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Sensory transduction is required for development of hair cell synapses

Citation

Lee, John; Holt, Jeffrey; Geleoc, Gwenaelle (2021), Sensory transduction is required for development of hair cell synapses, Dryad, Dataset, https://doi.org/10.5061/dryad.fxpnvx0sb

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.

Methods

ABR acquisition.  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 sound pressure levels (SPL) of 10 to 115 dB in 5 dB increment steps.  512 responses of alternating stimulus polarity were collected and averaged for each SPL.  ABR potentials were amplified (10,000x), 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.  Temporal bones were dissected and fixed in 4% paraformaldehyde for 1 h at room temperature.  Temporal bones were then decalcified in 120 mM EDTA for 2 h for 7-day-old mice and up to 20 h 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 h 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 h 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 10x air objective on a 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 (Muller 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 x 1.4 NA oil objective lens (Carl Zeiss, z step = 0.36 μm, scaling per pixel: 0.068 μm x 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 63x oil objective with AiryScan processing (Carl Zeiss, z step = 0.18 μm, scaling per pixel: 0.034 μm x 0.034 μm). 

Ribbon Synapse Counts and Volume Distribution Measurements.  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-TMC1ex1-WPRE 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. The Wilcoxon rank sum test was used for comparison of P2 WT and Tmc1Δ/Δ;Tmc2Δ/Δ CtBP2+ puncta counts. Mixed-model ANOVA was used to compare synapse counts across different genotypes and timepoints. Six paired comparisons were made between the four groups in Figures 3A-C and three paired comparisons were made between the three groups in Figures 4A-C and 6A. The Bonferroni correction was applied to correct for the multiple comparisons and the reported p values are the original p values divided by the number of paired comparisons made. The Wilcoxon rank sum test was used to compare average ABR thresholds in WT and Tmc1Δ/Δ mice injected with Tmc1 gene therapy. 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.  Exact p values are reported.  Statistical analyses were performed in GraphPad Prism and figures were created using OriginLab, OriginPro. 

Usage Notes

Original raw data files from Figures 1,2,3,4 and 6 are confocal images acquired using Zeiss, Zen software.  Zen software or any image analysis software that can open *.CZI files can be used to view these images.  Original raw data files from Figure 5 include ABR data acquired using custom data acquisition software from the Eaton-Peabody Laboratories Cochlear Function Test Suite.  The data can be viewed using Excel or other data analysis software.

Funding

National Institutes of Health, Award: DC018233

National Institutes of Health, Award: DC013521

National Institutes of Health, Award: DC008853