Name: morphological and functional evaluation of ribbon synapses at specific frequency regions of the mouse cochlea
Uploaded: 2019-05-10T13:30:00.000+00:00
Duration: 9 min 54 s
Description: Watch this Scientific Journal Video about Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea at JoVE.com

This work was supported by the National Natural Science Foundation of China (81770997, 81771016, 81830030); the joint funding project of Beijing Natural Science Foundation and Beijing Education Committee (KZ201810025040); the Beijing Natural Science Foundation (7174291); and the China Postdoctoral Science Foundation (2016M601067).

All procedures were carried out in accordance with the NRC/ILAR Guide for the Care and Use of Laboratory Animals (8th Edition). The study protocol was approved by the Institutional Animal Care and Use Committee of Capital Medical University, Beijing, China.
1. Animal Selection
<ol>
	<li>For all experiments, use adult C57BL/6J male mice (8 weeks old) as the animal model. 
	NOTE: C57BL/6J mice carrying a splice variant of the Cdh23 exhibit accelerated senescence in the auditory system, reflected as a 40% loss of ribbon synapses at the basal turn of the cochlea and a 10% loss at the middle turn by 6 months of age, followed by a rapid increase of this loss in the whole cochlea with age18,19. Thus, we advise caution when using C57BL/6J mice older than 6 months for auditory research. Other strains of mice can be used depending on specific experimental aims.</li>
	<li>Inspect the mice using a professional diagnostic pocket otoscope to rule out outer or middle ear pathologies prior to hearing assessments. Potential signs may include fluid or pus in the external auditory canal, redness and swelling in local tissue, and tympanic membrane perforation. 
	NOTE: Although this is rarely encountered, once identified, mice with outer or middle ear diseases should be excluded.</li>
</ol>
2. Hearing Assessment
<ol>
	<li>Anesthetize mice using an intraperitoneal injection of a mixture of ketamine hydrochloride (100 mg/kg) and xylazine hydrochloride (10 mg/kg). Judge the depth of anesthesia via painful stimuli (e.g., toe-pinch reflex). 
	NOTE: When the toe-pinch reflex is completely absent, the animal has reached an adequate depth of anesthesia for auditory testing. If more time is required for bilateral ABR recordings, administer a lower dosage (one-fifth the original dosage) of anesthetics to restore the original anesthetic plane. Take care to avoid anesthetic overdose, as this may lead to death in mice.</li>
	<li>Maintain the anesthetized animal&#8217;s body temperature at 37.5 &#176;C using a thermoregulating heating pad. Place the anesthetized animal in an electrically and acoustically shielded room to avoid interference throughout the hearing test. 
	NOTE: Maintain the physiological temperature during the entire procedure until the animal is totally awake, in order to prevent death caused by post-anesthesia hypothermia.</li>
	<li>Place subdermal needle electrodes (20 mm, 28 G) at the vertex of the skull (recording electrode), in the ipsilateral parotid region below the pinna of measured ear (reference electrode), and in the contralateral parotid region (ground electrode), with a depth of 3 mm under the skin of the mouse head, respectively20.</li>
	<li>Use a closed-field speaker to perform acoustic stimulation via a 2 cm plastic tube with a cone-shaped tip. Fit the tip into the external ear canal21. 
	NOTE: Ensure that the electrical impedance in the recording and reference electrodes is less than 3 kOhm (usually 1 kOhm). If the impedance is high, change the insertion site of the electrode, clean the electrode with alcohol, or replace the electrode to avoid alterations in ABR wave amplitude.</li>
	<li>For ABR recording, generate tone pips (3 ms duration, 1 ms rise/fall times, at a rate of 21.1/s, frequency: 4&#8211;48 kHz) and present them at decreasing SPLs from 90 to 10 dB in 5&#8210;10 dB SPL steps20. During this step, the responses are amplified (10,000 times), filtered (0.1&#8211;3 kHz), and averaged (1,024 samples/stimulus level). 
	NOTE: ABRs are collected for each stimulus level in 10 dB steps, with additional 5 dB steps near the threshold.</li>
	<li>At each frequency, determine the ABR threshold, which refers to the minimal SPL resulting in a reliable ABR recording with one or more distinguishable waves that can be clearly identified by visual inspection (Figure 1). 
	NOTE: It is usually necessary to repeat the process for low SPLs around the threshold to ensure the consistency of the waveforms. The response threshold is the lowest stimulus level at which the waveform can be observed, when a decrease of 5 dB would lead to the disappearance of the waveform.</li>
</ol>
3. Cochlear Tissue Processing
<ol>
	<li>After ABR recording, euthanize the anesthetized mice via cervical dislocation, decapitate them, expose the bulla from the ventral side, and open with sharp scissors to gain access to the cochlea.</li>
	<li>Using a fine forceps, remove the temporal bones, sever the stapes artery, remove the stapes from the oval window, and rupture the round window membrane. Make a small hole at the apex of the cochlea by gently rotating the tip of a needle (13 mm, 27 G).</li>
	<li>Fix the isolated bones with 4% (wt/vol) paraformaldehyde in 0.1 M phosphate-buffered saline (PBS, pH 7.4) overnight at 4 &#176;C. Using a fine-tipped pipette, gently flush fixative through the perilymphatic spaces via application to the oval or round windows (as an inlet) and the opening at the apex (as an outlet). 
	NOTE: Some proteins require a short duration of fixation to avoid destruction of their epitopes for immunolabeling. In such cases, incubate bones in 4% paraformaldehyde at room temperature (RT) for 2 h, depending on the manufacturer&#8217;s instructions for immunohistochemistry. Fixation can also be performed via cardiac perfusion to remove the cochlear blood, avoiding background noise due to non-specific staining at later stages, particularly in mouse models of cochlear synaptopathy.</li>
	<li>Rinse the bones three times for 5 min with 0.1 M cold PBS to remove residual paraformaldehyde. Decalcify the bones with 10% ethylenediaminetetraacetic acid (EDTA) either at RT for 4 h or at 4 &#176;C for 24 h via gentle shaking in a horizontal shaker at 20 rpm. EDTA can be refreshed midway. 
	NOTE: Decalcification times are subject to the concentration of EDTA and users&#8217; preferences. Decalcified tissue should maintain a certain degree of toughness, which facilitates the manipulation of isolating cochlear whole-mounts at later steps. Temporal bones can be decalcified in 10% EDTA with rotation, allowing researchers to leave the laboratory following ABR tests and fixation experiments. The decalcification time is flexible within the range of 20 to 30 h at 4 &#176;C.</li>
	<li>Transfer one decalcified temporal bone from EDTA to 0.1 M PBS. Use #3, #5 Dumont forceps and the 27 G needle to dissect the apical, middle and basal cochlear regions in turn and then dissect the cochlea out of the bone under a stereo dissection microscope (as previously described22). Make a series of small cuts along the spiral ligament using a razor blade, and remove the tectorial membrane and Reissner&#8217;s membrane (Figure 2). 
	NOTE: As long as the dissected cochlea is intact, this process can be modified according to the individual operator&#39;s usual protocol.</li>
	<li>Further dissect the remaining auditory epithelium including the spiral limbus into individual cochlear turns (apex, middle, and base with hook region) for whole-mount preparations.</li>
	<li>Under a 40x oil objective of a light microscope, measure the basilar membrane length with a 250 &#956;m scale placed in the eyepiece, which can be adjusted along the stereocilia of the IHCs.</li>
	<li>Calculate the length of each cochlear turn by adding all segment lengths (250 &#956;m per segment), and concomitantly obtain the total length of the basilar membrane by summing the lengths of each turn.</li>
	<li>Convert the total length of the basilar membrane including the hook region into a percentage based on distance from the cochlear apex (0% refers to the cochlear apex, 100% to the cochlear base).</li>
	<li>Convert this distance into the cochlear characteristic frequency using a logarithmic function (d(%) = 1 - 156.5 + 82.5 &#215; log(f), with a slope of 1.25 mm/octave of frequency, where d is the normalized distance from the cochlear apex in percent, f is the frequency in kHz), as previously described5,6. Thus frequency range in a corresponding area of the basilar membrane on each cochlear turn can be acquired.</li>
</ol>
4. Immunofluorescence Staining
<ol>
	<li>After dissection, place each cochlear turn in a separate 2.5 mL centrifuge tube and incubate cochlear turns in 10% goat serum/PBS/0.1% Triton X-100 for 1 h at RT on a rotator.</li>
	<li>Remove the above blocking/permeabilization solution from each tube using a 200 &#956;L pipette tip under a dissection microscope and incubate the specimens with primary antibodies diluted in 5% goat serum/PBS/0.1% Triton X-100 overnight at 4 &#176;C on a rotator. 
	NOTE: For immunolabeling of cochlear synaptic ribbons, use the presynaptic marker mouse anti-carboxyl-terminal binding protein 2 IgG1 (CtBP2, labeling the B domain of the RIBEYE scaffolding protein, 1:400) and the postsynaptic marker mouse anti-glutamate receptor 2 IgG2a (GluR2, labeling a subunit of the AMPA receptor, 1:200)23.</li>
	<li>Rinse three times for 5 min with 0.1 M cold PBS to remove residual primary antibodies and incubate the specimens with secondary antibodies diluted in 5% goat serum/PBS/0.1% Triton X-100 at RT for 2&#8210;3 h in darkness on a rotator. 
	NOTE: Prepare appropriate secondary antibody mixtures using goat anti-mouse Alexa Fluor 568 (IgG1, 1:500) and goat anti-mouse Alexa Fluor 488 (IgG2a, 1:500), which are complementary to the primary antibodies used in step 4.2. To improve labeling efficiency for synaptic ribbons, we recommend selecting specific secondary antibodies. Some labs extend incubation with secondary antibodies to increase GluR2 immunolabeling24.</li>
	<li>Rinse three times for 5 min with 0.1 M PBS to remove residual secondary antibodies and transfer the specimens from 2.5 mL centrifuge tubes to 35 mm plates containing 0.1 M PBS.</li>
	<li>Place a drop of mounting medium containing 4&#8217;,6-diamidino-2-phenylindole (DAPI) onto the slide and transfer the specimens from PBS to the mounting medium. Place one edge of a coverslip on the slide and release to let the coverslip fall gently. 
	NOTE: To ensure that the hair cells face upward and that no folding or twisting of cochlear specimens occurs during the procedure, mount cochlear specimens under a stereo dissection microscope.</li>
	<li>Place the slides in a slide box at 4 &#176;C overnight to let slides dry and then image under a laser confocal microscope.</li>
</ol>
5. Morphological Evaluation of Cochlear Ribbon Synapses
<ol>
	<li>Image slides using a confocal microscope with three lasers&#8212;a 405 nm UV diode, a 488 nm argon laser, and a 561 nm diode-pumped solid-state (DPSS) laser to excite DAPI (Excitation spectrum 409&#8211;464 nm), Alexa Fluor 488 (Excitation spectrum 496&#8211;549 nm) and Alexa Fluor 568 (Excitation spectrum 573&#8211;631 nm), respectively.</li>
	<li>Acquire confocal z-stacks over a distance of 8 &#956;m from each cochlear turn using a 63x high-resolution oil immersion lens. 
	&#8203;NOTE: Once defined, all parameters for digitizing photomicrographs should be saved and applied uniformly to all slides.</li>
	<li>For synaptic punctum counts, set the z-stacks (0.3 &#956;m step size) to span the entire length of IHCs, thereby ensuring that all synaptic puncta can be imaged.</li>
	<li>Merge the images containing puncta in a z-stack to obtain the z-axis projection, and import to the image-processing software.</li>
	<li>Divide synaptic total counts in each z-stack at specific frequency regions by the number of IHCs (equal to the DAPI nuclear manual counts) to calculate the number of synaptic puncta for each IHC. At each specific frequency region, average all synaptic puncta in three images of different microscopic fields containing 9&#8211;11 IHCs.
	<ol>
		<li>Outline the region of interests (ROIs) including the basolateral regions of each IHC using freehand selections button. Use the measure function for automatic quantification of puncta, and the watershed function to distinguish between closely adjacent spots.</li>
		<li>After each automated counting, perform visual inspections with manual corrections to ensure puncta counting reliable. 
		NOTE: Experimenters should remain blinded as to whether the slide is from the apex, middle, or basal turn of the cochlea.</li>
	</ol>
	</li>
	<li>Visually assess synaptic structure and distribution, to manually isolate individual IHCs from their neighbors by the Pencil Tool (M)&#160;to better visualize the cytoskeletal architecture and synaptic localization.</li>
	<li>To inspect the juxtaposition of presynaptic ribbons (CtBP2) and postsynaptic receptor patches (GluR2), extract the voxel space around ribbon by the Rectangular Marquee Tool and isolate individual ribbon by Crop. Through clicking Image &#62; Image Size, acquire a thumbnail array of these miniature projections, which can then be used to identify paired synapses (appeared as closely juxtaposed pairs of CtBP2-positive and GluR2-positive puncta) versus orphan ribbons (lacking postsynaptic glutamate receptor patches) (Figure 3). 
	NOTE: Normal cochlear synapses appear as combined immunolabeling of the presynaptic ribbon within the hair cell (anti-CtBP2) and the postsynaptic glutamate receptor patch on the auditory nerve terminal (anti-GluR2)25. Some labs use confocal projections in conjunction with 3D modeling to quantify synaptic patch size or volume26,27. Prior to significant loss of ribbon synapses, ribbons exhibiting changes in size or without paired glutamate receptor patches are likely indicative of synaptic dysfunction27,28.</li>
</ol>
6. Functional Evaluation of Cochlear Ribbon Synapses
<ol>
	<li>Collect all ABR waves for each frequency stimulus presented at an SPL of 90 dB for the analysis of suprathreshold ABR wave I amplitudes. 
	NOTE: Neurophysiological and morphological studies have demonstrated that low spontaneous-rate, high-threshold fibers are especially vulnerable to aging and noise exposure29,30. Although the simple loss of ribbon synapses cannot affect ABR thresholds, it commonly results in significant reductions in ABR wave I amplitudes, because these afferents including low spontaneous-rate, high-threshold fibers and high spontaneous-rate, low-threshold fibers contribute heavily to the summed activities of cochlear nerve fibers28,29,31. A suprathreshold intensity of 90 dB SPL is selected here.</li>
	<li>Determine peak-to-peak wave I amplitude using an offline analysis program (Figure 4). Each wave I in the ABR test consists of a starting positive (p) deflection and the subsequent negative (n) deflection. ABR wave I amplitude is defined as the difference in voltage between Ip (the positive peak of wave I) and In (the negative peak of wave I)29. 
	NOTE: In pathological conditions, cochlear synaptopathy can be determined based on the suprathreshold amplitudes of ABR wave I, which reflect the summed onset responses of SGNs evoked by sound. However, cochlear sensitivity that is not compromised due to OHC dysfunction is a prerequisite for this method.</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adding+insult+to+injury:+cochlear+nerve+degeneration+after+"temporary"+noise-induced+hearing+loss.">Adding insult to injury: cochlear nerve degeneration after "temporary" noise-induced hearing loss.</a> Journal of Neuroscience. 29 (45), 14077-14085 (2009).</li><li>Valero, M. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Noise-induced+cochlear+synaptopathy+in+rhesus+monkeys+(Macaca+mulatta).">Noise-induced cochlear synaptopathy in rhesus monkeys (Macaca mulatta).</a> Hearing Research. 353, 213-223 (2017).</li><li>Viana, L. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cochlear+neuropathy+in+human+presbycusis:+Confocal+analysis+of+hidden+hearing+loss+in+post-mortem+tissue.">Cochlear neuropathy in human presbycusis: Confocal analysis of hidden hearing loss in post-mortem tissue.</a> Hearing Research. 327, 78-88 (2015).</li><li>Tong, M., Brugeaud, A., Edge, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regenerated+synapses+between+postnatal+hair+cells+and+auditory+neurons.">Regenerated synapses between postnatal hair cells and auditory neurons.</a> Journal of the Association for Research in Otolaryngology. 14 (3), 321-329 (2013).</li><li>Landegger, L. D., Dilwali, S., Stankovic, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neonatal+murine+cochlear+explant+technique+as+an+in+vitro+screening+tool+in+hearing+research.">Neonatal murine cochlear explant technique as an in vitro screening tool in hearing research.</a> Journal of Visualized Experiments. (124), (2017).</li><li>Takeda, S., Mannstr&#246;m, P., Dash-Wagh, S., Yoshida, T., Ulfendahl, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+Aging+and+Noise+Exposure+on+Auditory+Brainstem+Responses+and+Number+of+Presynaptic+Ribbons+in+Inner+Hair+Cells+of+C57BL/6J+Mice.">Effects of Aging and Noise Exposure on Auditory Brainstem Responses and Number of Presynaptic Ribbons in Inner Hair Cells of C57BL/6J Mice.</a> Neurophysiology. 49 (5), 316-326 (2017).</li><li>Mehraei, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Auditory+brainstem+response+latency+in+noise+as+a+marker+of+cochlear+synaptopathy.">Auditory brainstem response latency in noise as a marker of cochlear synaptopathy.</a> Journal of Neuroscience. 36 (13), 3755-3764 (2016).</li></ol>

The authors have no conflicts of interest to disclose.

Since cochlear synaptopathy was first characterized in adult mice with a temporary threshold shift (TTS) induced by 8&#8210;16 kHz octave band noise at 100 dB SPL for 2 h31, researchers have increasingly investigated the effects of synaptopathy in various mammals, including monkeys and humans32,33. In addition to noise exposure, several other conditions have been associated with cochlear synaptopathy (e.g., aging, the use of ototoxic drugs, and genetic mutations), leading to short-term disruption of suprathreshold audition, followed by irreversible degeneration of the auditory nerve. In the early phase of cochlear insult, synaptopathy often occurs at a specific frequency location, especially in experimental models in which damage to IHC synapses is restricted to select frequency regions24,31. Therefore, our protocol is significant in that it enables the investigation of synaptic morphology and function at a specific frequency region.
The cochlear place-frequency map can be used to discriminate normal and abnormal auditory function, further reflecting normal and abnormal area in the inner ear. Since surface preparation techniques were first used to plot intact and missing hair cells in the different cochlear turns, the cochleogram has become a routine method for quantifying hair cell loss6. Therefore, to correlate morphological insults in the cochlea to relevant physiological changes, it is reasonable to include a place-frequency map on the cochleogram. This method allows one to determine the number and structure of synapses based on their positions along the cochlear basilar membrane in relation to established cochlear place-frequency maps, thereby providing sufficient information for detailed comparisons between histological and physiological outcomes. However, some studies assess synaptopathy based on cochlear segment or distance along the cochlear duct, which does not allow for direct comparisons among individual cochleae due to intra-species variations in basilar membrane length. Thus, it is particularly important to standardize the cochleogram by converting individual cochlear lengths from millimeters to a relative percentage.
Synaptopathy can be confirmed via visualization of immunostaining for CtBP2 protein (the synaptic isoform &#8220;ribeye&#8221;) in the basolateral regions of IHCs. These CtBP2-positive patches can serve as a structural marker for quantification of presynaptic ribbons, and the presence of fewer patches usually indicates presynaptic loss. Although previous studies have reported that 98% of presynaptic ribbons are paired with post-synaptic terminals in the normal ear30, the simple number of CtBP2-positive patches may not be accurate for quantitative analysis of complete synapses, as this may lead to overestimation of synapse counts in the injured ear by involving &#8220;orphans&#8221; (presynaptic ribbons unpaired with postsynaptic terminals). To improve the accuracy of estimating synapse number, additional antibodies against postsynaptic structure such as GluA2, GluA2/3, or PSD-95 are used. The postsynaptic density protein PSD-95, a membrane-associated guanylate kinase (MAGUK) scaffolding protein, can be labeled using an antibody against PSD-95, which is mostly observed at the contacts between hair cells and SGN fiber endings34,35. However, the antibody against GluR2 is more specific for AMPA-type ionotropic glutamate receptors in the postsynaptic membrane, which can more reliably identify ribbon synapses (juxtaposed pairs of immunofluorescent puncta of presynaptic CtBP2 and postsynaptic GluR2)15. In addition to morphological analysis, histological analysis of complete synapses can be used as a functional indicator for synaptopathy. In adult mice with cochlear synaptopathy, reductions in ABR wave I amplitude following the presentation of moderate- to high-level tone stimuli occur at frequencies tonotopically related to regions of synaptic loss26,29. Synapse counts obtained using this method are limited in that it takes approximately 10 minutes to scan one site on the basilar membrane. Furthermore, to evaluate spatial variations in synaptic number and morphology based on IHC location, it is necessary to accurately identify the boundary of the ICH body (e.g., via myosin VIIa staining) and to perform particular image-processing steps26. Using such methods, previous studies have confirmed that synaptic loss is greater on the modiolar side of the IHC than on the pillar side in animal models of noise-induced degeneration26.
Previous studies have demonstrated that low spontaneous-rate, high-threshold fibers are more susceptible to noise damage than high spontaneous-rate, low-threshold fibers in animal models of restricted synaptopathy26,30. These studies provide the rationale for using suprathreshold amplitudes of ABR wave I (measured using moderate- to high-level tone stimuli) to assess synaptic function in animal models with normal ABR threshold and distortion product otoacoustic emissions (DPOAEs). Because wave I of the ABR reflects the summed neural responses of auditory nerve fibers, when synaptopathy is assessed via ABR wave I amplitude, DPOAE testing should also be performed to exclude OHC damage, which also can reduce ABR amplitudes due to the disruption of mechanoelectric transduction. Although the first wave of the compound action potential (CAP), which is measured from round-window electrodes, also represents summed activity of the cochlear nerve, this method is more invasive and complex than ABR measurements. If DPOAE responses return to normal after temporary threshold shifts in noise-exposed mice31 or they have not yet deteriorated in aging mice29, the suprathreshold amplitude of ABR wave I can strongly predict the degree of cochlear synaptopathy, since affected neurons are silenced when their synaptic connections to IHCs are disrupted. However, if cochlear sensitivity is reduced due to various factors (e.g., OHC dysfunction), decreases in ABR wave I amplitude can no longer be attributed solely to synaptic loss because they will reflect the combination of all cochlear damage. Therefore, controlled experimental conditions for synaptopathy-restricted model preparation are required to ensure that cochlear sensitivity is not compromised by other factors. Unfortunately, although ABR wave I amplitude provides an objective measure of auditory nerve fiber loss in animals, it is difficult to measure in humans. Moreover, mixed pathologies involving synaptopathy, loss of hair cells, and the presence of other abnormalities in the cochlea may co-occur in humans, limiting the use of ABR wave I amplitude measurements in clinical settings. ABR wave latency is calculated as the time in milliseconds from the onset of the stimulus to the positive peak of each wave, providing insight into the transmission times along the auditory pathway. No significant changes in wave I latency have been observed in mouse models of noise-induced or age-related cochlear synaptopathy36. However, some evidence suggests that the effects of masking noise on ABR wave V latency can be used to diagnose cochlear synaptopathy in humans37.
Several recent studies have supported the notion that cochlear synaptopathy is the primary initial event associated with hidden hearing loss, tinnitus, and hyperacusis. Although the concept of cochlear synaptopathy in inner ear diseases has now been firmly established, its detailed impact on hearing ability remains unknown. The protocol presented in the current study enables the investigation of morphology and function in cochlear ribbon synapses within a specific frequency region. Thus, this protocol can be used to investigate cochlear synaptopathy, its underlying mechanisms, and the efficacy of potential therapeutic interventions in various experimental animal models.

Frequencies in the range of approximately 20&#8210;20,000 Hz can be perceived as auditory stimuli by humans. Human hearing is normally most sensitive near 1,000 Hz, where average sound pressure level is 20 &#956;Pa in young adults (i.e., 0 decibels of sound pressure level [dB SPL]). In some pathological conditions, hearing loss is restricted to specific frequencies. For example, in the early stages of noise-induced hearing loss (NIHL), a &#8220;notch&#8221; (i.e., hearing threshold elevation) can be observed in the audiogram at 4 kHz1. Along the mammalian cochlear partition, its gradations of stiffness and mass produce an exponential frequency map, with high-frequency sound detection at the base of the cochlea and low-frequency detection at the apex2. Indeed, there is a cochlear place-frequency map along the basilar membrane, leading to what is known as tonotopic organization2,3. Each given place on the basilar membrane has the highest sensitivity to only one particular sound frequency, which is usually termed the characteristic frequency3,4, although responses to other frequencies can also be observed.
To date, various mouse models have been employed to investigate normal function, pathological processes, and therapeutic efficacy in the auditory system. Precise knowledge of physiological parameters in the mouse cochlea is a prerequisite for such studies of hearing loss. The mouse cochlea is anatomically divided into apical, middle, and basal turns, which correspond to different frequency regions. By labeling auditory nerve afferents at the cochlear nucleus to analyze their corresponding peripheral innervation sites in the cochlea, M&#252;ller et al. succeeded in establishing the cochlear place-frequency map in the normal mouse in vivo5. In the interval of 7.2&#8211;61.8 kHz, which corresponds to positions between 90% and 10% of the full length of the basilar membrane, the mouse cochlear place-frequency map can be described by a simple linear regression function, suggesting a relation between the normalized distance from the cochlear base and the logarithm of the characteristic frequency5. In laboratory mice, the place-frequency map can be used to explore the relationship between hearing thresholds within specific frequency ranges and cochleograms showing the numbers of missing hair cells in relative regions along the basilar membrane6. Importantly, the place-frequency map provides a positioning system for the investigation of minimal structural damage, such as damage to the ribbon synapses of hair cells at specific cochlear frequency locations in mice with peripheral auditory trauma7,8.
In the mammalian cochlea, ribbon synapses are comprised of a presynaptic ribbon, an electron-dense projection that tethers a halo of release-ready synaptic vesicles containing glutamate within the IHC, and a postsynaptic density on the nerve terminal of the SGN with glutamate receptors9. During cochlear sound transduction, deflection of the hair cell bundle results in IHC depolarization, which leads to glutamate release from IHCs onto the postsynaptic afferent terminals, thereby activating the auditory pathway. Activation of this pathway leads to the transformation of sound-induced mechanical signals into a rate code in the SGN10. Indeed, the IHC ribbon synapse is highly specialized for indefatigable sound transmission at rates of hundreds of Hertz with high temporal precision, and is of critical importance for presynaptic mechanisms of sound encoding. Previous studies have revealed that ribbon synapses vary greatly in size and number at different frequency regions in the adult mouse cochlea11,12, likely reflecting structural adaptation to the particular sound coding for survival needs. Recently, experimental animal studies have demonstrated that cochlear synaptopathy contributes to multiple forms of hearing impairments, including noise-induced hearing loss, age-related hearing loss, and hereditary hearing loss13,14. Thus, methods for identifying correlated changes in synaptic number, structure, and function at specific frequency regions have been increasingly employed in studies of auditory development and inner ear disease, using models generated via experimental manipulation of genetic or environmental variables15,16,17.
In the current report, we present a protocol for analyzing the synaptic number, structure, and function at a specific frequency region of the basilar membrane in adult mice. Cochlear frequency localization is performed using a given place-frequency map in combination with a cochleogram. The normal morphological characteristics of cochlear ribbon synapses are evaluated via presynaptic and postsynaptic immunostaining. The functional status of cochlear ribbon synapses is determined based on the suprathreshold amplitudes of ABR wave I. With minor alterations, this protocol can be used to examine physiological or pathological conditions in other animal models, including rats, guinea pigs, and gerbils.

<table><tbody><tr><td>Ketamine hydrochloride</td><td>Gutian Pharmaceutical Co., Ltd., Fujian, China</td><td>H35020148</td><td>100 mg/kg</td></tr><tr><td>Xylazine hydrochloride</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>X-1251</td><td>10 mg/kg</td></tr><tr><td>TDT physiology apparatus</td><td>Tucker-Davis Technologies, Alachua, FL, USA</td><td>Auditory Physiology System III</td><td></td></tr><tr><td>SigGen/BioSig software</td><td>Tucker-Davis Technologies, Alachua, FL, USA</td><td>Auditory Physiology System III</td><td></td></tr><tr><td>Electric Pad</td><td>Pet Fun</td><td>11072931136</td><td></td></tr><tr><td>Dumont forceps 3#</td><td>Fine Science Tools, North Vancouver, B.C., Canada</td><td>0203-3-PO</td><td></td></tr><tr><td>Dumont forceps 5#</td><td>Fine Science Tools, North Vancouver, B.C., Canada</td><td>0209-5-PO</td><td></td></tr><tr><td>Stereo dissection microscope</td><td>Nikon Corp., Tokyo, Japan</td><td>SMZ1270</td><td></td></tr><tr><td>Goat serum</td><td>ZSGB-BIO, Beijing,China</td><td>ZLI-9021</td><td></td></tr><tr><td>Anti-glutamate receptor 2, extracellular, clone 6C4</td><td>Millipore Corp., Billerica, MA, USA</td><td>MAB397</td><td>mouse&amp;nbsp;</td></tr><tr><td>Purified Mouse Anti-CtBP2</td><td>BD Biosciences, Billerica, MA, USA</td><td>612044</td><td>mouse&amp;nbsp;</td></tr><tr><td>Alexa Fluor 568 goat anti-mouse IgG1antibody</td><td>Thermo Fisher Scientific Inc., Waltham, MA, USA</td><td>A21124</td><td>goat</td></tr><tr><td>Alexa Fluor 488 goat anti-mouse IgG2a antibody</td><td>Thermo Fisher Scientific Inc., Waltham, MA, USA</td><td>A21131</td><td>goat</td></tr><tr><td>Mounting medium containing DAPI</td><td>ZSGB-BIO, Beijing,China</td><td>ZLI-9557</td><td></td></tr><tr><td>Confocal fluorescent microscopy</td><td>Leica Microsystems, Wetzlar, Germany</td><td>TCS SP8 II</td><td></td></tr><tr><td>Image Pro Plus software</td><td>Media Cybernetics, Bethesda, MD, USA</td><td>version 6.0</td><td></td></tr><tr><td>Professional diagnostic pocket otoscope</td><td>Lude Medical Apparatus and Instruments Trade Co., Ltd., Shanghai,China</td><td>HS-OT10</td><td></td></tr><tr><td>Needle electrode</td><td>Friendship Medical Electronics Co., Ltd., Xi&#039;an,China</td><td>1029</td><td>20 mm, 28 G</td></tr><tr><td>Closed-field speaker</td><td>Tucker-Davis Technologies, Alachua, FL, USA</td><td>CF1</td><td></td></tr></tbody></table>

morphological and functional evaluation of ribbon synapses at specific frequency regions of the mouse cochlea

Cochlear inner hair cells (IHCs) transmit acoustic signals to spiral ganglion neurons (SGNs) through ribbon synapses. Several experimental studies have indicated that hair cell synapses may be the initial targets in sensorineural hearing loss (SNHL). Such studies have proposed the concept of cochlear &#34;synaptopathy&#34;, which refers to alterations in ribbon synapse number, structure, or function that result in abnormal synaptic transmission between IHCs and SGNs. While cochlear synaptopathy is irreversible, it does not affect the hearing threshold. In noise-induced experimental models, restricted damage to IHC synapses in select frequency regions is employed to identify the environmental factors that specifically cause synaptopathy, as well as the physiological consequences of disturbing this inner ear circuit. Here, we present a protocol for analyzing cochlear synaptic morphology and function at a specific frequency region in adult mice. In this protocol, cochlear localization of specific frequency regions is performed using place-frequency maps in conjunction with cochleogram data, following which the morphological characteristics of ribbon synapses are evaluated via synaptic immunostaining. The functional status of ribbon synapses is then determined based on the amplitudes of auditory brainstem response (ABR) wave I. The present report demonstrates that this approach can be used to deepen our understanding of the pathogenesis and mechanisms of synaptic dysfunction in the cochlea, which may aid in the development of novel therapeutic interventions.

ABR hearing tests were performed for 10 C57BL/6J mice (8 weeks of age) under anesthesia. ABRs were elicited using tone burst stimuli at 4, 8, 16, 32, and 48 kHz. The hearing threshold of each animal was visually detected by distinguishing at least one clear waveform in the ABR. All mice exhibited ABR thresholds in response to tone bursts, ranging between 25 and 70 dB SPL depending on the frequency of the stimulus. Our results indicated that the hearing threshold was lowest at 16 kHz (Figure 1), corresponding to approximately 43% distance from the cochlear apex (Figure 2), suggesting that acoustic sensitivity is significantly reduced in other cochlear regions.
Cochlear whole-mounts were isolated from temporal bones in adult mice under a stereo dissection microscope (Figure 2A). Whole-mounts of the auditory epithelium were dissected into three pieces, the lengths of which were measured and eventually converted into percent distance from the cochlear apex. The frequency location on the basilar membrane of each cochlear turn was calculated using a logarithmic function, as previously described5,6 (Figure 2B).
To evaluate the morphological characteristics of cochlear ribbon synapses, antibodies against CtBP2 and GluR2 were used to label presynaptic and postsynaptic structures, respectively. In normal ears of adult mice, immunostaining revealed juxtaposed pairs of synaptic ribbons and glutamate receptor patches studding the surface of the basolateral membrane of IHCs, with 8&#8211;20 pairs per IHC (Figure 3A). Although the vast majority of puncta appeared as juxtaposed pairs in normal ears, orphan ribbons could be observed rarely at high magnification (Figure 3B). Counts of IHC ribbon synapses (immunopositive spots for both CtBP2 and GluR2) were highest at the 16 kHz region, significantly decreasing as the distance from this location increased (Figure 3C). The synaptic counts determined based on confocal projections provide an estimate of the maximum number of auditory nerve fibers that transmit information from the cochlea to the brain29.
The functional status of cochlear ribbon synapses was investigated in all adult mice based on ABR wave I amplitudes, which provide information regarding the functional integrity of auditory nerve fibers29,31. ABR wave I amplitudes at each frequency of the stimulus presented at a sound pressure level of 90 dB were measured from peak to the following trough, as shown in Figure 4A. ABR wave I amplitude was highest at a frequency of 16 kHz, corresponding to the lowest hearing threshold, and amplitude values significantly decreased as distance from this location increased (Figure 4B). This result is consistent with the observed alterations in ribbon synapse counts, indicating that synapses within this cochlear region may exhibit the most vivid synaptic function. Furthermore, in previous mouse studies of noise-induced and age-related cochlear neurodegeneration, suprathreshold amplitudes of ABR wave I decreased in proportion to ribbon loss, indicating that ABR wave I amplitude is highly correlated with the degree of cochlear synaptopathy29,31.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59189/59189fig1.jpg" /> 
Figure 1: Hearing Assessment. ABR threshold comparisons among different frequencies of tone burst stimuli demonstrated that the hearing threshold was lowest at 16 kHz in 10 adult C57BL/6J mice. The ABR response was significantly elevated at other frequency regions (one-way ANOVA with Dunnett&#8217;s multiple comparison post hoc test; *: P &#60; 0.01, n = 20 ears). Data are expressed as the mean &#177; SEM. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59189/59189fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59189/59189fig2.jpg" /> 
Figure 2: Cochlear frequency localization in mice. (A) A representative image of the full explanted cochlea, which has been dissected out of the temporal bone under a stereo dissection microscope. (B) The mouse cochlea is divided into apical, middle, and basal turns, where the cochlear lateral wall is removed. Red circles on the fragments of the cochlear basilar membrane indicate the frequency locations and their corresponding normalized positions in the cochlea (0% refers to the cochlear apex, 100% to the cochlear base). Scale bar = 250 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59189/59189fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59189/59189fig3.jpg" /> 
Figure 3: Confocal analysis of cochlear ribbon synapses in mice. (A) Representative images of ribbon synapses for the 4, 8, 16, 32, and 48 kHz regions, immunostained for presynaptic ribbons (CtBP2, red) and postsynaptic structures (GluR2, green). White dashed lines are used to outline inner hair cells for reference. Scale bar = 10 &#956;m. (B) High-power thumbnails from confocal z-stacks show that these ribbon synapses appeared as closely juxtaposed pairs of CtBP2-positive (red) and GluR2-positive (green) puncta (left and middle), whereas orphan ribbons (right) lacking postsynaptic glutamate receptor patches were very rare. Scale bar = 0.5 &#956;m. (C) Quantitative analysis of paired ribbon puncta with all presynaptic and postsynaptic elements revealed that the number of synaptic puncta per inner hair cell was significantly higher at the 16 kHz region than in other frequency regions (one-way ANOVA with Dunnett&#8217;s multiple comparison post hoc test; *: P &#60; 0.01, n=6 ears). Data are expressed as the mean &#177; SEM. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59189/59189fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59189/59189fig4.jpg" /> 
Figure 4: Analysis of ABR wave I amplitudes in mice. (A) Representative ABR waveform from an 8 weeks old C57BL/6J mouse exposed to a 16 kHz pure tone stimulus at an intensity of 90 dB (stimulus onset at 0 ms). Roman numerals mark the peaks of the ABR waves. Dotted lines mark the wave I peak and trough, indicating the amplitude. (B) Quantitative analysis of average wave I amplitudes in response to the 4, 8, 16, 32, and 48 kHz stimuli presented at a sound pressure level of 90 dB. ABR wave I amplitudes were highest at the frequency of 16 kHz and significantly lower at other frequencies (one-way ANOVA with Dunnett&#8217;s multiple comparison post hoc test; *: P &#60; 0.01, n = 20 ears). Data are expressed as the mean &#177; SEM. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59189/59189fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea

This manuscript describes an experimental protocol for evaluating the morphological characteristics and functional status of ribbon synapses in normal mice. The present model is also suitable for noise-induced and age-related cochlear synaptopathy-restricted models. The correlative results of previous mouse studies are also discussed.

Cochlear inner hair cells (IHCs) transmit acoustic signals to spiral ganglion neurons (SGNs) through ribbon synapses. Several experimental studies have ...

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11.7K Views.  Capital Medical University. Het primaire rapport is echt stress belang van het nauwkeurig lokaliseren van de specifieke frequentie regio van de basilaire membraan in de meeste modellen, waar synaps nummers veranderd en functie kan worden geanalyseerd. De cochleaire lokalisatie van de specifieke frequentiegebieden wordt betrouwbaar uitgevoerd met behulp van frequentiekaarten in combinatie met cochleogramgegevens. De microscoop geeft inzicht in verschillende onderzoeken.Het ondersteunt het idee dat de cochleaire ne...

Video: Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea

Postsynaptic Recordings at Afferent Dendrites Contacting Cochlear Inner Hair Cells: Monitoring Multivesicular Release at a Ribbon Synapse

The afferent synapse between the inner hair cell (IHC) and the auditory nerve fiber provides an electrophysiologically accessible site for recording the postsynaptic activity of a single ribbon synapse 1-4. Ribbon synapses of sensory cells release neurotransmitter continuously, the rate of which is modulated in response to graded changes in IHC membrane potential 5. Ribbon synapses have been shown to operate by multivesicular release, where multiple vesicles can be released simultaneously to evoke excitatory postsynaptic currents (EPSCs) of varying amplitudes 1, 4, 6-11. Neither the role of the presynaptic ribbon, nor the mechanism underlying multivesicular release is currently well understood.
The IHC is innervated by 10-20 auditory nerve fibers, and every fiber contacts the IHC with a unmyelinated single ending to form a single ribbon synapse. The small size of the afferent boutons contacting IHCs (approximately 1 &mu;m in diameter) enables recordings with exceptional temporal resolution to be made. Furthermore, the technique can be adapted to record from both pre- and postsynaptic cells simultaneously, allowing the transfer function at the synapse to be studied directly 2. This method therefore provides a means by which fundamental aspects of neurotransmission can be studied, from multivesicular release to the elusive function of the ribbon in sensory cells.

Whole-cell patch-clamp recordings from auditory nerve fiber dendrites at the inner hair cell ribbon synapse in the mammalian cochlea.

The afferent synapse between the inner hair cell (IHC) and the auditory nerve fiber provides an electrophysiologically accessible site for recording the ...

Cerebrovascular Casting of the Adult Mouse for 3D Imaging and Morphological Analysis

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). Animal models are necessary for studying the etiopathology and potential therapies of cerebrovascular diseases. Imaging the vasculature in large animals is relatively easy. However, developing vessel imaging methods of murine brain disease models is desirable due to the cost and availability of genetically-modified mouse lines. Imaging the murine cerebral vascular tree is a challenge. In humans and larger animals, the gold standard for assessing the angioarchitecture at the macrovascular (conductance) level is x-ray catheter contrast-based angiography, a method not suited for small rodents.
In this article, we present a method of cerebrovascular casting that produces a durable skeleton of the entire vascular bed, including arteries, veins, and capillaries that may be analyzed using many different modalities. Complete casting of the microvessels of the mouse cerebrovasculature can be difficult; however, these challenges are addressed in this step-by-step protocol. Through intracardial perfusion of the vascular casting material, all vessels of the body are casted. The brain can then be removed and clarified using the organic solvent methyl salicylate. Three dimensional imaging of the brain blood vessels can be visualized simply and inexpensively with any conventional brightfield microscope or dissecting microscope. The casted cerebrovasculature can also be imaged and quantified using micro-computed tomography (micro-CT)1. In addition, after being imaged, the casted brain can be embedded in paraffin for histological analysis.
The benefit of this vascular casting method as compared to other techniques is its broad adaptation to various analytic tools, including brightfield microscopic analysis, CT scanning due to the radiopaque characteristic of the material, as well as histological and immunohistochemical analysis. This efficient use of tissue can save animal usage and reduce costs. We have recently demonstrated application of this method to visualize the irregular blood vessels in a mouse model of adult BAVM at a microscopic level2, and provide additional images of the malformed vessels imaged by micro-CT scan. Although this method has drawbacks and may not be ideal for all types of analyses, it is a simple, practical technique that can be easily learned and widely applied to vascular casting of blood vessels throughout the body.

In this article, we present a simple, practical technique for cerebrovascular casting that is easy to perform and can be utilized to image the vascular tree of the adult mouse brain.

Vascular imaging is crucial in the clinical diagnosis and management of cerebrovascular diseases, such as brain arteriovenous malformations (BAVMs). ...

Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a spiral shape and have a tonotopic gradient for sound detection. The mouse cochlea is approximately 6 mm long and often divided into three turns (apex, middle, and base) for analysis. To investigate cell loss, cell division, or mosaic gene expression, the whole mount or surface preparation of the cochlea is useful. This dissection method allows visualization of all cells within the organ of Corti when combined with immunostaining and confocal microscopy to image cells at different planes in the z-axis. Multiple optical cross-sections can also be obtained from these z-stack images. In addition, the whole mount dissection method can be used for scanning electron microscopy, although a different fixation method is needed. Here, we present a method to isolate the organ of Corti as three intact cochlear turns (apex, middle, and base). This method can be used for mice ranging from one week of age through adulthood and differs from the technique used for neonatal samples where calcification of the cochlea is incomplete. A slightly modified version can be used for dissection of the rat cochlea. We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies.

We present a method to isolate the adult organ of Corti as three intact cochlear turns (apex, middle, and base). We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies. Together these techniques allow the study of hair cells, supporting cells, and other cell types found in the cochlea.

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a ...

Evaluation of Planar-Cell-Polarity Phenotypes in Ciliopathy Mouse Mutant Cochlea

In recent years, primary cilia have emerged as key regulators in development and disease by influencing numerous signaling pathways. One of the earliest signaling pathways shown to be associated with ciliary function was the non-canonical Wnt signaling pathway, also referred to as planar cell polarity (PCP) signaling. One of the best places in which to study the effects of planar cell polarity (PCP) signaling during vertebrate development is the mammalian cochlea. PCP signaling disruption in the mouse cochlea disrupts cochlear outgrowth, cellular patterning and hair cell orientation, all of which are affected by cilia dysfunction. The goal of this protocol is to describe the analysis of PCP signaling in the developing mammalian cochlea via phenotypic analysis, immunohistochemistry and scanning electron microscopy. Defects in convergence and extension are manifested as a shortening of the cochlear duct and/or changes in cellular patterning, which can be quantified following dissection from developing mouse mutants. Changes in stereociliary bundle orientation and kinocilia length or positioning can be observed and quantitated using either immunofluorescence or scanning electron microscopy (SEM). A deeper insight into the role of ciliary proteins in cellular signaling pathways and other biological phenomena is crucial for our understanding of cellular and developmental biology, as well as for the development of targeted treatment strategies.

Primary cilia influence various signaling pathways. The mammalian cochlea is ideal for examining planar cell polarity (PCP) signaling. Cilia dysfunction affects cochlear outgrowth, cellular patterning and hair cell orientation, readouts of PCP. Our goal is to analyze PCP signaling in mouse cochlea via phenotypic analysis, immunohistochemistry and scanning electron microscopy.

In recent years, primary cilia have emerged as key regulators in development and disease by influencing numerous signaling pathways. One of the earliest ...

Medicine

Cochlear Surface Preparation in the Adult Mouse

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous etiologies such as exposure to excessive noise, the use of ototoxic medications, bacterial or viral ear infections, head injuries, and the aging process. Loss of sensory hair cells is a common pathological feature of the varieties of acquired hearing loss. Additionally, the inner hair cell synapse can be damaged by mild insults. Therefore, surface preparations of cochlear epithelia, in combination with immunolabeling techniques and confocal imagery, are a very useful tool for the investigation of cochlear pathologies, including losses of ribbon synapses and sensory hair cells, changes in protein levels in hair cells and supporting cells, hair cell regeneration, and determination of report gene expression (i.e., GFP) for verification of successful transduction and identification of transduced cell types. The cochlea, a bony spiral-shaped structure in the inner ear, holds the auditory sensory end organ, the organ of Corti (OC). Sensory hair cells and surrounding supporting cells in the OC are contained in the cochlear duct and rest on the basilar membrane, organized in a tonotopic fashion with high-frequency detection occurring in the base and low-frequency in the apex. With the availability of molecular and genetic information and the ability to manipulate genes by knockout and knock-in techniques, mice have been widely used in biological research, including in hearing science. However, the adult mouse cochlea is miniscule, and the cochlear epithelium is encapsulated in a bony labyrinth, making microdissection difficult. Although dissection techniques have been developed and used in many laboratories, this modified microdissection method using cell and tissue adhesive is easier and more convenient. It can be used in all types of adult mouse cochleae following decalcification.

This article presents a modified cochlear surface preparation method that requires decalcification and use of a cell and tissue adhesive to adhere the pieces of cochlear epithelia to 10 mm round cover slips for immunohistochemistry in adult mouse cochleae.

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous ...

In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively thin to properly visualize and access neurons for patch-clamping or imaging, and in vitro examination of brain circuitry is limited to only what is physically present in the acute slice. To maintain the benefits of in vitro slice experimentation while preserving a larger portion of presynaptic nuclei, we developed a novel slice preparation. This &#8220;wedge slice&#8221; was designed for patch-clamp electrophysiology recordings to characterize the diverse monaural, sound-driven inputs to medial olivocochlear (MOC) neurons in the brainstem. These neurons receive their primary afferent excitatory and inhibitory inputs from neurons activated by stimuli in the contralateral ear and corresponding cochlear nucleus (CN). An asymmetrical brain slice was designed which is thickest in the rostro-caudal domain at the lateral edge of one hemisphere and then thins towards the lateral edge of the opposite hemisphere. This slice contains, on the thick side, the auditory nerve root conveying information about auditory stimuli to the brain, the intrinsic CN circuitry, and both the disynaptic excitatory and trisynaptic inhibitory afferent pathways that converge on contralateral MOC neurons. Recording is performed from MOC neurons on the thin side of the slice, where they are visualized using DIC optics for typical patch-clamp experiments. Direct stimulation of the auditory nerve is performed as it enters the auditory brainstem, allowing for intrinsic CN circuit activity and synaptic plasticity to occur at synapses upstream of MOC neurons. With this technique, one can mimic in vivo circuit activation as closely as possible within the slice. This wedge slice preparation is applicable to other brain circuits where circuit analyses would benefit from preservation of upstream connectivity and long-range inputs, in combination with the technical advantages of in vitro slice physiology.

Integration of diverse synaptic inputs to neurons is best measured in a preparation that preserves all pre-synaptic nuclei for natural timing and circuit plasticity, but brain slices typically sever many connections. We developed a modified brain slice to mimic in vivo circuit activity while maintaining in vitro experimentation capability.

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively ...

Immunolabeling and Counting Ribbon Synapses in Young Adult and Aged Gerbil Cochleae

The loss of ribbon synapses connecting inner hair cells and afferent auditory nerve fibers is assumed to be one cause of age-related hearing loss. The most common method for detecting the loss of ribbon synapses is immunolabeling because it allows for quantitative sampling from several tonotopic locations in an individual cochlea. However, the structures of interest are buried deep inside the bony cochlea. Gerbils are used as an animal model for age-related hearing loss. Here, routine protocols for fixation, immunolabeling gerbil cochlear whole mounts, confocal imaging, and quantifying ribbon synapse numbers and volumes are described. Furthermore, the particular challenges associated with obtaining good material from valuable aging individuals are highlighted. 
Gerbils are euthanized and either perfused cardiovascularly, or their tympanic bullae are carefully dissected out of the skull. The cochleae are opened at the apex and base and directly transferred to the fixative. Irrespective of the initial method, the cochleae are postfixed and subsequently decalcified. The tissue is then labeled with primary antibodies against pre- and postsynaptic structures and hair cells. Next, the cochleae are incubated with secondary fluorescence-tagged antibodies that are specific against their respective primary ones. The cochleae of aged gerbils are then treated with an autofluorescence quencher to reduce the typically substantial background fluorescence of older animals' tissues. 
Finally, cochleae are dissected into 6-11 segments. The entire cochlear length is reconstructed such that specific cochlear locations can be reliably determined between individuals. Confocal image stacks, acquired sequentially, help visualize hair cells and synapses at the chosen locations. The confocal stacks are deconvolved, and the synapses are either counted manually using ImageJ, or more extensive quantification of synaptic structures is carried out with image analysis procedures custom-written in Matlab.

A protocol for processing young adult and aged gerbil cochleae by immunolabeling the afferent synaptic structures and hair cells, quenching autofluorescence in aged tissue, dissecting and estimating the length of the cochleae, and quantifying the synapses in image stacks obtained with confocal imaging is presented.

The loss of ribbon synapses connecting inner hair cells and afferent auditory nerve fibers is assumed to be one cause of age-related hearing loss. The ...

Imaging the Aging Cochlea with Light-Sheet Fluorescence Microscopy

Deafness is the most common sensory impairment, affecting approximately 5% or 430 million people worldwide as per the World Health Organization1. Aging or presbycusis is a primary cause of sensorineural hearing loss and is characterized by damage to hair cells, spiral ganglion neurons (SGNs), and the stria vascularis. These structures reside within the cochlea, which has a complex, spiral-shaped anatomy of membranous tissues suspended in fluid and surrounded by bone. These properties make it technically difficult to investigate and quantify histopathological changes. To address this need, we developed a light-sheet microscope (TSLIM) that can image and digitize the whole cochlea to facilitate the study of structure-function relationships in the inner ear. Well-aligned serial sections of the whole cochlea result in a stack of images for three-dimensional (3D) volume rendering and segmentation of individual structures for 3D visualization and quantitative analysis (i.e., length, width, surface, volume, and number). Cochleae require minimal processing steps (fixation, decalcification, dehydration, staining, and optical clearing), all of which are compatible with subsequent high-resolution imaging by scanning and transmission electron microscopy. Since all the tissues are present in the stacks, each structure can be assessed individually or relative to other structures. In addition, since imaging uses fluorescent probes, immunohistochemistry and ligand binding can be used to identify specific structures and their 3D volume or distribution within the cochlea. Here we used TSLIM to examine cochleae from aged mice to quantify the loss of hair cells and spiral ganglion neurons. In addition, advanced analyses (e.g., cluster analysis) were used to visualize local reductions of spiral ganglion neurons in Rosenthal&#39;s canal along its 3D volume. These approaches demonstrate TSLIM microscopy&#39;s ability to quantify structure-function relationships within and between cochleae.

A light-sheet microscope was developed to image and digitize whole cochlea.

Deafness is the most common sensory impairment, affecting approximately 5% or 430 million people worldwide as per the World Health Organization1. Aging or ...

Modified Experimental Conditions for Noise-Induced Hearing Loss in Mice and Assessment of Hearing Function and Outer Hair Cell Damage

An animal model of noise-induced hearing loss (NIHL) is useful for pathologists, therapists, pharmacologists, and hearing researchers to thoroughly understand the mechanism of NIHL, and subsequently optimize the corresponding treatment strategies. This study aims to create an improved protocol for developing a mouse model of NIHL. Male C57BL/6J mice were used in this study. Unanesthetized mice were exposed to loud noises (1 and 6 kHz, presented simultaneously at 115-125 dB SPL-A) continuously for 6 h per day for 5 consecutive days. Auditory function was assessed 1 day and 1 week after noise exposure, using auditory brainstem response (ABR). After the ABR measurement, the mice were sacrificed, and their organs of Corti were collected for immunofluorescence staining. From the auditory brainstem response (ABR) measurements, significant hearing loss was observed 1 day after noise exposure. After 1 week, the hearing thresholds of the experimental mice decreased to ~80 dB SPL, which was still a significantly higher level than the control mice (~40 dB SPL). From the results of immunofluorescence imaging, outer hair cells (OHCs) were shown to be damaged. In summary, we created a model of NIHL using male C57BL/6J mice. A new and simple device for generating and delivering pure-tone noise was developed and then employed. Quantitative measurements of hearing thresholds and morphological confirmation of OHC damage both demonstrated that the applied noise successfully induced an expected hearing loss.

Here, we present a protocol for a mouse model of noise-induced hearing loss (NIHL). To induce NIHL, we developed a new and simple device using corrugated plastic, a rat trap cage, and a speaker. Auditory brainstem response and immunofluorescence imaging were employed to assess the hearing function and outer hair cell damage, respectively.

An animal model of noise-induced hearing loss (NIHL) is useful for pathologists, therapists, pharmacologists, and hearing researchers to thoroughly ...

Evaluation of Ribbon Synapses at Specific Frequency Regions of Mouse Cochlea

Source: Yu, S., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/59189">Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea.</a> J. Vis. Exp. (2019).
This video demonstrates a method for evaluating ribbon synapses in specific regions of the mouse cochlea. It utilizes immunostaining and confocal imaging techniques to identify functional synapses across different frequency regions in each cochlear turn.

Morphological and Functional Evaluation of Ribbon Synapses at Specific Frequency Regions of the Mouse Cochlea

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