;Tmc2Δ/Δ mice lacking sensory transduction are similar to those in wild-type (WT) mice at postnatal day 2 (P2).

https://cdn.elifesciences.org/articles/69433/elife-69433-fig1-data1-v2.xlsx

To assess the consequences of Tmc1 and Tmc2 deletion on synaptic development, ribbon synapses were further evaluated at P7, P14, and P28 in WT and Tmc1Δ/Δ;Tmc2Δ/Δ mice. These three timepoints were selected to encompass events crucial to the maturation of the inner ear with P7 following acquisition and maturation of IHC sensory transduction, P14 following hearing onset in mice, and P28 reflecting the nearly mature organ of Corti and auditory function. For each timepoint and cochlear region, the average number of synapses per IHC was estimated by counting the total number of presynaptic CtBP2 juxtaposed to postsynaptic GluA2 puncta, divided by the number of IHCs sampled in each region (Figure 2).

Synapse counts are elevated at postnatal day 7 (P7) and reduced at P28 in Tmc1Δ/Δ;Tmc2Δ/Δ mice relative to wild-type (WT) mice.

(A–B) Representative image of P7 WT and Tmc1Δ/Δ;Tmc2Δ/Δ inner hair cells (IHCs) from 16 kHz region immunostained for anti-Myosin7a (gray), anti-CtBP2 (red), and anti-GluA2 (green). Higher magnification images are shown below. (C–D) P28 WT and Tmc1Δ/Δ;Tmc2Δ/Δ IHCs from 16 kHz region. Scale bars: 10 µm (upper) and 5 µm (lower).

While ribbon counts were unaltered in P2 Tmc1Δ/Δ;Tmc2Δ/Δ mice, the lack of Tmc1 and Tmc2 resulted in elevated synapse counts at P7 across all frequency regions relative to those in WT mice (Figure 3A, Supplementary file 2A). There were ~43% more synapses in Tmc1Δ/Δ;Tmc2Δ/Δ mice, suggesting there may be a correlation between the developmental acquisition of sensory transduction in IHCs during the first postnatal week and the developmental increase in the number of ribbons and postsynaptic densities. In WT mice, ~50% of ribbon synapses are lost between the end of the first and second postnatal weeks (Sundaresan et al., 2016). This decrease is thought to result from pruning, refinement, and fusion of ribbons and postsynaptic densities (Michanski et al., 2019; Sundaresan et al., 2016; Wong et al., 2014; Huang et al., 2012; Sendin et al., 2007). Consistent with these findings, we found a 46–57% reduction in synapses counts in WT mice between P7 and P14 between 8 and 32 kHz. In Tmc1Δ/Δ;Tmc2Δ/Δ mice, a more drastic decrease in synapse counts was evident during the second postnatal week. The number of synapses decreased by 67 to 73% at P14 relative to P7 between 8 and 32 kHz (Figure 3A and B). As a result, synapse numbers at P14 did not differ significantly between WT and Tmc1Δ/Δ;Tmc2Δ/Δ groups except at the 32 kHz region where Tmc1Δ/Δ;Tmc2Δ/Δ mice exhibited ~27% fewer synapse counts (Figure 3B, Supplementary file 2B).

Synapse counts were elevated at postnatal day 7 (P7) in the absence of both Tmc1 and Tmc2 and diminished at P28 in the absence of Tmc1 and Tmc2 or Tmc1 alone.

(A–C) The mean number of synapses/inner hair cell (IHC) was calculated for each frequency region. Data from wild-type (WT) (black), Tmc1Δ/Δ;Tmc2Δ/Δ (red), Tmc1Δ/Δ (blue), and Tmc2Δ/Δ (dark yellow) groups are shown. Individual points represent counts from one mouse. Temporal changes in synapse counts differed by genotype (two-way ANOVA; p < 0.001 based on two-way interaction between genotype and timepoints for all frequencies; Supplementary file 3A). However, genotype-specific trajectories of synaptic development did not vary by frequency (three-way ANOVA; p = 0.73 based on three-way interaction between group, time, frequency; Supplementary file 3A). Bolded lines depict mean ± SD. Black horizontal bars and asterisks represent statistically significant differences between group means (multiple pairwise comparisons, *p < 0.05, **p < 0.01, ***p < 0.001; p values are listed in Supplementary file 2A-C). Number of cochleas: 4–9 WT, 5–12 Tmc1Δ/Δ;Tmc2Δ/Δ, 4 Tmc1Δ/Δ, 4–5 Tmc2Δ/Δ.

Figure 3—source data 1

Synapse counts were elevated at postnatal day 7 (P7) in the absence of both Tmc1 and Tmc2 and diminished at P28 in the absence of Tmc1 and Tmc2 or Tmc1 alone.

https://cdn.elifesciences.org/articles/69433/elife-69433-fig3-data1-v2.xlsx

Following hearing onset in WT mice, ribbon synapse counts remain stable into adulthood (Michanski et al., 2019; Huang et al., 2012). As expected, our WT synapse counts did not differ significantly between P14 and P28 in any of the frequency regions examined (Figure 3B and C). However, synapse counts were markedly decreased in P28 Tmc1Δ/Δ;Tmc2Δ/Δ mice relative to WT mice and relative to P14 Tmc1Δ/Δ;Tmc2Δ/Δ numbers at all frequencies (Figure 3C, Supplementary file 2C). Though synapse counts remain stable in WT mice post-hearing onset, changes in the distribution of ribbon sizes and continued reduction in the sizes of presynaptic/postsynaptic densities are observed until their full development around P34 (Payne et al., 2021). As with the abnormally elevated synapse counts at P7 in Tmc1Δ/Δ;Tmc2Δ/Δ mice, the rapid loss of more mature ribbon synapses further suggests a role for sensory transduction in the development of synapses.

Synapse counts in Tmc single knockouts differ from those of double knockouts

Since Tmc2 expression coincides with the developmental onset of sensory transduction in OHCs (Lelli et al., 2009; Kawashima et al., 2011) and IHCs (Pan et al., 2013) at P0 in the base and P2-P4 in the apex and is followed several days later by expression of Tmc1, we wondered whether genetic deletion of Tmc2 would cause a delay in the developmental pattern of synapse development. In addition, to determine whether the changes observed in Tmc1Δ/Δ;Tmc2Δ/Δ synapses were due to deletion of Tmc1, we examined synapses in single knockouts of Tmc1 or Tmc2. At P7 and P14, synapse counts in neither Tmc1Δ/Δ nor Tmc2Δ/Δ mice differed significantly from those of WT mice across all frequencies (Figure 3A and B, Supplementary file 2A and B), suggesting that expression of either Tmc1 or Tmc2 was sufficient for acquisition of normal synapse counts and juxtaposition of presynaptic ribbons and their postsynaptic densities. However, it remains possible that synaptic function and development are altered in mice lacking Tmc1, Tmc2, or both in ways that cannot be captured by immunohistochemistry and synapse counts alone.

At P28, there was a significant deviation in the number of synapses observed between Tmc1Δ/Δ and Tmc2Δ/Δ groups. Synapse counts in Tmc1Δ/Δ mice were significantly reduced compared to WT mice and similar to those of Tmc1Δ/Δ;Tmc2Δ/Δ mice at all frequencies (Figure 3C, Supplementary file 2C). Synapse counts in Tmc2Δ/Δ mice, on the other hand, were not different from those in WT mice at any of the frequencies (Figure 3C, Supplementary file 2C). Thus, it appears the absence of Tmc1 specifically, in Tmc1Δ/Δ and Tmc1Δ/Δ;Tmc2Δ/Δ mice, accounts for the loss of synapses after hearing onset.

Tmie deletion leads to loss of synapses

While the results obtained from Tmc1Δ/Δ, Tmc2Δ/Δ, and Tmc1Δ/Δ;Tmc2Δ/Δ mice implicated a role for sensory transduction in the development and maturation of ribbon synapses, the possibility of ribbon synapses being affected by targeted deletion of Tmc1, Tmc2, or both via a mechanism unrelated to inhibition of sensory transduction remained a possibility. To determine whether the change in synapse counts was a specific consequence of Tmc deletion or a general consequence of the loss of sensory transduction, we investigated deletion of a different gene known to cause loss of hair cell sensory transduction. Synapses were counted in Spinner mice, which carry a spontaneous nonsense mutation at the Tmie locus (Tmiesr) (Mitchem et al., 2002). TMIE encodes transmembrane inner ear protein, which is an essential component of the transduction channel complex (Gleason et al., 2009; Shen et al., 2008; Zhao et al., 2014). While loss of functional TMIE in homozygous Spinner mice causes complete absence of sensory transduction (Zhao et al., 2014) and mislocalization of TMC proteins (Pacentine and Nicolson, 2019; Cunningham et al., 2020), it does not affect TMC expression (Cunningham et al., 2020). As shown in Figure 4A, Tmiesr synapse counts at P7 were significantly elevated compared to WT mice. There were no significant differences between Tmiesr and Tmc1Δ/Δ;Tmc2Δ/Δ synapse counts in any frequency region (Figure 4A, Supplementary file 4A). At P28, Tmiesr mice showed decreased synapse counts at all frequencies, with values that did not differ significantly from those in Tmc1Δ/Δ;Tmc2Δ/Δ mice (Figure 4B, Supplementary file 4B). The similarities in synapse counts in Tmc1Δ/Δ;Tmc2Δ/Δ and Tmiesr mice at P7 and P28 provide compelling evidence that the loss of normal synapse development and maturation is a consequence of the absence of sensory transduction and not disruption of a specific gene.

Loss of sensory transduction, not of Tmc1 and Tmc2 specifically, results in the synaptic differences observed at postnatal day 7 (P7) and P28.

(A–C) The mean number of synapses/inner hair cell (IHC) at each frequency region. Data from wild-type (WT) (black), Tmc1Δ/Δ;Tmc2Δ/Δ (red), Tmc1Δ/Δ (blue), Tmiesr (purple), and Tmc1Bth (gold) groups are shown. WT, Tmc1Δ/Δ, and Tmc1Δ/Δ;Tmc2Δ/Δ data are the same as those depicted in Figure 3, reprinted here to facilitate comparison. Individual points represent counts from one mouse. In A and B two-way interactions between genotype and timepoints were statistically significant for all frequencies based on two-way ANOVA (p < 0.001; Supplementary file 3B), suggesting the trajectory of synaptic development varies by genotype. The genotype-specific trajectory of synaptic development did not vary by frequency (three-way ANOVA; p = 0.68 based on three-way interaction between genotype, timepoints, and frequency; Supplementary file 3B). Frequency-specific synapse counts differed by genotype in C (two-way ANOVA; p = 0.002 based on two-way interaction between genotype and frequency; Supplementary file 3B). Bolded lines depict mean ± SD. Black horizontal bars and asterisks represent statistically significant differences in group means (multiple pairwise comparisons, *p < 0.05, **p < 0.01, ***p < 0.001; exact p values listed in Supplementary file 4A-C). Number of cochleas: 4 WT, 5–12 Tmc1Δ/Δ;Tmc2Δ/Δ, 4 Tmc1Δ/Δ, 5–6 Tmiesr, 4–6 Tmc1Bth.

Figure 4—source data 1

Loss of sensory transduction, not of Tmc1 and Tmc2 specifically, results in the synaptic differences observed at postnatal day 7 (P7) and P28.

https://cdn.elifesciences.org/articles/69433/elife-69433-fig4-data1-v2.xlsx

Altered sensory transduction permeability is inconsequential for synapses

To explore whether the changes in synapses were the result of calcium entry through sensory transduction channels, we performed a similar analysis in Beethoven mice with a dominant mutation in Tmc1. IHCs in Tmc1Bth mice (Vreugde et al., 2002) have normal sensory transduction current amplitudes, but reduced calcium permeability relative to control IHCs (Pan et al., 2013; Beurg et al., 2015; Corns et al., 2016). Marcotti et al., 2006 reported Tmc1Bth hair cells have normal resting potentials but altered basolateral currents, leading Pan et al., 2013 to speculate that altered calcium permeability in Tmc1Bth hair cell may have consequences for basolateral hair cell functions, perhaps including synapse development and maturation. No significant differences in P28 synapse counts were evident between Tmc1Bth and WT groups except in the 32 kHz region (Figure 4C, Supplementary file 4C), suggesting reduced calcium entry via sensory transduction channels does not cause a significant loss of ribbon synapses. However, the difference observed in the 32 kHz region suggests a tonotopic role for calcium signaling cannot be discounted.

Restoration of sensory transduction preserves auditory synapses

Next, we wondered whether gene therapy restoration of sensory transduction could prevent synapse loss. Recent inner ear gene therapy studies have demonstrated the efficacy of a novel utricle injection technique and a synthetic AAV9-PHP.B capsid for transducing nearly 100% of cochlear hair cells with high specificity (Lee et al., 2020) and restoring ABR thresholds with WT Tmc1 injected into Tmc1Δ/Δ mouse inner ears (Wu et al., 2021). Although AAV-mediated gene therapy for otoferlin, a synaptic protein, has demonstrated recovery of auditory synapses (Akil et al., 2019; Al-Moyed et al., 2019), the consequences of Tmc1 gene therapy on auditory synapses has not been assessed. To characterize the consequences of Tmc1 gene therapy on IHC synapses, 12 Tmc1Δ/Δ mice were injected at P1 with AAV9-PHP.B-Tmc1 via the utricle and ABRs/synapses were assessed at P28.

At P28, Tmc1Δ/Δ mice are profoundly deaf with no measurable ABR thresholds up to 110 dB sound pressure level (SPL) (Wu et al., 2021). All 12 Tmc1Δ/Δ mice that were injected showed some degree of improvement in auditory function relative to uninjected Tmc1Δ/Δ mice (Figure 5A and B, Supplementary file 5). Recovery was variable across mice, but consistently greater at lower frequencies. On average, the injected mice showed a recovery of 59 dB SPL from 8 to 16 kHz (8 kHz: 61.7 ± 18.3 dB SPL; 11 kHz: 57.9 ± 18.4 dB SPL; 16 kHz: 56.3 ± 25.3 dB SPL). The four best performers out of the 12 injected mice demonstrated mean thresholds of 39 dB SPL across 8–16 kHz (Figure 5B, Supplementary file 5, 8 kHz: 46.3 ± 6.3 dB SPL; 11 kHz: 40.0 ± 7.1 dB SPL; 16 kHz: 31.3 ± 7.5 dB SPL).

AAV9-PHP.B-Tmc1 restores ABR thresholds in Tmc1Δ/Δ mice.

(A) Representative ABR waveforms recorded at postnatal day 28 (P28) using 16 kHz tone bursts at sound pressure levels of increasing 5 dB increments. Waveforms from uninjected Tmc1Δ/Δ mouse (left), three Tmc1Δ/Δ mice injected with AAV9-PHP.B-Tmc1, representing best (green), average (blue), and worst (red) recovery and one wild-type (WT) control (black) are shown. Thresholds determined by the presence of Peak 1 and indicated by bolded, colored traces. (B) ABR thresholds plotted as a function of stimulus frequency for 12 Tmc1Δ/Δ mice injected with AAV9-PHP.B-Tmc1 tested at P28 (gray traces). Mice with the best (green), median (blue), and worst (red) recovery are indicated and correspond to the best, average, and worst traces in (A). Black dotted lines show mean ± SD thresholds from eight previously tested WT mice. Green dotted line shows mean ± SD thresholds from the 12 injected mice. (C) Peak 1 amplitudes measured from 16 kHz ABR waveforms (A) for 12 Tmc1Δ/Δ mice injected with AAV9-PHP.B-Tmc1. Colors correspond to conditions indicated in B. (D) Peak 1 latencies measured from 16 kHz ABR waveforms (A) for 12 Tmc1Δ/Δ mice injected with AAV9-PHP.B-Tmc1. Colors correspond to conditions indicated in B.

Figure 5—source data 1

AAV9-PHP.

B-Tmc1 restores ABR thresholds in Tmc1Δ/Δ mice.

https://cdn.elifesciences.org/articles/69433/elife-69433-fig5-data1-v2.xlsx

Following ABR recording, injected cochleas were immediately harvested and stained for synapses. Average synapse counts were greater in injected Tmc1Δ/Δ mice at all frequency regions (11.3 kHz p = 0.008, 16 kHz p = 0.001, 22.6 kHz p = 0.008, 32 kHz p = 0.012) except at 8 kHz (p = 0.377), than in uninjected Tmc1Δ/Δ mice (Figure 6A, Supplementary file 6A). Synapse counts were variable across injected mice, similar to ABR threshold recoveries. However, we did note a correlation between the level of ABR threshold recovery and the number of synapses/IHC. We calculated pure tone average (PTA) thresholds from 8 to 22.6 kHz and plotted the values as a function of average number of IHC synapses across the corresponding cochlear region for 12 mice (Figure 6B). The data were fit with a linear regression (r = 0.86) which indicated a statistically significant correlation (p = 0.00032) with an improvement of PTA threshold by 13 dB/synapse. The increased synapse counts in Tmc1Δ/Δ mice injected with AAV9-PHP.B-Tmc1 provide a novel line of evidence supporting the potential for Tmc1 gene therapy in preserving ribbon synapses, a requirement for normal auditory function.

AAV9-PHP.B-Tmc1 preserves synapse counts and ribbon volume distributions in Tmc1Δ/Δ mice.

(A) The mean number of synapses/inner hair cell (IHC) was counted at each frequency region from postnatal day 28 (P28) wild-type (WT) (black), Tmc1Δ/Δ (blue), and injected Tmc1Δ/Δ (green) mice. WT and Tmc1Δ/Δ ribbon counts are the same as those depicted in Figure 3. Individual points represent counts from one mouse. Frequency-specific synapse counts did not differ by group (two-way ANOVA; p = 0.49 based on two-way interaction between genotype and frequency; Supplementary file 3C). Bolded lines indicate mean ± SD. Black horizontal bars and asterisks represent statistically significant differences in group means (multiple pairwise comparisons, *p < 0.05, **p < 0.01, ***p < 0.001). Number of cochleas: 4 WT, 4 Tmc1Δ/Δ cochleas, 11–12 injected Tmc1Δ/Δ cochleas. (B) Pure tone average (PTA) thresholds were calculated for frequencies between 8 and 22.6 kHz for 12 Tmc1Δ/Δ mice injected with 1 µL AAV9-PHP.B-Tmc1 at P1. PTA thresholds were plotted as a function of the mean synapses/IHC (circles) based on synapse counts from corresponding cochlear regions. The data were fitted with a linear regression (red line) that had a slope of 13 dB/synapse, a correlation coefficient of 0.86; p = 0.00032. (C) Histograms showing distributions of ribbon volumes from confocal z-stacks plotted for each frequency region. Scale bars indicate volume counts on the Y-axis and ribbon volumes in µm3 on the X-axis. Data were obtained from four P28 WT (gray) cochleas, four Tmc1Δ/Δ (blue) cochleas, and four injected Tmc1Δ/Δ (green) cochleas for each of the 8–32 kHz regions.

Figure 6—source data 1

AAV9-PHP.

B-Tmc1 preserves synapse counts and ribbon volume distributions in Tmc1Δ/Δ mice.

https://cdn.elifesciences.org/articles/69433/elife-69433-fig6-data1-v2.xlsx

Lastly, we wondered whether the distribution of ribbon volumes was altered by loss and recovery of sensory transduction. Confocal z-stacks of cochleas stained with GluA2 and CtBP2 were acquired using Zeiss Airyscan from four WT mice, four uninjected Tmc1Δ/Δ mice, and four injected Tmc1Δ/Δ mice with the best ABR threshold recovery and synapse count preservation. For each z-stack, ribbon volumes were calculated using the ‘Surfaces’ module in Imaris.

The distributions of ribbon volumes are illustrated in Figure 6C. Standard deviations of ribbon volumes were compared to evaluate differences in variation between the three groups. P28 uninjected Tmc1Δ/Δ mice showed larger distributions of ribbon volumes than WT mice at the same age (Supplementary file 6B). These differences were statistically significant at all five frequency regions (Supplementary file 6C). Injected Tmc1Δ/Δ mice showed narrower distributions more similar to those in WT mice, suggesting Tmc1 gene therapy preserves both synapse counts and the distribution of ribbon volumes. However, the standard deviations of injected mice were not significantly different from those of uninjected mice (Supplementary file 6C).

Discussion

IHC synapses undergo significant developmental changes during the first few postnatal weeks (Sobkowicz et al., 1982; Huang et al., 2012; Yu and Goodrich, 2014; Michanski et al., 2019; Voorn and Vogl, 2020). Sensory transduction is also maturing during this time frame and is required for maturation and maintenance of IHC morphology and a number of biophysical properties (Corns et al., 2018). Here, we report that sensory transduction also contributes to the development and maturation of IHC synapses during the first postnatal month.

We began with Tmc1Δ/Δ;Tmc2Δ/Δ mice, which fail to acquire sensory transduction (Kawashima et al., 2011), and found they had a similar number of CtBP2-positive puntca to WT mice at P2. This was consistent with prior work suggesting that formation of presynaptic ribbon complexes is an intrinsic property of immature hair cells (Sobkowicz et al., 1986). Despite the similarities at P2, synapse counts in Tmc1Δ/Δ;Tmc2Δ/Δ mice were significantly elevated at P7 relative to WT mice (Figure 7). These unexpected results are the first to show elevated synapse counts in neonatal mice lacking sensory transduction and suggest that acquisition of sensory transduction may help regulate synaptic pruning and proper synaptic numbers.

Developmental changes in inner hair cell (IHC) synapses in genetic models of sensory transduction dysfunction.

(A) Number of synapses/IHC from the 32 kHz region of wild-type (WT), Tmc1Δ/Δ;Tmc2Δ/Δ, Tmc1Δ/Δ, and Tmc2Δ/Δ cochleas as a function of age. (B) Schematic diagram summarizing changes in IHC synapses between postnatal day 7 (P7) and P28 in five genetic models.

In WT mice, the number of synapses progressively increases and then undergoes a reduction during the second postnatal week until the onset of hearing (Sundaresan et al., 2016; Wong et al., 2014; Huang et al., 2012; Sendin et al., 2007). While the reduction of synapses still occurred in our Tmc1Δ/Δ;Tmc2Δ/Δ mice, the lack of sensory transduction led to a larger decline in the number of synapses by P14. Tmc1Δ/Δ;Tmc2Δ/Δ mice also displayed reduced synapse counts at P28, in a manner similar to that reported in previous studies assessing synaptic consequences of genetic disruption of synaptic transmission. For example, mice lacking Cav1.3 Ca2+ channels, which are critical for calcium-dependent neurotransmission at IHC synapses (Platzer et al., 2000; Brandt et al., 2005), lost IHC synapses 4 weeks after birth (Nemzou N. et al., 2006; Brandt et al., 2003). Another study assessed the synaptic consequences of genetic deletion of vesicular glutamate transporter type 3 (Kim et al., 2019) and found roughly half of IHC synapses were lost by 9 weeks of age. Although the genetic targets are different, disruption of synaptic proteins or transduction proteins lead to similar loss of IHC-SGN synapses suggesting they may converge upon similar molecular pathways that regulate synaptic development and maturation.

Genetic deletion of Tmc1 or Tmc2 alone did not alter synapse counts prior to hearing onset; Tmc1Δ/Δ and Tmc2Δ/Δ single KO mice exhibited similar counts to those of WT mice at P7. At P28, Tmc1Δ/Δ mice synapse counts were reduced like those in Tmc1Δ/Δ;Tmc2Δ/Δ mice, while Tmc2Δ/Δ synapses were normal (Figure 7B). These differences between Tmc1Δ/Δ and Tmc2Δ/Δ mice are consistent with a developmental shift from Tmc2 to Tmc1 mRNA and TMC protein expression that occurs in the cochlea (Kawashima et al., 2011; Kurima et al., 2015). Tmc2 is transiently expressed in the cochlea until P8, while a rise in Tmc1 expression is observed between P2 and P22 (Kawashima et al., 2011). At P7, synapse counts in Tmc1Δ/Δ and Tmc2Δ/Δ single KO mice are similar to those of WT mice because both are expressed at that stage and thus available to compensate for the absence of the other. Only in the absence of both were significantly elevated synapse counts observed at P7. At P28, Tmc2 is no longer expressed and is thus unavailable to compensate for the absence of Tmc1 in Tmc1Δ/Δ mice. Thus, Tmc1Δ/Δ mice lack sensory transduction, are profoundly deaf, and we find, exhibit similar decreases in synapse counts as Tmc1Δ/Δ;Tmc2Δ/Δ mice at P28. The absence of native Tmc2 expression after P8 also explains why Tmc2Δ/Δ mice are phenotypically normal and have synapse counts similar to WT mice at more mature stages.

To verify the synaptic changes observed at P7 and P28 in the absence of Tmc1 and Tmc2 were due to the loss of sensory transduction, synapse counts were also assessed in Tmiesr mice and Tmc1Bth mice. At both P7 and P28, synapse counts in Tmiesr and Tmc1Δ/Δ;Tmc2Δ/Δ double KO mice were remarkably similar (Figure 7), suggesting sensory transduction in general, rather than Tmc1 or Tmc2 specifically, helps drive normal development and maturation of IHC synapses. Indeed, while localization of TMC proteins may be disrupted in the absence of TMIE, TMC expression level is not affected (Cunningham et al., 2020). Though P28 synapse counts only differed significantly from WT mice at 32 kHz, Tmc1Bth synapse counts were not consistently normal. Some mice exhibited counts that were lower than those in WT mice, but greater than those in Tmc1Δ/Δ mice. The differences in Tmc1Bth mice suggest calcium permeability may play a subtle role in regulating synapse numbers. Additional studies assessing the relative contributions of calcium entry via sensory transduction channels will be necessary.

We predict that loss of sensory transduction in Tmiesr, Tmc1Δ/Δ and Tmc1Δ/Δ;Tmc2Δ/Δ mice may lead to hyperpolarization of IHC resting potentials and a lack of depolarizing receptor potentials, leaving voltage-gated Cav1.3 channels largely deactivated and glutamatergic neurotransmission attenuated. While the hair cell sensory signaling cascade is well established, secondary consequences of disrupted sensory signaling remain obscure. Our data suggest a mechanistic connection between sensory and molecular signaling pathways in the auditory periphery. We propose that sensory signals, in addition to relaying auditory information to the brain, may also converge with molecular signaling pathways that govern development and maturation of IHC-SGN synapses. Disruption of sensory signals, whether genetic in origin or via environmental insults or aging, may disrupt a common pathway leading to synaptopathy and auditory dysfunction. While a number of studies have documented both pathological and normal developmental changes in IHC synapses (Yu and Goodrich, 2014; Voorn and Vogl, 2020; Sobkowicz et al., 1982; Nouvian et al., 2006; Moser et al., 2006; Michanski et al., 2019; Sergeyenko et al., 2013; Fernandez et al., 2020; Kujawa and Liberman, 2015), the molecular mechanisms that regulate development and maturation of IHC synapses and their afferent connections remain unclear.

To further investigate the relationship between sensory transduction and IHC synapses, we discovered that Tmc1 gene therapy in Tmc1Δ/Δ mice promoted restoration of sensory transduction, and consequently, preserved synapse counts and ribbon volume distributions. However, variability in both ABR threshold recovery and synapse counts were observed. While the variability in gene therapy recovery is consistent with prior reports (Nist-Lund et al., 2019; Wu et al., 2021), the source is unclear but may be due to variability in viral injection or viral distribution within the cochlea. Regardless, the variability in the extent of recovery permitted analysis of the relationship between ABR threshold recovery and synapse counts. We found a strong correlation between PTA thresholds and average number of synapses in Tmc1Δ/Δ mice injected with Tmc1 gene therapy reagents. Importantly, the data suggest Tmc1 gene therapy promotes recovery of sensory transduction (Nist-Lund et al., 2019) and preservation of synapses, both of which are necessary for normal auditory function. Whether Tmc1 gene therapy introduced at more mature stages can prevent or promote recovery from synaptopathy remains to be determined.

While our study demonstrates an important role for sensory transduction in development and maturation of IHC-SGN synapses, one caveat is that synapse counts alone are not indicative of the functional status of IHC synapses. Direct assays of synaptic function and synaptic transmission may help validate our results. Verification of normal synapse function, functional sensory transduction, and robust ABR waveforms following inner ear gene therapy may also help inform the continued development of inner ear therapeutics.

This study also raises additional questions regarding the absolute volume of CtBP2-positive ribbons. While our quantification methods allowed for relative comparison of the distribution of ribbon volumes, super resolution microscopy may be better suited to quantify absolute changes in ribbon volumes during development, in the absence of sensory transduction and following gene therapy recovery of sensory transduction. Furthermore, whether synapses are preferentially lost or recovered on the pillar or modiolar side of IHCs remains to be investigated.

In summary, our study provides novel insight into the role of sensory transduction in synapse development and maturation. While glutamate excitotoxicity has been implicated in the acute loss of synapses observed immediately following noise exposure (Liberman and Kujawa, 2017), the mechanisms underlying chronic synaptic changes and neural degeneration of SGNs are unknown. Since noise exposure, aging, and genetic mutations can all cause deficits in sensory transduction, we speculate that damage to the sensory transduction apparatus, regardless of the source of the insult, may affect a common molecular pathway that leads to chronic loss of IHC-SGN synapses and degeneration of SGNs. Identification of these molecular pathways may guide development of future therapeutic strategies that prevent synaptopathy and promote healthy synaptic function.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Genetic reagent (Mus musculus)C57B/L6-Cdh23753A>GDerived from Lentz et al., 2010C57BL6Lentz et al. Dev. Neurobiol (2010)
Genetic reagent (Mus musculus)019146 - B6.129-Tmc1tm1.1Ajg/JAvailable from Jackson Lab, obtained initially from Dr A Griffith (NIH/NIDCD)Tmc1 Targeted (Reporter, Null/Knockout)Kurima et al. Nat Genet. (2002)
Genetic reagent (Mus musculus)019147 - B6.129-Tmc2tm1.1Ajg/JAvailable from Jackson Lab, obtained initially from Dr A Griffith (NIH/NIDCD)Tmc2 Targeted (Reporter, Null/Knockout)Kawashima et al. J Clin Invest (2011)
Genetic reagent (Mus musculus)003929 - BXA4/Pgn-Tmiesr-J/JAvailable from Jax C57BL/6-Tmiesr (‘spinner’) miceSpontaneous mutation in TmieStock No. 000543Mitchem et al. Hum Mol Genet. (2002)
Genetic reagent (Mus musculus)Tmc1BthTmc1Bth/Bth mice were obtained from M Hrabé de Angelis and H Fuchs, Institute of Experimental Genetics, Neuherberg, GermanyPoint mutation at residue 412 (M412K)Vreugde et al. Nat Genet. (2002)
AntibodyAnti-CtBP2 (Mouse IgG1 monoclonal)BD Transduction LaboratoriesCat #: 612044Primary antibody, IF (1:200)
AntibodyAnti-GluA2 (Mouse IgG2a monoclonal)Millipore SigmaCat #: MABN1189Primary antibody, IF (1:2000)
AntibodyAnti-MyosinVIIA (Rabbit polyclonal)Proteus BiosciencesCat #: 25–6790Primary antibody, IF (1:200)
AntibodyAnti-Rabbit Alexa Fluor 647 (Donkey polyclonal)Thermo Fisher ScientificCat #: A-31573Secondary antibody, IF (1:200)
AntibodyAnti-Mouse IgG2a Alexa Fluor 488 (Goat polyclonal)Thermo Fisher ScientificCat #: A-21131Secondary antibody, IF (1:1000)
AntibodyAnti-Mouse IgG1 Alexa Fluor 546 (Goat polyclonal)Thermo Fisher ScientificCat #: A-21123Secondary antibody, IF (1:1000)
OtherVectashield AntifadeVector LaboratoriesCat #: H-1000–10Mounting medium
OtherAAV9-PHP.B- CMV-Tmc1e × 1Wu et al., 2021Nucleic acid, Titer: 3.9 E + 13 gc/mL
Software, algorithmImarishttps://imaris.oxinst.com/
Software, algorithmImageJ softwarehttp://imagej.nih.gov/ij/
Software, algorithmEaton-Peabody Laboratories Cochlear Function Test Suitehttps://www.masseyeandear.org/research/otolaryngology/eaton-peabody-laboratories/engineering-core

Mice

WT control mice were C57B/L6 – Cdh23753A>G with a corrected ahl allele as described by Lentz et al., 2010. Tmc mutant mice carried mutant alleles of Tmc1, Tmc2, or both on a C57BL/6J background (Tmc1Δ/Δ, Tmc2Δ/Δ, Tmc1Δ/Δ;Tmc2Δ/Δ) (Vreugde et al., 2002; Kawashima et al., 2011). Spinner mice carrying a spontaneous mutation at the Tmie locus (Tmiesr) on C57BL/6J backgrounds were obtained from Jackson Laboratories. Beethoven mice carrying a dominant mutation in Tmc1Bth associated with DFNA36 in humans were initially donated by Martin Hrabé de Angelis and Helmut Fuchs at the University of Munich. Genotyping was performed as previously described (Kawashima et al., 2011; Mitchem et al., 2002). Mice ages P2, P7, P14, and P28 were used for ribbon synapse characterizations. Tmc1Δ/Δ mice ages P0-P1 were used for in vivo delivery of AAV vectors. Mice of both sexes were used in similar numbers and in accordance with protocols approved by the Institutional Animal Care and Use Committee (Protocols #20-02-4149R and #00001240) at Boston Children’s Hospital.

Viral vector preparation

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Tmc1ex1 was cloned into an AAV2 vector driven by a CMV promoter and followed with a woodchuck hepatitis virus post-transcriptional regulatory element (WPRE) site, as previously described (Wu et al., 2021). The AAV2 vector was then packaged into the AAV9-PHP.B capsid by the Viral Core at Boston Children’s Hospital and purified by iodixanol gradient ultracentrifuge followed by ion-exchange chromatography. The titer of genome-containing particles for the AAV2-PHP.B vector was determined using TaqMan quantitative PCR to detect amplicons located in inverted terminal repeats, as previously described (D’Costa et al., 2016). The titer of the AAV2/9-PHP.B-CMV-Tmc1ex1 WPRE was calculated to be 3.91E+13 gc/mL. The vector was aliquoted, stored at –80°C, and thawed immediately before use. Generation and use of AAV vectors were approved by the BCH Institutional Biosafety Committee (Protocol #IBC-P00000447).

Inner ear injections

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Utricle injections were approved by the Institutional Animal Care and Use Committees at BCH (Protocols #20-02-4149R and #00001240) and performed as previously described (Lee et al., 2020). Briefly, P1 mice were anesthetized with hypothermia and a postauricular incision was made to expose the semicircular canals. A small puncture into the temporal bone surrounding the utricle was made and a glass micropipette was inserted into the puncture to manually inject 1 µL AAV. After the injection, standard postoperative care was applied.

ABR acquisition

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ABR recordings were performed, as previously described (Nist-Lund et al., 2019). Mice were anesthetized with 0.5 mg of ketamine and 0.15 mg of xylazine per 10 g body weight via intraperitoneal injection. Subcutaneous needle electrodes were inserted at the vertex (reference electrode), pinna (active electrode), and rump (ground electrode). Acoustic stimuli were delivered directly into the ear through a custom probe tube speaker/microphone system (EPL PXI Systems) consisting of two electrostatic earphones (CUI Miniature Dynamics) to generate primary tones and a Knowles microphone (Electret Condenser) to record sound pressures from the ear canal. In a sound-proof chamber, mice were presented 5 ms pure tone stimuli of 8, 11.3, 16, 22.6, and 32 kHz at SPL of 10–115 dB in 5 dB increment steps; 512 responses of alternating stimulus polarity were collected and averaged for each SPL. ABR potentials were amplified (10,000×), band-pass filtered (0.3–10 kHz), and digitized using custom data acquisition software from the Eaton-Peabody Laboratories Cochlear Function Test Suite.

Waveforms with peak to trough amplitudes greater than 15 µV were discarded by an artifact-reject function. Sound stimuli and electrode voltages were sampled at 40 µs increments using a National Instruments digital input-output board and stored for offline analyses.

Tissue dissection, immunohistochemistry, and imaging

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Temporal bones were dissected and fixed in 4% paraformaldehyde for 1 hr at room temperature. Temporal bones were then decalcified in 120 mM EDTA for 2 hr for 7-day-old mice and up to 20 hr for 4-week-old mice. Following decalcification, the entire length of the organ of Corti was microdissected in PBS for whole-mount processing. Tissues were then permeabilized by freezing on dry ice in 30% sucrose and blocked for 1 hr at room temperature in PBS with 0.3% Triton X + 5% normal horse serum. Tissues were then stained with the following primary antibodies and incubated at 37°C overnight: (1) mouse isotype IgG1 anti-C-terminal binding protein 2 (CtBP2, 1:200, BD Transduction Laboratories #612044), (2) mouse isotype IgG2a anti-glutamate receptor 2 (GluA2, 1:2000, Millipore #MABN1189), and (3) rabbit anti-myosin VIIa (Myo7a, 1:200: Proteus Biosciences #25–6790). Tissues were washed in PBS and incubated for 2 hr at 37°C with the following secondary antibodies diluted in 1% normal horse serum + 0.3% Triton X: (1) goat anti-mouse IgG1 Alexa Fluor 546 (1:1000, Thermo Fisher #A-21123), (2) goat anti-mouse IgG2a Alexa Fluor 488 (1:1000, Thermo Fisher #A-21131), and (3) donkey anti-rabbit Alexa Fluor 647 (1:200, Thermo Fisher #A-31573). Finally, samples were mounted on glass coverslips with Vectashield mounting medium (Vector Laboratories).

Using the 10× air objective on an LSM 800 (Carl Zeiss), low-power images of the myosin channel were obtained from each microdissected piece. Using a custom ImageJ plugin, a cochlear frequency map was generated by measuring the full length of the cochlea from apex to base using all microdissected pieces (Müller et al., 2005). Full z-stacks were then acquired at cochlear regions corresponding to five frequencies (8, 11.3, 16, 22.6, 32 kHz) using a 63 × 1.4 NA oil objective lens (Carl Zeiss, z step = 0.36 μm, scaling per pixel: 0.068 μm × 0.068 μm); 9–12 IHCs per field were imaged with z-stacks spanning the entire length of the hair cells. For volume estimations, confocal z-stacks were acquired using the 63× oil objective with AiryScan processing (Carl Zeiss, z step = 0.18 μm, scaling per pixel: 0.034 μm × 0.034 μm).

Ribbon synapse counts and volume distribution measurements

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Confocal z-stacks were ported to Imaris, an image analysis software, for creation of 3D projections and quantitative analyses of synapse counts and volumes. The ‘Spots’ module in Imaris was used for automated identification and counting of all ribbons in a given z-stack. All counts were manually reviewed and verified. Synapses were defined as juxtaposition of presynaptic ribbons labeled with anti-CtBP2 with postsynaptic AMPA receptor puncta labeled with anti-GluA2. Juxtaposition was verified manually for every ribbon identified using the ‘Spots’ module. The total number of synapses was divided by the number of IHCs in the image to calculate the average number of synapses/IHC. For estimation of ribbon volume distributions, confocal z-stacks of ribbon synapses were obtained from P28 control, Tmc1Δ/Δ, and Tmc1Δ/Δ mice injected with AAV2/9-PHP.B-CMV- Tmc1ex1WPRE using Airyscan processing and ported to Imaris. 3D projections of Airyscan z-stacks were generated and ribbon volumes were segmented from each projection using the ‘Surfaces’ module in Imaris. Identical settings were applied across image stacks, including the threshold for ‘background subtraction (local contrast)’. This threshold was fixed to ensure digital analyses in Imaris were consistent across images (despite potential variability in immunostaining quality and image acquisition confocal settings) and to prevent subjective biases from affecting ribbon volume calculations. As with the synapse counts, ribbons identified using the ‘Surfaces’ module were manually verified to be juxtaposed with anti-GluA2 staining. Calculated volumes from each z-stack for each mouse were normalized to the median volume of all ribbons in the z-stack to account for differences in immunostaining quality and image acquisition settings across mice, genotypes, and cochlear regions. Standard deviations of normalized ribbon volumes from each z-stack for each mouse were compared to determine if significant differences in ribbon volume distributions were evident between groups.

Experimental design and statistical analyses

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The Wilcoxon rank sum test was used for comparison of P2 WT and Tmc1Δ/Δ;Tmc2Δ/Δ CtBP2+ puncta counts in Figure 1C. For the synapse counts in Figure 3A-C and Figure 4A-C, three-way ANOVAs were first conducted to evaluate whether genotype differences in the trajectory of synaptic development varied by frequency. In Figure 3, the interaction effect between four genotypes (WT, Tmc1Δ/Δ;Tmc2Δ/Δ, Tmc1Δ/Δ, Tmc2Δ/Δ), three timepoints (P7, P14, P28), and five frequency regions (8, 11.3, 16, 22.6, 32 kHz) was evaluated. In Figure 4A–B, the interaction effect between three genotypes (WT, Tmc1Δ/Δ;Tmc2Δ/Δ, Tmiesr), two timepoints (P7, P28), and five frequency regions (8, 11.3, 16, 22.6, 32 kHz) was evaluated. For Figure 3A-C and Figure 4A-B, two-way ANOVAs were also used to examine the effects of genotype on synapse development at each frequency region. For Figures 4 and 6, two-way ANOVAs were used to examine the interaction effect between three genotypes (Figure 4C: WT, Tmc1Δ/Δ, Tmc1Bth; Figure 6A: WT, Tmc1Δ/Δ, injected Tmc1Δ/Δ) and five frequency regions (8, 11.3, 16, 22.6, 32 kHz). Following three-way and two-way ANOVAs, multiple pairwise comparisons were conducted to determine which specific genotype groups differed from one another at each timepoint and frequency region. Six paired comparisons were made between the four groups in Figure 3A–C and three paired comparisons were made between the three groups in Figures 4A–C ,6A. The Bonferroni correction was applied to correct for the multiple comparisons and the reported p values are the original p values multiplied by the number of paired comparisons made. The Wilcoxon rank sum test was also used to compare average ABR thresholds in WT and Tmc1Δ/Δ mice injected with Tmc1 gene therapy in Figure 5B. To evaluate whether the size of ribbon volume distribution differed between control, Tmc1Δ/Δ and Tmc1Δ/Δ mice injected with Tmc1 gene therapy, standard deviations of normalized ribbon volumes from each z-stack were compared using Kruskal-Wallis tests followed by Dunn’s multiple comparisons tests using Bonferroni correction. Exact p values are reported. Statistical analyses were performed in GraphPad Prism, R, and SAS statistical software. Figures were created using OriginLab, OriginPro.

Data availability

All data generated or analysed during this study are included in the manuscript and supporting files. Original raw data files have been uploaded to Dryad and are freely available here: https://doi.org/10.5061/dryad.fxpnvx0sb.

The following data sets were generated
    1. Lee J
    2. Kawai K
    3. Holt J
    (2021) Dryad Digital Repository
    Data from: Sensory transduction is required for normal development and maturation ofcochlear inner hair cell synapses.
    https://doi.org/10.5061/dryad.fxpnvx0sb

References

Decision letter

  1. Tobias Reichenbach
    Reviewing Editor; Friedrich-Alexander-University (FAU) Erlangen-Nürnberg, Germany
  2. Barbara G Shinn-Cunningham
    Senior Editor; Carnegie Mellon University, United States
  3. Elisabeth Glowatzki
    Reviewer; Johns Hopkins University School of Medicine, United States
  4. Mark A Rutherford
    Reviewer; Washington University at St. Louis, United States
  5. Andreas Neef
    Reviewer; Max Planck Institute for Dynamics and Self-Organization, Germany

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Decision letter after peer review:

Thank you for submitting your article "Sensory Transduction is Required for Normal Development and Maintenance of Cochlear Inner Hair Cell Synapses" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Barbara Shinn-Cunningham as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Elisabeth Glowatzki (Reviewer #1); Mark A Rutherford (Reviewer #2); Andreas Neef (Reviewer #3).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission.

All reviewers found that the study is overall well executed and that the results are of significant interest both for auditory neuroscientists as well as potentially for the broader neuroscience community. However, the reviewers also raised a number of criticisms and comments that are listed below.

Figure 6:

There are some issues with panel C. When normalizing to the median, the distribution should be around the value of 1. Clearly, something else was done here but it is not clear what was done. Let's step back for a minute. The reason why Charlie normalizes to the median is two fold – (1) because of the difference in brightness across images, as you stated, and (2) because all of his comparisons are between modiolar and pillar groups in the same images, pooled for different images from the same group. Comparisons between groups are made only in relative terms. Here, you could compare modiolar and pillar and perhaps you should, since the data are already in hand. However, here you are not comparing modiolar and pillar, so, why are you normalizing to the median? The reason to normalize to the median in this case would be to compare the shapes of the distributions, which is basically what you described – a change in the shape where the dKO has a broader distribution. Please mention that you are not detecting potential difference in absolute volume, only differences in the shape of the distribution. Comparing modiolar and pillar would strengthen the paper as well. Another appropriate addition would be to look at GluA2 volumes as well.

The conclusion "suggesting Tmc1 gene therapy preserves both synapse counts and ribbon morphology" should not be based on distributions of normalized ribbon volumes, but absolute ribbon volumes. Is there no better way to address the question? Why is the volume estimate so strongly dependent on the staining? Does Imaris not use a threshold adapted to the dynamic range of the data? What do the raw data look like?

Statistical analysis:

The scarce information about the statistical evaluation is contradictory and unclear:

1. The combination of "assumed to be not normally distributed and "use ANOVA" could be disputed. The assumption of ANOVA is normal distribution, but it may work with samples drawn from distributions violating this assumption.

2. The sentence on line 549 indicates the use of multiple ANOVAs. This probably references the use of one ANOVA for Figure 3 WT vs double mut and one ANOVA across all variants in figure 3 and one ANOVA for figure 4. It would be much clearer if the authors stated that precisely and also made clear that the correction was done for triple testing and also state that the reported p values are one-third the original ones – just to be clear.

3. line 551: "Wilcoxon matched-pairs signed rank test was used for comparison of P2 WT and Tmc1Δ/Δ;Tmc2Δ/Δ ribbon counts".

The source of the "matched" is unclear to me. Obviously, within each WT group and each mutant group, data at different cochlear sections are matched. However, the comparison reported is between WT and mutant across all frequency bands. The data are not matched across the two groups, and therefore it is not clear why the matched-pairs test was warranted.

The statistical description ends as it begins, only that here the word "Gaussian" replaces "normal". Again, it is unclear what is meant by this and why the assumption "Gaussian" is associated with Wilcoxon tests. There is the faint possibility that the authors refer to a specific detail of the implementation of Wilcoxon rank tests. Sometimes, the test uses the assumption of a gaussian distribution of the test statistics to evaluate the probability of the observed rank counts much more quickly. In some programs, this behaviour can be toggled by a parameter switch. Should the authors indeed have this "assumption of Gaussian distribution" in mind, they can safely remove the sentence. For sample sizes of 9 this is never invoked. The speed improvement only kicks in for larger sample sizes. Should they refer to some other assumption, it was not clear.

Given the low number of independent data points, the authors might want to include a statement about the certainty with which they report "no difference". A possibility would be an effect size range they can safely exclude at 90% statistical power.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Sensory Transduction is Required for Normal Development and Maturation of Cochlear Inner Hair Cell Synapses" for further consideration by eLife. Your revised article has been evaluated by Barbara Shinn-Cunningham (Senior Editor) and a Reviewing Editor.

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined below:

Required changes:

1) Figure 6C:

– Maybe it is the log scale that is confusing. Please confirm, for each violin plot, that one-half of the data points are above the median of 1 and one-half of the data points are below the median of 1. How does this look on a linear scale (please show in the response to the reviewers, does not need to be included in the revised manuscript)?

– Figure 6C contains very many individual data points, which cannot be visually separated. Those do not contribute to the reader's information. Almost all detectable information lies in the shape of the violin plots. Those indicate Gaussian shapes for all data groups, but they appear Gaussian on a logarithmic scale. The ordinate values of maximum density are between 0.1 and 0.3 for the KO and around 0.8 to 1 for the WT and rescued KO.

This impression has nothing to do with the actual distribution of the data. I have tried very patiently to reproduce anything resembling those violin plots from the provided data for 8kHz, and I failed.

Just how wrong the impression given by the violin plots is, can readily be appreciated from histograms (of the logarithm of the normalized data). The highest density of data in the blue (KO) cloud is above 1, not below. Interestingly, for the accumulated normalized data BEFORE taking the logarithm, the density is highest below 1, because the distribution of the KO data is very skewed (but not for the WT or KOrescue data).

My recommendation: Please find a program, that correctly reproduces density distributions. If in doubt, whether the algorithms will work with the logarithmic scale, please take the logarithm of your data and then use the violin plot. Then plot on a linear scale and create ticks that reflect the logarithmic scaling.

– It is not per se ok to normalize distributions, then to accumulate the results treat the result as if it represents the original distributions (just with less noise). Only if the samples came from the same distribution (plus a linear scaling), this approach will always yield correct conclusions.

The reason named for normalizing is "variability between stainings". Under some strong assumptions (no offset, i.e. no background staining, and most importantly: a LINEAR relation between staining intensity and detected volume) the data distribution could be meaningfully normalized by division by the median. It is not clear to me that those assumptions have been understood and discussed.

– Raw volume values can be surprisingly small. Zeiss states that the Airyscan can at best resolve 0.14 x 0.14 x 0.35 um. An ellipsoid of these dimensions has a volume of 0.0035um^3. Some of the values in the dataset are smaller. That is counter-intuitive.

– On a positive note, I do not think the discussion about normalization and violin plots is discussion is necessary at all. The unnormalized data (at 8kHz), fully support the following claims about the measured volumes:

mean(Tmc1KO) < mean(WT) < mean(Tmc1rescued)

sd(Tmc1KO) > sd(WT) and no significant difference between the variance of WT and Tmc1rescued.

If I understood the current version correctly, the authors argue that the distributions are different. The statistics of the unnormalized data seems to support this claim.

My advice: work with raw data, accumulate across cochleas, plot histograms rather than violin plots and find a way to capture the vastly different shape of the distributions.

2) In response to a previous reviewer critique the authors say: "The Bonferroni correction was applied following these paired comparisons and the reported p values are 1/6 of the original ones." Let's be careful not to confuse people. For the Bonferroni correction, you need to multiply the original p values, not divide them. In other words, you must divide 0.05 by 3 or 6 to obtain the critical p value to get below to obtain significance. I assume this is a typo, and not a realization that changes the results of the statistical tests. In any case, it should be clarified, and the sentence changed.

3) Reporting of statistical Analysis

This will require some work of the authors or external experts. The standards are pretty clear. eLife has supported the push towards more reproducible research and published, among other resources, papers on typical pitfalls and reviews on inadequate reporting:

Meta-Research: Why we need to report more than 'Data were Analyzed by t-tests or ANOVA' Weissgerber et al. 2018 https://doi.org/10.7554/eLife.36163.001

Please read those and change the methods section but also the Results section accordingly. The problem is simply that it is not possible to understand what analysis has been performed, even though the data is available, and all variables are clear to me.

Please report the ANOVA design and results.

I am not a statistician, but here is a quick run-down of how ANOVA results are reported – see for instance the eLife review cited above.

– Naming the model type (mixed model) – that has been done.

– Naming the dimensionality of the model 3 factors with X levels (4 genotype x 3 Times x 5 Frequency ranges), that's very unclear here.

– Naming the significant effects observed with the model(e.g. effect of time, effect of frequency, effect of genotype, interactions: genotype by time, genotype by frequency), again, nothing is reported about which ones were observed.

By my training, only after the ANOVA shows an effect of factor X, the direction of the effect is checked with t-Tests (as the ANOVA already assumes normality).

What is clear from the numbers in the supplemental data is that 90 separate t-Tests were performed to test for the significance of individual pairs of synapse per IHC counts (5 frequencies x 6 genotype combinations x 3 time points). It is necessary to clarify how the relevant correction factor becomes 6 and not 90.

The authors implicitly report an effect of time at P7 we see this, at P28, we see this. Hence an omnibus ANOVA (freq x genotype x time) would have to show an interaction of time and genotype.

As a side-note: Once more it is unclear how the comparisons are "paired" (line 583). In my reading, 4 values from wt are compared to 4 values from the double KO at the same position of the basilar membrane from animals of the same age. That does not make those comparisons "paired". Truly 'paired' are comparisons of data from the same basilar membrane, i.e. counts at 8, 11.3, 16, … kHz. However, it seems the effect of frequency was not tested (which I presume leads to missing the Tmc2 KO vs wt x Frequency interaction).

I would like to suggest that the authors use less conservative ways of multiple-comparison correction (e.g. Holm). On several occasions (e.g. wt vs Tmc2 KO at P7 8 and 11.3 kHz), this would yield significant differences, where currently there is no effect detected.

Interestingly, this inability of Tmc1 to compensate for the Tmc2 KO at the apex seems to be consistent with the gradient of TMC2 --> TMC1 replacement reported in Kawashima 2011.

Suggested changes:

– If not done already, suggest to indicate how many images were analyzed per cochlea and how many total synapses are in each group.

– It appears the statistical unit for assessment of significance is # of cochlea, which I would agree is most appropriate. Suggest to state explicitly.

– On line 344: "However, the differences in standard deviations were not statistically significant relative to those uninjected mice" ….. perhaps better: "However, the standard deviations of injected mice were not significantly different from those of uninjected mice"

– Supplementary Table 5b Prism Stats: If you are going to include this, should there not be some description? It seems like you are now comparing the medians. Is this meaningful? Maybe I missed it, but I don't find any description of this in the text. Your response to reviewers agrees it is not meaningful to compare absolute volumes. When it says, "Do the medians vary signif (P<0.05)?" …. are those medians of volumes or medians of SD's? Please include description of Tables. Again, sorry if I just missed it. If the Supplementary files contain irrelevant things, perhaps the irrelevant things should be removed.

– I found some of the supplemental tables difficult to read, like #2 and #3.

– In the Transparent reporting file, what does it mean to say: "Samples were allocated into groups randomly based on genotype and ages."

– Regarding figure 2, I'm not sure I agree that synapse position "could not be assessed due to abnormal IHC morphology." At least, I don't see it in the paper. Maybe I missed it. If IHC morphology is abnormal in these mice, please mention this in the paper if not already. In the images and the schematics, they look normal. I accept that synapse position and morphology are outside the scope of this work.

- Line 587, "divided" should be "multiplied".

https://doi.org/10.7554/eLife.69433.sa1

Author response

Essential revisions:

Figure 6:

There are some issues with panel C. When normalizing to the median, the distribution should be around the value of 1.

Normalized ribbon volumes in the original Figure 6C were centered around 100% as percentages of the median. As recommended, Figure 6C has been revised so distributions are normalized with the median value set to 1.

Clearly, something else was done here but it is not clear what was done. Let's step back for a minute. The reason why Charlie normalizes to the median is two fold – (1) because of the difference in brightness across images, as you stated, and (2) because all of his comparisons are between modiolar and pillar groups in the same images, pooled for different images from the same group. Comparisons between groups are made only in relative terms. Here, you could compare modiolar and pillar and perhaps you should, since the data are already in hand. However, here you are not comparing modiolar and pillar, so, why are you normalizing to the median? The reason to normalize to the median in this case would be to compare the shapes of the distributions, which is basically what you described – a change in the shape where the dKO has a broader distribution. Please mention that you are not detecting potential difference in absolute volume, only differences in the shape of the distribution. Comparing modiolar and pillar would strengthen the paper as well. Another appropriate addition would be to look at GluA2 volumes as well.

We feel it is inappropriate to consider immunohistochemistry and confocal microscopy as accurate tools for measuring absolute volumes of synaptic elements, including presynaptic ribbons. There is too much variability in tissue quality, immunostaining quality, etc., which prevents accurate comparison of volumes across samples. However, we do feel that it is accurate to measure volumes within a single image and then normalize for each image. This yields a normalized distribution for each image to itself and allows comparison of volume distributions for each image. We show the normalized data, which we feel provide a valid distribution of CtBP2 volumes that can be compared across samples and experimental conditions.

Comparison of modiolar vs pillar synapses was not part of our study design as we had no reason to hypothesize that manipulation of sensory transduction might have different regional effects within a single cell. Future studies utilizing high resolution techniques methods (i.e. STED, STORM, FIB-SEM) may provide more accurate estimations of ribbon volumes and allow for more precise comparisons.

We have expanded our discussion of these points in the revised manuscript.

The conclusion "suggesting Tmc1 gene therapy preserves both synapse counts and ribbon morphology" should not be based on distributions of normalized ribbon volumes, but absolute ribbon volumes. Is there no better way to address the question? Why is the volume estimate so strongly dependent on the staining? Does Imaris not use a threshold adapted to the dynamic range of the data? What do the raw data look like?

We agree and have revised the text to read “suggesting Tmc1 gene therapy preserves both synapse counts and the distribution of ribbon volumes”. As noted above, we feel super resolution microscopy techniques could be used to allow better estimates of absolute ribbon volumes. Those measurements are beyond the scope of the current study but may be well suited for future studies.

Statistical analysis:

The scarce information about the statistical evaluation is contradictory and unclear:

1. The combination of "assumed to be not normally distributed and "use ANOVA" could be disputed. The assumption of ANOVA is normal distribution, but it may work with samples drawn from distributions violating this assumption.

Thank you for the comment. We agree the combination of ANOVA and assumption of non-normal distribution are contradictory. The contradictory sentence was removed from the “Experimental Design and Statistical Analyses” paragraph in the methods section. As detailed in our responses below and clarified in the methods section, ANOVA was used for Figures 3A-C, Figures 4A-C, and Figure 6A. We met with Boston Children’s Hospital’s senior biostatistician, Kosuke Kawai (see acknowledgements) and he confirmed it was reasonable to assume normality for our synapse count datasets. However, for CtBP2+ counts (Figure 1B) and ribbon volumes (Figure 6C), we used non-parametric tests because these data may not be normally distributed.

2. The sentence on line 549 indicates the use of multiple ANOVAs. This probably references the use of one ANOVA for Figure 3 WT vs double mut and one ANOVA across all variants in figure 3 and one ANOVA for figure 4. It would be much clearer if the authors stated that precisely and also made clear that the correction was done for triple testing and also state that the reported p values are one-third the original ones – just to be clear.

Thank you for the suggestion. For each panel in Figure 3 (i.e. each timepoint), a total of six paired comparisons was made using ANOVA to analyze the four groups (i.e. A-B, A-C, A-D, B-C, B-D, C-D). The Bonferroni correction was applied following these paired comparisons and the reported p values are 1/6 of the original ones. In Figure 4 and 6A, 3 paired comparisons were made using ANOVA to analyze the three groups (i.e. A-B, A-C, B-C) so reported p values are 1/3 of the original values. We now describe the statistical tests in more detail in the methods section.

3. line 551: "Wilcoxon matched-pairs signed rank test was used for comparison of P2 WT and Tmc1Δ/Δ;Tmc2Δ/Δ ribbon counts".

The source of the "matched" is unclear to me. Obviously, within each WT group and each mutant group, data at different cochlear sections are matched. However, the comparison reported is between WT and mutant across all frequency bands. The data are not matched across the two groups, and therefore it is not clear why the matched-pairs test was warranted.

Thank you for the comment. We agree the data are not matched across the two groups and the sentence/statistical test were erroneously included in our description. Statistical analyses of the P2 WT vs. Tmc double KO CtBP2+ puncta counts were repeated using the Wilcoxon rank sum test. P values in Supplemental Table 1 were updated and the sentence was also updated in our “Experimental Design and Statistical Analyses” paragraph. No statistically significant differences between the two groups were evident at any of the frequency regions.

The statistical description ends as it begins, only that here the word "Gaussian" replaces "normal". Again, it is unclear what is meant by this and why the assumption "Gaussian" is associated with Wilcoxon tests. There is the faint possibility that the authors refer to a specific detail of the implementation of Wilcoxon rank tests. Sometimes, the test uses the assumption of a gaussian distribution of the test statistics to evaluate the probability of the observed rank counts much more quickly. In some programs, this behaviour can be toggled by a parameter switch. Should the authors indeed have this "assumption of Gaussian distribution" in mind, they can safely remove the sentence. For sample sizes of 9 this is never invoked. The speed improvement only kicks in for larger sample sizes. Should they refer to some other assumption, it was not clear.

Thank you again for the detailed comments/suggestions regarding our statistical analyses. As with the “assumed to be not normally distributed” and “use ANOVA” comment above, we have removed these contradictory sentences from the revised methods sections to clarify the statistical analyses/descriptions.

Given the low number of independent data points, the authors might want to include a statement about the certainty with which they report "no difference". A possibility would be an effect size range they can safely exclude at 90% statistical power.

We recognize that large standard deviations may mask statistically significant differences in means, especially with low numbers of independent data points. We have edited our supplemental tables to reflect the new/updated statistical analysis and to include additional data (i.e. mean, mean difference, SD, SEM, 95% confidence interval, p values). We carefully examined effect size (mean difference and 95% CI) and believe that any significant differences between groups were captured in the statistical analyses.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Essential revisions:

1) Figure 6C:

– Maybe it is the log scale that is confusing. Please confirm, for each violin plot, that one-half of the data points are above the median of 1 and one-half of the data points are below the median of 1. How does this look on a linear scale (please show in the response to the reviewers, does not need to be included in the revised manuscript)?

– Figure 6C contains very many individual data points, which cannot be visually separated. Those do not contribute to the reader's information. Almost all detectable information lies in the shape of the violin plots. Those indicate Gaussian shapes for all data groups, but they appear Gaussian on a logarithmic scale. The ordinate values of maximum density are between 0.1 and 0.3 for the KO and around 0.8 to 1 for the WT and rescued KO.

This impression has nothing to do with the actual distribution of the data. I have tried very patiently to reproduce anything resembling those violin plots from the provided data for 8kHz, and I failed.

Just how wrong the impression given by the violin plots is, can readily be appreciated from histograms (of the logarithm of the normalized data). The highest density of data in the blue (KO) cloud is above 1, not below. Interestingly, for the accumulated normalized data BEFORE taking the logarithm, the density is highest below 1, because the distribution of the KO data is very skewed (but not for the WT or KOrescue data).

My recommendation: Please find a program, that correctly reproduces density distributions. If in doubt, whether the algorithms will work with the logarithmic scale, please take the logarithm of your data and then use the violin plot. Then plot on a linear scale and create ticks that reflect the logarithmic scaling.

– It is not per se ok to normalize distributions, then to accumulate the results treat the result as if it represents the original distributions (just with less noise). Only if the samples came from the same distribution (plus a linear scaling), this approach will always yield correct conclusions.

The reason named for normalizing is "variability between stainings". Under some strong assumptions (no offset, i.e. no background staining, and most importantly: a LINEAR relation between staining intensity and detected volume) the data distribution could be meaningfully normalized by division by the median. It is not clear to me that those assumptions have been understood and discussed.

– Raw volume values can be surprisingly small. Zeiss states that the Airyscan can at best resolve 0.14 x 0.14 x 0.35 um. An ellipsoid of these dimensions has a volume of 0.0035um^3. Some of the values in the dataset are smaller. That is counter-intuitive.

– On a positive note, I do not think the discussion about normalization and violin plots is discussion is necessary at all. The unnormalized data (at 8kHz), fully support the following claims about the measured volumes:

mean(Tmc1KO) < mean(WT) < mean(Tmc1rescued)

sd(Tmc1KO) > sd(WT) and no significant difference between the variance of WT and Tmc1rescued.

If I understood the current version correctly, the authors argue that the distributions are different. The statistics of the unnormalized data seems to support this claim.

My advice: work with raw data, accumulate across cochleas, plot histograms rather than violin plots and find a way to capture the vastly different shape of the distributions.

We appreciate the editor’s attention to this issue. Indeed, we also struggled with how to best represent the data before we settled on the violin plots. To address the editors concerns we replotted the data 24 different ways. In the end, we have followed the editor’s advice and plot the raw data in five histograms, one for each frequency region of the cochlea. We agree this presentation provides the most accurate visual representation of the data for the three conditions, WT, Tmc1 KO and recovery with Tmc1 gene therapy.

2) In response to a previous reviewer critique the authors say: "The Bonferroni correction was applied following these paired comparisons and the reported p values are 1/6 of the original ones." Let's be careful not to confuse people. For the Bonferroni correction, you need to multiply the original p values, not divide them. In other words, you must divide 0.05 by 3 or 6 to obtain the critical p value to get below to obtain significance. I assume this is a typo, and not a realization that changes the results of the statistical tests. In any case, it should be clarified, and the sentence changed.

This was a mistake as noted below in “Suggested changes”. The error was corrected and p values were verified in Prism, R, and SAS. Results of statistical tests were unaltered.

3) Reporting of statistical Analysis

This will require some work of the authors or external experts. The standards are pretty clear. eLife has supported the push towards more reproducible research and published, among other resources, papers on typical pitfalls and reviews on inadequate reporting:

Meta-Research: Why we need to report more than 'Data were Analyzed by t-tests or ANOVA' Weissgerber et al. 2018 https://doi.org/10.7554/eLife.36163.001

Please read those and change the methods section but also the Results section accordingly. The problem is simply that it is not possible to understand what analysis has been performed, even though the data is available, and all variables are clear to me.

Please report the ANOVA design and results.

I am not a statistician, but here is a quick run-down of how ANOVA results are reported – see for instance the eLife review cited above.

– Naming the model type (mixed model) – that has been done.

– Naming the dimensionality of the model 3 factors with X levels (4 genotype x 3 Times x 5 Frequency ranges), that’s very unclear here.

– Naming the significant effects observed with the model(e.g. effect of time, effect of frequency, effect of genotype, interactions: genotype by time, genotype by frequency), again, nothing is reported about which ones were observed.

By my training, only after the ANOVA shows an effect of factor X, the direction of the effect is checked with t-Tests (as the ANOVA already assumes normality).

What is clear from the numbers in the supplemental data is that 90 separate t-Tests were performed to test for the significance of individual pairs of synapse per IHC counts (5 frequencies x 6 genotype combinations x 3 time points). It is necessary to clarify how the relevant correction factor becomes 6 and not 90.

The authors implicitly report an effect of time at P7 we see this, at P28, we see this. Hence an omnibus ANOVA (freq x genotype x time) would have to show an interaction of time and genotype.

The Methods section has been rewritten to describe and report statistical analyses done more accurately. Additional supplemental tables (Supplementary file 6A-6C created to highlight findings of three-way and two-way ANOVAs). A statistician at Boston Children’s Hospital, Kosuke Kawai, was consulted and is now included as a co-author. In the revised manuscript we have rewritten the Methods section and verified the statistical analyses using R and SAS software.

As a side-note: Once more it is unclear how the comparisons are "paired" (line 583). In my reading, 4 values from wt are compared to 4 values from the double KO at the same position of the basilar membrane from animals of the same age. That does not make those comparisons "paired". Truly 'paired' are comparisons of data from the same basilar membrane, i.e. counts at 8, 11.3, 16, … kHz. However, it seems the effect of frequency was not tested (which I presume leads to missing the Tmc2 KO vs wt x Frequency interaction).

As noted above, the methods section was rewritten to clarify the statistical analyses done on our datasets.

I would like to suggest that the authors use less conservative ways of multiple-comparison correction (e.g. Holm). On several occasions (e.g. wt vs Tmc2 KO at P7 8 and 11.3 kHz), this would yield significant differences, where currently there is no effect detected.

Interestingly, this inability of Tmc1 to compensate for the Tmc2 KO at the apex seems to be consistent with the gradient of TMC2 --> TMC1 replacement reported in Kawashima 2011.

We decided to stick with the Bonferroni correction and not modify correction to be less conservative.

Suggested changes:

– If not done already, suggest to indicate how many images were analyzed per cochlea and how many total synapses are in each group.

Done.

– It appears the statistical unit for assessment of significance is # of cochlea, which I would agree is most appropriate. Suggest to state explicitly.

This is now stated in the figure legends.

– On line 344: "However, the differences in standard deviations were not statistically significant relative to those uninjected mice" ….. perhaps better: "However, the standard deviations of injected mice were not significantly different from those of uninjected mice"

Sentence was changed per editors’ suggestion.

– Supplementary Table 5b Prism Stats: If you are going to include this, should there not be some description? It seems like you are now comparing the medians. Is this meaningful? Maybe I missed it, but I don't find any description of this in the text. Your response to reviewers agrees it is not meaningful to compare absolute volumes. When it says, "Do the medians vary signif (P<0.05)?" …. are those medians of volumes or medians of SD's? Please include description of Tables. Again, sorry if I just missed it. If the Supplementary files contain irrelevant things, perhaps the irrelevant things should be removed.

Thank you for the suggestion. We agree the Supplementary Table 5 Prism Stats tab was contained irrelevant data and the tab was removed. The “Do the medians vary signif” referred to the medians of SDs. The updated Supplementary file 5 now contains three tabs, each corresponding to a supplemental table.

– I found some of the supplemental tables difficult to read, like #2 and #3.

Supplemental Tables 2 and 3 were re-structured to be easier to read. Each table was broken down into sub tables corresponding to their respective figures (i.e. Supplementary file 2A-C Figure 3A-C; Supplementary file 3A-C Figure 4A-C). Supplementary file legends were updated to reflect these new tables.

– In the Transparent reporting file, what does it mean to say: "Samples were allocated into groups randomly based on genotype and ages."

There was no randomization. We state “Samples were allocated into groups based on genotypes and ages”. The word random does not appear. Groups contained mice of the same genotype and age. Cochlea samples from WT, P7 mice were grouped together while cochleas from Tmc1 KO, p7 were placed in a separate group. i.e. groups were based on genotype and age.

– Regarding figure 2, I'm not sure I agree that synapse position "could not be assessed due to abnormal IHC morphology." At least, I don't see it in the paper. Maybe I missed it. If IHC morphology is abnormal in these mice, please mention this in the paper if not already. In the images and the schematics, they look normal. I accept that synapse position and morphology are outside the scope of this work.

We have not quantified that IHC morphology. At this point it is just an impression. In the absence of data, we’d rather not go on record stating it is abnormal. We agree, IHC morphology and synapse position are beyond the scope of the current manuscript and are better left for a future publication.

- Line 587, "divided" should be "multiplied".

Thank you for noting this typo. The sentence has been corrected.

https://doi.org/10.7554/eLife.69433.sa2

Article and author information

Author details

  1. John Lee

    1. Speech and Hearing Bioscience & Technology Program, Division of Medical Sciences, Harvard University, Boston, United States
    2. Department of Otolaryngology, Boston Children’s Hospital and Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Visualization, Writing - original draft
    Competing interests
    No competing interests declared
  2. Kosuke Kawai

    Department of Otolaryngology, Boston Children’s Hospital and Harvard Medical School, Boston, United States
    Contribution
    Formal analysis, Software, Writing – review and editing
    Competing interests
    No competing interests declared
  3. Jeffrey R Holt

    1. Department of Otolaryngology, Boston Children’s Hospital and Harvard Medical School, Boston, United States
    2. Department of Neurology, Boston Children’s Hospital and Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Formal analysis, Funding acquisition, Project administration, Supervision, Visualization, Writing – review and editing
    For correspondence
    jeffrey.holt@childrens.harvard.edu
    Competing interests
    holds a patent (62/638,697) on use of AAV9-PHP.B for gene therapy in the inner ear, is a scientific founder of Audition Therapeutics and an advisor to several biotech companies focused on inner ear therapeutics. The authors declare no other conflicts of interest.
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7182-8011
  4. Gwenaëlle SG Géléoc

    Department of Otolaryngology, Boston Children’s Hospital and Harvard Medical School, Boston, United States
    Contribution
    Conceptualization, Funding acquisition, Project administration, Supervision, Writing – review and editing
    Competing interests
    No competing interests declared

Funding

National Institute on Deafness and Other Communication Disorders (RO1 DC013521)

  • Jeffrey R Holt

National Institute on Deafness and Other Communication Disorders (RO1 DC008853)

  • Gwenaëlle SG Géléoc

National Institute on Deafness and Other Communication Disorders (F32 DC018233)

  • John Lee

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

This work was supported by the NIH/NIDCD grants F32 DC018233 (JL), R01 DC013521 (JRH), and R01 DC008853 (GSGG), Foundation Pour L'Audition, the Jeffrey and Kimberly Barber Fund and the Imaging and Vector Cores at Boston Children’s Hospital (BCH IDDRC P30 HD18655). The authors would like to thank Carl Nist-Lund for assistance with injections and ABRs and Stephanie Mauriac and Irina Marcovich for critical review of the manuscript.

Ethics

This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All of the animals were handled according to approved institutional animal care and use committee (IACUC) protocols (#20-02-4149R and #00001240) at Boston Children's Hospital.

Senior Editor

  1. Barbara G Shinn-Cunningham, Carnegie Mellon University, United States

Reviewing Editor

  1. Tobias Reichenbach, Friedrich-Alexander-University (FAU) Erlangen-Nürnberg, Germany

Reviewers

  1. Elisabeth Glowatzki, Johns Hopkins University School of Medicine, United States
  2. Mark A Rutherford, Washington University at St. Louis, United States
  3. Andreas Neef, Max Planck Institute for Dynamics and Self-Organization, Germany

Publication history

  1. Received: April 15, 2021
  2. Accepted: November 1, 2021
  3. Accepted Manuscript published: November 4, 2021 (version 1)
  4. Version of Record published: November 17, 2021 (version 2)

Copyright

© 2021, Lee et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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